The Protein Protocols Handbook, 3rd Edition (Springer Protocols Handbooks)

The Protein Protocols Handbook

SPRINGER PROTOCOLS HANDBOOKS

For other titles published in this series, go to www.springer.com/series/8623

The

Protein Protocols Handbook THIRD EDITION Edited by

John M. Walker University of Hertfordshire, Hatfield, UK

Editor John M. Walker University of Hertsfordshire Hatfield, Hertsfordshire UK

ISBN 978-1-60327-474-6 DOI 10.1007/978-1-59745-198-7

e-ISBN 978-1-59745-198-7

Library of Congress Control Number: 2009932604 ã Humana Press, a part of Springer Science+Business Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is for-bidden. 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 re-spect to the material contained herein. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface Since the second edition of this book was published in 2002 there have, of course, been continual methodological developments in the field of protein chemistry. Consequently, for this third edition, I introduced 57 chapters/protocols not present in the second edition, significantly updated a number of chapters remaining from the second edition and increased the overall length of the book from 164 to 208 chapters. The new chapters are generally spread across the various sections of the book, but in particular we have expanded the section on post-translational modifications to reflect the increasing importance of these modifications in understanding of protein function. As in the earlier editions, The Protein Protocols Handbook aims to provide a cross-section of analytical techniques commonly used to study proteins and peptides, thus providing a benchtop manual and guide for both those who are new to the protein chemistry laboratory and for more established workers who wish to use a technique for the first time. All chapters are written in the same format as that used in the Methods in Molecular Biology series. Each chapter opens with a description of the basic theory behind the method being described. The Materials section lists all the chemicals, reagents, buffers, and other materials necessary for carrying out the protocol. Since the principal goal of the book is to provide experimentalists with a full account of the practical steps necessary for carrying out each protocol successfully, the Methods section contains detailed step-by-step descriptions for every protocol that should result in the successful execution of each method. The Notes section complements the Methods material by indicating how best to deal with any problem or difficulty that may arise when using a given technique, by providing useful hints and tips, and by explaining how to go about making appropriate modifications or alterations to the protocol. John M. Walker

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Contents Preface .............................................................................................................. v Contributors ...................................................................................................xxiii

PART I 1 2 3 4 5

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7 8 9

10

QUANTITATION OF PROTEINS

Protein Determination by UV Absorption ................................................ 3 Alastair Aitken and Michèle P. Learmonth The Lowry Method for Protein Quantitation ............................................ 7 Jakob H. Waterborg The Bicinchoninic Acid (BCA) Assay for Protein Quantitation .............. 11 John M. Walker The Bradford Method For Protein Quantitation ..................................... 17 Nicholas J. Kruger Ultrafast Protein Determinations Using Microwave Enhancement ........................................................................................ 25 Robert E. Akins and Rocky S. Tuan The Nitric Acid Method for Protein Estimation in Biological Samples ............................................................................ 35 Scott A. Boerner, Crescent R. Isham, Yean Kit Lee, Jennifer D. Tibodeau, Scott H. Kaufmann, and Keith C. Bible Quantitation of Tryptophan in Proteins .................................................. 47 Alastair Aitken and Michèle P. Learmonth Kinetic Silver Staining of Proteins ......................................................... 51 Douglas D. Root and Kuan Wang Quantitation of Cellular Proteins by Flow Cytometry ............................ 55 Thomas D. Friedrich, F. Andrew Ray, Ralph L. Smith, and John M. Lehman Quantitation of Cellular Proteins by Laser Scanning Cytometry ............................................................................. 63 Thomas D. Friedrich, Ralph L. Smith, and John M. Lehman

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PART II

ELECTROPHORESIS OF PROTEINS AND PEPTIDES DECTECTION IN GELS

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Protein Solubility in Two-Dimensional Electrophoresis: Basic Principles and Issues .................................................................. 73 Thierry Rabilloud Mouse and Human Tissue Sample Preparation for 2-D Electrophoresis ......................................................................... 85 Claus Zabel and Joachim Klose Plant Protein Sample Preparation for 2-DE ........................................ 109 Sebastien Christian Carpentier, Rony Swennen, and Bart Panis Preparation of Bacterial Samples for 2-D PAGE ................................. 121 Brian B. Vandahl, Gunna Christiansen, and Svend Birkelund Preparation of Bodily Fluids for 2-D PAGE.......................................... 131 Anthony G. Sullivan, Henry Brzeski, Jeban Ganesalingam, and Manuel Mayr Immunoaffinity Depletion of High Abundance Plasma and Serum Proteins ............................................................... 139 Lynn A. Echan and David W. Speicher Preparation of Yeast Samples for 2-D PAGE ...................................... 155 Joakim Norbeck Membrane Protein Preparation Using Aqueous Polymer Two-Phase Systems ........................................................................... 159 Jens Schindler and Hans Gerd Nothwang Subcellular Fractionation of Small Sample Amounts .......................... 165 Hans Gerd Nothwang, Isabelle Guillemin, and Jens Schindler Nondenaturing Polyacrylamide Gel Electrophoresis of Proteins ................................................................. 171 John M. Walker SDS Polyacrylamide Gel Electrophoresis of Proteins ......................... 177 John M. Walker Gradient SDS Polyacrylamide Gel Electrophoresis of Proteins .......... 187 John M. Walker SDS-Polyacrylamide Gel Electrophoresis of Peptides ........................ 193 Ralph C. Judd Separation of Proteins by Blue Native Electrophoresis (B N-PAGE) ................................................................ 201 Elke A. Dian, Joachim Rassow, and Christian Motz

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Separation of Proteins by Gel Electrophoresis in the Tris-Taurine-HCl System............................................................ 211 Sylvie Luche, Mireille Chevallet, Cécile Lelong, and Thierry Rabilloud 26 Cetyltrimethylammonium Bromide Discontinuous Gel Electrophoresis of Proteins: Mr-Based Separation of Proteins with Retained Native Activity ............................................ 221 Robert E. Akins and Rocky S. Tuan 27 Acetic Acid–Urea Polyacrylamide Gel Electrophoresis of Basic Proteins ................................................................................. 239 Jakob H. Waterborg 28 Acid–Urea–Triton Polyacrylamide Gel Electrophoresis of H istones .......................................................................................... 251 Jakob H. Waterborg 29 Isoelectric Focusing of Proteins in Ultra-Thin Polyacrylamide Gels ........................................................................... 263 John M. Walker 30 Serial Immobilized pH Gradient Isoelectric Focusing over pH 4–9 ......................................................................... 269 Slobodan Poznanovi´c, Wojciech Wozny, Helmut Zengerling, Gerhard P. Schwall, and Michael A. Cahill 31 Radiolabeling of Eukaryotic Cells and Subsequent Preparation for 2-Dimensional Electrophoresis................................... 279 Nick Bizios 32 Two-Dimensional Polyacrylamide Gel Electrophoresis of Proteins Using Carrier Ampholyte pH Gradients in the First Dimension ......................................................................... 285 Patricia Gravel 33 Vertical Agarose Electrophoresis and Electroblotting of High-Molecular-Weight Proteins ..................................................... 293 Marion L. Greaser and Chad M. Warren 34 Two-Dimensional PAGE of High-Molecular-Mass Proteins ................. 299 Masamichi Oh-Ishi and Tadakazu Maeda 35 Casting Immobilized pH Gradients ..................................................... 305 Elisabetta Gianazza 36 A Modified Nonequilibrium pH Gradient Electrophoresis (NEPHGE) Technique for Protein Separation ..................................... 323 James A. Carroll

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Microchip Capillary Electrophoresis: Application to Peptide Analysis ........................................................... 329 Barbara A. Fogarty, Nathan A. Lacher, and Susan M. Lunte Protein Separations in Microfluidic Chips ........................................... 361 Andrea W. Chow and Bahram Fathollahi Difference Gel Electrophoresis (DIGE) ............................................... 379 David B. Friedman and Kathryn S. Lilley Comparing 2D Electrophoretic Gels Across Internet Databases: An Open Source Application .............................. 409 Peter F. Lemkin, Gregory C. Thornwall, and Jai A. Evans Quantification of Radiolabeled Proteins in Polyacrylamide Gels ....................................................................... 443 Wayne R. Springer Differential ProteoTope Radioactive Quantification of Protein Abundance Ratios .............................................................. 449 Wojciech Wozny, Gerhard P. Schwall, Chaturvedula S. Sastri, Slobodan Poznanovi´c,Werner Stegmann, Christian Hunzinger†, Karlfried Groebe, and Michael A. Cahill Quantification of Proteins on Polyacrylamide Gels ............................. 479 Bryan John Smith Using SDS-PAGE and Scanning Laser Densitometry to Measure Yield and Degradation of Proteins.................................... 487 Aaron P. Miles and Allan Saul Rapid and Sensitive Staining of Unfixed Proteins in Polyacrylamide Gels with Nile Red ................................................. 497 Joan-Ramon Daban, Salvador Bartolomé, Antonio Bermúdez, and F. Javier Alba 2+ Zn -Reverse S taining Technique ........................................................ 505 Carlos Fernandez-Patron Protein Staining with Calconcarboxylic Acid in Polyacrylamide Gels................................................................ 515 Wei-Tao Cong, Sun-Young Hwang, Li-Tai Jin, and Jung-Kap Choi Detection of Proteins in Polyacrylamide Gels by Silver Staining ................................................................................ 521 Michael J. Dunn Background-Free Protein Detection in Polyacrylamide Gels and on Electroblots Using Transition Metal Chelate Stains..................................................................................... 529 Wayne F. Patton

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Detection of Proteins in Polyacrylamide Gels by Fluorescent Staining ...................................................................... 547 Michael J. Dunn Detection of Glycoproteins in Gels and Blots...................................... 555 Nicolle H. Packer, Malcolm S. Ball, Peter L. Devine, and Wayne F. Patton Staining of Glycoproteins/Proteoglycans on SDS-Gels ...................... 569 Holger J. Møller and Jørgen H. Poulsen Detection of Proteins and Sialoglycoproteins in Polyacrylamide Gels Using Eosin Y Stain ....................................... 575 Fan Lin and Gary E. Wise A Modified Pro-Q Diamond Staining Protocol for Phosphoprotein Detection in Polyacrylamide Gels ........................ 579 Ganesh Kumar Agrawal and Jay J. Thelen Electroelution of Proteins from Polyacrylamide Gels .......................... 587 Paul Jenö and Martin Horst Autoradiography and Fluorography of Acrylamide Gels...................... 595 Antonella Circolo and Sunita Gulati Proteolytic Activity Detection by Two-Dimensional Zymography ........................................................................................ 605 Jeff Wilkesman

PART III 58 59 60 61

62 63 64

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BLOTTING AND DETECTION METHODS

Protein Blotting by Electroblotting ....................................................... 617 Mark Page and Robin Thorpe Protein Blotting by the Semidry Method.............................................. 621 Patricia Gravel Protein Blotting by the Capillary Method ............................................. 631 John M. Walker Western Blotting of Basic Proteins Electrophoretically Resolved on Acid-Urea-Triton-Polyacrylamide Gels ........................... 635 Geneviève P. Delcuve and James R. Davie Immunoblotting of 2-DE Separated Proteins....................................... 641 Barbara Magi and Laura Bianchi High-Efficiency Blotting of High-Molecular Weight Proteins................ 663 Marion L. Greaser and Darl R. Swartz Alkaline Phosphatase Labeling of IgG Antibody ................................. 673 G. Brian Wisdom

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Contents β-Galactosidase Labeling of IgG Antibody.......................................... 677 G. Brian Wisdom Horseradish Peroxidase Labeling of IgG Antibody ............................. 681 G. Brian Wisdom Digoxigenin Labeling of IgG Antibody ................................................. 685 G. Brian Wisdom Conjugation of Fluorochromes to Antibodies ...................................... 687 Su-Yau Mao Coupling of Antibodies with Biotin....................................................... 693 Rosaria P. Haugland and Wendy W. You Preparation of Avidin Conjugates........................................................ 705 Rosaria P. Haugland and Mahesh K. Bhalgat MDPF Staining of Proteins on Western Blots ..................................... 717 F. Javier Alba and Joan-Ramon Daban Copper Iodide Staining of Proteins and Its Silver E nhancement ............................................................................ 723 Douglas D. Root and Kuan Wang Detection of Proteins on Blots Using Direct Blue 71 ........................... 729 Wei-Tao Cong, Sun-Young Hwang, Li-Tai Jin, and Jung-Kap Choi Detection of Proteins on Western Blots Using Colorimetric and Radiometric Visualization of Secondary Ligands ......................................................................... 737 Nicholas J. Kruger Identification of Glycoproteins on Nitrocellulose Membranes Using Lectin Blotting ....................................................... 755 Patricia Gravel A Sensitive Method to Quantitatively Detect Total Protein on Membranes after Electrophoretic Transfer Using Avidin- or Streptavidin-Biotin ............................................................... 771 William J. LaRochelle Detection and Quantification of Proteins on Immunoblots using Enhanced Chemiluminescence ................................................. 779 Jennifer J. Young Reutilization of Western Blots After Chemiluminescent or Autoradiographic Detection............................................................. 789 Scott H. Kaufmann The Use of Quantum Dot Luminescent Probes for Western Blot Analysis .................................................................... 807 Savvas C. Makrides, Christina Gasbarro, and Job M. Bello

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The Use of Infrared Fluorescent Dyes in Quantitative Immunoblotting............................................................ 819 Ching-Hui Yang, Christopher Kasbek, and Harold A. Fisk The Use of Infrared Fluorescent Dyes in Immunofluorescence Microscopy .................................................... 827 Christopher Kasbek, Ching-Hui Yang, and Harold A. Fisk

PART IV 82

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CHEMICAL MODIFICATION OF PROTEINS AND PEPTIDE PRODUCTION AND PURIFICATION

Carboxymethylation of Cysteine Using lodoacetamide/lodoacetic Acid ................................................. 837 Alastair Aitken and Michèle Learmonth Performic Acid Oxidation..................................................................... 841 Alastair Aitken and Michèle Learmonth Succinylation of Proteins..................................................................... 845 Alastair Aitken and Michèle Learmonth Pyridylethylation of Cysteine Residues ............................................... 847 Malcolm Ward Side Chain Selective Chemical Modifications of Proteins ................... 851 Dan S. Tawfik Nitration of Tyrosines........................................................................... 855 Dan S. Tawfik Ethoxyformylation of Histidine ............................................................. 859 Dan S. Tawfik Modification of Arginine Side Chains with p-Hydroxyphenylglyoxal ............................................................... 863 Dan S. Tawfik Amidation of Carboxyl Groups ............................................................ 865 Dan S. Tawfik Amidination of Lysine Side Chains ..................................................... 867 Dan S. Tawfik Modification of Tryptophan with 2-Hydroxy5-Nitrobenzylbromide .......................................................................... 871 Dan S. Tawfik Modification of Sulfhydryl Groups with DTNB ..................................... 873 Dan S. Tawfik Chemical Cleavage of Proteins at Methionyl-X Peptide B onds ..................................................................................... 875 Bryan John Smith

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Contents Chemical Cleavage of Proteins at Tryptophanyl-X Peptide B onds ..................................................................................... 883 Bryan John Smith Chemical Cleavage of Proteins at Aspartyl-X Peptide Bonds ............. 891 Bryan John Smith Chemical Cleavage of Proteins at Cysteinyl-X Peptide B onds ..................................................................................... 895 Bryan John Smith Chemical Cleavage of Proteins at Asparaginyl-Glycyl Peptide B onds ..................................................................................... 899 Bryan John Smith Enzymatic Digestion of Proteins in Solution and in SDS Polyacrylamide Gels ........................................................ 905 Kathryn L. Stone, Erol E. Gulcicek, and Kenneth R. Williams On-PVDF Protein Digestions for N-terminal Sequencing and Peptide Mass Fingerprinting ........................................................ 919 Victoria Pham, William Henzel, and Jennie R. Lill Enzymatic Digestion of Membrane-Bound Proteins for Peptide Mapping and Internal Sequence Analysis ........................ 927 Joseph Fernandez and Sheenah M. Mische Reverse-Phase HPLC Separation of Enzymatic Digests of Proteins .............................................................................. 941 Kathryn L. Stone and Kenneth R. Williams Peptide Mapping by Two-Dimensional Thin-Layer Electrophoresis-Thin-Layer Chromatography ..................................... 951 Ralph C. Judd Peptide Mapping by Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis ....................................... 963 Ralph C. Judd Peptide Mapping for Protein Characterization .................................... 969 Peter Højrup Production of Protein Hydrolysates Using Enzymes........................... 989 John M. Walker and Patricia J. Sweeney Amino Acid Analysis by Precolumn Derivatization with 1-Fluoro-2,4-Dinitrophenyl-5-I-Alanine Amide (Marfey’s R eagent) .............................................................................. 995 Sunil Kochhar and Philipp Christen Amino Acid Analysis of Protein Hydrolysates Using Anion Exchange Chromatography and IPAD Detection .............................. 1001 Petr Jandik, Jun Cheng, and Nebojsa Avdalovic

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Validation of Amino Acid Analysis Methods ...................................... 1015 Andrew J. Reason Molecular Weight Estimation for Native Proteins Using High-Performance Size-Exclusion Chromatography ............................................................................... 1029 G. Brent Irvine Detection of Disulfide-Linked Peptides by HPLC .............................. 1039 Alastair Aitken and Michèle Learmonth Detection of Disulfide-Linked Peptides by Mass Spectrometry .................................................................................... 1043 Alastair Aitken and Michèle Learmonth Diagonal Electrophoresis for Detecting Disulfide Bridges................. 1047 Alastair Aitken and Michèle Learmonth Estimation of Disulfide Bonds Using Ellman’s Reagent .................... 1053 Alastair Aitken and Michèle Learmonth Quantitation of Cysteine Residues and Disulfide Bonds by Electrophoresis ................................................................. 1057 Alastair Aitken and Michèle Learmonth N-Terminal Sequencing of N-Terminally Modified P roteins .............................................................................. 1063 Roza Maria Kamp and Hisashi Hirano Deblocking of Proteins Containing N-Terminal Pyroglutamic A cid ............................................................................. 1075 Jacek Mozdzanowski Detection and Characterization of Protein Mutations by Mass Spectrometry...................................................... 1081 Yoshinao Wada Peptide Sequencing by Nanoelectrospray Tandem Mass Spectrometry ............................................................. 1095 Ole Nørregaard Jensen and Matthias Wilm Protein Identification by Peptide Mass Fingerprinting using MALDI-TOF Mass Spectrometry ............................................. 1117 Judith Webster and David Oxley Protein Ladder Sequencing .............................................................. 1131 Rong Wang and Brian T. Chait Sequence Analysis with WinGene/WinPep ....................................... 1141 Lars Hennig HPLC and Mass Spectrometry of Integral Membrane Proteins........................................................................... 1149 Julian P. Whitelegge

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Contents Enrichment of Serum Peptides and Analysis by MALDI-TOF Mass Spectrometry .................................................. 1167 Yanming An, Habtom W. Ressom, and Radoslav Goldman Computational Methods for Analysis of MALDI-TOF Spectra to Discover Peptide Serum Biomarkers ............................... 1175 Habtom W. Ressom, Rency S. Varghese, and Radoslav Goldman

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POSTTRANSLATIONAL MODIFICATIONS

Simple Tools for Complex N-Glycan Analysis ....................................1187 Anne-Catherine Fitchette, Meriem Benchabane, Thomas Paccalet, Loïc Faye, and Véronique Gomord A Lectin-Binding Assay for the Rapid Characterization of the Glycosylation of Purified Glycoproteins .................................. 1205 Mohammad T. Goodarzi, Angeliki Fotinopoulou, and Graham A. Turner Chemical Methods of Analysis of Glycoproteins ............................... 1215 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith Monosaccharide Analysis by HPAEC ............................................... 1219 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith Monosaccharide Analysis by Gas Chromatography (GC) ................ 1223 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith Determination of Monosaccharide Linkage and Substitution Patterns by GC-MS Methylation Analysis ......................................... 1227 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith Sialic Acid Analysis by HPAEC-PAD ................................................. 1231 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith Chemical Release of O-Linked Oligosaccharide Chains .................. 1233 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith O-Linked Oligosaccharide Profiling by HPLC ................................... 1235 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith O-Linked Oligosaccharide Profiling by HPAEC-PAD ......................... 1237 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith

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Release of N-Linked Oligosaccharide Chains by H ydrazinolysis .............................................................................. 1239 Tsuguo Mizuochi and Elizabeth F. Hounsell Enzymatic Release of O-and N-Linked Oligosaccharide C hains .................................................................... 1243 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith N-Linked Oligosaccharide Profiling by HPLC on Porous Graphitized Carbon (PGC) .............................................. 1247 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith N-Linked Oligosaccharide Profiling by HPAEC-PAD ......................... 1249 Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith HPAEC-PAD Analysis of Monosaccharides Released by Exoglycosidase Digestion Using the CarboPac MA1 C olumn ..................................................................................... 1253 Michael Weitzhandler, Jeffrey Rohrer, James R. Thayer, and Nebojsa Avdalovic Microassay Analyses of Protein Glycosylation ................................. 1261 Nicky K. C. Wong, Nnennaya Kanu, Natasha Thandrayen, Geert Jan Rademaker, Christopher I. Baldwin, David V. Renouf, and Elizabeth F. Hounsell Polyacrylamide Gel Electrophoresis of Fluorophore-Labeled Carbohydrates from Glycoproteins ................................................... 1273 Brian K. Brandley, John C. Klock, and Christopher M. Starr HPLC Analysis of Fluorescently Labeled Glycans ............................ 1289 Tony Merry and Sviatlana Astrautsova Glycoprofiling Purified Glycoproteins Using Surface Plasmon Resonance ................................................. 1313 Angeliki Fotinopoulou and Graham A. Turner Sequencing Heparan Sulfate Saccharides ....................................... 1321 Jeremy E. Turnbull Analysis of Glycoprotein Heterogeneity by Capillary Electrophoresis and Mass Spectrometry .......................................... 1335 Andrew D. Hooker and David C. James Affinity Chromatography of Oligosaccharides and Glycopeptides with Immobilized Lectins .................................... 1347 Kazuo Yamamoto In-Gel Enzymatic Release of N-Glycans........................................... 1363 David J. Harvey

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Analysis of N-Linked Glycans by Mass Spectrometry ...................... 1371 David J. Harvey MS Analysis of Protein Glycosylation ............................................... 1387 Nobuaki Takemori, Naoka Komori, and Hiroyuki Matsumoto Mapping protein N-Glycosylation by COFRADIC ............................. 1395 Bart Ghesquière, Joël Vandekerckhove, and Kris Gevaert Mass Spectrometric Analysis of O-Linked Glycans Released Directly from Glycoproteins in Gels Using β-Elimination ............................................................... 1403 Adrian M. Taylor and Jane Thomas-Oates Glycopeptide Analysis Using LC/MS and LC/MSn: Site-Specific Glycosylation Analysis of a Glycoprotein ..................... 1419 Satsuki Itoh, Daisuke Takakura, Nana Kawasaki, and Teruhide Yamaguchi Identification of Vitamin K-Dependent Proteins Using a Gla-Specific Monoclonal Antibody ....................................... 1431 Karin Hansson The Identification of Protein S-Nitrosocysteine ................................. 1451 Todd M. Greco, Sheryl L. Stamer, Daniel C. Liebler, and Harry Ischiropoulos Detection of Nitrotyrosine-Containing Proteins ................................. 1467 Xianquan Zhan and Dominic M. Desiderio Mass Spectrometric Determination of Protein Ubiquitination .................................................................................... 1491 Carol E. Parker, Maria Warren Hines, Viorel Mocanu, Susanna F. Greer, and Christoph H. Borchers Detection of Sumoylated Proteins ..................................................... 1519 Ok-Kyong Park-Sarge and Kevin D. Sarge Efficient Enrichment of Intact Phosphorylated Proteins by Modified Immobilized Metal-Affinity Chromatography ............................................................................... 1531 Anna Dubrovska Analyzing Protein Phosphorylation ................................................... 1547 John Colyer Mass Spectrometric Analysis of Protein Phosphorylation ................ 1555 Stefan Gander, Alessio Cremonesi, Johana Chicher, Suzette Moes, and Paul Jenö

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Protein Microarrays for Phosphorylation Studies .............................. 1567 Birgit Kersten and Tanja Feilner Two-Dimensional Phosphopeptide Mapping ..................................... 1579 Hikaru Nagahara, Robert R. Latek, Sergei A. Ezhevsky, and Steven F. Dowdy Identification of Proteins Modified by Protein (D-Aspartyl/L-Isoaspartyl) Carboxyl Methyltransferase ...................... 1589 Darin J. Weber and Philip N. McFadden Analysis of Tyrosine-O-Sulfation ....................................................... 1601 Jens R. Bundgaard, Jette W. Sen, Anders H. Johnsen, and Jens F. Rehfeld Analysis of Protein Palmitoylation by Metabolic Radiolabeling Methods ................................................ 1623 Katherine H. Pedone, Leah S. Bernstein, Maurine E. Linder, and John R. Hepler Incorporation of Radiolabeled Prenyl Alcohols and Their Analogs into Mammalian Cell Proteins: A Useful Tool for Studying Proteins Prenylation ................................ 1637 Alberto Corsini, Christopher C. Farnsworth, Paul McGeady, Michael H. Gelb, and John A. Glomset The Metabolic Labeling and Analysis of Isoprenylated Proteins .................................................................. 1657 Douglas A. Andres, Dean C. Crick, H. Peter Spielmann, and Charles J. Waechter

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ANTIBODY TECHNIQUES

Antibody P roduction .......................................................................... 1679 Robert Burns Production of Antibodies Using Proteins in Gel Bands ..................................................................................... 1687 Sally Ann Amero, Tharappel C. James, and Sarah C. R. Elgin Raising Highly Specific Polyclonal Antibodies Using Biocompatible Support-Bound Antigens ........................................... 1693 Monique Diano and André Le Bivic Production of Antisera Using Peptide Conjugates ............................ 1705 Thomas E. Adrian Small-Molecule–Protein Conjugation Procedures ............................ 1715 Stephen Thompson

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Contents The Chloramine T Method for Radiolabeling Protein ........................ 1727 Graham S. Bailey The Lactoperoxidase Method for Radiolabeling Protein ................................................................... 1731 Graham S. Bailey The Bolton and Hunter Method for Radiolabeling Protein ................................................................... 1733 Graham S. Bailey Preparation of 125I-Labeled Peptides and Proteins with High Specific Activity Using IODO-GEN .................................... 1735 J. Michael Conlon Purification and Assessment of Quality of Radioiodinated Protein.................................................................. 1743 Graham S. Bailey Purification of IgG by Precipitation with Sodium Sulfate or Ammonium Sulfate ........................................................... 1749 Mark Page and Robin Thorpe Purification of IgG Using Caprylic Acid ............................................. 1753 Mark Page and Robin Thorpe Purification of IgG Using DEAE-Sepharose Chromatography ............................................................................... 1755 Mark Page and Robin Thorpe Purification of IgG Using Ion-Exchange HPLC ................................. 1757 Carl Dolman, Mark Page, and Robin Thorpe Purification of IgG by Precipitation with Polyethylene Glycol (PEG) ........................................................ 1759 Mark Page and Robin Thorpe Purification of IgG Using Protein A or Protein G ............................... 1761 Mark Page and Robin Thorpe Analysis and Purification of IgG Using Size-Exclusion High Performance Liquid Chromatography (SE-HPLC) .................... 1765 Carl Dolman and Robin Thorpe Purification of IgG Using Affinity Chromatography on Antigen-Ligand Columns .................................. 1769 Mark Page and Robin Thorpe Purification of IgG Using Thiophilic Chromatography ....................... 1773 Mark Page and Robin Thorpe Analysis of IgG Fractions by Electrophoresis.................................... 1775 Mark Page and Robin Thorpe

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Purification of Immunoglobulin Y (IgY) from Chicken Eggs ............................................................................ 1779 Christopher R. Bird and Robin Thorpe Affinity Purification of Immunoglobulins Using Protein A Mimetic (PAM) ......................................................... 1783 Giorgio Fassina, Giovanna Palombo, Antonio Verdoliva, and Menotti Ruvo Detection of Serological Cross-Reactions by Western Cross-Blotting ................................................................ 1799 Peter Hammerl, Arnulf Hartl, Johannes Freund, and Josef Thalhamer Enzymatic Digestion of Monoclonal Antibodies ................................ 1811 Sarah M. Andrew How to Make Bispecific Antibodies ................................................... 1819 Ruth R. French Antigen Measurement Using ELISA ................................................. 1827 William Jordan Enhanced Chemiluminescence Immunoassay ................................. 1835 Richard A. W. Stott Immunoprecipitation and Blotting: The Visualization of Small Amounts of Antigens Using Antibodies and Lectins.............................................................1845 Stephen Thompson Determination of Epitopes by Mass Spectrometry............................ 1859 Christine Hager-Braun and Kenneth B. Tomer Immunogen Preparation and Immunization Procedures for Rats and Mice .......................................................... 1873 Mark Page and Robin Thorpe Making H ybridomas .......................................................................... 1877 Robert Burns Growing Hybridomas ........................................................................ 1887 Gary Entrican, Catherine Jepson, and David Deane Mouse Hybridomas as an Entryway to Monoclonal Antibody Design and Production....................................................... 1901 Eugene Mechetner Potential Pitfalls in Monoclonal Antibody Development and Applications.......................................................... 1911 Eugene Mechetner

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Recombinant Antibody Expression and Purification ......................... 1929 Achim Knappik and Ralf Brundiers 204 Screening Hybridoma Culture Supernatants Using Solid-Phase Radiobinding Assay ............................................ 1945 Mark Page and Robin Thorpe 205 Screening Hybridoma Culture Supernatants Using ELISA ............... 1947 Mark Page and Robin Thorpe 206 Growth and Purification of Murine Monoclonal A ntibodies ...................................................................... 1949 Mark Page and Robin Thorpe 207 Affinity Purification Techniques for Monoclonal Antibodies ......................................................................................... 1951 Alexander Schwarz 208 A Rapid Method for Generating Large Numbers of High-Affinity Monoclonal Antibodies from a Single Mouse ......................................................................... 1961 Nguyen Thi Man and Glenn E. Morris Index ............................................................................................................1975

Contributors Department of Physiology, United Arab Emirates University, Faculty of Medicine and Health Sciences, Al Ain, United Arab Emirates Ganesh Kumar Agrawal University of Missouri-Columbia, Division of Biochemistry, Christopher S. Bond Life Sciences Center, Columbia, MO Alastair Aitken School of Biological Sciences, University of Edinburgh, Edinburgh, UK Robert E. Akins Tissue Engineering and Regenerative Medicine Research, Nemours Biomedical Research, A.I. duPont Hospital for Children, Wilmington, DE F. Javier Alba Department of Biochemistry and Molecular Biology, University Autonoma de Barcelona, Faculty of Biosciences, Bellaterra, Barcelona, Spain Sally Ann Amero Center for Scientific Review, National Institutes of Health, Bethesda, MD Yanming An Department of Oncology, Georgetown University, Washington, DC Douglas A. Andres Department of Molecular and Cellular Biochemistry, University of Kentucky College of Medicine, Lexington, KY Sarah M. Andrew Chester College of Higher Education, UK Sviatlana Astrautsova Department of Microbiology, Medical University of Grodno, Belarus Nebojsa Avdalovic Dionex Corporation, Sunnyvale, CA Graham S. Bailey Department of Biological Sciences, University of Essex, Wivenhoe Park, Colchester, UK Christopher I. Baldwin Department of Biochemistry, University of Hong Kong, Pofulam, Hong Kong Malcolm S. Ball Co-operative Research Centre for Eye Research Technology, Sydney, Australia Salvador Bartolomé Department of Biochemistry and Molecular Biology, University Autonoma de Barcelona, Faculty of Biosciences, Bellaterra, Barcelona, Spain Job M. Bello EIC Laboratories, Inc., Norwood, MA Meriem Benchabane CNRS UMR, Faculté des Sciences, Université de Rouen, Rouen, France Thomas E. Adrian

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Department of Biochemistry and Molecular Biology, University Autonoma de Barcelona, Faculty of Biosciences, Bellaterra, Barcelona, Spain Leah S. Bernstein Solvay Pharmaceuticals Inc., Marietta, GA Mahesh K. Bhalgat Molecular Probes. Inc., Eugene, OR Laura Bianchi Department of Molecular Biology, University of Siena, Siena, Italy Keith C. Bible Department of Medical Oncology, Mayo Clinic, Rochester, MN Christopher R. Bird Division of Immunobiology, Ntional Institute for Biological Standards and Control, Potters Bar, UK Svend Birkelund Department of Medical Microbiology and Immunology, University of Aarhus, Denmark Nick Bizios AGI Dermatics, Freeport, NY Scott A. Boerner Division of Medical Oncology, Mayo Clinic, Rochester, MN Christoph H. Borchers Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill, Chapel Hill, NC USA. Current Address: University of Victoria Genome BC Proteomics Centre, Vancouver Island Technology Park, Victoria, BC, Canada Brian K. Brandley Glyko Inc., Navato, CA Ralf Brundiers MorphoSys AG, Martinsried, Germany Henry Brzeski University of Hertfordshire, Hatfield, Hertfordshire, UK Jens R. Bundgaard Department of Clinical Biochemistry, Rigshospitalet, Copenhagen University Hospital, Copenhagen, Denmark Robert Burns Scottish Agricultural Science Agency, Edinburgh Scotland, UK Michael A. Cahill School of Biomedical Sciences, Charles Sturt University, Wagga Wagga, NSW, Australia Sebastien Christian Carpentier Faculty of Bioscience Engineering, Division of Crop Biotechnics, Leuven, Belgium James A. Carroll University of Pittsburgh School of Medicine, Department of Molecular Genetics and Biochemistry, Pittsburgh, PA Brian T. Chait Rockefeller Univerisity, New York, NY Jun Cheng Dionex Corporation, Sunnyvale, CA Mireille Chevallet CEA/DSV/iRTSV/LBBSI, CEA Grenoble, Grenoble, France Johana Chicher Department of Biochemistry, Biozentrum of the University of Basel, Basel, Switzerland Jung-Kap Choi Laboratory of Analytical Biochemistry, College of Pharmacy, Chonnam National University, Gwangju, South Korea Antonio Bermúdez

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Andrea W. Chow Philipp Christen

Caliper Life Sciences, Mountain View, CA Biochemisches Institut, Universität Zürich, Zürich,

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Switzerland Department of Medical Microbiology and Immunology, University of Aarhus, Denmark Antonella Circolo Maxwell Finland Lab for Infectious Diseases, Boston, MA John Colyer Department of Biochemistry and Molecular Biology, University of Leeds, UK Wei-tao Cong College of Pharmacy and Research Institute of Drug Development, Chonnam National University, Gwangju, South Korea J. Michael Conlon Department of Biochemistry, U.A.E. University, United Arab Emirates Alberto Corsini Department of Pharmacological Sciences, University of Milan, Italy Alessio Cremonesi Department of Biochemistry, Biozentrum of the University of Basel, Basel, Switzerland Dean C. Crick Department of Microbiology, Immunology, and Pathology, Colorado Sate University, Fort Collins, CO Joan-Ramon Daban Department of Biochemistry and Molecular Biology, University Autonoma de Barcelona, Faculty of Biosciences, Bellaterra, Barcelona, Spain James R. Davie Manitoba Institute of Cell Biology, Winnipeg, Manitoba, Canada Michael J. Davies School of Biological and Chemical Sciences, Birkbeck University of London, UK David Deane Moredun Research Institute, Pentlands Science Park, Edinburgh, UK Geneviève P. Delcuve Manitoba Institute of Cell Biology, University of Manitoba, Winnipeg, Manitoba, Canada Dominic M. Desiderio Charles B. Stout Neuroscience Mass Spectrometry Laboratory, Department of Neurology, Univiversity of Tennessee Health Science Center, Memphis, TN Peter L. Devine Proteome Systems Ltd., Sydney, Australia Elke A. Dian Institute for Physiological Chemistry, Ruhr University Bochum, Bochum, Germany Monique Diano IBDM, Faculté des Sciences de Luminy, Marseille, France Carl Dolman Division of Immunobiology, National Institute for Biological Standards and Control, Potters Bar, UK Steven F. Dowdy Howard Hughes Medical Institute, Department of Cellular and Molecular Medicine, George Palade Laboratories, UCSD School of Medicine, La Jolla, CA Gunna Christiansen

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Contributors

Anna Dubrovska

Genomics Institute for the Novartis Research Foundation,

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San Diego, CA Proteome Research Centre, Conway Institute of Biomolecular and Biomedical Research, University College Dublin, Ireland Lynn A. Echan Systems Biology Division, Proteomics Laboratory, The Wistar Institute, Philadelphia, PA Sarah C. R. Elgin Washington University, Department of Biology, St. Louis, MO Gary Entrican Bacteriology/Moredun Research Institute, Pentlands Science Park, Edinburgh, UK Jai A. Evans Lockheed-Martin Corp, Washington, DC Sergei A. Ezhevsky San Diego, CA, USA Christopher C. Farnsworth Department of Protein Chemistry, IMMUNEX Corporation, Seattle, WA Giorgio Fassina Biopharmaceuticals, Tecnogen SCPA, Parco Scientifico, Piana di Monte Verna, Italy Bahram Fathollahi Caliper Life Science, Mountain View, CA Loïc Faye CNRS UMR, UFR des SCIENCES, Université de Rouen, France Tanja Feilner Genetics and Biotechnology Laboratory, Department of Biochemistry, UCC, Cork, Ireland Joseph Fernandez Protein/DNA Technology Center, Rockefeller University, New York, NY Carlos Fernandez-Patron Canadian Institutes of Health Research and Heart and Stroke Foundation of Canada, Department of Biochemistry, University of Alberta, Edmonton, Alberta, Canada Harold A. Fisk Department of Molecular Genetics, The Ohio State University, Columbus, OH Anne-Catherine Fitchette CNRS UMR, IFRMP 23, Université de Rouen, Faculté des Sciences, Mont Saint Aignan, France Barbara A. Fogarty Life Sciences Interface, Tyndall National Institute, Cork, Ireland Angeliki Fotinopoulou Department of Clinical Biochemistry, The Medical School, University of Newcastle, Newcastle upon Tyne, UK Ruth R. French Tenovus Research Laboratory, Cancer Sciences Division, Southampton University School of Medicine, Southampton, UK Johannes Freund Institute of Chemistry and Biochemistry, Unnology Group, University of Salzburg, Austria David B. Friedman Proteomics Laboratory, Mass Spectrometry Research Center, Department of Biochemistry, Vanderbilt University School of Medicine, Nashville, TN Thomas D. Friedrich Center for Immunology and Microbial Disease, Albany Medical College, Albany, NY Michael J. Dunn

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Department of Biochemistry, Biozentrum of the University of Basel, Basel, Switzerland Jeban Ganesalingam Department of Clinical Neurology, Institute of Psychiatry, King’s College, London, UK Christina Gasbarro EIC Laboratories, Inc., Norwood, MA Michael H. Gelb Departments of Chemistry and Biochemistry, University of Washington, Seattle, WA Kris Gevaert Department of Biochemistry, Faculty of Medicine and Health Sciences, Ghent University, Ghent, Belgium Bart Ghesquière Department of Biochemistry, Faculty of Medicine and Health Sciences, Ghent University, Ghent, Belgium Elisabetta Gianazza Dipartimento di Scienze Farmacologiche, Università degli Studi di Milano, Milan, Italy John A. Glomset Howard Hughes Medical Institute, University of Washington, Seattle, WA Radoslav Goldman Department of Oncology, Georgetown University, Washington, DC Véronique Gomord CNRS UMR, Faculté des Sciences, Université de Rouen, France Mohammad T. Goodarzi Department of Clinical Biochemistry, The Medical School, University of Newcastle, Newcastle upon Tyne, UK Patricia Gravel Triskel Integrated Services SA, Geneva, Switzerland Marion L. Greaser Muscle Biology Laboratory, University of WisconsinMadison, Madison, WI Todd M. Greco Stokes Research Institute, Children’s Hospital of Philadelphia, Philadelphia, PA Susanna F. Greer Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill, Chapel Hill, NC. Current address: Department of Biology, Georgia State University, Atlanta, GA Karlfried Groebe ProteoSys AG, Mainz, Germany Isabelle Guillemin Tierphysiologie, TU Kaiserslauternm, Germany Sunita Gulati Maxwell Finland Lab for Infectious Diseases, Boston, MA Erol E. Gulcicek Keck Protein Chemistry and Mass Spectrometry, Yale University, New Haven, CT Christine Hager-Braun NIEHS, Research Triangle Park, NC Peter Hammerl Institute of Chemistry and Biochemistry, Immunology Group, University of Salzburg, Austria Karin Hansson Department of Clinical Chemistry, University Hospital, Lund University, Malmö, Sweden Arnulf Hartl Institute of Chemistry and Biochemistry, Immunology Group, University of Salzburg, Austria Stefan Gander

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Contributors

Oxford Glycobiology Institute, Department of Biochemistry, University of Oxford, Oxford, UK Rosaria P. Haugland Molecular Probes, Inc., Eugene, OR Lars Hennig ETH Zurich, Zurich, Switzerland William Henzel Genentech Inc., South San Francisco, CA John R. Hepler Department of Pharmacology, Emory University School of Medicine, Atlanta, GA Hisashi Hirano Yokohama City University, International Graduate School of Arts and Sciences, Tsurumi Yokohama, Japan Peter Højrup Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odensk, Denmark Andrew D. Hooker Sittingbourne Research Centre, Pfizer Ltd, Analytical Research and Development (Biologics), Sittingbourne, UK Martin Horst STRATEC Medical, Oberdorf, Switzerland Elizabeth F. Hounsell School of Biological and Chemical Sciences, Birbeck University of London, UK Chrstian Hunzinger† ProteoSys AG, Mainz, Germany. Current Address: Analytical Development API and Bioanalytics, Merck KGaA, Darmstadt, Germany. Sun-Young Hwang College of Pharmacy and Research Institute of Drug Development, Chonnam National University, Gwangju, South Korea G. Brent Irvine Division of Psychiatry and Neuroscience, School of Medicine and Dentistry, Queen’s University Belfast, Belfast, UK Harry Ischiropoulos Stokes Research Institute and Department of Pharmacology, Children’s Hospital of Philadelphia, Philadelphia, PA Crescent R. Isham Department of Medical Oncology, Mayo Clinic, Rochester, MN Satsuki Itoh Division of Biological Chemistry and Biologicals, National Institute of Health Sciences, Kamiyoga, Setagaya-ku, Tokyo, Japan David C. James Sittingbourne Research Centre, Pfizer Ltd, Analytical Research and Development (Biologics), Sittingbourne, UK Tharappel C. James Dublin, Ireland Petr Jandik Dionex Corporation, Sunnyvale, CA Paul Jenö Department of Biochemistry, Biozentrum of the University of Basel, Basel, Switzerland Catherine Jepson Moredun Research Institute, Pentlands Science Park, Edinburgh, UK Li-Tai Jin School of Pharmacy, Wenzhou Medical College, Wenzhou, Zhejiang, South Korea Anders H. Johnsen Department of Clinical Biochemistry, Rigshospitalet, Copenhagen University Hospital, Copenhagen, Denmark David J. Harvey

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William Jordan

Department of Immunology, ICSM, Hammersmith Hospital,

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London, UK Division of Biological Science, University of Montana, Missoula, MT Roza Maria Kamp Technische Fachhochschule Berlin, University of Applied Sciences, FBV/Studiengang Biotechnologie, Berlin, Germany Nnennaya Kanu Department of Biochemistry, University of Hong Kong, Pofulam, Hong Kong Christopher Kasbek Department of Molecular Genetics, The Ohio State University, Columbus, OH Scott H. Kaufmann Oncology Research, Department of Molecular Pharmacology, Mayo Clinic,Rochester, MN Nana Kawasaki Division of Biological Chemistry and Biologicals, National Institute of Health Sciences, Kamiyoga, Setagaya-ku, Tokyo, Japan Birgit Kersten RZPD German Resource Center for Genome Research GmbH, Berlin, Germany John C. Klock Glyko Inc, Navato, CA Joachim Klose Institute for Human Genetics, Charite University Medicine Berlin, Berlin, Germany Achim Knappik MorphoSys AG, Martinsried, Germany Sunil Kochhar Nestlé Research Center, Lausanne, Switzerland Naoki Komori Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK Nicholas J. Kruger Department of Plant Sciences, University of Oxford, Oxford, UK Nathan A. Lacher Pfizer Research, St. Louis, MO William J. LaRochelle CuraGen Corporation, Branford, CT Robert R. Latek Whitehead Institute for Biomedical Research, Cambridge, MA Ralph C. Judd

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Michèle P. Learmonth School of Biological Sciences, University of Edinburgh, Edinburgh, UK André Le Bivic IBDM, Faculté des Sciences de Luminy, Marseille, France Yean Kit Lee Division of Medical Oncology, Mayo Clinic, Rochester, MN John M. Lehman Department of Pathology and Laboratory Medicine, Brody ●

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School of Medicine, East Carolina University, Greenville, NC Cécile Lelong CEA/DSV/iRTSV/LBBSI, CEA Grenoble, Grenoble, France Peter F. Lemkin North Potomac, MD Daniel C. Liebler Department of Biochemistry and Mass Spectrometry ●

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Research Center, Vanderbilt University School of Medicine, Nashville, TN Microchemistry and Proteomics Laboratory, Genentech Inc., South San Francisco, CA

Jennie R. Lill

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Holger J. Møller Aarhus University Hospital, Aarhus, Denmark Glenn E. Morris Wolfson Centre for Inherited Neuromuscular Disease, ●

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RJAH Orthopaedic Hospital, Oswestry, Shropshire, UK; Department of Biochemistry, Institute for Science and Technology in Medicine, Keele University, UK Chrisian Motz Institute for Physiological Chemistry, Ruhr University Bochum, Bochum,, Germany Jacek Mozdzanowski GlaxoSmithKline, King of Prussia, PA Hikaru Nagahara International University of Health and Welfare, Sanno Hospital, Minato-ku, Tokyo, Japan Joakim Norbeck Department of Chemical and Biological Engineering/ Molecular Imaging and Biotechnology, Chalmers University of Technology, Gothenburg, Sweden Ole Nørregard Jensen Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odense, Denmark Hans Gerd Nothwang Abt. Neurogenetik, Institut für Biologie und Umweltwissenschaften, Carl von Ossietzky Universität Oldenburg, Oldenburg, Germany Masamichi Oh-Ishi Kitasato University School of Science, Sagamihara, Kanagawa, Japan David Oxley Proteomics Research Group, Babraham Institute, Babraham, Cambridge, UK Thomas Paccalet CNRS UMR, Faculté des Sciences, Université de Rouen, France Nicolle H. Packer CORE of Functional Proteomics and Cellular Networks, Department of Chemistry and Biomolecular Sciences, Macquarie University, North Ryde, Sydney, NSW, Australia Mark Page Apovia, Inc., San Diego, CA Giovanna Palombo Biopharmaceuticals, Tecnogen SCPA, Parco Scientifico, Piana di Monte Verna, Italy Bart Panis K.U. Leuven, Faculty of Bioscience Engineering, Department of Biosystems, Leuven, Belgium Carol E. Parker UNC-Duke Michael Hooker Proteomics Center, University of North Carolina at Chapel Hill, Chapel Hill, NC Ok-Kyong Park-Sarge Department of Physiology, University of Kentucky, Lexington, KY Wayne F. Patton Molecular Probes Inc., Eugene, OR Katherine H. Pedone Department of Pharmacology, Emory University, Atlanta, GA Victoria Pham Genentech Inc, South San Francisco, CA ●

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Department of Clinical Biochemistry, Aarhus University Hospital, Aarhus, Denmark Slobodan Poznanovic´ ProteoSys AG, Mainz, Germany Thierry Rabilloud iRTSV/LBBSI, CEA Grenoble, Grenoble, France Geert Jan Rademaker Department of Biochemistry, University of Hong Kong, Pofulam, Hong Kong Joachim Rassow Institute for Physiological Chemistry, Ruhr University Bochum, Bochum, Germany F. Andrew Ray Department of Environmental and Radiological Health Sciences, Colorado State University, Fort Collins, CO Andrew J. Reason Managing Director, M-Scan Ltd - Mass Spectrometry Consultants and Analysts, Fishponds Close, Wokingham, Berkshire, UK Jens F. Rehfeld Department of Clinical Biochemistry, Rigshospitalet, Copenhagen University Hospital, Copenhagen, Denmark David V. Renouf Department of Biochemistry, University of Hong Kong, Pofulam, Hong Kong Habtom W. Ressom Department of Oncology, Georgetown University, Washington, DC Jeffrey Rohrer Dionex Corporation, Life Science Research Group, Sunnyvale, CA Douglas D. Root Department of Biological Sciences, University of North Texas, Denton, TX Menotti Ruvo Biopharmaceuticals, Tecnogen SCPA, Parco Scientifico, Piana di Monte Vernan, Italy Kevin D. Sarge Department of Molecular and Cellular Biochemistry, University of Kentucky, Lexington, KY Chaturvedula S. Sastri ProteoSys AG, Mainz, Germany Allan Saul Laboratory of Malaria and Vector Research, NIAID, NIH, Bethesda, MD Jens Schindler Institut für Physiologie II, Albert-Ludwigs-Universität Freiburg, Freiburg, Germany Gerhard P. Schwall ProteoSys AG, Mainz, Germany Alexander Schwarz Biosphere Medical Inc., Rockland, MA Jette W. Sen Department of Clinical Biochemistry, Rigshospitalet, Copenhagen University Hospital, Copenhagen, Denmark Bryan John Smith CellTech, Slough, Berkshire, UK Kevin D. Smith School of Biological and Chemical Sciences, Birkbeck University of London, UK Ralph L. Smith Department of Pathology and Laboratory Medicine, Brody School of Medicine, East Carolina University, Greenville, NC David W. Speicher Systems Biology Division, Proteomics Laboratory, The Wistar Institute, Philadelphia, PA Jørgen H. Poulsen

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Jennifer D. Tibodeau

Department of Medical Oncology, Mayo Clinic,

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Rochester, MN Mass Spectrometry Group, Laboratory of Structural Biology, Research Triangle Park, NC Rocky S. Tuan Department of Orthopaedic Surgery, Department of Biochemistry and Molecular Pharmacology, Cell and Tissue Engineering Graduate Program, Thomas Jefferson University, Philadelphia, PA Jeremy E. Turnbull University of Liverpool, Liverpool, UK Graham A. Turner Department of Clinical Biochemistry, The Medical School, University of Newcastle, Newcastle upon Tyne, UK Brian B. Vandahl Novo Nordisk A/S, Maaloev, Denmark Joël Vandekerckhove Department of Biochemistry, Faculty of Medicine and Health Sciences, Ghent University, Ghent, Belgium Rency S. Varghese Department of Oncology, Lombardi Comprehensive Cancer Center, Georgetown University, Washington DC Antonio Verdoliva Biopharmaceuticals, Tecnogen SCPA, Parco Scientifico, Piana di Monte Verna, Italy Yoshinao Wada Research Institute, Osaka Medical Center for Maternal and Child Health, Osaka University Graduate School of Medicine, Osaka, Japan Charles J. Waecheter Department of Molecular and Cellular Biochemistry, University of Kentucky College of Medicine, Lexington, KY John M. Walker School of Life Sciences, University of Hertfordshire, Hatfield, Hertfordshire, UK Kuan Wang Laboratory of Muscle Biology, NIH, NIAMS/LPB, Bethesda, MD Rong Wang Department of Genetics and Genomic Sciences, Mount Sinai School of Medicine, New York, NY Malcolm Ward Proteome Sciences plc, King’s College London, UK Chad M. Warren Department of Physiology, University of Illinois at Chicago, Chicago, IL Maria Warren Hines Program in Molecular Biology and Biotechnology, University of North Carolina at Chapel Hill, Chapel Hill, NC Jakob H. Waterborg School of Biological Sciences, UMKC, Kansas City, MO Darin J. Weber Department of Biochemistry and Biophysics, Oregon State University, Corvallis, OR Judith Webster Proteomics Group, Babraham Institute, Babraham, Cambridge, UK Michael Weitzhandler Dionex Corporation, Life Science Research Group, Sunnyvale, CA Kenneth B. Tomer

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The Pasarow Mass Spectrometry Laboratory, The NPI-Semel Institute, David Geffen School of Medicine, University of California Los Angeles, Los Angeles, CA Jeff Wilkesman Departmento Química, Facultad de Ciencias y Tecnología, Universidad de Carabobo, Carabobo, Venezuela Kenneth R. Williams Keck Protein Chemistry and Mass Spectrometry, Yale University, New Haven, CT Matthias Wilm UCD Conway Institute of Biomolecular and Biomedical Research, University College Dublin, Dublin, Ireland G. Brian Wisdom School of Medicine and Dentistry, The Queen’s University of Belfast, Belfast, Northern Ireland, UK Gary E. Wise Department of Pathology, Temple University Hospital, Philadelphia, PA Nicky K. C. Wong Department of Biochemistry, University of Hong Kong, Pofulam, Hong Kong Wojciech Wozny ProteoSys AG, Mainz, Germany Teruhide Yamaguchi Division of Biological Chemistry and Biologicals, National Institute of Health Sciences, Kamiyoga, Setagaya-ku, Tokyo, Japan Kazuo Yamamoto Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa, Chiba, Japan Ching-Hui Yang Department of Molecular Genetics, The Ohio State University, Columbus, OH Wendy W. You Department of Biochemistry and Biophysics, Oregon State University, Corvallis, OR Jennifer J. Young School of Life Sciences, University of Hertfordshire, Hatfield, Hertfordshire, UK Claus Zabel Institute for Human Genetics, Charite– niversity Medicine Berlin, Berlin, Germany Helmut Zengerling ProteoSys AG, Mainz, Germany Xianquan Zhan Department of Ophthalmic Research, Cole Eye Institute, Cleveland Clinic, Cleveland, OH Julian P. Whitelegge

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1 Protein Determination by UV Absorption Alastair Aitken and Michèle P. Learmonth 1. Introduction 1.1. Near UV Absorbance (280 nm) Quantitation of the amount of protein in a solution is possible in a simple spectrometer. Absorption of radiation in the near UV by proteins depends on the Tyr and Trp content (and to a very small extent on the amount of Phe and disulfide bonds). Therefore the A280 varies greatly between different proteins (for a 1 mg/mL solution, from 0 up to 4 [for some tyrosine-rich wool proteins], although most values are in the range 0.5–1.5 [1]). The advantages of this method are that it is simple, and the sample is recoverable. The method has some disadvantages, including interference from other chromophores, and the specific absorption value for a given protein must be determined. The extinction of nucleic acid in the 280-nm region may be as much as 10 times that of protein at their same wavelength, and hence, a few percent of nucleic acid can greatly influence the absorption. 1.2. Far UV Absorbance The peptide bond absorbs strongly in the far UV with a maximum at about 190 nm. This very strong absorption of proteins at these wavelengths has been used in protein determination. Because of the difficulties caused by absorption by oxygen and the low output of conventional spectrophotometers at this wavelength, measurements are more conveniently made at 205 nm, where the absorbance is about half that at 190 nm. Most proteins have extinction coefficients at 205 nm for a 1 mg/mL solution of 30–35 and between 20 and 24 at 210 nm (2). Various side chains, including those of Trp, Phe, Tyr, His, Cys, Met, and Arg (in that descending order), make contributions to the A205 (3). The advantages of this method include simplicity and sensitivity. As in the method outlined in Subheading 3.1. the sample is recoverable and in addition there From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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is little variation in response between different proteins, permitting near-absolute determination of protein. Disadvantages of this method include the necessity for accurate calibration of the spectrophotometer in the far UV. Many buffers and other components, such as heme or pyridoxal groups, absorb strongly in this region. 2. Materials 1. 2. 3. 4. 5. 6.

0.1 M K2SO4 (pH 7.0). 5 mM potassium phosphate buffer, pH 7.0. Nonionic detergent (0.01% Brij 35) Guanidinium-HCl. 0.2-µm Millipore (Watford, UK) filter. UV-visible spectrometer: The hydrogen lamp should be selected for maximum intensity at the particular wavelength. 7. Cuvets, quartz, for 0.01 mM >1 mM >0.01 mM >1 mM >0.0005% >1 mM >1 mM >0.01 mM >1 mM >0.01 mM >0.01 mM >0.01 mM

5. Kinetic silver staining is based on measurements of light scattering. Thus, other wavelengths may be used, but the corresponding optical density reading will be different and may require optimization. 6. The cellulose may require washing to remove solvents or buffers, if present prior to staining. References 1. Somerville, L. L. and Wang, K. (1981) The ultrasensitive silver “protein” stain also detects nanograms of nucleic acids. Biochem. Biophys. Res. Commun. 102, 53–58. 2. Gottlieb, M. and Chavko, M. (1987) Silver staining of native and denatured eucaryotic DNA in agarose gels. Analyt. Biochem. 165, 33–37. 3. Root, D. D. and Reisler, E. (1990) Copper iodide staining and determination of proteins adsorbed to microtiter plates. Analyt. Biochem. 186, 69–73. 4. Root, D. D. and Wang, K. (1993) Kinetic silver staining and calibration of proteins adsorbed to microtiter plates. Analyt. Biochem. 209, 354–359.

9 Quantitation of Cellular Proteins by Flow Cytometry Thomas D. Friedrich, F. Andrew Ray, Ralph L. Smith, and John M. Lehman 1. Introduction Quantitation of specific proteins in complex mixtures is simplified by the use of antibodies directed against the protein of interest. If the specific protein is differentially expressed within a population of cells, quantitation of the protein in cell lysates by immunoblotting will provide an average quantity of the protein per cell. As a result, when lysates of different cell populations are compared, large changes in the amount of a protein in a small percentage of cells cannot be distinguished from small changes in the amount of a protein in a large percentage of cells. Flow cytometric analysis solves this problem by providing a means to measure the amount of a specific protein within each individual cell in a population. A fluorescently labeled probe, usually an antibody, is required for detection of the protein. A single cell suspension is reacted with an antibody and passed through a flow cytometer, which focuses the cell suspension into a stream that intersects a laser beam. As each cell passes through the beam, the fluorescent probe in each cell is excited and photomultiplier tubes register the degree of fluorescence (1). Multivariate analysis, the simultaneous measurement of multiple parameters, is a powerful feature of flow cytometry. When determining the quantity of a specific protein in a cell, the quantity of other molecules in the same cell can also be measured by using additional fluors specifically targeted to other molecules. Examples of second molecules measured simultaneously include DNA (2–4) or additional proteins (5). In addition, measurement of light scatter can provide information about cell size and granularity. Most flow cytometers are capable of measuring six colors plus light scatter, but specialized flow cytometers that are capable of measuring as many as eighteen colors are now available.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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In this chapter we will describe techniques used to quantitate Simian Virus 40 (SV40) large T antigen (T Ag) in infected cells. By staining infected cells with antibodies against T Ag and with the DNA-binding dye propidium iodide it is possible to measure the amount of T Ag as a function of DNA content. 2. Materials 2.1 Cells and Antibodies The CV-1 line of African Green monkey kidney cells was obtained from the American Type Culture Collection (ATTC, CCL-70). Confluent cultures of CV-1 were infected with Simian Virus 40 strain RH-911 at 100 plaque forming units per cell. Pab101 is a monoclonal antibody specific to the carboxy-terminus of SV40 T Ag. Hybridoma cells producing Pab101 were obtained from ATCC (#TIB-117). Alexa conjugated secondary antibodies were obtained from Invitrogen. 2.2 Fixation and Staining 1. Ca2+/Mg2+-free phosphate-buffered saline (PBS): dissolve 8.0g NaCl, 0.2g KCl, 1.2g Na2HPO4 and 0.2g KH2PO4 in 1 liter of distilled water (dH2O). Adjust the pH to 7.4, sterilize by passing through a 0.2 µm filter and store at room temperature. 2. Trypsin-EDTA: dilute 10X trypsin/EDTA (Invitrogen) in PBS to give final concentrations of 0.05% trypsin, 0.53 mM EDTA. Filter-sterilize and store at 4°C. 3. Methanol. Store at −20°C. 4. Wash solution (WS): heat inactivate 100 ml of normal goat serum (Invitrogen) at 56°C for 1 hour. Mix with 900 ml PBS, 20 µl Triton X-100 and 1.0g sodium azide. Filter-sterilize and store at 4°C (see Note 1). 5. Propidium iodide (PI): dissolve 10.0 mg PI (Calbiochem) in 100 ml PBS. Add 20 µl Triton X-100 and 0.1 g sodium azide. Protect solution from light and store at 4°C. 6. Ribonuclease: dissolve 100 mg of RNAase A (Sigma) in 100 ml PBS. Boil for 1 hour. Add 20 µl Triton X-100 and 0.1g sodium azide. Store at 4°C.

3. Methods 3.1 Fixation 1. Remove the culture medium and rinse the cells with PBS. If floating and/or mitotic cells are of interest, medium and wash fractions should be saved, pelleted and pooled with attached cells after trypsinization. 2. Prewarm trypsin/EDTA to 37°C and add 1 ml per 60 mm dish. (Adjust proportionally for larger culture areas). Tilt the plate to thoroughly distribute solution and then remove. 3. Once the cells have detached (see Note 2), add 1 ml of WS to the plate and transfer the cells to a 1.5 ml microcentrifuge tube. Remove a small volume for the determination of cell number (see Note 3). Centrifuge remaining cells at 4000 × g for 15 sec. 4. Discard supernatant, resuspend the pellet in 1 ml PBS at 4°C and centrifuge as in step 3.

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5. Discard the supernatant and thoroughly resuspend cell pellet in 0.1 ml cold PBS. 6. Immediately add 0.9 ml methanol (−20°C) (see Note 4), mix and store at −20°C (see Note 5).

3.2 Titration of antibodies It is important that the primary antibody (see Note 6) is present in excess to ensure quantitative measurement of the specific protein (see Note 7). However, excessive antibody can lead to increased background. In order to determine the appropriate antibody concentration, stain parallel samples of cells with serial dilutions of antibody. If available, similar cells that don’t express the protein of interest are used for negative controls. Stain such cells in the same way as the experimental cells. 3.3 Staining 1. Transfer 1.5 × 106 cells to a 1.5 ml microcentrifuge tube and centrifuge at 4000 × g for 15 seconds (see Note 8). 2. Discard the supernatant and resuspend the cell pellet in 1 ml PBS at 4°C. Repeat centrifugation step. 3. Discard PBS and add 0.5 ml of diluted primary antibody. Mix gently. 4. Incubate with gentle agitation (see Note 9). 5. Pellet at 4000 × g for 15 sec and discard supernatant. 6. Resuspend the pellet in 0.5 ml WS and repeat step 5. 7. Resuspend the pellet in 0.5 ml of diluted fluorescently-labeled secondary antibody (see Note 10). Mix gently. 8. Incubate at 37°C for 2 h with gentle mixing; minimize exposure to light. 9. Add 0.5 ml RNAase and mix gently. 10. Incubate at 37°C for 30 min. 11. Add 0.5 ml PI, bringing the total volume to 1ml. Mix gently. 12. Filter each sample through a 53 µm mesh nylon grid (Nitex HC3-53, Tetko Inc.) (see Note 11).

3.4 Flow Cytometry and analysis Most commercially available flow cytometers are capable of multiparameter analysis (see Note 12). Using 20 mW of power tuned to 488 nm, minimize the coefficient of variation (CV) for the green photomultiplier (see Note 13) to G2 phase. Levels of T Ag expression in G1 phase cells cover a 10-fold range, but only cells expressing higher amounts of T Ag enter S phase. >G2 phase represents a virus-regulated override of normal cell cycle events. The cells in >G2 phase are infected cells that are replicating viral DNA and re-replicating cellular DNA (7). The G1 phase cells with fluorescence levels similar to the background levels of the uninfected cells are believed to be an uninfected subpopulation that conveniently serves as an internal control in this system. (C) The relative quantities of T Ag within each cell cycle gate, as determined by anti T Ag staining. The bar graph shows the average quantity of T Ag per cell in the T Ag expressing population.

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5. 6.

7.

8. 9.

10.

11.

12.

13. 14. 15.

Friedrich et al. dehyde followed by methanol has been used to retain antigens lost with methanol fixation alone (6). Fixed cells have been stored at −20°C for >1 year with negligible loss of T Ag. However stability during storage must be evaluated for each antigen. Monoclonal antibodies or affinity purified polyclonal antibodies are preferred. Polyclonal antisera may have additional antibodies of unknown specificity that will increase background staining. Antibody binding is actually a quantitative measure of the epitope recognized by the antibody and not necessarily the total amount of the protein antigen. If the epitope is masked through protein conformation/association or as an artifact of fixation, the epitope may not be detected. Use of a swinging bucket microcentrifuge reduces cell loss. Adjust time and temperature to maximize the signal over background. Incubation times generally range from 30 minutes to 2 hours at 37°C or overnight at 4°C. Gentle rocking is recommended. Primary antibodies that are directly conjugated to fluors are commercially available. In addition, kits that allow conjugation of Alexa fluors to antibodies can be obtained from Invitrogen. If particularly weak signals are encountered, the signal can be amplified by additional layers of fluorescently-tagged antibody. For example, cells stained with an FITC-labeled primary or secondary antibody can be reacted with Alexa 488-labeled rabbit anti-fluorescein followed by Alexa 488-labeled goat anti-rabbit IgG. Filtering of samples removes cell aggregates that may clog the tubing in the flow cytometer. If samples are stored and then reanalyzed, filtering should be repeated. We have used a Cytofluorograph Model 50 H-H with an air-cooled argon laser (model 532, Omnichrome). The Cyclops analysis program (Cytomation) was used for data analysis. More recently a BD FACScan has been utilized with similar settings and filters and identical results were obtained. Use a 535 nm band pass filter for this photomultiplier. Use a 640 nm long pass filter for this photomultiplier. PI stained lymphocytes are prepared by first making a stock of spleen cells from a healthy mouse. Spleens are removed and minced with fine surgical scissors in PBS. The cells in the PBS supernatant are removed and placed in a centrifuge tube. Remaining spleen fragments are minced in PBS until few cells are evident in the supernatants. Cells in the pooled supernatants are pelleted by centrifugation and washed 2X with PBS. The pellet is resuspended in 1ml of PBS, mixed with 9ml of −20° C methanol and stored at −20° C. Prior to each run, wash and stain 1 × 106 cells with PI/RNAse only.

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16. Values obtained from flow cytometric analysis do not provide an absolute quantitation (numbers per cell) of an epitope, but rather represent relative quantities of the epitope within cells in a population. A common method of determining the number of epitopes per cell is to compare the fluorescent intensity of the cell to standard beads that have a known number of primary antibody molecules per bead. The same fluorescently-labeled antibody that is used to stain cells is also reacted with the standard beads under the same conditions. Alternatively, if the number of fluorochrome molecules per antibody molecule is known, a comparison of a cell’s fluorescent intensity to that of standard beads with a known number of bound fluorochrome molecules is possible. Both types of beads are commercially available. References 1. Shapiro, H. M. (2003) Practical Flow Cytometry, 4th Ed. Wiley-Liss, New York 2. Jacobberger, J. W., Fogleman, D., and Lehman, J. M. (1986) Analysis of intracellular antigens by flow cytometry. Cytometry 7, 356–364 3. Laffin, J. and Lehman, J. M. (1994) Detection of intracellular virus and virus products. Meth. Cell Biol. 41, 543–557. 4. Lehman J.M., Friedrich T.D. and Laffin J. (2001). Analysis of viral infection and viral and cellular DNA and proteins by flow cytometry. Curr Protocols Cytometry. 7.17.1–7.17.9. 5. Stewart, C. C. and Stewart, S. J. (1994) Multiparameter analysis of leukocytes by flow cytometry. Meth. Cell Biol. 41, 61–79. 6. Sramkoski, R.M., Wormsley, S.W., Bolton, W.E., Crumpler, D.E. and Jacobberger, J.W. (1999) Simultaneous Detection of Cyclin B1, p105, and DNA Content Provides Complete Cell Cycle Phase Fraction Analysis of Cells That Endoreduplicate. Cytometry 35, 274–283. 7. Lehman, J. M., Laffin, J., and Friedrich, T. D. (2000) Simian virus 40 induces multiple S phases with the majority of viral DNA replication in the G2 and second S phase in CV-1 cells. Exp. Cell Res. 258, 215–222.

10 Quantitation of Cellular Proteins by Laser Scanning Cytometry Thomas D. Friedrich, Ralph L. Smith, and John M. Lehman 1. Introduction Flow Cytometry (FC) is a popular method for the simultaneous quantitation of multiple specific proteins and other molecules in each cell of a single cell suspension (1). Laser Scanning Cytometry (LSC) is a closely related technology that provides many of the benefits of FC and offers additional features that allow morphological and biochemical analysis of a cell population (2,3). In both methods, quantitation of endogenous cellular proteins generally requires antibodies against the protein of interest. These antibodies must be either directly labeled with a fluorochrome or detected by a fluorescently-labeled secondary antibody. Using fluorescently-labeled antibodies in combination with fluorescent dyes that quantitatively bind to DNA, both FC and LSC have been valuable in measuring the quantities of specific proteins in relation to cell cycle position (4). In contrast to FC, where a single cell suspension is passed through a fixed-position laser beam, LSC analyzes cells fixed to a surface such as a microscope slide or cover slip. Since the position of the cell on the slide is part of the recorded data, it is possible to visualize the cell after the initial recording event or to subsequently stain the cell with another fluorescent probe and make a second measurement. LSC also allows analysis of cells following biochemical treatments that are incompatible with analysis by FC. In this chapter we describe the value of LSC in analyzing chromatin-associated proteins. Specifically, we describe an in situ fractionation procedure that allows measurement of total and chromatin-associated MCM2, a DNA helicase subunit, as a function of cell cycle position.

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2. Materials 2.1. Cells and Antibodies The CV-1 line of African Green monkey kidney cells was obtained from the American Type Culture Collection (ATCC, CL-70). Cells were maintained in MEM/ 5% fetal bovine serum. BM28 (#610701), a monoclonal antibody that specifically recognizes MCM2 was obtained from BD Biosciences. Alexa-488 conjugated goat anti-mouse IgG (#A-11001) was obtained from Invitrogen. 2.2. Fixation and Staining 1. Ca2+/Mg2+-free phosphate-buffered saline (PBS): Dissolve 8.0g NaCl, 0.2g KCl, 1.2g Na2HPO4 and 0.2g KH2PO4 in 1 liter of distilled water (dH2O). Adjust the pH to 7.4, sterilize by passing through a 0.2 µm filter and store at room temperature. 2. Wash solution (WS): heat inactivate 100 ml of normal goat serum (Invitrogen) at 56°C for 1 hour. Mix with 900 ml PBS, 20 µl Triton X-100 and 1.0g sodium azide. Filter-sterilize and store at 4°C (see Note 1). 3. Propidium iodide (PI): dissolve 10.0 mg PI (Calbiochem) in 100 ml PBS. Add 20 µl Triton X-100 and 0.1 g sodium azide. Protect solution from light and store at 4°C. 4. Ribonuclease: dissolve 100 mg of RNAase A (Sigma) in 100 ml PBS. Boil for 1 hour. Add 20 µl Triton X-100 and 0.1 g sodium azide. Store at 4°C.

2.3. Cell Extraction 1. 2X CSK buffer (200 ml): bring 0.6 g PIPES, 20.5 g sucrose, (see Note 2), 4 ml 5 M NaCl and 0.6 ml of 1M MgCl2 to total volume of 190 ml with H2O. The solution will clear, with stirring, while adjusting to pH 7.0 with 10N NaOH. Add H2O to a final volume of 200 ml. (1X CSK= 10 mM PIPES pH7.0, 100 mM NaCl, 300 mM sucrose, 3 mM M gCl2 ). 2. 100X protease inhibitor cocktail: dissolve 2.5 mg leupeptin, 2.5 mg aprotinin, 15.0 mg benzamidine and 1.0 mg trypsin inhibitor (all protease inhibitors were from Sigma) in 1.0 ml H20. Store in 0.1 ml aliquots at −20° C. 3. CSK/0.5% Triton X-100. For 5 ml, combine 2.5 ml 2X CSK buffer, 2.15 ml H20, 0.25 ml 10% Triton X-100, 0.05 ml 100 mM PMSF and 0.05 ml 100X protease inhibitor cocktail.

3. Methods 3.1. CSK Extraction and Fixation This procedure was adapted from reports describing the use of a selective extraction procedure to demonstrate the release of MCM proteins from chromatin in S phase cells (6,7,8). 1. Cells can be grown in culture slides or on glass cover slips placed in tissue culture plates. For the purpose of this study, CV-1 cells were grown on 12 mm circular glass cover slips in 60 mm plates. 2. At time of harvest, aspirate medium and wash plate with 5 ml PBS.

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3. Remove cover slips with fine-tipped forceps and place cell-side up on a parafilm covered glass plate in a humidified chamber. (see Note 3) 4. Gently place 150 µl CSK buffer on each cover slip (see Note 4). 5. Aspirate CSK and replace with 100-150 µl CSK/0.5% Triton X-100; keep in humidified chamber for 5 min. 6. Aspirate CSK/0.5% Triton X-100 and gently wash 3X with CSK buffer (see Note 5). 7. Fix immediately by transferring cover slips to Alumina staining racks (Thomas Scientific) submerged in methanol at −20°C (see Note 6). 8. Allow cover slips to air dry; either store at −20°C or continue with antibody staining.

3.2. Titration of Antibodies Quantitative measurement of a specific protein requires that antibody be present in excess. Yet, the background of nonspecific antibody staining should be kept to a minimum. To determine the appropriate antibody concentration, stain parallel cover slips with serial dilutions of antibody. If similar cells not expressing the protein of interest are available, they can be stained at the same dilutions to determine levels of nonspecific staining. Background staining due to nonspecific binding by the primary antibody can also be measured in cells stained with nonspecific antibody of the same isotype. Background staining due to the fluorescent secondary antibody can be measured in cells stained with the secondary antibody only. 3.3. Protein and DNA Staining 1. Rehydrate dried cover slips by placing cell-side up on a parafilm covered glass plate in a humidified chamber and covering with 150 µl wash solution. 2. Remove WS and layer 20-50 µl of primary antibody diluted in WS onto each cover slip. Incubate 30 min at 37°C (see Note 7). 3. Rinse cover slips 3X by gentle addition and aspiration of 150 µl wash solution. 4. Layer 20-50 µl of fluorescently labeled secondary antibody onto each cover slip and incubate 30 min at 37°C (see Note 8). 5. Rinse cover slips 5X by gentle addition and aspiration of 150 µl wash solution. 6. Combine equal volumes of PI solution and RNAase solution. Incubate each cover slip with 20-50 µl of the mixture for 30 min at room temperature. 7. Briefly rinse cover slip by dipping in H20 and drain by touching edge of cover slip to a Kimwipe. Prepare mounting medium by mixing 90% glycerol/ 10% PBS and then diluting 1:1 with PI solution. Apply 5-10 µl of mounting media to a glass microscope slide and mount cover slip cell side down (see Note 9).

3.4. Laser Scanning Cytometry and Data Analysis The data was acquired with an iCys Research Imaging Cytometer manufactured by CompuCyte, Cambridge, MA, which is capable of scanning micro plates, slides, and other carriers. This microscope-based instrument uses three lasers (488 nm argon, 633 nm helium neon, and 405 nm violet diode), dichroic

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mirrors, optical filters (650 LP for CY5, 580/30 for PE, 530/30 for FITC, and 463/39 for DAPI), and photomultipliers (PMTs). Software utilized was the iGeneration Cytometric Analysis Software Version 3.2.5. Instrument settings were 5 mW laser power (488nm argon) with the detector voltages set at 35 V green channel with a 530/30 optical filter and 33 V long red channel with a 650 LP optical filter. For reanalysis, the contour primary minimum area of 20 µm2 was set to exclude cell fragments and a maximum area of 340 µm2 to allow doublet discrimination. The instrument will acquire the data and record the location of each cell. Therefore, after the data is acquired, stored and analyzed, the cells may be viewed in a gallery for their morphology and surrounding cells. Cells can be restained with other fluorochromes/antibodies if required. The procedure detailed above was used to examine the expression and nuclear association of the DNA helicase subunit, MCM2, in proliferating monkey kidney cells. Figure 1A shows MCM2 (y axis) vs DNA staining (x axis) of cells directly fixed in methanol. The greatest range in MCM2 expression is observed in G1 phase cells. Cells in S and G2 phases uniformly express higher levels of MCM2. Panel B shows cells that were extracted with CSK/0.5% Triton X-100 to remove soluble proteins prior to methanol fixation. A striking characteristic of extracted S phase cells is the decrease in MCM2 as cells increase in DNA content, whereas the directly fixed cells have consistently high levels of MCM2 throughout S phase. The conclusion from this analysis is that although the level of nuclear MCM2 remains constant throughout S phase, the increase in DNA content during S phase directly correlates with release of chromatin-bound MCM2. The release of MCM proteins as a consequence of replication origin firing has been described previously in biochemical and immunofluorescent microscopy studies (6-8). However, the multiparameter capabilities of LSC provide the advantage of simultaneous quantitation of MCM2 protein and DNA (5). Another powerful feature of LSC, which is not possible with FC, is the ability to visualize laser scan images of each event assembled in an image gallery. This is particularly useful in further defining specific events and eliminating undesirable events from the scattergrams. An example is seen in panel B where there appear to be subpopulations of extracted cells that retain MCM2 in G2/M phase and higher DNA contents. Examination of an image gallery revealed that these events were not single cells, but doublets and clusters of multiple nuclei. The acquired data (panels A and B), were reanalyzed to eliminate doublets and large nuclear clusters based on their area (see Note 10). Morphological identification of the cells from image galleries of the G2/M regions of panels B and D is presented is presented in panels E and F, respectively. Only nuclei are apparent due to visualization of the PI stain. Cells viewed in panel E prior to exclusion of events greater than 340 µm2 exhibit numerous doublets and multiple clusters, while the cells viewed following exclusion (panel F) are single cells. The scattergrams in panels C and D exclude events greater than 340 µm2 from panels A

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Fig. 1. Growing CV-1 monkey kidney cells stained for MCM2 (y axis) and DNA content (x axis). (A) Cells were directly fixed in methanol and then stained for MCM2 and DNA. (B) Parallel cover slips were extracted with CSK/0.5% Triton X-100 prior to fixation and staining. To eliminate doublet and larger nuclear clusters from the scattergrams, PI-stained events with an area greater than 340 µm2 were subtracted from the scattergrams A and B to generate the distributions in (C) directly fixed and (D) extracted. (E) A gallery of nuclei excluded by elimination of events greater than 340 µm2 from the G2/M phase region. (F) A gallery of nuclei with areas between 20 and 340 µm2 from the G2/M phase region.

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and B, respectively, and significantly reduce two artifactual results. First, cells that appear to have DNA contents greater than that of a G2/M phase cell are nearly eliminated. Second, a cell subpopulation that appears to retain MCM2 binding in G2/M and cells of higher DNA content is markedly decreased. The ability to perform in situ extractions brings elements of biochemical fractionation to the field of quantitative cytometry. Additional fractionation procedures or other biochemical manipulations should be feasible as long as primary cell morphology and the epitope being examined are not destroyed. In addition, the capacity of this instrumentation to reanalyze data provides a powerful tool to determine and characterize multiple parameters of the collected data. 4. Notes 1. Sodium azide is highly toxic. Consult the MSDS for proper handling instructions. 2. Use a molecular biology-grade sucrose, such as Sigma S0389. 3. A simple humidified chamber can be made from a plastic or glass 150 mm Petri dish. Filter paper or paper towel is cut to fit inside the lid and moistened with H20. A glass plate such as an 8 × 10 cm minigel plate is wrapped in parafilm and placed on the moistened paper. The base of the dish is used as the lid. 4. An alternative to individual treatment of cover slips is to leave cover slips in the tissue culture plate and treat with larger volumes of extraction and wash buffers. 5. Cells treated with CSK/0.5% Triton X-100 are more susceptible to detachment from the cover slip. Extra care must be taken in adding and removing buffer from the cover slip. 6. 70% acetone/30% methanol has also been successfully used for fixation of MCM2 (5). Appropriate fixation conditions will need to be optimized for each epitope. 7. Longer periods of incubation (1-2 hours) at room temp may also be used and should be optimized for each epitope. 8. To protect fluorescently labeled antibody from light, use aluminum foil to wrap the base and lid of the humidified chamber, described in Note 3. 9. If more permanent mounting is required, the cover slips can be mounted in ProLong (Invitrogen) supplemented with PI. 10. Doublets and clusters of multiple nuclei can be excluded based on the area (measured in square microns) of each event. For example, two G1 phase nuclei in close proximity may be recorded as a single event. Although the DNA content of the two G1 phase nuclei will be recorded as equivalent to a G2 phase nucleus, the area of the two G1 nuclei will be greater than that of the G2 nucleus. In this case, doublets and larger nuclear clusters

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were eliminated by excluding events with an area greater than 340 µm2. The optimal exclusion threshold should be determined empirically for each experimental group. References 1. Shapiro H.M. (2003) Practical Flow Cytometry, 4th Ed. Wiley-Liss, New York. 2. Luther E., Kamentsky, L., Henriksen, M., and Holden, E. (2004) Next-generation laser scanning cytometry. Meth. Cell Biol. 75, 185–218. 3. Kamentsky L.A. and Kamentsky L.D. (1991) Microscope-based multiparameter laser scanning cytometer yielding data comparable to flow cytometry data. Cytometry 12, 381–387. 4. Darzynkiewicz Z., Smolewski P., and Bedner E. (2001) Use of flow and laser scanning cytometry to study mechanisms regulating cell cycle and controlling cell death. Clin. Lab. Med. 21, 857–873. 5. Friedrich T.D., Bedner, E., Darzynkiewicz, Z., and Lehman J.M. (2005) Distinct Patterns of MCM protein binding in nuclei of S phase and rereplicating SV40infected monkey kidney cells. Cytometry 67A, 10–18. 6. Todorov I.T., Attaran A., and Kearsey S.E. (1995) BM28, a human member of the MCM2-3–5 family, is displaced from chromatin during DNA replication. J. Cell Biol. 129, 1433–1445. 7. Krude T., Musahl C., Laskey R.A., and Knippers R. (1996) Human replication proteins hCdc21, hCdc46 and P1Mcm3 bind chromatin uniformly before S-phase and are displaced locally during DNA replication. J. Cell Sci. 109, 309–318. 8. Stoeber K., Tlsty T.D., Happerfield L., Thomas G.A., Romanov S., Bobrow L., Williams E.D., and Williams G.H. (2001) DNA replication licensing and human cell proliferation. J. Cell Sci. 114, 2027–2041.

11 Protein Solubility in Two-Dimensional Electrophoresis Basic Principles and Issues Thierry Rabilloud 1. Introduction The solubilization process for two-dimensional (2-D) electrophoresis has to achieve four parallel goals: 1. Breaking macromolecular interactions in order to yield separate polypeptide chains. This includes denaturing the proteins to break noncovalent interactions, breaking disulfide bonds, and disrupting noncovalent interactions between proteins and nonproteinaceous compounds such as lipids or nucleic acids. 2. Preventing any artefactual modification of the polypeptides in the solubilization medium. Ideally, the perfect solubilization medium should freeze all the extracted polypeptides in their exact state prior to solubilization, both in terms of amino acid composition and in terms of posttranslational modifications. This means that all the enzymes able to modify the proteins must be quickly and irreversibly inactivated. Such enzymes include of course proteases, which are the most difficult to inactivate, but also phosphatases, glycosidases, and so forth. In parallel, the solubilization protocol should not expose the polypeptides to conditions in which chemical modifications (e.g., deamidation of Asn and Gln, cleavage of Asp-Pro bonds) may occur. 3. Allowing the easy removal of substances that may interfere with 2-D electrophoresis. In 2-D electrophoresis, proteins are the analytes. Thus, anything in the cell but proteins can be considered as an interfering substance. Some cellular compounds (e.g., coenzymes, hormones) are so dilute they go unnoticed. Other compounds (e.g., simple nonreducing sugars) do not interact with proteins or do not interfere with the electrophoretic process. However, many compounds bind to proteins and/or interfere with 2-D electrophoresis and must be eliminated prior to electrophoresis if their amount exceeds a critical interference threshold. Such compounds mainly include salts, lipids, polysaccharides (including cell walls), and nucleic acids.

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4. Keeping proteins in solution during the 2-D electrophoresis process. Although solubilization stricto sensu stops at the point where the sample is loaded onto the first dimension gel, its scope can be extended to the 2-D process per se, as proteins must be kept soluble till the end of the second dimension. Generally speaking, the second dimension is an SDS gel, and very few problems are encountered once the proteins have entered the SDS-polyacrylamide gel electrophoresis (SDS PAGE) gel. The one main problem is overloading of the major proteins when micropreparative 2-D electrophoresis is carried out, and nothing but scaling-up the SDS gel (its thickness and its other dimensions) can counteract overloading an SDS gel. However, severe problems can be encountered in the isoelectric fusing (IEF) step. They arise from the fact that IEF must be carried out in low ionic strength conditions and with no manipulation of the polypeptide charge. IEF conditions give problems at three stages: a. During the initial solubilization of the sample, important interactions between proteins of widely different pI and/or between proteins and interfering compounds (e.g., nucleic acids) may happen. This yields poor solubilization of some components. b. During the entry of the sample in the focusing gel, there is a stacking effect due to the transition between a liquid phase and a gel phase with a higher friction coefficient. This stacking increases the concentration of proteins and may give rise to precipitation events. c. At, or very close to, the isoelectric point, the solubility of the proteins comes to a minimum. This can be explained by the fact that the net charge comes close to zero, with a concomitant reduction of the electrostatic repulsion between polypeptides. This can also result in protein precipitation or adsorption to the IEF matrix.

Apart from breaking molecular interactions and solubility in the 2-D gel, which are common to all samples, the solubilization problems encountered will greatly vary from a sample type to another because of wide differences in the amount and nature of interfering substances and/or spurious activities (e.g., proteases). The aim of this outline chapter is not to give detailed protocols for various sample types, and the reader should refer to the chapters of this book dedicated to the type of sample of interest. I would rather like to concentrate on the solubilization rationale and to describe nonstandard approaches to solubilization problems. More detailed review on solubilization of proteins for electrophoretic analyses can be found elsewhere (1,2).

2. Rationale of Solubilization-Breaking Molecular Interactions Apart from disulfide bridges, the main forces holding proteins together and allowing binding to other compounds are noncovalent interactions. Covalent bonds are encountered mainly between proteins and some coenzymes. The

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noncovalent interactions are mainly ionic bonds, hydrogen bonds, and “hydrophobic interactions.” The basis for “hydrophobic interactions” is, in fact, the presence of water. In this very peculiar (hydrogen-bonded, highly polar) solvent, the exposure of nonpolar groups to the solvent is thermodynamically not favored compared to the grouping of these apolar groups together. Indeed, although the van der Waals forces give an equivalent contribution in both configurations, the other forces (mainly hydrogen bonds) are maximized in the latter configuration and disturbed in the former (solvent destruction). Thus, the energy balance in clearly in favor of the collapse of the apolar groups together (3). This explains why hexane and water are not miscible, and also that the lateral chain of apolar amino acids (L, V, I, F, W, Y) pack together and form the hydrophobic cores of the proteins (4). These hydrophobic interactions are also responsible for some protein-protein interactions and for the binding of lipids and other small apolar molecules to proteins. The constraints for a good solubilization medium for 2-D electrophoresis are therefore to be able to break ionic bonds, hydrogen bonds, hydrophobic interactions, and disulfide bridges under conditions compatible with IEF (i.e., with very low amounts of salt or other charged compounds) (e.g., ionic detergents). 2.1. Disruption of Disulfide Bridges Breaking of disulfide bridges is usually achieved by adding to the solubilization medium an excess of a thiol compound. Mercaptoethanol was used in the first 2-D protocols (5), but its use does have drawbacks. Indeed, a portion of the mercaptoethanol will ionize at basic pH, enter the basic part of the IEF gel, and ruin the pH gradient in its alkaline part because of its buffering power (6). Although its pK is approximately 8, dithiothreitol is much less prone to this drawback, as it is used at much lower concentrations (usually 50 mM instead of the 700 mM present in 5% mercaptoethanol). However, DTT is still not the perfect reducing agent. Some proteins of very high cysteine content or with cysteines of very high reactivity are not fully reduced by DTT. In these cases, phosphines are very often an effective answer. First, the reaction is stoichiometric, which allows in turn to use very low concentration of the reducing agent (a few mM). Second, these reagents are not as sensitive as thiols to dissolved oxygen. The most powerful compound is tributylphosphine, which was the first phosphine used for disulfide reduction in biochemistry (7). However, the reagent is volatile, toxic, has a rather unpleasant odor, and needs an organic solvent to make it water-miscible. In the first uses of the reagent, propanol was used as a carrier solvent at rather high concentrations (50%) (7). It was however found that DMSO or DMF are suitable carrier solvents, which enable the reduction of proteins by 2 mM tributylphosphine (8). All these drawbacks have disappeared with the introduction of a water-soluble phospine, tris (carboxyethyl) phosphine

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(available from Pierce), for which 1M aqueous stock solutions can be easily prepared and stored frozen in aliquots. The successful use of tributylphosphine in two-dimensional electrophoresis has been reported (9). However, the benefits over DTT do not seem obvious in many cases (10), although th use of phosphines allows in turn to use thiol-blocking agents (e.g., dithiodiethanol), resulting in iproved resolution of the basic proteins (11,12). 2.2. Disruption of Noncovalent Interactions The perfect way to disrupt all types of noncovalent interactions would be the use of a charged compound that disrupts hydrophobic interactions by providing a hydrophobic environment. The hydrophobic residues of the proteins would be dispersed in that environment and not clustered together. This is just the description of SDS, and this explains why SDS has been often used in the first stages of solubilization (13–16). However, SDS is not compatible with IEF, and must be removed from the proteins during IEF. The other way of breaking most noncovalent interactions is the use of a chaotrope. It must be kept in mind that all the noncovalent forces keeping molecules together must be taken into account with a comparative view on the solvent. This means that the final energy of interaction depends on the interaction per se and on its effects on the solvent. If the solvent parameters are changed (dielectric constant, hydrogen bond formation, polarizability, etc.), all the resulting energies of interaction will change. Chaotropes, which alter all the solvent parameters, exert profound effects on all types of interactions. For example, by changing the hydrogen bond structure of the solvent, chaotropes disrupt hydrogen bonds but also decrease the energy penalty for exposure of apolar groups and therefore favor the dispersion of hydrophobic molecules and the unfolding of the hydrophobic cores of a protein (1,17). Unfolding the proteins will also greatly decrease ionic bonds between proteins, which are very often not very numerous and highly dependent of the correct positioning of the residues. As the gross structure of proteins is driven by hydrogen bonds and hydrophobic interactions, chaotropes decrease dramatically ionic interactions both by altering the dielectric constant of the solvent and by denaturing the proteins, so that the residues will no longer positioned correctly. Nonionic chaotropes, as those used in 2-D electrophoresis, however, are unable to disrupt ionic bonds when high charge densities are present (e.g., histones, nucleic acids) (18). In this case, it is often quite advantageous to modify the pH and to take advantage of the fact that the ionizable groups in proteins are weak acids and bases. For example, increasing the pH to 10 or 11 will induce most proteins to behave as anions, so that ionic interactions present at pH 7 or lower turn into electrostatic repulsion between the molecules, thereby promoting solubilization. The use of a high pH results therefore in dramatically improved

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solubilizations, with yields very close to what is obtained with SDS (19). The alkaline pH can be obtained either by addition of a few mM of potassium carbonate to the urea-detergent-ampholytes solution (19), or by the use of alkaline ampholytes (16), or by the use of a spermine-DTT buffer which allows better extraction of nuclear proteins (20). For 2-D electrophoresis, the chaotrope of choice is urea. Although urea is less efficient than substituted ureas in breaking hydrophobic interactions (17), it is more efficient in breaking hydrogen bonds, so that its overall solubilization power is greater. It has been found that thiourea in addition to urea results in superior solubilization of proteins (21) and also in improved protein focusing. This may be explained by the fact that thiourea is a superior to urea for protein denaturation, being overwhelmed only by guanidine (22). However, the solubility of thiourea in water is not sufficient to draw full benefits from its denaturing power, so that it must be used in admixture with urea. However, denaturation by chaotropes induces the exposure of the totality of the proteins hydrophobic residues to the solvent. This increases in turn the potential for hydrophobic interactions, so that chaotropes alone are often not sufficient to quench completely the hydrophobic interactions, especially when lipids are present in the sample. This explains why detergents, which can be viewed as specialized agents for hydrophobic interactions, are almost always included in the urea-based solubilization mixtures for 2-D electrophoresis. Detergents act on hydrophobic interactions by providing a stable dispersion of a hydrophobic medium in the aqueous medium, through the presence of micelles for example. Therefore, the hydrophobic molecules (e.g., lipids) are no longer collapsed in the aqueous solvent but will disaggregate in the micelles, provided the amount of detergent is sufficient to ensure maximal dispersion of the hydrophobic molecules. Detergents have polar heads that are able to contract other types of noncovalent bonds (hydrogen bonds, salt bonds for charged heads, etc.). The action of detergents is the sum of the dispersive effect of the micelles on hydrophobic part of the molecules and the effect of their polar heads on the other types of bonds. This explains why various detergents show very variable effects varying from a weak and often incomplete delipidation (e.g., Tweens) to a very aggressive action where the exposure of the hydrophobic core in the detergent-containing solvent is no longer energetically unfavored and leads to denaturation (e.g., SDS). Of course, detergents used for IEF must bear no net electrical charge, and only nonionic and zwitterionic detergents may be used. However, ionic detergents such as SDS may be used for the initial solubilization prior to isoelectric focusing to increase solubilization and facilitate the removal of interfering compounds. Low amounts of SDS can be tolerated in the subsequent IEF (13) provided that high concentrations of urea (23) and nonionic (13) or zwitterionic detergents (24) are present to ensure complete removal of the SDS from the

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proteins during IEF. Higher amounts of SDS must be removed prior to IEF, by precipitation (13) for example. It must therefore be kept in mind that SDS will only be useful for solubilization and for sample entry, but will not cure isoelectric precipitation problems. The use of nonionic or zwitterionic detergents in the presence of urea presents some problems due to the presence of urea itself. In concentrated urea solutions, urea is not freely dispersed in water but can form organized channels (see [20]). These channels can bind linear alkyl chains, but not branched or cyclic molecules, to form complexes of undefined stoichiometry called inclusion compounds. These complexes are much less soluble than the free solute, so that precipitation is often induced upon formation of the inclusion compounds, precipitation being stronger with increasing alkyl chain length and higher urea concentrations. Consequently, many nonionic or zwitterionic detergents with linear hydrophobic tails (24,25) and some ionic ones (28) cannot be used in the presence of high concentrations of urea. This limits the choice of detergents mainly to those with nonlinear alkyl tails (e.g., Tritons, Nonidet P40, CHAPS) or with short alkyl tails (e.g., octyl glucoside), which are unfortunately less efficient in quenching hydrophobic interactions. However, sulfobetaine detergents with long linear alkyl tails have received limited applications, as they require low concentrations of urea. However, good results have been obtained in certain cases for sparingly soluble proteins (27–29), although this type of protocol seems rather delicate owing to the need for a precise control of all parameters to prevent precipitation. Thus, protocols using detergents with a good urea compatibility are generally preferred, both because of their simpler use and because synergistic effects between the urea-thiourea chaotrope mixture and some detergents have been observed (32). Apart from the problem of inclusion compounds, the most important problem linked with the use of urea is carbamylation. Urea in water exists in equilibrium with ammonium cyanate, the level of which increases with increasing temperature and pH (33). Cyanate can react with amines to yield substituted urea. In the case of proteins, this reaction takes place with the α-amino group of the N-terminus and the e-amino groups of lysines. This reaction leads to artefactual charge heterogeneity, N-terminus blocking and adduct formation detectable in mass spectrometry. Carbamylation should therefore be completely avoided. This can be easily made with some simple precautions. The use of a pure grade of urea (p.a.) decreases the amount of cyanate present in the starting material. Avoidance of high temperatures (never heat urea-containing solutions above 37°C) considerably decreases cyanate formation. In the same trend, urea-containing solutions should be stored frozen (−20°C) to limit cyanate accumulation. Last, but not least, a cyanate scavenger (primary amine) should be added to urea-containing solutions. In the case of isoelectric focusing, carrier ampholytes are perfectly

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suited for this task. If these precautions are correctly taken, proteins seem to withstand long exposures to urea without carbamylation (34). 3. Methods 3.1 Solubility During IEF Additional solubility problems often arise during the IEF at sample entry and solubility at the isoelectric point. 3.1.1 Solubility During Sample Entry Sample entry is often quite critical. In most 2-D systems, sample entry in the IEF gel corresponds to a transition between a liquid phase (the sample) and a gel phase of higher friction coefficient. This induces a stacking of the proteins at the sample-gel boundary, which results in very high concentration of proteins at the application point. These concentrations may exceed the solubility threshold of some proteins, thereby inducing precipitation and sometimes clogging of the gel, with poor penetration of the bulk of proteins. Such a phenomenon is of course more prominent when high amounts of proteins are loaded onto the IEF gel. The sole simple but highly efficient remedy to this problem is to include the sample in the IEF gel. This process abolishes the liquid-gel transition and decreases the overall protein concentration, as the volume of the IEF gel is generally much higher than the one of the sample. This process is however rather difficult for tube gels in carrier ampholytebased IEF. The main difficulty arises from the fact that the thiol compounds used to reduce disulfide bonds during sample preparation are strong inhibitors of acrylamide polymerization, so that conventional samples cannot be used as such. Alkylation of cysteines and of the thiol reagent after reduction could be an answer, but many neutral alkylating agents (e.g., iodoacetamide, ethyl maleimide) also inhibit acrylamide polymerization. Owing to this situation, most workers describing inclusion of the sample within the IEF gel have worked with nonreduced samples (35,36). Although this presence of disulfide bridges is not optimal, inclusion of the sample within the gel has proven of great, but neglected, interest (35,36). It must however be pointed out that it is now possible to carry out acrylamide polymerization in an environment where disulfide bridges are reduced. The key is to use 2 mM tributylphosphine as the reducing agent in the sample and using tetramethylurea as a carrier solvent. This ensures total reduction of disulfides and is totally compatible with acrylamide polymerization with the standard Temed/persulfate initiator (T. Rabilloud, unpublished results). This modification should help the experimentators trying sample inclusion within the IEF gel when high amounts of proteins are to be separated by 2D.

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The process of sample inclusion within the IEF gel is however much simpler for IPG gels. In this case, rehydration of the dried IPG gel in a solution containing the protein sample is quite convenient and efficient, provided that the gel has a sufficiently open structure to be able to absorb proteins efficiently (20). Coupled with the intrinsic high capacity of IPG gels, this procedure enables to easily separate milligram amounts of protein (20,37). 3.1.2. Solubility at the Isoelectric Point This is usually the second critical point for IEF. The isoelectric point is the pH of minimal solubility, mainly because the protein molecules have no net electrical charge. This abolishes the electrostatic repulsion between protein molecules, which maximizes in turn protein aggregation and precipitation. The horizontal comet shapes frequently encountered for major proteins and for sparingly soluble proteins often arise from such a near-isoelectric precipitation. Such isoelectric precipitates are usually easily dissolved by the SDS solution used for the transfer of the IEF gel onto the SDS gel, so that the problem is limited to a loss of resolution, which however precludes the separation of high amounts of proteins. The problem is however more severe for hydrophobic proteins when an IPG is used. In this case, a strong adsorption of the isoelectric protein to the IPG matrix seems to occur, which is not reversed by incubation of the IPG gel in the SDS solution. The result is severe quantitative losses, which seem to increase with the hydrophobicity of the protein and the amount loaded (38). The sole solution to this serious problem is to increase the chaotropicity of the medium used for IEF, by using both urea and thiourea as chaotropes (21). The benefits of using thiourea-urea mixtures to increase protein solubility can be transposed to conventional, carrier ampholyte-based focusing in tube gels with minor adaptations. Thiourea strongly inhibits acrylamide polymerization with the standard temed/persulfate system. However, photopolymerization with methylene blue, sodium toluene sulfinate and diphenyl iodonium chloride (39) enables acrylamide polymerization in the presence of 2M thiourea without any deleterious effect in the subsequent 2D (40) so that higher amounts of proteins can be loaded without loss of resolution (40). 3.1.3 The Epitome in Solubilization Problems: Membrane Proteins Biological membranes represent a prototype of difficult samples for 2-D electrophoresis. They contain high amounts of lipids, which are troublesome compounds for 2-D electrophoresis. In addition, many membrane proteins are highly hydrophobic, and therefore very difficult to keep in aqueous solution, even with the help of chaotropes and detergents. Thus, membrane proteins

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have often been reluctant to 2-D electrophoretic analysis (41). In fact, it has been shown that typical membrane proteins with multiple transmembrane helices are resistant to solubilization by chaotrope-detergent mixtures, even using thiourea (42). For such proteins, the development of new detergents proved necessary (42,43). Even if some successes have been obtained (42–45), it is still obvious that many membrane proteins are not properly solubilized and focused with the chemicals described up to now (46). Therefore, further development is still needed to improve the representation of membrane proteins in 2-D maps. However, the problem of isoelectric precipiation may prevent definitively proper solubilization of membrane proteins in IEF-based 2-D electrophoretic systems 4. Concluding Remarks Although this outline chapter has mainly dealt with the general aspects of solubilization, the main concluding remark is that there is no universal solubilization protocol. Standard urea-reducer-detergent mixtures usually achieve disruption of disulfide bonds and noncovalent interactions. Consequently, the key issues for a correct solubilization is the removal of interfering compounds, blocking of protease action, and disruption of infrequent interactions (e.g., severe ionic bonds). These problems will strongly depend on the type of sample used, the proteins of interest and the amount to be separated, so that the optimal solubilization protocol can vary greatly from a sample to another. However, the most frequent bottleneck for the efficient 2-D separation of as many and as much proteins as possible does not lie only in the initial solubilization but also in keeping the solubility along the IEF step. In both fields, the key feature is the disruption of hydrophobic interactions, which are responsible for most, if not all, of the precipitation phenomena encountered during IEF. This means improving solubility during denaturing IEF will focus on the quest of ever more powerful chaotropes and detergents. In this respect, the use of thiourea has proven to be one of the keys to increase the solubility of proteins in 2-D electrophoresis. It would be nice to have other, even more powerful chaotropes. However, testing of the chatropes previously described as powerful (22) has not led yet to improvement in protein solubilization (T. Rabilloud, unpublished results). One of the other keys for improving protein solubilization is the use of as powerful detergent or detergent mixtures as possible. Among a complex sample, some proteins may be well denatured and solubilized by a given detergent or chaotrope, while other proteins will require another detergent or chaotrope (e.g., in [42]). Consequently, the future of solubilization may still be to find mixtures of detergents and chaotropes able to cope with the diversity of proteins encountered in the complex samples separated by 2-D electrophoresis.

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References 1. Rabilloud, T. (1996) Solubilization of proteins for electrophoretic analyses. Electrophoresis 17, 813–829. 2. Rabilloud, T. and Chevallet, M. (1999) Solubilization of proteins in 2-D electrophoresis, in Proteome Research: Two-Dimensional Gel Electrophoresis and Identification Methods (Rabilloud, T., ed.), Springer-Verlag, Heidelberg, pp 9–30. 3. Tanford, C. The Hydrophobic Effect, 2nd ed., Wiley, New York, 1980. 4. Dill, K.A. (1985) Theory for the folding and stability of globular proteins. Biochemistry 24, 1501–1509. 5. O’Farrell P.H. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–4021. 6. Righetti, P.G., Tudor, G., and Gianazza, E. (1982) Effect of 2 mercaptoethanol on pH gradients in isoelectric focusing. J. Biochem. Biophys. Meth. 6, 219–227. 7. Ruegg, U.T. and Rüdinger, J. (1977) Reductive cleavage of cystine disulfides with tributylphosphine. Meth. Enzymol. 47, 111–116. 8. Kirley, T.L. (1989) Reduction and fluorescent labeling of cyst(e)ine containing proteins for subsequent structural analysis. Anal. Biochem. 180, 231–236. 9. Herbert, B., Molloy, M.P., Gooley, A.A., Walsh, B.J., Bryson, W.G., and Williams, K.L. (1998) Improved protein solubility in two-dimensional electrophoresis using tri butyl phosphine as a reducing agent. Electrophoresis 19, 845–851. 10. Hoving, S., Voshol, H., and Van Oostrum, J. (2000) Towards high performance two-dimensional gel electrophoresis using ultrazoom gels. Electrophoresis 21, 2617–2621. 11. Olsson, I., Larsson, K., Palmgren, R., and Bjellqvist, B. (2002) Organic disulfides as a means to generate streak-free two-dimensional maps with narrow range basic immobilized pH gradient strips as first dimension. Proteomics 2, 1630–1632. 12. Luche, S., Diemer, H., Tastet, C., Chevallet, M., Van Dorsselaer, A., Leize-Wagner, E., and Rabilloud, T. (2004) About thiol derivatization and resolution of basic proteins in two-dimensional electrophoresis. Proteomics 4, 551–561. 13. Wilson, D., Hall, M.E., Stone, G.C., and Rubin, R.W. (1977) Some improvements in two-dimensional gel electrophoresis of proteins. Anal. Biochem. 83, 33–44. 14. Hari, V. (1981) A method for the two-dimensional electrophoresis of leaf proteins. Anal. Biochem. 113, 332–335. 15.Ames, G.F.L. and Nikaido, K.(1976) Two-dimensional electrophoresis of membrane proteins. Biochemistry 15, 616–623. 16. Hochstrasser, D.F., Harrington, M.G., Hochstrasser, A.C., Miller, M.J., and Merril, C.R. (1988) Methods for increasing the resolution of two dimensional protein electrophoresis. Anal. Biochem. 173, 424–435. 17. Herskovits, T.T., Jaillet, H., and Gadegbeku, B. (1970) On the structural stability and solvent denaturation of proteins. II. Denaturation by the ureas. J. Biol. Chem. 245, 4544–4550. 18. Sanders, M.M., Groppi, V.E., and Browning, E.T. (1980) Resolution of basic cellular proteins including histone variants by two-dimensional gel electrophoresis: evaluation of lysine to arginine ratios and phosphorylation. Anal. Biochem. 103, 157–165.

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19. Horst, M.N., Basha, M.M., Baumbach, G.A., Mansfield, E.H., and Roberts, R.M.(1980) Alkaline urea solubilization, two-dimensional electrophoresis and lectin staining of mammalian cell plasma membrane and plant seed proteins. Anal. Biochem. 102, 399–408. 20. Rabilloud, T., Valette, C., and Lawrence, J.J.(1994) Sample application by in-gel rehydration improves the resolution of two-dimensional electrophoresis with immobilized pH gradients in the first dimension. Electrophoresis 15, 1552–1558. 21. Rabilloud, T., Adessi, C., Giraudel, A., and Lunardi, J. (1997) Improvement of the solubilization of proteins in two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 18, 307–316. 22. Gordon, J.A. and Jencks, W.P. (1963) The relationship of structure to the effectiveness of denaturing agents for proteins. Biochemistry 2, 47–57. 23. Weber, K. and Kuter, D.J. (1971) Reversible denaturation of enzymes by sodium dodecyl sulfate. J. Biol. Chem. 246, 4504–4509. 24. Remy, R., and Ambard-Bretteville, F.(1987) Two-dimensional electrophoresis in the analysis and preparation of cell organelle polypeptides. Meth. Enzymol. 148, 623–632. 25. March, J. (1977) Advanced Organic Chemistry, 2nd ed., McGraw-Hill London, pp. 83–84. 26. Dunn, M.J. and Burghes, A.H.M.(1983) High resolution two-dimensional polyacrylamide electrophoresi. I. Methodological procedures. Electrophoresis 4, 97–116. 27. Rabilloud, T., Gianazza, E., Catto, N., and Righetti, P.G. (1990) Amidosulfobetaines, a family of detergents with improved solubilization properties: application for isoelectric focusing under denaturing conditions. Anal. Biochem. 185, 94–102. 28. Willard, K.E., Giometti,C., Anderson, N.L., O’Connor, T.E., and Anderson, N.G. (1979) Analytical techniques for cell fractions. XXVI. A two-dimensional electrophoretic analysis of basic proteins using phosphatidyl choline/urea solubilization. Anal. Biochem., 100, 289–298. 29. Clare Mills, E.N. and Freedman, R.B. (1983) Two-dimensional electrophoresis of membrane proteins. Factors affecting resolution of rat liver microsomal proteins. Biochim. Biophys. Acta 734, 160–167. 30. Satta,D., Schapira,G., Chafey, P., Righetti, P.G., and Wahrmann, J.P. (1984) Solubilization of plasma membranes in anionic, non ionic and zwitterionic surfactants for iso-dalt analysis: a critical evaluation. J. Chromatogr. 299, 57–72. 31. Gyenes,T. and Gyenes, E. (1987) Effect of stacking on the resolving power of ultrathin layer two-dimensional gel electrophoresis. Anal. Biochem. 165, 155–160. 32. Luche, S., Santoni, V., and Rabilloud, T. (2003) Evaluation of nonionic and zwitterionic detergents as membrane protein solubilizers in two-dimensional electrophoresis.. Proteomics 3, 249–253. 33. Hagel, P., Gerding, J.J.T., Fieggen, W., and Bloemendal, H. (1971) Cyanate formation in solutions of urea. I. Calculation of cyanate concentrations at different temperature and pH. Biochim. Biophys. Acta 243, 366–373. 34. Bjellqvist, B., Sanchez, J.C., Pasquali, C., Ravier, F., Paquet, N., Frutiger, S., Hughes, G.J., and Hochstrasser, D.F. (1993) Micropreparative two-dimensional

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36.

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38.

39. 40. 41. 42.

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44.

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46.

Rabilloud electrophoresis allowing the separation of samples containing miiligram amounts of proteins. Electrophoresis 14, 1375–1378. Chambers, J.A.A., Degli Innocenti, F., Hinkelammert, K., and Russo, V.E.A (1985) Factors affecting the range of pH gradients in the isoelectric focusing dimension of two-dimensional gel electrophoresis: the effect of reservoir electrolytes and loading procedures. Electrophoresis 6, 339–348. Semple-Rowland, S.L., Adamus, G, Cohen, R.J., and Ulshafer, R.J. (1991) A reliable two-dimensional gel electrophoresis procedure for separating neural proteins. Electrophoresis 12, 307–312. Sanchez, J.C., Rouge, V., Pisteur, M., Ravier, F., Tonella, L., Moosmayer, M., Wilkins, M.R., and Hochstrasser, D.F. (1997) Improved and simplified in-gel sample application using reswelling of dry immobilized pH gradients. Electrophoresis 18, 324–327. Adessi, C., Miege, C., Albrieux, C., and Rabilloud, T. (1997) Two-dimensional electrophoresis of membrane proteins: a current challenge for immobilized pH gradients. Electrophoresis 18, 127–135. Lyubimova, T., Caglio, S., Gelfi, C., Righetti, P.G., Rabilloud, T. (1993) Photopolymerization of polyacrylamide gels with methylene blue. Electrophoresis 14, 40–50. Rabilloud, T. (1998) Use of thiourea to increase the solubility of membrane proteins in two-dimensional electrophoresis. Electrophoresis 19, 758–760. Rubin, R.W. and Milikowski, C (1978) Over two hundred polypeptides resolved from the human erythrocyte membrane.Biochim. Biophys. Acta 509, 100–110 Rabilloud, T., Blisnick, T., Heller, M., Luche, S., Aebersold, R., Lunardi, J., and Braun-Breton, C. (1999) Analysis of membrane proteins by two-dimensional electrophoresis: comparison of the proteins extracted from normal or Plasmodium falciparum - infected erythrocyte ghosts. Electrophoresis 20, 3603–3610. Chevallet, M., Santoni, V., Poinas, A., Rouquié, D., Fuchs, A., Kieffer, S., Rossignol, M., Lunardi, J., Garin, J., and Rabilloud, T. (1998) New zwitterionic detergents improve the analysis of membrane proteins by two-dimensional electrophoresis. Electrophoresis 19, 1901–1909. Santoni, V., Rabilloud, T., Doumas, P., Rouquié, D., Mansion, M., Kieffer, S., Garin, J., and Rossignol, M. (1999) Towards the recovery of hydrophobic proteins on two-dimensional gels. Electrophoresis 20, 705–711. Friso, G. and Wikstrom, L. (1999) Analysis of proteins from membrane-enriched cerebellar preparations by two-dimensional gel electrophoesis and mass spectrometry Electrophoresis 20, 917–927. Santoni, V., Molloy, M.P., and Rabilloud, T. (2000) Membrane proteins and proteomics: un amour imposssible? Electrophoresis 21, 1054–1070.

12 Mouse and Human Tissue Sample Preparation for 2-D Electrophoresis Claus Zabel and Joachim Klose 1. Introduction The protocol for extracting proteins from mouse and human tissues (organs) described in this chapter adheres to a strategy that is based on a rationale to include all protein species of a particular tissue in a set of samples which are suitable for two dimensional electrophoresis (2-DE), particularly for large gel 2-DE described in Chapter 32 (1). The extraction procedure is one of the most important steps to maintain reproducibility in 2-D gel based proteomics (2). The ultimate goal is resolution and visualization of all these protein species in 2-DE gels. This aim determines some features of our tissue extraction procedure for retrieving proteins. Since the last publication of our comprehensive protein extraction protocol (3) we found that an extraction protocol for total cell or tissue proteins may be sufficient for many more users than a protocol for fractionated extraction (4). Still, fractionated extraction has also a number of applications, especially when only certain classes of proteins such as cytoplasmic, membrane or nuclear proteins are of interest. 1.1. Total Extraction of all Cellular Proteins Extraction of total proteins at once insures that no distribution artefacts occur as may be seen when proteins are fractionated into different subcellular fractions. Therefore the total amount of a given protein is determined within each protein sample investigated. In addition, the total extraction procedure is much simpler, takes less time and is highly compatible with new protein visualization techniques such as differential in gel electrophoresis (DIGE). This extraction procedure can be applied to almost any tissue, cell culture or subcellular components. No ultracentrifugation steps are involved which avoids protein precipitation and

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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elimination during sample preparation. The total extraction procedure is based on adding protease inhibitors, detergent (CHAPS), urea/thiourea in very close succession to solubilze proteins in a tissue/cell as much as possible and keep them from degradation. Grinding the deep frozen tissue and sonication are the second instrumental part in the solubilization process. DNA-bound proteins are recovered by DNAse treatment. In this way all proteins are solubilized in one fraction without precipitation and fractionation steps. 1.2. Fractionated Extraction of Cellular Proteins On the other hand, using the fractionation procedure the many different protein species of a tissue can be distributed over several 2-DE gels and this certainly increases resolution. However, a postulate is that fractionation of tissue proteins results in fraction-specific proteins. Usually, cell fractionation is performed with the aim of isolating special cell organelles (nuclei, mitochondria) or cell structures (membranes). Proteins are then extracted from these subcellular fractions. This procedure, however, includes washing steps to purify cell fractions and eliminate cell components which are not of interest and cell residues which are rejected. Using fractionation this way, an uncontrolled loss of proteins is unavoidable. The tissue fractionation procedure described in this chapter renounces the isolation of defined cell components using washing and precipitation steps. This allows us to avoid any selective loss of proteins which is of utmost importance for all protein extraction protocols. Mouse (human) tissues (liver, brain, heart) are fractionated into three fractions: (1) The “supernatant I+II” (SI+II) containing the proteins soluble in buffer (cytoplasmic and nucleoplasmic proteins), (2) the “pellet extract” (PE) containing the proteins soluble in the presence of urea and CHAPS (proteins from membranes and other structures of the cells and cell organelles), and (3) the “pellet suspension” (PS) containing proteins released by DNA digestion (histones and other chromosomal proteins). The SI+II fraction is obtained by homogenization, sonication and centrifugation of tissue and reextraction of pellet (I) and the combination of the two supernatants gained in this way. The solution thus obtained is the first protein sample. The pellet (II) that remained is extracted with urea and CHAPS and the homogenate is centrifuged. The supernatant is the PE fraction and constitutes the second protein sample. The final pellet (III) is suspended into buffer containing benzonase, a DNA digesting enzyme. The pellet suspension (PS fraction) is the third protein sample. It is applied to 2-DE without further centrifugation. Care is taken during the whole procedure to avoid any loss of material. In spite of this caution, some material may be lost, for example by transfer of the pulverized, frozen tissue from the mortar to a tube or by removing the glass

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beads from the sonicated homogenate. However, this does not lead to a preferential loss of certain protein species or protein classes. 1.3. Important Considerations When Using Protein Extraction Procedures Generally, it is evident that the best conditions for keeping proteins stable and soluble are present in living cells (1,2). Therefore, an important principle of our tissue extraction procedure is to extract proteins, using conditions as close to the natural environment in the cell as possible. That means keeping ionic strength of the tissue homogenate at 150-200 mM, the pH in a range of 7.0-7.5, protein concentration high, and protecting the proteins against water by adding glycerol to the buffer. Generally, the best conditions for the first tissue extraction step would be if an addition of diluent that disturbs the natural protein concentrations and milieu of the cell were avoided. We prepared a pure cell sap from a tissue by homogenizing it without additives (except for protease inhibitor solutions added in small volumes) followed by high speed centrifugation and extracted the pellet that resulted successively in increasing amounts (0.5, 1 or 2 parts) of buffer. The series of protein samples obtained were separated by 2-DE and the patterns compared. The results showed that by increasing the dilution of proteins the number of spots and their intensities decreased in the lower part of the 2-DE patterns and increased in the upper part. The same phenomenon, but less pronounced, was observed even when the cell sap was diluted successively. Apparently, low molecular weight proteins are best dissolved in the pure cell sap and, presumably, tend to precipitate in more diluted extracts. In contrast, high molecular weight proteins dissolve better in more diluted samples. This effect was most pronounced in protein patterns from liver and not obvious in patterns from heart muscle. This is probably due to the high protein concentration in the liver cell sap that is not reached in extracts of other organs. The dependence of protein solubility on the molecular weight of proteins is obscured in 2-DE patterns by another effect that causes a similar phenomenon. An increasing protein concentration of the first tissue extract leads to a higher activity of proteases released by breaking cellular structure by homogenization. Protein patterns from pure liver cell sap, extracted without protease inhibitors (but even with inhibitors) showed an enormous number of spots in the lower part of the gel and a rather depleted pattern in the upper part. In the pH range around pH 6 protein spots disappeared almost completely, in the upper as well as in the lower part, suggesting that these proteins are most sensitive to degradation by proteases. By extracting tissue or the first pellet with increasing amounts of buffer, the 2-DE pattern (spot number and intensity) shifted from the lower part to the upper part of the gel. Again, this observation was made particularly in liver.

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The consequence of these observations for our total as well as our fractionated protein extraction procedure was to maintain the total extract or to obtain the supernatant I and II at concentrations that keep all the soluble proteins in solution but do not reach a level where proteases cannot be inhibited effectively anymore. As a consequence, we introduced buffer factors which adjust the concentration of the different extracts of each organ. The optimum protein concentrations in the extracts and therefore the buffer factors were determined experimentally. An optimum was reached when a maximum number of spots was present in the upper as well as in the lower part of a 2-DE protein pattern. In addition, the region close to pH 6 should not be depleted of spots, a process starting from the top.The methods described in the following sections were developed with mouse tissues but were found to be applicable to corresponding human tissues, cell culture cells and subcellular fractions as well. 2. Materials 2.1. Equipment 1. A small apparatus is preferred for performing sonication in a water-bath (Transsonic 310, FAUST, Singen, Germany). 2. Glass beads added to tissue samples for sonication: Size (diameter) of glass beads should be 2.0- 2.5 mm. A factor 0.034 was calculated for this bead size (see Note 1). 3. Mortar and pestle: Build and size of this equipment are shown in Fig. 1. Mortar and pestle are manufactured of agate or glass (WITA GmbH, Teltow, Germany). Glass

Fig. 1. Special equipment for pulverization of frozen tissue. (A) Glass mortar and plastic pestle. (B) Spatula used to transfer frozen tissue from a mortar to the test tube. A regular spatula was moulded into a small shovel.

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89

was found to be more stable in liquid nitrogen. Alternatively an electronic mortar grinder can be used (Mortar Grinder RM 200; RETSCH, Haan, Germany). 4. A small spatula is formed into a shovel by wrought-iron work (Fig. 1) and used to transfer tissue powder from a mortar to tubes.

2.2. Reagents 1. Buffer A: 20% glycerol, 100 mM KCl, 50 mM Tris-HCl, pH 7.1, filtered and aliquoted into 1,000 µL portions, and stored at −70°C. 2. Phosphate buffer, pH 7.1: Add 67 mL 200 mM Na2HPO4 to 33 mL 200 mM NaH2PO4. 3. Buffer B: 20 % glycerol, 200 mM KCl, 100 mM phosphate buffer, pH 7.1, filtered, aliquoted precisely into 900 µL portions and stored at −70°C. When used, 100 µL of an aqueous CHAPS solution is added. The CHAPS concentration in this aqueous solution is calculated (see Table 1) so that the pellet II/buffer homogenate (see Table 1) contains 4.5% CHAPS. This concentration was found to be best when 2-DE patterns are compared. The concentration was determined empirically using protein samples containing different amounts of CHAPS. 4. Buffer C: 1 mM MgSO4, 50 mM Tris-HCl, pH 8.0 (I mg of pellet III homogenised in 1 mL of this buffer gives a concentration of 1 mM MgSO4. The final solution is filtered, aliquoted into 1 ml portions, and stored at −70°C. 5. Buffer P: 7.7 % glycerol (final concentration in sample if buffer factor is 1.6), 50 mM KCl, 50 mM Tris-HCl, pH 7.5. The final solution is filtered and aliquoted into 900 µL units and stored at −70°C. 6. Buffer P-MgCl2: 5 mM MgCl2 in buffer P. Final solution is aliquoted into 20 µL units and stored at −70°C. 7. Protease inhibitor 1A: One tablet of Complete™ (ROCHE, Mannheim, Germany) is dissolved in 2 ml buffer A (according to the manufacturer’s instructions) and the resulting solution aliquoted into 50, 80 and 100 µL units. Inhibitor 1B was prepared in the same way but using buffer B (900 µL buffer + 100 µL H20) and aliquoted into 30 and 50 µL units. Inhibitor 1C was prepared in the same way as Inhibitor 1A but using buffer P. Inhibitor 2 (pepstatin A): Prepared as a stock solution (9.603mg/100mL ethanol) and aliquoted into 100 µL portions. All inhibitor solutions are stored at −70°C. 8. DTT-solution: 2.16 g DTT is dissolved in 10 mL bidistilled water. The solution is aliquoted into 100 µL portions and stored at −70°C. 9. Sample diluent: Buffer P-MgCl2. The solution is aliquoted into 250 µL units and stored at −70°C.

3. Methods 3.1. Extraction of Total Proteins 3.1.1. Dissection of Mouse Tissue To obtain good results with protein extraction harvest organs rapidly and remove all contaminating material such as hair and blood.

Stirring 4°C; 15 min Benzonase treatment P-MgCl2 (Σ3 × 0.021) Benzonase (Σ3 × 0.025) 4 °C; 15 min Σ4 (weight of sample determined after benzonase treatment) Control factor Liver tissue/Σ4

Sonication Number of glass beads (Σ2 × 0.034) Sonication, 6 × 10 sec Σ3 (weight of sample determined after sonication)

Homogenization – Liver tissue (25–50 mg) – Buffer P-CHAPS 56 mg CHAPS 47 mg bidestilled water + 900 µL buffer P - P-CHAPS (2.5 × mg liver tissue) Σ1 (mg liver tissue + mg P-CHAPS) Inhibitor 1C (Σ1 × 0.08) Inhibitor 2 ( (Σ1 × 0.01) Σ2 (mg Σ1 + mg Inhibitor 1C + mg Inhibitor 2)

Liver

Table 1 Total Tissue Protein Extraction Protocol

4°C; 30 min P-MgCl2 (Σ3 × 0.021) Benzonase (Σ3 × 0.025) 4 °C; 15 min Σ4 (weight of sample determined after benzonase treatment) Heart tissue/Σ4

4°C; 15 min P-MgCl2 (Σ3 × 0.021) Benzonase (Σ3 × 0.025) 4 °C; 15 min Σ4 (weight of sample determined after benzonase treatment) Brain tissue/Σ4

Number of glass beads (Σ2 × 0.034) Sonication, 12 × 10 sec Σ3 (weight of sample determined after sonication)

- P-CHAPS (1.6 mg heart tissue) Σ1 (mg heart tissue + mg P-CHAPS) Inhibitor 1C (Σ1 × 0.08) Inhibitor 2 ( (Σ1 × 0.01) Σ2 (mg Σ1 + mg Inhibitor 1C + mg Inhibitor 2)

- P-CHAPS (1.6 × mg brain tissue) Σ1 (mg brain tissue + mg P-CHAPS) Inhibitor 1C (Σ1 × 0.08) Inhibitor 2 ( (Σ1 × 0.01) Σ2 (mg Σ1 + mg Inhibitor 1C + mg Inhibitor 2) Number of glass beads (Σ2 × 0.034) Sonication, 6 × 10 sec Σ3 (weight of sample determined after sonication)

– Heart tissue (25–150 mg) – Buffer P-CHAPS 65 mg CHAPS 38 mg bidestilled water

Heart

– Brain tissue (25–150 mg) – Buffer P-CHAPS 65 mg CHAPS 38 mg bidestilled water

Brain

90 Zabel and Klose

IEF (1-D) 5 µL/gel

DTT (Σ6 × 0.1) RT, 5 min Ampholin (Σ6 × 0.1) Σ7 (weight of sample after DTT + Ampholin addition)

Urea/thiourea Urea 6M (Σ5 × 0.78) Thiourea 2M (Σ5 × 0.3) Stirring RT, 30 min Aliquot DIGE (min. 20 µL) Σ6 (weight of sample determined after aliquot DIGE)

Protein concentration Aliquot 5 µL Σ5 (weight of sample determined after aliqout)

5 µL/gel

Urea 6M (Σ5 × 0.78) Thiourea 2M (Σ5 × 0.3) Stirring RT, 30 min Aliquot DIGE (min. 20 µL) Σ6 (weight of sample determined after aliquot DIGE) DTT (Σ6 × 0.1) RT, 5 min Ampholin (Σ6 × 0.1) Σ7 (weight of sample after DTT + Ampholin addition)

Aliquot 5 µL Σ5 (weight of sample determined after aliqout)

5 µL/gel

Urea 6M (Σ5 × 0.78) Thiourea 2M (Σ5 × 0.3) Stirring RT, 30 min Aliquot DIGE (min. 20 µL) Σ6 (weight of sample determined after aliquot DIGE) DTT (Σ6 × 0.1) RT, 5 min Ampholin (Σ6 × 0.1) Σ7 (weight of sample after DTT + Ampholin addition)

Aliquot 5 µL Σ5 (weight of sample determined after aliqout)

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3.1.1.1. DISSECTION OF MOUSE LIVER 1. Kill the mouse by decapitation. Thereby, the body is allowed to bleed. The following steps are performed in a cold room: 2. Cut open the abdomen and through the vena femoralis on both sides and perfuse the liver with 5 ml saline (0.9% NaCl solution). 3. Dissect the complete liver from the body, remove the gall bladder without injuring it and separate the liver into its different lobes. The central part of each lobe, i.e. the region where blood vessels enter the liver lobe, is cut off as are the remainders of other tissues (diaphragm, fascia). 4. Cut the liver lobes into 2-4 pieces, rinse in ice-cold saline and briefly immerse it there. Immediately after this step the next organ (e.g. brain) is prepared from the same animal, if desirable, and processed to the same stage of preparation as liver. 5. Cut the liver pieces into smaller pieces (about 5 × 5 mm) and transfer each piece to filter paper, immerse into liquid nitrogen, and put into a screw-cap tube where all pieces are collected. During preparation time, tubes are kept in a liquid nitrogen containing box and afterwards stored at −70°C. 3.1.1.2. DISSECTION OF MOUSE BRAIN 1. Kill mouse by decapitation. The following steps are performed in a cold room. If several organs have to be taken from the same animal, start with the brain. 2. Cut off the skin of the head and open the cranium starting from the spinal canal proceeding in a frontal direction. Break the cranial bones apart to expose the brain. Remove the brain including both bulbi olfactorii and a small part of the spinal cord. Transfer the brain into a petri dish containing ice-cold saline. Remove any blood vessels and blood from the outside of the brain. 3. As desired, cut the brain to retrieve the desired brain regions or in half along the corpus callosum, place the pieces on filter paper and then individually immerse into liquid nitrogen and collect in a screw-capped tube. Keep the tubes in liquid nitrogen during tissue harvest and finally store at −70°C. 3.1.1.3. DISSECTION OF MOUSE HEART 1. Kill mouse by decapitation. The following steps are performed in a cold room. 2. Open the thorax and remove the heart. Place the heart into a petri dish containing ice-cold saline. Cut off both atria and open the ventriculi to remove any blood and blood clots. 3. Dry the heart on filter paper, freeze in liquid nitrogen and store in a screw-capped tube at −70°C.

3.1.2. Total Extraction of Liver Proteins 1. Fill frozen liver pieces into a small plastic tube of known weight and weigh quickly without thawing. The amount of liver tissues used for extraction should be between 50 and 150 mg.

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2. Place a mortar, pestle and a small metal spoon into a styrofoam box that contains liquid nitrogen to a level not exceeding the height of the mortar. The mortar should be pre-cooled for at least 3–4 min. Pre-cool pestle and spatula. 3. Put frozen liver pieces into the mortar, and add buffer P, inhibitor 1C and 2. The required volumes of each of the solutions are calculated as indicated in Table 1. The precise amount of each solution is pipetted as a droplet onto a small spoonlike spatula that was kept in the N2-box before use. The solution is immediately frozen into an ice bead that can easily be transferred into the mortar. 4. Slowly grind all frozen components in the mortar to powder. Care should be taken that small pieces of material do not jump out of the mortar when starting to break up the solid frozen material with a pestle. 5. Transfer the powder into a pre-frozen 2 ml Eppendorf tube using a special spatula (Fig. 1). Forceps are used to freeze the tube briefly in N2 and then to hold the tube close to the mortar in the N2-box. Care is taken that no powder is left in the mortar or on the pestle. For collection of this powder always use the same type of plastic tube. This contributes to reproducibility of the subsequent sonication step. Compress the powder collected in the tube by gently knocking the tube against the mortar. The powder can be stored at −70°C or immediately subjected to sonication. 6. For sonication a predetermined number of glass beads (see Table 1 and Note 1) is added to the sample and the powder is then thawed and kept immersed in ice. Sonication is performed in an ice-cold water-bath. The fill-height of the water is critical for the sonication effect and should always be at the level indicated in the instruction manual of the manufacturer. Furthermore, when dipping the sample tube into the water, it is important to do this at the “sonication center” visible by a concentric water surface motion which appear when holding the tube into the water. We prefer a small sonication apparatus that forms only one sonication center (see Subheading 2.1). Sonication is performed for 10 sec. Immediately afterwards the sample is stirred with a thin wire for 50 sec with the tube still remaining in the ice water. The tube is then kept immersed in ice for 1 min. Now the next sonication circle is started, until a total of 6 two-minute steps (see Note 1). After sonication, the tubes are turned upside down and punctured with a steel needle to generate a 2 mm diameter opening. The holes should not be larger than that to keep the glass beads from crossing over. A second tube is attached to the bottom of the first. Both tubes are transferred into a centrifuge using other samples or dummy tubes for balancing. Tubes are quickly centrifuged at 2,000 g for 1 min. The sample is thereby transferred in to the lower tube whereas the glass beads remain in the upper. 7. Add a small bar magnet and slowly stir homogenate at 4°C for 15 min. 8. Add benzonase (Merck, Darmstadt, Germany) and P-MgCl2. Slowly stir the homogenate for 15 min at 4°C (DNA digestion). 9. Determine control factor (Table 1 and Note 2). 10. Remove aliquot for determination of protein concentration. (Note: When sample volume is very low, remove aliquot only after addition of urea/thiourea.) 11. Add urea and thiourea to the homogenate. The amounts indicated in Table 1 are transferred into a capped 1.5 mL tube. The tube is sealed and turned up side down. Now the bottom of the tube is cut away by scissors and the tube is put on top of

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the 2 mL tube containing the homogenate. A quick-spin transfers the urea/thiourea completely into the homogenate (1 min, 1,000g). The homogenate is stirred for about 30 min until the urea/thiourea is dissolved. Now the aliquot for DIGE can be sampled (Table 1 and Note 3). 12. Add DTT and stir homogenate for at least 5 min then add Ampholin 2-4. Now, carefully remove the small bar magnet. Now aliquot samples to omit freeze and rethaw cycles. Usually samples need to be diluted (most commonly 1:2) by sample dilution solution (Buffer P-MgCl2) before application to isoelectric focusing (IEF). 13. Samples are quick-frozen and stored at −70°C.

3.1.3. Extraction of Total Proteins from Other Tissues In general, the total extraction protocol differs only in buffer factors and the repeats of sonication (Table 1). Only very special tissue such as lens would show deviations. In our experience lens does not require DNAse treatment. 3.2. Fractionated Extraction of Proteins 3.2.1. Fractionated Extraction of Liver Proteins 3.2.1.1. EXTRACTION OF LIVER PROTEINS SOLUBLE IN BUFFER (SUPERNATANT I + II) 1. Fill frozen liver pieces into a small plastic tube of known weight and weigh quickly without thawing. The weight of the liver tissues should be between 25 and 260 mg (for lower amounts see Note 4). 2. Place a mortar, pestle and a small metal spoon into a styrofoam box that contains liquid nitrogen to a level not exceeding the height of the mortar. 3. Put frozen liver pieces into the mortar, and add buffer A, inhibitor 1A and 2. The required volumes of each of the solutions are calculated as indicated in Table 2. The precise amount of each solution is pipetted as a droplet onto the spoon that was kept in the N2-box before use. The solution immediately is immediately frozen into an ice bead that can easily be transferred into the mortar. 4. Grind all frozen components in the mortar to powder. Care should be taken that small pieces of material do not jump out of the mortar when starting to break up the hard frozen material with the pestle. 5. Transfer the powder into a 2 mL Eppendorf tube using a special spatula (Fig. 1). Forceps are used to freeze the tube briefly in N2 and then to hold the tube close to the mortar in the N2-box. Care is taken that no powder is left in the mortar or on the pestle. For collection of this powder always use the same type of plastic tube. This contributes to reproducibility of the subsequent sonication step. Compress the powder collected in the tube by gently knocking the tube against the mortar. The powder can be stored at −70°C or immediately subjected to sonication. 6. For sonication a predetermined number of glass beads (see Table 2 and Note 1) is added to the sample and the powder is then thawed and kept immersed in ice. Sonication is performed in an ice-cold water-bath. The fill-height of the water is critical

– Stirring – Centrifugation Supernatant II add to I Supernatant I+II weigh – Aliquot of supernatant I+II (store rest of supernatant I+II frozen) Urea (50 µL × 1.08)

– Liver pieces Buffer A (liver mg × 1.5)1 Σ1 Inhibitor 1A (Σ1 × 0.08) Inhibitor 2 (Σ1 × 0.01) Σ2 – Liver powder – Sonication, 6 × 10 sec Number of glass beads (Σ2 × 0.034) – Centrifugation Supernatant I store frozen – Pellet I weigh Buffer A (pellet I mg × 2) Σ3 Inhibitor 1A (Σ3 × 0.08) Inhibitor 2 (Σ3 × 0.01) – Pellet powder

SI+II fraction: Liver

54 mg

628 mg 5 (B)3 50 µL

110 mg4 220 µL 330 26.4 µL 6.6 µL

23

250 mg2 (A)3 375 µL 625 50 µL 12.5 µL 688

Inhibitor 1A (Σ3 × 0.08) Inhibitor 2 (Σ3 × 0.01) – Pellet powder

Inhibitor 1A (Σ3 × 0.08) Inhibitor 2 (Σ3 × 0.01) – Pellet powder – Sonication, 6 × 10 sec Number of glass beads [Σ3 + (Σ3 × 0.08) + (Σ3 × 0.01)] × 0.034 – No stirring – Centrifugation Supernatant II add to I Supernatant I+II weigh – Aliquot of supernatant I+II (store rest of supernatant I+II frozen) Urea (50 µL × 1.08)

(continued)

– Stirring – Centrifugation Supernatant II add to I Supernatant I+II weigh – Aliquot of supernatant I+II (store rest of supernatant I+II frozen) Urea (50 µL × 1.08)

– Centrifugation Supernatant I store frozen – Pellet I weigh Buffer A (pellet I mg × 1.0)

– Heart powder – Sonication, 12 × 10 sec

– Brain powder – No sonication – Centrifugation Supernatant I store frozen – Pellet I weigh Buffer A (pellet I mg × 0.5)

Inhibitor 1A (Σ1 × 0.08) Inhibitor 2 (Σ1 × 0.01)

Inhibitor 1A (Σ1 × 0.08) Inhibitor 2 (Σ1 × 0.01)

Heart – 100-130 mg (total heart) Buffer A (liver mg × 1.0)

Brain – 25-250 mg fine pieces No buffer

Table 2 Fractionated Tissue Protein Extraction Protocol

Mouse and Human Tissue Sample Preparation 95

– 2-D electrophoresis PE fraction: Liver – Pellet II weight Buffer B/CHAPS (pellet II mg × 1.6) 900 µL buffer B 73 mg CHAPS (displace 69 µL) 31 µL bidistilled water 1000 µL buffer B/CHAPS Σ4 Inhibitor 1B (Σ4 × 0.08) Σ5 – Pellet powder Stirring – Urea [(pellet II mg × 0.3) + Σ5] × 1.08 DTT solution [(pellet II mg × 0.3) + Σ5] × 0.1 Stirring – Centrifugation

DTT solution (50 µL × 0.1) Ampholyte pH 2-4 (50 µL × 0.1) Final volume of 50 µL supernatant plus additives Diluent Supernatant I+II, ready for use, store frozen in aliquotes

SI+II fraction: Liver

Table 2 (continued)

9 µL/gel Brain – Pellet II weight Buffer B/CHAPS (pellet II mg × 1.4) 900 µL buffer B 77 mg CHAPS 27 µL bidistilled water 1000 µL buffer B/CHAPS

8 µL/gel

19 µL

– DTT solution [(pellet II mg × 0.56) + Σ5] × 0.1 Stirring – Centrifugation

239 19 µL Inhibitor 1B (Σ4 × 0.08) 19+1476 = 166 – Pellet powder Stirring 207 mg – Urea [(pellet II mg × 0.56) + Σ5] × 1.08

92 mg 4 (C)3 147 µL

100 µL 200 µL

DTT solution (50 µL × 0.1) Ampholyte pH 2-4 (50 µL × 0.1) Final volume of 50 µL supernatant plus additives No diluent Supernatant I+II, ready for use, store frozen in aliquotes

5 µL 5 µL 100 µL

Brain

– DTT solution [(pellet II mg × 0.25) + Σ5] × 0.1 Stirring – Centrifugation

– Pellet powder Stirring – Urea [(pellet II mg × 0.25) + Σ5] × 1.08

Inhibitor 1B (Σ4 × 0.08)

6 µL/gel Heart – Pellet II weight Buffer B/CHAPS (pellet II mg × 2.2) 900 µL buffer B 65 mg CHAPS 38 µL bidistilled water 1000 µL buffer B/CHAPS

DTT solution (50 µL × 0.1) Ampholyte pH 2-4 (50 µL × 0.1) Final volume of 50 µL supernatant plus additives Diluent Supernatant I+II, ready for use, store frozen in aliquotes

Heart

96 Zabel and Klose

9 µL/gel

9.4 µL

79 mg 7.3 µL

3.5+697 = 73

3.5 µL

69 mg4 69 µL 138

8 µL/gel

375 mg5 (D)3 20 µL

8 µL/gel

8 µL/gel

– Urea (Σ7 × 1.08) DTT solution (Σ7 × 0.1) Stirring Ampholyte pH 2-4 [(pellet III mg × 0.25) + Σ7] × 0.1

– Pellet powder – Benzonase (Merck) (Σ6 × 0.025) Stirring

– Pellet powder – Benzonase (Merck) (Σ6 × 0.025) Stirring – Urea (Σ7 × 1.08) DTT solution (Σ7 × 0.1) Stirring Ampholyte pH 2-4 [(pellet III mg × 0.56) + Σ7] × 0.1

Supernatant III weigh Ampholyte pH 2 – 4 (Serva) (supernatant III mg × 0.0526) Pellet extract, ready for use, store frozen in aliquots 7 µL/gel Heart – Pellet III Buffer C (pellet III mg × 1.0)

Supernatant III weigh Ampholyte pH 2 – 4 (Serva) (supernatant III mg × 0.0526) Pellet extract, ready for use, store frozen in aliquots 8 µL/gel Brain – Pellet III Buffer C (pellet III mg × 1.0)

2

All factors used in this table are explained in Note 8. This figure is given as an example. The amount of starting material may vary from 25 to 250 mg (see Note 4). 3 B ÷ A = control value; D ÷ C = control value (see Note 2). 4 This figure is provided as an example and is not the result of a calculation. The pellet weight varies because slight losses of material are unavoidable, even during very precise work. 5 See Footnote 4 for pellets; this also holds true for supernatants. 6 Buffer B/CHAPS volume. 7 Buffer C.

1

– Urea (Σ7 × 1.08) DTT solution (Σ7 × 0.1) Stirring Ampholyte pH 2-4 [(pellet III mg × 0.3) + Σ7] × 0.1 Pellet suspension, ready for use, store frozen in aliquots – 2-D electrophoresis

Supernatant III weigh Ampholyte pH 2 – 4 (Serva) (supernatant III mg × 0.0526) Pellet extract, ready for use, store frozen in aliquots – 2-D electrophoresis PS fraction: Liver – Pellet III Buffer C (pellet III mg × 1.0) Σ6 – Pellet powder – Benzonase (Merck) (Σ6 × 0.025) Stirring Σ7

Mouse and Human Tissue Sample Preparation 97

98

7.

8.

9.

10.

11. 12. 13.

Zabel and Klose for the sonication effect and should always be at the level indicated in the instruction manual of the manufacturer. Furthermore, when dipping the sample tube into the water, it is important to do this at a “sonication center” visible by a concentric water surface motion which appear when holding the tube into the water. We prefer a small sonication apparatus that forms only one sonication center (see Subheading 2.1). The water must be kept ice-cold. Sonication is performed for 10 sec. Immediately afterwards the sample is stirred with a thin wire for 50 sec with the tube still remaining in the ice water. The tube is then kept immersed in ice for 1 min. Now the next sonication circle is started, until a total of 6 two-minute steps (see Note 1). After sonication, the tubes are turned upside down and punctured with a steel needle to generate a 2 mm diameter opening. The holes should not be larger than that to keep the glass beads from crossing. A second tube is attached to the bottom of the first. Both tubes are transferred into a centrifuge using other samples or dummy tubes for balancing. Tubes are quickly centrifuged at 2,000 g for 1 min. The sample is thereby transferred into the lower tube whereas the glass beads remain in the upper. The homogenate is then frozen in liquid nitrogen and can be stored at −70°C. Be careful to hold the tube at an angle of about 45° to decrease the difficulty of retrieving the frozen homogenate. Detach the frozen homogenate in the tube from the wall by quickly knocking at the tube with a large screw driver. Transfer the frozen piece of sample into a preweighed centrifuge tube and briefly thaw the homogenate. Centrifuge for a maximum of 226,000 g for 30 min at 4°C. Completely withdraw the supernatant (I) with a pasteur pipette and fill into a preweighed small test tube. The centrifuge tube is kept on ice and the pipette is put back into the tube again with the tip at the center of its bottom (Note: the pellet sticks to the wall if a fixed-angle rotor was used). In this position, remainders of the supernatant accumulate at the bottom of the tube and inside the pipette. Withdraw residual supernatant with the pipette and transfer it to the test tube. Now the supernatant is frozen in liquid nitrogen and stored at −70°C. Weigh the pellet (I) left in the centrifuge tube on ice and add buffer A at an amount calculated as indicated in Table 2. Mix the pellet and buffer by vortexing, collect the homogenate at the bottom by a short spin, freeze in liquid nitrogen and store at −70°C, or process further immediately. Transfer the homogenate from the centrifuge tube to the mortar after detaching the frozen homogenate from the wall by knocking onto the bottom of the tube. Grind the homogenate together with inhibitors 1A and 2 to powder as described above for the liver pieces. Transfer the powder back into the used centrifuge tube, taking care that no powder remains in the mortar. Thaw the powder and slowly stir the homogenate for 45 min in a cold room. Centrifuge the homogenate as described in step 7 above. Completely withdraw the supernatant (II) from the pellet. This is done in such a way that a white layer which partially covers the surface of the supernatant sinks unaffected onto the pellet. Collect the remainder of the supernatant as mentioned in step 8. Add supernatant II to supernatant I and thoroughly mix both solutions. Determine the weight of the total supernatant.

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14. Take a 50 µL-aliquot from the total supernatant and mix with urea, DTT-solution and ampholyte pH 2-4, as indicated in Table 2. The final concentrations of these components are: 9 M urea, 70 mM DTT and 2% ampholytes. These three components should be added to the supernatant in the order indicated here and each component should be mixed and dissolved in the supernatant before adding the next. The final volume of this supernatant mixture is 100 µL. Add 100 µL sample diluent (see Table 3) and mix before IEF run. 15. The resulting solution is the final sample (”supernatant I + II”, SI + II). Divide the sample into several portions, freeze each portion in liquid nitrogen and store at −70°C. Usually 8 µL of sample are applied to an IEF gel, if the large gel 2-D electrophoresis technique described in Chapter 11 is used (see Note 5). Freeze the remaining portion of the pure supernatant and store at at −70°C. 16. Determine the weight of the pellet (II). Collect the pellet at the bottom of the tube by a short spin, then freeze in liquid nitrogen and store at −70°C. 3.2.1.2. EXTRACTION OF THE PELLET PROTEINS SOLUBLE IN THE PRESENCE OF UREA AND CHAPS (PELLET EXTRACT) 1. Grind pellet II, buffer B/CHAPS and inhibitor 1B to powder in a mortar placed in liquid nitrogen (see Subheading 3.2.1.1., steps 2-4). The calculations for buffer and inhibitor volumes are provided in Table 2. Transfer the powder back to the centrifuge tube, avoid leaving any remainders in the mortar or on the pestle. 2. Thaw the powder mix and stir slowly for 60 min in a cold room (CHAPS solubilizes structural proteins). 3. Add urea (for the amount see Table 2) to the homogenate and stir the mixture for 45 min at room temperature (urea reaction). Some minutes after adding urea a large part of it is already dissolved. At this point add DTT solution (for the amount see Table 2). 4. Remove the magnet rod from the homogenate. At this step, also avoid any loss of homogenate. Centrifuge the homogenate at 17°C for 30 min at 50,000 rpm. 5. Completely withdraw the supernatant (III) with a Pasteur pipette and fill it into a small pre-weighed test tube. Collect the remainders of the supernatant as mentioned in Subheading 3.2.1.1., step 8. Determine the weight of the supernatant. 6. Add ampholytes pH 2-4 (for amount see Table 2) to the supernatant and immediately mix solution.

Table 3 Sample Diluent Component Urea DTT-solution a Servalyt pH 2 – 4 (Serva) Bidistilled water Sample diluent a

See Subheading 2.2, item 5.

Mixture 1.08 0.10 0.10 1.00 2.00

Final concentration g ( =0.80 ml) ml ml ml ml

9.000 M 0.070 M 2.000 % 50.000 %

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7. The resulting solution is the final sample (”pellet extract”, PE). Divide the sample into several portions, freeze each portion in liquid nitrogen and store at −70°C. Usually the volume of sample applied per IEF gel is 8 µL, if large gel 2-D electrophoresis (see Chapter 18) is used (see Note 5). 8. Determine the weight of the pellet (III). Collect the pellet on the bottom of the tube by a short spin, then freeze in liquid nitrogen and store at −70°C. 3.2.1.3. SUSPENSION OF THE REMAINING PELLET (PELLET SUSPENSION) 1. Grind pellet III and buffer C to powder in a mortar as described in Subheading 3.2.1.1., steps 2-4. The buffer volume is calculated as indicated in Table 2. The powder is transferred into a test tube, avoiding any loss of material. 2. Thaw powder and add benzonase (Merck, Darmstadt, Germany; for the amount see Table 2). Slowly stir the homogenate for 30 min in a cold room (DNA digestion). 3. Add urea (for amount see Table 2) and stir the homogenate at room temperature for another 30 min. During this time add DTT solution (for amount see Table 2) once the largest part of urea is dissolved. At the end of this period add ampholytes pH 2-4 (for amount see Table 2) and quickly mix with the homogenate. 4. The resulting solution is the final sample (”pellet suspension,” PS). It is frozen in liquid nitrogen and stored at −70°C. Usually 9 µL of sample is applied per IEF gel (if large gel 2-D electrophoresis is used; see Chapter 11 and Note 6). The sample still contains some fine undisolved material and is therefore transferred to the gel with a thin Pasteur pipette instead of a microliter syringe.

3.2.2. Fractionated Extraction of Brain Proteins Frozen brain tissue is transferred into a mortar that was placed into a box containing liquid nitrogen, and crushed with the pestle to fine pieces. The crushed material is transferred completely back into test tubes so that one tube contains 25-250 mg of frozen tissue (weigh the tube without thawing the tissue). This material is used to prepare the supernatant I + II, the pellet extract and the pellet suspension. The procedure follows that of liver extraction with some differences indicated in Table 2. One exception is that the tissue powder is produced without buffer and subjected to centrifugation without sonication so that a rather small amount of supernatant I results. Sonication is only performed with pellet I homogenate. 3.2.3. Fractionated Extraction of Heart Proteins The supernatant I + II, the pellet extract and the pellet suspension are prepared from a single heart. The procedure is as described for liver with some modifications which are indicated in Table 2. 4. Notes 1. Sonication: Conditions for sonicating mouse tissue homogenates (liver, brain, heart) were optimized to break membranes of all cells and cell nuclei

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of a tissue. Three parameters were varied in the experiments: the time of sonication, the number of sonication repeats and the number of glass beads added per volume of homogenate. The effect of the various parameters was determined by inspection of sonicated material under the microscope. Sonication for 10 sec increases the temperature of the homogenate from 0°C to 11-12°C. Therefore, sonication was not performed for more than 10 sec. Glass beads are essential for breaking cellular structures (membranes). Since most of the homogenates are rather viscuous fluids, the beads cannot flow freely and their addition is limited. For a given homogenate volume a certain number of glass beads is necessary to expose it evenly to the disrupting power of sonication. Due to viscosity, an increase in this number has not much effect. This number can be calculated. By standardization experiments we determined that a factor 0.034 is a good approximation for the optimal number of glass beads for a given volume of homogenate. It can be concluded from the above-mentioned that the only parameter which can be varied without harming the sample while still increasing the effect of sonication, was the number of repeats of the 10 sec sonication period. Under the conditions described in the Methods section the membranes of all cells were broken and no longer visible under the microscope. However, a certain number of intact nuclei were still detectable. Still, this shortcoming was not compensated for by increasing the number of sonication repeats because we assume that homogenization, stirring and high-speed centrifugation may eventually extract all nucleoplasm proteins. A more aggressive sonication procedure using a metal tip cannot be recommended. We observed heavily disturbed 2-DE patterns as a result of employing this technique: many protein spots disappeared depending on the extent of sonication and new spot series occurred in the upper part of the gel, apparently as a result of aggregation of protein fragments. 2. Control values: Control values were calculated for each sample prepared by the fractionated extraction procedure to monitor correctness and reproducibility of the preparation. The calculation of control values is shown in Tables 1 and 2. To visualize the calculation procedure an example from a real experiment is provided: From a series of 73 individual mouse hearts the SI+II fractions were prepared and the control values B÷A (see Table 2) were calculated: 60 samples yielded values between 1.97-2.10, three samples between 1.94-1.96, and six samples between 2.11-2.13. Four samples with largest deviations (1.90, 1.91, 2.16 and 2.24) were excluded from the investigation. The range 1.97-2.10 (mean 2.04 ± 0.04) was taken as reference value for preparation of mouse heart SI+II samples. 3. The extraction procedure was adapted for labeling with CyDyes. It is important that only urea and thiourea should be present in the sample for

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labeling. DTT and ampholines are added only afterwards since both bind CyDyes. In addition, determination of the protein concentration is now mandatory since the amount of CyDyes for labeling is calculated based on the protein concentration. Therefore a 5 µL aliquot of the sample is specifically used for this purpose (Table 1, protein concentration). 50 µg protein labeled by 400 pMol CyDye works well for brain samples. 4. The tissue amount indicated (25 mg − 150 mg) yields enough sample to run a large number of 2-DE gels so that less rather than more material may be used. In cases in which only a very small amount of tissue is available (e.g. 2-5 mg heart biopsy samples, 10-12 mg of two mouse eye lenses, early mouse embryos) a total protein extract should be prepared instead of SI+II, PE and PS fractions. Small plastic tubes and a glass rod with a rough surface at the well fitting tip may serve as mortar and pestle. 5. Amount of protein applied per gel: The protein amount applied to IEF gels (see Tables 1 and 2), contains about 100 µg protein. There is, however, no need to determine the protein concentration of each sample prepared in order to get protein patterns of reproducible intensities. The concept of the procedure described here for extracting tissues was to keep the volume of the extracts in strong correlation to the amount of the starting material (tissue or pellet) which was extracted. Therefore, by working precisely, the final sample should always contain nearly the same protein concentration. This was confirmed by determining the protein concentrations of a large number of protein samples. Accordingly, the reproducibility of the pattern intensity depends on the precise sample volume applied to the gel – and, of course, on the protein staining procedure. The sample volumes per gel given in Table 2 are adapted for silver staining protocols. As a general guideline one should take into account: decreasing the protein amount per gel and increasing the staining period is better than the vice versa; diluted samples using reasonable volumes are better than concentrated protein samples at a small volume since in the first case clogging is avoided. 6. Maximum resolution of tissue protein fractions by 2-D electrophoresis: Fig. 2 shows 2-DE patterns of three protein fractions SI+II, PE and PS of mouse liver. The SI+II fraction reveals the highest number of protein spots.

Fig. 2. (continued) pattern and vice versa. Pellet-specific spots occur predominantly in the basic half, supernatant-specific spots more on the acid half of the pattern. The pellet suspension pattern reveals very basic proteins of a tissue extracts. Since IEF gels do not cover the entire basic pH range, these proteins cannot reach their isoelectric points. To prevent these proteins from accumulating at the basic end of a gel, the IEF run was shortened by 2 h at the 1000 V level. Consequently, the very basic proteins form streaks instead of focused spots. For evaluation of patterns in terms of spot number see Notes 6 and 7.

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Fig. 2. 2-DE protein patterns from mouse liver. Tissue was fractionated into supernatant I + II (a), pellet extract (b), and pellet suspension (c) as described in Subheading 3.1. Three fractions were subjected to large gel 2-D electrophoresis (see Chapter 11). In the pellet extract pattern many protein spots were visible which are not present in the supernatant

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When spots were counted visually, i.e. by placing a 2-DE gel on a light box and dotting each spot with a pencil (3), about 9200 proteins were detected in this fraction. The SI+II pattern of the brain revealed about 7700 proteins, that of the heart about 4800 protein spots. The high spot numbers reflect the high resolution of the large gel 2-DE (see Chapter 11), which reveals many weak spots between major spots. All these spots were counted with high precision and great care. 7. Effect of fractionated extraction of tissue proteins: The purpose of fractionating the proteins of a tissue was to increase the number of proteins detectable by 2-DE. This goal, however, would only be achievable if each fraction contained a notable number of proteins which are strongly fraction-specific, so that the total tissue proteins can be distributed over several gels. Comparison of the 2-DE patterns from the SI+II and PE fractions of the liver (Fig. 2) showed that the PE pattern revealed about 2000 protein spots not detectable among the 9200 spots of the SI+II pattern. The PS pattern revealed only about 70 additional spots. The PS protein spots represent classes of the most basic proteins (Fig. 2) and belong mainly to the chromosomal proteins (e.g. histones). Therefore, the PS fraction is only of interest when a class of very basic proteins is subject of the investigation. Considering the 2-DE patterns of the SI+II and PE fraction in more detail (Fig. 3), quite a number of very prominent spots can be observed which are visible in one pattern but do not occur, not even in trace amounts, in the others. At the same time, other spots revealing only low intensities are present in both patterns. This suggests that the phenomenon where many protein spots of the SN I+II pattern also occur in the PE pattern is not due to impurities of the pellet fraction by supernatant proteins caused by technical problems. We found, using centrifugation at different speeds and samples prepared with different degrees of viscosity, that an overlap in 2-DE patterns is due to the fact that most proteins exist solitarily as well as in protein-complexes. Apparently, protein complexes sink to the pellet at high speed and are later resolved again by detergent and urea extraction in the PE fraction. Taking into account all three fractions, the total liver proteins could be resolved into about 11,270 different proteins (polypeptide spots). This, however, does not mean that the protein sample preparation procedure described here – in combination with the 2-DE technique in Chapter 32 – revealed all proteins of the liver. Many proteins may exist in a tissue in undetectable amounts, and special fractionation procedures with subsequent protein concentration steps would be required to detect these proteins. We isolated liver and brain cell nuclei and extracted the nuclear pellet in a similar way as was done for the tissues. The 2-DE patterns showed that the nuclear extracts add a large number of new proteins

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Fig. 3. Sections from 2-DE patterns shown in Fig. 2. The supernatant I + II (a) and the pellet extract (b) of liver are compared. (i) Some of the prominent protein spots present in the supernatant pattern () but completely absent in the pellet extract pattern(), (ii) the reverse situation(), and (iii) some spots of low intensity present in both patterns() are indicated. Other spots show a high intensity in one pattern but low intensity in the other. These spots may reflect naturally occurring unequal distributions of proteins between the two different fractions due to a formation of protein complexes rather than contaminations of one fraction by the other (see Note 7).

to those already known from the tissue extract patterns. However, protein spots present in both the nuclear and the tissue extracts occur as well, particularly in the acidic halves of the supernatant patterns. In general, the proteins represented by the SI+II, PE, and PS patterns can be considered as the main population of protein species of a tissue to which further species may be added by analyzing purified and concentrated subfractions. There-

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fore, the amount of spots detectable with fractionated extraction is larger than by total extracts but the effort involved is also exceptionally higher. 8. Explanation of correction factors used in Table 2: Factors were calculated to determine the amounts of urea, DTT-solution and ampholytes necessary to transmute any volume of a solution (theoretically water) into a mixture containing 9 M urea, 70 mM DTT and 2% ampholytes. Calculations of factors: 500 µL water (aqueous protein extract) + 540 mg urea (displaces 400 µL) + 50 µL DTT-solution (see Subheading 2.2 item 5) + 50 µL ampholyte solution (commercial solutions which usually contain 40% ampholytes) = 1000 µL. If the volume of a protein solution to be mixed with urea, DTT and ampholytes is n µL, the amounts of the components to be added are: (540 ÷ 500) × n = 1.08 × n mg urea, (500 ÷ 50) × n = 0.1 × n µL DTT solution and 0.1 × n µL ampholyte solution. If the protein solution already contains urea and DTT (see Table 2: PE preparation), the factor 0.0526 is used to calculate the ampholyte volume for this solution. Calculation of this factor: (500 µL extract + 400 µL urea + 50 µL DTT solution) ÷ 50 µL ampholyte solution = 0.0526. If a protein solution of n µL includes a cell pellet, i.e. insoluble material, n µL volume should be reduced by the volume of insoluble material (theoretically by the volume of the dry mass of this material). For this reason a pellet factor (e. g. 0.3) was introduced. This factor was determined experimentally using urea as an indicator. The factor reduces the volume n µL of a protein solution (containing a pellet) to volume n µL; n µL × 1.08 yields an amount of urea that is added to the n µL of the solution, at the border of solubility, i.e. about 9 M. Note that pellet III contains urea and DTT by the foregoing steps. Therefore, in this case the pellet volume was not taken into account when calculating the amounts for urea and DTT to be added to the final pellet suspension. In all these calculations no distinction was made between values measured in volumes (µL) and values measured in weights (mg). This makes the calculation somewhat incorrect but more practicable and reproducible. The inhibitor 2 solution was prepared as concentrated as possible to keep the volume of this solution small (1/50th of the homogenate volume, i.e. factor 0.01). This allows for ignoring the volume determination error that results when this inhibitor solution is added instead of including it to the total volume of the homogenate. The factor 0.08 for all inhibitor 1 (A, B and C) solutions was derived from the dilution requirements according to manufacturers instruction (CompleteTM tablets, ROCHE, Mannheim, Germany): 1 tablet should be dissolved in 2 mL buffer and this volume added to 25 mL of the homogenate, i.e. the volume of the inhibitor 1 solution to be added to n µL homogenate is (2 ÷ 25) × n = 0.08 n µL. The volume of benzonase solution (ready-made solution from MERCK, Darmstadt, Germany) necessary to

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digest DNA in the pellet III suspension was determined experimentally: a chromatin pellet was prepared from isolated liver cell nuclei and found to change from a gelatinous clot to a fluid if treated as follows: 1 g chromatin pellet + 1 mL buffer + 0.050 mL benzonase solution (= 0.025 mL Benzonase / mL homogenate), stirred for 30 min at 4°C. If benzonase from other sources is used the amount added has to be adjusted. Buffer factors used to calculate the volumes of buffer added to tissues or pellets are explained in the Introduction. Note that tissue to be homogenized must be free of any wash solutions otherwise the calculated buffer volumes will be too high. Acknowledgment The authors appreciate critical comments and support from Marion Herrmann and Yvonne Kläre. References 1. Klose, J. and Kobalz, U. (1995) Two-dimensional electrophoresis of proteins: an updated protocol and implications for a functional analysis of the genome. Electrophoresis 16, 1034–1059. 2. Challapalli, K. K., Zabel, C., Schuchhardt, J., Kaindl, A. M., Klose, J., and Herzel, H. (2004) High reproducibility of large-gel two-dimensional electrophoresis. Electrophoresis 25, 3040–3047. 3. Klose, J. (1999) Fractionated extraction of total tissue proteins from mouse and human for 2-D electrophoresis. Meth. Mo.l Biol. 112, 67–85. 4. Zabel, C., Sagi, D., Kaindl, A. M., Steireif, N., Klare, Y., Mao, L., Peters, H., Wacker, M. A., Kleene, R., and Klose, J. (2006) Comparative proteomics in neurodegenerative and non-neurodegenerative diseases suggest nodal point proteins in regulatory networking. J Proteome Res. 5, 1948–1958.

13 Plant Protein Sample Preparation for 2-DE Sebastien Christian Carpentier, Rony Swennen, and Bart Panis 1. Introduction Most plant tissues are not a ready source for protein extraction and need specific precautions. The cell wall and the vacuole make up the majority of the cell mass, with the cytosol representing only 1 to 2 % of the total cell volume. Subsequently, plant tissues have a relatively low protein content compared to bacterial or animal tissues. The cell wall and the vacuole are associated with numerous substances responsible for irreproducible results such as proteolytic breakdown, streaking and charge heterogeneity. Most common interfering substances are phenolic compounds, proteolytic and oxidative enzymes, terpenes, pigments, organic acids, inhibitory ions, and carbohydrates. Some species contain extremely high levels of interfering compounds. Banana (Musa spp.), for example, contains extremely high levels of oxidative enzymes (polyphenol oxidase) (1–3) and phenol compounds (simple phenols [dopa], flavonoids, condensed tannins, and lignin). Moreover, it contains high amounts of latex and carbohydrates (4). Phenolic compounds reversibly combine with proteins by hydrogen bonding and irreversibly by oxidation followed by covalent condensations (5), leading to charge heterogeneity and streaks in the gels. It is known that carbohydrates can block gel pores causing precipitation and extended focusing times, resulting in streaking and loss. In the 1980s, much effort had been invested in the establishment of twodimensional (2-D) gel electrophoresis sample preparation methods for plant tissue (6–8). The original 2-DE protocol consists of only one step, i.e., denaturing extraction in a lysis buffer (9). This one step protocol is restricted to “clean” samples and is rarely used for plant material. The majority of the plant protocols introduce a precipitation step to concentrate the proteins and to separate them from the interfering compounds. Proteins are usually precipitated by the addition

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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of high concentrations of salts (10), extreme pH (11), organic solvents (12–14) or a combination of organic solvents and ions (15,16). Typical ions used for precipitation are ammonium sulfate and ammonium acetate. Ions effectively compete with water molecules on the protein surface, favoring protein-protein interactions over protein-solvent interactions and causing protein aggregation and precipitation. Organic solvents also decrease protein solubility by dehydrating the protein surface. They lower the solution dielectric constant, promoting charge-charge interactions between proteins. Typical organic solvents used for precipitation are methanol, ethanol, and acetone. Organic solvent precipitation is generally performed at low temperatures, since proteins are then less “soluble” (17). The protein precipitation step can be preceded by a denaturing or non-denaturing extraction step and is combined with one or two washing steps to remove introduced salt ions and other remaining interfering substances. The most commonly used method for extraction of plant proteins is the TCA/acetone precipitation method (15). Proteins are very sensitive to denaturation at low pH. Trichloroacetic acid (TCA) is a strong acid (pKa 0.7) that is soluble in organic solvents. The extreme pH and negative charges of TCA together with the addition of acetone realizes an immediate denaturation of the protein along with precipitation, thereby arresting instantly the action of proteolytic enzymes (18). However, a disadvantage of TCA precipitated proteins is that they are difficult to redissolve (19) and the extreme low pH might create problems with basic chemical labeling methods. The development of succinimidyl ester derivatives of different cyanine fluorescent dyes that modify free amino groups of proteins prior to separation (20) was a major achievement for 2-DE in terms of reproducibility and throughput. The difference gel electrophoresis (DIGE) minimal labeling approach uses fluorophores that have a different absorption optimum, making it possible to run multiple samples simultaneously in the same gel. The different dyes were designed to assure that a protein acquires the same relative mobility irrespective of the dye used to tag them. The difference in MW introduced by the different length linkers is compensated by the different alkyl moieties opposite the linker moiety. Succinimidyl ester derivatives react with the nucleophilic primary amines, subsequently releasing the N-hydroxysuccinimide group. At a specific pH (8.5), these reagents react almost exclusively with the ε-amino group of lysine to form stable amide linkages that are highly resistant to hydrolysis. The pH of the protein solution is hence of utmost importance. We have tested in the past different protocols to extract proteins from plant samples (21). In our hands, the phenol extraction protocol proved to be the most powerful. The protein precipitation step is in this method achieved by ammonium acetate and methanol and is preceded by a denaturing phenol extraction. We optimized this method for small amounts of banana fresh weight (21) and lyophilized tissues (22). This “micro” phenol protocol is applicable for a wide

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variety of plant species since we successfully applied it to apple, banana, pear (23), potato and stevia. We describe here in detail this successful “micro” phenol extraction protocol. 2. Materials 1. All the materials that were used were clean and the chemicals are of the highest purity in order to not interfere with the electrophoresis process and with the mass spectrometry. 2. DTT (dithiothreitol) has a limited shelf life and should not be added to the different stock solutions. Make always a fresh 10% (w/v) solution and add prior to use. 3. All consumables were purchased from Acros Organics (Geel, BE) unless stated otherwise.

2.1. Protein Extraction 1. 2. 3. 4. 5.

6. 7. 8. 9.

10.

Liquid nitrogen. Pestle and mortar (washed and baked in an oven at 180°C overnight prior to use). Phenol, acetone and methanol compatible tubes and pipettes. 10 % (w/v) DTT (GE Healthcare, Diegem, BE). Make freshly in MQ H2O. Extraction buffer: 50 mM Tris-HCl pH 8.5, 5 mM EDTA, 100 mM KCl, 1% (w/v) DTT, 30% (w/v) sucrose and complete protease inhibitor cocktail (according to the manufacturer, Roche Applied Science) in MQ H2O. Prepare freshly (see Note 1). Tris buffered phenol (pH 8.0) (see Notes 2 and 3). Precipitation buffer: 100 mM ammonium acetate in methanol. Store at −20°C. Cold acetone. Store at −20°C. Lysis buffer: 7 M urea (GE Healthcare, Diegem, BE), 2 M thiourea (Fluka, Bornem, BE), 4% (w/v) CHAPS (GE Healthcare), 0.8% (v/v) IPG-buffer (GE Healthcare), 1% (v/v) DTT (GE Healthcare) and 30mM Tris in MQ H2O. Store aliquots at −80°C (see Notes 4 and 5). Protein Quantification kit 2D Quant (GE Healthcare).

2.2. Protein Separation 1. 1% (w/v) bromophenol blue: bromophenol blue (GE Healthcare) and 50 mM Trisbase in MQ H2O. 2. Rehydration buffer: 6 M urea, 2 M thiourea, 0.5% (w/v) CHAPS, 10% (v/v) glycerol, 0.002% (v/v) bromophenol blue, 0.5% (v/v) IPG-buffer and 0.28% (v/v) DTT in MQ H2O (see Note 6). 3. 24 cm IPG-strips pI 4-7 (GE Healthcare) (see Note 7). 4. Reswelling tray (GE Healthcare). 5. Isoelectric focusing apparatus IPGphor II (GE Healthcare). 6. Sample cups (GE Healthcare). 7. Ceramic 12-strip sample tray (GE Healthcare). 8. Filter paper electrode pads (GE Healthcare). 9. Mineral cover oil (GE Healthcare).

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10. 11. 12. 13. 14.

Gel casting chamber (GE Healthcare). SDS-PAGE apparatus Ettan Dalt 6 (GE Healthcare). Cooling bath type 1140S (VWR international, Leuven, BE). Power supply Power PAC 3000 (Bio-Rad, Nazareth, BE). Equilibration buffer I: 6 M urea, 30% (v/v) glycerol, 2% (v/v) SDS, 0.002% (v/v) bromophenol blue, 50 mM Tris-HCl pH 8.8 and 1% (w/v) DTT in MQ H2O. Store aliquots at −20°C. Equilibration buffer II: 6 M urea, 30% (v/v) glycerol, 2% (v/v) SDS, 0.002% (v/v) bromophenol blue, 50 mM Tris-HCl pH 8.8 and 4.5% (w/v) iodoacetamide in MQ H2O. Store aliquots at −20°C (see Note 8). SDS 10%. Store at room temperature (see Note 9). Laemmli running buffer: 25mM Tris, 0.1% (v/v) SDS, 192 mM glycine in demineralized water. Do not adjust the pH. Store a 10x buffer at room temperature. Agarose sealing solution: 0.5% (w/v) agarose (ultra pure, GibcoBRL, Merelbeke, BE) 0.002% (v/v) bromophenol blue in Laemmli running buffer. 10% (w/v) Ammonium persulfate. Make freshl in MQ H2O. Thirty percent acrylamide/bis solution (37.5:1 with 2.6% C) (Bio-Rad). Store at 4°C (see Note 10). N,N,N,N -Tetramethyl-ethylenediamine (TEMED) (Bio-Rad). Store at room temperature (see Note 11). Tris 1.5M pH 8.8 in MQ H2O. Store at 4°C. Water-saturated isobutanol. Shake equal volumes of MQ H2O and isobutanol and allow separation. Use the top layer. Store at room temperature.

15.

16. 17. 18. 19. 20. 21. 22. 23.

2.3. Protein Labeling and Scanning 1. 2. 3. 4. 5.

10mM lysine. CyDye DIGE fluors (Cy2, Cy3 and Cy5) (GE Healthcare). Store at −20°C. N,N-dimethylformide (DMF). Store at room temperature (see Note 12). Typhoon™ imager (GE Healthcare). Low fluorescent glass plates (GE Healthcare).

3. Methods 3.1. Protein Extraction 1. Transfer fresh plant tissue to a liquid nitrogen pre-cooled mortar and grind in liquid nitrogen (see Note 13). 2. Transfer 50 to 100 mg of frozen tissue powder to an extraction tube (2 mL), add 500 µL extraction buffer and vortex 30 sec (see Note 14). 3. Add 500 µL of buffered phenol and vortex 10 min at 4°C. 4. Centrifuge for 3 min, 6000 g at 4°C, collect the upper phase, i.e., the phenolic phase, transfer to a new tube and discard the lower phase (see Note 15). 5. Re-extract by adding 500 µL of new extraction buffer. 6. Centrifuge for 3 min, 6000 g at 4°C. 7. Transfer the phenolic phase into a new 2 mL tube and precipitate the proteins overnight (or at least 2.5 h) with 5 volumes 100 mM ammonium acetate in methanol at −20°C.

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8. Centrifuge 60 min, 16 000 g, at 4°C. 9. Remove the supernatant and rinse the pellet twice (do not resuspend) in 2 mL rinsing solution (cold acetone/0.2 % DTT). After the first rinse leave in rinsing solution for 1 hour at −20° C. After rinsing, centrifuge 30 min, 13 000 rpm, at 4°C. 10. Dry the pellet (see Note 16). 11. Suspend the pellet in 100 mL lysis buffer (optimum concentration is 1–5 µg/µL) (see Note 17). 12. Clear samples by centrifugation (16 000 g, 2 × 30 min at 18° C) (see Notes 18 and 19). 13. Quantify the samples (see Note 20).

3.2. Protein Labeling 1. Briefly spin the fluor stock solutions. 2. To 4.2 µL DMF add 2.8 µL DMF reconstituted fluor stock solution (1 nmol/ µL) in a microfuge tube (200 µL). Ensure all fluor is removed from the pipette tip by pipetting up and down. This is the working fluor sample solution. 3. Add 50 µg of protein sample (without DTT and IPG buffer) to a microfuge tube (200 µL) and add 1 µL of working fluor sample solution. 4. Mix fluor and protein sample thoroughly and leave on ice for 30 min in the dark. 5. Repeat this for all samples (see Note 21). 6. Add 1 µL of 10 mM lysine to stop the reaction. Mix and leave for 10 min on ice in the dark. The labeled samples can be stored for at least 3 months at −80°C. 7. Dilute the samples with an equal volume of Lysis Buffer (with 2x DTT and 2x IPG buffer) and leave at room temperature for 10 min.

3.3. Protein Separation: First Dimension 1. Rehydrate IPG dry strips with rehydration buffer in a reswelling tray for at least 8 h and cover with mineral oil. 2. Apply the rehydrated IPG gel strips onto the cup-loading tray, gel side upward and acidic ends facing toward the anode. 3. Moisten two filter paper electrode pads with demi water and apply the moistened filter paper pads on the surface IPG gel covering the anodic and cathodic ends of the IPG strip. 4. Position the movable electrodes at the extremes of the electrode filter paper pads. 5. Overlay the IPG strips (and empty lanes) with several mL of cover fluid ( 120 mL) (see Note 22). 6. Position the movable sample cup either near the anode, and gently press the sample cup onto the surface of the IPG gel strip (see Notes 23 and 24). 7. Dilute your sample in rehydration buffer (final volume 150 µL) and pipette the sample into the cup. 8. Program the instrument (desired volthours, voltage gradient, temperature, etc.) and run IEF according to the settings recommended in Table 1. Current and power settings should be limited to 0.05 mA and 0.2 W per IPG gel strip, respectively. Optimum focusing temperature is 20°C. 9. After IEF, IPG gel strips can be stored between two sheets of plastic film at −78°C up to 3 months.

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Mode

Voltage

Step and hold Gradient Gradient Step and hold

300 1000 8000 8000

3h 6h 3h 32 kVh

3.4. Protein Separation: Second Dimension 1. Cast the polyacrylamide gels. For 600 mL (6 gels) 12.5% acrylamide gels: add 250 mL 30% acryl/bisacryl, 150 mL Tris buffer pH8.8, (1.5 M), 187 mL MQ H2O, 300 µL TEMED, 6 mL 10 % SDS and 2.4 mL 10% APS, under continuous stirring (see Note 25). 2. Cover with water saturated isobutanol and cover to prevent drying out. 3. Allow polymerization overnight (see Note 26). 4. Remove the isobutanol prior to use by rinsing with MQ (see Note 27). 5. Place the IPG strips in an individual tube and add 7 mL of equilibration buffer I. Shake for 15 min. 6. Remove the first equilibration solution and replace by 7 mL of equilibration buffer II. Shake for 15 min. 7. Prepare the running buffers for upper (2x) and lower chamber (1x) and fill the lower chamber. 8. Rinse the strips with running buffer. 9. Place the gel cassettes in an upright position and apply the strips and fill with 2–3 mL of hot (60°C) 0.5 % agarose solution. Make sure that the strip is in close contact with the second dimension gel without air bubbles. 10. Place the cassettes in the running tank, place the upper buffer chamber and fill it further with running buffer. 11. Run the gels at 2 W per gel overnight.

3.5. Scanning 1. Prescan the gels at the 3 wavelengths at a low resolution and adjust the PMT (photomultiplier tube) to ensure a maximum pixel intensity between 70,000 and 100,000 pixels using a Typhoon™ imager (GE Healthcare). 2. Scan the gels at the optimal pixel intensity. See Fig. 1 for a representative DIGE gel. A protein spot map realized in (24) containing MS/MS identified spots can be found at http://www.pdata.ua.ac.be/musa/

4. Notes 1. Our extraction buffer has been designed to minimize enzymatic reactions and to remove as much interfering compounds as possible. Sucrose is added to the buffer to create a phase inversion. The buffer forms the aqueous

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Fig. 1. Representative gel of the banana meristem proteome labelled with Cy3. 24 cm IPG strips pI 4–7.

2.

3.

4.

5. 6.

lower phase containing carbohydrates, nucleic acids and cell debris and the upper phenol rich phase contains cytosolic and membrane proteins, lipids, and pigments. The high pH of the buffer inhibits most common proteases, assures that the abundant phenolic compounds are mainly ionized (inhibiting H bonding with the proteins) and neutralizes acids that are released by disrupted vacuoles. KCl facilitates the extraction of proteins (salting in effect) and EDTA inhibits metalloproteases and polyphenol oxidase by chelating the metal ions. DTT is a powerful reducing agent that does not actually prevent the oxidation of plant (poly)-phenols (quinones) but reduces them to form thio-ethers. The protease inhibitor cocktail is added to inhibit proteases that are released upon cell rupture. The pH of the phenol is important. We opted to buffer the phenol to pH 8.0 instead of water buffered phenol (25–27) to assure that the interfering nucleic acids are partitioned to the buffer phase and not to the phenol rich phase. Phenol is toxic and needs to be handled with great care. Wear gloves and work in a dedicated room. Use only under the hood and make sure that the waste (tips, pipettes, tubes) is disposed safely. Tris is added to the lysis buffer to get the pH stable between 8 and 9 to assure proper labeling and to avoid pH adjusting with other salts afterward since salts affect the electrophoresis process. If the proteins are visualized with a post staining procedure the Tris can be omitted. DTT and IPG-buffer are added freshly prior to use. For the DIGE labeling they must be added after labeling since they are subject to labeling. Glycerol is added to the buffer in order to obtain an even reswelling of the strips and to minimize electro-osmotic flow (counter flow of water ions).

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7. The size of the strips plays an important role in the resolution of the gel. The longer the strip, the better the resolution. 8. As for the DTT, the iodoacetamide is not added to the stock solution but prior to use. 9. SDS is irritating. The powder should be handled with care since it can be inhaled very easily and should always be weighted under a hood. Make always a stock solution of 10% (w/v). 10. Acrylamide is a neurotoxin when unpolymerized and so care should be taken not to receive exposure. Wear gloves, work under a hood in a dedicated room. 11. TEMED has a limited shelf life. Buy therefore small amounts. The quality is important for a good polymerization. 12. The DMF must be high quality anhydrous (specification: 0.005% H2O, ≥ 99.8% pure) and every effort should be taken to ensure it is not contaminated with water. DMF, once opened will start to degrade to produce amine compounds. Those amines will react with the succinimidyl ester derivatives of the cyanine dyes, reducing the concentration of dye available for protein labeling. DMF is toxic and any contact should be avoided. Wear gloves. 13. The grinding process is very important. Do not stop grinding until the power is very fine and homogeneous. 14. It is important to keep this period as short as possible in order to keep enzymatic reactions like oxidation of polyphenols and protease activity to the strict minimum. All enzymes are considered to be irreversibly inactivated after adding phenol as is the case with TCA precipitation. 15. Take care to avoid transferring parts of the lower phase and the inter phase since they contain a lot of interfering compounds. 16. The drying of the pellet is very important. Take care that the pellet is dry enough (indicated by a slight color change) but not over dry. A pellet that is too dry will often be very difficult to resuspend. It is recommended to dry the pellet under a hood and to follow up the drying process. 17. A recalcitrant (difficult-to-redissolve) pellet can elegantly be forced to redissolve by freezing the solution at −20°C. The formed crystals will break the pellet upon vortexing. Repeated freezing and thawing of the sample must be avoided! 18. A lower temperature of the centrifuge can result in crystallization of urea. 19. Storage in a freezer at −78°C is preferred. Repeated freezing and thawing of the sample must be avoided. Make aliquots of the desired quantity and thaw only once! 20. The quantification of the proteins is very important but challenging. Each quantification method has its limitations since the concentration of a complicated protein mixture is estimated based on the quantification of one

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21.

22.

23.

24. 25. 26.

27.

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reference protein usually BSA. It is therefore important to have sufficient replications and to stay within the optimal linear range of the reference protein. Each sample has to be equally labeled and loaded. The power of the DIGE approach arises from the use of an internal standard, which is a common representative mixture of all analyzed samples. The internal standard is a mixture of an equal amount of all analyzed samples and is labeled with Cy2. This internal standard is used to calculate a standardized abundance of each spot and to match the spots across the gels. In order to anticipate any dye specific effect, the samples should be labeled at random with Cy3 and Cy5. All pre-culture sample points should be randomized over the gels to anticipate gel-specific effects. It is very important that all the strips are covered. Uncovered strips will dry out and urea will start to crystallize. If less than 12 strips are used also the remaining lanes should be filled to prevent crystallization. The oil can move from one lane to the other. Cup loading involves applying the samples at a specific experimentally determined zone of a rehydrated strip. All proteins approach their pI only from the same side. However, the application point of the cup should be carefully chosen. Proteins tend to precipitate at their pI and can lead to aggregation at the sample entry point. Therefore, the placement of the cup is sample specific and depends on the pI range of the IEF-strip. The cup is placed at the cathodal side in case of an acidic zoom strip and at the anodal side in case of a basic zoom strip. It is very important that the cups do not leak. Leakage can be checked by adding a small volume of rehydration buffer if desired. In order to better control the start of the polymerization reaction use acrylamide, water and Tris that is stored at 4°C. Although the gels look polymerized after a few hours, there is still silent polymerization going on. For reproducible gels, it is therefore very important to perform the polymerization overnight and always at the same room temperature. The shelf life of the Laemmli buffered gels is limited and the quality is guaranteed only for a few days.

Acknowledgments

Dr. S.C. Carpentier is supported by a postdoctoral fellowship of the K.U.Leuven. References 1. Wuyts, N., De Waele, D., and Swennen, R. (2006) Extraction and partial characterization of polyphenol oxidase from banana (Musa acuminata Grande naine) roots. Plant Physiol. Biochem. 44, 308–314.

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2. Wuyts, N., De Waele, D., and Swennen, R. (2006) Activity of phenylalanine ammonialyase, peroxidase and polyphenol oxidase in roots of banana (Musa acuminata AAA, cvs Grande Naine and Yangambi km5) before and after infection with Radopholus similis. Nematology 8, 201–209. 3. Gooding, P. S., Bird, C., and Robinson, S. P. (2001) Molecular cloning and characterization of banana fruit polyphenol oxidase. Planta 213, 748–757. 4. Simmonds, N. W. (1966) Bananas. Longmans, London. 5. Loomis, W. D. and Bataille, J. (1966) Plant phenolic compounds and the isolation of plant enzymes. Phytochemistry 5, 423–438. 6. Damerval, C., Zivy, M., Granier, F., and de Vienne, D. (1988) Two-dimensional electrophoresis in plant biology. in Advances in electrophoresis (Chrambach, A., Dunn, M. J., & Radola, B.J., eds.) New York, VCH, pp. 265–280. 7. Granier, F. (1988) Extraction of plant-proteins for two-dimensional electrophoresis. Electrophoresis 9, 712–718. 8. Meyer, Y., Grosset, J., Chartier, Y., and Cleyetmarel, J. C. (1988) Preparation by two-dimensional electrophoresis of proteins for antibody-production - antibodies against proteins whose synthesis is reduced by auxin in tobacco mesophyll protoplasts. Electrophoresis 9, 704–712. 9. O’Farrell, P. H. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–4021. 10. Cremer, F. and Vandewalle, C. (1985) Method for extraction of proteins from green plant-tissues for two-dimensional polyacrylamide-gel electrophoresis. Anal. Biochem. 147, 22–26. 11. Wu, F. S. and Wang, M. Y. (1984) Extraction of proteins for sodium dodecyl-sulfate polyacrylamide-gel electrophoresis from protease-rich plant-tissues. Anal. Biochem. 139, 100–103. 12. 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, 141–143. 13. Hari, V. (1981) A method for the two-dimensional electrophoresis of leaf proteins. Anal. Biochem. 113, 332–335. 14. Flengsrud, R. and Kobro, G. (1989) A method for two-dimensional electrophoresis of proteins from green plant-tissues. Anal. Biochem. 177, 33–36. 15. Damerval, C., Devienne, D., Zivy, M., and Thiellement, H. (1986) Technical improvements in two-dimensional electrophoresis increase the level of geneticvariation detected in Wheat-seedling proteins. Electrophoresis 7, 52–54. 16. Van Etten, J. L., Freer, S. N., and Mccune, B. K. (1979) Presence of a major (storage?) protein in dormant spores of the fungus Botryodiplodia theobromae. J. Bacteriol. 139, 650–652. 17. Englard, S. and Seifter, S. (1990) Precipitation techniques. Methods Enzymol. 182, 285–300. 18. Wu, F. S. and Wang, M. Y. (1984) Extraction of proteins for sodium dodecyl-sulfate polyacrylamide-gel electrophoresis from protease-rich plant-tissues. Anal. Biochem. 139, 100–103.

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19. Nandakumar, M. P., Shen, J., Raman, B., and Marten, M. R. (2003) Solubilization of trichloroacetic acid (TCA) precipitated microbial proteins via NaOH for twodimensional electrophoresis. J. Proteome Res.. 2, 89–93. 20. Unlu, 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. 21. Carpentier, S. C., Witters, E., Laukens, K., Deckers, P., Swennen, R., and Panis, B. (2005) Preparation of protein extracts from recalcitrant plant tissues: An evaluation of different methods for two-dimensional gel electrophoresis analysis. Proteomics 5, 2497–2507. 22. Carpentier, S. C., Dens, K., Van den houwe, I., Swennen, R., and Panis, B. (2007) Lyophilization, a practical way to store and transport tissues prior to protein extraction for 2-DE analysis? Proteomics Supl S, 64–69. 23. Pedreschi, R., Vanstreels, E., Carpentier, S., Hertog, M., Lammertyn, J., Robben, J. et al. (2007) Proteomic analysis of core breakdown disorder in Conference pears (Pyrus communis L.). Proteomics 7, 2083–2099. 24. Carpentier, S. C., Witters, E., Laukens, K., Van Onckelen, H., Swennen, R., and Panis, B. (2007) Banana (Musa spp.) as amodel to study the meristem proteome: acclimation to osmotic stress. Proteomics 7, 92–105. 25. Van Etten, J. L., Freer, S. N., and McCune, B. K. (1979) Presence of a major (storage?) protein in dormant spores of the fungus Botryodiplodia theobromae. J. Bacteriol. 139, 650–652. 26. Schuster, A. M. and Davies, E. (1983) Ribonucleic-acid and protein-metabolism in Pea epicotyls.1. The aging process. Plant Physiol. 73, 809–816. 27. Usuda, H. and Shimogawara, K. (1995) Phosphate deficiency in maize: changes in the 2-dimensional electrophoretic patterns of soluble-proteins from 2nd-leaf blades associated with induced senescence. Plant Cell Physiol. 36, 1149–1155.

14 Preparation of Bacterial Samples for 2-D PAGE Brian B. Vandahl, Gunna Christiansen, and Svend Birkelund 1. Introduction Sample preparation is a very crucial step in two-dimensional (2-D) gel electrophoresis in which the proteins of the sample are brought into a state where they can be separated by isoelectric focusing in the first dimension. The proteins must be denatured, reduced, and solubilized, and they must be kept so during electrophoresis without changing their pI. The buffer for this purpose is traditionally called the lysis buffer. Most bacterial studies aim to solubilize as many proteins as possible to obtain the best possible representation of the total protein content or protein expression under the investigated biological circumstances. However, prefractionation, or the successive application of different chemical reagents, can be used to investigate proteins with certain characteristics. Note that the gels will reflect the proteome of the bacteria at the time the proteins are solubilized and that preceding centrifugations or other manipulations may stress the bacteria, thereby influencing the protein profile. The solubilization procedure is highly dependent on the nature of the sample. Some bacteria are readily lysed by the constituents of the lysis buffer. Others must be disrupted mechanically and, for some, it may be necessary to remove the cell wall by enzymatic digestion prior to mechanical disruption. Subheading 3.1 provides a general protocol for solubilization. All reagents are described in the notes together with common alternatives. Some samples may contain nonprotein substances in amounts that are incompatible with especially first dimensional electrophoresis and, thus, have to be removed from the sample. Salts and most other compounds can be removed by protein precipitation, as described in Subheading 3.2.1. If nucleic acids are present in high amounts, these may have to be removed by enzymatic digestion, which is described in Subheading 3.2.2. From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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In addition, highly abundant proteins may cause problems for 2-D PAGE by preventing optimum focusing in the first dimension or by masking large areas of the gel. In such cases, it may be necessary to carry out prefractionations or to specifically remove the abundant proteins by immunoprecipitation. A novel method of enrichment as a result of low abundant proteins is the ProteoMiner technology from BioRad. By the use of a peptide labrary coupled to column material, the amount of highly abundant proteins is decreased because of binding site saturation and, by passing large volumes of sample over the column, the relative amount of low abundant proteins is simultaneously increased. Prefractionation can be obtained by various means such as chemical extractions or chromatographic or electrophoretic techniques (1), but these methods fall beyond the scope of this chapter. All procedures should be kept as simple as possible to ensure reproducibility and because proteolytic degradation must be considered a risk. When bacteria are disrupted, proteases will be released and degrade the proteins of the sample, if not inhibited. As folded proteins are less susceptible to proteolysis than denatured proteins and as proteases are often more resistant to denaturation than most other proteins, solubilization in urea will often make the problem worse. However, most proteases will be inactivated by disruption in lysis buffer containing thiourea, in 2% SDS or in precipitation solution containing 10% TCA. Still, all handling of the sample after disruption of the bacteria should be carried out as quickly as possible and on ice to minimize proteolytic degradation. It may also be necessary to add protease inhibitors. Several reagents used for the sample preparation are toxic and/or carcinogenic. For safety reasons use protective gloves and glasses and work in a fume hood. 2. Materials 1. Lysis buffer (see Note 1): 7 M Urea (2.10 g) (see Note 2), 2 M thiourea (0.76 g) (see Note 3), 65 mM DTE (650 mL of a 0.5 M stock) (see Note 4), 4% (W/V) CHAPS (0.2 g) (see Note 5), 2% (v/v) Pharmalyte pH 3–10 (see Note 6), 40 mM Tris base (see Note 7), a trace of bromphenol blue (see Note 8). 2. Presolubilization solution: 2% SDS, 65 mM DTE. 3. Precipitation solution: 10% Trichloracetic acid (TCA) in acetone, 20 mM DTE. 4. DNase/RNase solution: 1 mg/mL DNase I, 0.25 mg/mL RNase A, 50 mM MgCl2.

3. Methods 3.1. General Solubilization Protocol It is crucial that all bacteria are disrupted so that the lysis buffer gains access to all proteins. In studies where multiple extractions with different chemical reagents are employed sequentially, it will mask the result if more and more bacteria are disrupted during the procedure. The best method for disruption is dependent on the type of bacteria. In most cases, disruption by sonication will do, but it may

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be necessary to add lysosyme to break down cell walls. It is advantageous to perform the disruption in SDS or in lysis buffer containing thiourea in which most proteases are denatured (2). If prolonged manipulation, such as fractionation by different steps of centrifugation, must be performed, protease inhibitors should be added (see Note 9). In the procedure described below, the bacteria are sonicated in 2% SDS, 65 mM DTE and boiled to enhance the protein solubilization in general (3). It has been suggested (4) that SDS used for presolubilization does not interfere with first dimensional electrophoretic separation because it forms micelles with the nonionic detergent of the lysis buffer and migrates out of the strip. Still, the amount of SDS should be kept low compared to the amount of detergent in the lysis buffer (5) (see Note 10). Figure 1 shows a silver stained gel loaded with 100 mg of Chlamydia pneumoniae protein that was presolubilized by boiling in

Silver stained gel of 100 mg Chlamydia Pneumoniae Protein kDa 100

50

30

15

5.0

6.0

7.0

pI

Fig. 1. A pellet of purified Chlamydia pneumoniae elementary bodies (12) was resuspended in 1% SDS, 50mM Tris-HCl pH 7.0, sonicated and boiled for 5 min. Cooled sample was diluted 1:10 in lysis buffer, sonicated briefly, left at room temperature for one hour and centrifuged at 20.000 x g for 15 minutes. 350 µL of the supernatant containing 100 µg protein was loaded onto a pH 3–10 non-linear immobilized pH gradient strip (Amersham Biosciences) and focused for 120.000 Volt hours. Second dimension was 9–16% linear gradient SDS PAGE. The gel was silver stained.

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1% SDS and 50mM Tris-HCl and then diluting it in the described lysis buffer to a final concentration of 0.1% SDS (see Note 11). The protocol below describes the solubilization of proteins from pelleted bacteria. If proteins have been precipitated, for example, for desalting, the lysis buffer should be added directly (step 8) in the highest possible amount (see Note 12). 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Start out with an appropriate amount of bacteria as pellet (see Note 13). Add four times the pellet volume of 2% SDS, 65mM DTE (see Note 14). Sonicate three times for 2–20 sec depending on sample size (see Note 15). Spin shortly to collect the sample (see Note 16). Resuspend any pellet that may have formed. Boil for five min (see Note 17). Allow the sample to cool Add 8 volumes of lysis buffer to one volume of extract (see Note 18). Sonicate three times for 5 sec, cool between sonications (see Note 19). Leave the sample on a rocking table for 30 min. Spin at 20,000 × g for 15 min and collect the supernatant (see Note 20). Asses the protein concentration (see Note 21). Run first dimension immediately or store the sample at −70°C for several months (see Note 22).

3.2. Sample Purification Common contaminants in 2-D PAGE studies are salts, small ionic compounds, polysaccharides, nucleic acids and lipids. Salt is the most likely reason if bad first dimensional focusing is observed. Enhanced conductivity and water migration in the strip due to high concentrations of salts will cause horizontal streaks. The concentration of salt should preferably be below 10 mM when samples are loaded by strip rehydration. Small charged substances may likewise disturb the isoelectric focusing, especially at the acidic end, as such are often negatively charged. Polysaccharides may clog the gel of the strip and may complex proteins by electrostatic interactions. Lipids may also clog the gel but are mainly a problem due to complexing of hydrophobic proteins and binding of detergent. Nucleic acids may clog the gel, bind proteins through electrostatic interactions, and cause streaking in itself, especially in silver staining. Buffer shift using spin of drip columns, dialysis, and precipitation (Subheading 3.2.1) are straightforward and effective ways to reduce the concentration of salt and small ionic compounds to an acceptable level. Dialysis causes a minimal loss of sample, but requires relative large volumes of solute and is rather time consuming. Spin dialysis using, for instance, Amicon Ultra from Millipore is faster and requires no extra volume of solute, but protein may be lost by adsorption onto the dialysis membrane. Similar issues exist for buffer shifting. Precipitation may also be used to remove polysaccharides and, to some

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extent, lipids. Large polysaccharides can be removed by ultracentrifugation. If lipids are causing major problems, the amount and nature of detergent must be optimized for the particular sample. High amounts of nucleic acids may require treatment with DNase/RNase (Subheading 3.2.2). The presence of proteases is likely to be a problem during sample purification and in such case protease inhibitors must be added (see Note 9). It must be stressed that sample purification preceding addition of lysis buffer should only be carried out if necessary and not as a standard part of the sample preparation. 3.2.1. Precipitation Precipitation is very efficient for removal of most contaminants including salts, but no precipitant will precipitate all proteins, and some proteins will be difficult to resuspend following precipitation. This is a particular problem when a picture of the total protein content is desired. A combination of TCA and acetone is the most common precipitant in 2-D PAGE studies, as it is more effective than either of these reagents alone. Besides, very few proteases are active in 10% TCA. Resolubilization is easier after precipitation with acetone alone (75% final concentration), but precipitation wil not be as efficient as with TCA. The TCA/acetone precipitation described here is essentially as in (6). 1. 2. 3. 4. 5. 6. 7.

Add 10% TCA in ice-cold acetone with 20mM DTE to the sample (see Note 23). Leave at −20°C for two hours (see Note 24). Centrifuge at 10.000 × g for 10 min. Wash with cold acetone containing 20mM DTE. Repeat wash. Let the pellet dry to remove residual acetone. Resuspend pellet in lysis buffer (see Subheading 3.1).

3.2.2. DNase/RNase Treatment If nucleic acids are present in high amounts, the sample will appear viscous and a smear will be seen after silver staining. If ultracentrifugation does not solve the problem, enzymatic digestion will. 1. Add 1/10 of the sample volume of a solution containing 1 mg/mL DNase I, 0.25 mg/mL RNase A and 50 mM MgCl2 (see Note 25). 2. Incubate on ice for 20 min.

4. Notes 1. Absolute amounts are to make 5 mL. All reagents must be analytical grade. Use doubly distilled water. The solution is best mixed in a 10 mL tube on a rotating device. The solution should be made fresh before use or alterna-

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tively frozen in aliquots at −70°C and only thawed once. The solution must not be heated above 37°C. The composition of the lysis buffer is essential for the final result of 2-D PAGE and different lysis buffers may be optimum for different samples and for different proteins in one sample. The function of the lysis buffer described here is to bring as many proteins in the sample as possible into solution and keep them in solution during electrophoresis. As isoelectric focusing is best carried out under denaturing and reducing conditions, the lysis buffer should solubilize, denature, and reduce the proteins of the sample. At the same time, the lysis buffer must not change the pI of the proteins, and it must not be highly conductive. Thus, uncharged components are preferred. Most lysis buffers are still based on that introduced by O’Farrel in 1975 (4) containing urea as denaturing agent, a detergent, a reducing agent and carrier ampholytes. The standard lysis buffer described here is based on (3) and the characteristics of each reagent are described in the following notes. 2. Urea, (NH2)2CO, is a non-charged chaotrope that disrupts non-covalent bonds and thereby denatures proteins. It is the main denaturant in all lysis buffers used in 2-D PAGE and it can be brought into solution in concentrations up to 9.8 M if no thiourea is added. Urea in solution is in equilibrium with ammonium cyanate, which in the form of isocyanic acid will react with amino groups of lysine and arginine residues and the amino terminus of proteins causing carbamylation. The carbamylation of an amine group removes a positive charge from the protein causing a shift toward the acidic side in the gel. Furthermore, it prevents N-terminal sequencing and some enzymatic digests. To avoid carbamylation use only freshly prepared urea solutions. A urea solution should not be left at room temperature for longer periods and should never be heated above 37°C (7). 3. Thiourea, (NH2)2CS, improves the solubilization of especially hydrophobic proteins during first dimension (8) and, in combination with urea, it can be used in concentration up to 2.5 M. The addition of thiourea reduces the solubility of urea and combinations of 7 M urea and 2 M thiourea or 8 M urea and 0.5 M thiourea are most common. The addition of thiourea to the lysis buffer has a pronounced inhibitory effect on proteases, which may still be active in high concentrations of urea alone (2). As thiourea can hinder the binding of SDS to proteins, it should not be included in the buffers used to equilibrate strips prior to second dimension (8). 4. DTE, dithioerythritol, MW: 154.3, make a 0.5 M stock solution and store at −20°C. DTE has the same strong reducing power as dithiothreitol (DTT) and both can be used in concentrations from 10–100 mM. At alkaline pH both DTE and DTT are charged and migrate toward the anode during first dimen-

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sion, which may leave the basic end of the strip without reducing agent and thus cause streaking as a result of reoxidation and precipitation. An alternative and very strong reducing agent is tributyl phosphine (TBP). It can be used in concentrations as low as 2 mM and is noncharged, which means that it keeps all proteins reduced throughout the first dimension thereby enhancing the resolution (9). TBP is stable, but spontaneously inflammable in air. Make a 200mM stock in anhydrous isopropanol and store under nitrogen at 4°C (9). Other alternatives are to alkylate the cysteines prior to first dimension or to oxidize the disulfides by performing the isoelectric focusing in the presence of excess hydroxyethyldisulphide (HED), which is commercially available as DeStreak (GE Healthcare) and includes detailed protocols for its use. 5. CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate, is a zwitterionic detergent that is used in many 2-D PAGE studies. Zwitterionic or nonionic detergents are preferred to anionic detergents like SDS that interferes with the isoelectric focusing in first dimension (see Note 1). The efficiency of many zwitterionic detergents has been investigated and sulfobetaines with a hydrophobic tail of 12-16 alkyl carbons and an empirically determined linker in between have been found to be good alternatives to CHAPS and superior for some samples (10). Which detergent is best for a given sample can still not be predicted. The zwitterionic agent ASB-14 (C22H46N2O4S, Calbiochem) with a 14-carbon alkyl tail, the nonionic Triton X-100 (C14H22O(C2H4O)n with an average number (n) of ethylene oxide of 9 to 10) or the maltoside n-Dodecyl β-D-maltoside (C24H46O11, Sigma-Aldrich) would be good first-choice alternatives if CHAPS does not give satisfactory results (11). 6. Pharmalyte 3–10 can be used for most immobilized pH gradient strips, but if narrow strips or very basic strips are used, carrier ampholytes that match the pH range of the strip should be chosen. Ask the strip supplier if in doubt. Pharmalyte 3–10 is a mixture of carrier ampholytes with pI between 3 and 10. These are small amphoteric compounds with a molecular weight below 1 kDa that have a high buffering capacity at their pI but do not bind proteins due to their high hydrophilicity. When we use 2% v/v of Pharmalyte 3–10 (Amersham Pharmacia) it gives a final concentration of carrier ampholytes of 0.72% in the lysis buffer since “Pharmalyte 3-10” is 36% (w/v). Carrier ampholytes help keep proteins in solution during first dimension and especially prevent hydrophobic interactions between proteins and the IPG in the basic end of the strip. Furthermore, the precipitation of nucleic acids is improved by carrier ampholytes. 7. Tris base is added to raise pH of the lysis buffer to 8.5. Without the addition of base, pH of the lysis buffer would be about 5.5. At alkaline pH more proteins will be anionic and thus not bind to DNA. However, the pH for

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9.

10.

11. 12. 13.

14.

15.

Vandahl, Christiansen, and Birkelund optimal solubilization will vary among samples, and Tris base is left out in many studies. Note that if the sample is to be labbeled before electrophoresis, pH may be improtant for the labelling reaction, which for instance is the case for CyDye labelling in the DIGE (GE Healtcare) approach. Bromphenol blue should be added in a small amount to color the solution lightly blue. The color will move toward the anode during first dimension, which can be used to check that the isoelectric focusing is ongoing. However, the color will disappear before the first dimension is finished and cannot be used as an indicative of when to stop. Protease inhibitor cocktails are available from several commercial laboratory reagent suppliers but most proteases will be inhibited by adding 2 mM EDTA, 1 mM PMSF, 1 mM Pepstatin A, and 13 mM Bestatin. EDTA chelates free metal ions, thereby inhibiting metalloproteases, make a 0.5 M stock solution in water, pH 8.0. PMSF (phenylmethylsulfonyl fluoride), inhibits serine proteases and some cysteine proteases, make a 100 mM stock solution in methanol. Pepstatin A inhibits aspartic proteases, make a 1 mM stock stock solution in methanol. Bestatin inhibits aminopeptidases, make a 13 mM stock solution in water. When NP-40 is used as detergent in the lysis buffer it has been reported (5) that the ratio of NP-40 to SDS should be at least 8 to avoid streaking. NP-40 (Nonidet P-40) is a nonionic detergent that is very similar to Triton X-100 and the properties are often reported as being identical. NP-40 (Roche) is C15H24O(C2H4O)n, where n = 9–10 on average. No reducing agent was added during presolubilization. If the sample is applied by strip rehydration of 18 cm strips, the maximum amount is 350 mL per strip. For most bacteria 25–100 mg of protein is appropriate for silver staining, 100–150 mg for Western blotting, and 0.5–2 mg for preparative gels, when 18 cm immobilized pH gradient strips in the pH range of 3–10 or similar broad range intervals are used. Be sure not to add more SDS than it can be diluted to 0.25% in lysis buffer. If all the sample is to be used for one gel using 350 mL lysis buffer to rehydrate an immobilized pH gradient strip, this means that no more than 40 mL of 2% SDS should be added. However, if streaking is observed, try lowering the amount of SDS used for presolubilization. For some samples pH must be buffered to optimize solubilization. Use for instance 50–100 mM Tris-HCl, pH 7.0. The optimal pH may vary from sample to sample. Be aware that the final concentration of salt should not exceed 10 mM when samples are loaded by strip rehydration (see Subheading 3.2). Adjust the amplitude of the sonicator so that microbubbles are formed, and keep the tip of the probe deep in the sample to avoid too much foam formation. The sample should be cooled between sonications.

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16. It cannot be avoided that some foam is formed during sonication and this should be spun down before the sample is boiled, in order to avoid protein coagulation in drying bubbles. 17. DTE develops toxic gas upon heating. Boil in a fume hood. 18. Dilute the sample as much as possible in lysis buffer (see Notes 13 and 14). 19. When the sample is in lysis buffer containing urea it is important to keep the temperature below 37°C in order to avoid carbamylation of proteins. If boiling in SDS is left out, the sonication may have to be extended. 20. The centrifugation step is important to remove cell debris and precipitated DNA and it should not be left out. 21. Several constituents of the lysis buffer may cause problems for assessment of protein concentration. Carrier ampholytes, CHAPS, and other detergents will bind most dyes and reduction of cupric ion cannot be employed in the presence of thiourea and DTE. Thus, the protein must be selectively precipitated and then measured. This may be done with the 2-D Quant Kit form Amersham Pharmacia. Alternatively, the protein may be estimated in a parallel sample which is not solubilized in lysis buffer. This may not give the actual protein concentration in the lysis buffer but will in most cases provide adequate information to determine the load. 22. Repeated freeze thaw cycles should be avoided due to the risk of carbamylation and because the solubility of some proteins may be changed by the process. 23. The bacteria must be disrupted beforehand by an appropriate method (see Subheading 3.1). If the bacteria are disrupted directly in the precipitation buffer most proteases will be inactivated by the TCA. 24. The precipitation time should not be longer than the minimal time required for satisfactory precipitation. For some samples 15 min will do while others must be incubated overnight. Be aware that prolonged exposure to the very acidic solution may cause protein degradation. 25. The bacteria must be disrupted beforehand by an appropriate method (see Subheading 3.1). If active proteases are present it may be necessary to add protease inhibitors (see Note 9) even if the DNase/RNase treatment is carried out on ice. References 1. Righetti, P. G., Castagna, A., Herbert, B., Reymond, F., and Rossier, J. S. (2003) Prefractionation techniques in proteome analysis. Proteomics 3, 1397–1407. 2. Castellanos-Serra, L., and Paz-Lago, D. (2002) Inhibition of unwanted proteolysis during sample preparation: evaluation of its efficiency in challenge experiments. Electrophoresis 23, 1745–1753.

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3. Harder, A., Wildgruber, R., Nawrocki, A., Fey, S. J., Larsen, P. M., and Gorg, A. (1999) Comparison of yeast cell protein solubilization procedures for two-dimensional electrophoresis. Electrophoresis 20, 826–829. 4. O’Farrell, P. H. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol Chem. 250, 4007–4021. 5. Ames, G. F. and Nikaido, K. (1976) Two-dimensional gel electrophoresis of membrane proteins. Biochemistry 15, 616–623. 6. Jacobs, D. I., van Rijssen, M. S., van der Heijden, R., and Verpoorte, R. (2001) Sequential solubilization of proteins precipitated with trichloroacetic acid in acetone from cultured Catharanthus roseus cells yields 52% more spots after two-dimensional electrophoresis. Proteomics 1, 1345–1350. 7. McCarthy, J., Hopwood, F., Oxley, D., Laver, M., Castagna, A., Righetti, P. G., Williams, K., and Herbert, B. (2003) Carbamylation of proteins in 2-D electrophoresis–myth or reality? J Proteome Res. 2, 239–242. 8. Rabilloud, T., Adessi, C., Giraudel, A., and Lunardi, J. (1997) Improvement of the solubilization of proteins in two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 18, 307–316. 9. Herbert, B. R., Molloy, M. P., Gooley, A. A., Walsh, B. J., Bryson, W. G., and Williams, K. L. (1998) Improved protein solubility in two-dimensional electrophoresis using tributyl phosphine as reducing agent. Electrophoresis 19, 845–851. 10. Tastet, C., Charmont, S., Chevallet, M., Luche, S., and Rabilloud, T. (2003) Structure-efficiency relationships of zwitterionic detergents as protein solubilizers in two-dimensional electrophoresis. Proteomics 3, 111–121. 11. Luche, S., Santoni, V., and Rabilloud, T. (2003) Evaluation of nonionic and zwitterionic detergents as membrane protein solubilizers in two-dimensional electrophoresis. Proteomics 3, 249–253. 12. Vandahl, B. B., Birkelund, S., and Christiansen, G. (2002) Proteome analysis of Chlamydia pneumoniae. Meth. Enzymol. 358, 277–288.

15 Preparation of Bodily Fluids for 2-D PAGE Anthony G. Sullivan, Henry Brzeski, Jeban Ganesalingam, and Manuel Mayr 1. Introduction The importance of bodily fluids as a source of clinically relevant biomarkers/ surrogate markers of human disease has increased significantly over the last decade (1,2), and modern proteomic methods have evolved and been adapted to meet the demand. The specific challenges facing serum analysis include the wide dynamic range in the concentration of individual components and the tremendous number of potential variants of glycosylated proteins (3). The most dominant plasma proteins, albumin and immunoglobulin (Ig)G, typically comprise up to 70% of the plasma proteome in abundance. To enable the majority of the remaining, far less abundant proteins to be better visualized by two-dimensional gel electrophoresis (2-DE), these two proteins must first be removed or, at least, depleted in relative concentration. There are a number of currently available commercial products from a range of suppliers that enable albumin depletion by chemical affinity, exploiting the remarkable albumin-binding ability of structures closely related to the reactive dye molecule Cibacron blue 3GA (4), and the IgG binding properties of protein G (5). The blue dye has been shown to have a special affinity for proteins containing the dinucleotide fold, a structural feature that is common to several classes of proteins (4). Albumin can be separated from other plasma proteins using lectin affinity, as it is not normally glycosylated, while the majority of classical plasma proteins are. This approach allows both enrichment of lower-abundance proteins, and the study of differences in glycoprotein profiles (6). Highly effective depletion of albumin using monoclonal antibody selection has also been demonstrated, and coupled with protein G/IgG depletion (7).Recent research has identified 325 distinct proteins from 1800 2-D gel protein features following multi-component immunoaffinity extraction and further comprehensive chromatographic fractionation (8). Depletion of IgG by From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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a protein G resin can also be coupled with NaCl/Ethanol precipitation to deplete albumin (9). However, depletion can cause problems in itself. The concentration of the high abundant proteins varies considerably within bodily fluids. Therefore once the samples are depleted, there is a significant change in the relative concentration of the protein constituents from the original sample. As an example in our own studies the percentage protein remaining after depletion in cerebrospinal fluid has an average of 17.9% with a standard deviation of 6.6%. This could be overcome by loading according to sample volumes rather than protein loading, which is often the norm in proteomic experiments (10). A major drawback is that the depletion not only adds further steps in sample processing, but undoubtedly alters the protein constituents within the sample as high abundant proteins bind other proteins. Consequently, the depletion may complicate quantitative comparisons and result in the loss of potential biomarkers (11). 2. Materials 1. Blue dye affinity resin: Affi-gel, (Biorad), Mimetic Blue SA (Prometic). 2. 20 mM Na2HPO4, pH adjusted to 7.0 with HCl. 3. Lysis buffer: 7 M urea, 2 M thiourea, 30 mM Tris, 5 mM magnesium acetate, 4% CHAPS, 1% NP-40. 4. Trichloroacetic acid (TCA)/acetone protein precipitation: 2-D Cleanup Kit (Amersham Biosciences, Piscataway, NJ). 5. Biomax 10K NMWL membrane centrifugal filter (Millipore, Bedford, MA). 6. WGA-agglutinin lectin (Sigma, St. Louis, MO). 7. Handee Mini Spin Columns (Pierce Biotechnology, Inc., Rockford, IL) or Macro Spin Columns (NEST Group, Southborough, MA). 8. Phosphate buffer: 50 mM Na2HPO4, 0.2 M NaCl, pH adjusted to 7.0 with HCl. 9. Phosphate buffer with SDS: 50 mM Na2HPO4, 0.2 M NaCl, 0.05% SDS, pH adjusted to 7.0 with HCl. 10. Sugar solution: 0.3 M N-acetyl glucosamine or neuraminic acid (Sigma, St. Louis, MO), in phosphate buffer. 11. Resin slurry: Albumin and IgG Removal Kit (Amersham Biosciences, Piscataway, NJ). 12. POROS Affinity depletion cartridges (Applied Biosystems, Framingham, MA). 13. AccuGENE 10X PBS solution (Cambrex, East Rutherford, NJ).

3. Methods 3.1. Affinity Depletion of Serum Albumin 3.1.1. Batch Use of Dye-Agarose Affinity Resins Here we describe a generic method, based on manufacturer’s instructions, adaptable for the majority of available dye resin slurries. The steps involved are equilibration, binding, washing, and stripping.

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1. Add 250 µL of resin (see Note 1) to a 1.5-mL microcentrifuge tube (see Note 2), centrifuge for 2 min at 0.2 g, and remove liquid using a gel-loading pipet tip. 2. Add 200 µL of 20 mM sodium phosphate, pH 7.0, and shake gently for 10 min. Centrifuge at 1000 rpm for 2 min and remove liquid. Repeat the equilibration twice. 3. Take 25 µL of clarified plasma/serum, dilute it with 175 µL of the 20 mM sodium phosphate, pH 7.0 buffer, and mix thoroughly. 4. Add the diluted plasma to the conditioned resin, vortex briefly, and shake gently for 10 min to allow serum albumin to bind to the dye resin. Centrifuge for 2 min at 1000 rpm and remove supernatant using a gel-loading pipet tip. Transfer solution, containing unbound albumin-depleted proteins, to a clean 1.5-mL microcentrifuge tube. 5. Wash the resin three times with 150 µL 20 mM sodium phosphate, pH 7.0 buffer, shake for 10 min, centrifuge for 2 min at 1000 rpm, remove the supernatant, and combine with the previous solution. 6. Elute the proteins bound to the resin by washing three times with lysis buffer (see Note 3). Combine stripped fractions into a separate 1.5-mL tube. 7. Concentrate the albumin-depleted fraction (approx vol 650 µL) to a final vol of approx 100 µL using a 10,000 MWCO (molecular weight cut-off) membrane filter. Desalt and prepare for 2-DE by TCA/acetone precipitation (see Note 4). For the albumin-rich fraction, remove a 150-µL aliquot from the stock solution and perform TCA/acetone precipitation directly.

3.1.2. Lectin Affinity Serum Albumin Removal Using WGA/Agarose 1. Prepare the Handee Mini Spin column for use by ensuring the frit at the bottom of the main chamber is firmly in place by pushing down with a paper clip. Resuspend the lectin in the buffer supplied by the manufacturer to give a homogeneous lectin slurry, then pipet 200 µL into the chamber, centrifuge briefly, and remove liquid. 2. Add 500 µL of phosphate buffer and shake gently for 10 min. Centrifuge at 1000 rpm for 2 min, remove liquid, and repeat twice. 3. Prepare plasma by diluting 50 µL of plasma with 750 µL of phosphate buffer containing 0.05% SDS. This amount will typically yield 50 µg of albumin-free protein. 4. Transfer diluted plasma to the column containing the beads. Shake gently using a rotary mixer, ensuring good mixing, for 5 min. Centrifuge briefly and collect eluent. Wash beads three times with 200 µL of phosphate buffer. Combine wash eluents (albumin-rich fraction). 5. Elute the glycosylated proteins with 200 µL sugar solution see Note 6), shaking for 5 min on a rotary shaker. Wash three times with 150 µL of sugar solution and combine, total volume approx 650 µL. 6. Reduce the albumin-depleted fraction to a final vol of approx 100 µL using a 10,000 MWCO membrane filter. Desalt and prepare for 2-DE by TCA/acetone

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Fig. 1. An example of a two-dimensional gel showing serum prepared using the method outlined in Subheading 3.1.2.

precipitation (see Note 4). An example of a 2-D gel showing serum prepared by the method in Subheading 3.1.2. is shown in Fig. 1.

3.1.3 Antibody Serum Albumin/Immunoglobulin G Removal 3.1.3.1. USE OF ANTIBODY/PROTEIN G RESIN 1. Pipet 20 µL of plasma/serum into a 1.5-mL microcentrifuge tube and add 750 µL of resin slurry. Close cap and mix on a rotary shaker, ensuring strong mixing is taking place, for 30 min at room temperature. 2. Transfer entire slurry to the upper chamber of a microspin column and insert column into a suitable microcentrifuge tube. Centrifuge at 1000 rpm for 5 min. Remove liquid (approx 500 µL) and desalt/concentrate as described in Subheading 3.1.1., step 7. 3.1.3.2. USE OF ANTIBODY/PROTEIN G CARTRIDGES (SEE NOTE 7) 1. Connect a syringe to the anti-SA cartridge using a needle-port adapter and equilibrate packing material with at least five cartridge volumes of PBS solution, pH 7.4. 2. Dilute the serum/plasma 10-fold with PBS solution and inject an amount less than the maximum binding capacity into the cartridge at a steady flow rate (see Note 8). 3. Collect the flowthrough in a clean 0.5-mL microcentrifuge tube. Wash cartridge with three column vols of PBS solution and combine wash fractions in a separate tube.

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4. Connect the needle-port adapter to the protein G cartridge and condition the packing material with at least five cartridge vols of PBS solution, pH 7.4. 5. Inject the albumin-depleted fraction from step 3 at a steady flow rate and collect the flowthrough in a fresh 1.5-mL tube. 6. Inject the combined wash fractions from step 3 at a steady flow rate and combine with liquid from step 6. 7. Concentrate the albumin-IgG-depleted fraction to a final volume of approx 100 µL using a 10,000 MWCO membrane filter. Desalt and prepare for 2-DE by TCA/ acetone precipitation (see Note 4).

3.2. Use of Commercial Depletion Kits 3.2.1 Commercial Kits Available There are many different commercial depletion kits available which include Aurum Serum Protein Minikit (Bio-Rad, Hercules, CA), Multiple Affinity Removal Column (Agilent Technologies, San Diego, CA), POROS Affinity depletion Cartridges (Applied Biosystems, Framingham, MA) and Albumin and IgG Removal kit (Amersham Biosciences, Uppsala, Sweden). The relative performance of these columns has been compared by Bjorhall et al. (12). 3.2.2 Protocol Using the Agilent Multiaffinity Depletion Column 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Add an appropriate volume of bodily fluid to a 4 ml spin column. Centrifuge sample at 4000 rpm for 30 min.. Add 1 ml of buffer A (Agilent Technologies). Centrifuge sample at 4000 rpm for 30 min. Repeat steps 3 and 4 a further two times. Filter CSF solution through a 0.22 micron spin filter. Centrifuge sample through the column at 100 g for 1.5 min and collect elute. Add 400 mcl Buffer A and centrifuge through column at 100 g for 2.5 min and collect elute. Repeat step 8. Keep total elute (1 ml) which contains the depleted sample. Elute bound fraction by passing 2 ml Buffer B, (Agilent Techonologies) slowly through column. Buffer exchange samples into an appropriate buffer for downstream analysis. Representative 2-D gel images of cerebrospinal fluid, before and after depletion with the above described column, are shown in Fig. 2.

4. Notes 1. The amount of resin used here ensures that the total albumin loaded to the resin is less than its albumin-binding capacity (typically approx 11 mg/mL). 2. As an alternative to the procedure listed here, the resin may be applied to the upper chamber of a spin column, and the removal of liquid facilitated

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B

Albumin

Transferrin

Alpha 1 antitrypsin

Haptoglobulin

Ig Heavy chains

Ig Light Chains

Fig. 2. (continued)

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C

Fig. 2. 2-D gel images of cerebrospinal fluid before and after antibody depletion. Depletion was carried out using commerical depletion columns (Agilent Technologies). Representative images of the original sample (A), the bound fraction (B), and the depleted fraction (C).

3. 4.

5. 6. 7.

by centrifugation. In our hands, this variation led to less efficient albumin removal due to inconsistent residence times. Alternatives to lysis buffer include solutions of sodium chloride or guandinium chloride (5–6 M). Desalting/concentrating is absolutely required for the isoelectric focusing step. The use of MWCO membrane filters followed by protein precipitation was found in our hands to be the most effective method combination. The initial preconcentration to volumes of 100 µL depletes salt levels and reduces the amount of protein precipitation solutions required. Columns are available in several sizes and therefore total protein loading capacities. N-acetyl glucosamine can be replaced by N-acetyl neuraminic acid depending on the type of glycosylation under investigation. These cartridges can be used separately as outlined here, or in series, either manually with the aid of a syringe, or automatically using a solvent delivery system on a liquid chromatograph.

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8. The albumin-binding capacity depends on the column size and will be supplied by the manufacturer. The albumin content of the plasma can only be estimated, unless specific albumin assay methods are available. References 1. Steel, L. F., Trotter, M. G., Nakajima, P. B., Mattu, T. S., Gonye, G., and Block, T. (2003) Efficient and specific removal of albumin from human serum samples. Mol. Cell Proteomics 2, 262–270. 2. Tabar, L., Dean, P. B., Kaufman C. S., Duffy, S. W., and Chen, H. H. (2000) A new era in the diagnosis of breast cancer. Surg. Oncol. Clin. N. Am. 9, 233–277. 3. Anderson, N. L. and Anderson, N. G. (2002) The human plasma proteome, history character and diagnostic prospects. Mol. Cell Proteomics 1, 845–867. 4. Thompson, S. T., Cass, K. H., and Stellwagen, E. (1975) Blue dextran-sepharose: an affinity column for the dinucleotide fold. Proc. Nat. Acad. Sci. USA 72, 669–672. 5. Reis, K. J., Ayoub, E. M., and Boyle, M. D. P. (1984) Streptococcal Fc receptors. II. Comparison of the reactivity of a receptor from a group C streptococcus with staphylococcal protein A. J. Immunol. 132, 3098–3102. 6. Brzeski, H., Katenhusen, R. A., Sullivan, A. G., et al. (2003) Albumin depletion method for improved plasma glycoprotein analysis by 2-dimensional difference gel electrophoresis. Biotechniques 35, 1128–1132. 7. Steel, L. F., Trotter, M. G., Nakajima, P. B., Mattu, T. S., Gonye, G., and Block, T. (2003) Efficient and specific removal of albumin from human serum samples. Mol. Cell Proteomics 2, 262–270. 8. Pieper, R., Gatlin, C. L., Makusky A. J., et al. (2003) The human serum proteome: display of nearly 3700 chromatographically separated protein spots on two-dimensional electrophoresis gels and identification of 325 distinct proteins. Proteomics 3, 1345–1364. 9. Fu, Q., Garnham, C.P., Elliott, S.T., Bovenkamp, D.E., and Van Eyk, J.E. (2005) A robust, streamlined, and reproducible method for proteomic analysis of serum by delipidation, albumin and IgG depletion, and two-dimensional gel electrophoresis. Proteomics 5(10), 2656–2664. 10. Huang, J.T.J, McKenna, T., Hughes, C., Markus Leweke F., Schwarz, E., and Bahn, S. (2007) CSF biomarker discovery using label-free nano-LC-MAS based proteomic profiling:Technical aspects. J. Separation Sci. 30, 214–225. 11. Hye, A., Lynham, S., Thambisetty, M., Causevic, M., Campbell, J., Byers, H.L., Hooper, C., Rijsdijk, F., Tabrizi, S.J., Banner, S., Shaw, C.E., Foy, C., Poppe, M., Archer, N., Hamilton, G., Powell, J., Brown, R.G., Sham, P., Ward, M., and Lovestone, S. (2006) Proteome-based plasma biomarkers for Alzheimers disease. Brain 1(29), 3042–3050. 12. Bjorhall, K., Miliotis, T., and Davidsson, P. (2005) Comparison of different depletion strategies for improved resolution in proteomic analysis of human serum samples. Proteomics 5, 307–317.

16 Immunoaffinity Depletion of High Abundance Plasma and Serum Proteins Lynn A. Echan and David W. Speicher

1. Introduction Analysis of human serum and plasma has been a major focus of many recent proteomics studies because blood is a promising, routinely collected biological fluid that is likely to contain multiple novel biomarkers for many human diseases, such as cancers and cardiovascular disease (1). Most cells in the body are thought to secrete or shed proteins into the blood, and therefore plasma is very likely to contain information concerning the physiological status of all tissues and organs in the body (2–4). While serum and plasma hold great potential as sources of new disease biomarkers, systematic discovery of novel biomarkers in patient cohorts is greatly complicated by the presence of a small number of highly abundant (mg/ml range) proteins in blood (2–6). In particular, serum albumin constitutes about 65% of total serum protein, while immunoglobulins contribute an additional 15%. Most of the remaining 20% of total protein content is comprised of a modest number of additional high- and medium-abundance (µg/ml range) proteins, which tend to obstruct discovery of novel, specific disease biomarkers present at ng/ml to pg/ml levels. Until approximately 2003, the only available technology for depletion of major proteins was represented by commercial kits that combined Cibacron blue dye for removal of albumin with Protein A or G for removal of immunoglobulins. Unfortunately, these columns incompletely removed albumin and removed many non-targeted proteins, including potential biomarkers (7). Recently, several companies have produced immunoaffinity depletion resins that are more effective and more specific than the blue dye approach for depleting multiple blood proteins. Depletion of at least six or more abundant proteins has become standard practice as the first step in multidimensional From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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plasma or serum proteome analyses. Early research studies explored use of monoclonal or polyclonal antibodies to attempt to remove one or multiple abundant proteins. The development (8) and commercial production of the Multiple Affinity Removal System (Agilent) in 2003 was particularly noteworthy as it was the first widely available product that removed more than one or two proteins. Specifically, it removed six abundant proteins and reduced the protein content by 85%. This improves protein profiling capacities substantially, but the next level of abundant proteins continue to limit volumes of plasma that can be analyzed. Hence, ideally additional abundant proteins should be depleted so that at least 98–99% of total protein is removed (4,7). More recently, several manufacturers have produced antibody columns against 7, 12, 14, or 20 abundant blood proteins in both HPLC/FPLC and spin column formats. In addition, most manufacturers will produce custom larger scale FPLC or spin column formats to fit the needs of individual laboratories. The choice of FPLC or spin column format depends upon several factors. First, FPLC columns require relatively expensive instrumentation not available in every laboratory, and antibody LC columns are usually larger and more expensive than spin columns. However, the FPLC columns are highly robust, more completely remove the targeted proteins, and are more time efficient because they usually deplete larger volumes of plasma or serum in a single pass. In our experience, the available commercial multiprotein immunodepletion columns (Table 1) perform similarly. That is, they are highly efficient at depleting the targeted proteins, typically >98–99.5% removal of most targets, while simultaneously not removing significant amounts of untargeted minor proteins (7). Of course, if serum albumin is present at 40 mg/ml and 99% is removed, the residual 0.4 mg/ml of albumin is still a major protein. Hence, for the most critical experiments, it is necessary to optimize removal to greater than 99%, at least for the most abundant proteins. In this regard, it is not surprising that polyclonal antibodies are usually used for these depletion matrices rather than monoclonal antibodies. Monoclonal antibodies are sometimes more specific; each antibody has a single binding affinity, and it is easier to consistently produce a uniform product. But because they have single epitopes, they will usually be less effective than polyclonal antibodies in removing proteolytic fragments, some posttranslationally modified forms of the target protein, oxidized forms of the protein, etc. An unknown challenge is whether these commercial products can be consistently produced over the long term, since polyclonal antibody properties can vary from batch-to-batch due to variations in host immune response to injected antigens. Factors that may influence decisions concerning the type of column to use include: FPLC or spin format, column size and capacity options, moderate

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Table 1 Commercially Available Multiprotein Immunoaffinity Depletion Columnsa Manufacturer/Product Sigma ProteoPrep 20 (Human) Agilent MARS Human-14

# Proteins Depleted 20 14

# Runsb

Format

Capacity

10 ml L C Spin Column 4.6 × 50 mm LC 4.6 × 100 mm LC 10 × 100 mm LC Std. Spin Column

100 µl 8 µl

100

20–40 µl 20–40 µl 160 µl 8–10 µl

200

MARS Human-7

7

4.6 × 50 mm LC 4.6 × 100 mm LC 10 × 100 mm LC Std. Spin Column

30–70 µl 30–70 µl 250–300 µl 12–14 µl

200

MARS Human-6

6

4.6 × 50 mm LC 4.6 × 100 mm LC 10 × 100 mm LC Std. Spin Column HCc Spin Column

15–80 µl 15–80 µl 340 µl 7–10 µl 14–16 µl

200

MARS Mouse-3

3

4.6 × 50 mm LC 4.6 × 100 mm LC Std. Spin Column

37–100 µl 37–100 µl 25–30 µl

200

12

2 ml L C 10 ml LC Spin Column

50 µl 250 µl 20 µl

100

7

2 ml L C 10 ml LC Spin Column

20–40 µl 100–200 µl 10–15 µl

100

Beckman IgY- 12 (Human) IgY-R7 (Mouse/Rat) a

Only products that deplete at least six abundant human proteins and the only available mouse columns are summarized here. Some columns only deplete one or two proteins. b Stated maximum lifetime as reported by manufacturer. c High capacity

price-to-capacity differences, whether buffers are defined or proprietary, and most importantly, the proportion of the total plasma protein removed by the column. As a general rule, it is beneficial to deplete as many abundant proteins as possible, because the greater the reduction in total protein content per volume of original plasma, the greater the improvement in detection of lower abundance proteins. The following protocols describe common HPLC/FPLC and spin

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column immunoaffinity depletion using ProteoPrep 20 columns, which currently remove the largest number of proteins and remove approximately 95% of the total plasma protein content. The same methods can be used for either plasma or serum. For simplicity, we will generally refer to plasma only when describing the protocols. 2. Materials 2.1. FPLC Affinity Depletion of Plasma or Serum 1. Human plasma or serum. 2. HPLC or FPLC system capable of operating at low pressure. 3. ProteoPrep 20 LC Plasma Immunodepletion Column Kit (Sigma Catalog No. PROT20LC). Storage: 2–8 °C. Individual components are: a. ProteoPrep 20 LC Plasma Immunodepletion Column; 9.4 ml resin, 100 µl plasma (50–70 mg) capacity (Sigma Catalog No. P9874). b. ProteoPrep 20 Equilibration Buffer, 10x concentrate; (Sigma Catalog No. P1749). c. ProteoPrep 20 Elution Solution, 10x concentrate; (Sigma Catalog No. P9749). d. ProteoPrep Preservative Concentrate; (Sigma Catalog No. K3889). 4. Stericup GP Express Plus Membrane, 0.22 µm filter units (500 ml capacity); (Millipore Catalog No. SCGPU05RE). 5. Amicon Ultrafree-MC 0.22 µm microcentrifuge filters; (Millipore Catalog No. UFC30GVNB). 6. Milli-Q water or equivalent.

2.2. Spin Column Affinity Depletion of Plasma or Serum 1. Human plasma or serum. 2. ProteoPrep 20 Plasma Immunodepletion Kit (3 column kit: Sigma Catalog No. PROT20; Single column kit: Sigma Catalog No. PROT20S); Storage: 2–8 °C, Individual components are: a. ProteoPrep 20 Plasma Immunodepletion Spin Column (300 µl resin, 8 µl plasma capacity). b. ProteoPrep 20 Equilibration Buffer, 10× concentrate (Sigma Catalog No. P1749). c. ProteoPrep 20 Elution Solution, 10× concentrate (Sigma Catalog No. P9749). d. ProteoPrep Preservative Concentrate (Sigma Catalog No. K3889). 3. Luer Lock caps. 4. Several 5 or 10 ml syringes. 5. Amicon Ultrafree-MC 0.22 µm microcentrifuge filters (Millipore Catalog No. UFC30GVNB). 6. Amicon Ultra-4, 5 kDa MWCO centrifugal filter device (15 ml capacity) (Millipore Catalog No.UFC800524). 7. Milli-Q water or equivalent.

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2.3. Large-Scale Spin Column Affinity Depletion of Plasma or Serum 1. Human plasma or serum. 2. ProteoPrep 20 Custom Plasma Immunodepletion Kit; Storage: 2–8 °C, Individual components are: a. ProteoPrep 20 Jumbo Plasma Immunodepletion Spin Column (3.7 µl resin, 100 µl plasma capacity). b. ProteoPrep 20 Equilibration Buffer, 10x concentrate (Sigma Catalog No. P1749). c. ProteoPrep 20 Elution Solution, 10x concentrate (Sigma Catalog No. P9749). d. ProteoPrep Preservative Concentrate (Sigma Catalog No. K3889). 3. Luer Lock caps. 4. Several 20 ml syringes. 5. Amicon Ultrafree-CL 0.22 µm microcentrifuge filters, 2 ml capacity (Millipore Catalog No.UFC40GV00). 6. Amicon Ultra-4, 5 kDa MWCO centrifugal filter device (Millipore Catalog No.UFC800524). 7. Milli-Q water or equivalent.

2.4. Ethanol Precipitation 1. Unbound or bound fraction from Prot-20LC or spin column depletion. 2. 200 proof Ethanol, chilled to −20°C (Sigma). 3. Speed-vac centrifuge.

3. Methods 3.1. FPLC Affinity Depletion of Plasma or Serum Many HPLC systems require pressures greater than 100 psi to operate properly. The ProteoPrep 20 LC column packing has a rated pressure maximum of 30 psi. Hence, an FPLC-type system, capable of operating at low pressures and where a high pressure limit of 30 psi can be accurately set, is strongly preferred. If the available instrument requires higher minimum operating pressures, one option is to generate necessary backpressure in front of the column with a flow restrictor such as an appropriate length of small internal diameter PEEK tubing (e.g. 1–2 feet, 0.005 or 0.007 inch i.d.) and larger ID PEEK tubing (0.030 or 0.040 inch i.d.) after the column to avoid exposing the column to high pressures. Care must be taken to set the maximum pressure limit only slightly above the operating pressure to reduce the risk of compressing the column bed. Fortunately, the column itself has a much higher pressure rating than the column matrix. Hence, if an obstruction in the column or downstream does occur, there is minimal risk of a column explosion as long as the maximum pressure is set well below the pressure limit of the column housing.

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1. Prepare Buffer A (1x Equilibration Buffer): Dilute one part of ProteoPrep 20 Equilibration Buffer, 10x concentrate with nine parts of room temperature ultrapure water. Filter the diluted buffer through a 0.22 µm filter unit and degas for 5 min. at room temperature using vacuum filtration with stirring. The final volume needed for each depletion is approximately 55 ml. 2. Prepare Buffer B (1x Elution Solution): Dilute one part of ProteoPrep 20 Elution Solution, 10x concentrate with nine parts of room temperature ultrapure water. Filter the diluted buffer through a 0.22 µm filter and degas for 5 min. at room temperature using vacuum filtration with stirring. The final volume needed for each depletion is approximately 30 ml. 3. Before connecting the column, flush Buffer A through the system to the column inlet line for 15 min at 1 to 3 ml/min. Transfer column to room temperature at this time. 4. Connecting column: to prevent air from being introduced into the column, after flushing the system as described in step 3, stop flow, remove the plug from the top of the column only, start the pump at 0.5 ml/min, and while holding the column inlet tubing over the top end of the column, fill the column end fitting with Equilibration Buffer. Stop flow and connect tubing to column inlet. 5. Remove the screw plug from the bottom of the column, connect the detector inlet solvent line to the column bottom, and equilibrate with Buffer A for 5 min at 1 ml/ min (see Note 1). 6. Filter undiluted plasma (generally 100 µl) with a prerinsed 0.22 µm microcentrifuge filter (see Note 2). Keep filtered sample on ice until ready for use (see Note 3). 7. Prior to injection of the first sample of the day, run a blank run (no injection) and reequilibrate the column using Buffer A (see Note 4). 8. Inject 100 µl of filtered plasma at 0.3 ml/min. 9. Elute the depleted plasma (unbound fraction) from the column with Buffer A at 0.3 ml/ min. Collect the flow-through fractions and pool the unbound fractions containing protein as it comes off the column (see Note 5). Keep the pooled sample on ice. 10. Elute bound proteins with Buffer B at 1 ml/min. Collect and pool bound proteins and store on ice (see Note 6). 11. Reequilibrate column with Buffer A for 10 min at 3 ml/min. 12. Inject the next sample and repeat steps 8 to 11 until all samples are separated or a sufficient volume of a single sample is depleted. 13. Disconnect and store the column at 4°C in Buffer A. (see Note 7). 14. Replace Buffers A and B with ultrapure water and flush system, including detector, for 30 min at 1 ml/min followed by 50% isopropanol for 30 min at 1 ml/min (see Note 8).

3.2. Spin Column Affinity Depletion of Plasma or Serum There are several strategies for spin-column usage. First, to maximize column load, it is possible to moderately overload the column, which results in depletion of about 90–95% of the targeted proteins. The unbound fraction is then repurified on either the same column or a duplicate column to “polish”

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the purification. The repurification step can be conducted on a single unbound fraction after it has been concentrated, or multiple unbound fractions can be pooled, concentrated and repurified in a single repurification run. This twopass method has the advantage of being able to deplete larger final volumes of a single sample per day. An alternative strategy is to load the spin column at or slightly below the recommended capacity and conduct one or more single-pass purifications. If the plasma concentration is not too high (column capacities are rated by vendors based on volume, not protein concentration), the final sample or pool of multiple samples may not need to be redepleted. However, they will usually need to be concentrated and one is likely to only achieve 95% depletion of targeted proteins using this method. A third and most conservative strategy is to both underload the column and redeplete the initial unbound fraction in a final polishing step. This should provide at least 99% removal of most targeted proteins, but volume of plasma purified per day will be somewhat reduced. The method presented here is the manufacturer’s recommended approach to maximize volume of plasma depleted per given time period. Multiple aliquots of a plasma sample (up to 10) are depleted, pooled, and concentrated, and following elution of the bound proteins, the unbound fraction from all 10 samples is redepleted in a polishing step. While not all vendors recommend a polishing step, the principle of removing small amounts of residual targeted proteins in the unbound pool using a repurification is generally applicable and is a good idea for the most critical work, especially when spin columns are used. 1. Prepare Buffer A (1x Equilibration Buffer): Dilute one part of ProteoPrep 20 Equilibration Buffer, 10x concentrate with nine parts of room temperature ultrapure water. The final volume necessary for each plasma application is 5 ml. 2. Prepare Buffer B (1x Elution Solution): Dilute one part of ProteoPrep 20 Elution Solution, 10x concentrate with nine parts of room temperature ultrapure water. The final volume necessary for each plasma application is 2 ml. 3. Remove the bottom plug and screw cap from the spin column and place into a 2 ml collection tube. Centrifuge the spin column at 1000–2000 × g for 60 sec to remove the storage buffer using either a speed-vac without vacuum or a microfuge. 4. Attach a Luer Lock fitting to the spin column, draw 4 ml of 1x Equilibration Buffer into a syringe, attach the syringe to the Luer Lock cap, and slowly push the buffer through the column to equilibrate the resin. 5. Remove the Luer Lock fitting, loosely recap the column, and place into a collection tube. Centrifuge the spin column at 1000–2000 × g for 60 sec to remove the excess Equilibration Buffer. Discard the buffer and place the column into a clean collection tube. 6. Dilute plasma sample (generally 8 µl per depletion) to 100 µl with Equilibration Buffer (see Note 9). Filter diluted plasma with a prerinsed 0.22 µm microcentrifuge filter. Keep filtered sample on ice until ready for use. 7. Add 100 µl diluted sample to the top of the packed resin bed, and incubate at room temperature for 20 min.

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8. Centrifuge the spin column and collection tube at 1000–2000 × g for 60 sec. 9. Wash the remaining depleted plasma from the column with 100 µl 1x Equilibration Buffer. Collect wash by centrifuging the spin column at 1000–2000 × g for 60 sec. This wash may be collected in the same tube used in step 8. 10. Repeat step 9 for an additional 100 µl wash of the resin, the final pooled volume of the unbound fraction should be approximately 300 µl. 11. Elute bound proteins with 1x Elution Solution. Attach a Luer Lock fitting to the spin column, draw 2 ml of 1x Elution Solution into a syringe, and slowly push the buffer through the column to remove proteins bound to the resin. 12. Neutralize bound proteins with 1M NaOH to bring pH to 8.0 (see Note 6). 13. Reequilibrate the spin column: Attach a Luer Lock fitting to the spin column, draw 4 ml of 1x Equilibration Buffer into a syringe, and slowly push the buffer through the column to equilibrate the resin. 14. Remove the Luer Lock fitting and place the column into a clean collection tube. Centrifuge the spin column at 1000–2000 × g for 60 sec to remove the excess Equilibration Buffer. Discard the buffer and place the column into a new collection tube. 15. Repeat steps 7–14 for up to 10 depletions. 16. After all samples have been depleted, begin concentrating the unbound fraction (see Note 10). Add up to 4 ml of unbound pool to a prerinsed Amicon-Ultra 5K MWCO concentrator. Centrifuge for 15 min at 1500 × g, 4°C. 17. Repeat step 16 until the unbound pool is concentrated to 100–200 µl. 18. Rinse the concentrator unit with 100 µl Equilibration Buffer, and pool this with the concentrated unbound sample to recover proteins adhering to the concentrator membrane. 19. Add 100 µl of the unbound sample to the reequilibrated and packed resin bed. Incubate for 20 min and centrifuge spin column and collection tube at 1000–2000 × g for 60 sec. 20. Repeat step 19 until all of the concentrated unbound plasma has been reapplied to the column. 21. Wash the column twice with 100 µl Equilibration Buffer, centrifuge, and pool washes with the twice-depleted plasma. The final volume should be about 500 µl. 22. Repeat steps 11–14 from this section to elute remaining bound proteins and reequilibrate the column. 23. Store column in 300 µl Equilibration Buffer containing Preservative Solution, with the bottom plug securely in place, and tightly recap the column (see Note 7).

3.3. Large-Scale ProteoPrep 20 Spin Column Depletion Using a Jumbo Custom Column As mentioned above, some manufacturers will create custom HPLC or spin columns to meet the needs of investigators who want to deplete larger volumes of plasma or serum more efficiently. This protocol utilizes a custom ProteoPrep 20 Jumbo Plasma Immunodepletion Column (3.7 ml resin). Similar to the

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smaller spin column, there are several alternative loading/depletion strategies. For example, this column can be used to deplete 100 µl human plasma without a polishing step, and in one day it is practical to perform approximately 10 first pass depletions for a total volume of 1 ml per day. Alternatively, one can purify 250 µl per cycle and four first pass depletions followed by one polishing cycle of the combined and concentrated unbound fractions (also 1 ml but only 5 total cycles rather than 10 using the single-pass strategy). 1. Prepare Buffer A (1x Equilibration Buffer): Dilute one part of ProteoPrep 20 Equilibration Buffer, 10x concentrate with nine parts of room temperature ultrapure water. The final volume necessary for each plasma application is 60 ml. 2. Prepare Buffer B (1x Elution Solution): Dilute one part of ProteoPrep 20 Elution Solution, 10x concentrate with nine parts of room temperature ultrapure water. The final volume necessary for each plasma application is 24 ml. 3. Remove the bottom plug and screw cap from the spin column and place into a collection tube. Centrifuge the spin column at 1000–2000 × g for 60 sec to remove the storage buffer. 4. Attach a Luer Lock fitting to the spin column, draw 16 ml of 1x Equilibration Buffer into a syringe, attach the syringe to the Luer Lock cap, and slowly push the buffer through the column to equilibrate the resin. 5. Remove the Luer Lock fitting, loosely recap the column, and place into a collection tube. Centrifuge the spin column at 1000–2000 × g for 60 sec to remove the excess Equilibration Buffer. Discard the buffer and place the column into a clean collection tube. 6. Dilute plasma sample (100 or 250 µl per depletion) to 1.25 ml with Equilibration Buffer. Filter diluted plasma with a prerinsed 0.22 µm centrifuge filter. Keep filtered sample on ice until ready for use. 7. Add 1.25 ml diluted sample to the top of the packed resin bed, and incubate at room temperature for 20 min 8. Centrifuge the spin column and collection tube at 1000–2000 × g for 60 sec. 9. Wash the remaining depleted plasma from the column with 1.25 ml 1x Equilibration Buffer. Collect wash by centrifuging the spin column at 1000–2000 × g for 60 sec. This wash may be collected in the same tube used in step 8. 10. Repeat step 9 for an additional 1.25 ml wash of the resin, the final pooled volume should be about 3.75 ml. 11. Elute bound proteins with 1x Elution Solution. Attach a Luer Lock fitting to the spin column, draw 12 ml of 1x Elution Solution into a syringe, and slowly push the buffer through the column to remove proteins bound to the resin. 12. Centrifuge the spin column in collection tube at 1000–2000 × g for 60 sec to remove any additional Elution Solution from the column. 13. Neutralize bound proteins with 1M NaOH to bring pH to 8.0 (see Note 6). 14. Reequilibrate the spin column: Attach a Luer Lock fitting to the spin column, draw 16 ml of 1x Equilibration Buffer into a syringe, and slowly push the buffer through the column to equilibrate the resin.

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15. Remove the Luer Lock fitting and place the column into a clean collection tube. Centrifuge the spin column at 1000–2000 × g for 60 sec to remove the excess Equilibration Buffer. Discard the buffer and place the column into a new collection tube. 16. Repeat steps 7–14 for up to 10 depletions of 100 µl or 4 depletions of 250 µl. Unbound fractions can be concentrated while subsequent depletions are being performed. 17. After all samples have been depleted, complete concentration of the unbound pool (see Note 10). Add up to 4 ml of unbound to a prerinsed Amicon-Ultra 5K MWCO concentrator. Centrifuge for 15 min at 1500 × g, 4°C. 18. Repeat step 17 until the unbound sample is concentrated to approximately 1 ml. 19. Rinse the concentrator unit with 250 µl Equilibration Buffer, and pool this with the concentrated unbound sample to recover any proteins adhering to the concentrator membrane. 20. If 250 µl aliquots were purified per cycle, or if maximum depletion of 100 ml per cycle purifications are desired, add 1.25 ml of the unbound sample to the reequilibrated and packed resin bed. Incubate for 20 min and centrifuge spin column and collection tube at 1000–2000 × g for 60 sec. 21. Repeat step 20 until all of the concentrated unbound plasma has been reapplied to the column. 22. Wash the column twice with 1.25 ml Equilibration Buffer, centrifuge, and pool washes with the twice-depleted plasma. The final volume should be 3.75 ml. 23. Repeat steps 11–14 from this section to elute remaining bound proteins and reequilibrate the column. 24. Store column in 1.25 µl Equilibration Buffer containing Preservative Solution, with the bottom plug securely in place, and tightly recap the column (see Note 7).

3.4. Ethanol Precipitation We have found that ethanol precipitation is an effective alternative to ultrafiltration for final unbound and bound fractions from either FPLC or spin columns that will be used in subsequent steps that use denaturants such as either 8 M urea or SDS. The ethanol precipitation requires less operator time and more effectively removes salts and detergent. The ethanol should be high quality, absolute proof, and must be chilled to −20°C to facilitate protein precipitation. 1. 2. 3. 4.

Add 9 volumes of 200 proof ethanol (−20°C) to the pooled unbound fraction. Incubate overnight at 0°C. Centrifuge sample for 30 min at 1500 × g, 4°C. Carefully remove supernatant and air dry pellet briefly in a fume hood or speed vac (see Note 11).

4. Notes 1. The ProteoPrep LC column comes packed in a storage solution containing glycerol. For the first use of the column, attach the top of the column to the tubing as described in Subheading 3.1, step 4. Do not connect the

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column outlet line to the bottom of the column. Instead, place the column over a beaker to collect the glycerol solution. Allow the column to equilibrate with 20–25 ml of 1x Equilibration Buffer at no higher than 0.5 ml/min. Carefully monitor pressure and decrease the flow rate as needed to not exceed 30 psi. Increased pressures at this step may cause bed compression and a void at the top of the resin bed. When the resin has changed from opaque to white, and the buffer runs clear, the column outlet line may be connected. The column should be conditioned with two complete blank cycles prior to injecting a sample. It is also preferable to run a few non-critical control plasma samples before depleting plasma samples that will be used in comparative studies. 2. Earlier versions of this method recommended diluting the plasma sample 5x with Buffer A prior to filtering. To enhance depletion of apolipoproteins and recovery of unbound proteins, the manufacturer recommended omitting the dilution step. However, we have not observed any major changes in apolipoprotein removal nor does a five-fold dilution of the plasma significantly increase the final volume of the unbound pool. Additionally, diluting the sample makes it more convenient to add protease inhibitors to the sample before injection if desired. 3. The manufacturer describes this column’s capacity as 100 µl of plasma with a concentration of 40–50mg/ml. However, this is a very low estimate for protein concentration, and most human serum and plasma samples are likely to have substantially higher protein concentrations. For example, the range for total blood protein in normal human serum is 60–85 mg/ml, and plasma samples will have similar or slightly higher values. Furthermore, as indicated above, the normal range is quite broad and ranges in patients with diseases are often even more variable. Total protein concentration will also be affected by the use of different anti-coagulants and collection tubes. Therefore, it is advisable to determine the protein concentration of each sample prior to depletion and base loads upon total protein, not volume. When preparing samples, other considerations to note are: the number of depletions and/or different plasma samples that will be run in a single day. If multiple runs of the same sample are being depleted, it is best to thaw and prepare the entire sample at once and store it on ice until the entire sample is depleted, for optimal reproducibility. If several samples from different donors are being depleted sequentially, it is recommended that a single sample is thawed and filtered shortly before its injection, and while one sample is running, thaw and prepare the next sample. This will minimize proteolysis of the sample. A related option is to add protease inhibitors to the plasma sample prior to injection, although the value of adding protease inhibitors is ambiguous. Plasma and serum intrinsically contain many proteases as well as protease inhibitors, and some proteolysis

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4.

5. 6.

7.

Echan and Speicher of susceptible proteins usually occurs in vivo. While further proteolysis may occur during sample processing, some protease inhibitors may react indiscriminately with plasma proteins (9) and other inhibitors are proteins or large peptides. Instead of adding protease inhibitors to plasma or serum, we prefer to minimize exposure of samples to conditions that favor proteolysis. The one step where it is not practical to avoid protease favorable conditions is during binding of the targeted proteins to the immunoaffinity resin. This is because in order to optimize binding affinity, conditions close to physiological pH without denaturants must be used, and in order to achieve reasonable binding kinetics, low temperatures should be avoided. However, it should be noted that plasma and serum are far more stable to proteolysis than cell lysates, and moderate length exposure to physiological conditions at room temperature appear to have only very minor effects on proteomes (9). Regardless of the precise sample handling methods to be used, the most critical factor is to ensure that all samples to be compared are processed in as consistent a manner as possible. We have found that running an initial “quick blank gradient” at 3 ml/min for the entire gradient with no injection is useful for several reasons. First, it will ensure that the column is equilibrated to room temperature prior to the sample being injected. Also, as the column warms up, solvent outgassing may occur; but the increased flow rate forces any small air bubbles out of the column before large pockets can form and disrupt the column packing. Finally, a blank run at the beginning of the day removes any residual protein that may be bound to the column from previous depletions. Any protein peaks observed during the blank run should be run on 1-D SDSPAGE and analyzed with silver stain. It is useful to reserve a small aliquot of each fraction before pooling the unbound to analyze individual fractions on 1-D SDS-PAGE. It is recommended that bound proteins typically be collected and frozen for potential future analysis. Prior to storage, the bound pool should be neutralized by adding 1M NaOH to bring the pH to approximately 8.0. Proper storage and monitoring of antibody columns is essential for maintaining optimal column performance. Storage in Equilibration Buffer at 4°C is suitable for short-term storage, i.e., if the column will be reused within a few days. Long-term column storage (1 week or more) requires the addition of a Preservative Solution, included with the ProteoPrep 20LC and spin columns. Alternatively, flushing the column with Buffer A containing 0.01% sodium azide should protect the antibody column from microbial growth. It is important to keep track of the total number of cycles run on each column. Periodically and especially when the upper limit of recommended uses has been reached, column performance should be critically evaluated. We have found the lifetime of well managed columns is

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usually greater than the recommended lifetimes. In addition, as column capacity slowly and inevitably declines, it is usually adequate to simply reduce protein loads slightly to achieve the same degree of depletion as was obtained with a new column. To monitor antibody loss from the column, bound fractions from sample purifications and bound fractions from blank cycles after extended column storage should be periodically monitored by digesting an aliqout of the bound pool with trypsin followed by LC-MS/MS analysis. The resulting data should ideally be searched against an all species database. Alternatively, a human sequence database supplemented with all immunoglobulin sequences from the species of origin for the antibodies on the column should be used. In this manner, one can detect antibody bleed and/or proteolysis of the coupled antibodies. 8. After each use, the FPLC or HPLC system should be thoroughly flushed with water, followed by 50% Isopropanol. This will prevent corrosion due

Fig. 1. Depletion of human plasma using spin columns. (Left) Results from depletion of ten 8 µl aliquots of human plasma using a standard spin column. Lanes: 1-undepleted plasma; 2-pool of depleted plasma from the ten runs after first cycle of depletion; 3-depleted plasma pool after polishing step; 4-pool of bound proteins eluted from column and concentrated using an Amicon-ultra 5K MWCO centrifugal filters. (Right) Results from depletion of a single 250 µl aliquot of human plasma using a large custom spin column. Lanes: 5-undepleted plasma; 6-unbound pool after first cycle of depletion; 7-unbound pool after polishing step; 8-pool of bound proteins after elution and concentration using an Amicon-ultra 5K MWCO centrifugal filters.

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Fig. 2. Depletion of human plasma on a large LC column. Results from depletion of 100 µl human plasma after a single pass through a ProteoPrep 20 LC column. P-undiluted human plasma, UB-pooled unbound fraction after ethanol precipitation, B-pooled and neutralized bound fraction after ethanol precipitation.

to halogen salt-containing buffers or clogging of lines due to bacterial growth. 9. The spin column capacity is based on 8 µl of plasma with a concentration of 40–50mg/ml. If plasma concentration is higher, sample volume should be adjusted accordingly. If multiple aliquots of the same sample are to be depleted in one day, prepare enough sample for all depletions and store sample on ice. If proteolysis is a concern for downstream applications, it is acceptable to add protease inhibitors at this step (Fig. 1) (see Note 3). 10. Following each depletion, the manufacturer recommends continuous concentration of the unbound in the same micro-concentrator unit until the desired number of depletions has been completed. We prefer to pool the unbound, and begin concentration after all the depletions are completed, using a larger capacity concentrator. This will provide a total unbound sample before concentration that can be analyzed in parallel on 1-D

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SDS-PAGE with the concentrated sample to check for protein losses during concentration. 11. To completely dry pellet, without risk of airborne contaminants or sample oxidation, it is preferable to dry the pellet under a gentle stream of argon or in a speed vac for a few minutes. The pellet is then resuspended in the desired resolubilization buffer, or the dried pellet may be stored at −20–80°C until it is needed. To check for potential sample loss during the ethanol precipitation, dry 10% of the ethanol supernatant in a speed vac, and run in parallel with proportional volumes of sample before and after precipitation on 1-D SDS-PAGE. References 1. Jacobs, J.M., Adkins, J.N., Qian, W.J., Liu, T., Shen, Y., Camp, D.G. et. al. (2005) Utilizing human blood plasma for proteomic biomarker discovery. J. Proteome Res. 4, 1073–1085. 2. Anderson, N.L. and Anderson, N.G. (2002) The human plasma proteome: history, character, and diagnostic prospects. Mol. Cell. Proteomics 1, 845–867. 3. 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 2, 1096–1103. 4. Hoffman, S.A., Joo, W.A., Echan, L.A., and Speicher, D.W. (2007) Higher dimensional (Hi-D) separation strategies dramatically improve the potential for cancer biomarker detection in serum and plasma. J. Chromatogr. B Analyt Technol Biomed Life Sci. 849, 43–52. 5. Omenn, G.S., States, D.J., Adamski, M., Blackwell, T.W., Menon, R., Hermjakob, H. et al. (2005) Overview of the HUPO Plasma Proteome Project: Results from the pilot phase with 35 collaborating laboratories and multiple analytical groups, generating a core dataset of 3020 proteins and a publicly-available database. Proteomics 5, 3226–3245. 6. States, D.J., Omenn, G.S., Blackwell, T.W., Fermin, D., Eng, J., Speicher, D.W., and Hanash, S.M., (2006) Challenges in deriving high-confidence protein identifications from data gathered by a HUPO plasma proteome collaborative study. Nat. Biotech. 24, 1038–1183. 7. Echan, L.A., Tang, H.-Y., Ali-Khan, N., Lee, K.B., and Speicher, D.W. (2005) Depletion of multiple high abundance proteins improves protein profiling capacities of human serum and plasma. Proteomics 5, 3292–3303. 8. Pieper, R., Su, Q., Gatlin, C.L., Huang, S.T., Anderson, N.L., Steiner, S. (2003) Multicomponent immunoaffinity subtraction chromatography: an innovative step towards a comprehensive survey of the human plasma proteome. Proteomics 3, 422–32. 9. Rai, A.J., Gelfand, C.A., Haywood, B.C., Warunek, D.J., Yi, J., Schuchard, M.D. et al. (2005) HUPO Plasma Proteome Project specimen collection and handling: toward the standardization of parameters for plasma proteome samples. Proteomics 5, 3262–3277.

17 Preparation of Yeast Samples for 2-D PAGE Joakim Norbeck

1. Introduction Yeasts are the focus of much research, both in their role as pathogens and as biotechnically important organisms, and not the least in their role as model systems for eukaryotic cells. In particular, Saccharomyces cerevisiae has also been the object of several proteomics-related efforts. However, the preparation of protein extract from yeast is complicated by the presence of a cell wall of mainly chitin and glucans, which needs to be disrupted in the extraction process. Several methods presenting solutions to this problem, in connection with sample preparation for two-dimensional polyacrylamide gel electrophoresis (2-D PAGE), have been described (1,2), in which the cell wall is broken either by vortexing in the presence of glass beads or by sonication. We have found the method described below to be robust and to yield reproducible results in several studies (3–6). It is furthermore easy to perform and does not require any specialized equipment. 2. Materials 1. Sample buffer I: sodium dodecyl sulfate (SDS) (0.3 g), b-mercaptoethanol (5.0 mL), Tris-HCl (0.444 g), Tris base (0.266 g), MilliQ (or equivalent) water (to a final volume of 10 mL). 2. Sample buffer II (see Note 1): 1.5 M Tris base (80 mL), 1.5 M Tris-HCl (1585 mL), 1 M MgCl2 (250 mL), DNase I (Worthington Biochemical Corp., NJ) (5 mg), RNase A (Worthington Biochemical Corp., NJ) (1.25 mg), MilliQ (or equivalent) water (to a final volume of 5 mL).

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Immobilized pH gradient (IPG)-rehydration buffer (urea/thiourea buffer): urea (4.8 g; gives a final concentration of 8 M; see Note 2), Triton X-100 or Nonidet P40 (100 mL; see Note 3), 1 M DTT (100 mL), IPG-buffer/ampholine (50 mL; see Note 4), bromophenol blue (trace amount approx 0.01% w/v), MilliQ (or equivalent) water (to a final volume of 10 mL).

3. Method The method described in the following sections can be divided into (1) an initial cell disruption step; (2) a protein solubilization step; (3) a nuclease treatment step; and (4) a final phase in which protein extract is diluted in IPG-rehydration buffer immediately prior to application on the first dimension of 2-D PAGE. The procedure is carried out in 1.5-mL microcentrifuge tubes. A suitable starting material is a pellet of yeast cells from 10 mL of culture with a density of 5–10 million cells/mL, corresponding to an optical density (at 610 nm) of approx 0.5, which will typically yield a pellet of 5–10 mL of cells. The method described below is adjusted to this amount of cells. The protocol can be scaled up or down; however, care should be taken to not use a final extract volume of more than 500 mL or less than 50 mL, since this will reduce the efficiency of the cell-disruption step. 3.1. Cell Disruption 1. Add 160 mL of ice-cold milliQ-quality water containing protease inhibitors (e.g., Complete™, Roche, Inc.) to the cell pellet. 2. Add 0.25 g of chilled glass beads (diameter 0.5 mm) to the sample. 3. Vortex 4 • 30 s on a table shaker at maximum speed (approx 2500 rpm) with intermittent placement of samples on ice for at least 1 min.

3.2. Protein Solubilization 1. Add 20 mL of sample buffer I and vortex the tube(s) briefly to mix. 2. Place tube(s) at 95°C for 5 min. Make sure to secure the lid of the tube, alternatively to make a small hole in the lid, prior to the heating step. 3. Cool samples on ice for 5 min.

3.3. Nuclease Treatment 1. 2. 3. 4.

Add 20 mL of sample buffer II and vortex the tube(s) briefly to mix. Incubate on ice for 10 min. Centrifuge samples at full speed (approx 15,000g) in a microcentrifuge at 4°C. Aspire the supernatant (constituting the protein extract) to a new microcentrifuge tube and freeze at −20°C, or use immediately.

3.4. Dilution in IPG-Rehydration Buffer 1. Dissolve sample in required volume of IPG-rehydration buffer (see Note 5). 2. Incubate the sample at 37°C for 10 min.

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3. Spin down sample (15,000g, 10 min, room temperature) to remove any particles that might remain.

4. Notes 1. The DNase and RNase should be dissolved in the buffer as the final step. 2. 8 M Urea can be substituted by a combination of 7 M urea and 2 M thiourea. The chaotropic agent thiourea is the most highly beneficial addition to the IPGrehydration buffer; this addition can strongly improve the solubility of many proteins which may produce “streaking” or which are completely absent on 2-D PAGE gels run with normal urea-based buffer in the first dimension (7). However, thiourea requires a special permit from inspecting authorities in many countries due to its suspected carcinogenic properties. 3. Triton X-100 can be substituted for 2% (w/v) of CHAPS. CHAPS is considered to be the preferred detergent for the IPG-rehydration buffer (7). Unfortunately, it is also considerably more expensive than Triton X-100 and Nonidet P-40. We normally use Triton X-100, since in our hands the choice of detergent in the rehydration buffer has only a minor influence on the final protein resolution. 4. IPG buffer or ampholine should be chosen for each type of IPG strip, or for the desired separation interval. 5. The final volume to which the sample is diluted is determined by the system used for running this first dimension. We most often use the precast pH-gradient strips (supplied by, for example, Amersham Biosciences and BioRad, Inc.), for which the rehydration volume is commonly in the range of 125–500 mL. However, the protein extract is also compatible with the original glass tube-based system for running 2-D PAGE (8), or for application on IPG strips using sample cups, in which the sample volume will typically be 7) proteins. Yeast 13, 1519–1534. 5. Norbeck, J. and Blomberg, A. (1997) Metabolic and regulatory changes associated with growth of Saccharomyces cerevisiae in 1.4 M NaCl. Evidence for osmotic induction of glycerol dissimilation via the dihydroxyacetone pathway. J. Biol. Chem. 272, 5544–5554. 6. Blomberg, A., Blomberg, L., Norbeck, J., et al. (1995) Interlaboratory reproducibility of yeast protein patterns analyzed by immobilized pH gradient two-dimensional gel electrophoresis. Electrophoresis 16, 1935–1945. 7. Rabilloud, T., Adessi, C., Giraudel, A., and Lunardi, J. (1997) Improvement of the solubilization of proteins in two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 18, 307–316. 8. O’Farrell, P. H. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–4021. 9. Ozols, J. (1990) Amino acid analysis. Methods Enzymol. 182, 587–601.

18 Membrane Protein Preparation Using Aqueous Polymer Two-Phase S ystems Jens Schindler and Hans Gerd Nothwang 1. Introduction Membrane proteins serve many fundamental functions for cells such as cell-cell interactions, signal transduction, and molecular transport both within and between cells. Bioinformatic analysis has estimated that transmembrane domain containing proteins represent about 20-30% of all open reading frames in sequenced genomes (1). In addition, many proteins are anchored to the membranes via lipidic posttranslational modifications such as glycosylphosphatidyl inositol anchors or via interaction with other membrane proteins. In total, embrane anchored proteins represent a considerable portion of all cellular proteins. They also represent 70% of all known drug targets, which prioritize them in biomedical research (2). Consequently, profiling membrane proteomes will aid in both understanding basic biological processes and discovering novel targets for therapeutic agents. Membrane proteomics, however, is rendered difficult on several levels. Notorious difficulties in membrane proteomics include the solubilization, digestion, and separation of membrane proteins. They contain both hydrophilic and hydrophobic domains, which preclude their routine analysis by conventional two-dimensional polyacrylamide gel electrophoresis due to solubilization difficulties. Furthermore, membrane proteins contain stretches of highly hydrophobic transmembrane domains that lack an adequate number of arginine and lysine residues, which are required for in gel digestion with the commonly used endoproteinase trypsin. The sequence coverage of membrane proteins can hence be relatively poor when only this endoproteinase is used. Some of these problems have been overcome by advances using nongelbased liquid chromatography techniques applying a wide variety of stationary

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and mobile phases and chromatography types (3). Recent separation protocols made use of size-exclusion chromatography (SEC), affinity chromatography (AC), ion-exchange chromatography (IEX) and reversed-phase chromatography (RPC) (3). In the multidimensional protein identification technology (Mud-PIT), for instance, peptide mixtures are loaded onto a biphasic microcapillary column packed with reversed-phase and strong-cation-exchange material. Sequentially, eluted peptides are detected and identified by MS/MS (4). This approach was recently taken a step forward by combining it with methanol-facilitated solubilization and digestion of the protein sample. This combination allowed identification of 1,500-2,500 proteins starting with 100-200 µg of membrane proteins (5). Another serious, but often underestimated challenge in membrane proteomics is the complexity of a eukaryotic cell. A typical eukaryotic cell contains many different subcellular compartments each with its own subset of membrane proteins. In addition, the subcellular distribution pattern of a given protein might change under different conditions and this might alter its function. Striking example are integral plasma membrane proteins such as the glucose transporter Glut4 in muscle and fat cells (6) or the glutamate receptors (7) in the brain, which can shuttle between intracellular stores and the plasma membrane under different physiological conditions. Therefore, it is mandatory to characterize these subproteomes separately in order to identify compartment-specific proteins, to describe the physiological state of a tissue under defined conditions, and to gain insight into pathophysiological mechanisms. Unfortunately, most of the approaches mentioned before do not address this issue or fail to perform sufficiently well in separating the different membraneous compartments (8). There is hence a clear need for the reproducible production of well-characterized membrane protein samples. In this chapter, we provide a robust and efficient protocol to isolate plasma membranes from brain tissue (9). It is based on an aquous polymer two-phase system exploiting the principle of countercurrent distribution. Advantages of this protocol include its ease and efficiency as well as the low costs. Furthermore, two phase systems are very versatile allowing also isolation of substructures of membranes such as caveolae (for further details on membrane partitioning see [10]). The protocol can be adapted to other tissues as well. 2. Materials Due to the strong influence of ions on membrane partitioning in the two-phase systems, double distilled water should be used throughout the experiments. 1. 2. 3. 4.

Glass-Teflon homogenizer. Dextran stock solution: Dextran T500 (20% w/w) (see Note 1). PEG stock solution: PEG 3350 (40% w/w). Tris-H2SO4: 200 mM Tris, pH 7.8 adjusted with H2SO4.

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3. Methods 3.1 Two-Phase Partitioning All steps of the affinity two-phase partitioning protocol should be performed at 4 °C. Working at room temperature prevents phase separation. The procedure is illustrated in Fig. 1. The numbers in Fig. 1 correspond to the numbered twophase systems given in Table 1, and the letters refer to the phases as indicated in the protocol given below.

A

I.

II.

III.

two-phase system C D E

B

fresh top 2) phase

1)

fresh top 3) phase

2)

1)

fresh top 4) phase

3)

2)

F

G

F

G

1)

… VII. A

B

C D E two-phase system

Fig. 1. Scheme of countercurrent distribution. In CD experiments, the top phase of the first two-phase system A is transferred to a fresh bottom phase B and the bottom phase of two-phase system A is reextracted with a fresh top phase. After six iterations, biomaterial is efficiently separated.

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Table 1 Composition of Two-Phase Systems Dextran stock solution PEG stock solution Tris-H2SO4 Water

Two-phase system “A”

Two-phase systems “B-G”

1.035 g 0.518 g 0,750 g 0.598 g

1.035 g 0.518 g 0.750 g 0.698 g

1. Prepare seven two-phase systems with the compositions indicated in Table 1 one day prior to use. Mix them by 20 invertations, vortexing for 10 sec, another 20 invertations and store the mixtures at 4°C overnight. Two-phase systems will form over night with the top phase enriched in PEG and the bottom phase enriched in dextran. 2. On the next day, remove all top phases from two phase systems “B-G” and store them separately. 3. Homogenize 0.100 g brain tissue (see Note 2) in two-phase system “A” using a glass-teflon homogenizer followed by 45 sec of sonication. Centrifuge at 700 × g for 5 min to accelerate phase separation. 4. Transfer the top phase (Top) of two-phase system “A” onto the bottom-phase (Bot) of two-phase system “B” (TopA -> BotB) (see Note 3). Add an equal amount of fresh top-phase (stored in step 2) onto bottom-phase “A” (I. in Fig. 1). Mix both two phase systems by 20 invertations, vortex for 10 s, then mix again by another 20 invertations. Centrifuge at 700 × g for 5 min to accelerate phase separation. 5. Transfer top phases in the following order (II. in Fig. 1): 1) TopB -> BotC; 2) TopA -> BotB. Add an equal amount of fresh top-phase (stored in step 2) onto bottom-phase “A.” Mix all two-phase systems by 20 invertations, vortexing for 10 sec, and another 20 invertations. Centrifuge at 700 × g for 5 min to accelerate phase separation. 6. Transfer top phases in the following order (III. in Fig. 1): (1) TopC -> BotD; (2) TopB -> BotC; (3) Top A -> BotB. Add an equal amount of fresh top-phase (stored in step 2) on to bottom-phase “A.” 7. Mix all two-phase systems by 20 invertations, vortexing for 10 sec, and another 20 invertations. Centrifuge at 700 × g for 5 min to accelerate phase separation. 8. Transfer top phases in the following order: (1) Top D -> BotE; (2) TopC -> BotD; (3) TopB -> BotC; (4) Top A -> BotB. Add an equal amount of fresh top-phase (stored in step 2) onto bottom-phase “A.” Mix all two-phase systems by 20 invertations, vortexing for 10 sec, and another 20 invertations. Centrifuge at 700 × g for 5 min to accelerate phase separation. 9. Transfer top phases in the following order: (1) TopE -> BotF; (2) Top D -> BotE; (3) TopC -> BotD; (4) TopB -> BotC; (5) Top A -> BotB. Add an equal amount of fresh top-phase (stored in step 2) onto bottom-phase “A”. Mix all two-phase systems by 20 invertations, vortexing for 10 sec, and another 20 invertations. Centrifuge at 700 × g for 5 min to accelerate phase separation.

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10. Transfer top phases in the following order: (1) TopF -> BotG; (2) TopE -> BotF; (3) Top D -> BotE; (4) TopC -> BotD; (5) TopB -> BotC; 6) Top A -> BotB. Add an equal amount of fresh top-phase (stored in step 2) onto bottom-phase “A.” 11. Mix all two phase systems by 20 invertations, vortexing for 10 sec, and another 20 invertations. Centrifuge at 700 × g for 5 min to accelerate phase separation. After phase separation you end up with 7 two-phase systems (VII. in Fig. 1). Plasma membranes are enriched in top phases “F+G.” 12. To proceed further, isolate membranes out of TopG (maximum purity) or from TopG combined with TopF (maximum yield) by diluting the phases 1:10 with water and spin for 1 h at 150,000 × g (see Note 4). 13. Proteins in the pellet are solubilized by sonication in Laemmli-sample buffer. After incubation for 15 min at 37 °C the sample is separated by SDS-PAGE (see Note 5).

4. Notes 1. Dextran can contain up to 10% water and for that reason the 20% stock solution has to be prepared from freeze-dried dextran. For freeze drying, dissolve dextran in distilled water in a plastic dish with a large surface (e.g. Petri dish), freeze it at −80 °C and dry it by sublimating the water under vacuum. Store the freeze-dried dextran in closed plastic tubes sealed tightly with parafilm at −20 °C. Let it come to room temperature before opening to protect it from humidity. 2. The protocol can also be applied to other tissues. Applicants should take care that the tissue is thoroughly homogenized (e.g., sceletal muscle and heart tissue are much harder to homogenize than brain tissue) or the yield of plasma membranes will decrease. Altering the polymer concentration or the buffer conditions might be considered to optimize the protocol for the fractionation of other tissues (for review see Schindler and Nothwang, 2006) 3. In two-phase systems, interphases are always assigned to the bottom phase. 4. The amount of peripheral membrane proteins in the pellet can be reduced by extractions with Na2CO3. The procedure solubilizes peripheral membrane proteins whereas integral proteins remain in the pellet (10). For this purpose, resuspend the pellet in ice cold Na2CO3 (0.2 M), incubate for 15 min on ice and pellet the membranes by ultracentrifugation at 233,000 × g for 1 h. This step might be repeated twice. 5. Proteins can also be solubilized in different buffers, depending on further processing (e.g., 16-BAC-SDS-PAGE, BN-PAGE, immunoprecipitation) Acknowledgments This work was funded in part by the Nano+Bio-Center of the University of Kaiserslautern.

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References 1. Lehnert, U., Xia, Y., Royce, T.E., Goh, C.S., Liu, Y., Senes, A. et al. (2004) Computational analysis of membrane proteins: genomic occurrence, structure prediction and helix interactions. Q. Rev. Biophys. 37, 121–146. 2. Hopkins, A.L. and Groom, C.R. (2002) The druggable genome. Nat. Rev. Drug Discov. 1, 727–730. 3. Wang, H. and Hanash, S. (2003) Multi-dimensional liquid phase based separations in proteomics. J. Chromatogr. B. Analyt. Technol. Biomed. Life Sci. 787, 11–18. 4. Washburn, M.P., Wolters, D., and Yates, J.R.(2001) Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 19, 242–247. 5. Blonder, J., Chan, K.C., Issaq, H.J., and Veenstra, T.D. (2006) Identification of membrane proteins from mammalian cell/tissue using methanol-facilitated solubilization and tryptic digestion coupled with 2D-LC-MS/MS. Nat. Protoc. 1, 2784–2790. 6. Bryant, N.J., Govers, R., and James, D.E. Regulated transport of the glucose transporter glut4. Nat. Rev. Mol. Cell Biol. 3, 267–277. 7. Malinow, R. and Malenka, R.C. )2002) AMPA receptor trafficking and synaptic plasticity. Ann. Rev. Neurosci. 25, 103–126. 8. Issaq, H.J. (2001) The role of separation science in proteomics research. Electrophoresis 22, 3629–3638. 9. Schindler, J., Lewandrowski, U., Sickmann, A., and Friauf, E. (in press) Aqueous Polymer Two-Phase Systems for the Proteomic Analysis of Plasma Membranes from Minute Brain Samples. J. Proteome Res. 10. 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.

19 Subcellular Fractionation of Small Sample Amounts Hans Gerd Nothwang, Isabelle Guillemin, and Jens Schindler

1. Introduction A central issue in proteomics is the generation of qualitative and quantitative protein inventories for a given tissue, organ, or organism. In many eukaryotes, this amounts to a formidable task. Mammals may express more than 200,000 different transcripts with a quarter of them encoding proteins (1). In addition, there exist several hundred different (2) posttranslational modifications, increasing the number of different protein species by an estimated factor of 10, compared to the number of different mRNAs (3,4). Consequently, higher organism will display more than 1,000,000 different proteins (5). Considering as a lower, very conservative limit, the expression of 5–10% of the transcriptome in an organ, about 50,000–100,000 different protein species will be present in such a sample. Even in a single cell, > 10,000 different protein species likely exist. This number easily exceeds the potential of any known proteomic approach. Another important issue in protein analysis is the high range of protein abundance. Protein concentrations likely vary between 106–1010, depending on the tissue analyzed (6,7). This renders the identification of low abundant proteins impossible in the bulk of total protein extracts. To overcome these hurdles, many novel sample fraction approaches have been developed. These include SDS-PAGE combined with liquid chromatography coupled mass spectrometry or multidimensional liquid chromatography techniques (8,9). However, when aiming at proteomics as a bottom-up process to understand a biological system, most of these approaches fall short by neglecting the importance of the compartmentalized organization of any living cell. A cell is highly structured and organized. Most, if not all proteins have precise subcellular locations and mistargeting might have severe pathophysiological consequences. It matters, whether a transcription factor such as NF-kappaB is in the cytosol From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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or in the nucleus. Furthermore, posttranslational modifications depend on the localization of a protein. A striking example is the addition and modification of sugar residues during the passage of plasma membrane proteins through the biosynthetic secretory pathway. Therefore, everything in proteomics makes only sense in the light of subcellular organization and this information should be retained during sample preparation. Here, we present a subcellular fractionation protocol, developed for minute amounts of tissue or cell cultures (10). The protocol is based on hypotonic cell lysis, followed by differential centrifugation. It yields three fractions, a nuclei-enriched fraction, a combined membrane and organelle enriched fraction, and a cytoplasmic fraction. 2. Materials 1. CLB: 10 mM HEPES, 10 mM NaCl, 1 mM KH2PO4, 5 mM NaHCO3, 5 mM EDTA,1 mMCaCl2, 0.5 mM MgCl2, and Complete protease inhibitor (Roche, Penzberg, Germany). 2. TSE: 10 mM Tris, 300 mM sucrose, 1 mM EDTA, 0.1% IGEPAL-CA 630 (v/v), pH 7.5 (see Note 1). 3. 2.5 M s ucrose. 4. PBS: 130 mM NaCl, 7 mM Na2HPO4, 3 mM NaH2PO4, pH 7.4. 5. Motorized Teflon/glass homogenizer. 6. Table top centrifuge (e.g. 5417R, Eppendorf, Hamburg, Germany). 7. Beckman 70Ti rotor and Beckman ultracentrifuge (Beckman Coulter, Krefeld, Germany). 8. Micro-BCA assay (Pierce, Bonn, Germany). 9. 2-D gel clean-up kit (GE Healthcare, Freiburg, Germany). 10. Rehydration buffer: 5 M urea, 2 M thiourea, 2% CHAPS, 2% SB-3-10, 40 mM Tris, trace of bromophenol blue.

3. Methods 3.1. Fractionation of Cultures Cells All steps of the protocol should be performed on ice. The procedure is illustrated in Fig. 1. 1. Harvest ∼8 × 106 cells of a monolayer culture (75 cm2) by trypsination. 2. Incubate the cells in 1 ml of CLB for 5 min on ice. 3. Homogenize the cells with 50 strokes in a motor-driven homogenizer at 250 rpm. Keep the sample on ice. 4. Add 100 ml of 2.5 M sucrose to the sample to restore isotonic condition. 5. Centrifuge the sample at 6,300 × g for 5 min at 4°C. 6. Transfer the supernatant to a new tube. It will be processed in step 11. 7. Homogenize the pellet in 500 µl of CLB /0.25 M sucrose with a 1 ml pipette tip. 8. Centrifuge the sample at 6,300 × g for 5 min at 4°C. 9. Combine the supernatant with the supernatant of step 6.

Subcellular Fractionation of Small Sample Amounts Cultured cells

167

Tissue sample 2 strokes 10 min in hypotonic medium 6 strokes Isotonic restoration

5 min in hypotonic medium 50 strokes Isotonic restoration

Homogenate 10 min*

Pellet 1 (Nuclei & debris)

6,300 × g

Supernatant 1 (Membranes, organelles, cytoplasm) 30 min

5 min

107,000 × g

4,000 × g +

Pellet 2 (Nuclei)

Pellet 3 (Membranes & organelles)

Supernatant 2 (Cytoplasm)

Fig. 1. Schematic illustration of the subcellular fractionation protocol. Conditions of homogenization depend on the starting material, that is, either cultured cells or tissue. After homogenization, two rounds of differential centrifugation with increasing centrifugal force yield three final fractions. * for cultured cells, the first round of centrifugation lasted only 5 min; + for freshly prepared tissue, nuclei should be pelleted at only 1000 × g to prevent cosedimentation with other subcellular compartments.

10. Homogenize the pellet in 1 ml TSE with 30 strokes in a motor-driven homogenizer at 250 rpm. 11. Centrifuge the suspension at 4,000 × g for 5 min at 4°C (see Note 2). 12. Discard the supernatant and repeat steps 10 and 11 until the supernatant remains clear. Resuspend the final pellet in 200 ml of TSE and store it at −20°C until further usage. This suspension represents the nuclei enriched fraction. 13. Sediment the combined supernatants from step 6 and 9 at 107,000 × g for 30 min at 4°C min using a Beckman 70Ti rotor (or equivalent) in a Beckman ultracentrifuge (see Note 3). 14. Transfer the supernatant to a new tube and store it at −20 °C until further usage. It represents the cytosolic fraction. 15. Resuspend the pellet in 80 µl of PBS. It represents the membrane and organelle enriched fraction. Store it at −20°C until further usage.

3.2. Fractionation of Tissue 1. Transfer 500 mg of frozen tissue (e.g. brain) to 1 ml of ice cold CLB. 2. Prehomogenize the tissue by 2 strokes in a glass/Teflon homogenizer on ice.

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3. Incubate the pre-homogenate on ice for 10 min. 4. Homogenize the prehomogenate by 6 strokes in a motorized glass/Teflon homogenizer at 250 rpm (see Note 4). 5. Add 100 ml of 2.5 M sucrose to restore isotonic condition. 6. Centrifuge the sample at 6,300 × g for 10 min at 4°C (see Notes 5 and 6). 7. Transfer the supernatant to a new tube. It will be processed in step 11. 8. Homogenize the pellet in 500 ml of CLB /0.25 M sucrose with a 1 ml pipette tip. 9. Centrifuge the sample at 6,300 × g for 5 min at 4°C 10. Combine the supernatant with the supernatant of step 7. 11. Homogenize the pellet in 1 ml of TSE by 30 strokes in a motor-driven homogenizer at 250 rpm. 12. Centrifuge the sample at 4,000 × g for 5 min at 4°C (see Note 2). 13. Discard the supernatant and repeat steps 7 and 8 until the supernatant remains clear. 14. Resuspend the final pellet in 200 ml of TSE and store it at −20°C until further usage. This suspension represents the nuclei enriched fraction. 15. Sediment the supernatant from steps 7 and 10 at 107,000 × g for 30 min at 4°C min using a Beckman Ti70 rotor (or equivalent) in a Beckman ultracentrifuge (see Note 3). 16. Transfer the supernatant to a new tube and store it at −20°C until further usage. It represents the cytosolic fraction. 17. Resuspend the pellet in 80 ml of PBS. It represents the membrane and organelle enriched fraction. Store it at −20°C until further use.

3.3. Protein Quantification and Clean-Up The following procedure is applied to analyze the samples by conventional two-dimensional gel electrophoresis. 1. Determine the protein concentration of the cytosolic fraction by the Micro BCA Protein Assay Reagent Kit according to the manuracturer’s instructions. Use 1, 2, and 5 ml of the sample and BSA as standard. 2. If required for further analysis, concentrate the sample by acetone precipitation, as outlined in steps 3 to 8. 3. Add 4 times the sample volume of cold (−20°C) acetone to the sample. 4. Mix well and incubate at −20°C for 1 h. 5. Centrifuge the sample 10 min at 14,000 rpm. 6. Decant the supernatant and remove all remaining liquids carefully with a pipette tip. 7. Allow the acetone to evaporate from the open tube for 30 min (see Note 7). 8. Resuspend the pellet in the appropriate volume of the desired buffer (e.g., rehydration buffer). 9. Clean-up nuclear and membrane & organelle fractions using a clean-up kit (e.g., 2-D clean up kit, GE Healthcare, Freiburg, Germany) according to the manufacturer’s instructions. 10. Dissolve the resulting pellets in 200 µl rehydration buffer (see Note 8).

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11. Estimate the protein content of the nuclei and of the membrane and organelles enriched fraction by comparing an aliquot of these fractions with a dilution series of the cytosolic proteins quantified in step 1, after separation in a SDS-PAGE and staining with Coomassie or fluorescent dyes (see Note 9).

4. Notes 1. IGEPAL-CA 630 is identical to Nonidet P40 substitute. 2. This washing step purifies the nuclear pellet by removing nonnuclear membrane vesicles. Omitting this step will increase the yield but will reduce the purity of the fraction in parallel. 3. Alternatively, this step can be performed for 150 min at 14,000 rpm (corresponding to ~21,000 × g) at 4°C in a tabletop centrifuge. This by-passes the requirement of an ultracentriguge without affecting yield or purity. 4. Cultured cells require a higher number of strokes compared to tissue samples, as their cytoskeleton differs from the cytoskeleton in brain tissue. Tissues such as heart, liver or placenta will also need harsher conditions for homogenization. 5. When fresh tissue is used, centrifuge only at 3,000 rpm. 6. The prolonged centrifugation time compared to cultured cells results in an increased recovery of nuclear marker proteins. 7. Do not overdry, as the samples may not dissolve properly afterward. 8. This buffer with non-ionic detergents is required if samples will be separated by isoelectric focusing. for other downstream applications, an SDS containing buffer might be used. This will improve the resuspension of the pellets. Due to the presence of thiourea in the rehydration buffer, most colorimetric quantification methods will not work. References 1. Carninci, P., Kasukawa, T., Katayama, S., Gough, J., Frith, M.C., Maeda, N. et al. (2005) The transcriptional landscape of the mammalian genome. Science 309, 1559–1563. 2. Huber, L.A. (2003) Is proteomics heading in the wrong direction? Nat. Rev. Mol. Cell Biol. 4, 74–80. 3. Vuong, G.L., Weiss, S.M., Kammer, W., Priemer, M., Vingron, M., Nordheim, A. et al. (200) Improved sensitivity proteomics by postharvest alkylation and radioactive labeling of proteins. Electrophoresis 21, 2594–2605. 4. Fountoulakis, M.. (2004) Application of proteomics technologies in the investigation of the brain. Mass Spectrom. Rev. 23, 231–258. 5. Becker, M., Schindler, J., and Nothwang H.G. (2006) Neuroproteomics - the tasks lying ahead. Electrophoresis 27, 2819–2829. 6. Anderson, N.L. and Anderson, N.G. (2002) The human plasma proteome: history, character, and diagnostic prospects. Mol. Cell Proteomics 1, 845–867.

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7. Lottspeich, F. (1999) One genome-different proteomes. Angew Chem. Int. Ed. Engl. 38, 2476–2492. 8. Bisle, B., Schmidt, A., Scheibe, B., Klein, C., Tebbe, A., Kellermann, J. et al. (2006) Quantitative profiling of the membrane proteome in a halophilic archaeon. Mol. Cell Proteomics 5(9),1543–1558. 9. Washburn, M.P., Wolters, D., and Yates, J.R. (2001) Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 19, 242–247. 10. Guillemin, I., Becker, M., Ociepka, K., Friauf, E., and Nothwang, H.G. (2005) A subcellular prefractionation protocol for minute amounts of mammalian cell cultures and tissue. Proteomics 5, 35–45.

20 Nondenaturing Polyacrylamide Gel Electrophoresis of Proteins John M. Walker 1. Introduction SDS-PAGE (Chapter 21) is probably the most commonly used gel electrophoretic system for analyzing proteins. However, it should be stressed that this method separates denatured protein. Sometimes one needs to analyze native, nondenatured proteins, particularly if wanting to identify a protein in the gel by its biological activity (for example, enzyme activity, receptor binding, antibody binding, and so on). On such occasions it is necessary to use a nondenaturing system such as described in this chapter. For example, when purifying an enzyme, a single major band on a gel would suggest a pure enzyme. However this band could still be a contaminant; the enzyme could be present as a weaker (even nonstaining) band on the same gel. Only by showing that the major band had enzyme activity would you be convinced that this band corresponded to your enzyme. The method described here is based on the gel system first described by Davis (1). To enhance resolution a stacking gel can be included (see Chapter 21 for the theory behind the stacking gel system). 2. Materials 1. Stock acrylamide solution: 30 g acrylamide, 0.8 g bis-acrylamide. Make up to 100 mL in distilled water and filter. Stable at 4°C for months (see Note 1). Care: Acrylamide Monomer Is a Neurotoxin. Take care in handling acrylamide (wear gloves) and avoid breathing in acrylamide dust when weighing out. 2. Separating gel buffer: 1.5 M Tris-HCl, pH 8.8. 3. Stacking gel buffer: 0.5 M Tris-HCl, pH 6.8. 4. 10% Ammonium persulfate in water. 5. N,N,N ,N -tetramethylethylenediamine (TEMED).

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6. Sample buffer (5X). Mix the following: a. 15.5 mL of 1 M Tris-HCl pH 6.8; b. 2.5 mL of a 1% solution of bromophenol blue; c. 7 mL of water; and d. 25 mL of glycerol. Solid samples can be dissolved directly in IX sample buffer. Samples already in solution should be diluted accordingly with 5X sample buffer to give a solution that is 1X sample buffer. Do not use protein solutions that are in a strong buffer that is not near to pH 6.8 as it is important that the sample is at the correct pH. For these samples it will be necessary to dialyze against 1X sample buffer. 7. Electrophoresis buffer: Dissolve 3.0 g of Tris base and 14.4 g of glycine in water and adjust the volume to 1 L. The final pH should be 8.3. 8. Protein stain: 0.25 g Coomassie brilliant blue R250 (or PAGE blue 83), 125 mL methanol, 25 mL glacial acetic acid, and 100 mL water. Dissolve the dye in the methanol component first, then add the acid and water. Dye solubility is a problem if a different order is used. Filter the solution if you are concerned about dye solubility. For best results do not reuse the stain. 9. Destaining solution: 100 mL methanol, 100 mL glacial acetic acid, and 800 mL water. 10. A microsyringe for loading samples.

3. Method 1. Set up the gel cassette. 2. To prepare the separating gel (see Note 2) mix the following in a Buchner flask: 7.5 mL stock acrylamide solution, 7.5 mL separating gel buffer, 14.85 mL water, and 150 mL 10% ammonium persulfate. “Degas” this solution under vacuum for about 30 s. This degassing step is necessary to remove dissolved air from the solution, since oxygen can inhibit the polymerization step. Also, if the solution has not been degassed to some extent, bubbles can form in the gel during polymerization, which will ruin the gel. Bubble formation is more of a problem in the higher percentage gels where more heat is liberated during polymerization. 3. Add 15 µL of TEMED and gently swirl the flask to ensure even mixing. The addition of TEMED will initiate the polymerization reaction, and although it will take about 20 min for the gel to set, this time can vary depending on room temperature, so it is advisable to work fairly quickly at this stage. 4. Using a Pasteur (or larger) pipet, transfer the separating gel mixture to the gel cassette by running the solution carefully down one edge between the glass plates. Continue adding this solution until it reaches a position 1 cm from the bottom of the sample loading comb. 5. To ensure that the gel sets with a smooth surface, very carefully run distilled water down one edge into the cassette using a Pasteur pipet. Because of the great difference in density between the water and the gel solution, the water will

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6. 7.

8.

9.

10.

11.

12.

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spread across the surface of the gel without serious mixing. Continue adding water until a layer about 2 mm exists on top of the gel solution. The gel can now be left to set. When set, a very clear refractive index change can be seen between the polymerized gel and overlaying water. While the separating gel is setting, prepare the following stacking gel solution. Mix the following quantities in a Buchner flask: 1.5 mL stock acrylamide solution, 3.0 mL stacking gel buffer, 7.4 mL water, and 100 µL 10% ammonium persulfate. Degas this solution as before. When the separating gel has set, pour off the overlaying water. Add 15 µL of TEMED to the stacking gel solution and use some ( 2 mL) of this solution to wash the surface of the polymerized gel. Discard this wash, then add the stacking gel solution to the gel cassette until the solution reaches the cutaway edge of the gel plate. Place the well-forming comb into this solution and leave to set. This will take about 30 min. Refractive index changes around the comb indicate that the gel has set. It is useful at this stage to mark the positions of the bottoms of the wells on the glass plates with a marker pen. Carefully remove the comb from the stacking gel, remove any spacer from the bottom of the gel cassette, and assemble the cassette in the electrophoresis tank. Fill the top reservoir with electrophoresis buffer ensuring that the buffer fully fills the sample loading wells, and look for any leaks from the top tank. If there are no leaks, fill the bottom tank with electrophoresis buffer, then tilt the apparatus to dispel any bubbles caught under the gel. Samples can now be loaded onto the gel. Place the syringe needle through the buffer and locate it just above the bottom of the well. Slowly deliver the sample ( 5–20 µL) into the well. The dense sample solvent ensures that the sample settles to the bottom of the loading well. Continue in this way to fill all the wells with unknowns or standards, and record the samples loaded. The power pack is now connected to the apparatus and a current of 20–25 mA passed through the gel (constant current) (see Note 3). Ensure that the electrodes are arranged so that the proteins are running to the anode (see Note 4). In the first few minutes the samples will be seen to concentrate as a sharp band as it moves through the stacking gel. (It is actually the bromophenol blue that one is observing, not the protein but, of course, the protein is stacking in the same way.) Continue electrophoresis until the bromophenol blue reaches the bottom of the gel. This will usually take about 3 h. Electrophoresis can now be stopped and the gel removed from the cassette. Remove the stacking gel and immerse the separating gel in stain solution, or proceed to step 13 if you wish to detect enzyme activity (see Notes 5 and 6). Staining should be carried out, with shaking, for a minimum of 2 h and preferably overnight. When the stain is replaced with destain, stronger bands will be immediately apparent and weaker bands will appear as the gel destains. Destaining can be speeded up by using a foam bung, such as those used in microbiological flasks. Place the bung in the destain and squeeze it a few times to expel air bubbles and ensure the bung is fully wetted. The bung rapidly absorbs dye, thus speeding up the destaining process.

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13. If proteins are to be detected by their biological activity, duplicate samples should be run. One set of samples should be stained for protein and the other set for activity. Most commonly one would be looking for enzyme activity in the gel. This is achieved by washing the gel in an appropriate enzyme substrate solution that results in a colored product appearing in the gel at the site of the enzyme activity (see Note 7).

4. Notes 1. The stock acrylamide used here is the same as used for SDS gels (see Chapter 21) and may already be availabe in your laboratory. 2. The system described here is for a 7.5% acrylamide gel, which was originally described for the separation of serum proteins (1). Since separation in this system depends on both the native charge on the protein and separation according to size owing to frictional drag as the proteins move through the gel, it is not possible to predict the electrophoretic behavior of a given protein the way that one can on an SDS gel, where separation is based on size alone. A 7.5% gel is a good starting point for unknown proteins. Proteins of mol wt >100,000 should be separated in 3–5% gels. Gels in the range 5–10% will separate proteins in the range 20,000–150,000, and 10–15% gels will separate proteins in the range 10,000–80,000. The separation of smaller polypeptides is described in Chapter 23. To alter the acrylamide concentration, adjust the volume of stock acrylamide solution in Subheading 3., step 2 accordingly, and increase/decrease the water component to allow for the change in volume. For example, to make a 5% gel change the stock acrylamide to 5 mL and increase the water to 17.35 mL. The final volume is still 30 mL, so 5 mL of the 30% stock acrylamide solution has been diluted in 30 mL to give a 5% acrylamide solution. 3. Because one is separating native proteins, it is important that the gel does not heat up too much, since this could denature the protein in the gel. It is advisable therefore to run the gel in the cold room, or to circulate the buffer through a cooling coil in ice. (Many gel apparatus are designed such that the electrode buffer cools the gel plates.) If heating is thought to be a problem it is also worthwhile to try running the gel at a lower current for a longer time. 4. This separating gel system is run at pH 8.8. At this pH most proteins will have a negative charge and will run to the anode. However, it must be noted that any basic proteins will migrate in the opposite direction and will be lost from the gel. Basic proteins are best analyzed under acid conditions, as described in Chapters 27 and 28. 5. It is important to note that concentration in the stacking gel may cause aggregation and precipitation of proteins. Also, the pH of the stacking gel (pH

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6.8) may affect the activity of the protein of interest. If this is thought to be a problem (e.g., the protein cannot be detected on the gel), prepare the gel without a stacking gel. Resolution of proteins will not be quite so good, but will be sufficient for most uses. 6. If the buffer system described here is unsuitable (e.g., the protein of interest does not electrophorese into the gel because it has the incorrect charge, or precipitates in the buffer, or the buffer is incompatible with your detection system) then one can try different buffer systems (without a stacking gel). A comprehensive list of alternative buffer systems has been published (2). 7. The most convenient substrates for detecting enzymes in gels are small molecules that freely diffuse into the gel and are converted by the enzyme to a colored orfluorescent product within the gel. However, for many enzymes such convenient substrates do not exist, and it is necessary to design a linked assay where one includes an enzyme together with the substrate such that the products of the enzymatic reaction of interest is converted to a detectable product by the enzyme included with the substrate. Such linked assays may require the use of up to two or three enzymes and substrates to produce a detectable product. In these cases the product is usually formed on the surface of the gel because the coupling enzymes cannot easily diffuse into the gel. In this case the zymogram technique is used where the substrate mix is added to a cooled (but not solidified) solution of agarose (1%) in the appropriate buffer. This is quickly poured over the solid gel where it quickly sets on the gel. The product of the enzyme assay is therefore formed at the gel–gel interface and does not get washed away. A number of review articles have been published which described methods for detecting enzymes in gels (3–7). A very useful list also appears as an appendix in ref. 8. References 1. Davis, B. J. (1964) Disc electrophoresis II—method and application to human serum proteins. Ann. NY Acad. Sci. 121, 404–427. 2. Andrews, A. T. (1986) Electrophoresis: Theory, Techniques, and Biochemical and Clinical Applications. Clarendon, Oxford, UK. 3. Shaw, C. R. and Prasad, R. (1970) Gel electrophoresis of enzymes—a compilation of recipes. Biochem. Genet. 4, 297–320. 4. Shaw, C. R. and Koen, A. L. (1968) Starch gel zone electrophoresis of enzymes, in Chromatographic and Electrophoretic Techniques, vol. 2 (Smith, I., ed.), Heinemann, London, pp. 332–359. 5. Harris, H. and Hopkinson, D. A. (eds.) (1976) Handbook of Enzyme Electrophoresis in Human Genetics. North-Holland, Amsterdam. 6. Gabriel, O. (1971) Locating enymes on gels, in Methods in Enzymology, vol. 22 (Colowick, S. P. and Kaplan, N. O., eds.), Academic, New York, p. 578.

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7. Gabriel, O. and Gersten, D. M. (1992) Staining for enzymatic activity after gel electrophoresis. I. Analyt. Biochem. 203, 1–21. 8. Hames, B. D. and Rickwood, D. (1990) Gel Electrophoresis of Proteins, 2nd ed., IRL, Oxford and Washington.

21 SDS Polyacrylamide Gel Electrophoresis of Proteins John M. Walker 1. Introduction SDS-PAGE is the most widely used method for qualitatively analyzing protein mixtures. It is particularly useful for monitoring protein purification, and because the method is based on the separation of proteins according to size, the method can also be used to determine the relative molecular mass of proteins (see Note 14). 1.1. Formation of Polyacrylamide Gels Crosslinked polyacrylamide gels are formed from the polymerization of acrylamide monomer in the presence of smaller amounts of N,N ¢ -methylenebis-acrylamide (normally referred to as “bis-acrylamide”) (Fig. 1). Note that bisacrylamide is essentially two acrylamide molecules linked by a methylene group and is used as a crosslinking agent. Acrylamide monomer is polymerized in a head-to-tail fashion into long chains, and occasionally a bis-acrylamide molecule is built into the growing chain, thus introducing a second site for chain extension. Proceeding in this way, a crosslinked matrix of fairly well-defined structure is formed (Fig. 1). The polymerization of acrylamide is an example of free-radical catalysis, and is initiated by the addition of ammonium persulfate and the base N,N,N ¢,N ¢-tetramethylenediamine (TEMED). TEMED catalyzes the decomposition of the persulfate ion to give a free radical (i.e., a molecule with an unpaired electron): S2O82– + e– → SO42– + SO4– •

(1)

If this free radical is represented as R• (where the dot represents an unpaired electron) and M as an acrylamide monomer molecule, then the polymerization can be represented as follows: From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Polymerization of acrylamide.

R• + M → RM• RM• + M → RMM• • RMM + M → RMMM•, and so forth

(2)

In this way, long chains of acrylamide are built up, being crosslinked by the introduction of the occasional bis-acrylamide molecule into the growing chain. Oxygen “mops up” free radicals, and therefore the gel mixture is normally degassed (the solutions are briefly placed under vacuum to remove loosely dissolved oxygen) prior to addition of the catalyst. 1.2. The Use of Stacking Gels For both SDS and buffer gels samples may be applied directly to the top of the gel in which protein separation is to occur (the separating gel). However, in these cases, the sharpness of the protein bands produced in the gel is limited by the size (volume) of the sample applied to the gel. Basically the separated bands will be as broad (or broader, owing to diffusion) as the sample band applied to the gel. For some work, this may be acceptable, but most workers require better resolution than this. This can be achieved by polymerizing a short stacking gel on top of the separating gel. The purpose of this stacking gel is to concentrate the protein sample into a sharp band before it enters the main separating gel,

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thus giving sharper protein bands in the separating gel. This modification allows relatively large sample volumes to be applied to the gel without any loss of resolution. The stacking gel has a very large pore size (4% acrylamide) which allows the proteins to move freely and concentrate, or stack under the effect of the electric field. Sample concentration is produced by isotachophoresis of the sample in the stacking gel. The band-sharpening effect (isotachophoresis) relies on the fact that the negatively charged glycinate ions (in the reservoir buffer) have a lower electrophoretic mobility than the protein–SDS complexes. which in turn, have lower mobility than the Cl− ions if they are in a region of higher field strength. Field strength is inversely proportional to conductivity, which is proportional to concentration. The result is that the three species of interest adjust their concentrations so that [Cl−] > [protein-SDS] > [glycinate]. There are only a small quantity of protein–SDS complexes, so they concentrate in a very tight band between the glycinate and Cl− ion boundaries. Once the glycinate reaches the separating gel, it becomes more fully ionized in the higher pH environment and its mobility increases. (The pH of the stacking gel is 6.8 and that of the separating gel is 8.8.) Thus, the interface between glycinate and the Cl− ions leaves behind the protein–SDS complexes, which are left to electrophorese at their own rates. A more detailed description of the theory of isotachophoresis and electrophoresis generally is given in ref. 1. 1.3. SDS-PAGE Samples to be run on SDS-PAGE are first boiled for 5 min in sample buffer containing β-mercaptoethanol and SDS. The mercaptoethanol reduces any disulfide bridges present that are holding together the protein tertiary structure. SDS (CH3-[CH2]10 - CH2OSO3 – Na+) is an anionic detergent and binds strongly to, and denatures, the protein. Each protein in the mixture is therefore fully denatured by this treatment and opens up into a rod-shaped structure with a series of negatively charged SDS molecules along the polypeptide chain. On average, one SDS molecule binds for every two amino acid residues. The original native charge on the molecule is therefore completely swamped by the SDS molecules. The sample buffer also contains an ionizable tracking dye usually bromophenol blue that allows the electrophoretic run to be monitored, and sucrose or glycerol which gives the sample solution density, thus allowing the sample to settle easily through the electrophoresis buffer to the bottom when injected into the loading well. When the main separating gel has been poured between the glass plates and allowed to set, a shorter stacking gel is poured on top of the separating gel, and it is into this gel that the wells are formed and the proteins loaded. Once all samples are loaded, a current is passed through the gel. Once the protein samples have passed through the stacking gel and have entered the separating gel, the negatively charged protein–SDS complexes continue to move toward the

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anode, and because they have the same charge per unit length they travel into the separating gel under the applied electric field with the same mobility. However, as they pass through the separating gel the proteins separate, owing to the molecular sieving properties of the gel. Quite simply, the smaller the protein, the more easily it can pass through the pores of the gel, whereas large proteins are successively retarded by frictional resistance owing to the sieving effect of the gel. Being a small molecule, the bromophenol blue dye is totally unretarded and therefore indicates the electrophoresis front. When the dye reaches the bottom of the gel the current is turned off and the gel is removed from between the glass plates, shaken in an appropriate stain solution (usually Coomassie brilliant blue) for a few hours, and then washed in destain solution overnight. The destain solution removes unbound background dye from the gel, leaving stained proteins visible as blue bands on a clear background. A typical large format gel would take about 1 h to prepare and set, 3 h to run at 30 mA, and have a staining time of 2–3 h with an overnight destain. Minigels (e.g., Bio-Rad minigel) run at 200 V. Constant voltage can run in about 40 min, and require only 1 h staining. Most bands can be seen within 1 h of destaining. Vertical slab gels are invariably run since this allows up to 20 different samples to be loaded onto a single gel. 2. Materials 1. Stock acrylamide solution: 30% acrylamide, 0.8% bis-acrylamide. Filter through Whatman No. 1 filter paper and store at 4°C (see Note 1). 2. Buffers: a. 1.875 M Tris-HCl, pH 8.8. b. 0.6 M Tris-HCl, pH 6.8. 3. 10% Ammonium persulfate. Make fresh. 4. 10% SDS (see Note 2). 5. TEMED. 6. Electrophoresis buffer: Tris (12 g), glycine (57.6 g), and SDS (2.0 g). Make up to 2 L with water. No pH adjustment is necessary. 7. Sample buffer (see Notes 3 and 4):

β

0.6 M Tris-HCl, pH 6.8 SDS Sucrose -Mercaptoethanol Bromophenol blue, 0.5% stock

5.0 mL 0.5 g 5.0 g 0.25 mL 5.0 mL

Make up to 50 mL with distilled water. 8. Protein stain: 0.1% Coomassie brilliant blue R250 in 50% methanol, 10% glacial acetic acid. Dissolve the dye in the methanol and water component first, and then add the acetic acid. Filter the final solution through Whatman No. 1 filter paper if necessary.

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9. Destain: 10% methanol, 7% glacial acetic acid. 10. Microsyringe for loading samples. Micropipet tips that are drawn out to give a fine tip are also commercially available.

3. Method The system of buffers used in the gel system described below is that of Laemmli (2). 1. Samples to be run are first denatured in sample buffer by heating to 95–100°C for 5 min ( see Note 3). 2. Clean the internal surfaces of the gel plates with detergent or methylated spirits, dry, then join the gel plates together to form the cassette, and clamp it in a vertical position. The exact manner of forming the cassette will depend on the type of design being used. 3. Mix the following in a 250-mL Buchner flask (see Note 5):

1.875 M Tris-HCl, pH 8.8 Water Stock a crylamide 10% SD S Ammonium pe rsulfate ( 10%)

For 15% gels

For 10% gels

8.0 mL 11.4 mL 20.0 mL 0.4 mL 0.2 mL

8.0 mL 18.1 mL 13.3 mL 0.4 mL 0.2 mL

4. “Degas” this solution under vacuum for about 30 s. Some frothing will be observed, and one should not worry if some of the froth is lost down the vacuum tube: you are only losing a very small amount of liquid (see Note 6). 5. Add 14 µL of TEMED, and gently swirl the flask to ensure even mixing. The addition of TEMED will initiate the polymerization reaction and although it will take about 15 min for the gel to set, this time can vary depending on room temperature, so it is advisable to work fairly quickly at this stage. 6. Using a Pasteur (or larger) pipet transfer this separating gel mixture to the gel cassette by running the solution carefully down one edge between the glass plates. Continue adding this solution until it reaches a position 1 cm from the bottom of the comb that will form the loading wells. Once this is completed, you will find excess gel solution remaining in your flask. Dispose of this in an appropriate waste container not down the sink. 7. To ensure that the gel sets with a smooth surface very carefully run distilled water down one edge into the cassette using a Pasteur pipet. Because of the great difference in density between the water and the gel solution the water will spread across the surface of the gel without serious mixing. Continue adding water until a layer of about 2 mm exists on top of the gel solution (see Notes 7 and 8). 8. The gel can now be left to set. As the gel sets, heat is evolved and can be detected by carefully touching the gel plates. When set, a very clear refractive index change can be seen between the polymerized gel and overlaying water.

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9. While the separating gel is setting prepare the following stacking gel (4°C) solution. Mix the following in a 100-mL Buchner flask (see Notes 8 and 9): 0.6 M Tris-HCl, pH 6.8 Stock acrylamide Water 10% SDS Ammonium persulfate (10%)

1.0 mL 1.35 mL 7.5 mL 0.1 mL 0.05 mL

Degas this solution as before. 10. When the separating gel has set, pour off the overlaying water. Add 14 µL of TEMED to the stacking gel solution and use some (~2 mL) of this solution to wash the surface of the polymerized gel. Discard this wash, and then add the stacking gel solution to the gel cassette until the solution reaches the cutaway edge of the gel plate. Place the well-forming comb into this solution, and leave to set. This will take about 20 min. Refractive index changes around the comb indicate that the gel has set. It is useful at this stage to mark the positions of the bottoms of the wells on the glass plates with a marker pen to facilitate loading of the samples (see also Note 9). 11. Carefully remove the comb from the stacking gel, and then rinse out any nonpolymerized acrylamide solution from the wells using electrophoresis buffer. Remove any spacer from the bottom of the gel cassette, and assemble the cassette in the electrophoresis tank. Fill the top reservoir with electrophoresis buffer, and look for any leaks from the top tank. If there are no leaks fill the bottom tank with electrophoresis buffer, and then tilt the apparatus to dispel any bubbles caught under the gel. 12. Samples can now be loaded onto the gel. Place the syringe needle through the buffer and locate it just above the bottom of the well. Slowly deliver the sample into the well. Five- to 10-µL samples are appropriate for most gels. The dense sample buffer ensures that the sample settles to the bottom of the loading well (see Note 10). Continue in this way to fill all the wells with unknowns or standards, and record the samples loaded. 13. Connect the power pack to the apparatus, and pass a current of 30 mA through the gel (constant current) for large format gels, or 200 V (constant voltage) for minigels (Bio-Rad). Ensure your electrodes have correct polarity: all proteins will travel to the anode (+). In the first few minutes, the samples will be seen to concentrate as a sharp band as it moves through the stacking gel. (It is actually the bromophenol blue that one is observing not the protein, but of course the protein is stacking in the same way.) Continue electrophoresis until the bromophenol blue reaches the bottom of the gel. This will take 2.5–3.0 h for large format gels (16 cm × 16 cm) and about 40 min for minigels (10 cm × 7 cm) (see Note 11). 14. Dismantle the gel apparatus, pry open the gel plates, remove the gel, discard the stacking gel, and place the separating gel in stain solution. 15. Staining should be carried out with shaking, for a minimum of 2 h. When the stain is replaced with destain, stronger bands will be immediately apparent, and weaker bands will appear as the gel destains (see Notes 12 and 13).

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4. Notes 1. Acrylamide is a potential neurotoxin and should be treated with great care. Its effects are cumulative, and therefore, regular users are at greatest risk. In particular, take care when weighing out acrylamide. Do this in a fume hood, and wear an appropriate face mask. 2. SDS come out of solution at low temperature, and this can even occur in a relatively cold laboratory. If this happens, simply warm up the bottle in a water bath. Store at room temperature. 3. Solid samples can be dissolved directly in sample buffer. Pure proteins or simple mixtures should be dissolved at 1–0.5 mg/mL. For more complex samples suitable concentrations must be determined by trial and error. For samples already in solution dilute them with an equal volume of doublestrength sample buffer. Do not use protein solutions that are in a strong buffer, that is, not near pH 6.5, since it is important that the sample be at the correct pH. For these samples, it will be necessary to dialyze them first. Should the sample solvent turn from blue to yellow, this is a clear indication that your sample is acidic. 4. The β-mercaptoethanol is essential for disrupting disulfide bridges in proteins. However, exposure to oxygen in the air means that the reducing power of β-mercaptoethanol in the sample buffer decreases with time. Every couple of weeks, therefore, mercaptoethanol should be added to the stock solution or the solution remade. Similarly protein samples that have been prepared in sample buffer and stored frozen should, before being rerun at a later date, have further mercaptoethanol added. 5. Typically, the separating gel used by most workers is a 15% polyacrylamide gel. This give a gel of a certain pore size in which proteins of relative molecular mass (Mr) 10,000 move through the gel relatively unhindered, whereas proteins of 100,000 can only just enter the pores of this gel. Gels of 15% polyacrylamide are therefore useful for separating proteins in the range of 100,000–10,000. However, a protein of 150,000 for example, would be unable to enter a 15% gel. In this case, a larger-pored gel (e.g., a 10% or even 7.5% gel) would be used so that the protein could now enter the gel, and be stained and identified. It is obvious, therefore, that the choice of gel to be used depends on the size of the protein being studied. If proteins covering a wide range of mol-wt values need to be separated, then the use of a gradient gel is more appropriate (see Chapter 32). 6. Degassing helps prevent oxygen in the solution from “mopping up” free radicals and inhibiting polymerization although this problem could be overcome by the alternative approach of increasing the concentration of catalyst. However, the polymerization process is an exothermic one. For 15% gels, the heat liberated can result in the formation of small air bubbles

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7.

8.

9.

10.

11.

12.

13.

Walker in the gel (this is not usually a problem for gels of 10% or less where much less heat is liberated). It is advisable to carry out degassing as a matter of routine. An alternative approach is to add a water-immiscible organic solvent, such as isobutanol, to the top of the gel. Less caution is obviously needed when adding this, although if using this approach, this step should be carried out in a fume cupboard, not in the open laboratory. To save time some workers prefer to add the stacking gel solution directly and carefully to the top of the separating gel, i.e., the overlaying step (step 7) is omitted, the stacking gel solution itself providing the role of the overlaying solution. Some workers include a small amount of bromophenol blue in this gel mix. This give a stacking gel that has a pale blue color, thus allowing the loading wells to be easily identified. Even if the sample is loaded with too much vigor, such that it mixes extensively with the buffer in the well, this is not a problem, since the stacking gel system will still concentrate the sample. When analyzing a sample for the first time, it is sensible to stop the run when the dye reaches the bottom of the gel, because there may be low mol-wt proteins that are running close to the dye, and these would be lost if electrophoresis was continued after the dye had run off the end of the gel. However, often one will find that the proteins being separated are only in the top two-thirds of the gel. In this case, in future runs, the dye would be run off the bottom of the gel, and electrophoresis carried out for a further 30 min to 1 h to allow proteins to separate across the full length of the gel thus increasing the separation of bands. Normally, destain solution needs to be replaced at regular intervals since a simple equilibrium is quickly set up between the concentration of stain in the gel and destain solution, after which no further destaining takes place. To speed up this process and also save on destain solution, it is convenient to place some solid material in with the destain that will absorb the Coomassie dye as it elutes from the gel. We use a foam bung such as that used in culture flasks (ensure it is well wetted by expelling all air in the bung by squeezing it many times in the destain solution), although many other materials can be used (e.g., polystyrene packaging foam). It is generally accepted that a very faint protein band detected by Coomassie brilliant blue, is equivalent to about 0.1 µg (100 ng) of protein. Such sensitivity is suitable for many people’s work. However if no protein bands are observed or greater staining is required, then silver staining (Chapter 48) can be further carried out on the gel.

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14. Because the principle of this technique is the separation of proteins based on size differences, by running calibration proteins of known molecular weight on the same gel run as your unknown protein, the molecular weight of the unknown protein can be determined. For most proteins a plot of log10 molecular mass vs relative mobility provides a straight line graph, although one must be aware that for any given gel concentration this relationship is only linear over a limited range of molecular masses. As an approximate guide, using the system described here, the linear relationship is true over the following ranges: 15% acrylamide, 10,000–50,000; 10% acrylamide 15,000–70,000; 5% acrylamide 60,000–200,000. It should be stressed that this relationship only holds true for proteins that bind SDS in a constant weight ratio. This is true of many proteins but some proteins for example, highly basic proteins, may run differently than would be expected on the basis of their known molecular weight. In the case of the histones, which are highly basic proteins, they migrate more slowly than expected, presumably because of a reduced overall negative charge on the protein owing to their high proportion of positively-charged amino acids. Glycoproteins also tend to run anomalously presumably because the SDS only binds to the polypeptide part of the molecule. To determine the molecular weight of an unknown protein the relative mobilities (Rf ) of the standard proteins are determined and a graph of log molecular weight vs Rf plotted. Rf = (distance migrated by proteins/distance migrated by dye)

(3)

Mixtures of standard mol-wt markers for use on SDS gels are available from a range of suppliers. The Rf of the unknown protein is then determined and the logMW (and hence molecular weight) determined from the graph. A more detailed description of protein mol-wt determination on SDS gels is described in refs. 1 and 3. References 1. Deyl, Z. (1979) Electrophoresis: A Survey of Techniques and Applications. Part A Techniques. Elsevier, Amsterdam. 2. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 3. Hames, B. D. and Rickwood, D. (eds.) (1990) Gel Electrophoresis of Proteins—A Practical Approach. IRL, Oxford University Press, Oxford.

22 Gradient SDS Polyacrylamide Gel Electrophoresis of Proteins John M. Walker 1. Introduction The preparation of fixed-concentration polyacrylamide gels has been described in Chapters 20 and 21. However, the use of polyacrylamide gels that have a gradient of increasing acrylamide concentration (and hence decreasing pore size) can sometimes have advantages over fixed-concentration acrylamide gels. During electrophoresis in gradient gels, proteins migrate until the decreasing pore size impedes further progress. Once the “pore limit” is reached, the protein banding pattern does not change appreciably with time, although migration does not cease completely (1). There are two main advantages of gradient gels over linear gels. First, a much greater range of protein Mr values can be separated than on a fixed-percentage gel. In a complex mixture, very low-mol-wt proteins travel freely through the gel to begin with, and start to resolve when they reach the smaller pore size toward the lower part of the gel. Much larger proteins, on the other hand, can still enter the gel but start to separate immediately owing to the sieving effect of the gel. The second advantage of gradient gels is that proteins with very similar Mr values may be resolved, which otherwise cannot resolve in fixed percentage gels. As each protein moves through the gel, the pore size become smaller until the protein reaches its pore size limit. The pore size in the gel is now too small to allow passage of the protein, and the protein sample stacks up at this point as a sharp band. A similar-sized protein, but with slightly lower Mr, will be able to travel a little further through the gel before reaching its pore size limit, at which point it will form a sharp band. These two proteins, of slightly different Mr values, therefore separate as two, close, sharp bands.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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The usual limits of gradient gels are 3–30% acrylamide in linear or concave gradients. The choice of range will of course depend on the size of proteins being fractionated. The system described here is for a 5–20% linear gradient using SDS polyacrylamide gel electrophoresis. The theory of SDS polyacrylamide gel electrophoresis has been decribed in Chapter 21. 2. Materials 1. Stock acrylamide solution: 30% acrylamide, 0.8% bis-acrylamide. Dissolve 75 g of acrylamide and 2.0 g of N,N′ -methylene bis-acrylamide in about 150 mL of water. Filter and make the volume to 250 mL. Store at 4°C. The solution is stable for months. 2. Buffers: a. 1.875 M Tris-HCl, pH 8.8. b. 0.6 M Tris-HCl, pH 6.8. Store at 4°C. 3. Ammonium persulfate solution (10% [w/v]). Make fresh as required. 4. SDS solution (10% [w/v]). Stable at room temperature. In cold conditions, the SDS can come out of solution, but may be redissolved by warming. 5. N,N,N′ ,N′ -Tetramethylene diamine (TEMED). 6. Gradient forming apparatus (see Fig. 1). Reservoirs with dimensions of 2.5 cm id and 5.0 cm height are suitable. The two reservoirs of the gradient former should be linked by flexible tubing to allow them to be moved independently. This is necessary since although equal volumes are placed in each reservoir, the solutions differ in their densities and the relative positions of A and B have to be adjusted to balance the two solutions when the connecting clamp is opened (see Note 3).

Fig. 1. Gradient forming apparatus.

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3. Method 1. Prepare the following solutions: 1.875 M Tris-HCl, pH 8.8 Water Stock acrylamide, 30% 10% SDS Ammonium persulfate (10%) Sucrose

Solution A, mL 3.0 9.3 2.5 0.15 0.05 —

Solution B, mL 3.0 0.6 10.0 0.15 0.05 2.2 g (equivalent to 1.2 mL volume)

2. Degas each solution under vacuum for about 30 s and then, when you are ready to form the gradient, add TEMED (12 µL) to each solution. 3. Once the TEMED is added and mixed in, pour solutions A and B into the appropriate reservoirs (see Fig. 1.) 4. With the stirrer stirring, fractionally open the connection between A and B and adjust the relative heights of A and B such that there is no flow of liquid between the two reservoirs (easily seen because of the difference in densities). Do not worry if there is some mixing between reservoirs—this is inevitable. 5. When the levels are balanced, completely open the connection between A and B, turn the pump on, and fill the gel apparatus by running the gel solution down one edge of the gel slab. Surprisingly, very little mixing within the gradient occurs using this method. A pump speed of about 5 mL/min is suitable. If a pump is not available, the gradient may be run into the gel under gravity. 6. When the level of the gel reaches about 3 cm from the top of the gel slab, connect the pump to distilled water, reduce pump speed, and overlay the gel with 2–3 mm of water. 7. The gradient gel is now left to set for 30 min. Remember to rinse out the gradient former before the remaining gel solution sets in it. 8. When the separating gel has set, prepare a stacking gel by mixing the following: a. 1.0 mL 0.6 M Tris-HCl, pH 6.8; b. 1.35 mL Stock acrylamide; c. 7.5 mL Water; d. 0.1 mL 10% SDS; e. 0.05 mL Ammonium persulfate (10%). 9. Degas this mixture under vacuum for 30 s and then add TEMED (12 µL). 10. Pour off the water overlayering the gel and wash the gel surface with about 2 mL of stacking gel solution and then discard this solution. 11. The gel slab is now filled to the top of the plates with stacking gel solution and the well-forming comb placed in position (see Chapter 21). 12. When the stacking gel has set (~15 min), carefully remove the comb. The gel is now ready for running. The conditions of running and sample preparation are exactly as described for SDS gel electrophoresis in Chapter 21.

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Fig. 2. Diagrammatic representation of a method for producing a gradient using a two-channel peristaltic pump. Reservoir B has the high percentage acrylamide concentration, reservoir A the lower.

4. Notes 1. The total volume of liquid in reservoirs A and B should be chosen such that it approximates to the volume available between the gel plates. However, allowance must be made for some liquid remaining in the reservoirs and tubing. 2. As well as a gradient in acrylamide concentration, a density gradient of sucrose (glycerol could also be used) is included to minimize mixing by convectional disturbances caused by heat evolved during polymerization. Some workers avoid this problem by also including a gradient of ammienium persulfate to ensure that polymerization occurs first at the top of the gel, progressing to the bottom. However, we have not found this to be necessary in our laboratory.

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3. The production of a linear gradient has been described in this chapter. However, the same gradient mixed can be used to produce a concave (exponential) gradient. This concave gradient provides a very shallow gradient in the top half of the gel such that the percentage of acrylamide only varies from about 5–7% over the first half of the gel. The gradient then increases much more rapidly from 7–20% over the next half of the gel. The shallow part of the gradient allows high-mol-wt proteins of similar size to sufficiently resolve while at the same time still allowing lower mol-wt proteins to separate lower down the gradient. To produce a concave gradient, place 7.5 mL of solution B in reservoir B, then tightly stopper this reservoir with a rubber bung. Equalize the pressure in the chamber by briefly inserting a syringe needle through the bung. Now place 22.5 mL of solution A in reservoir A, open the connector between the two chambers, and commence pouring the gel. The volume of reservoir B will be seen to remain constant as liquid for reservoir A is drawn into this reservoir and diluted. 4. We have described the production of a linear gradient using a purpose built gradient mixer. However, it is not necessary to purchase this since the simple arrangement, shown in Fig. 2 using just flasks or beakers, a stirrer, and a dual channel peristaltic pump, can just as easily be used. Reference 1. Margolis, J. and Kenrick, K. G. (1967) Nature (London) 214, 1334.

23 SDS-Polyacrylamide Gel Electrophoresis of Peptides Ralph C. Judd 1. Introduction Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) has proven to be among the most useful tools yet developed in the area of molecular biology. The discontinuous buffer system, first described by Laemmli (1), has made it possible to separate, visualize, and compare readily the component parts of complex mixtures of molecules (e.g., tissues, cells). SDS-PAGE separation of proteins and peptides makes it possible to quantify the amount of a particular protein/peptide in a sample, obtain fairly reliable molecular mass information, and, by combining SDS-PAGE with immunoelectroblotting, evaluate the antigenicity of proteins and peptides. SDS-PAGE is both a powerful separation system and a reliable preparative purification technique (2; and see Chapter 21). Parameters influencing the resolution of proteins or peptides separated by SDS-PAGE include the ratio of acrylamide to crosslinker (bis-acrylamide), the percentage of acrylamide/crosslinker used to form the stacking and separation gels, the pH of (and the components in) the stacking and separation buffers, and the method of sample preparation. Systems employing glycine in the running buffers (e.g., Laemmli [1], Dreyfuss et al. [3]) can resolve proteins ranging in molecular mass from over 200,000 Daltons (200 kDa) down to about 3 kDa. Separation of proteins and peptides below 3 kDa necessitates slightly different procedures to obtain reliable molecular masses and to prevent band broadening. Further, the increased use of SDS-PAGE to purify proteins and peptides for N-terminal sequence analysis demands that glycine, which interferes significantly with automated sequence technology, be replaced with noninterfering buffer components.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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This chapter describes a modification of the tricine gel system of Schagger and von Jagow (4) by which peptides as small as 500 Daltons can be separated. This makes it possible to use SDS-PAGE peptide mapping (see Chapter 104); epitope mapping (5), and protein and peptide separation for N-terminal sequence analyses (6) when extremely small peptide fragments are to be studied. Since all forms of SDS-PAGE are denaturing, they are unsuitable for separation of proteins or peptides to be used in functional analyses (e.g., enzymes, receptors). 2. Materials 2.1. Equipment 1. SDS-PAGE gel apparatus. 2. Power pack. 3. Blotting apparatus.

2.2. Reagents 1. Separating/spacer gel acrylamide (1X crosslinker): 48 g acrylamide, 1.5 g N,N′ methylene-bis-acrylamide. Bring to 100 mL, and then filter through qualitative paper to remove cloudiness (see Note 1). 2. Separating gel acrylamide (2X crosslinker): 48 g acrylamide, 3 g N,N′ -methylenebisacrylamide. Bring to 100 mL, and then filter through qualitative paper to remove cloudiness (see Note 2). 3. Stacking gel acrylamide: 30 g acrylamide, 0.8 g N,N′ -methylene-bis-acrylamide. Bring to 100 mL, and then filter through qualitative paper to remove cloudiness. 4. Separating/spacer gel buffer: 3 M Tris base, 0.3% sodium dodecyl sulfate (see Note 3). Bring to pH 8.9 with HCl. 5. Stacking gel buffer: 1 M Tris-HCl, pH 6.8. 6. Cathode (top) running buffer (10X stock): 1 M Tris base, 1 M tricine, 1% SDS (see Note 3). Dilute 1:10 immediately before use. Do not adjust pH; it will be about 8.25. 7. Anode (bottom) buffer (10X stock): 2 M Tris base. Bring to pH 8.9 with HCl. Dilute 1:10 immediately before use. 8. 0.2 M tetrasodium EDTA. 9. 10% ammonium persulfate (make fresh as required). 10. TEMED. 11. Glycerol. 12. Fixer/destainer: 25% isopropanol, 7% glacial acetic acid in dH2O (v/v/v). 13. 1% Coomassie brilliant blue (CBB) (w/v) in fixer/destainer. 14. Sample solubilization buffer: 2 mL 10% SDS (w/v) in dH2O, 1.0 mL glycerol, 0.625 mL 1 M Tris-HCl, pH 6.8, 6 mL dH2O, bromphenol blue to color. 15. Dithiothreitol. 16. 2% Agarose.

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17. Molecular-mass markers, e.g., low-mol-wt kit (Bio-Rad, Hercules, CA), or equivalent, and peptide molecular-mass markers (Pharmacia Inc., Piscataway, NJ), or equivalent. 18. PVDF (nylon) membranes. 19. Methanol. 20. Blotting transfer buffer: 20 mM phosphate buffer, pH 8.0: 94.7 mL 0.2 M Na2HPO4 stock, 5.3 mL 0.2 M NaH2PO4 stock in 900 mL H2O. 21. Filter paper for blotting (Whatman No. 1), or equivalent. 22. Distilled water (dH2O).

2.3. Gel Recipes 2.3.1. Separating Gel Recipe Add reagents in order given (see Note 4): 6.7 mL water, 10 mL separating/ spacer gel buffer, 10 mL separating/spacer gel acrylamide (1X or 2X crosslinker), 3.2 mL glycerol, 10 µL TEMED, 100 µL 10% ammonium persulfate. 2.3.2. Spacer Gel Recipe Add reagents in order given (see Note 4): 6.9 mL water, 5.0 mL separating/ spacer gel buffer, 3.0 mL separating/spacer gel acrylamide (1X crosslinker only), 5 µL TEMED, 50 µL 10% ammonium persulfate. 2.3.3. Stacking Gel Recipe Add reagents in order given (see Note 4): 10.3 mL water, 1.9 mL stacking gel buffer, 2.5 mL stacking gel acrylamide, 150 µL EDTA, 7.5 µL TEMED, 150 µL 10% ammonium persulfate. 3. Methods 3.1. Sample Solubilization 1. Boil samples in sample solubilization buffer for 10–30 min. Solubilize sample at 1 mg/mL and run 1–2 µL/lane (1–2 µg/lane) (see Note 5). For sequence analysis, as much sample as is practical should be separated.

3.2. Gel Preparation/Electrophoresis 1. Assemble the gel apparatus (see Note 6). Make two marks on the front plate to identify top of separating gel and top of spacer gel (see Note 7). Assuming a well depth of 12 mm, the top of the separating gel should be 3.5 cm down from the top of the back plate, and the spacer gel should be 2 cm down from the top of the back plate, leaving a stacking gel of 8 mm (see Note 8). 2. Combine the reagents to make the separating gel, mix gently, and pipet the solution between the plates to lowest mark on the plate. Overlay the gel solution with

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3.

4.

5.

6.

7.

8.

Judd 2 mL of dH2O by gently running the dH2O down the center of the inside of the front plate. Allow the gel to polymerize for about 20 min. When polymerized, the water–gel interface will be obvious. Pour off the water, and dry between the plates with filter paper. Do not touch the surface of the separating gel with the paper. Combine the reagents to make the spacer gel, mix gently, and pipet the solution between the plates to second mark on the plate. Overlay the solution with 2 mL of dH2O by gently running the dH2O down the center of the inside of the front plate. Allow the gel to polymerize for about 20 min. When polymerized, the water–gel interface will be obvious. Pour off the water, and dry between the plates with filter paper. Do not touch the surface of spacer gel with the paper. Combine the reagents to make the stacking gel and mix gently. Place the well-forming comb between the plates, leaving one end slightly higher than the other. Slowly add the stacking gel solution at the raised end (this allows air bubbles to be pushed up and out from under the comb teeth). When the solution reaches the top of the back plate, gently push the comb all the way down. Check to be sure that no air pockets are trapped beneath the comb. Allow the gel to polymerize for about 20 min. When the stacking gel has polymerized, carefully remove the comb. Straighten any wells that might be crooked with a straightened metal paper clip. Remove the acrylamide at each edge to the depth of the wells. This helps prevent “smiling” of the samples at the edge of the gel. Seal the edges of the gel with 2% agarose. Add freshly diluted cathode running buffer to the top chamber of the gel apparatus until it is 5–10 mm above the top of the gel. Squirt running buffer into each well with a Pasteur pipet to flush out any unpolymerized acrylamide. Check the lower chamber to ensure that no cathode running buffer is leaking from the top chamber, and then fill the bottom chamber with anode buffer. Remove any air bubbles from the under edge of the gel with a benttip Pasteur pipet. The gel is now ready for sample loading. After loading the samples and the molecular-mass markers, connect leads from the power pack to the gel apparatus (the negative lead goes on the top, and the positive lead goes on the bottom). Gels can be run on constant current, constant voltage, or constant power settings. When using the constant current setting, run the gel at 50 mA. The voltage will be between 50 and 100 V at the beginning, and will slowly increase during the run. For a constant voltage setting, begin the electrophoresis at 50 mA. As the run progresses, the amperage will decrease, so adjust the amperage to 50 mA several times during the run or the electrophoresis will be very slow. If running on constant power, set between 5 and 7 W. Voltage and current will vary to maintain the wattage setting. Each system varies, so empirical information should be used to modify the electrophoresis conditions so that electrophoresis is completed in about 4 h (see Note 9). When the dye front reaches the bottom of the gel, turn off the power, disassemble the gel apparatus, and place the gel in 200–300 mL of fixer/destainer. Gently shake for 16 h (see Note 10). Pour off spent fixer/destainer, and add CBB. Gently shake for 30 min. Destain the gel in several changes of fixer/destainer until the background is almost clear. Then place the gel in dH2O, and gently mix until

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the background is completely clear. The peptide bands will become a deep purpleblue. The gel can now be photographed or dried. To store the gel wet, soak the gel in 7% glacial acetic acid for 1 h, and seal in a plastic bag.

Figure 1 demonstrates the molecular mass range of separation of a 1X crosslinker tricine gel. Whole-cell (WC) lysates and 1X and 2X purified (see Chapter 104) 44 kDa proteins of Neisseria gonorrhoeae, Bio-Rad low-mol-wt markers (mw), and Pharmacia peptide markers (pep mw) were separated and stained with CBB. The top of the gel in this figure is at the spacer gel–separating gel interface. Proteins larger than about 100 kDa remained trapped at the spacer gel–separating gel interface, resulting in the bulging of the outside lanes. Smaller proteins all migrated into the gel, but many remained tightly bunched at the top of the separating gel. The effective separation range is below 40 kDa. Comparison of this figure with Fig. 1 in Chapter 104, which shows gonococcal whole cells and the two mol-wt marker preparations separated in a standard 15% Laemmli gel (1), demonstrates the tremendous resolving power of low-mol-wt components by this tricine gel system.

Fig. 1. WC Iysates and 1X and 2X purified 44 kDa protein of Neisseria gonorrhoeae (see Chapter 104, Subheading 3.1.,), Bio-Rad mw (1 µg of each protein), and Pharmacia pep mw (to which the 1.3-kDa protein kinase C substrate peptide [Sigma, St. Louis, MO] was added) (3 µg of each peptide), separated in a 1X crosslinker tricine gel, fixed, and stained with CBB. Molecular masses are given in thousands of daltons.

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3.3. Blotting of Peptides Separated peptides can be electroblotted to PVDF membranes for sequencing or immunological analyses (see Note 11). 1. Before the electrophoresis is complete, prepare enough of the 20 mM sodium phosphate transfer buffer, pH 8.0 (see Note 12) to fill the blotting chamber (usually 2–4 L). Degas about 1 L of transfer buffer for at least 15 min before use. Cut two sheets of filter paper to fit blotting apparatus, and cut a piece of PVDF membrane a little larger than the gel. Place the PVDF membrane in 10 mL of methanol until it is wet (this takes only a few seconds), and then place the membrane in 100 mL of degassed transfer buffer. 2. Following electrophoresis, remove the gel from the gel apparatus, and place it on blotting filter paper that is submersed in the degassed transfer buffer. Immediately overlay the exposed side of the gel with the wetted PVDF membrane, being sure to remove all air pockets between the gel and the membrane. Overlay the PVDF membrane with another piece of blotting filter paper, and place the gel “sandwich” into the blotting chamber using the appropriate spacers and holders. 3. Connect the power pack electrodes to the blotting chamber (the positive electrode goes on the side of the gel having the PVDF membrane). Electrophorese for 16 h at 25 V, 0.8 A. Each system varies, so settings may be somewhat different than those described here. 4. Following electroblotting, disconnect the power, disassemble the blotting chamber, and remove the PVDF membrane from the gel (see Note 13). The PVDF membrane can be processed for immunological analyses or placed in CBB in fixer/destainer to stain the transferred peptides. Remove excess stain by shaking the membrane in several changes of fixer/destainer until background is white. Peptide bands can be excised, rinsed in dH2O, dried, and subjected to N-terminal sequencing.

3.4. Modifications for Peptide Sequencing Peptides to be used in N-terminal sequence analyses must be protected from oxidation, which can block the N-terminus. Several simple precautions can help prevent this common problem. 1. Prepare the separation and spacer gel the day before electrophoresing the peptides. After pouring, overlay the spacer gel with several milliliters of dH2O, and allow the gel to stand overnight at room temperature (see Note 14). 2. On the next day, pour off the water, and dry between the plates with filter paper. Do not touch the gel with the filter paper. Prepare the stacking gel, but use half the amount of 10% ammonium persulfate (see Note 15). Pipet the stacking gel solution between the plates as described above, and allow the stacking gel to polymerize for at least 1 h. Add running buffers as described above, adding 1–2 mg of dithiothreitol to both the upper and lower chambers to scavenge any oxidizers from the buffers and gel. 3. Pre-electrophorese the gel for 15 min. then turn off the power, and load the samples and molecular-mass markers.

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4. Run the gel as described above. 5. Blot the peptides to a PVDF membrane as described above, but add 1–2 mg of dithiothreitol to the blot transfer buffer. 6. Fix and stain as above, again adding 1–2 mg of dithiothreitol to the fixer/destainer, CBB, and dH2O used to rinse the peptide-containing PVDF membrane.

Acknowledgments The author thanks Joan Strange for her assistance in developing this system, Pam Gannon for her assistance, and the Public Health Service, NIH, NIAID (grants RO1 AI21236) and UM Research Grant Program for their continued support. 4. Notes 1. Working range of separation about 40 kDa down to about 1 kDa. 2. Working range of separation about 20 kDa down to less than 500 Dalton. 3. Use electrophoresis grade SDS. If peptide bands remain diffuse, try SDS from BDH (Poole, Dorset, UK). 4. Degassing of gel reagents is not necessary. 5. Coomassie staining can generally visualize a band of 0.5 µg. This may vary considerably based on the properties of the particular peptide (some peptides stain poorly with Coomassie). Some peptides do not bind SDS well and may never migrate exactly right when compared to mol-wt markers. Fortunately, these situations are rare. 6. Protocols are designed for a standard 13 cm × 11 cm × 1.5 mm slab gel. Dimensions and reagent volumes can be proportionally adjusted to accommodate other gel dimensions. 7. Permanent marks with a diamond pencil can be made on the back of the back plate if the plate is dedicated to this gel system. 8. The depth of the spacer gel can be varied from 1 to 2 cm. Trial and error is the only way to determine the appropriate dimension for each system. 9. It is wise to feel the front plate several times during the electrophoresis to check for over-heating. The plate will become pleasantly warm as the run progresses. If it becomes too warm, the plates might break, so turn down the power! 10. Standard-sized gels can be fixed in as little as 4 h with shaking. 11. It is best to blot peptides to PVDF membranes rather than nitrocellulose membranes, since small peptides tend to pass through nitrocellulose without binding. Moreover, peptides immobilized on PVDF membranes can be directly sequenced in automated instrumentation equipped with a “blot cartridge” (6). 12. The pH of the transfer buffer can be varied from 5.7 to 8.0 if transfer is inefficient at pH 8.0 (7). 13. Wear disposable gloves when handling membranes.

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14. Do not refrigerate the gel. It will contract and pull away from the plates, resulting in leaks and poor resolution. 15. Do not pour the stacking gel the day before electrophoresis. It will shrink, allowing the samples to leak from the wells. References 1. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–695. 2. Judd, R. C. (1988) Purification of outer membrane proteins of the Gram negative bacterium Neisseria gonorrhoeae. Analyt. Biochem. 173, 307–316. 3. Dreyfuss, G., Adam, S. A., and Choi, Y. D. (1984) Physical change in cytoplasmic messenger ribonucleoproteins in cells treated with inhibitors of mRNA transcription. Mol. Cell. Biol. 4, 415–423. 4. Schagger, H. and von Jagow, G. (1987) Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Analyt. Biochem. 166, 368–397. 5. Judd, R. C. (1986) Evidence for N-terminal exposure of the PIA subclass of protein I of Neisseria gonorrhoeae. Infect. Immunol. 54, 408–414. 6. Moos, M., Jr. and Nguyen, N. Y. (1988) Reproducible high-yield sequencing of proteins electrophoretically separated and transferred to an inert support. J. Biol. Chem. 263, 6005–6008. 7. Stoll, V. S. and Blanchard, J. S. (1990) Buffers: principles and practice, in Methods in Enzymology, vol. 182, A Guide to Protein Purification (Deutscher, M. P., ed.), Academic, San Diego, CA, pp. 24–38.

24 Separation of Proteins by Blue Native Electrophoresis (BN-PAGE) Elke A. Dian, Joachim Rassow, and Christian Motz 1. Introduction Proteins often work together in protein complexes. For the analysis of the function as well as the structure of proteins it is often of interest to analyze the assembly of such complexes. While protein complexes normally dissociate into their subunits when analyzed by SDS-PAGE (Fig. 1B), nondenaturing electrophoresis (so called native electrophoresis) methods leave the complexes intact. Blue native polyacrylamide gel electrophoresis (BN-PAGE, Fig. 1C), a method developed by Schägger and Jagow (1), has proven to be a valuable tool to analyze the respiratory chain complexes of different organisms (2,3) to investigate the complexes of the import machinery of mitochondria (4–6) or chloroplasts (7), or to analyze the proteome of isolated organelles or whole cellular lysates (8,9). Compared to other methods, BN-PAGE offers some major advantages: (1) It allows the determination of the molecular weight for both positively and negatively charged proteins as well as for membrane proteins. (2) In contrast to gel filtration, BN-PAGE offers the possibility of testing many samples on the same gel (i.e., under identical conditions). (3) Small samples (electrophoretic amounts down to 5 µg) are possible. (4) The electrophoresis can be performed using standard equipment for SDS-PAGE. With other native electrophoresis methods, the mobility of the proteins is mainly determined by their intrinsic charge but also to some extend by their size. Therefore neither the pI nor the molecular weight of the proteins can be exactly determined. The BN-PAGE overcomes this difficulty by binding of the anionic dye Coomassie Blue G-250 and so inducing a charge shift to the proteins. Thus the electrophoretic mobility depends only on the molecular weight.

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B

Sample

SDS-PAGE

Separation of subunits

C

BN-PAGE

Separation of intact complexes according to their molecular weight

Fig. 1. Principle of the separation with different electrophoretic techniques. (A) The proteins to be separated. (B) SDS-PAGE: the subunits are separated according to their size. (C) The intact protein complexes are separated by size.

Schägger et al. (10) showed that the BN-PAGE allows the analysis of molecular masses of most soluble as well as most membrane proteins. Only a few proteins that do not bind the Coomassie dye and have no or almost no negative charge at pH 7.5 showed a significantly different behavior. As a side effect, the negatively charged dye also prevents aggregation of the proteins. While in SDS-PAGE the SDS / protein ratio and thus the charge / mass ratio is constant for all proteins, the ratio of bound dye / protein shows some variation in BN-PAGE. Especially integral membrane proteins show an increase in their apparent molecular weights compared to soluble proteins due to differences in the affinity for detergent or dye molecules. Heuberger et al. (11) showed for different membrane proteins that bound detergents were replaced by dye molecules during electrophoresis. The apparent molecular weight was increased by the factor 1.8 compared to the molecular weight of soluble proteins that served as marker proteins. 2. Materials Prepare all solutions in distilled H2O unless otherwise stated. 1. Electrophoresis chamber: the gel has to be cooled during electrophoresis. Although running the gel in the cold room in a standard electrophoresis chamber may be

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Table 1 Composition of Separating and Stacking Gels Separating gel: acrylamide concentration Components

4%

6%

13 %

16 %

20 %

Stacking gel

AB-mix Gel b uffer Glycerol H 2O APS TEMED

0.75 mL 3 mL Fill t o 9 mL 38 µL 3.8 µL

1.15 mL 3 mL -

2.35 mL 3 mL 1 mL

3 mL 3 mL 1 mL

3.75 mL 3 mL 1 mL

38 µL 3.8 µL

30 µL 3 µL

30 µL 3 µL

30 µL 3 µL

0.3 mL 1.25 mL 2.2 mL 30 µL 3 µL

2. 3. 4. 5. 6. 7. 8. 9. 10.

11. 12. 13. 14.

sufficient, best cooling is achieved using a system with cooling in a water bath, such as the Hoefer SE600 system (see Note 1). A gradient mixer of the appropriate volume for a separating gel (see Table 1 and Note 7). Cathode buffer A: 50 mM Tricine, 15 mM Bis-Tris, 0.02 % coomassie brilliant blue G-250, pH 7.0 at 4°C, do not adjust the pH (see Notes 1 and 2). Cathode buffer B: 50 mM Tricine, 15 mM Bis-Tris, pH 7.0 at 4°C (see Note 1). Anode buffer: 50 mM Bis-Tris, adjust pH 7.0 with HCl (see Note 1). Gel Buffer (3x): 1,5 M ε-aminocaproic acid, 150 mM Bis-Tris (see Note 3). AB-Mix: 49.5 % acrylamide, 1,5 % bisacrylamide. APS: 10 % (w/v) ammoniumpersulfate in H2O. TEMED: N,N,N’,N’-tetrametylethylenediamine. Lysis buffer: 20 mM Tris-HCl, 0.1 mM EDTA, 500 mM ε-aminocaproic acid, 10 % glycerol, pH 7.0 (HCl). Complete with the desired detergent (see Notes 3 – 6) and add PMSF to 1 mM just before use. PMSF: phenylmethylsulfonylfluoride 200 mM in 2-propanol. Sample buffer: 500 mM ε-aminocaproic acid, 100 mM Bis-Tris, 5 % Serva Blue G, pH 7.0 (HCl) (see Note 2). 100 % methanol. Blot buffer: 20 mM Tris, 150 mM glycine, 0.02 % SDS, 20 % methanol, do not adjust the pH.

3. Methods 3.1. General Experiment 3.1.1. Preparing the Gel 1. Prepare the gel assembly according to the manufacturers instructions, 2. Mix the separating gel solutions (Table 1, see Notes 7 and 8) without APS and fill them into a gradient mixer, filling the higher concentration in the exit chamber. Add the APS and TEMED, then pour the gel. Overlay with 2-propanol or 1-butanol. 3. After the separating gel has polymerized, discard the alcohol, then mix and pour the stacking gel (Table 1). Place a comb in the gel to form the wells.

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3.1.2. Sample Preparation The sample preparation depends on whether the proteins to be analyzed are soluble, or membrane proteins. For first experiments start with or twice the amount you would use for a normal SDS-PAGE (see Note 9). Soluble proteins can be used without further preparation by dilution with lysis buffer and addition of 10 % (v/v) sample buffer and proceed with the electrophoresis. Already solubilized membrane proteins can be treated in a similar manner if the detergent does not interfere with the electrophoresis (see Note 10). Ideally, the proteins are precipitated or in membranes. Such samples are treated in the following way: Prior to the electrophoresis these samples are lysed in a buffer containing detergent. The choice of detergent is crucial because too harsh conditions will dissociate the complexes whereas too mild conditions may not be sufficient to solubilize the proteins (see Fig. 2B and Notes 4–6). 1. Reisolate the membranes (vesicles or organelles) or the protein precipitate by centrifugation. Discard the supernatant. 2. Resuspend the samples in 70 µL lysis buffer (i.e. 1 µL per µg protein) containing the detergent and 1 mM PMSF.

A

1 2

3

4

5

B

6

880-

Triton Digitonin 0,15 1 % 1 2 kDa ATP Synthase (dimer) Respiratory chain complexes 660 ATP Synthase (monomer)

-660 440198-

6630-

150-

440

Hsp60 complex

-200 200 150

66

Fig. 2. Separation of protein complexes by BN-PAGE (4–16 % acrylamide gradient), staining with Coomassie. (A) Marker proteins: lane 1: carbonic anhydrase (30 kDa), 2: BSA (66 kDa), 3: alcohol dehydrogenase (150 kDa), 4: β-amylase (200 kDa), 5: apoferritin (440 kDa), 6: thyroglobulin (660 kDa). Note that some of the markers give more than one band; therefore it is not recommended to mix all the markers in one lane. (B) Separation of proteins from isolated yeast mitochondria in BN-PAGE (4–16 % gradient): lane 1: lysis of mitochondria with Triton X-100, lane 2: lysis with digitonin. Note that the large complexes of the respiratory chain are visible only when the mitochondria were lysed in digitonin. The ATP synthase dimer is only visible in digitonin.

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3. Incubate 20 min on ice (see Note 6). 4. Centrifuge 20 min at 20000 g (minimum, use an ultracentrifuge if available) to pellet insoluble aggregates. 5. Discard the pellet (or keep it for lysis control using SDS-PAGE). 6. Add 7 µL of sample buffer (i.e. 10 % of the sample volume) to the supernatant.

3.1.3. Electrophoresis 1. Apply the samples to the gel. 2. Apply the markers solubilized in sample buffer (see Note 11). 3. Overlay the samples with cathode buffer A, assemble the precooled electrophoresis unit and fill the cathode buffer A in the upper buffer chamber. 4. Start the electrophoresis at 200 V until the blue front reaches the separating gel, then set the voltage to 500 V with the current limited to 15 mA per gel (see Note 12). 5. Removing excess dye can be achieved by changing the cathode buffer A to cathode buffer B after about one third of the run. This is recommended when blotting the gel but can be omitted if the gel will be stained after electrophoresis.

3.1.4. Staining The Serva Blue G will not stain all proteins, so it will be necessary to stain the gel according to your standard procedures, e.g. staining with Coomassie Blue R-250 or silver staining. 3.1.5. Western Blotting For western blotting it is best to use polyvinylidene (PVDF) membranes, which will allow removing the dye using organic solvents. Note that, unlike nitrocellulose, PVDF needs an extra preparation. 1. Prepare the PVDF membrane by immersing for a few seconds in 100 % methanol, until the membrane is translucent. Wash with water for another five min. 2. Incubate the membrane a few minutes in blot buffer. The membrane submerges when it is equilibrated. 3. Equilibrate the gel for a few seconds in transfer buffer. 4. Perform the western blotting according to your standard procedures. 5. After blotting, remove the blue dye by washing the membrane with methanol, then rinse with water. 6. The membrane can be stained or used for antibody detection following your standard procedures. Note that if the membrane once gets dry, you will have to repeat step 1 before going on.

3.2. Two-Dimensional Gels 1. After running the native gel, disassemble the glass plates and remove the stacking gel. 2. Cut out the lanes of interest. 3. Place the lane on a glass plate at the usual position of the stacking gel. Leave some space at the side for a marker or a control (see Fig. 3A and Note 13).

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Fig. 3. Two-dimensional gel. (A) Principle; in the first dimension (BN-PAGE, left to right), the protein complexes are separated, in the second dimension (SDS-PAGE, topdown), the subunits and monomers are separated. The subunits of one complex can be found one below the other. Monomers will be found on the dotted hyperbolic line. (B) Example: mitochondria (70 µg protein) lysed in digitonin: 1. ATP synthase (dimer), 2, 3: respiratory chain complexes, 4: ATP Synthase (monomer), 5: Hsp60 complex, 6: excess Coomassie dye

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4. Assemble the glass plates for a standard SDS gel. The excised gel strip will be held in its position by friction. 5. Pour the separating gel. Leave about 0.5–1 cm below the gel strip. Overlay with 2-propanol or 1-butanol (according to your standard procedures). 6. After polymerization, decant the propanol and pour the stacking gel, avoiding air bubbles under the gel strip. This can be achieved by tilting the gel assembly and slowly straighten it while pouring the gel. Place a small comb for one or two wells for marker and control (see Note 13). 7. Place the gel into the electrophoresis apparatus. Apply the marker. If you need reducing conditions, overlay the blue native strip with reducing sample buffer. 8. Perform the electrophoresis according to your standard protocols. The blue dye in the native gel strip will form a thin line. After the line has reached the separating gel, remove the gel strip with a thin spatula. This will avoid an uneven run. 9. The gel can be stained, blotted, and so on, according to your standard procedures.

3.3. Analysis of Organelles or Cell Lysates Blue Native electrophoresis offers the posibility to identify protein complexes in proteomic approaches. Camacho-Carvajal et al. (9) subjected whole cellular lysates to BN-PAGE with subsequent SDS-PAGE and identified various protein complexes using immunoblotting. Werhahn and Braun (8) used an elegant three-dimensional electrophoresis to identify the proteome of the mitochondrial complexes: in a first dimension they subjected mitochondria to BN-PAGE. The resulting bands of the protein complexes were cut out and each was subjected to first dimension isoelectric focusing and subsequent second dimension SDSPAGE, resulting in a set of “classic” two dimensional gels, each representing the proteome of one complex of mitochondrial proteins. 3.4. Analysis of Isolated Protein Complexes Figure 4A shows an assembly experiment with F1 ATPase from Escherichia coli monitored by Blue Native electrophoresis: isolated α, β, and γ subunits were mixed in a stoichometric ratio without (lane 2) and with (lane 3) MgATP and then analyzed by BN-PAGE. Lane 1 shows intact ATPase as a control. Whereas in the absence of MgATP, only a mixture of the subunits can be detected (lane 2), in the presence of MgATP, a larger complex is detected (lane 3, arrow) which corresponds to α3β3γ. Arnold et al. (12) analyzed isolated ATP synthase from yeast with Blue Native electrophoresis. They could show the existence of two different complexes: one with the molecular weight of the ATP synthase, one with the double weight. Using a subsequent second dimension SDS-PAGE, they could identify both complexes as ATP synthase and further identify some proteins which are only present in the dimer.

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Fig. 4. Examples for applications of BN-PAGE: A: BN-PAGE of an assembly experiment with F1 ATPase from Escherichia coli: 1: F1-ATPase, 2: isolated α, β und γ subunits (3:3:1) without MgATP, 3: isolated α, β und γ subunits (3:3:1) with MgATP. The arrow indicates the presence of a reconstituted α3β3γ complex. B: Autoradiogram of a BN-PAGE with mitochondria after the import of radiolabeled γ-subunit (Atp3) and lysis with digitonin. The right sample is treated with antiserum against Atp3. Due to binding of the IgG molecules to the ATP synthase complex, the corresponding bands are shifted from 600 kDa to 750 kDa.

3.5. Using an Antibody Shift to Determine Interaction Partners Antibody binding is not affected by BN-PAGE and can thus be detected by a molecular mass shift of about 150 kDa (the molecular weight of a IgG molecule). Wiedemann et al. (13) used this method to identify components of the assembly intermediates of the TOM complex. This method can be used as an alternative to coimmune precipitation. Dissolve the samples in lysis buffer containing 1 % digitonin and add 1–5 µL polyclonal antiserum against potential interaction partners. Use a sample with the same amount of preimmune serum as control. Incubate 20 min on ice. Centrifuge, then run the blue native gel as described in Subheading 3.1. Detect your protein of interest by western blotting or phosphor imager analysis. Protein bands showing an increase of the molecular weight by 150 or 300 kDa in comparison with the control indicate the binding of one or two IgG molecules and thus the presence of the protein the antibody was raised against in the protein complex of interest (Fig. 4B). 4. Notes 1. The Hoefer SE600 system needs 400 mL of cathode buffer and 4 L of anode buffer, both can be reused up to five times 2. Coomassie Brilliant Blue R-250 cannot be used.

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3. Schägger and Jagow (1) use a concentration of 500 mM ε-aminocaproic acid (EACA) in the gel and 750 mM in the lysis buffer. The concentration, however, can be varied in a wide range (100 mM to 1 M). 4. The choice of detergent is a critical step, as shown in Fig. 2B. For yeast mitochondria, 1 % digitonin will give a good lysis while keeping most protein complexes together. Also Triton X-100 in a range from 0,2 to 2 % may give good results. Any other detergent may be used; it should be selected to extract the proteins and preserve the protein complexes (see Note 6). 5. For determination of the protein pattern of the monomers, 1 % SDS or 8 M urea can be used. 6. The appropriate lysis conditions (lysis time, lysis buffer, detergent) should be determined in a pilot experiment: lyse the samples under different conditions, centrifuge at 20000 g (better: 100000 g) and analyze the pellet and supernatant on a SDS-gel for the protein of interest. Select conditions that keep the protein in the supernatant. 7. A linear gradient range from 4–16 % will give a good resolution from 1000 to 100 kDa, for other molecular weight ranges, gradients between 4 and 20% can be used. For separation of proteins in the range from 50 to 100 kDa, a homogenous gel with 10% acrylamide has proven suitable (14). 8. The volume of 18 mL for the separating gel is convenient for a gel with the dimensions 18 × 16 cm and 1 mm thickness, as purchased with the Hoefer SE600. For other gels, the appropriate volume has to be determined. The use of spacers with more than 1 mm thickness may result in broadening of the protein bands. 9. Protein amounts of 50 to 100 µg per sample for a heterogeneous mixture of proteins and 5 to 30 µg for purified protein are usually satisfactory detectable. For first experiments use twice the protein as you would for a normal experiment analyzed by SDS-PAGE. A high background or smearing on the gel, however, can be reduced by reducing the sample size. 10. It may be possible that some substances in the protein solution may interfere with the electrophoresis, so a precipitation or dialysis with lysis buffer may be necessary. 11. Molecular weight markers for gel filtration calibration (e.g. Sigma MW-GF1000, see Fig. 2A) can be used as markers. Some of the markers may give more than one band, so it is best to run each marker in a separate lane first to see which may be mixed without confusing the bands (e.g., BSA and Apoferritin). 12. The Gel will run approximately five hours. Set the Voltage to 50 V if you want to run the gel overnight. 13. For analyzing protein mixtures with two-dimensional gels, it is good to apply an aliquot from the same experiment in Laemmli buffer (SDS-Buffer) as control on the side of the gel. A control of a SDS denatured sample that contains the protein of interest will make it easier to find the corresponding spots on the 2-D Gel. It will also facilitate troubleshooting.

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References 1. Schägger, H. and von Jagow, G. (1991) Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form Anal. Biochem. 199, 223–231 2. Schägger, H. and Pfeiffer, K. (2000) Supercomplexes in the respiratory chains of yeast and mammalian mitochondria EMBO J. 19, 1777–1783 3. Jänsch, L., Kruft, V., Schmitz, U. K., and Braun, H.P. (1996) New insights into the composition, molecular mass and stoichiometry of the protein complexes of plant mitochondria Plant J. 9, 357–368 4. Dekker, P. J. T., Müller, H., Rassow, J., and Pfanner, N. (1996) Characterization of the preprotein translocase of the outer mitochondrial membrane by blue native electrophoresis Biol. Chem. 377, 535–538 5. Dekker, P. J. T., Martin, F., Maarse, A. C., Bömer, U., Müller, H., Guiard, B., Meijer, M., Rassow, J., and Pfanner, N. (1997) The Tim core complex defines the number of mitochondrial translocation contact sites and can hold arrested preproteins in the absence of matrix Hsp70/Tim44 EMBO J. 16, 5408–5419 6. Chacinska, A., Rehling, P., Guiard, B., Frazier, A. E., Schulze-Specking, A., Pfanner, N., Voos, W., and Meisinger, C. (2003) Mitochondrial translocation contact sites: separation of dynamic and stabilizing elements in formation of a TOM-TIMpreprotein supercomplex EMBO J. 22, 5370–5381 7. Jänsch, L., Kruft, V., Schmitz, U. K., and Braun, H.-P. (1998) Unique composition of the preprotein translocase of the outer mitochondrial membrane from plants J. Biol. Chem. 273, 17251–17257 8. Werhahn, W. and Braun, H.-P. (2002) Biochemical dissection of the mitochondrial proteome from Arabidopsis thaliana by three-dimensional gel electrophoresis Electrophoresis 23, 640–646 9. Camacho-Carvajal, M. M., Wollscheid, B., Aebersold, R., Steimle, V., and Schamel, W. W. A. (2004) Two-dimensional Blue Native/SDS gel electrophoresis of multi-protein complexes from whole cellular lysates Mol. Cell. Proteomics 3, 176–182 10. Schägger, H., Cramer, W. A., and von Jagow, G. (1994) Analysis of molecular masses and oligomeric states of protein complexes by blue native electrophoresis and isolation of membrane protein complexes by two-dimensional native electrophoresis Anal. Biochem. 217, 220–230 11. Heuberger, E. H. M. L., Veenhoff, L. M., Duurkens, R. H., Friesen, R. H. E., and Poolman, B. (2002) Oligomeric state of membrane transport proteins analyzed with blue native electrophoresis and analytical ultracentrifugation J. Mol. Biol. 317, 591–600 12. Arnold, I., Pfeiffer, K., Neuper, W., Stuart, R. A., and Schägger, H. (1998) Yeast mitochondrial F1Fo-ATP syntase exists as a dimer: identification of three dimerspecific subunits EMBO J. 17, 7170–7178 13. Wiedemann N., Kozjak V., Chacinska A., Schonfisch B., Rospert S., Ryan M.T., Pfanner N., and Meisinger C. (2003) Machinery for protein sorting and assembly in the mitochondrial outer membrane Nature 424, 565–571 14. Reif, S., Voos, W., and Rassow, J. (2001) Intramitochondrial dimerization of citrate synthase characterized by blue native electrophoresis Anal. Bíochem. 288, 97–99

25 Separation of Proteins by Gel Electrophoresis in the Tris-Taurine-HCl System Sylvie Luche, Mireille Chevallet, Cécile Lelong, and Thierry Rabilloud 1. Introduction SDS-PAGE (i.e., zone electrophoresis of proteins in the presence of SDS) is perhaps one of the most widely used protein separation tool. This is due to its relatively ease of setup, its robustness, in the sense that near all proteins are amenable to this type of separation, its compatibility with further protein analysis techniques (e.g., by blotting or by mass spectrometry), and its good resolution. The resolution of zone electrophoresis has been vastly improved by the use of discontinuous systems (1), so that almost only discontinuous systems are used nowadays, and the popular Laemmli system is just one of those (2). However, there is no universal solution in protein separation, so that there is no single gel that is able to exhibit optimal protein separation for all studies. Quite often, the resolution must be adjusted to the experimental needs. These can vary from a rather low resolution over a wide range of molecular masses to a high resolution in a limited molecular mass range. Most often, resolution is adjusted to the experimental needs by playing with the acrylamide concentration. By changing the pore size in the resulting gel, this results in changed sieving properties and thus to changed retardation of proteins in the gels. While very efficient, this way of adjusting resolution is not without practical drawbacks. Highly concentrated gels (15%acrylamide and over) are rather brittle, while low concentration gels (7.5% and lower) are very soft and tend to break. Moreover, highly concentrated gels pose often technical problems in the subsequent steps (e.g., in the mobilization of proteins for blotting or in the efficient penetration of trypsin in proteomics experiments).

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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However, there is another way to adjust resolution in SDS-PAGE, which is to play with the electrophoresis buffer system. In discontinuous electrophoresis, the migration is stopped when the ion boundary (i.e., the zone materializing the migration of the fastest ions) has reached the bottom of the gels. This means in turn that the resolution of the proteins is the ratio between their mobility in the gel and the mobility of this ion boundary, and this gives another mechanism to tune protein resolution. For example, at equal gel concentration, if the ion boundary mobility is slow, this means that all the small, fast-moving proteins will travel with the ion boundary with no resolution. Meanwhile, as the total electrophoretic process is slower and thus takes longer, the slow moving proteins will be given more time to travel in the gel and thus achieve greater resolution if the gel has adequate sieving properties. The overall result is an increased resolution of the high molecular weight proteins and a decreased resolution of low molecular weight proteins. Conversely, if the ion boundary is fast, even the fast-moving, low molecular weight proteins will not be able to travel as fast as the ion boundary, mainly because of gel sieving. As a result, separation will show. However, as in this case the migration time is short, high molecular weight proteins travel minimally in the gel and are therefore poorly resolved. Of course, both acrylamide concentration and ion boundary can be adjusted independently to provide the optimal resolution. While the Tris-glycine system originally described by Ornstein and Davis (1,3) is still by far the most popular, it is not a versatile system in resolution adjustment, and this illustrates how the adjustment of ion boundary speed works. Ion boundary mobilities can be calculated by using the system described by Jovin (4), but more intuitive rules can be used to understand what happens. Ion boundary mobility is just a subcase of isotachophoresis, and a consequence is that the boundary speed is the speed of the slowest-moving ion. Quite often, the low mobility of this trailing ion is tuned by choosing a zwitterionic compound with basic and acidic pKs, so that changing the pH will change the mobility of the ion. Going back to the example of a glycine-based anionic system, the basic pK of glycine is 9.7. This means at an operating pH of 9.7, the speed of the glycine ion wil be one half of its absolute mobility, as half of the molecules are charged. At 8.7, it will be one tenth of the absolute mobility, and 90% of the absolute mobility at 10.7. While this is not a problem per se, these values pose practical problems. For example, at pH 10.7, the acrylamide gel will start to hydrolyze, giving rise to electroosmosis and mechanical problems. Furthermore, while the operating pH is not the gel pH (it is higher in anionic systems), the operating pH is dictated by the pH of the gel, which itself is governed by the other, cationic buffering

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component—Tris in the case of Tris-glycine gels. As the pK of Tris is close to 8.1, this means that the buffering capacity becomes low at pH 9 and this alters the robustness of the system. It can be argued that other buffering components than Tris can be used. While this is in principle true, this buffer is the gel polymerization buffer. This means that this buffering component must not interfere with the acrylamide polymerization. As a result, choice of buffers that can be used is very limited. For example, only ammediol has been used to operate gels at high pH and thus allow for the resolution of low-molecular weight proteins in glycine systems (5). At the other end of the spectrum, decreasing the pH below the classical 8.8 value of the Ornstein-Davis system enables better resolution in the high molecular mass range (6,7). Thus, alternate systems have been proposed for the resolution of low-molecular weight proteins. As a result of the high performance of Tris as a polymerization buffer, these systems have used zwitterionic compounds which a much lower pK than glycine, for example Tricine (8), Bicine (9), or even MES (10). In addition, the use of high pH for gels electrophoresis is not without drawbacks. High pH gels have limited shelf-life because of slow hydrolysis of the acrylamide. Moreover, the reactivity of amino acids toward residual acrylamide monomer is increased at high pH, leading to modification of the side chains (11). It has been therefore proposed to use a neutral pH system using Tricine as a trailing ion (12). However, here again, the required pH of 7 is far from the pK of Tris, leading to robustness problems. The conclusion of these statements is that Tricine, pK 8, is well adapted to high velocity buffers while not adapted to low velocity buffers, while the reverse situation is encountered with glycine and its pK 9.7. We therefore reasoned that a system using a trailing ion of pK close to 8.8 should be adaptable to both high and low velocity buffers, while operating at moderately basic values and close to the pK of Tris—and therefore at maximal buffer capacity. Additional constraints in the possible trailing ions, such as cost, or the presence of a primary amine to ensure compatibility with all types of silver staining, further limited the choice, and we selected taurine as a trailing ion (13). Once this choice was made, we could devise a double gel system (stacking-separating gel) to further optimize the resolution. 2. Materials 2.1. Stock Solutions (All Stored at Room Temperature) 1. Stock acrylamide solution: 30% acrylamide, 0.8% bis-acrylamide. 2. Buffers (see Note 1): a. Sample stock buffer: 120 g/l Tris, 0.8 M HCl.

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Luche et al. b. Stacking buffer: 90 g/l Tris, 0.6 M HCl. c. Separating buffer, low speed: 110 g/l Tris, 0.6 M HCl. d. Separating buffer, medium speed: 130 g/l Tris, 0.6 M HCl. e. Separating buffer, high speed: 150 g/l Tris, 0.6 M HCl. 20% SDS. 10% ammonium persulfate, made fresh every week. Cathode buffer: 6.g/l Tris, 1.g/l SDS, 25.g/l taurine (see Note 2). Anode buffer: 6 g/l Tris, 1 g/l SDS, 30 g/l glycine (see Note 2). TEMED. 60% glycerol. Thioglycerol (see Note 3). Saturated bromophenol blue in water. Solid urea. 2-butanol, saturated with an equal volume of water, and kept as a two-phase system.

2.2. Gel Recipes and Sample Buffers 1. Concentrated sample buffer for 1D electrophoresis: Mix 250 ml of stacking buffer, 250 ml of 20% SDS, 350ml of glycerol, 100 ml of thioglycerol, and 5 ml of saturated bromophenol blue. Add water up to 1 ml. 2. Gel equilibration buffer for 2D electrophoresis: Mix 25 ml of stacking buffer, 25 ml of 20% SDS and 100 ml of glycerol. Add 72 g of urea. This makes directly 200 ml of gel equlibration buffer. 3. Stacking gel: Mix 1 ml of stacking gel buffer, 1 ml of stock acrylamide solution, and 4 ml of water. Add 6 ml of TEMED, and finally 60 ml of 10% ammonium persulfate. 4. Separating gel (see Note 4): Mix 10 ml of the selected separating gel buffer, 20 ml of stock acrylamide solution and 30 m of water. Add 20 ml of TEMED and finally 400 ml of 10% ammonium persulfate. 5. Agarose gel for two-dimensional electrophoresis: To 1 g of agarose, add 12.5 ml of sample stock buffer, 2 ml of 20% SDS, 2 ml of saturated bromophenol blue solution. Add water to 100 ml, and dissolve the agarose by boiling. Store in 10 ml aliquots and let the gel set.

3. Methods 3.1. Gel Casting for One-Dimensional Gel Electrophoresis 3.1.1. Sample Solubilization Mix an equal volume of sample with a volume of concentrated sample buffer. Incubate at 100°C (boiling water bath) for 5 min. Let cool at room temperature 3.1.2. Gel Casting and Loading Clean the glass plates and spacers with water and ethanol. Assemble the glass plates and spacers assembly and let dry. To orient and identify the gel, cut a small piece of filter paper to any recognizable shape (triangle, square, diamond, rectangle, pentagon, etc.) and drop it to the bottom of the gel assembly.

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Mix the components of the separating gel, and pour the mixture in the assembly, leaving adequate empty space at the top to cast the subsequent stacking gel (typically 2–5 cm, depending on the size of the gel and of the anticipated sample volume). Overlay with 2 mm of water-saturated butanol and let polymerize (see Note 5). Once the gel is polymerized (c. 30 min), remove the upper liquid layer (butanol plus gel exudate) and replace with 5 mm of water. Protect from evaporation with parafilm, or transfer in a closed humid box (see Note 6). When needed, remove the upper water layer, mix the components of the staking gel and pour the gel mixture in the gel assembly. Insert the sample comb and let polymerize for 30 min. Remove the comb, and remove the unpolymerized liquid with a needle. Add cathode buffer to fill the sample wells, and load the samples in each well with a syringe or with a sample loading micropipet tip. 3.2. Gel Casting for Two-Dimensional Gel Electrophoresis Gel casting proceeds as for one-dimensional gels, except that no stacking gel is needed. The separating gel mixture can the be cast up to 5 mm from the top, and overlaid with water-saturated butanol. The gel is then kept until use as described for one-dimensional gels. The IEF gels are equilibrated in the gel equilibration buffer for 5 min (carrier ampholytes tube gels) to 20 min (immobilized pH gradient strips). When ready to use, melt the agarose, remove the water layer from the top of the gel, remove the equilibration buffer from the IEF gel, and insert the IEF gel on the top of the second dimension gel. Seal in place with agarose and let the agarose gel set. 3.3. Gel Running Place the loaded gel in the electrophoresis tank. Fill the chambers with the adequate electrode buffer. The taurine-containing buffer must be at least in the cathode chamber (connected to the “minus” lead of the power supply). When separate electrode chambers with no buffer recirculation are present, a cheaper, glycine-containing anode electrode buffer can be used in the anode chamber (connected to the “plus” electrode of the power supply). Gels can be run at constant voltage, power or current. We recommend to run first the gels at 25 V constant voltage for 30 to 60 min to ensure slow and complete protein entry into the gel. The gels can then be run at the maximum power that does not lead to overheating. This depends on the gel size paramaters (length; width and thickness) as well as on the type of cooling used. We run 200 × 160 × 1.5 mm gels in a thermostated chamber with recirculating cooling water at 12 W/gel in this step.

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3.4. Subsequent Processing The migration is stopped when the bromophenol blue front, materializing the moving boundary, reaches the bottom of the separating gel. Gel made in this taurine system can be processed exactly as standard SDS gels cast in the glycine system. This includes staining with coomassie blue, fluorescent stains, silver nitrate or silver-ammonia stains, or blotting. Typical examples of the resolution obtained with the taurine gels, together with the tuning of the resolution window by buffer change, are shown in Figs. 1 and 2.

Fig. 1. Analysis of molecular weight markers. Molecular weight marker proteins were analysed by SDS-PAGE in various electrophoretic systems. The markers are: on the left lane, Bio-Rad wide range markers, in the middle lane, albumin + MyoglobinCNBr peptides, and in the right lane, albumin crosslinked with pyromellitic dianhydride (on gels A and D only). The molecular weight is indicated close to the corresponding band. Proteins were detected by silver staining (nonammoniacal). (A) 10% gel, Tris taurine system (low speed). (B) 10% gel, Tris taurine system (medium speed). (C) 10% gel, Tris taurine system (high speed). (D) 10% gel, Tris glycine system pH 8.8.

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Fig. 2. Effect of double alteration of buffer and acrylamide concentration. The same molecular weight markers as in Fig. 1 were used. (A) 7.5% gel, Tris taurine system (low speed). (B) 9% gel, Tris taurine system (low speed). (C) 10% gel, Tris taurine system (medium speed). (D) 10% gel, Tris taurine system (high speed). (E) 11.5% gel, Tris taurine system (high speed).

4. Notes 1. The versatility of the boundary speed means in turn that the pH of the gels buffers must be carefully controlled. This is especially difficult in the case of Tris buffers, because (i) many pH-meter electrodes have a sluggish response to Tris and (ii) Tris pK varies strongly with temperature (0.03 pH unit per °C). This means that the observed pH varies strongly with the laboratory temperature and/or the speed of addition of HCl. To avoid this variability, we recommend to make a buffer of constant composition, corresponding to the adequate pH. These buffers are most adequately made either by mixing Tris and Tris hydrochloride powders, or by mixing Tris powder with commercial titrated solutions of hydrochloric acid. The medium speed buffers has a moving boundary speed equivalent to the one of the Laemmli’s buffer.

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2. According to the theory of discontinuous electrophoresis, taurine is needed only in the cathode buffer, while Tris is required in the anode buffer. Thus, in an electrophoresis apparatus with separate buffer chambers, the cheaper glycine can replace taurine in the anode buffer. In an apparatus with a single chamber and/or with buffer recirculation, the taurine buffer must be used as electrode buffer. The electrode buffers are made the day of use, but can be stored for up to one week at room temperature 3. Thioglycerol is a substitute to the classical mercaptoethanol for disulfide reduction. Its efficiency is similar, but the odor is considerably weaker. 4. The recipe given yields a 10% separating gels. To adjust the acrylamide concentration, adjust the volume of stock acrylamide solution and complete to a total of 60ml (including the 10 ml of gel buffer) with water. 5. Polymerization is far from finished when the gel is set (i.e., converted from a liquid to a solid state). At this stage 50–70% of the monomer only is incorporated into the gel, depending on the temperature and time. Leaving the gel to “ripen” overnight at room temperature ensures >90% incorporation of monomers into the gel, leading to more consistent electrophoretic separations, less problems of reaction of proteins with acrylamide, and decreased safety problems with the toxic monomers 6. Once polymerized and “ripened,” gels can be stored in a humid box for up to one week in the cold room. References 1. Ornstein, L. (1964) Disc Electrophoresis, 1, Background and Theory. Ann. New York Acad. Sci. 121, 321–349 2. Laemmli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 3. Davis, B.J. (1964) Disc Electrophoresis. 2, Method and application to human serum proteins. Ann. New York Acad. Sci. 121, 404–427 4. Jovin, T.M. (1973) Multiphasic zone electrophoresis. I. Steady-state movingboundary systems formed by different electrolyte combinations. Biochemistry 12, 871–898. 5. Bury, A. F. (1981)Analysis of protein and peptide mixtures: evaluation of three sodium dodecyl sulphate-polyacrylamide gel electrophoresis buffer systems. J. Chromatogr. A 213, 491–500. 6. Johnson, B.F. (1982) Enhanced resolution in two-dimensional electrophoresis of low-molecular-weight proteins while utilizing enlarged gels. Anal Biochem. 127, 235–246. 7. Fritz, J.D., Swartz, D.R., and Greaser, M.L. (1989) Factors affecting polyacrylamide gel electrophoresis and electroblotting of high-molecular-weight myofibrillar proteins. Anal Biochem. 180, 205–210.

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8. Schägger, H. and von Jagow, G. (1987) Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal Biochem. 166, 368–379. 9. Wiltfang, J., Arold, N., and Neuhoff, V. (1991) A new multiphasic buffer system for sodium dodecyl sulfate-polyacrylamide gel electrophoresis of proteins and peptides with molecular masses 100,000–1000, and their detection with picomolar sensitivity.Electrophoresis 12, 352–366. 10. Kyte, J,. and Rodriguez, H. (1983)A discontinuous electrophoretic system for separating peptides on polyacrylamide gels. Anal Biochem. 133, 515–522. 11. Chiari, M., Righetti, P.G., Negri, A., Ceciliani, F., and Ronchi, S. (1992) Preincubation with cysteine prevents modification of sulfhydryl groups in proteins by unreacted acrylamide in a gel. Electrophoresis 13, 882–884. 12. Patton, W.F., Chung-Welch, N., Lopez, M.F., Cambria, R.P., Utterback, B.L., and Skea, W.M. (1991)Tris-tricine and Tris-borate buffer systems provide better estimates of human mesothelial cell intermediate filament protein molecular weights than the standard Tris-glycine system. Anal. Biochem. 197, 25–33. 13. Tastet, C., Lescuyer, P., Diemer, H., Luche, S., van Dorsselaer, A., and Rabilloud, T. (2003) A versatile electrophoresis system for the analysis of high- and lowmolecular-weight proteins. Electrophoresis 24, 1787–1794

26 Cetyltrimethylammonium Bromide Discontinuous Gel Electrophoresis of Proteins Mr-Based Separation of Proteins with Retained Native Activity Robert E. Akins and Rocky S. Tuan 1. Introduction This chapter describes a novel method of electrophoresis that allows the fine separation of proteins to be carried out with the retention of native activity. The system combines discontinuous gel electrophoresis in an arginine/N-Tris (hydroxymethyl methylglycine) (Tricine) buffer with sample solubilization in cetyltrimethylammonium bromide (CTAB). Because the components that distinguish this system are CTAB, arginine, and Tricine and because CTAB is a cationic detergent, we refer to this method as CAT gel electrophoresis (1,2). Proteins separated on CAT gels appear as discrete bands, and their mobility is a logarithmic function of Mr across a broad range of molecular weights. After CAT electrophoresis, many proteins retain high enough levels of native activity to be detected, and gel bands may be detected by both Mr and protein-specific activities. In this chapter, we provide a description of the procedures for preparing and running CAT gels. We also provide some technical background information on the basic principles of CAT gel operation and some points to keep in mind when considering the CAT system. 1.1. Technical Background The electrophoretic method of Laemmli (3) is among the most common of laboratory procedures. It is based on observations made by Shapiro et al. (4) and Weber and Osborn (5), which showed that sodium dodecyl sulfate (SDS) could be used for the separation of many proteins based on molecular size. In Laemmli’s method, SDS solubilization was combined with a discontinuous gel system using a glycine/Tris buffer, as detailed by Ornstein (6) and Davis (7)

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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(see Chapter 21). Typically, SDS-discontinuous gel electrophoresis results in the dissociation of protein complexes into denatured subunits and separation of these subunits into discrete bands. Since the mobility of proteins on SDS gels is related to molecular size, many researchers have come to rely on SDS gels for the convenient assignment of protein subunit Mr. Unfortunately, it is difficult to assess the biological activity of proteins treated with SDS: proteins prepared for SDS gel electrophoresis are dissociated from native complexes and are significantly denatured. Several proteins have been shown to renature to an active form after removal of SDS (8,9); however, this method is inconvenient and potentially unreliable. A preferred method for determining native protein activity after electrophoresis involves the use of nonionic detergents like Triton X 100 (Tx-100) (10); however, proteins do not separate based on molecular size. The assignment of Mr in the nonionic Tx-100 system requires the determination of mobilities at several different gel concentrations and “Ferguson analysis” (11–13). The CAT gel system combines the most useful aspects of the SDS and Tx-100 systems by allowing the separation of proteins based on Mr with the retention of native activity. Previous studies have described the use of CTAB and the related detergent tetradecyltrimethylammonium bromide (TTAB), in electrophoretic procedures for the determination of Mr (14–18). In addition, as early as 1965, it was noted that certain proteins retained significant levels of enzymatic activity after solubilization in CTAB (19). A more recent report further demonstrated that some proteins even retained enzymatic activity after electrophoretic separation in CTAB (14). Based on the observed characteristics of CTAB and CTAB-based gel systems, we developed the CAT gel system. In contrast to previous CTAB-based gel methods, the CAT system is discontinuous and allows proteins to be “stacked” prior to separation (see refs. 6 and 7). CAT gel electrophoresis is a generally useful method for the separation of proteins with the retention of native activity. It is also an excellent alternative to SDS-based systems for the assignment of protein Mr (see Note 1). 1.2. Basic Principles of CAT Gel Operation The CAT gel system is comprised of two gel matrices and several buffer components in sequence. A diagram of the CAT gel system is shown in Fig. 1. In an applied electric field, the positive charge of the CTAB–protein complexes causes them to migrate toward the negatively charged cathode at the bottom of the system. The arginine component of the tank buffer also migrates toward the cathode; however, arginine is a zwitterion, and its net charge is a function of pH. The arginine is positively charged at the pH values used in the tank buffer, but the pH values of the stacking gel and sample buffer are closer to the pI of arginine, and the arginine will have a correspondingly lower net

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Fig. 1. Diagram of a CAT gel. CAT gels begin at the top with the anode immersed in tank buffer and end at the bottom with the cathode immersed in additional tank buffer. The tank buffer solution contains CTAB, Arginine, and Tricine. Between the tank buffers are the stacking gel and the separating gel. The gels are made up of acrylamide polymers in a Tricine-NaOH-buffered solution. Prior to electrophoresis, protein samples are solubilized in a sample buffer that contains CTAB, to solubilize the protein sample, TricineNaOH, to maintain pH, and carry current and glycerol, to increase specific gravity. Proteins solubilized in sample buffer are typically layered under the upper tank buffer and directly onto the stacking gel. See Note 3 for a listing of some physical characteristics of the CAT gel components.

positive charge as it migrates from the tank buffer into these areas. Therefore, the interface zone between the upper tank buffer and the stacking gel/sample buffer contains a region of high electric field strength where the sodium ions in the stacking gel/sample buffer (Tricine-NaOH) move ahead of the reduced

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mobility arginine ions (Tricine-arginine). In order to carry the electric current, the CTAB-coated proteins migrate more quickly in this interface zone than in the sodium-containing zone just below. As the interface advances, the proteins “stack,” because the trailing edge of the applied sample catches up with the leading edge. When the cathodically migrating interface zone reaches the separating gel, the arginine once again becomes highly charged owing to a drop in the pH relative to the stacking gel. Because of the sieving action of the matrix, the compressed bands of stacked proteins differentially migrate through the separating gel based on size. Two features of CTAB-based gels set them apart from standard SDS-based electrophoretic methods. First, proteins separated in CTAB gels migrate as a function of log Mr across a much broader range of molecular weights than do proteins separated in SDS gels. As shown in Fig. 2, a plot of relative migration distance, as a function of known log Mr of standard proteins, results in a straight line. Because of the consistent relationship between Mr and distance migrated,

Fig. 2. Mobility of proteins in a CAT gel as a function of Mr. A mixture of proteins fractionated in a CAT gel with a 6% T acrylamide separator and a 0.7% agarose stacker was visualized by CBB R-250 staining. Relative mobilities (Rf) were calculated as distance, migrated divided by total distance to the salt/dye front and were plotted against the known Mr values for each protein band. The plot is linear across the entire range (R2 > 0.99). Protein bands included trypsinogen (24 kDa), carbonic anhydrase (29 kDa), glyceraldehyde-3-phospate dehydrogenase (36 kDa), ovalbumin (45 kDa monomer and 90 kDa dimer), bovine serum albumin (66 kDa monomer and 132, 198, and 264 kDa multimers), phosphorylase-B (97.4 kDa), and β-galactosidase (116 kDa). See Note 2 concerning the comparison of Rf values from different gels.

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the relative molecular weights of unknown proteins can be determined. CAT gels may be especially useful for the assignment of Mr to small proteins or for the comparison of proteins with very different molecular weights. Second, the retention of significant levels of native activity in CAT gels allows electrophoretic profiles to be assessed in situ for native activities without additional steps to ensure protein renaturation (see Note 2). Taken together, these two characteristics of CAT gels make them an attractive alternative to standard electrophoretic systems. 2. Materials 1. CAT tank buffer: One liter of 5X tank buffer may be prepared using CTAB, Tricine, and arginine free base. First, prepare 80 mL of a 1 M arginine free base solution by dissolving 13.94 g in distilled water. Next, dissolve 22.40 g of Tricine in 900 mL of distilled water; add 5 g of CTAB, and stir until completely dissolved. Using the 1 M arginine solution, titrate the Tricine/CTAB solution until it reaches pH 8.2. Approx 75 mL of 1 M arginine solution will be required/L of CAT tank buffer. Since Tricine solutions change pH with changes in temperature, the tank buffer should be prepared at the expected temperature of use (typically 10–15°C). Finally, add distilled water to 1000 mL. Store the CAT tank buffer at room temperature. Prior to use, prepare 1X tank buffer by diluting 200 mL of the 5X stock to 1000 mL using distilled water of the appropriate temperature (usually 10–15°C); filter the 1X tank buffer through #1 Whatman filter paper to remove any particulate material. The 1X CAT tank buffer may be stored cold, but it should not be reused (see Note 5). Note that CTAB is corrosive, and care should be taken when handling CTAB powder or CTAB solutions: avoid inhalation or skin contact as advised by the supplier. 2. CAT stacking gel buffer: Prepare a 500 mM Tricine-NaOH by dissolving 22.4 g of Tricine in 200 mL of distilled water. Add NaOH until the pH of the solution reaches 10.0. Bring the solution to a total volume of 250 mL using distilled water. As with all Tricine solutions, the pH of CAT stacking gel buffer should be determined at the expected temperature of use. The CAT stacking gel buffer should be stored at room temperature to avoid any precipitation that may occur during longterm cold storage. 3. CAT separating gel buffer: Prepare a 1.5 M Tricine-NaOH solution by dissolving 134.4 g of Tricine in 400 mL of distilled water. Add NaOH until the pH of the solution reaches 8.0. Bring the solution to a total volume of 500 mL using distilled water. As with all Tricine solutions, the pH of CAT separating gel buffer should be determined at the expected temperature of use. The CAT separating gel buffer should be stored at room temperature to avoid any precipitation that may occur during long-term cold storage. 4. CAT sample buffer: Dilute 0.67 mL of CAT separating gel buffer to approx 80 mL with distilled water; to this add 10 mL of glycerol and 1 g of CTAB. Mix the solution until all the components are dissolved, and adjust the pH to 8.8 using NaOH.

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Akins and Tuan Bring the solution to a final volume of 100 mL using distilled water. In some cases, it may be helpful to add a low-mol-wt cationic dye that will be visible during electrophoresis: 10 µL of a saturated aqueous solution of crystal violet may be added/ mL of sample buffer. Note that CTAB is corrosive, and care should be taken when handling CTAB powder or CTAB solutions: avoid inhalation or skin contact as advised by the supplier. Store CAT sample buffer at room temperature to avoid precipitation of the components. Acrylamide stock solution: A 40% acrylamide stock solution may be prepared by combining 38.93 g of ultrapure acrylamide with 1.07 g of bis-acrylamide in a total of 100 mL of distilled water. The final solution is 40%T (w/v) and 2.67%C (w/w). The “%T” and “%C” values indicate that the total amount of acrylamide in solution is 40 g/100 mL and that the amount of bis-acrylamide included is 2.67% of the total acrylamide by weight. The acrylamide stock solution should be stored in the refrigerator. Unpolymerized acrylamide is very toxic, and great care should be taken when handling acrylamide powders and solutions: Follow all precautions indicated by the supplier, including the wearing of gloves and a particle mask during preparation of acrylamide solutions. Agarose stock solution: A ready-to-use agarose stacking gel solution may be prepared by combining 25 mL of CAT stacking gel buffer, 0.1 g CTAB, and 0.7 g of electrophoresis-grade agarose distilled to a final volume of 100 mL. Mix the components well, and, if necessary, adjust the pH to 10.0. Heat the solution in a microwave oven to melt the agarose, and swirl the solution to mix thoroughly. Divide the agarose stock solution into 10 aliquots, and store at 4°C until ready to use. 10% Ammonium persulfate (AP): Dissolve 0.1 g of ammonium persulfate in 1 mL of distilled water. Make just prior to use. Water saturated isobutanol: Combine equal volumes of isobutanol and distilled water. Mix well, and allow the two phases to separate: the water-saturated isobutanol will be the upper layer. Store at room temperature in a clear container so that the interface is visible. CAT gel fixative: Combine 40 mL of distilled water, 10 mL of acetic acid, and 50 mL of methanol; mix well. Store CAT gel fixative in a tightly sealed container at room temperature. Coomassie brilliant blue stain (CBB): Combine 40 mL of distilled water with 10 mL of acetic acid and 50 mL of methanol. Add 0.25 g of CBB R-250, and dissolve with stirring (usually overnight). Filter the solution through #1 Whatman paper to remove any particulate material. Store at room temperature in a tightly sealed container. CBB Destain: Combine 437.5 mL of distilled water, 37.5 mL of acetic acid, and 25 mL of methanol. Mix well, and stored in a closed container at room temperature. Electrophoresis apparatus: A suitable electrophoresis apparatus and power supply are required to run CAT gels. It is desirable to set aside combs, spacers, gel plates, and buffer tanks to use specifically with CAT gels; however, if the same apparatus

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is to be used alternately for CAT gels and SDS gels, it is necessary to clean it thoroughly between each use. Often, the first CAT gel run in an apparatus dedicated to SDS gels will have a smeared appearance with indistinct bands. This smearing is the result of residual SDS, and subsequent CAT electrophoretic runs will resolve protein bands distinctly. This smearing may be somewhat avoided by soaking the gel apparatus and gel plates in CAT tank buffer prior to a final rinse in distilled water at the final step in the cleaning process. The selection of an electrophoresis apparatus to be used for CAT gels should be based on a consideration of the electrical configuration of the system. Because molecular bromine (Br2) will form at the anode, the anode should be located away from the top of the gel (see Note 5). In addition, it is important to realize that CAT gels are “upside-down” relative to SDS gels: proteins migrate to opposite electrodes in the two systems. Some electrophoresis apparatus are intentionally designed for use with SDS, and the anode (usually the electrode with the red lead) may be fixed at the bottom of the gel, whereas the cathode (usually the electrode with the black lead) is fixed at the top of the gel. If such an apparatus is used, the red lead wire should be plugged into the black outlet on the power supply, and the black lead should be plugged into the red outlet on the power supply. Crossing the wires in this fashion ensures that the CTAB-coated proteins in the CAT system will run into the gel and not into the tank buffer.

3. Method The methods for the preparation and running of CAT gels are similar to other familiar electrophoretic techniques. In this section, we will describe the basic methods for preparing samples, casting gels, loading and running gels, visualizing protein bands, and transferring proteins to nitrocellulose (or other) membranes. We will emphasize the differences between CAT gels and other systems. To provide the best results, the recommendations of the manufacturer should be followed concerning the assembly of the apparatus and the casting of discontinuous gels. 3.1. Preparing Samples 1. Protein samples should be prepared at room temperature immediately prior to loading the gel. Typically, tissue fragments, cells, or protein pellets are resuspended in 1.5-mL microfuge tubes using CAT sample buffer (see Note 6). CAT sample buffer may also be used to solubilize cultured cells or minced tissues directly. In each case, the samples should be spun in a microfuge for 0.5 min at 16,000 g to pellet any debris or insoluble material prior to loading the gel. Good results have been obtained when the final concentration of protein in CAT sample buffer is between 1 and 5 mg/mL; however, the preferred concentration of protein will vary depending on the sample and the particular protein of interest. A series of protein dilutions should be done to determine the optimal solubilization conditions for a particular application.

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3.2. Casting CAT Separating Gels 1. Assemble the gel plates and spacers in the gel casting stand as described by the manufacturer. 2. Prepare a separating gel solution by combining the 40%T acrylamide, CAT Separating gel buffer, and distilled water in the ratios indicated in Table 1. Mix the solution by swirling with the introduction of as little air as possible (oxygen inhibits the reactions necessary to accomplish acrylamide polymerization, see Note 7). 3. Degas the solution by applying a moderate vacuum for 5–10 min: the vacuum generated by an aspirator is generally sufficient. 4. Add 10% AP and TEMED to the solution as indicated in Table 1, and swirl the solution gently to mix. Note that insufficient mixing will result in the formation of a nonhomogeneous gel, but that vigorous mixing will introduce oxygen into the mixture. 5. Carefully pour the gel mixture into the gel plates to the desired volume; remember to leave room for the stacking gel and comb. 6. Finally, layer a small amount of water-saturated isobutanol onto the top of the gel. The isobutanol layer reduces the penetration of atmospheric oxygen into the surface of the gel and causes the formation of an even gel surface. Allow polymerization of the separating gel to proceed for at least 60 min to assure complete crosslinking; then pour off the isobutanol, and rinse the surface of the separating gel with distilled water.

3.3. Casting CAT Stacking Gels Two different types of gel stackers are routinely used with CAT gels. For gel histochemical analyses, or where subsequent protein activity assays will be performed, stacking gels made from agarose have provided the best results. 1. Slowly melt a tube of agarose stock solution in a microwave oven; avoid vigorous heating of the solution, since boiling will cause foaming to occur and may result in air pockets in the finished gel.

Table 1 Preparation of Acrylamide Solutions for CAT Gels Regent 40%T Acrylamide Tricine buffer Distilled water Degas solution 10% AP TEMED

4%T,mL

6%T,mL

8%T, mL

10%T,mL

1.00 2.50 6.39 0.10 0.01

1.50 2.50 5.89 0.10 0.01

2.00 2.50 5.39 0.10 0.01

2.50 2.50 4.89 0.10 0.01

Volumes indicated are in milliliters required to prepare 10 mL of the desired solution. Solutions should be degassed prior to the addition of the crosslinking agents, AP and TEMED.

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2. Insert the gel comb into the apparatus, and cast the stacking gel directly onto the surface of the acrylamide separating gel. Allow the agarose to cool thoroughly before removing the comb (see Note 8). 3. As an alternative to agarose stacking gels, low%T acrylamide stackers may also be used. To prepare an acrylamide stacking gel, combine the 40%T acrylamide stock, CAT separating gel buffer (0.5 M Tricine-NaOH, pH 10.0), and distilled water in the ratios indicated in Table 1. Typically, a 4%T stacking gel is used. Degas the solution by applying a moderate vacuum for 5–10 min. Next, add 10% AP and TEMED to the solution as indicated in Table 1, and swirl the solution gently to mix. Insert the gel comb and cast the stacking gel directly onto the surface of the acrylamide separating gel. Do not use water-saturated isobutanol with stacking gels! It will accumulate between the comb and the gel, and cause poorly defined wells to form. Allow the stacking gel to polymerize completely before removing the comb.

3.4. Loading and Running CAT Gels 1. After the stacking and separating gels are completely polymerized, add 1X CAT tank buffer to the gel apparatus so that the gel wells are filled with buffer prior to adding the samples. 2. Next, using a Hamilton syringe (or other appropriate loading device), carefully layer the samples into the wells. Add the samples slowly and smoothly to avoid mixing them with the tank buffer, and fill any unused wells with CAT sample buffer. The amount of sample to load on a given gel depends on several factors: the size of the well, the concentration of protein in the sample, staining or detection method, and so forth. It is generally useful to run several dilutions of each sample to ensure optimal loading. Check that the electrophoresis apparatus has been assembled to the manufacturer’s specifications, and then attach the electrodes to a power supply. Remember that in CAT gels, proteins run toward the negative electrode, which is generally indicated by a black-colored receptacle on power supplies. 3. Turn the current on, and apply 100 V to the gel. For a single minigel (approx 80 mm across, 90 mm long, and 0.8 mm thick), 100 V will result in an initial current of approx 25 mA. Excessive current flow through the gel should be avoided, since it will cause heating. 4. When the front of the migrating system reaches the separating gel, turn the power supply up to 150 V until the front approaches the bottom of the gel. The total time to run a CAT gel should be around 45–60 min for minigels or 4–6 h for full-size gels.

3.5. Visualization of Proteins 1. As with any electrophoretic method, proteins run in CAT gels may be visualized by a variety of staining techniques. A simple method to stain for total protein may be carried out by first soaking the gel for 15 min in CAT gel fixative, followed by soaking the gel into CBB stain until it is thoroughly infiltrated. Infiltration can take

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as little as 5 min for thin (0.8 mm), low-percentage (6%T) gels or as long as 1 h for thick (1.5 mm), high-percentage (12%T) gels. When the gel has a uniform deep blue appearance, it should be transferred to CBB destain. 2. Destain the gel until protein bands are clearly visible (see Note 9). It is necessary to observe the gel periodically during the destaining procedure, since the destain will eventually remove dye from the protein bands as well as the background. Optimally, CBB staining by this method will detect about 0.1–0.5 µg of protein/ protein track; when necessary, gels containing low amounts of total protein may be silver-stained (see Note 10). Note that the CBB stain may be retained and stored in a closed container for reuse. 3. In addition to total protein staining by the CBB method, enzyme activities may be detected by a variety of histochemical methods. The individual protocol for protein or enzyme detection will, of course, vary depending on the selected assay (see Note 11). Generally, when CAT gels are to be stained for enzyme activity, they should be rinsed in the specific reaction buffer prior to the addition of substrate or detection reagent. The CAT gel system provides an extremely flexible method for the analysis of protein mixtures by a variety of direct and indirect gel staining methods.

3.6. Electrophoretic Transfer Blotting Similar to other methods of electroblotting (e.g., see Chapters 57 and 58), proteins can be transferred from CAT gels to polyvinylidene difluoride (PVDF) membranes. Blotting CAT gels is similar to the standard methods used for SDSbased gels with the notable difference that the current flow is reversed. We have successfully transferred proteins using the method described in Chapter 58 with the following changes: 1) A solution of 80% CAT gel running buffer and 20% MeOH was used as the transfer buffer, 2) the transfer membrane was PVDF, and 3) the polarity of the apparatus was reversed to account for the cationic charge of the CTAB. (2) As with electrophoresis, the tank and apparatus used for electroblotting should be dedicated to CTAB gels (see Note 9). 3.7. Conclusion Gel electrophoresis of proteins is a powerful and flexible technique. There are many excellent references for general information concerning the theory behind electrophoresis. One good source of information is Hames and Rickwood (25). The CAT gel electrophoresis system presented here efficiently stacks and separates a wide range of proteins as a function of Mr and preserves native enzymatic activity to such a degree that it allows the identification of protein bands based on native activity. The nature of the interaction between CTAB and native proteins allows the formation of complexes in which the amount of CTAB present is related to the size of the protein moiety. Based on characterizations of detergent/protein interactions that indicate consistent levels of detergent binding without

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massive denaturation (10,26–28), the retention of activity in certain detergent/ protein complexes is expected, and CTAB is likely to represent a class of ionic detergents that allow the electrophoretic separation of native proteins by Mr. Since the level of retained activity after solubilization in CTAB varies depending on the protein of interest, and a given protein will retain varying amounts of measurable activity depending on the detergent used (see ref. 1, for example), a battery of detergents may be tried prior to selecting the desired electrophoresis system. In general, cationic detergents (e.g., the quartenary ammoniums like CTAB, TTAB, and so forth) may be used in the arginine/Tricine buffer system described above; anionic detergents (e.g., alkyl sulfates and sulfonates) may be substituted for SDS in the familiar glycine/Tris buffer system (29). The CAT gel system and its related cationic and anionic gel systems provide useful adjuncts to existing biochemical techniques. These systems allow the electrophoretic separation of proteins based on log Mr with the retention of native activity. The ability to detect native protein activities, binding characteristics, or associations and to assign accurate Mr values in a single procedure greatly enhances the ability of researchers to analyze proteins and protein mixtures. Acknowledgment This work has been supported in part by funds from Nemours Research Programs (to R. E. A.), NASA (SBRA93-15, to R. E. A.), the Delaware Affiliate of the American Heart Association (9406244S, to R. E. A.), and the NIH (HD29937, ES07005, and DE11327 to R. S. T.). 4. Notes 1. SDS vs CTAB: CTAB and SDS are very different detergents. CTAB is a cationic detergent, and proteins solubilized in CTAB are positively charged; SDS is an anionic detergent, and proteins solubilized in SDS are negatively charged. In terms of electrophoretic migration, proteins in CTAB gels migrate toward the cathode (black electrode), and proteins in SDS gels run toward the anode (red electrode). SDS is not compatible with the CAT gel system, and samples previously prepared for SDS-PAGE are not suitable for subsequent CAT gel electrophoresis. Also, the buffer components of the typical SDS-PAGE system, Tris and glycine, are not compatible with the CAT system: Tricine and arginine should be used with CAT gels. Although the detergents are different, protein banding patterns seen in CAT gels are generally similar to those seen when using SDS-PAGE. Rf values of proteins fractionated by CAT electrophoresis are consistently lower than Rf values determined on the same%T SDS-PAGE, i.e., a particular protein will run nearer the top of a CAT gel than it does in a similar

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%T SDS gel. As a rule of thumb, a CAT gel with a 4%T stacker and a 6%T separator results in electrophoretograms similar to an SDS gel with a 4%T stacker and 8%T separator. Differences between the protein banding patterns seen in CAT gels and SDS gels are usually attributable to subunit associations: multisubunit or self-associating proteins are dissociated to a higher degree in SDS than in CTAB, and multimeric forms are more commonly seen in CAT gels than in SDS gels. 2. Detergent solubilization and protein activity: Many proteins separated on CAT gels may be subsequently identified based on native activity, and under the conditions presented here, CTAB may be considered a nondenaturing detergent. Denaturants generally alter the native conformation of proteins to such an extent that activity is abolished; such is the case when using high levels of SDS. Sample preparation for SDS-based gels typically results in a binding of 1.4 g of SDS/1 g of denatured protein across many types of proteins (20), and it is this consistent ratio that allows proteins to be electrophoretically separated by log Mr. Interestingly, at lower concentrations of SDS, another stable protein binding state also exists (0.4 g/1 g of protein) which reportedly does not cause massive protein denaturation (20). In fact, Tyagi et al. (21) have shown that low amounts of SDS (0.02%) combined with pore-limit electrophoresis could be used for the simultaneous determination of Mr and native activity. The existence of detergent– protein complexes, which exhibit consistent binding ratios without protein denaturation, represents an exciting prospect for the development of new electrophoretic techniques: protein Mr and activity may be identified by any of a variety of methods selected for applicability to a specific system. 3. Comparing CAT gels: The comparison of different CAT gel electrophoretograms depends on using the same separating gel, stacking gel, and sample buffer for each determination. An increase in the acrylamide%T in the separating gel will cause bands to shift toward the top of the gel, and high%T gels are more suitable for the separation of low Mr proteins. In addition, the use of acrylamide or high-percentage agarose stackers will lead to the determination of Rf values that are internally consistent, but uniformly lower than those determined in an identical gel with a low-percentage agarose stacker. This effect is likely owing to some separation of proteins in the stacking gel, but, nonetheless, to compare Rf values among CAT gels, the stacker of each gel should be the same. Similarly, any changes in sample preparation (for example, heating the sample before loading to dissociate protein subunits) or the sample buffer used (for example, the addition of salt or urea to increase sample solubilization) often precludes direct comparisons to standard CAT gel electrophoretograms. 4. Characteristics of system components: The CAT gel system is designed around the detergent CTAB. The other system components were selected

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based on the cationic charge of CTAB and the desire to operate the gel near neutral pH. Some of the important physical characteristics of system components are summarized here; the values reported are from information supplied by manufacturers and Data for Biochemical Research (22). In solution, CTAB exists in both monomer and micelle forms. CTAB has a monomer mol wt of 365 Dalton. At room temperature and in low-ionicstrength solutions ( H3 > H2B > H4 (Fig. 1)—depend on sequence differences at residues whose side chains are mapped to the inside of the histone fold (12, 13). To date, many proteins involved in DNA organization and transcription have been shown to share the histone fold, creating homo- and hetero-dimer complexes (14). However, none have yet been analyzed by AUT gradient gel electrophoresis to confirm the predicted interaction with Triton X-100. 2. All acrylamide and bis-acrylamide solutions are potent neurotoxins and should be dispensed by mechanical pipetting devices.

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3. Storage of urea solutions at −20°C minimizes creation of ionic contaminants such as cyanate. The mixed-bed resin assures that any ions formed are removed. Care should be taken to exclude resin beads from solution taken, for example, by filtration through Whatman no. 1 paper. 4. The stacking ions between which the positively charged proteins and peptides are compressed within the stacking gel during the initial phase of gel electrophoresis are NH4+ within the gel compartment and glycine+ in the electrophoresis buffer. Chloride ions interfere with the discontinuous stacking system. This requires that protein samples should (preferably) be free of chloride salts, and that glycine base rather than glycine salt should be used in the electrophoresis buffer. 5. The separating and stacking gels contain 15 vs 4% acrylamide and 0.1 vs 0.16% N,N′-methylene bis-acrylamide, respectively, in 1 M acetic acid, 0.5% TEMED, 50 mM NH4OH, 8 M urea, and 0.0004% riboflavin 5 -phosphate. The concentration of Triton X-100 in the separating gel of the regular system is 9 mM, optimal for the separation desired in our research for histone H3 variant forms of plants and algae. In the gradient system, glycerol and Triton concentrations change in parallel: 10% (v/v) glycerol and 10 mM Triton in the heavy gel (Fig. 2B) and a gradient between 10 and 0% (v/v) glycerol and between 10 and 0 mM Triton in the gradient from heavy to light (Fig. 2B). 6. Acrylamide is photopolymerized with riboflavin or riboflavin 5′ -phosphate as initiator because the ions generated by ammonium persulfate initiated gel polymerization, as used for SDS polyacrylamide gels, interfere with stacking (see Note 4). 7. The height of stacking gel below the comb determines the volume of samples that can be applied and fully stacked before destacking at the surface of the separating gel occurs. In our experience, a 2.5 cm stacking gel height suffices for samples that almost completely fill equally long sample wells. In general, the single blue line of completely stacked proteins and methylene blue dye should be established 1 cm above the separating gel surface. Thus, 1–1.5 cm stacking gel height will suffice for small volume samples. 8. Salt-free samples are routinely prepared by exhaustive dialysis against 2.5% (v/v) acetic acid in 3500 mol wt cutoff dialysis membranes, followed by freezing at −70°C and lyophilization in conical polypropylene tubes (1.5-, 15-, or 50-mL, filled up to half of nominal capacity) with caps punctured by 21-gauge needle stabs. This method gives essentially quantitative recovery of histones, even if very dilute. (For alternative methods, see Chapter 27). 9. The amount of protein that can be analyzed in one gel lane depends highly on the complexity of the protein composition. As a guideline, 5–50 µg of total calf thymus histones with five major proteins (modified to varying extent) represents the range between very lightly to heavily

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Coomassie-stained individual protein bands in 1-mm-thick, 30-cm-long gels using 5-mm-wide comb teeth. The optimal amount of protein for a gradient gel also depends on the number of protein species that must be analyzed. In general, a gradient using the block comb used, equivalent to the width of 30 gel lanes, should be loaded with 30 times the sample for one lane. As an alternative to step 22, electrophoresis buffer is added to the upper buffer reservoir and all sample wells are filled. Samples are layered under the buffer when dispensed by a Hamilton microsyringe near the bottom of each well. The preparative well of a gradient gel should not be loaded with sample prior to the addition of electrophoresis buffer. Uneven distribution of sample cannot be avoided when buffer is added. For unknown reasons, the polyacrylamide gel below the buffer front tends to stick tightly to the glass in an almost crystalline fashion. This may cause tearing of a gradient gel at the heavy gel side below the buffer front. Since attachment to one of the gel plates is typically much stronger, one can release the gel without problems by immersing the glass plate, with gel attached, in the staining solution. A standard method to record the protein staining pattern in Coomassiestained polyacrylamide gels has been Polaroid photography, using an orange filter to increase contrast, with the gel placed on the fluorescent light box, covered by a glass plate to prevent Coomassie staining of the typical plastic surface. Polaroid negatives can be scanned but suffer from a nonlinear response of density, even within the range where careful Coomassie staining leads to near-linear intensity of protein band staining. With the advent of 24+ bit color flatbed scanners with transmitted light capabilities and linear density capabilities in excess of three optical densities, it has become easy to record, and quantitate, intensity of protein staining patterns. Care should be taken to develop a standard scanning setup, using the full dynamic range (typically all three colors used at their full range, 0–255 for a 24-bit scanner; excluding automatic adjustments for density and contrast). A standard gamma correction value should be determined, using an optical density wedge (Kodak), to assure that the density response is linear. Placing a destained polyacrylamide gel on the gel scanner, one should cover the top of the gel with an acetate film to prevent surface reflection abnormalities and prevent touching of the transilluminating light source surface to the film, avoiding moiré interference patterns. Recording gel patterns at 300 dpi in full color and saving loss-less tiff files facilitates faithful replication of experimental results (Fig. 1). Quantitative densitometry programs can use the image files. Triton X-100 interferes with native-mode electrotransfer of histones to nitrocellulose (15). Exchange of Triton by SDS under acidic conditions allows Western blotting (16) (see Chapter 61).

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References 1. Zweidler, A. (1978) Resolution of histones by polyacrylamide gel electrophoresis in presence of nonionic detergents. Meth. Cell Biol. 17, 223–233. 2. Manca, L., Cherchi, L., De Rosa, M. C., Giardina, B., and Masala, B. (2000) A new, electrophoretically silent, fetal hemoglobin variant: Hb F-Calabria [Ggamma 118 (GH1) Phe→Leu]. Hemoglobin 24, 37–44. 3. Urban, M. K., Franklin, S. G., and Zweidler, A. (1979) Isolation and characterization of the histone variants in chicken erythrocytes. Biochemistry 18, 3952–3960. 4. Schwager, S. L. U., Brandt, W. F., and Von Holt, C. (1983) The isolation of isohistones by preparative gel electrophoresis from embryos of the sea urchin Parechinus angulosus. Biochim. Biophys. Acta 741, 315–321. 5. Waterborg, J. H., Harrington, R. E., and Winicov, I. (1987) Histone variants and acetylated species from the alfalfa plant Medicago sativa. Arch. Biochem. Biophys. 256, 167–178. 6. Bonner, W. M., West, M. H. P., and Stedman, J. D. (1980) Two-dimensional gel analysis of histones in acid extracts of nuclei, cells, and tissues. Eur. J. Biochem. 109, 17–23. 7. Waterborg, J. H. (1992) Existence of two histone H3 variants in dicotyledonous plants and correlation between their acetylation and plant genome size. Plant Mol. Biol. 18, 181–187. 8. Waterborg, J. H. (1991) Multiplicity of histone H3 variants in wheat, barley, rice and maize. Plant Physiol. 96, 453–458. 9. Waterborg, J. H., Robertson, A. J., Tatar, D. L., Borza, C. M., and Davie, J. R. (1995) Histones of Chlamydomonas reinhardtii. Synthesis, acetylation and methylation. Plant Physiol. 109, 393–407. 10. Waterborg, J. H. (2000) Steady-state levels of histone acetylation in Saccharomyces cerevisiae. J. Biol. Chem. 275, 13,007–13,011. 11. Waterborg, J. H. and Matthews, H. R. (1983) Patterns of histone acetylation in the cell cycle of Physarum polycephalum. Biochemistry 22, 1489–1496. 12. Arents, G., Burlingame, R. W., Wang, B. C., Love, W. E., and Moudrianakis, E. N. (1991) The nucleosomal core histone octamer at 3.1 Å resolution. A tripartite protein assembly and a left-handed superhelix. Proc. Natl. Acad. Sci. USA 88, 10,138–10,148. 13. Luger, K., Mäder, A. W., Richmond, R. K., Sargent, D. F., and Richmond, T. J. (1997) Crystal structure of the nucleosome core particle at 2.8 Å resolution. Nature 389, 251–260. 14. Sullivan, S. A., Aravind, L., Makalowska, I., Baxevanis, A. D., and Landsman, D. (2000) The histone database: a comprehensive WWW resource for histones and histone fold-containing proteins. Nucleic Acids Res. 28, 320–322. 15. Waterborg, J. H. and Harrington, R. E. (1987) Western blotting from acid–urea– Triton– and sodium dodecyl sulfate-polyacrylamide gels. Analyt. Biochem. 162, 430–434. 16. Delcuve, G. P. and Davie, J. R. (1992) Western blotting and immunochemical detection of histones electrophoretically resolved on acid–urea–Triton- and sodium dodecyl sulfate-polyacrylamide gels. Analyt. Biochem. 200, 339–341.

29 Isoelectric Focusing of Proteins in Ultra-Thin Polyacrylamide Gels John M. Walker 1. Introduction Isoelectric focusing (IEF) is an electrophoretic method for the separation of proteins, according to their isoelectric points (pI), in a stabilized pH gradient. The method involves casting a layer of support media (usually a polyacrylamide gel but agarose can also be used) containing a mixture of carrier ampholytes (low-mol-wt synthetic polyamino-polycarboxylic acids). When using a polyacrylamide gel, a low percentage gel ( 4%) is used since this has a large pore size, which thus allows proteins to move freely under the applied electrical field without hindrance. When an electric field is applied across such a gel, the carrier ampholytes arrange themselves in order of increasing pI from the anode to the cathode. Each carrier ampholyte maintains a local pH corresponding to its pI and thus a uniform pH gradient is created across the gel. If a protein sample is applied to the surface of the gel, where it will diffuse into the gel, it will also migrate under the influence of the electric field until it reaches the region of the gradient where the pH corresponds to its isoelectric point. At this pH, the protein will have no net charge and will therefore become stationary at this point. Should the protein diffuse slightly toward the anode from this point, it will gain a weak positive charge and migrate back towards the cathode, to its position of zero charge. Similarly diffusion toward the cathode results in a weak negative charge that will direct the protein back to the same position. The protein is therefore trapped or “focused” at the pH value where it has zero overall charge. Proteins are therefore separated according to their charge, and not size as with SDS gel electrophoresis. In practice the protein samples are loaded onto the gel before the pH gradient is formed. When a voltage difference is applied, protein migration and pH gradient formation occur simultaneously.

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Traditionally, 1–2 mm thick isoelectric focusing gels have been used by research workers, but the relatively high cost of ampholytes makes this a fairly expensive procedure if a number of gels are to be run. However, the introduction of thin-layer isoelectric focusing (where gels of only 0.15 mm thickness are prepared, using a layer of electrical insulation tape as the “spacer” between the gel plate) considerably reduced the cost of preparing IEF gels, and such gels are therefore described in this chapter. Additional advantages of the ultra-thin gels over the thicker traditional gels are the need for less material for analysis, and much quicker staining and destaining times. Also, a permanent record can be obtained by leaving the gel to dry in the air, i.e., there is no need for complex gel-drying facilities. The tremendous resolution obtained with IEF can be further enhanced by combinations with SDS gel electrophoresis in the form of 2D gel electrophoresis. 2. Materials 1. Stock acrylamide solution: acrylamide (3.88 g), bis-acrylamide (0.12 g), sucrose (10.0 g). Dissolve these components in 80 mL of water. This solution may be prepared some days before being required and stored at 4°C (see Note 1). 2. Riboflavin solution: This should be made fresh, as required. Stir 10 mg riboflavin in 100 mL water for 20 min. Stand to allow undissolved material to settle out (or briefly centrifuge) (see Note 2). 3. Ampholytes: pH range 3.5–9.5 (see Note 3). 4. Electrode wicks: 22 cm × 0.6 cm strips of Whatman No. 17 filter paper. 5. Sample loading strips: 0.5 cm square pieces of Whatman No. 1 filter paper, or similar. 6. Anolyte: 1.0 M H3PO4. 7. Catholyte: 1.0 M NaOH. 8. Fixing solution: Mix 150 mL of methanol and 350 mL of distilled water. Add 17.5 g sulfosalicylic acid and 57.5 g trichloroacetic acid. 9. Protein stain: 0.1% Coomassie brilliant blue R250 in 50% methanol, 10% glacial acetic acid. N.B. Dissolve the stain in the methanol component first. 10. Glass plates: 22 cm × 12 cm. These should preferably be of 1 mm glass (to facilitate cooling), but 2 mm glass will suffice. 11. PVC electric insulation tape: The thickness of this tape should be about 0.15 mm and can be checked with a micrometer. The tape we use is actually 0.135 mm. 12. A bright light source.

3. Methods 1. Thoroughly clean the surfaces of two glass plates, first with detergent and then methylated spirit. It is essential for this method that the glass plates are clean. 2. To prepare the gel mold, stick strips of insulation tape, 0.5 cm wide, along the four edges of one glass plate. Do not overlap the tape at any stage. Small gaps at the

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join are acceptable, but do not overlap the tape at corners as this will effectively double the thickness of the spacer at this point. To prepare the gel solution, mix the following: 9.0 mL acrylamide, 0.4 mL ampholyte solution, and 60.0 µL riboflavin solution. N.B. Since acrylamide monomer is believed to be neurotoxic, steps 4–6 must be carried out wearing protective gloves. Place the glass mold in a spillage tray and transfer ALL the gel solution with a Pasteur pipet along one of the short edges of the glass mold. The gel solution will be seen to spread slowly toward the middle of the plate. Take the second glass plate and place one of its short edges on the taped edge of the mold, adjacent to the gel solution. Gradually lower the top plate and allow the solution to spread across the mold. Take care not to trap any air bubbles. If this happens, carefully raise the top plate to remove the bubble and lower it again. When the two plates are together, press the edges firmly together (NOT the middle) and discard the excess acrylamide solution spilled in the tray. Place clips around the edges of the plate and thoroughly clean the plate to remove excess acrylamide solution using a wet tissue. Place the gel mold on a light box (see Note 4) and leave for at least 3 h to allow polymerization (see Note 5). Gel molds may be stacked at least three deep on the light box during polymerization. Polymerized gels may then be stored at 4°C for at least 2 mo, or used immediately. If plates are to be used immediately they should be placed at 4°C for 15 min, since this makes the separation of the plates easier. Place the gel mold on the bench and remove the top glass plate by inserting a scalpel blade between the two plates and slowly twisting to remove the top plate. (N.B. Protect eyes at this stage.) The gel will normally be stuck to the side that contains the insulation tape. Do not remove the tape. Adhesion of the gel to the tape helps fix the gel to the plate and prevents the gel coming off in the staining/destaining steps (see Note 6). Carefully clean the underneath of the gel plate and place it on the cooling plate of the electrophoresis tank. Cooling water at 10°C should be passing through the cooling plate. Down the full length of one of the longer sides of the gel lay electrode wicks, uniformly saturated with either 1.0 M phosphoric acid (anode) or 1.0 M NaOH (cathode) (see Note 7). Samples are loaded by laying filter paper squares (Whatman No. 1, 0.5 cm × 0.5 cm), wetted with the protein sample, onto the gel surface. Leave 0.5 cm gaps between each sample. The filter papers are prewetted with 5–7 µL of sample and applied across the width of the gel (see Notes 8–10). When all the samples are loaded, place a platinum electrode on each wick. Some commercial apparatus employ a small perspex plate along which the platinum electrodes are stretched and held taut. Good contact between the electrode and wick is maintained by applying a weight to the perspex plate. Apply a potential difference of 500 V across the plate. This should give current of about 4–6 mA. After 10 min increase the current to 1000 V and then to 1500 V after

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a further 10 min. A current of ~4–6 mA should be flowing in the gel at this stage, but this will slowly decrease with time as the gel focuses. 14. When the gel has been running for about 1 h, turn off the power and carefully remove the sample papers with a pair of tweezers. Most of the protein samples will have electrophoresed off the papers and be in the gel by now (see Note 11). Continue with a voltage of 1500 V for a further 1 h. During the period of electrophoresis, colored samples (myoglobin, cytochrome c, hemoglobin in any blood samples, and so on) will be seen to move through the gel and focus as sharp bands (see Note 12). 15. At the end of 2 h of electrophoresis, a current of about 0.5 mA will be detected (see Note 13). Remove the gel plate from the apparatus and gently wash it in fixing solution (200 mL) for 20 min (overvigorous washing can cause the gel to come away from the glass plate). Some precipitated protein bands should be observable in the gel at this stage (see Note 14). Pour off the fixing solution and wash the gel in destaining solution (100 mL) for 2 min and discard this solution (see Note 15). Add protein stain and gently agitate the gel for about 10 min. Pour off and discard the protein stain and wash the gel in destaining solution. Stained protein bands in the gel may be visualized on a light box and a permanent record may be obtained by simply leaving the gel to dry out on the glass plate overnight (see Note 16). 16. Should you wish to stain your gel for enzyme activity rather than staining for total protein, immediately following electrophoresis gently agitate the gel in an appropriate substrate solution (see Note 7, Chapter 20).

4. Notes 1. The sucrose is present to improve the mechanical stability of the gel. It also greatly reduces pH gradient drift. 2. The procedure described here uses photopolymerization to form the polyacrylamide gel. In the presence of light, riboflavin breaks down to give a free radical which initiates polymerization (Details of acrylamide polymerization are given in the introduction to Chapter 21). Ammonium persulphate /TEMED can be used to polymerize gels for IEF but there is always a danger that artefactual results can be produced by persulfate oxidation of proteins or ampholytes. Also, high levels of persulfate in the gel can cause distortion of protein bands (see Note 10). 3. This broad pH range is generally used because it allows one to look at the totality of proteins in a sample (but note that very basic proteins will run off the gel). However, ampholytes are available in a number of different pH ranges (e.g., 4–6, 5–7, 4–8, and so on) and can be used to expand the separation of proteins in a particular pH range. This is necessary when trying to resolve proteins with very similar pI values. Using the narrower ranges it is possible to separate proteins that differ in their pI values by as little as 0.01 of a pH unit.

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4. Ensure that your light box does not generate much heat as this can dry out the gel quite easily by evaporation through any small gaps at the joints of the electrical insulation tape. If your light box is a warm one, stand it on its side and stand the gels adjacent to the box. 5. It is not at all obvious when a gel has set. However, if there are any small bubbles on the gel (these can occur particularly around the edges of the tape) polymerization can be observed by holding the gel up to the light and observing a “halo” around the bubble. This is caused by a region of unpolymerized acrylamide around the bubble that has been prevented from polymerizing by oxygen in the bubble. It is often convenient to introduce a small bubble at the end of the gel to help observe polymerization. 6. If the gel stays on the sheet of glass that does not contain the tape, discard the gel. Although usable for electrofocusing you will invariably find that the gel comes off the glass and rolls up into an unmanageable “scroll” during staining/destaining. To ensure that the gel adheres to the glass plate that has the tape on it, it is often useful to siliconize (e.g., with trimethyl silane) the upper glass plate before pouring the gel. 7. The strips must be fully wetted but must not leave a puddle of liquid when laid on the gel. (Note that in some apparatus designs application of the electrode applies pressure to the wicks, which can expel liquid.) 8. The filter paper must be fully wetted but should have no surplus liquid on the surface. When loaded on the gel this can lead to puddles of liquid on the gel surface which distorts electrophoresis in this region. The most appropriate volume depends on the absorbancy of the filter paper being used but about 5 µL is normally appropriate. For pure proteins load approx 0.5–1.0 µg of protein. The loading for complex mixtures will have to be done by trial and error. 9. Theoretically, samples can be loaded anywhere between the anode or cathode. However, if one knows approximately where the bands will focus it is best not to load the samples at this point since this can cause some distortion of the band. Similarly, protein stability is a consideration. For example, if a particular protein is easily denatured at acid pH values, then cathodal application would be appropriate. 10. The most common problem with IEF is distortions in the pH gradient. This produces wavy bands in the focused pattern. Causes are various, including poor electrical contact between electrode and electrode strips, variations in slab thickness, uneven wetting of electrode strips, and insufficient cooling leading to hot spots. However, the most common cause is excessive salt in the sample. If necessary, therefore, samples should be desalted by gel filtration or dialysis before running the gel. 11. Although not absolutely essential, removal of sample strips at this stage is encouraged since bands that focus in the region of these strips can be

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Walker distorted if strips are not removed. Take care not to make a hole in the gel when removing the strips. Use blunt tweezers (forceps) rather than pointed ones. When originally loading the samples it can be advantageous to leave one corner of the filter strip slightly raised from the surface to facilitate later removal with tweezers. It is indeed good idea to include two or three blood samples in any run to act as markers and to confirm that electrophoresis is proceeding satisfactorily. Samples should be prepared by diluting a drop of blood approx 1:100 with distilled water to effect lysis of the erythrocytes. This solution should be pale cherry in color. During electrophoresis, the red hemoglobin will be seen to electrophorese off the filter paper into the gel and ultimately focus in the central region of the gel (pH 3.5–10 range). If samples are loaded from each end of the gel, when they have both focused in the sample place in the middle of the gel one can be fairly certain that isoelectrofocusing is occurring and indeed that the run is probably complete. Theoretically, when the gel is fully focused, there should be no charged species to carry a current in the gel. In practice there is always a slow drift of buffer in the gel (electro-endomosis) resulting in a small ( 0.5 mA) current even when gels are fully focused. Blood samples (see Note 9) loaded as markers can provide additional confirmation that focusing is completed. It is not possible to stain the IEF gel with protein stain immediately following electrophoresis since the ampholytes will stain giving a uniformly blue gel. The fixing step allows the separated proteins to be precipitated in the gel, while washing out the still soluble ampholytes. This brief wash is important. If stain is added to the gel still wet with fixing solution, a certain amount of protein stain will precipitate out. This brief washing step prevents this. If you wish to determine the isoelectric point of a protein in a sample, then the easiest way is to run a mixture of proteins of known pI in an adjacent track (such mixtures are commercially available). Some commercially available kits comprise totally colored compounds that also allows one to monitor the focusing as it occurs. However, it is just as easy to prepare ones own mixture from individual purified proteins. When stained, plot a graph of protein pI vs distance from an electrode to give a calibration graph. The distance moved by the unknown protein is also measured and its pI read from the graph. Alternatively, a blank track can be left adjacent to the sample. This is cut out prior to staining the gel and cut into 1 mm slices. Each slice is then homogenized in 1 mL of water and the pH of the resultant solution measured with a micro electrode. In this way a pH vs distance calibration graph is again produced.

30 Serial Immobilized pH Gradient Isoelectric Focusing over pH 4–9 Slobodan Poznanovic¢ , Wojciech Wozny, Helmut Zengerling, Gerhard P. Schwall, and Michael A. Cahill 1. Introduction It has long been apparent that the separation capacity of conventional 18 cm IPGs as the first separation step in two dimensional polyacrylamide gel electrophoresis (2D-PAGE) was theoretically incapable of separating the complexity that is to be expected by the protein species of a mammalian cell line or tissue (1). We first attempted to improve the resolution of 2D-PAGE by developing IPGs of 54 cm in length (2). While this strategy was successful, it required the laborintensive preparation and batchwise validation of own-caste IPGs for each experiment. Subsequently we found that qualitatively similar excellent results could be obtained by placing commercially available IPGs in series (end to end) during IEF. We submitted this method as a patent application on September 12, 2003 (WO2005026715, but since allowed to lapse in all countries but Germany) and published it in the scientific literature in August 2005 (3). The main advantage of this analytical strategy is that less sample is consumed per high resolution protein separation as compared to the alternative use of conventional combinations of individual narrow pH range IPGs for IEF. Thus, the system is well suited to the optimal analysis of samples that are limited in amount, as we have since demonstrated in a number of analytical investigations, including cases where samples from small human clinical biopsies were employed (4–6). Farhoud and colleagues independently published an essentially identical serial IEF method and device (7), also emphasizing the maximal resolution and yield of minimally consumed scarce samples. That paper was submitted for publication in July 2005, four months after the publication of our patent application.

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270 2. Materials 2.1. Casting Polyacrylamide Bridges

1. Glass plates for casting polyacrylamide gels. 200 × 260 × 0.5 mm, U-frame and 200 × 260 × 4 mm glass plate (GE Healthcare 80-1106-87; 18-1102-47). 2. GelBond PAG film, 203 × 260 mm (GE Healthcare 80-1129-37). 3. PlusOne Acrylamide IEF (GE Healthcare 17-1300-01). 4. PlusOne N,N′-Methylene-bisacrylamide (GE Healthcare 17-1304-01). 5. PlusOne TEMED (GE Healthcare 17-1312-01). 6. PlusOne Ammonium Persulfate (GE Healthcare 17-1311-01). 7. PlusOne Repel-Silane ES (GE Healthcare 17-1332-01). 8. Kimtech Science Precision Wipes, Kimberly Clark Product Code: 75512.

2.2. Rehydration of IPG Strips 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Immobiline DryStrip pH 4.0–5.0, 18 cm (GE Healthcare 17-6001-84). Immobiline DryStrip pH 5.0–6.0, 18 cm (GE Healthcare 17-6001-86). Immobiline DryStrip pH 6–9, 18 cm (GE Healthcare 17-6001-88). IEF buffer: 7 M urea, 2 M thiourea, 1% Triton X-100, 10% glycerol, 4% CHAPS, 1% DTT. IPG Buffer pH 3.5–5.0 (GE Healthcare 17-6002-02). IPG Buffer pH 5.0–6.0 (GE Healthcare 17-6002-05). Servalyte 6–9 (Serva Heidelberg 42913.01). Immobiline DryStrip Reswelling Tray, for 7–18 cm IPG strips (GE Healthcare 80-6371-84). Flat laboratory forceps suitable for IPG strip manipulation. Sharp laboratory scissors (Stainless steel).

2.3. Assembly of Daisy Chain IPGs 1. Immobiline DryStrip Cover Fluid (GE Healthcare 17-1335-01). 2. Acrylamide bridges (see Note 1).

2.4. Isoelectric Focussing 1. Self-made IEF device (see Note 2).

3. Methods In this methodical report only the serial IEF methodology is thoroughly described. We assume that the experimenter is already able to perform conventional 2D-PAGE using IPGs. Introductory material and references for this area can be found in the series of articles by Görg and colleagues (8–10); see also Chapter 35. The individual IPGs are best subsequently electrophoresed by SDSPAGE and stained according to the protocols already established/optimized in your own laboratory for conventional single IPG-IEF.

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3.1. Casting Polyacrylamide Bridges 1. Prior to use, wash the glass plates thoroughly with diluted detergent, followed by acetone, and then rinse with deionized water and air dry (see Note 3). 2. Siliconize glass plate bearing U-frame (see Note 4). 3. Wash Gel Bond PAG films 3 × 10 min with deionized water prior to use (see Note 5). 4. Assemble gel casting cassette prior to use and leave it in vertical position (see Note 6). 5. Prepare 4% acrylamide solution according to the given recipe (see Note 7). Fill the assembled casting cassette with approximately 23 mL (or less) of acrylamide solution and polymerize for 15 min at room temperature (see Note 8). Subsequently polymerize the gel for 1 hour at 50°C in a heating cabinet (see Note 9). 6. Cool down the gel (see Note 10) and cut them into 4 mm wide strips (see Note 11). 7. Store Acrylamide bridges at −20°C until use (see Note 12).

3.2. Rehydration of IPG Strips 1. Prepare IEF rehydration buffer or thaw from −20°C previously prepared buffer. 2. Rehydrate IPG strips pH 4–5, 5–6 and 6–9 according to standard protocol as you would for individual IPGs of these pH ranges (see Note 13). 3. Rehydrate bridges in IEF buffer with IPG buffer mixture (see Note 14).

3.3. Assembly of Daisy Chain IPGs 1. Assemble the IEF chamber as you routinely do with Multiphor IEF chamber (Fig. 1). 2. Fill the IEF chamber with Immobiline DryStrip Cover Fluid or Paraffin oil you use for standard IEF of IPG strips (see Note 15). 3. Assemble IPG strips from acidic to basic (from left to right) (see Note 16). 4. Cut bridges in 1 – 1.5 cm long pieces. 5. Gently apply the bridging gels to span between the IPG strips, briefly applying slight pressure to facilitate electrical contact between the aqueous zones (Fig. 2).

3.4. Isoelectric Focusing 1. Voltages employed for IEF are parameterized in Table 1. 2. After IEF the IPGs are handled identically to IPGs in conventional single gel 2D-PAGE.

4. Notes 1. Casting of “bridges” should be done at least one day before one plans to start daisy chain. The best is to cast bridges in advance, few days before, in sufficient amount and to store them at −20°C until use. 2. Our description of the self-made IEF device has been published (3). Farhoud and colleagues independently published an essentially identical serial IEF method and device (7). We recommend consulting both of these cited references before attempting to construct an own high voltage IEF device, and certainly not to even consider this without the assistance of suitably qualified

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Fig.1. Working prototype long IEF chamber (for IEF up to 96 cm). (A) The IEF chamber and power supply. The safety lid is open and the temperature regulating water cooler is behind the device. (B) The power supply, running at 20 kV (for more rapid IEF, alternatively to Table 1). (C) Detail view of the IEF chamber. The safety lock is connected via a timedelayed circuit to an unlocking mechanism on the power supply, and can be opened only 5 minutes after the power has been switched off. Other features include: variable IEF length: 18 – 100 cm, integrated cooling surface, silicon oil coolant is an electrical insulator, 35 kV power supply, 0.05 mA adjustable steps, high voltage cables for 35 kV, custom designed electrodes for 35 kV, and electrically insulating safety chamber surrounds IEF chamber.

electrical and/or engineering skill. All electrical circuitry in the self-made IEF chamber was configured according to German regulation VDE 0411 concerning the use of voltages over 10,000 V. The IEF chamber was based upon a water-cooled aluminum surface of about 1100 mm × 240 mm. A shallow glass tray was placed onto the metal surface, and cooled by application of mineral oil between metal and glass to permit heat transfer. The glass tray itself is 940 mm × 145 mm, and has walls that are 18 mm high, resembling a long Pharmacia Multiphore DryStrip tray. The IPGs are placed into this tray as described below and electrodes are placed at each end. These electrodes have cables rated >10,000 V and lead to a HCN 140–35000 power supply which is programmable at 0–4 mA in 50 µA steps (F.u.G. Elektronic, Rosenheim Germany). The cooled aluminum surface with glass tray and electrodes were completely contained in a large Plexiglas (acrylic glass/polymethyl methacrylate) case with a hinging top panel that serves as a locking lid. Plexiglas was

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Fig. 2. (A) Schematic “Daisy Chain” IPG serial IEF design. 1, Anode; 2, Cathode; 3 and 4, Water-soaked paper wick; 5, IPG pH 4.0 – 5.0; 6, IPG pH 5.0 – 6.0; 7, IPG pH 6.0 – 9.0; 8, Bridge between 5 and 6; 9, Bridge between 6 and 7. (B-D) Silver stained 2D-PAGE gels of 250 µg of swine liver proteins from commercial IPGs (Amersham) electrophoresed according to A in 12% SDS-PAGE gels. Molecular weight markers: kDa 220; 160; 120; 100; 90; 80; 70; 60; 50 (fat); 40; 30; 25; 20 (fat/double); 15.

chosen for its excellent properties of electrical insulation. The hinged lid was additionally fitted with an opening safety mechanism whose catch was electronically linked to the power supply. This had been internally custom modified during self-construction to prevent the lid from opening until voltage levels had dropped to safe levels. The joints of the Plexiglas case were also waterproofed with silicon and the case was of sufficient volume that any leakage from the water-cooling system would be retained inside the Plexiglas vessel, and would not come into contact with the electrodes. These aspects are mentioned to emphasize that the personal safety of the experimenter is dependent

Table 1. Serial IEF Regime 150 V 300 V 600 V 3000 V 10000 V 1000 V

30 min 30 min 60 min up t o 1000 0 Vh up t o 1200 00 Vh for 1440 min (stand by)

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5.

6.

Poznanovic¢ et al. upon the technical design of the IEF device. Because commercially manufactured devices are not available, local governmental safety regulations (and common sense) should be strictly adhered to as the minimum standard in the manufacture of a suitable apparatus. All glass plates must be absolutely clean and free of fat and detergents. Treatment of plates with Repel-Silane enables an easier removal of glass plate from cast gels. Repel-Silane ES is a 2% solution of dimethyldichlorosilane dissolved in octamethyl cyclo-octasilane, a low toxicity, and environmentally safe solvent. Pipette 1–2 ml of repel silane on the glass plate with the U-frame and distribute it evenly over whole surface with a lint-free paper (e.g., Kimwipe). Let it dry for a few minutes, rinse with ethanol and repeat procedure once again. This procedure must be repeated occasionally in order to prevent the gels from sticking to the glass plates. Repel silane glass plates can be re-used, but never mix them with normal glass plates! When not in use siliconized glass plates should be stored separately from non-siliconized ones. Pack the plates face to face with a piece of tissue paper between them to avoid contamination or scratching of the glass. To facilitate efficient siliconization of the surface and thereby avoid sticking of gels to the glass plates, it is recommended that the procedure should be repeated at least twice in succession. Always wash with the hydrophilic side upwards. Gel Bond PAG film is packed with the treated side up, with paper interleaving to protect the treated surface. If uncertain, the hydrophilic side can be confirmed by applying a drop of water and noting that it spreads on the hydrophilic side, but beads up on the hydrophobic side. Always handle Gel Bond PAG film by the edges and with gloves to avoid getting fingerprints on the treated surface. Note that PAG film is both light and heat sensitive. It should be stored under cool dark conditions, and you should be aware that after prolonged or inappropriate storage the acrylamide groups may be degraded, and your gels will not be able to polymerize to the surface satisfactorily. Our casting cassette consists of two glass plates of 200 × 260 mm. One of these is covered with the Gel Bond PAG film and the other contains a 0.5 mm U-frame. The frame is made of silicon rubber enclosing 0.5 mm metal balls. The cassette is assembled as originally described by Görg and colleagues (http://www.weihenstephan.de/blm/deg/2D-Manual_06.pdf) (8–10). The cleaned glass plate without the U-frame is moistened with a few drops of water, and the Gel Bond PAG film, hydrophilic side upwards, is placed onto the wet glass surface. The surface of the Gel Bond PAG film is covered by a piece of Kimtech tissue paper and excess water is expelled with a roller as originally described by Görg and colleagues. Place the glass plate with the U-frame (0.5 mm thick) on top of the Gel Bond PAG film and clamp the cassette together.

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7. Chemicals used for preparation of gel should be of the highest possible quality. We use IEF grade acrylamide purchased in powder form. A stock solution (30% T; 2,7% C) is always freshly prepared or occasionally stored at +4°C for a maximum of one week. We have found it unnecessary to use ion exchange resin for preparation of such stock solutions. Omission of ion exchange avoids potential contamination of the acrylamide solution with impurities from the resin preparations. Stock solution (30% T; 2,7% C): Dissolve 29,18 g of Acrylamide and 0,81 g of bisacrylamide in MilliQ water and fill up to 100 mL. Solution should be filtered and stored at +4 °C. To make the 4% gels add 4 ml acrylamide stock to 26 mL H2O, then add 30 µL of freshly prepared 40% ammonium persulfate and 20 µL TEMED. 8. The gel can be caste using several different methods. The most important point is to fill slowly to avoid the formation of bubbles. We apply the acrylamide mixture slowly and steadily to one side of the casting chamber with a 10 mL pipette, initially tilting the cassette about 40° to run the gel down that side. This should obviously not be performed too slowly because of the advancing polymerization reaction itself. 9. Although the acrylamide mixture should have gelled by this stage, during polymerization in the oven the caste gel in its cassette should remain in the vertical position, this way avoiding any possible movement and leakage of the gel. 10. After polymerization at 50°C allow the gel to cool down to RT for at least 15 min. Disassemble the cassette by inserting a spatula between gel and glass plate and gently but firmly levering the plates apart. Remove the support film with attached gel carefully and wash the PAG film and gel 4 × 15 min with 500 ml MilliQ water to remove unpolymerized acrylamide monomers. Finally incubate the gel in 2% Glycerol for 30 min. Dry the gel overnight in a hood or with a fan or hair-dryer for a few hours. We prefer drying overnight, avoiding potential dust (and keratin) contamination which may occur by drying with a forced air flow, such as caused by a hair dryer or electric fan. 11. The gel can be cut into strips using an office paper cutter, as suggested by Görg. Cleaner edges can be achieved by scission with an industrial paper cutting machine. 12. The bridge gels, consisting only of polyacrylamide without buffers at this stage, are quite stable. We have used precast bridging gels stored sealed at −20°C for more than one year, after which the supply was exhausted. 13. Rehydration of strips should be done according to the instruction for the individual strips. There can be different methods of sample application for daisy chain experiments, e.g.: – sample application by in-gel rehydration. – sample application via wick- or cup-loading – sample application via bridge loading

Poznanovic¢ et al.

276 Table 2 Buffers for IPG Rehydration pH of IPG strip Case A (in-gel rehydration)

Case B (wick application)

4–5

IEF buffer with 0.8 % of IPG buffer pH 3.5–5.0 IEF buffer with 0.8 % of IPG buffer pH 5.0–6.0 IEF buffer with 0.8 % of Servalyte CA pH 6.0–9.0 IEF buffer with 0.8% mix of IPG buffers 3.5–5.0; 5.0–6.0 and Servalyte 6.0–9.0, 1:1:1

5–6 6–9 Bridges

Sample + IEF buffer with 0.8% of IPG buffer pH 3.5–5.0 IEF buffer with 0.8 % of IPG buffer pH 5.0–6.0 IEF buffer with 0.8 % of Servalyte CA pH 6.0–9.0 IEF buffer with 0.8% mix of IPG buffers 3.5–5.0; 5.0–6.0 and Servalyte 6.0–9.0, 1:1:1

The best result depend upon the nature of the proteins in your sample. In the first case the sample is loaded by the rehydration method into the IPG strip pH 4–5 or 5–6 depending on the sample. The other IPG strips for the daisy chain are rehydrated with IEF buffer containing the appropriate corresponding IPG buffer. The optimum method of loading the strips by rehydration will also be sample-specific, with better 2D-PAGE samples obtained by rehydration loading into the basic IPG, the acidic IPG, or all IPGs for different samples. When we apply sample via the wick then all single gels are rehydrated in their appropriate IEF buffer with corresponding individual pH-specific IPG buffer. In all cases for each IPG strip pH gradient we used 0.8% of corresponding IPG buffer. For specific samples better results can be obtained by bridge loading. The optimum position of the loading bridge is sample dependent and should be empirically determined. A tabular presentation of appropriate conditions is depicted in Table 2. 14. Bridges were always rehydrated in IEF buffer containing 0.8% of mixture all used IPG buffers and Servalyte in relation 1:1:1. In all cases rehydration was done on the lab bench overnight at room temperature. Shorter rehydration steps can be employed, although we advise rehydration for more than 5 hours. 15. The IPG holding tray chamber should be filled with oil sufficiently that the IPG strips and paper wicks are submerged. It may be more convenient to first fill the IPG tray with oil and then lay the strips gently down into the oil. 16. To simplify assembling of a daisy chain in the IEF tray we use laminated grid paper positioned under the tray. Try to position the IPGs as closely as possible end-to-end, avoiding any overlap. To position the IPGs we suggest cutting the plastic ends of the IPG strips before alignment, according to the following schema:

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– IPG strip pH 4–5: Cut the plastic end close to the gel at the cathodic (“−” or basic) end. – IPG strip pH 5–6: Cut the plastic ends close to the gel at both the anodic (“+” or acidic) and cathodic (“−” or basic) ends. – IPG strip pH 6–9: Cut the plastic end close to the gel at the anodic (“+” or acidic) end. References 1. Vuong, G. L., Weiss, S. M., Kammer, W., Priemer, M., Vingron, M., Nordheim, A., and Cahill, M. A. (2000) Improved sensitivity proteomics by postharvest alkylation and radioactive labelling of proteins. Electrophoresis 21, 2594–2605. 2. Poland, J., Cahill, M. A., and Sinha, P. (2003) Isoelectric focusing in long immobilized pH gradient gels to improve protein separation in proteomic analysis. Electrophoresis 24, 1271–1275. 3. Poznanovic, S., Schwall, G., Zengerling, H., and Cahill, M. A. (2005) Isoelectric focusing in serial immobilized pH gradient gels to improve protein separation in proteomic analysis. Electrophoresis. 26, 3185–3190. 4. Hunzinger, C., Wozny, W., Schwall, G. P., Poznanovic, S., Stegmann, W., Zengerling, H., Schoepf, R., Groebe, K., Cahill, M. A., Osiewacz, H. D., Jagemann, N., Bloch, M., Dencher, N. A., Krause, F., and Schrattenholz, A. (2006) Comparative profiling of the mammalian mitochondrial proteome: multiple aconitase-2 isoforms including N-formylkynurenine modifications as part of a protein biomarker signature for reactive oxidative species. J. Proteome. Res. 5, 625–633. 5. Neubauer, H., Clare, S. E., Kurek, R., Fehm, T., Wallwiener, D., Sotlar, K., Nordheim, A., Wozny, W., Schwall, G. P., Poznanovic, S., Sastri, C., Hunzinger, C., Stegmann, W., Schrattenholz, A., and Cahill, M. A. (2006) Breast cancer proteomics by laser capture microdissection, sample pooling, 54-cm IPG IEF, and differential iodine radioisotope detection. Electrophoresis 27, 1840–1852. 6. Poznanovic, S., Wozny, W., Schwall, G. P., Sastri, C., Hunzinger, C., Stegmann, W., Schrattenholz, A., Buchner, A., Gangnus, R., Burgemeister, R., and Cahill, M. A. (2005) Differential radioactive proteomic analysis of microdissected renal cell carcinoma tissue by 54 cm isoelectric focusing in serial immobilized pH gradient gels. J. Proteome. Res. 4, 2117–2125. 7. Farhoud, M. H., Wessels, H. J., Wevers, R. A., van Engelen, B. G., van den Heuvel, L. P., and Smeitink, J. A. (2005) Serial isoelectric focusing as an effective and economic way to obtain maximal resolution and high-throughput in 2D-based comparative proteomics of scarce samples: proof-of-principle. J. Proteome. Res. 4, 2364–2368. 8. Gorg, A., Postel, W., and Gunther, S. (1988) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 9, 531–546. 9. Gorg, A., Obermaier, C., Boguth, G., Harder, A., Scheibe, B., Wildgruber, R., and Weiss, W. (2000) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresi. 21, 1037–1053. 10. Gorg, A., Weiss, W., abd Dunn, M. J. (2004) Current two-dimensional electrophoresis technology for proteomics. Proteomics 4, 3665–3685.

31 Radiolabeling of Eukaryotic Cells and Subsequent Preparation for 2-Dimensional Electrophoresis Nick Bizios 1. Introduction Two dimensional polyacrylamide gel electrophoresis (2-DE) provides not only the ability to resolve and quantify thousands of proteins, but it also gives those in research and industry the ability to monitor in-process protein purification, quickly and easily (1). It can become a valuable tool in process analytical technology (PAT) in which the goal is to understand and control a process in order to build in quality. 2-DE is also used to identify variability in protein expression in a variety of cell lines (2,3). It is known that many parameters and laboratory conditions can influence the resolution of proteins on 2-DE, such as pH range of carrier ampholytes used, the quality of reagents and equipment used, temperature, voltage, and the skill of the researcher or technician. Even though there are a multitude of vendors that supply premade reagents, there is pride bestowed to the technician or researcher performing their work. One of the biggest obstacles one may encounter prior to performing 2-DE is the proper documentation and validation of a Standard Operating Procedure, or SOP. This falls under the category of current Good Laboratory Practices (cGLPs) and or current Good Manufacturing Practices (cGMPs). Without the proper documentation of reagents used, such as Certificates of Analysis (readily supplied by the supplier’s Quality Control/Assurance department), adherence to expiration dates of commercially prepared reagents, implementation of expiration dates of reagents prepared in the laboratory and the adherence of SOPs (see Note 1) the ability of generating repeatable and consistent results falls dramatically. However, the radiolabeling and preparation or eukaryotic cells for 2-D PAGE may be considered paramount in obtaining consistent results. Without proper

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radiolabeling and preparation, how would one find subsequent spots of interest and perform quantitative analysis? This chapter will describe a general and robust method for the labeling methionine-containing proteins, phosphorylation labeling, and subsequent lysate preparation for 2-DE that have been modified from Garrels (4) and Garrels and Franza (5). The Jurkat T-lymphoblast cell line is used as an example. A larger number of protocols have been published for the solubilization and sample preparation of eukaryotic cell lines and tissues for 2-DE. One of the best places for additional protocols is found at the Geneva University Hospital’s Electrophoresis laboratory, which can be accessed at http;//expasy.hcuge.ch/ ch2d/technical-info.html. 2. Materials 2.1. Equpiment 1. 0.2µm filters (see Note 2) 2. Heat block or 100°C water bath

2.2. Reagents 1. Complete culture medium: 90% RPMI-1640, 10% fetal bovine serum (FBS), streptomycin-penicillin: Mix 990 mL of RPMI-1640 with 100 mL of FBS, and add 10 mL of streptomycin-penicillin (100X). Cold-filter-sterilize using a 0.2µm filter. Store at 4°C. Maintain the Jurkat T-lymphoblast at a concentration of 105–106 cells/mL (see Note 3). 2. Methionine-free medium: 90% methionine-free RPMI-1640, 10% dialyzed FBS (dFBS): Mix 990 mL of methionine-free RPMI-1640 with 100 mL of dFBS. Coldfilter-sterilize using 0.2µm filter. Store at 4°C. 3. Sodium phosphate-free medium: 90% sodium phosphate-free RPMI-1640, 10% FBS: Mix 90% sodium phosphate free RPMI with 10% FBS. Cold-filter-sterilize. Store at 4°C. 4. Redivue L-[35S]methionine (cell labeling grade) (GE Healthcare) 5. 32P-orthophospahte (GE Healthcare). 6. Phosphate buffered saline (PBS): Mix 8 g NaCl, 0.2 KCl, 1.44 g Na2HPO4, and 0.24 grams KH2PO4 in 800 mL of dH2O) and adjust to 7.4 with HCl. Add dH2O to a final volume of 1000 mL and autoclave. Store at room temperature. 7. Dilute SDS (dSDS): 0.3% SDS, 1% β-mercaptoethanol (β-ME), 0.05 M Tris-HCl, pH 8.0. In a cold room, mix 3.0 g of SDS, 4.44 g of Tris-HCl, 2.65 g Tris base, 10 mL of β-ME in dH2O, and adjust the final to 1 L with dH2O. Aliquot 500 mL into microcentrifuge tubes and store at -70°C. 8. DNase/RNase solution: 1 mg/mL DNase I, 0.5 mg/mL RNase A, 0.5 M Tris, 0.05 M MgCl2, pH 7.0. Thaw RNase, Tris, and MgCl2 stocks, and thoroughly mix 2.5 mg of RNase A (Worthington Enzymes), 1585 µL of 1.5 M Tris-HCl, 80 µL of 1.5 Tris base, 250 µL of 1.0 M MgCl2, and 2960 µL of dH2O. Mix the liquids, and add 5 mg

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9.

10. 11. 12.

13.

281

DNase I (Worthington Biochemical). Do not filter. Keep cool while dispensing into microcentrifuge tubes. Makes 50 µL aliquots, and store at −70°C. 1.5 M Tris-HCl solution: Weigh out 11.8 g of desiccated Tris-HCl, and add 41.4 g dH2O. Mix well and filter through 0.2 µm filter. Aliquot into microcentrifuge tubes, and store at −70°C. 1.5 M Tris base solution: Weigh out 9.09 g of Tris base, and add 41.4 g of dH2O. Mix and filter through 0.2 µm filter. Aliquot into microcentrifuge tubes, and store at −70°C. 1.0 M MgCl2 solution: Weigh out 30.3 g of MgCl2, and add 85.9 g of dH2O. Mix and filter through 0.2 µm filter. Aliquot into microcentrifuge tubes, and store at −70°C. Sample buffer solution (SB): 9.95 M urea, 4.0% Igepal CA-630 (Sigma), 2% 6.0–8.0 ampholytes, 100 mM dithiothreitol (DTT). Mix 59.7 g urea, 44.9 g dH2O, 4.0 g Igepal CA-630, 5.5 g of pH 5.0 to 8.0 ampholytes (GE Healthcare), 1.54 g of DTT (Calbiochem) in this order in a 30° to 37° water bath just long enough to dissolve the urea. Filter through a 0.2 µm filter, and aliquot 1 mL into microcentrifuge tubes. Snap-freeze in liquid nitrogen, and store at −70°C. Sample buffer with SDS solution (SBS): 9.95 M urea, 4.0% Igepal CA-630 (Sigma), 0.3 % SDS, 2% 6.0–8.0 ampholytes, 100 mM dithiothreitol (DTT). Mix 59.7 g urea, 44.9 g dH2O, 4.0 g Igepal CA-630, 0.3 g SDS, 5.5 g of pH 5.0 to 8.0 ampholytes, 1.54 g of DTT in this order in a 30° to 37° water bath just long enough to dissolve the urea. Filter through a 0.2 µm filter, and aliquot 1 mL into microcentrifuge tubes. Snap-freeze in liquid nitrogen, and store at −70°C.

3. Methods 3.1. 35S-Labeling (see Note 4) 1. Jurkat T-lymphocytes are labeled for 3–24 h in methionine-free media containing 50–250 µCi/mL of 35S. 2. Follow cell lysate protocol (Subheading 3.3.,).

3.2. 32P-Labeling (see Note 3) 1. Add 100 µCi/mL of 32P for up to 3 h to cells that are in phosphate-free medium (see Note 5).

3.3. Whole-Cell Lysate Preparation 1. 2. 3. 4. 5. 6.

Wash cells with PBS three times in a microcentrifuge tube. Add an equal volume of hot (100°C) dSDS solution to the pellet. Boil tube (100°C) for 1–3 min. Cool in an ice bath (see Note 6). Add one-tenth (1/10) volume of DNase/RNase solution. Gently vortex for several minutes to avoid foaming. The sample should lose its viscosity, and the solution should look clear. If not, then add more dSDS and DNase/RNase solution (see Note 7). 7. Snap-freeze in liquid nitrogen, and store at −70°C. Samples may be kept for up to 6 mo at −70°C.

282

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3.4. Preparing the Sample for 2D-PAGE: Vacuum Drying 1. Lyophilize sample (frozen at −70°C) in a Speed Vac using no or low heat until dry see Note 8). 2. Add SB solution to the sample equal to that of the original dSDS sample volume, and mix thoroughly. 3. Heat sample to 37°C for a short period if necessary (see Note 9). 4. Store at −70°C. Samples can be kept for up to 6 mo at −70°C. 5. Radioisotope incorporation in the sample may now be determined by TCA precipitation. 6. Recommended first-dimension load is 500,000 dpm for 35S-labeled proteins and 200,000 dpm for 32P-labeled proteins. 7. If necessary, the sample is diluted and mixed thoroughly with SBS solution before loading onto the first-dimension gel (see Note 9).

4. Notes 1. cGLPs and cGMPs will help with all aspects in a research facility. Industry will have Quality Assurance/Control departments to oversee the implementation of these practices. Smaller research labs may wish to assign or hire a person to oversee these practices and adhere to. 2. Small syringe type 0.2 µm filters are best acquired from Nalgene while larger scale filters requiring a pump are best supplied from Sartorius-Stedim Biotech, Pall or Millipore. 3. Maintain Jurkat T-lymphocytes in complete culture medium supplemented with streptomycin/penicillin, in a humidified incubator with 95% air and 5% CO2 at 37°C at a concentration of 105-106 cells/mL. 4. It is extremely important that all radioactive work be performed with the utmost care, and according to your institutional and local guidelines. 5. It may be necessary to preincubate the cells. Preincubating the cells at a density of 1–10 × 106 cells/mL for 30 min in sodium-free, phosphate-free medium works well. 6. Sample preparation should be done quickly once to avoid degradation by proteases. 7. Keep salt concentrations as low as possible, high concentrations (>150 mM) of NaCl, KCl, and other salts cause streaking problems, as do lower concentrations of phosphate and charged buffers. Dialyzing samples to remove salts and other low-mol-wt substances is recommended. 8. Larger sample volumes may be prepared using a large scale commercial lyophilizer in order to prepare “internal standards” for each run. 9. After dissolving the sample in SB solution or diluting in SBS solution, it is imperative that the sample is not subjected to temperatures above 37°C. At extreme temperature (>40°C), the urea in SB and SBS will

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cause carbamylation. Charged isoforms will be generated by isocyanates formed by the decomposition of urea. References 1. Bizios, N. (2007) Unpublished data, AGI Dermatics. 2. Yu, L. R., Zeng, R., Shao, X. X., Wang, N., Xu, Y. H., and Xia, Q. C. (2000) Identification of differentially expressed proteins between human hepatoma and normal liver cell lines by two-dimensional electrophoresis and liquid chromatography-ion trap mass spectrophotometry. Electrophoresis 14, 3058–3068. 3. Carrol, K., Ray, K., Helm, B., and Carey, E. (2000) Two-dimensional electrophoresis reveals differential protein expression in high- and low-secreting variants of the rat basophilic leukemia cell line. Electrophoresis 12, 2476–2486. 4. Garrels, J. I. (1983) Quantitative two-dimensional gel electrophoresis of proteins, in Methods of Enzymology, vol. 100 (Grossman, L. Moldave, K. and Wu, R., eds.) Academic, New York, pp 411–423. 5. Garrels, J. I. And Franza, B.R. Jr. (1989) The REF52 protein database. J. Biol. Chem. 264, 5238–5298.

32 Two-Dimensional Polyacrylamide Gel Electrophoresis of Proteins Using Carrier Ampholyte pH Gradients in the First Dimension Patricia Gravel 1. Introduction Two-dimensional polyacrylamide gel electrophoresis (2-D PAGE) is the only method currently available for the simultaneous separation of thousands of protein. This method separates individual proteins and polypeptide chains according to their isoelectric point and molecular weight. The 2-D PAGE technology can be used for several applications, from which; separation of complex protein mixture into their individual polypeptide components; comparison of protein expression profiles of sample pairs (normal versus transformed cells, cells at different stages of growth or differentiation, etc.). This chapter describes the protocol for 2-D PAGE using carrier ampholyte pH gradient gel for the isoelectric focusing separation (pH gradient 3.5–10) and a polyacrylamide gradient gel for the second dimension (1,2). For many years the 2-D PAGE technology relied on the use of carrier ampholytes to establish the pH gradient. This traditional technique has proven to be difficult in the hands of many because of the lack of reproducibility created by uncontrolable variations in the batches of ampholytes used to generate the pH gradients. This problem was solved by using immobilised pH gradients (IPG) (see Chapter 35), where the compounds used to set up the pH gradient are chemically immobilized and so the gradient is stable. Another advantage is that a larger amount of protein could be used in the separation for micropreparative runs. The availability of commercial precast IPG gels in a variety of narrow and broad pH ranges, as well as precast SDS-PAGE gels has lead also to major advances in protein separation, display and protein characterization (3). 2-D PAGE reference maps are now available over the World-Wide Web (4, 5).

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2. Materials 2.1. Preparation of Samples 1. Lysis solution A:10 % (w/v) SDS (sodium dodecyl sulfate) and 2.32 % (w/v) DTE (1,4- dithio-erythritol) in distilled water (dH2O). 2. Lysis solution B:5.4 g urea (9.0 M) (must be solubilized in water at warm temperature, around 35°C), 0.1 g DTE (65 mM), 0.4 g CHAPS (cholamidopropyldimethylhydroxypropane sulfonate) (65 mM), 0.5 ml of Ampholines pH range 3.5–10 (5% v/v) made to 10 ml with dH2O. These solutions can be aliquoted and stored at −20°C for many months.

2.2. Isoelectric Focusing (IEF) 1. IEF is performed with the Tube Cell Model 175 (Bio-Rad) (see Fig. 1) and with glass capillary tubes (1.0 – 1.4 mm internal diameter and 210 mm long).

Fig. 1. Tube Cell Model 175 (Bio-Rad) for the isoelectric focusing with carrier ampholyte pH gradient capillary gels.

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Ampholytes pH 4–8 and Ampholytes pH 3.5–10 are from BDH (Poole, England). 2. Stock solution of acrylamide: 30% (w/v) acrylamide, 0.8% (w/v) PDA (piperazine diacrylamide) in dH2O. This solution should be stored in the dark at 4°C for 1 to 2 months. 3. Cathodic buffer: 20 mM NaOH in dH2O. This solution should be made fresh. 4. Anodic buffer: 6 mM H3PO4 in dH2O. This solution should be made fresh. 5. Ammonium persulfate (APS) stock solution: 10% (w/v) APS in dH2O. This solution should be stored at 4°C, protected from light and made fresh every 2 to 3 weeks. 6. SDS stock solution: 10% (w/v) SDS in dH2O. This solution can be stored at room temperature. 7. Bromophenol blue stock solution: 0.05% (w/v) bromophenol blue in dH2O. This solution can be stored at room temperature. 8. Capillary gel equilibration buffer: 20 ml 0.5 M Tris-HCl pH 6.8 40 ml 10% (w/v) SDS stock solution 8 ml 0.05% (w/v) bromophenol blue stock solution 72 ml dH 2O

This solution can be stored at room temperature for 2 to 3 months. 2.3. SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. The Protean II chamber (Bio-Rad) is employed by us for SDS-PAGE. The gels (160 × 200 × 1.5 mm) are cast in the Protean II casting chamber (Bio-Rad). The gradient former is model 395 (Bio-Rad). 2. Running buffer: 50 mM Tris base, 384 mM glycine, 0.1% (w/v) SDS in dH2O For 1 L:

6 g Tris base, 28.8 g glycine and 1 g SDS. Do not adjust pH. Fresh solution is made up for the upper tank. The lower tank running buffer can be retained for more than 6 months with the addition of 0.02% (w/v) sodium azide. 3. Sodium thiosulfate stock solution: 5% (w/v) sodium thiosulfate anhydrous in dH2O. This solution should be stored at 4°C. (See Note 1.) 4. Silver staining: Proteins in the 2-D gel are stained with silver. We used the ammoniacal silver nitrate method modified by Hochstrasser et al. (6) and Rabilloud (7).

3. Method 3.1. Preparation of Protein Samples Pellets of cells or tissue should be resuspended in 100 µl of 10 mM TrisHCl pH 7.4 and sonicated on ice for 30 s. Add one volume of lysis solution B and mix. The mixed solution should not be warmed up above room temperature. Samples can be loaded directly onto the gels or stored at −80°C until needed. (See Note 2 for plasma sample preparation and Note 3 for optimal sample loading).

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3.2. Isoelectric Focusing (IEF) 1. Draw a line at 16 cm on clean and dry glass capillary tubes (capillary should be cleaned with sulfochromic acid to eliminate all deposits). The remaining 0.5 cm of the capillary tubes is used to load the sample. Place each tube in a small glass test tube (tube of 5 ml) which will be filled with the isoelectric focusing gel solution. Connect the tops of each glass capillary tube to flexible plastic tubes joined together with a 1 ml plastic syringe. 2. At least twelve capillary gels can be cast with 11.5 ml of isoelectric focusing gel solution (800 µL of solution is needed per capillary). a. Prepare one milliliter of CHAPS 30% (w/v) and NP-40 (nonidet P-40) 10% (w/v) (0.3 g CHAPS and 0.1 g NP-40 ) and degase for 5 min. b. Separately, prepare a second solution: 10 g urea (7 ml of water is added to dissolve the urea at warm temperature, around 35°C), 2.5 ml of acrylamide stock solution, 0.6 ml of ampholytes pH 4–8, 0.4 ml of ampholytes pH 3.5–10 and 20 µL of TEMED. c. Mix the first solution (CHAPS, NP-40) with the second one. Degase the mixture and add 40 µL of APS 10% (w/v) stock solution. Pipette one ml of this isoelectric focusing gel solution into the glass test tubes (along the side walls in order to prevent the formation of air bubbles in the solution). Fill up the capillary tubes by slowly pulling the syringe (up to the height of 16 cm). d. After 2 h of polymerization at room temperature, pull the capillary tubes out of the glass test tubes. Clean and gently rub the bottom of each capillary gel with parafilm. 3. Fill the lower chamber with the anodic solution. Wet the external faces of the capillary tubes with water and insert them in the isoelectric focusing chamber. Load the samples on the top of the capillary (cathodic side). Generally, thirty to 40 µL of the final diluted sample is loaded using a 25 µL Hamilton syringe (see Notes 3 and 4). 4. Lay down the cathodic buffer solution at the top of the sample in the capillary tube and then fill the upper chamber. Connect the upper chamber to the cathode, and the lower chamber to the anode. 5. Electrical conditions for IEF are 200 V for 2 h, 500 V for 5 h and 1000 V overnight (16 h) at room temperature (see Note 5). 6. After IEF, remove the capillary tubes from the tank and force the gels out from the glass tube with a 1 ml syringe which is connected to a pipet tip and filled with water (see Note 6). Put the extruded capillary gels on the higher glass plate of the polyacrylamide gel (see Subheading 3.3). Residual water around the capillary gel should be soaked up with a filter paper. Put 140 µL of IEF equilibration buffer down on the capillary gel. Immediatly push the capillary gels between the glass plates of the polyacrylamide gel using a small spatula. Place the cathodic side (basic end) of the capillary gel at the right side of the polyacrylamide gel. Care should be taken to avoid the entrapment of air bubbles between the capillary and the polyacrylamide gels. It is not necessary to seal the capillary gels with agarose solution but the contact between the capillary gel and the 9–16% polyacrylamide gradient gel should be very tight.

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3.3. SDS Polyacrylamide Gel Electrophoresis 1. In order to separate the majority of proteins, a 9–16% (w/v) polyacrylamide gradient gel is used for the second dimension. The gel is made as follows with 60 ml solution for a 9–16% (w/v) gel: 30 ml of 9% (w/v) acrylamide solution and 30 ml of 16% (w/v) acrylamide solution. Sixty milliliters are necessary to cast one gel of 1.5 mm × 200 mm × 160 mm (see Note 7).

Light solution: 9 ml of acrylamide stock solution (30% v/v) 7.5 ml 1.5 M Tris-HCl pH 8.8 (25% v/v) 150 µL 5% sodium thiosulfate solution (0.5% v/v) 15 µL TEMED (0.05% v/v) 13.2 ml water. This solution is degased and then 150 µL of 10% APS solution (0.5% v/v) is added.

Heavy solution: 16 ml of acrylamide stock solution (53% v/v) 7.5 ml 1.5 M Tris-HCl pH 8.8 (25% v/v) 150 µL of 5% sodium thiosulfate solution (0.5% v/v) 15 µL TEMED (0.05% v/v) 6.2 ml water. This solution is degased and then 150 µL of 10% APS solution (0.5% v/v) is added.

The gradient gel is formed using the Model 395 gradient Former (Bio-Rad) and a peristaltic pump. Immediately after the casting, the gels are gently over layered with a water-saturated 2-butanol solution using a 1 ml syringe. This procedure avoids the contact of the gel with air and allows to obtain a regular surface of the gel. Caution should be taken to avoid mixing the 2-butanol with the gel solution. The gels are stored overnight at room temperature to assure complete polymerization. 2. Prior to the second dimension separation, wash extensively the top of the gels with deionized water to remove any remaining 2-butanol. Remove the excess of water by suction with a syringe. 3. After the transfer of the capillary gel on top of the gradient polyacrylamide gel (see Subheading 3.2., step 6), fill the upper and the lower reservoirs with the running buffer and apply a constant current of 40 mA per gel. The separation usually requires 5 h. The temperature in the lower tank buffer is maintained at 10°C during the run. 4. At the end of the run, turn off the power, rinse briefly the gel in distilled water for a few seconds and process for gel staining. A typical gel pattern of plasma proteins stained with the ammoniacal silver nitrate method is shown in Fig. 2 (6, 7).

4. Notes 1. The addition of thiosulfate to the gel delays the appearance of background staining with the ammoniacal silver nitrate method (6, 7). 2. Human plasma proteins have been efficiently separated with the following sample preparation: 5 µL of plasma (containing approximately 400 µg

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Fig. 2. Silver stained plasma proteins separated by two-dimensional polyacrylamide gel electrophoresis using carrier ampholyte pH gradient in the first dimension. 1, albumin; 2, transferrin; 3, α1-antichymotrypsin; 4, IgA α-chain; 5, α1-antitrypsin; 6, fibrinogen γ-chain; 7, haptoglobin β-chain; 8, haptoglobin cleaved β-chain; 9, α2-HS-glycoprotein; 10, fibrinogen β-chain; 11, IgG γ-chain; 12, Ig light chain; 13, Apolipoprotein A-1; 14, haptoglobin α2-chain; 15, apolipoprotein J.

of proteins) are solubilized in 10 µL of lysis solution A (SDS-DTE) and heated in boiling water for 5 min. After cooling for 2 min at room temperature, 485 µL of lysis solution B is added. For silver stained gel, 30 to 40 µL of the final diluted sample is loaded on the top (cathodic side) of the capillary gel. 3. The best separation of complex protein mixtures is performed when less than 100 µg of proteins are loaded onto the capillary gel. Overloading may

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cause streaking and inadequate resolution of spots. The use of a highly sensitive staining method (silver staining) allow to applied a low amount of proteins onto the IEF gel. For silver stained gel, concentrations up to 25–40 µg per gel are enough. During the sample loading, it is very important to avoid the formation of air bubbles between the gel and the sample. After the loading, the Hamilton syringe should be withdrawn slowly along the side wall. If more than 20 kVh is applied during the isoelectic focusing, more cathodic drift occurs and the protein pattern is not stationary. Low pressure on the 1 ml syringe should be exerted to extrude the capillary gel. If too much pressure is applied, small lumps will be formed on the gel. Precise determination of the volume of solution necessary to cast the gels should first be done by measuring the volume of distilled water required to fill one resolving gel or the casting chamber.

References 1. Hochstrasser, D.F., Harrington, M.G., Hochstrasser, A.C., Miller, M.J., and Merril, C.R. (1988) Methods for increasing the resolution of two-dimensional protein electrophoresis. Anal. Biochem. 173, 424–435. 2. Golaz, O.G., Walzer, C., Hochstrasser, D., Bjellqvist, B., Turler, H., and Balant, L. (1992) Red blood cell protein map: a comparison between carrier-ampholyte pH gradient and immobilized pH gradient, and identification of four red blood cell enzymes. Appl. Theor. Electrophoresis. 3, 77–82. 3. Herbert B.R., Sanchez J.C., and Bini L. (1997) Two-dimensional electrophoresis : the state of the art and future directions, in Proteomic Research: New Frontiers in Functional Genomics (Principles and Practice) (Wilkins MR, Williams KL, Appel RD, Hochstrasser DF, eds), Springer Verlag, Berlin, pp. 13–33. 4. Appel R.D., Bairoch A., and Hochstrasser D.F. (1994) A new generation of information retrieval tools for biologists : the example of the expansy WWW server. Trends Biochem. Sci. 19, 258–260. 5. Pietrogrande, M.C., Marchetti, N., and Dondi, F. (2006) Decoding 2D-PAGE complex maps: relevance to proteomics. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 833, 51–62. 6. Hochstrasser, D.F. and Merril, C.R. (1988) Catalysts for polyacrylamide gel polymerization and detection of proteins by silver staining. Appl. Theor. Electrophoresis. 1, 35–40. 7. Rabilloud, T. (1992) A comparison between low background silver diammine and silver nitrate protein stains. Electrophoresis 13, 429–439.

33 Vertical Agarose Electrophoresis and Electroblotting of High-Molecular-Weight Proteins Marion L. Greaser and Chad M. Warren 1. Introduction Very large molecular weight proteins are difficult to separate by electrophoresis because of their poor penetration into gels using the Laemmli SDS polyacrylamide system (1). Protein migration in SDS gels has been found to be linear with the log of the molecular weight (2), so the larger the protein, the more poorly it is resolved from other big proteins. Many workers have attempted to solve this problem by using very low concentration acrylamide gels (3), acrylamide mixed with agarose (4), or acrylamide gradients (5) to better separate large proteins. Low concentration acrylamide gels are difficult to use because of their mechanical fragility and distortion during handling; these problems are magnified when blotting is attempted. An additional difficulty in blotting very large proteins is their poor transfer to the membrane. Inclusion of 2-mercaptoethanol in the transfer buffer improves transfer efficiency, but acrylamide gels stained after transfer typically still contain most of the giant muscle protein titin (6). A new electrophoresis system using SDS agarose for protein electrophoresis and blotting has been described (7). An example showing the resolution for several muscle samples containing large proteins is shown in Fig. 1. Migration distance shows a linear relationship with the log of the molecular weight (7). This system allows quantitative transfer of proteins from the gel with much higher reproducibility than can be achieved with methods using low percentage acrylamide. 2. Materials 2.1. Apparatus 1. SE 600 Slab Gel Unit with 16 × 18 cm glass plates (Hoefer) or a similar commercial gel unit (see Note 1).

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Fig. 1. SDS 1% agarose gel stained with silver. A centimeter ruler is shown on the left and the sizes of the various protein bands in KDa are listed on the right. DV, dog ventricle; RS, rat soleus; HV, human ventricle; HS, human soleus; CF, crayfish claw muscle. Human soleus titin is 3700 KDa and human ventricle has two titin bands of 3300 and 3000 KDa. The bands at 780 and 850 KDa are rat and human nebulin respectively. The myosin heavy chain is 223 KDa. Blotting proteins this size from acrylamide gels usually results in incomplete transfer, but full transfer can be achieved with agarose (7).

2. 3. 4. 5.

65°C oven. A constant current power supply. Circulating cooler. TE62 Tank Blotting Unit, Hoefer.

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2.2. Reagents 1. Acrylamide gel for plug; 38.5 % acrylamide. Weight 37.5 g of acrylamide (Biorad) and 1 g DATD (Biorad) into a beaker, add about 50 ml of water, stir till dissolved, dilute to 100 ml. Filter through a 0.45 micron filter (such as a Millex-HA, Millipore Corporation). Store in a brown bottle in the cold room (4°C). Danger! Avoid skin contact. 2. Reservoir and agarose gel buffer concentrate (5X); 0.25 M Tris, 1.92 M glycine, 0.5% SDS. Store at room temperature. 3. Ammonium persulfate; Prepare a 100 mg/ml solution in water; store frozen in 0.5 ml aliquots (stable indefinitely at −20°C). 4. Sample buffer; 8 M urea, 2 M thiourea, 0.05 M Tris-HCl (pH 6.8), 75 mM DTT, 3% SDS, 0.05% bromophenol blue (adapted from 8). (Dissolve urea and thiourea and treat with mixed bed resin to remove ionic constituents; then add remaining ingredients. Store at −20 °C). 5. 50% v/v glycerol. 6. Transfer buffer; 20 mM Tris, 150 mM glycine, 20% v/v methanol (7, 8) or 10 mM CAPS, pH 11 (Sigma Chemical Company) (9). For high molecular weight proteins, add SDS and 2-mercaptoethanol to 0.1% and 10 mM respectively to the transfer buffer. 7. SeaKem Gold agarose (Lonza Group Ltd.)

3. Methods 3.1. Gel Preparation 1. Volumes listed will provide enough solution for two 16 × 18 cm gels with 1.5 mm spacers. One is used for staining either with Coomassie blue or with silver with a special procedure for agarose (7), the other for blotting. 2. Clean plates and spacers with soap, rinse with distilled water and finally with ethanol. 3. Assemble gel plates. Place plate on clean bench top. Place spacers hanging half the way off each side of plate. Place second plate on top. Stand up plates and place one side into the clamp. Align spacer with side of plates and clamp and push spacer down so that bottom is flush with the glass plates (top buffer will leak if spacers are not flush with plates). 4. Pour acrylamide plugs in bottom of gel plate assembly (see Note 2): In a 15 ml plastic beaker add: 1.924 ml de-ionized water, 1.7 ml 50% glycerol, 2.12 ml 3 M Tris (pH 9.3), 2.72 ml acrylamide (40%), 24 µL 10% ammonium persulfate, and 13 µL TEMED. (see Note 2) Mix by pipeting a few times. Immediately add 2.5 ml to each gel assembly. Add a small amount of water on top of each plug to level the upper surface and provide an oxygen barrier. Allow gel to polymerize for 20–30 minutes. Drain off water layer by inverting gel plate assembly on a paper towel. 5. Place assembly, 20 lane sample combs, and 60 ml plastic syringe in a 65°C oven for 10 minutes (see Note 3).

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6. Weigh 0.8 g of SeaKem Gold agarose (see Note 4) into a 600 ml beaker (see Note 5). To a 100 ml graduated cylinder add 48 ml of 50% v/v glycerol (see Note 6), 16 ml 5X electrophoresis buffer, and bring volume up to 80 ml with deionized water. Place parafilm over top of the graduated cylinder, mix by inverting a few times, and pour solution into the 600 ml beaker containing the agarose. Place saran wrap over top of beaker and poke a few holes in the saran wrap. Weigh beaker with contents. Place beaker in a microwave oven along with a separate beaker of de-ionized water. Heat for a total of 2 minutes (stop every 30 seconds to swirl – protect hand with an insulated glove) (see Note 7). 7. Allow agarose to cool for a few minutes at room temperature. Reweigh, and add sufficient heated de-ionized water to replace that lost by evaporation. 8. Draw up about 40 ml of agarose in the prewarmed 60 ml Luer-Lock syringe and pour each gel slowly until it just overflows the top of the plates. Try to avoid formation of bubbles (if bubbles present, bring them to the top of the gel and pinch them with the sample comb). Insert sample combs and allow unit to cool at room temperature for about 45 minutes (see Note 8),

3.2. Electrophoresis Setup and Sample Loading 1. Add 4 liters of buffer to lower chamber (3200 ml de-ionized water plus 800 ml 5X electrophoresis buffer). Start cooling unit and stir bar (gels run at 6°C). 2. Prepare 600 ml upper chamber electrophoresis buffer (same concentration as lower chamber buffer). Add 2-mercaptoethanol (final concentration of 10 mM). Buffer will be poured into top chamber after samples are loaded and assembly placed in unit. 3. Take combs out of gels by bending them back and forth to detach from gel and slowly pull them up. Pour a small amount of upper chamber buffer into a 15 ml beaker and pipette buffer into first and last wells (the rest will fill over). Add buffer to remove any trapped bubbles. Insert pipette tip to deposit sample in bottom of the sample well. Skip the first and last lanes (see Note 9). 4. Running gels. Once samples are loaded, put upper chamber on the assembly. Pour upper chamber buffer into upper chamber from corners (do not pour buffer directly over wells). Place lid on unit, and connect to power supply. Turn electrophoresis unit on and run at 30 mAmps (2 gels) for 3 hours.

3.3. Staining and Western Blotting 1. After tracking dye reaches the bottom of the acrylamide plug, turn off the power and disassemble the plates. Cut off sample wells and acrylamide plug and discard. Soak the remaining agarose gel in 10 mM CHAPS (pH 11.0), 0.1% SDS, and 10 mM 2-mercaptoethanol for 30 minutes with gentle shaking. 2. The gel is then placed on top of either a sheet of PVDF or nitrocellulose, assembled into the transfer unit, and the protein electrophoretically transferred using 40 V constant voltage for 2–3 hours (see Note 10). 3. Blotted proteins can then be treated using conventional procedures with either colorimetric (horseradish peroxidase or alkaline phosphatase substrates) or ECL (enhanced chemiluminescence) methods.

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4. Notes 1. The agarose gel procedure works equally well with small format gels (i.e., 8 × 10 cm). 2. The acrylamide plug is used to prevent the agarose from slipping out of the vertical gel plate assembly. Use of DATD as the cross-linker provides a stickier bond of the acrylamide to the glass plates than if a conventional bisacrylamide cross-linker is used. Plugs can be poured a day before making the gel (place tape or parafilm over the top of the plates to prevent drying and store in cold room). 3. Preheating the glass plate assembly, well comb, and syringe prevents premature agarose gelling when the solution touches the colder surfaces. In addition the plates are less likely to crack during pouring if they are closer to the temperature of the hot agarose. 4. The supplier for SeaKem Gold agarose has changed twice since 2003. Biowhittaker was succeeded by Cambrex who was followed by Lonza Group Ltd, Muenchensteinerstrasse 38, CH-4002 Basel, Switzerland. 5. It is essential to use SeaKem Gold agarose for optimal migration of high molecular weight proteins. This type has large pore size and excellent mechanical stability. Other types of agarose may be used, but the protein mobility will be significantly reduced. 6. Glycerol is included in the mixture to increase the solution viscosity inside the gel and thus sharper protein bands. 7. Periodic swirling during the heating step eliminates non-hydrated agarose granules in the final gel. 8. Sample combs should extend no longer than 1 cm into agarose; otherwise they may be difficult to remove. Gels can be used right away or stored overnight in cold room. 9. Conventional sample buffers may not be dense enough for the sample to stay at the bottom of the well. If necessary add additional glycerol (up to 30% v/v final concentration) to increase sample density. 10. The disulfide bond formation of large proteins during electrophoresis also retards their migration out of the gel onto blots during transfer. Thus inclusion of 2-mercaptoethanol in the transfer buffer improves efficiency of transfer of high molecular weight proteins. The use of the agarose electrophoresis system with inclusion of 2-mercaptoethanol in the transfer buffer results in complete transfer of all high molecular weight proteins out the gel, including titin (Mr 3000 to 3700 KDa subunit size) (7). Acknowledgments This work was supported by the College of Agricultural and Life Sciences, University of Wisconsin-Madison, Purdue University College of Agriculture, and from grants (MLG- NIH HL77196 and Hatch NC1131.

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References 1. Laemmli, U. K.(1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 2. Weber, K. and Osborn, M. (1969) The reliability of molecular weight determinations by dodecyl sulfate-polyacrylamide gel electrophoresis. J. Biol. Chem. 244, 4406–4412. 3. Granzier, H. and Wang, K. (1993) Gel electrophoresis of giant proteins: solubilization and silver-staining of titin and nebulin from single muscle fiber segments. Electrophoresis 14, 56–64. 4. Tatsumi, R and Hattori, A. (1995) Detection of giant myofibrillar proteins connectin and nebulin by electrophoresis in 2% polyacrylamide slab gels strengthened with agarose. Anal. Biochem. 224, 28–31. 5. Cazorla, O., Freiburg, A., Helmes, M., Centner, T., McNabb, M., Wu, Y., Trombitas, K., Labeit, S., and Granzier, H. (2000)Differential expression of cardiac titin isoforms and modulation of cellular stiffness. Circ. Res. 86, 59–67. 6. Fritz, J. D., Swartz, D. R., and Greaser, M.L. (1989) Factors affecting polyacrylamide gel electrophoresis and electroblotting of high-molecular-weight myofibrillar proteins. Anal. Biochem.180, 205–210. 7. Warren, C. M., Krzesinski, P.R., and Greaser, M.L. (2003) Vertical agarose gel electrophoresis and electroblotting of high-molecular-weight proteins. Electrophoresis 2, 1695–1702. 8. Yates, L.D. and Greaser, M.L. (1983) Quantitative determination of myosin and actin in rabbit skeletal muscle. J. Mol. Biol. 168, 123–141. 9. Matsudaira, P. (1987) Sequence from picomole quantities of proteins electroblotted onto polyvinylidene difluoride membranes. J. Biol. Chem. 262, 10035–10038.

34 Two-Dimensional PAGE of High-Molecular-Mass Proteins Masamichi Oh-Ishi and Tadakazu Maeda 1. Introduction Many high-molecular-mass proteins (MW>100 kDa) are known to be involved in cytoskeleton, defense and immunity, transcription and translation in higher eukaryotic organisms. Though a variety of protein separation techniques have been described, at the moment purification of a high-molecular-mass protein remains a difficult task. O’Farrell (1) was the first to devise a two-dimensional gel electrophoresis (2-DE) technique which could detect more than 1000 spots in a gel. Though it is evident that this method was quite powerful in his days, the method is not particularly suitable in analyzing high-molecular-mass proteins larger than 200 kDa. When skeletal muscle, for example, is simultaneously analyzed by sodium dodecyl sulfite (SDS)-polyacrylamide gel electrophoresis (PAGE) and O’Farrell’s 2-DE method, myosin heavy chain (approx 200 kDa) is clearly seen in the former gel, but not in the latter. Hirabayashi (2) was the first to develop a 2-DE method which could analyze high-molecular-mass proteins, including myosin heavy chain and dystrophin, as large as 500 kDa (3). His trick was the use of agarose gel instead of polyacrylamide gel for the first-dimensional isoelectric focusing (IEF). Agarose gel, when used for IEF, can analyze much larger proteins than the polyacrylamide gel can. Oh-Ishi and Hirabayashi (4) further improved the method by adding 1 M thiourea and 5 M urea in an agarose IEF medium. Thiourea is a potent protein solubilizing reagent especially effective for high-molecular-mass proteins that could enter the first-dimensional agarose IEF gel. Here, we describe the 2-DE method with agarose gels in the first dimension (agarose 2-DE) that is compatible with analyzing high-molecularmass proteins.

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2. Materials 2.1. Equipment 1. Apparatus for first-dimensional agarose IEF (AE-6300 electrophoresis unit) ATTO (Tokyo, Japan). 2. Glass tubes (260 mm × 3.4 mm ID) for agarose IEF (ATTO). 3. Dialysis membranes. 4. Rubber bands. 5. Syringe to fit thin 30-cm-long polyethylene tubing. 6. Electrophoresis power supply (Crosspower 3500) (ATTO). 7. Horizontal electrophoresis apparatus (Multiphor II, GE Healthcare). 8. Immobiline DryStrips pH 3–10 L (GE Healthcare). 9. Perista pump (SJ-1211) (ATTO). 10. Glass tubes (300 mm × 5 mm ID) for protein fixation on agarose IEF gels. 11. Polyvinyl chloride (PVC) tubing (400 mm × 2 mm ID). 12. Vertical type apparatus for second-dimensional SDS-PAGE (NA-1200 electrophoresis unit) Nihon Eido (Tokyo, Japan). 13. Glass plates with a 1.5-mm thick frame (size 165 mm × 240 mm) for seconddimensional SDS-PAGE (Nihon Eido). 14. Plain glass plates (size 165 mm × 240 mm) for second-dimensional SDS-PAGE (Nihon Eido).

2.2. Reagents 1. Agarose IEF gels: Agarose IEF and pharmalyte were purchased from GE Healthcare. D-sorbitol, urea, and thiourea were from Wako Pure Chemicals (Osaka, Japan). Agarose IEF gels were prepared according to the protocols in section 3.1.1 Gel preparation and in Table 1. 2. Extraction medium: 7 M urea, 2 M thiourea, 2% CHAPS, 0.1 M dithiothreitol (DTT), 2.5% Pharmalyte, and protease inhibitors (Complete Mini ethylenediaminetetraacetic acid [EDTA]-free; Roche). 3. Overlaying solution: 4 M urea, 1 M thiourea. 4. Cathode buffer: 0.2 M NaOH. 5. Anode buffer: 0.04 M DL-aspartic acid. 6. Protein fixing solution: 10% trichloroacetic acid, and 5% sulfosalycylic acid. 7. Incubation medium: 2% SDS, 10% glycerol, 5% 2-mercaptoethanol, 0.02% bromophenol blue, and 0.05 M Tris-HCl, pH 6.5. 8. Staining solution: 0.02% PhastGel Blue R (coomassie brilliant blue R-350; GE Healthcare), 30% methanol, and 10% acetic acid. 9. Destaining solution: 30% methanol, and 10% acetic acid.

3. Methods 3.1. Agarose Gels for IEF 3.1.1. Gel preparation 1. We prepared the first-dimensional agarose IEF gels following the same procedure as described in our former report (5), to which we added the modifications described

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here. Three agarose IEF gels (180 mm in length and 3.4 mm in diameter) containing 1 M thiourea and 6 M urea were prepared by the present protocol. Agarose IEF (0.10 g) and D-sorbitol (1.2 g) were put into a 50 mL beaker, and dissolved in 5.8 mL distilled water at room temperature (solution A) (see Note 1). Solution A was boiled 10 times for 15 s in a microwave oven until the solution became clear, and was kept in a 70°C water bath for 5 min. A mixture of urea (3.6 g) and thiourea (0.76 g) powder was put into the solution A at 70°C, which we shall call solution B hereafter (see Note 2 and Table 1). The volume of solution B was then adjusted to 9.0 mL with distilled water, and was kept at room temperature. Solution B was divided into three test tubes: 1.0 mL for acidic, 4.5 mL for neutral, and 2.5 mL for basic solutions. Four kinds of Pharmalyte (pH 2.5–5 for acidic, pH 3–10 and pH 4–6.5 for neutral and pH 8–10.5 for basic solutions) were added to the respective tubes according to the protocol shown in Table 1. A glass tube (standard size: 260 mm × 3.4 mm ID) was prepared beforehand, the bottom of which was covered with a piece of dialysis membrane and tied with a rubber band. The glass tube was set to AE-6300 electrophoresis unit (apparatus for agarose IEF produced by ATTO). The acidic, neutral, and basic solutions were sucked in with one syringe each through thin 30-cm-long polyethylene tubing. First, the acidic solution was softly injected into the glass tube until the meniscus reached a height of 20 mm from

Table 1 Protocols for Making Agarose Isoelectric Focusing (IEF) gels

Solution B Pharmalyte pH 2.5–5 pH 3–10 pH 4–6.5 pH 8–10.5

Agarose I EF 0.10 g D-sorbitol 1.20 g Distilled w ater 5.80 mL Dissolved at 100°C—–> Solution A —————— Set a t 70°C ———————Urea 3.60 g Thiourea 0.76 g Dissolved at 50°C—–> Solution B Addition of distilled water until solution B comes to 9.0 mL ———— Set at room temperature —————– 1.0 mL 4.5 mL ↓ ↓ 100 mL — — 300 mL — 150 mL — — ↓ ↓ Acidic solution Neutral solution

2.5 mL ↓ — — — 250 mL ↓ Basic solution

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3.1.2. Sample Preparation 1. Freshly obtained mammalian tissues were cut into small blocks, quickly frozen in liquid nitrogen, and stored at −80°C until use. Each frozen tissue piece (about 10 mg) was homogenized with a Teflon glass homogenizer in an extraction medium (20 times the volume of the tissue pieces) (e.g. brain, muscle, kidney, and liver). Homogenates of the tissues were centrifuged at 112,000 g for 20 min, and the clear supernatant was subjected to the agarose IEF gel.

3.1.3. Sample Application 1. We added 10–200 mL of protein sample solution at the cathodic end of the gel, and gently filled the overlaying solution above the sample solution to the top of the glass tube. We added anode buffer to the lower reservoir, and cathode buffer to the upper reservoir. First-dimensional IEF was conducted at 600 V for 18 h at 4°C. 2. Then the agarose gel was transferred onto the top of the second-dimensional SDS gel, either directly or after proteins in the gel were fixed (see Note 4).

3.2. Second-Dimensional SDS-PAGE 1. Slab gels for second-dimensional electrophoresis were 12% polyacrylamide gels (200 mm × 120 mm × 1.5 mm). Second-dimensional SDS-PAGE was carried out according to the stacking system of Laemmli (6) with the slight modification of adding 1% SDS both in the stacking and separation gels. 2. The first-dimensional agarose IEF gel was loaded on top of the stacking gel without SDS equilibration (see Note 5) and covered with 1% agar solution to keep the agarose IEF gel in place. An incubation medium was overlaid on the IEF gels. 3. The second-dimensional gel electrophoresis was started with a constant current at 40 mA for 1 h and continued at 70 mA until the end of the run. 4. The slab gels were first soaked and shaken overnight in a destaining solution, which removed Pharmalytes from the gel (see Note 6). The slab gels were then stained with a staining solution containing PhastGel Blue R and destained with the destaining solution.

3.3. Comparison of IPG-Dalt with Agarose 2-DE 1. In this study, we compared the agarose 2-DE system with IPG-Dalt (immobilized pH gradients for IEF in the first dimension, SDS-PAGE in the second dimension) to assess their capability of analyzing high-molecular-mass proteins. 2. The first-dimensional IEF with IPGs was performed essentially as described by Gorg et al. (7). IPG dry strips (Immobiline DryStrips pH 3–10 L) and Multiphor II (GE Healthcare) were used in this study.

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Fig.1. Comparison of IPG-Dalt and agarose two-dimensional gel electrophoresis (2-DE). Tissues used are rat duodenum. (A) A 2-DE gel with immobilized pH gradients (IPGs) in the first dimension. (B) A 2-DE gel with agarose gels in the first dimension. Proteins loaded on an agarose isoelectric focusing gel and an IPG gel were 740 mg. Note that spot densities are different in the high-molecular-mass regions. A, actin; MHC, myosin heavy chain. The gels were stained with PhastGel Blue R.

3. Coomassie-stained 2-DE patterns of rat duodenum (Fig. 1) revealed that highmolecular-mass proteins larger than 150 kDa could be analyzed with the agarose 2-DE but not with IPG-Dalt.

4. Notes 1. Shake the beaker gently until a mixture of agarose IEF and D-sorbitol powder is completely suspended. 2. Immediately after the mixture of urea and thiourea powder was put into the beaker, solution B was stirred with a magnetic stirrer until the urea and thiourea were completely dissolved. 3. As a side effect of thiourea, a thiourea-urea agarose IEF solution does not gel at room temperature but at 4°C, and the gel formed at 4°C does not melt even when the gel is returned to room temperature. From the practical point of view of an experimental scientist, the 1 M thiourea/5 M ureaagarose IEF gel was a tremendous improvement over the 7 M agarose IEF gel originally used (2), because the agarose solution temperature no longer needed to be kept above 40°C when preparing agarose IEF gels. 4. When proteins in the agarose gel were fixed prior to the second dimensional electrophoresis, the gel was extruded into a 300-mm-long, 5-mm-diameter glass tube filled with a protein fixing solution containing 10% trichloroacetic acid and 5% sulfosalycylic acid. Both ends of the 5 mm diameter tube were connected to a Perista pump with PVC tubing to form a closed

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circuit filled with the fixing solution. When more than two gels were to be fixed, each of the gels was respectively extruded in a 5-mm-diameter glass tube, which was serially connected to one another to form a bigger closed circuit filled with the fixing solution. The proteins in the gel were fixed by 1 h circulation of the fixing solution in the 5-mm-diameter glass tube with a Perista pump, followed by 1 h circulation of 500 mL distilled water. 5. Avoid SDS equilibration step because the step easily spreads proteins out of the agarose gel. 6. Thorough removal of Pharmalytes from first-dimensional agarose IEF gel is useful in two respects: one is to reduce spot deformations by Pharmalyte pH 8–10.5 in the basic pI (>8) and low molecular mass (3000 V. We recommend using a power supply with an amperometer sensitive to µe), all species in solution. regardless of charge will eventually pass the detector. Separations on microchips are faster and more efficient, and higher sample throughput can be realized than with conventional systems. The ability to move small volumes of fluids around the chips makes it possible to analyze samples of limited volumes as well as integrate sample handling and preparation steps onto a single platform. The planar nature of microchip devices also facilitates the use of 2-D separation mechanisms for the resolution of complex mixtures. This will be discussed in further detail later. Microchip devices have been fabricated in a number of different substrates. Glass devices fabricated in soda lime, borosilicate, and quartz substrates have surface chemistries similar to that of the fused silica capillaries used in conventional systems. This allows the direct transfer of conventional methods to microchip systems. However, glass microchips have limitations, including the time required for fabrication, high cost, and fragile nature of the devices. In addition, integration of functional elements, such as separation and detection electrodes, is challenging due to the high temperatures needed for the glass bonding process. Inexpensive polymer substrates, including poly(methylmethacrylate) (PMMA) and poly(dimethylsiloxane) (PDMS), have been employed for the production of low-cost microchips (6). These disposable devices eliminate issues of sample carryover and cross-contamination. However adsorption of peptides to hydrophobic polymers such as PDMS can be an issue (7). In addition, PDMS generates a reduced EOF relative to that of glass, which can influence the efficiency of separation (see Note 1). Approaches to minimize peptide adsorption to channel walls include the use of dynamic and permanent modifications. Dynamic coatings have been used to minimize adsorption of hydrophobic peptides to channel surfaces. Polymers such as polyvinylpyrrolidine have been used to modify channels in a glass microchip, resulting in improved resolution of small test peptides (8). PDMS devices have also been modified with different monomers using UV grafting techniques (9). Reduced peak adsorption and improved peak efficiencies were observed for two model peptides, F-PKC and F-src. Poly(ethylene glycol) has also been grafted onto the surface of PMMA using atom-transfer radical polymerization (10). The modification resulted in a substantial reduction

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in EOF and nonspecific adsorption of proteins on the surface of the PMMA. The separation of a tryptic digest of BSA was demonstrated with the device. Dynamic coating of a PMMA microchip with sodium dodecyl sulphate (SDS) allowed the separation of proteins and peptides up to 116 kDa in size (11). Cathodic EOF was used as the driving force for the separation. For a more detailed look at approaches to th e permanent surface modification of polymer-based electrophoretic devices, the reader is referred to a recent review of the area (12). The major advantages of using planar microchip devices for peptide analysis include the ability to integrate sample preparation, injection, separation, and detection steps onto a single platform. Approaches to each of these stages of peptide analysis with respect to microchip technology will be examined in further detail. 2. Methods 2.1. Sample Preparation Preparation of peptide samples prior to analysis can involve dialysis, filtering, or extraction techniques to ensure that the sample is as clean as possible, preferably free from proteins and salts. Techniques such as solid-phase extraction can be used for preconcentration of peptides present at low concentrations in complex mixtures. While sample preparation is generally carried out offline, the integration of extraction, dialysis, and digestion steps for the development of a true miniaturized total analysis system has been implemented with some success. However, much of the work in this area has been proof-of-concept with limited application to real-world samples thus far (13). 2.1.1. Sample Preconcentration and Desalting Extraction and preconcentration of peptides for electrophoretic analysis is traditionally achieved using conventional chromatographic stationary phase materials. To use conventional stationary phase particles in microfluidic devices, a mechanism for effective particle containment is needed to prevent clogging of microchannels. This is typically achieved using tapered reaction chambers to promote keystone packing of particles or the use of weir-like structures. Packed C18 particles have been used to preconcentrate and to remove salt and urea from peptide mixtures (14). Limitations of particle-based systems include the generation of high backpressures and the potential for clogging. One solution to these problems involves the use of polymer monolith materials. Advantages of these continuous bed systems for sample extraction and preconcentration include low back-pressures, tailored chemistries and easy control of pore size. For microfluidic applications, monoliths can be formed in situ and polymerized using heat or UV light. Methacrylate-based monolith systems have been employed for preconcentra-

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tion of the hydrophobic tetrapeptide Phe-Gly-Phe-Gly by a factor of 1000 (15). Lauryl methacrylate monoliths have been polymerized in SU8 channels for desalting and subsequent separation of a cytochrome c digest peptide digest (16). Further optimization of column dimension was needed to improve the separation of the fragments. A PMMA substrate was modified with zeolite nanoparticles for the digestion and enrichment of protein samples (17). Trypsin immobilized in a sol-gel matrix was used to form a bioreactor for the digestion of cytochrome c and bovine serum albumin with a reaction time of under 5 s. A 2-D approach to sample preparation and desalting has also been described (18). The chip device contains two channels with integrated conductivity detectors. A combination of isotachophoresis followed by capillary zone electrophoresis was used for the preconcentration of myoglobin and for the desalting of a protein mixture. The salt was removed during the ITP step, allowing subsequent separation of proteins in the second channel. 2.1.2. Microdialysis Microdialysis is an in vivo sampling technique used for continuous monitoring applications. Probes are implanted in biological tissues and fluids to monitor biological events such as neurotransmitter release. A membrane with a predetermined molecular weight cutoff is used to exclude large molecules such as proteins while allowing smaller molecules such as amino acids and peptides to pass. This technique has recently been coupled to microchip electrophoresis (19). Fig. 2 shows a microchip CE separation of two fluorescently labeled peptides following on-line recovery through a microdialysis probe (19). This system has been further improved by the incorporation of an online labeling protocol for the detection of a peptide mixture that included Leu-enkephalin (19). 2.1.3. Protein Digestion: Proteomic Applications Proteolytic enzymes such as trypsin are used for the cleavage and digestion of protein molecules into their respective peptide fragments prior to identification/sequencing. Trypsin-coated beads have been employed for the on-chip digestion of proteins in microchip CE devices. High bead-to-peptide mass ratios and efficient mass transfer can permit rapid digestions of proteins including melitten, cytochrome c, bovine serum albumin, and β-casein. Digestion times ranged between 5 sec and 6 min at room temperature (20). This is in contrast to conventional protein digestion procedures that can often take up to 24 hours. Pressure or electroosmotic flow can be used to pump proteins through trypsin reactors (21). Localization of trypsin by enzyme adsorption to integrated membranes such as poly(vinylidene fluorine) has been used for the on-chip digestion of cytochrome c (22, 23). Trypsin has also been immobilized in monolithic supports and applied to the digestion of myoglobin (24). More recently, sol-gel

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encapsulation has been investigated and used for the digestion of arginine ethyl ester (ArgOEt) and bradykinin (25, 26) (see Note 2). Fig. 3 shows a microchip device with integrated digestion, separation and postcolumn derivatization for the detection of labeled peptide fragments of insulin B-chain (27). More recently, an on-chip enzymatic reactor was demonstrated for rapid protein digestion (28). Protease was encapsulated in a silica sol-gel in a poly(ethylene terephthalate) PET microchannel. The digestion of cytoplasmic proteins from a human liver was achieved in the silica-gel based reactor within a few seconds. Protein digestion followed by selective phosphopeptide enrichment has been demonstrated on a glass microchip device (29). The microchip consisted of two channels connected by a weir structure. The first channel contained agarose beads with immobilized trypsin while agarose beads with ferric ion were used in the second channel for immobilized metal ion affinity chromatography (IMAC). The device was also used for the enrichment of glycopeptides by replacing the IMAC beads with beads containing immobilized concanavalin A.

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Fig. 3. (A) Cross-sectional view of a CE microchip, heating element, and the thermocouples. (B) Schematic of the microchip used for on-chip reactions, incorporating separation and postcolumn labeling. The fluid reservoirs are: (1) substrate, (2) enzyme or DTT, (3) buffer, (4) sample waste, (5) NDA, and (6) waste. Reprinted with permission from (27).

2.2. Injection Once sample cleanup has been performed there are a number of different options for loading into the separation channel. Generally, this necessitates voltage control over multiple reservoirs (30). Popular methods include pinched and gated injections. Pinched injections demand simultaneous voltage control over all microchip reservoirs. A double T injector configuration reproducibly defines the volume of the sample plug. During separation, sample leakage into the channel is prevented by the application of a “pinching” or pushback voltage. Limitations of this approach include the need for multiple power supplies and the inability to change the volume injected using a double T design.

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In gated injection schemes, the sample plug is not volume defined and can be altered. A voltage is applied to the sample reservoir with the sample waste reservoir held at ground. A second voltage is applied at the buffer reservoir with the buffer waste reservoir held at ground. Two separate flowing streams are established within the device. For injection, the sample reservoir is floated and a small plug of sample enters the separation channel. Both flows are then reestablished for separation. This approach uses a simple T injector configuration and requires two power supplies. While injection volumes can be varied, sample and buffer streams must be of similar ionic strength. Further information on injection methods is included in a recent review (31). 2.3. Separation Methods 2.3.1. Capillary Zone Electrophoresis Capillary zone electrophoresis (CZE) has been investigated for peptide separations on microchip systems due to the simplicity of the buffer system (see Note 3). The separation of Gly-Leu and Gly-Gly on a glass microchip was demonstrated using a 20 mM borate buffer, pH 9.5 (32). A mixture of bradykinin, substance P, luteinizing hormone, bombasin, oxytocin, and enkephalins was separated on a PMMA device in 100 µM phosphate at pH 5.0 (33). A simple CZE buffer consisting of 15 mM boric acid, pH 9.2, was used for the separation of neuropeptides on PDMS microchip devices (34). Angiotensin peptides have been separated on both PDMS and glass microchip devices using 20 mM boric acid and 100 mM tris(hydroxymethyl)aminomethane, pH 9.0 (7). Fig. 4 shows separations of these peptides performed on both glass and PDMS microchips with the polymer devices displaying significantly lower separation efficiencies. This is most likely due to peptide adsorption to the channel surface (see Note 4). Methods to improve detection limits for microchip CZE of peptides include the use of polarity switching to initiate sample stacking (35). 2.3.2. Micellar Electrokinetic Chromatography (MEKC) In micellar electrokinetic chromatography (MEKC), separation is based on differences in the extent to which analytes partition between surfactant micelles and the surrounding aqueous buffer solution. When the concentration of a charged surfactant added to the buffer exceeds a value known as the critical micelle concentration (CMC), an aggregate, termed a micelle, is formed (36). The hydrophobic tails of the surfactant orient toward the center of the aggregate, and the polar headgroups orient toward the aqueous solution (37). A hydrophobic core is formed into which neutral hydrophobic analytes can partition (38). This method, initially developed for the separation of neutral molecules, is especially useful for peptides with a blocked N- or C-terminus. MEKC can also be used to separate charged peptides with similar electrophoretic mobilities but different

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Fig. 4. Separation and laser induced fluorescence detection of FITC-labeled angiotensin peptides (angiotensin I, angiotensin II, and angiotensin III) at a concentration of 500 nM using an injection voltage of 2000 V. The buffer consisted of boric acid, Tris (20 mM, 100 mM, pH 9.0). (A) Separation obtained using a Pyrex microchip. The separation voltage used is 3200 V with an anti-leak voltage of 1600 V. (B) Separation obtained using a PDMS microchip. The separation voltage used is 4000 V with an antileak voltage of 2000 V. Reprinted with permission from (113).

hydrophobicites. A microchip MEKC separation of lysozyme, trypsin inhibitor, and carbonic anyhydrase was accomplished on a PMMA microchip with 1% SDS in the TRIS-HCl run buffer (33). The separation of fluorescently labeled bovine serum albumin from other model proteins (cytochrome c, lysozyme, ribonculease A, myoglobin, and lactalbumin) in approximately 1.5 seconds on PDMS-based devices was accomplished by microchip-based MEKC (39). The separation efficiencies achieved were higher than those reported previously on native and modified PDMS microchip devices. MEKC has also been used in conjunction with 2-D separation schemes, which will be discussed later. 2.3.3. Isotachophoresis (ITP) Isotachophoresis (ITP) is a separation technique based on electrophoresis occurring at a uniform speed where analyte migration is independent of mobility. It has been termed a “moving boundary” electrophoretic technique (40). ITP employs a leading and a terminating electrolyte with the sample zone sandwiched between them. For successful implementation of ITP, careful control of sample conductivity is needed. This can be difficult to obtain for biological

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samples containing high salt concentrations. In contrast to CZE, ITP will not separate anions and cations in the same run because EOF is generally suppressed. For successful implementation of ITP, the leading electrolyte must have a greater mobility than any of the analytes in the sample, while the terminating electrolyte must have a lower mobility. When the electric field is generated, the ions of interest will migrate toward the electrode of opposite charge in zones determined by their electrophoretic mobility. Zone width adjusts so that all zones have same conductivity. After the initial zone formation, equilibrium is reached inside the channel, and the analyte ions then migrate past the detector at the same velocity. Narrowing of the analyte band can be accomplished by adding a high concentration of both the leading and terminating electrolyte. Microchip-ITP has been used as a preconcentration technique to improve detection limits for the determination of fluorescently labeled peptide substrates following enzyme assay (41). Other applications have included the preconcentration of cytochrome c following on-line protein digestion (22) and characterization of peptide phosphorylation by protein kinase (42). 2.3.4. Isoelectric Focusing Isoelectric focusing (IEF) separates peptides and proteins based on differences in their isoelectric point (pI). IEF can be used to separate proteins differing in pI by 0.0005 units or less (40, 43–46). An advantage of IEF is that the whole channel can be filled with sample, improving detection sensitivity. The sample is dissolved in a mixture of carrier ampholytes prepared in different pH ranges, and the channel is filled (47). Loading is followed by focusing and mobilization steps (Fig. 5). Once the sample has been loaded, a voltage is applied and the carrier ampholytes separate into individual bands, establishing a pH gradient inside of the microchip. There are three different types of behavior that the peptide can exhibit once the voltage is applied. If the pH of the band is lower than the pI, the peptide will be positively charged and will migrate toward the cathode. If the pH is higher than the pI, the peptide will be negatively charged and will migrate toward the anode and, if the pI is equal to the pH, the peptide is neutral and will not migrate. All peptides will migrate until the pH is equal to the pI. At this point the current will approach zero, the resistance in the channel will be high, and focusing will cease. Eventually, all of the peptides will be focused into bands where their pI is equal to the pH and they will stop migrating. One drawback of IEF is that, because there is a pH gradient formed along the channel, the extent of surface ionization varies. This can produce an uneven generation of EOF and cause band broadening. Therefore, the channel is usually modified to eliminate the EOF and to ensure that focusing is achieved prior to detection. This is accomplished by coating the microchip walls with a neutral coating, which also limits peptide adsorption (48).

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Fig. 5. Principle of capillary isoelectric focusing by simultaneous pressure/voltage mobilization. (A) Catholyte is backflushed past the detection point and a sample plug is introduced into the coated capillary (no high voltage). (B) Focusing of sample is complete and the sample components are driven toward the detector by a low-pressure rinse. High voltage is applied during this step. Reprinted with permission from (47).

For single point detection systems, analytes must be mobilized past the detector region following the focusing step. The two most common approaches to analyte mobilization are hydraulic or chemical (37). Hydraulic mobilization of the focused zones occurs by applying either pressure or a vacuum to one end of a channel. Chemical mobilization is accomplished through the addition of a salt, such as NaCl, to one of the buffer reservoirs, followed by application of a high voltage. The actual mobilization in this case occurs due to the presence of a competing ion such as Cl− that competes with OH−. This lowers the pH, and the peptides become charged and migrate toward the cathode. The Na+ competes with H+, causing an increase in pH and making more negative analytes migrate toward the anode. This pH shift occurs across the entire length of the microchip channel (37). The end result of this pH shift is the mobilization of the previously focused analytes toward the cathode. An alternative approach termed “one-step IEF” achieves focusing and mobilization simultaneously. As the pH gradient is not stationary, the EOF mobilizes the sample zones during focusing. One-step IEF can be accomplished using either coated or uncoated devices (49). EOF-driven mobilization is useful for microchip systems due to the speed of analysis and minimal instrumentation requirements. This approach was used for the separation of Cy5-labeled peptides; however,

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higher separation efficiencies and reproducibilities were achieved using chemical and hydraulic mobilization strategies (49). The requirement for sample mobilization past a single detection point can be obviated through the use of whole column imaging. This technique has been applied to the separation of test peptides using a UV absorbance detector (50). Due to reduced detection sensitivity, high concentrations of peptide were required, and glycerol was added to maintain peptide solubility. A more sensitive approach used fluorescence detection for microchip IEF of rhodamine green-labeled peptides on a PMMA device (51). Real-time imaging of the channel was performed using a scanning detection system that facilitated the determination of peptide migration times along with the estimation of EOF and pressure-driven flows. A scanning detection system based on acousto-optical deflection has also been developed and used for imaging of proteins during IEF (52). Advantages of this technology include fast response times, the absence of mechanical parts, and minimal background noise. It is also possible to perform on-chip mobilization with an on-chip diaphragm following isoelectrofocusing in the microchip channel (53). 2.3.5. Electrochromatography The development of capillary electrochromatography (CEC), a hybrid of CE and liquid chromatography (LC), has allowed highly efficient separations of peptide mixtures (54). In CEC, the channel is packed with a stationary phase for chromatographic analyte retention similar to LC, but electroosmotic flow is also generated. The main advantage of this technique over LC is improved peak efficiencies due to the flat plug flow profile generated by the EOF (48). The use of conventional bead-based retention systems in microfluidic devices requires methods (weirs, channels) to localize the beads and prevent device clogging. A 200-µm channel packed with octadecylsilyl (ODS) stationary phase particles was used to separate a fluorescently labeled angiotensin peptide from excess labeling agent (55). In chip-based electrochromatography, the stationary phase, the channel, inlets, and outlets can all be produced in one device using lithographic patterning techniques that are already employed for the fabrication of microfluidic devices. Collocated monolithic support structures (COMOSS) that are molded directly into the channel have been created in both quartz and PDMS (56, 57). Advantages of this approach include a uniform distribution of support structures defined during fabrication and homogenous channel dimensions. Issues involved with packing and retention of stationary phase particles are also eliminated as the reversed-phase stationary phase is bonded directly to the structures, and there is no need for the production of frits. Fig. 6 shows a COMOSS fabricated in PDMS. Applications of COMOSS modified with C18 phases include the

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Fig. 6. Scheme of a COMOSS separation column. Reprinted with permission from (57).

separation of tryptic digests of ovalbumin (56) and fluoroisothiocyanate-labeled bovine serum albumin (58). Monolithic polymer stationary phase materials are proving useful for microchip CEC applications (Fig. 7). Advantages of these phases include in situ polymerization, high flow rate tolerance, and functionalities that can be tailored to suit a specific application (59). In situ photopolymerization can be achieved by selective exposure to UV light. The location of the polymer can be accurately defined using a photomask. Acrylate-based porous polymers have been cast in glass channels for the separation of fluorescently labeled peptides, including angiotensins (60). The glass microchips could be reused following thermal incineration of the monolith at 550°C for 2 h and overnight incubation in 0.2 N NaOH.

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Fig. 7. Schematic of the microchip used for electrochromatography. B, S, BW, and SW denote reservoirs containing buffer, sample, buffer waste, and sample waste, respectively. The inset shows a scanning electron micrograph (SEM) of a channel cross section filled with photoinitiated acrylate polmer monolith. The mean pore diameter is 1 mm. Reprinted with permission from (60).

Sol-gels modified with polyelectrolyte multilayers are also being investigated for potential application as cation-exchange materials for microchip CEC separations of peptides including enkephalins (61). 2.3.6. Electrophoretic Bioaffinity Assays Electrophoretic bioaffinity assays combine the specificity of an immunological or enzymatic response with the separation power of electrophoresis. Separation and quantification of enzymatic conversion or antibody/antigen binding can be achieved using microchip CE. Potential applications of affinity separation technologies include biochemical, clinical, and drug development operations (62). 2.3.6.1. IMMUNOASSAYS

Immunoassay techniques are employed for the determination of binding constants for antibodies and antigens in competitive and noncompetitive studies. The separation of human IgM and antihuman immunoglobulin M has been achieved using a PMMA microchip device with conductivity detection (63). Tween surfactant was added to the run buffer to reduce the adsorption of the proteins to the channel wall. A PMMA device was also used for the separation of fluorescently labeled human anti-goat antibody and unlabeled goat immunoglobulin G (IgG). In this case, no significant adsorption of IgG protein was observed (64). MEKC has been employed for an immunoassay monitoring serum levels of theophylline (65). A competitive immunoassay to monitor theophylline in serum

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samples and a direct assay for the determination of monoclonal mouse immunoglobulin G in mouse ascites fluid were demonstrated on-chip using a tricine buffer containing Tween 20 and NaCl (66). A subsequent microchip system incorporating mixing, reaction, separation, and detection steps was developed for the same application (67). A microchip CE device that performed electrokinetic mixing of antibody to bovine serum albumin (BSA) and a diluting buffer was demonstrated for the on-chip preparation of calibration standards. These standards were then reacted with flourescein-labeled BSA, allowing the on-line generation of a standard curve. The device was used to assay the BSA antibody prepared from diluted mouse ascites fluid (68). A competitive chip-based immunoassay has been developed for the determination of insulin secretion from single islet cells (69). Online mixing of reagents allowed continuous sampling, and the simple device design needed only one voltage power supply for operation. A competitive immunoassay using rabbit polyclonal anti-cortisol antiserum and a fluorescently labeled cortisol derivative was demonstrated on a glass microchip device (70). The working range of the device was determined to be within clinical range (1–60 µg/dL cortisol). A heterogeneous competitive immunoassay of human immunoglobulin G (IgG) was demonstrated on a hybrid PDMS and glass device (71). Cy5-labeled human IgG was used as a tracer and Cy3-mouse IgG was used as internal standard. The microchip system was used for the determination of IgG levels in human serum with fluorescence detection. The determination of binding efficiencies between diketopiperazine receptors and Arg-containing peptides was demonstrated on a commercial Shimadzu microchip system (72). The separation times achieved with the microchip system were 50 times shorter than what could be achieved on a conventional system. 2.3.6.2. ENZYME ASSAYS

The determination of endogenous extracellular signal-regulated protein kinase was achieved using microchip CE to separate the substrate and product of the enzyme assay from an internal standard within 20 s. The device was used to measure the activity of endogenous levels of the enzyme in cell lysates (73). Fig. 8 shows a separation of unphosphorylated and phosphorylated fluoresceinlabeled “Kemptide” following on-chip enymatic conversion in a reagent well by protein kinase (74). The Km for “Kemptide” was determined using Lineweaver-Burk plots. A capillary array was used to demonstrate a multiplexed approach to the analysis of kinases (75). The assay was performed by the addition of multiple enzymes to a reaction tube. Simultaneous resolution of four product and three substrate peaks was demonstrated within 30 s. The multiplexed approach was

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Fig. 8. Time course of the Protein Kinase A (PKA) reaction on a 12A chip. Electrophoretic separation of unphosphorylated and phosphorylated fluorescein labeled Kemptide upon enzymatic conversion by PKA in a reagent well of the 12A microchip. Conditions: 13.3 mM Fl-Kemptide, 24.5 mM PKA in 100 mM Hepes, pH 7.5, 1 M NDSB-195, 5 mM MgCl2, 100 mM ATP, 50 mM cAMP, 0.1% Triton X-100, 10 mM DTT. Reprinted with permission from (74).

also tested using a phospholipase and another kinase, demonstrating the ability to screen different types of enzymes and potential inhibitors in one analysis. Another advantage of microchip systems is the ability to fabricate multiple channels for high throughput analysis. Microchip array electrophoresis has also been used to monitor the kinetics of a phosphate substrate and alkaline phosphatase conjugate as a function of reaction time (76). The enzymatic products were detected using laser-induced fluorescence with a miniature semiconductor laser. An electrochemical enzyme immunoassay was developed incorporating post-column reactions of alkaline phosphatase-labeled antibody (77). Amperometric detection was then used to identify the free antibody and the antibody-antigen complex. Microchip CE has been used for the separation of four different inhibitors of acetylcholinesterase following on-chip mixing of enzyme, substrate, inhibitor and derivatizing reagents (78). Acetylthiocholine is hydrolyzed to thiocholine by acetylcholinesterase. The reaction of thiocholine and coumarinylphenylmaleimide forms a thioether that can be detected using fluorescence. Modulation of enzyme activity was identified by a decrease in fluorescence intensity.

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2.3.7. Multidimensional separations Chromatographic and electrophoretic separation systems have a maximum peak capacity that is dependent on column length and separation efficiency. With LC the peak capacity typically ranges from 20 to 100, while for CE values of 50–100 can be obtained (79). For the analysis of complex biological samples, a one-dimensional system will have a limited peak capacity; therefore, the use of two-dimensional separations can extend the applicability of the technique. For 2-D separations, the output of the first separation technique serves as the input to the second. Therefore to successfully analyze each peak, the separation in the second dimension must be faster than in the first. A conventional capillary HPLC system has been interfaced to microchip electrophoresis for the separation of fluorescently labeled peptides from tryptic digests of bovine serum albumin (80). Effluent from the HPLC was injected into the CE channel every 20 s. On- chip coupling of 2-D separation systems is facilitated by the planar nature of the microchip device. A combination of MEKC (10 mM sodium dodecyl sulfate) and CZE was demonstrated on a microchip platform for the separation of tryptic peptides (79). Output from the first dimension was introduced to the second using a gated injection scheme. A similar approach was adopted for the coupling of open channel electrochromatography and CE on a glass microchip (81). Fig. 9a shows the device used with the spiral channel employed for the CEC, which was coated with a C18 phase connected

Fig. 9. CEC-CE chip. (A) The CEC serpentine channel extends from V1 to V2 (25 cm), the CE channel from V2 to the detection point y (0.8 cm), (B) 2-D contour plot of a β-casein tryptic digest. Reprinted with permission from (81).

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to the straight channel employed for CZE in the second dimension. Fig. 9b shows the application of the device to a 2-D separation of fluorescently labeled tryptic peptides of β-caesin. Acrylic devices have been employed for the on-chip coupling of IEF and CZE (82). The acrylic microchip supported a reduced EOF relative to that of glass, and this property was exploited as a slow mobilization method. When the output of the first dimension (IEF channel) is introduced to the second (CZE channel), the voltage is switched and applied to the CE channel only. As a result, the analysis time for the entire system is dominated by that of the second dimension (0.5%, either under reducing or non-reducing conditions. For reducing condition, β-mercaptoethanol or dithiothreitol is typically added to the denaturing solution. The following steps are recommended for denaturing protein samples: 1. The protein samples mixed with the denaturing solution according to the recommended protocol, are heat-denatured at 90–100°C for 3–5 min. 2. The samples are then further diluted with deionized water and/or sample buffer, usually by ~10–15 fold before use.

The protein samples should be analyzed in the same day after denaturization. 3.2. Planar Microfluidic Chips The procedures to perform protein separations on planar chips consist of chip preparation, sample loading, and placing the chip onto the instrument. Chip preparation requires the complete filling of the separation channels of a dry chip with the polymer matrix mixed with SDS and dye (gel-dye mix) as follows: 1. First, 12 µL of the gel-dye mix is pipetted into the A4 well in the lower right corner of the well pattern of the chip (see Fig. 5 for chip layout). 2. A priming station consisting of a syringe is used to fill the channels with the polymer solution by applying pressure of 3 atm at well A4 for 60 s. 3. Once the channels are filled, 12 µL of gel-dye mix is pipetted into wells B4, C4 and D3. 4. Next, 12 µL of the destaining solution (gel without SDS-dye) is pipetted into well D4. 5. Six µL each of sample diluted into the sample buffer containing lower and upper protein markers is pipetted in all 10 sample wells. 6. Six µL of the diluted ladder is pipetted into the ladder well D2.

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The chip is now ready for loading onto the instrument. Inside the instrument, the chip rests on a heater block and there is an electrode block located on the lid that matches the well pattern on the chip. After the lid is closed, each electrode pin makes contact with the liquid in each of the 16 wells, and voltage and current can be applied to the channels through the reagent wells. The optics is located below the chip to excite and record the fluorescence of the intercalating dye. The sizing experiment commences by activating the start button on screen. The following procedures have been programmed into the software script controlling the voltage and current, so they are transparent to the users. The script starts with a warm and focus step, which allows the chip to come to temperature equilibration with the heater block at 30°C and the auto focus function to locate and focus the light source onto the separation channel. During this time, the instrument also checks for electrical connections in all the wells before starting the separation analysis. The protein ladder is first loaded by applying an electric field between the source “ladder” well (D2) towards well B4. After the upper protein marker reaches the injection intersection, the voltage is switched well A4 to wells C4 and D4 for separation. A small aliquot of ladder protein with a band on the order of 50 µm is injected into the separation channel, and the protein species are separated by size through the SDS-dye containing polymer matrix followed by destaining and detection as described in Section 1. Overlap loading and separation of subsequent samples is performed to minimize the total analysis time. All the steps after the start of the experiment take about 30 min to complete. The results are digital recordings of fluorescence intensity as a function of transit time for the ladder and samples as shown in Fig. 7. The electropherograms can also be displayed as a virtual gel image by software, and a result table. The protein size is computed from the transit time as compared to a calibration curve determined by the transit time of the ladder after the upper and lower markers in the samples and ladder are aligned by the software. Relative protein concentration is computed from the area under each peak as compared to the known concentrations of protein in the ladder. 3.3. Sipper Microfluidic Chips Unlike the planar chip, which is shipped dry, a sipper chip comes in a wet container in which the microchannels, the sipper, and the wells of the chip are filled with storage buffer. Before starting an experiment, the chip should be prepared using the following steps: 1. After the chip is removed from the container, all the wells are rinsed with filtered deionized water. 2. Contents in wells 5, 6, and 8 are replaced with 75 µL of polymer matrix containing SDS and dye, contents in wells 2 and 7 are replaced with 75 µL of polymer matrix

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Fig. 7. Data output of the 2100 Bioanalyzer protein run.

without SDS, and addition of 120 µL of wash buffer to well 3 (see Fig. 6 for chip layout). 3. The chip is next placed in a priming station where the electrophoresis channels are filled with the polymer-dye matrix solution by applying a pressure of 3 atm to wells 2, 5, 6, 7, and 8 for 6 min. 4. The fluid in well 3 is then replaced with 75 µL of the polymer matrix. 5. The fluid in well 4 is replaced with 120 µL of marker dye solution.

The chip is now ready to be loaded in the instrument for high throughput analysis. When the sipper chip is placed onto instrument at the bottom of the optics module, the sipper capillary hangs out from the enclosed module where it can interface with the microtiter plate placed on the plate holder. The plate holder sits on a XYZ robotic arm and can accommodate the microtiter plate containing denatured protein samples and two well strips, one for the ladder and one for sample buffer. The buffer well is used to rinse the capillary between samples to eliminate cross-contamination. Inside the instrument, the sipper chip rests on a heater block which maintains the chip at a constant temperature of 30°C throughout the experiment. The initial chip warm up and focus step consists

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of sipping sample buffer by applying a vacuum in the waste well followed by multiple injection and separation. The excitation laser is focused onto the separation channel using an autofocus mechanism. At the start of an experiment, the sipper samples the ladder well to introduce onto the main channel about 100 nL of protein ladder by applying a small vacuum in the waste well. Once on chip, the protein sample from the sipper is mixed with the marker and flow toward the waste well 1. The mixture of protein and marker is next loaded onto the separation channel network electrokinetically by applying a voltage gradient between wells 3 and 6. Subsequently, the voltage is switched to wells 5 and 8 to inject a small aliquot of the mixture into the separation channel, which separates the protein species by size through the polymer matrix. Voltage is also applied to wells 2 and 7 for destaining as described in Subheading 1. The excitation laser and detector are located downstream of the destain intersection to record the fluorescence intensity of the protein fractions. During electrophoretic separation of the first sample, a second sample is simultaneously sipped from the plate to be ready for sample injection. The sipping operation of one sample does not affect the separation resolution of another due to the high matrix viscosity and shallower channel depth in the separation channel, which prevent significant pressure driven flow to occur in the separation channel. Such overlap sipping and separation processes are repeated sequentially until all the samples on the plate are analyzed. The protein ladder is sipped repeatedly after every 12 samples. A 96-well plate full of samples takes appropriately 1 hour and 20 min to analyze. The system can be used to analyze full or partially filled plates. It is also possible to do continuous or intermittent analysis throughout the day with a single chip that has been primed once. The sizing of protein species in each sample is computed after the lower marker is aligned by software with that in the ladder sipped prior to the sample, and the transit times are compared with a calibration curve of ladder transit time versus known protein ladder size. The protein concentration for each peak is computed using the relative area under the peak as discussed before after the peak areas are normalized by the area of the marker dye peak. Alternatively, the protein concentration can be more accurately determined using an internal standard added to each sample. The separated proteins can be viewed as electrophoregrams and a virtual gel. The computed sizes, relative concentrations, and % purity for each protein within each well are presented in a table once two bracketing ladders per row have been analyzed (see Fig. 8). The software has an “expected protein” feature to mark protein peaks detected at a predetermined size range. Automated calculation of protein purity can also be enabled. The results from the tables can be exported for further postprocessing analysis.

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Fig. 8. Data output of a LabChip® 90 protein run.

4. Notes 1. In microfluidics chips, keeping bubbles and debris away from the channels is important for good device performance and lifetime. As in capillary electrophoresis, bubbles and debris in microchannels are usually detrimental to the separation performance and longevity of a microfluidic device. Before introducing any fluid onto the chip, it is highly recommended that it should be filtered with a micron- or submicron-pore filter first. During manual pipetting of solutions into chip wells, it is important to touch the tip of the pipette to the bottom of the small opening of the glass plate inside the well, not on top of the glass plate or to the side wall of the well, to avoid introducing or trapping bubbles in the well. Samples placed in tubes and microplates should be centrifuged in order to settle particulates to the bottom. Subsequent manual pipetting or automated sipping should be taken from the top half of the tube or microplate wells to minimize introducing particulates into the channels of the chip. The planar chips should always be run with all the sample wells filled with the recommended volume of fluid. If there are fewer than 12 samples to be used in one device, the remaining wells should be filled with an

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equivalent amount of buffer to balance the hydrodynamics. The chip will not run properly if any wells are left empty. It is extremely important that the buffer and sample in the wells are thoroughly mixed using the vortex mixer at the conditions specified before loading the chip onto the instrument, otherwise protein concentration determination will not be reproducible. The speed of the vortex mixer should be set to the indicated setting at 2400 RPM. To avoid cross contamination, the electrodes in the instrument should be cleaned periodically as recommended in the user manuals. As mentioned in Subheading 2, the intercalating dye is light sensitive. Therefore, it is important to keep solutions containing the dye in the dark when not in use. Otherwise, when the dye degrades significantly, signal intensity and thus detection sensitivity could be affected. Because new sipper chips and many reagents are refrigerated before use, adequate time should be allowed for the chip and reagents to reach room temperature before starting the chip preparation steps outlined in Subheading 3.2. If cold reagents are introduced into the chip, warming up of an aqueous solution to room temperature in an enclosed microchannel could cause outgassing especially under slight vacuum condition. 2. Thus far, the detection sensitivity of proteins in commercial microfluidic chips is about the same as Coomassie Blue to mid-range of Colloidal Coomassie. It is advisable not to decrease protein sample dilution from the recommended ratio to try to increase sensitivity. Because protein buffers vary a lot in composition and conductivity due to different protein processing conditions, decreasing denatured protein sample dilution from the recommended value would result in more ionic species associated with the protein sample being injected into the separation column. It has been found that certain components of protein buffers, such as some detergents, in relatively high concentration in the injected sample band can affect the baseline flatness of the electrophoregram or the migration rate of the lower marker, potentially causing degradation in accuracy of sizing and relative concentration determination. 3. Microfluidic devices are now being more widely used for sizing and quantitation of proteins by research laboratories that previously had used SDSPAGE and to a lesser degree SDS-CGE. With greater use of microfluidic platforms, one obvious question is how well the measured molecular weight and relative concentration compares between the different methods. The experimental data up to now show these approaches provide similar answers for most proteins. For several proteins, minor migration variation relative to other proteins that result in differences in the determined size has been reported. The differences appear to be due to thevariations in the staining efficiencies by the fluorescent dye. It has been accepted by the SDS-PAGE user community that migration rates and dye staining of

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Electrophoretic Mobility, cm2 / V.S

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reasonable to propose that protein-SDS complexes migrate very similarly in the conventional and microfluidic equivalent of SDS-PAGE for protein sizing. The small discrepancies observed in a gel image generated by a SDS-PAGE run versus the virtual gel image of a microfluidic sizing run are potentially due to the differences in the polymer matrix and dye used. Since the gel images of two SDS-PAGE runs generated by reagents made by two different manufacturers do not necessarily match, one should not expect a perfect match of separation resolution and relative staining intensities between a conventional gel image and one generated by a microfluidic chip on the same protein sample. 4. Analysis of whole cell lysates is essential for protein expression profiling studies. Figure 11 shows a comparison of a lysate sample analyzed by a sipper chip and by SDS-PAGE. The over-expressed protein around 48 kD is detected well by bothmethods, although the data reproducibility appears better in the chip-based method. In analyzing lysates, it is important that the sample concentration to be within the upper limit of the recommended range, about 2000 ng/µL, in order to avoid the potential problem of sipper clogging by potential unsolubilized protein aggregates. Development and production of antibody therapeutics require a significant amount of characterization and monitoring of stability and aggregation of these

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Expressed protein 48 kDa

Fig. 11. A comparison of a LabChip® 90 virtual gel view and SDS-PAGE for a lysate sample (Data provided as the courtesy of Structural GenomiX of San Diego).

Fig. 12. Separation of reduced Humira (Abbott Laboratories) monoclonal antibody on Caliper LabChip 90. Abbreviations: light chain (LC), heavy chain (HC), and non-glycosylated heave chain (NGHC).

biological molecules. Microfluidic systems have been increasingly used for this application. Figure 12 shows an electropherogram of separation of monoclonal antibody measured in the sipper chip system. The separation of light chain, heavy chain, and non-glycosylated heavy chain is comparable with data generated by SDS-CGE, but at a 60-fold reduction in separation time on a microchip (10). Another protein separation application demonstrated in chips is cereal protein characterization. Fig. 13 shows an electrophoregram of wheat glutenin fractions. The composition of glutenin-subunit is an important predictor for the genetic potential of breeding lines, because these proteins are associated with differences in bread-making quality. Rapid separation and quantification of

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Fig. 13. (A) An electrophoregarm of wheat glutenin fraction under reduced conditions and (B) its corresponding virtual gel. Abbreviations: low molecular weight glutenin subunits (LMW-GS), high molecular weight glutenin subunits (HMW-GS), low marker (LM), and system peak (SP).

glutenin subunits have been achieved using microfluidic chip-based electrophoresis (11). However, the planar chips (Agilent 2100 Bioanalyzer platform) are limited to analysis of 10 samples at a time and thus not practical for large scale analysis. Glutenin subunits have also been analyzed on a high throughput integrated microfluidic-based platform (LabChip 90) (12). The electropherogram in Figure 13 illustrates the separation of the HMW-GS and LMW-GS into two distinguished fractions with high resolution and fast analysis time. It is well known in SDS-PAGE analysis that glycosylated proteins migrate slower than their expected molecular weights. Qualitatively similar phenomenon has been observed in microfluidic chip separation of glycosylated proteins. Therefore, similar precautions in data interpretation should be taken in chipbased analysis as in SDS-PAGE. 5. With some proper care and good laboratory practices, these microfluidics systems have been proven to be robust and labor-saving compared to the traditional SDS-PAGE. Perhaps more important, the output data is of high quality and available in a digital form which can be easily archived and shared with other laboratories across the company and around the world. Protein detection sensitivity down to silver-stain equivalence on microfluidic chips is not yet commercial, but approaches explored so far look promising (13). Moreover,

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Chow and Fathollahi many research studies have also been reported on interfacing microfluidic chips with electrospray and MADLI mass spectrometry (14) and a polyimide LC/MS chip has recently been commercialized by Agilent. Microfluidicsbased protein separation technologies have the potential to revolutionize the productivity of proteomics research in this era of increasingly more global community with lightning-speed demand for information sharing.

References 1. Shapiro, A.L., Vinuela, E., Maizel, J.V. Jr. (1967) Molecular weight estimation of polypeptide chains by electrophoresis in SDS-polyacrylamide gels. Biochem. Biophys. Res. Commun. 28, 815–820. 2. Laemmli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 3. Weber, K. and Osborn, M.J. (1969) The Reliability of Molecular Weight Determinations by Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis. J. Biol. Chem. 244, 4406–4412. 4. Bousse, L. Mouradian, S., Minella, A., Lee, H., William, K., and Dubrow, R. (2001) Protein sizing on a microchip. Anal. Chem. 73, 1207–1212. 5. Smith, D.E., Perkins, T.T. and Chu, S. (1996) Dynamical scaling of DNA diffusion coefficients. Macromolecules 29, 1372–1373. 6. Reynolds, J.A. and Tanford, C. (1970) The gross conformation of protein sodium sodecyl sulfate complex. J. Biol. Chem. 254, 5161–5165. 7. Guo, X. H. and Chen, S. H. (1990) The structure and thermodynamics of protein-SDS complexes in solution and the mechanism of their transport in gel electrophoresis proecess. Chem. Phys. 149, 129–139. 8. Narenberg, R., Kliger, J., Horn, D. (1999) Angew. Chem., Int. Ed. Engl. 38, 1626. 9. Viovy, J.-L. (2000) Electrophoresis of DNA and other polyelectrolytes: physical mechanisms. Rev. Modern Phys. 72, 813–872. 10. Sala-Solano, O.,Tomlinson, B., Du, S., Parker, M., Strahan, A., and Ma, S. (2006) Optimization and Validation of a Quantitative Capillary Electrophoresis Sodium Dodecyl Sulfate Method for Quality Control and Stability Monitoring of Monoclonal Antibodies. Anal. Chem. 78, 6583–6594. 11. Uthayakumaran, S., Listiohadi, Y., Baratta, M., Batey, I.L., Wrigley, C.W. (2006) J. Cereal Sci., 44, 34–39. 12. Ben Abda, O., Rhazi, L., Fathollahi, B., Mejri, S., and Aussenac T. (2007) High throughput microfluidic system for separation and quantification of high-molecularweight glutenin subunits, AACC International Annual Meeting, October 7–10, San Antonio, TX. 13. Molho, J.I., Park, C., Price, K., Phan, H., Simeonov, A., Mouradian, S. and Spaid, M.A. (2004) Ultrasensitive protein sizing using integrated isotachophoresis - gel electrophoresis,” in Proceedings of the 8th International Conference on Miniaturized Systems for Chemistry and Life Sciences (Northrup, M.A. ed.), Royal Society of Chemistry, Cambridge, UK, 387–389. 14. Mouradian, S. (2002) Lab-on-a-chip: applications in proteomics. Curr. Opin. Chem. Biol. 6, 51–56.

39 Difference Gel Electrophoresis (DIGE) David B. Friedman and Kathryn S. Lilley 1. Introduction The proteome is a dynamic entity, constantly changing both in levels of protein expression as well as in posttranslational modification and localization, all of which are hidden in the static DNA code. The repertoire of techniques and associated instrumentation which is now being applied to proteomics experiments is expanding exponentially, and although a complete visualization of the proteome is still beyond reach of any single technique, several technology platforms exist that can provide complementary datasets. Proteomics in the clinical setting is also developing, providing a major impact on the way in which diseases will be diagnosed, treated and monitored (1). In particular, approaches involving two-dimensional (2-D) gel electrophoresis (2-DE) separations and subsequent protein identification using mass spectrometry (MS) have been useful for imaging thousands of intact proteins (including modified/proteolyzed forms) in a single run, but quantification has been challenging, particularly over a large sample set. Difference gel electrophoresis (DIGE) has proven to be a powerful quantitative technology for differential-expression proteomics on a global level, where the individual abundance changes for thousands of intact proteins can be simultaneously monitored in replicate samples over multiple variables with statistical confidence (see Note 1). This includes quantitative information on protein isoforms that arise due to a charged post translational modification that results in a change in the isoelectric point of the protein (such as acetylation or phosphorylation), and splice variants or protein processing events, all of which can be resolved for individual quantification and subsequent analysis by mass spectrometry. DIGE is based on conventional 2-D gel technology that is capable of resolving several thousand intact proteins first by charge using isoelectric focusing (IEF)

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and then by apparent molecular mass using SDS-PAGE (2, 3) (see Note 2). Importantly, DIGE overcomes many of the limitations commonly associated with 2-D gels such as analytical (gel-to-gel) variation and limited dynamic range that can severly hamper a quantitative differential-expression study. This is accomplished using up to three spectrally resolvable fluorescent dyes (Cy2, Cy3 and Cy5, refered to as CyDyes) that enable low- to sub-nanogram sensitivities with >104 linear dynamic range, and then by multiplexing the prelabeled samples into the same analytical run (2-D gel). Multiplexing in this way allows for direct quantative measurements between the samples coresolved in the same gel, and is therefore beyond the limitations imposed by between-gel comparisons with conventional 2-D gels. The highest statistical power of this multiplexing approach stems from the utilization of a pooled-sample internal standard comprised of an equal aliquot of every sample in the experiment (see Subheadings 1.2.1., and 3.3.3.,). With this method, two dyes (Cy3 and Cy5) are used to individually label two independent samples from a much larger experiment, and the Cy2 dye is used to label an internal standard which is comprised of an equal aliquot of proteins from every sample in the experiment. This pooled-sample internal standard is labeled only once in bulk to avoid additional technical variation, and enough is prepared and labeled to allow for an equal aliquot to be coresolved on each gel. The three differentially labeled samples are then coresolved on the same 2-D gel, after which direct measurements can be made for each resolved protein using the spectrally exclusive dye channels without interference from technical variation of the separation (gel-to-gel variation). Rather than making direct quantitative measurements between the two samples in the gel, the measurements are instead made relative to the Cy2 signal from the internal standard for each resolved protein. This is done for each protein form under investigation separately, and so each feature is quantified relative to its own unique internal standard. The Cy2 signal should be the same for a given protein across different gels because it came from the same bulk mixture/ labeling; therefore, any difference represents gel-to-gel variation, which can be effectively neutralized by normalizing all Cy2 values for a given protein across all gels. Using the Cy2 signal to normalize ratios between gels then allows for the Cy3:Cy2 and Cy5:Cy2 ratios for each protein within each gel to be normalized to the cognate ratios from the other gels, encompassing all samples. Each gel may contain different (and/or replicate) samples in the Cy3 and Cy5 channels, but all can be quantified relative to each other because each is measured to the cognate Cy2 signal from the internal standard present on each gel. With the use of sufficient replicates, a plethora of advanced statisitcal tests can be applied to highlight proteins of interest whose change in expression are related to the biology under investigation. Since the technical noise is low, these vital replicates should be independent (biological) replicates. In a final step, specific proteins of interest are then identified using standard mass spectrometry approaches on gel-resolved proteins that have been excised

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and proteolyzed into a discrete set of peptides. Briefly, excised proteins are subjected to in-gel digestion with trypsin protease (typically), and mass spectrometry (MS) is used to acquire accurate mass determinations on the resulting peptides and peptide fragments generated using tandem mass spectrometric approaches. The mass spectral data are then used to identify statistically significant candidate protein matches through sophisticated computer search algorithms that compare the observed MS data with theoretical peptide masses (using data generated by peptide mass fingerprinting) or collision-induced fragmentation patterns (obtained from tandem mass spectrometry) generated in silico from protein sequences present in databases. 1.1. Optimizing Sensitivity and Resolution There are currently two forms of CyDye labelling chemistries available: minimal labeling involving the use of N-hydroxy succinimidyl ester reagents for low-stoichiometry labeling of proteins largely via lysine residues, and saturation labeling which utilize maleimide reagents for the stoichiometric labeling of cysteine sulfhydyls. The most established DIGE chemistry is the “minimal labeling” method, which has been commercially available since July 2002. Here the CyDye DIGE Fluors are supplied as N-hydroxy succinimidyl (NHS) esters, which react with the ε-amine groups of lysine side-chains. The three fluors are mass matched (c. 500 Da), and carry an intrinsic +1 charge to compensate for the loss of each proton-accepting site that becomes labeled (thereby maintaining the pI of the labeled protein). Each dye molecule also adds a hydrophobic component to proteins, which along with MW, influences how proteins migrate in SDS-PAGE. Minimal labeling reactions are optimized such that only 2–5% of the total number of lysine residues are labeled, such that on average a given labeled protein would contain only one dye molecule. This is necessary because lysine is very prevalent in protein sequences, and multiple labeling events may affect the hydrophobicity of some proteins such that they may no longer remain soluble under 2-DE conditions. Although a given protein form may exhibit specific labeling efficiencies, these will be the same for labeling with all three dyes, allowing for direct relative quantification. Minimal labeling with CyDye DIGE fluors is very sensitive, comparable to silver-staining or many post electrophoretic fluorescent stains (ca. 1 ng, see Subheading 2.2.6), but with a linear response in protein concentration over four orders of magnitude (4) (see Note 3). For maleimide-labeling of the cysteine sulfhydryls, the overall lower cysteine content in proteins allows for labeling of these residues to saturation without increasing the overall hydrophobicity of the proteins to cause insolubility problems. Saturation labeling is ultimately more sensitive (150–500 picograms, and even more so for proteins with high cysteine content). Its use is not as commonplace, most

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likely due to the availability of only Cy3 and Cy5 with this chemistry (see Note 4), the fact that it is blind to the small but significant population of non-cysteine containing proteins, and the additional optimization of complete cysteine reduction necessary for reproducible labeling. For these reasons, saturation DIGE is usually reserved for experiments where samples are limited, where the advantage of the increased sensitivity out-weigh these additional considerations. To maximize the information that can be gained from experiments which utilize the DIGE technology, it is imperative that resolution of protein species within gels is optimized. Although single 2-DE runs can resolve proteins with pI ranges between pH 3–11 and apparent molecular mass ranges between 10–200 kDa, higher resolution and sensitivity can be obtained by running a series of medium range (e.g. pH 4–7, 7–11) and narrow range (e.g., pH 5–6) isoelectric focusing gradients with increasing protein loads, leading to an overall more comprehensive proteomic analysis (2, 3, 5) (see Note 5). This is analogous to gaining increased resolution and sensitivity in an LC/MS-based strategy by using multiple HPLC columns with different affinity chemistries (e.g., MudPIT (6) ). Much of the sensitivity limitation associated with 2-D gels can be attributed to the analysis of unfractionated, whole-cell and whole-tissue extracts. Additional sensitivity can be gained via enrichment for the proteins of interest, such as by analyzing prefractionated or subcellular samples, or immune complexes. However, the additional experimental manipulations required for prefractionation introduce more technical variation into the samples and necessitates increased independent (biological) replicates (which of course can be accommodated with the DIGE internal standard methodology). The identification of proteins of interest using mass spectrometry can be performed directly from the DIGE gels when protein amounts have been optimized in this way (see Subheading 3.5.,). Alternatively, some experimental approaches perform the DIGE analysis using “analytical” gels with lower protein amounts, followed by protein excision from a secondary, “preparative” gel with higher protein amounts. This approach has its advantages when dealing with small sample amounts, such is often the case using the Saturation dye chemistries, but is also prone to uncertainties that arise due to the disproportionate amount of protein loading (see Note 6). The methods presented in this protocol are for optimization of both the DIGE data as well as material for subsequent mass spectrometry using high protein loads. 1.2. Optimizing Statistical Significance 1.2.1. Using the Internal Standard The ability to coresolve and compare two or three samples in a single gel is attractive because it allows for direct relative quantification for a given protein without any interference from gel-to-gel variations in migration and resolution, removing the need for running replicate gels for each sample (similar to stable

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isotope LC/MS-based strategies) (44–47). This approach has limited statistical power, however, since confidence intervals are determined based on the overall variation within a population (see Subheading 3.6.2.,). Many researchers new to DIGE technology are not immediately aware of the increased statistical advantage and multiplexing capabilities of DIGE when combining this approach with a pooled-sample mixture as an internal standard for a series of coordinated DIGE gels (7). This design will allow for repetitive measurements (vital to any type of experimental investigation), and in such a way as to control both for gel-to-gel variation and provide increased statistical confidence. Importantly, these replicates should be from independent experiments (biological replicates) rather than multiple samples from the same experiment (technical replicates) (see Subheading 1.2.2.,). In this way, statistical confidence can be measured for each individual protein based on the variance between multiple measurements, independent of the variation in the population. Incorporating independently prepared replicate samples into the experimental design also controls for unexpected variation introduced into the samples during sample preparation. This more complex and statistically powerful experimental design is accomplished by using one of the three dyes (usually Cy2) to label an internal standard which is comprised from equal aliquots of protein from all of the samples in an experiment. The total amount of the Cy2-labeled internal standard is such that an equal aliquot can be coresolved within each DIGE gel that also contains an individual Cy3- and Cy5-labeled sample from the experiment. Since this standard is composed of all of the samples in a coordinated experiment, each protein in a given sample should be represented in the mixture and thus serve as an unique internal standard for that protein (see Note 7). Direct quantitative comparisons are made individually for each resolved protein between the Cy3or Cy5-labeled samples and the cognate protein signal from the Cy2-labeled standard for that gel (without interference from gel-to-gel variation), enabling the calculation of standardized abundance for every spot matched across all gels within a multigel experiment. The individual signals from the internal standard are also used to normalize and compare between each in-gel direct quantitative comparison for that particular protein from the other gels. Using the Cy2-labeled standard in this fashion, therefore, allows for more precise and complex quantitative comparisons between gels, including independent (biological) sample repetition (Fig. 1). Importantly, the internal standard experimental design allows for the identification of significant changes that would not have been identified if the analyses were performed separately, even when using Cy3- and Cy5-labeled samples on the same DIGE gel (8). This experimental design also allows for multivariable analyses to be performed in one coordinated experiment, whereby statistically

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Fig. 1. Illustration of DIGE and experimental design using the mixed-sample internal standard. (A) Representative gel from a 6-gel set containing three differentially labeled samples: Cy2-labeled internal standard, Cy3-labeled sample #1, and Cy5-labeled sample #2. The individual protein forms all coresolve in this one gel, but these three independently labeled populations of proteins can be individually imaged using mutually exclusive exitation/emission properties of the CyDyes. (B) Schematic of the sample loading matrix indicating gel number, Cydye labeling and three replicates (indicated as “1, 2, and 3”) of the four conditions being tested (A, B, C, and D). Within the boxed regions representing each labeled sample is depicted a theoretical protein that is upregulated in condition D. Dotted lines illustrate how the protein signals from each sample are directly quantified relative to the Cy2 internal standard signal for that protein without interference from gel-to-gel variation, and how the Cy3:Cy2 and Cy5:Cy2 intra-gel ratios are normalized between the 6 gels. (C) A graphical representation of the normalized abundance ratios for this theoretical protein change. Adapted from (5).

significant abundance changes can be quantitatively measured simultaneously between several sample types (e.g., different genotypes, drug treatments, disease states), with repetition and without the necessity for every pair-wise comparison to be made within a single DIGE gel (9, 10) (see Note 8).

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1.2.2. Assessing Intersample Variation Even under the best-controlled situations there is always room for technical variation in sample preparation. Moreover, there will always be some degree of biological “noise” that represents fluctuations in protein expression that fall into the normal range. In the majority of cases, however, biological variation usually outweighs technical variation. Clinical proteomic experiments are especially hampered by biological noise that describes the variation between patient samples that is not associated with disease. The largest proportion of this variation comes from biological diversity, but a significant amount may also come from variable collection and storage of biological samples. It is of vital importance to identify changes in protein expression that describe the biological processes under investigation rather than variation derived from technical or biological noise. In order to gain the more robust data sets necessary to be able to draw accurate conclusions, it is therefore of paramount importance to collect and store samples using very stringent and closely adhered to protocols. For clinically related samples it is also necessary to assess the biological variation within the population being tested and also within a single individual. Interindividual variation has been the focus of several studies (11, 12) and determining a typical diversity within a single patient (i.e. taking longitudinal samples and assessing variability in protein abundance) and between patients will determine the minimum number of patient samples required for an experiment. This is an essential step before embarking on any large-scale and potentially costly DIGE experiment. Without this type of pretest, the results of underpowered experiments run the risk of being peppered with false information (both false positives and negatives). As with all complex technologies, the DIGE technique itself is subject to technical variation which will be laboratory specific to a greater or lesser extent. However, the amplitude of this variation is generally outweighed by the biological variation associated with a typical sample set (13). 1.2.3. Univariate Statistical Analyses To date, the majority of published quantitative proteomics studies using the DIGE technology have applied a univariate test, such as a Student’s t-test or Analysis of Variance (ANOVA), to identify protein species with significant changes in expression (14). These tests calculate the probability (p) that the samples being compared are the same and therefore any apparent change in expression occurs by chance alone. An expression change is considered significant if the calculated p-value falls below a prescribed significance threshold, typically 0.05 (whereby 1 in 20 tests may give a change in expression by chance). For more stringent analyses, a p-value of 0.01 is often used as the significance threshold.

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When employing these tests on DIGE datasets there are several factors that must be considered if correct assumptions are to be made from ensuing analyses. Student’s t-tests and ANOVA assume that the data achieved is normally distributed and that any variance is homogeneous. The measurement and correction of systematic bias within DIGE experiments has been the subject of several studies which chart methods to optimize normalization of data sets (15–17). Another important consideration is that of false discovery rate (FDR) which could arise as a result of statistical tests such as the ones described above. These tests involve the simultaneous and independent testing of thousands of protein features. The probability of a false-positive being recorded for each test is such that a substantial number of false positives may accumulate. There are several approaches to determine the FDR and adjust p-values to compensate for this, the most widely used to date being the Benjamini and Hochberg method, whose use in conjunction with DIGE data has been described by Fodor et al. (15). 1.2.4. Multivariate Statistical Analyses Discovery phase proteomics often produce large lists of proteins that are identified as changing significantly in the experiment, many of which may well be false-positives. Another complementary approach to overcome issues surrounding univariate tests is the application of additional multivariate statistical analyses to these datasets, which can help to filter out false-positives that result from wholesample outliers (i.e., sample mis-classification and/or poor sample preparation technique). These analyses, such as Principal Component Analysis (PCA), partial least squares discriminate analysis (PLS-DA) and unsupervised hierarchical clustering (HC) (see Figs. 2 and 3) have recently been applied to DIGE datasets (5, 18–26). Raw and normalized data can be exported from most DIGE software solutions (e.g., DeCyder, Progenesis), and several multivariate analyses are now part of an extended data analysis software module as part of the DeCyder suite of software tools (GE Healthcare) which was specifically developed for DIGE analysis (see Subheading 3.6.,). Similar capabilities can also be found in newer versions of the Progenesis software (Non-Linear Dynamics) and Delta2D [Decodon]. These multivariate analyses work essentially by comparing the expression patterns of all (or a subset of) proteins across all samples, using the variation of expression patterns to group or cluster individual samples. Technical noise (poor sample prep, run-to-run variation) and biological noise (normal differences between samples, especially present in clinical samples) is almost always associated with any analytical dataset of this nature, and may well override any variation that arises due to actual differences related to the biological questions being tested. Unsupervised clustering of related samples, therefore, adds additional confidence that a “list of proteins” changing in a DIGE experiment are not arising stochastically (5).

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Dfur

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PC1 Fig. 2. Illustration of the use of Principal Component Analysis. DIGE was used to analyze changes in Staphylococcus proteins in reponse to genetic and chemical alterations affecting iron utilization. Adapted from (18).

1.3. DIGE in the Clinical Setting Although the potential for DIGE to address clinical studies is only beginning to be addressed (for example, see (23, 24) ), many studies have been published demonstrating the feasibility and benefit of DIGE/MS using small patient cohorts for preliminary studies in colon (8), liver (27–29), breast (30, 31), esophageal (32, 33), and pancreatic cancers (34), as well as other important clinical studies such as SARS (35). Many studies also explore the important benefit of procuring samples using laser capture microdissection (LCM) for a highly enriched population of the cells under study (10, 24, 36–38). These LCM studies necessitate the use of the saturation chemistry owing to the increased sensitivity but limited multiplexing power, and typically require secondary preparative gels with higher protein loads to enable protein identification by mass spectrometry. The study of Suehara et al. (23) represents the utility of a multivariable DIGE/MS analysis with an extended sample set pertinent for a clinical study. Eighty soft-tissue sarcoma samples comprising seven different histological backgrounds were analyzed. Using the saturation DIGE fluors, individual samples were labeled with Cy5 and multiplexed with a pooled-sample internal standard (labeled in bulk with Cy3) for each DIGE gel. Using high-resolution 2-D gel separations and a combination of multivariate statistical tools (support vector machines, leave-one-out cross-validation, PCA and HC), these studies identified a small subset of proteins including tropomyosin and HSP27 that were able to discriminate between the different classes of tumors. HSP27 in particular was part of a subclass of discriminating proteins that could distinguish between leiomyosarcoma and malignant fibrous histiocytoma (MFH), as

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well as correlate with patient survival between low-risk and high-risk groups. HSP27 has long been associated with prognosis in MFH as well as in other human carcinomas (39). This chapter assumes a solid understanding in 2-D gel electrophoresis, and will focus on the design and implementation of the DIGE method using the pooled-sample internal standard methodology and the minimal dye chemistry for Cy2, Cy3 and Cy5, with notes provided for saturation labeling chemistry. 2. Materials 2.1. Cell lysis buffers 1. TNE: 50 mM Tris-HCl pH 7.6, 150 mM NaCl, 2 mM EDTA pH 8.0, 2 mM DTT, 1% (v/v) NP-40. 2. RIPA buffer: 50 mM Tris-HCl pH 8.0, 150 mM NaCl, 1% NP-40, 0.5% deoxycholic acid, 0.1% SDS. 3. 2-D gel electrophoresis lysis buffer: 7 M urea, 2 M thiourea, 4% CHAPS, 2 mg/mL DTT, 50 mM Tris-HCl pH 8.0. 4. ASB14 lysis buffer: 7 M urea, 2 M thiourea, 2% amidosulfobetaine 14, 50 mM Tris-HCl pH 8.0.

Note: Depending on the sample, it may also be necessary to add protease inhibitors (Complete protease inhibitor cocktail tablets. Roche Diagnostics) and phosphatase inhibitors (sodium pyrophosphate (1 mM), sodium orthovanadate (1 mM), betaglycerophosphate (10 mM) and sodium fluoride (50 mM) ) to the chosen lysis buffer (see Subheading 3.1). 2.2. SDS-Polyacrylamide Gel Electrophoresis (PAGE) 1. Immobilized pH gradient strips and accompanying ampholyte mixures can be purchased from a number of commercial vendors. Strip lengths vary from 7 cm to high-resolution 24 cm strips, and pH ranges vary from wide-range (e.g. pH 3–11) to high-resolution narrow-range (e.g. pH 5–6) strips. 2. Bind silane working solution (50 mL): 40 mL ethanol, 1 mL acetic acid, 50 µL bind silane solution (GE Healthcare), 9 mL water (see Note 9). 3. 4x separating gel buffer: 1.5 M Tris-HCl pH 8.8. 4. 30% acrylamide:bis-acrylamide (37.5:1), N,N,N,N -Tetramethyl-ethylenediamine (TEMED), and ammonium persulfate. 5. 10x SDS-PAGE running buffer (1 L): 30.25 g Tris-base, 144.13 g glycine, 10 g SDS (0.1%). 6. Fixing solution for SyproRuby staining (1 L): 100 mL methanol, 70 mL acetic acid, 830 mL water. SyproRuby stain is available form several commercial sources and can be substituted by other fluorescent total-protein stains such as Deep Purple (GE Healthcare), LavaPurple (Fluorotechnics), or Flamingo Pink (BioRad). 7. 2nd dimensional equilibration buffer: 6 M urea, 50 mM Tris-HCl pH8.8, 30% glycerol, 2% SDS, trace bromophenol blue.

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8. Water-saturated butanol (see Note 10). 9. Dithiolthreitol (store dessicated). 10. Iodoacetamide (store dessicated, keep in the dark).

2.3. DIGE Labeling 1. N,N-dimethyl formamide (DMF) (see Note 11). 2. Labeling (L) buffer: 7 M urea, 2 M thiourea, 4% CHAPS, 30 mM Tris-base (do not pH, but ensure that pH of final solution is between 8.0–9.0), 5 mM magnesium acetate (see Note 12). Alternatively, 4% CHAPS can be replaced with 2% ASB14, especially in cases where membrane rich samples are being utilized. 3. Rehydration (R) buffer: 7 M urea, 2 M thiourea, 4% CHAPS, 2 mg/mL DTT (13 mM; 2 %). 4. Cyanine dyes with NHS-ester chemistry for minimal labeling (Cy2, Cy3, Cy5), and with maleimide chemistry for saturation labeling (Cy3 and Cy5) are available from GE Healthcare as dry solids. 5. Quenching solution (for minimal labeling): 10 mM lysine. 6. Dithiothreitol reduction stock solution: 200 mg/mL DTT.

3. Methods DIGE is a powerful technique for quantitative multivariable differentialexpression proteomics. However, the quality of the data will only be as good as the quality of the underlying 2-D gel electrophoresis technology upon which it is based. The main focus of this chapter is to provide detailed notes on the DIGE technology, however, some key considerations to successful high-resolution 2-D gel electrophoresis are also provided. This section describes methods associated with labeling using minimal CyDyes. 3.1. Sample Preparation The key to success for any analytical measurement begins with robust sample preparation. This not only includes the buffers and materials used, but also the nature of the samples and the way in which they are procured. The addition of exogenous materials (such as DNAse, RNAse), or allowing for uncontrolled manipulation of the sample (such as conditions that may lead to proteolysis) can severely hamper and sometimes completely prevent an analysis. Care should be taken to ensure against common laboratory contaminants (e.g., mycoplasma for tissue culture) that if present may be detected as significant changes using DIGE, either due to the presence in a subset of samples, or by responding to the experimental perturbation. 1. Prepare protein extracts using any method of preference. The appropriate amount of protein can be subsequently precipitated prior to resuspension in the CyDye labeling buffer (see Subheading 3.2). Ensure against proteolysis and loss of posttranslational modifications (e.g., phosphorylation) as this is of monumental importance.

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2.

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4.

5.

Friedman and Lilley Care should be taken not to use reagents that will resolve on the 2-D gel, such as soybean trypsin inhibitor. Small molecule inhibitors such as aprotinin, leupeptin, pepstatinA, antipain, AEBSF, sodium orthovanadate, okadaic acid and microcystin, among others, are far better choices. Lyse cells using standard lysis buffers such as TNE, RIPA buffers, or even the buffers used for 2-D gel electrophoresis. All of these buffers have the capability of producing high-resolution samples for 2-DE. In most cases, the presence of reagents that would otherwise interfere with CyDye labeling (such as those that contain primary amines) will be removed prior to labeling by protein precipitation (see Subheading 3.2). Sonicate cells if necessary to improve sample quality. Sonication improves sample quality by disrupting nucleic acids which are subsequently removed by sample cleanup (see Subheading 3.2) along with phospholipids. Both of these non-proteinaceous ionic components can obliterate the resolution during isoelectric focusing. Short bursts with a tip-sonicator are suggested. It is important to keep the system chilled, especially in the presence of urea-containing samples that should never be heated (see Note 12). Determine the protein concentration of the sample using a system that is compatible for the buffer that the proteins are extracted in. CHAPS and thiourea in the buffers used for DIGE, although adequately chaotropic, interfere with either the Bradford or BCA assays, making the data inaccurate and unreliable. In these cases, aliquots should be precipitated prior to quantification in a suitable buffer, or the use of a detergent compatible assay should be utilized. Aim to use a protein concentration between 1–10 mg/mL. Too dilute and it will be difficult to quantitatively recover proteins following precipitation clean-up (see Subheading 3.2); too concentrated and it will be difficult to accurately dispense the appropriate volume for the experiment. Freeze/thawing should also be kept to a minimum; freezing samples in 1 mL aliquots or less will usually suffice.

3.2. Sample Clean Up. The desired amount of sample to be used in the experiment should be precipitated prior to labeling. This both removes non-proteinaceous ions from the sample (e.g., nucleic acids, phospholipids) that can interfere with IEF, and also transfers the proteins into a labeling buffer optimized for CyDye labeling and subsequent IEF. Determine how much total protein will be on each gel, and precipitate ½ of that amount for each sample to be run on that gel. This is straightforward for a two-component separation, but also works out for the multigel experiments where 1/3 of the total protein amount on each gel comes from the pooled-sample internal standard (see Table 1). Precipitate only what is needed for each sample for the experiment; too much material may create pellets that are difficult to resolubilize completely. Many precipitation methods are available, the following is a MeOH/CHCl3 protocol that works well for DIGE, and can be easily performed in 1.5 mL tubes (adapted from (40)):

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Table 1 Experimental Design for CyDye Labeling Using a Pooled-Sample Internal Standard Samples Gel 1

Gel 2

Gel 3

Control-1 Treated-1 Control-2 Treated-2 Control-3 Treated-3 Precipitated amount L-buffer Aliquot Cy2 Cy3 Cy5 Lysine (quench)

150 µg

150 µg

150 µg

150 µg

150 µg

150 µg

24 µL 16 µL

24 µL 16 µL

24 µL 16 µL

24 µL 16 µL

24 µL 16 µL

24 µL 16 µL

2 µL

2 µL

2 µL

2 µL

2 µL

2 µL 2 µL 30 min on ice in the dark 2 µL 2 µL 2 µL

2 µL 2 µL

10 min on ice in the dark 20 µL 20 µL 20 µL 20 µL 20 µL 20 µL For each gel, combine the quenched Cy3-and Cy5-labeled samples and add 1/3 of the quenched Cy2-labeled pooled mixture 20 + 20 + 20 + 20 + 20 + 20 + 20 µL 20 µL 20 µL 2x R-buffer 60 µL 60 µL 60 µL total 120 µL 120 µL 120 µL R-buffer to Vf to Vf to Vf Total vol

pool

8 µL (x6) 6 µL

6. µL 60 µL

This table illustrates a typical DIGE labeling experiment, as described in Subheadings 3.2 and 3.3.

1. 2. 3. 4. 5. 6.

7. 8. 9. 10.

Bring up predetermined amount of protein extract to 100 µL with water. Add 300 µL (3-volumes) water. Add 400 µL (4-volumes) methanol Add 100 µL (1 volume) chloroform Vortex vigorously and centrifuge; the protein precipitate should appear at the interface. Remove the water/MeOH mix on top of the interface, being careful not to disturb the interface. Often the precipitated proteins do not make a visibly white interface, and care should be taken not to disturb the interface. Add another 400 µL methanol to wash the precipitate. Vortex vigorously and centrifuge; the protein precipitate should now pellet to the bottom of the tube. Remove the supernatant and briefly dry the pellets in a vacuum centrifuge. Resuspend the pellets in a suitable amount of CyDye labeling buffer (L-buffer, see Table 1).

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An alternative widely used precipitation method is as follows: 1. 2. 3. 4. 5. 6. 7. 8. 9.

Add 5 volumes of cold 0.1 M Ammonium acetate in methanol. Leave at −20°C for 12 hr or overnight Centrifuge at ~3000 rpm (1400 × g) for 10 min at 4°C and remove the supernatant. A pellet of protein should be visible at this stage To wash the pellet, add 80% 0.1 M ammonium acetate in methanol and mix to resuspend the protein. Centrifuge at 3000 rpm (1400 × g) for 10 min at 4°C and remove the supernatant. To dehydrate the pellet add 80% acetone and resuspend the pellet by mixing. Centrifuge at 3000 rpm (1400 × g) for 10 min at 4°C and remove the supernatant. Dry pellet for 15 min by leaving open tube in a laminar flow cabinet.

3.3. DIGE Experimental Design. 1. Start with a preliminary gel. All experiments should start with a preliminary gel on representative samples to ensure equivocal protein amounts between samples, and that the highest resolution and sensitivity are obtained before embarking on a multigel DIGE experiment (see Notes 13 and 6). The preliminary gel will also show any problems with the sample preparation that may be corrected by adjusting the procurement methods (see Subheading 3.1). This step can also be used to optimize the maximal amount of protein can be loaded without adversely affecting resolution. The preliminary gel needs only to test one or two of the samples of a much larger experiment. This gel can simply be stained with a total protein stain (see Subheading 2.2.6) to visually inspect the resolution and sensitivity. Alternatively, the gel can contain two different samples prelabeled with Cy3 and Cy5 and coresolved (see Note 14). 2. Choose a suitable pH gradient for IEF. Precast isoelectric focusing strips are commercially available from several vendors. The widest length is currently 24 cm, providing the highest resolving power for a given pH range. Medium-range isoelectric focusing gradients (e.g., pH 4–7) offer the best trade-off between overall resolution and sensitivity. Subsequent experiments can then be designed to resolve proteins in the basic range (pH 7–11) and in narrow pI ranges with commensurate increases in protein loading to gain access to the lower abundant proteins in a given sample (see Note 5). In this way a more comprehensive picture of the proteomes under study can be obtained. 3. Incorporate a pooled-sample mixture internal standard on every DIGE gel in a coordinated experiment. This internal standard, usually labeled with Cy2, is composed of an equal aliquot of every sample in the entire experiment, and therefore represents every protein present across all samples in an experiment. The use of this pooledsample internal standard on every DIGE gel in a coordinated experiment allows for the facile comparison of independent sample replicates with increased statistical confidence. This experimental design also enables the simultaneous quantitative comparison between multiple variables in a coordinated experiment (Fig. 1). 4. Plan out which samples will be labeled with which dyes ahead of time. For minimal dye-labeling chemistry (see Subheading 3.4), each gel will contain two individual

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samples labeled with either Cy3 or Cy5, and an equal amount of the pooled-sample internal standard. The example outlined in Table 1 is for a two-component comparison repeated in triplicate, with 300 µg total protein loaded onto each of three gels. In this case, 150 µg of each sample should be precipitated (see Subheading 3.2), resuspended in L-buffer and then split 2:1. Two-thirds of each sample (100 µg) will be individually labeled with either Cy3 or Cy5. The remaining 1/3 of each sample will be pooled together and labeled en masse with Cy2 to serve as an internal standard. By following this, there will be enough of the Cy2-labeled internal standard to have an equal amount as the Cy3 or Cy5 samples loaded onto each gel (see Note 15).

3.4. CyDye Labeling All steps are performed on ice. The following protocol is for sample loading via rehydration of IPG strips, and assumes incorporation of a pooled-sample internal standard to coordinate many samples across multiple DIGE gels simultaneously. The steps are summarized in Table 1 (see Note 16). 1. Resuspend precipitated sample in 24 µL labeling (L) buffer. Remove 8 µL (1/3 of sample) and place into a new tube that will contain the pooled-sample internal standard (8 µL from all of the other individual samples will be pooled into this tube) (see Note 17). 2. CyDyes are purchased as dry solids and should be reconstituted to 10x stock solutions (1 nmol/µL) in fresh DMF. Dilute stock solutions of CyDyes 1:10 in fresh DMF to a final working concentration of 100 pmol/µL. (see Note 11). 3. Label each sample (50–250 µg) with 2–4 µL (200–400 pmol) of either Cy3 or Cy5 working dilution for 30 min on ice in the dark. Label the pooled-sample mixture with 2–4 µL (200–400 pmol) of Cy2 working dilution for every equivalent amount of sample present in the pooled standard as compared with the individually labeled samples. That is, if 100 µg of each sample is labeled with 200 pmol of Cy3 or Cy5, then 50 µg of each of these samples is present in the pooled standard, and 200 pmol of Cy2 is used for every 100 µg of pooled standard (see Table 1 and Note 18). 4. Quench reactions with 2 µL of 10 mM lysine for 10 min on ice in the dark. 5. For each gel, combine the quenched Cy3-and Cy5-labeled samples and add 1/3 of the quenched Cy2-labeled pooled mixture. 6. To each tripartite mixture, add an equal volume of 2x R-buffer and incubate on ice 10 min. 2x R-buffer is R-buffer supplemented with an additional 2 mg/mL DTT using the 200 mg/mL DTT stock solution. DTT is omitted from the L-buffer to prevent unfavorable interaction with the CyDyes. Adding an equal volume of 2x R-buffer to the quenched reactions provides the reducing agents to the total reaction volume at 1x final concentration. 7. Add R-buffer (1x DTT concentration) to a final volume suggested by the manufacturer for the given IPG strip length (e.g., 450 µL for 24 cm strips). Add the appropriate volume of IPG buffer ampholines to 0.5% final (v/v) for isoelectric focusing. Proceed with rehydration of dehydrated immobilized pH gradient (IPG)

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Friedman and Lilley strips for >16 hr and proceed with isoelectric focusing (see Subheading 3.5.3 and Note 19).

3.5. 2-D Gel Electrophoresis and Poststaining As a result of the minimal labeling, quantification with the CyDyes is carried out on only 2–5% of the proteins that are labeled, and the labeled portion of the protein may migrate at a higher apparent molecular mass than the majority of the unlabeled protein due to the added mass and hydrophobicity of the dyes (exacerbated in lower Mr species). To ensure that the maximum amount of protein is excised for subsequent in-gel digestion and mass spectrometry, minimally labeled 2-D DIGE gels are poststained with a total protein stain (see Subheading 2.2.6). Accurate excision is also ensured by preferentially affixing the 2nd dimension gel to a presilanized glass plate during gel casting so that the gel dimensions do not change during the analysis. (see Notes 20 and 21) These methods assume the use of the Ettan 2-D electrophoresis system (GE Healthcare), but are easily adaptable to other commercially available systems. It also assumes usage of high resolution 24 cm × 20 cm gels. 1. Special gels for 2nd dimension SDS-PAGE. Using low-fluorescence glass plates, pretreat one plate for each gel with 3–5 mL bind silane working solution, carefully wiping the entire surface of the plate with a lint-free wipe. Leave treated plates covered with lint-free wipes for several hours to allow for sufficient out-gassing of fumes (that may contain bind silane) before assembling gel plates and casting of 2nd dimensional SDS-PAGE gels (see Note 22). 2. Assemble plates and pour 12% homogeneous SDS-PAGE gel(s) using the appropriate amount of 30% stock acrylamide and 4x separating gel buffer for the volumes needed for the number of gels being poured (see Note 23). Overlay the gels with water-saturated butanol for several hours to provide a straight and level surface to place the focused IPG strip (see Note 10). 3. Perform isoelectric focusing (for example using an IPGphor III isoelectric focusing unit, GE Healthcare) of the combined tripartite-labeled samples, brought up to final volume with 1x R-buffer and passively rehydrated into immobilized pH gradient (IPG) strips for >16 hr (see Subheading 3.4, step 7) (see Note 24). 4. Equilibrate the focused IPG strips into the 2nd dimensional equilibration buffer. During this step, the cysteine sulfhydryls in the focused proteins are reduced and carbamidomethylated by supplementing the equilibration buffer with 1% DTT for 20 min at room temperature, followed by 2.5% iodoacetamide in fresh equilibration buffer for an additional 20 min room temperature incubation (see Note 25). 5. Place equilibrated IPG strip on top of the SDS-PAGE gels that were precast with low-fluorescence glass plates. Use a thin card or ruler to carefully tamp down the IPG strip to the SDS-PAGE gel, removing air bubbles at the interface (see Notes 26 and 27).

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6. Perform 2nd dimensional SDS-PAGE at constant wattage, using 3) are handled by the BVA (Biological Variation Analysis) module of DeCyder. In a BVA experiment, the signals emanating from the internal standard are used both for direct quantification within each DIGE gel in a coordinated set (using DIA module), as well as for normalization and protein spot-pattern matching

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between gels (see Note 31). This allows for the calculation of Student’s t-test and ANOVA statistics for individual abundance changes (see Subheading 3.6.2, and Table 2). BVA is also used to match patterns between SyproRubyand CyDye-stained images to facilitate protein excision for subsequent mass spectrometry (see Notes 20, 21, and 30). 3.6.2. Experimental Design and Statistical Confidence. In the simplest form of a DIGE experiment, two or three samples are separately labeled with one of the three dyes and separated in the same gel for direct pair-wise comparisons. In this case, the software first normalizes the entire signal for each Cy-dye channel and then calculates the protein spot volume ratio for each protein pair. A normal distribution is modeled over the actual distribution of protein pair volume ratios, and two standard deviations of the mean of this normal distribution represent the 95th percent confidence level for significant abundance changes. This N = 1 type of experiment has limited statistical power, since the 95th percentile confidence interval is determined based on the overall distribution of changes within the population (see Note 32). Many more changes in abundance of much lesser magnitude can be detected with much greater statistical confidence (Student’s t-test and ANOVA, Table 2) by incorporating independent replicate samples into the experiment (see Note 33). The number of replicates required in a study depends on the amount of variation in the system being investigated. Increasing the number of replicates will increase confidence in smaller changes in expression. The number of gel replicates that are needed for the experiment to have sufficient sensitivity to detect expression changes can be determined using power calculations (41). With replicate samples, the Student’s t-test and ANOVA statistics are measuring the significance of the variation of a specific protein change, independent of the overall distribution of abundance changes in the population. Incorporating replicate samples into the experimental design also controls for unexpected variation introduced into the samples during sample preparation. This design not only allows for the identification of abundance changes that are consistent across multiple replicates of an experiment, but can also identify significant abundance changes that would not have been identified even if the analyses were performed using Cy3- and Cy5-labeled samples on the same gels, but without the pooled-sample internal standard to coordinate them (8). 3.6.3. Multivariate Statistical Analysis Univariate analyses such as the Student’s t-test and ANOVA (analysis of variants) have traditionally been used in DIGE experiments to provide a list of statistically significant changes in protein abundance. The application of multivariate

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Table 2 Statistical Applications of DeCyder Biological Variation Analysis (BVA) and Extended Data Analysis (EDA) modules Average ratio

Student’s T-test

One-Way ANOVA Two-Way ANOVA Principal Component Analysis (EDA only) Hierarchical Clustering (EDA only) K-means (EDA only) Self Organizing Maps (EDA only) Gene Shaving (EDA only) Discrimnant Analysis (EDA only)

Calculated for each protein spot-feature between two groups or experimental conditions. Derived from the log standardized protein abundance changes that were directly quantified within each DIGE gel relative to the internal standard for the protein spot-feature. Univariate test of statistical significance for an abundance change between two groups or experimental conditions. P-values reflect the probability that the observed change has occurred due to stochastic chance alone. With DIGE, p-values of 50 V-hr have been reached. Check recommendations from specific vendors. Using the ceramic manifold tray adaptor is recommended. 25. Volume of equilibration buffer should be large to ensure sufficient removal of ampholines and other components of the 1st dimensional run. 26. Carefully wash out any remaining liquid on top of the SDS-PAGE gel. Prewet the IPG strip with1x running buffer and place the strip between the gel plates with the plastic-backing adhering to the inside surface of one of the glass plates. The prewetted running buffer will facilitate the manipulation of the IPG strip down the inside surface of the plate and on top of the SDS-PAGE gel. 27. An agarose overlay, used by many protocols, is not absolutely necessary to ensure proper contact between the IPG strip and the 2nd dimensional SDSPAGE gel. Using a thin card or ruler to carefully tamp down the IPG strip to the gel is usually sufficient and removes the added problems associated with the overlay, such as trapped air bubbles in the solidified agarose.

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28. Running gels at less than 1 W/gel can improve resolution in the high molecular weight regions of the 2nd dimension gel. Use wattage appropriate for the 2nd dimensional unit being used. Many different gel units can accommodate increased power by compensating for the increased heat. 29. Absorption/emission maxima in DMF are 491/506 for Cy2, 553/572 for Cy3, and 648/669 for Cy5; although care must be taken to scan in regions of each spectrum that do not contain absorbance or emission in the other spectra, which may mean using a non-maximal region of a given spectrum. 30. Comparison of the 2-D spot maps between saturation-labeled samples and minimal labeled or unlabeled samples is impossible, as proteins containing multiple cysteine residues may appear as significantly larger Mr species when labeled with the saturation dyes, which of course cannot be predicted without first knowing the protein identity. 31. Almost all software packages for 2-D electrophoresis involve matching of protein spot-patterns between gels. For DeCyder, it is used in the BVA module to match the quantitative data obtained from the triply coresolved protein signals from each gel in the DIA module (where gel-to-gel variation does not come into play). Manual verification of the matching is almost always required with any software package. 32. There are many “all-or-none” type of experiments where the single gel comparison may be valid, and subtle changes are not expected. Nevertheless, using independent replicates and the pooled-sample internal standard methodology is still needed to control for non-biological sample preparation error. 33. The multigel approach allows many data points to be collected for each group to be compared. Protein features of interest can be selected by looking for significant change across the groups. Student’s t-test and Analysis Of Variance (ANOVA) probability scores (p) indicate the probability that the observed change occurred due to stochastic, random events (null hypothesis). Probability values < 0.05 are traditionally used to determine a statistically significant difference from the null hypothesis. As this represents 50 potential false-positives for 1,000 resolved proteins, confidence intervals within the 99th percentile (p < 0.01) are arguably more valid, and can be attained using DIGE (5, 8, 18, 50–54). References 1. Petricoin, E., Wulfkuhle, J., Espina, V., and Liotta, L. A. (2004) Clinical proteomics: revolutionizing disease detection and patient tailoring therapy. J. Proteome Res. 3, 209–217. 2. Gorg, A., Obermaier, C., Boguth, G., Harder, A., Scheibe, B., Wildgruber, R., and Weiss, W. (2000) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 21, 1037–1053.

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40 Comparing 2D Electrophoretic Gels Across Internet Databases: An Open Source Application Peter F. Lemkin, Gregory C. Thornwall, and Jai A. Evans 1. Introduction In (1–4) and in the second edition of this book (5), we described a Webbased computer-assisted visual method called Flicker for comparing two two-dimensional (2D) protein gel images across the Internet using a Java applet. In the second edition of this book (6), Flicker was described as a stand-alone application. The flicker method was originally developed in the GELLAB 2D-gel analysis system (7–11). The applet version was used for Web-based analysis using a Web-browser. The Java stand-alone application can run on a user’s computer, where it can access images from the Web but can easily access the user’s images data on their local computer. Flicker is available as open source on http://open2Dprot.sourceforge.net/Flicker where the executable program and the source code is also available. Some of the code was derived from the old Flicker applet program and some from the MicroArray Explorer (12) –an open-source data-mining tool for microarray analysis http://maexplorer.sourceforge.net/. The Open2Dprot protein expression analysis system http://open2Dprot.sourceforge.net/ was partly derived from Flicker, MicroArray Explorer, and GELLAB code and concepts. Because Flicker can analyze a user’s data on his or her computer independent of the Internet, this gives the user the ability to perform real-time comparisons of 2D-gel image data with gel images residing on the user’s local file system, on remote Internet databases on the Internet, or a combination of both sources. This approach may be useful for comparing similar protein samples created in different laboratories to help putatively identify or suggest possible protein spot identification. The gels should be run under similar pH and molecular weight ranges if possible. Although available for over three decades, 2D polyacrylamide gel electrophoresis (2D-PAGE) is still used (13–14) even considering From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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the now common use of mass spectrometry (13,25–26) and protein microarrays (13,21–22) for protein identification and biomarker discovery as part of the researcher’s toolkit (13). Advances, such as IEF “zoom” fractionation gels (23), are commonly used (24) to divide the protein sample by pH range or immunoaffinity subtraction with LC (18), greatly increase the resolution and numbers of spots able to be discriminated by subsequent 2D-gel electrophoresis. Another increasingly common image comparison technique uses 2 to 6 cyanine dyes using dye multiplexing to label multiple control and experimental samples run in the same gel such as GE Healthcare’s (formerly Amersham’s) DIGE (13,25–26) and scanned with system’s like Perkin Elmer’s ProExpress (27). Multiple scans of the same gel using different color filters can then be color mapped to see the contributions of the different samples. This is useful if one has control over the experimental design when determining the reference gel, set of control gels, and experimental gels. Advances in DIGE (DeCyder™) EDA software now allow comparison of multiple images run on the same apparatus (26). However, these advances do not completely solve the problem of trying to putatively compare one’s own sample against an Internet reference gel where they have identified protein spots. A number of 2D-gel databases that contain gel images are available on the Web for various types of tissues. Proteins are identified for some of the spots in a subset of these databases. Both WORLD-2DPAGE and 2D Hunt on the http:// www.expasy.org/ server can be used to find Web URL addresses for a number of 2D protein gel databases. Google searches are also used and we link to these sites in the Help menu. Many of these databases contain 2D-gel images with identified proteins. Some of these databases let you identify spots in their gels by clicking on a spot in their gel image shown in your Web browser. It then queries their Web server database to determine if the spot you pointed to is in that database and report its identity if found. These “clickable” 2D-gel map images are often published using a common federated database (DB) schema (28–29). One of the more interesting databases is SWISS-2DPAGE (28–31), accessible from the Expasy site. It has a large number of tissues with over 30 gel databases including a wide range of human tissues, mouse, E. coli, aribidopsis, dictyostelium, and yeast. Their site also has a series of IEF zoom fractionated gels for E. coli. We have incorporated links to these SWISS-2DPAGE database gel map images so they may be loaded and accessed directly from Flicker after having putatively matched a spot in your gel with one in the SWISS-2DPAGE gels. If you have loaded one of these active map gel images in Flicker and enabled the database access, then clicking on a spot in that image will pop up a Web page as it tries to look it up the spot in the SWISS-2DPAGE database. If a SWISS2DPAGE data entry exists for the spot coordinates you have selected, then it will report the corresponding protein or tell you it cannot be found.

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By comparing one’s own experimental 2D-gel image data with gel images of similar biological material from these from Internet reference databases, it opens up the possibility of using the spots in these reference gels to suggest the putative identification of apparently corresponding spots in your gels. The image analysis method described here allows scientists to more easily collaborate and compare their gel image data over the Web. How Can We Compare Two Gels? When two 2D gels from different laboratories are to be compared, simple techniques may not suffice. There are several methods for comparing two gel images: (1) put the images side by side and visually compare them; or (2) slide one gel (autoradiograph or stained gel) over the other while backlighted; or (3) build a 2D-gel quantitative computer database from both gels after scanning and quantitatively analyzing these gels using a 2D-gel database system; and (4) run DIGE dye multiplexing (25) to label different samples in the same gel or across gels (26). A variant of this last method is to spatially warp two gels to the same geometry and then pseudocolor them differentially. These methods may be impractical for many investigators since in the first case the physical gel or autoradiograph from another laboratory may not be locally available. The first method may work for very similar gels with only a few differences. The second method will work better for gels that are not so similar but that have local regions that are similar. The third method may be excessive if only a single visual comparison is needed because of the costs (labor and equipment) of building a multigel database solely to answer the question of whether one spot is probably the same spot in the two gels. The fourth method may also not be practical if you want to compare your sample against an existing reference gel. The Flicker Program We describe a computer-based image comparison technique called flicker that has been used for years in finding differences in star maps in astronomy. The Flicker program runs on most computers. It is started as one would any program after it is downloaded from the Flicker server and installed (see Method 1). One gel image may be read from any Internet 2D-gel database (e.g., SWISS-2DPAGE, etc.), the other may reside on the investigator’s Web server where they were scanned or copied, or the two gel images may be from either Web server source. Figure 1 shows the Flicker application after it has been started with some demo gels. You interact with the program by clicking or dragging the mouse in the left or right images, adjust parameter scrollers (upper right), set interaction modes (checkboxes upper left), keyboard short-cut commands, and primarily pull-down menu commands. Notation in This Paper We use the notation (<menu name> | ) throughout this paper to indicate the these menu commands. The <menu name> indicates one of the

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Fig. 1. Screen view of initial Flicker program. (A) shows the pull-down menus at the top used to invoke file operations, editing, view selection, landmarking, image transforms, spot quantification, and help. A set of scroll bars on the right determines various parameters used in some of the transforms. The File menu options include opening a new gel image. Checkboxes on the left activate flickering and active gel map access if the data supports it. A set of status lines appear below the checkboxes and indicate the state of operation and various other messages. The flicker image is in the upper middle of the frame when it is enabled. The two labeled human blood plasma gel images are shown in the bottom scrollable windows that may be positioned to the region of interest. These windows also have associated flicker time-delays used when flickering. Image plasmaH is an IPG non-linear gradient gel from SWISS-2DPAGE in Geneva and plasmaL is a carrier-ampholyte linear gradient gel from the Merril Lab at NIMH. Transformed image results are shown in the same scrollable windows. The four checkboxes are: Flicker (C-F) to enable/disable flickering; Click to access DB checkbox enables/disables access to a Web server that is associated with a clickable image DB if it exists for the selected image; Allow transforms checkbox enables/disables image transforms; Sequential transforms checkbox enables/disables using the last image transform output as input for the next image transform. The parameters used in various transforms

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Fig. 1. (continued) are adjusted directly by first selecting an image and then adjusting its values. You can popup the scrollable report window using the Report scroller values button. These parameters are saved when you save the state. You can change the size of the three image windows by using the “+” and “−” buttons. (B) The popup scrollable report window shows a log of all text output that appears in the status lines. It may be saved to a text file on the local disk.

pull-down menus: File, Edit, View, Landmark, Transform, Quantify, Help. The indicates one of the commands in that menu. Table 1 summarizes the menu commands. Some of the commands have alternative keyboard shortcuts

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Table 1 Summary of Commands Available in Flicker Nenu Submenus are indicated by underlining the name of the menu and adding the ‘->’ symbol. Checkboxes are indicated by the prefix to the command. Shortcuts are indicated by a (C-) at the end of the command. File menu – to load images, load demo images, active map urls, to load/save the Flicker .flk state, update (from the server): program, DB/Flk*DB.txt database files Open image File – pop up gel image file browser Open image URL – pop up gel image URL dialog Open demo images -> – load pairs of demonstration gel images from 3 data sets Open user images -> – load pairs of demonstration gel images Pairs of images -> – load pairs of user gel images Single images -> – load single user gel image to selected image List user’s images by directory Open active map image -> – load active gel image from Internet(Swiss-2DPAGE) Open recent images -> – load an image you have used recently Assign active image URL – to one of the open images to make it active —— Open state file – restore the Flicker state of previously saved session Save state file – save the Flicker state in current .flk state file SaveAs state file – save the Flicker state in new .flk state file —— Update -> – download and update your program and data from server Flicker program – to get the most recent release Active Map Image database – get latest active maps database Demo Images database – get latest demo images database Add user’s active images DB by URL —— Save Transformed image – of selected image as .gif file if transformed SaveAs Overlay image – the current overlay image Reset images – to the initial state when they were loaded Abort transform – abort any active image transforms —— Quit – exit the program, saving the .flk state of Flicker in the process Edit menu - to change various defaults Canvas size -> – change the size of 3 image canvases and overall Flicker window Increase size (C-Numpad ‘+’) - increase the canvas size Decrease size (C-Numpad ‘−’) - decrease the canvas size Set colors -> – set default colors for the overlays Target colors – to change the target color Trial object colors – to change the trial object color Landmarks colors – to change the color of landmarks Measurement color – to change the color of measurements Guard Region color – to change the color of the guard region (continued)

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—— Use linear else log of TIFF files > 8-bits – take log of tiff data if > 8-bits Enable saving transformed images when do a ‘Save(As) state’ Use protein DB browser, else lookup ID and name on active images —— Auto measure, protein lookup in active server and Web page popup – set access Select access to active DB – select protein report for active protein queries Use SWISS-2DPAGE DB access Use PIR UniProt DB access Use PIR iProClass DB access Use PIR iProLink DB access —— Reset default view – sets all view options to the defaults Clear all ‘Recent’ images entries – clears the list of recently accessed images View menu - to change the display overlay and popup report window options m Flicker images (C-F) – toggle flickering on and off Set overlay options – select one or more image overlay options View landmarks – add landmarks to the overlay display in images View target – add target to the overlay display images View trial object – add trial object to the overlay display images View Region Of Interest (ROI) - add ROI to the overlay display images Set measurement options – select one or more image measurement options View measurement circle – add measurement circles to overlay display images Use ‘Circle’ for measurement spot locations Use ‘+’ for measurement spot locations Use ‘spot number’for spot annotations – e.g., #1, #2, …, etc. Use ‘spot ID’for spot annotations – e.g., actin, APO-A1, etc. Set gang options – select one or more ganged images (left and right) options Multiple popups – make multiple popup windows instead of reusing one Gang scroll images – move left and right images scrolling together Gang zoom images – zoom left and right images scrolling together Add guard region to edges of images – this allows comparing edges of images Display gray values (C-G) – show gray values of cursor trial object Show report popup – display the report popup window again if needed Landmark menu - to define landmarks for spatial image warping Add landmark (C-A) – add trial objects (in images) as landmark Delete landmark (C-D) – delete the last landmark defined Show landmarks similarity – compute a LSQ error similarity measure of the two sets of landmarks Set 3 predefined landmarks for demo images (C-Y) Set 6 predefined landmarks for demo images (C-Z) Transform menu - contains various image processing transforms for selected image(s) Affine Warp – warp selected image using 3 pairs of landmarks (continued)

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Pseudo 3D transform – do pseudo 3D scaling based on image intensity —— Sharpen Gradient – gradient + gray scale sharpen selected image Sharpen Laplacian – Laplacian + gray scale sharpen selected image Gradient – gradient of the selected image Laplacian – Laplacian of the selected image Average – average selected image Median – median of selected image Max 3×3 - max of 3×3 neighborhood of selected image Min 3×3 - min of 3×3 neighborhood of selected image —— Complement – complement selected image Threshold – threshold the selected image by gray values in (T1:T2) Contrast Enhance – Contrast enhance selected image Histogram equalize – histogram equalize selected image —— Original color - Restore original data for selected image Pseudo color - compute pseudo color scaling for selected image Color to grayscale - compute NTSC RGB to grayscale transform for selected image (gray = red*0.33 + green*0.50 + blue*0.17) —— Flip Image Horizontally – flip image horizontally selected image Flip Image Vertically – flip image vertically selected image —— Repeat last transform (C-T) - repeat last transform, if any Use threshold inside (T1:T2) filter - filter by pixels inside the range (T1:T2), otherwise pixels outside of (T1:T2) ) Quantify menu - contains OD calibration, background and foreground measurements Measure circle -> – measure intensity/density within circle Capture background (C-B) – background measurement at current position Capture measurement (C-M) – report object measurement at current position Circle size -> – define the circlular mask size by radius in m 1×1 to m 11×11 pixels subregions —— Clear measurement – clear measurement data Edit selected spot(s) ‘id’ fields from spot list(s) (C-I) Edit selected spot(s) from spot list(s) (C-E) Delete selected spot from spot list (C-K) —— List spots in the spot list for selected image – list measured spots List spots in the spot list (tab-delimited) – same but can cut& paste data List ‘id’-paired annotated mean norm. spots in both spot lists (tab-delim.) List ‘id’-paired annotated spots in both spot lists (tab-delimited) (continued)

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—— Lookup Protein Ids & Names in spot list from active map server (sel’td image) —— Clear spot list (ask first) for selected image Print data-window -> - print numeric pixel data around image pixel Show data window of selected pixel (C-V) Set print window size – either 5×5, 10×10, … or 40×40 Set print data radix – either decimal, octal, hex, or optical density Calibrate -> - Calibrate grayscale as optical density or other units Optical density by step wedge – set calibration by ND step wedge in gel image Use demo Leukemia ND step wedge preloaded data – for use in demo —— Optical density by spot list – set OD values by mean spot list measurements Region of Interest (ROI) -> - region of interest operations Set ROI ULHC (C-U) - define upper left hand corner of ROI Set ROI LRHC (C-L) - define lower right hand corner of ROI Clear (ROI) (C-W) - delete ROI Show ROI grayscale histogram (C-H) – show grayscale histogram of ROI pixels Capture measurement by ROI (C-R) - measure integrated density inside the computation region ROI. Use the circular mask mean background for background correction Use sum density else mean density - use sum of the density else mean density within the region List-of-spots else trial-spot measurement mode (C-J) Help menu - popup Web browser documentation on Flicker from the server Flicker Home – pop up Flicker home page open2Dprot.sourceforge.net/Flicker Reference Manual – pop up the reference manual for Flicker application How-to use controls -> – pop up the reference at particular sections Vignettes -> – pop up short vignettes showing how-to-do tutorials Version on the web site – show current version available on the Web site About – show details on Flicker application —— Book chapter on Flicker (2005) – popup 2005 PDF of book chapter Old flicker applet documentation -> Flicker applet home page – popup the home page for old Flicker applet EP97 Paper – popup the Electrophoresis ‘97 paper Flicker applet Poster 1 – pop up poster describing Flicker applet Poster 2 – pop up poster describing Flicker applet Poster 3 – pop up poster describing Flicker applet Poster 4 – pop up poster describing Flicker applet 2D gel web resources -> SWISS-2DPAGE – pop up SWISS-2DPAGE home page WORLD-2DPAGE – pop up Expasy World-2DPAGE of federated gel databases 2D-HUNT – pop up Expasy’s 2D-Hunt for finding gel Web sites Google 2D search – pop up Google search for finding gel Web sites

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activated by using the Control key with another key and are indicated as Control or C- (e.g., C-A ). The checkbox menu commands are indicated with a prefix. Checkbox commands may be toggled on and off. The gels in the two lower left and right images are specified by the user with the FlickerFiles menu. Gel images may be loaded from: the local computer (File | Open image file), or any Internet site with GIF, JPEG, TIFF or PPX images (with .gif, .jpg, .tiff or .tif, or .ppx GELLAB-II format (9) file extensions) using the (File | Open image URL) command. In addition, the installation provides a few demonstration images (File | Open demo images) that loads pairs of comparable images. You may also specify active gel images from Web servers as described below. When Flicker starts, it creates additional submenu entries in the (File | Open user images | Pairs of images | …) and (File | Open user images | Single images | …) submenus that are the names of the user’s images they copied to the Flicker Images/ subdirectory. The user can load images using those commands. Flicker is also capable of interacting with federated 2D-gel databases to retrieve data on individual protein spots if one of the gels is a federated gel having an associated clickable gel map database. After aligning gels in Flicker, you enable federated database access in Flicker and then click on a spot in the gel belonging to the federated database (see Fig. 2). This causes a Web page to pop up with information from the federated server describing that protein. We provide menu entries (File | Open active map image | …) to let you load one of the SWISS-2DPAGE gel images. You may load a gel image in the lower left or right image windows. First click on the image you want to load the new image. Then select the active gel image to you want using the entry (File | Open active map image | …) pull-down menu (e.g., select (… | SWISS-2DPAGE Human | Plasma) ). Next, click on the other image and then use the other gel image you want to compare it with using either (File | Open image file) or (File | Open image URL) commands. The Flicker program is written in Java, a general purpose, object-oriented programming language developed by Sun Microsystems (32) http://java.sun. com/. Java has become a standard for portable Internet Web applications. Most often, the original images may be compared directly. However, occasionally, the comparison may be made visually easier by first applying enhancement transforms such as spatial warping, brightness, contrast or other image transforms. Adjusting image brightness and contrast so the two gels have similar ranges will make the image fusion easier for the user when flickering. For gels with a lot of geometric distortion, it is useful to adjust the geometry of one gel so that the geometry of the local region being compared approximates that of the other gel. By local geometry, we mean the relative positions, distances, and angles of a set of spots in corresponding regions.

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Fig. 2. Screen view of the landmarks used for the affine transform of the human plasma gel images. (A) The transform warps the geometry of a local region defined by the three landmarks so it more closely resembles the geometry of the corresponding local region in the other gel. Scrollable image windows with 3 “active” landmarks defined in both gel images that were selected interactively in preparation for doing the affine image transform. Corresponding landmark spots are selected so they are defined unambiguously in both gel images. For demonstration purposes, the command (Landmarks | Set 3 predefined landmarks for demo gels) will set up the 3 landmarks shown in this figure. (B) After defining the 3 landmarks, use the (Transform | Affine warp) command.

One technique to correct geometry differences is called spatial warping. When performing spatial warping, corresponding regions of interest are (1) first marked by the user (we call this “landmarking”) with several corresponding points in each gel image (3 for affine warping and 6 for poly warping), and (2) then one of the two gel images is warped to the geometry of the other gel

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Fig. 2. (continued)

(see equations 1 and 2). A landmark is a corresponding spot that is present in both gels. Landmarks are defined by clicking on the spot to mark and selecting the (Landmark | Add Landmark (C-A) ) command. The warping is performed by first selecting the image to warp by clicking on it, and then and selecting the (Transform | Affine Warp) command. Landmarking and warping are described in more detail below. Spatial warping doesn’t change the underlying grayscale values of the synthesized warped image to the extent that would cause local structural objects to appear and disappear and thus spot artifacts might be created. Instead, it samples pixels from the original image to be transformed and places them in the output image according to the geometry of the other input image. After warping is finished, gels may then be compared visually by flickering.

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1.1. Image Flickering The basic concept of using flickering as a dynamic visualization technique is simple. If two images may be perfectly aligned then one could simply align them by overlaying one over the other and shifting one image until they line up. However, many images such as 2D PAGE gels have rubber-sheet distortion (i.e., local translation, rotation, and magnification). This means there is more distortion in some parts of the image than in others. Although it is often impossible to align the two whole images at one time, they may be locally aligned piece-by-piece by matching the morphology of local regions. If it appears that a spot and the surrounding region do match, then one has more confidence that the objects are the same. This putative visual identification is our definition of matching when doing a comparison. Full identification of protein spots requires further work such as cutting spots out of the gels and subjecting them to sequence analysis, amino-acid composition analysis, mass spectrometry, testing them with monoclonal antibodies, or other methods. 2.2. Image Enhancement It is well-known that 2D gels often suffer from local geometric distortions making perfect overlay impossible. Therefore, making the images locally morphologically similar while preserving their grayscale data may make them easier to compare. Even when the image subregions are well aligned, it is still sometimes difficult to compare images that are quite different. Enhancing the images using various image transforms before flickering may also help. Some of these transforms involve spatial warping, which maps a local region of one image onto the geometry of the local region of another image while preserving its grayscale values. Another useful operation is contrast enhancement that helps when comparing light or dark regions by adjusting the dynamic range of image data to the dynamic range of the computer display. Other transforms include image sharpening and contrast enhancement. Image sharpening is performed using edge enhancement techniques such as adding a percentage of the gradient or Laplacian edge detection functions to the original grayscale image. The gradient and Laplacian have higher values at the edges of objects. In all cases, the transformed image replaces the image previously displayed. Other functionality is available in Flicker and is described in the Methods and Notes subheadings of this chapter, Table 1, and on the Web server. 1.3. Image Processing Transforms As mentioned, there are a number of different image transforms that may be invoked from the Transform menu. You may display the transformed image, use it as input to another transform, or save it as a .gif file on your local computer. When you save the state, you may also save the transformed images.

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1.3.1. Affine Spatial Warping Transform The spatial warping transforms require defining several corresponding landmarks in both gels. As we mentioned, one gel image can be morphologically transformed to the geometry of the other using the affine or other spatial warping transformations. These transforms map the selected image to the geometry of the other image. It does not interpolate the gray scale values of pixels – just their position in the transformed image. As described in (1–2, 4–6), this might be useful for comparing gels that have some minor distortion, comparing local regions, gels of different sizes or gels run under slightly different conditions. Flicker uses the affine transform as an inverse mapping as described in (33). Let (uxy,vxy) = f(x,y), where (x,y) are in the output image, and corresponding (u,v) are in the input image. Then, in a raster sweep through the output image, pixels are copied from the input image to the output image. The affine transformation is given in Eqs. 1–2: uxy = ax + by + c

(1)

nxy = dx + ey + f

(2)

When the affine transform is invoked, Flicker solves the system of 6 linear equations for coefficients (a,b,c,d,e,f) using three corresponding landmarks in each gel. 1.3.2. Pseudo-3-D Transform As described in (1–2, 4–6), the Pseudo-3-D transform is a forward mapping that generates a pseudo 3-D relief image to enhance overlapping spots with smaller spots seen as side peaks. The gel size is width by height pixels. The gray value determines the amount of y shift scaled by a percentage zscale (in the range of 0 to 50%). Pseudo perspective is created by rotating the image to the right (left) by angle theta (in the range of −45 to +45 degrees). The transform is given in Eqs. 3–5 for image of size width X height, shift in the horizontal dimension computed as dx. dx = width sin(q)

(3)

x' = (dx (hight - y)/hight) + x

(4)

y' = y-zscale * g(x‚y)

(5)

where g(x,y) is in the original input image and (x´ ,y´ ) is the corresponding position in the output mapped image. Pixels outside of the image are clipped to white.

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The Pseudo-3-D transform is applied to both images so that one can flicker the transformed image. 1.3.3. Edge Sharpening Edge sharpening may be useful for sharpening the edges of fuzzy spots. The sharpened image function g´ (x,y) is computed by adding a percentage of a 2-dimensional edge function of the image to original image data g(x,y) as shown in equation (6). The edge function increases at edges of objects in the original image and is computed on a pixel by pixel basis. Typical “edge” functions include the 8-neighbor gradient and Laplacian functions that are described in (1–2, 4–6) in more detail. The escale value (in the range of 0 to 50%) is used to scale the amount of edge detection value added. g'(x‚y) = (escale * edge(x‚y) + (100 - escale) * g(x‚y))/ 100.

(6)

2. Materials The following lists all items necessary for carrying out the technique. Since it is a computer technique, the materials consist of computer hardware, software and an Internet connection. We assume the user has some familiarity with computers and the World Wide Web. 1. A Windows PC, MacIntosh with MacOS-X, a Linux computer or a Sun Solaris computer having a display of at least 1024 × 768 resolution. At least 30 Mb of memory is required and more is desirable for comparing large images or performing many transforms. If there is not enough memory, it will be unable to load the images, the transforms may crash the program or other problems may occur. An Internet connection is required to download the program from the http:// open2Dprot.sourceforge.net/Flicker Web site (see Note 1). New versions of the program will become available on this Web site and can be uploaded to your computer using the various Update commands described in the Notes section or from the Downloads Web page. If you will be using the active gel image maps associated with federated 2D-gel databases, then you will need the Internet connection for accessing those (e.g., SWISS-2DPAGE) databases. You do not need the Internet for local image comparisons. 2. You should have Java (JDK or JRE version 1.5.0 or later) installed already on your computer. If you do not, you can download it from java.sun.com for free and install it yourself. 3. When you install Flicker, it creates several subdirectories: Images/ (containing the demonstration images), DB/ (containing startup database files), and FlkStartups/ (containing any startup files you create when you do a (File | SaveAs state file). The DB/ files are: FlkDemoDB.txt which describes the demo images, FlkMapDB.txt which describes the gel images and their corresponding active image map URLs, and

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FlkRecentDB.txt which lists recently accessed images. An empty database file FlkRecentDB.txt contains the file names and active gel map URLs, if any, of the recently accessed images. The program is in Flicker.jar is the Java Archive File for Flicker that is executed when you run Flicker. It also uses jai_core.jar, the core Java runtime from SUN’s Java Advanced Imaging (JAI at java.sun.com) and jai_codec.jar that is the JAI tiff file reader from SUN’s Java Advanced Imaging JAI at reader. For Windows users, there is a Flicker.exe file that you can click on to run the program. Otherwise, there are two script files Flicker-startup.bat (for Windows) and Flicker-startup.sh for Unix/ Linux or Mac OS-X that run it through the Java interpreter. 4. The Internet is a good source of 2D-gel images. You can find them by searching WORLD-2DPAGE and 2D Hunt on the http://www.expasy.org/ server or a Google search to find other Web 2D protein gel image databases. Links to these databases are available in the (Help | 2D gel Web resources) submenu. 5. We currently distribute Flicker so that it uses up to 96 megabytes of memory. If you want to run it with more or less memory, you will need to edit the startup files Flicker-startup.bat or Flicker-startup.sh you use to set it to a value in the range of 30 Mb to 1784 Mb. Both of these startup files contain the command

java -Xmx96M -jar Flicker.jar So to increase the startup memory, change the 96 M to some larger value (e.g., to increase it to 500 megabytes, change -Xmx96M to −Xmx500M). 3. Method We now describe the operation of the Flicker from the user’s point of view. You first install Flicker. Then run it with either the demonstration images, your own images or images from the Internet. Then you simply flicker the gel images. If necessary, to improve the image comparability, use image enhancement transforms first before then flickering the two images. 3.1. Installing Flicker from the Web Server Click on the Download link on the http://open2Dprot.sourceforge.net/Flicker Web site. The following method can be used to download the Zipped Flicker-dist installation package and install it as described below. Go to the Web site’s Files mirror under the Flicker releases. Look for the most recent release named “Flicker-V.XX.XX-dist.zip” such as Flicker-V0.87.2dist.zip. These zipped files include the program, required jar libraries, demo data, and Windows batch and Unix shell scripts. Download the zip file and put the contents where you want to install the program. Note that there is a Flicker.exe for Windows (created with launch4j.sourceforge.net). You might make a short-cut to this file to make it easier to find when starting the program. Alternatively, you can use the sample Flicker-startup.bat or Flicker-startup.sh scripts to run the program explicitly via the java interpreter. Note that this method assumes that you have Java installed on your computer and that it is at least JDK (Java Development

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Kit) or JRE (Java Runtime Environment) version 1.5.0. If you don’t have this, you can download the latest version free from the java.sun.com Website. The following is a short procedure that summarizes the procedure for downloading and installing Flicker. 1. In the Table of Contents on the left on the home page, click on “Files mirror” under “Source Code”. 2. Under latest file releases, where it has the header “Package”, click on the “open2Dprot” below that. 3. This will refresh the page and if you scroll down, it will show “Flicker files”. 4. Select the “+” on this to list the files. Pick the one with the highest version number called something like Flicker-V0.87.2-dist.zip file. 5. Click on that to download it. 6. Put it where you want to install it and unpack it. There is a Flicker.exe file (for Windows) as well as .bat and .sh file scripts. 7. If you have images to compare, you can copy them or subfolders of images into the Image folder in the distribution directory.

To start Flicker in Windows, click on the startup icon for Flicker.exe. You can also start it in Windows by clicking on the Flicker-startup.bat file. For Unix systems including MacOS-X, you can start Flicker from a command line file by specifying the path to Flicker-startup.sh. Normally it comes up with the two demonstration human plasma 2D-gel images (plasmaH.gif - an IPG gel from SWISS-2DPAGE on the left) and (plasmaL.gif – a carrier ampholyte gel from Dr. Carl Merril/NIMH on the right). If you have your own gels (JPEG, GIF or TIFF formats), you can try loading them. But first you must copy the files or folders of files to the Images/ folder where you installed Flicker. Then use the (File | Open user images | …) commands to load the images. There is currently no way to load images stored in a relational database. You may want to limit resolution by first decreasing their size using an image editing program like Adobe PhotoShop, shareware program ThumbsPlus (www.cerious.org), or open source programs like Gimp (www. gimp.org). Large very high-resolution images that are 20 Mb to 40 Mb will not work well. We suggest reducing the size to about 1 K×1 K for good interactivity if you have any problems with running out of memory or very sluggish response. These image-editing programs can also be used for converting other formats to JPEG, GIF or TIFF formats that Flicker can read. 3.2. Graphical User Interface for Flickering Fig. 1 shows the initial screen of the Flicker program. Pull-down menus at the top invoke file operations, edit preferences, view overlay options, landmarking, image transforms, and help commands. Scroll bars on the side determine various parameters used in the transforms. The two images to be compared are

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Fig. 3. Screen views of clickable active gel compared with another gel. (A) We have loaded the active SWISS-2DPAGE human plasma gel (PLASMA_HUMAN_id) in the left image and the plasmaL gel in the right image. Spots that appear in SWISS-2DPAGE are indicated with red “+” symbols. We then aligned the images using flickering. We then selected the Click to access DB checkbox. Finally, we clicked on the indicated spot in the left gel to determine the putative identification of the corresponding spot in the right gel. (B) The SWISS-2DPAGE window then popped up as a result of clicking on that spot in the left image and indicates the putative protein identification of the visually corresponding spot in the right gel. The plasmaH image is the same gel as PLASMA_HUMAN_id but without the graphic overlays. You can load these same gels using the (File | Open demo images | Human Plasma gels | (SWISS-2DPAGE vs Merril) - clickable) command, which should be used when you are connected to the Internet.

loaded into the lower scrollable windows. A flicker window appears in the upper middle of the screen. Checkboxes on the left activate flickering and control display options. A group of status lines below the checkboxes indicate the state

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Fig. 3. (continued)

of operations. Table 1 shows the summary of the commands in the pull-down menus. Only part of an image is visible in the scrollable windows. These subregions are determined by setting horizontal and vertical scroll bars. Another, preferred, method of navigating the scrollable images is to click on the point of interest while the CONTROL key is pressed. This will re-center the scrollable image around that point. This lets the user view any subregion of the image at high resolution. These images may be navigated using either the scroll bars or by moving the mouse with the button pressed in the scrollable image window. Then, each image in the flicker window is centered at the point last indicated in the corresponding scrollable image window. Note that if you are near the edge of the image when you do this, it will may not scroll the image properly. To fix this problem you can add a “guard region” around the edge of the image using the (View | Add guard region to edges of images). This is useful for aligning spots that are along the edge of the images while flickering.

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The guard region’s color can be changed via the Edit menu (Edit | Set colors | Guard region color). The guard region requires more memory that is why the default is to have the guard region default to off. A flicker window is activated in the upper-middle of the screen when the “Flicker” checkbox is selected. Images from the left and right scrollable images are alternatively displayed in the flicker window. The flicker delay for each image is determined by the adjusting the scroll bar below the corresponding scrollable image window. Various graphic overlays may be turned on and off using the various view “overlays” selected in the (View | <sub-menus> …) checkbox menu commands. Clicking on either the left or right image selects it as the image to use in the next transform. However, clicking on the flicker image window indicates the next transform you might use should be applied to both left and right images. You can change the selected image by just clicking on any of the images. You can increase or decrease the size of the three image windows by using the (Edit | Canvas size | Increase size (C-keypad ‘+’) ) and (Edit | Canvas size | decrease size (C-keypad ‘−’) ) commands. This will resize the main window accordingly. Alternatively, and easier to do, is to click on either the left-arrow (shrink) or right-arrow (enlarge) buttons on the Canvas size scroller on the middle right edge of the Flicker application window. 3.3. Loading Images As mentioned in the introduction, gel images may be loaded into the left or right selected image from: (a) the local computer using the (File | Open image file) command or (b) any Internet site using the (File | Open image URL) command. You may load pairs of demonstration images that come with Flicker, and are installed in the Images/ directory. Use the (File | Open demo images | …) command to load them into the left and right images. The demos include a few samples that may be useful for initially learning the system. They include: two human plasma gels - an IPG SWISS-2DPAGE gel vs a carrier ampholyte gel (Merril/NIMH); some human leukemias (AML, ALL, CLL, HCL) from Lester et.al. (10–11). You may specify active gel images from the Web using the (File | Open active map image | …) to let you load one of the Swiss-2DPAGE gel images into the left or right selected image. This list of active images is defined by the tabdelimited FlkMapDB.txt file read by Flicker when it is started. “Power users” could edit this file (use Excel and save as tab-delimited) to add active map entries pointing to other federated 2D-gel Web databases. The FlkMapDB.txt file is provided with your download installation in the DB/ directory. Gel images are loaded into the lower left or right images. First click on the left or right image you want to replace. Then, select the active gel image to you want using the entry (File | Open active map image | …) pull-down menu (e.g., select (… | SWISS-2DPAGE Human | Human Plasma) ). Next, click on the other image

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and then open the other gel image you want to compare it with using either (File | Open image file) or either (File | Open image URL) commands. 3.4. Adding Your Own Image Data to the User Images/Database There is another way for users to add many of their gel images without editing the DB/FlkDemoDB.txt file. When you place your image data directories in the Images/ directory, Flicker will discover them when it starts and add them to the demo menu. It works as follows: 1. You copy or move one or more of your directories of with the images you want to use with Flicker in the Images/ folder. 2. When Flicker starts, it creates additional submenu entries in the (File | Open user images | Pairs of images | …) and (File | Open user images | Single images | …) submenus that are the names of the user’s directories. 3. The first submenu contains unique combinations of pairs of all images within each of the user’s directories. Selecting one of these entries will load the pair of images into the left and right Flicker image windows. 4. The second menu command lets you select the right or left Flicker image, and then load a single image from any of the user image directories into that Flicker image window. This would be useful if you wanted to compare one of your images with one of the Internet reference gels.

3.4.1. Flickering When flickering two images with the computer, one aligns putative corresponding subregions of the two rapidly alternating images. The flicker window overlays the same space on the screen with the two images and is aligned by interactively moving one image relative to the other using the cursor in either or both of the lower images. Using the mouse, the user initially selects what they suspect is the same prominent spot or object in similar morphologic regions in the two gel images. The images are then centered in the flicker window at these spots. When these two local regions come into alignment, they appear to pulse and the images fuse together. At this point, differences are more apparent and it is fairly easy to see which spots or objects correspond, which are different, and how they differ. We have found that the user should be positioned fairly close to the flicker window on the screen to optimize this image-fusion effect (i.e., it does not work as well standing back more than a few feet from the screen). 3.4.2. Selecting the Proper Time Delays When Flickering The proper flicker delays, or time each image is displayed on the screen, is critical for the optimal visual integration of image differences. We have also found that optimal flicker rates are dependent on a wide variety of factors including: amount of distortion, similarity of corresponding subregions,

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complexity and contrast of each image, individual viewer differences, phosphor decay-time of the display, ambient light, distance from the display, and so on. We have found the process of flickering images is easier for some people than for others. When comparing a light spot in one gel with the putative paired darker spot in the other gel one may want to linger longer on the lighter spot to make a more positive identification. Because of this, we give the user the ability to set the display times independently for the two images (typically in the range of 0.01 second to 1.0 second with a default of 0.30 second) using separate Delay scroll bars located under each image. If the regions are complex and have a lot of variation, longer display times may be useful for both images. Differential flicker delays with a longer delay for the light gel are useful for comparing light and dark sample gels. This lets you stare at the lighter spots to have more verification that they are actually there. 3.5. Image Processing Methods As mentioned, there are a number of different image transforms that may be invoked from the menus. These are useful for changing the geometry, sharpness, or contrast making it easier to compare potentially corresponding regions. As we go through the transforms we will indicate how they may be used. Some affect one image while some affect both. Flickering is deactivated during image transforms to use most computational power for doing the transforms. The TRANSFORM menu has a number of commands that include warping, grayscale transforms and contrast functions. The two warp method selections: “Affine Warp and “Poly Warp” are performed on only one image (the last one selected by clicking on an image). The “Pseudo-3-D” transform makes a 3-D image with the “peaks” created proportional to gray level. Unlike the warp transforms, the grayscale transforms are performed on both images. These include: “SharpenGradient”, “SharpenLaplacian”, “Gradient”, “Laplacian”, “Average”, “Median”, “Max3×3” and “Min3×3”. The contrast functions are “Complement” and “ContrastEnhance”. You can transform image color images to grayscale using the “Color to grayscale” command, and generate a false color image from a grayscale using the “Pseudo color” command. You can flip the image using “Flip image horizontally” or “Flip image vertically” commands. 3.5.1. Landmarks: Trial and Active The affine transform requires that three active landmarks be defined before it can be invoked. A trial landmark is defined by clicking on an object’s center anywhere in a scrollable image window. This landmark would generally be placed on a spot. Clicking on a spot with or without the CONTROL key pressed still defines it as a trial landmark. After defining the trial landmark in both the

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left and right windows, selecting the (Landmark | Add Landmark (C-A) ) command to define them as the next active landmark pair and identifies them with a red letter label (+A, +B, +C, …) in the two scrollable image windows. The (Landmark | Delete Landmark (C-D) ) command is used for deleting the last landmark you defined. 3.5.2. The Affine Transform for Spatial Warping The two warping transforms, affine (see Eqs. 1 and 2) and polynomial, require 3 and 6 landmarks respectively. Attempting to run the transform with insufficient landmarks will cause Flicker to notify you that additional landmarks are required. The image to be transformed is the one last selected. You must select either the left or right image. Fig. 2a shows the landmarks the user defined in the two gels before the affine transform. Fig. 2b shows the transformed image. Then, recenter the transformed image before you flicker. After the transform, the landmarks can be lined up perfectly, and adjacent spots will line up better. 3.5.3. Pseudo-3-D transform As described in (1–2, 4–6) and as shown in Eqs. 3–5, the Pseudo-3-D transform generates a pseudo 3D relief image to enhance overlapping spots with smaller spots seen as side peaks. The gray value determines the amount of “y” shift scaled by a percentage (set by scroll bar zscale (in the range of 0 to 50%). Pseudo perspective is created by shifting the image to the right or left by setting by scroll bar “angle” degrees (in the range of −45 to +45 degrees). Negative angles shift it to the right and positive angles to the left. The image to be transformed is the one last selected. If neither was selected (i.e., you clicked on the flicker window), then both images are transformed. 3.5.4. Edge Sharpening Edge sharpening may be useful for improving the visibility of the edges of fuzzy spots. You can select either a Gradient or Laplacian edge sharpening function using the “SharpenGradient” or “SharpenLaplacian” operation in the TRANSFORM menu where the image to be transformed is the one last selected. The Laplacian filter generates a “softer” edge than the Gradient. You can set the scroll bar escale value (in the range of 0 to 50%) to scale the amount of edge detection value added. The image to be transformed is the one last selected. If neither was selected (i.e., you clicked on the flicker window), then both images are transformed. 3.5.5. Putative Identification of a Spot in One Gel by Comparison with Federated Database Gel Map Open an active gel image in the lower left or right window. First click on the window you want to load the new image. Then, select the active gel image to

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you want using the entry (File | Open active map image | …) pull-down menu (e.g., select (… | SWISS-2DPAGE Human | Plasma) ). Next, click on the other image and then use the other gel image you want to compare it with using one of the other (File | Open …) commands At this point flicker the two images so that you can make a putative guess on which spot you are interested in your gel corresponds to which spot in the active map gel. Then shut off flickering by turning off the “Flicker” checkbox. Then turn on the “Click to access DB” checkbox. Then click on the spot in the active map image which will pop up a Web browser window indicating the SWISS2DPAGE web page for that spot if it is in their database. 4. NOTES 1. Status of the Flicker application. As of the time of the submission of this chapter, most of the functionality available in the former Java applet (1–2, 4–6) is fully functional in the standalone application. The current state of Flicker is documented on the Web server. A future release of Flicker will contain a spot quantification capability with the ability to calibrate the image (either from the calibration information in the image itself if available, or from a scanned neutral density step wedge scanned with the image), estimate background density, and estimate spot intensity with a background subtraction. Documentation is available on the Web site. This documentation may also be invoked from the “Help” submenus. The original Flicker program was converted from a Java applet to a Java application by Peter Lemkin and Greg Thornwall with help from Jai Evans. Code was added from the open source MicroArray Explorer (http://maexplorer. sourceforge.net/) program. The new Flicker program uses the Mozilla 1.1 open source license and is available on the open source Web server http://open2Dprot. sourceforge.net/Flicker/. You can update your program and data files using the various Update options in the Files menu. The (File | Update | Update Flicker Program) command downloads and installs the latest Flicker.jar file. The (File | Update | Update active Web maps DB) command downloads and installs the latest active Web maps database DB/FlkMapDB.txt file. The (File | Update | Update Demo images DB) command downloads the latest demo images into the Images/ directory. 2. Hints on measuring spots. There are some disadvantages in comparing gels visually. It is useful for doing a rough comparison and there is currently no simple way available to do adequate quantitative comparison (as can be done with existing 2D-gel computer database systems) that use automatic spot segmentation and global normalization methods. However, you can look at the gray value of the cursor in

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the left or right image if you enable the (View | m Display gray values in image title (C-G) ) menu option. The program also allows single-spot quantification with optical density calibration. These limitations should be kept in mind when using the technique. In the mean time, if your gels and scanner are reasonably linear so that grayscale approximates protein concentration, you can use the simple grayscale method that can be used for the ballpark estimates of density. You can do this either of two ways: by measuring the area under a circular mask (you can set the radius), or the area inside a rectangular Region of Interest (ROI). Note that unless the spot fits well inside of the mask or ROI, you will not get a very accurate measurement. Both methods can subtract an optional background value you can capture and so can give intensity corrected for background if defined. To set the measurement circle region size (1×1, 3×3, 5×5 …, or 11×11), select the image you will work on and the adjust the Meas Circle slider on the right.. Click on a background region near where you want to measure a spot’s density within a circular mask. Then select the (Quantify | Measure by circle | Capture background (C-B) ) command. Then click on the center of the spot you want to estimate and select the (Quantify | Measure by circle | Capture measurement (C-M) ) command. This will compute and display background corrected data that appears in the report window as: plasmaH.gif (201,392) totBkgrd: 1557gray, meanBkgrd: 21 gray CircleMask: 5X5 area: 73 pixels (1) plasmaH.gif (214,376) Tot(Meas-Bkgrd): 9418.000 ygra TotMeas: 10975.000 (39.000:227.000) gray TotBkgrd: 1557.000 (201,392) (16.000:28.000) gray mnDens: 154.577, mn(Dens-Bkgrd): 132.648, mnBkgrd: 21.930 gray CircleMask: 5X5 area: 73 pixels

The (View | Set measurement view options | View measurement circle) displays the background and foreground measurement circles if enabled with “B” and “M” labels. Set the (View | Use sum density else mean density) menu option to specify that it report either total region density or mean density. You also can measure intensity inside a rectangular ROI regions you set by using both the (Quantify | Region of Interest (ROI) | Set ROI ULHC (C-U) ) and the (Quantify | Region of Interest (ROI) | Set ROI LRHC (C-L) ) commands. Use the (View | View Region Of Interest (ROI) ) to display the ROI as a rectangle from the ULHC to the LRHC (upper left-hand corner and lower right-hand corner). You can measure the integrated density of the (Quantify | Region of Interest (ROI) | Capture measurement by ROI (C-R) ) command. If you set it, the (C-R) command will subtract the background computed by the area times the mean background using the (Quantify | Measure by circle | Capture background (C-B) ) command.

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This will compute and display background corrected data that appears in the report window as: Setting ULHC (163,370) of ROI: left image Setting LRHC (181,389) of ROI: left image (1) plasmaH.gif (181,389) Tot(Meas-Bkgrd): 44989.309 gray TotMeas: 49271.000 (13.000:238.000) TotBkgrd: 4281.690 at (133,429) (9.000:20.000) gray ROI: (163:1 81, 370:389 )

When grayscale calibration is added in a future version, then the measurements will be in terms of the calibration rather than grayscale. The intent of applying image transforms is to make it easier to compare regions having similar local morphologies but with some different objects within these regions. Image warping prior to flickering is intended to spatially warp and rescale one image to the geometric “shape” of the other image so that we can compare them at the same scale. This should help make flickering of some local regions on quite different gels somewhat easier. Of the two warping transforms, affine and polynomial, the latter method handles nonlinearities better. For those cases where the gels are similar, the user may be able to get away with using the simpler (affine) transform. For demonstration purposes, if you are using the demo plasmaH and plasmaL gels, the (Landmark | Set 3 predefined landmarks for demo gels (C-Y) ) and (Landmark | Set 6 predefined landmarks for demo gels (C-Z) ) define 3 and 6 corresponding landmarks for these gels that may be used with the affine and polynomial warping transforms respectively. In cases where there is a major difference in the darkness or lightness of gels, or where one gel has a dark spot and the other a very faint corresponding spot, it may be difficult to visualize the light spot. By differentially setting the flicker display-time delays, the user can concentrate on the light spot using the brief flash of the dark spot to indicate where they should look for the light spot. We have found differential-flicker to be very helpful for deciding difficult cases. Adjusting one image so that its brightness and contrast are approximately that of the other image also helps when flickering. You change the image brightness and contrast using the Shift/Drag mouse control described below. 3. Additional hints on image transforms. Other transforms including image sharpening may be useful in cases where spots are very fuzzy, as might be the case when comparing Southern blots. When two corresponding local regions of the two images are radically different so the local morphologies are not even slightly similar (e.g., when high MW regions of

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gels that are run differently such as: IPG vs. non-IPG, gradient vs. nongradient SDS), then even using these transforms may not help that much. 4. Saving and restoring the Flicker state. Flicker gives you the option of saving the current state of your session including the images your are looking at and the parameter values of the sliders, etc. To save the current state, use the (File | Save (or SaveAs) state file) command. This creates a file with a .flk file extension in the installation FlkStartups/ folder (default FlkStartup.flk). If you have used the Flicker Web site Java installer (ZeroG.com) for installing Flicker, then it lets you click on a specific .flk you have previously saved to restart it where you left off. While running Flicker, you can also use (File | Open state file) command to change it to another state. 5. Mouse control of images. The following mouse and key-modified mouse operations control various actions. Pressing the mouse in either the left or right image selects it. If flickering is active, then it will move the flicker image center for the selected image to that position. A little yellow “+” indicates the position you have selected. If the Click to access DB checkbox is enabled and the image has an associated active map database server associated with it, then it will request the spot identify when you click on a spot from the map database.. Dragging the mouse is similar to pressing it. However, only pressing it will invoke a clickable database. It also displays the cursor coordinates in the image title. Control/Press will position the selected image so that the point you have clicked on will be in the center of the crosshairs. If you are near the edge of the image, it will ignore this request. Shift/Drag activates the brightness/contrast filter with minimum brightness and contrast in the lower left hand corner. 6. Checkbox control of flickering and database access. There are four checkboxes in the upper left part of the window that control commonly used options. Flicker checkbox enables/disables flickering. Click to access DB checkbox enables/disables access to a Web database server that is associated with a clickable image - if it exists for the selected image. Turning on this option will disable flickering. Allow transforms checkbox enables/disables image transforms. Sequential transforms checkbox enables/disables using the last image transform output as input for the next image transform (image composition) if Allow transforms is enabled.

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7. Keyboard shortcut controls There are several short-cut key combinations that can be use to perform operations instead of selecting the command from the pull-down menus. The notation C- means to hold the Control key and then press the following . C-A add landmark (you must have selected both left and right image trial objects) (see Landmark menu) C-B capture background intensity value for current image under a circular mask (see Quantify menu) C-D delete landmark - the last landmark defined (see Landmark menu) C-E edit selected measured spot (click on spot to select it) (see Quantify menu) C-F toggle flickering the lower left and right images into the upper flicker window (see View menu or the Flicker (C-F)) C-G toggle displaying gray values in the left and right image titles as move the cursor (see View menu) C-H show grayscale ROI histogram. Popup a histogram of the computation region ROI (see Quantify menu) C-I define or edit selected measured spot(s) annotation ‘id’ field (see Quantify menu) C-J toggle the spot measurement mode between list-of-spots measurement mode and the single spot trial-spot measurement mode (see Quantify menu) C-K delete selected measured spot (click on spot to select it (see Quantify menu) C-L define the lower right hand corner (LRHC) of the region of interest (ROI) and assign that to the computing window (see Quantify menu) C-M measure and show intensity under a circular mask for current image. Report background-corrected value if background was defined (see C-B shortcut and Quantify menu). An alternative way to measure spots is to hold the ALTkey when you press the mouse to select the spot. This combines spot selection and measurement in one operation. C-R measure and show intensity under a the computing window defined by the ROI (see commands (C-U) and (C-L) ) for the current image. Report background-corrected value if circular mask background was defined (see (C-B) shortcut and Quantify menu) C-T repeat the last Transform used, if one was previously performed else no-op (see Transform menu) C-U define the upper left hand corner (ULHC) of the region of interest (ROI) and assign that to the computing window (see Quantify menu) C-V show data-window of selected pixel in the popup report window. C-W clear the region of interest (ROI) and computing window (see Quantify menu) C-Y set 3 predefined landmarks for demo gels for Affine transform (see Landmark menu)

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C-Z set 6 predefined landmarks for demo gels for Polywarp transform (see Landmark menu) C-Keypad “+” increase the canvas size for all three images (see Edit menu) C-Keypad “-” decrease the canvas size for all three images (see Edit menu) 8. Reporting the status in the popup status window. Information is display in several places in Flicker. a. There are two status lines in the upper left part of the main window. The output into these status lines is also appended to the Report window (c). b. The selected image (clicking on the left or right image) changes its title to blue from black. If neither image is selected, then both titles are black. c. A report popup window is created when Flicker is started. It may be temporarily removed by closing it. You can get it back at any time by selecting (View | Show report popup). All text output is appended to this window. The Clear button clears all text. The SaveAs button lets you save the text in the window into a local text file.

9. Sliders for defining parameters. The following sliders are in the upper right part of the window and are used for adjusting parameters in the various image transforms. ZoomMag (X) to zoom both left and right images from 1/10X to 10X by a transform (3D) angle (degrees) used in the pseudo 3D transform (3D) z-Scale (%)used in the pseudo 3D transform (Sharpen) e-Scale(%) used in the sharpening transforms Contrast (%) set by Shift/Drag to change the image contrast Brightness (%) set by Shift/Drag to change the image brightness Threshold1 (grayscale or od) is the minimum grayscale value to show pixels otherwise they are shown as whites Threshold2 (grayscale or od) is the maximum grayscale value to show pixels otherwise they are shown as white Meas circle (diameter in pixels) size of the measurement circle for selected image 10. Local database files. When Flicker is installed, several tab-delimited (spreadsheet derived) .txt files are available in the DB/ directory (located where the Flicker.jar file is installed). These DB/Flk*DB.txt files are read on startup and are used to setup the (File | Open … image | …) menu trees. DB/FlkMapDB.txt - contains instances of Web-based active image maps with fields: (MenuName, ClickableURL, ImageURL, BaseURL, DatabaseName) DB/FlkDemoDB.txt - contains instances of pairs of images in the local Images/ directory and contains fields:

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(SubMenuName, SubMenuEntry, ClickableURL1, ImageURL1, ClickableURL2, ImageURL2, StartupData) DB/FlkRecentDB.txt - contains instances of recently accessed non-demo images with fields: (DbMenuName, ClickableURL, ImageURL, DatabaseName, TimeStamp) 11. Files required that are included in the download. The following files are packaged in the distribution and installed when you install Flicker. Flicker.jar is the Java Archive File for Flicker that is executed when you run Flicker jai_core.jar is the core Java runtime from SUN’s Java Advanced Imaging (JAI) at sun.com jai_codec.jar is the JAI tiff file reader from SUN’s Java Advanced Imaging JAI at sun.com DB/ is a directory containing the set of tab-delimited DB files Flk*DB.txt read at startup Images/ is a directory holding demo .gif, .tif, and .ppx sample files FlkStartups/ is empty directory to put the startup FlkStartup.flk files Flicker-startup.bat is a Windows .bat batch script startup file Flicker-startup.sh is a Unix/MacOS-X command line script startup file 12. Image transform and brightness-contrast display model. There are several display models for combinations of using image transforms, zooming, and brightness contrast filtering. Zooming is an image transform and you can de-magnify as well as magnify. These transforms and filtering are applied to the left and right windows and also are shown in the flicker window. Two checkboxes in the upper left of the main window control transforms: “Allow transform” enables/disables transforms, and “Sequential transforms” allows using the previous transform as the input to the next transform, i.e., this lets you implement image composition. This description applies independently to the left and right images. The original image is denoted iImg. If you allow transforms and are also composing image transforms, you may optionally use the previous transformed output image (denoted oImg) as input to the next image transform. The output (either iImg or oImg) is then sent to the output1. Then output1 may be optionally zoomed to output2 by being sent to the zoom transform (iff the magnification is different from 1.0X). Then output2 may be optionally contrast-adjusted by being sent to the brightness-contrast filter (if it is active as specified by dragging the mouse in the selected window with the SHIFT-key pressed). The output2 of the brightness-contrast filter is denoted as bcImg. If you have never used the

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zoom or brightness-contrast filtering since loading an image, then zImg and bcImg are not generated and hence not used in the displayed image. This will speed up display refresh as you navigate the windows. a. If no transforms or brightness-contrast filtering is used on the selected image (no tr ansforms) iImg Æ output1 b. The image may be optionally transformed from the original image (iImg) (transform) iImg Æ oImg Æ output1 c. Image transforms may be optionally composed from the original image or from the sequential composition of image transforms on the selected image (sequential tr ansforms) iImg

Æ

Transform

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d. The image may be optionally zoomed if the magnification is not 1.0X (zoom) output1 Æ zImg Æ output2 or output1

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e) The brightness-contrast filter may be optionally applied to the image (B-C filter) output2 Æ bcImg Æ display or output2

(no B-C filter) Æ display

References 1. Lemkin, P. F. (1997) Comparing Two-Dimensional electrophoretic gels across the Internet. Electrophoresis 18, 461–470. 2. Lemkin, P. F., Thornwall, G. (1999) Flicker image comparison of 2D gel images for putative protein identification using the 2DWG meta-database. Mol. Biotechnol. 12, 159–172. 3. Lemkin, P. F. (1997) 2DWG meta-database of 2D electrophoretic gel images on the Internet. Electrophoresis 18, 2759–2773. 4. Lemkin, P. F. (1997) Comparing 2D Electrophoretic gels across Internet databases. In 2D Protocols for Proteome Analysis, Andrew Link (ed), Humana Press, Totowa, NJ, pp 339–410.

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5. Lemkin, P. F. and Thornwall, G. (2002) Comparing 2D Electrophoretic gels across databases. In Protein Protocol Handbook, 2nd edn. J. M. Walker (ed), Humana Press, Totowa, NJ, pp 197–214. 6. Lemkin, P. F., Thornwall, G. C., and Evans, J. (2005) Comparing 2D Electrophoretic gels across databases. In Protein Protocol Handbook, J. M. Walker (ed), Humana Press, Totowa, NJ, pp 279–306. 7. Lemkin, P. F., Merril, C., Lipkin, L., Van Keuren, M., Oertel, W., Shapiro, B., Wade, M., Schultz, M., and Smith, E. (1979) Software aids for the analysis of 2D gel electrophoresis images. Comput. Biomed. Res. 12, 517–544. 8. Lipkin, L.E. and Lemkin, P. F. (1980) Data-base techniques for multiple two-dimensional polyacrylamide gel electrophoresis analyses. Clin. Chem. 26, 1403–1412. 9. Lemkin, P. F. and Lester, E. P. (1989) Database and search techniques for twodimensional gel protein data: a comparison of paradigms for exploratory data analysis and prospects for biological modeling. Electrophoresis 10, 122–140. 10. Lester, E. P., Lemkin, P. F., Lowery, J. F., and Lipkin L. E. (1982) Human leukemias: a preliminary 2D electrophoretic analysis. Electrophoresis 3, 364–375. 11. Lemkin, P. (2006) The LECB 2D PAGE Gel Images Data Sets. These 2D-gel image data sets from GELLAB-II research were made available for public use. http:// bioinformatics.org/lecb2Dgeldb/ 12. Lemkin, P. F., Thornwall, G. C., Walton, K. D., and Hennighausen, L. (2000) The Microarray Explorer tool for data mining of cDNA microarrays - application for the mammary gland. Nucleic Acid Res. 28, 4452–4459. 13. Duncan, M. W. and Hunsucker, S. W. (2005) Proteomics as a tool for clinically relevant biomarker discovery and validation. Exp. Biol. Med. (Maywood) 230, 808– 817. 14. Herbert, B. R., Pederson, S. L., Harry, J. L., Sebastian, L., Traini, M. D., McCarthy, J. T., Wilkins, M. R., Gooley, A., Packer, N. H., and Williams K. (2003) Mastering genome complexity using two-dimensional gel electrophoresis. PharmaGenomics Sept, 22–36. 15, Hood, L. (2002) A Personal View of Molecular Technology and How It Has Changed Biology. J. Proteome Biol. 1, 399–409. 16. Herbert, B. R., Pedersen, S. K., Harry, J. L., Sebastian, L., Grinyer, J., Traini, M. D., McCarthy, J. T., Wilkins, M. R., Gooley, A. A., Righetti, P. G., Packer, N. H., and Williams, K. K. (2003) Mastering Proteome Complexity Using Two-Dimensional Gel Electrophoresis. PharmaGenomics 3, 21–36. 17. Pedersen, S. K., Harry, J. L., Sebastian, L., Baker, J., Traini, M. D., McCarthy, J. T., Manoharan, A., Wilkins, M. R., Gooley, A. A., Righetti, P. G., Packer, N. H., Williams, K. L., and Herbert, B. R. (2003) Unseen Proteome: Mining Below the Tip of the Iceberg To Find Low Abundance and Membrane Proteins. J. Proteome Biol. 2, 303–311. 18. 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, 422–432.

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19. Pieper, R., Gatlin, C. L., Makusky, A. J., Russo, P. S., Miller, S. S., Su, Q., McGrath, A. M., Estock, M. A., Parmar, P. P., Zhao, M., Huang, S.-T., Zhou, J., Wang, F., Esquer-Blasco, R., Anderson, N. L., Taylor, J., and Steiner, S. (2003) The human serum proteome: Display of nearly 3700 chromatographically separated protein spots on two-dimensional electrophoresis gels and identification of 325 distinct proteins. Proteomics 3, 1345–1364. 20. Anderson, N. L. and Anderson, N. G. (2002). J. Mol. Cell. Proteomics 1, 845–867; J. Mol. Cell. Proteomics 2, 50. 21. Liotta, L. A., Espina, V., Mehta, A. I., Calvert, V., Rosenblatt, K., Geho, D., Munson, P. J., Young, L., Wulfkuhle, J., and Petricoin, E. F. 3rd. (2003) Protein microarrays: meeting analytical challenges for clinical applications. Cancer Cell 3, 317–325. 22. Reid, J. D., Parker, C. E., and Borchers, C. H. (2007) Protein arrays for biomarker discovery. Curr. Opin. Mo.l Ther. 9, 216–221. 23. Hoving, S,. Gerrits, B., Voshol, H., Muller, D., Roberts, R. C., and van Oostrum J. (2002) Preparative two-dimensional gel electrophoresis at alkaline pH using narrow range immobilized pH gradients. Proteomics 2, 127–134. 24. Overbye, A., Fengsrud, M., and Seglen, P. O. (2007) Proteomic analysis of membrane-associated proteins from rat liver autophagosomes. Autophagy 3:300–322. 25. Von Eggeling, F., Gawriljuk, A., Fiedler, W., Ernst, G., Claussen, U., Klose, J., and Romer I. (2001) Fluorescent dual colour 2D-protein gel electrophoresis for rapid detection of differences in protein pattern with standard image analysis software. Int. J. Mol. Med. 8, 373–377. 26. GE Healthcare’s Ettan DIGE Extended Analysis software. http://www1.gelifesciences.com/APTRIX/upp00919.nsf/Content/Proteomics+DIGEProteomics+DIGE +Software+DeCyder+downloads+software 27. Lopez, M. F., Mikulskis, A., Golenko, E., Herick, K., Spibey, C. A., Taylor, I., Bobrow, M., and Jackson, P. (2003) High-content proteomics: Fluorescence multiplexing using an integrated, high-sensitivity, multi-wavelength charge-coupled device imaging system. Proteomics 3, 1109–1116. 28. Sanchez, J.-C., Appel, R. D., Golaz, O., Pasquali C., Rivier, F., Bairoch, A., Hochstrasser, D. F. (1995) Inside SWISS-2DPAGE database. Electrophoresis 16, 1131–1151. 29. Appel, R. D., Bairoch, A., Sanchez, J-C., Vargas, J. R., Golaz, O., Pasquali, C., and Hochstrasser, D. F. (1996). Federated two-dimensional electrophoresis database: A simple means of publishing two-dimensional electrophoresis data. Electrophoresis 17, 540–546. 30. Appel, R. D., Sanchez, J-C., Bairoch, A., Golaz, O., Miu, M., Vargas J. R., and Hochstrasser, D. F. (1993) SWISS-2DPAGE: A database of two-dimensional gel electrophoresis images. Electrophoresis 14, 1232–1238. 31. Appel, R. D., Sanchez, J.-C., Bairoch, A., Golaz, O., Rivier, F., Pasquali, C., Hughes, G. J., and Hochstrasser, D. F. (1994) SWISS-2DPAGE database of two-dimensional polyacrylamide gel electrophoresis. Nucleic Acids Res. 22, 3581–3582. 32. Arnold, K., Gosling, J. (1996) The Java Programming Language. Addison Wesley, New York. 33. Wolberg, G. (1990) Digital Image Warping, IEEE Computer Press Monograph, Los Alamitos, CA.

41 Quantification of Radiolabeled Proteins in Polyacrylamide Gels Wayne R. Springer 1. Introduction Autoradiography is often used to detect and quantify radiolabeled proteins present after separation by polyacrylamide gel electrophoresis (PAGE) (see Chapter 56). The method, however, requires relatively high levels of radioactivity when weak β-emitters, such as tritium, are to be detected. In addition, lengthy exposures requiring the use of fluorescent enhancers are often required. Recent developments in detection of proteins using silver staining (1, 2) have added to the problem because of the fact that tritium emissions are quenched by the silver (2). Since for many metabolic labeling studies, tritium labeled precursors are often the only ones available, it seemed useful to develop a method that would overcome these drawbacks. The method the author developed involves the use of a cleavable crosslinking agent in the polyacrylamide gels that allows the solubilization of the protein for quantification by scintillation counting. Although developed for tritium (3), the method works well with any covalently bound label, as demonstrated here with 35S. Resolution is as good as or better than autoradiography (3 and Fig. 1), turnaround time can be greatly reduced, and quantification is more easily accomplished. 2. Materials Reagents should be of high quality, particularly the sodium dodecyl sulfate (SDS) to obtain the best resolution, glycerol to eliminate extraneous bands, and ammonium persulfate to obtain proper polymerization. Many manufacturers supply reagents designed for use in polyacrylamide gels. These should be purchased whenever practical. Unless otherwize noted, solutions may be stored indefinitely at room temperature.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Comparison of an autoradiogram, densitometry scan, and radioactivity in gel slices from a mixture of labeled proteins separated by PAGE. Identical aliquots of 35S-methioninelabeled proteins from the membranes of the cellular slime mold, D. purpureum, were separated using the methods described in Subheading 3.1. One lane was fixed, soaked in Fluoro-Hance (Research Products International Corp., Mount Prospect, IL), dried, and autoradiographed (photograph). Another lane was cut from the gel, sliced into 1-mm pieces, solubilized, and counted using a Tracor Mark III liquid scintillation counter with automatic quench correction (TM Analytic, Elk Grove Village, IL) as described in Subheading 3.3. The resultant disintigrations per minute (DPM) were plotted vs the relative distance from the top of the running gel (top figure). The autoradiogram (photograph) was scanned relative to the top of the running gel using white light on a Transidyne RFT densitometer (bottom figure).

2.1. SDS-Polyacrylamide Gels 1. Acrylamide-DATD: Acrylamide (45 g) and N,N′-dialyltartardiamide (DATD 4.5 g) are dissolved in water to a final volume of 100 mL. Water should be added slowly and time allowed for the crystals to dissolve. 2. Acrylamide-bis: Acrylamide (45 g) and N,N′-methylenebis(acrylamide) (bis-acrylamide, 1.8 g), are dissolved as in item 1 in water to a final volume of 100 mL.

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3. 4. 5. 6.

Tris I: 0.285 M Tris-HCl, pH 6.8. Tris II: 1.5 M Tris-HCl, pH 8.8, 0.4% SDS. Tris III: 0.5 M Tris-HCl pH 6.8, 0.4% SDS. Ammonium persulfate (APS) solution: A small amount of ammonium persulfate is weighed out and water is added to make it 100 mg/mL. This should be made fresh the day of the preparation of the gel. The ammonium persulfate crystals should be stored in the refrigerator and warmed before opening. 7. N,N,N′,N′-tetramethylethylenediamine (TEMED) is used neat as supplied by the manufacturer. 8. 10X Running buffer: Tris base (30.2 g), glycine (144.1 g), and SDS (10.00 g) are made up to 1 L with distilled water. 9. Sample buffer: The following are mixed together and stored at 4°C: 1.5 mL 20% (w/v) SDS, 1.5 mL glycerol, 0.75 mL 2-mercaptoethanol, 0.15 mL 0.2% (w/v) Bromophenol blue, and 1.1 mL Tris I.

2.2. Silver Staining of Gels All solutions must be made with good-quality water. 1. 2. 3. 4. 5.

Solution A: methanol:acetic acid:water (50:10:40). Solution B: methanol:acetic acid:water (5:7:88). Solution C: 10% (v/v) glutaraldehyde. 10X Silver nitrate: 1% (w/v) silver nitrate stored in brown glass bottle. Developer: Just before use, 25 µL of 37% formaldehyde are added to freshly made 3% (w/v) sodium carbonate. 6. Stop bath: 2.3 M Citric acid (48.3 g/100 mL).

2.3. Solubilization and Counting of Gel Slices 1. Solubilizer: 2% sodium metaperiodate. 2. Scintillation fluid: Ecolume (+) (ICN Biomedical, Inc., Irvine, CA) was used to develop the method. See Subheading 4. Notes. for other scintillants.

3. Methods The general requirements to construct, prepare, and run SDS-PAGE gels as described by Laemmli (4) are given in Chapter 11. Presented here are the recipes and solutions developed in the author’s laboratory for this particular technique. For most of the methods described a preferred preparation of a standard gel can be substituted, except for the requirement of the replacement of DATD for bis at a ratio of 1:10 DATD: acrylamide in the original formulation. 3.1. SDS-Polyacrylamide Gels Sufficient medium for one 8 × 10 × 0.75 to 1.0-cm gel can be made by combining stock solutions and various amounts of acrylamide-DATD and water to achieve the required percentage of acrylamide (see Note 1) by using the

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quantities listed in Table 1. The stacking gel is that described by Laemmli (4) and is formed by combining 0.55 mL of acrylamide-bis with 1.25 mL of Tris III, 3.2 mL of water, 0.015 mL of APS, and 0.005 mL of TEMED. 1. For both the running and the stacking gel, degas the solutions by applying a vacuum for approx 30 s before adding the TEMED. 2. After the addition of the TEMED, pour the gels, insert the comb in the case of the stacking gel, and overlay quickly with 0.1% SDS to provide good polymerization. 3. Prepare protein samples by dissolving two parts of the protein sample in one part of sample buffer and heating to 100°C for 2 min. 4. Run gels at 200 V constant voltage for 45 min to 1 h or until the dye front reaches the bottom of the plate.

3.2. Silver Staining The method used is that of Morrissey (1). 1. Remove gels from the plates, and immerse in solution A for 15 min. 2. Transfer to solution B for 15 min and then solution C for 15 min, all while gently shaking. 3. At this point, the gel can be rinsed in glass-distilled water for 2 h to overnight (see Note 2). 4. After the water rinse, add fresh water and enough crystalline dithiotreitol (DTT) to make the solution 5 µg/mL. 5. After 15 min, remove the DTT, and add 0.1% silver nitrate made fresh from the 1% stock solution. Shake for 15 min. 6. Rapidly rinse the gel with a small amount of water followed by two 5–10 mL rinses with developer followed by the remainder of the developer. 7. Watch the gel carefully, and add stop bath to the gel and developer when the desired darkness of the bands is reached. 8. Store the stopped gel in water until the next step.

Table 1 Recipe for Various Percentages of SDS-PAGE Gels Percentage of Acrylamide Stock Solution

7.5%

10%

12.5%

Acrylamide-DATD Tris II Water APS TEMED

0.96 ml 1.50 ml 3.50 ml 0.03 ml 0.003 ml

1.33 ml 1.50 ml 3.13 ml 0.03 ml 0.003 ml

1.66 ml 1.50 ml 2.80 ml 0.03 ml 0.003 ml

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3.3. Slicing and Counting of Gel Slices 1. Remove individual lanes from the gel for slicing by cutting with a knife or spatula. 2. Cut each lane into uniform slices, or cut identified bands in the gel (see Note 3). 3. Place each slice into a glass scintillation vial, and add 0.5 mL of 2% sodium metaperiodate solution. 4. Shake the vials for 30 min to dissolve the gel. 5. Add a 10-mL aliquot of scintillation fluid to the vial, cool the vial, and count in a refrigerated scintillation counter (see Notes 4–6).

4. Notes 4.1. SDS-Polyacrylamide Gels 1. Acrylamide-DATD gels behave quite similarly to acrylamide-bis gels, except for the fact that for a given percentage of acrylamide, the relative mobility of all proteins are reduced in the DATD gel. In other words, a DATD gel runs like a higher percentage acrylamidebis gel. 4.2. Silver Staining 2. The water rinse can be reduced to 1 h, if the water is changed at 10–15 min intervals. The author found in practice that the amount of DTT was not particularly critical, and routinely added the tip of a microspatula of crystals to approx 30 mL of water. 4.3. Slicing and Counting of Gel Slices 3. The method seems relatively insensitive to gel volume, as measured by changes in efficiency (3) over the range of 5–100 mm3 for tritium and an even larger range for 14C or 35S. Larger pieces of gel can be used if one is comparing relative amounts of label, such as in a pulse chase or other timed incorporation, but longer times and more metaperiodate may be required to dissolve the gel. Recovery of label from the gel is in the 80–90% range for proteins from 10–100 kDa (3). In most cases, the label will remain with the protein, but in the case of periodate-sensitive carbohydrates on glycoproteins, it may be released from the protein. This, however, does not prevent the quantification. It just does not allow the solubilized labeled protein to be recovered for other manipulations. 4. It is necessary to cool the vials in the counting chamber before counting in order to eliminate occasional chemiluminescence. The cause of this phenomenon was not explored, but one should determine whether this occurs when using other scintillants or counters. 5. Figure 1 shows the results of a typical experiment using 35S-methionine to label proteins metabolically from the cellular slime mold, Dictyostelium

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purpureum, and quantify them using the method described or by autoradiography. As can be seen, the resolution of the method is comparable to that of the autoradiogram (see ref. 3 for similar results using tritium). The time to process the lane by the method described here was approx 8 h, whereas, the results of the autoradiogram took more than 3 d to obtain. Examination of the stained gel suggests that, if one were interested in a particular protein, it would be fairly easy to isolate the slice of gel containing that protein for quantification. This makes this method extremely useful for comparing incorporation into a single protein over time as in pulse/chase experiments (3). 6. The author has found that as little as 400 dpm of tritium associated with a protein could be detected (3), which makes this method particularly useful for scarce proteins or small samples. References 1. Morrissey, J. H. (1981) Silver stain for proteins in polyacrylamide gels: a modified procedure with enhanced uniform sensitivity. Analyt. Biochem. 117, 307–310. 2. Van Keuren, M. L. Goldman, D., and Merril, C. R. (1981) Detection of radioactively labeled proteins is quenched by silver staining methods: quenching is minimal for 14 C and partially reversible for 3H with a photochemical stain. Analyt. Biochem. 116, 248–255. 3. Springer, W. R. (1991) A method for quantifying radioactivity associated with protein in silver-stained polyacrylamide gels. Analyt. Biochem. 195, 172–176. 4. Laemmli, U. K. (1970) Cleavage of structural proteins during assembly of the head of bacteriophage T4. Nature 227, 680–685.

42 Differential ProteoTope Radioactive Quantification of Protein Abundance Ratios Wojciech Wozny, Gerhard P. Schwall, Chaturvedula S. Sastri, Slobodan Poznanovic¢, Werner Stegmann, Christian Hunzinger†, Karlfried Groebe, and Michael A. Cahill 1. Introduction 2D-PAGE remains a widely employed proteomics method because it provides the highest separating resolution of intact polypeptide isoforms that are amenable to subsequent characterization (1, 2). ProteoTope is a differential radioactive detection method that we developed based upon temporally separated exposures of iodinated proteins on 2D-PAGE gels (3, 4). We have also developed systems for extremely high resolution 2D-PAGE over 54 cm or longer, which were designed for optimal quantification of radioactively labeled proteins (5, 6). These methods provide synergistic improvement in data quality for obtaining quality results from limited samples, such as typical valuable clinical tissue samples (7–9). Here we provide a comprehensive technical description of differential 2D-radioactive ProteoTope quantification using highly standardized samples from the ABRF PRG2006 study (10). Our method is a novel application of the widespread Imaging Plate radioactive detection method (11, 12). The twofold study aims for multiple ABRF study participants were to blindly identify the eight unknown standard proteins in the two sample mixtures, and to determine a best estimate of their relative amounts in each sample by multiple measurements of mixtures of the two samples at a ratio of 1:1 (10). 2D-PAGE and ProteoTope differential radioactive imaging provided excellent relative quantification data of even subtle differences between the two samples. However our inverse symmetrical replicate experimental design and precise relative quantifications revealed that the quantities of proteins in both samples were apparently altered during sample resolubilization in our laboratory. Thus this case study demonstrates

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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both the excellent quantification of limited protein samples achievable by differential radioactive imaging, and the requirement for stringently optimized sample-specific handling protocols which were not developed for this one-off time-restricted experiment. 2. Materials 2.1. Samples, Resolubization, and Labeling 1. The study was performed blind on unknown standard lyophilized samples provided by the ABRF (see Note 1), and as summarized in Table 1. 2. Resolubilization buffer: 2% SDS, 10 mM Tris pH 7.4 (see Note 2).

2.2. Protein Iodination 1. 2. 3. 4. 5. 6. 7. 8. 9.

Chloramine T. 125 I specific activity >600 GBq/mg Iodide, no added carrier. 131 I specific activity ~ 740 GBq/mg Iodide, no added carrier. Zeba Desalting Spin Columns, 500 µL, Pierce. Tributylphosphine (TBP), Fluka. Bromophenol blue (BPB), Sigma. Radioactive counters suitable for measuring 125I & 131I. Lead shielding suitable to protect the operator from radiation. Iodination Buffer: 8 M Urea, 4% CHAPS, 0.1 M Tris pH 7.4 (conveniently stored in 500 µL aliquots at −20°C). 10. Elution Buffer: 8 M urea, 2 M thiourea, 4% CHAPS, 4 µg/µL Tryptone (trypsinized yeast extract proteins). 11. Radiation safety equipment as specified by your authorized radiation safety officer, such as lead blocks with sufficient holes bored to hold all labeling reactions.

2.3. Isoelectric Focussing (IEF) 1. Isoelectric focusing (IEF) buffer: 7 M urea, 1% Triton-X-100, 10% glycerin, 2 M thiourea, 4% CHAPS, 1% DTT, 0,8% IPG-Buffer pH 3–10 (GE Healthcare). 2. Immobiline DryStrips (Immobilized pH gradients: IPGs), pH 3–10 linear gradient (GE Healthcare). 3. 1 % BPB solution in MilliQ water (stored in 100 µL – 200 µl aliquots at −20°C).

2.4. Equilibration 1. Equilibration buffer: 6 M urea, 30% glycerol, 4% SDS, 0.05 M Tris-HCl pH 8.8, 2% iodoacetamide. 4 mM TBP is added immediately prior to use. 2. Disposable rehydration/equilibration tray with Lid, 24 cm, Biorad.

2.5. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. 30% Acrylamide/ 0,8% Bis-acrylamide-stock solution (C. Roth, Catalogue Nr 3029.1). 2. 1.5 M Tris-HCl, pH 8.8. (May be autoclaved for long term storage).

5.0 6.4

6.8 6.6

5.6 5.8

5.6 8.4

23.4 57.5

97.1 28.9

35.6 13.5

66.3 80.4

pI

MW µg

µg

12.9 24.1 83.9

10.6 4

0.3 0.8

12.9 24.1 83.9

10.6 4

22.8 2.6

3.5 3.4

Mixture B

Mixture A

14 17.2

4*

3*

6*

7*

8

195 300

298 296

3 28

598 299

195 300

298 296

235 90

150 59

pmol/vial pmol/vial

1:1 1:1

1:1 1:1

1:76 1:3

4:1 5:1

Mixture B

9 Mixture A

10

Mixture B

11

12.2 18.8 126.1

18.6 18.5

0.2 1.8

37.4 18.7

12.2 18.8 101.4

18.6 18.5

14.7 5.6

9.4 3.7

1.37 2.11

2.10 2.08

0.02 0.20

4.20 2.10

1.37 2.11

2.10 2.08

1.65 0.63

1.05 0.41

pmol/‘5 µg’ pmol/‘5 µg’ pmol/gel pmol/gel

Mixture Mixture Nominal Mixture A B Ratio A

5*

*SSNominal values from the ABRF (10). Columns 1–7 (asterisked) were taken from information supplied by the ABRF after the results were announced (10). Columns 8 and 9 show the nominal amount of each protein present in the 5 µg radiolabeling reactions, and columns 10 and 11 show the nominal amounts of each protein loaded per 2D PAGE gel, assuming 100% yields.

Beta Casein Catalase Glycogen Phosphorylase Carbonic Anhydrase Horseradish Peroxidase Ribonuclease A Bovine Serum Albumin Lactoperoxidase

2*

1*

Table 1 Nominal sample description

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3. 10% sodium dodecyl sulfate (SDS) stock solution in MilliQ water. 4. 10% ammonium persulfate (APS) solution. (Mix fresh each time by weighing crystalline APS and adding 9 times that weight of MilliQ water).

2.6. Gel drying & lamination 1. 2. 3. 4.

4 × vacuum Gel Dryers (Speed Gel SG210D, ThermoSavant). Plastic kitchen foil. 3 mM blotting paper. Laminating Pouch Film large enough for your gels (for ISO-DALT gel we used 216 × 303 mm, 80 microns; Bürobedarf Keller, Marktredwitz, Germany) 5. A laminating device and foils large enough for your gels (Filux LM1250HC DIN A3, Topversand24 GbR, 26135 Oldenburg, Germany) 6. Radiation safety equipment as approved by your radiation safety officer for the process.

2.7. Image Acquisition 1. 20× Fuji Imaging Plates (IPs) for 21–22 h (BAS-MS 2325; raytest GmbH, Straubenhard, Germany). 2. 20 × IP imaging cassettes for IP exposure (raytest GmbH, Straubenhard, Germany). 3. Fuji FLA-3000 R Bioimager and accompanying computer (raytest GmbH, Straubenhard, Germany). 4. IP erasing capacity for 20 IP plates (we use an own-made apparatus). 5. Suction device to remove IPs from the cassettes (optional) (raytest GmbH, Straubenhard, Germany). 6. Copper and lead lined cabinet (safe) to expose the IPs (optional) (raytest GmbH, Straubenhard, Germany).

2.8. Image Deconvolution 1. A computer program capable of image deconvolution. We use the own-developed “select_deconv_v1.1.pl” program (ProteoSys AG)

3. Methods 3.1. Sample Resolubilization and Protein Labeling 1. The desiccated samples were dissolved directly into 40 µL of 10 mM Tris, 2% SDS, pH7.4 with 5 min shaking at 95°C (see Note 2). Protein content was assumed to be 80 µg/40 µl based upon the ABRF sample description.

3.2. Protein iodination 1. Into eight reaction tubes place 60 µL Iodination Buffer. 2. Design your own experimental replicates as appropriate. For the described ABRF study experiment: Transfer 2.5 µL (5 µg) of the redissolved sample A into four of the reaction tubes, and transfer 2.5 µL of sample B into the other four reaction tubes. This establishes inverse duplicates samples for eight iodination reactions:

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125

3.

4. 5. 6. 7. 8. 9. 10. 11.

12. 13. 14.

15. 16. 17. 18. 19.

I-A1, 131I-A1, 125I-A2, 131I-A2, 125I-B1, 131I-B1, 125I-B2 and 131I-B2 (see Notes 3 and 4). Transfer 3,7 MBq of 125I to each tube of sample A or B that is to be labeled with 125I from the previous step (125I-A1, 125I-A2, 125I-B1, and 125I-B2). Then transfer 3,7 MBq of 131I to the other tubes (131I-A1, 131I-A2, 131I-B1, and 131I-B2). (see Note 5). Start the reaction by adding 3.5 µL of 100 mM Chloramine T (20X concentrated stock solutions: reaction conditions 5 mM). Reaction time is 5 min at RT. The reaction is stopped by adding 2 µL of 1 M TBP in 2-propanol (see Note 6). The labeled proteins are purified by Spin columns (see Note 3). Remove the bottom seal of the column and loosen (do not remove) the cap. Place the column in a 1.5–2.0 mL microcentrifuge collection tube. Centrifuge at 1,500 × g for 1 min to remove storage solution. Place a mark on the side of the column where the compacted resin is slanted upward. Place the column in the microcentrifuge with the mark facing outward in all subsequent centrifugation steps. Add 300 µL of elution buffer on top of the resin bed. Centrifuge at 1,500 × g for 1 min to remove buffer. Repeat the previous step three additional times, discarding buffer from the collection tube. Place the column in a new collection tube, remove the cap, and apply the sample to the top of the compacted resin bed. Load sample directly to the centre of the resin bed. Carefully touch the pipette tip to the resin to expel all sample. Centrifuge at 1,500 × g for 2 min to collect the sample. Discard the desalting column after use. Measure radioactivity of the resulting protein solutions. Calculate the amount of iodine incorporated into proteins (see Notes 7 and 8) to assess the labeling yield (see Note 9). All procedures should be approved by your radiation safety officer.

3.3. Isoelectric Focusing (IEF) The methods of 2D-PAGE are so well established that these are described only in general reference to the steps employed. It is assumed that each laboratory has existing 2D-PAGE methods and equipment available and operating. For general methodical 2D-PAGE references see (13–16) or reference (17) from the Methods in Molecular Biology series. Samples were analyzed by 2D-PAGE according to the regime of Fig. 1A (see Note 4). 1. Prepare 20 Eppendorf tubes and 20 loading mixtures as follow: 2. Add 200 µL IEF buffer to each vessel. 3. Add 90 kBq of the appropriate 125I-labelled protein and 131I-labelled to each well according to the scheme of Table 2. Adjust the volume with IEF buffer up to 350 µL. Add 2.8 µL of IPG buffer pH 3–10 linear. Be sure to use a new disposable tip each time.

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Iodination replicate 2

Iodination replicate 1

Samples

Sample A1

Sample B1

Sample A2

Sample B2

Mixture (I-125 & I-131) 125-A

2D-PAGE

131-A 131-B 125-B

125-A 131-A 131-B 125-B

ProteoTope Imaging

Gel-1 125-A 131-B

Image-1 125-A

Image-2 131-B

Gel-2 125-B 131-A

Image-3 131-A

Image-4 125-B

Gel-3 125-A 131-B

Image-5 125-A

Image-6 131-B

Gel-4 125-B 131-A

Image-7 131-A

Image 8 125-B

Performed in quadruplicate 16 x 2D-PAGE gels

B % Activity of t0

100

Integral 1

50

Integral 2

t0

t 1 t2

t3 t4

Time

Fig. 1. Schematic depiction of ProteoTope analysis in this study. (A) Experimental ProteoTope Inverse Replicate and Tracer Gel Design employed for quantitative analytical multiplex. In this study the design for boxed gels 1–4 was performed in quadruplicate to generate 16 × 2D-PAGE gels, pH 3–10 linear. Each sample is separately iodinated in duplicate with each of two radioactive iodine isotopes: 125I (dark) and 131I (light). The radioactively labeled proteins were mixed together and coelectrophoresed on inversely labeled replicate 2D-PAGE gels as shown. The individual signals from each radioisotope were measured by ProteoTope and mutually calibrated to give equal BSA intensities in each channel of a respective gel. Double headed arrows represent multiplex image generation. The experimental design provides 16 independent estimates of the abundance ratio of proteins in particular spots between the samples, and includes all variables introduced by sample handling and electrophoretic conditions. (B) A schematic depiction of the integrals between t1 and t2, and t3 and t4 (being time intervals after t0), used to calculate decay factors that are provided in Table 2.

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4. Prepare 20 IEF Rehydration tubes. The tubes should be made of 5 mL disposable pipettes (606107, Greiner Bio-One GmbH). Cut each pipette at the ends to obtain 22 cm long tubes. 5. Place into each tube Immobiline dry strip 18 cm, pH 3–10, gel side down. 6. Put the cup (Caps, 12504, Moss Plastic Parts) at one end of each tube. Label the tubes as Loading mixture. 7. Apply the Loading mixtures into appropriate strip. Carefully pipette the mixture between the gel side of the strip and the wall of the tube. Avoid introducing bubbles (see Fig. 2). Close the tube with the cap. 8. Incubate horizontally (gel side down) overnight at RT. 9. Rinse the rehydrated IPG strips with water, blot lightly onto filter paper soaked with water and place them gel side up in the strip aligner tray of a Multiphor IEF device. 10. Cut paper electrode strips (ca 15–20 mm wide and approx. 130 mm long for the IPG-Phor from Macherey-Nagel Filterpaper MN 440. Soak the paper strips with water and blot excess water with clean blotting paper, then place the wet paper electrode strips over the ends of IPG strips so that the electrode strip overlaps onto about 5 mm of the IPG gel. 11. Install electrodes so that they press down roughly in the middle of the paper electrode strips. 12. Switch on the water cooling system to maintain constant temperature at 25°C. 13. With the power supply limited to 2 mA and 5 W, electrophorese the samples for 1 h at 150 V, 1 h at 300 V, 0.5 h at 600 V, 0.5 h 1500 V, 0.5 h at 2000 V, 1 h at 3500 V, 10 h at 6000 V (see Note 5). 14. All procedures should be approved by your radiation safety officer.

3.4. Equilibration 1. After the completion of IEF, incubate each individual IPG for 30 min, in 5 mL Equilibration Buffer, while gently shaking. Use disposable equilibration trays (see Note 3). 2. All procedures should be approved by your radiation safety officer.

Fig. 2. Loading of rehydration mixture.

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3.5. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. 20 × 12% acrylamide SDS gels were electrophoresed in one ISO-DALT chamber, loading two gels per ISO-DALT slot, for 3 h at 50 V, and 110 V overnight with temperature regulated to 20°C. 2. All procedures should be approved by your radiation safety officer.

3.6. Gel Drying and Llamination 1. Separate the 2D-PAGE gels from one of the glass plates, systematically removing the glass plate from the same side of all gels. The aim is to process all gels in the same orientation so that all resulting gel images will have the same orientation. If some gels are processed in the wrong orientation the images can still be later manually rotated or mirrored by computer. 2. Place a piece of blotting paper on top of the gel, conveniently precut to about 23 × 30 cm ( see Notes 3 and 10). 3. Invert the glass plate and paper, so that the paper is underneath the glass and gel, then gently remove the glass plate. 4. Place plastic kitchen foil over the gel, fully covering all potential sources of radioactivity. 5. Place the gel, paper down and plastic up, on top of two sheets of blotting paper on a gel dryer. (Up to four ISO-DALT gels can fit onto one gel dryer.) 6. Dry the gels at 80°C under vacuum for at least 45 min up to 90 min (see Note 11) on the Savant gel dryer (see Note 5). 7. Place the gels, dried onto one sheet of paper, into 0.08 mm plastic protective layer laminating foils, paying careful attention that the orientation of all gels is the same. Obviously, avoid contaminating the outside of the plastic! These will later be in direct contact with the IPs and you cannot afford a contamination of the IPs with 125 I, which has a half-life of 60 days. 8. Seal the gel in the laminating foil using the lamination device. The seal should ideally be perfect and airtight. At this stage the gels are ready for imaging. 9. All procedures should be approved by your radiation safety officer.

3.7. Image Acquisition 1. Erase 20 IP plates (see Note 12). 2. Place the gels into an IP exposure cassette, paper side down and gel side up, once more paying attention that the orientation of all gels is identical (see Note 13). 3. Gently place a freshly pre-erased IP into the cassette, white/phosphor side down and contacting the upturned gel surface of the laminated gel, also paying attention that the edges are flush with the edges of the exposure cassette in one corner. Be careful not to bend the IP or touch the white surface with your fingers (see Note 12). 4. Close the cassette, noting the exact time for each different IP to give the individual t1 values analogous to those from Table 2 (see Note 14). 5. Expose for 21–22 h (see Note 15).

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6. After the exposure is complete, open the IP exposure cassette under conditions of low light (see Note 16). Record the exact time the exposure is terminated to give t2 from Table 2. 7. Carefully remove the exposed IP from the cassette containing the laminated gel and transfer the IP to the imaging device for scanning (see Note 17). Pay attention to the consistent and exact orientation of all IPs with respect to the IP stage (see Note 18). 8. Place the IP stage with IP into the Imager, which has been switched on for sometime beforehand to warm up the lasers (see Note 19). 9. Scan on a Fuji FLA-3000 R Bioimager at 0.2 mm resolution using the IP filter selection and grid setting height = 1125, width = 1250 of the BASReader software (Version 3.01) (see Note 20). 10. When the images are acquired the IP may be removed. Remove the IP stage from the imager and take the IP off it by sticking your gloved finger through the hole in the stage taking care once more not to touch the white surface with your bare fingers or to bend the IP. 11. The IP may then be erased. 12. Acquisition of the first scan images is complete. 13. After waiting at least two weeks (see Note 21) to permit differential decay of the 125 I and 131I (half lives of 59.41 days for 125I and 8.04 days for 131I) on the labeled proteins, repeat this entire exposure and imaging process to generate the second scan images (see Note 22). Remember to record t3 (commencement of the second exposure) and t4 (termination of the second exposure) from Table 2 for each individual exposure.

3.8. Image deconvolution 1. Expositions in this study were commenced on 10 November 2005 and 1 December 2005 according to the regime of Table 2. 2. The signal distribution of 125I and 131I on the gel images is deconvoluted by solving simultaneous equations with a computer algorithm to generate images of the two individual isotopes on each gel (see Notes 23 and 24). An example set of images for gel 1 from Table 2 is depicted in Fig. 3.

3.9. Image processing and spot detection Quantitative image processing was performed with the PIC/GREG software package (Fraunhofer-Institut für Angewandte Informationstechnik, Sankt Augustin, see http://www.fit.fraunhofer.de/projekte/greg/index_en.xml). There are multiple commercially available software products suitable for 2D-PAGE image processing. The signal intensities of deconvoluted images for each isotopic channel were calibrated to produce a best-fit 1:1 ratio for albumin spots for each gel according to sample information provided by the ABRF (10). Therefore final abundance ratio estimates were relative to the assumed albumin ratio. (Note that this is not our standard normalization method. We usually normalize

B1 B1 A1 A1 B2 B2 A2 A2 B1 B1 A1 A1 B2 B2 A2 A2 A1 A2 B1 B2

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

A1 A1 B1 B1 A2 A2 B2 B2 A1 A1 B1 B1 A2 A2 B2 B2 A1 A2 B1 B2

I-131

Gel-No. I-125

cold protein 4.90 4.90 4.90 4.90 4.90 4.90 4.90 4.90 4.90 4.90 4.98 4.98 4.98 4.98 4.98 4.98 4.98 4.98 4.98 4.98

t1a 5.75 5.76 5.77 5.78 5.78 5.80 5.82 5.86 5.87 5.88 5.89 5.90 5.92 5.93 5.94 5.95 5.96 5.97 5.97 5.98

t2 25.82 25.82 25.82 25.82 25.82 25.82 25.82 25.82 25.82 25.82 28.88 28.88 28.88 28.88 28.88 28.88 28.88 28.88 28.88 28.88

t3 26.75 26.76 26.77 26.78 26.78 26.79 26.80 26.81 26.81 26.82 29.77 29.78 29.79 29.80 29.80 29.81 29.82 29.83 29.85 29.85

t4

I-131

0.8038 0.8141 0.8209 0.8271 0.8333 0.8460 0.8684 0.9089 0.9155 0.9228 0.8467 0.8579 0.8731 0.8838 0.8947 0.9014 0.9147 0.9215 0.9280 0.9344

0.6847 0.6907 0.6968 0.7038 0.7086 0.7136 0.7189 0.7238 0.7286 0.7336 0.6363 0.6433 0.6484 0.6534 0.6583 0.6634 0.6692 0.6739 0.6880 0.6931

1.174 1.179 1.178 1.175 1.176 1.186 1.208 1.256 1.257 1.258 1.331 1.334 1.346 1.352 1.359 1.359 1.367 1.367 1.349 1.348

0.5406 0.5473 0.5518 0.5558 0.5599 0.5681 0.5826 0.6088 0.6131 0.6178 0.5648 0.5720 0.5817 0.5886 0.5957 0.5999 0.6085 0.6129 0.6170 0.6211

0.0965 0.0973 0.0982 0.0991 0.0998 0.1004 0.1012 0.1018 0.1025 0.1032 0.0715 0.0723 0.0728 0.0734 0.0739 0.0745 0.0751 0.0756 0.0771 0.0777

5.601 5.624 5.621 5.608 5.612 5.656 5.759 5.979 5.983 5.989 7.897 7.914 7.987 8.022 8.060 8.058 8.104 8.107 8.001 7.997

Integral Integral Integral Integral 1 2 DF 125 1 2 DF 131

I-125

Table 2 2D-PAGE gels performed in this study, and deconvolution parameters

458 Wozny et al.

A2 B1 B1 A1 A1 B2 B2 A2 A2 A1

B2 A1 A1 B1 B1 A2 A2 B2 B2 A1

8 9 10 11 12 13 14 15 16 17

B1 B1 A1 A1 B2 B2

I-131

A1 A1 B1 B1 A2 A2

Gel-No. I-125

1 2 3 4 5 6

I-131

Text tabular version

A1 A1 B1 B1

Gel-No. I-125

21 22 23 24

cold protein

cold protein

A1 A1 B1 B1

4,90 4,90 4,90 4,98 4,98 4,98 4,98 4,98 4,98 4,98

t1

a

4,90 4,90 4,90 4,90 4,90 4,90

t1a

5,86 5,87 5,88 5,89 5,90 5,92 5,93 5,94 5,95 5,96

t2

5,75 5,76 5,77 5,78 5,78 5,80

t2

25,82 25,82 25,82 28,88 28,88 28,88 28,88 28,88 28,88 28,88

t3

25,82 25,82 25,82 25,82 25,82 25,82

t3

26,81 26,81 26,82 29,77 29,78 29,79 29,80 29,80 29,81 29,82

t4

26,75 26,76 26,77 26,78 26,78 26,79

t4

0,9089 0,9155 0,9228 0,8467 0,8579 0,8731 0,8838 0,8947 0,9014 0,9147

Integral 1

I-125

0,8038 0,8141 0,8209 0,8271 0,8333 0,8460

Integral 1

0,7238 0,7286 0,7336 0,6363 0,6433 0,6484 0,6534 0,6583 0,6634 0,6692

Integral 2

0,6847 0,6907 0,6968 0,7038 0,7086 0,7136

Integral 2

1,256 1,257 1,258 1,331 1,334 1,346 1,352 1,359 1,359 1,367

DF 125

1,174 1,179 1,178 1,175 1,176 1,186

DF 125

0,6088 0,6131 0,6178 0,5648 0,5720 0,5817 0,5886 0,5957 0,5999 0,6085

Integral 1

I-131

0,5406 0,5473 0,5518 0,5558 0,5599 0,5681

Integral 1

5,601 5,624 5,621 5,608 5,612 5,656

DF 131

0,1018 0,1025 0,1032 0,0715 0,0723 0,0728 0,0734 0,0739 0,0745 0,0751

(continued)

5,979 5,983 5,989 7,897 7,914 7,987 8,022 8,060 8,058 8,104

Integral DF 131 2

0,0965 0,0973 0,0982 0,0991 0,0998 0,1004

Integral 2

Differential ProteoTope Radioactive Quantification 459

A2 B1 B2

18 19 20 21 22 23 24

A1 A1 B1 B1

cold protein

4,98 4,98 4,98

t1

a

5,97 5,97 5,98

t2 28,88 28,88 28,88

t3 29,83 29,85 29,85

t4

Integral Integral DF 125 1 2 0,9215 0,6739 1,367 0,9280 0,6880 1,349 0,9344 0,6931 1,348

I-125 Integral Integral 1 2 0,6129 0,0756 0,6170 0,0771 0,6211 0,0777

I-131

8,107 8,001 7,997

DF 131

a The actual time point of t1 for Gel no. 1 was 10 Nov. 2005, 14:16 (hh.mm). Samples A1, A2, B1, and B2 correspond to replicates of Sample A and Sample B. Gels 1–16 were used to quantify the differential abundances of proteins between samples A and B. Gels 17–20 were labeling controls, and gels 21–24 were preparative gels, loaded with 10 µg of the indicated non-radioactively labeled proteins (cold protein) per gel to be used for mass spectrometric protein identification (not described here). t1 and t2 are given in days after the calibration date of 125I. These represent the start and finish of the first exposure, and are used to calculate Integral 1 for each isotope 125I and 131I, according to the theoretical fraction of radioactive decay between t1 and t2 and as schematically depicted in Fig. 1B. Integral 2 is similarly calculated from t3 and t4. Integrals 1 and 2 are then used to calculate the decay factors (DF) for deconvolution for each isotope as indicated. The decay factors shown correspond to 1/M and 1/N from equation 2 (see Note 23).

A2 B1 B2 A1 A1 B1 B1

I-131

Gel-No. I-125

Table 2 (continued)

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Fig. 3. Deconvolution of gel 1 from Table 2. (A) The signal obtained from the initial exposure, made for 21 h starting on day 1. (B) The signal obtained from the second exposure, made for 22 h starting on day 21. (C) The deconvoluted signal from the 125I-labeled sample. (D) The deconvoluted signal from the 131I-labeled sample. Deconvolution was performed according to Eqs. 1 and 2 (see Note 23) using parameters shown in Table 2. Our deconvolution process produces non-linearly scaled TIFF images from the floating point FLA3000 data files.

the total signal volumes for both isotopic channels to each other.) The composite average gel images for sample A and sample B from all 16 gels are shown in Fig. 4, as are data depicting the reproducibility of data acquisition for the replicates of different 2D spots for the test proteins. 3.10. Statistical analysis Quantitative values from PIC/GREG were exported in a spreadsheet and statistics were performed using Excel (Microsoft). The mean values and standard errors are modeled from the log2(Isample2/Isample1) intensity ratio values, taking advantage of the gel replicates. P-values are calculated using a t-test based on the null hypothesis of identical populations unless otherwise specified. The overall result for all spots used for quantitative analysis is shown in Fig. 6,

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A

GP LP

BSA

C P

BC

CA1

R

B

log2 (I1/I2) - log2 (I1/I2)SpotAvg

BA_2369 BA_2292

3

glycogen phosphorylase b

2

lactoperoxidase albumin

1

catalase

0

horse radish peroxidase C1

-1

beta casein CA1 protein

-2

ribonuclease a

-3

3

4

5

6

7

8

9

10

11

12

13

14

15

16

17

log2(min(I1,I2))

C A/B Ratio/Overall Mean Ratio (n=16)

1.4 mean 16 rplcts. +/- SEM

1.2 1

mean 8 rplcts. +/- SEM

0.8

mean 4 rplcts. +/- SEM

0.6

Replicates:

16

8

4

Spot BA_2292

16

8

4

Spot BA_2369

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and the comparison of ProteoTope-estimated abundance ratios with those given by the ABRF as nominally correct is shown in Fig. 6 (see Notes 25 and 26). 4. Notes 1. A technical description of the samples has been published (10). The samples (one vial of Sample A, one vial of Sample B) from the ABRF arrived in our premises on 31 October 2005. The deadline for submitting results was no later than 9 December 2005. ABRF-PRG2006 was composed of two mixtures, labeled A and B, that contained eight major unknown proteins each of various unknown amounts according to the sample description. The total amount of unknown protein in each sample vial was given as approximately 80 µg. Individual proteins were present on average at Fig. 4. Master gel and spot quantification. (A) A screen shot showing the spot detection made with the PIC/GREG software using the 32 images from all 16 measured 2D gels. The upper left panel shows a master synthetic differential image, with the positions of spots used to estimate the abundance ratios of the eight test proteins in Fig. 5. The proteins identified (with SwissProt Accession and number of spots quantified per protein in this study) in order of increasing SDS-PAGE mobility were: GP = Glycogen Phosphorylase B (P00489, n = 5); LP = Lactoperoxidase (P80025, n = 1); BSA = Bovine Serum Albumin (P02769, n = 8); C = Catalase (P00432, n = 8); P = Horse Radish Peroxidase (P00433, n = 4); BC = Beta Casein (P02666, n = 4); CA1 = Carbonic Anhydrase I (P00915, n = 4); R = Ribonulclease A (P61823, n = 4). Spots were selected for quantification only after unambiguous mass spectrometric identification. All identifications matched the identities supplied by the ABRF (10). BA_2369 and BA_2292 are the spots in C below. The lower two panels show the spot segmentation performed by PIC/GREG on the averaged gel images for Sample A (lower left panel) and Sample B (lower right panel). These averaged images are synthetically constructed using the 16 warped images generated with both isotopes of iodine for each sample. The upper right panel shows a user interface containing spot parameters for the selected most acidic BSA spot. (B) Spot intensity ratios over minimum spot intensity. Logarithms to base 2 of the ratios of spot intensities under conditions A over B minus log2 of the average intensity ratio of the respective spot log2(I1/I2) - log2(I1/I2)SpotAvg are displayed for each gel replicate. Data points are plotted as a function of the log2 of the minimum spot intensity for either Sample A or B log2(min(I1,I2) ). Symbols indicate the proteins that were identified in the respective spots. The diagram demonstrates that the measurement error in replicate spot quantifications becomes smaller as signal intensities increase. (C) Dependency of error on replicate number. The estimated mean spot volumes (with standard errors of means) for the most intense (Spot BA_2292) and least intense (Spot BA_2369) spots for Ribonuclease A are plotted using estimates made from four sets of n = 4, two sets of n = 8, or one set of n = 16 replicates (rplcts), plotted as fold overall mean ratio for the n = 16 set (A/B Ratio/Overall Mean Ratio (n = 16) ). All estimated mean abundance ratios for both spots were within 20% of the mean obtained for n=16 gels. Gel grouping was numerical from Table 2 (i.e., 1–4, 5–8, 9–12, 13–16, 1–8, 9–16, 1–16).

464

2.

3. 4.

5.

6.

7.

Wozny et al. 300 pmol, ranging from approximately 3 to 600 pmol, with ratios of proteins between mixtures A and B that could vary by up to 1:100. Some of the information as presented by the ABRF PRG2006 committee after completion of the study on the abundance of sample proteins is summarized in columns 1–7 of Table 1. The overall comparative results of the study for all participants using all experimental methods have been published, where ProteoTope is referred to as “2D radioactivity” (10). We were the only participant in this study to resolubilize the desiccated protein pellet in an SDS-buffer. This was a one-off real life experiment without method optimization. This method apparently did not resolubilize all desiccated proteins with equal efficiency. We would recommend lysing frozen cryogenically sliced tissue samples into a boiling SDS or a urea-based buffer. For desiccated protein samples a urea buffer is apparently more successful. Be sure to label all reaction vessels and other objects beforehand to avoid unnecessary exposure to radioactivity. The inverse replicate experimental design described here systematically controls for errors that may be introduced by radiolabeling or electrophoretic laboratory steps. Proteins were prepared for 2D-PAGE as described (8, 10–12) without cysteine alkylation prior to radiolabeling due to schedule constraint and the associated inability to optimize conditions. 1 mm lead shielding can be placed over apparatus to avoid exposure from 125 I, however note that gamma radiation from 131I is not effectively impeded by such thin lead sheeting, and so the best protection from radiation exposure is substantial shielding such as a lead brick wall, preferably combined with sufficient physical distance (e.g., leave the room when you are not working). Your radiation safety officer should approve the protocol that you implement. The concentrated form of TBP is explosive in the presence of oxygen, however a 200 mm stock solution is not explosive and can be stored at 4 degrees under inert gas. The stock solution is prepared by diluting the concentrated form (3.93 M) in solvent under an inert gas. TBP also is volatile, toxic, and unpleasant smelling (18), and should be handled in a fume hood. Briefly, iodination reactions with either 125I or 131I were conducted under identical chemical iodine concentrations, using 3.7 MBq of each isotope (125I specific activity >600 GBq/mg Iodide, no added carrier; 131I specific activity ~ 740 GBq/mg Iodide) per 5 µg sample aliquot in a reaction volume of 60 µL by the chloramine T method as described (19), taking all appropriate radiation safety precautions. Iodine incorporation into proteins was determined to be approximately 800 kBq per labeling reaction, with the yield of iodine incorporated into proteins ranging from 18.6% to 21.1%.

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Table 3 Theoretical level of labeling of an ‘ideal average’ 40 kDa protein and of average tyrosine residues at maximum theoretical specific activity of each isotope

I-125 I-131

1 fmol/kBq

2 fmol/800 kBq

3 mol 40 kDa/mol Iodine

4 mol Tyr/mol Iodine

12,39 1,67

9912 1336

13 94

151 1123

Column 1 shows the approximate maximum theoretical specific radioactivity per Becquerel of the iodine isotopes used to label proteins, and column two shows the corresponding approximate number of femtomoles of radioactive iodine atoms incorporated into proteins in ideal 5 µg radiolabeling reactions with yield of 800 kBq. Column 3 shows the ratio moles of an “ideal average” 40 kDa protein per mole of incorporated iodine. Column 4 shows the number moles of tyrosine per radioiodinated tyrosine residue in the “ideal sample” (assuming an “average amino acid residue of 110 Da” and a tyrosine amino acid frequency of 0.033 for the “ideal average” 40 kDa protein). In our case, the specific activities of the 125I and 131I isotopes employed were approximately equal, indicating that 131I had decayed through between two and three half-lives at the time of calibration (16% max. theoretical specific activity: a value which decreases by 50% each 8 days), and therefore the chemical labeling stoichiometry achieved for 131I in this experiment was within approximately 23% of that given for 125I above (i.e. 186 mol Tyr/mole iodine corresponding to column 4). Similarly, the specific activity of the 125I employed was within 7% of its maximum theoretical value, and so the labeling stoichiometry of 125I should have been within 7% of that given for 125 I in the table (i.e. >140 mol Tyr/mole iodine).

8. To elaborate upon the degree of labeling achieved in this experiment, it may be useful to consider the state of labeling of a population of “ideal average 40 kDa proteins” of random sequence with iodine isotopes of maximum theoretical specific activity as depicted in Table 3. Each 5 µg labeling reaction would contain 125 pmol of such “average 40 kDa proteins,” which accords well with the nominal sum tallies of 126 pmol and 101 pmol for the experimental sample proteins of columns 8 and 9 from Table 1. Assuming that all the iodine present in the yield of radioactive protein was present as labeled tyrosine (an approximately correct yet ideal assumption which ignores nonspecific side reactions and noncovalent affinity of iodine to the protein mixture), approximately every thirteenth “ideal 40 kDa protein molecule” would be iodinated with 125I and each ninetieth 40 kDa protein would be iodinated with 131I under these reaction conditions. This would correspond to approximately every one hundred and fiftieth tyrosine in the mixture being labeled with 125I or less than every thousandth tyrosine being labeled with 131I. Our actual chemical labeling conditions were similar for both isotopes because of

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relatively higher proportion of 127I (nonradioactive stable isotope) in the 131I preparation, as described in the legend to Table 3. Although these values are more qualitative than quantitative, they illustrate that the labeling conditions employed are conservative in order to reduce the possibility of generating double labeled proteins and corresponding protein chains. 9. As is evident from Table 3 and Note 8, we labeled only a small fraction of the protein molecules present. This is performed to provide a margin of safety for the labeling conditions. To understand this, it is necessary to consider the production of the isotopes in some detail. The iodine is usually produced by irradiating a target of Te in a cyclotron. For instance, in one method to produce 125I an isotopically enriched target of 125Te would be irradiated (125Te(p,n)125I). However since the target also contains an often undetermined and batch-specific amount of 128Te impurity, traces of (nonradioactive) 127I are produced simultaneously (128Te(p,2n)127I). Furthermore, after irradiation the iodine is washed from the surface of the blank using a solution which inevitably contains some iodine. While the specific activity of commercial iodine products is provided as “carrier free,” it is technically impossible to produce an isotopically pure preparation of the radioisotope, and “carrier free” means that no free iodine was deliberately added to the preparation in addition to these necessary and unknown sources, although a multiple molar excess of nonradioactive iodine may be present (this phenomenon is not restricted to iodine isotopes). For instance, whereas the maximum theoretical specific activity of 125I is 642.7 GBq/mg iodide, the product we purchased was rated as >600 GBq/mg. The corresponding values for 131I are 4587 GBq/mg (max. theor.) and 740 GBq/mg (product specification). For a technical description of the processes involved, see Bonardi et al. (20). For an estimate of the labeling stoichiometries obtained by us, see the legend to Table 3. For most researchers there is no practical way to quantify the isotopic purity of the small highly radioactive mixture that is commercially delivered. When dealing with very small amounts of protein, such as from clinical microdissection samples (7), we sometimes experienced smearing (not chain formation) in the basic region of 2D-PAGE patterns due to over-iodination caused by cold iodine in the “nocarrier-added” commercial stocks. Because these samples were extremely valuable, this led us to explore the strategy of sample pooling to generate a critical mass of protein in the labeling reactions, which produced more consistently reliable results over replicate samples (7). On the other hand, when sufficient sample is available the labeling conditions can be calibrated for particular batches of iodine and protein to achieve very much higher incorporated specific activity and thereby sensitivity than attained here. In hindsight we could have systematically iodinated the samples in

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10.

11.

12. 13.

14.

467

parallel with tenfold more specifically active iodine in the labeling reactions, which would probably have greatly improved the quantification of glycogen phosphorylase, whose signal was close to background levels in our Sample A gels and which consequently deconvoluted with errors only slightly better than background variation. While the indicative calculations of Table 3 assume that each tyrosine reacts identically, in practice the electron density contributed to the tyrosine ring by adjacent amino acids may produce sequence-specific differences in reactivities of perhaps up to 100fold or more in extreme cases (assuming a range of pka values of 2 units for protein tyrosine residues). These would nevertheless only contribute to overlabeling when two or more hyperreactive residues were present in one analyte protein molecule. It is recommended to place 1 µL of a diluted labeled protein from each reaction mixture used on that gel onto the border of the blotting paper adjacent to the gel but within the area that will be imaged. The amount of radioactivity required in this signal source will vary depending upon the sensitivity of your imager. It should not saturate the pixel distribution of the imager during the exposure period you choose, yet is useful when >50% saturation is achieved: 60 Bq is a useful guide value. Also place 1 µL of a 1:1 mixture of both labeled proteins (i.e. 30 Bq of protein mixture labeled with each isotope in 1 µL). These optional signals provide useful controls for the deconvolution process later. Unless the gels are perfectly dry they risk shattering into many pieces when the vacuum is removed, or slowly cracking after lamination. When the latter occurs between the first and second imaging scans the images are not superposable, and cannot be deconvoluted! Although these effects can affect every experiment, the consequences are especially severe when working with a large number of gels in a matrix-designed radioactive experiment. As a rule of thumb, the gels are dry when there is no temperature difference apparent between the metal surface of the gel dryer and the drying gel as judged by (disposable gloved) hand contact. DO NOT BEND IPs. The phosphor crystals of the IP are damaged when the IP bends, resulting in less sensitive regions. Because the gels will be exposed twice, and both images must be processed in parallel to generate the individual isotope images, pay attention that the corner of the laminated gel is pressed so that the edges are flush with the edges of the exposure cassette at one particular corner for both exposures. It is highly advisable to expose the same gel with the same IP in the exact same orientation for both scans in your experiment. In case the surface of the IP has been damaged and exhibits differing sensitivities, this can be largely eliminated by imaging the same gel region with the same IP surface in both

468

15.

16.

17. 18.

19.

20.

21.

Wozny et al. scans. Each IP has one corner cut away to help with systematic orientation of the IP with respect to the gel. Exposure in a lead- and copper-lined metal exposure cabinet (“safe”) significantly decreases background signals from environmental radiation, permitting the more accurate detection of low intensity signals from the gels. Our version was purchased from raytest GmbH, Straubenhard. You should have the imaging device in a room with low light level, such as drawn blinds or a room without windows. (You will require a few candle power of light to see what you are doing though, such as perhaps the light of the monitor of your PC that runs the FLA3000.) The signal on the IPs is extremely sensitive to light. Only remove the exposed IP from the cassette with the room lights off. The FLA3000 should be able to operate in light while scanning. However we have very occasionally had problems that the sealings of the scanner were not tight and some measurements were interrupted, destroying that exposure and potentially costing a repeat of the whole experiment. So we advise to also keep the room lights off while the imager is scanning. To avoid bending the IP while removing it, a suction device may be useful to lift the plate the first few millimeters (Fig. 7). For the FLA3000 the IP has a magnetic backing and is placed into an IP stage for scanning. The IP stage is marked with a grid of letters and numbers. To assist with image orientation, we suggest that the cut corner of the IP should go towards the A9 corner as shown (Fig. 8). Make sure that the IP sits squarely in the stage, and that the edges are as close as possible to both sides. Make sure that the IP stage is sitting exactly in the FLA3000. If it is crooked it can destroy the motor of the FLA3000 and/or the IP stage if the stage does not sit flatly as the FLA3000 is trying to move it into the scanning area. Make sure the arrow between “H” & “I” on the IP Stage lines up with the arrow on the FLA 3000 (Fig. 9). It may be advisable to make two or more successive measurements without removing the IP stage and IP from the imager if some regions of the gel exhibit saturated pixel intensity. You will need to establish an appropriate systematics of data structure and file nomenclature. We employed only a three week decay period here because of study schedule restraints for the ABRF PRG2006 study. The ability to accurately deconvolute signals containing low intensities of 125I will be increased by permitting longer decay times, whereby the actual number of randomly decayed 125I atoms is given time to approach the theoretically calculated expected number more precisely. Three to five weeks decay period is a practical compromise and should be empirically determined by each laboratory for itself.

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22. ProteoTope experiments resemble somewhat the activity of the warrior, with shorter periods of activity interspersed by longer periods of waiting (3–5 weeks) for radioactive decay. However different temporally overlapping experiments can be sequentially coordinated such that the overall throughput is continuous and considerable, and has proved to be adequate for our industrial applications. Although iodine is not a natural component of most proteins, it can be efficiently introduced into proteins from any source by the post harvest labeling method (19), including clinical human samples. Because the differential images produced for ProteoTope rely on the different decay rates for each isotope (half lives of 59.41 days for 125 I and 8.04 days for 131I), the same Imaging Plate can be used for both measurements without the requirement of an absorber as required by the method of Johnstone et al. (21), and thus there is no source of distortion of the spatial signal distribution or detection efficiency. The vast majority of signal from 125I captured by the Imaging Plate is from the 27 keV and 31 keV photons. The Auger electron is essentially too weak to be detected. For 131I the 364 keV photon does not efficiently interact with the plate crystals, so that the vast majority of signal from this isotope comes from the 810 keV beta particle. Because this beta particle interacts more efficiently with the phosphor crystals than the low energy photons from 125 I, the signal detected from 131I is slightly more localized than the more diffuse signal for 125I. However both are favorably comparable in resolution, as evident from our results in this and other cited studies. The precise composition of our imaging plates remains unpublished by the manufacturer, and there is no data available from the manufacturer or distributor on the detection efficiency of the Imaging Plates for the various types of radiation we used. Therefore the discussion of detection efficiencies in this note represents the outcome of empirical observation and deduction. We do not employ ProteoTope to obtain absolute quantitative values, but rather to obtain accurate estimates of differential protein abundances between two samples. There may be some concern that the slightly more diffuse signal pattern generated by 125I may lead to problems in differential quantification between isotope channels. We neglect this since the overwhelming majority of the signal is localized close to the source for both isotopes, and because the signal intensities of the quantified areas for particular spots are normalized to each other after deconvolution of the image channels. The latter effectively corrects any systematic distortion error that might arise because of the different signal distributions for each isotope (signal volumes for the whole gel are effectively normalized across detected spots). The major source of error is statistically related to signal counting acquisition, which is dictated by the amount of radio-

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activity present. Fluctuations from ideal exponential decay will result in the signal from one isotope being incorrectly deconvoluted into the channel of the other. Low intensity signals of 125I are most prone to this error. Results are effectively identical whether one employs the spot detection mask for the 125I or 131I channel, however it is advisable to employ the one mask for both channels for calibrated quantification (as was done to generate Fig. 4). One could also consider employing an intensity peak-based signal redistribution algorithm on the 125I signal to make it more closely resemble that of 131I, however we have not found that to be necessary and prefer to work with raw distributions which generate fewer opportunities for unforeseen data-processing artifacts to arise. Therefore the simple solution of equations (1) and (2) (see Note 23) is (A) technically correct to separate the radioactive signals originating from the two respective isotopes by deconvolution, and (B) pragmatically adequate to generate the data quality reported here without subsequently correcting for the marginally different signal distributions of those isotopes. 23. The two resulting images of each gel are used to generate deconvoluted images of the signals for each isotope using simultaneous equations. Briefly, the pixel value of any pixel of the first exposure is expressed in Eq. 1, A+B=X

(1)

and the pixel value of a given pixel of the second exposure is expressed in Eq. 2, MA + NB=Y

(2)

where A = the signal contribution of radiation type A (e.g., 125I) measured from the first exposure, B = the signal contribution of radiation type B (e.g., 131I) measured from the first exposure, M = decay factor for the set-specific radiation type A between the two measurements, N = decay factor for the set-specific radiation type B between the two measurements. X = pixel value corresponding to the radiation intensity in the first measurement, Y = pixel value corresponding to the radiation intensity in the second measurement. A and B are unknown and to be determined. X and Y are the measured intensities. M and N can be determined by measuring calibration samples if

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radioisotopic impurity is suspected, or by calculation based on theoretical decay rates and times of exposures, as performed in this study. The values for A and B can be calculated by simultaneously solving the equation system consisting of Eq. 1 and 2. This function was performed on a pixel by pixel basis using the aligned pixel matrices of both gel images by the owndeveloped “select_deconv_v1.1.pl” program (ProteoSys AG). 24. The use of the Imaging Plate to produce differential images is not new. Exposure to X-rays of different energies was already used to differentially visualize bone and soft tissue in clinical X-rays by some of the seminal Imaging Plate studies (3, 4 ). Johnstone et al. (21) used an absorbing sheet between source and Imaging Plate to discriminate between the energies of 32 P and either 35S or 14C tocreate differential images of proteins and nucleic acids in gels. In a related strategy,Monribot-Espagne et al. (22) made two different exposures using Imaging Plates sensitive to either 3H and 14C or only to 14C to obtain differential 2D-PAGE images of proteins labeled with 3H or 14C that had been blotted onto membranes. However these methods have disadvantages relative to the ProteoTope strategy. The use of an absorbing sheet is associated with back-scatter as the beta-particles are deflected by atomic collisions within the absorber leading to altered geometry, altered detection efficiency, and spatial diffusion of the signal. Therefore the size and intensity of the signal measured for a given source with or without absorber is different, resulting in markedly reduced resolution and unaesthetic images (data not shown). The use of an absorber would have been quite advantageous, however we were unable to generate acceptable 2D images with any of them. Despite testing multiple absorbers types of varying thickness including diverse low Z materials (e.g., plastics and carbohydrates) and metals with various Z values, we have never encountered one that would produce data comparable to that shown here, and we therefore continue to employ differential radioisotopic decay rates. Both of the analogous imaging methods mentioned above involving differential use of either 32P and 35S/14C, or 3H and 14C isotope pairs suffer from the serious drawback that the chemical labeling of different protein samples is dissimilar because the chemical distribution of the base elements in proteins differs when using 3H, 14C, 32P or 35S as labeling reagents. Therefore two images obtained from the same sample labeled by any two methods exhibits a different 2D-PAGE pattern, as discussed (19). Accordingly, slight differences between closely related samples as revealed in this study cannot be determined with those methods. By contrast, the ProteoTope strategy offers an efficient solution to the differential imaging of proteins, employing two chemically identical isotopes of radioactive iodine that produce identical 2D-PAGE patterns from the same sample

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Fig. 5. Statistical results for each individual spot. The average normalized photostimulated light units (PSL) for each spot in both samples is given as (“A av Norm” and “B av Norm”). Because individual gels vary in intensity these normalized intensities were derived by applying the average of individual abundance ratios from all gels to the average sum signal intensity for each spot in columns AO-AR of spreadsheet “SelectedSpots.” The corresponding ratio of A/B for relative protein abundance estimated from each spot is calculated (ratio A/B) with the accompanying 95% confidence interval for that ratio, as shown. Standard deviation (SD) and standard error of the means (SEM) are given in percentage total abundance, with the corresponding P-value for a null hypothesis of equal intensities. The bar diagrams at the right show the percentage signal for A (black bars) and B (light bars). Error bars show SEM.

in the polyacrylamide matrix of dried 2D-gels, with effectively zero postelectrophoretic sample loss. 25. Although our differential ratio estimates were the most accurate 2D-PAGE results submitted to the ABRF study by any participant, and comparable to differential isotope mass spec methods, unfortunately the abundance ratio estimates of some proteins corresponded to those given by the ABRF only within reasonably large error margins for our liking (Fig. 6). However the observations discussed below lead us to conclude that the major source of

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Fig. 6. Comparison of ProteoTope abundance ratios with those given by the ABRF. Abundance ratios submitted for weighted mean estimates from Fig. 5 (ProteoTope) compared to the abundance ratios published by the ABRF after the study. Error bars show Standard Errors of Means. Protein name abbreviations follow Fig. 4. ABRF ratios for A/B (with submitted ProteoTope ratio) were GP, 0.013 (0.015); LP, 1.000 (0.631); BSA, 1.000 (1.000*); C, 5.000 (6.106); P, 1.000 (1.287); BC, 4.000 (8.104); CA1 0.333 (0.385); R, 1.000 (1.129). The absolute percentage errors of the submitted ProteoTope estimates of these ratios relative to the ABRF values are presented underneath the protein name abbreviations (% Error). The asterisk associated with BSA values indicates perfect value of 1:1 because this protein was used to normalize the signals from each radio-isotope across all gels.

discrepancy arose at the point of sample resolubilization, rather than during the quantitative analysis itself. There were two estimates of abundance ratios between Samples A and B that warrant particular discussion in this context: Lactoperoxidase and Beta Casein. For Lactoperoxidase our estimated ratio A/B was 0.63:1 (P = 4×10−6), rather than the ABRF value of 1:1. (We explicitly and categorically have no reason to presume incorrect sample mixing by the ABRF.) This protein exhibited generally weak incorporation of radioactivity relative to that of a silver stained gel of the

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Fig. 7. A suction device to remove IPs from exposure cassettes without bending them. The handle is a hollow tube. Suction is created by hole at the blocking the upper (left in the image) end with a finger. This provides sufficient force to lift the IP high enough out of the cassette that its edges can be accessed with the fingers. The suction device was purchased from raytest GmbH.

45° marked corner

Fig. 8. Recommended orientation of the Fuji IP in the FLA3000 IP stage. The left panel shows an IP in the IP stage ready for scanning. The right panel shows an enlargement of the upper left corner, where the diagonally removed corner of the IP is oriented. Both edges of the IP should be exactly flush with the walls of the IP stage.

Fig. 9. Positioning the FLA3000 IP stage into the FLA3000 imager. The small arrow between H & I on the reverse of the IP stage should align with the large arrow on the FLA3000 imager.

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same samples (10), which was reflected by the rather large 95% confidence limits of Fig. 5 (0.473 – 0.808). However, in light of the result for Beta Casein, we conclude that the abundance of this protein had probably been altered in at least one sample in some manner prior to taking the sample aliquots for the radiolabeling step. For Beta Casein our estimate of the A/B abundance ratio contained >100% error relative to the nominal ABRF value (weighted average estimate 8.104:1 compared to ABRF value 4:1). None of the 95% confidence limits for any of our four quantified spots approach a ratio of 4 (Fig. 5), and a t-test against the null hypothesis of a 4:1 ratio shows a highly significant difference (P = < 0.00001). Because of our rigorous experimental design (Fig. 1A) we can therefore be statistically quite confident that the abundance of this protein was altered in at least one sample between the time the samples were mixed in the laboratory of the ABRF and the time that the two identical aliquots were taken from each resolubilised sample before iodination in our laboratory. Note that the alteration(s) did not occur during or after radio-iodination of the samples (during the ProteoTope analysis itself), which would have increased the measured variability and reduced the significance of the results. There is no other (persuasive) argument for the reproducible estimation of these observed abundance ratios being obtained after labeling the proteins with two different radioiodine isotopes used in eight labeling reactions across sixteen inverse replicate gels (Fig. 1A). It must be emphasized that although the estimates of abundance ratios that we submitted were associated with undesirably high deviations from the values provided by the ABRF (Fig. 6), we could identify the probable cause of the discrepancy because of our stringent experimental design. It is that process and its attributes, rather than the submitted abundance ratios, which provide the main technical features of the data quality generated by the ProteoTope method. The most probable source of this sample alteration is sample handling prior to labeling in our laboratory, or possibly during shipment. One obvious source of potential variability is the resolubilization of the desiccated pellet. There was no opportunity to optimize this process because of the imposed time limit, and the resolubilization of desiccated proteins is not standardized in our laboratory. We would normally recommend transport of proteins as frozen SDS solutions or tissue sections, although this is associated with higher costs. We were the only participant to resolubilize these desiccated proteins in boiling SDS buffer, suggesting that this approach may have been suboptimal for these samples, however further experiments would be necessary to draw a conclusion. The fact that at least one of the samples was apparently suboptimally resolubilized underlines that this was a real-life one-off experiment. This further emphasizes the requirement

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for scrupulously standardized sample handling procedures when working with physico-chemically labile protein species. Nevertheless, the confidence with which we may draw these conclusions stems from our rigorous experimental design and the high quality quantitative data generated. This introduces a further consideration of data quality assessment that merits brief discussion. In this study we employed the somewhat unusual number of 16 SDS-PAGE gels, generating 32 images, to compare two samples. The entire ProteoTope quantitative analysis consumed just 25% of each sample (nominally 20 µg = 2×2×5 µg), and therefore we could have generated 64 such differential 2D-PAGE gels using the same design with the nominally 80 µg protein in each sample. This high number of accurate measurements gave rise to extraordinarily precise and highly significant differential quantifications of the various abundance ratios in the samples we measured (after they had apparently been differentially solubilized). For instance, Ribonuclease A was a protein with a nominal 1:1 mixture ratio given by the ABRF. The ratios estimated for the four individual spots of this protein in Fig. 5 ranged from 1.112 to 1.204, generating P-values (n = 16 per spot) between 0.042 to 1 × 10−5. Once again, because of the meticulous experimental design, we can conclude that there was a slight difference (relative to BSA levels) in the abundance of this protein between the samples before the replicate aliquots were taken for iodination. A similar situation would apply for Horse Radish Peroxidase. Alternatively, there may have been a difference in BSA levels, against which signal intensities for both radio-isotopes were normalized. Consistent with the latter possibility, normalization of spot intensities to assumed 1:1 values of Peroxidase and Ribonuclease rather than 1:1 for BSA did give somewhat better approximation of abundance ratio estimations to those provided by the ABRF. In summary, although the extraordinarily stringent level of statistical certainty obtained here is seldom required, the availability of an analytical detection method with these technical parameters can be advantageous, especially when smaller numbers of replicates are performed on limited amounts of valuable material. 26. The weakest average protein spot intensity in this study (Spot BA_459 sample A) was 116 photo stimulated luminescence units (PSL) (11, 12). The highest was 94973 PSL (BA_2369), representing a dynamic range of 800-fold measured between protein signals in this experiment. The strongest signal was on gel 2 which had an intensity of 16800-fold above measurement background and occupied almost 60000 levels of the 16 bit data spectrum. The FLA3000 we used is a well characterized instrument whose technical specifications are available from the manufacturer (e.g. http://fujifilmlifescienceusa.com/assets/pdf/fla3000brochure.pdf).

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Essentially, laser induced PSL events are recorded by a photomultiplier detector. Data are then written to a floating point matrix file which stores data logarithmically over the 65536 bits. The linear dynamic range of the Imaging Plate, although intrinsically vastly superior to alternative methods, is presently limited by the imaging device (data not shown). Briefly the Imaging Plate has at least 250-fold better dynamic range than is currently exploited by the FLA3000 imaging device. References 1. Van den, B. G., Arckens, L. (2005) Recent advances in 2D electrophoresis: an array of possibilities. Expert. Rev. Proteomics 2, 243–252. 2. Zolg, J. W., Langen, H. (2004) How industry is approaching the search for new diagnostic markers and biomarkers. Mol. Cell Proteomics 3, 345–354. 3. Cahill, M. A., Wozny, W., Schwall, G., Schroer, K., Holzer, K., Poznanovic, S., Hunzinger, C., Vogt, J. A., Stegmann, W., Matthies, H., Schrattenholz, A. (2003) Analysis of relative isotopologue abundances for quantitative profiling of complex protein mixtures labelled with the acrylamide/D3-acrylamide alkylation tag system. Rapid Commun. Mass Spectrom. 17, 1283–1290. 4. Schrattenholz, A., Wozny, W., Klemm, M., Schroer, K., Stegmann, W., Cahill, M. A. (2005) Differential and quantitative molecular analysis of ischemia complexity reduction by isotopic labeling of proteins using a neural embryonic stem cell model. J. Neurol. Sci. 229–230, 261–267. 5. Poland, J., Cahill, M. A., Sinha, P. (2003) Isoelectric focusing in long immobilized pH gradient gels to improve protein separation in proteomic analysis. Electrophoresis 24, 1271–1275. 6. Poznanovic, S., Schwall, G., Zengerling, H., Cahill, M. A. (2005) Isoelectric focusing in serial immobilized pH gradient gels to improve protein separation in proteomic analysis. Electrophoresis 26, 3185–3190. 7. Neubauer, H., Clare, S. E., Kurek, R., Fehm, T., Wallwiener, D., Sotlar, K., Nordheim, A., Wozny, W., Schwall, G. P., Poznanovic, S., Sastri, C., Hunzinger, C., Stegmann, W., Schrattenholz, A., Cahill, M. A. (2006) Breast cancer proteomics by laser capture microdissection, sample pooling, 54-cm IPG IEF, and differential iodine radioisotope detection. Electrophoresis 27, 1840–1852. 8. Poznanovic, S., Wozny, W., Schwall, G. P., Sastri, C., Hunzinger, C., Stegmann, W., Schrattenholz, A., Buchner, A., Gangnus, R., Burgemeister, R., Cahill, M. A. (2005) Differential radioactive proteomic analysis of microdissected renal cell carcinoma tissue by 54 cm isoelectric focusing in serial immobilized pH gradient gels. J. Proteome. Res. 4, 2117–2125. 9. Wozny, W., Schroer, K., Schwall, G. P., Poznanovic, S., Stegmann, W., Dietz, K., Rogatsch, H., Schaefer, G., Huebl, H., Klocker, H., Schrattenholz, A., Cahill, M. A. (2007) Differential radioactive quantification of protein abundance ratios between benign and malignant prostate tissues: cancer association of annexin A3. Proteomics 7, 313–322.

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10. Turck, C. W., Falick, A. M., Kowalak, J. A., Lane, W. S., Lilley, K. S., Phinney, B. S., Weintraub, S. T., Witkowska, H. E., Yates, N. A. (2007) ABRF-PRG2006 study: Relative protein qQuantitation. Mol. Cell Proteomics 6, 1291–1298. 11. Amemiya, Y., Miyahara, J. (1988) Imaging plate illuminates many fields. Nature 336, 89–90. 12. Miyahara, J. (1999) The imaging plate: a new radiation image sensor. Chem. Today (Japan) 223, 29–36. 13. Gorg, A., Postel, W., Gunther, S. (1988) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 9, 531–546. 14. Gorg, A. (1993) Two-dimensional electrophoresis with immobilized pH gradients: current state. Biochem. Soc. Trans. 21, 130–132. 15. Gorg, A., Obermaier, C., Boguth, G., Harder, A., Scheibe, B., Wildgruber, R., Weiss, W. (2000) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 21, 1037–1053. 16. Gorg, A., Weiss, W., Dunn, M. J. (2004) Current two-dimensional electrophoresis technology for proteomics. Proteomics 4, 3665–3685. 17. Gorg, A., Weiss, W. (1999) Analytical IPG-Dalt. Methods Mol. Biol. 112, 189–195. 18. Rabilloud, T. (1996) Solubilization of proteins for electrophoretic analyses. Electrophoresis 17, 813–829. 19. Vuong, G. L., Weiss, S. M., Kammer, W., Priemer, M., Vingron, M., Nordheim, A., Cahill, M. A. (2000) Improved sensitivity proteomics by postharvest alkylation and radioactive labelling of proteins. Electrophoresis 21, 2594–2605. 20. Bonardi, M. L., Birattari, C., Groppi, F., Gini, L., Mainardi, H. S. C. (2004) Cyclotron production and quality control of “High Specific Activity” radionuclides in “No Carrier Added” form for radioanalytical applications in life sciences. J. Radioanalyt. Nucl. Chem. 259, 415–419. 21. Johnston, R. F., Pickett, S. C., Barker, D. L. (1990) Autoradiography using storage phosphor technology. Electrophoresis 11, 355–360. 22. Monribot-Espagne, C., Boucherie, H. (2002) Differential gel exposure, a new methodology for the two-dimensional comparison of protein samples. Proteomics 2, 229–240.

43 Quantification of Proteins on Polyacrylamide Gels Bryan John Smith 1. Introduction Quantification of proteins is a common challenge. There are various methods described for estimation of the amount of protein present in a sample, for example, amino acid analysis and the bicinchoninic acid (BCA) method (see Chapters 83 and 3). Proteins may be quantified in sample solvent prior to electrophoresis and can provide an estimate of total amount of protein in a sample loaded onto a gel (1,2). These methods quantify total protein present but not of one protein in a mixture of several. For this, the mixture must be resolved. Liquid chromatography achieves this, and the various proteins may be quantified by their absorbancy at 220 nm or 280 nm. Microgram to submicrogram amounts of protein can be analyzed in this way, using microbore high-performance liquid chromatography (HPLC). A method has been published whereby theoretical extinction coefficients may be calculated for peptides, based on their sequence, though it is probably less suited to intact proteins because of the influence on UV aborption of longer sequence and structure. (3). Problems may occur if particular species chromatograph poorly (such as hydrophobic polypeptides on reverse-phase chromatography). An alternative is polyacrylamide gel electrophoresis (as the mixture-resolving step) followed by protein staining and densitometry. Exceptions are small peptides that are not successfully resolved and stained in gels, and which are more suited to chromatography (3). Microgram to submicrogram amounts of protein (of more than a few kilodaltons in size) may be quantified in this way. Not every protein stain is best suited to quantification in gels, however. The popular noncolloidal Coomassie gel stains such as PAGE blue ‘83 are not suitable. As discussed by Neuhoff et al. (4), staining by Coomassie dye is difficult to control—it may not fully penetrate and stain dense bands of protein, it may demonstrate a metachromatic effect (whereby the protein–dye complex may From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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show any of a range of colors from blue through purple to red), and it may be variably or even completely decolorized by excessive destaining procedures (because the dye does not bind covalently). As a consequence, stoichiometric binding of Coomassie dye to protein is commonly not achieved. Sensitive silver stains are commonly used to detect rare proteins at levels at levels of the order of 0.05 ng/mm2 in two-dimensional (2-D) gels, for example, but they, too, are not suitable for quantification. For various reasons, proteins vary widely in their stainability by silver—some do not stain at all, others may show a metachromatic effect. Use of silver staining for quantification is complicated by other factors, too—such as the difficulty in obtaining staining throughout the gel, not just at the surface. This subject is reviewed at greater length in ref. 5, but in summary it may be said that while silver stain methods have great sensitivity, they are problematical for quantification purposes. One quantitative staining method generates negatively stained bands (6). The technique generates a white background of precipitated zinc salt, against which clear bands of protein (where precipitation is inhibited). This may be viewed by dark field illumination. The gel may be scanned at this stage, the negative staining being approximately quantitative (for horse myoglobin) in a range from about 100 ng/mm2 to 2 µg/mm2 or more. Sensitivity may be improved by staining of the background by incubation with tolidine. The method was described for 12% and 15% T gels, but the degree of zinc salt precipitation and subsequent toning is dependent upon %T and at closer to 20%T the toning procedure may completely destain the gel, restricting the usefulness of the method. A number of other quantitative staining methods have been discussed in the literature. Notably, Neuhoff et al. (4) have made a thorough study of various factors affecting protein staining by Coomassie Brilliant Blue G-250 (0.1% [w/v] Color Index number 42655 in 2% [w/v] phosphoric acid, 6% [w/v] ammonium sulfate). This colloidal stain does not stain the background in a gel and so washing can give crystal-clear backgrounds. A method derived from that of Reisner et al. (7) has proven useful for quantification of proteins on gels. The stain used is 0.4% (w/v) Brilliant Blue G250 in 3.5% (w/v) perchloric acid. This method is described below. O’Keefe (8) described a method whereby cysteine residues are labeled with the thiol-specific reagent monobromobimane. On transillumination (at λ = 302 nm) labeled bands fluoresce. However, the method requires quantitative reaction of cysteines, which may be totally lacking in some polypeptides, and a nominal detection limit of 10 pmol of cysteine is claimed. Better luminescent stains have been developed by Molecular Probes—the Sypro family of dyes. The Sypro Ruby stain is possibly the most sensitive of these, approaching the sensitivity of silver stains. The method for use of this stain is given below.

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2. Materials 2.1. Colloidal Coomassie Brilliant Blue G Method 1. A suitable scanning densitometer, for example, the Molecular Dynamics Personal Densitometer SI with Image Quant software or the Bio-Rad Fluor S MultiImager with Multi Analyst software. Such equipment can scan transparent objects such as wet gels, gels dried between transparent films, and photographic film, and digitize the image that may then be analyzed. 2. Protein stain: 0.4% (w/v) Coomassie Brilliant Blue G (C.I. 42655; Sigma, code B0770, 90% by elemental analysis) in 3.5% (w/v) perchloric acid (see Note 1). Stable for months at room temperature, in the dark. Beware the low pH of this stain. Use personal protective equipment (coat, gloves, safety glasses). Use fresh, undiluted stain, as supplied. For best results, do not re-use the stain. 3. Destaining solution: Distilled water.

2.2. Sypro Ruby Method 1. Equipment for scanning fluorescent bands on gels, for example, the Bio-Rad Fluor S MultiImager with Multi Analyst software, which can be used to photographically record or scan wet gels, allowing repeated scanning procedures (as staining or destaining proceeds). Simple viewing of stained bands may be done under a hand-held 300-nm UV lamp or on a transilluminator. Protect eyes with UV-opaque glasses. The Sypro dye may be excited at 280 or 450 nm and emits at 610 nm. 2. Protein stain: Sypro Ruby gel stain (Molecular Probes, product number S-12000 or S-12001) (see Note 2). This stain is stable at room temperature in the dark. Exact details of the stain are not revealed but according to the manufacturer, the stain contains neither hazardous nor flammable materials. Use fresh, undiluted stain only. 3. Destain: Background may be reduced by rinsing in water. 4. Clean dishes, free of dust that might contribute to background staining.

3. Method 3.1. Colloidal Coomassie Brilliant Blue G Method 1. At the end of electrophoresis, wash the gel for a few minutes with several changes of water (see Note 3), then immerse the gel (with gentle shaking) in the colloidal Coomassie Brilliant Blue G protein stain. This time varies with the gel type (e.g., 1.0–1.5 h for a 1–1.5 mm thick sodium dodecyl sulfate [SDS] polyacrylamide gel slab), but cannot really be overdone. Discard the stain after use, for its efficacy declines with use. 2. At the end of the staining period, decolorize the background by immersion in distilled water, with agitation, and a change of water whenever it becomes colored. Background destaining is fairly rapid, giving a clear background after a few hours (see Notes 4 and 5).

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3. Measure the extent of blue dye bound by each band by scanning densitometry. Compare the dye bound by a sample with those for standard proteins run and stained in parallel with the sample, on the same gel (see Notes 6–8 and 11–17).

3.2. Sypro Ruby Method 1. At the end of electrophoresis, rinse the gel in water briefly, put it into a clean dish and then cover it with Sypro Ruby gel stain solution. Gently agitate until staining is completed, which may take up to 24 h or longer (see Note 9). Overstaining will not occur during prolonged stained. Do not let the stain dry up on the gel during long staining procedures. Discard the stain after use, for it becomes less efficacious with use. During the staining procedure the gel may be removed from the stain and inspected under UV light to monitor progress. If the staining is insufficient, the gel may be replaced in the stain for further incubation. 2. Destain the background by washing the gel in a few changes of water for 15–30 min. 3. Measure the extent of dye bound, that is, the luminescence, by each band by scanning. Compare the dye bound by a sample with that for standard proteins run and stained in parallel with the sample, on the same gel (see Note 10).

4. Notes 1. The Coomassie Brilliant Blue G stain mixture may be readily made from the components but a commercially-available alternative is the GelCode blue stain reagent from Pierce (product no. 24590 or 24592). Details of the stain components are not divulged, other than they also include Coomassie (G250), but the stain is used in the same way as described for the reagent prepared as described above. Pierce recommend that their GelCode stain is agitated before use, to disperse any dye aggregates that may have formed therein. 2. Molecular Probes do not reveal the components of their reagent. Although more sensitive than the blue stain, it does require UV irradiation for detection. 3. The water wash that precedes staining by Coomassie Brilliant Blue G is intended to wash away at least some SDS from the gel, and so speed up destaining of the background. However, it should be remembered that proteins are not fixed in the gel until in the acidic stain mixture and consequently some loss of small polypeptides may occur in the wash step. Delete the wash step if this is of concern, or employ a fixing step immediately after electrophoresis, for example, methanol/glacial acetic acid/ water: 50:7:43, by volume for 15–30 min, followed by water washing to remove the solvent and acid. 4. Destaining of the background may be speeded up by frequent changes of the water, and further by inclusion in this wash of an agent that will absorb free dye. Various such agents are commercially available (e.g.,

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6.

7.

8.

9.

10.

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Cozap, from Amika Corp.), but a cheap alternative is a plastic sponge of the sort used to plug flasks used for microbial culture. The agent absorbs the stain and is subsequently discarded. The background can be made clear by these means, and the stained bands remain stained while stored in water for weeks. They may be restained if necessary. The Coomassie Brilliant Blue G-stained gel may be dried between transparent sheets of dialysis membrane or cellophane (available commercially, e.g., from Novex) for storage and later scanning. Beware that bubbles or marks in the membrane may add to the background noise upon scanning. Heavily loaded samples show up during staining with Coomassie Brilliant Blue G, but during destaining the blue staining of the proteins becomes accentuated. Bands of just a few tens of ng are visible on a 1 mm-thick gel (i.e., the lower limit of detection is < 10 ng/mm2). Variability may be experienced from gel to gel, however. For example, duplicate loadings of samples on separate gels, electrophoresed and stained in parallel, have differed in staining achieved by a factor of 1.5, for reasons unknown. Furthermore, different proteins bind the dye to different extents: horse myoglobin may be stained twice as heavily as is bovine serum albumin, although this, too, is somewhat variable. While this Coomassie Brilliant Blue G is a good general protein stain, It is advisable to treat sample proteins on a case by case basis. It is a requirement of this method that dye is bound stoichiometrically to polypeptide over a useful wide range of sample size. The staining by Coomassie Brilliant Blue G may be quantitative, or nearly so, from about 10 to 20 ng/mm2 up to large loadings of 1–5 µg/mm2. Concentrated protein solutions may be diluted to fall within the useful range. A general point may be made, that whatever method is used for quantification, it should be checked with the protein of interest, to confirm that staining is linear (quantitative) over the range of sample size being used. The stoichiometry of dye binding is subject to some variation, such that standard curves may be either linear or slightly curved, but even the latter case is acceptable provided standards are run on the same gel as samples. Because performance of the Coomassie Brilliant Blue G stain is variable it essential to run and stain standards and samples on the same gel, and to do so in duplicate. The Sypro Ruby method allows for periodic observation and further staining if required. It can be the case that for small loadings, however, prolonged staining (24 h or more) may be required to obtain best sensitivity. The stained gel may be dried for storage but can then lose luminescence. The Sypro Ruby method is subject to the same sort of variation as the Coomassie Brilliant Blue G method, from gel to gel with standard curves being linear or slightly curved. Generally the Sypro Ruby method is more

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sensitive than the Coomassie Brilliant Blue G method, for instance about fivefold more so for bovine serum albumin. However, protein to protein variation may apparently be greater in the case of the Sypro Ruby stain. For instance, horse myoglobin binds about 10-fold less dye than bovine serum albumin does. Thus in one experiment, the minimum amount of bovine serum albumin detectable after Sypro Ruby staining was about 5 ng/mm2 (about four- to fivefold more sensitive than samples stained in parallel by the Coomassie Brilliant Blue G method), whereas the minimum amount of horse myoglobin detectable was about 50 ng/mm2 (similar to that detectable by the Coomassie Brilliant Blue G stain). It is therefore recommended that samples and standards are run and stained in parallel on the same gel, in duplicate. 11. Best quantification is achieved after having achieved good electrophoresis. Adapt the electrophoresis as necessary to achieve good resolution, lack of any band smearing or tailing, and lack of retention of sample at the top of the gel by aggregation. For stains where penetration of dense bands may be a problem, avoid dense bands by reducing the size of the loaded sample and/ or electrophoresing the band further down the length of the gel (to disperse the band further) and/or use lower %T gel. Thin gels, of 1 mm thickness or less, allow easier penetration of dye throughout their thickness. Gradient gels have a gradient of pore size that may cause variation of band density and background staining. Use nongradient gels if this is a problem. 12. For absolute quantification of a band on a gel, accurate pipetting is required, as is a set of standard protein solutions that cover the concentration range expected of the experimental sample. Ideally, adjust the concentrations of the various standards and a sample so that the volume of each that is loaded onto the gel is the same and the need for pipet adjustment is minimized. Run and stain these standard solutions at the same time and if possible on the same gel, as the experimental sample in order to reduce possible variations in band resolution or staining, background destaining and so on. Ideally the standard protein should be the same as that to be quantified but if, as is commonly the case, this is impossible then another protein may be used (while recognizing that this protein may bind a different amount of dye from that by the experimental protein, so that the final estimate obtained may be in error). The standard protein should have similar electrophoretic mobility to the proteins of interest, so that any effect such as dye penetration, due to pore size, is similar. Make the standard protein solution by dissolving a relatively large and accurately weighed amount of dry protein in water or buffer, and dilute this solution as required. If possible, check the concentration of this standard solution by alternative means (say amino acid analysis). Standardize treatment of samples in preparation for electrophoresis—treating of sample

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solutions prior to SDS-PAGE may cause sample concentration by evaporation of water for instance. To minimize this problem heat in small (0.5 mL) capped Eppendorf tubes, cool, and briefly centrifuge to take condensed water back down to the sample in the bottom of the tube. When analyzing the results of scanning, construct a curve of absorbency vs protein concentration from standard samples and compare the experimental sample(s) with this. Construct a standard curve for each experiment. If comparing the abundance of one protein species with others in the same sample, then standards are not required, provided that no species is so abundant that dye binding becomes saturated. Be aware that such relative estimates are approximate, as different proteins bind dye to different extents. It is sometimes observed in electrophoresis that band shape and width are irregular—heavily loading a gel can generate a broad band that may interfere with the running of neighboring bands, for instance. It is necessary to include all of such a band for most accurate results. Avoid damage to the gel—a crack can show artificially as an absorbing band (or peak) on the scan. Gels of low %T are difficult to handle without damage, but they may be made tougher (and smaller) by equilibration in aqueous ethanol, say, 40% (v/v) ethanol in water, 1 h or so. Too much ethanol may cause the gel to become opaque, but if this occurs merely rehydrate the gel in a lower % (v/v) ethanol solution. A gradient gel may assume a slightly trapezoid shape upon shrinkage in ethanol solutions— this makes scanning tracks down the length of the gel more difficult. When scanning, eliminate dust, trapped air bubbles, and liquid droplets, all of which contribute to noisy baseline. Methods have been described for quantification of submicrogram amounts of proteins that have been transferred from gels to polyvinylidene difluoride or similar membrane (e.g., 9). A Sypro dye blot stain equivalent to the method described above for gels is available from Molecular Probes. Note, however, that transfer from gel to membrane need not (indeed, usually does not) proceed with 100% yield, so that results do not necessarily accurately reflect the content of the original sample.

References 1. Draber, P. (1991) Quantification of proteins in sample buffer for sodium dodecyl sulfate-polyacrylamide gel electrophoresis using colloidal silver. Electrophoresis 12, 453–456. 2. Henkel, A. W. and Bieger, S. C. (1994) Quantification of proteins dissolved in an electrophoresis sample buffer. Analyt. Biochem. 223, 329–331. 3. Kuipers, B. J. H,. and Gruppen, H. (2007) Prediction of molar extinction coefficients of proteins and peptides using UV absorption of the constituent amino acids

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5. 6.

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at 214 nm to enable quantitative reverse phase high-performance liquid chromatography-mass spectrometry analysis, J. Agric. Food Chem 55, 5445–5451. Neuhoff, V., Stamm, R., Pardowitz, I., Arold, N., Ehhardt, W., and Taube, D. (1990) Essential problems in quantification of proteins following colloidal staining with Coomassie Brilliant Blue dyes in polyacrylamide gels, and their solution. Electrophoresis 11, 101–117. Syrovy, I. and Hodny, Z. (1991) Staining and quantification of proteins separated by polyacrylamide gel electrophoresis. J. Chromatogr. 569, 175–196. Ferreras, M., Gavilanes, J. G., and Garcia-Segura, J. M. (1993) A permanent Zn2+ reverse staining method for the detection and quantification of proteins in polyacrylamide gels. Analyt. Biochem. 213, 206–212. Reisner, A. H., Nemes, P., and Bucholtz, C. (1975) The use of Coomassie Brilliant Blue G250 perchloric acid solution for staining in electrophoresis and isoelectric focusing on polyacrylamide gels. Analyt. Biochem. 64, 509–516. O’Keefe, D. O. (1994) Quantitative electrophoretic analysis of proteins labeled with monobromobimane. Analyt. Biochem. 222, 86–94. Patton, W. F., Lam, L., Su, Q., Lui, M., Erdjument-Bromage, H., and Tempst, P. (1994) Metal chelates as reversible stains for detection of electroblotted proteins: application to protein microsequencing and immune-blotting. Analyt. Biochem. 220, 324–335.

44 Using SDS-PAGE and Scanning Laser Densitometry to Measure Yield and Degradation of Proteins Aaron P. Miles and Allan Saul 1. Introduction A variety of techniques are available for the quantification of proteins, their degradation products and other impurities. Densitometry is particularly useful due to its sensitivity, accuracy and versatility, and it can be applied to proteins in gels or on membranes and used with numerous detection methods. In addition to being accurate, sensitive and reproducible, the technique is cost-effective, simple, and does not require a high degree of specialized training, yet provides technical advantages over other available tools. For example, Vuletich and Osawa showed that densitometry following SDS-PAGE and electroblotting was 20-fold more sensitive than HPLC at detecting oxidatively modified myoglobin (1). In addition, Morçöl and Subramanian reported the development of a sensitive Ponseau-S-stained dot blot/densitometric protein assay that was less subject to interference by pH, detergent and other reagents than the commonly used Bradford protein assay (2). More recently, Zhu et al. and Miles et al. reported a quantitative densitometric method that measures host cell derived impurity levels on immunoblots of recombinant proteins expressed in E. coli (3, 4). This method was shown to be at least 20 times more sensitive than a commercially available ELISA kit designed to measure host cell impurity levels (3). Other groups have reported on the utility of gel or fluorograph densitometry, using multiple detection methods, in carbohydrate analysis (5), taxonomic and forensic applications (6), as a clinical diagnostic tool in Balkan nephropathy (7) and in the analysis of high density lipoproteins (8), as a means of quantifying relative levels of excretorysecretory polypeptides synthesized in vitro by Schistosoma mansoni daughter sporocysts (9), and for analyzing adsorption of proteins and an oligodeoxynucleotide in Alhydrogel®-based malaria vaccine candidates (10).

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As a typical example we describe here the analysis of recombinant proteins by SDS-PAGE and scanning laser densitometry. This approach offers advantages over other techniques for two main reasons: 1) proteolytic degradation is common among these proteins, and this method is able to accurately quantify it in both minute and large amounts, and 2) the common constituents of fermentation and refolding tanks and purification processes do not cause interference, so it can be used at any stage throughout production and purification. Analysis of a recombinant malarial protein at two different stages of purity will be used to illustrate this method. 2. Materials 1. SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis) equipment. 2. Coomassie Blue staining solution: 40% methanol, 10% glacial acetic acid, two PhastGel Blue R tablets (Amersham Biosciences, Piscataway, NJ) per liter. 3. Coomassie Blue destaining solution: 5% methanol, 10% glacial acetic acid. 4. 0.45-micron filtration unit. 5. Concentrated, diafiltered test protein (Pvs25H) fermentation culture supernatant. 6. Purified test protein (Pvs25H) solution. 7. Personal Densitometer SI with ImageQuant software (Molecular Dynamics Inc., Sunnyvale, CA, [11]).

3. Methods The methods described below outline (1) the running of SDS-PAGE gels followed by visualization with Coomassie Blue, and (2) scanning and analyzing the gels using laser densitometry and ImageQuant software. 3.1. SDS-PAGE 1. Load the gel with samples as desired, and run according to the manufacturer’s instructions. 2. Filter the staining solution using a 0.45 µm filtration unit (see Note 1). 3. Add staining solution to the gel, bring to a boil in a microwave oven, and agitate for 30 min (see Notes 2 and 3). 4. Pour off staining solution, add destaining solution, bring to a boil, and agitate for 15 min ( see Note 4). 5. Change destaining solution, bring to a boil again, and continue agitation for 15 more min. 6. Change destaining solution once more, bring to a boil, and continue destaining until gel background is clear or almost clear, approximately 10 min. Take care not to overly destain the gel as minor protein bands can disappear (see Note 5). 7. Place the gel in water until scanned. Fig. 1 shows a nonreduced 4–20% Tris-glycine SDS-PAGE gel that contains standards of purified Pvs25H (a recombinant form of

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Fig. 1. SDS-PAGE of Pvs25H Fermentation Culture Supernatant with Purified Pvs25H Standards. Nonreduced 4–20% Tris-glycine SDS-PAGE Coomassie Blue-stained gel showing purified Pvs25H standards and concentrated, diafiltered fermentation culture supernatant. Purified Pvs25H was loaded in the indicated amounts for comparison to Pvs25H in the fermentation supernatant.

the 25 kDa ookinete suface protein from the malaria parasite Plasmodium vivax, [12]) and a concentrated, diafiltered fermentation culture supernatant of the same protein. Fig. 2 shows a reduced 15% acrylamide gel loaded with the same purified protein that was subjected to 37°C storage for up to 14 days to monitor stability (see Note 6). This protein is heavily disulfide bonded and must be reduced in order to visualize the degradation occurring over time. These gels will serve as examples of two applications of scanning laser densitometry: (1) monitoring low levels of degradation in a highly purified protein, and (2) quantifying yields of a protein recovered in a fermentation broth not yet subject to purification.

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Fig. 2. SDS-PAGE of Purified Pvs25H Stored at 37°C. Reduced 15% SDS-PAGE Coomassie Blue-stained gel showing samples taken at the indicated time points during storage at 37°C. All protein loads are 2 µg, separated by blank lanes. *A minor band accumulating with time whose sequence corresponds to cleavage at a single site within Pvs25H. **A minor band (Pvs25Hderived) present in final purified Pvs25H, not increasing upon storage. This type of gel was chosen to better resolve Pvs25H, which migrates diffusely in a gradient gel (see Fig. 1).

Fig. 3. Gel from Fig. 2 with Lines Drawn in ImageQuant for Analysis. Shown is the same gel as in Fig. 2, with lines drawn down two lanes using ImageQuant. These lines define the length and width of the areas being analyzed. Lengths and widths are adjusted to encompass all protein in each lane. These lines give rise to the line graphs shown in Fig. 5.

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3.2. Laser Densitometry 1. Scan the gel according to manufacturer’s instructions, ensuring that the lanes of the gel run straight from the top to the bottom of the scanning bed. 2. Draw an encompassing line for analysis down each lane of interest and widen the line appropriately. Include in the analyzed area of each lane enough length from top to bottom and enough width to envelope all visible protein bands. Figs. 3 and 4 illustrate this, using the same gels as shown in Figs. 2 and 1, respectively. 3. Create a line graph for each line drawn. Examples are shown in Fig. 5, which shows representations (in optical density units) of purified Pvs25H stored for one (A) and 14 (B) days at 37°C, and Fig. 6, which shows Pvs25H in concentrated, diafiltered

Fig. 4. Gel from Fig. 1 with Lines Drawn in ImageQuant for Analysis. Shown is the same gel as in Fig. 1, with lines drawn down each lane containing protein. The line drawn down the fermentation supernatant lane gives rise to the line graph shown in Fig. 6.

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fermentation culture supernatant. Note in Fig. 5B the appearance of a minor degradation product over time (peak 2), and it’s corresponding band in Fig. 2. 4. Adjust all relevant settings to give the most accurate representation of the baseline signal. Options for this within the ImageQuant software may include “automatic” A 1.0 1 0.8

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Fig. 5. Line Graphs of Pvs25H at 37°C, Days 1 and 14. The line graphs for the lines drawn in Fig. 3 are shown, with (A) showing day 1 and (B) showing day 14. These are graphical representations of the optical density vs. millimeters measured by the scanner at each point from the top to the bottom of the drawn lines. In these cases, the baselines were detected by simply making minor adjustments to the peak detection parameters (baselines not shown). Note the increase in the area of peak 2, a minor degradation product, from day 1 to day 14.

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(the baseline is detected by the software based on manually adjusted settings to appropriately identify peaks), “lowest point” (the most transparent part of the lane is taken as the baseline), and point-by-point manipulation. There are other software programs available that allow more customized identification and tuning of the baseline and peaks (e.g., PeakFit). It is then usually necessary to redefine the peaks by hand, rather than letting the software do it, in order to fine tune where each peak begins and ends (see Note 7). Once the peaks are defined, they are integrated (i.e., the area under each peak is calculated by the software). In this way one quantifies the proportion of one or several peaks in relation to all others, and determines both the total protein in the lane and the percent of each protein of interest. 6. Using the areas generated from the integration of each Pvs25H standard peak, create a graph of area counts vs. known protein load using the appropriate software (e.g., Excel, SigmaPlot). Determine the linear range, fit a line for the data points of that range, and obtain an equation for the line. Using this method, the amount of Pvs25H in the fermentation supernatant lane of Fig. 1 was found to be 1.49 µg. This corresponds to a total production of 500 mg from a 60 L fermentation (see Note 8). Likewise, the amount of full-length (nondegraded) purified Pvs25H was found to be decreasing with time, while a minor degradation product was increasing.

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4. Notes 1. It is especially important when using this method to filter the Coomassie Blue stain prior to use. This eliminates large particles that otherwise stick to the gel and give rise to high background readings that interfere with quantification. Filtered stain leads to much cleaner, more transparent gels. To accelerate the filtering process and to help prevent clogging of the filter, use a conveniently sized filtration unit (e.g., 1 L) and pour the solution in slowly, allowing it to filter through, before pouring more. If too much solution is poured in all at once, large particles quickly block the filter. It is not necessary to use a 0.22 µm filter, which will become clogged more quickly anyway. 2. Microwaving allows staining and destaining to proceed much more quickly than at room temperature, with no loss in sensitivity. 3. Standard staining and destaining times must be adhered to, especially when gel-to-gel comparisons are to be made. Staining for less than 30 minutes can result in non-uniformity in the binding of the Coomassie Blue dye to the proteins, especially in bands containing large amounts of protein. This can in turn cause some underestimation in the analysis of those bands. 4. A sponge or paper towel can be added to soak up stain from the gel, but this is not necessary, especially when microwaving, as the gel will quickly destain. 5. It is unnecessary to destain the gel until the background is completely clear. A small amount of background stain will generally not be detected as a protein peak or peaks by the software, and will become part of the baseline. Higher concentrations of acrylamide in gradient gels require slightly more time to destain, but with more extended destaining times protein bands (especially low molecular weight bands) can disappear. This will affect quantification of yield and purity and should be avoided. 6. After purification of Pvs25H, it was determined that this protein is better resolved on a 15% SDS-PAGE gel than on a 4–20% gradient gel (compare Fig. 1, 3.8 µg lane, to Fig. 2, 1 Day lane). This serves to illustrate an important point to consider when performing densitometry: protein quantification is more difficult to perform when the protein runs as a diffuse band. Consequently, the error in this technique increases with poorer resolution. While a good estimate can still be achieved, peak identification can be problematic. If necessary, different gel types should be tested to find the one that best resolves each protein of interest. 7. One is often interested in the proportion of one peak (the protein of interest) in relation to all others. Defining the peaks by hand allows the peak of

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interest to be isolated while, for instance, treating all other peaks as one. A percent yield is then easily calculated. 8. Scanning laser densitometry was found to be especially useful during the production of Pvs25H. This is because Pvs25H is produced as two populations of molecules that migrate well separated by SDS-PAGE (see large diffuse band beneath Pvs25H in Fig. 1). They do, however, co-purify during the Ni-NTA capture step. Traditional protein assays (e.g., BCA, Lowry, A280) performed at this point would overestimate the yield of the target protein (labeled Pvs25H in Fig. 1), since these other methods are incapable of distinguishing between the two forms of Pvs25H that are produced. Densitometry makes this distinction, with the added benefit that no capture step is required before an estimate of yield is given. 9. When analyzing a sample that is not yet purified, determining the baseline is usually more difficult than when working with purified material. Each sample must be analyzed on a case-by-case basis as appropriate. Acknowledgments The authors wish to thank Anthony W. Stowers, Richard L. Shimp, Vu Nguyen, Jacqueline Glen, Brian Henderson, and Jin Wang for their contributions in producing and purifying the recombinant Pvs25H protein, and Holly McClellan for expert technical assistance.

References 1. Vuletich, J. L. and Osawa, Y. (1998) Chemiluminescence assay for oxidatively modified myoglobin. Analyt. Biochem. 265, 375–380. 2. Morçöl, T. and Subramanian, A. (1999) A red-dot-blot protein assay technique in the low nanogram range. Analyt. Biochem. 270, 75–82. 3. Zhu, D., Saul, A., and Miles, A. P. (2005) A quantitative slot blot assay for host cell protein impurities in recombinant proteins expressed in E. coli. J. Immunolog. Methods 306, 40–50. 4. Miles, A. P., Zhu, D., and Saul, A. (2005) Determining residual host cell antigen levels in purified recombinant proteins by slot blot and scanning laser densitometry, in Therapeutic Proteins: Methods and Protocols (C. Mark Smales and David C. James, eds.) Humana, Totowa, NJ, pp. 233–242. 5. Westfall, D. A., Flores, R. R., Negrete, G. R., Martinez, A. O., and Haro, L.S. (1998) High-resolution polyacrylamide gel electrophoresis of carbohydrates derivatized with a visible dye. Analyt. Biochem. 265, 232–237. 6. Khawar, S. L., Watson, K., and Jones, G. L. (1997) A comparative electrophoretic analysis of mammalian hair and avian feather proteins. Int. J. Biochem. Cell Biol. 29, 367–380. 7. Ikonomov, V., Melzer, H., Nenov, V., Stoicheva, A., Stiller, S., and Mann, H. (1999) Importance of sodium dodecyl sulfate pore-graduated polyacrylamide gel electro-

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Miles and Saul phoresis in the differential diagnostic of Balkan nephropathy. Artificial Organs 23, 75–80. Li, Z., McNamara, J. R., Ordovas, J. M., and Schaefer, E. J. (1994) Analysis of high density lipoproteins by a modified gradient gel electrophoresis method. J. Lipid Res. 35, 1698–1711. Crews-Oyen, A. E. and Yoshino, T. P. (1995) Schistosoma mansoni: Characterization of excretory-secretory polypeptides synthesized in vitro by daughter sporocysts. Exp. Parasitol. 80, 27–35. Aebig, J., Mullen, G. E. D., Dobrescu, G., Rausch, K., Lambert, L., Ajose-Popoola, O., Long, C. A., Saul, A., and Miles, A. P. (2007) Formulation of vaccines containing CpG oligonucleotides and alum. J. Immunolog. Methods 323. 139–146. ImageQuant User’s Guide Version 5.0. Molecular Dynamics, Inc., 1998. Hisaeda, H., Stowers, A. W., Tsuboi, T., Collins, W. E., Sattabongkot, J. S., Suwanabun, N., Torii, M., and Kaslow, D. C. (2000) Antibodies to malaria vaccine candidates Pvs25 and Pvs28 completely block the ability of Plasmodium vivax to infect mosquitoes. Infection Immunity 68, 6618–6623.

45 Rapid and Sensitive Staining of Unfixed Proteins in Polyacrylamide Gels with Nile Red Joan-Ramon Daban, Salvador Bartolomé, Antonio Bermúdez, and F. Javier Alba 1. Introduction Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) is one of the most powerful methods for protein analysis (1,2). However, the typical procedures for the detection of protein bands after SDS-PAGE, using the visible dye coomassie blue and silver staining, have several time-consuming steps and require the fixation of proteins in the gel. This chapter describes a rapid and very simple method for protein staining in SDS gels developed in our laboratory (3–5). The method is based on the fluorescent properties of the hydrophobic dye Nile red (9-diethylamino-5H-benzo[α]phenoxazine-5-one; see Fig. 1), and allows the detection of < 10 ng of unfixed protein per band about 5 min after the electrophoretic separation. Furthermore, it has been shown elsewhere (6,7) that, in contrast to the current staining methods, Nile red staining does not preclude the direct electroblotting of protein bands and does not interfere with further staining, immunodetection and sequencing (see Chapter 71). Nile red was considered a fluorescent lipid probe, because this dye shows a high fluorescence and intense blue shifts in presence of neutral lipids and lipoproteins (8). We have shown that Nile red can also interact with SDS micelles and proteins complexed with SDS (3). Nile red is nearly insoluble in water, but is soluble and shows a high increase in the fluorescence intensity in nonpolar solvents and in presence of pure SDS micelles and SDS–protein complexes. In the absence of SDS, Nile red can interact with some proteins in solution but the observed fluorescence is extremely dependent on the hydrophobic characteristics of the proteins investigated (8,9). In contrast, Nile red has similar fluorescence properties in solutions containing different kinds of proteins associated with SDS, suggesting that this detergent induces the formation of structures having From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Structure of the noncovalent hydrophobic dye Nile red.

equivalent hydrophobic properties independent of the different initial structures of native proteins (3). In agreement with this, X-ray scattering and cryoelectron microscopy results have shown that proteins having different properties adopt a uniform necklace-like structure when complexed with SDS (10). In these structures the polypeptide chain is mostly situated at the interface between the hydrocarbon core and the sulfate groups of the SDS micelles dispersed along the unfolded protein molecule. The enhancement of Nile red fluorescence observed with different SDS–protein complexes occurs at SDS concentration lower than the critical micelle concentration of this detergent in the typical Tris–glycine buffer used in SDS-PAGE (3). Thus, for Nile red staining of SDS-polyacrylamide gels (4), electrophoresis is performed in the presence of 0.05% SDS instead of the typical SDS concentration (0.1%) used in current SDS-PAGE protocols. This concentration of SDS is high enough to maintain the stability of the SDS–protein complexes in the bands, but is lower than SDS critical micelle concentration and consequently precludes the formation of pure detergent micelles in the gel (4,5). The staining of these modified gels with Nile red produces very high fluorescence intensity in the SDS–protein bands and low background fluorescence (see Fig. 2). Furthermore, under these conditions (see details in Subheading 3.), most of the proteins separated in SDS gels show similar values of the fluorescence intensity per unit mass. A review about the physicochemical basis of Nile red staining has been published elsewhere (11). 2. Materials All solutions should be prepared using electrophoresis-grade reagents and deionized water and stored at room temperature (exceptions are indicated). Wear gloves to handle all reagents and solutions and do not pipette by mouth. Collect and dispose all waste according to good laboratory practice and waste disposal regulations. 1. Nile red. Concentrated stock (0.4 mg/mL) in dimethyl sulfoxide (DMSO). This solution is stable for at least 3 months when stored at room temperature in a glass bottle wrapped in aluminum foil to prevent damage by light. Handle this solution

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Fig. 2. Example of Nile red staining of different proteins and peptides in 0.05% SDS–15% polyacrylamide gels. The protein molecular weight markers (lane 6, from top to bottom: BSA, ovalbumin, glyceraldehyde-3-phosphate dehydrogenase, trypsinogen, and lysozyme), and BSA digested with increasing amounts of trypsin (lanes 1–5), were stained for 5 min with a solution of Nile red in water prepared by quick dilution of a stock solution of this dye in DMSO (see Subheading 3.).

2. 3.

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with care, DMSO is flammable, and, in addition, this solvent may facilitate the passage of water-insoluble and potentially hazardous chemicals such Nile red through the skin. Nile red can be obtained from Sigma Chemical (St. Louis, MO). The acrylamide stock solution and the resolving and stacking gel buffers are prepared as described in Chapter 21. 2× Sample buffer: 4% (w/v) SDS, 20% (v/v) glycerol, 10% (v/v) 2-mercaptoethanol, 0.125 M Tris-HCl, pH 6.8; bromophenol blue (0.05% [w/v]) can be added as tracking dye. 10× Electrophoresis buffer: 0. 5% (w/v) SDS, 0.25 M Tris, 1.92 M glycine, pH 8.3 (do not adjust the pH of this solution). Plastic boxes for gel staining. Use opaque polypropylene containers (e.g., 21 × 20 × 7.5 and 12 × 7.5 × 7 cm for large [20 × 16 × 0.15 cm] and small [8 × 6 × 0.075 cm] gels, respectively) with a close-fitting lid to allow intense agitation without spilling the staining solution. Orbital shaker for the agitation of the plastic boxes during gel staining. Transilluminator equipped with midrange ultraviolet (UV) bulbs (∼300 nm) to excite Nile red (3,4). A transilluminator with a cooling fan (e.g., Foto UV 300 [Fotodyne Inc., Harland, WI] or similar) is very convenient to prevent thermal

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Daban et al. damage of the gel when long exposures are necessary (see Subheading 3., step 11). UV light is dangerous to skin and particularly to eyes. UV-blocking goggles and a full face shield, and protective gloves and clothing, should be worn when the stained gel is examined using the transilluminator. For the photography, we place the transilluminator with the gel and the photographic camera inside a home-made cabinet with opaque (UV-blocking) curtains to prevent operator exposure to UV light. Alternatively, the gel can be transilluminated inside a compact darkroom of a documentation system provided with a charge-coupled device (CCD) camera (e.g., Gel Doc 1000 from Bio-Rad Laboratories [Hercules, CA]). To avoid the problems associated with UV light, we have constructed a transilluminator that works in the visible region (∼540 nm; green-light) and can also be used for the excitation of Nile red stained bands (12). Photographic camera (e.g., Polaroid [Cambridge, MA] MP-4 camera) or a CCD camera. Optical filters. With the Polaroid camera and UV transilluminator use the Wratten (Kodak [Rochester, NY]) filters number 9 (yellow) and 16 (orange) to eliminate the UV and visible light from the transilluminator. Place the two filters together in the filter holder of the camera so that filter 16 is on top of filter 9 (i.e., filter 16 should be facing towards the camera lens). Store the filters in the dark and protect them from heat, intense light sources and humidity. With the CCD camera and UV transilluminator we have used the filter 590DF100 (Bio-Rad). Wratten filter number 26 (red) is required for both Polaroid and CCD cameras when the bands are visualized with the green-light transilluminator. Photographic films. The following Polaroid instant films can be used for the photography of the red bands (maximum emission at ∼640 nm [3]) seen after staining: 667 (ISO 3000, panchromatic, black-and-white positive film), 665 (ISO 80, panchromatic, black-and-white positive/negative film), and 669 (ISO 80, color, positive film). Store the films at 4°C. Negative-clearing solution: 18% (w/v) sodium sulfite. Wetting agent for Polaroid 665 negatives (Kodak Photo-Flo diluted at 1:600). Different densitometers available commercially can be used for the densitometry of negative films. Alternatively, the images obtained with a CCD camera can be directly analyzed with the appropriate software (e.g., Molecular Analyst [Bio-Rad]).

3. Method The method described in this part gives all the details for the staining SDS-polyacrylamide gels with Nile red. See Note 1 for Nile red staining of gels without SDS. Unless otherwise indicated, all operations are performed at room temperature. 1. Typically we prepare 15% acrylamide–0.4% bis-acrylamide separating gel containing 0.05% SDS (see Note 2), 0.375 M Tris-HCl, pH 8.8. The stacking gel contains 6% acrylamide–0.16% bis-acrylamide, 0.05% SDS (see Note 2), 0.125 M Tris-HCl, pH 6.8 2. Dissolve the proteins in water, add one volume of the 2× sample buffer (Subheading 2.3., see Note 3), and incubate the resulting samples in a boiling-water bath for

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3 min. The samples are kept at room temperature before loading them into the gel (see Note 4). Place the gel sandwiched between the glass plates in the electrophoresis apparatus, fill the electrode reservoirs with 1 × electrophoresis buffer (see Note 2), and rinse the wells of the gel with this buffer. Load the protein samples (about 20 and 10 µL in large and small gels, respectively), carry out electrophoresis, and at the end of the run remove the gel sandwich from the electrophoresis apparatus and place it on a flat surface. Wearing gloves, remove the upper glass plate (use a spatula) and excise the stacking gel and the bottom part of the separating gel (see Note 5). Add quickly 2.5 mL of the concentrated Nile red (0.4 mg/mL) staining solution in DMSO to 500 mL of deionized water previously placed in a plastic box (see Note 6). These volumes are required for staining large gels; for small gels, use 0.25 mL of the concentrated Nile red solution in DMSO and 50 mL of water. Immediately after the addition of concentrated solution of Nile red to water, agitate the resulting solution vigorously for 3 s (see Note 6). Immerse the gel very quickly in the staining solution, put the lid on, and agitate vigorously (see Note 6) using an orbital shaker (at about 150 rpm) for 5 min (see Note 7). Discard the staining solution and rinse the gel with deionized water (4×; about 10 s) to remove completely the excess Nile red precipitated during the staining of the gel (see Note 8). Wearing gloves, remove the gel from the plastic box and place it on the UV or green-light transilluminator. Turn off the room lights, turn on the transilluminator and examine the protein bands, which fluoresces light red (see Note 9). Turn off the transilluminator immediately after the visualization of the bands (see Note 10). Focus the camera (Polaroid or CCD) with the help of lateral illumination with a white lamp, place the optical filters described in Subheading 2., item 9, in front of the camera lens and, in the dark, turn on the transilluminator and photograph the gel (see Note 10). Finally, turn off the transilluminator. Develop the different Polaroid films for the time indicated by the manufacturer (see Note 11). Spread the Polaroid print coater on the surface of the 665 positive immediately after development. (The positive prints of the 667 and 669 films do not require coating.) The 665 negatives can be stored temporary in water but, before definitive storage, immerse the negatives in 18% sodium sulfite and agitate gently for about 1 min, wash in running water for 5 min, dip in a solution of wetting agent (about 5 s), and finally dry in air (see Note 12). Scan the photographic negatives to determine the intensity of the Nile red stained protein bands if a quantitative analysis is required. The quantitative analysis can also be performed directly from the image obtained with a CCD documentation system (see Note 13).

4. Notes 1. Isoelectric focusing gels do not contain SDS and should be treated with this detergent after the electrophoretic run to generate the hydrophobic SDS–protein complexes specifically stained by Nile red (6). In general, for systems

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Daban et al. without SDS we recommend an extensive gel washing (20 min in the case of 0.75-mm thick isoelectric focusing gels) with 0.05% SDS, 0.025 M Tris, 0.192 M glycine, pH 8.3, after electrophoresis. The gel equilibrated in this buffer can be stained with Nile red following steps 6–9 of Subheading 3. To reduce the background fluorescence after the staining with Nile red it is necessary to preclude the formation of pure SDS micelles in the gel (see Subheading 1.). Thus, use 0.05% SDS to prepare both the separating and stacking gels and the electrophoresis buffer. This concentration is lower than the critical micelle concentration of this detergent (∼0.1% [3]), but is high enough to allow the formation of the normal SDS–protein complexes that are specifically stained by Nile red (4). Use 2% SDS in the sample buffer to be sure that all protein samples are completely saturated with SDS. Lower concentrations of SDS in the sample buffer can produce only a partial saturation of proteins (in particular in highly concentrated samples), and consequently the electrophoretic bands could have anomalous electrophoretic mobilities. The excess SDS (uncomplexed by proteins) present in the sample buffer migrates faster than the proteins and form a broad band at the bottom of the gel (see Note 5 and ref. 4). Storage of protein samples prepared as indicated in Subheading 3.2. at 4°C (or at lower temperatures) causes the precipitation of the SDS present in the solution. These samples should be incubated in a boiling-water bath to redissolve SDS before using them for electrophoresis. The bottom part of the gel (i.e., from about 0.5 cm above the bromophenol blue band to the end) should be excised before the staining of the gel. Otherwise, after the addition of Nile red, the lower part of the gel produces a broad band with intense fluorescence. This band is presumably caused by the association of Nile red with the excess SDS used in the sample buffer (see Note 3). In the case of long runs, bromophenol blue and the excess SDS band diffuse into the buffer of the lower reservoir and it is not necessary to excise the gel bottom. Nile red is very stable when dissolved in DMSO (see Subheading 2.1.), but this dye precipitates in aqueous solutions. Since the precipitation of Nile red in water is a rapid process and this dye is only active for the staining of SDS–protein bands before it is completely precipitated (4,5), to obtain satisfactory results, it is necessary: (a) to perform all the agitations indicated in these steps (in order to favor as much as possible the dispersion of the dye); (b) to work very rapidly in the steps 6–9 of Subheading 3. Furthermore, to obtain a homogeneous staining the gel has to be completely covered with the staining solution. The staining time is the same for large and small gels. After the water rinsing indicated in the step 9 of Subheading 3., the staining process is completely finished and the gel can be photographed immedi-

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ately. It is not necessary, however, to examine and photograph the gel just after staining. Nile red stained bands are stable and the gel can be kept in the plastic box immersed in water for 1–2 h before photography. About 100 ng of protein per band can be seen by naked eye (when using the green-light transilluminator it is necessary to place a Wratten filter number 26 in front of the eyes to visualize the bands). Faint bands that are not visible by direct observation of the transilluminated gel can be clearly seen in the photographic image. Very faint bands (containing as little as 10 ng of protein) can be detected by long exposure using Polaroid film. Long integration times with a CCD camera allow the detection of 5 ng of protein per band (using both UV and green-light transilluminators). To obtain this high sensitivity it is necessary to have sharp bands; broad bands reduce considerably the sensitivity. Nile red is sensitive to intense UV irradiation (3), but has a photochemical stability high enough to allow gel staining without being necessary to introduce complex precautions in the protocol (see Subheading 3., steps 6–9). For long-term storage, solutions containing this dye are kept in the dark. Transillumination of the gel for more than a few minutes produces a significant loss of fluorescence intensity. Thus, transillumination time must be reduced as much as possible both during visualization and photography. The relative short exposure times (2–12 s) required for the Polaroid film 667 and the CCD documentation system allow to make several photographs with different exposure times if necessary. With films (Polaroid 665 and 669) requiring longer exposures (4–5 min) only the first photograph from each gel shows the maximum intensity in the fluorescent bands. The development time of Polaroid films is dependent on the film temperature. Store the films at 4°C (see Subheading 2., item 10), but allow them to equilibrate at room temperature before use. The relatively large area (7.3 × 9.5 cm) of the Polaroid 665 negative is very convenient for further densitometric measurements (see Note 13). Furthermore, this negative can be used for making prints and digital images with an adequate level of contrast (see, e.g., the photograph presented in Fig. 2). The images obtained with the CCD documentation system can be stored in the computer and printed using different printers (inkjet printers with glossy paper yield photo quality images). In quantitative analyses care has to be taken to ensure that the photographic film or the CCD documentation system have a linear response for the amounts of protein under study. Use different amounts (in the same range as the analyzed protein) of an internal standard to obtain an exposure time producing a linear response. Furthermore, the internal standard is necessary to normalize the results obtained with different gels, under different electrophoretic conditions and with different exposure and development times.

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Daban et al. Nile red can be considered as a general stain for proteins separated in SDS gels (4). However, proteins with prosthetic groups such as catalase, and proteins having anomalous SDS binding properties such as histone H5, show atypical values of fluorescence intensity after staining with Nile red (4).

Acknowledgment This work was supported in part by grant BFU2005–3883 (Ministerio de Educatión y Ciencia).

References 1. Pennington, S. R. and Dunn, M. J. (2001) Proteomics: From Protein Sequence to Function. Bios Scientific Publishers, Oxford, UK. 2 Patton, W. F. (2000) A thousand points of light: The application of fluorescence detection technologies to two-dimensional gel electrophoresis and proteomics. Electrophoresis 21, 1123–1144. 3. Daban, J.-R., Samsó, M., and Bartolomé, S. (1991) Use of Nile red as a fluorescent probe for the study of the hydrophobic properties of protein–sodium dodecyl sulfate complexes in solution. Analyt. Biochem. 199, 162–168. 4. Daban, J.-R., Bartolomé, S., and Samsó, M. (1991) Use of the hydrophobic probe Nile red for the fluorescent staining of protein bands in sodium dodecyl sulfatepolyacrylamide gels. Analyt. Biochem. 199, 169–174. 5. Alba, F. J., Bermúdez, A., Bartolomé, S., and Daban, J.-R. (1996) Detection of five nanograms of protein by two-minute Nile red staining of unfixed SDS gels. BioTechniques. 21, 625–626. 6. Bermúdez, A., Daban, J.-R., Garcia, J. R., and Mendez, E. (1994) Direct blotting, sequencing and immunodetection of proteins after five-minute staining of SDS and SDS-treated IEF gels with Nile red. BioTechniques. 16, 621–624. 7. Alba, F. J. and Daban, J.-R. (1998) Rapid fluorescent monitoring of total protein patterns on sodium dodecyl sulfate-polyacrylamide gels and Western blots before immunodetection and sequencing. Electrophoresis 19, 2407–2411. 8. Greenspan, P. and Fowler, S. D. (1985) Spectrofluorometric studies of the lipid probe Nile red. J. Lipid Res. 26, 781–789. 9. Sackett, D. L. and Wolff, J. (1987) Nile red as polarity-sensitive fluorescent probe of hydrophobic protein surfaces. Analyt. Biochem. 167, 228–234. 10. Sansó, M., Daban, J.-R., Hansen, S., and Jones, G. R. (1995) Evidence for sodium dodecyl sulfate/protein complexes adopting a necklace structure. Eur. J. Biochem. 232, 818–824. 11. Daban, J.-R. (2001) Fluorescent labeling of proteins with Nile red and 2-methoxy2,4-diphenyl-3(2H)-furanone: Physicochemical basis and application to the rapid staining of sodium dodecyl sulfate polyacrylamide gels and Western blots. Electrophoresis 22, 874–880. 12. Alba, F. J., Bermúdez, A., and Daban, J.-R. (2001) Green-light transilluminator for the detection without photodamage of proteins and DNA labeled with different fluorescent dyes. Electrophoresis 22, 399–403.

46 Zn2+-Reverse S taining Technique Carlos Fernandez-Patron 1. Introduction With the advent of “proteomics,” many previously unidentified proteins will be isolated in gels for subsequent structural and functional characterization. It is, therefore, important that methods are available for detecting these proteins with minimal risk of modification. This chapter describes a “reverse” staining technique that facilitates the sensitive detection of unmodified proteins (1–9). The Zn2+ reverse staining technique exploits the ability of biopolymers (in particular, proteins and protein-SDS complexes) to bind Zn2+ (9–11) and that of imidazole (ImH, C3N3H4, see Note 1) to react with unbound Zn2+ to produce insoluble zinc imidazolate (ZnIm2). Deposition of ZnIm2 along the gel surface results in the formation of a deep white stained background, against which unstained biopolymer bands contrast (as biopolymer bands bind Zn2+, they locally inhibit deposition of ZnIm2 and are not stained; see Notes 2 and 3). With regard to protein analysis, reverse staining is a very gentle method of detection that does not require strong acid solutions, organic dyes, chemical modifiers or protein sensitizers. Protein interactions with Zn2+, established during the reverse staining step, are abrogated by metal chelation. As such, the risk of protein modification during reverse staining is minimal and reverse stained proteins can be efficiently eluted and used in biological and enzymatic assays (see Note 4 and refs. 7–9, 15). Proteins can also be processed for microsequencing or mass spectrometric analysis at any time after detection (see Note 5 and refs. 5–9, 13, 14). Another benefit of the reverse staining technique is its high sensitivity of detection (~ 10 ng protein / band in SDS-PAGE gels), which is greater than that of the Coomassie blue stain (~100 ng protein / band) and approaches the

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sensitivity of the silver stains (1–10 ng protein / band). Indeed, reverse staining often reveals many proteins that display low affinity for Coomassie blue and are thus not detected with this stain (see Note 6 and ref. 11). Other advantages of the reverse staining technique include the speed of procedure, as it consumes ~15 min, thus being significantly faster than the Coomassie blue (> 1 hr) and silver (> 2 hr) stains. The reverse stained gels can be kept in water for several hours to years without loss of image or sensitivity of detection. Similar to other stains, reverse stained patterns can be analyzed densitometrically, and a gel “toning” procedure has been recently developed to preserve the image upon gel drying (12). The Zn2+ reverse staining technique can be applied to detect virtually any gelseparated biopolymer that binds Zn2+ (i.e., proteins/peptides, glycolipids, oligonucleotides, and their multimolecular complexes) (ref. 9 and references therein). Moreover, these biopolymers are detected regardless of the electrophoresis system used for their separation (see Notes 7, 8, and 9 and ref. 9), which was not possible with the previously developed metal salt stains (1–3). Therefore, Zn2+ reverse staining is a widely applicable detection method. The SDS-PAGE method of protein electrophoresis is probably the most popular. Thus, the author chose to describe a standard reverse staining method that works very well with SDS-PAGE. The detection of proteins, peptides and their complexes with glycolipids in less commonly used gel electrophoresis systems is addressed as well (see Notes 7 to 9). 2. Materials Reagent- and analytical-grade zinc sulfate, imidazole, acetic acid, sodium carbonate are obtained from Sigma (St. Louis, MO). 1. Equilibration solution (1x): 0.2 M imidazole, 0.1% (w/v) SDS (see Note 1). 2. Developer (1x): 0.3 M zinc sulfate. 3. Storage solution for reverse stained gels (1x): 0.5 % (w/v) sodium carbonate.

All solutions are prepared as 10 times concentrated stocks, stored at room temperature, and diluted (1:10) in distilled water, to yield the working concentration (1x), just before use. 3. Methods 3.1. Polyacrylamide gel electrophoresis SDS-PAGE is conducted following the method of Laemmli (16). Native PAGE is conducted following the protocol of Laemmli, except that SDS is not included in the gel and electrophoresis solutions. Conventional agarose gel electrophoresis is conducted in 0.8% agarose gels and using Tris-acetate, pH 8.0; as gel and running buffer (17).

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3.2. Standard Reverse Staining of SDS-PAGE and Native PAGE Gels The following reverse staining method detects proteins in standard polyacrylamide gels (see Note 2, Fig. 1). All incubations are performed under continuous gentle agitation in a plastic or glass tray with a transparent bottom. The volume of the corresponding staining/storage solutions must be enough to cover the gel (typically 50 ml for 1 minigel (10 cm × 7 cm × 0.75 mm) ). 1. Following electrophoresis, the gel is incubated for 15 min in the equilibration solution (see Note 2). 2. To develop the electropherogram, the imidazole solution is discarded and the gel soaked for 30–40 s in developer solution. Caution!: This step must not be extended longer than 45 s or band overstaining and loss of the image will occur. Overstaining is prevented by pouring off the developer solution and rinsing the gel 3–5 times ( 1 min) in excess water (see Note 3).

At this point, the reverse stained gel can be photographed (Fig. 1). Photographic recording is best conducted with the gel placed on a glass plate held a few centimeters above a black underground and under lateral illumination. While SDS-PAGE is a popular, high-resolution method for separating complex protein mixtures, sometimes it is desired to avoid either protein denaturation or disruption of macromolecular complexes during electrophoresis. In this case, the proteins are separated in the absence of SDS (native PAGE or agarose gel electrophoresis). Therefore, it is desirable to avoid the use of SDS during the reverse 2D-PAGE gel 116 kDa -

14 kDa pH: 3-10

Fig. 1. Reverse staining of rat brain homogenate proteins (70 µg load) after 2D-PAGE. Proteins in a wide range of molecular weights and isoelectric points are detected as transparent spots. These spots contain protein-SDS complexes that bind Zn2+ and thereby inhibit the precipitation of ZnIm2 locally.

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native PAGE gel

2M -

M2500 1250 625

312

156

78 39 ng

Fig. 2. Reverse staining of serial dilutions of human serum albumin (HSA) after native PAGE. HSA migrates yielding two main bands corresponding to its monomer (M) and its dimer (2 M) and is detected under native conditions (no SDS) due to its natural ability to complex with Zn2+.

Agarose gel Coomassie blue

Reverse stain

(-)

(+) Peptide : glycolipid 2.5 (molar ratio)

5

10

50

0

2.5

5

10

50

0

Fig. 3. Reverse staining of complexes between a synthetic cationic peptide with lipopolysaccharide binding properties and the glycolipid of a N. meningitidis strain. Formation of complexes was promoted by incubating (37°C) peptide and glycolipid for 30 min in 25 mM Tris-HCl, 0.1% Triton X-100, pH 8.0; at indicated peptide:glycolipid molar ratios. Reverse staining revealed bands of both: peptide.glycolipid complexes and uncomplexed glycolipid. Coomassie blue stained mainly the uncomplexed peptide migrating towards the negative (−) electrode. Coomassie blue failed to stain the glycolipid bands and stained peptide.glycolipid complexes very weakly.

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staining step. Two procedures have been developed to resolve this problem (see Notes 8 and 9, Figs. 2 and 3). 4. Notes 1. Imidazole is a five-membered heterocyclic ring containing a tertiary (“pyridine”) nitrogen at position 3, and a sondary (“pyrrole”) nitrogen at position 1. It is a monoacidic base whose basic nature is due to the ability of pyridine nitrogen to accept a proton. The pyrrole nitrogen can lose the hydrogen atom producing imidazolate anion (Im−), at high pH values (pKa ~14.2). However, deprotonation of imidazole’s pyrrolic nitrogen may also occur at lower pH values, upon complexation of Zn2+ at the pyridinium nitrogen (ref. 9 and references therein). As a result, ZnIm2 can form and precipitate at pH > 6.2. Upon treatment of a polyacrylamide or agarose electrophoresis gel with salts of zinc(II) and imidazole, a complex system is generated. Complexity is due to the presence of amide groups in the polyacrylamide matrix and sulfate groups in the agarose gel as well as Tris, glycine and dodecylsulfate, hydroxyle and carbonate anions in the electrophoresis buffers. These groups can coordinate with Zn(II), act as counteranions in the complexes of zinc with imidazole, and lead to the formation of complex salts and hydroxides. Diffusion phenomena (reflected in the times required for optimal gel “equilibration” and “development” during reverse staining) also critically influence the reverse staining reactions between Zn2+ and imidazole (see Notes 2 and 3). Nevertheless, when the protocol described in this chapter is followed, ZnIm2 is the major component of the precipitate that stains the gels treated with zinc sulfate and imidazole (9). 2. This reverse staining protocol is optimized for use with any standard PAGE system regardless of whether PAGE is the first (1D-PAGE) or the sond (2D-PAGE) dimension in the separation strategy. The equilibration step assures that proteins in the gel are all uniformly coated with SDS. Therefore, protein’s ability to bind Zn2+ is modulated by the protein-SDS complex and limits of detection ( 10 ng protein / band) are similar for gels cast with and without SDS (i.e., SDS-PAGE and native PAGE, respectively). The larger the gel thickness or acrylamide concentration, the longer the equilibration step. A 15 min long equilibration is enough for gels with ≤ 15% acrylamide and ≤ 1 mm thickness. Preparative gels are often as thick as 3–5 mm; these gels should be equilibrated for 30–60 min. Insufficient equilibration may result in faint reverse stained patterns, which may fade upon prolonged storage. 3. Development time must be between 30 and 45 s as insufficient development results in pale background staining and over-development causes overstaining. An overstained gel can be re-stained. For this, the gel is treated with 10 mM EDTA or 100 mM glycine for 5–10 min to redissolve the

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white ZnIm2 precipitate that has deposited on the gel surface, rinsed in water (30 s) and, finally, soaked in the “storage” sodium carbonate solution. If the reverse stained pattern does not restore in ~5 min, the reverse staining procedure can be repeated as indicated in Methods. Usually, the reverse stained pattern will restore during the equilibration step due to the precipitation of traces of Zn2+ already present in the gel with the imidazole from the equilibration solution. If the above suggestions do not lead to a homogenous reverse stained pattern of suitable quality, the gel can be “positively” restained with Coomassie blue or silver (5). In this case, it is recommended that the reverse-stained gel be treated with EDTA (50 mM, pH 8.0; 30 min incubation) to free proteins of Zn2+. Then, the gel can be processed with Coomassie blue or silver stains. 4. Protein elution from reverse stained gels can be performed following any conventional method such as electroelution or passive elution; however, a highly efficient procedure has been developed (7, 8). The reverse stained band of interest is excised, placed in a (1 ml) plastic vial and incubated with EDTA (50 mM, 2 × 5 min and 10 mM, 1 × 5 min) to chelate protein bound zinc ions. Supplementation of EDTA with non-ionic detergents is optional but convenient if protein is to be in-gel refolded; e.g., Triton X-100 was useful when refolding proteins that were to be bioassayed (9, 15). EDTA (or EDTA, detergent mixture) is replaced by an appropriate “assay” buffer (e.g., phosphate saline solution), in which the protein band is equilibrated. Finally, the band is homogenized and protein is eluted into an appropriate volume of the assay buffer (7, 8, 14). The slurry is centrifuged and the supernatant filtered to collect a clean, transparent solution of protein ready for subsequent analysis (7–9, 14, 15). 5. An important application of reverse staining is in structural/functional proteomics (13, 14). Identification of proteins separated by electrophoresis is a prerequisite to the construction of protein databases in proteome projects. Matrix assisted laser desorption/ionization-mass spectrometry (MALDIMS) has a sensitivity for peptide detection in the lower fmol range. In principle, this sensitivity should be sufficient for an analysis of small amounts of proteins in silver-stained gels. However, a variety of known factors (e.g., chemical sensitizers such as glutaraldehyde) and other unknown factors modify the silver-stained proteins, leading to low sequence coverage (13). Low-abundance proteins have been successfully identified after ZnIm2reverse-staining (13, 14). 6. Reverse staining of gels that have been already stained with Coomassie blue reveals proteins undetected with Coomassie blue, thus improving detection. Double staining can enhance sensitivity of detection in silverstained gels as well. Before subjecting a Coomassie (or silver)-stained

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gel to double staining, the gel should be rinsed in water (2–3 × 10 min). This step assures a substantial removal of acetic acid from Coomassie and silver stains. Then, the gel is subjected to the standard reverse staining protocol. The resultant double stained patterns consist of the previously seen Coomassie blue (brown, in the case of silver)–stained bands and new unstained (reverse-stained) bands that contrast against the deep-white, ZnIm2, stained background. 7. Reverse staining can be used in combination with other “specific” protein stains to indicate the presence of both protein and glycolipid. As with proteins, the biological properties of the presumed glycolipid bands are testable in a functional experiment following elution from the reverse stained gel (9, 18). After the electrophoresis step, the gel is rinsed for 30 min (2x, 15 min) in aqueous solution of methanol (40%, v/v). This step presumably removes excess SDS from glycolipid molecules that co-migrate with the proteins in SDS-PAGE, making the glycolipid adopt a conformation with high affinity for Zn2+. Next, the gel is reverse stained (Methods). Similar to proteins, glycolipid bands show up transparent and unstained. Sensitivity of detection is ~10 ng glycolipid / band. If the same (or a parallel gel) is treated with Coomassie blue (silver) stain, that does not detect glycolipids; protein and glycolipid bands can be distinguished by their distinct staining and migration properties (9, 18). A replicate protein sample treated with proteinase K is a useful control, as this protease digests protein, but not glycolipid. 8. An alternative method of precipitating ZnIm2 facilitates the reverse staining of native PAGE gels without the use of SDS (9). A quick and a slow reverse staining procedure were implemented (9). The quick version makes use of the characteristic neutral to basic pH of the gels immediately after electrophoresis. When the gel is incubated (3–6 min) in an slightly acidic solution containing zinc sulfate and imidazole (pre-mixed to yield the molar ratio Zn(II) : ImH = 15 mM : 30 mM, pH 5.0 – 5.5), the gel stains negatively as ZnIm2 deposits along its surface. Blotchy deposits are prevented by intense agitation and resolubilized by lowering the pH of the zinc-imidazole solution; this also provides a basis for a slow version of this method. In the slow version (Fig. 2), the gel is incubated (10–20 min) in a solution of zinc sulfate (15 mM) and imidazole (30 mM) adjusted to pH 4.0. No precipitation occurs at this pH. The solution is poured off, the gel is rinsed with water (30 s); the electropherogram is developed by incubating the gel during ~5 min in 1% sodium carbonate. Sodium carbonate increases the pH first at the gel surface. Therefore, Zn2+ and imidazole, already present in the gel, react at the gel surface to form ZnIm2. Again, protein bands are not stained as they complex with Zn2+ and locally prevent the precipitation of ZnIm2.

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9. Many proteins and peptides as well as glycolipids and their complexes with certain proteins/peptides separate well on agarose gels under nondenaturing conditions (9). To reverse stain an agarose gel (9); following electrophoresis, the gel is incubated for 25 – 30 min in a zinc sulfateimidazole solution (Zn(II) : ImH = 15 mM: 30 mM, adjusted to pH 5.0 with glacial acetic acid). During this step, Zn2+ and imidazole diffuse into agarose matrix. The gel is then rinsed in water for 5–8 min to remove any excess of the staining reagents from the gel surface. The reverse stained electropherogram is developed by incubating the gel for 5–8 minutes in 1% Na2CO3 (Fig. 3). Of note, due to poorly understood factors, agarose gels are not as amenable to reverse staining as polyacrylamide gels. Patterns of positive and negative bands are often seen in agarose gels. Acknowledgments The author expresses his gratitude to his former colleagues, Drs. L. Castellanos-Serra and E. Hardy from the Centre for Genetic Engineering and Biotechnology (CIGB), Havana, Cuba for their contributions to the development of the reverse staining concept, and to the CIGB for early support. Currently, the author is a New Investigator of the Canadian Institutes of Health Research and the Heart and Stroke Foundation of Canada as well as funded by grants from Natural Sciences and Engineering Council and the Canadian Institutes of Health Research.

References 1. Lee, C., Levin, A., and Branton, D. (1987) Copper staining: a five-minute protein stain for sodium dodecyl sulfate-polyacrylamide gels. Analyt. Biochem. 166, 308–312. 2. Dzandu, J. K., Johnson, J. F., and Wise, G. E. (1988) Sodium dodecyl sulfate-gel electrophoresis: staining of polypeptides using heavy metal salts. Analyt. Biochem. 174, 157–167. 3. Adams, L. D. and Weaver, K. M. (1990) Detection and recovery of proteins from gels following zinc chloride staining. Appl. Theor. Electrophor. 1, 279–282. 4. Fernandez-Patron, C. and Castellanos-Serra, L. (1990) Eight International Conference on Methods in Protein Sequence Analysis. Kiruna, Sweden, July 1–6, Abstract booklet. 5. Fernandez-Patron, C., Castellanos-Serra, L., and Rodriguez P. (1992) Reverse staining of sodium dodecyl sulfate polyacrylamide gels by imidazole-zinc salts: sensitive detection of unmodified proteins. Biotechniques 12, 564–573. 6. Fernandez-Patron, C., Calero, M., Collazo, P. R., Garcia, J. R., Madrazo, J., Musacchio, A., Soriano, F., Estrada, R., Frank, R., Castellanos-Serra, L. R., and Mendez, E. (1995) Protein reverse staining: high-efficiency microanalysis of unmodified proteins detected on electrophoresis gels. Analyt. Biochem. 224, 203–11.

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7. Castellanos-Serra, L. R., Fernandez-Patron, C., Hardy, E., and Huerta, V. (1996) A procedure for protein elution from reverse-stained polyarcylamide gels applicable at the low picomole level: An alternative route to the preparation of low abundance proteins for microanalysis. Electrophoresis 17, 1564–1572. 8. Castellanos-Serra, L. R., Fernandez-Patron, C., Hardy, E., Santana, H., and Huerta, V. (1997) High yield elution of proteins from sodium dodecyl sulfate-polyacrylamide gels at the low-picomole level: Application to N-terminal sequencing of a scarce protein and to in-solution biological activity analysis of on-gel renatured proteins. J. Protein Chem. 16, 415–419. 9. Fernandez-Patron, C., Castellanos-Serra, L., Hardy, E., Guerra, M., Estevez, E., Mehl, E., Frank, R. W. (1998) Understanding the mechanism of the zinc-ion stains of biomacromolecules in electrophoresis gels: generalization of the reverse-staining technique. Electrophoresis 19, 2398–2406. 10. Kaim, W. and Schwerderki, B. (eds.). (1994) Bioinorganic Chemistry: Inorganic Elements in the Chemistry of Life. John Wiley, Chichester-New York-BrisbaneToronto-Singapore. 11. Fernandez-Patron, C., Hardy, E., Sosa, A., Seoane, J., and Castellanos, L. (1995) Double staining of coomassie blue-stained polyacrylamide gels by imidazolesodium dodecyl sulfate-zinc reverse staining: sensitive detection of coomassie blue-undetected proteins. Analyt. Biochem. 224, 263–9. 12. Ferreras, M., Gavilanes, J. G., and Garcia-Segura, J. M. (1993) A permanent Zn2+ reverse staining method for the detection and quantification of proteins in polyacrylamide gels. Analyt. Biochem. 213, 206–12. 13. Scheler, C., Lamer, S., Pan, Z., Li, X. P., Salnikow, J., and Jungblut, P. (1998) Peptide mass fingerprint sequence coverage from differently stained proteins on two-dimensional electrophoresis patterns by matrix assisted laser desorption/ ionization-mass spectrometry (MALDI-MS). Electrophoresis 19, 918–27. 14. Castellanos-Serra, L., Proenza, W., Huerta, V., Moritz, R. L., and Simpson, R. J. (1999) Proteome analysis of polyacrylamide gel-separated proteins visualized by reversible negative staining using imidazole-zinc salts. Electrophoresis 20, 732–7. 15. Hardy, E., Santana, H., Sosa, A., Hernandez, L., Fernandez-Patron, C., and Castellanos-Serra, L. (1996) Recovery of biologically active proteins detected with imidazole-sodium dodecyl sulfate-zinc (reverse stain) on sodium dodecyl sulfate gels. Anal. Biochem. 240, 150–2. 16. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 17. Maniatis, T., Fritsch, E. F., and Sambrook, J. (eds.). (1982) Molecular Cloning: A Laboratory Manual. CHS, US. 18. Hardy, E., Pupo, E., Castellanos-Serra, L., Reyes, J., and Fernandez-Patron, C. (1997) Sensitive reverse staining of bacterial lipopolysaccharides on polyacrylamide gels by using zinc and imidazole salts. Analyt. Biochem. 244, 28–32.

47 Protein Staining with Calconcarboxylic Acid in Polyacrylamide Gels Wei-Tao Cong, Sun-Young Hwang, Li-Tai Jin, and Jung-Kap Choi 1. Introduction Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) has become a highly reliable separation technique for protein characterization. Broad application of electrophoresis techniques has required the development of detection methods that can be used to visualize the proteins separated on polyacrylamide gels. A few of these methods are Coomassie blue (CB) staining (1), silver staining (2), fluorescent staining (3–5), specific enzyme visualization (6), and radioactive detection (7). CB staining is the most commonly used method owing to its proven reliability, simplicity, and economy (8, 9), but it lacks sensitivity compared with the silver-staining method. In addition, the staining/destaining process is time consuming (10). Silver staining is the most sensitive nonradioactive protein detection method currently available, and can detect as little as 10 fg of protein (11, 12). However, it has several drawbacks, such as high-background staining, multiple steps, high cost of reagent, and toxicity of formaldehyde (10). In this chapter a protein staining method using calconcarboxylic acid (1-[2-hydroxy-4-sulfo-l-naphthylazo]-2-hydroxy-3-naphthoic acid) is described (see Note 1; Fig. 1; ref. 13). This method can be performed by both simultaneous and postelectrophoretic staining techniques. Simultaneous staining using 0.01% of NN in upper reservoir buffer eliminates the poststaining step, and thus enables detection of the proteins more rapidly and simply. In poststaining, proteins can be stained by a 30 min incubation of a gel in 40% methanol/7% acetic acid solution of 0.05% NN and destaining in 40% methanol/7% acetic acid for 40 min with agitation. NN staining can detect as little as 10 ng of bovine serum albumin (BSA) by poststaining and 25 ng by simultaneous staining, compared to 100 ng detectable by CB poststaining (see Note 2). These techniques produce From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Structure of NN.

protein-staining patterns identical to the ones obtained by the conventional CB staining and also work well in nondenaturing PAGE, like in SDS-PAGE. The bands stained with NN present purple color. In addition, NN staining gives better linearity than CB staining, although the slopes (band intensity/amount of protein) of it are somewhat lower (see Table 1, Fig. 2). It suggests that NN staining is more useful than CB staining for quantitative work of proteins. 2. Materials 1. All working solutions should be freshly prepared with distilled water and clean glassware or plastic ware. All steps were carried out at room temperature with shaking. 2. Destaining solution (1.0 L): Mix 530 mL distilled water with 400 mL methanol and 70 mL glacial acetic acid. 3. Simultaneous staining solution (1.0% [w/v] NN): Dissolve 1.0 g NN (pure *NN without K2SO4) in 100 mL reservoir buffer. Stir until fully dissolved at 50–60°C (stable for months at 4°C) (see Note 3). 4. Poststaining solution (0.05% [w/v] NN): Dissolve 0.05 g NN (pure *NN) in 100 mL destaining solution. Stir until dissolved thoroughly (store at room temperature).

3. Methods 3.1. Simultaneous Staining The method is based on the procedure of Borejdo and Flynn (14), and is described for staining proteins in a 7.5% SDS-polyacrylamide gel. 1. Electrophorese the samples for 10 min to allow protein penetration into the upper gel phase. 2. Turn off the power, and then add 1% NN dissolved in reservoir buffer to the upper reservoir buffer to give a final concentration of 0.01–0.015% NN (see Notes 3 and 4). 3. Stir the reservoir buffer sufficiently to ensure homogeneity. 4. Resume electrophoresis. 5. Immediately after electrophoresis, remove the stained gel from the apparatus. 6. Destain the stained gels in 40% methanol/7% acetic acid for 30 min. To destain completely, change destaining solution several times and agitate (see Notes 5–7). *NN diluted with 100- to 200-fold K SO has been used as an indicator for the determination of calcium 2 4 in the presence of magnesium with EDTA.

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Table 1 Linearity of CB and NN Staining for Four Purified Proteins Slopeb

y-interceptb

Correlation coefficientb

Proteina

A

B

A

B

A

B

BSA OVA G-3-P DHase CA

14.1 8.7 8.0 15.4

10.1 7.1 5.8 13.5

13.2 2.7 5.3 12.5

6.4 1.4 2.2 8.8

0.986 0.993 0.997 0.986

0.994 0.998 0.999 0.997

a Proteins were separated on 12.5% polyacrylamide gel, densities and band area were determined with computerized densitometer. Some of the data are illustrated in Fig. 2. The range of amount of proteins was 0.25–12.5 µg. The number of points measured was six (0.25, 0.5, 1.0, 2.5, 5.0, and 12.5 µg). b Slopes, y-intercepts, and correlation coefficients were determined by linear regression analysis. A, CB staining; B, NN staining.

Fig. 2. Densitometric comparison of CB and NN staining. Proteins were separated on 12.5% gels. (A) poststaining with 0.1% CB; (B) simultaneous staining with 0.015% NN. Densitometric scanning was performed at 585 nm (CB) and 580 nm (NN). The curves are fitted by the method of least squares. Each point represents the mean of three determinations. BSA, bovine serum albumin; OVA, ovalbumin; G-3-P DHase, glyceraldehyde-3-phosphate dehydrogenase; CA, carbonic anhydrase

3.2. Postelectrophoretic Staining 1. Agitate the freshly run gel in 0.05% NN staining solution for 30 min. 2. Pour off the staining solution and rinse the gel with changes of destaining solution (two to three times). Staining solution can be reused several times. 3. Destain the stained gels in 40% methanol/7% acetic acid for 40 min with agitation (see Notes 8 and 9).

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4. Notes 1. NN has several functional groups, such as hydroxyl, diazoic, carboxyl, and sulfonate groups (see Fig. 1). At acidic pH, NN probably forms electrostatic bonds with protonated amino groups, which are stabilized by hydrogen bonds and Van der Waals forces, as CB does (1). 2. The protocol for poststaining is the same as that for CB staining, except for staining/destaining times and dye used. 3. The preparation of staining solution requires stirring and warming at 50–60°C since NN is poorly soluble. 4. The simultaneous staining method allows one to control the intensity of stained bands reproducibly by adjusting the concentration of the dye in the upper reservoir. More than 0.02% NN in the upper reservoir buffer does not increase sensitivity but requires more destaining time. 5. Gel staining/destaining with NN is pH dependent. Intense staining occurs at pH 1.6–4.4, and weak staining with blue-purple color is observed at alkaline pH. In excessively strong acidic solution (pH55%), gels are opaqued and shrunken. Addtionally, increasing the temperature of destaining solution is a great help in removing background (at 60–70°C, in 5 min), although sensitivity is a little reduced.

Fig. 3. Mechanism of protein–dye interaction in acidic solution.

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7. Destaining can be completed within 30 min in 7.5% polyacrylamide gels, but destaining time should be increased for 10 and 12.5% gels (50– 60 min). 8. Bands stained with NN are indefinitely stable when gels are stored in a refrigerator wrapped up in polyethylene film or dried on Whatmann No. 1 filter paper. 9. Throughout the staining/destaining processes, it is necessary to agitate the gel container using a shaker. Acknowledgment This work was supported by the Korean Research Foundation in 2004 (Grant No. E00211).

References 1. Kang, D., Gho, Y, S., Suh, M., Kang, C. (2002) High sensitive and fast protein detection with Coomassie brilliant blue in sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Bull. Korean Chem. Soc. 23, 1511–1512. 2. Merril, C. R., Goldman, D., Sedman, S., and Ebert, M. (1980) Ultrasensitive stain for proteins in polyacrylamide gels shows regional variation in cerebrospinal fluid proteins. Science 211, 1437–1438. 3. Jakowski, G. and Liew, C. C. (1980) Fluorescamine staining of nonhistone chromatin proteins as revealed by two-dimensional polyacrylamide gel electrophoresis. Analyt. Biochem. 102, 321–325. 4. Chevalier, F., Rofidal, V., Vanova, P., Bergoin, A., Rossignol, M. (2004) Proteomic capacity of recent fluorescent dyes for protein staining. Phytochemistry 65, 1499–1506. 5. Grus, F. H., Sabuncuo, P., Augustin, A. J. (2001) Analysis of tear protein patterns of dry-eye patients using fluorescent staining dyes and two-dimensional quantification algorithms. Electrophoresis 22, 1845–1850. 6. Havemeister, W., Rehbein, H., Steinhart, H., Gonzales-Sotelo, C., KrogsgaardNielsen, M., Jorgensen, B. (1999) Visualization of the enzyme trimethylamine oxide demethylase in isoelectric focusing gels by an enzyme-specific staining method. Electrophoresis 20, 1934–1938. 7. Nakamura, H., Saito, K., Shiro, Y. (2001) Quantitative measurement of radioactive phosphorylated proteins in wet polyacrylamide gels. Analyt. Biochem. 294, 187–188. 8. Pal, J. K., Sarkar, A., Katoch, B. (2001) Detergent-mediated destaining of Coomassie Brilliant Blue-stained SDS polyacrylamide gels. Indian J. Exp. Biol. 39, 95–97. 9. Wei, Y, J., Li, K. A., Tong, S, Y. (1997) A linear regression method for the study of the Coomassie brilliant blue protein assay. Talanta 44, 923–930. 10. Hames, B. D. and Rickwood, D. (1990) Analysis of gels following electrophoresis, in Gel Electrophoresis of Proteins: A Practical Approach, IRL, Oxford, UK, pp. 52–81. 11. Jin, L, T., Hwang, S,Y., Yoo, G, S., Choi, J, K. (2006) A mass spectrometry compatible silver staining method for protein incorporating a new silver sensitizer in sodium dodecyl sulfate-polyacrylamide electrophoresis gels. Proteomics 6, 2334–2337.

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12. Ohsawa, K. and Ebata, N. (1983) Silver stain for detecting 10-femtogram quantities of protein after polyacrylamide gel electrophoresis. Analyt. Biochem. 135, 409–415. 13. Hong, H. Y., Yoo, G. S., and Choi, J. K. (1993) Detection of proteins on polyacrylamide gels using calconcarboxylic acid. Analyt. Biochem. 214, 96–99. 14. Borejdo, J. and Flynn, C. (1984) Electrophoresis in the presence of Coomassie Brilliant Blue R-250 stains polyacrylamide gels during protein fractionation. Analyt. Biochem. 140, 84–86.

48 Detection of Proteins in Polyacrylamide Gels by Silver Staining Michael J. Dunn 1. Introduction The versatility and resolving capacity of polyacrylamide gel electrophoresis has resulted in this group of methods becoming the most popular for the analysis of patterns of protein expression in a wide variety of complex systems. These techniques are often used to characterise protein purity and to monitor the various steps in a protein purification process. Moreover, two-dimensional polyacrylamide gel electrophoresis (2-DE) remains the core technology of choice for separating complex protein mixtures in the majority of proteome projects (1). This is due to its ability to separate simultaneously thousands of proteins and to indicate post-translational modifications that result in alterations in protein pI or Mr. Additional advantages are the high-sensitivity visualization of the resulting 2-DE separations, compatibility with quantitative computer analysis to detect differentially regulated proteins (2), and the relative ease with which proteins from 2-DE gels can be identified and characterised by mass spectrometry (MS) (3). After 2-DE, the separated proteins must be visualised at high sensitivity, and the detection method should ideally combine the properties of an extended dynamic range, a linear staining response, reproducibility, and compatibility with post-electrophoretic protein identification procedures, especially those based on the use of MS. Unfortunately, no current staining method can fulfil all of these requirements. Staining methods based on the use of the organic dye, Coomassie Brilliant Blue (CBB), have been used extensively for the detection of proteins on 2-DE gels, due to their low price, ease of use and compatibility with downstream

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protein identification by MS. However, methods using CBB are limited by their lack of sensitivity (around 200–500 ng protein per spot), such that only a few hundred protein spots can usually be visualised on a 2-DE gel, even if milligram amounts of protein sample are loaded (4). The need to detect small amounts of protein within 2-DE gel spots has resulted in the development of a range of more sensitive detection methods (4). The first method that was able to meet this requirement and which is still used in many proteomic projects is silver staining. The ability of silver to develop images was discovered in the mid-17th century and this property was exploited in the development of photography, followed by its use in histological procedures. Silver staining for the detection of proteins following gel electrophoresis was first reported in 1979 by Switzer et al (5), resulting in a major increase in the sensitivity of protein detection. More than 100 publications have subsequently appeared describing variations in silver staining methodology (4, 6). This group of procedures is generally accepted to be around 100 times more sensitive than methods using CBB R-250, being able to detect low ng amounts of protein per gel band or spot. All silver staining procedures depend on the reduction of ionic silver to its metallic form, but the precise mechanism involved in the staining of proteins has not been fully established. It has been proposed that silver cations complex with protein amino groups, particularly the ε-amino group of lysine (7), and with sulphur residues of cysteine and methionine (8). However, Gersten and his colleagues have shown that “stainability” cannot be attributed entirely to specific amino acids and have suggested that some element of protein structure, higher than amino acid composition, is responsible for differential silver staining (9). Silver staining procedures can be grouped into two types of method depending on the chemical state of the silver ion when used for impregnating the gel. The first group is alkaline methods based on the use of an ammoniacal silver or diamine solution, prepared by adding silver nitrate to a sodium-ammonium hydroxide mixture. Copper can be included in these diamine procedures to give increased sensitivity, possibly by a mechanism similar to that of the Biuret reaction. The silver ions complexed to proteins within the gel are subsequently developed by reduction to metallic silver with formaldehyde in an acidified environment, usually using citric acid. In the second group of methods, silver nitrate in a weakly acidic (approximately pH 6) solution is used for gel impregnation. Development is subsequently achieved by the selective reduction of ionic silver to metallic silver by formaldehyde made alkaline with either sodium carbonate or NaOH. Any free silver nitrate must be washed out

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of the gel prior to development, as precipitation of silver oxide will result in high background staining. Silver stains are normally monochromatic, resulting in a dark brown image. However, if the development time is extended, dense protein zones become saturated and colour effects can be produced. Some staining methods have been designed to enhance these colour effects, which were claimed to be related to the nature of the polypeptides detected (10). However, it has now been established that the colours produced depend on: (a) the size of the silver particles, (b) the distribution of silver particles within the gel, and (c) the refractive index of the gel (11). Rabilloud has compared several staining methods based on both the silver diamine and silver nitrate types of procedure (12). The most rapid procedures were found to be generally less sensitive than the more time-consuming methods. Methods using glutaraldehyde pretreatment of the gel and silver diamine complex as the silvering agent were found to be the most sensitive. However, it should be noted that the glutaraldehyde and formaldehyde present in many silver staining procedures results in alkylation of α- and ε-amino groups of proteins, thereby interfering with their subsequent chemical characterisation by MS. To overcome this problem, silver staining protocols compatible with mass spectrometry in which glutaraldehyde is omitted have been developed (13, 14) but these suffer from a decrease in sensitivity of staining and a tendency to a higher background. This problem can be overcome using post-electrophoretic fluorescent staining techniques (4, 15). One the most commonly used stains of this type is SYPRO Ruby (see Chapter 50), which has a sensitivity approaching that of standard silver staining and is fully compatible with protein characterisation by MS (16). The method of silver staining we describe here is recommended for analytical applications and is based on that of Hochstrasser et al (17, 18), together with modifications and technical advice that will enable an experimenter to optimise results. An example of a one-dimensional SDS-PAGE separation of the total proteins of human heart proteins stained by this procedure is shown in Fig. 1. The power of 2-DE combined with sensitive detection by silver staining to display the complex protein profile of a whole tissue lysate is shown in Fig. 2. 2. Materials 1. All solutions should be freshly prepared, and overnight storage is not recommended. Solutions must be prepared using clean glassware and deionised, distilled water. 2. Gel fixation solution: Trichloroacetic acid (TCA) solution, 20% (w/v). 3. Sensitisation solution: 10% (w/v) glutaraldehyde solution.

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Fig. 1. One-dimensional 12%T SDS-PAGE separation of human heart proteins (lanes b-g) visualised by silver staining. Lane (m) contains the Mr marker proteins and the scale at the left indicates Mr × 10−3. The sample protein loadings were (b) 1 µg, (c) 5 µg, (d) 10 µg, (e) 25 µg, (f) 50 µg, (g) 100 µg.

Fig. 2. 2-DE separation of human heart proteins visualised by silver staining. A loading of 100 µg protein was used. The first dimension was pH 3–10 NL IPG IEF and the second dimension was 12% T SDS-PAGE. The scale at the top indicates the non-linear pH gradient used in the first IPG 3–10 NL IEF dimension, while the scale at the left indicates Mr × 10−3.

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4. Silver diamine solution: 21 mL of 0.36% (w/v) NaOH is added to 1.4 mL of 35% (w/v) ammonia and then 4 mL of 20% (w/v) silver nitrate is added drop-wise with stirring. When the mixture fails to clear with the formation of a brown precipitate, further addition of a minimum amount of ammonia results in the dissolution of the precipitate. The solution is made up to 100 mL with water. The silver diamine solution is unstable and should be used within 5 min. 5. Developing solution: 2.5 mL of 1% (w/v) citric acid, 0.26 mL of 36% (w/v) formaldehyde made up to 500 mL with water. 6. Stopping solution: 40% (v/v) ethanol, 10% (v/v) acetic acid in water. 7. Farmer’s reducer: 0.3% (w/v) potassium ferricyanide, 0.6% (w/v) sodium thiosulphate, 0.1% (w/v) sodium carbonate.

3. Method Note: All incubations are carried out at room temperature with gentle agitation 3.1. Fixation 1. After electrophoresis, fix the gel immediately (see Note 1) in 200 mL (see Note 2) of TCA (see Note 3) for a minimum of 1 h at room temperature. High-percentage polyacrylamide and thick gels require an increased period for fixation, and overnight soaking is recommended. 2. Place the gel in 200 mL of 40% (v/v) ethanol, 10% (v/v) acetic acid in water and soak for 2 × 30 min (see Note 4). 3. Wash the gel in excess water for 2 × 20 min, facilitating the rehydration of the gel and the removal of methanol. An indication of rehydration is the loss of the hydrophobic nature of the gel.

3.2. Sensitisation 1. Soak the gel in a 10% (w/v) glutaraldehyde solution for 30 min at room temperature (see Note 5). 2. Wash the gel in water for 3 × 20 min to remove excess glutaraldehyde.

3.3. Staining 1. Soak the gel in the silver diamine solution for 30 min. For thick gels (1.5 mm), it is necessary to use increased volumes so that the gels are totally immersed. Caution should be exercised in disposal of the ammoniacal silver reagent, since it decomposes on standing and may become explosive. The ammoniacal silver reagent should be treated with dilute hydrochloric acid (1 N) prior to disposal. 2. Wash the gel (3 × 5 min) in water.

3.4. Development 1. Place the gel in developing solution. Proteins are visualised as dark brown zones within 10 min (see Note 6), after which the background will gradually increase (see Note 7). It is important to note that the reaction displays inertia, and that

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staining will continue for 2–4 min after removal of the gel from the developing solution. Staining times in excess of 20 min usually result in an unacceptable high background (see Note 8). 4. Terminate staining by immersing the gel in stopping solution. 5. Wash the stained gel in water prior to storage or drying.

3.5. Destaining Partial destaining of gels using Farmer’s reducing reagent is recommended for the controlled removal of background staining that obscures proper interpretation of the protein pattern. 1. Wash the stained gel in water for 5 min to remove the stop solution. 2. Place the gel in Farmer’s reducer for a time dependent upon the intensity of the background. 3. Terminate destaining by returning the gel to the stop solution.

4. Notes 1. Gloves should be worn at all stages when handling gels, since silver staining will detect keratin proteins from the skin. 2. Volumes of the solutions used at all stages should be sufficient such that the gel is totally immersed. If the volume of solution is insufficient for total immersion, staining will be uneven and the gel surface can dry out. 3. A mixture of alcohol, acetic acid and water (9:9:2) is recommended for gel fixation in many published protocols, but TCA is a better general protein fixative and its use is compatible with silver staining provided that the gel is washed well after fixation to remove the acid. 4. In addition to removing TCA, the washing step also effectively removes reagents such as Tris, glycine and detergents (especially SDS) which can bind silver and result in increased background staining. 5. Treatment of the gel with reducing agents such as glutaraldehyde prior to silver impregnation results in an increase in staining sensitivity by increasing the speed of silver reduction on the proteins. 6. If image development is allowed to proceed for too long, dense protein zones will become saturated and negative staining will occur, leading to serious problems if quantitative analysis is attempted. In addition, certain proteins stain to give yellow or red zones regardless of protein concentration, and this effect has been linked to the post-translational modification of the proteins. 7. An inherent problem with the staining of gradient SDS-PAGE gels is uneven staining along the concentration gradient. The less concentrated polyacrylamide region develops background staining prior to the more concentrated region. A partial solution to this problem is to increase the time of staining in silver diamine (see Subheading 3.3., step 1).

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8. Various chemicals used in one- and two-dimensional electrophoresis procedures can inhibit staining, whereas others impair resolution or produce artefacts. Acetic acid will inhibit staining and should be completely removed prior to the addition of silver diamine solution (Subheading 3.3., step 1). Glycerol, used to stabilise SDS gradient gels during casting, and urea, used as a denaturing agent in isoelectric focusing (IEF), are removed by water washes. Agarose, often used to embed rod IEF gels onto SDSPAGE gels in 2-D PAGE procedures, contains peptides that are detected by silver staining as diffuse bands and give a strong background. Tris, glycine and detergents (especially SDS) present in electrophoresis buffers can complex with silver and must be washed out with water prior to staining. The use of 2-mercaptoethanol as a disulphide bond reducing agent should be avoided since it leads to the appearance of two artefactual bands at 50 and 67 kDa on the gel (19). 9. Radioactively labelled proteins can be detected by silver staining prior to autoradiography or fluorography for the majority of the commonly used isotopes (14C, 35S, 32P, 125I). In the case of 3H, however, silver deposition will absorb most of the emitted radiation. Acknowledgment MJD is the recipient of a Science Foundation Ireland Research Professorship and work in his laboratory is supported by SFI under Grant No. 04/RP1/B499.

References 1. Görg, A., Weiss, W., and Dunn, M. J. (2004) Current two-dimensional electrophoresis technology for proteomics. Proteomics 4, 3665–3685. 2. Dowsey, A. W., Dunn, M. J., and Yang, G. Z. (2003) The role of bioinformatics in two-dimensional gel electrophoresis. Proteomics 3, 1567–1596. 3. Cañas, B., López-Ferrer, D., Ramos-Fernández, Camafeita, E., and Calvo, E. (2006) Mass spectrometry technologies for proteomics. Brief. Funct. Genomic. Proteomic. 4, 295–320. 4. Miller, I. Crawford, J.,and Gianazza, E. (2006) Protein stains for proteomic applications: which, when and why? Proteomics 6, 5385–5408. 5. Switzer, R. C., Merril, C. R., and Shifrin, S. (1979) A highly sensitive stain for detecting proteins and peptides in polyacrylamide gels. Analyt. Biochem. 98, 231–237. 6. Rabilloud, T. (1990) Mechanisms of protein silver staining in polyacrylamide gels: a 10-year synthesis. Electrophoresis 11, 785–794. 7. Don, A. S. and Pomenti, A. A. (1983) Ammoniacal silver staining of proteins: mechanism of glutaraldehyde enhancement. Analyt. Biochem. 129, 490–496. 8. Heukeshoven, J. and Dernick, R. (1985) Simplified method for silver staining of proteins in polyacrylamide gels and the mechanism of silver staining. Electrophoresis 6, 103–112.

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9. Gersten, D. M., Rodriguez, L. V., George, D. G., Johnston, D. A., and Zapolski, E. J. (1991) On the relationship of amino acid composition to silver staining of protein in electrophoresis gels: II. Peptide sequence analysis. Electrophoresis 12, 409–414. 10. Sammons, D. W., Adams, L. D., and Nishizawa, E. E. (1982) Ultrasensitive silver-based color staining of polypeptides in polycarylamide gels. Electrophoresis 2, 135–141. 11. Merril, C. R., Harasewych, M. G., and Harrington, M. G. (1986) Protein staining and detection methods, in Gel Electrophoresis of Proteins (Dunn, M.J., ed.), Wright, Bristol, pp. 323–362. 12. Rabilloud, T. (1992) A comparison between low background silver diamine and silver nitrate protein stains. Electrophoresis 13, 429–439. 13. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. (1996) Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Analyt. Chem. 68, 85–858. 14. Yan, J. X., Wait, R., Berkelman, T., Harry, R., Westbrook, J.A., Wheeler, C.H., and Dunn, M.J. (2000) A modified silver staining protocol for visualization of proteins compatible with matrix-assisted laser desorption/ionization and elctrospray ionization-mass spectrometry. Electrophoresis 21, 3666–3672. 15. Patton, W. F. (2000) A thousand points of light: The application of fluorescence detection technologies to two-dimensional gel electrophoresis and proteomics. Electrophoresis 21, 1123–1144. 16. Yan, J.X., Harry, R.A., Spibey, C., Dunn, M.J. (2000) Postelectrophoretic staining of proteins separated by two-dimensional gel electrophoresis using SYPRO dyes. Electrophoresis 21, 3657–3665. 17. Hochstrasser, D. F., Patchornik, A., and Merril, C. R. (1988) Development of polyacrylamide gels that improve the separation of proteins and their detection by silver staining. Analyt. Biochem. 173, 412–423. 18. Hochstrasser, D. F. and Merril, C. R. (1988) “Catalysts” for polyacrylamide gel polymerization and detection of proteins by silver staining. Appl. Theor. Electrophoresis 1, 35–40. 19. Guevarra, J., Johnston, D. A., Ramagli, L. S., Martin, B. A., Capitello, S., and Rodriguez, L. V. (1982) Quantitative aspects of silver deposition in proteins resolved in complex polyacrylamide gels. Electrophoresis 3, 197–205.

49 Background-Free Protein Detection in Polyacrylamide Gels and on Electroblots Using Transition Metal Chelate Stains Wayne F. Patton 1. Introduction Electrophoretically separated proteins may be visualized using organic dyes such as Ponceau Red, Amido Black, Fast Green, or most commonly Coomassie Brilliant Blue (1,2). Alternatively, sensitive detection methods have been devised using metal ions and colloids of gold, silver, copper, carbon, or iron (3–12). Metal chelates form a third class of stains, consisting of transition metal complexes that bind avidly to proteins resolved in polyacrylamide gels or immobilized on solid-phase membrane supports (13–27). In recent years, metal chelate stains have been designed and optimized specifically for compatibility with commonly used microchemical characterization procedures employed in proteomics. The metal chelate stains are simple to implement, and do not contain extraneous chemicals such as glutaraldehyde, formaldehyde, or Tween-20 that are well known to interfere with many downstream protein characterization procedures. Metal chelates can be used to detect proteins on nitrocellulose, poly(vinylidene difluoride) (PVDF), and nylon membranes as well as in polyacrylamide and agarose gels. The metal complexes do not modify proteins, and are compatible with immunoblotting, lectin blotting, mass spectrometry, and Edman-based protein sequencing (13–17,22–27). Metal chelate stains are suitable for routine protein measurement in solid-phase assays owing to the quantitative stoichiometry of complex formation with proteins and peptides (15,16). Such solid phase protein assays are more sensitive and resistant to chemical interference than their solution-based counterparts (15). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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A variety of metal ions and organic chelating agents may be combined to form metal chelate stains but only a few have been evaluated extensively for protein detection in electrophoresis. Ferrene S has been utilized for the specific detection of iron-containing proteins, such as cytochrome c, transferrin, ferritin, and lactoferrin in polyacrylamide gels (18). In this situation the metal complex only forms when the chelate interacts with the native metal ion bound within the protein itself. Copper phthalocyanine 4, 4' ,4", 4'" -tetrasulfonic acid has been shown to stain total protein in electrophoresis gels and on nitrocellulose membranes (19). In addition, we demonstrated the use of Ferrene S-ferrous, Ferrozine-ferrous, ferrocyanide-ferric, and Pyrogallol Red-molybdate complexes for colorimetric detection of electrophoretically separated proteins immobilized on membranes (13–16). In 1978, a pink bathophenanthroline disulfonate-ferrous complex was reported as a nonspecific protein stain for polyacrylamide gel electrophoresis (20). The stain is rather insensitive and was later modified by substituting [59Fe] into the complex in order to detect proteins by autoradiography (21). Although increasing sensitivity substantially, the hazards associated with working with radioactivity and the burden of license application for an infrequently used radioisotope have precluded routine utilization of bathophenanthroline disulfonate-[59Fe] as a general protein stain. Measuring light emission is intrinsically more sensitive than measuring light absorbance, as the later is limited by the molar extinction coefficient of the colored complex (28). Thus, luminescent protein detection systems utilizing chelates complexed to transition metal ions such as europium, or ruthenium should offer greater sensitivity than their colorimetric counterparts without the accompanying hazards associated with radioactivity. The organic component of the complex absorbs light and transfers the energy to the transition metal ion, which subsequently emits light at longer wavelength. This is demonstrated by substituting europium into the bathophenanthroline– disulfonate complex (17). This luminescent reagent has been commercialized as SYPRO Rose protein blot stain (Molecular Probes, Eugene, OR). The bathophenanthroline disulfonate-europium complex can detect as little as 8 ng of protein immobilized on nitrocellulose or PVDF membranes. By comparison, the original bathophenanthroline disulfonate–ferrous complex is capable of detecting 600 ng of protein, while the modification employing [59Fe] is capable of detecting 10–25 ng of protein (20,21). The luminescent stain is readily removed by incubating blots in mildly alkaline solution, is highly resistant to photobleaching and is compatible with popular downstream biochemical characterization procedures including immunoblotting, lectin blotting, and mass spectrometry (17). Disadvantages of the bathophenanthroline disulfonate–europium stain are that the dye can only be adequately visualized using 302 nm UV-B epi-illumination and the dye exhibits intense 430 nm (blue) fluorescence emission as well as the desired red emission maxima of 595 and 615 nm.

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Subsequently, SYPRO Rose Plus protein blot stain, an improved europium-based metal chelate stain roughly 10 times brighter than the original bathophenanthroline disulfonate-europium stain, was introduced (25,26). The intense blue fluorescence from uncomplexed ligand, observed in the original stain, was eliminated by employing a thermodynamically more stable europium complex. Due to improved absorption properties, the stain could now be readily visualized with UV-A UV-B or UV-C epi-illumination. Just as with the bathophenanthroline disulfonate–europium stain, SYPRO Rose Plus stain is easily removed by increasing solution pH. The stain is fully compatible with biotin–streptavidin and immunoblotting detection technologies that use a wide variety of visualization strategies. Neither of the europium-based stains is compatible with laser-based gel scanners as they lack visible excitation peaks. SYPRO Ruby dye is a proprietary ruthenium-based metal chelate stain developed to address the limitations of the SYPRO Rose and SYPRO Rose Plus dyes. SYPRO Ruby protein blot stain visualizes electroblotted proteins on nitrocellulose and PVDF membranes with a detection sensitivity of 0.25–1 ng of protein/mm2 in slot-blotting applications. Approximately 2–8 ng of protein can routinely be detected by electroblotting, which side-by-side comparisons demonstrate is as sensitive as colloidal gold stain (22). While colloidal gold staining requires 2–4 h, SYPRO Ruby dye staining is complete in 15 min. The linear dynamic range of SYPRO Ruby protein blot stain is vastly superior to colloidal gold stain, extending over a 1000-fold range. The dye can be excited using a standard 302 nm UV-B transilluminator or using imaging systems equipped with 450-, 473-, 488-, or even 532- nm lasers. Unlike colloidal gold stain, SYPRO Ruby stain does not interfere with mass spectrometry or immunodetection procedures (22). SYPRO Ruby protein gel stain and SYPRO Ruby IEF protein gel stains allow one-step, low background staining of proteins in polyacrylamide or agarose gels without resorting to lengthy destaining steps (see Fig. 1). The linear dynamic range of these dyes extends over three orders of magnitude, thus surpassing silver and Coomassie Blue stains in performance. An evaluation of 11 protein standards ranging in isoelectric point from 3.5 to 9.3 indicates that SYPRO Ruby IEF gel stain is 3–30 times more sensitive than highly sensitive silver stains (24). Proteins that stain poorly with silver stain techniques are often readily detected by SYPRO Ruby dye (27). Similar to colloidal Coomassie Blue stain but unlike silver stain, SYPRO Ruby dye stains are end point stains. Thus, staining times are not critical and staining can be performed over night without gels overdeveloping. A potential disadvantage to detection of proteins using luminescent compared to colorimetric metal chelate stains is the requirement for ancillary equipment such as a laser gel scanner, UV light box, bandpass filters, and photographic or charge-coupled device (CCD) camera. The tremendous improvement in detection sensitivity and linear dynamic range certainly justifies the investment in equipment. Procedures for the detection of electrophoretically

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Fig. 1. Protein detection sensitivity in 1-D gels using SYPRO Ruby Protein gel stain: Twofold serial dilutions of solution-quantified bovine serum albumin (P-7656, Sigma Chemical, Saint Louis, MO) electrophoretically separated on 13% Duracryl SDS-polyacrylamide gels (Genomic Solutions, Ann Arbor, MI). After staining with SYPRO Ruby protein gel stain (Molecular Probes, Eugene, OR), gels were imaged using a Lumi-Imager-F1 scanner (Roche Biochemicals, Indianapolis, IN). The 302-nm UV-B transilluminator was used in conjunction with the system’s 600-nm emission filter. As little as 250 pg of serum albumin is detectable, with a linearly increasing signal extending to 1000 ng.

separated proteins utilizing colorimetric and luminescent metal chelate stains are presented in this chapter. Methods for the elution of the metal chelate stains are also presented. The procedures are applicable for detection of proteins or peptides in polyacrylamide or agarose gels as well as on nitrocellulose, PVDF, or nylon membranes. Owing to the electrostatic mechanism of the protein visualization methods, metal chelate stains are unsuitable for detection of proteins and peptides immobilized on cationic membranes. 2. Materials 2.1. Colorimetric Detection of Electroblotted Proteins on Membranes 1. Block buffer: 0.1% polyvinylpyrrolidone-40 (PVP-40) in 2% glacial acetic acid. 2. Ferrozine–ferrous stain (stable for at least 6 mo at room temperature): 0.75 mM 3-(2-pyridyl)-5,6-bis (4-phenylsulfonic acid)-1,2,4-triazine disodium salt

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(Ferrozine), 30 mM ferric chloride, 15 mM thioglycolic acid in 2% glacial acetic acid. Alternatively, commercially prepared stain solutions of Ferrozine–ferrous (Rev–Pro stain kit; Genomic Solutions, Ann Arbor, MI) or Pyrogallol Red-molybdenum (Microprotein-PR kit; Sigma Chemical Company, St. Louis, MO) can be used. 3. 2% Glacial acetic acid. 4. Ferrocyanide–ferric stain (stable for at least 6 mo at room temperature): 100 mM sodium acetate, pH 4.0; 100 mM potassium ferrocyanide, 60 mM ferric chloride.

2.2. Luminescent Detection of Electroblotted Proteins Using Bathophenanthroline Disulfonate-europium (SYPRO Rose Stain) 1. Formate buffer: 100 mM formic acid, pH 3.7, 100 mM sodium chloride. 2. Bathophenanthroline–europium blot stain (stable for at least 6 mo at room temperature): 1.5 mM bathophenanthroline disulfonic acid disodium salt, 0.5 mM europium chloride, and 0.2 mM EDTA (added from 1000× stock, pH 7.0).

2.3. Luminescent Detection of Electroblotted Proteins Using SYPRO Rose Plus Protein Blot Stain 1. SYPRO Rose Plus protein blot stain kit (Molecular Probes, Inc. cat. #S-12011) The kit contains the following components: SYPRO Rose Plus blot wash solution (component A), 200 mL SYPRO Rose Plus blot stain solution (component B), 200 mL SYPRO Rose Plus blot destain solution (component C), 200 mL

The kit contains sufficient material for staining 10–40 minigel electroblots or four large-format electroblots (20 × 20 cm). The SYPRO Rose Plus solutions may be reused up to four times with little loss in sensitivity. 2.4. Luminescent Detection of Electroblotted Proteins Using SYPRO Ruby Protein Blot Stain 1. SYPRO Ruby protein blot stain (Molecular Probes, Inc.) is provided in a unit size of 200 mL. The 200-mL volume is sufficient for staining 10–40 minigel electroblots or four large-format electroblots (20 × 20 cm). SYPRO Ruby protein blot stain may be reused up to four times with little loss in sensitivity. 2. 7% Acetic Acid, 10% methanol.

2.5. Luminescent Detection of Proteins in SDS-Polyacrylamide Gels Using SYPRO Ruby Protein Gel Stain 1. SYPRO Ruby protein gel stain (Molecular Probes, Inc.) is provided ready to use, in either 200 mL volume (will stain ~four minigels) or 1 L volume (~ 20 minigels or 2–3 large-format gels). 2. 7% Acetic Acid, 10% methanol.

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2.6. Luminescent Detection of Proteins in Isoelectric Focusing (IEF) Gels Using SYPRO Ruby IEF Protein Gel Stain 1. SYPRO Ruby IEF protein gel stain (Molecular Probes, Inc.) is supplied as a 400-mL ready-to-use staining solution, sufficient to stain ~10 IEF minigels or two standard flatbed IEF gels. Use caution when handling the SYPRO Ruby IEF protein gel stain as it contains a strong acid that can cause burns.

2.7. The Elution of Metal Chelate Stains 1. 50 mM Tris-HCl, pH 8.8, 200 mM NaCl, 20 mM EDTA (for the Ferrozine–ferrous stain). 2. 200 mM Sodium carbonate, 100 mM EDTA, pH 9.6 (for the Ferrozine–ferrous stain enhanced with the ferrocyanide–ferric stain and for the bathophenanthroline– europium blot stain). 3. 200 mM Sodium carbonate; 100 mM EDTA, pH 9.6 in 30% methanol (for the bathophenanthroline-europium gel stain). 4. SYPRO Rose Plus blot destain solution (component C) (for the SYPRO Rose Plus protein blot stain.)

3. Methods 3.1. Colorimetric Detection of Electroblotted Proteins on Membranes The colorimetric metal chelate stains allow rapid visualization of proteins on solid-phase supports with detection sensitivities that are comparable to Coomassie Brilliant Blue staining (13–16). Detection sensitivity of the Ferrozine–ferrous stain can be enhanced to a level comparable to silver staining by further incubating membranes in ferrocyanide-ferric stain (13,15). The colorimetric metal chelate stains are fully reversible and compatible with Edman-based protein sequencing, lectin blotting, mass spectrometry and immunoblotting (13–16). 1. After electroblotting, vacuum slot blotting, or dot blotting, membranes are completely immersed in block buffer for 10 min. Blocking and staining steps are performed on a rotary shaker (50 rpm). 2. Thoroughly immerse membranes in Ferrozine-ferrous stain for 10–15 min until purple bands or spots appear. 3. Unbound dye is removed by several brief rinses in 2% glacial acetic acid until the membrane background appears white. Shaking can be performed manually using wash volumes roughly 2× greater than in the blocking and staining steps. 4. If increased sensitivity is desired, the blot can be double stained by subsequently incubating in the ferrocyanide-ferric stain for 10–15 min (see Notes 1–3). 5. Unbound dye is removed by several brief washes in 100 mM sodium acetate, pH 4.0 (see Note 4). Shaking can be performed manually using wash volumes roughly 2× greater than in the blocking and staining steps. 6. Stained proteins are visualized by eye and quantified using a CCD camera (see Note 5).

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3.2. Luminescent Detection of Electroblotted Proteins Using Bathophenanthroline Disulfonate-Europium (SYPRO Rose Stain) Bathophenanthroline disulfonate–europium complex (SYPRO Rose protein blot stain) is a medium sensitivity metal chelate stain that offers the same advantages as colorimetric stains, but with the additional benefits of a 500-fold linear dynamic range and detection sensitivity of α-Glc (Con A) β-(1,4)GlcNAc Terminal GlcNAc α-(2,3)Neu-N-Ac complex N-linked Gal (1,3)GalNAc Terminal Gal (RCA120) α-(2,6)Neu-N-Ac (GlcNAc)2 > GlcNAc Terminal GalNAc α-Man, α-Glc GalNAc Gal (1,4)GlcNAc

a

Many lectins are available commercially as free (unlabeled); digoxigenin (DIG) labeled; biotinylated; peroxidase labelled; or conjugated to agarose beads from companies such as Roche Diagnostics, Sigma Chemical Company, and Amersham-Pharmacia. b GalNAc, N-acetyl galactosamine; GlcNAc, N-acetylglucosamine; Gal, galactose; Glc, glucose; Man, mannose; Fuc, fucose; NeuNAc, N-acetylneuraminic acid (sialic acid). c D-sugars are the preferred sugars.

or AP-labeled lectins can be detected directly. Alternatively, lectins can be detected using anti-DIG peroxidase (if DIG labeled) or streptavidin peroxidase (if biotin labeled), followed by an insoluble substrate. Sensitivity is generally increased with these indirect methods. The detection can be carried out with blots on nitrocellulose or PVDF (see Note 13). 1. Fix membrane for 5 min with 1% KOH. (See Note 12). 2. Rinse for 1 min with distilled water. 3. Block unbound sites with a 1-h incubation at room temperature with blocking reagent. 4. Rinse away blocking reagent with 3 × 1 min washes with TBS-Tween. 5. Add DIG-labeled lectins diluted in TBS-Tween containing 0.1 mM CaCl2, 0.1 mM MgCl2, 0.1 mM MnCl2. Leave overnight at 4°C (see Notes 14 and 15). 6. Remove unbound lectin by washing with 6 × 5 min in TBS-Tween. 7. Incubate nitrocellulose with 10 µL of anti-DIG AP diluted in 10 µL of TBS for 1 h at room temperature. 8. Repeat washing step 6.

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9. Immerse the membrane without shaking into 10 µL of buffer C containing 37.5 µL of solution 4 and 50 µL of solution 3 (mix just before use). 10. Stop reaction when gray to black spots are seen (few minutes to overnight) by rinsing with water.

3.4. Pro-Q Emerald 300 Dye Staining Pro-Q™ Emerald 300 Glycoprotein Gel Stain Kit (Molecular Probes, OR) provides a method for differentially staining glycosylated and nonglycosylated proteins in the same gel. The technique combines the green fluorescent Pro-Q™ Emerald 300 glycoprotein stain with the red fluorescent SYPRO Ruby total protein gel stain. A related product, Pro-Q™ Emerald 300 Glycoprotein Blot Stain Kit, allows similar capabilities for proteins electroblotted to PVDF membranes. Using this stain allows detection of 10%) or gradient gels (acrylamide concentration 5–15%) may crack when dried. This problem is reduced by adding glycerol (1–5%) before drying. When acid-based fluorography enhancers are used, the glycerol should be added during the fluors precipitation step in water after removal of the enhancer. When watersoluble enhancers are used, the glycerol is added during equilibration of the gel in water, before addition of the enhancer. If the concentration of glycerol is too high, gels are difficult to dry, and they may stick to the film. Addition of the enhancer does not increase the cracking. We currently use water-based enhancers for our experiments, because they give sharp autoradiography images and good sensitivity, and can be reused for several gels. Fluorography with commercially available enhancers is simpler and less tedious than the traditional method with PPO-DMSO, and the results are as good as, or better than, those obtained with this method. 3. Enhancers are also used to increase the sensitivity of the autoradiograpy of DNA and RNA of agarose, acrylamide, or mixed gels. When enhancers are used, the gels must be dried at the suggested temperature, because excessive heat will cause damage of the fluors crystal of the enhancer and formation of brown spots on the surface of the gel. 4. Cracking may also be owing to the formation of air bubbles between the gel and the rubber cover of the dryer. This is generally caused by a weak vacuum that is insufficient to maintain the gel well adherent to the paper filter and to the dryer’s tray. Air bubbles can be eliminated by rolling a pipet over the rubber cover while the vacuum is being applied.

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5. Also, excessive stretching of the gel during the transfer to the filter paper may contribute to cracking, particularly for gels that contain high acrylamide concentration. A filter paper should be placed under the gel when removing it from the solution, and the method described in Subheading 3.1.2. should be used for larger gels. 6. It is always necessary to cover the gel with a plastic wrap to prevent sticking to the cover of the dryer. In addition, the vacuum should never be released during the drying procedure until the gel is completely dry, since this will cause the gel to shatter. 7. Staining of the gels with Coomassie blue before fluorography may quench the effect of the enhancer, particularly when low amounts of radioactivity are used. Therefore, the gels must be destained thoroughly, until the background is clear and only the protein bands are stained. 8. X-ray films that are not pre-exposed to light respond to radiation in a sigmoidal fashion, because the halide crystals of the emulsion are not fully activated (10). In contrast, in a pre-exposed film, the response becomes linear and proportional to the amount of radioactivity, therefore, allowing precise quantitative measurement of radioactivity by scanning the autoradiography (11). In addition, pre-exposure (preflashing) of the film results in a two-to threefold increase in sensitivity, for levels of radioactivity near the minimum level of detectability (7), enabling autoradiography of gels containing 14C and 3H radioisotopes to be performed at room temperature, instead of −70°C. To pre-expose a film, a stroboscope or a flash of light (106 Da) as several molecules of each component are linked. This is the most simple labeling procedure to carry out and, although the yields of enzyme activity and immunoreactivity are small, the conjugates obtained are stable and practical reagents. Alkaline phosphatase may also be coupled using heterobifunctional reagents containing the N-hydroxysuccinimide and maleimide groups, for example, succinimidyl 4-(N-maleimidyl)-cyclohexane-1-carboxylate. However, because the enzyme has no free thiol groups this approach is usually used for the labeling of F(ab’)2 fragments of IgG via their thiols (2).

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2. Materials 1. Alkaline phosphatase from bovine intestinal mucosa. 2000 U/mg (with 4-nitrophenyl phosphate as substrate). There are numerous commercial sources of labelinggrade enzyme; it is usually supplied at a concentration of 10 mg/mL in phosphate or triethanolamine buffer (this avoids interference with the conjugation). If ammonium sulfate, Tris or primary or secondary amine is present, it must be removed (see Note 1). 2. IgG antibody. This should be the pure IgG fraction or, better, affinity-purified antibody from serum or pure monoclonal antibody (see Chapters 179–190). 3. Phosphate-buffered saline (PBS): 20 mM sodium phosphate buffer, pH 7.2, containing 0.15 M NaCl. 4. Glutaraldehyde. 5. 50 mM Tris-HCl buffer, pH 7.5, containing 1 mM MgCl2, 0.02% NaN3, and 2% bovine serum albumin (BSA).

3. Methods 1. Add 0.5 mg of IgG antibody in 100 µL of PBS to 1.5 mg of alkaline phosphatase. 2. Add 5% glutaraldehyde (about 10 µL) to give a final concentration of 0.2% (v/v) and stir the mixture for 2 h at room temperature. 3. Dilute the mixture to I mL with PBS and dialyze against PBS (2L) at 4°C overnight (see Note 1). 4. Dilute the solution to 10 mL with 50 mM Tris-HCl buffer, pH 7.5, containing 1 mM MgCl2, 0.02% NaN3, and 2% BSA, and store at 4°C.

4. Notes 1. Dialyses of small volumes can be conveniently done in narrow dialysis tubing by placing a short glass tube, sealed at both ends, in the tubing so that the space available to the sample is reduced. Transfer losses are minimised by carrying out the subsequent steps in the same dialysis bag. There are also various microdialysis systems available commercially such as the Slide-A-Lyzer units from Pierce Biotechnology (Rockford, IL). 2. The conjugates are stable for several years at 4°C as the NaN3 inhibits microbial growth and the BSA minimizes denaturation and adsorption loses. The conjugates should not be frozen. 3. Purification of the conjugates is usually unnecessary; however, if there is evidence of free antibody it can be removed by size-exclusion chromatography in Sepharose CL-6B (GE Healthcare Bio-sciences, Uppsala, Sweden and Piscataway, NJ) or a similar gel with PBS as solvent. 4. The efficacy of the enzyme-labeled antibody can be tested by immobilizing the appropriate antigen on the wells of a microtiter plate or strip, incubating various dilutions of the conjugate for a few hours, washing the wells, adding substrate, and measuring the amount of product formed. This

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approach may also be used for monitoring conjugate purification of chromatographic fractions. References 1. Engvall, E. and Perlmann, P. (1972) Enzyme-linked immunosorbent assay. III. Quantitation of specific antibodies by enzyme-labeled anti-immunoglobulin in antigen-coated tubes. J. Immunol. 109, 129–135. 2. Mahan, D. E., Morrison, L., Watson, L., and Haugneland, l.S. (1987) Phase change enzyme immunoassay. Analyt. Biochem. 162, 163–170.

65 b-Galactosidase Labeling of IgG Antibody G. Brian Wisdom

1. Introduction The Eschericia coli β-galactosidase (EC 3.2.1.23) is a large enzyme (465 kDa) with a high turnover rate and wide specificity which allows it to be measured conveniently in several different ways. Unlike several other enzyme labels β-galactosidase is not found in mammalian tissues or fluids hence background contributions are negligible when this label is measured at a neutral pH. This can be advantageous as peroxidase activities (due to various heme proteins) and alkaline phosphatase are frequently present in these samples. The heterobifunctional reagent, m-maleimidobenzoyl-N-hydroxysuccinimide ester (MBS), is of value when one of the components of a conjugate has no free thiol groups eg. IgG. In this method (1) the IgG antibody is first modified by allowing the N-hydroxysuccinimide ester group of the MBS to react with amino groups in the IgG; the β-galactosidase is then added and the maleimide groups on the modified IgG react with thiol groups in the enzyme to form thioether links. This procedure produces conjugates with molecular weights in the range 0.6-1 ×106. F(ab′)2 fragments of IgG have been labeled via the β-galactosidase’s amino groups with sulfo-succinimidyl 4-(N-maleimidomethyl) cyclohexane-1-carboxylate (2) and there are various other heterobifunctional reagents that may be used for β-galactosidase labelling (3). 2. Materials 1. β -Galactosidase from E. coli, 600 U/mg or greater (with 2-nitrophenyl-β -galactopyranoside as substrate) (see Note 1).

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2. IgG antibody. This should be the pure IgG fraction or, better, affinity-purified antibody from an antiserum or pure monoclonal antibody. 3. 0.1 M Sodium phosphate buffer, pH 7.0, containing 50 mM NaCl 4. MBS. 5. Dioxan. 6. Sephadex G25 (GE Healthcare Bio-sciences, Uppsala, Sweden and Piscataway, NJ) or an equivalent gel. 7. 10 mM Sodium phosphate buffer, pH 7.0, containing 50 mM NaCl and 10 mM MgCl2. 8. 2-Mercaptoethanol. 9. DEAE-Sepharose (GE Healthcare Bio-sciences) or an equivalent ion-exchange medium. 10. 10 mM Tris-HCl buffer, pH 7.0, containing 10 mM MgCl2 and 10 mM 2-mercaptoethanol. 11. Item 10 containing 0.5 M NaCl. 12. Item 10 containing 3% bovine serum albumin (BSA) and 0.6% NaN3.

3. Methods 1. Dissolve 1.5 mg of IgG in 1.5 mL of 0.1 M sodium phosphate buffer, pH 7.0, containing 50 mM NaCl. 2. Add 0.32 mg of MBS in 15 µL of dioxan, mix, and incubate for 1 h at 30°C. 3. Fractionate the mixture on a column of Sephadex G25 (approx. 0.9 × 20 cm) equilibrated with 10 mM sodium phosphate buffer, pH 7.0, containing 50 mM NaCl and 10 mM MgCl2, and elute with the same buffer. Collect 0.5 mL fractions, measure the A280 and pool the fractions in the first peak (about 3 mL in volume). 4. Add 1.5 mg of enzyme, mix, and incubate for 1 h at 30°C. 5. Stop the reaction by adding 1 M 2-mercaptoethanol to give a final concentration of 10 mM (about 30 µL). 6. Fractionate the mixture on a column of DEAE-Sepharose (approx. 0.9 × 15 cm) equilibrated with 10 mM Tris-HCl buffer, pH 7.0, containing 10 mM MgCl2 and 10 mM 2-mercaptoethanol; wash the column with this buffer (50 mL) followed by the buffer containing 0.5 M NaCl. (50 mL). Collect 3 mL fractions in tubes with 0.1 mL of Tris buffer containing 3% BSA and 0.6% NaN3. Pool the major peak (this is eluted with 0.5 M NaCl), and store at 4°C (see Notes 2 and 3).

4. Notes 1. The thiol groups of β-galactosidase may become oxidized during storage thus diminishing the efficacy of the labeling. It is relatively easy to measure these groups using 5,5 -dithiobis(2-nitrobenzoic acid) (see Chapter 93); about 10 thiol groups per enzyme molecule allow the preparation of satisfactory conjugates.

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2. The conjugates are stable for a year at 4°C as the NaN3 inhibits microbial growth and the BSA minimizes denaturation and adsorption losses. 3. The activity of the conjugate can be checked by the method described in Note 4 of Chapter 64. References 1. O’Sullivan, M. J., Gnemmi, E., Morris, D., Chieregatti, G., Simmonds, A. D., Simmons, M., Bridges, J. W., and Marks, V. (1979) Comparison of two methods of preparing enzyme-antibody conjugates: Application of these conjugates to enzyme immunoassay. Anal. Biochem. 100, 100–108. 2. Luz, Z., Gurlo, T., and von Grafenstein, H. (2000) Cell-ELISA using β-galactosidase conjugated antibodies. J. Immunol. Methods 234, 153–167. 3. Hermanson, G. T. (1996) Bioconjugate Techniques (Academic Press, San Diego), pp 229–248.

66 Horseradish Peroxidase Labeling of IgG Antibody G. Brian Wisdom

1. Introduction Horseradish peroxidase (HRP; EC 1.11.1.7) is probably the most widely used enzyme label. This protein is relatively small (44 kDa), stable, and has a broad specificity which allows it to be measured by absorption, fluorescence or luminescence. Some of its products are intensely coloured, which makes this enzyme suitable for immunocytochemical or immunoblotting applications. The most popular method (1) for labeling IgG antibody molecules with HRP exploits the glycoprotein nature of the enzyme. The saccharide residues are oxidized with sodium periodate to produce aldehyde groups that can react with the amino groups of the IgG molecule, and the Schiff bases formed are then reduced to give a stable conjugate of high molecular weight (0.5–1.0 × 106). The enzyme has few free amino groups so self-coupling is not a significant problem. Other methods have been described for using this enzyme as a horseradish peroxidase as a label. IgG may be labeled with HRP using glutaraldehyde in a two-step proecure (2) and, when partially reduced, IgG has been labelled using the heterobifunctional succinimidyl 4-(N-maleimidyl)-cyclohexane1-carboxylate (3). 2. Materials 1. Horseradish peroxidase, 1000 U/mg or greater (with 2,2´ -azino-bis-[3ethylbenzthiazoline-6-sulfonic acid] as substrate) (see Note 1). 2. IgG antibody. This should be the pure IgG fraction or, better, affinity purified antibody from an antiserum or pure monoclonal antibody (see Chapters 179–189). 3. 0.1 M Sodium periodate, freshly prepared. 4. 1 mM Sodium acetate buffer, pH 4.4. 5. 10 mM Sodium carbonate buffer, pH 9.5.

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6. 0.2 M Sodium carbonate buffer, pH 9.5. 7. Sodium borohydride, 4 mg/mL (freshly prepared). 8. Sepharose CL-6B (GE Healthcare Bio-sciences, Uppsala, Sweden and Piscataway, NJ) or a similar gel. 9. PBS: 20 mM sodium phosphate buffer, pH 7.2, containing 0.15 M NaCl. 10. Bovine serum albumin (BSA).

3. Method 1. Dissolve 2 mg of peroxidase in 500 µL of water. 2. Add 100 µL of freshly prepared 0.1 M sodium periodate, and stir the solution for 20 min at room temperature. (The color changes from orange to green). 3. Dialyze the modified enzyme against 1 mM sodium acetate buffer, pH 4.4 (2 L) overnight at 4°C (see Note 2). 4. Dissolve 4 mg of IgG in 500 µL of 10 mM sodium carbonate buffer, pH 9.5. 5. Adjust the pH of the dialyzed enzyme solution to 9.0–9.5 by adding 10 µL of 0.2 M sodium carbonate buffer, pH 9.5, and immediately add the IgG solution. Stir the mixture for 2 h at room temperature. 6. Add 50 µL of freshly prepared sodium borohydride solution (4 mg/mL), and stir the mixture occasionally over a period of 2 h at 4°C. 7. Fractionate the mixture by size-exclusion chromatography in a column (approx. 1.5 × 85 cm) of Sepharose CL-6B in PBS. Determine the A280 and A403 (see Note 3) 8. Pool the fractions in the first peak (both A280 and A403 peaks coincide), add BSA to give a final concentration of 5 mg/mL, and store the conjugate in aliquots at −20°C (see Notes 4 and 5).

4. Notes 1. Preparations of horseradish peroxidase may vary in their carbohydrate content and this can affect the oxidation reaction. Free carbohydrate can be removed by size exclusion chromatography. Increasing the sodium periodate concentration to 0.2 M can also help, but further increases lead to inactivation of the peroxidase. 2. Dialysis of small volumes can be conveniently done in narrow dialysis tubing by placing a short glass tube, sealed at both ends, in the tubing so that the space available to the sample is reduced. There are also various microdialysis systems available commercially such as the Slide-A-Lyzer units from Pierce Biotechnology (Rockford, IL). 3. The absorbance at 403 nm is caused by the peroxidase’s heme group. The enzyme is often specified in terms of its RZ value; this is the ratio of A280 and A403, and it provides a measure of the heme content and purity of the preparation. Highly purified peroxidase has an RZ of about 3. Conjugates with an RZ of 0.4 perform satisfactorily. 4. BSA improves the stability of the conjugate and minimizes loses due to adsorption and denaturation. NaN3 should not be used with peroxidase

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conjugates because it inhibits the enzyme. If an antimicrobial agent is required, 0.2% sodium merthiolate (thimerosal) should be used. 5. The activity of the conjugate can be checked by the method described in Note 4 of Chapter 64. References 1. Wilson, M. B. and Nakane, P. P. (1978) Recent developments in the periodate method of conjugating horse radish peroxidase (HRPO) to antibodies, in Immunofluorescence and Related Staining Techniques (Knapp, W., Holubar, K., and Wick, G., eds.), Elsevier/North Holland Biomedical, Amsterdam, pp. 215–224. 2. Avrameas, S. and Ternynck, T. (1971) Peroxidase labeled antibody and Fab conjugates with enhanced intracellular penetration. Immunochemistry, 8, 1175–1179. 3. Yoshitake, S., Imagawa, M., Ishikawa, E., Niitsu, Y., Urushizaki, I., Nishiura, M., Kanazawa, R., Kurosaki, H., Tachibana, S., Nakazawa, N., and Ogawa, H. (1982) Mild and efficient conjugation of rabbit Fab’ and horseradish peroxidase using a maleimide compound and its use in enzyme immunoassay. J. Biochem. 92, 1413–1424.

67 Digoxigenin Labeling of IgG Antibody G. Brian Wisdom 1. Introduction Digoxigenin (DIG) is a plant steroid (390 Daltons) which can be used as a small, stable label of IgG molecules. It is a valuable alternative to biotin as the biotin-streptavidin system can sometimes give high backgrounds due, for example, to the presence of biotin-containing enzymes in the sample. There is a range of commercially available mouse and sheep anti-DIG Fab antibody fragments labeled with various enzymes and fluorescent molecules for the detection of the DIG-labeled IgG antibody in many applications (1). The IgG molecule is labeled, via its amino groups, with an N- hydroxysuccinimide ester derivative of the steroid containing the 6-aminocaproate spacer. 2. Materials 1. Digoxigenin-3-O-succinyl-ε-aminocaproic acid-N-hydroxysuccinimide ester (Roche Applied Science, Indianapolis, IL). 2. IgG antibody. This should be the pure IgG fraction or, better, affinity purified antibody from an antiserum or pure monoclonal antibody. 3. Dimethylsufoxide (DMSO). 4. PBS: 20 mM sodium phosphate buffer, pH 7.2, containing 0.15 M NaCl. 5. 0.1 M ethanolamine, pH 8.5. 6. PBS containing 0.1% bovine serum albumin (BSA). 7. Sephadex G-25 (GE Healthcare Bio-sciences, Uppsala, Sweden and Piscataway, NJ) or similar gel (see Note 1).

3. Method 1. Dissolve 1 mg of the antibody in 1 mL of PBS. 2. Prepare the digoxigenin-3-O-succinyl-O-minocaproic acid-N-hydroxysuccinimide ester immediately prior to use by dissolving it at a concentration of 2 mg/mL in DMSO. From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Add 24 µL of the DIG reagent to the antibody solution slowly with stirring and incubate at room temperature for 2 h. 4. Terminate the reaction by adding 0.1 mL of 0.1 M ethanolamine and incubate for 15 min. 5. Remove the excess DIG reagent by size-exclusion chromatography in a small column of Sephadex G-25 equilibrated with PBS containing 0.1% BSA. The IgG is in the first A280 peak. 6. Store the labeled antibody at 4°C with 0.05% NaN3 or in aliquots at −20°C or lower.

4. Note 1. Suitable ready-made columns of cross-linked dextran are available from GE Biosciences (PD-10 or HiTrap columns) and from Pierce Biotechnology (Rockford, IL) (Presto and Kwik columns). Reference 1. Kessler, C. (1991) The digoxigenin-anti-digoxigenin (DIG) technology: a bioanalytical indicator system. Molec. Cell. Probes 5, 161–205.

68 Conjugation of Fluorochromes to Antibodies Su-Yau Mao 1. Introduction The use of specific antibodies labeled with a fluorescent dye to localize substances in tissues was first devised by A. H. Coons and his associates. At first, the specific antibody itself was labeled and applied to the tissue section to identify the antigenic sites (direct method) (1). Later, the more sensitive and versatile indirect method (2) was introduced. The primary, unlabeled, antibody is applied to the tissue section, and the excess is washed off with buffer. A second, labeled antibody from another species, raised against the IgG of the animal donating the first antibody, is then applied. The primary antigenic site is thus revealed. A major advantage of the indirect method is the enhanced sensitivity. In addition, a labeled secondary antibody can be used to locate any number of primary antibodies raised in the same animal species without the necessity of labeling each primary antibody. Four fluorochromes are commonly used: fluorescein, rhodamine, Texas red, and phycoerythrin (3). They differ in optical properties, such as the intensity and spectral range of their absorption and fluorescence. Choice of fluorochrome depends on the particular application. For maximal sensitivity in the binding assays, fluorescein is the fluorochrome of choice because of its high quantum yield. If the ligand is to be used in conjunction with fluorescence microscopy, rhodamine coupling is advised, since it has superior sensitivity in most microscopes and less photobleaching than fluorescein. Texas red (4) is a red dye with a spectrum that minimally overlaps with that of fluorescein; therefore, these two dyes are suitable for multicolor applications. Phycoerythrin is a 240-kDa, highly soluble fluorescent protein derived from cyanobacteria and eukaryotic algae. Its conjugates are among the most sensitive fluorescent probes available (5) and are frequently used in flow cytometry and immunoassays (6). In addition, the Alexa fluorochromes are a series of fluorescent dyes with

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excitation/emission spectra similar to those of commonly used ones, but are more fluorescent and more photostable (7). Thiols and amines are the only two groups commonly found in biomolecules that can be reliably modified in aqueous solution. Although the thiol group is the easiest functional group to modify with high selectivity, amines are common targets for modifying proteins. Virtually all proteins have lysine residues, and most have a free amino terminus. The ε-amino group of lysine is moderately basic and reactive with acylating reagents. The concentration of the free-base form of aliphatic amines below pH 8.0 is very low. Thus, the kinetics of acylation reactions of amines by isothiocyanates, succinimidyl esters, and other reagents is strongly pH-dependent. Although amine acylation reactions should usually be carried out above pH 8.5, the acylation reagents degrade in the presence of water, with the rate increasing as the pH increases. Therefore, a pH of 8.5–9.5 is usually optimal for modifying lysines. Where possible, the antibodies used for labeling should be pure. Affinitypurified, fluorochrome-labeled antibodies demonstrate less background and nonspecific fluorescence than fluorescent antiserum or immunoglobulin fractions. The labeling procedures for the isothiocyanate derivatives of fluorescein and sulfonyl chloride derivatives of rhodamine are given below (8). The major problem encountered is either over- or undercoupling, but the level of conjugation can be determined by simple absorbance readings. 2. Materials 1. IgG. 2. Borate buffered saline (BBS): 0.2 M boric acid, 160 mM NaCl, pH 8.0. 3. Fluorescein isothiocyanate (FITC) or Lissamine rhodamine B sulfonyl chloride (RBSC). 4. Sodium carbonate buffer: 1.0 M NaHCO3-Na2CO3 buffer, pH 9.5, prepared by titrating 1.0 M NaHCO3 with 1.0 M Na2CO3 until the pH reaches 9.5. 5. Absolute ethanol (200 proof) or anhydrous dimethylformamide (DMF). 6. Sephadex G-25 column. 7. Whatman DE-52 column. 8. 10 mM Sodium phosphate buffer, pH 8.0. 9. 0.02% Sodium azide. 10. UV spectrophotometer.

3. Methods 3.1. Coupling of Fluorochrome to IgG 1. Prior to coupling, prepare a gel-filtration column to separate the labeled antibody from the free fluorochrome after the completion of the reaction. The size of the column should be 10 bed volumes/sample volume (see Note 1).

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The molar ratio (F/P) is calculated by Eq. 4. F/P = (A575 nm /73,000) × (150,000/Rho IgG conc.)

(4)

4. Notes 1. Sephadex G-25 resin is the recommended gel for the majority of desalting applications. It combines good rigidity, for easy handling and good flow characteristics, with adequate resolving power for desalting molecules down to about 5000 Dalton mol wt. If the volume of the reaction mixture is 9.5; at lower pH, the side reactions proceed very rapidly. Hence it is important to add the methyl acetimidine solution to the buffered protein solution without causing a change of pH (6). The methyl acetimidine is purchased as the hydrochloride salt, which is neutralized by dissolving it in 1 M NaOH (see Subheading 3., step 2). The pH of the resulting solution should be approx 10; if necessary the pH may be adjusted before the addition to the protein solution with 1 M NaOH or 1 M HCl. As acetimidates are rapidly hydrolyzed at basic pH, the entire process should be performed very rapidly. It is therefore recommended, in a preliminary experiment, to dissolve the methyl acetimidine hydrochloride and determine the exact amount of 1 M NaOH that yields a solution of pH 10. The same process is then repeated with a freshly prepared methyl acetimidine solution which is rapidly added to the protein. 2. The acetimidyl group may be removed by treatment of the modified protein with an ammonium acetate buffer prepared by adding concentrated ammonium hydroxide solution to acetic acid to a pH of 11.3 (Caution! Preparation of this buffer must be done carefully and in a well-ventilated chemical hood). 3. Amidination is obviously unsuitable for the modification of proteins that are sensitive to basic pH. Reductive alkylation, using formaldehyde and sodium cyanoborohydride (see ref. 7), can be performed at neutral pH and is recommended for the modification of such proteins.

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References 1. Means, G. E. and Feeney, R. E. (1971) Chemical Modifications of Proteins. Holden-Day, San Francisco. 2. Hirs, C. H. N. and Timasheff, S. N. (eds.) (1972) Enzyme structure B. Meth. Enzymol. 25. 3. Lundblad, R. L. and Noyes, C. M. (1984) Chemical Reagents for Protein Modifications, Vols. 1 and 2. CRC Press, Boca Raton, FL. 4. Feeney, R. E. (1987) Chemical modification of proteins: comments and perspectives. Int. J. Pept. Protein Res. 27, 145–161. 5. Hunter, M. J. and Ludwig, M. L. (1962) The reaction of imidoesters with small proteins and related small molecules. J. Amer. Chem. Soc. 84, 3491–3504. 6. Wallace, C. J. A. and Harris, D. E. (1984) The preparation of fully N-e-acetimidylated cytochrome c. Biochem. J. 217, 589–594. 7. Jentoft, N. and Dearborn, D. G. (1979) Labeling of proteins by reductive methylation using sodium cyanoborohydride. J. Biol. Chem. 254, 4359–4365. 8. Walter, T. S. et al (2006) Lysine methylation as a routine rescue strategy for protein crystallization Structure 14, 1617–1622. 9. Fields, R. (1972) The rapid determination of amino groups with TNBS. Meth. Enzymol. 25, 464–468.

92 Modification of Tryptophan with 2-Hydroxy-5Nitrobenzylbromide Dan S. Tawfik 1. Introduction 2-Hydroxy-5-nitrobenzylbromide, Koshland’s reagent (1), reacts rapidly and under mild conditions with tryptophan residues. At low pH (< 7.5) this reagent exhibits a marked selectivity for tryptophan; under more basic pH or at higher reagent concentrations, cysteine, tyrosine, and even lysine residues can be modified as well (see Note 1). The reaction is extremely rapid either with the protein or with water. The reagent is relatively insoluble in water; it is therefore necessary to first dissolve it in an organic solvent (e.g., dioxane) and then to add it to the protein solution in buffer. Unlike that of most other modifying reagents, the final organic solvent concentration in the reaction mixture is relatively high (5–15%). Determination of the number of modified tryptophans is done spectrophotometrically. 2. Materials 1. 2. 3. 4. 5.

2-Hydroxy-5-nitrobenzylbromide. Dioxane (water free) (see Note 2). Protein for modification: 1 mg/mL in 0.1 M phosphate buffer, pH 7.0. 1 M NaOH. A Sephadex G-25 column.

3. Method 1. Prepare a fresh solution of 200 mM 2-hydroxy-5-nitrobenzylbromide in dioxane (keep the solution in the dark). 2. Dilute this solution in dioxane to 10× the final reagent concentration (see Note 3). 3. Add 10 µL of the 2-hydroxy-5-nitrobenzylbromide solution to a 90-µL aliquot of the protein solution and shake for 2 min (e.g., on a Vortex).

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4. Centrifuge (for few minutes at 10,000 rpm) if a precipitate forms (see Note 4). 5. Filter on a Sephadex G-25 and then dialyze against an appropriate buffer (see Note 4). 6. Determine the number of modified tryptophans by measuring the absorbance of the purified protein at 410 nm at pH ≥ 10 (ε = 18,450 M−1 cm−1). 7. Determine the activity of the protein.

4. Notes 1. At neutral and slightly acidic pH the reaction is generally selective. At higher pH, the major side reaction is the benzylation of sulfhydryl groups; to prevent the modification of free cysteine residues the protein can be carboxymethylated prior to the modification with 2-hydroxy-5-nitrobenzylbromide. 2. Avoid the use of acetone, methanol, or ethanol as organic co-solvents for this reagent. Water-free dioxane is commercially available or can be prepared by drying dioxane over sodium hydroxide pellets. 3. At high reagent concentrations a precipitate of 2-hydroxy-5-nitrobenzyl alcohol is sometimes observed and can be removed by centrifugation. Purification of the modified protein from all the remaining alcohol product is sometimes difficult and may require extensive dialysis in addition to gel filtration. 4. The reaction of hydroxy-5-nitrobenzylbromide is accompanied by the release of hydrobromic acid. The pH of the reaction should be maintained if necessary by the subsequent addition of small aliquots of 1 M NaOH solution. References 1. Horton, H. R. and Koshland, D. E., Jr. (1972) Modification of proteins with active benzyl halides. Meth. Enzymol. 25, 468–482.

93 Modification of Sulfhydryl Groups with DTNB Dan S. Tawfik 1. Introduction 5-5 -Dithio-bis (2-nitrobenzoic acid) (DTNB or Ellman’s reagent; ref. 1) reacts with the free sulfhydryl side chain of cysteine to form an S–S bond between the protein and a thionitrobenzoic acid (TNB) residue. The modification is generally rapid and selective. The main advantage of DTNB over alternative reagents (e.g., N-ethylmaleimide or iodoacetamide) is in the selectivity of this reagent and in the ability to follow the course of the reaction spectrophotometrically. The reaction is usually performed at pH 7.0–8.0 and the modification is stable under oxidative conditions. The TNB group can be released from modified protein by treatment with reagents that are routinely used to reduce S–S bonds, for example, mercaptoethanol, or by potassium cyanide (2) (see Note 1). In addition, the often highly pronounced differences in reactivity of different cysteine side chains in the same protein or even active site, and the availability of a variety of thiol-modifying reagents can be exploited to selectively modify cysteine side chains in proteins in the presence of other, more reactive cysteine residues (3). 2. Materials 1. DTNB. 2. 0.1 M Tris-HCl, pH 8.0. 3. Protein for modification: approx 5 µM in 0.1 M Tris-HCl, pH 8.0.

3. Method 1. Prepare fresh solutions of DTNB (0.5–5 mM; see Note 2) in 0.1 M Tris-HCl, pH 8.0. 2. Add 20-µL aliquots of the DTNB solution to 180 µL of the protein solution and incubate for 30 min.

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3. Determine the number of modified cysteines by measuring the absorbance of the released TNB anion at 412 nm ( = 14,150 M−1 cm−1) (see Note 3). 4. Dialyze against an appropriate buffer (see Note 4). 5. Determine the activity of the protein.

4. Notes 1. Release of the TNB modification can be achieved by treatment of the modified protein (preferably after dialysis) with thiols (e.g., mercaptoethanol) or by potassium cyanide (20 mM final concentration; 10–60 min). The release of the TNB anion can be followed spectrophotometrically at 412 nm (2). 2. A molar excess of 10–100 of DTNB and an incubation time of 30 min is usually sufficient with most proteins. The modification of a particular cysteine residue may require a higher DTNB concentration, a longer reaction time, or a higher pH. 3. The extent of modification is determined by measuring the absorbance of the released TNB anion and hence can be followed during the reaction and in the presence of an excess of unreacted DTNB. 4. Dialysis of the reaction mixture is not always necessary; in many cases the activity of the protein (enzymatic or binding) can be determined directly after the reaction (for an example see ref. 2). References 1. Ellman, G. L. (1959) Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82, 70–77. 2. Fujioka, M., Takata, Y., Konishi, K., and Ogawa, H. (1987) Function and reactivity of sulfhydryl groups of rat liver glycine methyltransferase. Biochemistry 26, 5696–5702. 3. Altamirano, M. M., Garcia, C., Possani, L. D., and Fersht, A. R. (1999) Oxidative refolding chromatography: folding of the scorpion toxin Cn5 [see comments]. Nat. Biotechnol. 17, 187–191.

94 Chemical Cleavage of Proteins at Methionyl-X Peptide Bonds Bryan John Smith

1. Introduction One of the most commonly used methods for proteolysis uses cyanogen bromide to cleave the bond to the carboxy-(C)-terminal side of methionyl residues. The reaction is highly specific, with few side reactions and a typical yield of 90–100%. It is also relatively simple and adaptable to large or small scale. Because methionine is one of the least abundant amino acids, cleavage at that residue tends to generate a relatively small number of peptides of large size—up to 10,000–20,000 Da. For this reason the technique is usually less useful than some other methods (such as cleavage by trypsin) for identification of proteins by mass mapping, which is better done with a larger number of peptides. Cleavage at Met-X can be useful for other purposes, however: 1. 2. 3. 4.

Generation of internal sequence data, from the large peptides produced (1). Peptide mapping. Mapping of the binding sites of antibodies (2) or ligands (3). Generation of large, functionally distinct domains (e.g., from hirudin, by Wallace et al. [4]) or proteins of interest from fusion proteins (5). 5. Confirmation of estimates of methionine content by amino acid analysis, which has a tendency to be somewhat inaccurate for this residue (6). This is by determination of the number of peptides produced by cleavage at an assumed 100% efficiency.

2. Materials 1. Ammonium bicarbonate (0.4 M) solution in distilled water (high-performance liquid chromatography [HPLC] grade). Stable for weeks in refrigerated stoppered bottle. 2. 2-Mercaptoethanol. Stable for months in dark, stoppered, refrigerated bottle.

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3. Trifluoroacetic acid (TFA), Aristar grade. Make to 70% v/v by addition of distilled water (HPLC grade), and use fresh (see Notes 1 and 2). 4. CNBr. Stable for months in dry, dark, refrigerated storage. Warm to room temperature before opening. Use only white crystals, not yellow ones. Beware of the toxic nature of this reagent: hydrogen cyanide is a breakdown product. Use in a fume cupboard (see Note 3). 5. Sodium hypochlorite solution (domestic bleach). 6. Equipment includes a nitrogen supply, fume hood, and suitably sized and capped tubes (e.g., Eppendorf microcentrifuge tubes).

3. Methods 3.1. Reduction 1. Dissolve the polypeptide in water to between 1 and 5 mg/mL, in a suitable tube. Add one volume of ammonium bicarbonate solution, and add 2-mercaptoethanol to between 1% and 5% (v/v). 2. Blow nitrogen over the solution to displace oxygen, seal the tube, and incubate at room temperature for approx 18 h.

3.2. Cleavage 1. Dry down the sample under vacuum, warming if necessary to help drive off all of the bicarbonate. Any remaining ammonium bicarbonate will form a nonvolatile salt on subsequent reaction with the TFA that follows. If a white salty deposit remains, redissolve in water and dry down again. 2. Redissolve the dried sample in 70% v/v TFA, to a concentration of 1–5 mg/mL. 3. Add excess white crystalline CNBr to the sample solution, to 10-fold or more molar excess over methionyl residues. Practically, this amounts to approx equal weights of protein and CNBr. To very small amounts of protein, add one small crystal of reagent. Carry out this stage in the fume hood (see Notes 3 and 4). 4. Seal the tube and incubate at room temperature for 24 h. 5. Terminate the reaction by drying down under vacuum. Store samples at −10°C or use immediately (see Note 5). 6. Immediately after use decontaminate equipment that has contacted CNBr by immersion in hypochlorite solution (bleach) until effervescence stops (a few minutes). Wash decontaminated equipment thoroughly.

4. Notes 1. The mechanism of the action of cyanogen on methionine-containing peptides is shown in Fig. 1. For further details, see the review by Fontana and Gross (7). The methioninyl residue is converted to homoseryl or homoseryl lactone. The relative amounts of these two depend on the acid used, but when 70% TFA is the solvent, homoserine lactone is the major derivative. Peptides generated are suitable for peptide sequencing by Edman

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Fig. 1. Mechanism of cleavage of Met-X bonds by CNBr.

chemistry. Methionine sulfoxide does not take part in this reaction and the first step in the method is intended to convert any methionyl sulfoxide to methionyl residues, and so maximize cleavage efficiency. If the reduction is not carried out, the efficiency of cleavage may not be greatly diminished. If virtually complete cleavage is not necessary, partial cleavage products are desired (see Note 6), the sample is small and difficult to handle without loss, or speed is critical, the reduction step may be omitted. An acid environment is required to protonate basic groups and so prevent reaction there and maintain a high degree of specificity. Met-Ser and Met-Thr bonds may give significantly less than 100% yields of cleavage and simultaneous conversion to methionyl to homoseryl residues within the uncleaved polypeptide. This is because of the involvement of the β-hydroxyl groups of seryl and threonyl residues in alternative reactions, which do not result in cleavage (7). Morrison et al. (8) however, have found that use of 70% v/v TFA gives a better yield of cleavage of a Met-Ser bond in apolipoprotein A1 than does use of 70% formic acid (see Note 2). Using model peptides, Kaiser and Metzka (9) analyzed the cleavage reaction at Met-Ser and Met-

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Thr and concluded that cleavage that efficiency is improved by increasing the amount of water present, and for practical purposes 0.1 M HCl is a good acid to use, giving about 50% cleavage of these difficult bonds. Remaining uncleaved molecules contained either homoserine or methionyl sulfoxide instead of the original methionyl. Cleavage efficiency improved with increasing strength of acid, but there was an accompanying risk of degradation in the stronger acids. Utilization of C-terminal homoseryl lactone for linkage to solid phase is discussed in Note 7. 2. Acid conditions are required for the reaction to occur. In the past, 70% v/v formic acid (pH 1) was commonly used because it is a good protein solvent and denaturant, and also volatile. However, it may damage tryptophan and tyrosine residues (8) and also cause formation of seryl and threonyl side chains (showing up during analysis by mass spectroscopy as an increase of 28 amu per modification [9,10]). Use of other acids avoids this problem. TFA (also volatile) may be used in concentrations in the range 50%–100% (v/v). The pH of such solutions is approx pH 0.5 or less. The rate of cleavage in 50% TFA may be somewhat slower than in 70% formic acid, but similar reaction times of hours, up to 24 h will provide satisfactory results. Caprioli et al. (11) and Andrews et al. (12) have illustrated the use of 60% and 70% TFA (respectively) for cyanogen bromide cleavage of proteins. Acetic acid (50%–100% v/v) may be used as an alternative but reaction is somewhat slower than in TFA. Alternatively, 0.1 M HCl has been used (9,10). To increase solubilization of proteins, urea or guanidine·HCl may be added to the solution. Thus, in 0.1 M HCl, 7 M urea, for 12 h at ambient temperature, a Met-Ala bond was cleaved with 83% efficiency, and the more problematical Met-Ser and Met-Thr bonds with 56% and 38% efficiency, respectively (9). 3 Although the specificity of this reaction is excellent, some side reactions may occur. This is particularly so if colored (yellow or orange) CNBr crystals are used, when there may be destruction of tyrosyl and tryptophanyl residues and bromination may also be detected by mass spectroscopy. Treatment of samples in other formats is discussed in Notes 8 and 10. 4. The above protocol describes addition of solid CNBr to the acidic protein solution, to give a molar excess of CNBr over methionyl residues. This has the advantage that pure white crystals may be selected in favor of pale yellow ones showing signs of degradation (see Note 3). It does not allow accurate estimation of the quantity of reagent used, however. The work of Kaiser and Metzka (9) suggests that more than a 10-fold molar excess of CNBr over methionyl residues does not increase the extent of cleavage. If in doubt as to the concentration of methionyl residues, however, err on the side of higher cyanogen concentration. If accurate quantification of CNBr is required, solid cyanogen bromide may be weighed out and dissolved to a given concentration by addition of the appropriate volume of 70% v/v

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TFA, and the appropriate volume of that solution added to the sample. The CNBr will start to degrade once in aqueous acid, so use when fresh. An alternative is to dissolve the CNBr in acetonitrile, in which it is more stable. CNBr in acetonitrile solution is available commercially, for instance, at a concentration of 5 M (Aldrich). While such a solution may be seen to be degrading by its darkening color, this is not so obvious as it is with CNBr in solid form. For use, sufficient acetonitrile solution is added to the acidic protein solution to give the desired excess of cyanogen bromide over protein (e.g., 1/20 dilution of a 5 M CNBr solution to give a final 250 mM solution). The data of Kaiser and Metzka (9) indicate that high concentrations (70–100%) of acetonitrile can interfere with the cleavage reaction by decreasing the amount of water present, but below a concentration of 30% (in 0.1 M HCl) the effect is noticeable in causing a small decrease of MetSer and Met-Thr bond cleavage, but negligible for the Met-Ala bond. 5. The reagents used are removed by lyophilization, unless salt has formed following failure to remove all the ammonium bicarbonate. The products of cleavage may be fractionated by the various forms of electrophoresis and chromatography currently available. If analyzed by reverse phase HPLC, the reaction mixture may be applied to the column directly without lyophilization. Since methionyl residues are among the less common residues, peptides resulting from cleavage at Met-X may be large and therefore in HPLC, use of wide-pore column materials may be advisable (e.g., 30 µm pore size reverse-phase column, using gradients of acetonitrile in 0.1% v/v TFA in water). Beware that some large peptides that are generated by this technique may prove to be insoluble (fore instance, if the solution is neutralized after the cleavage reaction) and therefore form aggregates and precipitates. 6. Incomplete cleavage, generating combinations of (otherwise) potentially cleaved peptides, may be advantageous, for ordering peptides within a protein sequence. Mass spectrometric methods are suitable for this type of analysis (10). Such partial cleavage may be achieved by reducing the duration of reaction, even to less than 1 h (10). The acid conditions employed for the reaction may lead to small degrees of deamidation of glutamine and asparagine side chains (which occurs below pH 3) and cleavage of acid-labile bonds, for example, Asp-Pro. A small amount of oxidation of cysteine to cysteic acid may occur, if these residues have not previously been reduced and carboxymethylated. Occasional cleavage of Trp-X bonds may be seen, but this does not occur with good efficiency, as it does when the reduction step of this technique is replaced by an oxidation step (for a description of this approach to cleavage of Trp-X bonds). Rosa et al. (13) cleaved both Met-X and Trp-X bonds simultaneously by treatment of protein with 12 mM CNBr in 70% TFA solution, plus 240 µM potassium bromide.

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7. As described in Note 1, the peptide to the N-terminal side of the point of cleavage, has at its C-terminus a homoserine or homoserine lactone residue. The lactone derivative of methionine can be coupled selectively and in good yield (17) to solid supports of the amino type, for example, 3-amino propyl glass. This is a useful technique for sequencing peptides on solid supports. The peptide from the C-terminus of the cleaved protein will, of course, not end in homoserine lactone (unless the C-terminal residue was methionine!) and so cannot be so readily coupled. Similarly, the C-terminal peptide carboxyl can react (if not amidated) with acidic methanol, to become a methyl ester (with a corresponding mass increment of 14 amu). Homoserine lactone, present as the C-terminal residue on other peptides in a CNBr digest, will react with acidic methanol and show a mass increase of 32 amu. With account made for side chain carboxyl residues, this is a means to identify C-terminal peptides by mass spectroscopy (18). 8. Frequently, the protein of interest is impure, in a preparation containing other proteins. Polyacrylamide gel electrophoresis (PAGE) is a popular means by which to resolve such mixtures. Proteins in gel slices may be subjected to treatment with CNBr (14), as follows: The piece of gel containing the band of interest is cut out, lyophilized, and then exposed to vapor from a solution of CNBr in TFA for 24 h, at room temperature in the dark. The vapor is generated from a solution of 20 mg CNBr in 1 mL of 50% v/v TFA by causing it to boil by placing it under reduced pressure, in a sealed container together with the sample. The gel piece is then lyophilized again, and the peptides in it analyzed by PAGE. 9. Proteins that have been transferred from polyacrylamide gel to polyvinylidene difluoride (PVDF) membrane may be cleaved in situ, as described by Stone et al. (15). The protein band (of a few micrograms) is first cut from the membrane on the minimum size of PVDF (as excess membrane reduces final yield). The dried membrane is then wetted with about 50 µL of CNBr solution in acid solution—Stone et al. (15) report the use of CNBr applied in the ratio of about 70 µg per 1 g of protein. Note that although PVDF does not wet directly in water, it does do so in 70% formic acid, or in the alternative of 50% or 70% v/v TFA. Cleavage is achieved by incubation at room temperature, in the dark for 24 h. Oxidation of methionine during electrophoresis and blotting was not found to be a significant problem in causing reduction in cleavage yield, being about 100% in the case of myoglobin (15). The peptides generated by cleavage may be extracted for further analysis, first in the solution of CNBr in the acid solution, second in 100 µL of acetonitrile (40% v/v, 37°C, 3 h), and thirdly in 100 µL of TFA (0.05% v/v in 40% acetonitrile, 50°C. All extracts are pooled and dried under vacuum before any subsequent analysis.

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10. Analysis of protein samples in automated peptide sequencing may sometimes yield no result. The alternative causes (lack of sample or amino-(N)terminal blockage) may be tested by cleavage at methionyl residues by CNBr. Generation of new sequence(s) indicates blockage of the original N-terminus. The method is similar if the sample has been applied to a glass fiber disk, or to a piece of PVDF membrane in the sequencing cartridge. The cartridge containing the filter and/or membrane is removed from the sequencer, and the filter and/or membrane saturated with a fresh solution of CNBr in acid solution. The cartridge is wrapped in sealing film to prevent drying out, and then incubated in the dark at room temperature for 24 h. The sample is then dried under vacuum, replaced in the sequencer and sequence started again. Yields tend to be poorer than in the standard method described in the preceding. If the sample contains more than one methionine, more than one new N-terminus is generated, leading to a complex of sequences. This may be simplified by subsequent reaction with orthophthaladehyde which blocks all N-termini except those bearing a prolyl residue (16). For success with this approach, prior knowledge of the location of proline in the sequence is required, the reaction with orthophthaladehyde being conducted at the appropriate cycle of sequencing. 11. The reagents used are removed by lyophilization, unless salt has formed following failure to remove all of the ammonium bicarbonate. The products of cleavage may be fractionated by the various forms of electrophoresis and chromatography currently available. If analyzed by reverse-phase HPLC, the reaction mixture may be applied to the column directly without lyophilization. As methionyl residues are among the less common residues, peptides resulting from cleavage at Met-X may be large, and so in HPLC use of wide-pore column materials may be advisable (e.g., 30-µm pore size reverse-phase columns, using gradients of acetonitrile in 0.1% v/v TFA in water). Beware that some large peptides that are generated by this technique may prove to be insoluble (e.g., if the solution is neutralized after the cleavage reaction) and so form aggregates and precipitates. References 1. Yuan, G, Bin, J. C., McKay, D. J., and Snyder, F. F. (1999) Cloning and characterization of human guanine deaminase. Purification and partial amino acid sequence of the mouse protein. J. Biol. Chem. 274, 8175–8180. 2. Malouf, N. N., McMahon, D., Oakeley, A. E., and Anderson, P. A. W. (1992). A cardiac troponin T epitope conserved across phyla. J. Biol. Chem. 267, 9269–9274. 3. Dong, M., Ding, X.-Q., Pinon, D. I., Hadac, E. M., Oda, R. P., Landers, J. P., and Miller, L. J. (1999) Structurally related peptide agonist, partial agonist, and antagonist occupy a similar binding pocket within the cholecystokinin receptor. J. Biol. Chem. 274, 4778–4785.

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4. Wallace, D. S., Hofsteenge, J., and Store, S. R. (1990). Use of fragments of hirudin to investigate thrombin-hirudin interaction. Eur. J. Biochem. 188, 61–66. 5. Callaway, J. E., Lai, J., Haselbeck, B., Baltaian, M., Bonnesen, S. P., Weickman, J., et al. (1993). Antimicrob. Agents Chemother. 17, 1614–1619. 6. Strydom, D. J., Tarr, G. E., Pan, Y.-C, E., and Paxton, R. J. (1992). Collaborative trial analyses of ABRF-91AAA, in Techniques in Protein Chemistry III (Angeletti, R. H., ed.) Academic Press, San Diego, New York, Boston, London, Sydney, Tokyo, Toronto, pp. 261–274. 7. Fontana, A. and Gross, E. (1986) Fragmentation of polypeptides by chemical methods in Practical Protein Chemistry A Handbook (Darbre, A., ed. John Wiley and Sons, Chichester, pp. 67–120. 8. Morrison, J. R., Fidge, N. H., and Greo, B. (1990) studies on the formation, separation, and characterisation of cyanogen bromide fragments of human A1 apolipoprotein. Analyt. Biochem. 186, 145–152. 9. Kaiser, R. and Metzka, L. (1999) Enhancement of cyanogen bromide cleavage yields for methionyl-serine and methionyl-threonine peptide bonds. Analyt. Biochem. 266, 1–8. 10. Beavis, R. C. and Chait, B. T. (1990) Rapid, sensitive analysis of protein mixtures by mass spectrometry. Proc. Natl. Acad. Sci. USA 87, 6873–6877. 11. Caprioli, R. M., Whaley, B., Mock, K. K., and Cottrell, J. S. (1991). Sequenceordered peptide mapping by time-course analysis of protease digests using laser description mass spectrometry in Techniques in Protein Chemistry II (Angeletti, R. M., ed.) Academic Press, San Diego, pp. 497–510. 12. Andrews, P. C., Allen, M. M., Vestal, M. L., and Nelson, R. W. (1992) Large scale protein mapping using infrequent cleavage reagents, LD TOF MS, and ES MS, in Techniques in Protein Chemistry II (Angeletti, R. M., ed.) Academic Press, San Diego pp. 515–523. 13. Rosa, J. C., de Oliveira, P. S. L., Garrat, R., Beltramini, L., Roque-Barreira, M.-C., and Greene, L. J. (1999) KM+, a mannose-binding lectin from Artocarpus integrifolia: amino acid sequence, predicted tertiary structure, carbohydrate recognition, and analysis of the beta-prism fold. Protein Science 8, 13–24. 14. Wang, M. B., Boulter, D., and Gatehouse, J. A. (1994) Characterisation and sequencing of cDNA clone encoding the phloem protein pp2 of Cucurbita pepo Plant Mol. Biol. 24, 159–170. 15. Stone, K. L., McNulty, D. E., LoPresti, M. L., Crawford, J. M., DeAngelis, R., and Williams K. R. (1992). Elution and internal amino acid sequencing of PVDF blotted proteins, in Techniques in Protein Chemistry III (Angeletti, R. M., ed.) Academic Press, San Diego, pp. 23–34. 16. Wadsworth, C. L., Knowth, M. W., Burrus, L. W., Olivi, B. B., and Niece, R. L. (1992) Reusing PVDF electroblotted protein samples after N-terminal sequencing to obtain unique internal amino acid sequence, in Techniques in Protein Chemistry III (Angeletti, R. M., ed.) Academic Press, San Diego, pp. 61–68. 17. Horn, M. and Laursen, R. A. (1973) Solid-phase Edman degradation. Attachment of carboxyl-terminal homoserine peptides to an insoluble resin. FEBS Lett. 36, 285–288. 18. Murphy, C. M. and Fenselau, C. (1995) Recognition of the carboxy-terminal peptide in cyanogen bromide digests of proteins. Analyt. Chem. 67, 1644–1645.

95 Chemical Cleavage of Proteins at Tryptophanyl-X Peptide Bonds Bryan John Smith

1. Introduction Tryptophan is represented in the genetic code by a single codon and has proven useful in cloning exercises in providing an unambiguous oligonucleotide sequence as part of a probe or primer. It is also one of the less abundant amino acids found in polypeptides, and cleavage of bonds involving tryptophan generates large peptides. This may be convenient for generation of internal sequence information (the usual purpose to which this technique is put), but is less useful for identification of proteins by mass mapping, where a larger number of smaller peptides makes for more successful database searching. Development of mass spectrometric approaches (determination of short sequences from short peptides) in recent years has meant that methods of cleavage at Tryptophanyl residues have become used more rarely. While there is no protease that shows specificity for tryptophanyl residues, various chemical methods have been devised for cleavage of the bond to the carboxy-(C)-terminal side of tryptophan. These are summarized in Table 1. Some show relatively poor yields of cleavage, and/or result in modification of other residues (such as irreversible oxidation of methionine to its sulfone), or cleavage of other bonds (such as those to the C-terminal side of tyrosine or histidine). The method described in this chapter is one of the better ones, involving the use of cyanogen bromide (CNBr). Cleavage of bonds to the C-terminal side of methionyl residues (see Chapter 94) is prevented by prior reversible oxidation to methionine sulphoxide. Cleavage by use of N-bromosuccinimide or N-chlorosuccinimide remains a popular method, however, despite some chance of alternative reactions (see Table 1).

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Apart from its use in peptide mapping and generation of peptides for peptide sequencing, cleavage at tryptophan residues has also been used to generate peptides used to map the binding site of an antibody (1), or the sites of phosphorylation (2). Another application has been generation of a recombinant protein from a fusion protein (3): tryptophan was engineered at the end of a β-galactosidase leader peptide, adjacent to the N-terminus of phospholipase A2. Reaction with N-chlorosuccinimide allowed subsequent purification of the enzyme without leader peptide. 2. Materials 1. Oxidizing solution: Mix together 30 vol of glacial acetic acid, 15 vol of 9 M HCl, and 4 vol of dimethyl sulfoxide (DMSO). Use best grade reagents. Although each of the constituents is stable separately, mix and use the oxidizing solution when fresh. 2. Ammonium hydroxide (15 M) (see Note 4). 3. CNBr solution in formic acid (60% v/v): Bring 6 mL of formic acid (minimum assay 98%, Aristar grade) to 10 mL with distilled water. Add white crystalline cyanogen bromide to a concentration of 0.3 g/mL. Use when fresh (see Note 5.) Store CNBr refrigerated in the dry and dark, where it is stable for months. Use only white crystals. Beware of the toxic nature of this reagent. Use in a fume hood. 4. Sodium hypocholorite solution (domestic bleach). 5. Equipment includes a fume hood and suitably sized capped tubes (e.g., Eppendorf microcentrifuge tubes).

3. Methods 1. Oxidation: Dissolve the sample to approx 0.5 nmol/µL in oxidizing solution (e.g., 2–3 nmol in 4.9 µL of oxidizing solution). Incubate at 4 °C for 2 h (see Notes 6 and 7). 2. Partial neutralization: To the cold sample, add 0.9 volume of ice-cold NH4OH (e.g., 4.4 µL of NH4OH to 4.9 µL of oxidized sample solution). Make this addition carefully so as to maintain a low temperature (see Note 7.) 3. Cleavage: Add 8 vol of CNBr solution. Incubate at 4 °C for 30 h in the dark. Carry out this step in a fume hood. 4. To terminate the reaction, lyophilize the sample (all reagents are volatile). (See Note 8). 5. Decontaminate equipment such as spatulas that have contacted CNBr, by immersion in bleach until the effervescence stops (a few minutes) and thorough washing.

4. Notes 1. The method described is that of Huang et al. (4). Although full details of the mechanism of this reaction are not clear, it is apparent that tryptophanyl residues are converted to oxindolylalanyl residues in the oxidation step, and the bond to the C-terminal side at each of these is readily

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cleaved in excellent yield (approaching 100% in ref. 4) by the subsequent CNBr treatment. The result is seemingly unaffected by the nature of the residues surrounding the cleavage site. During the oxidation step, methionyl residues become protected by conversion to sulfoxides, bonds at these residues not being cleaved by the cyanogen bromide treatment. Cysteinyl residues will also suffer oxidation if they have not been reduced and alkylated beforehand. Rosa et al. (5) cleaved both Trp-X and Met-X bonds simultaneously by omission of the oxidation step and inclusion of 240 µM potassium iodide in the reaction of protein with 12 mM CNBr in 70% TFA solution. The peptide to the C-terminal side of the cleavage point has a free amino-(N)-terminus and so is suitable for amino acid sequencing by Edman chemistry. 2. Methionyl sulfoxide residues in the peptides produced may be converted back to the methionyl residues by incubation in aqueous solution with thiols (e.g., dithiothreitol, as described in ref. 3, or see use of 2-mercaptoethanol above). 3. The acid conditions used for oxidation and cleavage reactions seem to cause little deamidation (4), but one side reaction that can occur is hydrolysis of acid-labile bonds. The use of low temperature minimizes this problem. If a greater degree of such acid hydrolysis is not unacceptable, speedier and warmer alternatives to the reaction conditions described above can be used as follows: a. Oxidation at room temperature for 30 min, but cool to 4 °C before neutralization. b. Cleavage at room temperature for 12–15 h. 4. As alternatives to the volatile base NH4OH, other bases may be used (e.g., the nonvolatile potassium hydroxide or Tris base). 5. Formic acid is a good protein denaturant and solvent, and is volatile and so relatively easy to remove. However, it has been noted that use of formic acid can cause formulation of seryl and threonyl residues (6) in the polypeptide (seen as an 28 amu increase in molecular mass) and damage to tryptophan and tyrosine (evidenced by spectral changes [7]). As an alternative to formic acid, 5 M acetic acid may be used, or as in the use of CNBr in cleaving methionyl-X bonds, 70% (v/v) trifluoroacetic acid may prove an acceptable alternative (5). 6. Samples eluted from sodium dodecylsulfate (SDS) gels may be treated as described, but for good yields of cleavage, Huang et al. (4) recommend that the sample solutions are acidified to pH 1.5 before lyophilization in preparation for dissolution in the oxidizing solution. Any SDS present may help to solubilize the substrate and, in small amounts at least, does not interfere

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with the reaction. However, nonionic detergents that are phenolic or contain unsaturated hydrocarbon chains (e.g., Triton, Nonidet P-40) and reducing agents are to be avoided. 7. The method is suitable for large-scale protein cleavage, requiring simple scaling up. Huang et al. (4) made two points, however: a. The neutralization reaction generates heat. As this might lead to protein or peptide aggregation, cooling is important at this stage. Ensure that the reagents are cold and are mixed together slowly and with cooling. A transient precipitate is seen at this stage. If the precipitate is insoluble, addition of SDS may solubilize it (but will not interfere with the subsequenttre atment). b. The neutralization reaction generates gases. Allow for this when choosing a reaction vessel. 8. At the end of the reaction, all reagents may be removed by lyophilization and the peptide mixture analyzed, for example, by polyacrylamide gel electrophoresis or by reverse-phase high-performance liquid chromatography (HPLC). Peptides generated may tend to be large, ranging up to a size of the order of 10,000 Da or more. Some of these large peptides may not be soluble, for example, if the solution is neutralized following the cleavage reaction, and consequently they aggregate and precipitate. 9. Note that all reactions are performed in one reaction vial, eliminating transfer of sample between vessels, and so minimizing peptide losses that can occur in such exercises. 10. Alternative methods for cleavage of tryptophanyl-X bonds are outlined in Table 1. The method (vi) that employs N-chlorosuccinimide is the most specific but shows only about 50% yield. BNPS-skatole is a popular TrpX-cleaving reagent whose reaction and products have been studied in some detail (e.g., see refs. 16 and 17) 11. Both BNPS-skatole and N-chlorosuccinimide methods (Table 1, iv and vi, respectively) have been adapted to cleave small amounts (micrograms or less) of proteins on solid supports or in gels. Thus, proteins bound to glass fiber (as used in automated peptide sequences) may be cleaved by wetting the glass fiber with 1 µg/mL BNPS-skatole in 70%, (v/v) acetic acid, followed by incubation in the dark for 1 h at 47 °C. After drying, sequencing may proceed as normal. Alternatively protein blotted to polyvinylidene difluoride membrane may be similarly treated and resulting peptides eluted for further analysis (11). Alternatively, protein in slices of polyacrylamide gel (following polyacrylamide gel electrophoresis [SDS–PAGE]) may be cleaved by soaking for 30 min 0.015 M in N-chlorosuccinimide, in 0.5 g/ mL of urea in 50%, (v/v) acetic acid. Following washing, peptides may be electrophoresed to generate peptide maps (14).

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References 1. Kilic, F. and Ball, E. H. (1991). Partial cleavage mapping of the cytoskeletal protein vinculin. Antibody and talin binding sites. J. Biol. Chem. 266, 8734–8740. 2. Litchfield, D. W., Lozeman, F. J., Cicirelli, M. F., Harrylock, M., Ericsson, L. H., Piening, C. J., and Krebs. E. G. (1991) Phosphorylation of the beta subunit of casein kinase II inhuman A431 cells. Identification of the autophosphorylation site and a site phosphorylated by p34cdc2. J. Biol. Chem. 266, 20,380–20,389. 3. Tseng, A., Buchta, R., Goodman, A. E., Loughman, M., Cairns D., Seilhammer, J., et al. (1991) A strategy for obtaining active mammalian enzyme from a fusion protein expressed in bacteria using phospholipase A2 as a model. Pro. Express. and Purificat. 2, 127–135. 4. Huang, H. V., Bond, M. W., Hunkapillar, M. W., and Hood, L. E. (1983) Cleavage at tryptophanyl residues with dimethyl sulfoxide-hydrochloric acid and cyanogen bromide. Meth. Enzymol. 91, 318–324. 5. Rosa, J. C., de Oliveira, P. S. L., Garrat, R., Beltramini, L., Roque-Barreira, M.-C. and Greene, L. J. (1999) KM+, a mannose-binding lectin from Artocarpus integrifolia: amino acid sequence, predicted tertiary structure, carbohydrate recognition, and analysis of the beta-prism fold. Prot. Sci. 8, 13–24. 6. Beavis, R. C. and Chait. B. T. (1990). Rapid, sensitive analysis of protein mixtures by mass spectrometry. Proc. Natl. Acad. Sci. USA 87, 6873–6877. 7. Morrison, J. R., Fidge, N. H., and Grego, B. (1990) Studies on the formation, separation, and characterisation of cyanogen bromide fragments of human A1 apolipoprotein. Analyt. Biochem. 186, 145–152. 8. Ozols, J. and Gerard, C. (1977). Covalent structure of the membranous segment of horse cytochrome b5. Chemical cleavage of the native hemprotein. J. Biol. Chem. 252, 8549–8553. 9. Savige, W. E. and Fontana, A. (1977) Cleavage of the tryptophanyl peptide bond by dimethyl sulfoxide-hydrobromic acid. Meth. Enzymol. 47, 459–469. 10. Fontana, A. (1972) Modification of tryptophan with BNPS skatole (2-(2-nitrophenylsulfenyl)-3-methyl-3′-bromoindolenine). Meth. Enzymol. 25, 419–423. 11. Crimmins, D. L., McCourt, D. W., Thoma, R. S., Scott, M. G., Macke, K. and Schwartz, B. D. (1990) In situ cleavage of proteins immobilised to glass-fibre and polyvinylidene difluoride membranes: cleavage at tryptophan residues with 2-(2´-nitropheylsulfenyl)-3-methyl-3′ -bromoindolenine to obtain internal amino acid sequence. Analyt. Biochem. 187, 27–38. 12. Ramachandran, L. K. and Witkop, B. (1976). -Bromosuccinimide cleavage of peptides. Meth. Enzymol. 11, 283–299. 13. Lischwe, M. A. and Sung, M. T. (1977). Use of -chlorosuccinimide/urea for the selective cleavage of tryptophanyl peptide bonds in proteins. J. Biol. Chem. 252, 4976–4980. 14. Lischwe, M. A. and Ochs, D. (1982). A new method for partial peptide mapping using chlorosuccinimide/urea and peptide silver staining in sodium dodecyl sulphatepolyacry-lamide gels. Analyt. Biochem. 127, 453–457. 15. Fontana, A., Dalzoppo, D., Grandi, C., and Zambonin, M. (1983) Cleavage at tryptophan with iodosobenzoic acid. Meth. Enzymol. 91, 311–318.

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16. Vestling, M. M., Kelly, M. A., and Fenselau, C. (1994) Optimization by mass spectrometry of a tryptophan-specific protein cleavage reaction. Rapid Commun. Mass Spectr. 8, 786–790. 17. Rahali, V. and Gueguen, J. (1999) Chemical cleavage of bovine (β-lactoglobulin by BNPS-skatole for preparative purposes: comparative study of hydrolytic procedures and peptide characterization. J. Prot. Chem. 18, 1–12.

96 Chemical Cleavage of Proteins at Aspartyl-X Peptide Bonds Bryan John Smith

1. Introduction Some methods for chemically cleaving proteins, such as those described in Chapters 94 and 95, are fairly specific for a particular residue, show good yields, and generate usefully large peptides (as reaction occurs at relatively rare amino acid residues). There may be cases, however, when such peptides (or indeed, proteins) lacking these rarer residues need to be further fragmented, and in such instances cleavage at the more common aspartyl residue may prove useful. The method described in this chapter (and in ref. 1) for cleavage to the carboxy-(C)terminal side of aspartyl residues is best limited to smaller polypeptides rather than larger proteins because yields are 100 kDa are typically electrophoresed in 7–10% polyacrylamide gels, whereas smaller proteins are electrophoresed in 10–17.5% polyacrylamide. The in gel digestion procedure is suitable for both 1 and 2 dimensional gels with the latter digests often performed on a digester robot. The protein of interest is excised using a razor blade and tweezers, and the gel spots are then placed in an eppendrof tube that has been pre-washed with 0.1% TFA/ 60% CH3CN (to minimize contaminants). Alternatively, a robotic gel picker such as the GE Healthcare Ettan Spot Picker can be used to excise the proteins of interest. This spot picker cuts out a 2mm disc and deposits the gel piece in a 96 well plate. Care must always be taken to avoid keratin contamination which can confound any protein identification analysis. Figure 1 contains a typical MALDI-Tof/Tof spectra of an in gel tryptic digest. 3.2.2. In-Gel Digestion of Proteins with Trypsin and Lysyl Endopeptidase 1. Determine the approximate volume of gel to be digested (length × width × thickness). 2. Cut the gel band(s) containing the protein of interest into approx 1 × 2 mm pieces, and place them in an Eppendorf tube or a 96 well plate for robotic digestion. Repeat for the “blank” section of gel.

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Fig. 1. In-gel trypsin digestion of an unknown protein spot from a 2D gel. Human plasma was analyzed by 2D gel electrophoresis following IgY12 depletion (Beckman-Coulter) and Cy dye labeling as per the GE Healthcare Differential (fluorescence) gel electrophoresis protocol. Protein spots of interest were determined using GE Healthcare DeCyder software of the fluorescent gel image, and excised robotically (GE Healthcare Ettan Spot Picker) as described in Subheading 3. 100% of the digest was loaded onto an AB 4800 MALDI-Tof/ Tof plate after dissolving in 0.6nl of the alpha-cyano-4-hydroxy cinnamic acid matrix containing 1 fmol of bradykinin and 2 fmols of ACTH clip as internal standards. The sample was analyzed using both peptide mass fingerprint and MS/MS of 10 peptides. The inset panel shows the MS/MS spectra obtained from the mass at 1694.9. A Mascot (Matrix Science) database search identified this sample as Vitamin D-binding protein precursor. The estimated amount of sample analyzed is 0.5 to 1 fmol based on the bradykinin internal standard.

3. Add 250 µL (or more, if necessary to cover the gel pieces) 50% CH3CN/50% water to the gel pieces. 4. Wash for 5 min at room temperature on a rocker table. 5. Remove wash. 6. Repeat steps 3–5 using 250 µl 50% CH3CN/50mM NH4HCO3 7. Repeat steps 3–5 using 250 µl 50% CH3CN/10mM NH4HCO3 8. Dry the washed gel pieces in a SpeedVac 9. Make up the enzyme solution by diluting 5 µL 0.1 mg/mL enzyme stock solution with 10 µL 10mM NH4HCO3 for every 15 mm3 gel that is to be digested.

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10. Rehydrate gel pieces with the enzyme solution from step 9, which should be equal in volume to that of the gel pieces and should provide a final enzyme ratio of about 0.5 µg/15-mm3 gel volume. 11. If the gel pieces are not totally immersed in the enzyme solution, add an additional volume of 10mM NH4HCO3 to submerge the gel pieces totally. 12. Incubate at 37 °C for 5 to 16 h. 13. After incubation, the supernatant can be removed and injected on to a HPLC column or LC-MS/MS system. Gels pieces can also be further extracted as in steps 19–20 and Cysteine modification, if desired, can be performed as follows in steps 14 – 18 14. Estimate the total volume of the sample plus gel. 15. Calculate the volume of 45 mM DTT needed to give a final (DTT) of about 1 mM in the sample. 16. Add the above volume of 45 mM DTT and then incubate at 50 °C for 20 min. 17. Remove samples from the incubator; cool to room temperature and add an equal volume of 100 mM IAA or 200 mM methyl 4-nitrobenzene sulfonate (MNS) (see Note 10) as the volume of DTT added in step 16. 18. For IAA alkylation, incubate at room temperature in the dark for 20 min; for MNS treatment, incubate at 37 °C for 40 min. 19. Extract peptides by adding at least 100 µL 0.1% TFA, 60% CH3CN (or, if it is greater, a volume equal to the gel volume estimated in step 2), and shake on a rocker table at room temperature for at least 40 min. 20. Sonicate for 5 min in a water bath sonicator. 21. Remove and save the supernatant, which contains the released peptides, and repeat steps 19 and 20. 22. SpeedVac dry the combined washes from step 21. 23. Redissolve the dried samples in 3 to 5 µL 70% formic acid (to dissolve the peptides) and bring to proper injection volume using 0.1% TFA. 24. The sample is now ready for reverse-phase HPLC peptide separation as described in Chapter 102 or mass spectrometric analysis.

3.2.3. In-Gel Robotic Digestion of Proteins with Trypsin A digester robot such as the GE Healthcare Ettan Ta digester is required for robotic digestion. 1. Excise the protein of interest by hand or using a gel picker (e.g. Ettan spot picker) and place in an 96 well plate (see Note 11). 3. Add 50 µL 50% CH3CN/50% water to the gel pieces. 4. Wash for 3 min at room temperature. 5. Remove wash. 6. Repeat steps 3–5. 7. Repeat steps 3–5 using 50 µl 5mM NH4HCO3. 8. Air dry gel pieces for 5 min. 9. Make up the enzyme solution by diluting 100 µL 0.1 mg/mL enzyme stock solution with 2.9 mL 5mM NH4HCO3. 10. Rehydrate gel pieces with 15 µl of the enzyme solution from step 9.

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11. Incubate at 37 °C for 5 to 16 h. 12. After incubation, the supernatant can be removed and injected on to a HPLC column or LC-MS/MS system. For MALDI-MS analysis, speedvac the digest to dryness and re-dry two times from 20 µl water. The sample is now ready to be dissolved in matrix and analyzed.

4. Notes 1. In many instances, large losses occur during the final purification steps when the protein concentrations are invariably lower. Hence, although ultrafiltration or dialysis of a 5 mg/mL crude solution of a partially purified enzyme may lead to nearly 100% recovery of activity, similar treatment of a 25 µg/mL solution of the purified protein might well lead to significant, if not total loss of activity owing to nonspecific adsorption. Similarly, the effectiveness of organic and acid-precipitation procedures often decreases substantially as the final protein concentration is decreased below about 100 µg/mL. Whenever possible, therefore, the final purification step should be arranged such that the resulting protein solution is as concentrated as possible and, ideally, can simply be dried in a SpeedVac prior to enzymatic digestion. In this regard, it should be noted that a final NaCl concentration of 1 M does not significantly affect the extent of trypsin digestion (8). When it is necessary to carry out an organic or acid precipitation to remove salts or detergents, the protein should first be dried in a SpeedVac (in the 1.5-mL tube in which it will ultimately be digested) prior to redissolving or suspending in a minimum volume of water and then adding the acetone or TCA. In this way, the protein concentration will be as high as possible during the precipitation and losses will be minimized. Two common contaminants that are extremely deleterious to enzymatic cleavage are detergents (as little as 0.005% SDS will noticeably decrease the rate of tryptic digests carried out in the presence of 2 M urea [8]) and ampholines. Since detergent removal is often associated with protein precipitation, and since many detergents (such as SDS) form large micelles, which cannot be effectively dialyzed, it is usually preferable to extract the detergent from the protein (that has been dried in the tube in which it will be digested) rather than to dialyze it away from the protein. In the case of ampholines, our experience is that even prolonged dialysis extending over several days with a 15,000-Dalton cutoff membrane is not sufficient to decrease the ampholine concentration to a level that permits efficient trypsin digestion. Rather, the only effective methods we have found for complete removal of ampholines are TCA precipitation, hydrophobic or reverse-phase chromatography, or SDS-PAGE followed by staining and destaining.

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2. It is particularly important to quantify samples using amino acid analysis if protein profiling (i.e. comparative analysis) is to be done. For samples in solution, the aliquot for amino acid analysis should be taken either immediately prior to drying the sample in the tube in which it will be digested or after redissolving the sample in 8 M urea, 0.4 M NH4HCO3. Although up to 10 µL of 8 M urea is compatible with acid hydrolysis/ion-exchange amino acid analysis, this amount of urea may not be well tolerated by PTC amino acid analysis. Hence, in the latter case, the amino acid analysis could be carried out prior to drying and redissolving the sample in urea. Samples destined for comparative 2D gel analysis such as DIGE should also be quantified using amino acid analysis prior to running the 2D gel. The aliquot for analysis can be taken after the samples have been dissolved in the gel loading buffer. By having an accurate protein concentration, comparative protein profiling experiments are themselves more accurate. 3. Since many native proteins are resistant to enzymatic cleavage, it is usually best to denature the protein prior to digestion. Although some proteins may be irreversibly denatured by heating in 8 M urea, this treatment is not sufficient to denature transferrin. In this instance, prior carboxymethylation, which irreversibly modifies cysteine residues, brings about a marked improvement in the resulting tryptic peptide map (8). Another advantage of carboxymethylating the protein is that this procedure enables cysteine residues to be identified during edman amino acid sequencing. Cysteine residues have to be modified in some manner prior to edman sequencing to enable their unambiguous identification. Under the conditions that are described in Subheading 3., the excess dithiothreitol and iodoacetic acid do not interfere with subsequent digestion. Although carboxymethylated proteins are usually relatively insoluble, the 2 M urea that is present throughout the digestion is frequently sufficient to maintain their solubility. However, even in those instances where the carboxymethylated protein precipitates following dilution of the 8 M urea to 2 M, trypsin and chymotrypsin will usually still provide complete digestion. Often, the latter is evidenced by clearing of the solution within a few minutes of adding the enzyme. Alternatively, the newer denaturants RapiGest™ and PPS Silent™ Surfactant can be used to try to better solubilize the proteins of interest. If carboxymethylation is insufficient to bring about complete denaturation of the substrate, an alternative approach is to cleave the substrate with cyanogen bromide (1000-fold molar excess over methionine, 24 h at room temperature in 70% formic acid). The resulting peptides can then either be separated by SDSPAGE (since they usually do not separate well by reverse-phase HPLC) or, preferably, they can be enzymatically digested with trypsin or lysyl endopeptidase and then separated by reverse phase HPLC or LC-MS/MS.

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If this approach fails, the protein may be digested with pepsin, which, as described above, is carried out under very acidic conditions or can be subjected to partial acid cleavage (9). However, the disadvantage of these latter two approaches is that they produce an extremely complex mixture of overlapping peptides. Finally, extensive glycosylation (i.e., typically >10– 20% by weight) can also hinder enzymatic cleavage. In these instances, it is usually best to remove the carbohydrate prior to beginning the digest. In the case of in gel digests, this may often be best carried out immediately prior to SDS-PAGE, which thus prevents loss (owing to insolubility) of the deglycosylated protein and effectively removes the added glycosidases. 4. Every effort should be made to be used as a high substrate and enzyme concentration as possible to maximize the extent of cleavage. Although the traditional 1:25, weight:weight ratio of enzyme to substrate provides excellent results with milligram amounts of protein, it will often fail to provide complete digestion with low nanogram amounts of protein. For instance, using the procedures outlined above, this weight:weight ratio is insufficient to provide complete digestion when the substrate concentration falls below about 20 µg/mL (8). The only reasonable alternative to purifying additional protein is either to decrease the final digestion volume or to compensate for the low substrate concentration by increasing the enzyme concentration. The only danger in doing this, of course, is the increasing risk that some peptides may be isolated that are autolysis products of the enzyme. Assuming that only enzymes, such as trypsin, chymotrypsin, lysyl endopeptidase, and Protease V8 are used, whose sequences are known, it is usually better to risk sequencing a peptide obtained from the enzyme (which can be quickly identified via a data base search) than it is to risk incomplete digestion of the substrate. Often, protease autolysis products can be identified by comparative HPLC peptide mapping of an enzyme (i.e., no substrate) control that has been incubated in the same manner as the sample and by subjecting candidate HPLC peptide peaks to matrix-assisted laser desorption mass spectrometry prior to edman sequencing. The latter can be extremely beneficial both in identifying (via their mass) expected protease autolysis products and in ascertaining the purity of candidate peptide peaks prior to edman sequencing. For sequencing by LC-MS/MS analysis, it is not as critical to eliminate the trypsin/enzyme autolysis products since a database search will identify these peptides. To promote more extensive digestion, we have sometimes used enzyme:substrate mole ratios that approach unity. 5. One approach to identifying disulfide-linked peptides is to comparatively HPLC peptide map a digest that has been reduced/carboxymethylated vs one that has only been carboxymethylated (thus leaving disulfide-linked

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7.

8.

9.

10.

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Stone et al. peptides intact). In this instance, pepsin offers an advantage in that the digest can be carried out under acidic conditions where disulfide interchange is less likely to occur. As in the case of in-solution digests, care must be exercised to guard against sample loss during final purification. Whenever possible, SDS (0.05%) should simply be added to the sample prior to drying in a SpeedVac and subjecting to SDS-PAGE. Oftentimes, however, if the latter procedure is followed, the final salt concentration in the sample will be too high (i.e., >1 M) to enable it to be directly subjected to SDS-PAGE. In this instance, the sample may either be concentrated in a SpeedVac and then precipitated with TCA (as described above) or it may first be dialyzed to lower the salt concentration. If dialysis is required, the dialysis tubing should be rinsed with 0.05% SDS prior to adding the sample, which should also be made 0.05% in SDS. After dialysis versus a few micromolar NH4HCO3 containing 0.05% SDS, the sample may be concentrated in a SpeedVac and then subjected to SDS PAGE (note that samples destined for SDS PAGE may contain several % SDS). Another approach that works extremely well is to use a SDS polyacrylamide gel containing a funnel-shaped well that allows samples to be loaded in volumes as large as 300 µL (10). In general, the sample should be run in as few SDS-PAGE lanes as possible to maximize the substrate concentration and to minimize the total gel volume present during the digest. Whenever possible, a 0.5–0.75mm thick gel should be used. In general, we recommend using 0.5 µg enzyme/15 mm3 of gel with the only caveat being that we use a corresponding lower amount of enzyme if the mole ratio of protease/substrate protein would exceed unity. Since high concentrations of Coomassie blue interfere with digestion, it is best to use the lowest Coomassie blue concentration possible and to stain for the minimum time necessary to visualize the bands of interest. In addition, the gel should be well destained so that the background is close to clear. Although most estimates of protein amounts are based on relative staining intensities, our data suggest there is a 5–10-fold range in the relative staining intensity of different proteins. Obviously, when working in the low pmol/fmol levels, such a 5–10-fold range could well mean the difference between success and failure. For edman sequencing, modification of cysteine residues with MNS provides the advantage that the resulting S-methyl cysteine phenylthiohydantoin derivative elutes in a favorable position (i.e., between PTH-Tyr and PTH-Pro) using an Applied Biosystems PTH C-18 column. Due to the manner in which the robot operates, it is necessary to not digest too large a gel volume. Recommended gel volumes are 2mm discs.

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Acknowledgments Much of the methods described above were used in studies funded with Federal funds from NHLBI/NIH contract N01-HV-28186, NIDA/NIH grant 1 P30 DA018343-01, and NIAD/NIH grant 5 U54 AI057158-02 (Northeast Biodefense Center - Regional Centers of Excellence).

References 1. Henzel, W.J., Watanabe, C., and Stults J.T., (2003) Protein identification: the origins of peptide mass fingerprinting. J. Am. Soc. Mass Spectrom. 14, 931–942. 2. Yates, J.R., 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, 1426–1436. 3. Perkins, D.N., Pappin, D, J.C., Creasy, D.M. and Cottrell, J. S. (1999) Probabilitybased protein identification by searching sequence databases using mass spectrometry data. Electrophoresis 20, 3551–3567. 4. Williams. K. R. and Stone, K. L. (1995) In gel digestion of SDS PAGE-separated protein: observations from internal sequencing of 25 proteins in Techniques in Protein Chemistry VI (Crabb, J. W., ed.), Academic, San Diego, pp. 143–152. 5. Rosenfeld, J., Capdevielle, J., Guillemot, J., and Ferrara, P. (1992) In gel digestions of proteins for internal sequence analysis after 1 or 2 dimensional gel electrophoresis. Analyt. Biochem. 203, 173–179. 6. Fernandez, J., DeMott, M., Atherton, D., and Mische, S. M. (1992) Internal protein sequence analysis: enzymatic digestion for less than 10 µg of protein bound to polyvinylidene difluoride or nitrocellulose membranes. Analyt. Biochem. 201, 255–264. 7. Aebersold, R. H., Leavitt, J., Saavedra, R. A., Hood, L. E., and Kent, S. B. (1987) Internal amino acid sequence analysis of proteins separated by one- or two-dimensional gel electrophoresis after in situ protease digestion on nitrocellulose. Proc. Natl. Acad. Sci. USA 84, 6970–6974. 8. Stone, K. L., LoPresti, M. B., and Williams, K. R. (1990) Enzymatic digestion of proteins and HPLC peptide isolation in the subnanomole range in Laboratory Methodology in Biochemistry: Amino Acid Analysis and Protein Sequencing (Fini, C., Floridi, A., Finelli, V., and Wittman-Liebold, B., eds.), CRC, Boca Raton, FL, pp. 181–205. 9. Sun, Y., Zhou, Z., and Smith, D. (1989) Location of disulfide bonds in proteins by partial acid hydrolysis and mass spectrometry in Techniques in Protein Chemistry (Hugli, T., ed.), Academic Press, San Diego, pp. 176–185. 10. Lombard-Platet, G. and Jalinot, P. (1993) Funnel-well SDS-PAGE: a rapid technique for obtaining sufficient quantities of low-abundance proteins for internal sequence analysis. BioTechniques 15, 669–672.

100 On-PVDF Protein Digestions for N-terminal Sequencing and Peptide Mass Fingerprinting Victoria Pham, William Henzel, and Jennie R. Lill

1. Introduction Two common analytical techniques employed for the characterization of proteins are N-terminal sequencing and peptide mass fingerprinting (PMF). (1) N-terminal sequencing employing the chemistries first described by Pehr Edman (2) is routinely used in many laboratories for deciphering if correct translational and post-translational processing of a protein has occurred. Peptide mass fingerprinting on the other hand, is used to determine if the correct protein has been cloned and translated, to look for mutations within a known protein sequence, and to look for posttranslational modifications. A typical format for the N-terminal sequencing of protein is to transfer the sample onto a PVDF membrane. This is achieved by first separating proteins by SDS-PAGE, followed by electro-blotting onto PVDF membrane where they remain immobilized and stable until analysis. Edman chemistries require a free α-amino group at the N-terminus of the protein in order for phenylisothiocyante (PITC) to couple with the amino group of the terminus. However, many proteins expressed in mammalian cell lines are N-terminally blocked with co- or post translational modifications such as pyroglutamyl, acetyl or formyl groups, which render Edman chemistry inaccessible. Several protocols are available to remove these N-terminal modifications for example (deacetylation and pyroglutamyl). However, in some cases, e.g. for proteins with a pyroglutamyl group followed by a proline residue, removal of the modified group is not possible due to hindered stereochemistry. Under such situations, PVDF-imbedded N-terminally blocked proteins can be identified from their internal fragments generated from on-PVDF chemical or proteolytic cleavage.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Peptide mass fingerprinting (PMF) involves the enzymatic or chemical proteolytic digestion of a protein into constituent peptides. Each protein will produce a unique signature of resultant peptides that can be compared against a list of theoretical masses to identify the protein, and also to look for alterations in sequence such as mutations and post translational modifications (3). For characterization purposes, depending on the availability of the protein and the information that one needs to obtain, both Edman degradation and PMF analysis can be utilized since the 2 techniques are complimentary with each other. Here we describe protocols for the rapid on-PVDF enzymatic cleavage of proteins which can subsequently be analyzed either by Edman degradation or by PMF. 2. Materials 2.1. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. Running buffer (10X): 247 mM Tris, 1.92 M glycine, 34.7 mM SDS, store at room temperature (RT). 2. Precast 4-20% Tris-glycine (Invitrogen, Carlsbad, CA). 3. Sample buffer (2X): 0.5 M Tris-HCl, pH 8.0, 20% SDS, 0.5% bromophenol blue, 26 % glycerol (in-house), store at RT. 4. Dithiothreitol DTT (BioVectra, Oxford, CT). 5. N-isopropyl iodoacetamide (NIPIA) (synthesized in-house).

2.2. Electroblotting 1. CAPS buffer: 10 mM, pH 11.0 containing 10 mM thioglycolic acid and 20% methanol, store at RT. 2. Problott PVDF membrane (Applied Biosystems, Foster City, CA). 3. Methanol at HPLC-grade (Burdick & Jackson, Muskegon, MI).

2.3. Staining/Destaining of PVDF Membrane 1. Staining solution: 0.1% Coomasie brilliant blue R250 in 50% methanol/10% acetic acid (in-house), store at RT. 2. Destaining solution: 50% methanol/10% acetic acid/MQ water (in-house), store at RT.

2.4. On-Membrane Chemical Cleavage for Edman Degradation 1. Mouse antihuman OX40 ligand antibody (R & D Systems, Minneapolis, MN). 2. Asp/Pro cleavage solution: 10% acetic acid containing 7 M guanidine, pH 2.5.

2.5. Rapid on-Membrane Digestion for Automated Edman Degradation 1. Human IL-20 receptor alpha poly his (purified in-house). 2. Clostripain (Calbiochem, San Diego, CA), trypsin (Promega, Madison, WI) store at −80 °C. 3. Zwittergent 3–16 (Calbiochem, San Diego, CA). 4. 0.1 M Tris-HCl, pH 8.0, 2 mM DTT, 1 mM CaCl2.

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2.6. On-Membrane Digestion for Peptide Mass Fingerprinting (PMF) 1. Recombinant human growth hormone (rec-HGH) (purified in-house), store at −80 °C. 2. Matrix α-cyano-4-hydroxycinnaminic acid (CHCA): nitrocellulose (NC) (20:5 w/w) in acetone: isopropyl alcohol at 1:1 ratio. 3. α-Cyano-4-hydroxycinnamic Acid (CHCA) (Sigma, St Louis, MO), NC (BioRad, Hercules, CA). 4. Acetone, isopropyl alcohol, acetonitrile at HPLC-grade (Burdick & Jackson, Muskegon, MI).

2.7. Instrumentation 1. Procise N-terminal sequencer, model 494 (Applied Biosystems, Foster City, CA). 2. Voyager-DE-STR MALDI-TOF (Applied Biosystems, Foster City, CA).

3. Methods On-PVDF chemical cleavage has been widely used without pretreatment of the membrane. However, when on-PVDF enzymatic digestion is performed, the unoccupied sites on the membrane must be blocked. If these sites are not blocked enzyme activity maybe hindered, and also the ratio of enzyme to analyte upon analysis will be overwhelming. PVP-360 has previously been reported as a blocking agent for on-membrane digestion (4) and more recently we reported the use of Zwittergent 3–16, a sulfobetaine type detergent, with a hydrophobic un-branched 16-C chain, which binds strongly to the PVDF membrane, as an alternative surfactant (3). Our results indicated that Zwittergent 3–16 is a better blocking agent than PVP-360, and as a consequence more efficient on-PVDF digestion for N-terminal sequencing or PMF was observed. 3.1. SDS-PAGE and Electroblotting 1. Proteins are reduced with DTT (10 mM in 2X sample buffer) at 95 °C for 10 min and alkylated with NIPIA at a final concentration of 20 mM at RT for 20 min. 2. Proteins are then separated on a precast 4–20% Tris-glycine gel for 120 min at 120 V.

3.2. Electroblotting 1. Following separation, equilibrate gel in CAPS buffer (see Note 1) at RT for 10 min. 2. Electro-blot gel onto a prewetted PVDF membrane in CAPS buffer for 45-60 min at 250 mA (see Note 2)

3.3. Staining & destaining 1. Visualize proteins by staining in coomasie blue R250 staining solution for 30–60 s. 2. Destain PVDF membrane with 50% MeOH/ 10% acetic acid/H2O for 2 min and extensively rinse with MQ H2O (10 × 1.0 mL). 3. Store membrane at RT until further analysis.

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3.4. On-PVDF Chemical Cleavage for Edman Degradation 1. Excise the band of interest and wet with 1–2 µL of methanol, rinsed with MQ water (10 × 1.0 mL). 2. Add 30 µL of diluted acid (2.4) and incubate in a thermocycler tube at 90 °C for 1 h (see Note 3) 3. After incubation, wash the protein band with MQ water (10 × 1.0 mL) and dry in the Speed Vac. 4. Peptides can then be analyzed by N-terminal sequencing to identify the protein (see Note 4).

3.5. Rapid on-Membrane Digestion for Automated Edman Degradation 1. Protein bands are wetted with 1-2 µL of methanol, rinsed with MQ water (10 × 1.0 mL). 2. Treat PVDF membranes with 100 µL of Zwittergent 3-16, 0.5% in Tris (0.1 M, pH 8.0) and incubate on a shaker at RT for 5 min to block the unoccupied sites on the membrane. 3. Wash membranes extensively with MQ water (10 × 1.0 mL). 4. After blocking the membranes, add appropriate digestion buffer (20 µL) and enzyme of choice (0.2 µg) and start digestion at 37 °C for 5–60 min. Table 1 lists the digestion conditions of different enzymes. 5. After incubation, add 3 µL of 4% TFA/water to quench the digestion. 6. Rinse membranes with MQ water (10 × 1.0 mL) and dry in the Speed Vac.

Table 1 Digestion conditions of various enzymes (*) Enzymes Trypsin Lys-C

Buffers Tris (0.1 M, pH 8.0)

Ammonium bicarbonate (50 mM, pH 8.0) Clostripain Tris (0.1 M, pH 8.0) containing 2 mM DTT, 1 mM CaCl2. Glu-C Ammonium bicarbonate (0.1 M, pH 8.) Glu-C Sodium phosphate (50 mM, pH 8.0)

Time (min) 10 10–30

Specificity C-terminal side of K&R C-terminal side of K

10–20

C-terminal side of R

10–30 10–30

C-terminal side of E C-terminal side of E&D Lys-N Sodium phosphate (50 mM, pH 8.0) 10–20 N-terminal side of K Asp-N Sodium phosphate (50 mM, pH 8.0) 10–20 N-terminal side of D PGAP Sodium phosphate (50 mM, pH 7.0) 60 Removal of containing 10 mM DTT, 1mM EDTA pyroglutamyl group (*) Pyroglutamyl aminopeptidase (PGAP) digestion was performed at 90 °C and the rest was performed at 37 °C

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7. Analyze the resultant mixture of peptides as mentioned in Subheading 3.4 (see Note 5).

3.6 On-Membrane Digestion for Peptide Mass Fingerprinting (PMF) 1. Block protein bands with Zwittergent solution as described in Subheading 3.5. 2. Carefully remove Zwittergent solution and rinse the membrane thoroughly with MQ water (10 × 1.0 mL) (see Note 6). 3. Heat protein bands to 95 °C for 5-10 min in the appropriate digestion buffer in the presence of 10% acetonitrile (see Notes 7 and 8). 4. Add the enzyme of choice to the digestion buffer and incubate at 37 °C for 1-3 h. 5. After incubation, quench the reaction with 3 µL of 4% TFA/water. 6. Spot the resultant peptide mixture (0.2-1.0 µL) onto the MALDI target plate using a nitrocellulose-base method (see Note 9) and air dry. 7. After the organic matrix is dried, wash sample spots with 1 µL of 0.1% TFA/water prior to MALDI analysis to remove any trace amount of salt. 8. Perform PMF on a MALDI-TOF instrument in reflectron mode. (Note, typically we employ the following conditions: mass range is set to 600-6000 Da at a laser power of 1800-1900 AU, acceleration voltage is set to 25 kV.) 9. Mass spectral analysis may be performed using any search algorithm that can perform PMF, for example Mascot (Matrix Sciences, London, UK) or by generating an in silico proteolytic digest peak list and manually comparing.

4. Notes 1. If proteins being electroblotted from the gel onto PVDF membrane have a high molecular weight, (> 80 kDa), use CAPS buffer containing 10% of methanol to improve transfer efficiency. 2. When assembling the sandwich cassette, remove all air bubbles from the gel, paper and PVDF, to ensure proper transfer of proteins onto the membrane. 3. Digestion of proteins using dilute acid usually cleaves the bonds between Asp and Pro residues resulting in peptides with a Proline N-terminal residue. However, cleavage between Asp and Gly residues may also be observed. In order to simplify the mixture of generated peptides, especially for proteins with a molecular weight over 80 kDa, one can block peptides starting with a primary amino acid with 30 µL of sodium borate 0.1 M containing 30 mM o-phthaladehyde (OPA) and 57 mM β-mercaptoethanol at RT for 5 min (5). 4. Fig.1 represents an on-PVDF chemical digestion of anti-OX40 ligand heavy chain using dilute acid solution. Under mild acidic condition, proteins are cleaved at bonds between Asp/Pro residues as shown by proline residue in the first cycle. Treatment of the membrane prior to chemical

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Fig.1. N-terminal sequence analysis of an on-PVDF Asp/Pro cleavage of anti-OX40 ligand heavy chain at 90 °C for 1h.

Fig. 2. On-PVDF clostripain digestion of human IL-20 receptor at 37 °C for 20 min. Underlined are peptides generated, which were identified by Edman degradation analysis.

cleavage is not needed. Sequencing data matches the variable and constant region of the heavy chain. 5. Human IL-12 receptor has part of an STII tag, a poly His tag and an enterokinase cleavage site at the N-terminus. Totally the tag consists of 21 amino acids. In order to confirm the identity of this protein by Edman sequencing, one needs to sequence out the first 30 residues in order to get pass the tag, i.e. it would take 10 h (20 min × 30 cycles). Using rapid clostripain digestion, one can confirm the presence of the tag as well as the identity of the protein in much less time (Fig. 2). 6. For peptide mass fingerprinting, it is necessary to remove Zwittergent solution as completely as possible in order to prevent signal suppression in MALDI-TOF analysis. This can be performed by taking a glass Pasteur pipette and repeatedly rinsing the membrane in MQ water. 7. Some researchers have shown that proteolysis of proteins in the presence of mixed organic-aqueous solvents, such as methanol or acetonitrile in

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Fig. 3. PMF mass spectrum of B7H3 receptor after on-PVDF tryptic digestion at 37 °C for 1 h in Tris buffer (0.1 M, pH 8.0) containing 10% acetonitrile.

digestion buffer, helps denature the proteins, which allows more access of the enzyme to the substrate proteins. As a consequence digestion is more effective (6). 8. Acetonitrile is included in the digestion buffer to help extract generated peptides from the hydrophobic membrane into the solution which can be subsequently analyzed by mass spectrometry analysis without the further clean-up that is necessary if other detergents are used for the extraction step. (Fig. 3) 9. If the digestion buffer contains a high concentration of salt (i.e. Asp-N digestion), it is necessary to desalt using a C18 ziptip prior to loading the sample onto the MALDI plate for MALDI-TOF analysis. (High salt concentration leads to ion suppression during ionization). References 1. Henzel, W. J., Watanabe, C., and Stults, J. T. (2003). Protein identification: the origins of peptide mass fingerprinting. J. Am. Soc. Mass Spectrom. 14, 931–42. 2. Edman, P. (1967). A protein sequenator. Eur. J. Biochem. 1, 80–91. 3. Pham, V. C., Henzel, W. J., and Lill, J. R. (2005). Rapid on-membrane proteolytic cleavage for Edman sequencing and mass spectrometric identification of proteins. Electrophoresis 26, 4243–51. 4. Henzel,W. J., Grimley, C., Bourell, J. H., Billeci, T. M., Wong, S. C., and Stults, J. T. Methods: A Companion to Methods in Enzymology, Vol. 6, Elsevier 1994, pp. 239–247. 5. Kishiyama, A., Zhang, Z., and Henzel, W. J. (2000). Cleavage and identification of proteins: a modified aspartyl-prolyl cleavage. Anal. Chem. 72, 5431–5436. 6. Russell, W.K, Park, Z.Y, and Russell, D.H. (2001). Proteolysis in mixed organic-aqueous solvent systems: Applications for peptide mass mapping using mass spectrometry. Anal. Chem. 73, 2682–2685.

101 Enzymatic Digestion of Membrane-Bound Proteins for Peptide Mapping and Internal Sequence Analysis Joseph Fernandez and Sheenah M. Mische

1. Introduction Enzymatic digestion of membrane-bound proteins is a sensitive procedure for obtaining internal sequence data of proteins that either have a blocked amino terminus or require two or more stretches of sequence data for DNA cloning or confirmation of protein identification. Since the final step of protein purification is usually SDS-PAGE, electroblotting to either PVDF or nitrocellulose is the simplest and most common procedure for recovering protein free of contaminants (SDS, acrylamide, and so forth) with a high yield. The first report for enzymatic digestion of a nitrocellulose-bound protein for internal sequence analysis was by Aebersold et al. in 1987, with a more detailed procedure later reported by Tempst et al. in 1990 (1, 2). Basically, these procedures first treated the nitrocellulose-bound protein with PVP-40 (polyvinyl pyrrolidone, Mr 40,000) to prevent enzyme adsorption to any remaining nonspecific protein binding sites on the membrane, washed extensively to remove excess PVP-40, and the sample was enzymatically digested at 37°C overnight. Attempts with PVDF-bound protein using the above procedures (3, 4) give poor results and generally require >25 µg of protein. PVDF is preferred over nitrocellulose because it can be used for a variety of other structural analysis procedures, such as amino-terminal sequence analysis and amino acid analysis. In addition, peptide recovery from PVDF-bound protein is higher, particularly from higher retention PVDF (ProBlott, Westran, Immobilon Psq). Finally, PVDF-bound protein can be stored dry as opposed to nitrocellulose, which must remain wet during storage and work up to prevent losses during digestion. Enzymatic digestion of both PVDF- and nitrocellulose-bound protein in the presence of 1% hydrogenated Triton X-100 (RTX-100) buffers as listed in From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Table 1 was first performed after treating the protein band with PVP-40 (5). Unfortunately, the RTX-100 buffer also removes PVP-40 from the membrane, which can interfere with subsequent reverse-phase HPLC. Further studies (6, 7) demonstrate that treatment with PVP-40 is unnecessary when RTX-100 is used in the digestion buffer. It appears that RTX-100 acts as both a blocking reagent and a strong elution reagent. PVDF-bound proteins are visualized by staining and subsequently excised from the blot. Protein bands are immersed in hydrogenated Triton X-100 (RTX100), which acts as both a reagent for peptide extraction and a blocking reagent for preventing enzyme adsorption to the membrane during digestion. Remaining cystine bonds are reduced with dithiotreitol (DTT) and carboxyamidomethylated with iodoacetamide. After incubation with the enzyme of choice, the peptides are recovered in the digestion buffer. Further washes of the membrane remove the remaining peptides, which can be analyzed by microbore HPLC. Purified peptides can then be subjected to automated sequence analysis. Additional studies have been reported that have enhanced this procedure. Best et al. (8) have reported that a second aliquot of enzyme several hours later improves the yield of peptides. Reduction and alkylation of cysteine is possible directly in the digestion buffer, allowing identification of cysteine during sequence analysis (9). Finally, octyl- or decylglucopyranoside can be substituted for RTX-100 in order to obtain cleaner mass spectrometric analysis of the digestion mixture (10). 2. Materials The key to success with this procedure is cleanliness. Use of clean buffers, tubes, and staining/destaining solutions, as well as using only hydrogenated Triton X-100 as opposed to the nonhydrogenated form, greatly reduces contaminant peaks obscured during reverse-phase HPLC. A corresponding blank piece of membrane must always be analyzed at the same time as a sample is digested, as a negative control. Contaminants can occur from may sources and particularly protein contaminants are a cocnern. Human Kertain may contaminate samples if gloves are not worn. Proteins from Western blotting can contaminate the PVDF membrane if previously used dishes are used for staining. UV absorbing contaminants can arise from dirty tubes and sub quality detergents. All solutions should be prepared with either HPLC-grade water or double-glassdistilled water that has been filtered through an activated charcoal filter, and passed through a 0.22-µm filter (11). 2.1. Preparation of the Membrane-Bound Sample Protein should be analyzed by SDS-PAGE or 2D-IEF using standard laboratory techniques. Electrophoretic transfer of proteins to the membrane should

10 mg OGP,

100 µL acetonitrile, 400 µL HPLC-grade water, and 500 µL 200 mM Tris stockb 10 mg OGP,

500 µL HPLC-grade water, and 500 µL 200 mM Tris stockb 10 mg OGP, 100 µL acetonitrile, 45 µL 45 mM DTT, 10 µL 100 mM CaCl2,

100 µL 10% RTX-100 stock,

100 µL acetonitrile, 300 µL HPLC-grade water, and 500 µL 200 mM Tris stockb 100 µL 10% RTX-100 stock,

400 µL HPLC-grade water, and 500 µL 200 mM Tris stockb 100 µL 10% RTX-100 stock, 100 µL acetonitrile, 45 µL 45 mM DTT, 10 µL 100 mM CaCl2,

1% Detergent/

10% acetonitrile/

Clostripain

Glu-C

1% Detergent/

10% acetonitrile/

2 mM DTT/

1 mM CaCl2/

100 mM Tris, pH 8.0

1% Detergent/

100 mM Tris, pH 8.0

Trypsin or Lys-C

Recipe (using OGP)c

Digestion buffer

Enzyme

Recipe (using RTX- 100)a

Table 1 Digestion Buffers Recipes for Various Enzymes

(continued)

DTT and CaCl2 are necessary for Clostripain activity

Acetonitrile decreases digestion efficiency of Glu-C

Detergent prevents enzyme adsorption to membrane and increases recovery of peptides

Comments

Enzymatic Digestion of Membrane-Bound Proteins 929

100 mM Tris, pH 8.0

Digestion buffer

Recipe (using OGP)c 345 µL HPLC-grade water, and 500 µL 200 mM Tris stockb

Recipe (using RTX- 100)a 245 µL HPLC-grade water, and 500 µL 200 mM Tris stockb

Comments

b

Hydrogenated Triton X-100 (RTX-100) as described in Tiller et al. (13). 200 mM Tris stock (pH 8.0) is made up as follows: 157.6 mg Tris-HCl and 121.1 mg trizma base to a final volume of 10 mL with HPLCgrade water. c Octyl glucopyranoside can be substituted for RTX-100.

a

Enzyme

Table 1 (continued)

930 Fernandez and Mische

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be performed in a full immersion tank rather than a semidry transfer system to avoid sample loss and obtain efficient transfer (12). PVDF membranes with higher protein binding capacity such as Immobilon Psq (Millipore, Bedford, MA), Problott (Applied Biosystems, Foster City, CA), and Westran (Bio-Rad, Hercules, CA), are preferred owing to greater protein recovery on the blot, although all types of PVDF and nitrocellulose can be used with this procedure. The following stains are compatible with the technique: Ponceau S, Amido black, india ink, and chromatographically pure Coomassie brilliant blue with a dye content >90%. A blank region of the membrane should be excised to serve as a negative control. 2.2. Enzymatic Digestion Buffers Digestion buffer should be made as described in Table 1. Make up 1 mL of buffer at a time and store at −20°C for up to 1 wk. Hydrogenated Triton X-100 (RTX-100) (protein grade, Calbiochem, LaJolla, CA) is purchased as a 10% solution, which should be stored at −20°C. Note: Only hydrogenated Triton X-100 should be used since UV-absorbing contaminants are present in ordinary Triton X-100, making identification of peptides on subsequent HPLC impossible (see Fig. 1). Alternately, octyl glucopyranoside (OGP) (Ultrol-grade, Calbiochem) or decyl glucopyranoside (DGP) (Ultrol-grade, Calbiochem) can be substituted for RTX-100 as described in Table 1. 2.3. Reduction and Carboxyamidomethylation 1. 45 mM DTT: bring 3.5 mg DTT (Ultrol- grade, Calbiochem) up in 500 µL HPLCgrade water. This can be stored at −20°C for up to 3 mo. 2. 100 mM iodoacetamide solution: bring 9.25 mg iodoacetamide (reagent-grade) up in 500 µL HPLC-grade water. This solution must be made fresh just prior to use. Dry DTT and iodoacetamide should be stored at 4°C or −20°C.

2.4. Enzyme Solutions and Inhibitors Enzymes should be stored as small aliquots at −20°C, and made up as 0.1 µg/µL solutions immediately before use. These aliquots can be stored for at least 1 mo at −20°C without significant loss of enzymatic activity. 1. Trypsin (25 µg, sequencing-grade, Boehringer Mannheim, Indianapolis, IN): Solubilize trypsin in 25 µL of 0.01% trifluoroacetic acid (TFA), and let stand ~10 min. Aliquot 5 µL (5 µg) quantities to clean microcentrifuge tubes, dry in a SpeedVac, and store at −20°C. Reconstitute the dry enzyme in 50 µL 0.01% TFA for a 0.1 µg/µL working solution, which is good for 1 d. Trypsin cleaves at arginine and lysine residues. 2. Endoproteinase Lys-C (3.57 mg, Wako Pure Chemicals, Osaka, Japan): Solubilize enzyme in 1000 µL HPLC- grade water, and let stand ~10 min. Make nine 100-µL

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Fig. 1. Peptide maps of trypsin digestion of beta galactosidase bound to PVDF. Panels A–D represent varying amounts of proteins, 40 pmol, 20 pmol, 10 pmol, and 5 pmol respectively. Proteins were analyzed by SDS-PAGE, transferred to PVDF, and stained with Coomassie Brilliant Blue G-250.

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quantities to clean tubes, and store at −20°C. Disperse remaining 100 µL into 20 × 5 µL aliquots (17.85 µg each), and store at −20°C. When needed, take one 5-µL aliquot and add 173 µL of HPLC-grade water to establish a 0.1 µg/µL working solution, which can be used for up to 1 wk if stored at −20°C between uses. When 5-µL aliquots are used up, disperse another 100-µL aliquot. Endoptoteinase Lys-C cleaves only at lysine residues. 3. Endoproteinase Glu-C (50 µg, sequencing- grade, Boehringer Mannheim) : Solubilize in 100 µL of HPLC- grade water, and let stand ~10 min. Aliquot 10-µL (5 µg) quantities to clean tubes, dry in a SpeedVac, and store at −20°C. Reconstitute the enzyme in 50 µL HPLC-grade water for a 0.1 µg/µL solution, which is good for only 1 d. Under these conditions, endoproteinase Glu-C cleaves predominantly at glutamic acid residues, but can sometimes cleave at aspartic acid residues. 4. Clostripain (20 µg, sequencing-grade, Promega, Madison, WI): Solubilize enzyme in 200 µL of manufacturer’s supplied buffer for a concentration of 0.1 µg/µL. Enzyme solution can be stored at −20°C for 1 mo. Clostripain cleaves at arginine residues only. 5. 1% Diisopropyl fluorophosphate (DFP) solution: DFP is a dangerous neurotoxin that must be handled with double gloves in a chemical hood. Please follow all precautions listed with this chemical. Add 10 µL of DFP to 990 µL absolute ethanol in a capped microcentrifuge tube. Store at −20°C.

3. Method 3.1. Preparation of the Membrane-Bound Sample Protein should be electrophoresed in one or two dimensions, followed by electrophoretic transfer to the membrane and visualization of the bands according to the following suggestions. 1. Electroblotting of proteins will be most efficient with a tank transfer system, rather than a semidry system. Concentrate as much protein into a lane as possible; however, if protein resolution is a concern, up to 5 cm2 of membrane can be combined for digestion (see Notes 3 and 5). 2. Stain PVDF membrane with either Ponceau S, Amido black, india ink, or chromatographically pure Coomassie brilliant blue with a dye content of 90%. Destain the blot until the background is clean enough to visualize the stained protein. Complete destaining of the blot is unnecessary, but at least three washes (~5 min) with distilled water should be done to remove excess acetic acid, which is used during destaining (see Note 5). 3. Excise protein band(s), and place into a clean 1.5-mL microcentrifuge tube. In addition, excise a blank region of the membrane approximately the same size as the protein blot to serve as a negative control (see Note 4). 4. Air dry PVDF-bound protein dry at room temperature and store at −20°C or 4°C.

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3.2. Digestion of the Membrane-Bound Protein NOTE: Gloves should be worn during all steps to avoid contamination of sample with skin keratin. 1. Place ~100 µL of HPLC-grade water onto a clean glass plate, and submerge the membrane-bound protein into the water. Transfer the wet membrane to a dry region of the plate and with a clean razor blade, cut the membrane first lengthwise into 1-mm wide strips, and then perpendicular so that the membrane pieces are 1 × 1 mm. Treat the negative control under the same conditions as the sample. Keeping the membrane wet will simplify manipulation of the sample as well as minimize static charge, which could cause PVDF to “jump” off the plate. The 1 × 1 mm pieces of membrane will settle to the bottom of the tube and require less digestion buffer to immerse the membrane completely (see Note 7). 2. Slide the cut membrane onto the forceps with the razor blade, and return it to a clean 1.5-mL microcentrifuge tube. Use the cleanest tubes possible to minimize contamination during peptide mapping. Surprisingly, many UV-absorbing contaminants can be found in microcentrifuge tubes, and this appears to vary with supplier and lot number. Tubes can be cleaned by rinsing with 1 mL of HPLC-grade methanol followed by 2 rinses of 1 mL HPLC-grade water prior to adding the protein band. 3. Add 50 µL of the appropriate digestion buffer (Table 1), and vortex thoroughly for 10–20 s. Optionally, add 50 µL digestion buffer to an empty microcentrifuge tube to serve as a digestion blank for HPLC analysis. The amount of digestion buffer can be increased or decreased depending on the amount of membrane; however, the best results will be obtained with a minimum amount of digestion buffer. PVDF membrane will float in the solution at first, but will submerge after a short while, depending on the type and amount of PVDF. 4. Add 5 µL of 45 mM DTT, vortex thoroughly for 10–20 s, seal the tube cap with parafilm, and incubate at 55°C for 30 min. DTT will reduce any remaining cystine bonds. DTT should be of the highest grade to reduce uv absorbing contaminants that might interfere with subsequent peptide mapping. 5. Allow the sample to cool to room temperature. Add 5 µL of 100 mM iodoacetamide, vortex thoroughly for 10–20 s, and incubate at room temperature for 30 min in the dark. Iodoacetamide alkylates cysteine residues to generate carboxyamidomethyl cysteine, allowing identification of cysteine during sequence analysis. Allowing the sample to cool prior to adding iodoacetamide and incubating at room temperature are necessary to avoid side reaction to other amino acids. 6. Add enough of the required enzyme solution to obtain an estimated enzyme to substrate ratio of 1:10 (w/w) and vortex thoroughly for 10–20 s. Incubate the sample (including digestion buffer blank) at 37°C for 22–24 h. The amount of protein (substrate) can be estimated by comparison of staining intensity to that of known quantities of stained standard proteins. The 1:10 ratio is a general guideline. Ratios of 1:2 through 1:50 can be used without loss of enzyme efficiency or peptide recovery. An enzyme should be selected that would likely produce peptides of > 10 amino acids long. Amino acid analysis of the protein would be informative for estimating the number of cleavage sites. A second aliquot of enzyme can be added after 4–6 h (see Note 6).

Enzymatic Digestion of Membrane-Bound Proteins

935

3.3. Extraction of the Peptides 1. After digestion, vortex the sample for 5–10 s, sonicate for 5 min by holding in a sonicating water bath, spin in a centrifuge (~1800g) for 2 min, and transfer the supernatant to a separate vial that will be used directly for HPLC analysis. 2. Add a fresh 50 µL of digestion buffer to the sample, repeat step 11, and pool the supernatant with the original buffer supernatant. 3. Add 100 µL 0.1% TFA to the sample, and repeat step 1. The total volume for injection onto the HPLC is 200 µL. Most of the peptides (~80%) are recovered in the original digestion buffer; however, these additional washes will ensure maximum recovery of peptides from the membrane. 4. Terminate the enzymatic reaction by either analyzing immediately by HPLC or adding 2 µL of the DFP solution.

CAUTION: DFP is a dangerous neurotoxin and must be handled with double gloves under a chemical fume hood. Please follow all precautions listed for this chemical. 3.4. Analysis of Samples by Reverse-Phase HPLC and Storage of Peptide Fractions 1. Prior to reverse-phase HPLC, inspect the pooled supernatants for small pieces of membrane or particles that could clog the HPLC tubing. If membrane or particles are observed, either remove the membrane with a clean probe (such as thin tweezers, a thin wire, thin pipet tip, and so forth), or spin in a centrifuge for 2 min and transfer the sample to a clean vial. A precolumn filter will help increase the life of HPLC columns, which frequently have problems with clogged frits. 2. Sample is ready to be fractionated by HPLC (see Chapter 102). Fractions can be collected in capless 1.5-mL plastic tubes, capped, and stored at −20°C until sequenced (see Note 9). A typical fractionation is shown in Fig. 1.

4. Notes 1. This procedure is generally applicable to proteins that need to have their primary structure determined and offers a simple method for obtaining internal sequence data in addition to amino terminal sequence analysis data. The procedure is highly reproducible and is suitable to peptide mapping by reverse-phase HPLC. Proteins 12–300 kDa have been successfully digested with this procedure with the average size around 100 kDa. Types of proteins analyzed by this technique include DNA binding, cystolic, peripheral, and integral-membrane proteins, including glycosylated and phosphorylated species. The limits of the procedure appear to be dependent on the sensitivity of both the HPLC used for peptide isolation and the protein sequencer. 2. There are several clear advantages of this procedure over existing methods. First, it is applicable to PVDF (especially high-retention PVDF

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membranes), which is the preferred membrane owing to higher recovery of peptides after digestion, as well as being applicable to other structural analysis. The earlier procedures (1,2) have not been successful with PVDF. Second, because of the RTX-100 buffer, recovery of peptides from nitrocellulose is higher than earlier nitrocellulose procedures (5). Third, the procedure is a onestep procedure and does not require pretreatment of the protein band with PVP-40. Fourth, since the procedure does not require all the washes that the PVP-40 procedures do, there is less chance of protein washout. Fifth, the time required is considerably less than with the other procedures. Overall, the protocol described here is the simplest and quickest method to obtain quantitative recovery of peptides. 3. The largest source of sample loss is generally not the digestion itself, but rather electroblotting of the sample. Protein electroblotting should be performed with the following considerations. Use PVDF (preferably a higher binding type, i.e., ProBlott, Immobilon Psq) rather than nitrocellulose, since peptide recovery after digestion is usually higher with PVDF (5,8). If nitrocellulose must be used, e.g., protein is already bound to nitrocellulse before digestion is required, never allow the membrane to dry out since this will decrease yields. Always electroblot protein using a transfer tank system, since yields from semidry systems are not as high (12). Using stains such as Ponceau S, Amido black, or chromatographically pure Coomassie brilliant blue, with a dye content >90% will increase detection of peptide fragments during reverse-phase HPLC. Note: Most commercial sources of Coomassie brilliant blue are extremely dirty and should be avoided. Only chromatographically pure Coomassie brilliant blue with a dye content of 90% appears suitable for this procedure. 4. The greatest source of failure in obtaining internal sequence data is not enough protein on the blot, which results in the failure to detect peptide during HPLC analysis. An indication of insufficient protein is that either the intensity of the stain is weak, i.e., cannot be seen with Amido black even though observable with india ink (about 10-fold more sensitive), or possibly detectable by radioactivity or immunostaining, but not by protein stain. Amino acid analysis, amino-terminal sequence analysis, or at the very least, comparison with stained standard proteins on the blot should be performed to help determine if enough material is present. When 3000 Dalton. Smaller peptides are best separated in the tricine gel system of Schagger and von Jagow (7) (see also Chapter 23). The methods described in Chapter 97, Subheading 3.3 can be used to generate peptides. Cleavage technique varies slightly depending on the form of the purified proteins to be compared. 3.2.1. In-Gel Cleavage/Separation 3.2.1.1. LYOPHILIZED/SOLUBLE PROTEIN 1. Boil the purified protein (~1 mg/mL) for 5 min in enzyme buffer for Cleveland “ingel” digestion. 2. Load 10–30 µL (10–30 µg) of each protein to be compared into three separate wells of an SDS-PAGE gel (see Note 1). 3. Overlay samples of each protein with 0.005, 0.05, and 0.5 µg enzyme in 10–20 µL of Cleveland “in-gel” digestion buffer to separate wells. V8 protease (endoproteinase Glu-C) works very well in this system (see Note 2). SDS hinders the activity of trypsin, α-chymotrypsin, and themolysin, so cleavage with these enzymes may be very slow. In-gel cleavage with chemical reagents is not generally recommended, since they can be inefficient in neutral, oxygenated environments. Gently fill wells and top chamber with running buffer. 4. Subject the samples to electrophoresis until the dye reaches the bottom of the stacking gel. Turn off the power, and incubate for 2 h at 37°C to allow the enzyme to digest the protein partially. Following incubation, continue electrophoresis until the dye reaches the bottom of the gel. Fix, stain, and destain, or electroblot onto NCP for immunoanalysis (see Note 3). 3.2.1.2. PROTEIN IN-GEL-SLICE

Peptides of proteins purified by SDS-PAGE usually retain adequate SDS to migrate into a second gel without further treatment. 1. Run SDS-PAGE gels, fix, stain with Coomassie brilliant blue, and destain. 2. Excise protein bands to be compared with a razor blade. 3. Soak excised gel slices containing proteins to be compared in ethanol–1 M TrisHCl, pH 6.8, for 30 min to shrink the gel, making loading on the second gel easier. Place the gel slices into the wells of a second gel. 4. Continue from step 3, Subheading 3.2.1.1.

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3.2.1.3. PROTEIN ON NCP STRIP 1. Run SDS-PAGE gels, and electroblot to NCP. 2. Stain blot with Ponceau S, NBB, or block with 0.05% Tween-20 in PBS for 30 min, and then add three drops India ink (see Note 4). 3. Excise protein bands to be compared with a razor blade. 4. Push NCP strips to bottom of the wells of a second SDS-PAGE gel. 5. Continue from step 3, Subheading 3.2.1.1.

Once the separation conditions, protein concentrations, and enzyme concentrations have been established, a single digestion lane for each sample can be used for comparative purposes. 3.2.2. Protein Cleavage Followed by SDS-PAGE It is often preferable to cleave the proteins to be compared prior to loading onto an SDS-PAGE gel for separation. This generally gives more complete, reproducible cleavage and allows for the use of chemical cleavage reagents not suitable for in-gel cleavages. 3.2.2.1. LYOPHILIZED/SOLUBLE PROTEIN 1. Rehydrate the lyophilized proteins in the appropriate buffer at 1 mg/mL (less concentrated samples can be used successfully). Soluble proteins may need to be dialyzed against the proper buffer. 2. Place 10–30 µL (10–30 µg) of each protein to be compared in 1.5-mL microfuge tubes. 3. Add up to 25 µL of the appropriate chemical cleavage reagent (1 mg/mL) to suspended protein. For enzymes, use only as much enzyme as needed to achieve complete digestion (1:50 enzyme to sample maximum) (see Note 5). 4. Incubate with shaking at 37°C for 4 h (for enzymes) or at room temperature for 24–48 h in dark under nitrogen (for chemical reagents). 5. Add equal volume of 2X SDS-PAGE solubilization buffer and boil for 5 min (see Note 6). 6. Load entire sample on SDS-PAGE gel and proceed with electrophoresis. 3.2.2.2. PROTEIN IN-GEL SLICE 1. Run SDS-PAGE gels, fix, stain with Coomassie brilliant blue, and destain. 2. Excise protein bands to be compared with a razor blade. 3. Dry gel slices containing proteins using a Speed-Vac concentrator or other drying system, such as heat lamps, warm air, and so forth. 4. Put the dry gel slice containing the protein in a 1.5-mL microfuge tube. 5. Add up to 10 µL of the appropriate chemical cleavage reagent (1 mg/mL) directly to the gel slice protein, and then and 90 µL of appropriate buffer. For enzymes, use only as much enzyme as needed to achieve complete digestion (1:50 enzyme to sample maximum) (see Note 5). 6. Incubate with shaking at 37°C for 4 h (for enzymes) or at room temperature for 24–48 h in dark under nitrogen (for chemical reagents).

Peptide Mapping by SDS-PAGE

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Fig. 1. Example of peptides separated in an SDS-PAGE gel. Whose cells (WC), a sarkosyl insoluble pellet, and a periplasmic extract of Neisseria gonorrhoeae were separated in “preparative” 15% SDS-PAGE gels and blotted to NCP as described in Chapter 103, Subheading 3.1.1. The 37,000-Dalton major outer membrane protein (POR) and two 44,000-Dalton (44-kDa) proteins, one isolated from a sarkosyl insoluble (membrane) extract (44-kDa Mem.), and the other isolated from a periplasmic extract (44-kDa Peri.), were located on the NCP by Ponceau S staining, excised, and cleaved with BNPS-skatole as described in Chapter 96, Subheading 3.3.1. Approximately 30 µg of peptides of each protein were solubilized and separated in an SDS-PAGE gel along with whole-cells (WC), BioRad low-mol-wt markers, and Pharmacia peptide mol-wt markers (mw) (expressed in thousands of Dalton [k]). The gel was stained with Coomassie brilliant blue (CBB) to visualize peptides.

7. Aspirate peptide-containing supernatant. 8. Completely dry-down the supernatant in a Speed-Vac, and wash the sample several times by adding 50 µL of H2O, vortexing, and redrying in a Speed-Vac. Alternate drying systems will work. 9. Add 10–20 µL SDS-PAGE solubilizing solution to samples and boil for 5 min (see Note 6). 10. Load samples onto SDS-PAGE gel and proceed with electrophoresis. 3.2.2.3. PROTEIN ON NCP STRIP 1. Run SDS-PAGE gels, and electroblot to NCP. 2. Stain blot with Ponceau S, NBB, or block with 0.05% Tween-20 in PBS for 30 min, and then add three drops India ink (see Note 3). 3. Excise protein bands to be compared with a razor blade. 4. Put the NCP strip containing the protein in a 1.5-mL microfuge tube. 5. Continue from step 5, Subheading 3.2.2.1.

Figure 1 is presented to demonstrate the separation of peptides generated by cleavage with BNPS-skatole in an SDS-PAGE gel. These peptide maps indicate that the porin protein (POR) was structurally unrelated to the 44-kDa proteins, whereas the 44-kDa proteins from a sarkosyl insoluble (membrane) extract (44-kDa Mem.) and a periplasmic extract (44-kDa Peri.) appeared to have similar primary structures.

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4. Notes 1. Between 5 × 104 cpm and 105 cpm should be loaded in each lane if radioiodinated samples are to be used. Autoradiography should be performed on unfixed gels. Fixation and staining may wash out small peptides. Place gel in a plastic bag and overlay with XAR-5 film, place in a cassette with a Lightening Plus intensifying screen, and expose for 16 h at −70°C. 2. Control lanes containing only enzyme must be run to distinguish enzyme bands from sample bands. 3. PVDF nylon membrane is preferable to NCP when blotting small peptides. 4. Do not compare proteins stained with Ponceau S with those stained with NBB or India ink. Use the same staining procedure for all proteins to be compared. 5. To ensure complete digestion, it is advisable to incubate the samples for increasing periods of time or to digest with increasing amounts of enzyme. Once optimal conditions are established, a single incubation time and enzyme concentration can be used. 6. It is often advisable to solubilize protein in SDS prior to cleavage, since some peptides do not bind SDS well. Acknowledgments The author thanks Pam Gannon for her assistance and the Public Health Service, NIH, NIAID (grant RO1 A121236), and UM Research Program for their continued support. References 1. Cleveland, D. W., Fischer, S. G., Kirschner, M. W., and Laemmli, U. K. (1977) Peptide mapping by limited proteolysis in sodium dodecyl sulfate and analysis by gel electrophoresis. J. Biol. Chem. 252, 1102–1106. 2. Judd, R. C. (1986) Evidence for N-terminal exposure of the PIA subclass of protein I of Neisseria gonorrhoeae. Infect. Immunol. 54, 408–414. 3. Judd, R. C. (1987) Radioiodination and 125I-labeled peptide mapping on nitrocellulose membranes. Analyt. Biochem. 160, 306–315. 4. Moos, M., Jr. and Nguyen, N. Y. (1988) Reproducible high-yield sequencing of proteins electrophoretically separated and transferred to an inert support. J. Biol. Chem. 263, 6005–6008. 5. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–695. 6. Caldwell, H. D. and Judd, R. C. (1982) Structural analysis of chlamydial proteins. Infect. Immunol. 38, 960–968. 7. Schagger, H. and von Jagow, G. (1987) Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Analyt. Biochem. 166, 368–397.

105 Peptide Mapping for Protein Characterization Peter Højrup

1. Introduction Analysis of intact proteins is hampered by the fact that even when using high resolution techniques, like electrospray mass spectrometry; you will only be able to tell whether the protein has the expected mass. If you find a deviation from the expected you will in most cases be left wondering where (and perhaps what) the difference is. Recent methods, such as top-down sequencing (1,2), can give you invaluable information on terminal trimming or even simple posttranslational modifications, but generally works best for “simple” changes to the protein. Observations on intact proteins are compounded when using low-resolution techniques like gel filtration or SDS gel electrophoresis. Although 2-D gel electrophoresis is able to show a single charge difference, in the absence of additional information you are still left wondering where and what the differences are. Performing Edman degradation on an intact protein will in most cases yield enough information for you to identify a protein and perhaps enable the construction of oligonucleotide probes, but many proteins are N-terminally blocked, and the need to purify enough protein is often prohibitive. The solution is to perform peptide mapping. This approach is essential for the total characterization of a protein, and the approach also enables quick identification of a protein (if present in a protein database) and to locate differences between proteins. The steps involved in peptide mapping can be listed as: 1. Purify the protein. Depending on the objective of the peptide mapping, it may be sufficient with a single spot from a 2-D gel, or you may have to purify several microgram of material using traditional protein chemical methods.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2. Cleave the protein using a suitable proteolytic enzyme or by chemical means. A software tool like GPMAW (see Note 1) greatly facilitates the selection of cleavage method and analysis of the resulting peptides. 3. Fractionate the resulting peptides, generally by reversed phase HPLC, but other methods may be applicable depending on the final method of analysis. 4. Analyze the purified peptide using mass spectrometry, Edman degradation, or amino acid analysis.

1.1. Peptide Mapping Peptide mapping is usually carried out for two quite different reasons: 1.1.1. Protein Identification Currently the most sensitive identification of an unknown protein is mass spectrometric peptide mapping (peptide mass fingerprinting, PMF). Using the rapidly expanding protein databases based on nucleotide sequencing, it is possible to identify a protein based on the molecular masses of its constituent (tryptic) peptides if they are determined with sufficient precision. The beauty of the method is that it is not necessary to determine the mass of all peptides; usually 6-7 peptides are sufficient for accurate protein identification. As it is not necessary to separate the peptides before analysis, the method is very sensitive; with routine determination of proteins in the low nanogram range (e.g. a single silver stained spot from an SDS gel). Two alternative approaches are liquid chromatography coupled to electrospray mass spectrometry (LC-MS/MS) and Edman degradation. Both methods will generate a peptide sequence thus making it possible to do a homology search in case the protein is not present in the database. However, while the sensitivity of LC-MS/MS approaches that of MALDI-MS, the sensitivity of Edman degradation is 2-3 orders of magnitudes lower and further needs a purification step prior to analysis. The topic of LC-MS/MS is beyond the scope of this chapter and will not be treated further. For an introduction please see ref. (3). 1.1.2. Obtaining the Maximum Sequence Coverage Unlike protein identification where even a limited number of peptides are able to identify the protein, you will try to cover the whole sequence when comparing proteins for identity/similarity or when performing complete chemical characterization. 1.1.2.1. COMPARISON OF PROTEINS

Comparison of two proteins is done by performing an identical enzymatic digest on each protein followed by identification of each fragment. In some cases it may be sufficient to perform mass spectrometric mapping of the peptide mixture, but in most cases you will not be able to identify sufficient fragments to

Peptide Mapping for Protein Characterization

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cover the whole sequence. You will thus have to isolate each fragment by HPLC and perform peptide identification. Due to the high resolution of reversed phase HPLC it will in many cases be sufficient to compare the two chromatograms carefully in order to identify differences. When comparing proteins you should note that choosing an enzyme, which generates long peptide fragments, tends to emphasis similarities while short peptide fragments tends to emphasize differences (see Note 2). 1.1.2.2. Determine the Exact Chemical Structure of a Protein

Even though the number of known protein sequences increases exponentially over time through nucleotide sequence analysis, and you are thus likely to find your protein to be already known, the result of these analyses is the ‘naked’ amino acid sequence of the protein. However, the vast majority of proteins are posttranslationally modified after synthesis, resulting in a structure that deviates from the nucleotide-derived sequence. The preferred strategy to solve the actual protein structure is to perform peptide mapping followed by an appropriate analytical techniques like mass spectrometry, Edman degradation and/or amino acid analysis. Here you always aim for total sequence coverage, and in order to obtain this you will in most cases have to perform at least two separate enzymatic digests. The amount of material needed for maximum sequence coverage analysis is usually much higher than for PMF analysis, usually in the low to middle microgram range. In special cases where you search only for specific modifications using specific extraction of modified peptides (e.g. purification of phosphorylated peptides using titanium oxide), the sensitivity approach is similar to that of PMF (4). 2. Materials 2.1. Peptide Mapping for Protein Identification 1. Trypsin (modified trypsin, Promega,). This is most easily handled in aliquots of 1 µg trypsin in 10 µL 0.01 N HCl per tube at –20 °C. Dilute with 70 µL 50 mM ammonium bicarbonate before use (final concentration 12.5 ng /µL). Remember always to use UHQ (ultrahigh quality) water. 2. 50 mM ammonium bicarbonate. You can store aliquots of 100 mM ammonium bicarbonate at –20 °C, it is not necessary to adjust the pH. Dilute 1 : 1 for use. 3. Neat acetonitrile (HPLC quality). 4. 10 mM dithiothreitol (DTT). A 1 M DTT solution is stable for month at -20 °C. Store as 25 µL aliquots and dilute 1:100 with 100 mM ammonium bicarbonate for use. 5. 55 mM iodoacetamide (Sigma,) in 100 mM ammonium bicarbonate. Has to be prepared freshly (iodoacetamide: 10.2 µg / µL). 6. 0.5 mL polypropylene tubes. 7. Zip-tips (Waters) or homemade micro-purification tips based on Poros 50 R2 reversed phase material (Applied BioSystems, #1-1159-05) and Eppendorf GELoader tips (Eppendorf, #0030 001.222) (see Note 3).

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8. Vacuum centrifuge. 9. Concentrated solution of alpha-cyano-4-hydroxycinnamic acid (Aldrich) in 70% acetonitrile, 0.1% TFA in water. 10. MALDI mass spectrometer.

2.2. Alkylation of Protein 1. 8 M urea, 100 mM ammonium bicarbonate (or Tris-HCl), pH 8. 2. Reduction agent: typically dithiothreitol (DTT) is used for reduction, but mercaptoethanol or tris(carboxyethyl)phosphine (TCEP – Sigma) may be used instead. Mercaptoethanol is volatile while TCEP is also active at low pH. 3. Alkylating reagent: iodoacetate, iodoacetamide, 4-vinylpyridine and N-isopropyliodoacetamide are all commonly used reagents. 4-vinylpyridine has the disadvantage of being prone to polymerization and thus has a limited shelf life.

2.3. Peptide Mapping with Purification of Peptides 1) Suitable enzyme or chemistry (see Table 1). 2) Buffer for digestion (see Note 4) 3) HPLC system capable of binary gradient formation (see Note 5) equipped with an in-line UV detector and a strip-chart recorder and/or computer integrator. 4) Acetonitrile (HPLC grade, UV cutoff < 210 nm). Neat trifluoroacetic acid (TFA) (analytical grade is acceptable, sequencing grade is optimal) (see Note 6). 5) Automatic or manual fraction collection into polypropylene tubes. 6) Vacuum centrifuge

As many of the solvents and reagents are used in very small quantities (e.g. ≤ 1 mL) it can be advantageous to prepare stock solutions that are stored at –20 °C, even of those that are not specifically mentioned in the text. Most proteolytic enzymes are supplied freeze dried. When first opened you may dissolve the enzyme in pure water, make 0.5-2 µg aliquots in small polypropylene tubes, and lyophilize the aliquots. Endo Glu-C protease may be stored frozen in pure water. 3. Methods 3.1. Selection of Proteolytic Enzyme The selection of an enzyme for digestion is one of the most important considerations when performing peptide mapping, and should be carefully considered in relation to the task you want to perform. For peptide mass fingerprinting (PMF) the most common choice is trypsin. This is one of the most active enzymes, it is well characterized for PMF analysis (see Notes 8 and 15), and cleaves with high specificity (C-terminal to Lys and Arg). Most tryptic peptides will have molecular masses in the range 800–2500 Da, which is well suited for mass spectrometry, Furthermore, you are certain that

23,463

25,600 25,700

27,000

3.4.21.4

Chymotryp 3.4.21.1 sin A B

3.4.21.9

3.4.99.30

Trypsin

Endo Glu-C

Endo Lys-C

30,000

23,293

3.4.21.4

Trypsin

P00766 P00767

P00761

P00760

Serine Lysobacter P15636 enzymogenes

Serine StaphyP04188 lococcus aureus V8

Serine Bovine

Serine Porcine modified

Serine Bovine

Size Da. Type

EC no.

Name

Accession Organism number

Table 1 Proteolytic Enzymes Suitable for Peptide Mapping.

4–9

7–9

5.5 – 10.5

5.5 – 10.5

Stabilized by small amounts of Ca++

Stabilized by small amounts of Ca++

pH Requirerange ments

8.5–8.8 7–9

7.8

8

8

8

pH optimum High

High

Arg, Lys 2 M urea >0.1% SDS 50% acetonitrile

Arg, Lys 2 M urea >0.1% SDS 50% acetonitrile

0.1% SDS Lys 1 M urea

(continued)

Medium R,S,PS to high

19

18

R,A,P, W,S, PS

Glu (Asp) 2 M urea -Glu 0.2% SDS 10% acetonitrile

High

17

16

16

Note

0.1% SDS Phe, Trp, Tyr Medium R,A, to high W,S, 1 M urea PS 10–30% acetonitrile

A,S R, P,W, S, PS

R,W,S

Specifi- Manucity facturer

Tolerates detergents Cleavage

Peptide Mapping for Protein Characterization 973

27.000

30,000

50,000

3.4.24.33

Endo Asp-N

3.4.22.8

3.4.21.40

Endo Arg-C

46.000

Clostripain

3.4.21.50

Lysyl endopeptidase

22,000

23.000

3.4.24

Endo Lys-N

Grifola frondesa

-SH

-SH

Metallo

Clostrid- P09870 ium histolyticum

Jack Bean

Pseudomonas fragi

Serine Mouse submaxillary

P15636

Accession Organism number

Serine Achromabacter lyticus

Metallo

Size Da. Type

Endo Asn-C

EC no.

Name

Table 1 (continued)

7.6

5.0

8

8

8

10

pH optimum

4.5– 6.5

6.0– 8.5

7.5– 8.5

6– 10.5

1–10 mM dithiothreitol

1–10 mM dithiothreitol

Avoid EDTA

High

4 M urea Arg (Lys) 10% acetonitrile

Asn

19

19

19

Note

T

19

19

T,R,A,S, 19 PS

T,S,PS

A

Medium R,W,S

High

N-terminal to High 0.1% Asp SDS 1 M urea 10% acetonitrile

2 M urea Arg 10% acetonitrile

High

SE

Specifi- Manucity facturer

N-terminal to High Lys

Tolerates detergents Cleavage

Lys Inhibited >0.1% by ammo- SDS 4 M nium salts urea 40% acetonitrile

Zn Avoid EDTA

++

pH Requirerange ments

974 Højrup

3.4.24.4

3.4.21.36

Thermolysin

Elastase

25.900

37,500

34,500

8.5

8

P00800 Bacillus thermoproteolyticus P00772

2

Pig gastric P00791 mucosa

Serine Pig pancreas

Metallo

Acid

7–9

7–9

1–5

Ca++ Avoid EDTA Low

N-terminal to Low hydrophobic Not if followed by Pro

0.1% SDS Uncharged 2 M urea (small) aliphatic

8 M urea

N-terminal to Low hydrophobic Preferentially if preceded by Phe, Met, Leu, Trp

R,A, W,S

A,S

R,A, W,S, PS

15

15

15

Chemical cleavage procedures are not included in the table as experience shows that they tend to be rather unspecific for general peptide mapping, however, for those special hard to cleave proteins see relevant chapters in this book. Accession number: Numbers refer to the Swiss-Prot/Uniprot database (http://www.ebi.ac.uk/). Cleavage: All enzymes cleave C-terminal to the indicated residue except for except where noted. Minor, but still significant, secondary cleavages are listed in parenthesis. Cleavage does not take place if the cleavage position is followed by proline or a residue indicated by “-” (e.g. a -Lys-Pro- sequence is not cleaved by trypsin). See also notes for individual enzymes. Specificity: Please see Note 15 and notes for the individual enzymes. Detergents: Even very small amounts of SDS is detrimental to mass spectrometric analysis, while reversed phase HPLC is capable of handling small amounts of SDS (see Note 4). When using urea you may dissolve the protein in a small amount of 8 M urea you can dilute to the required strength with buffer containing the relevant enzyme. Manufacturer: R) Roche Biomolecular, S) Sigma Inc., P) Promega, T) Takara, W) Worthington, A) Wako, C) Calbiochem, PS) Princeton Separations

3.4.21.7

Pepsin

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all peptides will contain a basic residue (except the C-terminal peptide), which means they will ionize better when analyzed by mass spectrometry in positive polarity mode. Endo-Lys and in particular lysyl endopeptidase (higher activity and specificity) are also well suited for PMF, but the peptides generated will, on average, be much larger, which may impede their identification. However, in many cases you will increase the sequence coverage, particularly in basic regions of the protein. Other potential enzymes are Endo Asp-N and Endo Glu-C, but the results are usually much more unpredictable. If the protein sequence of your target protein is known, you should look at the distribution of residues when aiming for the largest sequence coverage before selecting an enzyme. Particularly the charged residues are important, as most of the specific enzymes will target these residues (Table 1). A few computer programs for this type of analysis are available (see Note 1). Important aspects for enzyme selection are: ●

●

●

●

●

●

Short peptides. Very short peptides are difficult to handle, both by MALDI mass spectrometry (< 7 residues) and HPLC (< 5 residues if they are hydrophilic). Long peptides. The mass resolution and sensitivity decreases on the mass spectrometer as you get above 30–35 residues. Likewise large, hydrophobic peptides can be difficult to purify by reversed phase HPLC (> 50 residues). If your prime objective is to obtain amino acid sequence by Edman degradation, long peptides are generally most informative. Hydrophobic peptides tend to stick to the wall of polypropylene tubes, particularly if the sample is dried completely (see Note 7). If you want to do protein comparison, you should note that small peptides tend to emphasize differences (e.g. making it easier to identify small differences between proteins) while large peptides will emphasize similarity (e.g. make it easier to identify similar regions). Enzyme specificity, activity and buffer compatibility (see table 1) should be chosen relative to the intended analysis method (mass spectrometry, HPLC separation, Edman degradation, amino acid analysis etc.). For most proteins you will need at least two different enzymes (digests) in order to get a good coverage (> 90%).

3.2. Peptide Mass Mapping 1. Cut out the stained gel band or gel spot using a sharp scalpel or a properly sized puncher (2–4 mm diameter). Cut the spot into smaller pieces (1 mm square) and place them in a small polypropylene tube. Try to cut as close as possible to the actual spot. A small concentrated spot yields better results than a large diffuse one. If you have a one-dimensional gel band, you may not need the whole band, but can reserve part of it for additional experiments. 2. Wash the gel pieces carefully in 50 µL, 50 mM ammonium bicarbonate. Discard the supernatant.

Peptide Mapping for Protein Characterization

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3. Add 50 µL neat acetonitrile. Let stand for 5 min until the gel pieces has fully shrunken and become white. Withdraw the acetonitrile and discard. Dry the gel pieces in a vacuum centrifuge for 20 min. 4. Add 50 µL 10 mM dithiothreitol and incubate for 30 min at 56 °C. 5. Cool to room temperature and remove excess solvent. 6. Add 55 mM iodoacetamide in 100 mM ammonium bicarbonate so the gel pieces are just covered. 7. Let stand for 30 min in the dark before discarding the supernatant. 8. Wash the gel pieces in 20 µL 50% acetonitrile in water. 9. Add 30 µL neat acetonitrile. Mix until the gel pieces are fully shrunken and white. Remove the acetonitrile. 10. Add 20 µL 100 mM ammonium bicarbonate and mix for 5 min. 11. Add additional 30 µL neat acetonitrile and mix until the gel pieces are shrunken and white. 12. Remove the supernatant and dry the gel pieces in a vacuum centrifuge. 13. Add 12.5 ng/µL modified trypsin until the gel pieces are just covered with solvent. If you are using a low sensitivity stain like coomassie blue, the trypsin concentration should be doubled (see Note 8). Let stand for 5 min before removing excess trypsin containing solvent. 14. Add 50 mM ammonium bicarbonate until the gel pieces are just covered. 15. Let the digestion proceed overnight at 37 °C. 16. Extract the excess solvent, which will contain the peptides. Purify the peptides on a Zip tip (Waters) or for higher sensitivity use a home-packed GelLoader tip (3) using the recommended procedures (see Note 3). 17. Run the peptide map on a MALDI mass spectrometer in alpha cyano hydroxy cinnamic acid. 18. Analysis of the resulting mass spectra can easily be carried out using World Wide Web based tools (see Note 9), but be careful to check the results (see Note 8).

3.3. Reduction and Carboxymethylation of Proteins in Solution 1. Dried sample (20–100 µg) is dissolved in 20 µL 8 M urea, 0.4 M ammonium bicarbonate buffer (see Note 10). 2. Add 5 µl 45 mM dithiothreitol. 3. Stand at 50 °C for 15 min. Cool to room temperature. 4. Add 5 µL 100 mM iodoacetamide. 5. Stand at room temperature for 15 min in the dark.

Purify the protein by reversed phase HPLC (a short Poros R1 column, PerSeptive Biosystems, is well suited), gel filtration or reversed phase solid phase extraction. For digestion with trypsin, endo Glu-C or lysyl endoproteinase (Table 1) you can proceed as follows without prior purification: 1. Add 140 µL water to dilute the urea to less than 2 M. 2. Add 2–4 % w/w of the selected enzyme. For prolonged digestion you can add the enzyme in two equal portions with an interval of 4–6 h.

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3. Stand for 16–24 h. at 37 °C. 4. Freeze the sample or inject directly onto reversed phase HPLC column. When programming the separation gradient you should start with 2–5 min of isocratic elution in order to let the urea elute from the column before starting the separation of the peptides.

3.4. Digesting in Solution If your protein contains disulfide bridges you should reduce and alkylate the protein prior to digestion, see above. 1. Dissolve the protein in an appropriate buffer (50 mM ammonium bicarbonate, see Note 4) to a concentration of 1–5 µg/µL. If you have less than 20 µg, you should dissolve in 20 µL. 2. Add the enzyme of your choice (Table 1) to a final concentration of 2–4% w/w. Hard to digest proteins and dilute solutions will need a higher enzyme to substrate ratio. For limiting the cleavage to a single or a few positions you can try to lower the concentration to 1% or lower. 3. Most proteins will be digested in 6–8 hours at 37 °C, but conditions may have to be varied to suit each individual protein. For hard to digest proteins you can use extended digestion times (e.g. 24–48 hours or longer) (see Note 4). For long digestion times it is advantageous to add the enzyme in two equal portions separated by 4–8 hours. 4. Digestions may be stopped by lowering the pH with the addition a small amount of 2% TFA (raising the pH to above 5 in the case of pepsin), by injecting the sample onto the HPLC system or by adding an appropriate enzyme inhibitor. 5. Separate the peptide map by reversed phase HPLC. Prior to separation you may want to extract ½ µL of your sample for MALDI analysis in order to check the extent of proteolysis.

A control reaction run under identical conditions as the main digest should be run in parallel to show which peptides comes from the sample and which are autodigest products or other contaminants. A control digest of a known protein (e.g. beta-lactoglobulin, Fig. 1) can also be run to check the activity of the protease. A special case of peptide mapping is when you want to determine the disulfide bridge structure of a protein. 1. Verify the complete protein structure using standard peptide mapping techniques 2. Use a software tool to determine the optimal cleavage pattern of the target protein (see Note 1). Remember that disulfide interchange is minimized at low pH. Enzymes like trypsin, Endo Glu-C, Endo Asp-N, Endo Asn-C and pepsin are all active below a pH of 7 and are thus to be preferred. 3. Digest your protein as described above and separate half the sample by reversed phase HPLC. 4. Make the other half of the sample 10 mM in dithiothreitol or TCEP (will reduce you sample even at low pH). Let stand for at least 30 min before separation the sample under conditions identical to the separation in step 3).

Peptide Mapping for Protein Characterization mAU 100

14

15

11

80 60

979

7

8 9

0

5 3

40

1

2

12

6 10

4 13

20 214 nm

0

280 nm

-20

254 nm

-40 5.0

10.0

15.0

20.0

25.0

30.0

35.0

min

Fig. 1. Tryptic digest of 250 pmol bovine beta lactoglobulin. The traces of 214 nm, 254 nm and 280 nm are shown. The 254 nm and 280 nm traces are expanded x5 compared to the 214 nm trace. The stippled line show the gradient used. The identification of each peak is shown in figure 2. Peak 1 and peak 13 were not identified. Peak 13 is likely to be a partial digest product. The peaks at ‘0’ are a mixture of small peptides and the buffer salts from the injection.

10 20 30 40 50 LIVTQTMKGLDIQKVAGTWYSLAMAASDISLLDAQSAPLRVYVEELKPTP

6

4

15

60 70 80 90 100 EGDLEILLQKWENGECAQKKIIAEKTKIPAVFKIDALNENKVLVLDTDYK

2 9 3 8 10 11 110 120 130 140 150 KYLLFCMENSAEPEQSLACQCLVRTPEVDDEALEKFDKALKALPMHIRLS 14

12 160 FNPTQLEEQCHI

5

9

7

11

Fig. 2. Identification of the peaks separated in figure 1. Aromatic residues are shown in bold. The two peptides labeled 11 are linked by a disulphide bond (Cys66–Cys160). Peptide 12 has an internal disulphide bond. The peptides labeled 9 elute very close together (fig. 1).

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5. Compare the two chromatograms. Peaks that disappear from the first chromatogram and appear in the second chromatogram must arise from Cys containing peptides and will thus have to be identified by mass spectrometry, Edman degradation or amino acid analysis (see Note 10). 6. Closely spaced Cys residues in the primary structure presents a special problem as you will have to cleave between them for a positive identification of the disulfide bridge. In this case you will often have to isolate a large peptide containing multiple disulfide bridges followed by subdigestion using a less specific enzyme (Table 1) (5).

Mass spectrometry is particularly suited to disulfide bridge determination and several methods have been developed (7). 3.5. Peptide Map Purification by HPLC Other chapters in this book deals in detail with the purification of proteins and peptides by HPLC, so the following only emphasis the features relevant to peptide mapping. When purifying a peptide map by HPLC, the objective is usually to obtain the highest possible resolution, sensitivity and yield, but not to handle large amounts. The following description should be read in this context. For general peptide mapping use a large pore (300 Å), reversed phase column packed with the smallest possible particle size and the longest column length. These parameters are conflicting as backpressure in the system rises with both smaller particle size and longer column length. A compromise is to use a long 250 mm column packed with 5 µm particles or a shorter 150 mm column packed with 3 µm particles. Backpressure in the running system should not exceed 100 bar (increasing the temperature to 45-50 °C using a column oven will lower the back pressure and will additionally increase the resolution slightly). Using smaller diameter columns will increase the sensitivity of separation, if the HPLC system is optimized for the lower flow, at the cost of lower capacity (see Note 5). Furthermore, the peak volume from a 2.1 mm column is about 100 µL compared to 400-500 µL from a 4.6 mm column. This makes the volumes more compatible for further downstream processing (e.g. mass spectrometry or Edman degradation) and speeds up lyophilization (see Note 7). An in-line UV detector is essential for separating peptide mixtures. As we cannot be certain that all peptides contain aromatic residues, detection has to be carried out at a wavelength low enough to observe the absorbance of the peptide bond. However, as solvents like 0.1% TFA and acetonitrile also absorbs at low wavelength we need to find the wavelength with the highest signal to noise ratio. On most systems this is at 214-215 nm. If you do not operate with the highest purity of solvents, you may have to increase the wavelength to 218-220 nm, but your sensitivity will be reduced by a factor of three.

Peptide Mapping for Protein Characterization

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Multiple wavelength detection (e.g. 254 nm and 280 nm) can greatly increase the confidence of peptide identification (see Note 11). 3.5.1. Separating a Peptide Map 1. Prepare 1 liter of A-solvent (5% acetonitrile/0.1% TFA in ultra high quality water) and ½ liter of B-solvent (90% acetonitrile in water, 0.85% TFA) (see Notes 6 and 12). Degas solvents carefully before use or, better, use in-line degassing, as degassed solvents will re-absorb oxygen from the air within a few hours. 2. Program your HPLC system to a gradient of 1%/min from 0% to 40% B-solvent, followed by 2%/min from 40 to 70% B-solvent. Finally step to 100% B-solvent for 2 min before equilibrating at 5% for at least 10 column volumes until you have a stable baseline. If your peptide map is not too complex, you may shorten the run time by increasing the gradient to 1½%/min between 0 and 40% and 2.5–3%/min between 40 and 70%. 3. Make a blind run each morning to equilibrate the column. The first run of a day is usually atypical with a rising or falling baseline that may not be present in subsequent analyses. You can most easily check the condition of your HPLC system and reversed phase column by making a separation run using a standard peptide mixture (e.g. a tryptic digest of β-lactoglobulin) (see Note 13). 4. Dissolve your sample in A-solvent and inject. Keep the injector in the “inject” position for two injection loop volumes before you switch back to “load” position. If your sample is already dissolved in a different solvent (i.e. urea, tris or phosphate) you should allow 2–3 min of isocratic elution before starting the gradient. 5. Collect the isolated peaks in polypropylene tubes. If you run at low flow rates, you may have to take a delay between the detector and the effluent from the capillary into account (see Note 14). 6. Dry down your collected fractions in a vacuum centrifuge (see Note 7). For several applications you may not have to dry the sample; just evaporate the major part of the organic modifier.

For problems in peptide map separations, see Note 2. 4. Notes 1. When the primary structure of the protein being analyzed is known, a couple of software tools are available for planning the peptide map. These tools can predict the peptides generated by various proteolytic enzymes or chemical cleavages. One freeware tool, PAWS, is available for download (http://bioinformatics.genomicsolutions.com/Paws.html), while a somewhat more advanced tool, GPMAW (http://www.gpmaw.com) (7), will allow you to work with multiple sequences and, in addition to the prediction of peptides, provides additional physical/chemical characteristics of each peptide (e.g. charge, theoretical pI, hydrophobicity and size). GPMAW can even generate a simulated HPLC RP chromatogram. Although this chromatogram is only approximate (separation varies with column, system and

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running conditions), it does give valuable indications for problems during separation. The Expasy web site (http://www.expasy.ch/tools/) contains links to a large number of tools that are useful for peptide mapping (most are included in GPMAW). 2. If your peptide map contains small hydrophilic peptides they will elute very early in the reversed phase chromatogram, perhaps even as part of the solvent peak. Early eluting peaks may also have a bad peak shape, particularly if the column has not been equilibrated sufficiently before the start of the gradient. You may collect these early eluting peaks, dry them sufficiently to get rid of the organic solvent, and then re-separate on a 100–120Å pore size, high carbon load column (e.g. Hypersil ODS2 or equivalent) using 0.1%TFA / 90% methanol as a solvent system. The gradient should be a rather shallow 1%/min. gradient from 0 to 30%. An alternative is to separate hydrophilic peptides using HILIC (hydrophilic interaction chromatography), particularly glycopeptides can be efficiently separated in this way. Running HILIC chromatography is very similar to reversed phase chromatography, but the elution order of components is reversed. You dissolve your sample in high concentration of organic solvent and separate by running the gradient from high to low concentration of organic solvent. If on the other hand you have large, hydrophobic peptides you should use a 300Å, C4 or C8 column and elute with either acetonitrile or, for the really hydrophobic peptides, use 2-propanol. 3. The use of reversed phase purification as the last step before analysis of peptide mixtures is necessary to obtain the highest sensitivity and sequence coverage in mass spectrometric peptide mapping. You may use Zip-tips (Waters Inc.), but when analyzing peptide maps isolated from silver stained gel spots you can obtain considerable higher sensitivity by using homemade micro purification tips based on Eppendorf GELoader tips packed with Poros R2 50 µm reversed phase material (Applied BioSystems, Framingham, MA). For the purification of proteins you should use Poros R1 material. For details on how to construct these devices please see (8) The GELoader tip is constricted at the end using the blunt end of a pair of tweezers or a pencil. Column material is dispersed in acetonitrile and sufficient material is applied to make a 3–5 mm length column. The GELoader tip is initially washed with 10–20 µL of neat acetonitrile, and then it is equilibrated in 2 × 10 µL of 0.1% TFA. The sample is added in 1–20 µL of solvent followed by washing with 2 × 10 µL of 0.1% TFA. Finally the sample is eluted with 0.8–1 µL of alpha-cyano-4-hydroxycinnamic acid in 70% acetonitrile/0.1% TFA (4 mg/mL to concentrated depending on instrument) directly onto the MALDI target.

Peptide Mapping for Protein Characterization

983

4. Most buffers without an excessive amount of detergent or salt are suitable for proteolytic digestion. 50 mM ammonium bicarbonate is convenient because it can be removed by evaporation (do not use with lysyl endopeptidase). If you have large quantities or high concentrations of ammonium bicarbonate you may need to add a bit of acetic acid to help completely remove the ammonium bicarbonate. This buffer may be readily substituted for other buffers, e.g. 50 mM Tris-HCl, pH 8, or 50 mM phosphate buffer, pH 8.0. However, these buffers are not directly compatible with mass spectrometry. Hard to dissolve proteins can often be brought into solution by first dissolving the protein in small amount of 6–8 M urea (e.g. 5–10 µL) and then diluting the solution to the appropriate level (e.g. below 2 M for trypsin, see Table 1) with a buffer containing the enzyme for digestion. Another very efficient method is to use detergents, however, beware of downstream processing incompatibilities (e.g. SDS is incompatible with mass spectrometry and HPLC). RapiGest SF (Waters) and PPS Silent™ (Protein Discovery) are detergents that upon acidification degrade into smaller components which makes them compatible with both mass spectrometry and HPLC. Organic solvents (see Table 1) or heat denaturation may denature proteins, thus leading to faster, more efficient digestions (9). 5. The sensitivity of HPLC separations decreases with the square of the column diameter, thus a 2 mm column shows four times the sensitivity of a 4 mm column, if the HPLC system is built to handle the smaller diameter. As the column diameter decreases from 4 mm to 2 and 1 mm the optimal flow rate decreases from 1 mL/min to 0.25 mL/min and 0.06 mL/min. Unless you perform flow splitting you will need special HPLC pumps to be able to perform a gradient in 0.06 mL/min. In addition, the rest of the HPLC equipment has to be optimized for the lower flow rate in order to minimize extra column band broadening. The inner diameter of capillaries has to be decreased (see Table 2). You also need to make connections as short as possible, particularly after Table 2 Parameters for HPLC System for Binary Gradient Formation Column diameter

Typical flow Flow cell mL/min volume

Normal

4–4.6 mm

1.0

< 12 µL

Narrow

2–2.1 mm

0.2–0.25

< 8 µL

Micro

1 mm

0.05–0.07

< 2 µL

Capillary diameter

Sensitivity increase

0.25 mm / 0.10” 0.18 mm / 0.07” 0.12 mm / 0.05”

×1 ×4 × 16

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the column. The size of the detector flow cell is particularly important; as a standard flow cell with a volume of 10–12 µL will significantly decrease the resolution of a 2 mm column, even though the rest of the system is optimized (see Table 2). When changing to a smaller diameter column you may also have to replace the dynamic mixer as you may otherwise experience very long delay times. For dual pump systems, each pump should have a minimum flow of no more than 5% of the running flow (e.g. a minimum flow of no more than 12.5 µL/min for a set total flow of 250 µL/min), and a minimum step of no more than 1% of the set flow rate. You may manage with slightly lower performance as the dynamic mixer will smooth out irregularities, but particularly at the beginning of the gradient you may observe irregular separation behavior. 6. TFA deteriorates over time. As the 0.1 % TFA used for buffer A ages you will see an increase in baseline during gradient runs. You may increase/decrease the content of TFA in the B-solvent to compensate. A ratio of 1:0.85 is usually a good starting point, but varies slightly from detector to detector. If you experience regular baseline noise, you can try to lower the content of TFA in buffer A to 0.06% and the content in buffer B correspondingly (0.05%). 7. For small amounts of peptide, the drying step can be critical for recovery as the peptides have a tendency to adhere to the walls of the polypropylene tube (10). For many analytical purposes it is not necessary to change solvent or you may only have to evaporate the organic solvent. 8. An accurate calibration of your mass spectra is essential for correct protein identification. The most accurate calibration is to use known peaks in each spectrum for internal calibration. When using Promega modified trypsin (porcine) you will find the following autolytic peptides (monoisotopic m/z) that are suited for internal calibration: 842.510, 1045.564 and 2211.105. If these peaks in general are too high or too low, you should adjust the concentration (not the volume) of trypsin added in step 13. When analyzing the results of a search, you should consider the following when differentiating between hits: How do the deviations between theoretical and experimental masses fit together (i.e. do the deviations lie on a straight line when plotted against the peptide mass you have a calibration error). If a peptide contains a methionine it is likely to have a +16 Da peak. The oxidized methionine seldomly occurs by itself. N-terminal glutamine containing peptides are always accompanied by a −17 Da satellite peak (deamidation to pyroglutamic acid). Multibasic sites often give rise to lysine/arginine ladders (tryptic cleavage).

Peptide Mapping for Protein Characterization

9.

10.

11.

12.

985

Missed cleavages (i.e. peptides with an internal Lys or Arg residue) are much more likely to occur if there is a neighboring acidic residue (tryptic cleavage). In a peptide mixture, (small) lysine-containing peptides are often suppressed relative to arginine containing peptides due to the more basic nature of arginine. The GPMAW program (see Note 1) is also useful for checking the validity of a hit as you can perform detailed analysis of likely/unlikely peptide identifications as well as search for unusual enzyme cleavage sites. Several Web sites cater for the analysis of peptide mass data. Some analyze MS data (PMF) others MS/MS data. The actual number changes over time, but you may want to check out the following: Mascot (http://www.matrixscience.com) ProteinProspector (MsFit) (http://prospector.ucsf.edu/). Aldente (http://www.expasy.org/tools/aldente/) PeptideSearch (http://www.mann.embl-heidelberg.de/GroupPages/pageLink/ peptidesearchpage.html) XTandem (http://www.thegpm.org/) VEMS (http://yass.sdu.dk/) Phenyx (http://phenyx.vital-it.ch/pwi/login/login.jsp) You will have to read the accompanying help files for each search engine for detailed instructions. You cannot usually identify reduced cysteine residues by Edman degradation or amino acid analysis, but the surrounding sequence will in most cases be sufficiently unique that positive identification of the peptide is no problem. When separating a peptide map, it is advantageous to use multiple wavelengths. Modern in-line UV detectors can often be set up for three simultaneous wavelengths. You should use 214 nm for separating peptides, as you will get the highest response at this wavelength (you are measuring the absorbance of the peptide bond). Only the aromatic amino acid residues have any significant absorption at higher wavelength. Tryptophan and tyrosine have absorption maxima at 278 nm and 280 nm respectively while phenylalanine has maximum absorbance at 254 nm. You should thus set your detector at 214 nm, 254 nm and 280 nm, Tryptophan is usually recognized by having an absorbance several times that of tyrosine. The inclusion of TFA in the B-solvent is in order to create a stable baseline as both acetonitrile and TFA absorbs at low wavelength. Depending on your system you may have to vary the concentration of TFA in the B-solvent between 0.75 and 0.85 % (see Note 6). Mixing a small amount of organic solvent in the A-solvent and some water in the B-solvent helps to give a better mixing of the solvents (mixing

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water and acetonitrile is quite difficult), gives a better separation (helps to “wet” the hydrophobic surface, it may “collapse” in pure water) and helps to degas the A-solvent. If you have hydrophilic peptides that are eluted in the solvent front by the initial 5% organic content, you can use an A-solvent without organic content, but the first part of the chromatogram tend not to be so reproducible. Column manufacturers recommend that you store the column in 50% methanol/water. However, experience has shown that most columns are stable for several years when stored in 0.1% TFA/acetonitrile (preferably with a high content of organic solvent). 13. Easy-to-make test mixture of peptides for checking reversed phase columns for peptide mapping: Dissolve 182 µg β-lactoglobulin (Sigma Inc, #L0130) in 100 µL, 50 mM ammoniumbicarbonate, pH 8.2. Add 3.6 µg trypsin (Roche Biochemicals, #109 819) and digest for 4 hours at 37 °C. Stop the digestion with 900 µL 0.1% TFA. Total solution is now 1000 µL at a concentration of 10 pmol /µL. Divide into 55 µL portions and store at −20 °C. Use 50 µL for 4 mm columns and 20 µL for 2 mm columns. The digest is also useful as calibrant/check for MALDI-MS. 14. If you use an automatic fraction collector you should be careful to calibrate the fraction collector for exact peak collection. In addition you should be careful that the collector adds a minimum of additional peak tubing to the separation system as this will result in delays and mixing of your sample resulting in a loss of resolution. Manual fraction collection is usually more precise but you may (initially) need an assistant to assist in tube handling and annotation. For manual collection a strip chart recorder is most convenient, as computer acquisition or integrators often have an unacceptable delay. 15. Most proteolytic enzymes do not have an absolute specificity, but are active toward several residues with varying activities. Activity towards a given type of residue is usually also dependent on the local environment (e.g. 3-D structure and neighboring residues). Excessive digestion times may thus lead to secondary cleavages that are not desired and may be difficult to predict. Most of the enzymes that have a single residue specificity (Lys, Arg, Glu, Asp, Asn) will show some level of activity towards “like” residues. While the selection of the specific enzymes can be clearly related to peptide predictions based on the primary structure (e.g. using the GPMAW program), the selection of unspecific enzymes like pepsin, thermolysin and elastase is usually based on other parameters like low solubility of the target protein. They have a difficult to predict, low specificity that is very dependent on the local structure of the substrate. Pepsin is active at pH 2.0, a pH where proteins often are more soluble than at neutral pH.

Peptide Mapping for Protein Characterization

16.

17.

18.

19.

987

Thermolysin is active at temperatures as high as 80 °C (even in the presence of 8 M urea) where most proteins are denatured. Elastase targets a group of small hydrophilic residues not targeted by any of the “specific” enzymes. These enzymes are usually used either for an initial cleavage where you hope for few cleavage positions to either make a protein soluble or to cleave a large protein into smaller, more manageable pieces. In these cases you have to experiment with short digestion times and/or very dilute solutions. Alternatively the enzymes are used as a final cleavage to identify closely spaced disulfide bridges (see step 4). During digestion, trypsin will autolyse and generate a small amount of ψ-trypsin, which has some chymotryptic activity. In certain cases, depending on the sequence or residual structure of the substrate, this may lead to complete cleavage at single or a few residues in a protein. The activity of trypsin is also affected by neighboring residues as acidic groups will slow cleavages and a following proline residue will in most cases completely inhibit cleavage. If you have neighboring basic residues (e.g. -Lys-Lys-) trypsin may cleave at either residue, but as the activity of trypsin towards terminal residues is considerably lower than towards internal residues, you may find all cleavage variants present in the peptide map even after prolonged digestion times. Porcine trypsin is usually preferred rather than bovine, due to less autolysis. Modified trypsin, which is even more resistant to autolysis, is available from a number of manufacturers (Table 1). Chymotrypsin has a high activity towards the C-terminal side of aromatic residues, but other residues like Leu and Met can also be cleaved, particularly if they are situated in a hydrophobic environment (mostly -X-X↓Y where X is hydrophobic). Full digestions of intact proteins often give rise to unsuspected peptides, while the specificity when sub-digesting peptides is usually quite high. Endoproteinase Glu-C, also called S. aureus protease V8, is a most useful enzyme in protein mapping. Its specificity nicely complements that of trypsin while still generating peptides of a suitable size. The main activity is directed towards the C-terminal of glutamic acid, but a reasonable activity towards aspartic acid means that you may have to control digestion times carefully in order to limit cleavage to Glu. In a few cases activity has also been seen towards Gly, but this is sequence dependent. If the following residue is Glu or Pro, cleavage will not take place (e.g. in a -Glu-Glusequence cleavage is only observed after the second Glu). Earlier reports that Endo Glu-C was specific towards Glu at pH 4.0 seem to be caused by the lower activity of the enzyme at this pH (11). Lysyl endopeptidase, Endo Lys-C, Endo Asp-N, Endo Lys-N, and Endo Asn-C all tend to generate very clean peptide maps with a low amount of secondary cleavages. However, some of the peptides generated may be quite large

988

Højrup and difficult to handle. Endo Asp-N, Endo Lys-N, and Endo Asn-C are considerably more expensive than the other enzymes mentioned but a single optimal digest is more likely to succeed than multiple rounds of digestion and separation. Endo Arg-C (submaxillary gland proteinase) and clostripain both target the C-terminal side of arginine, but with a significant amount of lysine cleavage. Furthermore, their activity is not very high making them problematic for digesting intact proteins.

References 1. Suckau, D. and Resemann, A. (2003) T3-sequencing: targeted characterization of the N- and C-termini of undigested proteins by mass spectrometry. Anal Chem. 75, 5817–5824. 2. Macek, B., Waanders, L. F., Olsen, J. V., and Mann, M. (2006) Top-down protein sequencing and MS3 on a hybrid linear quadrupole ion trap-orbitrap mass spectrometer. Mol Cell Proteomics 5, 949–958. 3. De Hoffmann, E. and Stroobant, V. (2007) Mass Spectrometry: Principles and Applications, 3rd ed. John Wiley, New York. 4. Thingholm, T. E., Jørgensen, T. J., Jensen, O. N., and Larsen, M. R. (2006) Highly selective enrichment of phosphorylated peptides using titanium dioxide. Nat. Protoc. 1, 1929–1935. 5. Højrup, P., Jensen, M. S., and Petersen, T. E. (1985) Amino acid sequence of bovine protein Z: a vitamin K-dependent serine protease homolog. FEBS Lett 184, 333–387. 6. Peri, S., Steen H., and Pandey A. (2001) GPMAW – a software tool for analyzing proteins and peptides. Trends Biochem. Sci. 26, 691–693. 7. Gorman, J. J., Wallis, T. P., and Pitt, J. J. (2002) Protein disulfide bond determination by mass spectrometry, Mass Spectrom. Rev, 21, 183–216 8. Kussmann, M., Nordhoff, E., Rahbek-Nielsen, H., Háebel, S., Rossel-Larsen, M., Jacobsen, L., Gobom, J., Mirgorodskya, E., Kroll-Kristensen, A., Palm, L., and Roepstorff, P. (1997) Matrix-assisted laser desorption/ionization mass spectrometry: sample preparation techniques designed for various peptide and protein analytes. J. Mass Spectrom. 32, 593–601. 9. Russell W.K., Park Z.Y., and Russell D.H. (2001) Proteolysis in mixed organicaqueous solvent systems: applications for peptide mass mapping using mass spectrometry. Anal Chem. 73, 2682–2685. 10. Speicher K.D., Kolbas O., Harper S., and Speicher D.W. (2000) Systematic Analysis of Peptide Recoveries from In-Gel Digestions for Protein Identifications in Proteome Studies. J. Biomol. Tech. 11, 74–86. 11. Sørensen, S.B., Sørensen, T. L., and Breddam, K. (1991) Fragmentation of proteins by S. Aureus strain V8 protease. FEBS Lett. 294, 195–197.

106 Production of Protein Hydrolysates Using Enzymes John M. Walker and Patricia J. Sweeney

1. Introduction Traditionally, protein hydrolysates for amino acid analysis are produced by hydrolysis in 6 N HCl. However, this method has the disadvantage that tryptophan is totally destroyed, serine and threonine partially (5–10%) destroyed, and most importantly, asparagine and glutamine are hydrolyzed to the corresponding acids. Digestion of the protein/peptide with enzymes to produce protein hydrolysate overcomes these problems, and is particularly useful when the concentration of asparagine and glutamine is required. For peptides less than about 35 residues in size, complete digestion can be achieved by digestion with aminopeptidase M and prolidase. For larger polypeptides and proteins, an initial digestion with the nonspecific protease Pronase is required, followed by treatment with aminopeptidase M and prolidase. Since it is important that all enzymes have maximum activity, the following sections will discuss the general characteristics of these enzymes. 1.1. Pronase Pronase (EC 3.4.24.4) is the name given to a group of proteolytic enzymes that are produced in the culture supernatant of Streptomyces griseus K-1 (1–3). Pronase is known to contain at least ten proteolytic components: five serine-type proteases, two Zn2+ endopeptidases, two Zn2+-leucine aminopeptidases, and one Zn2+ carboxypeptidase (4, 5). Pronase therefore has very broad specificity, hence its use in cases where extensive or complete degradation of protein is required. The enzyme has optimal activity at pH 7.0–8.0. However, individual components are reported to retain activity over a much wider pH range (6–9). The neutral components are stable in the pH range 5.0–9.0 in the presence of calcium, and have optimal activity at pH 7.0–8.0. The alkaline components are stable in the pH range

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3.0–9.0 in the presence of calcium, and have optimal activity at pH 9.0–10.0 (4). The aminopeptidase and carboxypeptidase components are stable at pH 5.0–8.0 in the presence of calcium (9). Calcium ion dependence for the stability of some of the components (mainly exopeptidases) was one of the earliest observations made of Pronase (2). Pronase is therefore normally used in the presence of 5–20 mM calcium. The addition of excess EDTA results in the irreversible loss of 70% of proteolytic activity (10). Two peptidase components are inactivated by EDTA, but activity is restored by the addition of Co2+ or Ca2+. One of these components, the leucine aminopeptidase, is heat stable up to 70°C. All other components of Pronase lose 90% of their activity at this temperature (5). The leucine aminopeptidase is not inactivated by 9 M urea, but is labile on dialysis against distilled water (2). Some of the other components of Pronase are also reported to be stable in 8 M urea (2), and one of the serine proteases retains activity in 6 M guanidinium chloride (11). Pronase retains activity in 1% SDS (w/v) and 1% Triton (w/v) (12). Among the alkaline proteases, there are at least three that are inhibited by diisopropyl phosphofluoridate (DFP) (10). In general, the neutral proteinases are inhibited by EDTA, and the alkaline proteinases are inhibited by DFP (4). No single enzyme inhibitor will inhibit all the proteolytic activity in a Pronase sample. 1.2. Aminopeptidase M Aminopeptidase M (EC 3.4.11.2), a zinc-containing metalloprotease, from swine kidney microsomes (13–16) removes amino acids sequentially from the N-terminus of peptides and proteins. The enzyme cleaves N-terminal residues from all peptides having a free α-amino or α-imino group. However, in peptides containing an X-Pro sequence, where X is a bulky hydrophobic residue (Leu, Tyr, Trp, Met sulfone), or in the case of an N-blocked amino acid, cleavage does not occur. It is for this reason that prolidase is used in conjunction with aminopeptidase M to produce total hydrolysis of peptides. The enzyme is stable at pH 7.0 at temperatures up to 65°C, and is stable between pH 3.5 and 11.0 at room temperature for at least 3 h (15). It is not affected by sulfhydryl reagents, has no requirements for divalent metal ions, is stable in the presence of trypsin, and is active in 6 M urea. It is not inhibited by PMSF, DFP, or PCMB. It is, however, irreversibly denatured by alcohols and acetone, and 0.5 M guanidinium chloride, but cannot be precipitated by trichloroacetic acid (15). It is inhibited by 1,10-phenanthroline (10M) (16). Alternative names for the enzyme are amino acid arylamidase, microsomal alanyl aminopeptidase, and α-aminoacyl peptide hydrolase. 1.3. Prolidase Prolidase (EC 3.4.13.9) is highly specific, and cleaves dipeptides with a prolyl or hydroxyprolyl residue in the carboxyl-terminal position (17, 18). It has no

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activity with tripeptides (19). The rate of release is inversely proportional to the size of the amino-terminal residue (19). The enzyme’s activity depends on the nature of the amino acids bound to the imino acid. For optimal activity, amino acid side chains must be as small as possible and apolar to avoid steric competition with the enzyme receptor site. The enzyme has the best affinity for alanyl proline and glycyl proline. The enzyme has optimal activity at pH 6.0–8.0, but it is normally used at pH 7.8–8.0 (20). Manganous ions are essential for optimal catalytic activity. The enzyme is inhibited by 4-chloromercuribenzoic acid, iodoacetamide, EDTA, fluoride, and citrate. However, if Mn2+ is added before iodoacetamide, no inhibition is observed (21). Alternative names for the enzyme are imidodipeptidase, proline dipeptidase, amino acyl l-proline hydrolyase, and peptidase D. 2. Materials 1. Buffer: 0.05 M ammonium bicarbonate, pH 8.0 (no pH adjustment needed) or 0.2 M sodium phosphate, pH 7.0 (see Note 1). 2. Pronase: The enzyme is stable at 4°C for at least 6 mo and is usually stored as a stock solution of 5–20 mg/mL in water at −20°C. 3. Aminopeptidase M: The lyophilized enzyme is stable for several years at −20°C. A working solution can be prepared by dissolving about 0.25 mg of protein in 1 mL of deionized water to give a solution of approx 6 U of activity/mL. This solution can be aliquoted and stored frozen for several months at −20°C. 4. Prolidase: The lyophilized enzyme is stable for many months when stored at −20°C and is stable for several weeks at 4°C if stored in the presence of 2 mM MnCl2 and 2 mM β-mercaptoethanol (18).

3. Methods 3.1. Digestion of Proteins (22) 1. Dissolve 0.2-µmol of protein in 0.2 mL of 0.05M ammonium bicarbonate buffer, pH 8.0, or 0.2 M sodium phosphate, pH 7.0 (see Note 1). 2. Add Pronase to 1% (w/w), and incubate at 37°C for 24 h. 3. Add aminopeptidase M at 4% (w/w), and incubate at 37°C for a further 18 h. 4. Since in many cases the X-Pro- bond is not completely cleaved by these enzymes, to ensure complete cleavage of proline-containing polypeptides, the aminopeptidase M digest should be finally treated with 1 µg of prolidase for 2 h at 37°C. 5. The sample can now be lyophilized and is ready for amino acid analysis (see Note 2).

3.2. Digestion of Peptides (22) This procedure is appropriate for polypeptides less than about 35 residues in length. For larger polypeptides, use the procedure described in Subheading 3.1. 1. Dissolve the polypeptides (1 nmol) in 24 µL of 0.2 M sodium phosphate buffer, pH 7.0, or 0.05 M ammonium bicarbonate buffer, pH 8.0 (see Note 1).

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2. Add 1 µg of aminopeptidase M (1 µL), and incubate at 37°C. 3. For peptides containing 2–10 residues, 8 h are sufficient for complete digestion. For larger peptides (11–35 residues), a further addition of enzyme after 8 h is needed, followed by a further 16-h incubation. 4. To ensure complete cleavage at proline residues, finally treat the digest with 1 µg of prolidase for 2 h at 37°C. 5. The sample can now be lyophilized and is ready for amino acid analysis (see Note 2).

4. Notes 1. Sodium phosphate buffer should be used if ammonia interferes with the amino acid analysis. 2. When using two enzymes or more, there is often an increase in the background amino acids owing to hydrolysis of each enzyme. It is therefore important to carry out a digestion blank to correct for these background amino acids. References 1. Hiramatsu, A. and Ouchi, T. (1963) On the proteolytic enzymes from the commercial protease preparation of Streptomyces griseus (Pronase P). J. Biochem. (Tokyo) 54(4), 462–464. 2. Narahashi, Y. and Yanagita, M. (1967) Studies on proteolytic enzymes (Pronase) of Streptomyces griseus K-1. Nature and properties of the proteolytic enzyme system. J. Biochem. (Tokyo) 62(6), 633–641. 3. Wählby, S. and Engström, L. (1968) Studies on Streptomyces griseus protease. Amino acid sequence around the reactive serine residue of DFP-sensitive components with esterase activity. Biochim. Biophys. Acta 151, 402–408. 4. Narahashi, Y., Shibuya, K., and Yanagita, M. (1968) Studies on proteolytic enzymes (Pronase) of Streptomyces griseus K-1. Separation of exo- and endopeptidases of Pronase. J. Biochem. 64(4), 427–437. 5. Yamskov, I. A., Tichonova, T. V., and Davankov, V. A. (1986) Pronase-catalysed hydrolysis of amino acid amides. Enzyme Microb. Technol. 8, 241–244. 6. Gertler, A. and Trop, M. (1971) The elastase-like enzymes from Streptomyces griseus (Pronase). Isolation and partial characterization. Eur. J. Biochem. 19, 90–96. 7. Wählby, S. (1969) Studies on Streptomyces griseus protease. Purification of two DFP-reactin enzymes. Biochim. Biophys. Acta 185, 178–185. 8. Yoshida, N., Tsuruyama, S., Nagata, K., Hirayama, K., Noda, K., and Makisumi, S. (1988) Purification and characterisation of an acidic amino acid specific endopeptidase of Streptomyces griseus obtained from a commercial preparation (Pronase). J. Biochem. 104, 451–456. 9. Narahashi, Y. (1970) Pronase. Meth. Enzymol. 19, 651–664. 10. Awad, W. M., Soto, A. R., Siegei, S., Skiba, W. E., Bernstrom, G. G., and Ochoa, M. S. (1972) The proteolytic enzymes of the K-1 strain of Streptomyces griseus obtained from a commercial preparation (Pronase). Purification of four serine endopeptidases. J. Biol. Chem. 257, 4144–4154.

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11. Siegel, S., Brady, A. H., and Awad, W. M. (1972) Proteolytic enzymes of the K-1 strain of Streptomyces griseus obtained from a commercial preparation (Pronase). Activity of a serine enzyme in 6M guanidinium chloride. J. Biol. Chem. 247, 4155–4159. 12. Chang, C. N., Model, P., and Blobel, G. (1979) Membrane biogenesis: cotranslational integration of the bacteriophage F1 coat protein into an Escherichia coli membrane fraction. Proc. Natl. Acad. Sci. USA 76(3), 1251–1255. 13. Pfleiderer, G., Celliers, P. G., Stanulovic, M., Wachsmuth, E. D., Determann, H., and Braunitzer, G. (1964) Eizenschafter und analytische anwendung der aminopeptidase aus nierenpartikceln. Biochem. Z. 340, 552–564. 14. Pfleiderer, G. and Celliers, P. G. (1963) Isolation of an aminopeptidase in kidney tissue. Biochem. Z. 339, 186–189. 15. Wachsmuth, E. D., Fritze, I., and Pfleiderer, G. (1966) An aminopeptidase occuring in pig kidney. An improved method of preparation. Physical and enzymic properties. Biochemistry 5(1), 169–174. 16. Pfleiderer, G. (1970) Particle found aminaopeptidase from pig kidney. Meth. Enzymol. 19, 514–521. 17. Sjöstrom, H., Noren, O., and Jossefsson, L. (1973) Purification and specificity of pig intestinal prolidase. Biochim. Biophys. Acta 327, 457–470. 18. Manao, G., Nassi, P., Cappugi, G., Camici, G., and Ramponi, G. (1972) Swine kidney prolidase: Assay isolation procedure and molecular properties. Physiol. Chem. Phys. 4, 75–87. 19. Endo, F., Tanoue, A., Nakai, H., Hata, A., Indo, Y., Titani, K., and Matsuela, I. (1989) Primary structure and gene localization of human prolidase. J. Biol. Chem. 264(8), 4476–4481. 20. Myara, I., Charpentier, C., and Lemonnier, A. (1982) Optimal conditions for prolidase assay by proline colourimetric determination: application to iminodipeptiduria. Clin. Chim. Acta 125, 193–205. 21. Myara, I., Charpentier, C., and Lemonnier, A. (1984) Minireview: Prolidase and prolidase deficiency. Life Sci. 34, 1985–1998. 22. Jones, B. N. (1986) Amino acid analysis by o-phthaldialdehyde precolumn derivitization and reverse-phase HPLC, in Methods of Protein Microcharacterization (Shively, J. E., ed.), Humana Press, Totowa, NJ, pp. 127–145.

107 Amino Acid Analysis by Precolumn Derivatization with 1-Fluoro-2,4-Dinitrophenyl-5-I-Alanine Amide (Marfey’s Reagent) Sunil Kochhar and Philipp Christen

1. Introduction Precolumn modification of amino acids and the subsequent resolution of their derivatives by reverse-phase high-performance liquid chromatography (RP-HPLC) is the preferred method for quantitative amino acid analysis. The derivatization step introduces covalently bound chromophores necessary not only for interactions with the apolar stationary phase for high resolution but also for photometric or fluorometric detection. Marfey’s reagent, 1-fluoro-2,4-dinitrophenyl-5-L-alanine amide or (S)-2[(5-fluoro-2,4-dinitrophenyl)-amino]propanamide, is used to separate and to determine enantiomeric amino acids (1). The reagent reacts stoichiometrically with the amino group of enantiomeric amino acids to produce stable diastereomeric derivatives, which can readily be separated by reverse-phase HPLC (Fig. 1). The dinitrophenyl alanine amide moiety strongly absorbs at 340 nm (ε = 30,000 M−1 cm−1), allowing detection in the subnanomolar range. With smaller columns, the sensitivity will correspondly increase. Pre-column derivatization with the reagent can also be used to quantify the 19 commonly analyzed L-amino acids (2). Major advantages of Marfey’s reagent over other pre-column derivatizations are: (1) possibility to carry out chromatography on any multipurpose HPLC instrument without column heating; (2) photometric detection at 340 nm that is insensitive to most solvent impurities; and (3) stable amino acid derivatives. For the occasional user, the simple methodology provides, an attractive and inexpensive alternative to the dedicated amino acid analyzer. Further development to on-line derivatization and microbore chromatography has been reported (3). The pre-column derivatization From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Marfey’s reagent (I) and l,l-diastereomer derivative from l-amino acid and the reagent (II).

with Marfey’s reagent has found applications in many diverse areas of biochemical research (4–16; for recent reviews, see 17, 18), including determination of substrates and products in enzymic reactions of amino acids (see Note 1). 2. Materials 2.1. Vapor-Phase Protein Hydrolysis 1. Pyrex glass vials (25–50 × 5 mm) from Corning. 2. Screw-capped glass vials. 3. 6 N HCl 1.from Pierce.

2.2. Derivatization Reaction 1. Amino acid standard solution H from Pierce. 2. Marfey’s reagent from Pierce. Caution: Marfey’s reagent is a derivative of 1- fluoro-2,4-dinitrobenzene, a suspected carcinogen. Recommended precautions should be followed in its handling (19). 3. HPLC/spectroscopic grade triethylamine, methanol, acetone, and dimethyl sulfoxide (DMSO) from Fluka.

2.3. Chromatographic Analysis 1. Solvent delivery system: Any typical HPLC system available from a number of manufacturers can be used for resolution of derivatized amino acids. We have used an HPLC systems from Bio-Rad (HRLC 800 equipped with an autoinjector AST 100), Hewlett-Packard 1050 system and Waters system. 2. Column: A silica-based C8 reverse-phase column, for example, LiChrospher 100 RP 8 (250 × 4.6 mm; 5 µm from Macherey-Nagel), Aquapore RP 300 (220 × 4.6 mm; 7 µm from Perkin-Elmer), Nucleosil 100-C8 (250 × 4.6 mm; 5 µm from MachereyNagel), or Vydac C8 (250 × 4.6 mm; 5 µm from The Sep/a/ra/tions group). 3. Detector: A UV/VIS HPLC detector equipped with a flow cell of lightpath 0.5–1 cm and a total volume of 3–10 µL (e.g., Bio-Rad 1790 UV/VIS monitor,

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5. a. b.

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Hewlett-Packard Photodiode Array 1050, Waters Photodiode Array 996). The derivatized amino acids are detected at 340 nm. Peak integration: Standard PC-based HPLC software with data analysis program can be employed to integrate and quantify the amino acid peaks (e.g., ValueChrom from Bio-Rad, Chemstation from Hewlett-Packard or Millennium from Waters). Solvents: The solvents should be prepared with HPLC grade water and degassed. Solvent A: 13 mM trifluoroacetic acid plus 4% (v/v) tetrahydrofuran in water. Solvent B: Acetonitrile (50% v/v) in solvent A.

3. Methods 3.1. Vapor-Phase Protein Hydrolysis 1. Transfer 50–100 pmol of protein sample or 20 µL of amino acid standard solution H containing 2.5 µmol/mL each of 17 amino acids into glass vials and dry under reduced pressure in a SpeedVac concentrator. 2. Place the sample vials in a screw cap glass vial containing 200 µL of 6 N HCl. 3. Flush the vial with argon for 5–15 min and cap it airtight. 4. Incubate the glass vial at 110 °C for 24 h or 150 °C for 2 h in a dry-block heater. 5. After hydrolysis, remove the glass vials from the heater, cool to room temperature, and open them slowly. 6. Remove the insert vials, wipe their outside clean with a soft tissue paper, and dry under reduced pressure.

3.2. Derivatization Reaction 1. Add 50 µL of triethylamine/methanol/water (1:1:2) to the dried sample vials, mix vigorously by vortexing, and dry them under reduced pressure (see Note 2). 2. Prepare derivatization reagent solution (18.4 mM) by dissolving 5 mg of Marfey’s reagent in 1 mL of acetone. 3. Dissolve the dried amino acid mixture or the hydrolyzed protein sample in 100 µL of 25% (v/v) triethylamine in water and add 100 µL of the Marfey’s reagent solution and mix by vortexing (see Note 3). 4. Incubate the reaction vial at 40 °C for 60 min with gentle shaking protected from light (see Note 4). 5. After incubation, stop the reaction by adding 20 µL of 2 N HCl (see Note 5). Dry the reaction mixture under reduced pressure. 6. Store the dried samples at −20 °C in the dark until used (see Note 6). 7. Dissolve the dry sample in 50% (v/v) DMSO.

3.3. Chromatographic Analysis 1. Prime the HPLC system according to the manufacturer’s instructions with solvent A and solvent B. 2. Equilibrate the column and detector with 90% solvent A and 10% solvent B. 3. Bring samples for analysis to room temperature; dilute, if necessary, with solvent A, and inject 20 µL (50–1000 pmol) onto the column (see Note 7).

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4. Elute with gradient as described in Table 1 (see Note 8). Resolution of 2,4-dinitrophenyl-5-L-alanine amide derivatives of 18 commonly occurring L-amino acids and of cysteic acid is achieved within 120 min (Fig. 2) (see Note 9). 5. Determine the response factor for each amino acid from the average peak area of standard amino acid chromatograms at 100, 250, 500, and 1000 pmol amounts. 6. When HPLC is completed, wash the column and fill the pumps with 20% (v/v) degassed methanol for storage of the system. Before re-use, purge with 100% methanol.

Table 1 HPLC Program for Resolution of Amino Acids Derivatized with Marfey’s Reagent Time (min)

% Solvent A

% Solvent B

0 15 15.1 115 165 170

90 90 90 50 0 0

10 10 10 50 100 100

Input Detector auto zero Inject

Fig. 2. Separation of 2,4-dinitrophenyl-5-l-alanine amide derivatives of l-amino acids by HPLC. A 20-µL aliquot from the amino acid standard mixture (standard H from Pierce Chemical Company) was derivatized with Marfey’s reagent. Chromatographic conditions: Column, LiChrospher 100 RP 8 (250 × 4.6 mm; 5 µm from Macherey-Nagel); sample, 100 pmol of diastereomeric derivatives of 18 l-amino acids; solvent A, 13 mM trifluoroacetic acid plus 4% (v/v) tetrahydrofuran in water; solvent B, 50% (v/v) acetonitrile in solvent A; flow rate, 1 mL/ min; detection at 340 nm; elution with a linear gradient of solvent B in solvent A (Table 1). The amino acid derivatives are denoted by single-letter code, cysteic acid as CYA and cystine as Cs. FFDA indicates the reagent peak; the peaks without denotations are reagent-related as shown in an independent chromatographic run of the reagent alone.

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4. Notes 1. For analysis of amino acids as substrates or products in an enzymic reaction, deproteinization is required. Add 4 M perchloric acid to a final concentration of 1 M and incubate on ice for at least 15 min. Excess perchloric acid is precipitated as KClO4 by adding an equal volume of ice-cold 2 M KOH. After centrifugation for 15 min at 4 °C, the supernatant is collected, dried, and used for derivatization. 2. Complete removal of HCl is absolutely essential for quantitative reaction between amino acids and Marfey’s reagent. 3. The molar ratio of the Marfey’s reagent to the total amino acids should not be more than 3:1. 4. During derivatization, the color of the reaction mixture turns from yellow to orange-red. 5. On acidification, the color of the reaction mixture turns to yellow. 6. The dried amino acids are stable for over one month when stored at −20 °C in the dark. In 50% (v/v) DMSO, derivatives are stable for 72 h at 4 °C and for > 6 wk at −20 °C. 7. To obtain good reproducibility, an HPLC autoinjector is highly recommended. 8. Silica-based C8 columns from different commercial sources produce baseline resolution of the 19 amino acid derivatives; nevertheless, the gradient slope of solvent B should be optimized for each column. S-carboxymethyl-l-cysteine and tryptophan, if included, are also separated in a single chromatographic run (see ref. 2). 9. The identity of each peak is established by adding a threefold molar excess of the amino acid in question. A reagent blank should be run with each batch of the Marfey’s reagent to identify reagent-related peaks. Excess reagent interferes with baseline separation of the arginine and glycine peaks. Lysine and tyrosine are separated as disubstituted derivatives. References 1. Marfey, P. (1984) Determination of D-amino acids. II. Use of a bifunctional reagent, 1,5-difluoro-2,4-dinitrobenzene. Carlsberg Res. Commun. 49, 591–596. 2. Kochhar, S. and Christen, P. (1989) Amino acid analysis by high-performance liquid chromatography after derivatization with 1-fluoro-2,4-dinitrophenyl-5-Lalanine amide. Analyt. Biochem. 178, 17–21. 3. Scaloni, A., Simmaco, M., and Bossa, F. (1995) D-L Amino acid analysis using automated precolumn derivatization with 1-fluoro-2,4-dinitrophenyl-5-alanine amide. Amino Acids 8, 305–313. 4. Kochhar, S. and Christen, P. (1988) The enantiomeric error frequency of aspartate aminotransferase. Eur. J. Biochem. 175, 433–438. 5. Martínez del Pozo, A., Merola, M., Ueno, H., Manning, J. M., Tanizawa, K., Nishimura, K., et al. (1989) Stereospecificity of reactions catalyzed by bacterial D-amino acid transaminase. J. Biol. Chem. 264, 17,784–17,789.

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6. Szókán, G., Mezö, G., Hudecz, F., Majer, Z., Schön, I., Nyéki, O., et al. (1989) Racemization analyses of peptides and amino acid derivatives by chromatography with pre-column derivatization. J. Liq. Chromatogr. 12, 2855–2875. 7. Adamson, J. G., Hoang, T., Crivici, A., and Lajoie, G. A. (1992) Use of Marfey’s reagent to quantitate racemization upon anchoring of amino acids to solid supports for peptide synthesis. Analyt. Biochem. 202, 210–214. 8. Goodlett, D. R., Abuaf, P. A., Savage, P. A., Kowalski, K. A., Mukherjee, T. K., Tolan, J. W., et al. (1995) Peptide chiral purity determination: hydrolysis in deuterated acid, derivatization with Marfey’s reagent and analysis using high-performance liquid chromatographyelectrospray ionization-mass spectrometry. J. Chromatogr. A 707, 233–244. 9. Moormann, M., Zähringer, U., Moll, H., Kaufmann, R., Schmid, R., and Altendorf, K. (1997) New glycosylated lipopeptide incorporated into the cell wall of smooth variant of Gordona hydrophobica. J. Biol. Chem. 272, 10,729–10,738. 10. Gramatikova, S. and Christen, P. (1997) Monoclonal antibodies against Na-(5´ phos-phopyridoxyl)-L-lysine. Screening and spectrum of pyridoxal 5 -phosphatedependent activities toward amino acids. J. Biol. Chem. 272, 9779–9784. 11. Vacca, R. A., Giannattasio, S., Graber, R., Sandmeier, E., Marra, E., and Christen, P. (1997) Active-site Arg→Lys substitutions alter reaction and substrate specificity of aspartate aminotransferase. J. Biol. Chem. 272, 21,932–21,937. 12. Wu, G. and Furlanut, M. (1997) Separation of DL-dopa by means of micellar electrokinetic capillary chromatography after derivatization with Marfey’s reagent. Pharmacol. Res. 35, 553–556. 13. Goodnough, D. B., Lutz, M. P., and Wood, P. L. (1995) Separation and quantification of D-and L-phosphoserine in rat brain using N-alpha-(2,4-dinitro-5-fluorophenyl)L-alanine amide (Marfey’s reagent) by high-performance liquid chromatography with ultra-violet detection. J. Chromatogr. B. Biomed. Appl. 672, 290–294. 14. Yoshino, K., Takao, T., Suhara, M., Kitai, T., Hori, H., Nomura, K., et al. (1991) Identification of a novel amino acid, o-bromo-L-phenylalanine, in egg-associated peptides that activate spermatozoa. Biochemistry 30, 6203–6209. 15. Tran, A. D., Blanc, T., and Leopold, E. J. (1990) Free solution capillary electrophoresis and micellar electrokinetic resolution of amino acid enantiomers and peptide isomers with L- and D-Marfey’s reagents. J. Chromatogr. 516, 241–249. 16. Montes-Bayón, M., B’Hymer, C., Ponce de Léon, C., and Caruso, J. A. (2001) Resolution of seleno-amino acid optical isomers using chiral derivatization and inductively coupled plasma mass spectrometric (ICP-MS) detection. J. Anal. At. Spectrom. 16, 945–950. 17. B’Hymer, C., Montes-Bayón, M., and Carruso, J. A. (2003) Marfey’s reagent: Past, present, and future uses of 1-fluoro-2,4-dinitrophenyl-5-l-alanine amide. J. Sep. Sci. 26, 7–19. 18. Bhushan, R., and Brückner, H. (2004) Marfey’s reagent for chiral amino acid analysis: A review. Amino Acids 27, 231–247. 19. Thompson, J. S. and Edmonds, O. P. (1980) Safety aspects of handling the potent allergen FDNB. Ann. Occup. Hyg. 23, 27–33.

108 Amino Acid Analysis of Protein Hydrolysates Using Anion Exchange Chromatography and IPAD Detection Petr Jandik, Jun Cheng, and Nebojsa Avdalovic

1. Introduction Following the commercial introduction as “AAA Direct,” amino acid analysis by anion exchange chromatography and integrated pulsed amperometric detection (IPAD) (1) has been applied in many different types of amino acid assays (1–16). The new method does not require any derivatization. It is thus easier to use and less costly than derivatizaton-based methods. Additional savings are achieved by the relatively inexpensive single-component mobile phases and by the low flow rate (0.25 mL/min). The limits of detection are in the femtomol range and linear calibration plots extend over three orders of magnitude for the most amino acids (1). Various protocols for analyzing protein and peptide hydrolysates are outlined in the literature (1,9–11). The results obtained for a collagen hydrolysate were compared with those generated by cation exchange / ninhydrin method (1). Simultaneous analysis of carbohydrates and amino acids was reported by several authors (12–16). The use of the technique for monitoring and control of metabolism in mammalian cell has been discussed (16). In this chapter, we provide suitable protocols for the validation of reproducibility and accuracy. Such validation should be performed not only during a start up but also in regular intervals during the routine use. Evaluation of reproducibility is made easier by a new report format of Dionex software. We expect many of the new users of AAA Direct to be experienced in one of the post- or pre-column derivatization techniques. In anticipation, we include guidelines for the selection of compatible buffers, surfactants and reagents.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Several of the compounds that are not compatible or difficult to use with the older methods do not cause any problems when utilized in conjunction with the AAA Direct (i.e. Triton X100, urea, ammonia from the hydrolysis of polyacrylamide, sodium hydroxide). On the other hand, some chemicals that are in widespread use in the pre- or post-column derivatization-based protocols do interfere, if utilized with our method. The most important examples of the latter category are tris(hydroxymethyl)aminomethane (Tris) or carbohydrates when present in high concentrations. However, both Tris and carbohydrates interfere only within the simplest format of AAA Direct. Recently, the authors have reported a modified version of AAA Direct overcoming Tris and carbohydrate interference (17,18). 2. Materials 2.1. Chromatography 1. Water 18 megohm (Millipore, Bedford, MA, Milli Q or equivalent, see Note 1) 2. Sodium hydroxide solution 50% w/w (Certified Grade, Fisher Scientific, Pittsburgh, PA) 3. Sodium acetate, anhydrous (Microselect Grade, FLUKA, Milwaukee, MI) 4. Helium or argon gas cylinder, pressure regulation valve, Tygon tubing connecting cylinder to eluent containers. 5. Sterile filtration units from Nalge (0.2 µm Nylon filter in covered funnels, vol = 1 L, VWR, West Chester, PA) 6. Serological glass pipets, vol =10 mL (Fisher Scientific) 7. Eluent A: Water 18 megohm filtered through a 0.2 µm Nylon filter (see Notes 2 and 3) 8. Eluent B: Filter 1 L of water through a 0.2 µm Nylon filter; using the serological pipets (see Note 4), add 13.1 mL of 50% sodium hydroxide to the filtered 1L-volume of water (see Note 3). 9. Eluent C: Dissolve 82 g of anhydrous sodium acetate in ca. 500 mL of water. Fill up to 1L and filter through the 0.2 µm Nylon filter. 10. AminoPac PA10 Guard Column (2 × 50mm) and AminoPac PA10 analytical column (2 × 250mm). 11. Ternary low-pressure gradient, microbore, HPLC system, Dionex BioLC or comparable with an autosampler (e.g. Dionex AS50) and column thermostat. The system must include Dionex ED40 electrochemical detector. 12. AAA certified gold cell for use with the ED40 detector. 13. PC with installed copy of Dionex PeakNet 6.1 or equivalent (see Note 5).

2.2. Reproducibility Testing Using Injections of Standards 1. Amino Acids in 0.1 M HCl, Standard Reference Material 2389 (National Institute of Standards and Technology, Gaithersburg, MD) 2. Norleucine (Sigma, St. Louis, MO) 3. Sodium azide (Sigma) 4. 0.1 M HCl (reagent grade)

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5. Diluent containing norleucine and sodium azide: Prepare 4 mM stock solution of norleucine (524.8 mg/L) in 0.1 M HCl. Dilute 500x with a solution of azide (20mg/L). 6. Reproducibility Standard: pipet 320 µL of SRM 2389 into a 100 mL volumetric flask and fill up to volume with the norleucine diluent. If stored in a refrigerator, the Reproducibility Standard is stable for up to 30 days. If used at room temperature, the Reproducibility Standard remains stable for up to three days (i.e. over a weekend). 7. The Reproducibility Standard from item 6 does not contain all of the standard compounds that are required for the analysis of certain types of hydrolysate samples. These standards can be purchased from Sigma and are listed below: δ-hydroxylysine, D(+)galactosamine hydrochloride, D(+)-glucosamine hydrochloride, hydroxy-L-proline, phospho-L-arginine sodium salt, o-phospho-D,L-serine, o-phospho-D,L-threonine, o-phospho-D,L-tyrosine, L-cysteic acid hydrate and L-tryptophan. 8. Prepare 1 mM, single component solutions of all standards from step 7 in 0.1 M HCl and store them in a refrigerator. As required, add 100 µL of one or more of the above single-component stock solution to the Reproducibility Standard (step 6). 9. Disposable transfer pipets (glass or polyethylene)

2.3. Accuracy of Amino Acid Analysis in a BSA Hydrolysate 1. Bovine serum albumin (7% solution) Standard Reference Material 927c (National Institute of Standards and Technology, Gaithersburg, MD) 2. Hydrolysate glass tubes with Teflon Plugs (Pierce, Rockford, IL) 3. Speedvac (Savant, Farmingdale, NY) 4. Heating Module with aluminum heating block (Pierce) 5. Constant Boiling HCl (Pierce)

2.4. Selecting Suitable Buffers, Surfactants and Reagents (see Note 6) 1. Buffers, surfactants, and reagents found to be compatible with AAA Direct: ACES (see Note 7); ADA; AMPSO; CAPS (see Note 7); CAPSO; CHES (see Note 7); citric acid (Fluka); EDTA (Aldrich); MES (Boehringer Manheim); MOBS; MOPS; MOPSO; n-octyl-β-glucoside (see Note 8); phenol; PIPES (see Note 7) sodium diphosphate; sodium monophosphate; sodium triphosphate; sodium azide; TES; TRITON X 100 (Aldrich). 2. Buffers, surfactants, and reagents producing interfering peaks in AAA Direct: BES (BDH Chemicals, Poole England); Bicine; BISTRIS (see Note 9); CHAPS; CHAPSO; DTE; EPPS; HEPES; imidazole; TEA (see Note 8); Tricine (see Note 9); TRIS (see Note 9).

3. Methods 3.1. Chromatography (see Note 10) The chromatogram in Fig. 1 was generated using the gradient conditions in Table 1 and detection conditions in Table 2. We recommend using those conditions for all reproducibility and accuracy testing described below.

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GalN P-Thr

GlcN

300

His

P-Ser

nC

Arg P-Arg

200

Hyp Ala

Gly

Met Pro

Val

100

Phe

Ser

Thr

Ile

Cys

P-Tyr Tyr

Glu Nleu Asp

Leu

Cya

Trp

0 0

10

20

30

40

Minutes Fig. 1. Simultaneous separation of amino acids (including hydroxy-proline and hydroxylysine, or Hyp and Hyl, respectively), amino sugars (GalN and GlcN), phospho amino acids (P-Arg, P-Ser and P-Thr) and cysteic acid (Cya) under the conditions of standard AAA Direct gradient.

Fig. 1 illustrates the capability of AAA Direct for protein and peptide hydrolysates. All common types of hydrolysates (HCl, HCl /propionic acid, MSA, NaOH,) can be analyzed under identical gradient and detection conditions. An important exception is the separation of methionine sulfone, that can be achieved only at the column temperature of 35 °C (See ref 9). In contrast, the correct column temperature for all other separations (including that in Fig. 1) is 30 °C. 1. Connect the fully assembled ED40 cell directly to the injector outlet. Running the initial gradient conditions (76%A, 24%B at 0.25 mL/min) switch the cell voltage on (Specify “Integrated Amperometry” and apply Waveform from Table 2). Verify that the signal is lower than 80 nC. 2. Install the AminoPac PA10 column set and check again that the signal background is at less than 80 nC. 3. Allow the column oven to reach 30 °C and run a blank gradient (see Table 1) injecting 25 µL of water. Verify that gradient rise is not exceeding 30 nC. 4. Prepare an aliquot of Reproducibility Standard (Subheading 2.2, item 6) and place it into the autosampler. Follow proper procedures in executing the first standard injection (vol = 25 µL) by not allowing any delay between the end of the blank gradient run and first injection of standard. 5. Verify the absence of extraneous peaks in the blank gradient and a correct value of peak height for arginine (>100 nC). Verify the presence of peaks for all amino

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acids. P-amino acids, hydroxylysine, hydroxyproline, amino sugars, cysteic acid and tryptophan are not present in the Reproducibilty Standard. Verify that the separation between alanine and threonine in the Reproducibility Standard chromatogram is comparable to that in Fig. 1.

3.2. Reproducibility Testing Using Injections of Standards 1. Verify the system status by carrying out steps 1-5, Subheading 3.1. 2. Transfer 1.5 mL of Reproducibility Standard (Subheading 2.2, step 6) into a glass autosampler vial. Place the full vial into the autosampler. Make sure there is also a vial with water inside the autosampler. 3. Create a “PeakNet Sequence” for running a blank gradient (vol = 25 µL injection of water, Table 1 gradient, Table 2 waveform) followed by nine injections of Reproducibility Standard (vol = 25 µL “Full Loop,” Table 1, Table 2). Do not perform more than nine vol =25 µL Full-Loop-injections out of a single 1.5 mL-vial. 4. Create or update a quantitation method (.qnt) and include it in the PeakNet sequence. A meaningful evaluation of reproducibility relies on correct identification and integration of all peaks of interest. The correct identification and integration is dependent on a correct user input of retention times and integration parameters (see Note 11). The retention times can be updated and integration parameters optimized during the

Table 1 Gradient Conditions for Hydrolysates (Flow rate: 0.25 mL/min) Time (min) Init. 0.00 2 8 11 18 21 23 42 42.1 44.1 44.2** 75***

%A* 76 76 76 64 64 40 44 14 14 20 20 76 76

%B* 24 24 24 36 36 20 16 16 16 80 80 24 24

%C* 0 0 0 0 0 40 40 70 70 0 0 0 0

Curve

8 8 5 8 5 5

* Eluents A,B,C: See Subheading 2.1., Steps7,8,9 **Start of reequilibration to initial conditions ***Complete reequilibration and column clean-up under normal conditions

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Potential (V)* vs. pH

0 40 50 110 210 220 460 470 560 570 580 590 600

0.13 0.13 0.33 0.33 0.33 0.55 0.55 0.33 0.33 −1.67 −1.67 0.93 0.13

Integration

Begin

End

*

Sequence of potentials (or waveform) applied to the Au working electrode and referenced vs. Glass/Ag/AgCl combination electrode.

5.

6.

7.

8.

first standard run. It is not necessary to input pmol amounts for amino acids and to carry out a calibration at this stage as reproducibility is evaluated from the relative standard deviations of peak areas. Following the completion of all standard runs, verify correct identification and integration of all peaks by inspecting all nine standard chromatograms separately. Close the files you opened for identification and integration verification. Select all nine standard chromatograms simultaneously (press Ctrl. and click on corresponding row numbers). In the menu that opens up by rightclicking on one of the selected rows, choose “compare>ECD1” and an overlay of all selected chromatograms appears on the screen. Rightclick anywhere in the upper-right portion of the chromatographic overlay. In the menu that now appears choose “Load Report Format>%RSD Peak Areas” (see Note 12) to generate a similar report as the one shown in Fig. 2. Rightclick in the upper-right portion of the screen, go to “Load Report Format” again and switch to %RSD Retention Times (actual report not shown).

3.3. Accuracy of Amino Acid Analysis in a BSA Hydrolysate The following protocol utilizes a data reduction procedure producing three numerical QC values, “Average % Error,” Recovery [g],”and “% Recovery,” characterizing the accuracy of results for a protein of known amino acid composition.

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Fig. 2. Reproducibility Report. Relative standard deviations for each amino acid are listed in the bottom row. The report is linked to all data files in a given PeakNet Sequence. Any change of integration parameters produces instant updates of the corresponding area, average, and %RSD values.

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(1)

Where: % Error = 100* ABS (analytical result for n – true value for n)/true value for n ABS (x): absolute value of x n = number of each amino acid per molecule Recovery [g] = {(Corrected Average Ratio) * molecular weight of protein}

(2)

Where: Corrected Average Ratio: mean value of those First Ratios values deviating by less than 20% from the Average of First Ratios (Corrected Average Ratio represents the analytical result for the mol-amount of protein injected) First Ratio = analytical result in pmol of amino acid/ true value of n % Recovery = {(Recovery [g] *8}/ g of Protein in 200 µL of hydrolysate)

(3)

This data reduction approach has been used, for example, in the “multicenter” (round robin) studies organized by the Association of Biomolecular Resource Facilities or ABRF (19). Evaluation of accuracy is simplified in AAA Direct by an automatic calculation of QC parameters from pmol results after calibration. As in the reproducibility evaluation (Subheading 3.2), the parameters are cal culated directly within the HPLC software. It is not necessary to export area or pmol results into another application such as Excel first (see Note 14). As a protein of known amino acid composition, we have chosen the 7 % BSA solution from the National Institute of Standard and Technology (Subheading 2.3, item 1). The amino acid composition of BSA can be obtained in the literature (20). The NIST documentation certifies that the SRM 927c is from bovine blood and has the correct molecular weight and spectral properties for BSA. It also states that the SRM 927c may be used for calibration in amino acid analysis. The SRM 927c documentation, however, does not certify the amino acid composition of the actual protein sample used to prepare the 7% solution. 1. Prepare a 500fold dilution of SRM 927c in water containing 20 mg/L sodium azide. 2. Transfer 33.6 µL of 500x dilute SRM 927c into a clean microvial (vol=1.5 mL) and carry out evaporative centrifugation (Speedvac) to dryness. 3. Add to the dry residue 100 µL of constant boiling HCl and vortex thoroughly. 4. Using glass Pasteur pipets transfer the solution to a clean hydrolysis tube (Subheading 2.3, step 2). 5. To the microvial emptied in step 4, add a new 50 µL aliquot of constant boiling HCl, vortex and subsequently transfer as much as possible of the 50 µL aliquot to the sample in the hydrolysis tube from step 4. Use the same transfer pipet as in step 4.

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6. Alternate vacuum and inert gas inside the hydrolysis tube three times. Place fully evacuated tube into a heating module (Subheading 2.3, item 4). Carry out hydrolysis for 16–24 hours at 110 °C. 7. Remove the hydrolysis tube from the heating module and allow it to cool down to the ambient temperature. 8. Using a clean glass Pasteur pipet transfer the hydrolyzed sample from the hydrolysis tube into a clean microvial. 9. Pipet another 50 µL aliquot of constant boiling HCl into the hydrolysis tube from step 8. Rinse and subsequently transfer as much as possible of the 50 µL aliquot to the first aliquot of hydrolysate in the microvial. Use the same transfer pipet for steps 8 and 9. 10. Carry out evaporative centrifugation of the combined hydrolysate aliquots from steps 8 and 9. 11. To the completely dry residue from step 10, add 200 µL of norleucine/azide diluent (Subheading 2.2, item 5) and vortex thoroughly. 12. Transfer an aliqot of the reconstituted hydrolysate into an autosampler vial (Polyethylene vial, (vol=0.3 mL, with a split septum, Dionex). Make sure that vials with water and calibration standard have also been placed into the autosampler tray. 13. Create a “PeakNet Sequence” starting with an injection of water blank and followed by at least three injections of both, the calibration standard (Subheading 2.2, item 6) and hydrolyzed sample. Use a program file (.pgm) that includes the gradient and waveform from Tables 1 and 2, respectively. Input correct pmol amounts (see Note 15) into the Amount Table of the quantitation method (.qnt, Subheading 3.2, step 4). Use 25 µL Fool-Loop injections for all samples and standards. In the quantitation method, specify the sequence line containing the calibration standard. Start the automatic execution of the entire sequence. 14. After executing all runs, inspect each standard and sample chromatogram. Verify correct identification and integration of all amino acid peaks.

In a hydrolysate chromatogram, rightclick in the upper-right portion of the chromatogram. In the menu that now appears choose “Load Report Format>QCBSA (see Note 14).” A similar report as the one in Fig. 3 appears on the screen. 15. Make sure to input the actual concentration of BSA in the hydrolysate as specified in the report subtitle (line 3). Switch to “Layout Mode” (Edit>Layout Mode) to enable input of BSA concentration. The calculated values of Average % Error, Protein Recovery [g] and % Protein Recovery appear instantaneously in the lower half of the report page. The BSA QC report is now ready to be saved or printed out. 16. Additional QC Reports for the other BSA hydrolysate injections can be generated by moving from one sample file to another using the corresponding two icons on the screen (left and right arrow, top bar, third row). To complete the documentation, it is also possible to generate the %RSD Area , %RSD Retention Times reports for the standard injections of the same sequence by carrying out the steps 6 to 8 of the Subheading 3.2.

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Fig. 3. Accuracy Report. The upper table is the usual “Amount Table” of PeakNet HPLC software. The pmol amounts in the sample are calculated from the pmol/area coefficients in the calibration file. The lower of the two tables evaluates equations (1–3) in the Subheading 3.3. This report is also linked to all data files of a given sequence. Any change of integration parameters produces an instant change of all calculated results.

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3.4. Selecting Suitable Buffers, Surfactants, and Reagents 1. Generate chromatograms of 1 mM solutions of all buffers, surfactants and reagents present in your protein or peptide samples. 2. Hydrolyse (Subheading 3.3., steps 6–12) 20 µL aliquots of 10 mM solutions of all buffers, surfactants and reagents in 180 µL of constant boiling HCl. 3. Generate chromatograms of hydrolyzed and reconstituted (azide/norleucine diluent) samples from steps 1 and 2 using the chromatographic conditions of Subheadings 3.2 and 3.3. 4. Avoid using chemicals causing peaks interfering with amino acids or carry out desalting prior to hydrolysis of protein solutions containing such additives. See Subheading 2.4. for guidelines in the selection of new additives. 5. Repeat steps 1-4 for all new batches of additives.

4. Notes 1. Minimize extraneous plastic tubing connected to the laboratory water unit and use only filters supplied by the manufacturers. Polymeric surfaces support bacterial growth. Bacterial contamination of water used in preparation of eluents causes high backgrounds and spurious peaks in amino acid analysis. 2. The main purpose of eluent filtration is to eliminate bacterial contamination stemming from the water purification unit and from the atmosphere. Wear powder-free gloves whenever preparing or replenishing mobile phases to avoid contamination by skin contact. 3. Minimize carbonate contamination of the eluents. Follow the guidelines in the Installation Instructions and Troubleshooting Guide for AAA Direct Amino Acid Analysis System (Dionex Document No. 031481). 4. Most vol=10 mL pipets can be used for the prescribed volume of 13.1 mL. Usually this larger volume is at or near the markings for the maximum possible volume. Verify the correct height of 13.1 mL by pipeting and weighing out water. Do not re-use serological pipets. 5. Make sure that the “Report Publisher” option is part of the installed software and it is switched on. (In the Pull-down menu: Help/About PeakNet) 6. Except where indicated otherwise, all of the buffers, reagents and surfactants were purchased from Sigma. 7. Four of the buffers that we evaluated (ACES, CAPS, CHES, PIPES) gave well-defined peaks, sufficiently separated from those of hydrolysate amino acids under the gradient conditions in Table 1. Users are advised to run control blanks when planning to utilize those buffers. Some of the peaks appear to be impurities unrelated to the actual buffer compound and may vary depending on the source and age of the chemical. 8. Glucoside-based surfactants produce a large interference peak when injected directly. However, they are completely hydrolyzed after 16–24 hours and

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10.

11.

12.

13.

14.

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110 °C in constant boiling HCl and do not thus interfere in a completely hydrolysed sample. Several of the non-anionic buffers (BISTRIS, TEA, TricineTRIS) produce a large peak interfering with arginine. This interference can be resolved using the modified AAA Direct technique described in ref. (17). A copy of the Installation Instructions and Troubleshooting Guide for the AAA Direct Amino Acid Analysis (Dionex Document No. 031481) is included with each AminoPac PA10 column shipped by Dionex. The text under Heading 3 describes only a subset of AAA Direct methodology used for reproducibility and accuracy testing. Optimization of integration parameters and other general guidelines for reproducible quantitation are described in “PekNet 6 User’s Guide.” (Dionex Document No. 031630). Specify the version of PeakNet you are running when ordering the document from Dionex. The “Excel-like” environment in the report part of Dionex HPLC software makes it possible for a user to create many different report formats executing simple or complex mathematical formulas. However, the user has to purchase the “Report Publisher” option to be able to generate a custom report format (see Note 5 for verification of presence of Report Publisher in the installed software). The electronic files for reproducibility evaluation without installing the Report Publisher can be obtained from Dionex Customer Support. Specify “Reproducibility Evaluation of AAA Direct” when ordering the report format file. A comprehensive evaluation of AAA Direct in conjunction with hydrolysis of peptides in HCl and MSA is presented in ref. (10). Reference (9) contains detailed protocols for HCl, MSA (with and without performic acid oxidation) and NaOH hydrolysates. Also described in ref. (9) is a new method for using the same set of chromatographic conditions for automatic analysis of carbohydrates, amino sugars and amino acids in different types of TFA and HCl hydrolysates of glycoproteins. As in the reproducibility evaluation (Subheading 3.2 and Note 12), it is possible for a user to create highly customized report formats for the QC parameters of accuracy evaluation. The user has to have either the “Report Publisher” option installed on the PC (see Note 5) or obtain a suitable report format file from Dionex customer support. Specify “Accuracy of Amino Acid Analysis” when ordering the report format file from Dionex. For a 321.5-fold dilution of SRM 2389 and 25 µL injection, divide all values in Table 1 of SRM 2389 by 0.0125. For example: the original concentration of 2.50 mmol/L of aspartic acid yields 200 pmol amount to be entered into the Amount Table.

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References 1. Clarke, A. P., Jandik, P., Rocklin, R. D., Liu, Y., and Avdalovic, N. (1999) An integrated amperometry waveform for the direct, sensitive detection of amino acids and amino sugars following anion-exchange chromatography. Anal. Chem. 71, 2774–2781. 2. Jandik, P., Cheng, J., Evrovski, J., and Avdalovic, N. (2001) Simultaneous analysis of homocysteine and methionine in plasma. J. Chromatogr. B, 759, 145–151. 3. Hanko, V. P. and Rohrer, J. S. (2002) Direct determination of tryptophan using high-performance anion-exchange chromatography with integrated pulsed amperometric detection. Anal. Biochem. 308, 204–209. 4. Lasater, A., Cruz, N. F., Cheng, J., Jandik, P., and Dienel, G. A. (2005) Scaling up a microbore separation for semipreparative analysis: differential recoveries of radiolabeled amino acids. Anal. Biochem. 342, 28–33. 5. Liang, L., Mo, S., Cai, Y., Mou, S. Jiang, G., and Wen, M. (2006) Direct amino acid analysis method for speciation of selenoamino acids using high-performance anion-exchange integrated pulsed amperometric detection. J. Chromatogr. A 1118, 134–138. 6. Ding, Y., Yu, H., and Mou, S. (2002) Direct determination of free amino acids and sugars in green tea by anion-exchange chromatography with integrated pulsed amperometric detection. J. Chromatogr. A 982, 237–244. 7. Hanko, V. P. and Rohrer, J. S. (2004), Determination of amino acids in cell culture and fermentation broth media using anion-exchange chromatography with integrated pulsed amperometric detection. Anal. Biochem. 324, 29–38. 8. Mo, S., Liang, L.Cai, Y., Mou , S., and Wen, M. (2006) Analysis of amino acids in soy sauce with high performance anion-exchange chromatography and electrochemical detector. Chin. J. Anal. Lab. 25, 36–39. 9. Jandik, P., Pohl, C., Barreto V., and Avdalovic, N. Anion Exchange Chromatography and Integrated Amperometric Detection of Amino Acids, in Methods in Molecular Biology, Vol. 159: Amino Acid Analysis Protocols (Cooper, C., Packer, N., and Williams, K., eds.). Humana Press, Totowa, NJ. 10. Dionex Corporation (2000), Determination of the Amino Acid Content of Peptides by AAA-Direct. Technical Note 50, pp. 1–20. 11. Dionex Corporation (2004), Determination of Protein Concentration Using AAA-Direct. Application Note 163, pp. 1–15. 12. Jandik, P., Clarke, A. P., Avdalovic, N., Andersen, D. C., and Cacia, J. (1999) Analyzing mixtures of amino acids and carbohydrates using bi-modal integrated amperometric detection. J. Chromatogr. B 732, 193–201. 13. Yu, H., Ding, Y. S., Mou, S. F., Jandik, P., and Cheng, J. (2002) Simultaneous determination of amino acids and carbohydrates by anion-exchange chromatography with integrated pulsed amperometric detection. J. Chromatogr. A 966, 89–97. 14. Jandik, P. Cheng, J. Avdalovic, N. (2004) Analysis of amino acid-carbohydrate mixtures by anion exchange chromatography and integrated pulsed amperometric detection. J. Biochem. Biophys. Methods 60, 191–203. 15. Hanko, V. P., Roher, J. S. (2006) Accurate determination of amino acids in highcarbohydrate-containing samples. Am. Biotech. Lab. 24, 131–141.

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16. Larson, T.M., Gawlitzek, M., Evans, H., Albers, U., and Cacia, J. (2002) Chemometric evaluation of on-line high-pressure liquid chromatography in mammalian cell cultures: Analysis of amino acids and glucose. Biotech. Bioeng 77, 553–563. 17. Jandik, P., Cheng, J., Jensen, D., Manz, S., and Avdalovic, N. (2000) New technique for increasing retention of arginine on an anion-exchange column. Anal. Biochem. 287, 38–44. 18. Jandik, P., Cheng, J., Jensen, D., Manz, S., and Avdalovic, N. (2001) Simplified inline sample preparation for amino acid analysis in carbohydrate containing samples. J. Chromatogr. B 758, 189–196. 19. Hunziker, P., Andersen, T. T., Bao, Y., Cohen, S. A., Denslow, N. D., Hulmes, J. D., Mahrenholz, A. M., Mann, K., Schegg, K. M., West, K. A., and Crabb, J. W. (1999) Identification of proteins electroblotted to polyvinylidene difluoride membrane by combined amino acid analysis and bioinformatics: An ABRF Multicenter Study. J. Biomolec. Techn. 10, 129–136. 20. Brown, J. R., (1975) Structure of Bovine Serum Albumine. Fed. Proc. 34, 591.

109 Validation of Amino Acid Analysis Methods Andrew J. Reason

1. Introduction (see Notes 1–4) This chapter presents a discussion of the points to consider during the validation of analytical methods and more specifically amino acid analysis. Amino acid analyses are utilised in various areas of research for free amino acids (in, for example, foodstuffs) and for analysis of products and components of interest comprising peptides, polypeptides and peptide/protein conjugates. In this Chapter, I have concentrated on the validation of amino acid analyses that are to be submitted in a package to obtain a regulatory authority licence for a peptide, protein, or conjugate product. In such studies, amino acid analyses can be used to provide various pieces of information generally required for biotechnological and/or biological products. For example quantitative amino acid analysis can be used to determine protein quality and quantity. Such data, combined with Optical Density (OD) measurements, can be used to determine the extinction coefficient for a protein. Analyses that are to be submitted in a package to obtain a regulatory authority licence for a particular product should be validated in accordance with the International Conference on Harmonisation of Technical Requirements for Registration of Pharmaceuticals for Human Use (ICH) Harmonised Tripartite Guidelines “Validation of Analytical Procedures: Text and Methodology” (1) except where there are specific issues for unique tests used for analyzing biotechnological and biological products (2). The terms and definitions within these documents are meant to bridge the differences that often exist between the regulators in Europe, Japan and USA. The validation criteria discussed below can be assessed using both precolumn and postcolumn amino acid analysis derivatization methods. The available techniques are described in detail in other chapters of this book (10–15). Requirements of speed and sensitivity have led

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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M-Scan to utilize a method involving acid hydrolysis followed by precolumn derivatization using phenylisothiocyanate and separation by reverse-phase highperformance liquid chromatography (RP-HPLC) for amino acid analysis. This method is used as a background to the discussion of validation of amino acid analysis methods. 2. Materials 2.1. Equipment for Hydrolysis of Proteins and Precolumn Derivatization of the Released Amino Acids (See Note 5) ? 4.6 mm, 5 1. An RP-HPLC system equipped with a Supelcosil LC-18-DB (250 

µm; Supelco, Poole, UK) HPLC column, and capable of tertiary solvent/buffer supply, gradient control, autoinjection, UV detection at 254 nm and data acquisition (see Note 6). 2. Vacuum hydrolysis tubes (Pierce, Rockford, IL) 5 mL internal volume. 3. Vacuum pump system capable of evacuating hydrolysis tubes to 1–2 Torr. 4. Oven or dry heating block capable of heating vacuum hydrolysis tubes to hydrolysis temperature (usually 110°C).

2.2. Chemicals (see Note 5) 2.2.1. Hydrolysis of Proteins and for Precolumn Derivatization of the Released Amino Acids 1. 2. 3. 4.

Constant boiling 6 M HCl (Pierce). Coupling buffer, acetonitrile:pyridine:water:triethylamine (10:5:3:2 v/v). Phenylisothiocyanate (Pierce). Amino Acid Standard H (Pierce), containing a mixture of 17 amino acids at 2.5 mM (cystine at 1.25 M) in 0.1 M HC1 (see Note 7). 5. Internal standard, L-Norleucine (Pierce).

2.2.2. RP-HPLC Separationof Derivatized Amino Acids 1. Solvent A: Ultra High Quality (UHQ) water (1 L). 2. Solvent B: Acetonitrile (HPLC grade):UHQ water (80:20 v/v), 1 L. 3. Buffer C, Weigh out 28.7g of anhydrous sodium acetate and dissolve in 500 mL of UHQ water ensuring that all the solid dissolves. Add 1.25 mL of triethylamine and adjust the pH of the buffer to 6.4 using glacial acetic acid. 4. Sample loading solvent, Solvent A:Solvent B:Buffer C (75:5:20 v/v).

3. Methods 3.1. Preparation of Amino Acid Standard Mixture 1. Weigh out a known amount of norleucine internal standard (∼1 mg). 2. Dissolve the weighed internal standard (Norleucine) in 5% (v/v) acetic acid in water to generate a 1 nanomole/µL solution. This solution is stable for up to 12 mo at −20° C ± 5°C and can be used as the internal standard bulk solution.

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3. An aliquot (50 µL) of the resulting solution, i.e., 50 nmoles is added to each sample being prepared for amino acid analysis and to the tube to be used for the amino acid standard mixture. 4. Add 20 µL of Amino Acid Standard H to 50nmoles of the norleucine (internal standard) in the tube set aside for the standard mixture. 5. The standard mixture of amino acids including the internal standard (norleucine) is derivatized in parallel with the samples to be studied (see Subheading 3.3).

3.2. Hydrolysis (see Notes 8 and 9) 1. Aliquot the desired amount of peptide, protein or conjugate (ideally 50–100 µg) to be hydrolyzed into a clean vacuum hydrolysis tube and label the tube(s) to identify the sample. 2. Add a known amount (ideally 50 nmoles) of norleucine (internal standard) to the sample tube. 3. Lyophilize resulting sample under vacuum using a Savant or similar drying device. 4. Add 200 µL of 6 M (constant boiling) hydrochloric acid. 5. Evacuate the vacuum hydolysis tube(s) for 2 min with agitation. 6. Incubate samples in a heating block or oven at 110°C for 24 h. 7. Allow samples to cool and lyophilize the products. 8. Derivatize released amino acids with phenylisothiocyanate.

3.3. Derivatization of Released Amino Acids and Amino Acid Standard Mixture 1. Add 100 µL of coupling reagent to the dried hydrolyzed or amino acid standard sample (see Note 10). 2. Lyophilize (see Note 10). 3. Add a further 100 µL of coupling reagent and 5 µL of phenylisothiocyanate (PITC) to the dried sample. 4. Incubate the mixture at room temperature for 5 min. 5. Lyophilize the products. 7. Add 100 µL UHQ water to the dried products (see Note 11). 8. Lyophilize (see Note 11). 9. Resuspend products in 200 µL of sample loading buffer and analyze an aliquot by RP-HPLC as described in Subheading 3.4.

3.4. RP-HPLC Separation of Derivatized Amino Acids 1. Connect the flow to the Supelcosil LC-18-DB HPLC column within the column heater compartment equilibrated at 45°C. 2. Wash the column with solvent B until a stable baseline is observed (see Note 12). 3. Load the amino acid analysis gradient parameters and allow the system to equilibrate at the initial conditions (see Table 1 and Note 13). 4. Once the baseline has stabilized inject 10 µL of the sample loading buffer and run the gradient shown in Table 1.

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Reason Table 1 Reversed Phase-High Performance Liquid Chromatography Conditions for Separation of PTC-Derivatized Amino Acids Time (min) 0 25 28 33 34

% Solvent

% Solvent

% Solvent

A

B

C

75 30 20 20 75

5 50 80 80 5

20 20 0 0 20

Flow rate (mL/min) 1.0 1.0 1.0 1.0 1.0

5. Visually examine the chromatogram obtained and ensure that there is no significant UV absorption in the region of interest (between 5 and 25 min following injection; see Fig. 1). If the chromatogram contains no significant absorbance in the region of interest proceed to step 6. If UV peaks are observed the column should be cleaned or a new column used and a second aliquot of sample loading buffer should be analyzed. 6. Inject sequentially six separate 10 µL aliquots of the derivatized amino acid standard mixture. 7. Examine the resulting chromatograms for peak area variation, elution time variation, peak resolution, and tailing to determine that the system is suitable for use. Peak resolution should be calculated using the equation:

r=2(t2−t1) W2+W1 Where, t2 and t1 are the corresponding widths at the bases of the peaks obtained by extrapolating the relatively straight sides of the peaks to the baseline (see Fig. 2). Peak tailing should be calculated using the equation: W

0.05

2f Where W0.05 is the width of the peak at 5% height and, f is the distance from the peak front to the peak center at 5% peak height (see Fig. 3). If the system is suitable for use (see Note 14) analysis of samples may continue. If the data do not pass the set criteria, a test regime must be established to determine the reason for the failure (see Note 15). 8. Inject sequentially duplicate aliquots (10 µL) of the samples of interest. 9. Following each run sample peaks should be integrated taking into account the internal standard. The response ratio for each amino acid in each sample run is calculated by dividing the peak area for each amino acid by the peak area obtained for the internal standard (norleucine) in the same run.

Fig. 1. UV chromatogram (254 nm) obtained from analysis of a standard mixture of derivatized amino acids.

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Fig. 2.

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Fig. 3.

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Calculated by dividing the peak area for each amino acid by the peak area obtained for the internal standard (norleucine) in the same run. The response ratio is in turn divided by the response ratio for the same amino acid in the amino acid standard mixture. The figure generated is multiplied by the known amount of the relative amino acid present in the standard mixture (i.e., 50 nmoles) to produce the molar amount of that amino acid in the sample. This may be summarized in the equations, Rr = Rs/Ris (in the sample data) Rm = Rs/Ris (in the standard mixture data) Rc = Rr/Rm nmoles of amino acid = Rc × 50 Where Rr is the response ratio for a particular amino acid in the sample or standard run of interest having a peak area Rs and an internal standard peak area (in the same run) Ris and, Rm is the response ratio for the same amino acid in the relevant standard mixture having a peak area Rs and an internal standard peak area (in the same run) Ris. and, Rc is the overall response ratio. 3.5. Validation Parameters (see Note 16) The validation parameters that should be assessed during a full validation to ICH guidelines are outlined in Subheadings 3.5.1.–3.5.9. Samples should be prepared as outlined in Subheadings 3.1.–3.4. or using another suitable amino acid analysis procedure. 3.5.1. Specificity Specificity is defined as the ability to assess unequivocally the analyte(s) in the presence of other components. A suitable study should be designed to establish retention times and responses of known amino acids released from the study sample and the data obtained should be compared to that obtained from a standard mixture containing derivatized amino acids. Generally, a standard aliquot of the peptide, protein, or conjugate for which the validation is being carried out is hydrolyzed and analyzed in duplicate and the data generated is compared with the data obtained from derivatized phenylthiocarbamyl (PTC)-amino acid standards. The retention time of each of the PTC-amino acids released from the peptide, protein, or conjugate sample should normally be within ±3% of the same PTCamino acid present in the standard mixture analyzed in the same session. 3.5.2. System Suitability Test (see Note 17) System suitability is a test designed to ascertain the effectiveness of the operating system.

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System suitability can be determined in a number of ways. Generally a series of replicate injections (n = 6) of the derivatized amino acid standards is performed to assess the relative standard deviations for peak area measurements and retention times. In addition, peak tailing (T) and peak resolution (R) can be determined (see Subheading 3.3). The retention time variance should normally be within 3% and the relative standard deviation of peak areas taken from the six sets of data should normally be less than 5%. The peak tailing factor (T) should be less than or equal to two and the resolution factor (R) should be greater than one for the amino acids of interest. 3.5.3. Linearity Linearity is defined as the ability (within a given range) of the analytical method to obtain results that are directly proportional to the concentration of analyte(s) in the sample. Linearity may be demonstrated on the peptide, protein, or conjugate and standard mixture of amino acids via dilution of a standard stock solution or on separate weighings out of the peptide, protein, or conjugate. Linearity of response of the amino acid standard mixture can be demonstrated by choosing at least 5 points in duplicate in a range to cover that anticipated for the peptide, protein, or conjugate. This study should be repeated with aliquots of the peptide, protein, or conjugate (5 points in duplicate; 0, 50, 75, 100, and 150% of the usual aliquot analyzed). Linearity should initially be determined using a visual inspection of a plot of peak area ratio (area of amino acid signal over the area of internal standard signal) against relative amount of the peptide, protein, or conjugate taken. If a linear relationship exists the plot obtained should be statistically evaluated. For example the sum of least squares around the regression line (r2) should be calculated. From the final plot, the correlation coefficient, y-intercept, slope of the regression line, and residual sum of squares should be determined and entered in the validation document. The linearity data obtained should obey the equation y = mx + C, where C is zero within 95% confidence limits, and the sum of least squares around the regression line (r2) is greater than 0.98. 3.5.4. Range The range is derived from the linearity study and is established by confirming that the analytical procedure provides an acceptable degree of linearity, accuracy, and precision when applied to samples containing amounts of peptide, protein, or conjugate within or at the extremes of the specified range of the analytical procedure.

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Following the linearity procedure described above, the working range for the analysis would be 50–150% of the concentration of the batch used for the analysis. If the working range required is likely to be wider, additional concentrations may be added to the linearity study to take into account the future expected concentration range. 3.5.5. Accuracy/Recovery An assay method should provide a true (accurate) result for a test peptide, protein, or conjugate sample. Accuracy should be established across the specified range of the analytical procedure and can often be inferred once precision, linearity, and specificity have been established. Generally percentage recoveries for the amino acids of interest should fall within the range of 90–110% and the percentage correlation of variance (%CV; ? 100 / mean) for multiple measurements should normally standard deviation  be less than 10%. 3.5.6. Repeatability Repeatability is measured by carrying out analysis (in six replicates) in one laboratory by one operator, using one instrument over a relatively short time span. A standard aliquot of the peptide, protein or conjugate is hydrolyzed and the derivatized products are injected six times onto the RP-HPLC system. In general, the percentage correlation of variance (%CV) for the released amino acids should be less than or equal to 5%. 3.5.7. Intermediate Precision Intermediate Precision is defined as long term variability of the measurement process, which may be determined for a method run on different days and or by different operators. Six separate aliquots of the peptide, protein, or conjugate should be hydrolyzed, derivatized and analyzed by RP-HPLC on different days. The analyses should be carried out using different operators and different instruments in the same facility (see Note 18). In general, the percentage correlation of variance (%CV) for the released amino acids should be less than or equal to 10%. 3.5.8. Reproducibility Reproducibility expresses the precision between laboratories and can be assessed using an extension of the intermediate precision study described in Subheading 3.5.7 to include analyses at different laboratories.

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Ideally, the percentage correlation of variance (%CV) for the released amino acids should be less than or equal to 10%. 3.5.9. Robustness/Ruggedness Robustness/ruggedness is defined as a review of the critical parameters in the method and the steps taken to minimise method variability. Generally experiments should be designed to assess the following criteria: 1. 2. 3. 4. 5. 6.

Stability of analytical solutions and derivatized products. Influence of variations of pH in a mobile phase. Influence of variations in mobile-phase composition. Different columns (different lots and/or suppliers). Temperature. Flow rate.

For most validation studies it is impractical to carry out analyses to study all the factors previously outlined. The use of experimental matrix software can often be used to design a set of experiments to test the robustness/ruggeddness of the method being validated. Acceptance criteria are set prior to the start of the analyses and should adhere to the criteria set for linearity and percentage correlation of variance (%CV) described earlier. Unfortunately there are no actual in-depth guidelines to aid us in designing robustness/ruggeddness analyses. 3.5.10. Detection and Quantification Limit (i.e., Minimum Amount of Amino Acid Needed for Reliable Detection and Quantification) Several approaches for determining the detection limit are possible (2). However, during a validation of amino acid analysis detection limits for each amino acid can be most easily determined based on the standard deviation of the response and the slope of the linearity plot. The detection limit (DL) may be expressed as: DL = 3.3 αS ?  ? the standard deviation of the response about the line of best fit Where  ? the slope of the calibration curve The quantification limit (QL) may be S determined in the same way and is expressed as:

QL = 10 αS ?  ? the standard deviation of the response, and S  ? the slope of the where  calibration curve.

4. Notes For other methods of amino acid analysis some details may differ, according to equilibration times, column lifetime, sensitivity (e.g., the amount of standard

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required), and so forth, but the general principles of validation outlined above apply. Ideally, the HPLC system used should meet the basic criteria specified in the liquid chromatography Pharmacopoeias (3–5). 1. It is important to remember that the main objective of validation of an analytical procedure is to demonstrate that the procedure is suitable for its intended purpose. 2. All of the analytical data collected during a validation and any calculations made should be discussed in the resulting validation document as appropriate. 3. Well-characterized reference materials, with documented purity, should be used throughout a validation study. 4. The experimental work should be designed so that the appropriate validation characteristics can be considered simultaneously to provide a sound, overall knowledge of the capabilities of the analytical procedure. 5. Analyst information, performance validation, and/or calibration certificates for the equipment used should also be recorded and stored along with the raw data. All reagents should be approved for use prior to any validation study that is to be included in a regulatory authority application and their details (supplier, catalog number, lot number and expiry dates) recorded. 6. A Hewlett-Packard HP 1100 or HP 1200 with quaternary switching pump is ideal. Reagents of the highest quality should be used during these analyses. 7. If validation is being carried out for a protein or amino acid mixture that contains unusual amino acids or amino acids that are not present in the standard mixture, the unusual amino acid should be purchased, accurately weighed, and added to the standard mixture. If the amino acid is not commercially available, the amino acid can either be synthesized, rigorously characterized, accurately weighed, and added to the standard mixture or quantitated relative to a similar amino acid. 8. Ideally prior to hydrolysis peptide, protein and conjugate samples should be salt-, amine-, and detergent-free, because these may interfere with hydrolysis or subsequent chromatography. 9. Inter-amino acid bonds differ in their susceptibility to acid hydrolysis. For example, longer acid hydrolyses are more likely to yield accurate results for hydrophobic amino acid residues, such as Valine, Isoleucine, and Phenylalanine. Other factors also have to be taken into account when utilizing amino acid analysis data. Asparagine and Glutamine are converted to their corresponding acids, Aspartic acid and Glutamic acid, and therefore cannot be directly quantified. Losses of Serine, Threonine, and to an extent Tyrosine, can be expected during acid hydrolysis through hydrolytic breakdown. More accurate estimates of Serine and Threonine can often be determined

Validation of Amino Acid Analysis Methods

10. 11.

12. 13. 14.

15.

16.

17.

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either by assuming 10% and 5% losses, respectively, over 24 h at 110°C, or by performing a series of hydrolyses for various times and extrapolating the amounts of these amino acids detected back to zero time. Determination of Cysteine is also problematic when using standard hydrolysis conditions, due to oxidation. Tryptophan is completely destroyed during normal hydrolyses and is not detected. Also, the quantitation of Aspartic and Glutamic acid, in precolumn derivatization methods, can also be compromised by the presence of buffer salts. Proteins are routinely hydrolyzed at 110°C for 24 h. Higher recoveries of more hydrophobic amino acids from membrane proteins can be achieved using higher-tempera-ture hydrolyses for example 160°C for 6 h. This matter is discussed further in Chapter 10. Addition of coupling buffer and subsequent lyophilization ensures neutralization of the hydrolysed sample and the amino acid standard mixture. The addition of UHQ water to the derivatized amino acids and subsequent lyophilization ensures optimal removal of derivatizing reagents and prevents variable chromatography. The column routinely needs to be flushed for 30 min with solvent B to ensure that the column and UV lamp have equilibrated. Equilibration at the initial conditions for amino acid analysis normally requires 5–10 min. The system can be considered suitable if the retention time variance is ±3.0% and the relative standard deviation in peak areas for the six runs is < 5%. Peak tailing (T) should be less than or equal to two and Resolution (R) should be greater than one. If the system fails a suitability test, the liquid chromatography system being used including hardware (pump, UV detector, switching valves, column, etc.) and solvents should be fully examined, for instance for leaks or partial blockages. Columns have a finite lifetime. For instance the type of RP-HPLC column used in the system described in Subheading 2.1.1. would normally be expected to allow 150–300 sample injections before peak broadening and loss of resolution result in excessively poor performance. Validation of amino acid analysis for a particular sample should only be contemplated following optimization of the method or methods to be employed. In general the optimized method, the sample formulation and, of course, amino acid sequence for each peptide, protein, and conjugate will differ. Therefore, amino acid analysis should be validated for each product rather than using a generic validated method for analysis of various products. Following validation the system suitability test should ideally be performed prior to each analysis of peptide, protein, or conjugate, providing that the

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data obtained passes the criteria set the system is deemed to be suitable for analysis. 18. Precision across different instruments need only be assessed if more than one instrument is to be used for analysis of the sample of interest. References 1. The European Agency for the Evaluation of Medicinal Products Human Medicines Evaluation Unit ICH Topic Q2(R1) Validation of Analytical Procedures: Text and Methodology (CPMP/ICH/381/95) (2005). 2. The European Agency for the Evaluation of Medicinal Products Human Medicines Evaluation Unit ICH Topic Q6B Specifications: Test Procedures and Acceptance Criteria for Biotechnological/Biological Products (CPMP/ICH/ 365/96) (1999). 3. High Pressure Liquid Chromatography USP (1995) Ninth Supplement, USP-NF Section 621 pp. 4648–1805. 4. Liquid Chromatography BP (1993) A100 Appendix IIIc – A102 Appendix IIIe. 5. Liquid Chromatography Ph. Eur (1997) 2.2.29, pp. 32–34.

110 Molecular Weight Estimation for Native Proteins Using High-Performance Size-Exclusion Chromatography G. Brent Irvine

1. Introduction The chromatographic separation of proteins from small molecules on the basis of size was first described by Porath and Flodin, who called the process “gel filtration” (1). Moore applied a similar principle to the separation of polymers on crosslinked polystyrene gels in organic solvents, but named this “gel permeation chromatography” (2). Both terms came to be used by manufacturers of such supports for the separation of proteins, leading to some confusion. The term size-exclusion chromatography is more descriptive of the principle on which separation is based and has largely replaced the older names, although the expression “gel filtration” is still commonly used by biochemists to describe separation of proteins in aqueous mobile phases. Comprehensive reviews that cover both practical and theoretical aspects of the technique are available, with emphasis on both proteins (3,4) and peptides (5), and there is also much useful information in a technical booklet published by a leading manufacturer (6). Conventional liquid chromatography is carried out on soft gels, with flow controlled by peristaltic pumps and run times on the order of several hours. These gels are too compressible for use in high-performance liquid chromatography (HPLC), but improvements in technology have led to the introduction of new supports with decreased particle size (5–13 µm) and improved rigidity. This technique is usually called high-performance size-exclusion chromatography (HPSEC), but is sometimes referred to as size-exclusion high-performance liquid chromatography (SE-HPLC). Columns (about 1 × 30 cm) are sold prepacked with the support, and typically have many thousands of theoretical plates. They

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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can be operated at flow rates of about 1 mL/min, giving run times of about 12 min, 10–100 times faster than conventional chromatography on soft gels. A medium pressure high-performance system called fast protein liquid chromatography (FPLC) that avoids the use of stainless steel components was developed by one major supplier (Amersham Pharmacia Biotech, now GE Healthcare Life Sciences). Prepacked columns of Superose and Superdex developed for this system can also be operated on ordinary HPLC systems. As well as being a standard chromatographic mode for desalting and purifying proteins, size-exclusion chromatography can be used for estimation of molecular weights. This is true only for ideal size-exclusion chromatography, in which the support does not interact with solute molecules (see Note 1). Because the physical basis for discrimination in size-exclusion chromatography is size, one would expect that there would be some parameter related to the dimensions and shape of the solute molecule that would determine its Kd value (see Note 2). All molecules with the same value for this parameter should have identical Kd values on a size-exclusion chromatography column. Dimensional parameters that have been suggested for such a “universal calibration” include two terms related to the hydrodynamic properties of proteins, namely Stokes radius (RS) and viscosity radius (Rh). Current evidence appears to indicate that Rh is more closely correlated to size-exclusion chromatography behavior than is RS (7). However, even this parameter is not suitable for a universal calibration that includes globular proteins, random coils, and rods (8). Hence calibration curves prepared with globular proteins as standards cannot be used for the assignment of molecular weights to proteins with different shapes, such as the rodlike protein myosin. Several mathematical models that relate size-exclusion chromatography to the geometries of protein solutes and support pores have been proposed (see Note 3). However, in practical terms, for polymers of the same shape, plots of log molecular weight against Kd have been found to give straight lines within the range 0.1 < Kd < 0.9 (see Note 4). It has been found that the most reliable measurements of molecular weight by HPSEC are obtained under denaturing conditions, when all proteins have the same random coil structure. Disulfide bonds must be reduced, usually with dithiothreitol, in a buffer that destroys secondary and tertiary structure. Buffers containing guanidine hydrochloride (9,10) or sodium dodecyl sulfate (SDS) (11) have been used for this purpose. However, the use of denaturants has many drawbacks, which are described in Note 5. In any case, polyacrylamide gel electrophoresis in SDS-containing buffers is widely used for determining the molecular weights of protein subunits. This technique can also accommodate multiple samples in the same run, although each run takes longer than for HPSEC. More recently SEC has been used in combination with other more absolute methods for molecular weight determination, particularly mass spectrometry (12) and multiangle laser light scattering (13).

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The method described in this chapter is for the estimation of molecular weights of native proteins. The abilities to measure bioactivity, and to recover native protein in high yield, make this an important method, even though a number of very basic, acidic, or hydrophobic proteins will undergo non-ideal size exclusion under these conditions. 2. Materials 2.1. Apparatus An HPLC system for isocratic elution is required. This comprises a pump, an injector, a size-exclusion column and a guard column (see Note 6), a UVdetector, and a data recorder. There are many size-exclusion columns based on surface-modified silica on the market. The results described here were obtained using a Zorbax Bioseries GF-250 column (0.94 × 25 cm), of particle size 4 µm, sold by Agilent Technologies. This column can withstand very high back pressures (up to 380 Bar or 5500 psi) and can be run at flow rates up to 2 mL/min with little loss of resolution (14). It has a molecular weight exclusion limit for globular proteins of several hundred thousand. The exclusion limit for the related GF-450 column is about a million. Other suitable columns include TSK-GEL SW columns (Tosoh Biosep), and Superdex and Superose columns (Amersham Pharmacia Biotech). 2.2. Chemicals 1. 0.2 M Disodium hydrogen orthophosphate (Na2HPO4). 2. 0.2 M Sodium dihydrogen orthophosphate (NaH2PO4). 3. 0.2 M Sodium phosphate buffer, pH 7.0: Mix 610 mL of 0.2 M Na2HPO4 with 390 mL of 0.2 M NaH2PO4. Filter through a 0.22-µm filter (Millipore type GV). 4. Solutions of standard proteins: Dissolve each protein in 0.2 M sodium phosphate buffer, pH 7.0 at a concentration of about 0.5 mg/mL. Filter the samples through a 0.45-µm filter (Millipore type HV). Proteins suitable for use as standards are listed in Table 1 (see Note 7). 5. Blue dextran, average molecular weight 2,000,000 (Sigma), 1 mg/mL in 0.2 M sodium phosphate buffer, pH 7.0. Filter through a 0.45-µm filter (Millipore type HV). 6. Glycine, 10 mg/mL in 0.2 M sodium phosphate buffer, pH 7.0. Filter through a 0.45-µm filter (Millipore type HV).

3. Method 1. Allow the column to equilibrate in the 0.2 M sodium phosphate buffer, pH 7.0, at a flow rate of 1 mL/min (see Note 8) until the absorbance at 214 nm is constant. 2. Inject a solution (20 µL) of a very large molecule, such as blue dextran (see Note 9) to determine V0. Repeat with 20 µL of water (negative absorbance peak) or a solution of a small molecule, such as glycine, to determine Vt.

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Irvine Table 1 Protein Standards Protein

Molecular weight

Thyroglobulin Apoferritin β-Amylase Immunoglobulin G Alcohol dehydrogenase Bovine serum albumin Ovalbumin β-Lactoglobulin Carbonic anhydrase Trypsinogen Soybean trypsin inhibitor Myoglobin Ribonuclease A Insulin Glucagon

669,000 443,000 200,000 160,000 150,000 66,000 42,700 36,800 29,000 24,000 20,100 16,900 13,690 5900 3550

All proteins listed were obtained from Sigma, Poole, England.

3. Inject a solution (20 µL) of one of the standard proteins and determine its elution volume, Ve, from the time at which the absorbance peak is at a maximum. Repeat this procedure until all the standards have been injected. A chromatogram showing the separation of seven solutes during a single run on a Zorbax Bio-series GF-250 column is shown in Fig. 1. Calculate Kd (see Note 2) for each protein and plot Kd against log molecular weight. A typical plot is shown in Fig. 2. 4. Inject a solution (20 µL) of the protein of unknown molecular weight and measure its elution volume, Ve, from the absorbance profile. If the sample contains more than one protein, and the peaks cannot be assigned with certainty, collect fractions and assay each fraction for the relevant activity. 5. Calculate Kd for the unknown protein and use the calibration plot to obtain an estimate of its molecular weight.

4. Notes 1. Silica to which a hydrophilic phase such as a diol has been bonded still contains underivatized silanol groups. Above pH 3.0 these are largely anionic and will interact with ionic solutes, leading to non-ideal sizeexclusion chromatography. Depending on the value of its isoelectric point, a protein can be cationic or anionic at pH 7.0. Proteins that are positively charged will undergo ion exchange, causing them to be retarded. Conversely, anionic proteins will experience electrostatic repulsion from the pores, referred to as “ion exclusion,” and will be eluted earlier than

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Fig. 1. Separation of a mixture of seven solutes on a Zorbax Bio-series GF-250 column. Twenty microliters of a mixture containing about 1.5 µg of each protein was injected. The solutes were, in order of elution, thyroglobulin, alcohol dehydrogenase, ovalbumin, myoglobin, insulin, glucagon, and sodium azide. The number beside each peak is the elution time in minutes. The absorbance of the highest peak, insulin, was 0.105. The equipment was a Model 501 Pump, a 441 Absorbance Detector operating at 214 nm, a 746 Data Module, all from Waters (Millipore, Milford, MA, USA) and a Rheodyne model 7125 injector (Rheodyne, Cotati, CA, USA) with 20-µL loop. The column was a Zorbax Bio-Series GF-250 with guard column (Aligent Technologies). The flow rate was 1 mL/min and the chart speed was 1 cm/ min. The attenuation setting on the Data Module was 128.

expected on the basis of size alone. When size exclusion chromatography is carried out at a low pH, the opposite behavior is found, with highly cationic proteins being eluted early and anionic ones being retarded. To explain this behavior, it has been suggested that at pH 2.0 the column may have a net positive charge (15). To reduce ionic interactions it is necessary to use a mobile phase of high ionic strength. On the other hand, as ionic strength increases, this promotes the formation of hydrophobic interactions. To minimize both ionic and hydrophobic interactions, the mobile phase should have an ionic strength between 0.2 and 0.5 M (16). 2. The support used in size-exclusion chromatography consists of particles containing pores. The molecular size of a solute molecule determines the degree to which it can penetrate these pores. Molecules that are wholly excluded from the packing emerge from the column first, at the void volume, V0. This represents the volume in the interstitial space (outside the

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Fig. 2. Plot of Kd vs log molecular weight for those proteins listed in Table 1 with Kd values between 0.1 and 0.9 (i.e., all except glucagon and thyroglobulin). Chromatography was carried out as described in the caption to Fig. 1. V0 was determined to be 6.76 mL from the elution peak of blue dextran. Vt was determined from the elution peak of glycine and from the negative peak given by injecting water, both of which gave a value of 11.99 mL. The regression line y = −0.4234x + 2.4378, r2 = 0.9657 was computed using all the points shown.

support particles) and is determined by chromatography of very large molecules, such as blue dextran or DNA. Molecules that can enter the pores freely have full access to an additional space, the internal pore volume, Vi. Such molecules emerge at Vt, the total volume available to the mobile phase, which can be determined from the elution volume of small molecules. Hence Vt = V0 + Vi. A solute molecule that is partially restricted from the pores will emerge with elution volume, Ve, between the two extremes, V0 and Vt. The distribution coefficient, Kd, for such a molecule represents the fraction of Vi available to it for diffusion. Hence Ve = V0 + KdVi and Kd = (Ve − V0)/Vi = (Ve − V0)/(Vt − V0) 3. These models give rise to various plots that should result in a linear relationship between a function of solute radius, usually RS, and some sizeexclusion chromatography parameter, usually Kd (17). However, none has proven totally satisfactory. Recently it has been suggested that size-exclusion chromatography does not require pores at all, but rather that Kd can be calculated from a thermodynamic model for the free energy of mixing of the solute and the gel phase (18). 4. Manufacturers’ catalogs often show plots of Kd against log molecular weight, M. For most globular proteins, this plot is a sigmoidal curve that is approximately linear in the middle section, where

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Kd = a – b log M where a is the intercept on the ordinal axis, and b is the slope. From such plots one can estimate fractionation range (the working range lies in the linear portion, between about 0.9 > Kd > 0.1) and the selectivity. This latter parameter depends on the slope of the plot and is a function of the pore size distribution. A support with average pore size distribution in a narrow band gives high selectivity (large value of b) but a restricted separation range (the larger the value of b, the lower the range of values for M). If no estimate is available for the size of the protein under investigation it is better to use a support with a broad fractionation range. A support of higher selectivity in a more restricted fractionation range can then be used later to give a more precise value for solute size. 5. Problems arising when size-exclusion chromatography is carried out under denaturing conditions include: a. For a particular column, the molecular weight range in which separation occurs is reduced. This is because the radius of gyration, and hence the hydrodynamic size, of a molecule increases when it changes from a sphere to a random coil. For example, the separation range of a TSK G3000SW column operating with denatured proteins is 2000– 70,000, compared to 10,000–500,000 for native proteins (10,19). Of course this may actually be an advantage when working with small proteins or peptides. b. Proteins are broken down into their constituent subunits and polypeptide chains, so that the molecular weight of the intact protein is not obtained. c. Bioactivity is usually destroyed or reduced and it is not usually possible to monitor enzyme activity. This can be a serious disadvantage when trying to identify a protein in an impure preparation. d. The denaturants usually absorb light in the far UV range, so that monitoring the absorbance in the most sensitive region for proteins (200– 220 nm) is no longer possible. e. Manufacturers often advise that once a column has been exposed to a mobile phase containing denaturants it should be dedicated to applications using that mobile phase, as the properties of the column may be irreversibly altered. In addition, the denaturant, especially if it is SDS, may be difficult to remove completely. f. Because these mobile phases have high viscosities, flow rates may have to be reduced to avoid high back pressures. g. High concentrations of salts, especially those containing halide ions, can adversely affect pumps and stainless steel.

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6. Most manufacturers sell guard columns appropriate for use with their sizeexclusion columns. To protect the expensive size-exclusion column it is strongly recommended that a guard column be used. 7. Even in the presence of high ionic strength buffers several proteins show nonideal behavior and are thus unsuitable as standards. For example, the basic proteins cytochrome c (pI ≅10) and lysozyme (pI ≅11) have Kd values > 1.0, under the conditions described for Fig. 1, because ion-exchange interactions are not totally suppressed. On the other hand, the very acidic protein pepsin (pI ≅1) emerges earlier than expected on the basis of size, because of ion exclusion. One should be aware that such behavior might also occur when interpreting results for proteins of unknown pI. 8. Columns are often stored in ethanol–water mixtures or in 0.02% sodium azide to prevent bacterial growth. Caution: Sodium azide is believed to be a mutagen. It should be handled with care (see suppliers’ safety advice) and measures taken to avoid contact with solutions. It can also lead to explosions when disposed of via lead pipes. Solutions should be collected in waste bottles. When changing mobile phase some manufacturers recommend that the flow rate should not be greater than half the maximum flow rate. 9. Although V0 is most commonly measured using blue dextran, Himmel and Squire suggested that it is not a suitable marker for the TSK G3000SW column, owing to tailing under nondenaturing conditions, and measured V0 using glutamic dehydrogenase from bovine liver (Sigma Type II; mol. wt 998,000) (20). Calf thymus DNA is also a commonly used marker for V0. References 1. Porath, J. and Flodin, P. (1959) Gel filtration: a method for desalting and group separation. Nature 183, 1657–1659. 2. Moore, J. C. (1964) Gel permeation chromatography. I. A new method for molecular weight distribution of high polymers. J. Polymer Sci. A2, 835–843. 3. Hagel, L. (1989) Gel filtration, in Protein Purification: Principles, High Resolution Methods and Applications (Janson, J. C., and Ryden, L., eds.) VCH, New York, pp. 63–106. 4. Baker, J. O., Adney, W. S., Himmel, M. E., and Chen, M. (2002) in Handbook of Size Exclusion Chromatogaphy and Related Techniques, 2nd ed. (Wu, C. S., ed). Marcel Dekker, New York, pp. 439–462. 5. Irvine, G.B. (2003) High-performance size-exclusion chromatography of peptides. J. Biochem. Biophys. Meth. 56, 233–242. 6. Gel Filtration: Principles and Methods (2002) (edition A1) 18-1022-18, Amersham Biosciences, Uppsala. 7. Potschka, M. (1987) Universal calibration of gel permeation chromatography and determination of molecular shape in solution. Analyt. Biochem. 162, 47–64.

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8. Dubin, P. L. and Principi, J. M. (1989) Failure of universal calibration for size exclusion chromatography of rodlike macromolecules versus random coils and globular proteins. Macromolecules 22, 1891–1896. 9. Ui, N. (1979) Rapid estimation of molecular weights of protein polypeptide chains using high-pressure liquid chromatography in 6 M guanidine hydrochloride. Analyt. Biochem. 97, 65–71. 10. Kato, Y., Komiya, K., Sasaki, H., and Hashimoto, T. (1980) High-speed gel filtration of proteins in 6 M guanidine hydrochloride on TSK-GEL SW columns. J. Chromatogr. 193, 458–463. 11. Josic, D., Baumann, H., and Reutter, W. (1984) Size-exclusion high-performance liquid chromatography and sodium dodecyl sulphate-polyacrylamide gel electrophoresis of proteins: a comparison. Analyt. Biochem. 142, 473–479. 12. Goetz, H., Kuschel, M., Wulff, T., Sauber, C., Miller, C., Fisher, S., and Woodward, C. (2004) Comparison of selected analytical techniques for protein sizing, quantitation and molecular weight determination. J. Biochem. Biophys. Meth. 60, 281–293. 13. Ye, H. (2006) Simultaneous determination of protein aggregation, degradation, and absolute molecular weight by size exclusion chromatography-multiangle laser light scattering. Anal. Biochem. 356, 76–85. 14. Anspach, B., Gierlich, H. U., and Unger, K. K. (1988) Comparative study of Zorbax Bio Series GF 250 and GF 450 and TSK-GEL 3000 SW and SWXL columns in size-exclusion chromatography of proteins. J. Chromatogr. 443, 45–54. 15. Irvine, G. B. (1987) High-performance size-exclusion chromatography of polypeptides on a TSK G2000SW column in acidic mobile phases. J. Chromatogr. 404, 215–222. 16. Regnier, F. E. (1983) High performance liquid chromatography of proteins. Meth. Enzymol. 91, 137–192. 17. Ackers, G. K. (1970) Analytical gel chromatography of proteins. Adv. Protein Chem. 24, 343–446. 18. Brooks, D. E., Haynes, C. A., Hritcu, D., Steels, B. M., and Muller, W. (2000) Size exclusion chromatography does not require pores. Proc. Natl. Acad. Sci. USA 97, 7064–7067. 19. Kato, Y., Komiya, K., Sasaki, H., and Hashimoto, T. (1980) Separation range and separation efficiency in high-speed gel filtration on TSK-GEL SW columns. J. Chromatogr. 190, 297–303. 20. Himmel, M. E. and Squire, P. G. (1981) High pressure gel permeation chromatography of native proteins on TSK-SW columns. Int. J. Peptide Protein Res. 17, 365–373.

111 Detection of Disulfide-Linked Peptides by HPLC Alastair Aitken and Michèle Learmonth

1. Introduction Classical techniques for determining disulfide bond patterns usually require the fragmentation of proteins into peptides under low pH conditions to prevent disulfide exchange. Pepsin or cyanogen bromide are particularly useful (see Chapters 101 and 94 respectively). Diagonal techniques to identify disulphide-linked peptides were developed by Brown and Hartley (see Chapter 113). A modern micromethod employing reversephase high-performance liquid chromatography (HPLC) is described here. 2. Materials 1. 2. 3. 4. 5. 6. 7. 8.

1 M Dithiothreitol (DTT, good quality, e.g., Calbiochem). 100 mM Tris-HCl, pH 8.5. 4-Vinylpyridine. 95% Ethanol. Isopropanol. 1 M triethylamine-acetic acid, pH 10.0. Tri-n-butyl-phosphine (1% in isopropanol). HPLC system such as Dionex UltiMate Micro HPLC systems for LC and LC/MS. 9. Reverse phase HPLC columns such as Vydac C4, C8, C18 microbore (1.0 mm I.D. columns) for the purification of low picomole amounts (< 5 pmol) of peptides. (see Note 1).

3. Method (see refs. 1 and 2) 1. Alkylate the protein (1–10 mg in 20–50 mL of buffer) without reduction to prevent possible disulfide exchange by dissolving in 100 mM Tris-HCl, pH 8.5, and adding 1 µL of 4-vinylpyridine (see Note 2). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2. Incubate for 1 h at room temperature and desalt by HPLC or precipitate with 95% ice-cold ethanol followed by bench centrifugation. 3. The pellet obtained after the latter treatment may be difficult to redissolve and may require addition of 10-fold concentrated acid (HCl, formic, or acetic acid) before digestion at low pH. It may be sufficient to resuspend the pellet with acid using a sonic bath if necessary, then commence the digestion. Vortex-mix the suspension during the initial period until the solution clarifies. 4. Fragment the protein under conditions of low pH (see Note 3) and subject the peptides from half the digest to reverse-phase HPLC. Vydac C4, C8, or C18 columns give particularly good resolution depending on the size range of fragments produced.

Typical separation conditions are;- column equilibrated with 0.1% (v/v) aqueous trifluoro acetic acid (TFA), elution with an acetonitrile–0.1% TFA gradient. A combination of different cleavages, both chemical and enzymatic, may be required if peptide fragments of interest remain large after one digestion method. 5. To the other half of the digest (dried and resuspended in 10 µL of isopropanol) add 5 µL of 1 M triethylamine-acetic acid pH 10.0; 5 µL of tri-n-butyl-phosphine (1% in isopropanol) and 5 µL of 4-vinylpyridine. Incubate for 30 min at 37 °C, and dry in vacuo, resuspending in 30 µL of isopropanol twice. This procedure cleaves the disulfides and modifies the resultant –SH groups. 6. Run the reduced and alkylated sample on the same column, under identical conditions on reverse-phase HPLC. Cysteine-linked peptides are identified by the differences between elution of peaks from reduced and unreduced samples. 7. Collection of the alkylated peptides (which can be identified by rechromatography on reverse-phase HPLC with detection at 254 nm) and a combination of sequence analysis and mass spectrometry (see Note 1 and Chapter 112) will allow disulfide assignments to be made.

4. Notes 1. If the HPLC separation is combined with mass spectrometric characterization, the level of TFA required to produce sharp peaks and good resolution of peptide (approx 0.1% v/v) results in almost or complete suppression of signal. This does not permit true on-line HPLC-MS as the concentration of TFA in the eluted peptide must first be drastically reduced. However, the “low TFA,” 218MS54, reverse-phase HPLC columns from Vydac (300 Å pore size) are available in C4 and two forms of C18 chemistries. They are also supplied in 1 mm diameter columns that are ideal for low levels of sample eluted in minimal volume. We have used as little as 0.005% TFA without major loss of resolution and have observed minimal signal loss. There may be a difference in selectivity compared to “classical” reversephase columns; for example, we have observed phosphopeptides eluting approx. 1% acetonitrile later than their unphosphorylated counterparts (the

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opposite to that conventionally seen). This is not a problem, but it is something of which one should be aware, and could be turned to advantage. 2. The iodoacetic acid used must be colorless. A yellow color indicates the presence of iodine; this will rapidly oxidize thiol groups, preventing alkylation and may also modify tyrosine residues. It is possible to recrystallize from hexane. Reductive alkylation may also be carried out using iodo[14C]-acetic acid or iodoacetamide (see Chapter 82). The radiolabelled material should be diluted to the desired specific activity before use with carrier iodoacetic acid or iodoacetamide to ensure an excess of this reagent over total thiol groups. 3. Fragmentation of proteins into peptides under low pH conditions to prevent disulfide exchange is important. Pepsin, Glu-C, or cyanogen bromide are particularly useful (see Chapters 99 and 94 respectively). Typical conditions for pepsin are 25 °C for 1–2 h at pH 2.0–3.0 (10 mM HCl, 5% acetic or formic acid) with an enzyme:substrate ratio of about 1:50. Endoproteinase Glu-C has a pH optimum at 4 as well as an optimum at pH 8.0. Digestion at the acid pH (typical conditions are 37 °C overnight in ammonium acetate at pH 4.0 with an enzyme/substrate ratio of about 1:50) will also help minimize disulfide exchange. CNBr digestion in guanidinium 6 M HCl/ 0.1–0.2 M HCl may be more suitable acid medium due to the inherent redox potential of formic acid which is the most commonly used protein solvent. When analyzing proteins that contain multiple disulfide bonds it may be appropriate to carry out an initial chemical cleavage (CNBr is particularly useful), followed by a suitable proteolytic digestion. The initial acid chemical treatment will cause sufficient denaturation and unfolding as well as peptide bond cleavage to assist the complete digestion by the protease. If a protein has two adjacent cysteine residues this peptide bond will not be readily cleaved by specific endopeptidases. For example, this problem was overcome during mass spectrometric analysis of the disulfide bonds in insulin by using a combination of an acid proteinase (pepsin) and carboxypeptidase A as well as Edman degradation (3). References 1. Friedman, M., Zahnley, J. C., and Wagner, J.R. (1980) Estimation of the disulfide content of trypsin inhibitors as S-b-(2-pyridylethyl)-L-cysteine. Analyt. Biochem. 106, 27–34. 2. Amons, R. (1987) Vapor-phase modification of sulfhydryl groups in proteins. FEBS Lett. 212, 68–72. 3. Toren, P., Smith, D., Chance, R., and Hoffman, J. (1988). Determination of Interchain Crosslinkages in Insulin B-Chain Dimers by Fast Atom Bombardment Mass Spectrometry. Analyt. Biochem. 169, 287–299.

112 Detection of Disulfide-Linked Peptides by Mass Spectrometry Alastair Aitken and Michèle Learmonth

1. Introduction Mass spectrometry is playing a rapidly increasing role in protein chemistry and sequencing and is particularly useful in determining sites of co- and posttranslational modification (1,2), and application in locating disulfide bonds is no exception. This technique can of course readily analyze peptide mixtures; therefore it is not always necessary to isolate the constituent peptides. However, a cleanup step to remove interfering compounds such as salt and detergent may be necessary. Thus can be achieved using matrices such as 10-µm porous resins slurry-packed into columns 0.25 mm diameter. Polypeptides can be separated on stepwise gradients of 5–75% acetonitrile in 0.1% formic or acetic acid (3). On-line electrospray mass spectrometry (ES-MS) coupled to capillary electrophoresis or high-performance liquid chromatography (HPLC) has proved particularly valuable in the identification of modified peptides. If HPLC separation on conventional columns is attempted on-line with mass spectrometry, the level of trifluoroacetic acid (TFA) (0.1%) required to produce sharp peaks and good resolution of peptides results in almost or complete suppression of signal. In this case it is recommended to use the “low TFA,” 218MS54, reverse-phase HPLC columns from Vydac (300 Å pore size) which can be used with as little as 0.005% TFA without major loss of resolution and minimal signal loss (see further details in Chapter 111). Sequence information is readily obtained using triple quadrupole tandem mass spectrometry after collision-induced dissocation (4). Ion trap mass spectrometry technology (called LCQ) is now well established which also permits sequence information to be readily obtained. Not only can MS-MS analysis be

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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carried out, but owing to the high efficiency of each stage, further fragmentation of selected ions may be carried out to MS.” The charge state of peptide ions is readily determined by a “zoom-scan” technique that resolves the isotopic envelopes of multiply charged peptide ions. The instrument still allows accurate molecular mass determination to 100,000 Da at 0.01% mass accuracy. The recent development of the Orbitrap mass spectrometer (5) allows very high resolution and sensitivity. Selective detection of modified peptides is possible on ES-MS. For example, phosphopeptides can be identified from the production of phosphate-specific fragment ions of 63 Da (PO) and 79 Da (PO) by collision-induced dissociation during negative ion HPLC–ES-MS. This technique of selective detection of posttranslational modifications through collision-induced formation of lowmass fragment ions that serve as characteristic and selective markers for the modification of interest has been extended to identify other groups such as glycosylation, sulfation and acylation (6). 2. Materials Materials for proteolytic and chemical cleavage of proteins are described in Chapters 94–99. 3. Method 3.1. Detection of Disulfide-Linked Peptides by Mass Spectrometry 1. Peptides generated by any suitable proteolytic or chemical method that minimizes disulfide exchange (i.e., acid pH, see Note 1). Partial acid hydrolysis, although nonspecific, has been successfully used in a number of instances. The peptides are then analyzed by a variety of mass spectrometry techniques (7). The use of thiol and related compounds should be avoided for obvious reasons. Despite this, it is possible that disulfide bonds will be partially reduced during the analysis and peaks corresponding to the individual components of the disulfide-linked peptides will be observed. Control samples with the above reagents are essential to avoid misleading results (see Note 2). 2. The peptide mixture is incubated with reducing agents, such as mercaptoethanol and dithiothreitol (DTT), and reanalyzed as before. Peptides that were disulfide linked disappear from the spectrum and reappear at the appropriate positions for the individual components. For example, in the positive ion mode the mass (M) of disulfide-linked peptides (of individual masses A and B) will be detected as the pseudomolecular ion at (M+H)+, and after reduction this will be replaced by two additional peaks for each disulfide bond in the polypeptide at masses (A+H)+ and (B+H)+. Remember that A + B = M + 2, as reduction of the disulfide bond will be accompanied by a consistent increase in mass due to the conversion of cystine to two cysteine residues, that is, −S-S → −SH + HS- and peptides containing an intramolecular disulfide bond will appear at 2 amu higher. Such peptides, if they

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are in the reduced state can normally be readily reoxidized to form intramolecular disulfide bonds by bubbling a stream of air through a solution of the peptide for a few minutes (see Note 3). 3. Computer programs are readily available on the Internet that are supplied with the mass spectrometer software package and will predict the cleavage position of any particular proteinase or chemical reagent. Simple knowledge of the mass of the fragment will, in most instances, give unequivocal answers as to which segments of the polypeptide chain are disulfide linked. If necessary, one cycle of Edman degradation can be carried out on the peptide mixture and the mass spectroometry analysis repeated. The shift in mass(es) that correlates with loss of specific residues will confirm the assignment.

4. Notes 1. Fragmentation of proteins into peptides under low pH conditions is important (e.g., with pepsin, Glu-C, or cyanogen bromide; see Chapter 99 and 94 respectively) to prevent disulfide exchange. Analysis can also be carried out by electrospray mass spectrometers (ES-MS), which will give an accurate molecular mass up to 80–100,000 Da (and in favorable cases up to 150,000 Da). The increased mass of 2 Da for each disulfide bond will be too small to obtain an accurate estimate for a polypeptide of mass Ca 10,000 (accuracy obtainable is >0.01%). There has been a recent marked increase in resolution obtained with both electrospray mass spectrometers and laser desorption time-of-flight mass spectrometers that could now permit a meaningful analysis. On the other hand, oxidation with performic acid will cause a mass increase of 48 Da for each cysteine and 49 Da for each half-cystine residue. Note that Met and Trp will also be oxidized. Met sulfoxide, the result of incomplete oxidation and that is often found after gel electrophoresis, has a mass identical to that of Phe. Met sulfone is identical in mass to Tyr. Orbitrap mass spectrometers have ppm accuracy that is well within the necessary range of resolution even for medium to large proteins (8). 2. Mass analysis by ES-MS (3,9) and matrix-assisted laser desorption mass spectrometry with time-of-flight detection (MALDI-TOF) (10) is affected, seriously in some cases, by the presence of particular salts, buffers, and detergents. In some cases, using nonionic saccharide detergents such as n-dodecyl-β-D-glucopyranoside, improvements in signal-to-noise ratios of peptides and proteins were observed (10). The effect on ES-MS sensitivity of different buffer salts, detergents, and tolerance to acid type may vary widely with the instrument and particularly with the ionization source. Critical micelle concentration is not a good predictor of how well a surfactant will perform (9). 3. The difference of 2 Da may allow satisfactory estimation of the number of intramolecular disulfide bonds by mass spectrometry. If necessary a larger

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mass difference may be generated by oxidation with performic acid (11), and see Chapter 83). This will cause a mass increase of 48 Da for each cysteine and 49 Da for each half-cystine residue. (Remember that Met and Trp will also be oxidized.) (12) or by exopeptidase digestion from the N- and C-termini. This essentially a subtractive technique (one looks at the mass of the remaining fragment after each cycle). For example, when a phosphoserine residue is encountered, a loss of 167.1 Da is observed in place of 87.1 for a serine residue. This technique therefore avoids one of the major problems of analyzing posttranslational modifications. Although the majority of modifications are stable during the Edman chemistry, O- or S-linked esters, for example (which are very numerous), may be lost by β-elimination (e.g., O-phosphate) during the cleavage step to form the anilinothioazolidone or undergo O- (or S- in the case of palmitoylated cysteine) to N-acyl shifts which block further Edman degradation. Exopeptidase digestion may be difficult and the rate of release of amino acid may vary greatly. References 1. Costello, C. E. (1999) Bioanalytic applications of mass spectrometry. Curr. Opin. Biotechnol. 10, 22–28. 2. Burlingame, A. L., Boyd, R. K., and Gaskell, S. J. (1998) Mass spectrometry. Analyt. Chem. 70, 647R–716R. 3. Aitken, A., Howell, S., Jones, D., Madrazo, J., and Patel, Y. (1995) 14-3-3 α and δ are the phosphorylated forms of Raf-activating 14-3-3 β and ζ. In vivo stoichiometric phosphorylation in brain at a Ser-Pro-Glu-Lys motif. J. Biol. Chem. 270, 5706–5709. 4. Hunt, D. F., Yates, J. R., Shabanowitz, J., Winston, S., and Hauer, C. R. (1986) Protein sequencing by tandem mass spectrometry. Proc. Natl. Acad. Sci. USA 83, 6233–6237. 5. Hu, Q., Noll, R. J., Li, H., Makarov, A., Hardman, M., and Cooks R. G.(2005) The Orbitrap: a new mass spectrometer. J Mass Spectrom. 40, 430–443. 6. Bean, M. F., Annan, R. S., Hemling, M. E., Mentzer, M., Huddleston, M. J., and Carr, S. A. (1994) LC-MS methods for selective detection of posttranslational modifications in proteins, in Techniques in Protein Chemistry VI (Crabb, J. W., ed.), Academic Press, San Diego, pp. 107–116. 7. Morris, R. H. and Pucci, P. (1985). A new method for rapid assignment of S—S bridges in proteins. Biochem. Biophys. Res. Commun. 126, 1122–1128. 8. Scigelova, M. and Makarov, A.( 2006) Orbitrap mass analyzer - overview and applications in proteomics. Proteomics 6 Suppl 2, 16–21. 9. Loo, R., Dales, N., and Andrews, P. C. (1994) Surfactant effects on protein structure examined by electrospray ionisation mass spectrometry. Protein Sci. 3, 1975–1983. 10. Vorm, O., Chait, B. T., and Roepstorff, (1993) Mass spectrometry of protein samples containing detergents, in Proceedings of the 41st ASMS Conference on Mass Spectrometry and Allied Topics, pp. 621–622. 11. Sun, Y. and Smith, D. L. (1988) Identification of disulfide-containing peptides by performic acid oxidation and mass spectrometry. Analyt. Biochem. 172, 130–138.

113 Diagonal Electrophoresis for Detecting Disulfide Bridges Alastair Aitken and Michèle Learmonth

1. Introduction Methods for identifying disulfide bridges have routinely employed “diagonal” procedures using two-dimensional paper or thin-layer electrophoresis. This essentially utilizes the difference in electrophoretic mobility of peptides containing either cysteine or cystine in a disulfide link, before and after oxidation with performic acid. It was first described by Brown and Hartley (1). Peptides unaltered by the performic acid oxidation have the same mobility in both dimensions and, therefore, lie on a diagonal. After oxidation, peptides that contain cysteine or were previously covalently linked produce one or two spots off the diagonal, respectively. This method has also been adapted for HPLC methodology and is discussed in Chapter 111. First the protein has to be fragmented into suitable peptides containing individual cysteine residues. It is preferable to carry out cleavages at low pH to prevent possible disulfide bond exchange. In this respect, pepsin (active at pH 3.0) and cyanogen bromide (CNBr) are particularly useful reagents. Proteases with active-site thiols should be avoided (e.g., papain, bromelain). Before the advent of HPLC, paper electrophoresis was the most commonly used method for peptide separation (2). Laboratories with a history of involvement with protein characterization are likely to have retained the equipment, but it is no longer commercially available. Although it is possible to make a simple electrophoresis tank in house (3), thin-layer electrophoresis equipment is still commercially available, and it is advisable that this be used, owing to the safety implications. For visualization of the peptides, ninhydrin is the classical amino group stain. However, if amino acid analysis or sequencing is to be carried out, fluorescamine is the reagent of choice. From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2. Materials 2.1. Equipment 1. Electrophoresis tank. 2. Flat bed electrophoresis apparatus (e.g., Hunter thin-layer peptide-mapping apparatus (CBS Scientific, Del Mar, CA,. and VWR International Ltd, UK). 3. Whatman (Maidstone, UK) 3MM and Whatman No. 1 paper. 4. Cellulose TLC plates (Camlab [Cambridge, UK]

2.2. Reagents 1. Electrophoresis buffers (see Note 1): Commonly used volatile buffers are: pH 2.1 acetic acid/formic acid/water 15/5/80, v/v/v pH 3.5 pyridine/acetic acid/water 5/50/945 v/v/v pH 6.5 pyridine/acetic acid/water 25/1/225, v/v/v 2. Nonmiscible organic solvents: toluene, for use with pH 6.5 buffer: white spirit for use with pH 2.1 and 3.5 buffers. 3. Formic acid. Care! 4. 30% w/v Hydrogen peroxide. Care! 5. Fluorescamine: 1 mg/100 mL in acetone. 6. Marker dyes: 2% orange G, 1% acid fuschin, and 1% xylene cyanol dissolved in appropriate electrophoresis buffer. 7. 1% (v/v) Triethylamine in acetone.

3. Methods 3.1. First-Dimension Electrophoresis 3.1.1. Paper Electrophoresis 1. Dissolve the peptide digest from 0.1–0.3 µmol protein in 20–25 µL electrophoresis buffer. 2. Place the electrophoresis sheet on a clean glass sheet (use Whatman No. 1 for analytical work, Whatman 3MM for preparative work). Support the origin at which the sample is to be applied on glass rods. Where paper is used, multiple samples can be run side by side. Individual strips can then be cut out for running in the second dimension. 3. Apply the sample slowly without allowing to dry (covering an area of about 2 × 1 cm) perpendicular to the direction intended for electrophoresis. NB: for pH 6.5 buffer, apply near the center of the sheet, for acidic buffers, apply nearer the anode end. 4. Apply a small volume of marker dyes (2% orange G, 1% acid fuschin, 1% xylene cyanol, in electrophoresis buffer) on the origin and additionally in a position that will not overlap the peptides after the second dimension. 5. Once the sample is applied, wet the sheet with electrophoresis buffer slowly and uniformly on either side of the origin so that the sample concentrates in a thin line. Remove excess buffer from the rest of the sheet with blotting paper. 6. Place the wet sheet in the electrophoresis tank previously set up with electrophoresis buffer covering bottom electrode. An immiscible organic solvent (toluene

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where pH 6.5 buffer is used, white spirit for acidic buffers) used to fill the tank to the top. Upper trough filled with electrophoresis buffer. Care! 7. Electrophorese at 3 kV for about 1 h, with cooling if necessary. The progress of the electrophoresis can be monitored by the movement of the marker dyes (see Note 1). 8. Dry sheet in a well-ventilated place overnight, at room temperature, secured from a glass rod with Bulldog clips.

3.1.2. Thin-Layer Electrophoresis 1. Dissolve the peptide digest from 0.1–0.3 µmol protein in at least 10 µL electrophoresis buffer. 2. Mark the sample and dye origins on the cellulose side of a TLC plate with a cross using an extrasoft blunt-ended pencil, or on the reverse side with a permanent marker. 3. Spot the sample on the origin. This can be done using a micropipet fitted with a disposable capillary tip. To keep the spot small, apply 0.5–1 µL spots and dry between applications. 4. Apply 0.5 µL marker dye to the dye origin. 5. Set up electrophoresis apparatus with electrophoresis buffer in both buffer tanks. Prepare electrophoresis wicks from Whatman 3MM paper, wet with buffer, and place in buffer tanks. 6. Prepare a blotter from a double sheet of Whatman 3MM, with holes cut at the positions of the sample and marker origins. Wet with buffer and place over TLC plate. Ensure concentration at the origins by pressing lightly around the holes. 7. Place TLC plate in apparatus. 8. Electrophorese at 1.5 kV for about 30–60 min. 9. Dry plate in a well-ventilated place at room temperature, overnight.

3.2. Performic Acid Oxidation (see Note 2) 1. Prepare performic acid by mixing 19 mL formic acid with 1 mL 30% (w/v) hydrogen peroxide. The reaction is spontaneous. Care! 2. Place the dry electrophoresis sheet/plate in a container where it can be supported without touching the sides. 3. Place the performic acid in a shallow dish inside the container. Close the container and leave to oxidize for 2–3 h. (Note the marker dyes change from blue to green.) 4. Dry sheet thoroughly at room temperature overnight.

3.3. Second Dimension Electrophoresis 3.3.1. Paper Electrophoresis 1. To prepare for the second dimension, individual strips from the first dimension can be machine zigzag stitched onto a second sheet. The overlap of the second sheet should be carefully excised with a razor blade/scalpel. 2. Wet the sheet with electrophoresis buffer, applying the buffer along both sides of the sample, thus concentrating the peptides in a straight line. 3. Repeat electrophoresis, at right angles to the original direction. 4. Thoroughly dry sheet, as before.

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3.3.2. Thin-Layer Electrophoresis 1. Wet TLC plate with electrophoresis buffer using two sheets of prewetted Whatman 3 MM paper on either side of sample line. 2. Repeat electrophoresis at right angles to the original direction. 3. Thoroughly dry plate as before.

3.4. Visualization Peptide spots can be seen after reaction with fluorescamine. 1. The reaction should be carried out under alkaline conditions. The sheet or plate should be dipped in a solution of triethylamine (1% [v/v]) in acetone. This should be carried out at least twice if the electrophoresis buffer employed was acidic. Dry the sheet well. 2. Dip the sheet in a solution of fluorescamine in acetone (1 mg/100 mL). 3. Allow most of the acetone to evaporate. 4. View the map under a UV lamp at 300–365 nm (Care: Goggles must be worn). Peptides and amino-containing compounds fluoresce. Encircle all fluorescent spots with a soft pencil. 5. Interpretation of diagonal maps: Fig. 1 shows that peptides unaltered by the performic acid treatment have the same mobility in both dimensions and therefore lie

Fig. 1. Diagonal electrophoresis for identification and purification of peptides containing cysteine or disulfide bonds. This figure shows that peptides unaltered by the performic acid treatment (open circles) have the same mobility in both dimensions and therefore lie on a diagonal. Peptides that contain cysteine or were previously covalently linked (closed circles) produce one or two spots respectively, that lie off the diagonal after oxidation.

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on a diagonal. Peptides that contain cysteine or were previously covalently linked produce one or two spots, respectively, that lie off the diagonal after oxidation.

3.5. Elution Peptides may be eluted from the paper using, for example, 0.1 M NH3 or 25% acetic acid. Peptides may be extracted from TLC plates by scraping off the spot into an Eppendorf tube containing elution buffer. This should be vortexed for 5 min and then centrifuged for 5 min. The cellulose can then be re-extracted once or twice with the same buffer to ensure optimal recovery of peptide. 4. Notes 1. The buffer of choice for the initial analysis is pH 6.5. However, if the cysteine residues have already been blocked with iodoacetate (see Chapter 82), the pH 3.5 buffer is very useful, since peptides containing these residues lie slightly off the diagonal, being slightly more acidic in the second dimension after the performic acid oxidation. 2. The movement of the marker dyes will enable progress of the electrophoresis to be followed to ensure that the samples do not run off the end of the paper. 3. It is important to exclude halide ions rigorously, since these are readily oxidized to halogens, which will react with histidine, tyrosine, and phenylalanine residues in the protein. References 1. Brown, J. R. and Hartley, B. S. (1966) Location of disulphide bridges by diagonal paper electrophoresis Biochem. J. 101, 214–228. 2. Michl, H. (1951) Paper electrophoresis at potential differences of 50 volts per centimetre. Monatschr. Chem. 82, 489–493. 3. Creighton, T. E. (1997) Disulphide bonds between cysteine residues, in Protein Structure—a Practical Approach, 2nd ed. (Rickwood, D. and Hames B. D., eds.), IRL, Oxford, UK, pp. 155–167.

114 Estimation of Disulfide Bonds Using Ellman’s Reagent Alastair Aitken and Michèle Learmonth

1. Introduction Ellman’s reagent 5,5' -dithiobis(2-nitrobenzoic acid) (DTNB) was first introduced in 1959 for the estimation of free thiol groups (1). The procedure is based on the reaction of the thiol with DTNB to give the mixed disulfide and 2-nitro5-thiobenzoic acid (TNB) which is quantified by the absorbance of the anion (TNB2−) at 412 nm. The reagent has been widely used for the quantitation of thiols in peptides and proteins. It has also been used to assay disulfides present after blocking any free thiols (e.g., by carboxymethylation) and reducing the disulfides prior to reaction with the reagent (2,3). It is also commonly used to check the efficiency of conjugation (4) of sulfhydryl-containing peptides to carrier proteins (such as maleimide activated keyhole limpet haemocyanin, KLH) in the production of antibodies and to ensure that cysteine labeled sythetic peptides are coupled to affinity resins. 2. Materials 1. Reaction buffer: 0.1 M phosphate buffer, pH 8.0. 2. Denaturing buffer: 6 M guanidinium chloride, 0.1 M Na2HPO4, pH 8.0 (see Note 1). 3. Ellman’s solution: 10 mM (4 mg/mL) DTNB (Pierce, Chester, UK) in 0.1 M phosphate buffer, pH 8.0 (see Note 2). 4. Dithiothreitol (DTT) (Boerhinger or Calbiochem) solution: 200 mM in distilled water.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Methods 3.1. Analysis of Free Thiols 1. It may be necessary to expose thiol groups, which may be buried in the interior of the protein. The sample may therefore be dissolved in reaction buffer or denaturing buffer. A solution of known concentration should be prepared with a reference mixture without protein. Sufficient protein should be used to ensure at least one thiol per protein molecule can be detected; in practice, at least 2 nmol of protein (in 100 µL) are usually required. 2. Sample and reference cuvets containing 3 mL of the reaction buffer or denaturing buffer should be prepared and should be read at 412 nm. The absorbance should be adjusted to zero (Abuffer). 3. Add 100 µL of buffer to the reference cuvet. 4. Add 100 µL of Ellman’s solution to the sample cuvet. Record the absorbance (ADTNB). 5. Add 100 µL of protein solution to the reference cuvet. 6. Finally, add 100 µL protein solution to the sample cuvet, and after thorough mixing, record the absorbance until there is no further increase. This may take a few minutes. Record the final reading (Afinal). 7. The concentration of thiols present may be calculated from the molar absorbance of the TNB anion. (See Note 3.)

∆

A412 = E412TNB2−[RSH]

(1)

Where ∆A412 = Afinal – (3.1/3.2) (ADTNB – Abuffer) −1M−1 and E412 TNB2− = 1.415 × 104 cm. If using denaturing buffer, use the value E412 TNB2− = 1.415 × 104 cm −1M−1. 3.2. Analysis of Disulfide Thiols 1. Sample should be carboxymethylated (see Chapter 82) or pyridethylated (see Chapter 85) without prior reduction. This will derivatize any free thiols in the sample, but will leave intact any disulfide bonds. 2. The sample (at least 2 nmol of protein in 100 µL, is usually required) should be dissolved in 6 M guanidinium HCl, 0.1 M Tris-HCl, pH 8.0 or denaturing buffer, under a nitrogen atmosphere. 3. Add freshly prepared DTT solution to give a final concentration of 10–100 mM. Carry out reduction for 1–2 h at room temperature. 4. Remove sample from excess DTT by dialysis for a few hours each time, with two changes of a few hundred mL of the reaction buffer or denaturing buffer (see Subheading 3.1). Alternatively, gel filtration into the same buffer may be carried out. 5. Analysis of newly exposed disulfide thiols can thus be carried out as described in Subheading 3.1.

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4. Notes 1. It is not recommended to use urea in place of guanidinium HCl, since this can readily degrade to form cyanates, which will react with thiol groups. 2. Unless newly purchased, it is usually recommended to recrystallize DTNB from aqueous ethanol. 3. Standard protocols for use of Ellman’s reagent often give E412 TNB2− = 1.36 × 104 cm −1M−1. A more recent examination of the chemistry of the reagent indicates that these are more suitable values (5), and these have been used in this chapter. References 1. Ellman, G. L. (1959) Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82, 70–77. 2. Zahler, W. L. and Cleland, W. W. (1968) A specific and sensitive assay for disulfides. J. Biol. Chem.243, 716–719. 3. Anderson, W. L. and Wetlaufer, D. B. (1975) A new method for disulfide analysis of peptides. Analyt. Biochem. 67, 493–502. 4. Ellman assay, instruction book 77649, Pierce website: http://www.piercenet.com/ files/1371as4.pdf 5. Riddles P. W., Blakeley, R. L., and Zerner, B. (1983) Reassessment of Ellman’s reagent. Meth. Enzymol. 91, 49–60.

115 Quantitation of Cysteine Residues and Disulfide Bonds by Electrophoresis Alastair Aitken and Michèle Learmonth

1. Introduction Amino acid analysis quantifies the molar ratios of amino acids per mole of protein. This generally gives a nonintegral result, yet clearly there are integral numbers of the amino acids in each protein. A method was developed by Creighton (1) to count integral numbers of amino acid residues, and it is particularly useful for the determination of cysteine residues. Sulfhydryl and disulfide groups are of great structural, functional, and biological importance in protein molecules. For example, the Cys sulfhydryl is essential for the catalytic activity of some enzymes (e.g., thiol proteases) and the interconversion of Cys SH to Cystine S—S is directly involved in the activity of protein disulfide isomerase (2). The conformation of many proteins is stabilized by the presence of disulfide bonds (3), and the formation of disulfide bonds is an important posttranslational modification of secretory proteins (4). Creighton’s method exploits the charge differences introduced by specific chemical modifications of cysteine. A similar method was first used in the study of immunoglobulins by Feinstein in 1966 (5). Cys residues may be reacted with iodoacetic acid, which introduces acidic carboxymethyl (—O2CCH2S—) groups, or with iodoacetamide, which introduces the neutral carboxyamidomethyl (H2NCOCH2S—) groups. The reaction with either reagent is essentially irreversible, thereby producing a stable product for analysis. Using a varying ratio of iodoacetamide/iodoacetate, these acidic and neutral agents will compete for the available cysteines, and a spectrum of fully modified protein molecules having 0,1,2, … n acidic carboxymethyl residues per molecule is produced (where n is the number of cysteine residues in the protein). These species will

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Idealised electrophoretic pattern in an analysis of a protein with six cysteine residues, run with the low-pH system. Lanes 1–5 (left to right) contain (respectively) samples reacted with neutral iodoacetamide; 1:1; 1:3; 1:9 ratios of neutral to acidic reagent; acidic iodoacetate. The right hand lane contains a mixture of equal portions of the samples in lanes 1–5.

have, correspondingly, n, n − 1, n − 2, … 0 neutral carboxyamidomethyl groups. These species may then be separated by electrophoresis, isoelectric focusing, or by a combination of both (1,6,7). The theoretical analysis of a protein with 6 cysteines is shown in Fig. 1. Creighton used a low-pH discontinuous system (1). Takahasi and Hirose recommend a high-pH system (6), whereas Stan-Lotter and Bragg used the Laemmli electrophoresis system followed by isoelectric focusing (7). It may therefore be necessary to carry out preliminary experiments to find the best separation conditions for the protein under analysis. The commonly used methods are given below. In order to ensure that all thiol groups are chemically equivalent, the reactions must be carried out in denaturing (in the presence of urea) and reducing (in the presence of dithiothreitol, DTT) conditions. The electrophoretic separation must also be carried out with the unfolded protein (i.e., in the presence of urea) in order that the modification has the same effect irrespective of where it is in the polypeptide chain. The original method has been modified into a two-stage process to allow for the quantification of both sulfhydryl groups and disulfide bonds (see Notes 1 and 2) (6,8). The principle of the method has also been adapted to counting the numbers of lysine residues after progressive modification of the ε-amino groups with succinic anhydride, which converts this basic group to a carboxylic acid-containing moiety.

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2. Materials 2.1. Reaction Solutions 1. 2. 3. 4. 5. 6.

1 M Tris-HCl, pH 8.0. 0.1 M EDTA, pH 7.0. 1 M DTT (good-quality, e.g., Calbiochem, Nottingham, UK). 8 M Urea (BDH [Poole, UK], Aristar-grade, see Note 2). Solution A: 0.25 M iodoacetamide, 0.25 M Tris-HCl, pH 8.0. Solution B: 0.25 M iodoacetic acid, prepared in 0.25 M Tris-HCl, pH readjusted to 8.0 with 1 M KOH.

2.2. Solutions for Electrophoretic Analysis in the Low-pH System (pH 4.0)(9) 1. 30% Acrylamide solution containing 30 g acrylamide, 0.8 g bis-acrylamide (extreme caution: work in fume hood), made up to 100 mL with distilled water. 2. 10% Acrylamide solution containing 10 g acrylamide (extreme caution: work in fume hood) and 0.25 g bis-acrylamide made up to 100 mL with distilled water. 3. Low-pH buffer (eight times concentrated stock) for separating gel; 12.8 mL glacial acetic acid, 1 mL N,N,N N -tetramethylethylenediamine (TEMED), 1 M KOH (approx 35 mL) to pH 4.0, made up to 100 mL with distilled water. 4. Low pH buffer (8 times concentrated) for stacking gel; 4.3 mL glacial acetic acid, 0.46 mL TEMED, 1 M KOH to pH 5.0, to 100 mL with distilled water. 5. 4 mg ribofla vin/100 mL w ater. 6. Low-pH buffer for electrode buffer; dissolve 14.2 g β-alanine in 800 mL water then adjust to pH 4.0 with acetic acid. Make up to a final volume of 1 L with distilled water. 7. Tracking dye solution (five times concentrated); 20 mg methyl green, 5 mL water, and 5 g glycerol.

2.3. Gel Solution Recipes for Low-pH Electrophoresis (pH 4.0) (see Note 3) 1. 30 mL Separating gel (10% acrylamide, photopolymerized with riboflavin) is made up as follows: 10 mL 30% acrylamide stock, 4 mL pH 4.0 buffer stock, 3 mL riboflavin stock, 14.7 g urea, water (approx 2.5 mL) to 30 mL. Degas on a water vacuum pump (to removeoxygen which inhibits polymerization). 2. 8 mL Stacking gel (2.5% acrylamide, photopolymerized with riboflavin) is made up with: 2 mL 10% acrylamide stock, 1 mL pH 5.0 buffer stock, 1 mL riboflavin stock, 3.9 g urea, water (approx 1.2 mL) to 8 mL. Degas.

2.4. Electrophoresis Buffers for High-pH Separation (pH 8.9) 1. 30% Acrylamide solution containing 30 g acrylamide, 0.8 g bis-acrylamide (extreme caution: work in fume hood), made up to 100 mL with distilled water. 2. 10% Acrylamide solution containing 10 g acrylamide (extreme caution: work in fume hood) and 0.25 g bis-acrylamide made up to 100 mL with distilled water.

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3. High-pH buffer (four times concentrated stock) for separating gel; 18.2 g Tris base (in ~40 mL water), 0.23 mL TEMED, 1 M HCl to pH 8.9, made up to ~100 mL with distilled water. 4. High-pH buffer (four times concentrated) for stacking gel; 5.7 g Tris base (in ~40 mL water), 0.46 mL TEMED, 1 M H3PO4 to pH 6.9, made up to 100 mL with distilled water. 5. 4 mg Riboflavin in 100 mL water 6. 10% Ammonium persulfate solution (consisting of 0.1 g ammonium persulfate in 1 mL w ater). 7. High-pH buffer for electrode buffer; 3 g Tris base, 14.4 g glycine, distilled water to 1 L. 8. Tracking dye solution (five times concentrated): 1 mL 0.1% Bromophenol blue, 4 mL water and 5g glycerol.

2.5. Gel Solution Recipes for High-pH Electrophoresis (see Note 3) 1. 30 mL Separating gel (7.5% acrylamide, polymerized with ammonium persulfate) is made up with: 7.5 mL 30% acrylamide stock, 7.5 mL pH 8.9 buffer stock, 0.2 mL 10% ammonium persulfate (add immediately before casting), 14.7 g urea, water (approx 4.5 mL) to 30 mL. Degas. 2. 8 mL Stacking gel (2.5% acrylamide, photopolymerized with riboflavin) is made up with: 2 mL 10% acrylamide stock, 1 mL pH 6.9 buffer stock, 1 mL riboflavin stock solution, 3.9 g urea, water (approx 1.2 mL) to 8 mL. Degas.

3. Methods 3.1. Reduction and Denaturation 1. To a 0.2-mg aliquot of lyophilized protein add 10 µL of each of the solutions containing 1 M Tris-HCl, pH 8.0, 0.1 M EDTA, and 1 M DTT (see Note 4). 2. Add 1 mL of the 8 M urea solution (see Note 2). 3. Mix and incubate at 37°C for at least 30 min.

3.2. Reaction 1. Freshly prepare the following solutions using solutions A and B listed in Subheading 2. a. Mix 50 µL of solution A with 50 µL solution B (to give solution C). b. Mix 50 µL of solution A with 150 µL solution B (to give solution D). c. Mix 50 µL of solution A with 450 µL of solution B (to give solution E). 2. Label six Eppendorf tubes 1–6. 3. Add 10 µL of solutions A, B, C, D, and E to tubes 1–5. Reserve tube 6. 4. Add 40 µL of denatured, reduced protein solution prepared as in Subheading 3.1. to each of tubes 1–5. 5. Gently mix each tube and leave at room temperature for 15 min. Thereafter, store on ice. 6. After the 15 min incubation period, place 10 µL aliquots from each of tubes 1–5 into tube 6. Mix.

The samples are now ready for analysis (see Note 5).

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3.3. Electrophoretic Analysis 1. 50 µL aliquots of each sample, Labeled 1–6, mixed with 12 µL of appropriate tracking dye solution, are loaded onto successive lanes of a polymerized high- or low-pH gel, set up in a suitable slab gel electrophoresis apparatus. 2. Low-pH buffer system: Electrophoresis is carried out toward the negative electrode, using a current of 5–20 mA for each gel, overnight at 8°C. 3. High-pH buffer system (10); Electrophoresis is carried out toward the positive electrode at 10–20 mA per gel (or 100–180 V) for 3–4 h. 4. Electrophoresis is stopped when the tracking dye reaches bottom of the gel. 5. Proteins are visualized using conventional stains e.g., Coomassie blue (Pierce, Chester, UK), silver staining (see Chapter 48).

4. Notes 1. The method of Takahashi and Hirose (6) can be used to categorize the halfcystines in a native protein as: a. Disulfide bonded; b. Reactive sulfhydryls; and c. Nonreactive sulfhydryls. In the first step, the protein sulfhydryls are alkylated with iodoacetic acid in the presence and absence of 8 M urea. In the second step, the disulfide bonded sulfhydryls are fully reduced and reacted with iodoacetamide. The method described above is then used to give a ladder of half-cystines so that the number of introduced carboxymethyl groups can be quantified. 2. Urea is an unstable compound; it degrades to give cyanates which may react with protein amino and thiol groups. For this reason, the highest grade of urea should always be used, and solutions should be prepared immediately before use. 3. Electrophoresis in gels containing higher or lower percent acrylamide may have to be employed depending on the molecular weight of the particular protein being studied. 4. Where protein is already in solution, it is important to note that the pH should be adjusted to around 8.0, and the DTT and urea concentrations should be made at least 10 mM and 8 M, respectively. 5. Other ratios of iodoacetic acid to iodoacetamide may need to be used if more than about eight cysteine residues are expected, since a sufficiently intense band corresponding to every component in the complete range of charged species may not be visible. A greater ratio of iodoacetic acid should be used if the more acidic species are too faint (and vice versa). References 1. Creighton, T. E. (1980) Counting integral numbers of amino acid residues per polypeptide chain. Nature 284, 487, 488.

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2. Freedman R. B., Hirst, T. R., and Tuite, M. F. (1994) Protein disulphide isomerase: building bridges in protein folding. Trends Biochem. Sci. 19, 331–336. 3. Creighton, T. E. (1997) Disulphide bonds between cysteine residues, in Protein Structure—a Practical Approach, 2nd ed. (Rickwood, D. and Hames B. D., eds.), IRL, Oxford, UK, pp. 155–167. 4. Freedman, R. B. (1984) Native disulphide bond formation in protein biosynthesis; evidence for the role of protein disulphide isomerase. Trends Biochem. Sci. 9, 438–441. 5. Feinstein, A. (1966) Use of charged thiol reagents in interpreting the electrophoretic patterns of immune globulin chains and fragments. Nature 210, 135–137. 6. Takahashi, N. and Hirose, M. (1990) Determination of sulfhydryl groups and disulfide bonds in a protein by polyacrylamide gel electrophoresis. Analyt. Biochem. 188, 359–365. 7. Stan-Lotter, H. and Bragg, P. (1985) Electrophoretic determination of sulfhydryl groups and its application to complex protein samples, in vitro protein synthesis mixtures and cross-linked proteins. Biochem. Cell Biol. 64, 154–160. 8. Hirose, M., Takahashi, N., Oe, H., and Doi, E. (1988) Analyses of intramolecular disulfide bonds in proteins by polyacrylamide gel electrophoresis following twostep alkylation. Analyt. Biochem. 168, 193–201. 9. Reisfield, R. A., Lewis, U. J., and Williams, D. E. (1962) Disk electrophoresis of basic proteins and peptides on polyacrylamide gels. Nature 195, 281–283. 10. Davis, B. J. (1964) Disk electrophoresis II method and application to human serum proteins. Ann. N.Y. Acad. Sci. 21, 404–427.

116 N-Terminal Sequencing of N-Terminally Modified Proteins Roza Maria Kamp and Hisashi Hirano

1. Introduction A small amount of proteins separated by one-dimensional polyacrylamide gel electrophoresis (PAGE) or two-dimensional electrophoresis (2D-PAGE) and transferred from the gel onto a polyvinylidene difluoride (PVDF) membrane by electroblotting can be sequenced directly by a gas-phase protein sequencer (1). This blotting/sequencing technique has come to be widely used in protein de novo sequence analysis. However, even if proteins are successfully separated by PAGE and then transferred onto a PVDF membrane, N-terminal amino acid sequences of the proteins with a blocking group at the N-terminus cannot be determined by Edman degradation in the sequencer. Many proteins are N-terminally blocked (2,3). Brown and Roberts (4) found over 50% of soluble mammalian proteins to be N-terminally blocked. Thus, a simple and rapid technique for obtaining N-terminal sequence information on blocked proteins should be developed. Most techniques proposed so far for this purpose were applicable in order to obtain only the internal sequence of proteins. Acetyl modification is the most prevalent in the blocking groups identified so far (5), and the formyl and pyroglutamyl groups are also frequently detected (see Note 1). In this chapter, techniques for deblocking and N-terminal sequencing of proteins with an acetyl, formyl, or pyroglutamyl group at the N-terminus, which are electroblotted from a polyacrylamide gel onto a PVDF membrane (8), are described (see Note 2). Techniques for PAGE and protein electroblotting of proteins are presented elsewhere in this volume (see also 9,10) (see Note 3). On the other hand, recently, the alternative techniques using mass spectrometry have been rapidly developed, which allows the direct analysis of the N-terminally blocked peptides. The following three methods in combination with mass From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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spectrometry are available. (I) Sequencing of N-terminally blocked peptide by mass spectrometry after protease digestion of proteins (see Note 3): This technique will be described for acetylated and mirystoylated proteins in this chapter (5). (II) Enrichment of N-terminal modified peptides using DITC followed by sequencing of separated peptides using mass spectrometry: The blocked N-terminal peptides can be isolated by using diisothiothyanate (DITC)-glass, which reacts only with unmodified NH2-groups. Blocking groups, e.g. acetylated N-terminal groups do not react with DITC and can be easily and selectively isolated from all other peptides (6). Futhermore, the modified peptides can be analyzed by using mass spectrometry. (III) Specific sorting of N-terminal blocked peptides using chromatography (COFRADIC) and sequencing by mass spectrometry: Geavaert et al. described a new method, called COFRADIC, for isolation of modified N-terminal peptides. The peptides are separated using fractional diagonal chromatography. The proteins are cleaved by using endopeptidases, e.g. trypsin, and separated by using reversed phase chromatography. In the next step the internal and unblocked N-terminal peptides are treated by using 2,4,6-trinitrobenzene sulfonic acid (TNBS) and separated again by using reversed phase chromatography. The TNBS-modified peptides show strong hydrophobic shift and can be isolated from the unaltered N-terminal blocked peptides, which do not react with TNBS. The N-terminally modified peptides, e.g. acetylated peptides can be identified using mass spectrometry (7). 2. Materials 2.1. Deblocking of N-Formylated Proteins 1. Hydrochloric acid (0.1 M). 2. Acetonitrile (Sequencing grade).

2.2. Deblocking of Proteins with N-Terminal Pyroglutamic Acid 1. Acetonitrile (Sequencing grade). 2. Polyvinylpyrrolidone (PVP)-40: 0.5% w/v in acetic acid (100 mM). 3. Phosphate buffer (0.1 M, pH 8.0) containing 5mM dithiothreitol (DTT) and 10 mM ethylenediaminetetraacetic acid (EDTA). 4. Pyroglutamyl peptidase (Takara Shuzo, Japan).

2.3. Deblocking of N-Acetylated Proteins 2.3.1. Deblocking with Trifluoroacetic Acid (TFA) 1. Trifluoroacetic acid (TFA) (Sequencing grade). 2. Nitrogen Gas.

2.3.2. Deblocking with TFA in Methanol 1. TFA (Sequencing grade). 2. Methanol (Analytical grade).

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2.3.3. Deblocking with Acylamino Acid Releasing Enzyme 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Acetonitrile (Sequencing grade). PVP-40 (0.5% w/v) in acetic acid (100 mM). Trypsin (Promega, USA, modified by reductive methylation). Ammonium bicarbonate buffer (0.1 M, pH 8.0). Pyridine (50% v/v). Phenylisothiocyanate (Sequencing grade). Formic acid. Hydrogen peroxide (30% w/v). Phosphate buffer (0.2 M, pH 7.2). DTT (1 mM). Acylamino acid-releasing enzyme (AARE) (Takara Shuzo). Nitrogen gas.

2.4. Deblocking of Proteins with N-Acyl Groups by Pyrococcus Aminopeptidase 1. Ethylmorpholine buffer (50 mM, pH 8.0) containing CoCl2 (0.1%, v/v). 2. Pyrococcus furiosus aminopeptidase (Takara Shuzo).

2.5. Sequencing of the N-terminally blocked peptides by mass spectrometry 1. 2. 3. 4.

Trypsin (Promega, Wisconsin, USA). TFA (Sequencing grade). Acetonitrile (sequencing grade) Nanocapillary reversed phase column C18, 150 mm length, 0.75 µm diameter (LC Pacing, California, USA). 5. Trap column for CapLC System (Waters Corp., Massachussets, USA). 6. Electrospray ionization-quadrupole/time-of-flight mass spectrometry (ESI-Q/TOF MS)(Micromass, Manchester, UK).

3. Methods 3.1. Deblocking and Sequencing of N-Formylated Proteins 1. Separate the N-terminally blocked protein ( methoxy-acetoxy > acetoxy-acetoxy. The resulting spectra are diagnostic for the substitution pattern, and hence, the previous position of linkage, e.g., a 2-linked hexose will produce a different set of ions to a 3-linked hexose, and a 2,3-linked hexose being different again. Selected ions from the spectra of all commonly occurring linkages can be used to analyze across the chromatogram (selected ion monitoring). References 1. Hansson, G. C. and Karlsson, H. (1993) Gas chromatography and gas chromatography-mass spectrometry of glycoprotein oligosaccharides, in Methods in Molecular Biology, vol. 14: Glycoprotein Analysis in Biomedicine (Hounsell, E. F., ed.), Humana, Totowa, NJ, pp. 47–54. 2. Güther, M. L. and Ferguson, M. A. J. (1993) The microanalysis of glycosyl phosphatidylinositol glycans, in Methods in Molecular Biology, vol. 14: Glycoprotein Analysis in Biomedicine (Hounsell, E. F., ed.), Humana, Totowa, NJ, pp. 99–117. 3. Hounsell, E. F. (1994) Physicochemical analysis of oligosaccharide determinants of glycoproteins. Adv. Carbohyd. Chem. Biochem. 50, 311–350.

132 Sialic Acid Analysis by HPAEC-PAD Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith

1. Introduction The most labile monosaccharides are the family of sialic acids, which are usually chain-terminating substituents. These are therefore usually released first by either mild acid hydrolysis or enzyme digestion, and can be analyzed with great sensitivity by high pH anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD [1, 2]). The remaining oligosaccharide is analyzed as discussed in Chapter 131 to identify the position of linkage of the sialic acid. 2. Materials 1. 2. 3. 4.

HCl (0.01, 0.1, and 0.5 M). 100 mM NaOH. 1 M NaOAc. HPLC and PA1 columns as described in Chapter 129.

3. Method 1. Dry the glycoprotein into a clean screw-top vial with a Teflon-backed silicone lid insert (see Note 1). 2. Release the sialic acids by hydrolysis with 0.01 or 0.1 M HCl for 60 min at 70°C in an inert N2 atmosphere (see Note 2). 3. Dry down the hydrolysate, and wash three times with HPLC-grade H2O. 4. Prepare 500 mL of 100 mM NaOH, 1.0 M sodium acetate (eluant A). 5. Prepare 500 mL of 100 mM NaOH (eluant B). 6. Degas eluants by bubbling helium through them in their reservoirs (see Note 3). 7. Regenerate the HPLC column in 100% eluant B for 30 min at a flow rate of 1 mL/min. 8. Equilibrate the column in 95% eluant B for 30 min at a flow rate of 1 mL/min (see Note 4). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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9. Inject approx 0.2 nmol of sialic acid onto the column, and elute using the following gradient at a flow rate of 1 mL/min: a. Time = 0 min; 95% eluant B. b. Time = 4 min; 95% eluant B. c. Time = 29 min; 70% eluant B. d. Time = 34 min; 70% eluant B. e. Time = 35 min; 100% eluant B. f. Time = 44 min; 100% eluant B. g. Time = 45 min; 95% eluant B; 1 mL/min. 10. Quantitate the sialic acids by comparison with known standards run on the same day. 11. When the baseline has stabilized, the system is ready for the next injection. 12. When the analyses have been completed, regenerate the column in eluant B, and flush pumps with HPLC-grade H2O (see Note 5).

4. Notes 1. If problems with contaminants are encountered, it may be necessary to wash the vials with chromic acid overnight, wash them thoroughly with distilled water, and then treat with a hydrophobic coating, such as repelcoat (see Chapter 129). 2. 0.01 M HCl will release sialic acids with intact N- or O-acyl groups, but without quantitative release of the sialic acids. These can also be detected by HPAEC-PAD (3). At 0.1 M HCl, quantitive release is achieved, but with some loss of O- and N-acylation. At this concentration, some fucose residues may also be labile. Alternatively, the sialic acids can be released by neuraminidase treatment, which can be specific for α2–6 or α2–3 linkage, e.g., with α-sialidase of Arthrobacter ureafacians for α2–6 and α-sialidase of Newcastle disease virus for α2–3 using the manufacturer’s instructions. 3. Use polypropylene reagent vessels as far as possible for HPAEC-PAD because of the corrosive nature of the NaOH, and to minimize leaching of contaminants from the reservoirs. 4. For maximum efficiency of detection, always ensure that the PAD reference electrode is accurately calibrated, the working electrode is clean, and the solvents are thoroughly degassed. 5. Failure to wash out the eluants from the pumps at the end of an analysis may result in crystallization and serious damage to the pump heads. References 1. Townsend, R. R. (1995) Analysis of glycoconjugates using high-pH anion-exchange chromatography. J. Chromatog. Library 58, 181–209. 2. Manzi, A. E., Diaz, S., and Varki, A. (1990) HPLC of sialic acids on a pellicular resin anion exchange column with pulsed amperometry. Anal. Biochem. 188, 20–32.

133 Chemical Release of O-Linked Oligosaccharide Chains Elizabeth F. Hounsell, Michael J. Davies, and Kevin D. Smith

1. Introduction O-linked oligosaccharides having the core sequences shown below can be released specifically from protein via a β-elimination reaction catalyzed by alkali. The reaction is usually carried out with concomitant reduction to prevent peeling, a reaction caused by further β-elimination around the ring of 3-substituted monosaccharides (1). The reduced oligosaccharides can be specifically bound by solid sorbent extraction on phenylboronic acid (PBA) columns (2). O-linked protein glycosylation core structures linked to Ser/Thr: Galβ1-3GalNAcα1– GlcNAcβ1-3GalNAcα1– GlcNAcβ1-6 GalNAcα1– Galβ1-3 GlcNAcβ1-6 GalNAcα1– GlcNAcβ1-3 GalNAcα1-3GalNAcα1– GlcNAcβ1-6GalNAcα1– GalNAcα1-6GalNAcα1–

2. Materials 1. 1 M NaBH4 (Sigma, Poole, UK) in 50 mM NaOH. This is made up fresh each time from 50% (w/v) NaOH and HPLC-grade H2O. 2. Methanol (HPLC-grade containing 1% acetic acid). 3. Acetic acid. 4. 1 mL Dowex H+ (50 W × 12) strong cation ion-exchange column (Sigma, St. Louis, MO). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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5. Bond elute phenyl boronic acid column (Jones Chromatography, Hengoed, UK; activated with MeOH). 6. 0.1 M HCl. 7. 0.2 M NH4OH. 8. 0.1 M Acetic acid. 9. Methanol.

3. Method 1. Dry the glycoprotein (100 µg to 1 mg) in a screw-topped vial and resuspend in 100 µL 50 mM NaOH containing 1 M NaBH4 (see Notes 1 and 2). 2. Incubate at 55°C for 18 h. 3. Quench the reduction by the addition of ice-cold acetic acid until no further effervescence is seen. 4. Dry the reaction mixture down, and then wash and dry three times with a 1% acetic acid, 99% methanol solution to remove methyl borate. 5. Resuspend the alditols in H2O, and pass down a 1-mL H+ cation exchange resin. The alditols will not be retained and will elute by washing the column with water. 6. Dry the alditols and resuspend them in 100 µL of 0.2 M NH4OH. 7. Activate a phenyl boronic acid (PBA) column with 2 × 1 mL MeOH. 8. Equilibrate the PBA column with 2 × 1 mL 0.1 M HCl, 2 × 1 mL H2O, and 2 × 1 mL 0.2 M NH4OH. 9. Add the sample in 100 µL 0.2 M NH4OH and elute with 2 × 100 µL 0.2 M NH4OH, 2 × 100 µL H2O, and 6 × 100 µL 0.1 M Acetic acid. Collect these fractions and test for monosaccharide (Chapter 128) or combine and analyze according to Chapter 134. 10. Regenerate the PBA column with 0.1 M HCl and 2 × 1 mL H2O before storing and reactivation in 2 × 1 mL MeOH.

4. Notes 1. The NaBH4/NaOH solution is made up L R H K <TINVIR> D K R C R K R <SRVNLFSDR> F K C R V R F R E K V R E K S K S R V L R L R H K V K D

complex chain on N98KT

91.7

66–78

a

Sequencesb

R D

Relative hydrophobicity index on C18 HPLC. < > denotes the possible alternative tryptic cleavage sites.

b

Fx 8–9 Possible GPI Anchor at VKC- or high Man at N23 NT

Fx 10 High Man at N23 NT

Fx 13–14 Complex and hybrid at N74 FT

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7. To avoid unnecessary cross-contamination between samples, a 96-well dotblotter (Bio-Rad) was used instead of spotting the samples directly onto the membrane. Prior to activation or modification of the NCM, cut the membrane to size for placing onto the dot blotter cassette. Once the membrane is in a fixed position, apply respective samples as indicated in Subheading 3.5. After loading samples remove membrane from dot blotter and leave to air dry. Once dried, mark the membrane with a pencil to indicate the order of the samples. Also, draw a grid on paper to indicate the location of samples. 8. When applying respective samples onto the membrane, always use fresh Gilson tips for each sample to avoid contamination. 9. All incubations in the lectin overlay analysis are carried out on a rocking table at room temperature. 10. During the lectin overlay analysis, use gloves and avoid direct contact of the membrane with skin. Handle the membrane carefully with a flat-ended twiser to avoid damage. 11. For a clearer background, the blocking solution can be incubated with the membrane for overnight at 4°C. After, incubate the membrane with the blocking solution for a further 30 min at room temperature prior to washing. 12. For convenience, use large Petri dishes to incubate each membrane with respective lectin solutions. Make sure that the lectin solution always covers the membrane surface during incubation. 13. In contrast to the manufacturer’s instructions the staining reaction can be extended until significant staining could be observed (1–18 h). 14. Standard glycopeptides should also spotted onto the membrane to monitor the progress of staining. 15. To protect the membrane from dust and damage, place a clear plastic film over the membrane and store in the dark. 16. Table 2 presents a summary of results of a series of whole non-proteasetreated glycoproteins (at various concentrations) analyzed for lectin specificity and sensitivity by dot-blot lectin overlay. The results from this assay Table 2 The Sensitivity Levels of Commercial Lectins on Glycoproteins by Lectin Overlay Analysis Lectin

Fetuin

MAA SNA DSA GNA

10 ng 0.1 µg 0.1 µg 1 µg

Asialylated fetuin Bovine serum albumin Chicken ovalbumin RNase B 1 µg NB 0.1 µg 1 µg

NB, No binding of lectin detected to sample.

NB NB NB 2 µg

NB NB NB 1 µg

NB NB NB NB

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clearly show that three lectins (MAA, SNA, and DSA) recognize Bovine fetuin (BF) at high sensitivity protein levels ranging from 10 ng to 0.1 µg. Asialylated BF (ABF) were detected by lectins MAA and DSA but not SNA. As expected, all three lectins showed no binding to Bovine serum albumin (BSA), which is not glycosylated. According to the lectin specificity as published by the manufacturer, GNA should stain positive for RNase B and ovalbumin but not the other glycoproteins/proteins; however, negative results with RNase B were shown even after repeated tests such as using increased concentrations of RNase B (up to 20 µg). The sugar of RNase B was analyzed by hexose assay and shown to contain the expected 3%; thus the specificity of GNA is not as published, and the lectin can distinguish between ovalbumin and RNase B glycosylation. 17. Table 3 shows the binding preferences of the lectins used. From the results obtained, the sensitivity of lectins can detect protein concentrations down to 10 ng. The choice between the commercial digoxigenin-labeled and biotinylated lectins used relate to their recognition of a range of defined and limited sugar structures depending on the samples tested. One drawback to the procedure is that because a number of structurally distinct oligosaccharides interact identically with certain lectins, this suggests that the structural heterogeneity frequently encountered with glycoproteins may not always be reflected in the interactions of these moieties with lectins. 18. For pigeon intestinal mucin (2) 8 of the 10 lectins (AAL, LEL, LTL, MAL1, MPL, RCA-1, SBA, SJA, UEA-II, and WGA) reacted specifically with a different spectrum of PLL-oligosaccharide conjugates. LTL stained the entire blot, suggesting that this lectin interacted with the nitrocellulose. SJA reacted with all of the fractions and controls, indicative of nonspecific interaction with PLL itself. For the other lectins there was a wide range of activities with the various HPLC fractions. Difference in staining patterns between the lectins demonstrated that oligosaccharides of varying structure have been released from pigeon mucin by hydrazinolysis and have successfully been coupled to PLL, thus allowing binding to the NCM. Optimization of conditions at the macro-level included both neutral and sialylated oligosaccharides, with the chosen conditions giving maximum yield of both. The principle of this assay is derived essentially from enzyme-linked immunosorbent assay (ELISA) and commercial kits are available, where lectins are already linked to antibodies or other methods for detection. Acknowledgments The authors wish to thank Erminia Barboni for collaboration in a larger study that furnished the analytical amount of Thy-1 used in this study and to the following for support: E. B. PRIN97 program grant, University for Scientific and

Biotinylated lectins Aleuria aurantia lectin Lotus tetragonolobus lectin Lycopersicon esculentum (tomato) lectin Maclura pomifera lectin Ricinus communis agglutinin I Sophora japonica agglutinin Soybean agglutinin Ulex europaeus agglutinin II Wheat germ agglutinin

Sambucus nigra agglutinin

Maackia amurensis agglutinin

Galanthus nivalis agglutinin

Digoxigenin-labeled Lectins Datura stramonium agglutinin

Lectin

Fucose linked α1–6 to GlcNAc or to fucose linked α1–3 to N-acetyllactosamine α-Linked l-fucose containing oligosaccharides Trimers and tetramers of GlcNAc α-Linked GalNAc Terminal galactose and GalNAc Terminal GalNAc and galactose residues, with preferential binding to β-anomers Terminal α- or β-linked GalNAc, and to a lesser extent, galactose residues 2′ Flucosyllactose (fucosyl α1–2 galactosyl β1–4 glucose) Terminal GlcNAc or chitobiose and sialic acid

Galβ1–4GlcNAc in complex and hybrid N-glycans, in O-glycans and GlcNAc in O-glycans Terminal Man, α1–3, α1–6, or α1–2 linked to Man; suitable for identifying “high Man” N-glycan chains or O-glycosidically linked Man in yeast glycoproteins Sialic linked α2–3 to galactose; suitable for indentifying complex, sialylated carbohydrate chains and type of sialic acid linkage Sialic acid linked α2–6 to galactose; suitable for identifying complex, sialylated N-glycan chains in combination with lectin MAA

Specificity

Table 3 Binding Preferences of Digoxigenin-Labeled and Biotinylated Lectins

13 14 15 16 17 18 19 14, 20 21

12

11

10

9

References

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Technological Research, Rome, Italy; N. K. C. W., EU-BIOTECH research grant PL9765055; G. J. R., EU-HCM grant; C. I. B., Wellcome Trust grants 042462 and 052676; and E. F. H. and D. V. R., the UK Medical Research Council. References 1. Sharon, N. (1998) Lectins: from obscurity into the limelight. Protein Sci. 7, 2042– 2048. 2. Baldwin, C. I., Calvert, J. E., Renouf, D. V., Kwok, C., and Hounsell, E. F. (1999) Analysis of pigeon intestinal mucin allergens using a novel dot blot assay. Carbohydr. Res., in press. 3. Barboni, E., Pliego Rivero, B., George, A. J. T., Martin, S. R., Renouf, D. V., Hounsell, E. F., et al. (1995) The glycophosphatidylinositol anchor affects the conformation of the Thy-1 protein. J. Cell Sci. 108, 487–497. 4. Baldwin, C. I., Todd, A., Bourke, S. J., Allen, A., and Clavert, J. E. (1998) IgG subclass responses to pigeon intestinal mucin are related to development of pigeon fanciers’ lung. Clin. Exp. Allergy 28, 349–357. 5. Davies, M., Smith, K. D., Harbin, A.-M., and Hounsell, E. F. (1992) High performance liquid chromatography of oligosaccharide alditols and glycopeptides on a graphitised carbon column. J. Chromatogr. 609, 125–131. 6. Lauritzen, E., Masson, M., Rubin, I., and Holm, A. (1990) Dot immunobinding and immunoblotting of picogram and nanogram quantities of small peptides on activated nitro-cellulose. J. Immunol. Meth. 131, 257–267. 7. Browne, C. A., Bennett, H. P. I., and Solomon, S. (1982) The isolation of peptides by high performance liquid chromatography using predicted elution positions. Analyt. Chem. 124, 201–208. 8. Davies, M. J. and Hounsell, E. F. (1998) HPLC and HPAEC of oligosaccharides and glycopeptides, in Glycoanalysis Protocols (Hounsell, E. F., ed.), Humana Press, Totowa, NJ, pp. 79–107. 9. Green, E. D., Brodbeck, R. M., and Baenziger, J. U. (1987) Lectin affinity highperformance liquid chromatography: interactions of N-glycanase-released oligosaccharides with leukoagglutinating phytohemagglutinin, concanavalin A, Datura stramonium agglutinin, and Vicia villosa agglutinin. Analyt. Biochem. 167, 62–75. 10. Shibuya, N., Goldstein, I. J., Van Damme, E. J., and Peumans, W. J. (1988) Binding properties of a mannose-specific lectin from the snowdrop (Galanthus nivalis) bulb. J. Biol. Chem. 263, 728–734. 11. Wang, W. C. and Cummings, R. D. (1988) The immobilized leukoagglutinin from the seeds of Maackia amurensis binds with high affinity to complex-type Asnlinked oligosaccharides containing terminal sialic acid-linked alpha-2,3 to penultimate galactose residues. J. Biol. Chem. 263, 4576–4585. 12. Shibuya, N., Goldstein, J., Broeknaert, W. F., Nsimba-Luvaki, M., Peeters, B., and Peumanns, W. J. (1987) The elderberry (Sambucus nigra L.) bark lectin recognizes the Neu5Ac(alpha 2-6)Gal/GalNAc sequence. J. Biol. Chem. 262, 1596–1601. 13. Yazawa, S., Furukawa, K., and Kochibe, N. (1984) Isolation of fucosyl glycoproteins from human erythrocyte membranes by affinity chromatography using Aleuria aurantia lectin. J. Biochem. (Tokyo) 96, 1737–1742.

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14. Allen, H. J., Johnson, E. A., and Matta, K. L. (1977) A comparison of the binding specificities of lectins from Ulex europaeus and Lotus tetragonolobus. Immunol. Commun. 6, 585–602. 15. Zhu, B. C. and Laine, R. A. (1989) Purification of acetyllactosamine-specific tomato lectin by erythroglycan-sepharose affinity chromatography. Prep. Biochem. 19, 341–350. 16. Lee, X., Thompson, A., Zhang, Z., Ton-that, H., Biesterfeldt, J., Ogata, C., et al. (1998) Structure of the complex of Maclura pomifera agglutinin and the T-antigen disaccharide, Galbetal,3GalNAc. J. Biol. Chem. 273, 6312–6318. 17. Wu, J. H., Herp, A., and Wu, A. M. (1993) Defining carbohydrate specificity of Ricinus communis agglutinin as Gal 1–4GlcNAc>Gal 1–3GlcNAc>Gal 1– 3Gal>Gal 1–3GalNAc. Mol. Immunol. 30, 333–339. 18. Fournet, B., Leroy, Y., Wieruszeski, J. M., Montreuil, J., Poretz, R. D., and Goldberg, R. (1987) Primary structure of an N-glycosidic carbohydrate unit derived from Sophora japonica lectin. Eur. J. Biochem. 166, 321–324. 19. Dessen, A., Gupta, D., Sabesan, S., Brewer, C. F., and Sacchettini, J. C. (1995) X-ray crystal structure of the soybean agglutinin cross-linked with a biantennary analog of the blood group I carbohydrate antigen. Biochemistry 34, 4933–4942. 20. Akiyama, M., Hayakawa, K., Watanabe, Y., and Nishikawa, T. (1990) Lectin-binding sites in clear cell acanthoma. J. Cutan. Pathol. 17, 197–201. 21. Renkonen, O., Penttila, L., Niemela, R., Vainio, A., Leppanen, A., Helin, J., et al. (1991) N-Acetyllactosaminooligosaccharides that contain the βd-GlcpNac-(1→6)d-GlcpNAc-(1→6)-d-GalNAc sequences reveal reduction-sensitive affinities for wheat germ agglutinin. Carbohydr. Res. 213, 169–183.

142 Polyacrylamide Gel Electrophoresis of Fluorophore-Labeled Carbohydrates from Glycoproteins Brian K. Brandley, John C. Klock, and Christopher M. Starr

1. Introduction Prior to 1980, most methods for analysis of glycoprotein carbohydrates utilized column, thin-layer, and paper chromatography, gas chromatography, mass spectroscopy, and rarely nuclear magnetic resonance spectroscopy. These methods required relatively large amounts of materials (micromoles), specialized training and experience, and in some cases, significant capital equipment outlays. Because of these restrictions, convenient carbohydrate analysis on small samples was not available to most biologists. Recently, improvements in chromatographic methods, labeling methods for carbohydrates, carbohydratespecific enzymes, and higher resolution electrophoresis methods have allowed carbohydrate analysis to be done on nanomolar amounts of material. Because of these improvements, today’s biologist now has an improved ability to evaluate the role of carbohydrates in their research and development work. Polyacrylamide gel electrophoresis (PAGE) of fluorophore-labeled carbohydrates has also been referred to as fluorophore-assisted carbohydrate electrophoresis, or FACE®. The technique was first developed in England by Williams and Jackson (1–4) and utilizes reductive amination of carbohydrates by low molecular weight, negatively charged or neutral fluorophores and electrophoresis on 20–40% polyacrylamide slab gels. This method permits separation of charged or uncharged sugars or oligosaccharides with high resolution and can detect single hydroxyl anomeric differences between mono- and oligosaccharides of sugars with otherwise identical molecular weight, charge, and sequence. The separation of sugars by electrophoresis is largely empirical, and it is not always possible to predict relative mobilities of structures. For the most part the success From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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of this approach has been based on experimental observations and the use of highly specific reagents, enzymes, and standards. What follows in this chapter are descriptions of the materials and methods required to perform two of the most common manipulations of oligosaccharides used in biologic research today. The method with some modification is useful for analysis of a variety of carbohydrates including reducing and nonreducing sugars, substituted and unsubstituted monosaccharides, and oligosaccharides from glycoproteins, proteoglycans, glycolipids, and polysaccharides. In this chapter only profiling of N-linked and O-linked glycoprotein oligosaccharides is discussed. The method is also useful for a variety of other manipulations including structure–function studies, preparative work, and synthesis of oligosaccharides that are not discussed in detail here. Other reviews of this technique have been published previously (5,6) and a number of research studies have been performed using this method (7–31). 2. Materials 2.1. N-linked Oligosaccharide Analysis For profiling of Asn-linked oligosaccharides released by peptide N-glycosidase F. 1. N-linked Gels: 10 × 10 cm low fluorescence glass plates with 0.5 mm spacers and 8-well combs filled with 20% T acrylamide:bis in 0.448 M Tris-acetate buffer pH 7.0 and containing a 5-mm stack of 10% polyacrylamide in the same buffer (gels should be made fresh or obtained precast from a commercial source). 2. Releasing enzyme: peptide N-glycosidase F (from commercial sources). 3. Running buffer: 50 mM Tris tricine buffer (pH 8.2). 4. Enzyme buffer: 100 mM sodium phosphate buffer (pH 7.5). 5. Labeling dye: 1 M 8-aminonaphthalene-1,3,6-trisulfonic acid (ANTS) in 15% acetic acid (reagent is stable for 2 wk at–70°C in the dark). 6. Reducing agent: 1 M sodium cyanoborohydride (NaBH3CN) in dimethyl sulfoxide (DMSO) (reagent is stable for 2 wk at–70°C). 7. Sample loading solution: 25% glycerol with thorin 1 dye (store at 4°C). 8. Tracking dye: Mixture of Thorin 1, Bromphenol blue, Direct red 75, and Xylene cyanole in water. 9. Electrophoresis gel box (with 2-sided cooling). 10. Deionized or distilled water. 11. Assorted pipeting devices including a 0–10 µL positive displacement pipet (e.g., Hamilton syringe). 12. Centrifugal vacuum evaporator. 13. Oven or water bath at 45°C and 37°C. 14. Sodium dodecyl sulfate (SDS), 5%. 15. β-Mercaptoethanol (βME). 16. 7.5% nonidet P-40 (NP-40). 17. 100% Cold ethanol (undenatured).

Determination of Monosaccharide Linkage 18. 19. 20. 21.

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Microcentrifuge tubes (1.5 mL). Microcentrifuge. Chicken trypsin inhibitor control (optional). Maltotetraose or partially hydrolyzed starch standard (optional).

2.2. O-linked Glycoprotein Oligosaccharide Analysis For profiling of Ser/Thr-linked oligosaccharides released by hydrazine. 1. O-linked gels: 10 × 10 cm low fluorescence glass plates with 0.5-mm spacers and eight-well combs filled with 35% T acrylamide:bis in 0.448 M Tris-acetate buffer and containing a 5-mm stack of 16% polyacrylamide in the same buffer (gels should be made fresh or obtained as precast gels from a commercial source). 2. Gel running buffer: 50 mM Tris-glycine (pH 8.2). 3. O-linked cleavage reagent: anhydrous hydrazine, 1 mL amp. Hydrazine is toxic and flammable; discard ampoule and residual contents after using once; dispose of safely according to your institution’s regulations. 4. Re-N-acetylation reagent: acetic anhydride. 5. Re-N-acetylation buffer: 0.2 M ammonium carbonate, pH 9.4. 6. Desalting resin: Dowex AG50X8. 7. Tracking dye: Mixture of Thorin 1, Bromphenol blue, Direct red 75, and Xylene cyanole in water. 8. Sample loading solution: 25% glycerol with Direct red 75 (store at 4°C). 9. Labeling dye: 1 M 8-aminonaphthalene-1,3,6-trisulfonic acid (ANTS) in 15% acetic acid (reagent is stable for 2 wk if stored in the dark at –70°C). 10. Reducing agent: 1 M sodium cyanoborohydride (NaBH3CN) in DMSO (reagent is stable for 2 wk at –70°C). 11. Distilled-deionized water. 12. Microcentrifuge tubes (2-mL, glass-lined). 13. Assorted pipeting devices including a 0–100 µL capillary positive displacement pipet. 14. Centrifugal vacuum evaporator. 15. Oven or water bath set at 37°C. 16. Sand filled heat block set at 60°C. 17. Vacuum dessicator. 18. Microfuge. 19. Phosphorus pentoxide (P2O5). 20. Bovine submaxillary mucin control (optional). 21. Maltotetraose or partially hydrolyzed starch standard (optional).

3. Methods 3.1. Principle Using fluorescent PAGE, individual oligosaccharides can be quantified to obtain molar ratios, to obtain degree of glycosylation, and to detect changes in the extent or nature of glycosylation. Oligosaccharide profiling involves four steps:

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Release of the oligosaccharides from the glycoprotein enzymatically or chemically; Labeling of the mixture of released oligosaccharides with a fluorescent tag; Separation of the fluorophore-labeled oligosaccharides by PAGE; and Imaging of the gel either on a UV lightbox to obtain qualitative band information or using a commercial imaging system to determine the amount of oligosaccharide present in each band and the relative mobility of the bands.

Once separated on the gel, individual oligosaccharide bands can also be purified for further study. 3.2. Preparation of Glycoprotein 1. Isolate the glycoprotein according to your usual procedures. The sample should be relatively salt-free and contain no extraneous carbohydrates (e.g., sephadexpurified material contains large amounts of glucose) (see Note 1). 2. If the volume of the glycoprotein solution required is >100 µL, dry the glycoprotein in a 1.5-mL microcentrifuge tube. Generally 50–200 µg of glycoprotein is required for N-linked analysis (see Note 2). For analysis of O-linked oligosaccharides 100–500 µg of glycoprotein may be required. 3. For the N-linked oligosaccharide control (see Note 3) remove 100 µg of the chicken trypsin inhibitor in a 45 mL aliquot, and place it in a 1.5 mL microfuge tube. Proceed with the enzymatic digestion as described below. For the O-linked oligosaccharide control remove 50 µL (100 µg) of Bovine submaxillary mucin, and place it in a reaction vial. Proceed with lyophilization, P2O5 drying and process along with the sample glycoprotein (see Subheading 3.4.2.). Store remaining glycoproteins at 4°C for future use.

3.3. Release of Asn-Linked Oligosaccharides with Peptide N-Glycosidase F 1. Add an equal volume of enzyme buffer to the glycoprotein in solution or dissolve the dried glycoprotein in 22.5 µL of water and add 22.5 µL enzyme buffer. 2. SDS is often required to completely denature the glycoprotein prior to enzymatic digestion. To denature the protein add SDS to 0.1% (1.0 µL of 5% SDS to 45 µL reaction) and β-ME to 50 mM (1.5 µL of a 1:10 dilution of 14.4 M stock β-ME to 45 µL reaction). Boil for 5 min (see Note 4), cool to room temperature and add NP-40 to 0.75% (5 µL of 7.5% NP-40 into 45 µL). Mix with finger flicks. 3. Add 2.0 µL (5 U of peptide-N-glycosidase F or as specified by the manufacturer) of enzyme to the glycoprotein sample. Mix with finger flicks and centrifuge for 5 s. Store remaining enzyme at 4°C. 4. Incubate sample for 2 h at 37°C. 5. Precipitate protein by adding 3 vol of cold 100% ethanol. Keep samples on ice for 10 min. Spin samples in microcentrifuge for 5 min to pellet protein. 6. Remove the supernatant and transfer to a clean 1.5-mL microcentrifuge tube. IMPORTANT! Do not discard the supernatant. It contains the released carbohydrates!

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7. If a large amount of protein was digested (>250 µg) 5–10% of the released oligosaccharides may remain in the pellet. The recovery of these oligosaccharides can be accomplished by drying the pellet completely in a centrifugal vacuum evaporator or lyophilizer. Add 50 µL H2O to resuspend, then 150 µL 100% cold ethanol and precipitate on ice. Centrifuge and combine the supernatants. 8. Dry the supernatants in a centrifugal vacuum evaporator or lyophilize to a translucent pellet. At this point samples may be stored at −20°C or proceed with the fluorophore labeling procedure described in Subheading 3.4.2.

3.4. O-Linked Oligosaccharide Release Using Hydrazine 3.4.1. Isolation of Glycoprotein 1. Isolate glycoprotein according to your usual procedures. The purified glycoprotein should be prepared in a non-Tris buffer containing a minimum amount of salt. The presence of nonvolatile salts may cause the breakdown of the oligosaccharides during hydrazinolysis. If the glycoprotein is in a buffer containing salt it is recommended that the sample be dialyzed against distilled water to remove salts prior to chemical digestion.

3.4.2. Hydrazinolysis 1. Dry 100–500 µg of glycoprotein in a glass-lined reaction vial, using a centrifugal vacuum evaporator or lyophilizer. The actual amount of glycoprotein required will depend on the size of the protein and the extent of glycosylation (see Note 2). 2. The sample must be completely dry before hydrazinolysis. Following lyophilization, dry the sample overnight under vacuum in the presence of P2O5 to remove all traces of H2O. Place samples in a dessicator flask with a beaker containing a small amount of P2O5. Attach the dessicator directly to the pump without a cold-trap— any water remaining in the sample will be trapped by the P2O5. 3. Open a fresh ampoule of anhydrous hydrazine O-linked cleavage reagent. Add 50 µL of hydrazine to the dried sample using a glass transfer pipet, or a positive displacement capillary pipet (metal or plastic should not be used). Resuspend the dried sample. Overlay the sample with dry nitrogen and cap tightly. Hydrazine is very hygroscopic. Discard unused hydrazine according to your hazardous waste regulations. Do not reuse. 4. Incubate samples for 3 h in a sand bath or dry heat block set at 60°C (do not use a water bath) to release O-linked oligosaccharides (higher temperatures may result in the degradation of O-linked sugars or in the release of non-O-linked sugar chains, such as N-linked sugars, from the sample if they are present). 5. Dry samples in vacuum evaporator on low heat setting.

3.4.3. Re-N-Acetylation Procedure 1. Add 30 µL of Re-N-acetylation buffer to the dried pellet from step 5 in Subheading 3.4.2. 2. Resuspend by vortexing. Spin 2 s in a microfuge. 3. Add 2 µL of re-N-acetylation reagent to the solution. Mix well. Spin 2 s in a microfuge.

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4. Incubate tubes on ice for 15 min. 5. Following the 15 min incubation, stop the reaction by adding 60 µL of the desalting resin. Desalting resin is prepared by adding 0.5 g of Dowex AG50X8 to 0.7 mL water. Invert or vortex the resin immediately prior to removing the 60 µL for each tube. Incubate the resin with the sample at room temperature for 5 min mixing by placing the tube on a shaker or by continuously inverting the tube to keep the resin suspended. 6. Briefly centrifuge to pellet the resin, remove the supernatant (save supernatant) and wash the resin 2× with 120 µL of water for 2 min each (save supernatant). 7. Combine the resin supernatants in a 1.5-mL microcentrifuge tube. 8. Dry the supernatants in a vacuum evaporator on low heat setting.

3.5. Labeling Oligosaccharides 3.5.1. Preparation of Samples and Standards If quantitation of the oligosaccharide bands in the samples is required, then one must compare the intensity of an internal standard (e.g., maltotetraose) band with the intensity of sample bands and it is, therefore, essential that the standard is present on each gel used (see Note 5 for preparation and use of this material). 3.5.2. ANTS Labeling 1. Prepare the labeling dye as 1 M ANTS in 15% acetic acid (dye solution can be stored in the dark at −70°C for up to 2 wk). 2. Prepare 1 M solution of NaBH3CN in DMSO and mix well by vortexing until crystals are completely dissolved (this reducing agent can be stored for 2 wk at −70°C). 3. Add 5 µL of labeling dye to each dried oligosaccharide pellet. Mix well until the oligosaccharide pellet is dissolved. 4. Add 5 µL of reducing agent. Mix well by vortexing. Centrifuge 5 s in microcentrifuge. 5. Incubate samples at 45°C for 3 h (temperatures higher than 45°C or times longer than 3 h can destroy or modify carbohydrates, e.g., sialic acids). Greater than 90% of the oligosaccharides are labeled under these conditions. As a convenient alternative samples can be labeled at 37°C (not 45°C) overnight (or approx 16 h). These latter conditions result in labeling of >90% of the oligosaccharides (see Note 6). 6. After labeling, dry the samples in centrifugal vacuum evaporator for approx 15 min or until the sample reaches a viscous gel stage.

3.6. Electrophoresis 3.6.1. Preparation of a Sample for Electrophoresis 1. Resuspend the dried fluorophore “labeled” oligosaccharide in 5–20 µL H2O. The actual volume of H2O used to resuspend the sample will depend on the amount of oligosaccharide present in the sample (start with 10 µL, this will enable the sample to be diluted further if necessary).

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2. Remove an aliquot of the sample (generally 1–2 µL) and dilute it with an equal volume of sample loading solution. Load the entire aliquot into one lane of a gel. Best results are obtained by loading 4 µL/lane on a 10 × 10 cm gel with eight lanes.

3.6.2. Electrophoresis 1. For N-linked analysis chill the running buffer to 4–6°C prior to use. For N-linked analysis perform electrophoresis at a buffer temperature of 5–8°C. All O-linked gels are run at 15–20°C. 2. For N-linked analysis set up electrophoresis with a recirculating chiller and place the electrophoresis tank containing a stir bar on a mechanical stirrer. Connect the gel box cooling chamber to a refrigerating circulator. Turn on the circulator and stirrer and set the coolant temperature to 5°C. 3. For N-linked analysis pour the precooled running buffer into the electrophoresis tank up to the appropriate level. The temperature of the buffer should be monitored during the run using a thermometer inserted through the hole in the lid or other method. The temperature will probably increase a few degrees during electrophoresis, but should not exceed 10°C. For O-linked gels the temperature should not exceed 23°C. 4. Determine the number of gels required for the samples prepared. Each gel should contain eight lanes. The outside lanes should be used for the tracking dye and glucose polymer standard leaving the six inner lanes for samples and quantitation standard (maltotetraose). 5. Gently remove the comb(s) from the gel(s). To avoid distorting the wells, gently wiggle each comb to free the teeth from the gel, then lift up slowly until the comb is released. 6. Place the gel cassette(s), one on each side of the center core unit of the gel apparatus with the short glass plate against the gasket. Be sure the cassette is centered and that the cassette is resting on the “feet” at the bottom of the apparatus. If only one gel is being run place the buffer dam on the other side. 7. It is essential that the wells of the gel are thoroughly rinsed out with the running buffer from the upper buffer reservoir prior to sample loading. This is best accomplished by using a syringe with a blunt needle (a Pasteur pipet is not recommended because of the possibility of breakage into the wells). 8. With the core unit containing the gels placed securely on the bench, load samples into the wells by underlaying the upper buffer. Use flat sequencing pipet tips to load by delivering the sample to the bottom of each well. Optimal resolution will be achieved by using 4 µL of sample per lane. Note: For the most reliable quantitation of oligosaccharide bands the use of a positive displacement pipet (e.g., Hamilton syringe) is recommended. 9. Load 4 µL of the standard in a lane when prepared as described in Note 5. 10. Load 2 µL of tracking dye in a lane directly from the vial. 11. Load 4 µL of each labeled oligosaccharide sample in a lane. Samples should be diluted 1:1 in the sample loading solution (see Note 7).

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12. To prevent possible lane distortions as a result of different loading volumes it is recommended that 4 µL of Sample Loading Solution be loaded in any unused lanes. Best results are obtained when the same volume of sample is added to each lane. 13. Place the core unit containing the loaded gels into the electrophoresis tank and connect the power cords to the electrophoresis tank then connect the power supply. 14. Place the thermometer into the lower buffer chamber through the hole in the lid. For N-linked analysis the initial temperature of the lower buffer must be 5–8°C; for O-linked analysis 15–20°C is optimal. 15. Turn on the power supply and select the proper current. Gels should be run at a constant current of 15 mA/gel (30 mA for 2 gels; 15 mA for 1 gel and 1 buffer dam). Limits on the power supply should be set for 1000V and 60W. These run conditions will result in voltages of 100–400V at the beginning of the run and may approach 800–1000V at the end of the run. If the initial voltage is significantly different check to be sure that the leads are connected properly and that the buffers are at the recommended levels (see Notes 8 and 9). 16. Most N-linked oligosaccharides fall in the Glucose4–Glucose12 range (also referred to as G4–G12 or DP4–DP12), so the time of electrophoresis should be adjusted to optimize the separation of this region of the gel. Most O-linked oligosaccharides run in the G1–G6 range (DP1–DP6). 17. Monitor the electrophoresis by following the migration of the fast moving thorin dye (orange band). Generally, electrophoresis of N-linked oligosaccharides is complete when the orange dye just exits the bottom of the gel in approx 1 to 1¼ h. For the O-linked oligosaccharides the gel run is complete when the orange dye is 1 cm above the bottom of the gel. In a darkened room, the migration of the labeled oligosaccharides can be monitored directly during electrophoresis by turning off the power supply, removing the leads and the gel box cover and holding a handheld UV light over the gels. The run can be continued by repositioning the gel in the electrophoresis box and reconnecting the power supply as described in step 15. The amount of time the current is off should be as short as possible (20 nmol it is recommended that an internal labeling control is included. 7 Sample handling and storage: a. Always avoid exposing labeled samples and dyes to light or excess heat;

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b. Labeled samples are stable when stored for 3 mo at −70°C in the dark; c. Unused solutions of the dye and reducing agent can be stored for as long as 2 wk at −70°C. Thaw immediately before use. Band distortion in gels caused by vertical streaking or smearing may result if the sample is overloaded. Use a maximum of 1/5 of the volume of the labeling reaction for each lane. The sample may have a high concentration of salt. Remove salts by dialysis, desalting column, and so on, prior to enzymatic digestion. Distorted sample wells in gel may be caused by tearing of wells when the comb was removed. Remove the comb slowly using a gentle back and forth rocking motion and lift vertically. Alternatively, gels may have been in contact with upper buffer too long prior to sample loading. Samples should be loaded within 5 min of placing the gel in the upper buffer tank. Voltage and/or current leaks can result when high voltages are used. If at the beginning of the run the voltage is >400V or readings are unstable, turn the power off before checking the following: possible electrical leak, check for cracks in glass plates. Remove inner core assembly and check for buffer leak between gaskets and cassette plates. If leaks are evident check that the plates are clean and not cracked or chipped, and that they are installed properly. Buffers should not be reused as they have fluorophore contamination after use. Reuse of buffers may result in no bands being visible on the gel owing to “washout” of the fluorophore-labeled oligosaccharides. Accurate quantification is essential for detailed carbohydrate analysis. Although oligosaccharide patterns on PAGE gels can be viewed and photographed on a standard laboratory UV lightbox, it is not reliable for accurate quantification. Images of gels can be recorded using a Polaroid camera. The proper choice of light source, filters, and film must be made. A filter must be fitted to the camera lens that completely covers the glass of the lens (stray UV contacting the lens will cause it to fluoresce and subsequently lower the sensitivity of the film). A suitable filter will have no inherent fluorescence, peak transmission at approx 500 nm and bandwidth of 80 nm FWHM. A medium speed, medium resolution, Polaroid film is recommended. Use Polaroid 53 film for cameras which use single 4 × 5″sheet film; use Polaroid 553 film for cameras that use 8 sheet film cartridges. To visualize the carbohydrate banding patterns, the low fluorescent glass cassette containing the gel (or the gel removed from the casette) is placed on a longwave UV lightbox with a peak excitation output at approx 360 nm. Photograph the gel using the lowest practical f-stop setting on the lens with the gel filling as much of the frame as possible. E.g., exposures at f5.6

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13.

14.

15.

16.

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using Polaroid 53 film have ranged from 5–40 s using the equipment specified above. Keep UV exposure of the gel to a minimum to prevent bleaching. Develop the film according to the manufacturer’s instructions. For electronic archiving and quantitation, several types of imaging systems are available. To give best results these systems must have an illumination source with an excitation wavelength of 365 nm and a 520 nm emission filter placed in the light path between the gel and the image capturing device. The use of an internal standard in the gels is also required for quantitation. Following electrophoresis, the gel is inserted into the imager under longwave UV excitation, and an electronic image of the fluorescent carbohydrate banding pattern of the gel is acquired by the imager’s CCD as a digital image. The gel image is displayed on a computer screen using the imaging software. The imaging system should allow for detection and quantification of individual carbohydrate bands into the low picomole range of 1.6–300 pmol. In practice, the most useful and accurate range of the imager for band quantification is between 5 and 500 pmol of carbohydrate and this range was used for the experiments described in this chapter. You may have “smile effect” gel distortions at both sides of the gel. This can happen if the gel is not being cooled uniformly. Check that the cooling system is on and working properly. Check the buffer temperature. Check that the power supply is set for the proper current level. You may have band distortions or “fuzzy bands.” This can be caused by wells that may have not been rinsed thoroughly with electrophoresis upper buffer prior to loading samples, or the current may have not been set properly, i.e., the current was too high. Incomplete re-N-acetylation may result in little or no labeling of the released oligosaccharides, presumably by the hydrazide interfering with the reductive amination using ANTS. To check the re-N-acetylation reagents you can use glucosamine that will migrate at a DP of approx 2.5 on the gel when N-acetylated and below DPI when unacetylated. You can also use N-acetyl-glucosamine as an internal control at the beginning of the experiment and take it through hydrazinolysis and re-N-acetylation expecting the same migrations as stated. Oligosaccharides are small molecules that can diffuse rapidly in the gel matrix. Band diffusion and resulting broad or “fuzzy” bands can occur if: a. Electrophoresis is run too slowly; b. Electrophoresis is run at higher than optimal temperatures; c. Electrophoresis is stopped and started repeatedly; and d. The gel is removed for visualization for longer than 10 min and then re-electrophoresed.

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References 1. Williams, G. R. and Jackson, P. (1992) Analysis of carbohydrates. U.S. Patent 5, 508. 2. Jackson, P. (1990) The use of polyacrylamide-gel electrophoresis for the high resolution separation of reducing sugars labeled with the fluorophore 8-aminonaphtha lene-1,3,6-trisulfonic acid. Biochem. J. 270, 705–713. 3. Jackson, P. and Williams, G. R. (1991) Polyacrylamide gel electrophoresis of reducing saccharides labeled with the fluorophore 8-aminonaphthalene-1,3,6- trisulfonic acid: application to the enzymatic and structural analysis of oligosaccharides. Electrophoresis 12, 94–96. 4. Jackson, P. (1994) The analysis of fluorophore-labeled glycans by high-resolution polyacrylamide gel electrophoresis. Analyt. Biochem. 216, 243–252. 5. Starr, C., Masada, R. I., Hague, C., Skop, E., and Klock, J. (1996) Fluorophoreassisted-carbohydrate- electrophoresis, FACE® in the separation, analysis, and sequencing of carbohydrates. J. Chromatogr. A 720, 295–321. 6. Masada, R. I., Hague, C., Seid, R., Ho, S., McAlister, S., Pigiet, V., and Starr, C. (1995) Fluorophore-assisted-carbohydrate-electrophoresis, FACE®, for determining the nature and consistency of recombinant protein glycosylation. Trends Glycosci. Glycotech. 7, 133–147. 7. Hu, G. (1995) Fluorophore assisted carbohydrate electrophoresis technology and applications. J. Chromatogr. 705, 89–103 8. Higgins, E. and Friedman, Y. (1995) A method for monitoring the glycosylation of recombinant glycoproteins from conditioned medium, using fluorophore assisted carbohydrate electrophoresis. Analyt. Biochem. 228, 221–225. 9. Basu, S. S., Dastgheib-Hosseini, S., Hoover, G., Li, Z., and Basu, S. (1994) Analysis of glycosphingolipids by fluorophore-assisted carbohydrate electrophoresis using ceramide glycanase from Mercenaria mercenaria. Analyt. Biochem. 222, 270–274. 10. Flesher, A. R., Marzowski, J, Wang, W., and Raff, H. V. (1995) Fluorophore-labeled carbohydrate analysis of immunoglobulin fusion proteins: correlation of oligosaccharide content with in vivo clearance profile. Biotech. Bioeng. 46, 399–407. 11. Lee, K. B., Al-Hakim, A., Loganathan, D., and Linhard, R. J. (1991) A new method for sequencing carbohydrates using charged and fluorescent conjugates. Carbohydr. Res. 214, 155–162. 12. Stack, R. J. and Sullivan, M. T. (1992) Electrophoretic resolution and fluorescence detection of N-linked glycoprotein oligosaccharides after reductive amination with 8-aminonaphthalene-1,3,6-trisulfonic acid. Glycobiology 2, 85–92. 13. Roy, S. N., Kudryk, B., and Redman, C. M. (1995) Secretion of biologically active recombinant fibrinogen by yeast. J. Biol. Chem. 270, 23,761–23,767. 14. Denny, P. C., Denny, P. A., and Hong-Le, N. H. (1995) Asparagine-linked oligosaccharides in mouse mucin. Glycobiology 5, 589–597. 15. Qu, Z., Sharkey, R. M., Hansen, H. J., Goldenberg, D. M., and Leung, S.-O. (1997) Structure determination of N-linked oligosaccharides engineered at the CH1 domain of humanized LL2. Glycobiology 7, 803–809.

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16. Prieto, P. A., Mukerji, P., Kelder, B., Erney, R., Gonzalez, D., Yun, J. S., et al. (1995) Remodeling of mouse milk glycoconjugates by transgenic expression of a human glycosyltransferase. J. Biol. Chem. 270, 29,515–29,519. 17. Pirie-Shepherd, S., Jett, E. A., Andon, N. L., and Pizzo, S. V. (1995) Sialic acid content of plasminogen 2 glycoforms as a regulator of fibrinolytic activity. J. Biol. Chem. 270, 5877–5881. 18. Goss, P. E., Baptiste, J., Fernandes, B., Baker, M., and Dennis, J. W. (1994) A phase I study of swainsonine in patients with advanced malignancies. Cancer Res. 54, 1450–1457. 19. Sango, K., McDonald, M. P., Crawley, J. N., Mack, M. L., Tifft, C. J., Skop, E., et al. (1996) Mice lacking both subunits of lysosomal beta-hexosaminidase display gangliosidosis and mucopolysaccharidosis. Nature Genet. 14, 348–352. 20. Masada, R. I., Skop, E., and Starr, C. M. (1996) Fluorophore-assisted-carbohydrate-electrophoresis for quality control of recombinant protein glycosylation. Biotechnol. Appl. Biochem. 24, 195–205. 21. Kumar, H. P. M., Hague, C., Haley, T., Starr, C. M., Besman, M. J., Lundblad, R., and Baker, D. (1996) Elucidation of N-linked oligosaccharide structures of recombinant factor VII using fluorophore assisted carbohydrate electrophoresis. Biotechnol. Appl. Biochem. 24, 207–216. 22. Tsuchida, F., Tanaka, T., Matuo, Y., Moriyama, N., Okada, K., Jimbo, H., et al. (1997) Oligosaccharide Analysis Of Renal Cell Carcinoma by FluorophoreAssisted Carbohydrate Electrophoresis (FACE). Jpn J. Electroph. 41, 39–41, 81–83. 23. Hague, C., Masada, R. I., and Starr, C. (1998) Structural determination of oligosaccharides from recombinant iduronidase released with peptide N-glycase F, using Fluorophore-Assisted Carbohydrate Electrophoresis. Electrophoresis 19, 2612– 2620 24. Morimoto, K., Maeda, N., Foad, A., Toyoshima, S., and Hayakawa, T. (1999) Structural Characterization of Recombinant Human Erythropoietins by Fluorophoreassisted Carbohydrate Electrophoresis. Biol. Pharm. Bull. 22, 5–10. 25. Allen, A. C., Bailey, E. M., Barratt, J., Buck, K. S., and Feehally, J. (1999) Analysis of IgA1 O-glycans in IgA nephropathy by fluorophore-assisted carbohydrate electrophoresis. J. Am. Soc. Nephrol. 10, 1763–1771. 26. Bardor, M., Cabanes-Macheteau, M., Faye, L., and Lerouge, P. (2000) Monitoring the N-glycosylation of plant glycoproteins by fluorophore-assisted carbohydrate electrophoresis. Electrophoresis. 21, 2250–2256. 27. Calabro, A., Benavides, M., Tammi, M., Hascall, V. C., and Midura, R. J., (2000) Microanalysis of enzyme digests hyaluronan and chondroitin/dermatan sulfate by Fluorophore-Assisted Carbohydrate Electrophoresis (FACE). Glycohiology 10, 273–281. 28. Calabro, A. C., Hascall, V. J., and Midura, R., (2000) Structural characterization of oligosaccharides in recombinant soluble human interferon receptor 2 using Fluorophore-Assisted Carbohydrate Electrophoresis. Electrophoresis 21, 2296– 2308.

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29. Frado, L. Y. and Strickler, J. E. (2000) Relative abundance of oligosaccharides in candida species as determined by Fluorophore-Assisted Carbohydrate Electrophoresis. J. Clin. Microbiol. 38, 2862–2869. 31. Lemoine, J., Cabanes-Macheteau, M., Bardor, M., Michalaski, J. C., Faye, L., and Lerouge, P. (2000) Analysis of 8-aminoapthalene-1,3,6-trisulfonic acid labelled N-glycans by matrix assisted laser desorption/ionisation time-of-flight mass spectrometry. Rapid Commun Mass Spectrom, 14, 100–104.

143 HPLC Analysis of Fluorescently Labeled Glycans Tony Merry and Sviatlana Astrautsova

1. Introduction The study of the glycan chains (oligosaccharides) of glycoproteins presents a number of analytical problems that generally make their analysis more difficult than that of the peptides to which they are attached. A number of different analytical techniques have been applied to the study of glycans. Definitive characterization has traditionally been performed using nuclear magnetic resonance spectroscopy (NMR) and this still remains the only way to unequivocally assign the structure to novel glycans, but the techniques do require relatively large amounts of material (generally in the milligram range). In addition, access to the sophisticated equipment and the expertise required for the interpretation of data mean that the technique is available only to specialized dedicated laboratories. Another technique, which has also been applied extensively to studies of glycan structure, is that of mass spectrometry in various forms. A number of structures have been solved by means of fast atom bombardment (FABMS) (1,2) although generally further information from linkage analysis by gas chromatography-MS of permethylated alditol acetates (PMAA (3–5)) or by use of specific exoglycosidase enzymes (6) is required. Other techniques for mass spectrometric analysis are becoming more widely used as a result of recent developments in instrumentation (7–9). Both matrix-assisted laser desorption mass spectrometry with time-of-flight detection (8) (MALDI-TOF) and electrospray coupled to post source decay (9) to produce fragmentation ions are now becoming routinely used. The instrumentation is still expensive, however, and requires skilled operation and interpretation. Another technique that is also used is capillary electrophoresis, which can offer rapid characterization but that can also be difficult to optimize (10). Polyacrylamide gel electrophoresis of fluorescently labeled glycans, a technique From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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that requires no expensive instrumentation or specialized expertise, may also be used for screening or profiling. Although this is very useful for routine screening of a large number of samples, detailed characterization or sequencing of complex mixtures is problematic. High-performance liquid chromatography (HPLC) techniques with fluorescent detection offer a compromise in that they give the capability of full characterization of complex glycan mixtures relatively quickly, and although they do require good instrumentation this is considerably less expensive than mass spectrometry or NMR. HPLC is therefore the method of choice for routine analysis of glycosylation of protein, which have glycans of a type that have been previously characterized. Even when the glycan type is novel it is a technique that is compatible with others that can give further structural information, in particular mass spectrometry with LC-MS techniques now becoming fully integrated. The development of HPLC has presented several problems, which are unique to glycan analysis. In particular they generally do not possess a strong chromophore or fluorophore for detection. In addition they may not be charged and different glycans may have very similar composition and physicochemical properties. Therefore, their study has required the development of techniques for their derivatization and also for their separation, which differ from those used for peptides. When the glycans are released with a free reducing terminus, by either hydrazinolysis or endo-glycosidase, or endo-glycopeptidase enzymes, they may be derivatized by the relatively simple reaction of reductive animation (11). It is desirable that the derivatization is nonselective to achieve quantitative analysis and it should not cause structural changes such as desialylation. The incorporation of tritium into the C1 position of the reducing terminal monosaccharide fulfils these requirements most effectively, and many studies have been performed by this technique (12–14). This technique does, however, require the use of relatively large amounts or radioactivity and the use of scintillation counting for high sensitivity work and is not widely used now. Fluorescent labels may also be introduced into the C1 position by similar reductive amidation reactions and a number of these have now been described including 2-amino pyridine (15–17), 2-aminoacridone (18), 3-(acetylamino)-6-aminoacridine (AA-Ac) (19), 2-aminoanthanilic acid (20,21), and 2-aminobenzoic acid (2-AB) (20). A number of chromatographic systems have been developed for separation including gas–liquid chromatography (22) size-exclusion chromatography on polyacrylamide based beads, notably the BioGel P4 series (24,24), ion-exchange chromatography on standard matrices (25), and the development of specialized matrices for anion-exchange (high-performance anion-exchange chromatography [(HPAEC]) (26–29). They have been used in a large number of studies on protein glycosylation but these techniques have certain drawbacks. Lowpressure size-exclusion chromatography requires great care in column packing

HPLC Analysis of Fluorescently Labeled Glycans Monosaccharide composition

GLYCOPROTEIN

SDS-PAGE In gel release

I

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2-AB labeling

Anion Exchange

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Normal Phase

Reverse Phase

MALDI TOF

Enzymatic Sequencing

ESI MS/MS

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V

Fig. 1. The steps involved in complete analysis are shown diagrammatically. The Roman numerals relate to the stages given in the Introduction.

and the run times are long, HPAEC requires the use of high-pH and high-salt buffers that have to be removed before further analysis of separated glycans, such as exoglycosidase sequencing. The use of glycans labeled with a fluorescent tag such as 2-AB (20) and separated on suitable HPLC matrices (30,31) provides a convenient and sensitive means of profiling and sequencing of glycoprotein glycans (30,32,33) that can now be performed in many laboratories. The analysis of glycans by this technique proceeds in a number of stages: 1. 2. 3. 4. 5.

Release of glycans from the glycoprotein. 2-AB labelling of the glycans. Initial HPLC profiling. Enzymatic sequencing of glycans. Conformation of structures by mass spectrometry.

This approach is summarized in Fig. 1. The glycan release techniques may be adapted according to the source and purity of the proteins under study. 2. Materials 2.1. Equipment 1. HPLC system: High-pressure mixing two-solvent system capable of delivering 0.1–1.0 mL/min with shallow gradient (see Note 1). 2. Solvent degasser: Additional solvent degasser aids reproducibility.

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3. Detector fluorescent detector capable of excitation at λ330nm and emission at λ420nm (see Note 2). 4. Computer system for data collection and analysis. 5. Data acquisition software, for example, Waters Millenium™. 6. Curve-fitting software, Microsoft Excel™ or Proprietary software from Instrument Vendor (see Note 3).

2.2. Reagents 1. Anhydrous hydrazine was prepared by distillation from reagent grade hydrazine (Pierce Aldrich 21515-1) by mixture with calcium oxide and toluene as previously described (34) (see Note 4). This is available commercially from Ludger Ltd, Abingdon, UK 2. Acetic anhydride (ACS reagent, Sigma). 3. Anion-exchange resin: Bio-Rad AG50 X12. 4. Acetonitrile: HPLC grade with low background fluorescence such as E Chromosolv® (Reiedel-de-Haën, from Sigma). 5. Formic acid Aristar grade (BDH). 6. 26% Extra pure ammonia solution (Reiedel-de-Haëen, from Sigma). 7. Water MilliQ or equivalent, sub boiling point double distilled water (for mass spectrometric analysis). 8. Whatman no. 3 chromatography paper. 9. 2-Aminobenzamide labelling kit (see Note 5). 10. Pro-Mem Filter, 0.45 µm cellulose nitrate (R. B. Radley and Co. Ltd., Shire Hill, Saffron Walden, Essex, UK). 11. 3-([3-Cholamidopropyl)dimethylammonio)-1-propanesulfonate (CHAPS) detergent buffer: 50 mM ammonium formate, pH 8.6, 1.29. w/v CHAPS, 0.1 M EDTA.

2.3. Enzymes 1. “PNGase F” is available from Ludger Ltd, Abingdon, UK) PNGQA-BioTM (PNGase F Peptide-N4-(acetyl-ß-glucosaminyl)-asparagine amidaseN-Glycosidase F.) 2. Exoglycosidase enzymes

Sequencing grade enzymes were obtained from Prozyme (Palm Desert, California – distributed in Europe by Ludger Ltd (Abingdon) unless specified and were used with the buffers supplied as shown in Table 1. The following enzymes were used for exoglycosidase sequencing and given the abbreviations shown below along with their specificities. a) b) c) d) e) f)

ABS NDVS BTG SPG AMF BKF

α2–6 + 2,3 Specific sialidase (Arthrobacter ureafaciens). α2–3 Specific sialidase (Newcastle disease virus). β1–3 (>β1,4 or 6) Specific galactosidase (bovine testes). β1–4 Specific galactosidase (Streptococcus pneumoniae). α1–3 Specific fucosidase (almond meal). α1–6 (>α1,2 or 3 or 4) Specific fucosidase (bovine kidney).

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β1–2,3, 4 or 6 Specific hexosaminidase (GlcNAc or GalNAc, Jack Bean). β1,3 or 6 to Gal β1,2,3 or 6 to Man N-acetylglucosaminidase (Streptococcus pneumoniae). α1–2, 1–3, 1–6 Specific mannosidase (Jack Bean). β1–4 Specific mannosidase (Helix pomatia).

2.4. Standards 1. Partially hydrolyzed dextran (glucose oligomer standard) (Ludger Ltd, Abingdon UK). 2. Arabinose oligomers (monomer to octamer are available from Dextra Labs, Reading, UK). 3. 2-AB labeled glycan standards, all available from Ludger Ltd (Abingdon UK): a. Biantennary galactosylated. b. Biantennary galactosylated (with core 1,6-linked fucose). c. Biantennary galactosylated (with bisecting GlcNAc). d. Biantennary sialylated. e. Triantennary. f. Tetraantennary. g. Oligomannose (Man 5,6,7,8,9). h. Type 2 core O-glycan.

3. Methods 3.1. Glycan Release 3.1.1. Enzymatic Release with PNGaseF from Glycoproteins in Solution (Method from ref. 35 ) 1. PNGaseF solution at 1000 U/mL (Boehringer Mannheim). 2. Isolate the glycoprotein according to your usual procedures. 3. The sample should be relatively salt free and contain no extraneous carbohydrates (e.g., Sepharose-purified material contains large amounts of free carbohydrate that should be removed by dialysis using a 15–18,000 mol wt cut-off membrane) 4. If the volume of the glycoprotein solution required is >100 µL, dry the glycoprotein in a 1.5-mL microcentrifuge tube. Generally 50–200 µg of glycoprotein is required. 5. The incubation of a control glycoprotein with known glycosylation alongside experimental samples is recommended (see Note 6). 6. Proceed with the enzymatic digestion as described in Subheading 3.1.2. 7. Store remaining glycoprotein at 4°C for future use. 8. Dissolve sample in 50 µL of 50 mM ammonium formate, pH 8.6, 0.4% sodium dodecyl sulfate (SDS). 9. Incubate for 3 min at 100°C. 10. Cool and add 50 µL of CHAPS detergent buffer. 11. Add 1 U of PNGaseF (1 µL). 12. Incubate for 24 h at 37°C ( add 5 µL of toluene to prevent bacterial growth).

β1,3 or 4 GlcNAc or GalNAc

β1,3 4 GlcNAc

β-N-acetylglucosaminidase

β1,3 Gal

β1,3 or 4 Gal

α 2,3 sialic acid only

α2-3, 6, 8 sialic acid

Specificity

β-galactosidase Streptomyces Pneumoniae EC β-N acetylhexoseaminidase Recombinant

Sialidase Arthrobacter Ureafaciens EC Sialidase From Newcastle Disease virus Recombinant EC 3.2.1.26 β-galactosidase Bovine Testes EC

Enzyme

Table 1 Incubation Conditions for Exoglycosidase Enzymes

pH 6.0 100mM Na citrate/ phosphate

100mM Sodium Acetate pH 6.0 100mM Na citrate-phosphate

pH 5.0

100 mM citrate-phosphate

50mM sodium acetate pH 5.5 50mM sodium phosphate pH 6.0

Buffer

120µU/µl 16hrs pH 6.0

10µU/µl 16hrs

80µU/µl 16hrs

1mU/µl 3-16hrs

1µU/µl 16hrs

1µU/µl 3-16hrs

Concentration & Incubation Time*

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Man

β1-3,4 Gal in poly N-acetyl lactosamine

Jack Bean

Endo β-galactosidase Sodium Acetate pH 5.8

50mM

100 mM Na citrate pH 6.0 16hrs 100 mM sodium acetate, 2 mM Zn, pH 5.0

50 mM NaAc pH 5.0

3 hrs

Requires 2× 16hrs incubations 100µU/µl

67 mU/µ l

1mU/µl 16hrs 1mU/µl

It should be noted that for most of these exoglycosidases the enzyme concentration and the incubation times have not been optimised for the type of glycan or the amount used in studies with fluorescent glycans. The conditions are generally those which can be expected to give complete digestion of the particular linkages. In practice lower enzyme concentrations and shorter incubation times could often be used. It is advisable to check any enzymes against known standard glycans of different types before use in sequencing.

*

B. fragilis

α1-4, 6 Fuc α1–3, Fuc α1-2, 4

α-fucosidase Almond Meal α-fucosidase Bovine Kidney α-mannosidase

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13. Remove 5 µL and analyze the reaction mixture by SDS-polyacrylamide gel electrophoresis (SDS-PAGE). 14. If sample is completely deglycosylated proceed with step otherwise continue with incubation (see Note 8). 15. Filter samples through protein binding membrane or perform gel filtration. 16. Dry sample in vacuum centrifuge.

3.1.2. Enzymatic Release with PNGaseF from Glycoprotein Bands in SDS-PAGE Gels Suitable for analysis of low microgram amounts of protein or for unpurified proteins separated by SDS-PAGE or 2-dimensional electrophoresis. Method of Kuster et al. (36). See Royle et al. (37) for later modifications 1. Destain Coomassie Blue stained gels in 30% methanol–7.5% acetic acid. 2. Cut out gel pieces with band of interest using a washed scalpel blade, keeping pieces as small as possible. 3. Put into 1.5-mL tubes and wash with 1 mL of 20 mM NaHCO3, pH 7.0, twice using a rotating mixer. 4. Add 300 µL of NaHCO3, pH 7.0. 5. Add 20 µL 45 mM dithiothreitol (DTT). 6. Incubate at 60°C for 30 min. 7. Cool to room temperature and add 20 µL of 100 mM iodoacetamide. 8. Incubate for 30 min at room temperature in the dark. 9. Add 5 mL of acetonitrile–20 mM NaHCO3, pH 7.0. 10. Incubate for 60 min to wash out reducing agents and SDS. 11. Cut gel into pieces of 1 mm2. 12. Place in a vacuum centrifuge to dry. 13. Prepare PNGaseF solution at 1000 U/mL (Boehringer Mannheim): Add 30 U 30 µL of PNGaseF in 20 mM NaHCO3, pH 7.0. 14. Allow gel to swell and then add a further 100-µL aliquot of buffer. 15. Incubate at 37°C for 12–16 h.

3.1.3. Release by Hydrazinolysis Suitable for analysis of N- or O-linked glycans where the amount of protein is limited, where steric hindrance to enzymatic release is known, or where selective release of glycans by enzymatic means is suspected. 3.1.3.1. PREPARATION oF SAMPLES FOR HYDRAZINOLYSIS

Desalt and dry the samples completely as follows (see Note 8). 1. Dissolve the sample in 0.1% trifluorocetic acid (TFA) in as small a volume as possible. 2. Set up dialysis at 4°C. A microdialysis system is recommended. 3. Dialyze for a minimum of 48 h in a microdialysis apparatus fitted with a 6000–8000-Dalton cut-off membrane.

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4. Recover sample from dialysis membrane. Wash membrane with 0.1% TFA to ensure recovery. 5. Transfer to a suitable tube for hydrazinolysis (see Note 9). 6. Lyophilize sample for 48 h. 7. For O-glycan analysis further drying is recommended (see Note 10). 8. Remove sample from lyophilizer immediately prior to addition of hydrazine. 3.1.3.2. HYDRAZINOLYSIS PROCEDURE

Suitable for analysis of N- and O-linked glycans when expertise and equipment for the procedure are available (34). 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29.

Remove tubes from drying immediately prior to hydrazine addition. Flush tube with argon, taking care not to dislodge lyophilized protein. Rinse dried syringe fitted with stainless steel needle with anhydrous hydrazine. Take up fresh hydrazine and dispense onto sample. A 0.1-mL volume of hydrazine is sufficient to dissolve up to 2 mg of glycoprotein. For larger amounts add more hydrazine. Flame seal the tube. (note reaction may be performed in suitable air tight vials – e.g. Gently shake tube—the protein should dissolve. Place in an incubator (do not use water bath). For release of N-linked glycans incubate at 95°C for 5 h; for O-glycan release incubate at 60°C for 6 h. Allow to cool and remove hydrazine by evaporation. Add 250 µL of toluene and evaporate. Repeat 5×. Place tube on ice and add 100 µL of saturated sodium bicarbonate solution. Add 20 µL of acetic anhydride. Mix gently and leave at 4°C for 10 min. Add a further 20 µL acetic anhydride. Incubate at room temperature for 50 min. Pass solution through a column of Dowex AG50 X12 (H+ form)—0.5 mL bed volume. Wash tube with 4 × 0.5 mL of water and pass through a Dowex column. Evaporate solution to dryness. This should be done in stages by redissolving in decreasing volumes of water. Prepare 50 × 2.5 cm strips of Whatman no. 1 chromatography paper (prewashed in water by descending chromatography for 2 d). Spot sample on strip. Perform descending paper chromatography for 2 d in 4:4:1 by vol butanol–ethanol– water for N-glycans and 8:2:1 1 by vol butanol–ethanol–water for O-glycans. Remove strip from tank and allow all traces of solvent to evaporate. Cut out region of strip from −2 cm to +5 cm from application. Roll up cut chromatography paper and place in a 2.5-mL nonlubricated syringe. Fit a 0.45 µm PTFE filter to syringe. Add 0.5 mL of water and allow to soak into paper for 15 min. Fit syringe plunger and force solution through filter. Wash filter with water 4×, and pass through filter. Evaporate sample to dryness, dissolve in 50 µL of water, transfer to microcentrifuge tubes, and store at −20°C until required.

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3.2. Glycan Labeling Fluorescent labelling with 2-aminobenzamide was performed as described by Bigge et al. (20) using the kit provided by Ludger Ltd.(Abingdon, UK). The procedure is as follows: 1. Take an aqueous solution of glycan that should contain a minimum of 0.1 pmol and a maximum of 50 pmol of glycans. The volume should be no more than 50 µL. 2. Place in a 0.5-mL microcentrifuge tube and dry down in centrifugal evaporator. 3. Prepare labelling solvent by addition of 150 µL of glacial acetic acid to 100 µL of dimethyl sulfoxide (DMSO). Add 100 µL of the acidified dimethyl sulfoxide solvent to 2-aminobenzamide to make a 0.25 M solution. Mix well to dissolve the 2-AB (may require gentle warming). 4. Add all this solution to sodium cyanoborohydride to make a 1.0 M solution. 5. Mix well for 5 min to dissolve the reductant. 6. Add 5 µL of the labelling reagent to the dried sample. 7. Mix well and centrifuge briefly. 8. Incubate at 65°C for 1 h and mix well. Incubate for a further 2 h. 9. Cool the labelling mixture on ice. 10. Remove free label by either technique given in Subheadings 3.2.1 and 3.2.2.

3.2.1. Removal of Free 2-AB Label by Ascending Paper Chromatography 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Cool the 2-AB reaction mixture in freezer. Apply all 5 µL of sample to a point in centre of strip 1 cm from the bottom. Allow to dry for 2 h. Perform ascending chromatography for 1 h in acetonitrile. Examine strip under UV to see if the spot of free dye has migrated to the top of the strip. If it is not at the top continue chromatography until it is. Dry the paper completely. Cut out the origin on the strip. Place in a 2.5-mL syringe fitted with a 0.45-µm PTFE filter. Apply 0.5 mL of water and leave for 15 min. Push water through the filter. Wash twice with a further 0.5 mL of water. Dry down labeled sample in a vacuum centrifuge.

3.2.2. Removal of 2-AB Label on Filter Disks 1. Before use wash Whatman no. 1 filter paper in 500 mL of MilliQ water. Place filter paper in a beaker and add water. Leave for 15 min at room temperature. Decant water. Repeat this 4 times.

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2. Dry paper in a 65°C over for 2 h. The paper may be stored at room temperature after this step. 3. Cut 6-mm discs from washed paper. 4. Place discs in a Bio-Rad disposable column. For a sample of < 10 µg of glycoprotein use two discs; for more use five discs). 5. Add 2 mL of water and leave for 5 min with columns capped. 6. Uncap columns and wash with another 4 × 2 mL of water (see Note 11). 7. Wash with a further 5 × 2 mL of 30% acetic in water (see Note 11). 8. Cap column and add 2 mL of acetonitrile. 9. Leave for 5 min and uncap the column. 10. Wash with a further 2 mL of acetonitrile just before applying sample; cap column. 11. Remove incubation vial and put in freezer for 5 min to cool down. 12. Centrifuge the tube and spot all the sample onto the disc. 13. Leave for 15 min and rinse tube with 100 µL of acetonitrile and add to disc. Leave for 5 min. 14. Add 2 mL of acetonitrile—uncap column and discard wash. 15. Wash with a further 4 × 2 mL of acetonitrile. 16. Place syringes (2.5-mL Fortuna) fitted with a 0.45-µm filter and stoppers, under the filter. 17. Elute glycans with 4 × 0.25 mL of water (see Note 11). 18. Dry sample down for analysis.

Note: A kit employing a proprietary separation membrane for separation of free 2-AB from glycans (LudgerCleanTM EB Cartridges) is now available from Ludger Ltd (Abingdon, UK)

3.3. Normal Phase Chromatography The following conditions are recommended: 1. Buffers:

Ammonium formate: 50 mM formic acid adjusted to pH 4.4 with ammonia solution. Acetonitrile: HPLC grade; Column: GlycoSep N™ column (Ludger Ltd) or TokoHas TSK-amide 80. Gradient: Sample loading: The sample should be loaded in 80% acetonitrile–20% water (v/v). In practice take 20 µL of the sample in water and add 80 µL of acetonitrile. b. Inject 95 µL of the sample. c. A standard dextran should be included with all sample runs. d. Fig. 2 shows a typical separation of human serum IgG with dextran ladder calibration. 2. 3. 4. a.

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Fig. 2. Analysis of IgG N-glycans released by hydrazinolysis on normal phase HPLC. The top panel shows the calibration curve for conversion of retention time into glucose units based on a third order polynomial. The middle panel shows the separations of glucose oligomers from the dextran ladder. The lower panel shows the separation of the fluorescently labeled glycans from IgG with structures corresponding to the major peaks indicated. Software (GlycoBase) for this purpose is now available on the Internet as part of the EuroCarbDB project (http://www.eurocarbdb.org/) (see Note 3).

HPLC Analysis of Fluorescently Labeled Glycans a. Startup method run time, 30 min.

1301

b. Separating method run time, 180 min

Time (min)

Flow (ml/min)

A (%)

B (%)

Time (min)

Flow (ml/min)

A (%)

B (%)

0 04 08 13 16 25 26 60 61

0 1.4 1.4 1.4 1.4 1.4 0.4 0.4 0

20 20 95 95 20 20 20 20 20

80 80 05 05 80 80 80 80 80

0 152 155 157 162 163 177 178 260

0.4 0.4 0.4 10. 10. 10. 10. 0.4 0

20 058 100 100 100 020 020 020 2200

80 42 00 00 00 80 80 80 80

5. Data analysis: a. The elution times of all peaks in the dextran ladder should be recorded. b. To assign glucose unit values a polynomial fit should be applied to the data to generate a standard curve. A third-order polynomial will generally give a good fit (see Fig. 2). c. The glucose unit values of sample peaks may then be calculated (see also Note 3). d. Published values of glucose unit values for a wide series of glycans may be used to give an indication of possible structures present. These are now available on the Internet as part of the GlycoBase software developed by NIBRT, Dublin (http:// glycobase.ucd.ie/cgi-bin/public/glycobase.cgi) for the EUROCarbDB project (http://www.eurocarbdb.org/)

3.4. Reverse-Phase Chromatography 1. Buffers: a. ammonium formate–triethylamine 50 mM formic acid adjusted to pH 5.0 with triethylamine. b. Acetonitrile: HPLC grade; see Subheading 2.2. 2. Column: Reverse-phase column GlycoSep™ R—Ludger Ltd (Abingdon, UK) or equivalent. 3. Sample loading:

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a. Startup method run time, 30 min

b. Separating method run time, 180 min

Time

Flow

A

B

Time

Flow

A

B

00 05 10 20 21 28 29 90 91

0.05 100. 100. 100. 100. 100. 0.50 0.50 00.0

05 05 05 05 95 95 95 95 95

95 95 95 95 05 05 05 05 05

000 030 160 165 166 172 173 178 179 220 221

0.5 0.5 0.5 0.5 1.5 1.5 1.5 1.5 0.5 0.5

95 95 85 76 05 05 95 95 95 95 0

05 05 15 24 95 95 05 05 05 05 95

5

a. The sample should be loaded as a aqueous solution. A volume of 95 µL of sample should be loaded. b. A standard of an arabinose oligomers (available from Dextra Labs, Reading) should be included with all sample runs. 4. Data analysis: a. The data may be analyzed in a similar way to that of normal phase but using the values for the arabinose ladder. b. Typical values for the AU units for a number of structures are available (37). c. It can often be helpful to compare normal and reverse-phase profiles for the same glycans as shown in Fig. 3. The different principles for separation are illustrated by the differences in separation on the two systems. By combining both types of analysis all four structures can readily be identified, for example, the separation of A2G2F and A2G2FB.

3.5. Weak Anion-Exchange Chromatography (WAX)

Fig. 3. Analysis of fetuin on Weak Anion Exchange Chromatography. The separation of glycans possessing from 1 to 5 sialic acids is shown.

HPLC Analysis of Fluorescently Labeled Glycans 1. a. b. 2. 3. a.

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Buffers: 500 mM Formic acid adjusted to pH 9.0 with ammonia solution. Methanol water: 10:90 (v/v). Gradient Sample loading The sample should be loaded as an aqueous solution. A volume of 95 µL of sample should be loaded. a.Startup method run time, 30 min. Separating method run time, 180 min Time 000 05 20 21 28 29 90 91

Flow 0 10. 10. 10. 10. 0.5 0.5 00.

A

B

0 05 05 95 95 95 95 95

0 95 95 05 05 05 05 05

Time 0 12 50 55 65 66 77 78

Flow 1 1 1 1 1 2 2 1

A

B

00 005 080 100 100 000 000 000

100 095 020 000 000 100 100 100

b. No suitable standards are currently available although it is useful to run a well characterized glycoprotein such as bovine serum fetuin (Sigma). c. The glycans generally elute on the basis of charge although the size of the glycan also contributes to the position of elution. d. A typical separation of charged N-glycans from bovine serum fetuin is shown in Fig. 3.

3.6. Exoglycosidase Digestion of N-Glycans Digestions are generally performed on the pool of glycans by the application of exoglycosidases under the incubation conditions shown in Table 1 in a series of enzyme arrays of increasing complexity as shown in Table 2. 1. The amount required for each digestion can be judged from the previous profiling run, but in general 100 fmol of glycan is detectable. In practice more may Table 2 Typical Arrays for N-Glycan Sequencing Array

Vol enzyme

Vol buffer

Water

ABS ABS BTG ABS BTG BKF ABS BTG BKF AMF ABS BTG BKF AMF JBH JBM (high) JBM (low)

1 12 121 1211 12112 10 0.8

2 2 2 2 2

7 5 4 3 1 9.2

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Merry and Astrautsova

be used with the enzyme concentrations given below if sufficient material is available. Pipette no more than 50 µL of solution into a 0.5-mL microcentrifuge tube. Evaporate glycan solutions to dryness in a vacuum centrifuge. Add 2–5 µL of enzyme solution and 2 µL of incubation buffer. Make the total volume up to 10 µL with water. Incubate for 16–24 h at 37°C. Cool and load the digestion mixture onto a protein binding filter (Microspin 45). Leave for 15 min at room temperature.

Fig. 4. Exoglycosidase digestion of IgG glycans monitored on normal phase column normal phase HPLC. Exoglycosidase abbreviations ABS − α 2,6 + α 2,3 specific sialidase (Arthrobacter Ureafaciens), BTG 1,2 (+ α 1–4) specific galactosidase (Bovine Testes), AMF- α 2,3 specific fucosidase (Almond Meal) SPH- β1,3 (4,6) specific hexosaminidase (Streptococcus Pneumonae) JBM-α-mannosidase (Jack Bean).

HPLC Analysis of Fluorescently Labeled Glycans

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9. 10. 11. 12. 13. 14.

Centrifuge for 15 min at 5000g. Wash tube with 10 µL of freshly prepared 5% acetonitrile in water and load onto filter. Leave for 15 min at room temperature. Centrifuge for 15 min at 5000g. Repeat steps 10–12. Twenty microliters of sample may be directly analyzed by HPLC if sufficient material is available; if not, then dry down sample and redissolve in 20 µL of water. 15. Example of digestion of N-glycans from IgG is shown in Fig. 4.

3.7. Exoglycosidase Digestion of O-Glycans Digestions are generally performed on the pool of glycans; however, the strategy differs from that for N-glycans. Digestions should be performed by the following mixtures of glycans where the choice of glycosidases is dictated by the outcome of previous digestions. Since several O-glycans may co elute on any matrix the analysis on both normal and reverse phase is recommended. Incubation conditions recommended for common exoglycosidases are shown in Table 1. A detailed description of the procedure and reference values for digestion products may be found at the following web site; http://glycobase.ucd.ie/ 1. 2. 3. 4. 5. 6.

7. 8. 9. 10. 11. 12.

The samples are prepared for digestion as described in steps 1–3 (see Subheading 3.6). First perform ABS digestion to detect presence of sialic acid (in any linkage). Perform ABS + BTG digestion to show the presence of galactose. Perform ABS + BKF digestion to show the presence of fucose. Perform ABS + JBH digestion to show presence of N-acetylglucosamine or N-acetylgalac-tosamine. In each case digestion will be apparent from a shift in peaks. If the digests are analyzed by normal phase and reverse phase then possible identities of structures may be found from the GU and AU values. If digestion with ABS occurs then also digest pool with NDVS to determine if the sialic acid linkage is 2,3. If digestion with ABS + BTG occurs then digest with ABS + SPG to determine if linkage of galactose is 1,4. If digestion occurs with ABS + BKF then digest with ABS + AMF to determine if fucose linkage is 1,3 or 1,4. If digestion with ABS + JBH occurs then digest with ABS + SPH to determine if N-acetylglucosamine or N-acetylgalactosamine is present. Note that the complete characterization may require repeated digestions as there is no common core sequence as found for N-glycans. Example of O-glycan sequencing is shown Fig. 5.

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Fig. 5. Analysis of bi-anennary, fucosylated, bisected glycans by reverse phase HPLC – comparison with normal phase HPLC showing complimentarily of separation techniques.

3.7.1. Data Analysis (software for this analysis is now available as pert of the open access EUROCarbDB project (http://www.eurocarbdb.org/). 1. The movement of a peak in an exoglycosidase incubation indicates that it contains the monosaccharides in the linkages removed by that enzyme. 2. A shift in the position of peaks will be seen with decreasing complexity of the chromatogram with increase in the number of enzymes applied. 3. Reference to the last chromatogram in a series will allow identification of the basic core N-glycan structure as shown in Fig. 4. 4. The last digest in a series will indicate the core glycans present. 5. Working back through the digests, the structures from which a glycan has been removed are identified. a. In the example the presence of terminal N-acetylglucosamine in the structure is shown comparing digest 5 to 4. b. The presence of terminal galactose is shown by comparing digest 3 to 4. c. The presence of sialic acid is shown by comparing digest 3 to 1. d. In this way the structure shown can be assigned to peak A. 6. The presence of other peaks indicates that monosaccharides not covered by the array are present (such as oligomannose) or that residues are substituted by groups such as sulfate or phosphate. 7. An examples of O-glycan sequencing is shown in Fig. 6 with sequential application of glycosidases.

HPLC Analysis of Fluorescently Labeled Glycans

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Fig. 6. Example of exoglycosidase sequencing an O-glycan on normal phase HPLC. SPG -β1,4 specific galactosidase (Streptococcus pneumoniae), other exoglycosidse abbreviations as in Fig. 4.

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3.8. Mass Spectrometry Samples may be further analyzed by mass spectrometry (MALDI-TOF or ESI/ MS/MS) for confirmation of structures but generally require further desalting although online systems are now available and are particularly suitable for this kind of analysis. A recent International Study by HUPO (Human Disease Glycolic/Proteome Initiative) compared various techniques for HPLC analysis and showed that the combination of HPLC and Mass Spectrometry allowed sensitive and accurate analysis of N-glycans (38). This study is now being extended to O-glycans. The tools developed by the EuroCarbDB are particularly recommended 1. It is often useful to perform the MS analysis on the glycan pool alongside the HPLC analysis but approx 10× more material is generally required. 2. As all solvents used are volatile, fractions containing peaks from the HPLC run may be collected and dried in a vacuum centrifuge but cleanup may be required (see Note 12).

4. Notes 1. Waters System is recommended. Hewlett-Packard is also suitable. System with similar specification from other manufacturers may be suitable but must be capable of high-pressure mixing of acetonitrile and aqueous solvents. 2. Jasco or Waters is recommended but other detectors of similar specificity may be suitable. 3. Glycobase is a list of GU values and associated software for analysis of exoglycosidase digestion of glycans that was originally developed at the Oxford Glycobiology Institute and subsequently at NIBRT, Dublin, as part of the EuroCarbDB project. It will form one of the databases available at www.eurocarbdb.org/. 4. Great care should be taking in the handling of hydrazine. Dry hydrazine of suitable quality may be obtained from Ludger Ltd. (Abingdon, UK). 5. Available from Ludger Ltd. (Abingdon, UK). 6. Suitable glycoproteins include ribonuclease B, bovine serum fetuin, and human serum haptoglobulin. 7. The change in position of a band on SDS-PAGE indicates N-glycosylation. In some cases incomplete digestion may lead to the production of a series of bands corresponding to partially deglycosylated glycoprotein. Complete release is indicated by conversion to a single band at the lowest molecular weight. 8. The presence of most salts, dyes, or detergents will interfere with hydrazinolysis. The sample must also be completely anhydrous. 9. All glassware used should be soaked in 4 M nitric acid for 4 h and thoroughly washed in water before use.

HPLC Analysis of Fluorescently Labeled Glycans

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10. The use of cryogenic drying by placing under high vacuum connected to a trap of charcoal submerged in liquid nitrogen for a period of 72 h as described by Ashford et al. (34) is recommended. 11. It may be necessary to apply slight pressure to the top of the column or to use a suitable vacuum apparatus. 12. Double-distilled (sub boiling point) water should be used for mass spectrometric analysis. Suitable techniques for post-HPLC cleanup and for mass spectrometry by electrospray (8) and by MALDI (9) have been described recently. Acknowledgments Professor Sviatlana Astrautsova is at the Department of Microbiology, Medical University of Grodno, Belarus. The advice and help of the former members of the Glycoimmunology group at the Oxford Glycobiology Institute, Director Professor R. A. Dwek and Group Leader Prof. Pauline Rudd (now at NIBRT, Dublin), is gratefully acknowledged. In particular, I would like to thank Dr. Louise Royle (now at Ludger Ltd Abingdon, UK) and Dr Catherine Radcliffe for their valued help and comments. Thanks also to Dr Max Crispin (STRUBI, Oxford) for data in Fig. 4 and for his helpful comments on the manuscript. The techniques described here are the result of many years of development by previous members of the Oxford Glycobiology Institute, and in particular Dr David Harvey for his mass spectrometry expertise, Dr. Geoffrey Guile for his HPLC expertise and Dr. Taj Mattu who were all involved in the development of the protocols described in this chapter. Although the group no longer exists at the Glycobiology Institute in Oxford Prof Rudd is continuing development and in particular training all the procedures detailed here at the NIBRT Dublin and the EuroGlycosciences Forum (www.glycosciences.eu) is promoting training in this area.

References 1. Dell, A., Thomas Oates, J. E., Rogers, M. E., and Tiller, P. R. (1988) Novel fast atom bombardment mass spectrometric procedures for glycoprotein analysis. Bio chimie 70, 1435–1444. 2. Carr, S. A. and Roberts, G. D. (1986) Carbohydrate mapping by mass spectrometry: a novel method for identifying attachment sites of Asn-linked sugars in glycoproteins. Analyt. Biochem. 157, 396–406. 3. Karlsson, H., Karlsson, N., and Hansson, G. C. (1994) High-temperature gas chromatography and gas chromatography-mass spectrometry of glycoprotein and glycosphingolipid oligosaccharides. Mol. Biotechnol. 1, 165–180. 4. Anumula, K. R. and Taylor, P. B. (1992) A comprehensive procedure for preparation of partially methylated alditol acetates from glycoprotein carbohydrates. Analyt. Biochem. 203, 101–108. 5. Cumming, D. A., Hellerqvist, C. G., Harris Brandts, M., Michnick, S. W., Carver, J. P., and Bendiak, B. (1989) Structures of asparagine-linked oligosaccharides of

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6.

7. 8. 9.

10.

11.

12. 13.

14.

15.

16.

17.

18.

19.

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the glycoprotein fetuin having sialic acid linked to N-acetylglucosamine. Biochemistry 28, 6500–6512. Edge, C. J., Rademacher, T. W., Wormald, M. R., Parekh, R. B., Butters, T. D., Wing, D. R., and Dwek, R. A. (1992) Fast sequencing of oligosaccharides: the reagent-array analysis method. Proc. Natl. Acad. Sci. USA 89, 6338–6342. Dwek, R. A., Edge, C. J., Harvey, D. J., Wormald, M. R., and Parekh, R. B. (1993) Analysis of glycoprotein-associated oligosaccharides. Annu. Rev. Biochem. 62, 65–100. Harvey, D. J. (1999) Matrix-assisted laser desorption/ionization mass spectrometry of carbohydrates. Mass Spectrom. Rev. 18, 349–450. Harvey, D. J. (2000) Electrospray mass spectrometry and fragmentation of N-linked carbohydrates derivatized at the reducing terminus. J. Am. Soc. Mass Spectrom. 11, 900–915. Taverna, M., Tran, N. T., Merry, T., Horvath, E., and Ferrier, D. (1998) Electrophoretic methods for process monitoring and the quality assessment of recombinant glycoproteins. Electrophoresis 19, 2572–2594. Hase, S., Ikenaka, T., and Matsushima, Y. (1979) Analyses of oligosaccharides by tagging the reducing end with a fluorescent compound. I. Application to glycoproteins. J. Biochem. Tokyo 85, 989–994. Takasaki, S. and Kobata, A. (1978) Microdetermination of sugar composition by radioisotope labelling. Meth. Enzymol. 50, 50–54. Takasaki, S., Mizuochi, T., and Kobata, A. (1982) Hydrazinolysis of asparaginelinked sugar chains to produce free oligosaccharides. Meth. Enzymol. 83, 263– 268. Endo, T., Amano, J., Berger, E. G., and Kobata, A. (1986) Structure identification of the complex-type, asparagine-linked sugar chains of beta D-galactosyl-transferase purified from human milk. Carbohydr. Res. 150, 241–263. Takahashi, N., Hitotsuya, H., Hanzawa, H., Arata, Y., and Kurihara, Y. (1990) Structural study of asparagine-linked oligosaccharide moiety of taste-modifying protein, miraculin. J. Biol. Chem.265, 7793–7798. Nakagawa, H., Kawamura, Y., Kato, K., Shimada, I., Arata, Y., and Takahashi, N. (1995) Identification of neutral and sialyl N-linked oligosaccharide structures. Analyt. Biochem. 226, 130–138. Rice, K. G., Takahashi, N., Namiki, Y., Tran, A. D., Lisi, P. J., and Lee, Y. C. (1992) Quantitative mapping of the N-linked sialyloligosaccharides of recombinant erythropoietin: combination of direct high-performance anion-exchange chromatography and 2-aminopyridine derivatization. Analyt. Biochem. 206, 278–287. Camilleri, P., Harland, G. B., and Okafo, G. (1995) High resolution and rapid analysis of branched oligosaccharides by capillary electrophoresis. Analyt. Biochem. 230, 115–122. Charlwood, J., Birrell, H., Gribble, A., Burdes, V., Tolson, D., and Camilleri, P. (2000) A probe for the versatile analysis and characterization of N-linked oligosaccharides. Analyt. Chem. 72, 1453–1461. Bigge, J. C., Patel, T. P., Bruce, J. A., Goulding, P. N., Charles, S. M., and Parekh, R. B. (1995) Nonselective and efficient fluorescent labelling of glycans using 2-amino benzamide and anthranilic acid. Analyt. Biochem. 230, 229–238.

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21. Anumula, K. R. and Dhume, S. T. (1998) High resolution and high sensitivity methods for oligosaccharide mapping. Glycobiology 8, 685–694. 22. Endo, Y., Yamashita, K., Han, Y. N., Iwanaga, S., and Kobata, A. (1977) The carbohydrate structure of a glycopeptide release by the action of plasma kallikrein on bovine plasma high-molecular-weight kininogen. J. Biochem. Tokyo 82, 545–550. 23. Kobata, A., Yamashita, K., and Tachibana, Y. (1978) Oligosaccharides from human milk. Meth. Enzymol. 50, 216–220. 24. Kobata, A., Yamashita, K., and Takasaki, S. (1987) BioGel P-4 column chromatography of oligosaccharides: effective size of oligosaccharides expressed in glucose units. Meth. Enzymol. 138, 84–94. 25. Takamoto, M., Endo, T., Isemura, M., Kochibe, N., and Kobata, A. (1989) Structures of asparagine-linked oligosaccharides of human placental fibronectin. J. Biochem. Tokyo 105, 742–750. 26. Field, M., Papac, D., and Jones, A. (1996) The use of high-performance anionexchange chromatography and matrix-assisted laser desorption/ionization timeof-flight mass spectrometry to monitor and identify oligosaccharide degradation. Analyt. Biochem. 239, 92–98. 27. Townsend, R. R. and Hardy, M. R. (1991) Analysis of glycoprotein oligosaccharides using high-pH anion exchange chromatography. Glycobiology 1, 139–147. 28. Martens, D. A. and Frankenberger, W. T., Jr. (1991) Determination of saccharides in biological materials by high-performance anion-exchange chromatography with pulsed amperometric detection. J. Chromatogr. 546, 297–309. 29. Rohrer, J. S. and Avdalovic, N. (1996) Separation of human serum transferring isoforms by high-performance pellicular anion-exchange chromatography. Protein Exp. Purif. 7, 39–44. 30. Guile, G. R., Rudd, P. M., Wing, D. R., Prime, S. B., and Dwek, R. A. (1996) A rapid high-resolution high-performance liquid chromatographic method for separating glycan mixtures and analyzing oligosaccharide profiles. Analyt. Biochem. 240, 210–226. 31. Guile, G. R., Harvey, D. J., O’Donnell, N., Powell, A. K., Hunter, A. P., Zamze, S., et al. (1998) Identification of highly fucosylated N-linked oligosaccharides from the human parotid gland. Eur. J. Biochem. 258, 623–656. 32. Rudd, P. M., Mattu, T. S., Zitzmann, N., Mehta, A., Colominas, C., Hart, E., et al. (1999) Glycoproteins: rapid sequencing technology for N-linked and GPI anchor glycans. Biotechnol. Genet. Eng. Rev. 16, 1–21. 33. Rudd, P. M., Guile, G. R., Kuster, B., Harvey, D. J., Opdenakker, G., and Dwek, R. A. (1997) Oligosaccharide sequencing technology. Nature 388, 205–7. 34. Ashford, D., Dwek, R. A., Welply, J. K., Amatayakul, S., Homans, S. W., Lis, H., et al. (1987) The beta 1→2-D-xylose and alpha 1→3-L-fucose substituted N-linked oligosaccharides from Erythrina cristagalli lectin. Isolation, characterisation and comparison with other legume lectins. Eur. J. Biochem. 166, 311–320. 35. Goodarzi, M. T. and Turner, G. A. (1998) Reproducible and sensitive determination of charged oligosaccharides from haptoglobin by PNGase F digestion and HPAEC/PAD analysis: glycan composition varies with disease. Glycoconj. J. 15, 469–475.

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36. Kuster, B., Hunter, A. P., Wheeler, S. F., Dwek, R. A., and Harvey, D. J. (1998) Structural determination of N-linked carbohydrates by matrix-assisted laser desorption/ ionization-mass spectrometry following enzymatic release within sodium dodecyl sulphate-polyacrylamide electrophoresis gels: application to species-specific glycosylation of alpha1-acid glycoprotein. Electrophoresis 19, 1950–1959. 37. Royle, L., Campbell, M. P., Radcliffe, C. M., White, D. M., Harvey, D. J., Abrahams, J. L., Kim, Y. G., Henry, G. W., Shadick, N. A., Weinblatt, M. E., Lee, D. M., Rudd, P. M., Dwek, R. A. (2008) HPLC-based analysis of serum N-glycans on a 96-well plate platform with dedicated database software. Anal. Biochem. 376, 1–12. 38. Wada Y, et al (2007) Comparison of the methods for profiling glycoprotein glycans–HUPO Human Disease Glycomics/Proteome Initiative multi-institutional study. Glycobiology 17, 411–422.

144 Glycoprofiling Purified Glycoproteins Using Surface Plasmon Resonance Angeliki Fotinopoulou and Graham A. Turner 1. Introduction 1.1. Glycosylation and Its Investigation The carbohydrate part of glycoproteins can define several of their biological properties, including the clearance rate, immunogenicity, thermal stability, solubility, the specific activity, and conformation (1). Often, small differences in the composition of the sugar side chains of a glycoprotein can affect the biological properties (2). Characterization of oligosaccharide structures on glycoproteins is essential for glycoprotein therapeutic products, because inflammatory and systemic responses may arise if glycosylation of the product is different from that of the native substance. Determination of the precise oligosaccharide profile of a glycoprotein, however, cannot yet be considered as a routine laboratory task. Methods for glycosylation analysis can be generally divided in two major categories, direct methods and methods using lectins. This chapter describes a recently developed method that measures the binding of lectins to glycoproteins using surface plasmon resonance (SPR). Lectins are a class of proteins that bind to carbohydrates in a noncovalent reversible way that does not chemically modify the sugar molecule (3). The lectin specificity is not absolute, but usually there is one carbohydrate grouping to which a lectin binds with much higher affinity than to other carbohydrate structures. This property makes them ideal for recognizing oligosaccharide structures. Although lectin methods are indirect, they can be used when comparisons are needed. They provide many advantages compared to other methods for investigating carbohydrate structures. There are a large number of lectins available with different and distinct specificities. Lectin methods are also quick and simple (4). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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1.2. Surface Plasmon Resonance and Lectins SPR is an optical sensing phenomenon that allows one to monitor biomolecule interactions in real time (Fig. 1). The sensor device is composed of a sensor chip consisting of three layers (glass, a thin gold film, and a carboxymethylated dextran matrix); a prism placed on the glass surface of the chip; and a microfluidic cartridge, which controls the delivery of liquid to the sensor chip surface (5). When light illuminates the thin gold film, energy is transferred to the electrons in the metal surface causing the reflected light to have reduced intensity at a specific incident angle. This angle of nonreflectance changes as the refractive index in the vicinity of the metal surface changes. The refractive index depends upon the mass on the surface of the gold film. A response of 1000 resonance units (RUs) corresponds to a shift of 0.1° in the resonance angle and represents a change in the surface protein concentration of about 1 ng/mm2 (6). Because all proteins, independent of sequence, contribute the same refractive index, SPR can be used as a mass detector. A glycoprotein, therefore, can be immobilized onto a sensor chip surface and then be probed by a panel of lectins. The binding or nonbinding of the lectins provides information about the oligosaccharide structures found on the carbohydrate chains. For example, if a glycoprotein has a trimannose core on its N-linked chains, it will bind to the lectin concanavalin A (Con A), or if it has terminal α2–3 or α2–6 N-acetylneuraminic acid (Neu5NAc) on its oligosaccharide chains it will bind to the Neu5NAc specific lectins Maackia amurensis agglutinin (MAA) or Sambucus nigra agglutinin (SNA), respectively. Lectin binding can also be performed after treatment with glycosidases to gain more information about the oligosaccharide structures present. By sequentially treating the immobilized glycoprotein with glycosidases and measuring

Fig. 1. SPR detects changes in the angle of nonreflectance. This angle changes if the refractive index of the chip surface changes, for example, when a protein is immobilized on the chip. (A) A glycoprotein is immobilized resulting in angle 1; (B) a lectin is bound to the immobilized glycoprotein resulting in angle 2.

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the binding of a panel of lectins after every treatment it is possible to gain information on the oligosaccharide sequence. Some lectins will bind to the immobilized glycoprotein only if they are pretreated with sialidase to remove the terminal Neu5NAc. An example of this is peanut agglutinin (PNA), which binds to Galactose β1–3 N-acetylgalactosamine groupings on O-linked chains after the removal of Neu5NAc. SPR has been used for investigating the kinetics of the interaction between oligosaccharides and lectins (7–9), for characterization of fetuin glycosylation (10), for the determination of agalactoIgG in rheumatoid arthritis patients (11), and recently, we have developed a method for investigating the glycosylation of recombinant glycoproteins (12). 2. Materials 2.1. Reagents 1. 2. 3. 4. 5.

6. 7. 8. 9. 10. 11.

0.05 M N-Hydroxysuccinimide (NHS). 0.2 M N-ethyl-N' (dimethylaminopropyl)-carbodiimide (EDC). 1 M Ethanolamine hydrochloride, pH 8.5. 0.1 M HCl. Running buffer (HBS): 10 mM 2-[4-(2-hydroxyethyl)-1-piperazinyl] ethanesulfonic acid (HEPES), pH 7.4, 150 mM NaCl, 3.4 mM ethylenediaminetetraacetic acid (EDTA), 0.0005% Surfactant P20. 10 mM Sodium acetate, pH 4.0. 10 mM Sodium acetate, pH 4.5. 10 mM Sodium acetate, pH 5.0. 10 mM Sodium acetate, pH 5.5. Lectin buffer: 10 mM sodium acetate, pH 5.0 with cations 2 mM MgCl2, MnCl2, ZnCl2, CaCl2. Lectins used: MAA, SNA, Datura stramonium Agglutinin (DSA), ConA, PNA, aleuria aurantia Agglutinin (AAA).

2.2. Equipment BIAcore 1000 apparatus (Biacore AB, Uppsala) is fully automated and controlled by the manufacturer’s software. During the operation of the equipment the output from the chip is displayed on VDU as resonance units (RUs) vs time. 3. Method The procedure used for lectin/SPR can be divided into four steps. The first three are essential for any SPR experiments and the last one is optional. 1. 2. 3. 4.

“Preconcentration” step. Ligand immobilization step. Lectin binding. Enzyme treatment.

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3.1. Preconcentration Before immobilizing a ligand to the chip surface a procedure called “preconcentration” needs to be performed. The latter step is important for efficient chemical immobilization of the ligand. It is accomplished by passing the ligand over the chip surface and utilizing the electrostatic attraction between the negative charges on the surface matrix (carboxymethyl dextran on a CM chip) and positive charges on the ligand at pH values below the ligand pI. Preconcentration is performed at 25°C. The ligand is injected at different pHs and the pH that provides the steepest curve is used (see Note 1). The detailed sequence of steps is as follows: 1. 2. 3. 4. 5. 6.

A continuous flow of HBS buffer is applied at flow rate of 10 mL/min. A 2-min pulse of ligand in 10 mM sodium acetate, pH 4.0, is applied. A 2-min pulse of ligand in 10 mM sodium acetate, pH 4.5, follows. A 2-min pulse of ligand in 10 mM sodium acetate, pH 5.0, follows. A 2-min pulse of ligand in 10 mM sodium acetate, pH 5.5, follows. Two 2-min washes with 0.1 M HCl follow for regeneration.

3.2. Glycoprotein Immobilization Choosing the correct matrix for immobilization depends on properties of the glycoprotein to be used and the functional groups on the chip surface. There are various types of chips available, which utilize different immobilization chemistries. Amine coupling is used for neutral and basic proteins (13). This introduces N-hydroxysuccinimide esters into the surface matrix by modifying the carboxymethyl groups of the matrix with a mixture of NHS and EDC. These esters react with the amines and other nucleophilic groups on the ligand to form covalent links. Optimization of the amount of immobilized ligand is an important factor in SPR measurements and different buffers, ligand concentrations and injection times must be investigated. This procedure is performed at 25°C (Fig. 2). 1. A continuous flow of HBS buffer is applied at flow rate of 5 µL/min. 2. The sensor chip surface is activated with a 7-min pulse of NHS and EDC. 3. Immobilization of ligand is performed by a 7-min pulse of the ligand solution (see Notes 2 and 3). 4. Deactivation of excess reactive groups on the surface and removal of noncovalently bound material is performed by a 7-min pulse of ethanolamine. 5. Two 4-min pulses with HCl perform regeneration (complete removal of nonbound material).

3.3. Lectin Binding Lectin binding is performed at 37°C. A continuous liquid flow is applied to the chip and the lectin is injected as a short pulse. If the lectin recognizes a carbohydrate grouping on the glycoprotein then it will bind resulting in an increasing

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Fig. 2. (A) Baseline during continuous buffer flow; (B) injection of NHS/EDC to activate the surface; (C) baseline after activation; (D) injection of ligand; (E) immobilized ligand before deactivation; (F) deactivation of chip surface using ethanolamine; (G) immobilized ligand after deactivation; (H) final immobilization level after two HCl washes.

Fig. 3. Typical sensogram of ConA binding. AB, baseline; B, sample injection; BC, association; CD, dissociation; D, E, HCl injections; F, baseline.

RU value. A RU reading is taken approx 15 s after dissociation starts. Bound lectin is removed by passing HCl over the chip and then the binding of another lectin is investigated (Fig. 3). 1. A continuous flow of HBS buffer is applied at flow rate of 5 µL/min. 2. Lectin is injected by a 7-min pulse (see Note 4). 3. A reading is taken 15 s after dissociation starts (see Note 5).

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4. Lectin is removed by two 4-min pulses with HCl (see Note 6). 5. Steps 1–4 are repeated for each lectin and buffer solution.

3.4. Enzyme Treatment The specificity of lectin binding measured by SPR can be confirmed by enzyme treatment at 37°C with glycosidases. These are usually very specific for one carbohydrate structure. After enzyme treatment lectin binding should not be detected if the interaction was specific. Figure 4 shows the binding of MAA before and after treatment with neuraminidase. Immobilized glycoproteins can be treated with other enzymes such as galactosidase and PGNase F (see Note 7). 1. Neuraminidase treatment is performed by a 7-min injection of the enzyme followed by a stop in the flow for 6 h. 2. The flow rate restored back to 5 µL/mL. 3. Regeneration (removal of neuraminidase) is performed by two 4-min pulses with HCl. 4. Lectin injection, reading, and regeneration as in steps 1–5 of Subheading 3.3. 5. The same procedure as described in steps 1–4 can be repeated for other enzymes until all residues on a sugar chain are chopped off.

4. Notes 1. A preliminary choice of pH for immobilization can be made if the pI of the ligand is known according to the following rule of thumb: for pI > 7, use pH 6, for pI 5.5–7 use 1 unit pH below pI, for pI 3.5–5.5, use 0.5 pH units below pI. For pH range of 4–5.5 10 mM acetate buffer is recommended.

Fig. 4. MAA binding before (solid line) and after (dotted line) neuraminidase treatment (buffer plot subtracted from both sensograms).

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Table 1 Typical Results for Different Glycoproteins with Different Lectins

IgG Fetuin Rec1 Rec2

ConA

SNAb

SNAa

MAAb

MAAa

PNAb

PNAa

DSA

AAA

1680 776 2768 1203

344 1230 22 10

50 78 20 25

25 624 10 325

26 5 25 22

24 5 26 4

23 80 30 297

33 230 11 42

71 32 22 15

a,b

Before and after neuraminidase treatment. IgG and fetuin were commercial products, while rec1 and rec2 are recombinant glycoproteins. Buffer readings gave values of 20–35 RU. See Note 8.

2. The time of activation may be modified to regulate the amount of ligand immobilized. 3. For a given pH the ideal ligand concentration is the lowest value that gives maximum preconcentration. Usually, a suitable concentration will be in the range 10–200 µg/mL. A target response is a level of immobilization that gives about 0.07 pmol ligand/mm2. That corresponds to mol wt/15 RU. 4. Analyte concentration can be chosen between 5 µg/mL and 500 µg/mL. 5. A reading value above 30–40 RU is considered as real binding. 6. After regeneration the baseline should be at the same level as before lectin binding. A drift in the baseline may appear after repeated regeneration, which may be due to loss of Neu5Ac. 7. Different enzyme incubation times may apply depending on the enzyme properties. 8. Table 1 gives typical results for different glycoproteins with different lectins. The following conclusions can be drawn from these results: IgG has N-linked chains (ConA), α2–6 Neu5NAc (SNA) and fucose (AAA) and it does not have any O-linked chains (PNA) or any α2–3 Neu5NAc (MAA); Fetuin has N- and O-linked chains, α2–6 Neu5NAc and α2–3 Neu5NAc, galactose β1–4 N-acetylglucosamine (DSA) and no fucose; Rec1 has N-linked chains and no fucose, no Neu5NAc, no O-linked chains; Rec2 has N- and O-linked chains, α2–6 Neu5NAc, no α2–3 Neu5NAc and no fucose. Acknowledgments This work was supported by funds from Biomed Laboratories, Newcastle upon Tyne, UK, British Biotech Ltd., Oxford, UK and Cambridge Antibody Technology, Royston, UK.

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References 1. Imperiali, B. and O’Connor, S. E. (1999) Effect of N-linked glycosylation on glycopeptide and glycoprotein structure. Curr. Opin. Chem. Biol. 3, 643–649. 2. Varki, A. (1993) Biological roles of oligosaccharides: all of the theories are correct. Glycobiology 3, 97–130. 3. Singh, R. S., Tiwary, A. K., and Kennedy, J. F. (1999) Lectins: sources, activities, and applications. Crit. Rev. Biotechnol. 19, 145–178. 4. Turner, G. A. (1992) N-Glycosylation of serum proteins in disease and its investigation using lectins. Clin. Chim. Acta 208, 149–171. 5. Fivash, M., Towler, E. M., and Fisher, R. J. (1998) BIAcore for macromolecular interaction. Curr. Opin. Biotechnol. 9, 97–101. 6. (1994) The SPR signal, in BIAtechnology Handbook, Pharmacia Biosensor AB, Uppsala, Sweden, pp. 4–3. 7. Haseley, S. R., Talaga, P., Kamerling, J. P., and Vliegenthart, J. F. G. (1999) Characterization of the carbohydrate binding specificity and kinetic parameters of lectins by using surface plasmon resonance. Analyt. Biochem. 274, 203–210. 8. Shinohara, Y., Kimi, F., Shimizu, M., Goto, M., Tosu, M., and Hasegawa, Y. (1994) Kinetic measurement of the interaction between an oligosaccharide and lectins by a biosensor based on surface plasmon resonance. Eur. J. Biochem. 223, 189–194. 9. Satoh, A. and Matsumoto, I. (1999) Analysis of interaction between lectin and carbohydrate by surface plasmon resonance. Analyt. Biochem. 275, 268–270. 10. Hutchinson, M. (1994) Characterization of glycoprotein oligosaccharides using surface plasmon resonance. Analyt. Biochem. 220, 303–307. 11. Liljeblad, M., Lundblad, A., and Pahlsson, P. (2001) Analysis of agalacto-IgG in rheumatoid arthritis using surface plasmon resonance. Glycoconj. J. 17, 323–329. 12. Fotinopoulou, A., Cooke, A., and Turner, G. A. (2000) Does the ‘glyco’ part of recombinant proteins affect biological activity. Immunol. Lett. 73, 105. 13. (1994) Ligand immobilization chemistry, in BIA applications Handbook, Pharmacia Biosensor AB, Uppsala, Sweden, pp. 4–1.

145 Sequencing Heparan Sulfate Saccharides Jeremy E. Turnbull

1. Introduction The functions of the heparan sulfates (HSs) are determined by specific saccharide motifs within HS chains. These sequences confer selective protein binding properties and the ability to modulate protein activities (1,2). HS chains consist of an alternating disaccharide repeat of glucosamine (GlcN; N-acetylated or N-sulfated) and uronic acid (glucuronic [GlcA] or iduronic acid [IdoA]). The initial biosynthetic product containing N-acetylglucosamine (GlcNAc) and GlcA is modified by N-sulfation of the GlcN, ester (O)-sulfation (at positions 3 and 6 on the GlcN and position 2 on the uronic acids) and by epimerization of GlcA to IdoA. The extent of these modifications is incomplete and their degree and distribution varies in HS between different cell types. In HS chains N- and O-sulfated sugars are predominantly clustered in sequences of up to eight disaccharide units separated by N-acetyl-rich regions with relatively low sulfate content (3). Sequence analysis of HS saccharides is a difficult analytical problem and until recently sequence information had been obtained for only relatively short saccharides from HS and heparin. Gel chromatography and high-performance liquid chromatography (HPLC) methods have been used to obtain information on disaccharide composition (3,4). Other methods such as nuclear magnetic resonance (NMR) spectroscopy and mass spectroscopy (5–9) have provided direct sequence information, but are difficult for even moderately sized oligosaccharides and in the case of NMR requires large amounts of material (micromoles). This situation has changed rapidly in the last few years with the availability of recombinant exolytic lysosomal enzymes. These exoglycosidases and exosulfatases remove specific sulfate groups or monosaccharide residues from the nonreducing end (NRE) of saccharides (10). They can be employed in combination with polyacrylamide gel electrophoresis (PAGE) separations to From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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derive direct information (based on band shifts) on the structures present at the nonreducing end of GAG saccharides (11; see Fig. 1 for an example). Integral glycan sequencing (IGS), a PAGE-based method using the exoenzymes, was developed as the first strategy for rapid and direct sequencing of HS and heparin saccharides (11). Its introduction has been quickly followed by a variety of similar approaches using other separation methods including HPLC and matrix-assisted laser desorption (MALDI) mass spectrometry (12–14). An outline of the IGS sequencing strategy is given in Fig. 2. An HS (or heparin) saccharide (previously obtained from the polysaccharide by partial chemical or enzymatic degradation and purification) is labeled at its reducing terminus with a fluorescent tag. It is then subjected to partial nitrous acid treatment to give a ladder of evenly numbered oligosaccharides (di-, tetra-, hexa-, etc.) each having a fluorescent tag at their reducing end. Portions of this material are then treated

Fig. 1. Basic principles of integral glycan sequencing. (A) Fluorescence detection of different amounts of a 2-AA-tagged heparin tetrasaccharide run on a 33% minigel. (B) Exosequencing of a 2-AA-tagged heparin tetrasaccharide with lysosomal enzymes and separation of the products on a 33% minigel (15 pmol per track). Band shifts following the exoenzyme treatments shown reveal the structure of the nonreducing end disaccharide unit (track 1, untreated). I2Sase, iduronate-2-sulfatase; Idase, iduronidase; G6Sase, glucosamine-6-sulfatase; Nsase, sulfamidase. (C) Schematic representation of IGS of a hexasaccharide (pHNO2, partial nitrous acid treatment). (D) Actual example of IGS performed on a purified heparin hexasaccharide, corresponding to the scheme in C, using the combinations of pHNO2 and exoenzyme treatments indicated (track 1, untreated, 25 pmol; other tracks correspond to approx 200 pmol per track of starting sample for pHNO2 digest). The hexasaccharide (purified from bovine lung heparin) has the putative structure IdoA(2S)-GlcNSO3(6S)-IdoA(2S)GlcNSO3(6S)-IdoA(2S)-AMannR(6S). Electrophoresis was performed on a 16-cm 35% gel (from ref. 11, Copyright 1999 National Academy of Sciences, USA).

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Fig. 2. The IGS sequencing strategy.

with a variety of highly specific exolytic lysosomal enzymes (exosulfatases and exoglycosidases) that act at the nonreducing end of each saccharide (if it is a suitable substrate). The various digests are then separated on a high-density polyacrylamide gel and the positions of the fragments detected by excitation of the fluorescent tag with a UV transilluminator. Band shifts resulting from the different treatments permit the sequence to be read directly from the banding pattern (see Fig. 1 for an example). This novel strategy allows direct read-out sequencing of a saccharide in a single set of adjacent gel tracks in a manner analogous to DNA sequencing. IGS provides a rapid approach for sequencing HS saccharides, and has proved very useful in recent structure–function studies (15). It should be noted that this methodology is designed for sequencing purified saccharides, not whole HS preparations. An important factor in all sequencing methods is the availability of sufficiently pure saccharide starting material.

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HS and heparin saccharides can be prepared following selective scission by enzymic (or chemical) reagents and isolation by methods such as affinity chromatography (4). Final purification usually requires the use of strong anionexchange HPLC (11,15). 2. Materials 1. 2-Aminobenzoic acid (2-AA; Fluka Chemicals). 2. 7-Aminonaphthalene-1,3-disulfonic acid monopotassium salt (ANDSA; Fluka Chemicals). 3. Formamide. 4. Sodium cyanoborohydride (>98% purity). 5. Sodium triacetoxyborohydride (Aldrich). 6. Distilled water. 7. Oven or heating block at 37°C. 8. Desalting column (Sephadex G-25; e.g., HiTrap™ desalting columns, Pharmacia). 9. Centrifugal evaporator. 10. 200 mM HCl. 11. 20 mM Sodium nitrite (1.38 mg/mL in distilled water; prepare fresh). 12. 200 mM Sodium acetate, pH 6.0:27.2 g/L of sodium acetate trihydrate; adjust pH to 6.0 using acetic acid. 13. Enzyme buffer: 0.2 M Na acetate, pH 4.5. Make 0.2 M sodium acetate (27.2 g/L of sodium acetate trihydrate) and 0.2 M acetic acid (11.6 mL/L) and mix in a ratio of 45 mL to 55 mL, respectively. 14. Enzyme stock solutions (typically at concentrations of 500 mU/mL, where 1 U = 1 µmol substrate hydrolyzed per minute). Available from Glyko, Novato, CA. 15. Vortex tube mixer. 16. Microcentrifuge. 17. Acrylamide stock solution (T50%–C5%). Caution: Acrylamide is neurotoxic. Wear gloves (and a face mask when handling powdered forms). It is convenient to use premixed bis-acrylamide such as Sigma A-2917. Add 43 mL of distilled water to the 100-mL bottle containing the premixed chemicals and dissolve using a small stirrer bar (approx 2 h). The final volume should be approx 80 mL. Store the stock solution at 4°C. Note that it is usually necessary to warm gently to redissolve the acrylamide after storage. 18. Resolving gel buffer stock solutions: 2 M Tris-HCl, pH 8.8 (242.2 g/L of Tris base; adjust pH to 8.8 with HCl). 19. Stacking gel buffer stock solution: 1 M Tris-HCl, pH 6.8 (121.1 g/L of Tris base; adjust pH to 6.8 with HCl). 20. Electrophoresis buffer: 25 mM Tris, 192 mM glycine, pH 8.3.3 g/L of Tris base, 14.4 g/L of glycine; adjust pH to 8.3 if necessary with HCl. 21. 10% Ammonium persulfate in water (made fresh or stored at −20°C in aliquots). 22. N,N,N ,N -Tetramethylethylenediamine (TEMED). 23. Vertical slab gel electrophoresis system (minigel or standard size). 24. D.C. Power supply unit (to supply up to 500–1000 V and 200 mA).

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25. UV transilluminator (312 nm maximum emission wavelength). 26. Glass UV bandpass filter larger than gel size (type UG-11, or M-UG2). 27. Charge coupled device (CCD) imaging camera fitted with a 450-nm (blue) bandpass filter.

3. Methods 3.1. Tagging Saccharides with a Fluorophore HS (and heparin) saccharides can be endlabeled by reaction of their reducing aldehyde functional group with a primary amino group of a fluorophore (reductive amination). For sulfated saccharides anthranilic acid (2-AA; 11) has been found to be effective for the IGS methodology. 2-AA conjugates display an excitation maxima in the range 300–320 nm, which is ideal for visualization with a commonly available 312-nm UV source (e.g., transilluminators used for visualizing ethidium bromide stained DNA). Emission maxima are typically in the range 410–420 nm (bright violet fluorescence). It has also been found that approx 10-fold more sensitive detection is possible using an alternative fluorophore ANDSA, and this has been demonstrated to be effective for HS sequencing (16). ANDSA has an excitation maxima of 350 nm and emission maxima of 450 nm. Both approaches described in Subheadings 3.1.1. and 3.1.2. allow rapid labeling and purification of tagged saccharide from free tagging reagent, and give quantitative recoveries and products free of salts that might interfere with subsequent enzymic conditions. For saccharides in the size range hexa to dodecasaccharides, approx 2–3 nmol (approx 2–10 µg) of purified starting material is the minimum required using the 2-AA label and approx 5- to 10-fold less for ANDSA labeling. It should be noted that recently a novel label, BODIPY hydrazide, has been developed for highly sensitive detection of saccharides including those from HS (17). This label couples more efficiently to saccharides, has an excitation maxima at 503nm (making it ideal for detection using common 488nm lasers) and an extinction co-efficient of 71,000; this permits very sensitive detection of labelled saccharides (~100fmol on a standard HPLC fluorimeter). In the future it may prove possible to apply this label to highly sensitive IGS sequencing on gels using fluorescent gel scanners or alternatively HPLC or capillary electrophoresis separations with in-line fluorescence detection. 3.1.1. Labeling Saccharides with 2-AA 1. Dry down the purified saccharide (typically 2–10 nmol) in a microcentrifuge tube by centrifugal evaporation. 2. Dissolve directly in 10–25 µL of formamide containing freshly prepared 400 mM 2-AA (54.8 mg/mL) and 200 mM reductant (sodium cyanoborohydride; 12.6 mg/mL) and incubate at 37°C for 16–24 h in a heating block or oven. (Caution: The reductant is toxic and should be handled with care.) The volume used should be sufficient to provide a 500–1000-fold molar excess of 2-AA over saccharide (see Note 1).

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3. Remove free 2-AA, reductant, and formamide from the labeled saccharides by gel filtration chromatography (Sephadex G-25 Superfine). Dilute the sample (maximum 250 µL of reaction mixture) to a total of 1 mL with distilled water (see Note 2). 4. Load sample onto two 5-mL HiTrap™ Desalting columns (Pharmacia Ltd.) connected in series. Alternatively it is possible to use self-packed columns of other dimensions. 5. Elute with distilled water at a flow rate of 1 mL/min and collect fractions of 0.5 mL. Saccharides consisting of four or more monosaccharide units typically elute in the void volume (approx fractions 7–12). Note that the HiTrap™ columns can be eluted by hand with a syringe without need for a pump. 6. Pool and concentrate these fractions by centrifugal evaporation or freeze drying.

3.1.2. Labeling Saccharides with ANDSA 1. Dry down the purified saccharide (typically 2–10 nmol) in a microcentrifuge tube by centrifugal evaporation. 2. Dissolve directly in 10 µL of formamide. 3. Mix with 15 µL of formamide containing ANDSA at a concentration of 80 mg/mL (approaching saturation) and incubate at 25°C for 16 h. 4. Mix with 10 µL of formamide containing 1 mg of the reductant sodium triacetoxyborohydride (see Note 1) and incubate for 2 h at 25°C. 5. Remove free ANDSA by gel filtration as described in Subheading 3.1.1. for 2-AA.

3.2. Nitrous Acid Treatment of Saccharides Low pH nitrous acid cleaves HS only at linkages between N-sulfated glucosamine and adjacent hexuronic acid residues (18,19). Under mild controlled conditions nitrous acid cleavage creates a ladder of bands corresponding to the positions of internal N-sulfated glucosamine residues in the original intact saccharide (11). A series of different reaction stop points are pooled, resulting in a partial digest with a range of different fragment sizes. 1. Dry down 1–2 nmol of labeled saccharide by centrifugal evaporation. 2. Redissolve in 80 µL of distilled water and chill on ice. 3. Add 10 µL of 200 mM HCl and 10 µL of 20 mM sodium nitrite (both prechilled on ice) and incubate on ice. 4. At a series of individual time points (typically 15, 30, 60, 120, and 180 min), remove an aliquot and stop the reaction by raising the pH to approx 5.0 by the addition of 1/5 volume of 200 mM sodium acetate buffer, pH 6.0 (see Note 3). 5. Pool the set of aliquots and either use directly for enzyme digests or desalt as described in

3.3. Exoenzyme Treatment of Saccharides The approach for treatment of HS samples with exoenzymes is described below. Details of the specificities of the exoenzymes is given in Table 1. These enzymes have differing optimal pH and buffer conditions, but in general they

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Table 1 Exoenzymes for Sequencing Heparan Sulfate and Heparin Enzymea Substrate specificityb Sulfatases Iduronate-2-sulfatase IdoA(2S) Glucosamine-6-sulfatase GlcNAc(6S), GlcNSO3(6S) GlcNSO3 Sulphamidase (glucosamine N-sulfatase) Glucuronate-2-sulfatase GlcA(2S) Glucosamine-3-sulfatase GlcNSO3(3S) Glycosidases Iduronidase IdoA Glucuronidase GlcA GlcNAc α-N-Acetylglucosaminidase Bacterial exoenzymes ∆4,5-Glycuronate-2-sulfatase ∆UA(2S) ∆4,5-Glycuronidase ∆UA a Enzyme availability: Glucuronidase is widely available commercially as purified or recombinant enzyme. Recombinant iduronate-2-sulfatase, iduronidase, glucosamine-6sulfatase, sulfamidase, and α-N-acetylglucosaminidase were purchased from from the Glyko product range, now supplied by Prozyme (San Leandro, CA; www.prozyme.com). Glucuronate-2-sulfatase and glucosamine-3-sulfatase have only been purified from cell and tissue sources to date. The bacterial exoenzymes are available from Grampian Enzymes, Nisthouse, Harray, Orkney, Scotland; e-mail, [email protected]. b The specificities are shown as the nonreducing terminal group recognized by the enzymes.

can be used under the single set of conditions given here, which simplifies the multiple enzyme treatments required (see Note 4). Sulfatases remove only the sulfate group, whereas the glycosidases cleave the whole nonsulfated monosaccharide. 1. Dissolve the sample (typically 10–200 pmol of saccharide) in 10 µL of H2O in a microcentrifuge tube. 2. Add 5 µL of exoenzyme buffer (100 mM sodium acetate buffer, pH 4.5), 1 µL of 0.5 mg/mL bovine serum albumin, 2 µL of appropriate exoenzyme [0.2–0.5 mU], and distilled water to bring the final volume to 20 µL. 3. Mix the contents well on a vortex mixer, and centrifuge briefly to ensure that the reactants are at the tip of the tube. 4. Incubate the samples at 37°C for 16 h in a heating block or oven.

3.4. Separation of Saccharides by PAGE PAGE is a high-resolution technique for the separation of HS and heparin saccharides of variable sulfate content and disposition. Its resolution is generally superior to gel filtration or anion-exchange HPLC (20,21). Improved resolution can be obtained using gradient gels, although these are more difficult to prepare

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and use routinely. In most cases sufficient resolution can be obtained with isocratic gels (see Note 5). PAGE provides a simple but powerful approach for separating the saccharide products generated in the sequencing process. 3.4.1. Preparing the PAGE Gel 1. Assemble the gel unit (consisting of glass plates and spacers, etc). 2. Prepare and degas the resolving gel acrylamide solution without ammonium persulfate or TEMED. To make a 30% acrylamide gel solution for a 16 cm • 12 cm • 0.75 mm gel, 16 mL is required. Mix 9.6 mL of T50%–C5% acrylamide stock with 3 mL of 2 M Tris, pH 8.8, and 3.4 mL of distilled water. 3. Add 10% ammonium persulfate (30 µL) and TEMED (10 µL) to the gel solution, mix well, and immediately pour into the gel unit. 4. Overlay the unpolymerized gel with resolving gel buffer (375 mM Tris-HCl, pH 8.8, diluted from the 2 M stock solution) or water-saturated butanol. Polymerization should occur within approx 30–60 min. The gel can then be used immediately or stored at 4°C for 1–2 wk.

3.4.2. Electrophoresis 1. Immediately before electrophoresis, rinse the resolving gel surface with stacking gel buffer (0.125 M Tris-HC1 buffer, pH 6.8, diluted from the 1 M stock solution). 2. Prepare and degas the stacking gel solution (for 5 mL, mix 0.5 mL of T50%–C5% acrylamide stock with 0.6 mL of 1 M Tris, pH 6.8, and 3.9 mL of distilled water). 3. Add 10% ammonium persulfate (10 µL) and TEMED (5 µL). Immediately pour onto the top of the resolving gel and insert the well-forming comb. 4. After polymerization (approx 15 min) remove the comb and rinse the wells thoroughly with electrophoresis buffer. 5. Place the gel unit into the electrophoresis tank and fill the buffer chambers with electrophoresis buffer. 6. Load the oligosaccharide samples (5–20 µL dependent on well capacity, containing approx 10% [v/v] glycerol or sucrose in 125 mM Tris-HCl, pH 6.8) carefully into the wells with a microsyringe. Marker samples containing bromophenol blue and phenol red should also be loaded into separate tracks. 7. Run the samples into the stacking gel at 150–200 V (typically 20–30 mA) for 30–60 min, followed by electrophoresis at 300–400 V (typically 20–30 mA and decreasing during run) for approx 5–8 h (for a 16-cm gel). Heat generated during the run should be dissipated using a heat exchanger with circulating tap water, or by running the gel in a cold room or in a refrigerator. 8. Electrophoresis should be terminated before the phenol red marker dye is about 5 cm from the bottom of the gel. (At this point, disaccharides should be 3–4 cm from the bottom of the gel.)

3.5. Gel Imaging The most effective approach for gel imaging requires a CCD camera that can detect faint fluorescent banding patterns by capturing multiple frames. Systems

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commonly used for detection of ethidium bromide stained DNA can usually be adapted with appropriate filters as described below (see Note 6). 1. Place a UV filter (UG-1, UG-11, or MUG-2) onto the transilluminator, and fit a 450-nm blue filter onto the camera lens. 2. Remove the gel carefully from the glass plates after completion of the run and place on the UV transilluminator surface wetted with electrophoresis buffer. Wet the upper surface of the gel to reduce gel drying and curling. 3. Switch on the transilluminator and capture the image using the CCD camera. Exposure times are typically 1–5 s depending on the amount of labeled saccharide (see Note 7).

3.6. Data Interpretation The sequence of saccharides can be read directly from the banding pattern by interpreting the band shifts due to removal of specific sulfate or sugar moieties. Fig. 1 shows an actual example and a schematic representation. First, bands generated by the partial nitrous acid treatment indicate the positions of N-sulfated glucosamine residues in the original saccharide (Fig. 1C, track 2). A “missing” band in the ladder at a particular position indicates the presence of an N-acetylated glucosamine residue in the original saccharide at that position (an example of this is shown in Fig. 3). Such saccharides can be sequenced by the additional use of the exoenzyme N-acetylglucosaminidase, which removes this residue and allows further sequencing of an otherwise “blocked” fragment. Following the nitrous acid treatment, the “ladder” of bands is then subjected to various exoenzyme digestions. The presence of specific sulfate or sugar residues can be deduced from the band shifts that occur (Fig. 1C, tracks 3–5). Figure 4 shows an example of a decasaccharide from HS that has been purified by SAXHPLC and sequenced using IGS (11). Although the band shifts are usually downwards (because of to the lower molecular mass and thus higher mobility of the product) it should be noted that occasionally upward shifts occur, probably due to subtle differences in charge/ mass ratio (for examples, see Figs. 1B, 3B, and 4C). Note also that minor “ghost” bands sometimes appear after the nitrous acid treatment. They are probably due to loss of an N-sulfate group, and normally these do not affect interpretation of the shifts in the major bands (11). If the saccharide being sequenced was derived by bacterial lyase treatment, it will have a ∆4,5-unsaturated uronate residue at its nonreducing terminus. If this residue has a 2-O-sulfate attached, this can be detected by susceptibility to I2Sase (see Fig. 3B), but the sugar residue itself is resistant to both Idase and Gase. Its removal is required to confirm whether there is a 6-O-sulfate on the adjacent nonreducing end glucosamine (see Figs. 3B and 4C for examples). Bacterial enzymes that specifically remove the ∆4,5-unsaturated uronate residues (and the 2-O-sulfate groups that may be present on them) are now available

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Fig. 3. IGS of a heparin hexasaccharide. A heparin hexasaccharide with the structure ∆HexA(2S)-GlcNSO3(6S)-IdoA-GlcNAc(6S)-GlcA-GlcNSO3(6S) was 2-AA-tagged and subjected to sequencing on a 16-cm 33% gel. (A) IGS of hexasaccharide using the combinations of pHNO2 and exoenzyme treatments indicated (track 1, untreated, 20 pmol; other tracks correspond to approx 90 pmol per track of starting sample for pHNO2 digest). NAG, N-acetylglucosaminidase. (B) Determining the sequence of the nonreducing disaccharide unit of the hexasaccharide using the I2Sase, G6Sase, and mercuric acetate (MA) treatments shown (approx 20 pmol per track; track 1, untreated). (From ref. 11, Copyright 1999 National Academy of Sciences, USA.)

commercially (see Table 1). Alternatively, they can be removed chemically with mercuric acetate (22; see Figs. 3B and 4C). In addition to the basic sequencing experiment, it is wise to confirm agreement of the data with an independent analysis of the disaccharide composition of the saccharide (11). It can sometimes be difficult to sequence the reducing terminal monosaccharide owing to it being a poor substrate for the exoenzymes. In these cases it has proved more effective to analyze the terminal 2AA-labeled disaccharide unit in comparison to 2AA-labeled disaccharide standards (11). In the future it is anticipated that methods using HPLC or capillary electrophoresis separations and the fluorophore BODIPY hydrazide (17) will permit ultra-high sensitivity IGS sequencing of HS saccharides, superceding the standard electrophoretic methods described in this chapter.

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Fig. 4. HPLC purification and IGS of a HS decasaccharide. (A) SAX-HPLC of a pool of HS decasaccharides derived by heparitinase treatment of porcine mucosal HS (for details see ref. 11). The arrowed peak was selected for sequencing. (B) IGS of the purified HS decasaccharide on a 16-cm 33% gel using the combinations of pHNO2 and exoenzyme treatments indicated (track 1, untreated, 20 pmol; other tracks correspond to approx 400 pmol per track of starting sample for pHNO2 digest). (C) Determining the sequence of the nonreducing disaccharide unit of the HS decasaccharide using the mercuric acetate (MA) and G6Sase treatments shown (approx 40 pmol per track; track 1, untreated). (From ref. 11, Copyright 1999 National Academy of Sciences, USA.)

4. Notes 1. Using large excesses of reagent as described, saccharides derived from HS and heparin by bacterial lyase scission generally couple with 2-AA with efficiencies in the range of 60–70%. Note that saccharides derived from HS and heparin by low pH nitrous acid scissioning (i.e., having an anhydromannose residue at their reducing ends) label more efficiently (approx 70–80% coupling efficiency). Labeling with ANDSA achieves similar coupling efficiencies, and the alternative reducing agent, sodium triacetoxyborohydride, is less toxic than sodium cyanoborohydride. 2. Unwanted reactants and solvent can also be removed from labeled saccharides by methods such as dialysis but the rapid gel filtration chromatography step described above using the HiTrap desalting columns is convenient and usually allows good recoveries of loaded sample (typically 70–80%). 3. It is best to perform some trial incubations to test for optimal time points needed to generate a balanced mix of all fragments in the partial nitrous

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5.

6.

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acid digestion. With longer saccharides (octasaccharides and larger) it is observed that the largest products are generated quickly and thus a bias toward shorter incubation times is required as saccharide length increases (16). The enzyme conditions described should provide for complete digestion of all susceptible residues. This is important to the sequencing process, as incomplete digestion would create a more complex banding pattern and would give a false indication of sequence heterogeneity. It is useful to run parallel controls with standard saccharides to enable monitoring of reaction conditions. Where combinations of exoenzymes are required, these can be incubated simultaneously with the sample. If required, the activity of one enzyme can be destroyed prior to a secondary digestion with a different enzyme by heating the sample at 100°C for 2–5 min. Adequate separations, particularly over limited size ranges of saccharides, can be obtained using single concentration gels, typically in the range 25–35% acrylamide. Improvements in resolution can be made by using longer gel sizes. Different voltage conditions (usually in the range 200–600 V) and running times are required for different gel formats, and should be established by trial and error with the particular samples being analyzed. Gels up to 24 cm in length can usually be run in 5–8 h using high voltages, whereas for longer gels it is often convenient to use lower voltage conditions and overnight runs. Minigels can also be used effectively for separation of small HS–heparin saccharides up to octasaccharides in size (see Fig. 1). Note that it is also possible to run Tris-acetate gels with a Tris-MES electrophoresis buffer (see Fig. 1; 11). Because the emission wavelength of 2-AA tagged saccharides is 410– 420 nm, there is a need to filter out background visible wavelength light from the UV lamps. This can be done effectively with special glass filters that permit transmission of UV light but do not allow light of wavelengths >400 nm to pass. A blue bandpass filter on the camera also improves sensitivity. Suitable filters are available from HV Skan (Stratford Road, Solihull, UK; Tel: 0121 733 3003) or UVItec Ltd. (St. John’s Innovation Centre, Cowley Road, Cambridge, UK; www.uvitec.demon. co.uk). Required exposure times are strongly dependent on sample loading and the level of detection required. Over-long exposures will result in excessive background signal. Note that negative images are usually better for band identification (see figures). Under the conditions described the limit of sensitivity is approx 10–20 pmol per band of original starting material for 2-AA (see Fig. 1) and 2–5 pmol per band for ANDSA.

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References 1. Turnbull, J. E., Powell, A., and Guimond, S. E. (2001) Heparan sulphate: decoding a dynamic multifunctional cellular regulator. Trends Cell Biol. 11, 75–82. 2. Bernfield, M., Gotte, M., Park, P. W., et al. (1999) Functions of cell surface heparan sulfate proteoglycans. Annu. Rev. Biochem. 68, 729–777. 3. Turnbull, J. E. and Gallagher, J. T. (1991) Distribution of iduronate-2-sulfate residues in HS: evidence for an ordered polymeric structure. Biochem. J. 273, 553–559. 4. Turnbull, J. E., Fernig, D., Ke, Y., Wilkinson, M. C., and Gallagher, J. T. (1992) Identification of the basic FGF binding sequence in fibroblast HS. J. Biol. Chem. 267, 10,337–341. 5. Pervin, A., Gallo, C., Jandik, K., Han, X., and Linhardt, R. (1995) Preparation and structural characterisation of heparin-derived oligosaccharides. Glycobiology 5, 83–95. 6. Yamada, S., Yamane, Y., Tsude, H., Yoshida, K., and Sugahara, K. (1998) A major common trisulfated hexasaccharide isolated from the low sulfated irregular region of porcine intestinal heparin. J. Biol. Chem. 273, 1863–1871. 7. Yamada, S., Yoshida, K., Sugiura, M., Sugahara, K., Khoo, K., Morris, H., and Dell, A. (1993) Structural studies on the bacterial lyase-resistant tetrasaccharides derived from the antithrombin binding site of porcine mucosal intestinal heparin. J. Biol. Chem. 268, 4780–4787. 8. Mallis, L., Wang, H., Loganathan, D., and Linhardt, R. (1989) Sequence analysis of highly sulfated heparin-derived oligosaccharides using FAB-MS. Analyt. Chem. 61, 1453–1458. 9. Rhomberg, A. J., Ernst, S., Sasisekharan, R., and Bieman, K. (1998) Mass spectrometric and capillary electrophoretic investigation of the enzymatic degradation of heparin-like glycosaminoglycans. Proc. Natl. Acad. Sci. USA 95, 4176–4181. 10. Hopwood, J. (1989) Enzymes that degrade heparin and heparan sulfate, in Heparin (Lane and Lindahl, eds.), Edward Arnold Press, pp. 191–227. 11. Turnbull, J. E., Hopwood, J. J., and Gallagher, J. T. (1999) A strategy for rapid sequencing of heparan sulfate/heparin saccharides. Proc. Natl. Acad. Sci. USA 96, 2698–2703. 12. Merry, C. L. R., Lyon, M., Deakin, J. A., Hopwood, J. J., and Gallagher, J. T. (1999) Highly sensitive sequencing of the sulfated domains of heparan sulfate. J. Biol. Chem. 274, 18,455–18,462. 13. Vives, R. R., Pye, D. A., Samivirta, M., Hopwood, J. J., Lindahl, U., and Gallagher, J. T. (1999) Sequence analysis of heparan sulphate and heparin oligosaccharides. Biochem. J. 339, 767–773. 14. Venkataraman, G., Shriver, Z., Ramar, R., and Sasisekharan, R. (1999) Sequencing complex polysaccharides. Science 286, 537–542. 15. Guimond, S. E. and Turnbull, J. E. (1999) Fibroblast growth factor receptor signalling is dictated by specific heparan sulfate saccharides. Curr. Biol. 9, 1343–1346. 16. Drummond, K. J., Yates, E. A., and Turnbull, J. (2001) Electrophoretic sequencing of heparin/heparan sulfate oligosaccharides using a highly sensitive fluorescent end label. Proteomics 1, 304–310. 17. Skidmore, MA; Guimond, SE ; Dumax-Vorzet, AF ; Atrih, A ; Yates, EA & Turnbull JE. (2006). High sensitivity separation and detection of heparan sulphate disaccharides. J Chrom A. 1135, 52–56.

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18. Shively, J. and Conrad, H. (1976) Formation of anhydrosugars in the chemical depolymerisation of heparin. Biochemistry 15, 3932–3942. 19. Bienkowski and Conrad, H. (1985) Structural characterisation of the oligosaccharides formed by depolymerisation of heparin with nitrous acid. J. Biol. Chem. 260, 356–365. 20. Turnbull, J. E. and Gallagher, J. T. (1988) Oligosaccharide mapping of heparan sulfate by polyacrylamide-gradient-gel electrophoresis and electrotransfer to nylon membrane. Biochem. J. 251, 597–608. 21. Rice, K., Rottink, M., and Linhardt, R. (1987) Fractionation of heparin-derived oligosaccharides by gradient PAGE. Biochem. J. 244, 515–522. 22. Ludwigs, U., Elgavish, A., Esko, J., and Roden, L. (1987) Reaction of unsaturated uronic acid residues with mercuric salts. Biochem. J. 245, 795–804.

146 Analysis of Glycoprotein Heterogeneity by Capillary Electrophoresis and Mass Spectrometry Andrew D. Hooker and David C. James

1. Introduction The drive toward protein-based therapeutic agents requires both product quality and consistency to be maintained throughout the development and implementation of a production process. Differences in host cell type, the physio-logical status of the cell, and protein structural constraints are known to result in variations in posttranslational modifications that can affect the bioactivity, receptor binding, susceptibility to proteolysis, immunogenicity, and clearance rate of a therapeutic recombinant protein in vivo (1). Glycosylation is the most extensive source of protein heterogeneity, and many recent developments in analytical biotechnology have enhanced our ability to monitor and structurally define changes in oligosaccharides associated with recombinant proteins. Variable occupancy of potential glycosylation sites may result in extensive glycosylation macroheterogeneity in addition to the considerable diversity of carbohydrate structures that can occur at individual glycosylation sites, often referred to as glycosylation microheterogeneity. Variation within a heterogeneous population of glycoforms may lead to functional consequences for the glycoprotein product. Therefore, regulatory authorities such as the US Food and Drug Administration (FDA) and the Committee for Proprietary Medical Productions demand increasingly sophisticated analysis for biologics produced by the biotechnology and pharmaceutical industries (2). The FDA has described a “well-characterized biologic” as “a chemical entity whose identity, purity, impurities, potency and quantity can be determined and controlled.”

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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The glycosylation of a recombinant protein product can be examined by: 1. Analysis of glycans released by chemical or enzymatic means. 2. Site-specific analysis of glycans associated with glycopeptide fragments following proteolysis of the intact glycoprotein. 3. Direct analysis of the whole glycoprotein.

A number of techniques are currently available to provide rapid and detailed analysis of glycan heterogeneity: 1. High-pH anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD). 2. Enzymatic analysis methods such as the reagent array analysis method (RAAM; 3,4). 3. High-performance capillary electrophoresis (HPCE). 4. Matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS). 5. Electrospray ionization mass spectrometry (ESI-MS).

In particular, novel mass spectrometric strategies continue to rapidly advance the frontiers of biomolecular analysis, with technical innovations and methodologies yielding improvements in sensitivity, mass accuracy, and resolution. 1.1. High-Performance Capillary Electrophoresis Capillary electrophoresis has been employed in various modes in the high-resolution separation and detection of glycoprotein glycoforms, glycoconjugates, glycopeptides, and oligosaccharides, even though carbohydrate molecules do not absorb or fluorese and are not readily ionized (5–8). A number of approaches have been employed to render carbohydrates more amenable to analysis which include in situ complex formation with ions such as borate and metal cations (9) and the addition of ultraviolet (UV)-absorbing or fluorescent tags to functional groups (10). 1.2. MALDI-MS MALDI-MS has been extensively used to determine the mass of proteins and polypeptides, confirm protein primary structure, and to characterize posttranslational modifications. MALDI-MS generally employs simple time-of-flight analysis of biopolymers that are co-crystallized with a molar excess of a low molecular weight, strongly UV absorbing matrix, such as 2,5-dihydroxybenzoic acid, on a metal sample disk. Both the biopolymer and matrix ions are desorbed by pulses of a UV laser. Following a linear flight path the molecular ions are detected, the time between the initial laser pulse and ion detection being directly proportional to the square root of the molecular ion mass/charge (m/z) ratio. For maximum mass accuracy, internal and external protein or peptide calibrants of known molecular mass are required. In addition to this “linear” mode, many instruments offer a “reflectron” mode that effectively lengthens the flight path

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by redirecting the ions toward an additional ion detector that may enhance resolution, at the expense of decreased sensitivity. MALDI-MS is tolerant to low (micromolar) salt concentrations, can determine the molecular weight of biomolecules in excess of 200 kDa with a mass accuracy of ±0.1%, and is capable of analyzing heterogeneous samples with picomole to femtomole sensitivity. These properties combined with its rapid analysis time and ease of use for the nonspecialist have made it an attractive technique for the analysis of glycoproteins, glycopeptides, and oligosaccharides. 1.3. Electrospray Ionization Mass Spectrometry ESI-MS is another mild ionization method, where the covalent bonding of the biopolymer is maintained and is typically used in combination with a single or triple quadrupole. This technique is capable of determining the molecular weight of biopolymers up to 100 kDa with a greater mass accuracy (±0.01%) and resolution (±2000) than MALDI-MS. Multiply charged molecular ions are generated by the ionization of biopolymers in volatile solvents, the resulting spectrum being convoluted to produce noncharged peaks. ESI-MS has been extensively used for the direct mass analysis of glycopeptides and glycoproteins and is often interfaced with liquid chromatography (11–13), but has found limited application for the direct analysis of oligosaccharides (14). ESI-MS is better suited to the analysis of whole glycoprotein populations than MALDI-MS, its superior resolution permitting the identification of individual glycoforms (15, 16). 2. Materials 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

P/ACE 2100 HPCE System (Beckman Instruments Ltd., High Wycombe, UK). Phosphoric acid (Sigma Chemical Co., Poole, UK). Boric acid (Sigma). Trypsin, sequencing grade (Boehringer Mannheim, UK, Lewes, UK). Waters 626 Millenium HPLC System (Millipore Ltd., Watford, UK). Vydac 218TP52 reverse-phase column: C18, 2.1 × 250 mm (Hichrom Ltd., Reading, UK). HPLC grade water-acetonitrile (Fischer Scientific, Loughborough, UK). α-Cyano-4-hydroxy cinnaminic acid (Aldrich Chemical Co., Gillingham, UK). VG Tof Spec Mass Spectrometer (Fisons Instruments, Manchester, UK). Vasoactive intestinal peptide—fragment 1–12 (Sigma). Peptide-N-glycosidase F and glycosidases (Glyko Inc., Upper Heyford, UK). 2,5-Dihydroxybenzoic acid (Aldrich). 2,4,6-Trihydroxyacetophenone (Aldrich). Ammonium citrate (Sigma). VG Quattro II triple quadrupole mass spectrometer (VG Organic, Altrincham, UK). Horse heart myoglobin (Sigma).

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3. Methods This chapter describes some of the recent technological advances in the analysis of posttranslational modifications made to recombinant proteins and focuses on the application of HPCE, MALDI-MS, and ESI-MS to the monitoring of glycosylation heterogeneity. These techniques are illustrated by describing their application to the analysis of recombinant human γ-interferon (IFN-γ), a well-characterized model glycoprotein that has N-linked glycans at Asn25 and at the variably occupied site, Asn97 (17). 3.1. Glycosylation Analysis by HPCE Micellar electrokinetic capillary chromatography (MECC) can be used to rapidly “fingerprint” glycoforms of recombinant human IFN-γ produced by Chinese Hamster Ovary (CHO) cells (8) and to quantitate variable-site occupancy (macroheterogeneity; see Fig. 1). This approach allows glycoforms to be rapidly resolved and quantified without the need for oligosaccharide release, derivatization or labeling. 1. Separations are performed with a P/ACE 2100 capillary electrophoresis system using a capillary cartridge containing a 50 mm internal diameter (i.d.) × 57 cm length of underivatized fused silica capillary. 2. Buffer solutions are prepared from phosphoric and boric acids using NaOH to adjust the pH. 3. Capillaries are prepared for use by rinsing with 0.1 M NaOH for 10 min, water for 5 min, 0.1 M borate, pH 8.5, for 1 h, then 0.1 M NaOH and water for 10 min,

Fig. 1. Whole recombinant human IFN-γ analyzed by capillary electrophoresis. Recombinant human IFN-γ glycoforms were “fingerprinted” by micellar electrokinetic capillary chromatography. Peak groups represent IFN-γ variants with both Asn sites occupied (2N), one site occupied (1N), or no sites occupied (0N).

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respectively. Prior to use, capillaries are equilibrated with electrophoresis buffer (400 mM borate + 100 mM sodium dodecyl sulfate [SDS], pH 8.5) for 1 h. 4. Voltages are applied over a 0.2-min linear ramping period at a detection wavelength of 200 nm and operating temperature of 25°C. Recombinant human IFN-γ (1 mg/ mL in 50 mM borate, 50 mM SDS, pH 8.5) is injected for 5 s prior to electrophoresis at 22 kV. Between each separation, the capillary is rinsed with 0.1 M NaOH, water, and electrophoresis buffer for 5 min, respectively (see Note 1).

3.2. Glycosylation Analysis by MALDI-MS There are only a few reports of the analysis of whole glycoproteins due to the limited resolution of this technique (18). As a result, analysis of intact glycoproteins is generally limited to those proteins that contain one glycosylation site and are < 15–20 kDa (15). MALDI-MS has proved more useful for the identification and characterization of glycopeptides following their separation and purification by reverse-phase HPLC (19). The advantage of this approach over other methods is that sitespecific glycosylation data can be obtained (20,21; see Note 2). This approach has been successfully used to determine the differences in N-linked glycosylation for the Asn25 and Asn97 sites of recombinant human IFN-γ when produced in different expression systems (22), to monitor changes during batch culture (23), and to monitor intracellular populations (24; see Fig. 2). 1. Purified IFN-γ samples are digested with trypsin (1.5 µγ; 50 µg) for 24 h at 30°C. 2. The glycopeptides containing the N-glycan populations are isolated following their separation by reverse-phase HPLC. Samples are applied in 0.06% (v/v) trifluoroacetic acid (TFA) and the peptides separated with a linear gradient (0–70%) of 80% (v/v) aqueous CH3CN with 0.052% TFA over 100 min at a flow rate of 0.1 ml/min. Peptide peaks are detected at a wavelength of 210 nm and collected individually. 3. The glycopeptides are reduced to the aqueous phase in a Speed Vac concentrator, lyophilized overnight, and stored at −20°C. 4. A 0.5-µL aliquot of the digest samples is mixed with 0.5 µL of a saturated solution of α-cyano-4-hydroxy cinnaminic acid in 60% (v/v) aqueous CH3CN and allowed to co-crystallize on stainless steel sample discs. 5. MALDI-MS is performed with a N2 laser at 337 nm. Desorbed positive ions are detected after a linear flight path by a microchannel plate detector and the digitalized output signal adjusted to obtain an optimum output signal-to-noise ratio from 20 averaged laser pulses. Mass spectra are calibrated with an external standard, vasoactive intestinal peptide with an average molecular mass of 1425.5. 6. Digestion of glycopeptides with peptide-N-glycosidase F (PNGaseF) prior to analysis by mass spectrometry confirms the mass of the core peptide. For this determination, 0.5 µL of sample and 0.5 µL of PNGaseF are incubated at 30°C for 24 h and 0.5-µL aliquots are removed for MALDI-MS analysis. 7. Simultaneous digestion of purified glycopeptides with linkage-specific exoglycosidase arrays for the sequential removal of oligosaccharides permit the sequen-cing of

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Fig. 2. Site-specific N-glycosylation of recombinant human IFN-γ examined by MALDIMS analyses of glycopeptides. (A) The complete analytical protocol. The masses of individual N-glycans at a single glycosylation site were calculated by subtracting the known mass of the core peptide moiety from each component in a glycopeptide spectrum. (B) An N-glycan structure was then tentatively assigned, based on mass criteria alone. As ionization is entirely dependent on the core peptide moiety after desialylation, individual N-glycan structures can be quantified. Monosaccharide structures are schematically represented as: (▲), galactose (162.14); (), N-acetylglucosamine (203.20 Da); (●), mannose (162.14 Da); and (★), fucose (146.14 Da).

N-glycans at individual glycosylation sites by MALDI-MS (21–24). Sample (0.5 µL) and glycosidase (0.5 µL), or a combination of glycosidases, are incubated at 30°C for 24 h and 0.5-µL aliquots are removed for MALDI-MS analysis (see Note 3).

3.3. Glycosylation Analysis by ESI-MS ESI-MS has been used to aid the analysis of glycosylation macro- and microheterogeneity and proteolytic cleavage of the C-terminal in conjunction with information obtained by HPLC and MALDI-MS of released oligosaccharides (25; see Note 4).

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1. Spectra are obtained with a VG Quattro II triple quadrupole mass spectrometer having a mass range for singly charged ions of 4000 Da (Fig. 3). 2. Lyophilized proteins are dissolved in 50% aqueous acetonitrile, 0.2% formic acid to a concentration of 0.1 µg/µL and introduced into the electrospray source at 4 µL/min. The mass-to-charge (m/z) range of 600–1800 Da are scanned at 10 s/scan and data are summed for 3–10 min, depending on the intensity and complexity of the spectra.

Fig. 3. Heterogeneous glycoprotein populations directly analyzed by ESI-MS. This technique provides highly resolved mass analyses with a mass accuracy of 0.01% (1 Dalton/10 kDa). In this example, individual transgenic mouse derived recombinant human IFN-γ components were assigned a C-terminal polypeptide cleavage site and an overall monosaccharide composition.

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During each scan, the sample orifice-to-skimmer potential (cone voltage) are scanned from 30 V at m/z 600 to 75 V at m/z 1800. The capillary voltage is set to 3.5 kV. 3. Mass scale calibration employ the multiply charged ion series from a separate introduction of horse heart myoglobin (average molecular mass of 16,951.49). Molecular weights are based on the following atomic weights of the elements: C = 12.011, H = 1.00794, N = 14.00674, O = 15.9994, and S = 32.066 (see Note 5). 4. Background subtracted m/z data are processed by software employing a maximumentropy (MaxEnt) based analysis to produce zero-charge protein molecular weight information with optimum signal-to-noise ratio, resolution, and mass accuracy.

4. Notes 1. Attempts to separate IFN-γ with borate alone, as used by Landers et al. (26) for the separation of ovalbumin glycoforms (26), were unsuccessful, because SDS is required to disrupt the hydrogen-bonded dimers. Application of this technique to ribonuclease B and fetuin met with variable success. The glycoprotein microheterogeneity of a monoclonal antibody with a single glycosylation site has been mapped using a borate buffer at high pH; the glycans were enzymatically or chemically cleaved and the resulting profile used for testing batch-to-batch consistency in conjunction with MALDI-MS analysis (27). 2. MALDI-MS of free N-linked oligosaccharides, following chemical release with hydrazinolysis or enzymatic release with an endoglycosidase such as PNGaseF, is also popular as it requires no prior structural knowledge of the glycoprotein of interest and is ideal for the analysis of underivatized populations of oligosaccharides. However, there is a loss of glycosylation site-specific data. Enzymatic release of oligosaccharides is preferred where an intact deglycosylated protein product is required, as N-glycan release by hydrazinolysis may result in peptide bond cleavage and the oligosaccharide product requires reacetylation. A drawback to the enzymatic release of oligosaccharides is that the presence of SDS is often required to denature the glycoprotein and has to be removed prior to MALDI-MS analysis. MALDI-MS has also been used for the analysis of IFN-γ glycoforms separated by SDS-polyacrylamide gel electrophoresis (PAGE) (28). 3. Until recently, only desialylated oligosaccharides could be analyzed successfully by MALDI-MS using 2,5-dihydroxybenzoic acid as matrix, as negatively charged sialic acids interfere with the efficiency of ionization using this procedure (29,30). However, advances in matrix mixtures and sample preparation schemes now promise to further improve analytical protocols. For example, sialylated oligosaccharides have recently been shown to ionize effectively, with picomole to femtomole sensitivity, as deprotonated molecular ions using 2,4,6-trihydroxyacetophenone as matrix in the presence of ammonium citrate (31). Further improvements in sensitivity and resolution may be obtained by derivatization of oligosaccharides with

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fluorophores such as 2-aminoacridome (AMAC) or 2-aminobenzamidine (AB; Fig. 4). Integration of MALDI-MS peak areas obtained on analysis of sialylated glycans from IFN-γ provided quantitative information that compared favorably with analysis of the derivatized sialylated glycans by ion-exchange HPLC.

Fig. 4. Sialylated N-glycans associated with recombinant human IFN-γ released with PNGaseF, labeled with 2-aminobenzamide, and analyzed by MALDI-MS using 2,4,6trihydroxyacetophenone as matrix (A). The masses of individual N-glycans are used to assign an overall monosaccharide composition, including degree of sialylation (B). H, hexose; N, N-acetylhexosamine; D, deoxyhexose; and S, N-acetylneuraminic acid.

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4. Glycopeptides may be directly analyzed by ESI-MS (32,33), and the oligosaccharides sequenced following digestion with combinations of exoglycosidases (34) or glycoforms separated by liquid chromatography prior to analysis (35). Possibly the most powerful application of this technique has resulted from its interfacing with liquid chromatography which permits the separation and on-line identification of glycoproteins from protein digests (36). Although ESI-MS may be interfaced with capillary electrophoresis, the separation of glycoproteins and glycopeptides under acidic conditions makes the analysis of sialylated oligosaccharides unsuitable. 5. ESI-MS is not ideally suited for the analysis of neutral and anionic oligosaccharides that have been chemically or enzymatically released from a glycoprotein of interest as they do not readily form multiply charged ions. However, the characterization of methylated derivatives of oligosaccharides from recombinant erythropoietin by ESI-MS has been reported (37). References 1. Jenkins, N. and Curling, E. M. (1994) Glycosylation of recombinant proteins: problems and prospects. Enzyme Microb. Technol. 16, 354–364. 2. Liu, D. T. Y (1992) Glycoprotein pharmaceuticals—scientific and regulatory considerations and the United States Orphan Drug Act. Trends Biotechnol. 10, 114–120. 3. Dwek, R. A., Edge, C. J., Harvey, D. J., and Wormald, M. R. (1993) Analysis of glycoprotein associated oligosaccharides. Ann. Rev. Biochem. 62, 65–100. 4. Huberty, M. C., Vath, J. E., Yu, W., and Martin, S. A. (1993) Site-specific carbohydrate identification in recombinant proteins using MALDI-TOF MS. Analyt. Chem. 65, 2791–2800. 5. Frenz, J. and Hancock, W. S. (1991) High performance capillary electrophoresis. Trends Biotechnol. 9, 243–250. 6. Novotny, M. V. and Sudor, J. (1993) High performance capillary electrophoresis of glycoconjugates. Electrophoresis 14, 372–389. 7. Rush, R., Derby, P., Strickland, T., and Rohde, M. (1993) Peptide mapping and evaluation of glycopeptide microheterogeneity derived from endoproteinase digestion of erythropoietin by affinity high-performance capillary electrophoresis. Analyt. Chem. 65, 1834–1842. 8. James, D. C., Freedman,, R. B. Hoare, M., and Jenkins, N. (1994) High resolution separation of recombinant human interferon-γ glycoforms by micellar electrokinetic capillary chromatography. Analyt. Biochem. 222, 315–322. 9. Rudd, P. M., Scragg, I. G., Coghill, E., and Dwek, R. A. (1992) Separation and analysis of the glycoform populations of ribonuclease b using capillary electrophoresis. Glycoconjugate J. 9, 86–91. 10. El-Rassi, Z. and Mechref, Y. (1996) Recent advances in capillary electrophoresis of carbohydrates. Electrophoresis 17, 275–301. 11. Müller, D., Domon, B., Karas, M., van Oostrum, J., and Richter, W. J. (1994) Characterization and direct glycoform profiling of a hybrid plasminogen-activator by

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13. 14.

15.

16.

17. 18.

19.

20. 21.

22.

23.

24.

25.

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matrix-assisted laser-desorption and electrospray mass spectrometry—correlation with high performance liquid-chromatographic and nuclear-magnetic-resonance analyses of the released glycans. Biol. Mass Spectrom. 23, 330–338. Hunter, A. P. and Games, D. E. (1995) Evaluation of glycosylation site heterogeneity and selective identification of glycopeptides in proteolytic digests of bovine α 1-acid glycoprotein by mass spectrometry. Rapid Commun. Mass Spectrom. 9, 42–56. Mann, M. and Wilm, M. (1995) Electrospray mass spectrometry for protein characterization. Trends Biochem. Sci. 20, 219–224. Gu, J., Hiraga, T., and Wada, Y. (1994) Electrospray ionization mass spectrometry of pyridylaminated oligosaccharide derivatives: sensitivity and in-source fragmentation. Biol. Mass Spectrom. 23, 212–217. Tsarbopoulos, A., Pramanik, B. N., Nagabhushan, T. L., and Covey, T. R. (1995) Structural analysis of the CHO-derived interleukin-4 by liquid chromatography electrospray ionization mass spectrometry. J. Mass. Spectrom. 30, 1752–1763. Ashton, D. S., Beddell, C. R., Cooper, D. J., Craig, S. J., Lines, A. C., Oliver, R. W. A., and Smith, M. A. (1995) Mass spectrometry of the humanized monoclonal antibody CAMPATH 1H. Analyt. Chem. 67, 835–842. Hooker, A. D. and James, D. C. (1998) The glycosylation heterogeneity of recombinant human IFN-γ. J. Interf. Cytok. Res. 18, 287–295. Bihoreau, N., Veillon, J. F., Ramon, C., Scohyers, J. M., and Schmitter, J. M. (1995) Rapid Commun. Characterization of a recombinant antihaemophilia-A factor (factor-VIII-delta-II) by matrix-assisted laser desorption ionization mass spectrometry. Rapid Commun. Mass. Spectrom. 9, 1584–1588. Stone, K. L., LoPresti, M. B., Crawford, J. M., DeAngelis, R., and Williams, K. R. (1989) A Practical Guide to Protein and Peptide Purification for Microsequencing, Academic Press, San Diego, CA, p. 36. Treuheit, M. J., Costello, C. E., and Halsall, H. B. (1992) Analysis of the five glycosylation sites of human α1-acid glycoprotein. Biochem. J. 283, 105–112. Sutton, C. W., O’Neill, J., and Cottrell, J. S. (1994) Site specific characterization of glycoprotein carbohydrates by exoglycosidase digestion and laser desorption mass spectrometry. Analyt. Biochem. 218, 34–46. James, D. C., Freedman, R. B., Hoare, M., Ogonah, O. W., Rooney, B. C., Larionov, O. A., et al. (1995) N-Glycosylation of recombinant human interferon-gamma produced in different animal expression systems. BioTechnology 13, 592–596. Hooker, A. D., Goldman, M. H., Markham, N. H., James, D. C., Ison, A. P., Bull, A. T., et al. (1995) N-Glycans of recombinant human interferon-γ change during batch culture of Chinese hamster ovary cells. Biotechnol. Bioeng. 48, 639–648. Hooker, A. D., Green, N. H., Baines, A. J., Bull, A. T., Jenkins, N., Strange, P. G., and James, D. C. (1999) Constraints on the transport and glycosylation of recombinant IFN-γ in Chinese hamster ovary and insect cells. Biotechnol. Bioeng. 63, 559–572. James, D. C., Goldman, M. H., Hoare, M., Jenkins, N., Oliver, R. W. A., Green, B. N., and Freedman, R. B. (1996) Post-translational processing of recombinant human interferon-gamma in animal expression systems. Protein Sci. 5, 331–340.

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26. Landers, J. P., Oda, R. P., Madden, B. J., and Spelsberg, T. C. (1992) High-performance capillary electrophoresis of glycoproteins: the use of modifiers of electroosmotic flow for analysis of microheterogeneity. Analyt. Biochem. 205, 115–124. 27. Hoffstetter-Kuhn, S. H., Alt, G., and Kuhn, R. (1996) Profiling of oligosaccharidemediated microheterogeneity of a monoclonal antibody by capillary electrophoresis. Electrophoresis 17, 418–422. 28. Mortz, E., Sareneva, T., Haebel, S., Julkunen, I., and Roepstorff, P. (1996) Mass spectrometric characterization of glycosylated interferon-gamma variants by gel electrophoresis. Electrophoresis 17, 925–931. 29. Stahl, B., Steup, M., Karas, M., and Hillenkamp, F. (1991) Analysis of neutral oligosaccharides by matrix-assisted laser desorption/ionization mass spectrometry. Analyt. Chem. 63, 1463–1466. 30. Tsarbopoulos, A., Karas, M., Strupat, K., Pramanik, B., Nagabhushan, T., and Hilenkamp, F. (1994) Comparative mapping of recombinant proteins and glycoproteins by plasma desorption and matrix-assisted laser desorption/ionization mass spectrometry. Analyt. Chem. 66, 2062–2070. 31. Papac, D., Wong, A., and Jones, A. (1996) Analysis of acidic oligosaccharides and glycopeptides by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Analyt. Chem. 68, 3215–3223. 32. Rush, R. S., Derby, P. L., Smith, D. M., Merry, C., Rogers, G., Rohde, M. F., and Katta, V. (1995) Microheterogeneity of erythropoietin carbohydrate structure. Analyt. Chem. 67, 1442–1452. 33. Bloom, J. W., Madanat, M. S., and Ray, M. K. (1996) Cell-line and site specific comparative analysis of the N-linked oligosaccharides on human ICAM/-1DES454-532 by electrospray mass spectrometry. Biochemistry 35, 1856–1864. 34. Schindler, P. A., Settineri, C. A., Collet, X., Fielding, C. J., and Burlingame, A. L. (1995) Site-specific detection and structural characterization of the glycosylation of human plasma proteins lecithin:cholesterol acyltransferase and apolipoprotein D using HPLC/electrospray mass spectrometry and sequential glycosidase digestion. Protein Sci. 4, 791–801. 35. Medzihradszky, K. F., Maltby, D. A., Hall, S. C., Settineri, C. A., and Burlingame, A. L. (1994) Characterization of protein N-glycosylation by reverse-phase microbore liquid chromatography electrospray mass spectrometry, complimentary mobile phases and sequential exoglycosidase digestion. J. Am. Soc. Mass Spectrom. 5, 350–358. 36. Ling, V., Guzzetta, A. W., Canova-Davis, E., Stults, J. T., Hancock, W. S., Covey, T. R., and Shushan, B. I. (1991) Characterization of the tryptic map of recombinant DNA derived tissue plasminogen activator by high performance liquid chromatography-electrospray ionization mass spectrometry. Analyt. Chem. 63, 2909–2915. 37. Linsley, K. B., Chan, S. Y., Chan, S., Reinhold, B. B., Lisi, P. J., and Reinhold, V. N. (1994) Applications of electrospray mass-spectrometry to erythopoietin N-linked and O-linked glycans. Analyt. Biochem. 219, 207–217.

147 Affinity Chromatography of Oligosaccharides and Glycopeptides with Immobilized Lectins Kazuo Yamamoto

1. Introduction Sugar moieties on the cell surface play one of the most important roles in cellular recognition. In order to elucidate the molecular mechanisms of these cellular phenomena, assessment of the structure of sugar chains is indispensable. However, it is difficult to elucidate the structures of cell-surface oligosaccharides because of two technical problems. The first is the difficulty in fractionating various oligosaccharides that are heterogeneous in the number, type, and substitution patterns of the outer sugar branches. The second problem is that only very limited amounts of material may be available, making it difficult to perform detailed structural studies. Lectins are proteins with sugar-binding activity. Each lectin binds to a specific sugar sequence in oligosaccharides and glycopeptides. Therefore, lectins are very useful tools for overcoming the problems described above. Recently, many attempts have been made to fractionate oligosaccharides and glycopeptides on immobilized-lectin columns. The use of a series of immobilized-lectin columns, with sugar-binding specificities that have been precisely elucidated, has enabled us to fractionate very small amounts (~10 ng) of oligosaccharides or glycopeptides into structurally distinct groups. Here, we summarize our serial lectin-Sepharose affinity-chromatographic techniques for the rapid, sensitive, and specific fractionation and analysis of asparagine-linked oligosaccharides of glycoproteins. The structures of asparagine-linked oligosaccharides fall into three main categories, termed high mannose-type, complex-type, and hybrid-type oligosaccharides (1). They share the common core structure, Manα1-3(Manα1-6)

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Manβ1-4GlcNAcβ1-4GlcNAc-Asn, but differ in their outer branches (Fig. 1). High mannose-type oligosaccharides have two to six α-mannose residues added to the core structure. Typical complex-type oligosaccharides contain two to four outer branches with a sialyllactosamine sequence. Hybrid-type structures have the features of both high mannose-type and complex-type oligosaccharides and most contain a bisecting N-acetylglucosamine, which has a β1-4 linkage to the β-linked mannose residue of the core structure. A novel type of carbohydrate chain, called the poly-N-acetyllactosamine-type oligosaccharide, also has been described (2–5). Its outer branches have a characteristic structure composed of N-acetyllactosamine repeating units. Although it may be classified as a complextype oligosaccharide, it is antigenically and functionally distinct from standard complex-type sugar chains (4). Some poly-N-acetyllactosamine-type oligosaccharides have branched sequences containing Galβ1-4GlcNAcβ1-3(Galβ14GlcNAcβ1-6)Gal units (2, 3), which is the determinant of the I-antigen. Another novel complex-type sugar chain, having GalNAcβ1-4GlcNAc groups in its outer chain moieties, has been found (6, 7) and GalNAc residues are sometimes sulfated

high mannose-type Manα1-2Manα1 6 Manα1 3 Manα1-2Manα1

6 Manβ1-4GlcNAcβ1-4GlcNAc-Asn 3

Manα1-2Manα1-2Manα1 complex-type

Fucα1

NeuAcα2-6Galβ1-4GlcNAcβ1-2Manα1

6 6 3 Manβ1-4GlcNAcβ1-4GlcNAc-Asn

NeuAcα2-6Galβ1-4GlcNAcβ1-2Manα1

hybrid-type Manα1-2Manα1

GlcNAcβ1 6 Manα1 3 4 6 Manβ1-4GlcNAcβ1-4GlcNAc-Asn Manα1-2Manα1 3 Galβ1-4GlcNAcβ1-2Manα1

Fig. 1. Structures of major types of asparagine-linked oligosaccharides. The boxed area encloses the core structure common to all asparagine-linked oligosaccharides.

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at the C-4 position or sialylated at the C-6 position. Complex-type sugar chains with non-reducing Galα1-3(4)Gal structures have also been reported (8). Glycopeptides or oligosaccharides can be prepared from glycoproteins by enzymatic digestions or chemical methods, as discussed in subsequent chapters in this book. The most widely used means for preparing glycopeptides is to completely digest the material with pronase. Oligosaccharides can be prepared from glycoproteins or glycopeptides by treating samples with anhydrous hydrazine (9) or endoglycosidases. Since the released oligosaccharides retain their reducing termini, they can be radiolabeled by reduction with NaB3H4 (10) or fluorescently labeled by 2-aminopyridine (11). Primary amino groups of the peptide backbone of glycopeptides are labeled by acetylation with [3H]- or [14C]-acetic anhydride (12). Before employing columns of immobilized lectins for analyses, oligosaccharides or glycopeptides should be separated on a column of QAE- or DEAE-cellulose, based on the anionic charge derived from sialic acid, phosphate, or sulfate residues. Separated acidic oligosaccharides should be converted to neutral oligosaccharides to simplify the subsequent separations. For simplicity, the oligosaccharides discussed here do not contain sialic acid, phosphate, or sulfate residues, although these acidic residues, especially sialic acid residues, are found in many oligosaccharides. In most cases, the influence of these residues on the interaction of oligosaccharides with immobilized lectins is weak, but where documentation of the effects of these residues is available, it will be mentioned in the appropriate sections. Here we describe the general procedure for serial lectin affinity chromatography of glycopeptides and oligosaccharides using several well-defined immobilized lectins. 2. Materials 1. Mono Q HR5/50 GL, DEAE-Sepharose, Sephadex G-25 (GE Healthcare Bio-Sciences AB, Uppsala, Sweden). 2. High performance liquid chromatograph, two pumps, with detector capable of monitoring ultraviolet absorbance at 220 nm. 3. Neuraminidase: 1 unit/ml of neuraminidase from Streptococcus sp. (Seikagaku Kogyo, Tokyo, Japan) in 50 mM acetate buffer, pH 6.5. 4. Dowex 50W-X8 (50–100 mesh, H+ form). 5. Bio-Gel P-4 minus 400 mesh (Bio-Rad, Richmond, CA). 6. HPLC mobile phase for Mono Q: A, 2 mM Tris-HCl, pH 7.4; B, 2 mM Tris-HCl, pH 7.4, 0.5 M NaCl. 7. HPLC mobile phase for Bio-Gel P-4: distilled water. 8. HPLC standard for Bio-Gel P-4: partial hydrolysate of chitin, prepared according to Rupley (13); 10 µg mixed with 50 µl distilled water. Store frozen. 9. Concanavalin A, Ricinus communis lectin, wheat germ lectin, Datura stramonium lectin, Maackia amurensis leukoagglutinin, Wistaria floribunda lectin, Allomyrina dichotoma lectin, Amaranthus caudatus lectin, Griffonia (Bandeiraea) simplicifolia

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10. 11. 12. 13. 14. 15. 16. 17.

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isolectin B4 (EY Laboratories, San Mateo, CA), Phaseolus vulgaris erythroagglutinin, P. vulgaris leukoagglutinin (Seikagaku Kogyo). Immobilized lectins are prepared at a concentration of 1–5 mg lectin/ml of gel (see Notes 1 and 2). The following may be obtained commercially as immobilized lectins: Galanthus nivalis lectin, Lens culinaris lectin, Pisum sativum lectin, Vicia fava lectin, pokeweed mitogen, Sambucus nigra L. lectin (e.g., GE Healthcare Bio-Sciences AB, EY Laboratories, Bio-Rad, Seikagaku Kogyo). 3 H-NaB3H4: 3.7 × 109 Bq of 3H-NaBH4 (sp. 1.9–5.6 × 1011 Bq/mmol; PerkinElmer, Boston, MA) mixed with 2 ml 10 mM NaOH; store at −80°C. Tris-buffered saline (TBS): 10 mM Tris-HCl, pH 7.4, 0.15 M NaCl. Lectin column buffer: 10 mM Tris-HCl, pH 7.4, 0.15 M NaCl, 1 mM CaCl2, 1 mM MnCl2 (see Note 3). N-acetylgalactosamine (Sigma, St Louis, MO): 100 mM in lectin column buffer; store refrigerated. Methyl-α-mannoside (Sigma): 100 mM in TBS; store refrigerated. Methyl-α-glucoside (Sigma): 10 mM in TBS; store refrigerated. Lactose (Sigma): 50 mM in TBS; store refrigerated. N-acetylglucosamine (Sigma): 200 mM in TBS; store refrigerated.

3. Methods 3.1. Separation of Acidic Sugar Chains on Mono Q or DEAE-Sepharose and Removal of Sialic Acids. 3.1.1. Ion Exchange Chromatography 1. Equilibrate the Mono Q or DEAE-Sepharose column with 2 mM Tris-HCl, pH 7.4, at a flow rate of 1 ml/min at room temperature. 2. Dissolve the oligosaccharides or glycopeptides in 0.1 ml of 2 mM Tris-HCl, pH 7.4, and apply to the column. 3. Elute with 2 mM Tris-HCl, pH 7.4, for 10 min, followed by a linear gradient (0-20%) of 2 mM Tris-HCl, pH 7.4, 0.5 M NaCl for 60 min, at a flow rate of 1 ml/min. 4. Neutral oligosaccharides are recovered in the pass-through fraction. Acidic monosialo-, disialo-, trisialo-, and tetrasialooligosaccharides are eluted successively by the linear gradient of NaCl.

3.1.2. Removal of Sialic Acid Residues 1. To 10–100 µg oligosaccharides, free of buffers and salts, add 100 µl of neuraminidase buffer and 100 µl of neuraminidase and incubate at 37°C for 18 h. 2. Heat-inactivate the neuraminidase by immersion in a boiling-water bath for 3 min. 3. Apply to the Dowex 50W-X8 column (0.6 cm id × 2.5 cm), wash the column with 1 ml of distilled water, and concentrate the eluate under vacuum.

Alternatively, add 500 µl of 0.1 M HCl to 10–100 µg oligosaccharides, heat at 80°C for 30 min, and dry the sample using an evaporator.

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3.2. Separation of Poly-N-Acetyllactosamine-Type Sugar Chains from Other Types of Sugar Chains Poly-N-acetyllactosamine-type sugar chains vary as to the number of N-acetyllactosamine repeating units and their branching modes. Thus, the structural characterization of poly-N-acetyllactosamine-type sugar chains has been quite difficult (14). This type of sugar chain has a higher molecular weight than high mannose-type, complex-type, or hybrid-type chains. PolyN-acetyllactosamine-type sugar chains with molecular masses of more than 4,000 are excluded in the Bio-Gel P4 column chromatography (2, 15, 16) and thus are easily separated from other types. 1. Equilibrate two coupled Bio-Gel P-4 columns (0.8 cm id × 50 cm) in water at 55°C using a water jacket. 2. Elute the oligosaccharides at a flow rate of 0.3 ml/min and collect 0.5 ml fractions. Monitor absorbance at 220 nm. 3. Collect poly-N-acetyllactosamine-type oligosaccharides that elute in the void volume of the column. Other types of oligosaccharides are subjected to subsequent separations (Subheadings 3.3–3.6), as illustrated in Fig. 2. The specificities of the lectins are summarized in Fig. 3 and Table 1.

3.3. Separation of Complex-Type Sugar Chains Containing GalNAc 1-4GlcNAc Groups from Other Sugar Chains Novel complex-type oligosaccharides and glycopeptides with GalNAcβ14GlcNAcβ1–2 outer chains bind to a W. floribunda lectin (WFA)-Sepharose column (7). 1. Equilibrate the WFA-Sepharose column (0.6 cm id × 5.0 cm) in lectin column buffer. 2. Dissolve oligosaccharides or glycopeptides in 0.5 ml of lectin column buffer and apply to the column. 3. Elute 1.0-ml fractions with three column-volumes of lectin column buffer and then with three column-volumes of 100 mM N-acetylgalactosamine, at a flow rate of 2.5 ml/h at room temperature. 4. Collect other types of sugar chains, which pass through the column. 5. Collect complex-type sugar chains with GalNAcβ1-4GlcNAc outer chains, which elute after the addition of N-acetylgalactosamine.

3.4. Separation of High Mannose Type Sugar Chains from Complex Type and Hybrid Type Sugar Chains 3.4.1. Affinity Chromatography on Immobilized RCA After the separation of high-molecular-weight poly-N-acetyllactosaminetype oligosaccharides, a mixture of the other three types of sugar chains can

1352

Yamamoto Bio-Gel P-4 poly-N-acetyllactosamine-type PWM branched

WFA other types

complex-type with GalNAc-GlcNAc

unbranched

GS-IB4 complex-type with terminal α-Gal high mannose-type

RCA

hybrid-type complex-type WGA

hybrid-type

complex-type Con A

triantennary +bisecting GN

tri-,tetraantennary

biantennary

E-PHA

LCA

tri-, tetraantennary

biantennary +Fuc

biantennary -Fuc

L-PHA tri-2,6-branched tetraantennary

tri-2,4-branched

Fig. 2. Fractionation scheme for asparagine-linked sugar chains by affinity chromatography with immobilized lectins

be separated on a column of R. communis lectin (RCA), which recognizes the Galβ1-4GlcNAc sequence (17, 18). 1. Equilibrate the RCA-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Dissolve oligosaccharides or glycopeptides in 0.5 ml TBS and apply to the column. 3. Elute 1.0-ml fractions with three column-volumes of TBS, and then with three column-volumes of 50 mM lactose, at a flow rate of 2.5 ml/h at room temperature. 4. Bind both complex-type and hybrid-type sugar chains to the RCA-Sepharose column (see Note 4). 5. Collect high mannose-type oligosaccharides, which pass through the column.

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GlcNAcβ1

NeuAcα2-6Galβ1-4GlcNAcβ1-2Manα1

Fucα1

4 6 6 3 Manβ1-4GlcNAcβ1-4GlcNAc-Asn

NeuAcα2-3Galβ1-4GlcNAcβ1-2Manα1

triantennary complex-type Galβ1-4GlcNAcβ1 6 2 Manα1

Galβ1-4GlcNAcβ1

6 Manβ1-4GlcNAcβ1-4GlcNAc-Asn 3

Galβ1-4GlcNAcβ1-2Man α1

GlcNAcβ1 Galβ1-4GlcNAcβ1-2Manα1 4 6 Manβ1-4GlcNAcβ1-4GlcNAc-Asn 3

Galβ1-4GlcNAcβ1 4 2 Manα1

Galβ1-4GlcNAcβ1 tetraantennary complex-type Galβ1-4GlcNAcβ1

6 2 Manα1

Galβ1-4GlcNAcβ1

6 3 Manβ1-4GlcNAcβ1-4GlcNAc-Asn

Galβ1-4GlcNAcβ1 4 Manα1 2

Galβ1-4GlcNAcβ1 unbranched poly-N-acetyllactosamine-type

Galβ1-4GlcNAcβ1-(-3Galβ1-4GlcNAcβ1-)n -2Manα1 6 Manβ1-4GlcNAcβ1-4GlcNAc-Asn 3 Galβ1-4GlcNAcβ1-(-3Galβ1-4GlcNAcβ1-)n -2Manα1 branched poly-N-acetyllactosamine-type ---4GlcNAcβ1 6 Galβ1------4GlcNAcβ1-3Galβ1-4GlcNAcβ1-- 2Manα1 6 Manβ1-4GlcNAcβ1-4GlcNAc-Asn 3 ---4GlcNAcβ1-3Galβ1-4GlcNAcβ1-- 2Manα1 6 ---4GlcNAcβ1 n

complex-type with GalNAc-GlcNAc or terminal α-Gal −

SO3-4GalNAcβ1-4GlcNAcβ1-2Manα1

6 Manβ1-4GlcNAcβ1-4GlcNAc-Asn 3

Galα1-3(4)Galβ1-4GlcNAcβ1-2Manα1

Fig. 3. Structures of several complex-type oligosaccharides. The boxed areas indicate the characteristic structures that are recognized by immobilized lectins.

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6. Purify the oligosaccharides or glycopeptides from salts and haptenic sugar by gel filtration on a Sephadex G-25 column (1.2 cm id × 50 cm) equilibrated with distilled water.

3.4.2. Affinity Chromatography on Immobilized Snowdrop Lectin High mannose-type glycopeptides, which carry Manα1-3Man units are specifically retarded on immobilized snowdrop (G. nivalis) lectin (GNA) (19). 1. Equilibrate the GNA-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Dissolve oligosaccharides or glycopeptides in 0.5 ml TBS and apply to the column. 3. Elute 0.5-ml fractions with five column-volumes of TBS to collect sugar chains lacking Manα1-3Man units or hybrid-type sugar chains, which are not retarded. 4. Elute with three column-volumes of 100 mM methyl-α-mannoside at a flow rate of 2.5 ml/h at room temperature to obtain the specifically retarded high mannose-type glycopeptides that carry Manα1-3Man units.

3.5. Separation of Hybrid Type Sugar Chains from Complex Type Sugar Chains 3.5.1. Affinity Chromatography on Immobilized WGA Wheat germ lectin (WGA)-Sepharose has a high affinity for hybrid-type sugar chains. It has been demonstrated that the GlcNAcβ1-4Manβ1-4GlcNAcβ14GlcNAc-Asn structure is essential for tight binding of glycopeptides to a WGA-Sepharose column (20). 1. 2. 3. 4.

Equilibrate the WGA-Sepharose column (0.6 cm id × 5.0 cm) in TBS. Dissolve glycopeptides in 0.5 ml TBS and apply to the column. Elute 0.5-ml fractions with five column-volumes of TBS. Collect the typical complex-type (and also high mannose-type) sugar chains eluted in the void volume of the column. 5. Collect hybrid-type glycopeptides with a bisecting N-acetylglucosamine residue, which are retarded on the WGA column.

3.6. Separation of Complex Type Biantennary Sugar Chains 3.6.1. Affinity Chromatography on Immobilized Concanavalin A (Con A) Oligosaccharides and glycopeptides with tri- and tetraantennary complex-type sugar chains pass through Con A-Sepharose, whereas biantennary complex-type, hybrid-type, and high mannose-type sugar chains bind to Con A and can be differentially eluted from the column (21, 22). 1. Equilibrate the Con A-Sepharose column (0.6 cm id × 5.0 cm) in lectin column buffer. 2. Pass the oligosaccharide mixture of complex-type chains from the WGA column through the Con A-Sepharose column. 3. Elute 1-ml fractions with three column-volumes of lectin column buffer.

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4. Collect oligosaccharides with tri- and tetraantennary complex-type sugar chains, which pass through the column. Complex-type biantennary glycopeptides or oligosaccharides having bisecting GlcNAc also pass through the column. 5. Elute 1-ml fractions with three column-volumes of 10 mM methyl-α-glucoside and then with three column-volumes of 100 mM methyl-α-mannoside. 6. Collect complex-type biantennary sugar chains, which elute after the addition of methyl-α-glucoside. 7. Collect high mannose-type and hybrid-type oligosaccharides or glycopeptides eluted after the addition of methyl-α-mannoside.

3.6.2. Affinity Chromatography on Immobilized LCA, PSA, or VFA The biantennary complex-type sugar chains bound to the Con A-Sepharose column and eluted with 10 mM methyl-α-glucoside include two types of oligosaccharides, which can be separated on a column of lentil (L. culinaris) lectin (LCA), pea (P. sativum) lectin (PSA), or fava (V. fava) lectin (VFA) (23–25) (see Note 5). 1. Equilibrate the LCA-, PSA-, or VFA-Sepharose column (0.6 cm id × 5.0 cm) in lectin column buffer. 2. Pass the biantennary complex-type sugar chains from the Con A column through the LCA-, PSA-, or VFA-Sepharose column. 3. Elute 1.0-ml fractions with three column-volumes of lectin column buffer, followed by three column-volumes of 100 mM methyl-α-mannoside, at a flow rate of 2.5 ml/h at room temperature. 4. Collect biantennary complex-type sugar chains without fucose, which pass through the column. 5. Elute bound biantennary complex-type sugar chains having a fucose residue attached to the innermost N-acetylglucosamine.

3.6.3. Affinity Chromatography on Immobilized E-PHA Complex-type biantennary sugar chains having outer galactose residues and bisecting N-acetylglucosamine are retarded by P. vulgaris erythroagglutinin (E-PHA)-Sepharose (18, 26). 1. Equilibrate the E-PHA-Sepharose column (0.6 cm id × 5.0 cm) in lectin column buffer. 2. Apply the pass-through fraction from the Con A column to the E-PHA-Sepharose column. 3. Elute 0.5-ml fractions with five column-volumes of lectin column buffer at a flow rate of 2.5 ml/h at room temperature. 4. Collect biantennary complex-type sugar chains having a bisecting N-acetylglucosamine residue, which were retarded on the E-PHA column (see Note 6). When the column is eluted at 4°C, biantennary complex-type oligosaccharides without a bisecting N-acetylglucosamine are also retarded by the E-PHASepharose column.

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3.7. Separation of Complex Type Triantennary and Tetraantennary Sugar Chains 3.7.1. Affinity Chromatography on Immobilized E-PHA E-PHA-Sepharose interacts with high affinity with triantennary (having 2,4branched mannose) oligosaccharides or glycopeptides containing both outer galactose residues and a bisecting N-acetylglucosamine residue (26). 1. Equilibrate the E-PHA-Sepharose column (0.6 cm id × 5.0 cm) in lectin column buffer. 2. Apply the pass-through fraction from the Con A column to the E-PHA-Sepharose column. 3. Elute 0.5-ml fractions with five column-volumes of lectin column buffer at a flow rate of 2.5 ml/h at room temperature. 4. Collect retarded triantennary (having 2,4-branched mannose) oligosaccharides or glycopeptides containing both outer galactose and bisecting N-acetylglucosamine. Other tri- and tetraantennary oligosaccharides pass through the column (see Note 7).

3.7.2. Affinity Chromatography on Immobilized L-PHA P. vulgaris leukoagglutinin (L-PHA), which is an isolectin of E-PHA, interacts with triantennary and tetraantennary complex-type glycopeptides having an α-linked mannose residue substituted at positions C-2 and C-6 with Galβ14GlcNAc (27). 1. Equilibrate the L-PHA-Sepharose column (0.6 cm id × 5.0 cm) in lectin column buffer. 2. Apply the pass-through fraction from the Con A column to the L-PHA-Sepharose column. 3. Elute 0.5-ml fractions) with five column-volumes of lectin column buffer at a flow rate of 2.5 ml/h at room temperature.4. Collect retarded triantennary and tetraantennary complex-type glycopeptides having both 2,6-branched α-mannose and outer galactose (see Note 8). Other tri- and tetraantennary oligosaccharides pass through the column.

3.7.3. Affinity Chromatography on Immobilized DSA D. stramonium lectin (DSA) has high affinity for tri- and tetraantennary complex-type oligosaccharides. Triantennary complex-type oligosaccharides containing 2,4-substituted α-mannose are retarded by the DSA-Sepharose column. Triantennary and tetraantennary complex-type oligosaccharides having an α-mannosyl residue substituted at the C-2 and C-6 positions bind to the column and are eluted by GlcNAc oligomers (28, 29). 1. Equilibrate the DSA-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Apply the pass-through fraction from the Con A column to the DSA-Sepharose column.

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3. Elute 0.5-ml fractions with three column-volumes of TBS at a flow rate of 2.5 ml/h at room temperature to obtain the retarded triantennary complex-type sugar chains having 2,4-branched α-mannose. 4. Elute with three column-volumes of 5 mg/ml N-acetylglucosamine oligomer at a flow rate of 2.5 ml/h at room temperature to obtain bound triantennary and tetraantennary complex-type oligosaccharides having an α-mannose residue substituted at the C-2 and C-6 positions.

3.8. Separation of Poly-N-Acetyllactosamine Type Sugar Chains High-molecular-weight poly-N-acetyllactosamine-type oligosaccharides are classified into two groups: branched poly-N-acetyllactosaminoglycan containing a Galβ1-4GlcNAcβ1-3(Galβ1-4GlcNAcβ1-6)Gal unit and a linear poly-Nacetyllactosamine structure that lacks galactose substitutions at the C-3 and C-6 positions. 3.8.1. Affinity Chromatography on Immobilized PWM Branched poly-N-acetyllactosamine-type oligosaccharides can be separated using a pokeweed mitogen (PWM)-Sepharose column (30). Since the sugar sequence Galββ1-4GlcNAcβ1-6Gal firmly binds to the PWM-Sepharose column, the branched poly-N-acetyllactosamine chains can be retained and the unbranched chains recovered without any retardation (31) (see Note 9). 1. Equilibrate the PWM-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Apply the poly-N-acetyllactosamine-type sugar chains separated on Bio-Gel P-4 (see Subheading 3.2) to the PWM-Sepharose column. 3. Elute 1.0-ml fractions with three column-volumes of TBS, followed by three columnvolumes of 0.1 M NaOH, at a flow rate of 2.5 ml/h at room temperature. 4. Collect the unbranched poly-N-acetyllactosamine-type sugar chains, which pass through the column. 5. Collect the bound branched poly-N-acetyllactosamine-type sugar chains.

3.8.2. Affinity Chromatography on Immobilized DSA Immobilized DSA lectin interacts with high affinity with sugar chains having a linear, unbranched poly-N-acetyllactosamine sequence. For binding to DSA-Sepharose, more than two intact N-acetyllactosamine repeating units may be essential (29). 1. Equilibrate the DSA-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Apply the poly-N-acetyllactosamine-type sugar chains separated on Bio-Gel P-4 (see Subheading 3.2) to the DSA-Sepharose column. 3. Elute 1.0-ml fractions with three column-volumes of TBS, followed by three columnvolumes of 5 mg/ml GlcNAc oligomer, at a flow rate of 2.5 ml/h at room temperature. 4. Collect branched poly-N-acetyllactosamine-type sugar chains, which pass through the column and are separated from unbranched poly-N-acetyllactosamine-type sugar chains, which bind to the column.

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3.9. Separation of Terminally Sialylated or b-galactosylated sugar chains The basic Galβ1-4GlcNAc sequence present in complex-type sugar chains usually contains sialic acids in α2,6 or α2,3 linkages in the outer galactose residues. 3.9.1. Affinity chromatography on immobilized MAL M. amurensis leukoagglutinin (MAL) (32, 33) interacts with high affinity with complex-type tri- and tetraantennary glycopeptides containing an outer sialic acid residue linked α2,3 to the penultimate galactose. Glycopeptides containing sialic acid linked only α2,6 to galactose do not interact detectably with the immobilized MAL (see Note 10). 1. Equilibrate the MAL-Sepharose column (0.6 cm id × 5.0 cm) in lectin column buffer. 2. Apply the acidic oligosaccharides or glycopeptides separated on Mono Q or DEAE-Sepharose (see Subheading 3.1.1, step 1) to the MAL-Sepharose column. 3. Elute 0.5-ml fractions with five column-volumes of lectin column buffer at a flow rate of 2.5 ml/h at room temperature. 4. Collect glycopeptides or oligosaccharides containing α2,6-linked sialic acid(s), which pass through the column. 5. Collect retarded glycopeptides or oligosaccharides containing α2,3-linked sialic acid(s).

3.9.2. Affinity Chromatography on Immobilized Allo A A. dichotoma lectin (allo A) (34, 35) recognizes the other isomer of sialyllactosamine. Mono-, di-, and triantennary complex-type oligosaccharides containing terminal sialic acid(s) in α2,6 linkages bind to allo A-Sepharose, whereas complex-type sugar chains having isomeric α2,3-linked sialic acid(s) do not bind to immobilized allo A. 1. Equilibrate the allo A-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Apply the acidic oligosaccharides or glycopeptides separated on Mono Q or DEAESepharose (see Subheading 3.1.1, step 1) to the allo A-Sepharose column. 3. Elute 0.5-ml fractions with three column volumes of TBS and then with three column-volumes of 50 mM lactose at a flow rate of 2.5 ml/h at room temperature. 4. Collect glycopeptides or oligosaccharides containing α2,3-linked sialic acid(s), which pass through the column. 5. Elute bound glycopeptides or oligosaccharides having α2,6-linked sialic acid(s) (see Note 11).

3.9.3. Affinity Chromatography on Immobilized SNA Elderberry (S. nigra L.) bark lectin (SNA) (36, 37) has the same sugar-binding specificity as allo A. All types of oligosaccharides with at least one NeuAcα26Gal unit bind firmly to SNA-Sepharose.

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1. Equilibrate the SNA-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Apply the acidic oligosaccharides or glycopeptides separated on Mono Q or DEAE-Sepharose (see Subheading 3.1.1, step 1) to the SNA-Sepharose column. 3. Elute 0.5-ml fractions with three column-volumes of TBS, followed by three column-volumes of 50 mM lactose, at a flow rate of 2.5 ml/h at room temperature. 4. Collect glycopeptides or oligosaccharides containing α2,3-linked sialic acid(s), which pass through the column. 5. Bound glycopeptides or oligosaccharides having α2,6-linked sialic acid(s) elute in the 50 mM lactose.

3.9.4. Affinity Chromatography on Immobilized GS-IB4 Griffonia (Bandeiraea) simplicifolia isolectin B4 (GS-IB4) binds specifically and with high affinity to complex-type sugar chains containing terminal Galα13Gal or Galα1-4Gal sequences (8, 38). Although G. simplicifolia isolectin A4 (GS-IA4) and GS-IB4 bind α-galactosyl groups with equal affinity, B4 has a Ka for α-N-acetylgalactosaminyl groups that is three orders of magnitude smaller than that of A4. 1. Equilibrate the GS-IB4-Sepharose column (0.6 cm id × 5.0 cm) in TBS. 2. Apply the neutral oligosaccharides or glycopeptides onto the GS-IB4-Sepharose column. 3. Elute 0.5-ml fractions with three column-volumes of TBS, followed by three column volumes of 100 mM methyl α-galactoside, at a flow rate of 2.5 ml/h at room temperature. 4. Collect glycopeptides or oligosaccharides without the Galα1-3(4)Gal sequence, which pass through the column. 5. Retarded or bound glycopeptides or oligosaccharides having α-galactosyl residue(s) elute in the 100 mM methyl α-galactoside eluant.

4. Notes 1. During the coupling reactions, sugar-binding sites of lectins must be protected by the addition of the specific haptenic sugars. 2. Immobilized lectin is stored at 4°C. In most cases, immobilized lectin is stable for several years. 3. Some lectins, especially leguminous lectins, need Ca2+ and Mn2+ ions for sugar binding. Therefore, the buffers used for affinity chromatography on the lectin column should contain 1 mM CaCl2 and 1 mM MnCl2. 4. Complex-type and hybrid-type oligosaccharides are retarded on, rather than tightly bound to, a RCA-Sepharose column when their sugar sequences are masked by sialic acids or α-galactosyl residues. 5. Intact N-acetylglucosamine and asparagine residues at the reducing end are required for tight binding of complex-type oligosaccharides to LCA-, PSA-, or VFA-Sepharose columns.

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6. High-affinity interaction with E-PHA-Sepharose is prevented if both outer galactose residues on a bisected sugar chain are substituted at position C-6 by sialic acid. 7. Biantennary and triantennary complex-type sugar chains having bisecting GlcNAc can be separated on a Bio-Gel P-4 column. 8. L-PHA-Sepharose does not retard the elution of sugar chains lacking outer galactose residues. 9. WGA can be used instead of PWM. 10. M. amurensis hemagglutinin (MAH), which is an isolectin of MAL, binds strongly to sialylated Ser/Thr-linked Galβ1-3GalNAc, but not to sialylated Asn-linked sugar chains (38). 11. Mono-, di-, tri-, and tetraantennary complex-type oligosaccharides without sialic acid(s) are retarded by the allo A lectin column. 12. More detailed reviews on the separation of oligosaccharides and glycopeptides by means of affinity chromatography on immobilized lectin columns have been published (39, 40).

References 1. Kornfeld, R. and Kornfeld, S. (1985) Assembly of asparagine-linked oligosaccharides. Ann. Rev. Biochem. 54, 631–664. 2. Tsuji, T., Irimura, T., and Osawa, T. (1981) The carbohydrate moiety of Band 3 glycoprotein of human erythrocyte membrane. J. Biol. Chem. 256, 10497–10502. 3. Fukuda, M., Dell, A., Oates, J. E., and Fukuda, M. N. (1984) Structure of branched lactosaminoglycan, the carbohydrate moiety of Band 3 isolated from adult human erythrocytes. J. Biol. Chem. 259, 8260–8273. 4. Merkle, R. K. and Cummings, R. D. (1987) Relationship of the terminal sequences to the length of poly-N-acetyllactosamine chains in asparagine-linked oligosaccharides from the mouse lymphoma cell line BW5147. J. Biol. Chem. 282, 8179–8189. 5. Fukuda, M. (1985) Cell surface glycoconjugates as onco-differentiation markers in hematopoietic cells. Biochem. Biophys. Acta 780, 119–150. 6. Green, E. D. and Baenziger, J. U. (1988) Asparagine-linked oligosaccharides on lutropin, follitropin, and thyrotropin: I. Structural elucidation of the sulfated and sialylated oligosaccharides on bovine, ovine, and human pituitary glycoprotein hormones. J. Biol. Chem. 263, 25–35. 7. Nakata, N., Furukawa, K., Greenwalt, D. E., Sato, T., and Kobata, A. (1993) Structural study of the sugar chains of CD36 purified from bovine mammary epithelial cells: Occurrence of novel hybrid-type sugar chains containing the Neu5Acα26GalNAcβ1-4GlcNAc and the Manα1-2Manα1-3Manα1-6Man groups. Biochemistry 32, 4369–4383. 8. Suzuki, N., Khoo, K. -H., Chen, H. -C., Johnson, J. R., and Lee, Y. C. (2001) Isolation and characterization of major glycoproteins of pigion egg shite. J. Biol. Chem. 276, 23221–23229.

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9. Fukuda, M., Kondo, T., and Osawa, T. (1976) Studies on the hydrazinolysis of glycoproteins. Core structures of oligosaccharides obtained from porcine thyroglobulin and pineapple stem bromelain. J. Biochem. 80, 1223–1232 10. Takasaki, S. and Kobata, A. (1978) Microdetermination of sugar composition by radioisotope labeling. Methods Enzymol. 50, 50–54. 11. Kondo, A., Suzuki, J., Kuraya, N., Hase, S., Kato, I., and Ikenaka, T. (1990) Improved method for fluorescence labeling of sugar chains with sialic acid residues. Agric. Biol. Chem, 54, 2169–2170. 12. Tai, T., Yamashita, K., Ogata, M. A., Koide, N., Muramatsu, T., Iwashita, S., Inoue, Y., and Kobata, A. (1975) Structural studies of two ovalbumin glycopeptides in relation to the endo-β-N-acetylglucosaminidase specificity. J. Biol Chem. 250, 8569–8575. 13. Rupley, J. A. (1964) The hydrolysis of chitin by concentrated hydrochloric acid, and the preparation of low-molecular-weight substrates for lysozyme. Biochem. Biophys. Acta. 83, 245–255. 14. Krusius, T., Finne, J., and Rauvala, H. (1978) The poly(glycosyl) chains of glycoproteins. Characterization of a novel type of glycoprotein saccharides from human erythrocyte membrane. Eur. J. Biochem. 92, 289–300. 15. Yamamoto, K., Tsuji, T., Tarutani, O. and Osawa, T. (1984) Structural changes of carbohydrate chains of human thyroglobulin accompanying malignant transformations of thyroid grands. Eur. J. Biochem. 143, 133–144 16. Tsuji, T., Irimura, T. and Osawa, T. (1980) The carbohydrate moiety of Band-3 glycoprotein of human erythrocyte membranes. Biochem. J. 187, 677–686. 17. Baenziger, J. U. and Fiete, D. (1979) Structural determinants of Ricinus communis agglutinin and toxin specificity for oligosaccharides. J. Biol. Chem. 254, 9795–9799. 18. Irimura, T., Tsuji, T., Yamamoto, K., Tagami, S. and Osawa, T. (1981) Structure of a complex-type sugar chain of human glycophorin A. Biochemistry 20, 560–566 19. Shibuya, N., Goldstein, I. J., Van Damme, E. J. M. and Peumans, W. J. (1988) Binding properties of a mannose-specific lectin from the snowdrop (Galanthus nivalis) bulb. J. Biol. Chem. 263, 728–734 20. Yamamoto, K., Tsuji, T., Matsumoto, I. and Osawa, T. (1981) Structural requirements for the binding of oligosaccharides and glycopeptides to immobilized wheat germ agglutinin Biochemistry 20, 5894–5899 21. Ogata, S., Muramatsu, T. and Kobata, A. (1975) Fractionation of glycopeptides by affinity column chromatography on concanavalin A-Sepharose. J. Biochem. 78, 687–696. 22. Krusius, T., Finne, J. and Rauvala, H. (1976) The structural basis of the different affinities of two types of acidic N-glycosidic glycopeptides from concanavalin A-Sepharose. FEBS Lett. 71, 117–120. 23. Kornfeld, K., Reitman, M. L. and Kornfeld, R. (1981) The carbohydrate-binding specificity of pea and lentil lectins. J. Biol. Chem. 256, 6633–6640 24. Katagiri, Y., Yamamoto, K., Tsuji, T. and Osawa, T. (1984) Structural requirements for the binding of glycopeptides to immobilized vicia faba (fava) lectin. Carbohydr, Res. 129, 257–265 25. Yamamoto, K., Tsuji, T. and Osawa, T. (1982) Requirement of the core structure of a complex-type glycopeptide for the binding to immobilized lentil- and pea-lectins. Carbohydr. Res. 110, 283–289

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26. Yamashita, K., Hitoi, A. and Kobata, A. (1983) Structural determinants of Phaseolus vulgaris erythroagglutinating lectin for oligosaccharides. J. Biol. Chem. 258, 14753–14755. 27. Cummings, R. D. and Kornfeld, S. (1982) Characterization of the structural determinants required for the high affinity interaction of asparagine-linked oligosaccharides with immobilized Phaseolus vulgaris leukoagglutinating and erythroagglutinating lectins. J. Biol. Chem. 257, 11230–11234 28. Cummings, R. D. and Kornfeld, S. (1984) The distribution of repeating [Galβ1,4GlcNAcβ1,3] sequences in asparagine-linked oligosaccharides of the mouse lymphoma cell line BW5147 and PHAR2.1. J. Biol. Chem. 259, 6253–6260. 29. Yamashita, K., Totani, K. T., Ohkura, Takasaki, S., Goldstein, I. J. and Kobata, A. (1987) Carbohydrate binding properties of complex-type oligosaccharides on immobilized Datura stramonium lectin. J. Biol. Chem. 262, 1602–1607 30. Irimura, T. and Nicolson, G. L. (1983) Interaction of pokeweed mitogen with poly(Nacetyllactosamine)-type carbohydrate chains. Carbohydr. Res. 120, 187–195 31. Kawashima, H., Sueyoshi, S., Li, H., Yamamoto, K. and Osawa, T. (1990) Carbohydrate binding specificities of several poly-N-acetyllactosamine-binding lectins. Glycoconjugate J. 7, 323–334. 32. Wang, W. -C. and Cummings, R. D. (1988) The immobilized leukoagglutinin from the seeds of Maackia amurensis binds with high affinity to complex-type Asnlinked oligosaccharides containing terminal sialic acid-linked α-2,3 to penultimate galactose residues. J. Biol. Chem. 263, 4576–4585. 33. Kawaguchi, T., Matsumoto, I. and Osawa, T. (1974) Studies on hemagglutinins from Maackia amurensis seeds. J. Biol. Chem. 249, 2786–2792. 34. Sueyoshi, S., Yamamoto, K. and Osawa, T. (1988) Carbohydrate binding specificity of a beetle (Allomyrina dichotoma) lectin. J. Biochem. 103, 894–899. 35. Yamashita, K., Umetsu, K., Suzuki, T., Iwaki, Y., Endo, T. and Kobata, A. (1988) Carbohydrate binding specificity of immobilized Allomyrina dichotoma lectin II. J. BIol. Chem. 263, 17482–17489. 36. Shibuya, N., Goldstein, I. J., Broekaert, W. F., Lubaki, M. N., Peeters, B. and Peumans. W. J. (1987) Fractionation of sialylated oligosaccharides, glycopeptides, and glycoproteins on immobilized elderberry (Sambucus nigra L.) bark lectin. Arch. Biochem. Biophys. 254, 1–8. 37. Shibuya, N., Goldstein, I. J., Broekaert, W. F., Lubaki, M. N., Peeters, B. and Peumans, W. J. (1987) The elderberry (Sambucus nigra L.) bark lectin recognizes the Neu5Ac(α2–6)Gal/GalNAc sequence. J. Biol. Chem. 262, 1596–1601. 38. Konami, Y., Yamamoto, K., Osawa, T. and Irimura, T. (1994) Strong affinity of Maackia amurensis hemagglutinin (MAH) for sialic acid-containing Ser/Thrlinked carbohydrate chains of N-terminal octapeptides from human glycophorin A. FEBS Lett. 342, 334–338. 39. Osawa, T. and Tsuji, T. (1987) Fractionation and structural assessment of oligosaccharides and glycopeptides by use of immobilized lectins. Ann. Rev. Biochem. 56, 21–42. 40. Osawa, T. (1989) Recent progress in the application of plant lectins to glycoprotein chemistry. Pure & Appl. Chem. 61, 1283–1292.

148 In-Gel Enzymatic Release of N-Glycans David J. Harvey 1. Introduction Analysis of N-linked glycans, those attached to Asn in an Asn-Xxx-Ser(Thr) motif, where Xxx is any amino acid except proline, requires determination of which consensus sequence is occupied, the extent of occupancy and the structures of the glycans that are attached to each site. Structural analysis of the glycans is most commonly performed following their removal, either chemically (1,2) or enzymatically, and this chapter describes an enzymatic method for glycan removal using protein N-glycosidase F (PNGase F) (3) from within the SDS-PAGE gels commonly used to separate and purify the glycoproteins. The method is a modification of that described by Küster et al. (4) in 1997. It can be used for mammalian glycans but not for those that are substituted at the 3-position of the reducing terminal GlcNAc residue, commonly found in plants and insects. These glycans are resistant to PNGase F but can be released with PNGase A after trypsinolysis of the protein. Released glycans can then be examined directly by mass spectrometry, usually with MALDI or electrospray ionization, or they can be labeled at their reducing terminus for analysis by chromatographic or electrophoretic techniques. 2. Materials 2.1. Apparatus 2.1.1. Electrophoresis and Glycan Release 1. 2. 3. 4. 5. 6.

BioRad Mini-Protean II apparatus with 80 × 80 × 0.75 mm plates, or similar. 500 µL Eppendorf tubes. Temperature-controlled heating block. BioRad Spin column or equivalent. Plastic hypodermic-type syringe (1 mL). Millipore 0.5 µm FH-type filters. From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2.1.2. Optional Glycan Purification 1. GelLoader® pipette tips. 2. Supply of nitrogen.

2.2. Chemicals 2.2.1. Electrophoresis and Glycan Release 1. 2. 3. 4.

Protein molecular weight markers. Reference glycoprotein (e.g., ovalbumin. ribonuclease B) Aluminium foil. Protein N-glycosidase F e.g. 1000 U/mL recombinant glycerol-free lyophilisate (Roche Life Sciences) (see Note 1). 5. Dowex AG50Wx12 Resin (H+ form; 100 to 200 mesh; Bio-Rad). 6. pH Test paper (to check for neutrality).

2.2.2. Optional Glycan Purification 1. 2. 3. 4. 5.

Dowex AG3x4 resin C-18 resin (see Note 2). Dimethylsulfoxide (DMSO). Methyl iodide. Sodium hydroxide.

2.2.3. Solutions 1. 17.5% Resolving gel for proteins in the range 10–70 kDa (see Note 3): 87.5 mL of 30% acrylamide, 5.5 mL of 2% bis-acrylamide 50.5 mL of 1 M Tris buffer (pH 8.8, acetic acid), 0.75 mL of 20% SDS and add water to give a total volume of 150 mL. 2. 12.5% Resolving gel for proteins in the range 70–200 kDa: 62.5 mL of 30% acrylamide, 15.5 mL of 2% bis-acrylamide 56.0 mL of 1 M Tris buffer (pH 8.8, acetic acid), 0.75 mL of 20% SDS and add water to give a total volume of 150 mL. 3. Resolving gel polymerization mixture: 18 µL TEMED and 33 µL of 10% APS. 4. Stacking gel: 20.0 mL of 30% acrylamide, 7.8 mL of 2% bis-acrylamide 15.0 mL of 1 M Tris buffer (pH 8.8, acetic acid), 0.6 mL of 20% SDS and add water to give a total volume of 120 mL. 5. Stacking gel polymerization mixture: 12 µL TEMED and 25 µL of 10% APS. 6. Loading buffer: 5 mL 2 M ammonium formate, pH 4.4; 50 mL Milli-Q-purified water, Adjust pH to 8.6, Add 0.4 g SDS and water to a final volume of 100 mL. 7. Electrophoresis running buffer: 25 mM Tris, 190 mM glycine, 0.5% SDS. 8. Destain solution: 5% methanol/7% acetic acid. 9. Incubation buffer: 20 mM NaHCO3, pH 7.0; 336 mg NaHCO3 in 200 mL Milli-Q water, adjust to pH 7.0 with acetic acid. 10. 45 mM DTT solution: 6.9 mg of DDT in 1mL of NaHCO3 buffer. 11. Iodoacetamide solution: 18.5 mg of iodoacetamide in 1 mL of NaHCO3 buffer (see Note 4).

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12. PNGase-F solution: 50 Units of PNGase F powder in 0.25 mL Milli-Q water (200 Units/mL and dialyse into 20 mM NaHCO3 buffer (see Note 5). Add 50 µL distilled water to give a final concentration of 100 Units/mL.

3. Methods 3.1. Gel Electrophoresis 3.1.1. Preparation of Gel 1. Mix 5 mL of the resolving gel mixture with the polymerization mixture and pour to 1 cm below the top of the electrophoresis cell. 2. Overlay with about 2 mm of iso-butanol/water and leave the gel to set for about 30 min. 3. Wash off iso-butanol/water and remove excess water. 4. Mix 5 mL of the stacking gel mixture with the polymerization mixture and pour onto the resolving gel to fill the cell. 5. Insert the comb into the stacking gel, avoiding air bubbles, and allow the mixture to set for 30 mins.

3.1.2. Preparation of Sample 1. Mix the sample, which should contain about 50–100 pmol of glycoprotein with twice its volume of loading buffer and bromophenol blue. 2. Boil for 5 mins in a hot block (in an uncapped tube), cool and spin in a microcentrifuge.

3.1.3. Running of Gel 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Clip the cell containing the gel into its frame and place into the electrophoresis tank. Add running buffer to the tank and remove any air bubbles. Remove the comb from the stacking gel and wash the wells with Milli-Q water. Fill the spaces with running buffer. Add bromophenol blue dye to the outer lanes. Add 3 µL of molecular weight markers to the next lanes. Add a reference glycoprotein (e.g. ovalbumin 10 µL, 100 pmol, 4.5 mg) + dye to one lane. Load the samples (10 µL) into the remaining lanes (see Note 6). Fill empty lanes with running buffer. Connect to the power supply (200 V). Switch on and set the current to about 50 mA. Allow to run until the dye runs out at the bottom of the gel (see Note 7).

3.1.4. Gel Staining 1. 2. 3. 4.

Empty the tank, remove the gel in its cell and wash with tap water. Remove the spacers, lever apart the glass plates and remove the stacking gel. Mark the top of the gel by cutting off a corner. Wash the gel from the remaining glass plate with a stream of water into a plastic box.

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5. Cover the gel with Coomassie blue solution, cover and shake for 10–15 mins 6. Pour away the dye solution.

3.1.5. Gel Destaining 1. 2. 3. 4. 5.

6. 7. 8. 9. 10.

Wash the gel with tap water. Cover the gel with the destain solution (see Note 8) and shake for 10 min Change the destain solution and shake for a further 10 mins. Repeat step 3. Cut out the bands from the gel that contains the target glycoproteins. (see Note 9) and place into the Eppendorf tube with a small amount of destain solution. Place the tube in a shaker overnight at room temperature. Wash gel pieces with 300 µL of incubation buffer, vortex, centrifuge briefly, and soak for 30 min at room temperature. Discard the wash. Repeat step 7. Wash the gel pieces for 60 min in 300 µL of 1:1 acetonitrile: 20 mM NaHCO3 to remove residual SDS. Dry gel pieces in a vacuum centrifuge.

3.2. Glycan Removal 3.2.1. Reduction of Glycoprotein 1. Add 300 µL of incubation buffer (to cover gel bits). 2. Add 20 µL of the DTT solution. 3. Cap, spin and heat in a shaker at 60°C for 30 mins.

3.2.2. Alkylation of Glycoprotein Add 20 µL of the iodoacetamide solution to the gel-containing tube. Wrap the tube in aluminium foil and shake for 30 min at room temperature. Remove the buffer and iodoacetamide. Add about 0.5 mL of NaHCO3 buffer and 0.5 mL of acetonitrile and shake at room temperature for 1 hour. 5. Remove the buffer and acetonitrile. 6. For each sample, cut the gel piece into small bits and place the gel bits into a 500 µL Eppendorf tube that has holes pierced in the lid. 7. Place the tubes in a SpeedVac (continuous rotor) until they are dry (crisp). 1. 2. 3. 4.

3.2.3. Removal of Sugars with PNGase-F 1. Add 30 µL of the PNGase solution to each sample (the solution should just cover the gel bits). 2. Leave at room temperature until all liquid has been taken up by the gel (20–30 mins.) 3. Add 150 µL of NaHCO3 buffer. 4. Incubate at 37°C overnight.

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Extraction of N-Glycans from the Gel

1. Prepare AG50 (30 µL/sample) by adding the AG50 resin (see Note 10) to a BioRad BioSpin column, running approx. 10 col. volumes of HCl (1:1, v:v dilution of concentrated HCl) through the AG50 and then washing with Milli-Q water until the washings are neutral (about pH 5 as tested by pH paper). 2. Add 100 µL of distilled water to each sample and sonicate for 30 mins. 3. Remove the solution and add to a small screw-cap tube containing the AG50 resin. 4. Repeat steps 2 and 3 twice, adding the solution to the tube containing the resin. 5. Add 200–300 µL of acetonitrile to each sample and sonicate for 30 mins. (see Note 11). 6. Remove the acetonitrile and add to the tube containing the resin. 7. Shake the resin-containing tube and spin in a microcentrifuge. 8. Filter through a 0.5 µm FH-type Millipore filter attached to a 1 mL plastic syringe. 9. Wash the resin, filter the washings and add to the first filtrate 10. Dry the sample in a SpeedVac. 11. Add 20 mL 1% formic acid and allow to stand at room temperature for 40 mins (see Note 12). 12. Dry in a SpeedVac.

3.2.5. Removal of Contaminants from the Glycans The samples will still be contaminated with, possibly, peptides, salts, SDS etc. that must be removed before analysis by mass spectrometry or HPLC. Several methods are available. We prefer the Nafion membrane procedure (5) that is described in the chapter on the analysis of N-glycans by mass spectrometry. However, a second stage of preliminary cleaning may be required. For neutral glycans (see Note 13), this can be achieved as follows: 1. Fill a GelLoader tip with ethanol. 2. Add a few µL of G3 × 4 resin, followed by washed AG50 resin and C-18 resin to the GelLoader tip to form three layers. 3. With nitrogen from a cylinder, force ethanol through column until just to the top of the resin (see Note 14). 4. Add 100 µL of Milli-Q water to the top of the resin “sandwich” and force the water through the resins with the nitrogen. 5. Repeat the washing procedure twice. 6. Add the aqueous sample solution to the resin column. 7. Force the solution through the resins with nitrogen and collect into a small Eppendorf (PCR) tube. 8. Add 50 µL Milli-Q water to the resins, blow through and add to the glycan solution. 9. Dry the glycans in a Speedvac.

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3.2.6. Sialylated Glycans Sialylated glycans will be removed by the AG3 resin in the above procedure. However they can be neutralized (and stabilized for analysis by MALDI mass spectrometry) by formation of methyl esters prior to clean-up by the three-bed resin procedure. 1. Place about 100 µL AG5OW × 12 resin in a short column. 2. Wash with 1 mL of 1 M sodium hydroxide solution followed by water until the washings are approximately neutral. 3. Dissolve the glycan sample (100–500 pmole) in about 5–20 µL water and apply to the column. 4. Wash through with 5–100 µL of water and collect the washings in a small Eppendorf tube. 5. Dry in a rotary evaporator. 6. Dissolved the glycans in about 1 µL of dry DMSO and add 1 µL of methyl iodide (see Note 15). 7. Mix well and allow to stand for 2 h at room temperature. 8. Add about 10 mL of DMSO and evaporate the methyl iodide with a stream of nitrogen. 9. Evaporate to dryness (see Note 16).

4. Notes 1. Preparations containing glycerol should be avoided. 2. C18 resin can be obtained from a C18 SepPak cartridge 3. Higher bis-acrylamide concentrations lead to increased cross-linking and reduced glycan yield. 4. Iodoacetamide is light sensitive and solutions should be kept in the dark. 5. It is best not to use nonammonium-containing buffers at this stage. PNGase F releases the glycans as the glycosylamine that need to be converted into the free sugars before further analysis. Ammonium-containing buffers promote glycosylamine retention (6). 6. It is a good idea to load the samples asymmetrically so that it is possible to determine the left and right sides of gel. 7. The current normally drops to about 10 mA. 8. A low concentration of acetic acid is used to minimizes loss of sialic acid from the glycans. 9. The gel pieces should be small but wide enough to retain minor glycoforms. 10. The resin, which should be stored in water, can be handled with a Gilson pipette fitted with a blue or yellow tip that has been cut to widen the end. 11. The acetonitrile causes the gel to shrink, expelling the remaining water. 12. Because the glycans are released as glycosylamines, a considerable amount of the sample may be in this form at this stage. The glycosylamines can

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14.

15. 16.

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be detected by mass spectrometry by their mass being one unit less than that of the native glycan. If labelling of the reducing terminus is attempted at this stage, the glycosylamines will not react, resulting in a poor yield of labeled material. Furthermore, the rate and extent of glycosylamine hydrolysis to the glycan depends on whether or not there is a fucose residue at the reducing terminus. Consequently, samples must be fully converted into their native glycan (OH) form before analysis is attempted. Conversion can be achieved by leaving the glycans to stand in water or, better, in dilute formic acid. However, care must be taken to avoid hydrolysis of sensitive bonds such as those linking the sialic acids. The AG3 resin is included in this clean-up procedure mainly to remove any residual SDS. However, it will also remove acidic (sialylated, sulfated glycans). To prevent sialic acid removal, the sialic acids can be converted into methyl esters as described by Powell and Harvey (7) and in Subheading 3.2.6 before the 3-resin method is applied. However, sulfates cannot be methylated by this method. In this and subsequent steps, release the nitrogen pressure when the top surface of the liquid reaches the top of the resins in order to avoid introducing air bubbles to the resin. Methyl iodide is toxic; consequently, this procedure should be performed in a fume hood. DMSO is not very volatile, consequently, evaporation is slow.

Acknowledgments The author would like to thank Professor Bernhard Küster who originally developed the in-gel method and the Oxford Glycobiology endowment fund for financial support.

References 1. Patel, T., Bruce, J., Merry, A., Bigge, C., Wormald, M., Jaques, A. and Parekh, R. (1993) Use of hydrazine to release in intact and unreduced form both N- and O-linked oligosaccharides from glycoproteins. Biochemistry 32, 679–693. 2. Wing, D. R., Field, M. C., Schmitz, B., Thor, G., Dwek, R. A., Schachner, M. S. and Rademacher, T. W. (1992) The use of large-scale hydrazinolysis in the preparation of N-linked oligosaccharide libraries: application to brain tissue. Glycoconj. J. 9, 293–301. 3. Tarentino, A. L., Gómez, C. M. and Plummer, T. H., Jr. (1985) Deglycosylation of asparagine-linked glycans by peptide:N-glycosidase F. Biochemistry 24, 4665–5671. 4. Küster, B., Wheeler, S. F., Hunter, A. P., Dwek, R. A. and Harvey, D. J. (1997) Sequencing of N-linked oligosaccharides directly from protein gels: In-gel deglycosylation followed by matrix-assisted laser desorption/ionization mass spectrometry

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and normal-phase high performance liquid chromatography. Anal. Biochem. 250, 82–101. 5. Börnsen, K. O., Mohr, M. D. and Widmer, H. M. (1995) Ion exchange and purification of carbohydrates on a Nafion(R) membrane as a new sample pretreatment for matrix-assisted laser desorption-ionization mass spectrometry. Rapid Commun. Mass Spectrom. 9, 1031–1034. 6. Küster, B. and Harvey, D. J. (1997) Ammonium-containing buffers should be avoided during enzymatic release of glycans from glycoproteins when followed by reducing terminal derivatization. Glycobiology 7, vii–ix. 7. Powell, A. K. and Harvey, D. J. (1996) Stabilisation of sialic acids in N-linked oligosaccharides and gangliosides for analysis by positive ion matrix-assisted laser desorption-ionization mass spectrometry. Rapid Commun. Mass Spectrom. 10, 1027–1032.

149 Analysis of N-Linked Glycans by Mass Spectrometry David J. Harvey

1. Introduction Analysis of N-linked glycans, those attached to Asn in an Asn-Xxx-Ser(Thr) motif, where Xxx is any amino acid except proline, involves determination of which consensus sequence is in fact occupied by glycans, the extent of occupancy and the structures of the glycans that are attached. This chapter deals only with a small set of these procedures, namely the structural analysis of the carbohydrates using mass spectrometry as the main analytical technique. It is assumed that the glycans have been released from the glycoprotein or derived glycopeptide by techniques such as hydrazinolysis (1,2) or enzymolysis with an endoglycosidase such as protein-N-glycosidase F (PNGase F) (3,4) and that the released glycans are available in aqueous solution. Traditionally, such mixtures of glycans would be labeled with a chromophore or fluorophore such as 2-aminobenzamide (5) and profiled by gel-filtration or high-performance liquid chromatography (HPLC) with structures being deduced by subsequent profiling of the products of sequential exoglycosidase digestion with a library of exoglycosidases. Knowledge of the enzyme specificity and the number of constituent monosaccharide residues removed at each stage of the procedure allows the structure of the glycan(s) to be deduced. The method, although valuable to the mass spectroscopist for providing information on the nature of the constituent monosaccharides, is limited by the range, availability and purity of the available exoglycosidases which does not include enzymes appropriate for all structural features. Mass spectrometry, although capable of rapidly supplying much of the information that is difficult to obtain by the exoglycosidase technique, still relies on

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it for confirmation of the identity of the constituent monosaccharides. However, because of the very conserved nature of the N-glycan biosynthetic pathway and because much work on the analysis of N-glycans is performed with species with well characterized enzymology, mass measurements are often sufficient to provide the structural information that is required. This information relates to questions such as what type of glycan (high-mannose, hybrid, complex) is present and the branching pattern and composition of the antennae. However, for rigorous work, it must be remembered that a mass measurement will only classify monosaccharide constituents into groups such as hexose, HexNAc, and so on. The ideal mass spectrometric approach is one in which the initial spectrum produces the glycan profile, preferably quantitatively, with no fragmentation and subsequent spectra give extensive fragmentation to provide the detailed structural information required on each component. Glycan profiles are conveniently obtained by matrix-assisted laser desorption/ionization (MALDI) mass

Fig. 1. (a) Positive ion MALDI-TOF mass spectrum of N-glycans released from a mixture of glycoproteins (mainly ovalbumin) from chicken egg white. (b) Positive ion reflectron MALDI-TOF spectrum of a disialylated biantennary glycan. Peaks annotated with a black spot are metastable (PSD) ions. Key to symbols used for the constituent monosaccharides in this scheme and subsequent Figs. à = galactose,  = GlcNAc,  = mannose,  = fucose,  = N-acetylneuraminic (sialic) acid. Solid lines connecting the symbols are β-bonds and broken lines are α-bonds.

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spectrometry (6–9), a technique that gives a quantitative response and negligible fragmentation for neutral N-glycans of varying structure (10,11) (Fig. 1a). Sialylated glycans, however, tend to fragment by loss of sialic acid (Fig. 1b) but can be neutralized and stabilized by methyl ester formation (12). The somewhat milder technique of electrospray ionization (ESI) is being increasingly used but tends to produce ions in different charge states that limit its usefulness as a profiling technique. It is, however, an ideal method for introducing glycans into a mass spectrometer for fragmentation analysis. Older, less sensitive techniques such as fast-atom bombardment (FAB) mass spectrometry have also been widely used for glycan analysis (13,14) but produce extensive fragmentation, require derivatization of the glycans and are now mainly of historical interest. The mass of the glycan provides a direct indication of the constituent monosaccharide composition in terms of sugar type (hexose, deoxy-hexose, and so on) because most N-glycans consist of only a limited number of mass-different monosaccharides. Table 1 lists the residue masses for these monosaccharides. Suitable software is available on the internet at https://tmat.proteomesystems. Table 1 Residue Masses of Common Monosaccharides Found in N-Glycans Monosaccharide

Residue formula

Residue massa

Pentose

C5H8O4

Deoxy-hexose

C6H10O4

Hexose

C6H10O5

Hexosamine

C6H11NO4

HexNAc

C8H13NO5

Hexuronic-Acid

C6H8O6

N-Acetyl-neuraminic acid

C11H17NO8

N-glycoyl-neuraminic acid

C11H17NO9

132.042 132.116 146.078 146.143 162.053 162.142 161.069 161.157 203.079 203.179 176.032 176.126 291.095 291.258 307.090 307.257

a) Top Fig. = monoisotopic mass (based on C = 12.000000, H = 1.007825, N = 14.003074, O = 15.994915), Lower Fig. = average mass (based on C = 12.011, H = 1.00794, N = 14.0067, O = 15.9994. The masses of the intact glycans can be obtained by addition of the residue masses given above, plus the mass of the terminal group (H2O for an unmodified glycan) 18.011 (monoisotopic) and 18.152 (average) and the mass of any reducing-terminal or other derivative.

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com/glyco/glycosuite//glycodb (requires password) for deriving compositions from experimental masses. Once the monosaccharide composition is established, structural features can be assigned by fragmentation. Fragmentation from MALDI-generated ions produced in time-of-flight (TOF) instruments can be obtained by analysis of spontaneously produced post-source decay (PSD) ions (15) but resolution is generally poor and many commercial instruments require the spectra to be recorded in segments with consequent reduction in quantitative information. Greatly improved performance can be obtained by generating the fragments by collision in a tandem instrument such as a quadrupole/time-of flight (Q-Tof) mass spectrometer (16–20). Multiple successive fragmentations can be produced with a storage instrument such as an ion trap (21–23) and can be used to extract detailed structural information from small fragments of the glycan. The information that can be obtained from the spectrum depends to a large extent on the type of ion that is initially formed. MALDI Produces mainly [M+Na]+ ions that fragment predominantly by cleavage between the sugar rings (glycosidic cleavage) to provide sequence information but little linkage data. Linkage information is provided by cross-ring cleavage fragments that are usually present in only low relative abundance in low-energy spectra of the type produced by trap and Q-Tof-type instruments. Cross-ring fragments of higher abundance are found in the spectra recorded with instruments providing higher-energy collisions (24–26). Unfortunately, fragmentation spectra of [M+Na]+ ions contain ambiguous information because fragment ions of the same mass and composition often arise from different regions of the molecule. The situation is even more difficult with the [M+H]+ ions frequently encountered in ESI spectra (Fig. 2a); fragmentation of these ions rarely yield cross-ring cleavages and are prone to internal rearrangements (27–30) that again yield ambiguous information. Negative ion spectra, on the other hand, are much more informative because cross-ring cleavage reactions are frequently dominant (31–37) (Fig. 2) and the

Fig. 2. Nomenclature for describing fragment ions from carbohydrates according to Domon and Costello (39).

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redundancy seen in the positive ion spectra is largely absent. Spectra of isomeric compounds are usually different in that they produce ions of different masses unlike positive ion spectra where the presence of isomers is often only reflected in differences in ion abundance. Consequently, the spectra contain ions that are diagnostic of specific structural features that cannot be obtained easily from the positive ion spectra or from the more classical analytical methods and form the basis of this chapter. The protocol described below is one that is used in our laboratory to provide most of the structural information necessary to analyse complex mixtures of N-glycans that have been released with PNGase from glycoproteins separated in SDS-PAGE gels (4). 2. Materials 2.1. Purification of Glycans 1. Nafion 117 membrane (Aldrich Chemical Co. Ltd., Poole, UK). 2. 35% Nitric acid. 3. Milli-Q water.

2.2. MALDI Mass Spectrometry 1. Matrix solution (saturated solution of dihydroxybenzoic acid (DHB) in acetonitrile (see Note 1). 2. Ethanol.

2.3. Electrospray Mass Spectrometry (Negative Ion) 1. 2. 3. 4.

Ammonium phosphate. Nanospray capillaries (Proxeon). Methanol:Milli-Q water (1:1, v:v). Gel-Loader pipette tips (Eppenforf 20 µL).

3. Methods 3.1. Purification of Glycans Most glycan solutions contain residual salt, peptides etc. whose presence will degrade the quality of the mass spectrum or completely suppress the signals from the glycans. Normally neutral glycans give much weaker signals than peptides on account of their different ionization properties (formation of [M+Na]+ rather than [M+nH]n+(−) ions. If such contamination is expected, proceed with a clean-upstage as follows. If not, this stage can be omitted. The method is essentially as described by Börnsen et al. (38). 1. Condition small squares (about 3 cm square) of the Nafion 117 membrane by heating in 35% nitric acid for 4 hours at 80°C. 2. Wash each square thoroughly with Milli-Q water (see Note 2).

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3. Dry a square of membrane with filter paper and float it on the surface of Milli-Q water in a small shallow container. 4. Deposit 1 µL (or similar) drops of the aqueous glycan solutions (see Note 3) onto the membrane surface and allow it to sit for about 20 mins. (see Note 4). 5. Remove the glycan solution with a pipette and either deposit directly onto the MALDI target or dilute for ESI analysis as described below.

3.2. MALDI Mass Spectrometry 1. Deposit 0.5–1 µL glycan solution onto the MALDI target. 2. Using a separate pipette tip, add 0.3 µL matrix solution and allow the mixture to dry at room temperature. Inspect the dried sample deposit. (see Notes 5 and 6). 3. If good crystals are present, redissolve the spot in the minimum amount of ethanol and allow it to recrystalize (see Note 7). 4. Introduce the target into the mass spectrometer and acquire both positive and negative ion spectra in reflectron mode (see Note 8). Keep the laser power as low as practical in order to avoid peak broadening and consequent mass errors (see Note 9) and frequently move the laser spot to different areas of the target (see Note 10) until a satisfactory signal:noise ratio is obtained (see Note 11). 5. If sialic acids are detected, they can be stabilized by forming methyl esters using the protocol described in the chapter on in-gel release of glycans and reference (12) (see Note 12). 6. If desired, recover the remaining sample from the MALDI target after MS analysis by dissolving the spot in several repetitive additions of 2 µL of 1:1 (v:v) methanol:water and removing with a pipette. The matrix can then removed from the resultant solution by drop dialysis or use of a C18 ZipTip.

3.3. Electrospray Mass Spectrometry (Negative Ion) 1. Dilute the cleaned glycan solution with about 5 µL of 1:1 (v:v) methanol:water containing a trace of ammonium phosphate (see Note 13). 2. Introduce the glycan solution into the nanospray capillary with a 20 µL pipette fitted with a GelLoader pipette tip (see Note 14). 3. Introduce the capillary into the mass spectrometer with the voltage set to zero, introduce a small pressure on the capillary and start the flow by touching the tip of the capillary on the cone. If the spray does not start immediately, scrape the tip on the cone to try and break off the end. 4. Once the flow has started, turn up the capillary voltage to about 1.1 kV (WatersMicromass Q-Tof Ultima Global instrument. Other instruments may vary). 5. Record the MS spectrum and then the MS/MS spectra of each relevant ion (see Notes 15 and 16).

3.4. Spectral Interpretation 3.4.1. Neutral Glycans The N-glycans fragment by glycosidic (between the monosaccharide rings) and cross-ring cleavages. All glycosidic cleavages are accompanied by hydrogen

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migration to give the product even-electron ion and a neutral molecule. The nomenclature used to describe the fragmentation was proposed by Domon and Costello in 1988 (39) and has been universally accepted. It is outlined in Fig. 2. Ions retaining the charge on the reducing terminus are designated X, Y, and Z where X is a cross ring cleavage and Y and Z are glycosidic cleavages depending on which side of the linking oxygen the bond is broken. Subscript numbers designate the position along the chain where cleavage occurs. Unfortunately, for series of glycans of different chain length, this number can change for the same fragmentation such as loss of the terminal residue from the reducing terminus. In order to overcome this difficulty we have introduced a modification whereby the subscript R is used for this cleavage and R-1 etc. is proposed for cleavages further into the glycan chain. For branched structures, chains are further designated by Greek letters with the largest chain being alpha. Ions retaining the charge at the nonreducing terminus are named similarly using A, B and C in place of X, Y and Z. Cross ring cleavages are further defined by superscript numbers preceding the letter and consisting of the lowest number of the two carbon atoms of the bond cleaved or “O” if the O-1 bond is cleaved. Major ions in positive ion spectra tend to be formed by B and Y glycosidic cleavages whereas ions in the negative ion spectra are mainly products of cross-ring or C-type glycosidic cleavages. The high information content of the negative ion fragmentation spectra arises from abstraction of protons from individual and specific hydroxyl groups which catalyses specific electron movements leading to the diagnostic fragments as illustrated in Fig. 3 for formation of the 2,4AR ions representing loss from the reducing terminal GlcNAc residue. Unfortunately, acidic glycans, such as those containing sialic acid, loose the acidic protons in preference to the hydroxylic hydrogens, thus suppressing formation of the diagnostic ions. Much specificity can be restored by blocking the acidic group by formation of methyl esters but the resulting fragmentation spectra are complicated by many additional ions formed by losses of methanol.

Fig. 3. Proposed mechanism for the formation of the 2,4AR ion from N-linked glycans.

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Fig. 4. Negative ion CID fragmentation spectra of (a) the high-mannose glycan Man 5GlcNAc2 (b) the high-mannose glycan Man9GlcNAc2 (c) a fucosylated, bisected complex glycan (d) a 3-branched triantennary glycan (e) a 6-branched triantennary glycan.

A suggested spectral interpretation protocol could be as follows: typical spectra are shown in Fig. 4. 1. Determine the constituent monosaccharide composition of the glycans producing each peak using the information in Table 1. The mass of the glycans recorded by nano-electrospray as phosphate adducts will be 98 mass units higher than this calculated mass (mass of glycan + H2PO4−) and those in the MALDI-TOF spectra will be 23 mass units higher ([M+Na]+ ion). 2. From the negative ion MS/MS spectrum, verify that the adduct is phosphate. This can be done by locating the 2,4AR, BR and 2,4AR-1 triplet at 259, 319 and 462 mass units below the mass of the molecular ion (Fig. 4b) or at 405, 465 and 608 mass units below if the glycan contains a fucose residue attached to the reducing terminal GlcNAc residue (Fig. 4c). There is a possibility of other modifications and this must also be taken into account but most common N-linked glycans will fit this pattern. The most likely alternative adduct is chlorine which will add 35 and 37 mass units to the mass of the glycan and whose ions can often be identified by the characteristic chlorine isotope pattern. 3. Use the information in Table 2 and the four publications (40–43) to deduce the structure of the glycan. Some of the more important diagnostic ions are outlined below: 4. The masses of the 2,4AR, BR and 2,4AR-1 ions can be used to determine the presence of fucose at the 6-position of the reducing terminal GlcNAc residue; fucose at the 3-position will result in the absence of the 2,4AR ion because its formation involves abstraction of the hydrogen from this position in the initial loss of the adduct.

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5. The D-type ions in Table 2 (m/z 647 in Fig. 4a, 971 in Fig. 4b, 688 in Fig. 4d, and 1053 in Fig. 4e) consist of the β-linked core mannose residue together with the mannose attached to the 6-position together with its attached residues. Thus, they can be used to determine the composition of the 6-antenna and classify the glycan into complex or high-mannose/hybrid. The ion is accompanied by a second ion 18 mass units lower of comparable abundance. If a bisecting GlcNAc is present (Fig. 4c), the D ion is absent because the bisecting GlcNAc residue is readily expelled (221 mass units) to give a very abundant ion at the same mass as the D-18 ion from the un-bisected glycans. 6. Antenna composition is revealed by a 1,3A cleavage of the mannose residue in that antenna to produce what is normally a very abundant ion termed an F-type ion (e.g., antennae containing Gal-GlcNAc residues produce this ion at m/z 424 (Gal-GlcNAc + 59 mass units, m/z 424 in Figs. 4c, 4d, and 4e). If fucose is present on this antenna, the ion shifts to m/z 570. Shifts produced by other substituents are listed in Table 2. 7. An O,4A-cleavage of the antennae mannose residues from an unbranched antenna of composition Gal-GlcNAc gives an ion at m/z 466, an ion termed E. If the 3-antenna is branched (Fig. 4d), an abundant E-type ion is present at m/z 831 (m/z 466 + the mass of Gal-GlcNAc). Branching of the 6-antenna (Fig. 4e) does not produce this shift because the 6-position, the substitution position of the second branch in the 6-antenna is not present in the E-ion. Thus, identification of the branching pattern of triantennary glycans is very simple (44). 8. The residues at the nonreducing termini of the antennae are revealed by the C1 ion: m/z 179 when hexose is present or m/z 220 when chains terminate in GlcNAc. 9. Other diagnostic ions and masses are listed in Table 2. Table 2 Ions Defining Structural Features in the Negative Ion Spectra of N-Linked Glycans Structural feature

Ion

Ionic composition −

m/z

Composition

Molecular

[M+X]

-

Antenna sequence

C

Gal, Man, Glc GlcNAc, GalNAc [Fuc]Gal Gal-[Fuc]GlcNAc αGal-Gal, Man-Man Man-[Man]Man GalNAc-GlcNAc

179 220 325 528 341 503 423

Antenna composition

F

Man GlcNAc Gal-GlcNAc Gal-[Fuc]GlcNAc αGal-Gal GalNAc-GlcNAc (Gal-GlcNAc)2

221 262 424 570 586 465 789 (continued)

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Table 2 (continued) Structural feature

Ion

Ionic composition

m/z

(Gal-GlcNAc)2Fuc (Gal-GlcNAc)3

935 1154

Fucose at 6-position of reducing terminus

2,4

AR

[M-HCl-307]− [M-HNO3-307]− [M-H3PO4-307]−

[M-343]− [M-370]− [M-405]−

Absence of fucose at 6-position of reducing terminus

2,4

AR

[M-HCl-161]− [M-HNO3-161]− [M-H3PO4-161]−

[M-197]− [M-224]− [M-259]−

GlcNAc Gal-GlcNAc Gal-[Fuc]GlcNAc (Gal-GlcNAc)2 (Gal-GlcNAc)2Fuc Man3 Man4 Man5

526, 508 688, 670 834, 816 1053, 1035 (1017) 1199, 1181 (1163) 647, 629 809, 791 971, 953

Composition of 6-antenna D and [D-18]− ([D-36]−)

O,3

AR-2 and O,4AR-2

GlcNAc GalGlcNAc Gal-[Fuc]GlcNAc (Gal-GlcNAc)2 (Gal-GlcNAc)2Fuc Man3 Man4 Man5

292, 2621 454, 424 600, 570 819, 789 965, 935 251, 221 413, 383 575, 545

Composition of 3-antenna

O,4

AR-3 (E) ion

Gal-GlcNAc GlcNAc2 Gal-GlcNAc2 (Gal-GlcNAc)2 (Gal-GlcNAc)2-Fuc

466 507 669 831 977

Presence of bisect

Abundant [D-221]− ion. Ion at [M-221-18]− Missing Ion D

GlcNAc Gal-GlcNAc (Gal-GlcNAc)2-Fuc (Gal-GlcNAc)2 (Gal-GlcNAc)2Fuc Man3 Man4 Man5

508 670 816 1035 1181 629 791 953

Presence of sialic acid

B1

Neu5Ac Neu5Gc

290 306

Presence of α2→ 6-linked sialic acid

O,4

Neu5Ac Neu5Gc

306 322

A2-CO2

1, These ions are normally of relatively low abundance. An abundant ion at the mass of the O,4A ion is more likely to be an F-type ion.

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4. Notes 1. Several other matrices are available (see [ref. 7] ) that claim to produce advantages for certain types of carbohydrate. 2. The conditioned membranes can be stored in water for several months without noticeable loss of activity. Once used, they can be regenerated by repeating the nitric acid treatment. 3. Solutions containing organic solvents should be avoided. 4. The drops should remain at the same volume but often they increase slightly as water is drawn through the membrane. If the samples contains glycerol (found in some PNGase F preparations), considerable amounts of water can be drawn into the sample and the analysis usually fails. PNGase preparations containing glycerol should, consequently, be avoided. 5. The dried sample usually has long needle-shaped crystals projecting from the periphery of the spot towards the center. If the deposit is sticky, then either the sample is much too concentrated or it contains a high concentration of detergent or glycerol. Such deposits are unlikely to yield good spectra and the sample should be purified further. 6. Some investigators prefer to premix the matrix and sample solutions prior to application to the MALDI target. In the author’s hands, this method does not appear to offer any particular advantages. 7. Only ~0.2 µL ethanol is necessary, and care must be taken to prevent spreading the sample over a larger area. This procedure is used to produce a more homogeneous target than can be achieved when water is present. 8. Sialylated glycans frequently give abundant peaks in positive ion mode due to loss of sialic acid when their spectra are recorded with reflectron-TOF instruments. Consequently, their spectra are often recorded in linear mode in order to minimize observed sialic acid loss. However, it must be borne in mind that this technique only removes PSD ions and not those fragment ions produced within the ion source. Minimizing the in-source delay is the best method for reducing the abundance of these ions. Also, it must be borne in mind that the resulting mass measurement may well be of average rather than a monoisotopic mass because of the lower resolution. 9. This effect is less pronounced with delayed-extraction instruments, but increased laser power can still lead to reduced resolution. 10. The laser spot should be moved to different areas of the target during acquisition, because not all areas of the target will yield a signal. It is frequently found with DHB that an active spot will only give a signal for a few laser shots due either to depletion of the sample/matrix mixture or vertical inhomogeneities within the crystal. Recent work (45) has suggested

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11. 12.

13.

14.

15. 16.

17.

Harvey

that carbohydrates reside predominantly on the outside of the crystals, accounting for this phenomenon. Usuaully 60–70 laser shots will be sufficient but more will be required for weak signals. The mass spectrometer should be calibrated. Suitable calibration compounds are a mixture of dextran oligomers (e.g. dextran from Leuconostoc ssp. from Fluka,) for positive ion mode (m/z 203.05, 365.11, 527.16, 689.21, 851.26, 1013.32, 1175.37, 1337.42, 1499.48, 1661.53, 1823.58, 1985.63, 2147.69, 2309.74, 2471.79, 2633.85, 2795.90, 2957.95, 3120.00, 3282.06, 3444.11, 3606.16, 3768.22, 3930.27, 4092.32; and the same compounds derivatized with 2-aminobenzoic acid for calibrating negative ion spectra (m/z 300.11, 462.16, 624.21, 786.27, 948.32, 1110.37, 1272.43, 1434.48, 1596.53, 1758.58, 1920.64, 2082.69, 2244.74, 2406.80, 2568.85, 2730.90, 2892.95, 3055.01, 3217.06, 3379.11, 3541.17, 3703.22, 3865.27, 4027.32). Formation of methyl esters allows all compounds to be examined as neutral molecules in positive ion mode. Without methylation, as well as laser-induced decomposition of sialylated glycans, these compounds will form both positive and negative ions, thus splitting the signal and the sialic acids will give additional ions by formation of sodium and potassium salts. Although negative ion spectra are usually associated with the formation of [M-H]− ions, these tend to be unstable and fragment in the mass spectrometer ion source. Addition of an inorganic salt such as ammonium chloride, nitrate or phosphate produces [M+anion]− ions that are much more stable and do not fragment until they reach the collision cell of the mass spectrometer. Phosphate is used because it has been found that phosphate adducts are the major species observed from glycans released from gel-separated samples. Fragmentation of adducts with different anions is virtually identical because the first loss is that of the adduct together with a proton to give what is effectively the [M-H]− ion. Larger anions such as bromide and iodide should be avoided because these produce very few fragment ions. The tip should be long enough to reach to the end of the capillary. MS/MS spectra of major components can be obtained in a few seconds but those of weaker components can be accumulated for many minutes in order to achieve a satisfactory signal:noise ratio. The spray should last on average from 1–3 hours. It is assumed that the mass spectrometer has been set up and is operated according to the manufacturer’s instructions. Suitable calibration compounds for negative ion work are the 2-AA-derivatized glycans listed above or underivatized glycans released from bovine fetuin.

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Acknowledgments The author thanks the Biotechnology and Biological Sciences Research Council and the Wellcome Trust for equipment grants to purchase the two mass spectrometers used in this work and further funding from the Oxford Glycobiology endowment.

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27. Brüll, L. P., Heerma, W., Thomas-Oates, J., Haverkamp, J., Kovácik, V. and Kovác, P. (1997) Loss of internal 1–6 substituted monosaccharide residues from underivatized and per-O-methylated trisaccharides. J. Am. Soc. Mass Spectrom. 8, 43–49. 28. Brüll, L. P., Kovácik, V., Thomas-Oates, J. E., Heerma, W. and Haverkamp, J. (1998) Sodium-cationized oligosaccharides do not appear to undergo ‘internal residue loss’ rearrangement processes on tandem mass spectrometry. Rapid Commun. Mass Spectrom. 12, 1520–1532. 29. Franz, A. H. and Lebrilla, C. B. (2002) Evidence for long-range glycosyl transfer reactions in the gas phase. J. Am. Soc. Mass Spectrom. 13, 325–337. 30. Harvey, D. J., Mattu, T. S., Wormald, M. R., Royle, L., Dwek, R. A. and Rudd, P. M. (2002) “Internal residue loss”: rearrangements occurring during the fragmentation of carbohydrates derivatized at the reducing terminus. Anal. Chem. 74, 734–740. 31. Chai, W., Piskarev, V. and Lawson, A. M. (2001) Negative-ion electrospray mass spectrometry of neutral underivatized oligosaccharides. Anal. Chem. 73, 651–657. 32. Chai, W., Piskarev, V. and Lawson, A. M. (2002) Branching pattern and sequence analysis of underivatized oligosaccharides by combined MS/MS of singly and doubly charged molecular ions in negative-ion electrospray mass spectrometry. J. Am. Soc. Mass Spectrom. 13, 670–679. 33. Chai, W., Piskarev, V. E., Mulloy, B., Liu, Y., Evans, P. G., Osborn, H. M. I. and Lawson, A. M. (2006) Analysis of chain and blood group type and branching pattern of sialylated oligosaccharides by negative ion electrospray tandem mass spectrometry. Anal. Chem. 78, 1581–1592. 34. Pfenninger, A., Karas, M., Finke, B. and Stahl, B. (2002) Structural analysis of underivatized neutral human milk oligosaccharides in the negative ion mode by nano-electrospray MSn (Part 1: Methodology). J. Am. Soc. Mass Spectrom. 13, 1331–1340. 35. Pfenninger, A., Karas, M., Finke, B. and Stahl, B. (2002) Structural analysis of underivatized neutral human milk oligosaccharides in the negative ion mode by nano-electrospray MSn (Part 2: Application to isomeric mixtures). J. Am. Soc. Mass Spectrom. 13, 1341–1348. 36. Quéméner, B., Désiré, C., Lahaye, M., Debrauwer, L. and Negroni, L. (2003) Structural characterisation of both positive- and negative-ion electrospray mass spectrometry of partially methyl-esterified oligogalacturonides purified by semipreparative high-performance anion-exchange chromatography. Eur. J. Mass Spectrom. 9, 45–60. 37. Sagi, D., Peter-Katalinic, J., Conradt, H. S. and Nimtz, M. (2002) Sequencing of tri- and tetraantennary N-glycans containing sialic acid by negative mode ESI QTOF tandem MS. J. Am. Soc. Mass Spectrom. 13, 1138–1148. 38. Börnsen, K. O., Mohr, M. D. and Widmer, H. M. (1995) Ion exchange and purification of carbohydrates on a Nafion(R) membrane as a new sample pretreatment for matrix-assisted laser desorption-ionization mass spectrometry. Rapid Commun. Mass Spectrom. 9, 1031–1034.

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39. Domon, B. and Costello, C. E. (1988) A systematic nomenclature for carbohydrate fragmentations in FAB-MS/MS spectra of glycoconjugates. Glycoconj. J. 5, 397–409. 40. Harvey, D. J. (2005) Fragmentation of negative ions from carbohydrates: Part 1; Use of nitrate and other anionic adducts for the production of negative ion electrospray spectra from N-linked carbohydrates. J. Am. Soc. Mass Spectrom. 16, 622–630. 41. Harvey, D. J. (2005) Fragmentation of negative ions from carbohydrates: Part 2, Fragmentation of high-mannose N-linked glycans. J. Am. Soc. Mass Spectrom. 16, 631–646. 42. Harvey, D. J. (2005) Fragmentation of negative ions from carbohydrates: Part 3, Fragmentation of hybrid and complex N-linked glycans. J. Am. Soc. Mass Spectrom. 16, 647–659. 43. Harvey, D. J., Royle, L., Radcliffe, C. M., Rudd, P. M. and Dwek, R. A. (2008) Structural and quantitative analysis of N-linked glycans by MALDI and negative ion nanospray mass spectrometry. Anal. Biochem. 376, 44–60. 44. Harvey, D. J., Crispin, M., Scanlan, C., Singer, B. B., Lucka, L., Chang, V. T., Radcliffe, C. M., Thobhani, S., Yuen, C.-T. and Rudd, P. M. (2008) Differentiation between isomeric triantennary N-linked glycans by negative ion tandem mass spectrometry and confirmation of glycans containing galactose attached to the bisecting (b1-4-GlcNAc) residue in N-glycans from IgG. Rapid Commun. Mass Spectrom. 22, 1047–1052. 45. Bouschen, W. and Spengler, B. (2007) Artifacts of MALDI sample preparation investigated by high-resolution scanning microprobe matrix-assisted laser desorption/ionization (SMALDI) imaging mass spectrometry. Int. J. Mass Spectrom. 266, 129–137.

150 MS Analysis of Protein Glycosylation Nobuaki Takemori, Naoka Komori, and Hiroyuki Matsumoto 1. Introduction Post-translational modifications (PTM) are important molecular events because they generate functional diversities of proteins. A large number of proteins undergo multiple PTM in vivo and over 300 types of PTM have been identified to date (1). Among various PTM, phosphorylation and glycosylation have been intensely studied because of their physiological significance. Phosphorylation plays key regulatory roles in cellular signaling processes (2), and glycosylation mediates crucial cellular mechanisms such as protein folding and trafficking (3, 4). To confirm the presence of these modifications in vivo, the conventional analytical procedures require large quantities of samples and laborious biochemical steps. For this reason, small-scale analysis of PTM remains among the most challenging areas of biological science. This chapter describes our recent proteomic methodology using the combination of two-dimensional gel electrophoresis (2-DGE) and MALDI quadrupole ion trap time-of-flight mass spectrometer (MALDI-QIT-TOF MS), which enables a highly sensitive analysis of post-translational modifications (PTM). Two-DGE has been used extensively as a powerful tool to separate proteins from a limited amount of biological samples (5–8). We stained gels with Pro-Q™ Emerald/ Diamond dyes immediately after 2-DGE and prior to CBB staining, and because this allows us to perform direct detection of glycoproteins/phosphoproteins on the same 2-D gels (9,10). Gel-separated glycoproteins are further subjected to the structural analysis using MALDI-QIT-TOF MS. MALDI-QIT-TOF MS is capable of structural characterization of various PTMs through highly sensitive fragmentation, or MS/MS, analysis (11,12). In MALDI-QIT-TOF MS, two mass analyzers (i.e., QIT and TOF) are connected to MALDI ion source, and the combination of MALDI and QIT renders a unique opportunity to perform multistage fragmentation (MSn) analysis with high-energy collision-induced From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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dissociation (CID) (11,13). In this chapter, we will describe proteomic techniques that we use to analyze N-linked glycosylation. However, the protocol can be applied to other PTM analyses with some modifications. 2. Materials 1. 2. 3. 4. 5. 6. 7. 8. 9.

MALDI-QIT-TOF MS (AXIMA QIT; Shimadzu Biotech, Manchester, UK) MALDI-TOF MS MALDI stainless sample target Acetonitrile (HPLC grade) Ammonium bicarbonate Trifluoroacetic acid (TFA) Milli-Q water (Millipore, Bedford, MA, USA) Sequencing-grade trypsin (Promega Madison, WI, USA). Stock solutions for in-gel digestion: a. 50% (v/v) acetonitrile/100 mM ammonium bicarbonate (pH 8.9) b. 50% (v/v) acetonitrile/5% (v/v) TFA c. 0.2% (v/v) TFA solution d. 10 mM ammonium bicarbonate (pH 8.9) 10. Recrystalized 2,5 dihydroxybenzoic acid (DHB) 11. DHB solution for MS analysis: 2% (w/v) DHB in 50% (v/v) acetonitrile/0.1% (v/v) TFA (see Note 1)

3. Methods Figure 1 shows a schematic diagram of our glycoprotein analysis. Protein samples are first separated using 2-DGE, and glycoprotein spots are detected using a glycoprotein staining reagent (Subheading 3.1). Glycoprotein spots are in-gel digested with trypsin (Subheading 3.2) (see Note 2), and identified by peptide mass fingerprinting analysis (Subheading 3.3). In order to characterize their glycan structures and modification sites, tryptic glycopeptides are further subjected to fragmentation analysis in MALDI-QIT-TOF MS (Subheading 3.4). Because human keratin contamination interferes with MS analysis, it is recommended that powder-free latex gloves be worn throughout the procedure. 3.1. Gel Electrophoresis and Glycoprotein Staining 1. Perform 2-DGE to resolve glycoproteins. We perform isoelectric focusing for the 1st dimension and SDS-PAGE for the 2nd dimension (see Note 3). Instead of 2-DGE, one-dimensional gel electrophoresis (e.g., SDS-PAGE) can be used for this protocol. However, the resolution of proteins on a one-dimensional gel is generally inadequate for the separation of complex protein mixtures. 2. Following 2-DGE, gels are stained with a glycoprotein staining reagent for detection of glycoproteins (see Note 4) or followed by Coomassie Brilliant Blue (CBB) for visualization of all protein spots (see example in Fig. 2).

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B In-Gel Tryptic Digestion

CBB (Total Protein Staining)

SDS-PAGE

IEF

C Peptide Mass Fingerprinting by MALDI-TOF MS

Pro-Q Emerald (Glycoprotein Staining)

m/z

D Structural Characterization by MALDI-QIT-TOF MS

Mannose (162 Da) GlcNAc (203 Da) Fucose (146 Da)

Fig. 1. A schematic drawing of glycoprotein analysis using 2-DGE and MALDI-QITTOF MS. (A) Proteins are separated by 2-DGE based on their isoelectric focusing points and molecular weights in the first and second dimension, respectively. A glycosylated protein spots are detected by staining with Pro-Q Emerald dye. (B) After in-gel digestion of the glycoprotein spot with trypsin, the tryptic peptides are subjected to peptide mass fingerprinting analysis. (C) Peptide mass fingerprinting analysis provides the mass profiles and sequence information of the tryptic peptides. (D) A glycopeptide thus detected is further fragmented by MALDI-QIT-TOF MS. Using the information from fragment ions, the structure of a glycan moiety can be deduced.

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B CBB

Fig. 2. Differential staining of Drosophila proteins with a fluorescent glycosylation sensor dye, Pro-Q™ Emerald (A) and Coomassie blue (B). Drosophila eye proteins were separated on 2-D gel. Glycosylated protein spot (arrow) was identified by staining with Pro-Q Emerald dye. Our peptide mass fingerprinting analysis identified the spot as chaoptin.

3.2. In-Gel Tryptic Digestion 1. After detection of protein spots positive for glycoprotein staining, corresponding protein spots stained with CBB are excised by a razor blade and transferred into a 1.5 ml polypropylen tube. Excised gel pieces are shaken in 1 ml of 50% (v/v) acetonitrile/100 mM ammonium bicarbonate (pH 8.9) at room temperature to remove CBB. 2. Distained gel pieces are incubated with 1 ml of acetonitrile for 30 min to dehydrate the gels. After removing acetonitrile, the gel pieces are rehydrated with 2 µl of trypsin solution for 30 min at 4°C. Add 50 µl of 10 mM ammonium bicarbonate (pH 8.9) and incubate at 37°C for 5–10 hrs. During the incubation, trypsin continues

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to digest a protein in a gel piece and while some of the tryptic peptides come out of the gel piece. 3. Transfer the solution containing tryptic digests into a new 0.65 ml polypropylen tube. To extract the rest of the tryptic digests from the gel, vigorously shake the gel in 50 µl of 50% (v/v) acetonitrile/5% (v/v) TFA for 20 min. After a brief centrifuge, collect and transfer the supernatant into the new tube to mix with the original digest solution. 4. Concentrate the collected peptide solution down to near dryness using vacuum centrifuge (see Note 5).

3.3. Peptide Mass Fingerprinting Analysis using MALDI-TOF MS 1. Each peptide sample is reconstructed with 5 µl of 0.2% (v/v) TFA solution. Load the peptide solution (0.5 µl) onto a MALDI-TOF sample target plate with a fresh DHB solution (0.5 µl) and dry the mixture completely at room temperature. 2. Peptide samples on the MALDI sample target are subjected to peptide mass fingerprinting analysis using MALDI-TOF MS. After measuring the masses of the tryptic peptides by MALDI-TOF MS, the observed masses are submitted to MASCOT PMF search program (Matrix Science, London, UK) at http://matrix science.com. 3. We routinely perform MASCOT search against the latest version of the National Center for Biotechnology Information (NCBI) nonredundant database using the following parameters; (a) unlimited protein molecular weight and pI ranges, (b) presence of protein modifications including acrylamide modification of cysteine, methionine oxidation, protein N-terminus acetylation, and pyroglutamic acid, and (c) mass tolerance of ± 0.25 Da. We consider the confirmation to be positive when a significant MOWSE score (p < 0.05) is generated. 4. After the protein identification by peptide mass fingerprinting analysis, the masses of the observed peptides are compared to that of the theoretical tryptic peptides. The peptides which contain glycans will be detected as the unassigned ion peaks because glycosylation leads to a predictable increase in the mass of a modified amino acid residue. When unassigned ion peaks are detected in MS spectrum, the masses of the unassigned peaks are subjected to ExPASy-GlycoMod Tool (http://au.expasy.org/tools/glycomod) to predict possible glycan structures and modification sites. To calculate the theoretical fragment ions resulting from CID fragmentation of the predicted peptide, we use MS-Product program (http:// prospector.ucsf.edu/).

3.4. Structural Characterization of Protein Glycosylation Using MALDI-QIT-TOF MS 1. Load peptide solution (0.5 µl) onto a MALDI-QIT-TOF sample target with a fresh DHB solution (0.5 µl) and dry the mixture completely at room temperature. Each peptide sample is subjected to MALDI-QIT-TOF MS analysis. 2. In general, an ionized glycopeptide is fragmented without CID in MALDIQIT-TOF MS, and as a result, a precursor glycopeptide together with some of its fragment ions will be detected in MS mode. Sugar components are readily

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A MS

*

2154.9 Hexose (162 Da) HexNAc (203 Da)

*

*

2239.0

B

MS/MS 3978.5 P 1951.4

P 2154.9

P

∆203

3006.5 3168.5 3330.5 3492.6 3654.3 3816.7

2520.3

∆486 (162 x 3)

∆162

∆162 ∆162

y14 1120

Relative Intensity (%)

∆162 ∆162

y6-H2O

y9-H2O

y5

y7

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y4

L T N I T F S GP QF TN LNE R 1136 b10

y4

∆162 ∆162

3000

MS/MS 2154.9

y5-H2O

7

3978.5

2000

100

*

*

P 2357.8

∆203

C

*

*

3492.6 3654.3 3816.7 3978.5

3077.6

P [1120-1136]

b16

b16-H2O

y10-H2O

y8-H2O y9 y10 y11-H2O y11 b10

y12

y13

y14

b16

0 600

1000

1400 m/z

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2200

Fig. 3. Structural characterization of chaoptin glycopeptide using MALDI-QIT-TOF MS. The tryptic peptides of chaoptin (Drosophila glycoprotein) separated on 2-D gel (Fig. 2) were subjected to a series of CID fragmentation to deduce the glycan sequence and glycosylation site. (A) MS spectrum of the tryptic digests. *, peptide ions not assigned as tryptic peptides of chaoptin. (B) MS/MS spectrum of an unassigned ion peak at m/z 3978.5 observed in MS mode. CID fragmentation resulted in a series of fragment ions separated by 162, 203, or 486 (162 × 3) Da. (C) MS/MS spectrum of a fragment ion at m/z 2154.9. The y ion and b ion series derived from the peptide corresponded to the amino acid residues 1120 to 1136 of chaoptin. ≠, a putative N-linked glycosylation site. , a peptide ion with a cross-ring cleavage of HexNAc residue. in dark gray, hexose residue; in light gray, N-acetylhexosamine (HexNAc) residue. (With permission from ref. 11)

distinguishable from amino acid residues and other post-translationally modified residues based on their masses. As a result, we can easily distinguish glycopeptides from other nonglycopeptides.

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3. To determine the actual glycan structure, an ionized glycopeptide is further subjected to fragmentation analysis in MS/MS mode. As compared with peptide-bonds, glycosidic bonds are susceptible to CID fragmentation resulting in a sequential loss of sugar components from the terminal end of a glycan moiety (see example in Fig. 3). 4. Because some peptide-bonds seldom undergo total dissociation in a single run of MS/MS analysis, further fragmentation analysis by MSn analysis may be required to determine the amino acid sequence and modification site(s).

4. Notes 1. Prepare fresh DHB solution just before use. 2. Some glycoproteins are not susceptible to tryptic cleavage due to the sterical hindrance of their glycans. Digestion with other proteases may improve the recovery of glycopeptides. 3. In our laboratory, isoelectric focusing gel electrophoresis is carried out using IPGphor (GE Healthcare/Amersham, Buckinghamshire, UK) with a cup loading strip holder. Immobiline Dry Strip (pH 3–10, 13 cm in length) is rehydrated for 15 hr at room temperature with 250 µl of lysis solution (8.5 M urea, 2% (w/v) CHAPS, 0.2% (w/v) Bio-Lyte 3/10 (BIO-RAD, Hercules, CA, USA), and 5% (v/v) β-mercaptoethanol). The same lysis solution (100 µl) is used for sample homogenization. After centrifugation at 15,000 × g for 10 min, the supernatant is loaded into a sample cup. Electrophoresis is performed at 500 V for 3 min, 4,000 V for 2 hr, and 8,000 V up to a total of 22,000 Vhr. 4. Several glycoprotein staining reagents are commercially available. We use Pro-Q™ Emerald glycoprotein gel stain (Molecular Probes, Eugene, OR, USA) because of its high sensitivity. Regarding the details of the Pro-Q staining, refer to manufacture’s instructions. 5. Peptide samples can be stored at −20°C at least for several months. References 1. Delta Mass, a database of protein post translational modifications (http://www. abrf.org/index.cfm/dm.home). 2. Graves, J. D. and Krebs, E. G. (1999) Protein phosphorylation and signal transduction. Pharmacol. Ther. 82, 111–121, 3. Helenius, A. and Aebi, M. (2000) Intracellular functions of N-linked glycans. Science 291, 2364–2369. 4. Roseman, S. (2001) Reflections on glycobiology. J Biol Chem. 276, 41527–41542. 5. Matsumoto, H., O’Tousa, J. E., and Pak, W. L. (1982) Light-induced modification of Drosophila retinal polypeptides in vivo. Science 217, 839–841. 6. Matsumoto, H. and Pak, W. L. (1984) Light-induced phosphorylation of retinaspecific polypeptides of Drosophila in vivo. Science 223, 184–186.

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7. Matsumoto, H., Kurien, B. T., Takagi, Y., Kahn, E. S., Kinumi, T., Komori, N., Yamada, T., Hayashi, F., Isono, K., Pak, W. L., Jackson, K. W., and Tobin, S. L. (1994) Phosrestin I undergoes the earliest light-induced phosphorylation by a calcium/calmodulin-dependent protein kinase in Drosophila photoreceptors. Neuron 12, 997–1010. 8. Komori, N., Takemori, N., Kim, H. K., Singh, A., Hwang, S. H., Foreman, R. D., Chung, K., Chung, J. M., and Matsumoto, H. (2007) Proteomics study of neuropathic and nonneuropathic dorsal root ganglia: Altered protein regulation following segmental spinal nerve ligation injury. Physiol. Genomics. 29, 215–230. 9. Hart, C., Schulenberg, B., Steinberg, T. H., Leung, W. Y., and Patton, W. F. (2003) Detection of glycoproteins in polyacrylamide gels and on electroblots using Pro-Q Emerald 488 dye, a fluorescent periodate Schiff-base stain. Electrophoresis 24, 588–598. 10. Steinberg, T. H., Agnew, B. J., Gee, K. R., Leung, W. Y., Goodman, T., Schulenberg, B., Hendrickson, J., Beechem, J. M., Haugland, R. P., and Patton, W. F. (2003) Global quantitative phosphoprotein analysis using Multiplexed Proteomics technology. Proteomics 3, 1128–1144. 11. Takemori, N., Komori, N., and Matsumoto, H. (2006) Highly sensitive multistage mass spectrometry enables small-scale analysis of protein glycosylation from twodimensional polyacrylamide gels. Electrophoresis 27, 1394–1406. 12. Takemori, N., Komori, N., Thompson, J. N. Jr, Yamamoto, M-T., and Matsumoto, H. (2007) Novel eye-specific calmodulin methylation characterized by protein mapping in Drosophila melanogaster. Proteomics 7, 2651–2658. 13. Martin, R. L. and Brancia, F. L. (2003) Analysis of high mass peptides using a novel matrix-assisted laser desorption/ionisation quadrupole ion trap time-of-flight mass spectrometer. Rapid Commun Mass Spectrom. 17, 1358–1365.

151 Mapping protein N-Glycosylation by COFRADIC Bart Ghesquière, Joël Vandekerckhove, and Kris Gevaert 1. Introduction The majority of proteins require co- and posttranslational modifications to become functional entities. Such modifications fulfill a plethora of tasks (e.g., modulation of enzyme activity and directing proteins to cellular compartments). One of these protein modifications that, amongst many others, plays an essential role in determining a protein’s structure and functionality is N-glycosylation, which is the enzymatic attachment of a glycan to an asparagine in the consensus motif NXS/T/C (1,2). Next to asparagines, glycosylation can also occur on serine and threonine (O-glycosylation) (3–6) and tryptophan (C-glycosylation) (7). Protein N-glycosylation has intrinsic as well as extrinsic functions. Intrinsic functions brought about by this modification are the creation of structural modules and changes of the solubility, antigenicity and stability of the affected protein. Examples of extrinsic functions are the routing of proteins to a specific subcellular localization, and the guiding and modulation of cell adhesion and intracellular communication. Congenital Disorders of Glycosylation (CDG) are a group of diseases caused by glycan synthesis errors (8). Several CDG subtypes are distinguished based on the malfunctioning of a specific enzyme in the protein-N-glycosylation pathway. Other states such as the malignant transformation of cells to cancerous cells portray altered glycan chains on glycoproteins (9–14). Both examples show the importance of protein glycosylation in the normal physiology and pathophysiology of organisms and illustrate the value of monitoring protein N-glycosylation sites on a proteomic scale. Several methods for studying protein N-glycosylation have been described. Lectin based methods rely on the affinity of lectins for subsets of glycans to isolate glycosylated proteins or peptides (15,16). However, to identify the broadest possible range of N-glycosylated molecules, different lectins must be combined, From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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either in parallel or serial, to a multilectin affinity column (14,17–20). Hydrazide based techniques chemically enrich N-glycosylated peptides and are not biased towards any specific glycan structure (21–23). Although the hydrazide technology offers a straightforward analysis of N-glycosylation sites, all information about the actual nature of the N-glycan is lost. We recently developed a gel-free technique for analyzing protein-N-glycosylation (24) based upon combined fractional diagonal chromatography (COFRADIC 25) and we here describe a detailed protocol for isolating N-glycosylated peptides for MS/MS analysis. Briefly, proteins in a complex sample such as serum are digested with trypsin. These peptides are separated a first time on a RP-HPLC column (primary COFRADIC run) and time-based collected into several primary fractions. Sufficiently spaced primary fractions are combined, treated with PNGase F and reseparated by the same RP-HPLC setup (secondary COFRADIC runs). PNGase F cleaves N-glycans from asparagines and hereby converts these to aspartic acid, thus creating a marker (1 Da difference) that can be used to interpret results of database searches using MS/MS-spectra. Deglycosylated peptides display different retention on the reverse-phase column during the secondary HPLC separations and thus shift away from nonglycosylated peptides. These shifts finally lead to the specific collection of formerly in vivo N-glycosylated peptides which may then be analyzed by MS techniques. 2. Materials 2.1. Protein Preparation 1. 2. 3. 4.

Tris(2-carboxyethyl)phosphine (TCEP, Pierce, Rockford, IL). Iodoacetamide (IA). Hydrogen peroxide:(30% (w/w) in water. Disposable desalting columns packed with Sephadex™ G-25 (GE Healthcare BioSciences, Uppsala, Sweden). 5. Sequencing grade modified trypsin (Promega, Madison, WI, USA).

2.2. COFRADIC Protocol for the Isolation of N-Glycopeptides 1. Analytical RP-HPLC column: 2.1 mm internal diameter (I.D.) × 150 mm (length) 300SB-C18 column, Zorbax® (Agilent, Waldbronn, Germany). 2. HPLC grade water and acetonitrile. 3. PNGase F from Elizabethkingia (Chrysobacterium/Flavobacterium) meningosepticum (Sigma-Aldrich). 4. Acetic acid glacial, analytical reagent grade.

3. Methods The methods described below use a serum proteome for characterization of N-glycosylation sites. It should be clear that any proteome or peptidome in which N-glycosylated asparagines are present can be sampled by these methods.

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3.1. Preparation of Serum Proteins Prior to COFRADIC 1. 1 mL of mouse serum proteins (see Note 1) deprived of its three most abundant proteins is desalted on a NAP-10 column and eluted in 1.5 mL of 2 M guanidinium hydrochloride in 50 mM Tris-HCl pH 8.6. 2. Reduce the sample volume to 1 mL by vacuum drying. 3. Add freshly prepared TCEP.HCl and iodoacetamide solutions to 10 and 20 mM final concentrations respectively. Let the reduction and alkylation of cysteines proceed in the dark for 1 h at 37°C (see Note 2). 4. Desalt the mixture of modified proteins in 1.5 mL of freshly prepared 20 mM NH4HCO3 (pH 7.6) using a NAP-10 column. 5. Heat the protein mixture for 10 min at 95°C followed by cooling for 10 min on ice. 6. Add sequence grade modified trypsin (the enzyme/substrate ratio should be 1/50 to 1/100 (w/w) ) and incubate overnight at 37°C. 7. Reduce the volume to 500 µL by vacuum drying.

3.2. COFRADIC Sorting of N-Glycosylated Peptides 1. Acidify 98 µL of the peptide mixture with 2 µL of a 50% (w/v) acetic acid solution (f.c. of 1%). 2. Add 2 µL of a 30% (w/v) hydrogen peroxide solution (f.c. of about 0.6% of H2O2) to oxidize methionines to sulfoxides. The peptides are incubated for 30 min at 30°C (see Note 3). 3. Load the sample immediately on the reverse-phase column (see 3.3) for the primary COFRADIC separation and collect peptides in 56 consecutive fractions of 1 min each starting 25 min after sample injection (see Note 4). 4. Pool primary fractions that are spaced by 14 min and dry them under vacuum. 5. Redissolve the dried peptides in 97 µL of 50 mM NH4HCO3 (pH 7.6) and add 1 µL of a 0.5 U/µL stock solution of PNGase F. This mixture is incubated for 3 hours at 37°C to fully deglycosylate the N-glycosylated peptides. 6. Add 2 µL of a 50% (w/v) acetic acid solution and reseparate each PNGase F treated pool of primary fractions on the same RP-HPLC column under the same conditions. Per primary fraction, deglycosylated peptides are collected in two intervals: one 11 to 3 min prior to and one 2 to 10 min following the original collection interval. Thus, per secondary run, 5 such secondary intervals are collected, each consisting of 4 secondary fractions of 2 min (or 160 µL) (Fig. 1 and Table 1). 7. Such isolated secondary fractions are dried and stored at −20°C until further MS/ MS-analysis (see Note 5).

3.3. Reverse Phase HPLC Separation in COFRADIC The solvent gradient described below is applied both for the primary and the secondary runs. We found it important to use a HPLC solvent system buffering around a pH of about 5. In contrast to lower pH values, at pH 5 the extent of the chromatographic shift of deglycosylated versus glycosylated peptides is primarily based on the removal or gain of charges. Due to the nature of the glycans, both hydrophilic and hydrophobic shifts can be expected. When the

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mAU

1000 800 600 400 200 0 10

20

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80

min

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200 150

*

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Fig. 1. Example of a primary and a secondary COFRADIC RP-HPLC run for isolating N-glycopeptides. The upper panel shows the primary RP-HPLC separation (UV absorbance at 214 nm) of a tryptic digest of human alpha-1-acid-glycoprotein and bovine fetuin; primary fractions were collected between 25 and 81 min and pooled as described in Subheading 3.2. Such collected peptides were then treated with PNGase F to remove N-linked glycan chains. The lower panel shows the secondary RP-HPLC separation of the PNGase F treated primary fractions that eluted between 27–28, 41–42, 55–56 and 69–70 min (indicated in dark grey boxes). The secondary collection intervals containing the shifted, deglycosylated peptides (indicated in light grey boxes) were collected 11 to 3 min prior to and 2 to 10 min following each primary fraction. There is a clear shift of peptides (indicated with an asterisk) that elute in the secondary collection interval between the primary fractions collected at 41–42 and 55–56 min. Further MS and MS/MS analyses identified tryptic peptides carrying known N-glycosylation sites of bovine fetuin (N99 and N176) and human alpha1-acid glycoprotein (N93).

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Table 1 Fraction Pooling Scheme for COFRADIC Isolation of N-Glycosylated Peptides One example of the pooling of secondary fractions is given (see Figure 1) in which it is made clear that peptides undergoing a hydrophobic shift following PNGase F treatment of fraction X overlap with fractions undergoing a hydrophilic shift from fraction X + 14 following the same treatment.

POOLING SCHEME OF PRIMARY FRACTIONS

A B C D E F G H I J K L M N

prim. elution fraction interval

prim. elution fraction interval

prim. elution fraction interval

prim. elution fraction interval

1 2 3 4 5 6 7 8 9 10 11 12 13 14

15 16 17 18 19 20 21 22 23 24 25 26 27 28

29 30 31 32 33 34 35 36 37 38 39 40 41 42

43 44 45 46 47 48 49 50 51 52 53 54 55 56

25–26 min 26–27 min 27–28 min 28–29 min 29–30 min 30–31 min 31–32 min 32–33 min 33–34 min 34–35 min 35–36 min 36–37 min 37–38 min 38–39 min

39–40 min 40–41 min 41–42 min 42–43 min 43–44 min 44–45 min 45–46 min 46–47 min 47–48 min 48–49 min 49–50 min 50–51 min 51–52 min 52–53 min

53–54 min 54–55 min 55–56 min 56–57 min 57–58 min 58–59 min 59–60 min 60–61 min 61–62 min 62–63 min 63–64 min 64–65 min 65–66 min 66–67 min

67–68 min 68–69 min 69–70 min 70–71 min 71–72 min 72–73 min 73–74 min 74–75 min 75–76 min 76–77 min 77–78 min 78–79 min 79–80 min 80–81 min

POOLING SCHEME OF SECONDARY FRACTIONS FROM PRIMARY POOL C (cfr. Figure 1) fraction 3 C

16–24 min

fraction 17 30–38 min

44–52 min

fraction 31 58–66 min

fraction 45 72–80 min

glycan is not charged, the evoked shift will be hydrophilic by the deamidation of the asparagine to aspartic acid (gain of negative charge). If the glycan was charged (e.g., sialic acid, phosphate …) the glycopeptide looses charged residues and undergoes a hydrophobic shift. Lowering the pH of the solvent system (e.g., using TFA instead of ammonium acetate) will have minor effect on acidic residues (these will mainly be neutral) and was found to lead to insufficient chromatographic shifts and inefficient collection of deglycosylated peptides. 1. Prepare HPLC solvent A (10 mM ammonium acetate (see Note 6) (pH 5.5) in water/acetonitrile, 98/2 (v/v) ) and HPLC solvent B (10 mM ammonium acetate (pH 5.5) in water/acetonitrile, 30/70 (v/v) ). 2. The following RP-HPLC solvent gradient is applied: a. Following injection of the sample onto the column, apply a 10 min isocratic separation with 100% of solvent A at a constant flow rate of 80 µl/min. b. Apply a linear gradient to 100 % over 100 min (i.e. an increase of 1 % solvent B per minute).

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c. Apply a 10 min isocratic wash with 100% of solvent B, followed by a linear gradient over 5 min to 0% of solvent B (100% of solvent A). d. Reequilibrate the column for another 20 min with 100% of solvent A before injection of another sample.

4. Notes 1. For the studies mentioned above, we were specifically interested in identifying low-abundant N-glycosylated serum proteins. Therefore, we performed MARS (multiple affinity removal system) depletion of abundant serum proteins (albumin, transferrin and IgG’s) to increase the possibility of identifying low-abundant glycoproteins (26). MARS depletion (Agilent, Waldbronn, Germany, #5188–5218) was applied to 90 µL serum from a male C57 black strain mouse according to the manufacturer’s protocol and yielded a protein mixture with a concentration of about 1 mg/mL. We then typically load a peptide amount equivalent to about 200 µg of protein material on the RP-column for the primary COFRADIC run. 2. Blocking cysteine residues by iodoacetamide avoids the formation of disulfide bridges between different cysteinyl peptides further on in the experiment. For the alkylation of cysteines we use an estimated 750-fold molar excess of TCEP and a 1500-fold molar excess of IA over the proteins (an average molecular weight of 60,000 g/mol for proteins is considered here). Addition of TCEP-HCl can change the pH of the protein solution and it is therefore necessary to check the pH and correct it by adding the appropriate volumes of a 1 M stock solution of NaOH. It is crucial to keep the pH at around 8 since a more acidic pH may result in an incomplete alkylation. 3. Oxidation of methionines by hydrogen peroxide prior to the primary COFRADIC run prevents unwanted shifts of methionine peptides during the secondary runs. Spontaneous oxidation of methiones between the two runs will cause a hydrophilic shift of nonglycosylated methionyl peptides (25) that will finally get sampled by the mass spectrometers. Methionine oxidation should be carried out strictly as described in Subheading 3.2.2 since prolonged incubation lead to nonquantitative formation of methionine-sulphon, oxidation of tryptophan and cysteine. 4. Most of the peptides separated using the described RP-HPLC gradient and solvents will elute between 25 min and 81 min. Intuitively, depending on the peptide elution profile during the primary run, the collection scheme should be adapted. 5. Following COFRADIC, N-glycosylated peptides are characterized by deamidation of deglycosylated asparagines in NXS/T motif. However, spontaneous deamidation between the primary and secondary runs (especially at Asn-Gly and Asn-Ser sites (27)) also leads to hydrophilic shifts of non-glycosylated peptides into the secondary collection intervals.

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However, such peptides will be identified with a deamidated asparagine that resides outside the NXS/T motif and should be filtered out of the final list of identifications. 6. For the ammonium acetate buffer system we use a 500 mM ammonium acetate stock solution that is prepared as follows: for 2 L of this stock solution we titrate 57.19 mL of acetic acid with NH4OH to a pH of 5.5 (about 60 mL of NH4OH should be added). Acknowledgments The authors would like to acknowledge financial support by the Fund for Scientific Research – Flanders (Belgium) (project number 3G.0280.07).

References 1. Bause, E. (1983) Structural requirements of N-glycosylation of proteins. Studies with proline peptides as conformational probes. Biochem J 209, 331–6. 2. Kornfeld, R., and Kornfeld, S. (1985) Assembly of asparagine-linked oligosaccharides. Annu Rev Biochem 54, 631–64. 3. Ernst, J. F., and Prill, S. K. (2001) O-glycosylation. Med Mycol 39 Suppl 1, 67–74. 4. Hounsell, E. F., Davies, M. J., and Renouf, D. V. (1996) O-linked protein glycosylation structure and function. Glycoconj J 13, 19–26. 5. Jentoft, N. (1990) Why are proteins O-glycosylated? Trends Biochem Sci 15, 291–4. 6. Van den Steen, P., Rudd, P. M., Dwek, R. A., and Opdenakker, G. (1998) Concepts and principles of O-linked glycosylation. Crit Rev Biochem Mol Biol 33, 151–208. 7. Vliegenthart, J. F., and Casset, F. (1998) Novel forms of protein glycosylation. Curr Opin Struct Biol 8, 565–71. 8. Kjaergaard, S., Schwartz, M., and Skovby, F. (2001) Congenital disorder of glycosylation type Ia (CDG-Ia): phenotypic spectrum of the R141H/F119L genotype. Arch Dis Child 85, 236–9. 9. Kasbaoui, L., Harb, J., Bernard, S., and Meflah, K. (1989) Differences in glycosylation state of fibronectin from two rat colon carcinoma cell lines in relation to tumoral progressiveness. Cancer Res 49, 5317–22. 10. Multhaupt, H. A., Arenas-Elliott, C. P., and Warhol, M. J. (1999) Comparison of glycoprotein expression between ovarian and colon adenocarcinomas. Arch Pathol Lab Med 123, 909–16. 11. Rodriguez-Pineiro, A. M., de la Cadena, M. P., Lopez-Saco, A., and RodriguezBerrocal, F. J. (2006) Differential expression of serum clusterin isoforms in colorectal cancer. Mol Cell Proteomics 5, 1647–57. 12. Samadi, A. A., Davidson, S. D., Mordente, J. A., Choudhury, M. S., Tazaki, H., Mallouh, C., and Konno, S. (1999) Differential Glycosylation of Cellular Prostate Specific Antigen and the Regulatory Mechanism of Its Secretion. Mol Urol 3, 147–52. 13. Schulenberg, B., Beechem, J. M., and Patton, W. F. (2003) Mapping glycosylation changes related to cancer using the Multiplexed Proteomics technology: a protein differential display approach. J Chromatogr B Analyt Technol Biomed Life Sci 793, 127–39.

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14. Zhao, J., Patwa, T. H., Qiu, W., Shedden, K., Hinderer, R., Misek, D. E., Anderson, M. A., Simeone, D. M., and Lubman, D. M. (2007) Glycoprotein microarrays with multi-lectin detection: unique lectin binding patterns as a tool for classifying normal, chronic pancreatitis and pancreatic cancer sera. J Proteome Res 6, 1864–74. 15. Zhao, J., Simeone, D. M., Heidt, D., Anderson, M. A., and Lubman, D. M. (2006) Comparative serum glycoproteomics using lectin selected sialic acid glycoproteins with mass spectrometric analysis: application to pancreatic cancer serum. J Proteome Res 5, 1792–802. 16. Kaji, H., Saito, H., Yamauchi, Y., Shinkawa, T., Taoka, M., Hirabayashi, J., Kasai, K., Takahashi, N., and Isobe, T. (2003) Lectin affinity capture, isotope-coded tagging and mass spectrometry to identify N-linked glycoproteins. Nat Biotechnol 21, 667–72. 17. Patwa, T. H., Zhao, J., Anderson, M. A., Simeone, D. M., and Lubman, D. M. (2006) Screening of glycosylation patterns in serum using natural glycoprotein microarrays and multi-lectin fluorescence detection. Anal Chem 78, 6411–21. 18. Yang, Z., and Hancock, W. S. (2005) Monitoring glycosylation pattern changes of glycoproteins using multi-lectin affinity chromatography. J Chromatogr A 1070, 57–64. 19. Yang, Z., and Hancock, W. S. (2004) Approach to the comprehensive analysis of glycoproteins isolated from human serum using a multi-lectin affinity column. J Chromatogr A 1053, 79–88. 20. Thompson, S., Stappenbeck, R., and Turner, G. A. (1989) A multiwell lectinbinding assay using lotus tetragonolobus for measuring different glycosylated forms of haptoglobin. Clin Chim Acta 180, 277–84. 21. Liu, T., Qian, W. J., Gritsenko, M. A., Camp, D. G., 2nd, Monroe, M. E., Moore, R. J., and Smith, R. D. (2005) Human plasma N-glycoproteome analysis by immunoaffinity subtraction, hydrazide chemistry, and mass spectrometry. J Proteome Res 4, 2070–80. 22. Zhang, H., and Aebersold, R. (2006) Isolation of glycoproteins and identification of their N-linked glycosylation sites. Methods Mol Biol 328, 177–85. 23. Zhang, H., Li, X. J., Martin, D. B., and Aebersold, R. (2003) Identification and quantification of N-linked glycoproteins using hydrazide chemistry, stable isotope labeling and mass spectrometry. Nat Biotechnol 21, 660–6. 24. Ghesquiere, B., Van Damme, J., Martens, L., Vandekerckhove, J., and Gevaert, K. (2006) Proteome-wide characterization of N-glycosylation events by diagonal chromatography. J Proteome Res 5, 2438–47. 25. Gevaert, K., Van Damme, J., Goethals, M., Thomas, G. R., Hoorelbeke, B., Demol, H., Martens, L., Puype, M., Staes, A., and Vandekerckhove, J. (2002) Chromatographic isolation of methionine-containing peptides for gel-free proteome analysis: identification of more than 800 Escherichia coli proteins. Mol Cell Proteomics 1, 896–903. 26. 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. 27. Krokhin, O. V., Antonovici, M., Ens, W., Wilkins, J. A., and Standing, K. G. (2006) Deamidation of -Asn-Gly- sequences during sample preparation for proteomics: Consequences for MALDI and HPLC-MALDI analysis. Anal Chem 78, 6645–50.

152 Mass Spectrometric Analysis of O-Linked Glycans Released Directly from Glycoproteins in Gels Using b-Elimination Adrian M. Taylor and Jane Thomas-Oates 1. Introduction Extensive posttranslational modification of proteins through the glycosidic attachment of carbohydrate moieties to the backbone of polypeptides makes glycosylation an important area of study. The function of these glycan chains may be to affect the conformation and stability of a protein (1), or to perform essential roles in cell surface or intracellular recognition and cell adhesion and therefore play important roles in mediating biological activity (2). Analysis of glycoproteins is made difficult by the sheer structural diversity of glycans attached to proteins, and because each glycosylated polypeptide is generally associated with a population of different glycan structures frequently attached at more than one site, possibly via more than one different type of chemical linkage. Consequently, detailed structural analysis of one or more glycans is impractical while the glycans are still attached to the polypeptide; release of the glycan is therefore required. Traditional methods used for glycan release employ both chemical and enzymatic approaches. The most commonly used chemical methods for the release of O-linked glycans are based on β-elimination in solution. Once the glycans have been released from the protein, methods for the analysis of glycoprotein glycans generally require a combination of techniques that include mass spectrometry, nuclear magnetic resonance (NMR) spectroscopy, monosaccharide compositional analysis and linkage analysis. Protein characterisation has been revolutionised by proteomics approaches, in which mixtures of proteins and glycoproteins are routinely separated using SDSPAGE. The need to identify not only the polypeptide but also its posttranslational From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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modifications has driven a whole range of method developments aimed at identifying posttranslational modifications on proteins that have been separated by gel electrophoresis. We, and others, have described blotting glycoproteins from gels onto polyvinylidene fluoride (PVDF) membranes prior to either PNGase F release of N-linked glycans (3,4) or reductive β-eliminative release of O-linked glycans (5,6), and one paper has described the success of directly performing in-gel protein de-N-glycosylation using PNGase F (7). Kilz and coworkers have described an analogous in-gel HF-pyridine de-O-glycosylation (8). However, such reagents are toxic, so that alternative in-gel protein de-O-glycosylation methods were sought. One such procedure, a modification of the nonreductive basecatalysed β-elimination method developed by Rademaker et al. (9), has been used for in-gel de-O-glycosylation/ethylaminylation (10), but was employed for glycoprotein identification and localisation of O-glycosylation sites; no attempt was made to recover the released glycans for further analysis. In this chapter we detail a method (11) for reductive β-eliminative release of the intact O-linked glycans from SDS-PAGE separated glycoproteins, and their mass spectrometric analysis. The method leaves the deglycosylated polypeptide in the gel for identification using established proteomic approaches, based on in-gel tryptic digestion and mass spectrometric analysis of the released peptides. 2. Materials 2.1. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. Resolving buffer: 0.5 M Tris-HCl, pH 6.8, 0.1% SDS. Store at room temperature. 2. Stacking buffer: 0.5 M Tris ( (hydroxymethyl) methylamine)-HCl, pH 6.8, 0.1% SDS. Store at room temperature. 3. 30% degassed acrylamide/Bis (N,N-bis-methylene-acrylamide) (30% T, 2.67% C), where for acrylamide gels: %T = {g (acrylamide + bisacrylamide)/100 mL} 100 %C = {g (bisacrylamide)/g (acrylamide + bisacrylamide)} 100 This acrylamide solution is a neurotoxin when unpolymerised and precautions should be taken to limit direct exposure. 4. TEMED (N,N,N,N-tetra-methyl-ethylenediamine), best stored at room temperature in a desiccator. 5. Ammonium persulfate: Prepare 10% solutions in water and immediately freeze in aliquots appropriate for single use at −20°C. 6. Sample Buffer: DDI H2O (3.55 mL), 0.5 M Tris-HCl, pH 6.8 (1.25 mL), glycerol (2.5 mL), 10% w/v SDS (2.0 mL), 0.5% (w/v) bromophenol blue (0.2 mL): Total volume 9.5 ml. Add 50 µL β-mercaptoethanol to 950 µL sample buffer prior to its use.

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7. 10x Electrode (Running) Buffer, pH 8.3 (1 L) Stock Solution: Tris base (30.3 g), glycine (144.0 g), SDS (10 g). Prior to use in electrophoresis, a 1:10 dilution of this stock solution is performed. 8. Standard Marker: BioRad SDS-PAGE standard control 91621 is used as a low M.W. range marker. Contains the following: Standard M.W. /Da Phosphorylase 97, 000 Serum albumin 66, 200 *Ovalbumin 45, 000 *Carbonic anhydrase 31, 000 Trypsin inhibitor 21, 000 *Lysozyme 14, 400 * glycosylated – gives positive result on Con A-Blot

2.2. Protein Staining/Destaining 1. Bio-Safe G250 Coomassie Brilliant Blue stain (Bio-Rad, USA). 2. Rapid Silver Stain based on the Amersham Biosciences Plus One Silver Stain Kit. 3. 1:1 (v/v) solution of 30 mM potassium ferricyanide and 100 mM sodium thiosulfate.

2.3. Lectin Blotting 1. 10x Transfer buffer stock solution: 1.32 M glycine (144.1 g/L), 250 mM Tris (30.3 g/L). The stock solution is diluted 1:10 prior to use with 0.5:3.5:1 ratio, stock solution:DDI H2O:methanol. 2. Concanavalin A Lectin Blot: Concanavalin A is stored at −20°C (1 mg/mL). 3. Tris-buffered saline with Tween (TBST) buffer: 20 mM Tris (2.4 g/L), 150 mM NaCl (8.7 g/L), adjust to pH 7.5 with 25% HCl, 0.1 mM MnCl2 (12.6 mg/L), 0.1 mM CaCl2 (11.1 mg/L), 0.1% Tween 20 (1 mL/L). 4. Developing solution: 7 mL 4-chloro-1-naphthol (3 mg/mL in methanol, prepare freshly), 39 mL DDI H2O, 2 mL 0.2 M Tris-HCl, pH 7.5, 20 µL 30% H2O2 (add directly before use).

2.4. In-Gel Reductive b-Elimination 1. β-elimination reagent: a freshly made aqueous solution of 0.1 M NaBH4 in 0.3 M NaOH. 2. Glacial acetic acid. 3. Heating block. 4. Vacuum centrifuge.

2.5. In-Gel Nonreductive b-Elimination in Aqueous Ammonia 1. β-elimination reagent: 25% aqueous ammonium hydroxide solution. 2. Heating block. 3. Vacuum centrifuge.

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2.6. Acid-Catalyzed Acetylation 1. Round-bottomed glass tubes (11 × 100 mm) with Teflon™-lined screw top. 2. Acetylation reagent: a freshly made mixture of trifluoroacetic acid anhydride (2 vols), glacial acetic acid (1 vol) mixed in a round glass bottomed tube, capped and allowed to cool to room temperature. 3. Vacuum centrifuge. 4. Dichloromethane.

2.7. De-O-Acetylation 1. 1:1 (v:v) conc. NH4OH/methanol. 2. Heating block.

2.8. Per-O-Methylation 1. 2. 3. 4.

99.9 % A.C.S. dimethyl sulfoxide. NaOH pellets. Pestle and mortar. Iodomethane – as a methylating reagent, this has potential to be a carcinogen, and so should be used with caution; protective clothing should be worn and, since it is volatile, it should be used in a fume cupboard. 5. Freshly made sodium thiosulfate solution, 100 mg/mL in water. 6. Dichloromethane.

3. Methods 3.1. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) The instructions that follow are based on separating bovine submaxillary gland mucins Type I-S (BSM) (Sigma, USA) using NuPAGE™ 10% Bis-Tris precast gels (1.0 mm × 10 well) (Invitrogen, USA) in an Xcell Surelock™ MiniCell. Other formats are equally appropriate. 1. Insert a precast minigel into the Xcell Surelock™ Mini-Cell and remove the preinserted comb. 2. Dissolve the standard (glyco)protein mixture in NuPAGE® LDS sample buffer (4x) and NuPAGE® reducing agent (10x) with quantities between 5 and 25 µg of protein. 3. Heat the mixture at 70°C for 10 min. 4. Prepare SDS running buffer by adding 50 mL NuPAGE® 2-morpholinethanesulfonic acid (MES) to 950 mL of HPLC grade water. Fill the lower buffer chamber of the Invitrogen Xcell Surelock™ Mini-Cell. 5. Add 500 µL NuPAGE® antioxidant solution to 200 mL 1x SDS running buffer and fill the upper buffer chamber of the Xcell Surelock™ Mini-Cell. 6. Load 20 µL of each sample into separate wells (see Note 1). Include in one well a prestained molecular weight marker.

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7. Complete the assembly of the gel unit and connect to a power supply set at 200 V for 60 min. 8. The gel unit is disconnected from the power supply and disassembled. The stacking gel is removed and discarded. Cut a small triangle about 0.5 cm in from the top right corner of the resolving gel to allow the gel’s orientation to be tracked.

3.2. Protein Staining/Destaining (see Note 2) (Glyco)proteins can be visualised by staining with Bio-Safe G250 Coomassie Brilliant Blue (Bio-Rad, USA) or by silver staining. 3.2.1. Staining with Coomassie Brilliant Blue 1. In the case of Coomassie Brilliant Blue staining, simply add enough of the commercially preprepared stain to submerge the gel, which is in an appropriate-sized tray. 2. Place the tray on a rocking platform for 30 min. 3. Pour off the stain. 4. Wash the gel with water for 30 min using a rocking platform. Remove the water and repeat the wash step. 5. After visualisation, excise individual Coomassie-stained protein bands from the gel by manually cutting out the appropriate bands, and place into separate 1500 µL microcentrifuge tubes. 6. Dehydrate the gel piece in the microcentrifuge tube using a vacuum centrifuge.

3.2.2. Silver Staining To visualise the proteins with silver staining, mass spectrometry-compatible Rapid Silver Stain based on the Amersham Biosciences PlusOne Silver Stain Kit is an appropriate stain to use. (see Note 3) 1. Prepare the fixing solution by mixing 100 mL ethanol with 25 mL concentrated acetic acid and make up to 250 mL with HPLC grade water. Add to the gel and leave for 30 min at room temperature on a rocking platform. 2. Prepare the sensitizing reagent by mixing 75 mL ethanol, 10 mL sodium thiosulfate (5% w/v) and 17 g sodium acetate and make up to 250 mL with HPLC grade water. Add to the gel and leave for 30 min at room temperature on a rocking platform. 3. Pour off the sensitizing reagent solution. 4. Wash the gel for 5 min with HPLC-grade water. Repeat twice. 5. Prepare the silver reagent by mixing 25 mL silver nitrate (2.5% w/v) with HPLCgrade water to give a final volume of 250 mL and add to the gel for 20 min. 6. Pour off. 7. Wash the gel with water for 30 min using a rocking platform. Remove the water and repeat the wash step. 8. Prepare a developing solution by mixing 6.25 g sodium carbonate with 0.05 mL formaldehyde (37% w/v) and make up to 250 mL with HPLC-grade water. Add to the gel for 5–15 min while on a rocking platform.

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9. Prepare a stopping solution by dissolving 3.65 g EDTA-Na2.2H2O in 250 mL with HPLC-grade water. Add to the gel for 10 min. 10. After visualisation, excise individual silver stained protein bands from the gel by manually cutting out the appropriate bands and place into separate 1500 µL microcentrifuge tubes. 11. Destain using 1:1 (v/v) solution of 30 mM potassium ferricyanide and 100 mM sodium thiosulfate by adding 50 µL of this solution to each of the 1500 µL microcentrifuge tubes that contain the excised protein gel bands. Perform occasional vortexing. The colour of the gel bands is monitored until the brownish colour disappears. 12. The gel pieces are subsequently washed several times with HPLC-grade water. 13. Add 100 µL 200 mM ammonium bicarbonate solution to the gel pieces for 20 min and then discard. 14. Wash the gel pieces with HPLC-grade water and then dehydrate with changes of acetonitrile (Fischer Scientific, Loughborough) until the gel pieces turn opaque white.

3.3. Lectin Blotting To identify potential glycoprotein bands, prepare a second gel of the same samples. This second SDS-PAGE-separated sample is then transferred by electroblotting the separated proteins on to a polyvinylidene fluoride membrane support activated with methanol, and incubated with Concanavalin A lectin. These directions are based on the use of an Invitrogen Xcell II™ blot module to perform the western blot, which is placed in the Invitrogen Xcell SureLock™ Mini-Cell buffer chamber. 1. Prepare a 10x transfer buffer stock solution (1.32 M glycine, 250 mM Tris) and dilute 1:10 with 0.5:3.5:1 ratio stock solution:DDI H2O:methanol prior to use. 2. Prepare a tray of transfer buffer large enough to lay out a transfer cassette with its pieces of foam and with two sheets of 3 MM paper submerged on one side. Cut a sheet of PVDF membrane so that it is larger than the size of the separating gel and lay it on the surface of a separate tray of distilled water and allow the membrane to wet via capillary action. Then submerge the membrane in the transfer buffer on top of the 3 MM paper. Lay the resolving gel on top of the PVDF membrane. 3. Wet two further sheets of 3 MM paper in the transfer buffer and carefully lay them on top of the gel, ensuring that no bubbles are trapped in the resulting sandwich. Lay the second wet foam on top and close up the cassette. 4. Insert the cassette into the buffer chamber ensuring that the PVDF membrane is located between the gel and the anode. 5. Assemble the apparatus prior to connecting to the power supply. Transfer can be achieved using 30 V constant for 1 h using a Bio-Rad Power Pac 1000. 6. Once the transfer is complete, remove the cassette from the buffer chamber and disassemble it.

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7. Prior to incubating the blot with Concanavalin A lectin, wash the PVDF membrane 3 times with TBST buffer. 8. Incubate the membrane with 100 µg lectin/20 mL TBST buffer at room temperature with constant shaking for 1 h. 9. Wash the membrane 5 times with DDI H2O. 10. Prepare the developing solution and develop the blot by incubating the membrane for 15 min in the dark. 11. Wash the blot in H2O for 1 min and then dry the blot between two pieces of Whatman No. 1 paper.

3.4. In-Gel Release Using Reductive b-Elimination of O-Glycans 1. Place the excised dehydrated gel piece (see Note 4) into a 1500 µL microcentrifuge tube and add 70 µL 0.5 M NaBH4 in 0.3 M NaOH Incubate at 50°C for 16 h (see Note 5). Quench the reaction by adding glacial acetic acid dropwise, until the evolution of hydrogen gas ceases. 2. Remove the liquid surrounding the gel piece (see Note 6) and place in a roundbottomed glass tube with a Teflon™-lined screw cap. 3. Wash the gel piece several times with 100 µL HPLC grade water with a few seconds of vortexing. 4. Pool the water washes with the original liquid from around the gel piece. Dry the combined extracts using a vacuum centrifuge.

3.5. b-Elimination in Aqueous Ammonia (see Note 7) Place the excised dehydrated gel piece into a 1500 µL microcentrifuge tube and add 70 µL 25% NH4OH and incubate at 45°C for 16 h. 3.6. Acetylation of the Dried Extracted Material (see Notes 8 and 9) 1. Acetylate the dried extracted material by adding 250–500 µL acetylation reagent to the dry sample, cap tube and mix well. 2. Allow the reaction to proceed at room temperature for 15–20 min. 3. Evaporate to dryness in a vacuum centrifuge or under a stream of nitrogen. 4. Use 1 mL of dichloromethane to redissolve the residue, cap the tube and vortex. 5. Add ~1 mL deionised water, vortex and centrifuge at low speed to separate the phases. 6. Remove the upper aqueous phase with a Pasteur pipette and discard. 7. Repeat steps 5 and 6. 8. Evaporate the dichloromethane using a vacuum centrifuge or stream of air or nitrogen.

3.7. De-O-Acetylation (see Note 9) 1. Mix dry acetylated material in a round-bottomed glass tube with a Teflon™-lined screw cap with 250–500 µL freshly made 1:1 (v:v) conc. NH4OH/methanol and incubate at 20–22°C for 18 h. 2. Dry the reaction mixture under nitrogen.

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3.8. Per-O-Methylation (see Note 9) 1. Dissolve the dry de-O-acetylated sample in 0.5 mL 99.9 % A.C.S dimethyl sulfoxide (Sigma-Aldrich, St. Louis. USA) in a round-bottomed, Teflon™-screwcapped glass tube. 2. Rapidly grind a few pellets of NaOH in a pestle and mortar and add two microspatulas (~50 mg) of the ground NaOH to the glass tube and a few drops (50–100 µL) of iodomethane using a Pasteur pipette, and recap the tube. 3. Allow to stand for 10 min at room temperature, after which add a further 5–10 drops of iodomethane to the tube. 4. Allow to stand for a further 10 min at room temperature, after which add 20 more drops of iodomethane and recap the tube. 5. After 20 min, quench the reaction by cautiously adding 1 mL freshly made 100 mg/mL sodium thiosulfate solution to the sample. 6. Immediately add 1 mL dichloromethane. Recap the tube and mix the reagents thoroughly to form an emulsion. 7. Centrifuge the sample for 10–15 s. Remove the upper aqueous layer with a pipette and then wash the dichloromethane three times with water. During each wash, form an emulsion, followed by centrifuging for 10–15 s to separate the phases and then remove the upper aqueous phase. 8. After the third washing, dry the dichloromethane under nitrogen.

3.9. Mass Spectrometry of Per-O-Methylated and Per-O-Acetylated Released O-Glycans The following conditions are based on acquiring mass spectra on an Applied Biosystems/MDS SCIEX API QSTAR Pulsar i (Ontario, Canada) quadrupole orthogonal acceleration time of flight tandem mass spectrometer. Other mass spectrometers are equally appropriate to use. 3.9.1. Electrospray 1. For electrospray applications the ion source contains a microelectrospray arm mounted in an Applied Biosystems ion source housing. 2. A 100 µL Hamilton 1710N syringe (1.46 mm inside diameter) was used with the integral syringe driver to deliver the sample, dissolved in methanol (Fischer Scientific, Loughborough), to the ion source in a continuous flow at 1 µL/ min. 3. The ion source gas reading is set to 4, the curtain gas to 20 and the capillary is held at 5500 V. 4. The instrument is operated in the positive mode with declustering and focusing potentials of 65 and 265 V respectively. 5. For Pulsar operation the Ion Release Delay is set to 6 and the Ion Release Width to 5. 6. In MS mode, the collision gas is set to read 3 and increased to 6 during tandem MS. 7. The collision energy offset for tandem MS is varied between 25 V and 70 V. Nitrogen is used for both the collision gas and for the curtain gas.

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8. The detection of ions is performed using a microchannel plate detector system with a time to digital converter. 9. The data are recorded and analysed using the Applied Biosystems/ MDS SCIEX Analyst QS Software.

The electrospray mass spectrum (Fig. 1A) of per-O-acetylated glycans released using reductive β-elimination from a bovine submaxiliary mucin SDSPAGE band, following staining with Coomassie Brilliant Blue, contains an intense (M+Na)+ ion at m/z 456 for per-O-acetylated HexNAc-ol. A less intense ion at m/z 743 is also observed for (M+Na)+ of HexNAc-HexNAc-ol. The product ion spectrum of the ion at m/z 456 (Fig. 1B) contains intense fragment ions at m/z 314 (M+H+ − 2 × 60), 336 (M+Na+ − 2 × 60) and 396 (M+Na+ − 60) and less intense ions at m/z 374 (M + H+ − 60) and 254 (M+H+ − 3 × 60), the 60 mass unit loss corresponding to the elimination of the elements of acetic acid. Other confirmatory fragment ions (m/z 110 (M+H+ − (2 × 42) − (4 × 60) ), 152 (M+H+ − 42 − (4 × 60) ), 212 (M+H+ − 42 − (3 × 60) ), 230 (M+H+ − (2 × 42) – (2 × 60) ), 254, 272 (M+H+ − 42 − (2 × 60) ), 314, 336, 374 and 396) are also present. Tandem mass spectrometric analysis of the precursor ion at m/z 743 (Fig. 1C) contains intense fragment ions at m/z 683 (M + Na+ − 60), 623 (M+Na+ − 2 × 60) and 563 (M+Na+ − 3 × 60), and two less intense fragment ions at m/z 601 (M+H + − 2 × 60) and 541 (M+H+ − 3 × 60). There are less intense fragment ions at m/z 352, 370 and 414, which correspond to diagnostic sodiated carbohydrate fragment B1, C1 and Y1 ions respectively for a per-O-acetylated HexNAc-HexNAc-ol. Mass spectrometric analysis of the glycans released from a silver-stained BSM glycoprotein gel band also resulted in the detection of ions at m/z 456 and 743 corresponding to the HexNAc-ol and HexNAc-HexNAc-ol BSM glycans. Product ion analyses of these ions yielded fragment ions indistinguishable from those produced on CID of the (M+Na)+ per-O-acetylated HexNAc-ol and HexNAcHexNAc-ol species released from the Coomassie stained gel (Fig. 1B and 1C). 3.9.2. MALDI 1. When the QSTAR is being operated with the MALDI source a standard stainless steel PerSeptive flat microtiter target plate sample introduction system is used. 2. The dried sample is dissolved in methanol (typically 5–10 µL). 3. The dried-droplet method is used for preparing the sample-matrix spots. 1 µL sample solution is mixed with 1 µL of 10 mg mL−1 α-cyano-4-hydroxycinnamic acid (in 50 % aqueous acetonitrile 0.1 % TFA solution) in a plastic, 1500 µL microcentrifuge tube, in order to produce a matrix to sample ratio of about 5000:1. An aliquot of this mixture (0.5–2.0 µL) is then applied to the stainless steel sample target and allowed to air dry.

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Intensity (%)

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m/z 414

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Fig. 1. Electrospray Mass Spectra of Per-O-Acetylated Glycans Released In-Gel using Reductive β-Elimination from an SDS-PAGE-separated Bovine Submaxillary Mucin Glycoprotein. (A) Positive ion mode electrospray mass spectrum recorded in methanol; (B) CID product ion mass spectrum of (M+Na)+ Per-O-Acetylated HexNAc-ol ion at m/z 456; (C) CID product ion mass spectrum of (M+Na)+ Per-O-Acetylated HexNAc-HexNAc-ol ion at m/z 743 (Reproduced with permission from ref. 11).

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4. Ionisation of the matrix and sample is performed using a nitrogen UV laser with a wavelength of 337 nm, 1 to 30 Hz pulsing rate and 200 µJ laser power output, although for most applications a laser power of 25 µJ is sufficient. 5. The data are recorded and analysed using the Applied Biosystems/MDS SCIEX Analyst QS Software. The laser power and plate position are controlled using the Applied Biosystems oMALDI Server 2.2 software. Instrumental and detection parameters used are similar to those described above for electrospray operation.

4. Notes 1. The smallest amount of BSM protein that could be applied to a gel yielding a band from which a BSM glycan could be detected by ESI-MS after in-gel reductive β-elimination and extraction was 5 µg (Fig. 2). 2. Not performing destaining of the Coomassie stained bands does not seem to have any adverse effect on the chemical de-O-glycosylation and detection of the released glycans. On the other hand, destaining of silver stained

314 100% 95% 90%

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212

40% 35% 30%

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456

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5% 0% 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 400 420 440 460 480 500

m/z

Fig. 2. Electrospray CID Product Ion Mass Spectrum of HexNAc-ol from an SDS-PAGE BSM Glycoprotein Band, visualised using silver staining after In-Gel De-O-Glycosylation and Per-O-Acetylation. The BSM glycoprotein band was excised from a gel lane that had 5 µg BSM applied to it. (Reproduced with permission from ref. 11).

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bands is required prior to chemical de-O-glycosylation. The removal of SDS prior to chemical de-O-glycosylation does not appear to have an effect on the chemical release or detection of O-linked glycans. Silver staining provides more sensitive staining of proteins than does Coomassie Brilliant Blue and is a very widely used staining technique. In silver stained gels, glycan release appears not to be as efficient as that obtained from a BSM glycoprotein in a band that has been visualised by Coomassie staining, since poorer signal:noise ratios for the (M+Na)+ perO-acetylated HexNAc-ol and HexNAc-HexNAc-ol ions (m/z 456 and 743) were obtained. Silver staining relies on cross-linkage in the gel, which may restrict access of the reagent to the protein, or restrict the extraction of released glycans from the gel, as well as potentially modifying the glycans. We therefore propose that this may explain the poorer signal:noise of the ions observed on in-gel release using silver stained bands. The hydration state of the gel piece prior to the addition of the β-elimination reagent has the biggest impact of all experimental variables on the effectiveness of glycan release. It is preferable to have the gel piece dried (we use a vacuum centrifuge) prior to performing the in-gel de-O-glycosylation. This is understandable, as in this way there is no dilution of the reagent. In addition, if the gel piece is dried, the aqueous reagent readily enters the gel, so accessing the protein more easily. We have noted that the use of dried bands provides an increase in the glycan release over shorter reaction times. The strength of the alkali and the reaction time have less of an impact on the extent of glycan release; the intensity of glycan ions is not increased by increasing reaction time and reagent concentration. Indeed, longer reaction times may allow the alkaline conditions to destroy the glycan, or may produce something that interferes with either the extraction of the glycans or their mass spectrometric detection. A shorter reaction time provides for better detection of the glycan ions. A compromise between losses of disaccharide and failure to release it was achieved by carrying out glycan release reactions for no longer than 16 h. Released glycans were effectively extracted from the gel pieces by vortexing in water; no improvement in the amount of glycan extracted with use of further extractions with 1:1 v:v acetonitrile/water was achieved In-gel nonreductive β-elimination release has been performed on a gel band from a Coomassie Brilliant Blue stained SDS-PAGE separation of M. avium capsular proteins. This was then followed by in-gel tryptic digestion of the residual polypeptide following glycan release. Analysis of the

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tryptic peptides using a RP capillary monolithic column coupled off-line with MALDI-ToF/ToF analysis, yielded an NCBI database hit for an M. avium protein, by matching two peptides. The significance of this modified approach is that using ammonium hydroxide to release the glycans nonreductively from the protein in-gel is compatible with subsequent in-gel tryptic digestion of the protein, enabling identification of the protein from its released peptides. It is also a convenient means of labelling the amino acid in the protein to which the glycan had been attached, thus having the potential to allow glycosylation site analysis (9). This variant of our approach could therefore be used not only to profile the glycan moieties but also to identify the protein and determine the site of glycan attachment as part of a gel-based proteomics experiment, allowing comprehensive glycoprotein characterisation. 8. The extraction into dichloromethane acts as a sample clean-up step to remove water-soluble reagents and interferents (13). 9. While per-O-acetylation is readily carried out in the presence of a range of salts and other reagents (13), the mass spectrometric response of peracetylated glycan is not as good as that of the permethylated derivative, and the mass spectrometric fragmentation tends to be less informative. However, the permethylation reaction is less tolerant of noncarbohydrate impurities, and so is best carried out on a purer sample – peracetylation and organic-aqueous partition is a convenient way to produce an appropriately purified sample. Although permethylation is carried out in strongly basic conditions, we have observed that better data were obtained if the peracetylated glycan was first de-O-acetylated prior to permethylation. 10. To demonstrate the utility of the developed in-gel de-O-glycosylation procedure, the reductive β-elimination method has been applied to a protein gel band (Fig. 3A, band X) from a Coomassie-stained SDS-PAGE separation of capsular glycoproteins from M. avium strain 2151 with smooth transparent (SmT) morphotype (11). The band chosen had produced a positive result in a duplicate Concanavalin A lectin blot, which suggested that this ~32 kDa protein was glycosylated. The MALDI mass spectrum of the glycans obtained after per-O-acetylation did not contain any ions for per-O-acetylated glycans. The mixture was de-O-acetylated and per-O-methylated in order to improve the mass spectrometric response, and analysed using ESI-MS. An ion at m/z 493 for (M+Na)+ of Hex1Hex-ol was observed. Tandem mass spectrometric analysis of the ion at m/z 493 (Fig. 3B) yielded an intense ion at m/z 275 for the diagnostic sodiated Y1 carbohydrate fragment ion for Hex-Hexitol.

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A

97 kDa 66 kDa 45 kDa

X 31 kDa

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Fig. 3. (A) Coomassie Stained SDS-PAGE Separation of Proteins Fractionated from the Capsules of Two M. avium Morphotypes, Smooth Transparent (SmT) and Smooth Opaque (SmO). (B) Positive ion mode CID tandem electrospray mass spectrum of (M+Na)+ perO-methylated Hex1Hexitol ion at m/z 493 obtained from M. avium 2151 SmT Capsular Glycoprotein After In-Gel De-O-Glycosylation and Per-O-Methylation. (Reproduced with permission from ref. 11).

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Acknowledgments We are very grateful to Prof Otto Holst and Astrid Gruhl of the Research Center Borstel, Borstel, Germany, for the productive and very enjoyable Mycobacterium avium collaboration, and to Astrid for carrying out the SDS-PAGE shown in Figure 3A and providing band X. We gratefully acknowledge AMT’s studentship provided by the Royal Society of Chemistry (RSC) and the Engineering and Physical Sciences Research Council (EPSRC), and the British Council (ARC project 1126) and the Deutscher Akademischer Austauschdienst (DAAD) (ARC project ARC-Holst) for funding the Anglo-German collaboration. JTO gratefully acknowledges funding from the Analytical Chemistry Trust Fund, the RSC Analytical Division and EPSRC.

References 1. Kenne, L., Stromberg, S. (1990) A method for the microanalysis of hexoses in glycoproteins. Carbohydr. Res., 198, 173–179. 2. Perkel, J. M. (2002) Glycobiology goes to the ball. Scientist, 16 (9), 32–36. 3. Bulone, V., Rademaker, G. J., Pergantis, S., Krogstad-Johnsen, T., SmestadPaulsen, B., Thomas-Oates, J. (2000) Characterisation of horse dander allergen glycoproteins using amino acid and glycan structure analyses. a mass spectrometric method for glycan chain analysis of glycoproteins separated by two-dimensional electrophoresis, Int. Arch. Allergy Immunol., 123, 220–227. 4. Wilson, N. L., Schulz, B. L., Karlsson, H. G., Packer, N. H. (2002) Sequential Analysis of N- and O-Linked Glycosylation of 2D-PAGE Separated Glycoproteins, J. Proteome, Res., 1, 521–529. 5. Schulz, B. L., Oxley, D., Packer, N. H., Karlsson, H. G. (2002) Identification of two highly sialylated human tear-fluid DMBT1 isoforms: the major high-molecularmass glycoproteins in human tears, Biochem. J., 366, 511–520. 6. Schulz, B. L., Packer, N. H., Karlsson, H. G. (2002) Small-Scale Analysis of O-Linked Oligosaccharides from Glycoproteins and Mucins Separated by Gel Electrophoresis, Anal. Chem., 74, 6088–6097. 7. Küster, B., Wheeler, S. F., Hunter, A. P., Dwek, R. A., Harvey, D. J. (1997) Sequencing of N-Linked Oligosaccharides Directly from Protein Gels: In-Gel Deglycosylation Followed by Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry and Normal-Phase High-Performance Liquid Chromatography, Anal. Biochem., 250, 82–101. 8. Kilz, S., Budzikiewicz, H., Waffenschmidt, S. (2002) In-gel deglycosylation of sodiumdodecyl sulfate polyacrylamide gel electrophoresis-separated glycoproteins for carbohydrate estimation by matrix-assisted laser desorption/ionization time-offlight mass spectrometry, J. Mass Spectrom., 37, 331–335. 9. Rademaker, G. J., Pergantis, S. A., Blok-Tip, L., Langridge, J. I., Kleen, A., Thomas-Oates, J. E. (1998) Mass spectrometric determination of the sites of O-glycan attachment with low picomolar sensitivity. Anal. Biochem., 257, 149–160. 10. Hanisch, F.-G., Jovanovic, M., Peter-Katalinic, J. (2001) Glycoprotein Identification and Localization of O-Glycosylation Sites by Mass Spectrometric Analysis of Deglycosylated/Alkylaminylated Peptide Fragments, Anal. Biochem., 290, 47–59.

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11. Taylor A. M., Holst O., Thomas-Oates J. (2006) Mass Spectrometric Profiling of O-Linked Glycans released directly from Glycoproteins in Gels using In-Gel Reductive β-Elimination, Proteomics, 6, 2936–2946. 12. Laemmli U. K. (1970) Cleavage of Structural Proteins during the Assembly of the Head of Bacteriophage T4, Nature, 227, 680–689. 13. Huisman M.M.H., Brüll, L.P., Thomas-Oates, J.E., Haverkamp, J., Schols, H.A., Voragen, A.G.J. (2001) The occurrence of internal (1->5)-linked arabinofuranose and arabinopyranose residues in arabinogalactan side chains from soybean pectic substances, Carbohydr. Res. 330, 103–114.

153 Glycopeptide Analysis Using LC/MS and LC/MSn Site-Specific Glycosylation Analysis of a Glycoprotein Satsuki Itoh, Daisuke Takakura, Nana Kawasaki, and Teruhide Yamaguchi 1. Introduction A glycoprotein consists of heterogeneous molecules attached to diverse oligosaccharides at multiple glycosylation sites. Glycosylation analysis at each glycosylation site is important for understanding the function of carbohydrate moiety in glycoproteins. MS of glycopeptides is a commonly used method for the analyses of glycosylation sites and site-specific glycosylations (1–4). Since glycopeptide ions sometimes fail to be acquired by MS in the presence of excess peptides due to their lower ionization efficiency, separation or enrichment of glycopeptides is crucial for the MS of glycopeptides to be performed. LC/MS that allows for separation of glycopeptides from a mixture of peptides and acquisition of their mass spectra is one of the most effective means in use for the site-specific glycosylation analysis of glycoproteins. Fig. 1 outlines the site-specific glycosylation analysis by using LC/MS and LC/MSn. Disulfide bonds in the glycoproteins are reduced and alkylated with monoiodoacetic acid to facilitate the complete digestion with proteinases (see Subheading 3.1.1). The alkylated glycoprotein is incubated with an appropriate proteinase that provides glycopeptides containing a single glycosylation site (see Subheading 3.1.2). The complex mixture of peptides is separated by reversedphase LC and subjected to on-line electrospray ionization (ESI) MS (see Subheading 3.2.1). The predominant ions acquired are automatically subjected to MSn for sequencing of carbohydrates and peptides (see Subheading 3.2.2). After acquisition of the data, the MS/MS spectra of glycopeptides are picked out by matching the all acquired MS/MS spectra for the predicted MS/MS

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Flow chart of typical site-specific glycosylation analysis by LC/MS and LC/MSn.

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spectra of the object glycoprotein by using a search engine (see Subheding 3.3.1) (5). Even if database search analysis failed to locate the MS/MS spectra of glycopeptides, tracing carbohydrate-distinctive ions, such as HexNAc+ (m/z 204) and Hex-HexNAc+ (m/z 366), acquired in MS/MS provides a clue to locate the desired spectra (see Subheding 3.3.2) (6,7). The carbohydrate structure can be deduced from the fragment pattern appearing in the extracted MS/MS spectra (see Subheding 3.3.3). The peptide-related ions, which bear reducing-end HexNAc at peptides ( (Peptide-HexNAc + H)+), often arise from the glycopeptide molecular protonated ions. The peptide sequence can be estimated by further MSn acquired from the peptide-related ions as precursors (see Subheding 3.3.4). Since glycopeptides containing identical sequences are retained at close position, preliminary site-specific carbohydrate heterogeneity can be estimated from the integrated mass spectrum acquired at the glycopeptide peaks (see Subheding 3.3.5). Tissue-plasminogen activator (t-PA) is a secreted serine protease which converts the plasminogen to plasmin, a fibrinolytic enzyme. This protease consists of 527 amino acid residues and is supposed to be digested into 51 peptides by trypsin (Fig. 2). Previous studies suggest the fucosylation at Thr61 (peptide T8), the attachment of high-mannose type oligosaccharides to Ans117 (peptide T11),

Fig. 2. Amino acid sequence of t-PA and tryptic peptides T1-T51. Boldface type indicates the peptides identified by the database search analysis with the search engine: SEQUEST (Thermo Fisher Scientific). Potential N-glycosylation sites are underlined. Thr61 is fucosylated.

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and the linkage of sialylated complex type oligosaccharides to Asn 184 and 448 (peptide T17 and T45) (8). Using t-PA as a model glycoprotein, we here illustrate the protocol for the mass spectrometric peptide/glycopeptide mapping for the site-specific glycocylation analysis of a glycoprotein. 2. Materials 2.1. Sample Preparation 2.1.1. Reduction and Carboxymethylation 1. Buffer: 0.5 M Tris-HCl (pH8.6) containing 8 M guanidine hydrochloride and 5 mM EDTA. 2. Reducing reagent: 2-mercaptothanol, dithiothreitol, and Tris(carboxyethyl)phosphin (TCEP) are used instead. 3. Alkylation reagent: sodium iodoacetate (5 mg) is suspended in 40 µl of the reduction and carboxymethylation buffer. Iodoacetamide is used instead. 4. Gel filtration column for desalting: Sephadex G-25 column (e.g., NAP-10, and PD-10 column, GE Healthcare). 5. Freeze-drying device or vacuum centrifuge (e.g., SpeedVac concentrator).

2.1.2. Proteolytic Digestion 1. Proteinase: Trypsin, Lys-C, Glu-C and Asp-N are commonly used. For the tryptic digestion, Trypsin-Gold (Promega, Madison, WI, USA) is dissolved in 50 mM acetic acid at the concentration of 1 mg/ml (see Note 1). The solution is divided into several tubes and stored at −70°C. 2. Tryptic digestion buffer: 0.1 M Tris-HCl (pH 8.0).

2.2. LC/MSn 1. LC equipment: HPLC system capable of binary gradient formation at a flow rate of 0.3–5.0 µl/ml. 2. On-line cartridge: A reversed-phase cartridge guard column (e.g., Microcolumn, C18, 0.3 mm × 5 mm, particle size 5 µm, Chemicals Evaluation and Research Institute, Japan.). 3. Column: A reversed-phase column (0.075–0.3 mm i. d. × 50–150 mm length, 3 or 5 µm particle size). A C18 or C30 column is commonly used for the peptide/glycopeptide mapping (e.g., L-column, 0.075 mm i. d. × 150 mm length, particle size 3 µm, Chemicals Evaluation and Research Institute). A graphitized carbon column (e.g., Thermo Fisher Scientific, Waltham, MA, USA) can be used for the analysis of hydrophilic glycopeptides. 4. Nanospray tip and x-y-z translational stage: Commercially available from analytical instrument manufacturers (e.g., New Objective, Inc., Woburn, MA, USA). 5. Ion trap type mass spectrometer equipped with nanoESI source (e.g., LTQ, Thermo Fisher Scientific).

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6. LC mobile phase: HPLC grade H2O, acetonitrile and formic acid. Solvent A: 0.1% formic acid in 2% acetonitrile. Solvent B: 0.1% formic acid in 90% acetonitrile.

2.3. Data Analysis 1. Search engine: e.g., TurboSEQUEST (Thermo Fisher Scientific) or Mascot (Matrixscience, London, UK). 2. Database: Swiss-Prot or NCBI-nr.

3. Methods 3.1. Sample Preparation 3.1.1. Reduction and Alkylation of a Glycoprotein 1. A glycoprotein (10 µg) is dissolved in 68 µl of reduction and carboxymethylation buffer. 2. To the solution, 2-mercaptethanol (0.5 µl) is added. The mixture is incubated at room temperature for 2 h. 3. Sodium monoiodoacetate solution (11.3 µl) is added to the sample solution, and the mixture is incubated at room temperature for 2 h in the dark. 4. The reaction mixture is applied on a Sephadex G-25 column equilibrated with water. The excess reagent is passed through the filter to remove. 5. Elution was carried out with water, and the eluant containing the alkylated glycoprotein is collected. 6. The eluant is lyophilized.

3.1.2. Proteolytic digestion 1. The alkylated glycoprotein is dissolved in 100 µl of proteolytic digestion buffer. 2. Trypsin solution (0.5 µl) is added to the sample solution. The mixture is incubated at 37°C for 16 h. 3. The mixture is stored at −20°C until just before the analysis.

3.2. LC/MSn 3.2.1. LC/MS 1. The column is equilibrated by running 98% of solvent A through it at an appropriate flow rate for 20–30 min (see Note 2). 2. The nanospray tip is attached to the outlet of the column and set in the x-y-z translational stage. The spray tip is aligned with the capillary of the mass spectrometer for accurate spray direction by moving the x-y-z translational stage. 3. The mass spectrometer (ESI voltage, electron multiplier, gas pressure, and capillary temperature, etc.) is set up and the stable spray is checked with elution buffer. 4. A sample (1 µl) is injected onto the column and both the chromatography gradient and mass spectrometer data collection are simultaneously started (see Notes 3 and 4). The column is typically eluted by a linear gradient from 2% to 45% of

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Fig. 3. Mass spectrometric peptide map. (A) Total ion chromatogram (TIC) obtained by mass scan (m/z 450–2,000) in the positive ion mode. (B) TIC obtained by mass scan (m/z 1,000–2,000). (C–E) Mass chromatograms at m/z 366, 528 and 657 obtained by MS/MS

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solvent B in 100 min. A mass scan (m/z 450–2,000) is performed for the acquisition of both peptides and glycopeptides. Since glycopeptides are relatively large, the mass scan at m/z 1,000–2,000 is effective for the specific acquisition of glycopeptides (see Note 5).

As a typical mass spectrometric peptide map, the total ion chromatogram (TIC) obtained by mass scan of the tryptic digested t-PA at m/z 450–2,000 and 1,000–2,000 are shown in Fig. 3A and 3B, respectively. 3.2.2. LC/MSn The most intense ion at each mass scan is automatically subjected to MSn as a precursor ion. Molecular related ions once used as precursor ions are not subjected to MSn within 30 s. 3.3. Data Analysis 3.3.1. Database Search Analysis for Locating the MS/MS Spectra of Glycopeptides The MS/MS spectra of glycopeptides are located by matching the acquired MS/MS spectra for the predicted MS/MS spectra from objective glycoprotein in a database using a search engine. N-glycosylated peptides are often identified by the database search analysis by using the following parameters: carboxymethylation (58 Da) at Cys, and a possible modification of HexNAc (203 Da) at Asn, and dHex (146 Da) at Ser and Thr (5). Boldface type in Fig. 2 indicates the peptides identified by the database search analysis. Nearly 85% of amino acid residues were identified (The peak assignment is illustrated in Fig. 3A). Thr61-fucosylated peptide T8 was found around 78 min. Glycosylated and deglycosylated peptide T11b were located around 62 and 64 min. Peptide T17 was identified as the deglycosylation form but not the glycosylated form. The database search analysis did not provide any clues to locate either glycosylated or deglycosylated peptide T45.

Fig. 3. (continued) Sample: Tryptic digest of carboxymethylated recombinant human t-PA (Wako Pure Chemical Industries, Osaka, Japan); Amount of injection, 0.1 µg (1 µl) LC: Column, C18 (L-column, 0.075 mm i. d. × 150 mm length, particle 3 µm, Chemicals Evaluation and Research Institute, Japan); Instrument: Paradigm (Michrom Bioresources, Inc.); cartridge, Micro column (0.3 mm × 5 mm, particle size 5 µm, Chemicals Evaluation and Research Institute); Pump A: 0.1% formic acid/2% acetonitrile; Pump B: 0.1% formic acid/ 90% acetonitrile; flow rate, 300 nl/min; gradient program, 2–45% of B in 100 min. MS: Instrument, LTQ-FT (Thermo Fisher Scientific); Tip: Fortis tip (AMR, Tokyo, Japan); Ion mode, positive.

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3.3.2. Location of glycopeptides using carbohydrate-related ions The MS/MS spectra of glycopeptides can be located by tracing the carbohydrate-related fragment ions acquired in MS/MS. The peaks appearing in mass chromatograms at m/z 366 (Hex-HexNAc+) and 657 (NeuAc-Hex-HexNAc+) afford a clue to locate the glycopeptide MS/MS spectra. Fig. 3C, 3D, and 3E show the mass chromatograms of ions at m/z 366, 528 and 657, respectively. Glycopeptides T45 and T11a, which were overlooked by the database search analysis, were located by it. 3.3.3. Structural Estimation of Carbohydrate Moiety The mass and MS/MS spectra are extracted from the acquisition positions of glycopeptides located by Subheading 3.3.1 and/or 3.3.2. The sequences of carbohydrates can be deduced from the fragment ions in the MS/MS spectra. Fig. 4A and 4B show the mass spectrum of the glycopeptide peak at 62 min and the MS/MS spectrum acquired from one of the protonated ions (m/z 1,412) as a precursor, respectively. The carbohydrate-related fragment ions at m/z 528, 690, and 852, and the peptide-related ion, (Peptide-HexNAc + 2H)2+ (m/z 1,611) in Fig. 4B suggest the attachment of a high-mannose type oligosaccharide (HexHexNAc2) to peptide T11b (theoretical mass: 3016.33) (Fig. 4A). 5 3.3.4. Sequencing of Peptide Moiety The peptide sequence can be confirmed by the b- and y-ions that arise in MS/ MS/MS acquired from the peptide-related ions (9). Fig. 4C shows the MS/MS/MS spectrum acquired from (Peptide-HexNAc + 2H)2+ (m/z 1,611). N-glycosylation at Asn117 is proven by the presence of b- and y-ion series derived from peptide T11b bearing HexNAc at Asn. 3.3.5. Heterogeneity Analysis at Each Glycosylation Site The heterogeneity of glycosylation can be estimated from the m/z values and intensities of protonated ions in the integrated mass spectrum acquired at the elution positions of glycopeptides. The carbohydrate structure is deduced from the molecular masses of individual molecules. Fig. 5A shows the integrated mass spectrum of glycopeptide T45-SA2 which is attached to disialylated bi- and tri-antennary oligosaccharides. Glycopeptides T45 were separated in order of sialylation (T45-SA0, -SA1, -SA2, and -SA3). Fig. 5B is the integrated mass spectrum of the other glycopeptides and suggests the linkage of monosialylated biantennary form to peptide T17.

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Fig. 4. MS~MS/MS/MS of glycopeptide T11b. (A) Mass spectrum acquired at 62.48 min. (B) MS/MS spectrum acquired from (M + 3H)3+ (m/z 1411.94) as a precursor ion. Carbohydrate structure was deduced from the fragment pattern. (C) MS/MS/MS spectrum acquired from (Peptide-GlcNAc + 2H)2+ (m/z 1611.3) as a precursor ion. Peptide sequence and glycosylation site were identified by the database search analysis.

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Fig. 5. Integrated mass spectra of glycopeptides derived from t-PA. (A) Integrated mass spectrum of glycopeptide T45-SA2 and structural assignment of glycopeptides. (B) Integrated mass spectrum of glycopeptide T17.

4. Notes 1. Enzyme specificity is as follows: Trypsin, Lys/Arg-↓-X (X≠Pro); Lys-C, Lys-↓-X; Glu-C, Glu (Asp)-↓-X; AspN, X-↓-Asp (Glu). Glu-C digests without cleavage of Asp-X at a 1:50 enzyme-to-substrate ratio at pH 8.0, and Asp-X can be hydrolyzed by Glu-C at a 1:4 enzyme-to-substrate ratio. 2. Typical flow rate against the column diameter is as follows: 0.3 mm, 5 µl/ min; 0.2 mm, 3 µl/min; 0.1 mm, 0.5 µl/min; 0.075 mm, 0.3 µl/min. 3. It is recommended that a test run be conducted beforehand by using a standard sample, such as tryptic digested bovine serum albumin (BSA, commercially available. e.g., Michrom BioResources, Inc., Auburn, CA, USA). Database search analysis should be performed to check coverage of the amino acid sequence in BSA. 4. Positive ion mode is used for sequencing of both carbohydrate and peptide moieties. Negative ion mode is often used for the detection of highly sialylated or sulfated glycopeptides.

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5. For the location of both N- and O-glycosylated peptides, monitoring of the oxonium ions at m/z 204 (HexNAc+) produced in MS/MS is effective (10). However, in most cases, ion trap type mass spectrometer cannot detect small fragment ions, such as HexNAc+, due to its low mass cut-off system. Hex-HexNAc+ (m/z 366) and NeuAc-Hex-HexNAc+ (m/z 657) can be used instead of HexNAc+ (m/z 204). Monitoring HexNAc+ (m/z 204) produced by in-source CID and the neutral loss of (m/z 162, 204, 291) by CID-MS/ MS can be substituted for the locating the MS/MS spectra of glycopeptides (11, 12). References 1. Demelbauer, U. M. Zehl, M. Plematl, A. Allmaier, G. and Rizzi, A. (2004) Determination of glycopeptide structures by multistage mass spectrometry with lowenergy collision-induced dissociation: comparison of electrospray ionization quadrupole ion trap and matrix-assisted laser desorption/ionization quadrupole ion trap reflectron time-of-flight approaches, Rapid Commun. Mass Spectrom. 18, 1575–1582. 2. Morelle, W. Canis, K. Chirat, F. Faid, V. and Michalski, J. C. (2006) The use of mass spectrometry for the proteomic analysis of glycosylation, Proteomics, 6, 3993–4015. 3. Wuhrer, M. Catalina, M. I. Deelder, A. M. and Hokke, C. H. (2006) Glycoproteomics based on tandem mass spectrometry of glycopeptides, J Chromatogr B Analyt. Technol. Biomed. Life Sci,.849, 115–128. 4. Wuhrer, M. Deelder, A. M. and Hokke, C. H. (2005) Protein glycosylation analysis by liquid chromatography-mass spectrometry, J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 825, 124–33. 5. Itoh, S. Kawasaki, N. Harazono, A. Hashii, N. Matsuishi, Y. Kawanishi, T. & Hayakawa, T. (2005) Characterization of a gel-separated unknown glycoprotein by liquid chromatography/multistage tandem mass spectrometry: analysis of rat brain Thy-1 separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, J. Chromatogr. A. 1094, 105–117. 6. Domon, B. and Costello, C. E. (1988) A systematic nomenclature for carbohydrate fragmentation in FAB-MS/MS spectra of glycoconjugates, Glycoconjugate J. 5, 397–409. 7. Huddleston, M. J. Bean, M. F. and Carr, S. A. (1993) Collisional fragmentation of glycopeptides by electrospray ionization LC/MS and LC/MS/MS: methods for selective detection of glycopeptides in protein digests, Anal. Chem. 65, 877–884. 8. Ling, V. Guzzetta, A. W. Canova-Davis, E. Stults, J. T. Hancock, W. S. Covey, T. R. and Shushan, B. I. (1991) Characterization of the tryptic map of recombinant DNA derived tissue plasminogen activator by high-performance liquid chromatographyelectrospray ionization mass spectrometry, Anal. Chem. 63, 2909–2915. 9. Roepstorff, P. and Fohlman, J. (1984) Proposal for a common nomenclature for sequence ions in mass spectra of peptides, Biomed Mass Spectrom. 11, 601.

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10. Harazono, A. Kawasaki, N. Kawanishi, T. and Hayakawa, T. (2005) Site-specific glycosylation analysis of human apolipoprotein B100 using LC/ESI MS/MS, Glycobiology 15, 447–462. 11. Jiang, H. Desaire, H. Butnev, V. Y. and Bousfield, G. R. (2004) Glycoprotein profiling by electrospray mass spectrometry, J. Am. Soc. Mass Spectrom. 15, 750–758. 12. Sullivan, B. Addona, T. A. and Carr, S. A. (2004) Selective detection of glycopeptides on ion trap mass spectrometers, Anal. Chem. 76, 3112–3118.

154 Identification of Vitamin K-Dependent Proteins Using a Gla-Specific Monoclonal Antibody Karin Hansson 1. Introduction γ-Carboxyglutamic acid (Gla) is unique among naturally occurring amino acids in that it possesses two carboxyl groups on its side chain. The biosynthesis of Gla occurs post-translationally from specific glutamate (Glu) residues in vitamin K-dependent proteins and involves γ-glutamyl carboxylase, a resident enzyme of the endoplasmic reticulum (1–4). Gla was first discovered in the N-terminal so-called Gla domain of certain mammalian proteins involved in the haemostatic process (5,6,7). In these proteins the Gla residues chelate calcium ions and induce a conformational transition that endows the proteins with phospholipid membrane binding properties. Since its initial discovery Gla has been identified in a wide range of vertebrates including mammals, birds, reptiles and fish. In addition, Gla has been shown to be present in neurotoxic venom peptides of the invertebrate marine snails of genus Conus (8,9). These discoveries indicate that the role of this post-translational modification in blood coagulation represents only a subset of Gla functions in animal phyla and it appears that other Glacontaining proteins with novel functions remain to be identified in the future. Several approaches have been developed for the qualitative and quantitative identification of Gla in proteins including isotopic labelling, specific stains, modified protein sequencing, mass spectrometry, ion exchange chromatography of alkaline hydrolysates, and gas or thin layer methodologies (5,10–15). However, most of these procedures are restricted as they require purified proteins or highly enriched preparations since most currently used methods for the measurement of Gla are fairly insensitive. Furthermore, Gla is particularly difficult to identify as it easily decarboxylates to Glu. Thus Gla residues are decarboxylated to Glu when heated under acid conditions, the step that normally precedes

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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determination of amino acid composition. Alkaline hydrolysis is required to identify Gla and that is rarely done as several amino acids (but not Gla) are destroyed in this procedure. Also protein/peptide sequencing using the routine automated Edman degradation procedure does not provide straightforward identification of Gla. The phenylthiohydantoin (PTH) derivative of Gla is poorly extracted and it does not separate from other anionic amino acids during HPLC. Gla can routinely be identified during sequencing but this requires modification (methyl esterfication) of the peptide prior to sequencing to ensure adequate separation of Gla from other amino acids (12,16). Consequently, erroneous sequence assignments are likely unless the presence of Gla is expected and due precautions are taken. The approach described here circumvents these problems by identifying Gla-containing proteins prior to their analysis and allowing the protein to be traced easily during purification (see Fig. 1). A Complex protein extract Possibly prefractionation

Immunoaffinity purification of Gla-proteins

Amino acid composite analysis after acid and alkaline hydrolysis

Separation of eluted Gla-candidate proteins by gel electrophoresis

Western blot analysis of Gla-candidate proteins

B In-gel proteolysis of Gla-candidate protein band(s)

Protein identification by mass spectrometry and determination of amino acid sequence Database searching based on peptide sequence tags or generation of enough amino acid sequence information for cDNA cloning

Localization of Gla-residue(s) by mass spectrometry Prior and after methyl-esterfication

Fig. 1. Flow sheet showing the route for identification of Gla in proteins. Steps employing the Gla-specific antibody for the identification of vitamin K-dependent proteins are labeled with grey boxes. (A) Steps involved in the identification of Gla-candidate protein(s). (B) Steps involved in the localization of Gla-residue(s) in Gla-candidate protein(s).

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The method depicted here is based on the development of mouse monoclonal antibodies that specifically recognize and bind Gla in proteins (17). The antibodies work well not only in Western blots but also with solution phase antigens, as has been shown by competitive immunoassays and surface plasmon resonance spectroscopy. Immobilized, the antibodies work well for affinity chromatography to purify Gla-containing proteins from complex protein extracts. The antibodies are unique and a highly functional tool that can be used to identify proteins containing one or more Gla residues (17–21). This tool in combination with modern mass spectrometry technique makes it possible to identify and localize Gla with high sensitivity in peptides and proteins. 2. Materials 2.1. Immunoaffinity Purification of Gla-Proteins 1. Mouse monoclonal antibody against Gla (17) (available from American Diagnostica Inc., CT). 2. Sulfolink coupling gel kit (Pierce Chemical Co., Rockford, IL). 3. Loading buffer: 20 mM Tris–HCl, 50 mM NaCl, 10 mM EDTA, pH 7.4 (see Note 1). 4. Column preservative: 0.05% (w/v) NaN3 in Loading buffer. 5. Dialysis membrane (Spectra/Por 3, MW cutoff 3500, Spectrum Laboratories, Inc., CA) and 0.45-µm filter. 6. Wash buffer: 20 mM Tris–HCl, 300 mM NaCl, 10 mM EDTA, pH 7.4. 7. Elution buffer: 20 mM Tris–HCl, 50 mM CaCl2, pH 7.4. 8. Protein storage buffer: 50 mM Tris–HCl, 100 mM NaCl, pH 8.0.

2.2. Separation of Eluted Gla-Candidate Proteins by Gel Electrophoresis 1. Separating buffer: 1.5 M Tris–HCl, pH 8.8. 2. Thirty percent acrylamide/Bis solution (37.5:1 with 2.6% C) (Bio-Rad Laboratories, Inc. CA) (see Note 2). 3. Twenty percent SDS. 4. N, N, N’, N’-Tetramethyl-ethylendiamine (TEMED, Merck A/S, Oslo, Norway (see Note 3). 5. Ammonium persulfate: prepare 10% solution in water. Store at +4°C. 6. Stacking buffer: 0.5 M Tris–HCl, pH 6.8. 7. Electrophoresis buffer (5X): 250 mM Tris, 1.9 M glycine, 0.5% (w/v) SDS. Store at room temperature. 8. Sample buffer: 100 mM Tris–HCl pH 8.8, 0.015% (w/v) bromphenolblue, 36% (w/v) sucrose, 3% (w/v) SDS. Store in aliquots at −20°C. 9. 1 M dithiothreitol (DTT): prepare in water and immediately freeze in aliquots at −20°C. 10. Prestained molecular weight marker: MultiMark multi-colored standard (Invitrogen, Carlsbad, CA).

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11. Silver stain reagents: fixation solution (50% (v/v) methanol, 5% (v/v) acetic acid), sensitizing solution (0.02% (w/v) sodium thiosulfate), staining solution (0.1% (v/v) silver nitrate), developing solution (2% (v/v) sodium carbonate), quenching solution (1% (v/v) acetic acid).

2.3. Western Blot Analysis of Gla-Candidate Proteins 1. Immobilone–P transfer membrane (Millipore, Bedford, MA) and 3 MM Chr chromatography filter paper (Whatman. Maidstone UK), two fiber pads. 2. Transfer buffer: 20 mM Tris, 150 mM glycine, 20% (v/v) ethanol. 3. Blocking buffer: 10% (w/v) skim milk powder in TBS. 4. Tris–buffered saline (TBS): Prepare 10X stock solution with 200 mM Tris–HCl, 1.54 M NaCl, pH 7.4. Dilute 100 mL with 900 mL water for use. 5. TBS with Tween (TBS–T): TBS supplemented with 0.2% (v/v) Tween 20. 6. Primary antibody: Mouse monoclonal antibody against Gla; M3B (17) (available from American Diagnostica Inc., CT). 7. Antibody dilution buffer: TBS–T containing 1% (w/v) fraction V bovine serum albumin (BSA). 8. Secondary antibody: Polyclonal rabbit anti-mouse IgG conjugated to alkalinephosphatase (Dako A/S, Glostrup, Denmark). 9. 50 mg/mL BCIP (5-bromo-4-chloro-3-indolyl phosphate, Sigma chemical Co., MO) in DMSO. Prepare and store in the dark at +4°C. 10. 50 mg/ml NBT (2,2-di-p-nitrophenyl-5,5-diphenyl-3,3-[3,3-dimethoxy-4,4diphenylene] ditetrazolium chloride (Nitro blue tetrazolium, Duchefa biochemie, Harleem, The Netherlands) in 50% (v/v) DMSO. Prepare and store in the dark at +4°C. 11. BCIP/NBT dilution buffer: 0.1 M Tris-HCl, 0.1 M NaCl, 5 mM MgCl2, pH 9.0. 12. Stop solution: 0.02 M Tris-HCl, 5 mM EDTA, pH 8.0.

2.4. In-Gel Proteolysis of Gla-Candidate Protein Bands 1. 2. 3. 4. 5. 6.

Gel wash solution: 50% (v/v) acetonitrile. Gel dehydration solution: 100% acetonitrile. Reducing agent solution: 10 mM dithiothreitol (DTT), 0.1 M NH4CO3 pH 7.9. Alkylating agent solution: 55 mM iodoacetic acid, 0.1 M NH4HCO3 pH 7.9. Digestion buffer: 50 mM NH4HCO3 pH 7.9. Endoproteinases: Trypsin (sequencing grade modified, Promega, Madison, WI), Asp-N (sequencing grade, Roche Diagnostics GmbH, Penzburg, Germany), Lys-C (sequencing grade, Roche Diagnostics GmbH, Penzburg, Germany). 7. Extraction solution: 5% (v/v) formic acid.

2.5. Mass Spectrometry 1. Micro-purification and desalting of peptides for mass spectrometry: ZipTipC18 columns (Millipore Corporation, MA), Wetting solution: 50% (v/v) MeOH, Equilibration solution/Sample preparation solution/Wash solution: 1% (v/v) formic acid, Elution solution: 50% (v/v) MeOH, 1% (v/v) formic acid.

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2. Quadropole-TOF hybrid mass spectrometer; we use a QSTAR pulsar-i quadropole-TOF tandem mass spectrometer (Applied Biosystems/MDS Sciex, Toronto, Canada) equipped with a nanoelectrospray ion source (MDS Protana, Odense, Denmark). 3. Nanoelectrospray capillaries (New Objectives, MA).

2.6. Methyl-Esterfication to Confirm the Presence of Gla 1. Methylating reagent: Put 3 mL of methanol in a beaker. Place the beaker in a freezer and cool to −80°C. Take the beaker from the freezer and place in an ice bath and immediately add 510 µl acetyl chloride dropwise under constant agitation (see Note 4). 2. Nitrogen gas.

2.7. In-Solution Proteolysis of Purified Gla-Candidate Protein and HPLC Peptide Isolation 1. Denaturating solution: 6 M guanidine hydrochloride, 1 M Tris-HCl, 10 mM EDTA, pH 8.6. 2. Reducing agent: 1 M dithiothreitol (DTT). 3. Alkylating agent: 134 mg/ml iodoacetic acid in 0.5 M NaOH. Prepare fresh. 4. Quench solution: β-mercaptoethanol. 5. Dialysis membrane (Spectra/Por 3, MW cutoff 3500, Spectrum Laboratories, Inc., CA). 6. Twenty percent acetic acid. 7. Digestion buffer: 100 mM NH4HCO3 pH 7.9 (trypsin), 50 mM sodium phosphate buffer pH 8.0 (Asp-N), 25 mM Tris-HCl, 1 mM EDTA pH 8.5 (Lys-C). 8. Endoproteinases: Trypsin (sequencing grade modified, Promega, Madison, WI), Asp-N (sequencing grade, Roche Diagnostics GmbH, Penzburg, Germany), Lys-C (sequencing grade, Roche Diagnostics GmbH, Penzburg, Germany). 9. Peptide separation with reversed phase HPLC: 0.1% (v/v) trifluoroacetic acid (Buffer A), 0.1% (v/v) trifluoroacetic acid, acetonitrile (Buffer B). Degas both buffers before use.

2.8. Edman Degradation of Isolated Gla-Peptide(s) 1. Automatic sequence analyzer; we use a Procise 494 automatic sequencer (ABI, Foster City, CA) with standard reagents (ABI, Foster City, CA). 2. Biobrene Plus (Applied Biosystems, Foster City, CA). 3. Micro TFA filter (Perkin Elmer, Foster City, CA). 4. Cartridge seal (Applied Biosystems, Foster City, CA). 5. 0.1% (v/v) trifluoracetic acid.

2.9. Amino Acid Composite Analysis After Acid and Alkaline Hydrolysis 1. Dialysis membrane (Spectra/Por 3, MW cutoff 3500, Spectrum Laboratories, Inc., CA). 2. 20% acetic acid.

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3. Hydrolysis tubes (10 × 100 mm, Schott Duran, Mainz, Germany). Wash tubes and incubate at 400°C overnight. 4. Alkaline hydrolysis solution: 4 M lithium hydroxide. 5. Acid hydrolysis solution: 6 M hydrochloric acid, 0.1% (w/v) phenol, 100 µM norleucine. Prepare in water and store at +4°C. 6. Alkaline internal standard: 62 µM β-carboxyglutamic acid (β-Gla). Prepare in water and store at +4°C. 7. Precipitation solution: 6.7 M phosphorous acid. 8. Sample dilution solution: Na-S high performance hydrolyzate sample dilution buffer (Beckman Coulter Inc., CA). 9. Standard solution for alkaline hydrolysis; 0.5 mM γ−carboxyglutamic acid (γ-Gla). Prepare in water and store at +4°C. 10. Standard solution for acid hydrolysis; Std amino acid standard for hydrolyzate analysis, CM-cysteine 3.4 nmol/µl, Beckman Coulter Inc., CA). Prepare in NaS and store at +4°C. 11. Automatic amino acid analyzer; we use a Beckman 6300 amino acid analyzer (Beckman Coulter, Inc., CA) and System gold software (Beckman Coulter Inc., CA). 12. High performance buffers Na-A, Na-B, Na-C, Na-D, Na-R, Na-S, Fluo-R (Beckman Coulter, Inc., CA). 13. Sodium high performance column (12 cm) (Beckman Coulter, Inc., CA).

3. Methods As described in the introduction, the key to the specific and sensitive identification of vitamin K-dependent proteins in the method presented here is the recently developed mouse monoclonal antibodies that specifically recognize Gla in proteins. This unique tool permits the Gla-residue(s) to be traced during purification and analysis. The method description is mainly divided into two parts. The first part explains the identification and purification of Gla-proteins from biological fluids and tissues using the Gla-antibody for affinity chromatography and Western blot analysis (see Subheadings 3.1 to 3.3; Fig. 1A). The second part outlines the localization of Gla residues(s) in the sequence(s) by enzymatic digestion of the protein followed by peptide sequencing by mass spectrometry and/or Edman degradation prior and after methyl esterfication (see Subheadings 3.4 to 3.9; Fig. 1B). 3.1. Immunoaffinity Purification of Gla-proteins 1. Couple the Gla-specific monoclonal antibody M3B to Sulfolink coupling gel according to the manufacturer’s instructions. These instructions assume the use of ~4 mg of antibody coupled to 2 mL of coupling gel. 2. Use the M3B-coupled resin to prepare an immunoaffinity column. Store the column in loading buffer containing 0.05% (w/v) NaN3 at +4°C. Before use equilibrate the column with loading buffer at a flow rate of 3 mL/min.

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3. Prepare the protein extract from the relevant biological fluid, tissue or organism and possibly pre-fractionate the material (see Fig. 2A). (It is beyond the scope of the present paper to discuss the protein extract preparation step in detail). 4. Dialyze the protein extract against loading buffer and filter by passage through a 0.45-µm filter. Load the filtrate onto the immunoaffinity column at a flow rate of 3 mL/min and monitor the A280 of the effluent (see Fig. 2B). After the A280 decreases to near the baseline wash/elute the column with wash buffer at 10 mL/min and elute subsequently the column with elution buffer. (Proteins containing a single Gla residue might bind loosely and elute with the wash buffer while proteins that enclose several Gla residues, for instance Gla-domain proteins elute with the elution buffer).

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Fig. 2. Purification of a Gla-protein by immunoaffinity chromatography. (A) A complex protein extract prepared from the venom of the mollusc Conus marmoreus was prefractionated on a Sephadex G-50 Superfine gel filtration column. (B) The collected fractions denoted by a horizontal bar I (pool 1) were subjected to immunoaffinity chromatography on a column of monoclonal antibody M3B-coupled resin. The peak marked with a horizontal bar II contained the purified GlaCrisp protein containing a single Gla residue.

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5. Dialyze the collected NaCl-eluted (wash-buffer) and/or CaCl2-eluted (elution buffer) fractions extensively against protein storage buffer and store at −20°C. 6. Analyze the Gla-containing collected fractions by SDS-PAGE and Western blot analysis.

3.2. Separation of Eluted Gla-Candidate Proteins by Gel Electrophoresis The protein(s) retained by affinity chromatography is subsequently analyzed by SDS-PAGE and Western blot employing the Gla-specific antibody. Prepare one gel for protein visualization and one gel for Western blot analysis. 1. This protocol is for two gels and assumes the use of a BioRad Mini-PROTEAN®3 Cell minigel system. Rinse the glass plates with distilled water and then with 95% ethanol and let them dry. 2. Prepare two 0.75-mm thick 12% gels by mixing 2.5 mL separating buffer with, 4.0 mL thirty percent acrylamide/Bis solution, 50 µL twenty percent SDS solution, 5.0 µL TEMED, 3.3 mL water, and 50 µL ammonium persulfate solution. Pour the gels, leaving space for stacking gel, and overlay immediately with water. Add the water slowly and evenly to avoid mixing. Allow the gels to polymerize for about 45 min. 3. Pour off the water and insert a piece of filter paper to dry the area in between the glass plates above the separating gels. Take care not to touch the surface of the gels. 4. Prepare the stacking gels by mixing 1.2 mL stacking buffer with, 0.6 mL thirty percent acrylamide/Bis solution, 25 µL twenty percent SDS solution, 5 µL TEMED, 3 mL water, and 25 µL ammonium persulfate solution. Pour the stacking gels until the top of the plates is reached and insert the desired combs. Allow the stacking gels to polymerize for 30 min. 5. Prepare the electrophoresis buffer by diluting 400 mL of the 5X electrophoresis buffer with 1600 mL of water in a cylinder. Cover with Para-Film and invert to mix. 6. Once the stacking gels have set, carefully remove the combs and use a 10-mL syringe fitted with a needle to wash the wells thoroughly with electrophoresis buffer. 7. Add the electrophoresis buffer to the inner chamber assembly and the lower buffer chamber. 8. Prepare the samples by mixing 20 µl (50 pmol; reduce the volume of the sample if necessary in a centrifugal vacuum concentrator) of each collected fraction with, 5 µl sample buffer, and 2 µl DTT solution (see Note 5). Incubate the samples for 5 min at 99°C. The samples are now ready for separation by SDS-PAGE. 9. Load the samples into the wells with a Hamilton syringe. Include one well for prestained molecular weight marker. 10. Run the gel at 150 V for 50 min or until the dye front is reaching the bottom of the gel. 11. Visualize the protein bands on one of the gels by silver staining (see Fig. 3A) (see Note 6). Fix the gel in fixation solution for 30 min. Rinse the gel with water (two changes, 2 min per change) and wash the gel with water for one hour on a shaking platform. Incubate the gel in sensitizing solution for 2 min with gentle shaking and rinse quickly with two changes of water (10 sec per change). Incubate the gel

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Fig. 3. Gel electrophoresis and Western blot analysis for the identification of Gla residues in proteins. (A) Protein samples from the purification steps of GlaCrisp (Fig. 2) were reduced, and resolved by 12% SDS-PAGE and silver stained. (B) The Gla-specific antibody M3B was employed to detect the Gla-containing components in the Western blot experiments. Key: lane 1, molecular weight marker; lane 2, crude C. marmoreus extract; lane 3, pool 1 from the Sephadex G-50 Superfine gel filtration column (Fig. 2A); lane 4, eluted fraction from the M3B immunoaffinity column (Fig. 2B).

in pre-chilled staining solution for 30 min at 4°C. Discard the staining solution and quickly rinse the gel with two changes of water (30 sec per change). Incubate the gel in developing solution at room temperature until adequate staining occurs. Quench staining by replacing the development solution with quenching solution. The gel can be stored in quenching buffer at +4°C.

3.3. Western Blot Analysis of Gla-Candidate Proteins Transfer the protein bands on the second gel to an Immobilone–P transfer membrane electrophoretically. 1. Prepare a sheet of transfer membrane, cut just larger than the size of the separating gel by soaking it in 95% ethanol for a few seconds. Equilibrate the membrane, twelve pieces of 3 MM chromatography filter paper cut just larger than the transfer membrane, and two fiberpads in transfer buffer for a few minutes. 2. Remove the stacking gel from the separating gel with a razor blade. 3. Prepare the transfer sandwich by placing in the following order on the bottom anode electrode a fiber pad, six pieces of 3 MM chromatography filter paper, the transfer membrane, the gel, six pieces of 3 MM chromatography filter paper, a fiber pad, and on the top the cathode electrode. Avoid trapping of air bubbles between the gel and the transfer membrane. 4. Insert the transfer sandwich into the transfer tank. Fill the tank with transfer buffer. Attach the electrodes and electrophorese at 50 V overnight. 5. Once the transfer is complete take out the transfer sandwich of the tank and carefully disassemble it. Block the membrane in blocking buffer for 2 hrs at room temperature while shaking.

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6. Remove the blocking buffer and rinse the membrane with TBS-T (four changes, 10 sec per change). 7. Incubate the membrane with 20 mL primary antibody M3B (10 µg/mL mouse monoclonal antibody against Gla diluted in antibody dilution buffer) for 2 hrs while shaking (see Note 7). 8. Wash the membrane with TBS-T (two changes with 10 sec per change, and four changes with 5 min per change). 9. Incubate the membrane with 20 mL secondary antibody (polyclonal rabbit antimouse IgG diluted 1:1000 in antibody dilution buffer) for 1 hr while shaking. 10. Wash the membrane with TBS-T (two changes with 10 sec per change, three changes with 5 min per change, and once for 10 min). 11. Incubate the membrane in 15 mL of BCIP/NBT substrate (prepare by mixing 100 µL 50 mg/mL BCIP solution with 50 µL 50 mg/ml NBT solution in BCIP/NBT dilution buffer) until adequate staining occurs (see Fig. 3B). 12. Stop development by replacing the BCIP/NBT substrate with stop solution.

3.4. In-Gel Proteolysis of Gla-Candidate Protein Bands Wear gloves to reduce contamination with human keratins when preparing solutions and when handling samples. The described in-gel enzymatic digestion protocol is slightly modified from that described by Shevchenko et al. (22). 1. Rinse the silver stained gel with water two times for 10 min each. 2. Excise protein bands corresponding to immunoreactive protein bands in the Western blot analysis with a stainless steel cutting device. Cut as close to the edge of the band as possible to reduce the amount of background gel. Slice the gel band into small pieces (1 mm × 1 mm) and transfer them to a clean microcentrifuge tube. 3. Wash the gel with water (two times, 15 min each) and with gel wash solution (two times, 15 min each). The liquid volumes should be about five times the gel volume. 4. Remove the liquid and add 20 µL of gel dehydration solution. Incubate for 5 min until the gel pieces shrink and become white and stick together. Remove the gel dehydration solution. 5. Swell the gel pieces in 50 µL reducing agent solution and incubate for 1 h at 56°C to reduce disulfide bonds. 6. Cool the microcentrifuge tube to room temperature and remove the excess of reducing agent solution. Add quickly 50 µL alkylating agent solution and incubate for 30 min at room temperature in the dark. 7. Remove the alkylating agent solution and wash the gelpieces as described in step 3. Remove the liquid and add 20 µL of gel dehydration solution. 8. Rehydrate the gel pieces in enough digestion buffer containing 12.5 ng/µL trypsin to cover the gel pieces. Incubate on ice for 45 min. Add more digestion buffer if all liquid has been absorbed by the gel pieces. This protocol also applies to in-gel digestions with other commonly used endoproteinases such as Asp-N and Lys-C. 9. Remove the remaining liquid after 45 min and replace it with 20–25 µL of digestion buffer and incubate at 37°C overnight.

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10. After overnight digestion spin the microcentrifuge tube shortly and extract the peptides from the gel pieces by adding 15 µL extraction solution and incubate for 15 min. Add an equal volume of acetonitrile and incubate for 15 min. Collect the supernatant. Repeat the extraction once and pool the extracts. Split the extract into two micocentrifuge tubes. 11. Dry down the pooled extracts using a vacuum centrifuge.

3.5. Protein Identification of In-Gel Digested Protein Bands by Mass Spectrometry Localization of the Gla-residue(s) in the protein requires total coverage of the amino acid sequence followed by examination of the peptide masses, derived after enzymatic digestion, to target peptides that are carboxylated at their Glu residues. Mass spectrometric analysis is disturbed by the presence of salts and the sample must therefore be desalted. (It is beyond the scope of the present paper to discuss the identification of proteins by mass spectrometry analysis in detail). Analyze the extracted peptides by nano electrospray ionization (ESI) tandem mass spectrometry to obtain the entire amino acid sequence of the candidate Gla-protein. 1. Use a micro-purification and desalting microcolumn prior to the analysis of one half of the recovered peptides. Re-dissolve the dried peptides in 10 µL of sample preparation solution and desalt and concentrate the sample using a ZipTipC18-column according to the manufacturers instructions. Elute the peptides in 1–5 µL elution solution. 2. Analyze the peptides by nano ESI tandem mass spectrometry where the peptides are first analyzed by full-scan MS and then by MS/MS. Program the software of the mass spectrometer to perform one scan to determine the peptide masses and then to sequence the two to four most abundant peptides. 3. Scan the spectra against a protein sequence database using a search algorithm such as Mascot or Sequest, which assigns peptide identifications based on matched criteria. If the protein is present in a protein sequence database and is identified the peptide assignments should be manually verified. If the protein proves to be unknown (i.e., not present in a sequence database) use the amino acid sequence tags obtained by de novo sequencing for subsequent construction of oligonucleotides to predict the complete amino acid sequence of the protein by cloning and cDNA sequencing. If necessary, sequence additional peptides manually by nano ESI MS/MS to generate enough sequence tags.

3.6. Localization of Gla-Residue(s) by Mass Spectrometry 1. Use the obtained amino acid sequence of the Gla-candidate protein to calculate the theoretical masses of the predicted peptides generated by in-gel digestion with the endoproteinase used. 2. Nano ESI mass spectrometry normally provides multiply charged peptide ions. Determine the mass of the peptides in the MS experiment described in Subheading 3.5., by correcting the mass-to-charge values by the charge and the mass of added protons providing the charge.

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3. Compare the observed peptide masses deduced from the ESI mass spectrometry analysis with the predicted theoretical peptide masses and target peptides that are carboxylated at their Glu residues. These peptides have a mass increase of 44 Da for each Gla residue as compared to the mass of the predicted Glu residue (see Fig. 4A). The ion signals in the MS spectrum corresponding to the Gla-containing peptides are typically accompanied by signals corresponding to loss of 44 Da caused by ejection of the γ-CO2 by gas-phase decarboxylation or by loss of the γ-CO2 during isolation and purification. To localize the peptide(s) containing Gla, it might be necessary to perform mass spectrometry analysis of peptides generated by endoproteinase in-gel digestions using two or more different enzymes, for example trypsin, Asp-N, and Lys-C. 4. Perform MS/MS fragmentation of the doubly and/or triply charged peptide ions corresponding to the Gla-peptide(s) (if this was not already performed in the MS/ MS experiments described under Subheading 3.5.) to assign the position of the Gla residue(s) in the Gla-peptide(s). 5. Interpret the MS/MS fragment ion spectra and deduce the corresponding peptide sequences from the y and b ions. The peptidyl mass of Gla is 173 Da (see Fig. 5).

3.7. Methyl-Esterfication to Confirm the Presence of Gla Confirm the presence of Gla-residue(s) in the protein by chemical derivatization of carboxyl groups in the peptides recovered from the in-gel digestion. 1. Add 50 µl of the methylating reagent to the other half of the dried peptides recovered after the in-gel digestion of the protein (see Subheading 3.4.). 2. Incubate for 2 hrs under nitrogen at room temperature. 3. Dry down the reaction mixture using a vacuum centrifuge. 4. Micropurify the derivatized peptides on a ZipTipC18 column as described in Subheading 3.5. and analyze the peptides by nano ESI mass spectrometry. Acquire spectra in both MS and MS/MS mode and correlate peptide molecular ions corresponding to the Gla-peptide(s) in the unmodified (see Subheading 3.5. and 3.6.) and derivatized digests. Methanolic HCl adds one methyl group of 14 Da to Asp, Glu, S-carboxymethylated Cys, and to the C-terminus of the peptide and two methyl groups to Gla (see Fig. 4B).

3.8. In-Solution Proteolysis of Purified Gla-Candidate Protein and HPLC Peptide Isolation If a pure Gla-protein(s) in sufficient amounts is retained by affinity chromatography (see Subheading 3.1.) it can be enzymatically digested and the generated peptides fractionated by reversed-phase HPLC to isolate the Gla-containing peptide(s). 1. Dissolve the protein (1 nmol) in 500 µl of denaturating solution in a microcentrifuge tube. 2. Add 10 µl of reducing agent to a final concentration of 20 mM and incubate at 37°C for 2 hrs (see Note 8).

Identification of Vitamin K-Dependent Proteins A

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YSgVTPTHTMCLTDNANAVAVTLTQEVK GlaCrisp (M+3H) 1046.79

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Fig. 4. Positive ion nano-ESI mass spectrometry analysis of a Gla-containing tryptic peptide fragment prior and after methyl-esterfication. The nanoESI-MS spectra of the S-carboxymethylated tryptic peptide fragment covering amino acid residues 7–34 of GlaCrisp prior (A) and after (B) methyl-esterfication. The ions at m/z 785.41 (4+), 1046.79 (3+), 806.41 (4+) and 1074.83 (3+) are accompanied by losses of 44 Da (corresponding to the mass of CO2), which is consistent with a Gla residue in the peptide. After methyl esterfication an 84 Da increase in the monoisotopic molecular mass (corresponding to the methylation of all six carboxyl groups including the two at the Gla residue and the one at the C-terminus) was observed.

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Fig. 5. Positive ion nanoESI-MS/MS analysis of the Gla-containing peptide fragment derived from digestions of GlaCrisp with trypsin and Asp-N. (A) The b and y ions according to the proposed sequence of the S-carboxymethylated internal peptide that covers amino acids 7–19 of GlaCrisp are labeled. (B) The m/z region from 230 to 630 has been expanded to show the b2, b3, b4, and b5 ions from which the N-terminal sequence of the peptide could be determined to localize the Gla residue.

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3. Cool the microcentrifuge tube to room temperature and add 12.5 µl alkylating agent to a final concentration of 50 mM. Incubate in the dark at room temperature for 30 min. 4. Add 5 µl of quench solution to a final concentration of 1% (v/v). 5. Dialyze extensively against twenty percent acetic acid. 6. Dry down the reduced and alkylated protein using a vacuum centrifuge (see Note 9). 7. Add the endoproteinase in digestion buffer and incubate at 37°C for 18 hrs (see Note 10). 8. Dry down the digestion using a vacuum centrifuge. 9. Re-dissolve the dried peptides in 100 µl of buffer A and centrifuge for 1 min in a microcentrifuge at 10 000 rpm. 10. Remove the supernatant and inject onto an HPLC system equipped with a reversedphase column (2.1 mm i.d.). After washing with buffer A elute the peptides using a linear gradient of acetonitrile in 0.1% (v/v) trifluoroacetic acid at a flow rate of 150 µl/min. Monitor the A214 and A280 of the effluent. 11. Analyze a small portion of the collected fractions of the HPLC separation by nano ESI tandem mass spectrometry, first by full-scan MS and then by MS/MS, to identify the fraction(s) containing the Gla-peptide(s) of the protein.

3.9. Edman Degradation of Isolated Gla-Peptide(s) Verify the Gla residue(s) in the peptide sequence by Edman degradation. The current method for automated sequence analysis of Gla residues involves esterfication of the sample with methanolic HCl prior to sequencing. 1. Dry down two portions (100 pmol each) of the HPLC-isolated Gla-peptide(s) in microcentrifuge tubes using a vacuum centrifuge. 2. Perform methyl-esterfication as described in steps 1–3 in Subheading 3.7. of one of the portions of dried Gla-peptide(s). 3. Dry down the methyl-esterfied peptides using a vacuum centrifuge. 4. Re-dissolve the methylated and non-methylated portions of peptides in 0.1% trifluoracetic acid and load onto Biobrene Plus-pretreated micro TFA filters. 5. Analyze the Gla-peptide(s) both prior and after modification with methanolic HCl on an automatic sequence analyzer using the methods supplied by the manufacturer. 6. Evaluate the chromatograms and determine the amino acid sequence of the Glapeptide(s). Compare the chromatograms of the peptide prior and after methylation. Methylation of the PTH derivatives of Glu and Asp causes a shift in the chromatographic elution time. The dimethyl ester of PTH Gla elutes between PTH proline and PTH methionine (see Fig. 6).

3.10. Amino Acid Composite Analysis After Acid and Alkaline Hydrolysis Determine the amount of Gla in the isolated Gla-protein by performing amino acid analysis after acid and alkaline hydrolysis, respectively. Gla is extremely acid labile and is completely converted to glutamic acid during acid hydrolysis

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Fig. 6. C18 reversed phase HPLC chromatography of PTH amino acids. Separation of twenty-five picomoles of the standard mixture of PTH amino acids prior (A) and after (B) methyl-esterfication with methanolic HCl. The amino acid residues are designated by a one-letter code. γ denotes gamma-carboxyglutamic acid. DPTU and PMTC are by-products of the sequencing reaction and serve as useful reference peaks in the chromatogram. The arrows denote the PTH derivatives of Gla, Glu and Asp that show a shift in the chromatographic elution time caused by methylation. The absence of the Gla-derivative in the chromatogram prior to methyl esterfication is due to the low recovery from the extraction step prior to separation and inadequate separation from the other amino acid derivatives.

normally used for amino acid composite analysis. Alkaline hydrolysis provides quantitative release of Gla from peptide linkage without significant destruction. 1. Dialyze the purified protein (1 nmol as judged from A280) suspected of containing Gla extensively against twenty percent acetic acid. 2. Divide the dialyzed protein into two hydrolysis tubes and dry down the samples using a vacuum centrifuge. 3. Add 100 µL of alkaline hydrolysis solution to one hydrolysis tube for alkaline hydrolysis. Add 100 µL of acid hydrolysis solution to the other hydrolysis tube for acid hydrolysis.

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4. 5. 6. 7. 8.

Seal the hydrolysis tubes under vacuum with a torch. Heat the sealed tubes at 110°C for 24 hours. Etch and crack the tubes at the top after hydrolysis. Dry down the samples in a vacuum centrifuge. For the alkaline hydrolysate: Add 100 µL alkaline internal standard and mix on a vortex for 15 sec. Add 20 µL precipitation solution and mix the sample on a vortex until a white precipitate is formed. Centrifuge the sample for 15 min at 10000 rpm and collect the clear supernatant. For the acid hydrolysate: Add 100 µL sample dilution solution. 9. For the alkaline hydrolysate: Mix 25 µL of the recovered supernatant with 35 µL of sample dilution solution and transfer to sample coil. For the acid hydrolysate: Mix 35 µL of the sample with 35 µL of sample dilution solution and transfer to a sample coil. 10. Analyze the samples on an automatic amino acid analyzer using the methods supplied by the manufacturer. 11. Evaluate the chromatograms and determine the amount of Gla in the protein.

4. Notes 1. Unless stated otherwise, all solutions should be prepared in water that has a resistivity of 18.2 MΩ-cm. This standard is referred to as water in the text. 2. Caution! Acrylamide and Bis are toxic in the monomeric form. Wear gloves to avoid skin contact. 3. TEMED is best stored at +4°C in a desiccator. 4. Caution! Put on safety goggles and gloves. The mixture may boil up instantly. 5. For many Gla-proteins we have noticed that reduction of disulphide bonds generates more intense Western blot signals employing the Gla-specific antibody. This is probably due to increased exposure of Gla residues to the environment. 6. This silver-staining protocol in which glutardialdehyde is omitted is compatible with mass spectrometric analysis (22). 7. The primary antibody can be re-used for up to ten times only adjustment being increased length of incubation at the developing step with the BCIP/ NBT substrate. 8. Less DTT can be used, but a 20- to 100-fold molar excess over protein thiols is advised. 9. Drying should be performed in the same microcentrifuge tube that will be used for the enzymatic digestion. 10. The recommended amount of enzyme is for trypsin and Lys-C 1:100 and for Asp-N 1:200 of the protein by weight.

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Acknowledgments The author would like to thank Prof. Johan Stenflo for critically reading the manuscript. This work was supported by Grant K2001-03X-04487-27A from the Swedish Medical Research Council, the Swedish Foundation of Strategic Research, the Kock Foundations, and the Foundation of University Hospital, Malmö.

References 1. Stenflo, J. and Suttie, J. W. (1977) Vitamin K-dependent formation of gammacarboxyglutamic acid. Annu. Rev. Biochem. 46, 157–172. 2. Suttie, J. W. (1985) Vitamin K-dependent carboxylase. Annu. Rev. Biochem. 54, 459–477. 3. Furie, B., Bouchard, B. A., and Furie, B. C. (1999) Vitamin K-dependent biosynthesis of gamma-carboxyglutamic acid. Blood 93, 1798–1808. 4. Stenflo, J. (1999) Contributions of Gla and EGF-like domains to the function of vitamin K-dependent coagulation factors. Crit. Rev. Eukaryot. Gene Expr. 9, 59–88. 5. Stenflo, J., Fernlund, P., Egan, W., and Roepstorff, P. (1974) Vitamin K dependent modifications of glutamic acid residues in prothrombin. Proc. Natl. Acad. Sci. U S A 71, 2730–2733. 6. Magnusson, S., Sottrup-Jensen, L., Petersen, T. E., Morris, H. R., and Dell, A. (1974) Primary structure of the vitamin K-dependent part of prothrombin. FEBS Lett. 44, 189–193. 7. Nelsestuen, G. L., Zytkovicz, T. H., and Howard, J. B. (1974) The mode of action of vitamin K. Identification of gamma-carboxyglutamic acid as a component of prothrombin. J. Biol. Chem. 249, 6347–6350. 8. McIntosh, J. M., Olivera, B. M., Cruz, L. J., and Gray, W. R. (1984) Gammacarboxyglutamate in a neuroactive toxin. J. Biol. Chem. 259, 14343–14346. 9. Olivera, B. M., Rivier, J., Clark, C., Ramilo, C. A., Corpuz, G. P., Abogadie, F. C., Mena, E. E., Woodward, S. R., Hillyard, D. R., and Cruz, L. J. (1990) Diversity of Conus neuropeptides. Science 249, 257–263. 10. Hauschka, P. V. (1979) Specific tritium labeling of gamma-carboxyglutamic acid in proteins. Biochemistry 18, 4992–4999. 11. Kuwada, M. and Katayama, K. (1983) An improved method for the determination of gamma-carboxyglutamic acid in proteins, bone, and urine. Anal. Biochem. 131, 173–179. 12. Cairns, J. R., Williamson, M. K., and Price, P. A. (1991) Direct identification of gamma-carboxyglutamic acid in the sequencing of vitamin K-dependent proteins. Anal. Biochem. 199, 93–97. 13. Jie, K. S., Gijsbers, B. L., and Vermeer, C. (1995) A specific colorimetric staining method for gamma-carboxyglutamic acid-containing proteins in polyacrylamide gels. Anal. Biochem. 224, 163–165. 14. Kelleher, N. L., Zubarev, R. A., Bush, K., Furie, B., Furie, B. C., McLafferty, F. W., and Walsh, C. T. (1999) Localization of labile posttranslational modifications by electron capture dissociation: the case of gamma-carboxyglutamic acid. Anal. Chem. 71, 4250–4253.

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15. Vo, H. C., Britz-Mckibbin, P., Chen, D. D., and MacGillivray, R. T. (1999) Undercarboxylation of recombinant prothrombin revealed by analysis of gammacarboxyglutamic acid using capillary electrophoresis and laser-induced fluorescence. FEBS Lett. 445, 256–260. 16. Fernlund, P., Stenflo, J., Roepstorff, P., and Thomsen, J. (1975) Vitamin K and the biosynthesis of prothrombin. V. Gamma-carboxyglutamic acids, the vitamin K-dependent structures in prothrombin. J. Biol. Chem. 250, 6125–6133. 17. Brown, M. A., Stenberg, L. M., Persson, U., and Stenflo, J. (2000) Identification and purification of vitamin K-dependent proteins and peptides with monoclonal antibodies specific for gamma-carboxyglutamyl (Gla) residues. J. Biol. Chem. 275, 19795–19802. 18. Brown, M. A., Hambe, B., Furie, B., Furie, B. C., Stenflo, J., and Stenberg, L. M. (2002) Detection of vitamin K-dependent proteins in venoms with a monoclonal antibody specific for gamma-carboxyglutamic acid. Toxicon. 40, 447–453. 19. Stenberg, L. M., Brown, M. A., Nilsson, E., Ljungberg, O., and Stenflo J. (2001) A functional prothrombin gene product is synthesized by human kidney cells. Biochem. Biophys. Res. Commun. 280, 1036–1041. 20. Stenberg, L. M., Nilsson, E., Ljungberg, O., Stenflo, J., and Brown, M. A. (2001) Synthesis of gamma-carboxylated polypeptides by alpha-cells of the pancreatic islets. Biochem. Biophys. Res. Commun. 283, 454–459. 21. Hansson, K., Thämlitz, A. M., Furie, B., Furie, B. C., and Stenflo, J. (2006) A single gamma-carboxyglutamic acid residue in a novel cysteine-rich secretory protein without propeptide. Biochemistry 45, 12828–12839. 22. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. (1996) Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68, 850–858.

155 The Identification of Protein S-Nitrosocysteine Todd M. Greco, Sheryl L. Stamer, Daniel C. Liebler, and Harry Ischiropoulos 1. Introduction Nitric oxide (NO•) bioactivity regulates cellular function in most major mammalian organ systems. NO signaling either through soluble guanylate cyclase activation and cGMP production, or S-nitrosylation, the modification of cysteine residues to form S-nitrosocysteine, are currently the two most well studied nitric oxide signaling pathways. The emergence of S-nitrosylation as an intricate participant in cellular signaling is evidenced by the identification of endogenously S-nitrosylated proteins, including the NMDA receptor (1), caspase-3 (2), GAPDH (3), and NF-κB (4), which all participate in signal transduction pathways that regulate the fine balance between cell survival and cell death. The identification of novel S-nitrosylated proteins has traditionally stemmed from the observation that nitric oxide could modulate a protein’s function, e.g. its enzymatic activity, and that these effects were often recapitulated when thiol reactive compounds such N-ethylmaleimide (NEM) or organomercury derivatives such as p-choloromercuribenzoic acid (PCMB) were used. Moreover, if the observed effects of nitric oxide could be reversed by reducantants such as dithiothreitol (DTT), cysteine modification was suspected. Researchers would then undertake the task of mutational analysis to identify which critical cysteine residue(s) was responsible for the observed effects. While these methods are still critically important for establishing the functional role of S-nitrosylation, the development of proteomic techniques, namely the biotin switch method has accelerated the pace with which endogenously and potentially S-nitrosylated proteins have been identified. Here we describe an extension of the original biotin switch method (3) that incorporates peptide affinity capture, tandem mass spectrometry (MS/MS), and peptide sequence mapping,

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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permitting both the S-nitrosylated protein and the corresponding S-nitrosocysteine residue to be identified in a single experiment (5,6). 2. Materials 2.1. Cell Culture and Lysis 1. Dulbecco’s Modified Eagle’s Medium (DMEM, Gibco/Invitrogen, Carlsbad, CA) supplemented with 10% fetal bovine serum (FBS, Hyclone, Logan, UT). 2. Hydrochloric acid (0.5 M, HCl) containing 0.1 mM DTPA. Store at room temperature. 3. L-cysteine (1 M): Prepared fresh in 0.5 N hydrochloric acid/0.1 mM DTPA. 4. Sodium nitrite (1 M): Prepared fresh in water (see Note 1). 5. S-nitrosocysteine stock solution (CysNO, ~0.5 M): Prepared by mixing equimolar concentrations of L-cysteine solution (1 M) and sodium nitrite solution (1 M). Vortex immediately and incubate on ice for 5 min. Neutralize with 1 N NaOH to pH 7.2–7.4, immediately aliquot (20 uL) and store at −80°C. Aliquots are single use only. The concentration of each aliquot should be determined before use (see Note 2). 6. D-PBS-MC: Dulbecco’s Phosphate Buffered Saline (D-PBS, Gibco) supplemented with 1 mM diethylenetriaminepentaacetic acid (DTPA) and 0.1 mM neocuproine. 7. TE solution: trypsin (0.05%) and ethylenediamine tetraacetic acid (EDTA) (0.02%) (Gibco). 8. Isolation buffer: 250 mM HEPES-NaOH, pH 7.7, containing 1 mM DTPA, 0.1 mM neocuproine. Store at room temperature. Prepare fresh every 2–3 months. 9. Methyl methanethiosulfonate (MMTS, 10 M) (Sigma, St. Louis, MO). 10. Lysis buffer: Prepared fresh from isolation buffer adjusted to contain 1% Triton X-100 and 1 mM MMTS (see Note 3).

2.2. Biotin Switch Assay: Derivitization of Protein S-nitrosocysteine (Pr-SNO) 1. SDS stock solution: Prepared at 25% (12.5 g in 50 mL final volume of water). Store at room temperature. CAUTION: Respiratory irritant, wear appropriate protective equipment. 2. Acetone (Fisher Scientific, Waltham, MA). Store at −20°C. CAUTION: Flammable. 3. Resuspension buffer: Prepared fresh from 1 part isolation buffer, 8.6 parts water, and 0.4 parts SDS stock solution. 4. Blocking buffer: Prepared fresh from isolation buffer adjusted to 2.5% SDS and 20 mM MMTS. 5. Biotin-HPDP (Pierce, Rockland, IL): Stock solution prepared at 20 mM in dimethylformamide (DMF). Aliquot and store at −20°C. 6. Labeling buffer: Adjust 25 mM HEPES-NaOH, pH 7.7, containing 1 mM DTPA amd 0.1 mM neocuproine (prepared as a stock) to contain 0.2 mM biotin-HPDP and 1% SDS immediately before use. 7. Sodium ascorbate solution: Prepared fresh at 0.5 M in water. Store on ice and protect from light.

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8. Bicarbonate buffer: Freshly prepared at 100 mM ammonium bicarbonate containing 0.1% SDS. 9. Bradford reagent (Bio-rad) and BCA protein assay (Pierce).

2.3. SDS-PAGE and Western Blotting for Biotinylated Proteins 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.

MOPS SDS running buffer (20X, Invitrogen) NuPAGE Bis-Tris 4–2% mini gels (Invitrogen) (see Note 4). XCell SureLock™ Mini-Cell (Invitrogen). SeeBlue molecular weight marker (Invitrogen). Nonreducing loading buffer (6X): 375 mM Tris-HCl, pH 6.8, 7.5% SDS, 50% glycerol, and 0.1% bromophenol blue (w/v). Base buffer (10X): 0.25 M Tris base (do not adjust pH), 1.9 M glycine. Transfer buffer (1X): Mix 700 mL water, 200 mL methanol, and 100 mL 10X base buffer. Mini Trans-Blot cell (Biorad, Hercules, CA). Immobilon-FL (PVDF) transfer membrane (Millipore) and chromatography paper (3 MM Chr, Whatman, Maidstone, UK). Tris-buffered saline (TBS, 10X): 0.5 M Tris-HCl, pH 7.4, and 1.5 M NaCl. Tween 20 (Biorad). TBS-T wash buffer: Dilute 100 mL of 10X TBS with 900 mL water and adjust to 0.05% Tween 20. Membrane blocking buffer: 5% (w/v) nonfat dry milk in 1X TBS. Mouse antibiotin antibody (200 µg/mL, Spring Bioscience, Fremont, CA). Primary antibody solution: Membrane blocking buffer adjusted to 0.05% Tween 20 and 0.1 ug/mL mouse antibiotin antibody. Odyssey Infrared Imaging System (Licor Biosciences, Lincoln, NE) (see Note 5). Alexa Fluor 680 antimouse IgG secondary antibody (2 mg/mL, Molecular Probes). Secondary antibody solution: 1 part membrane blocking buffer, 4 parts TBS-t; adjust to 0.2 µg/mL secondary antibody. Protect from light.

2.4. Protein Digestion and Peptide Capture 1. Microcon YM-10 (10 kDa NMWL) filters (Millipore). 2. Trypsin gold, mass spectrometry grade (Promega, Madison, WI). Resuspend at 1 mg/mL in 50 mM acetic acid. Store at −20°C. Use within 5 freeze-thaw cycles. 3. Streptavidin-agarose suspension (Pierce). 4. Bicarbonate wash buffer: Prepared fresh at 1 M ammonium bicarbonate in water. 5. Formic acid, mass spectrometry grade (Sigma). 6. SpeedVac system.

2.5. Peptide Identification by Liquid Chromatography-Tandem Mass Spectrometry (LC-MS/MS) 1. HPLC-grade water. 2. Acetonitrile, HPLC grade.

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3. Thermo LTQ two-dimensional ion trap mass spectrometer equipped with a micro electrospray source and a Thermo Surveyor pump and autosampler (Thermo Scientific, San Jose, CA). 4. Capillary LC column (100 µm I.D. × 110 mm) packed with Jupiter C18 resin (5 µm, Phenomenex, Torrance, CA). 5. HPLC solvent A: 0.1% formic acid in HPLC-grade water. 6. HPLC solvent B: 0.1% formic acid in HPLC-grade acetonitrile. 7. Data analysis software: SEQUEST (Bioworks software, Thermo Scientific) and Scaffold (Proteome Software, Portland, OR) (see Note 6).

3. Methods Protein S-nitrosocysteine (S-nitrosothiols or Pr-SNOs) are labile posttranslational modifications, which are sensitive to reduction by ultraviolet (UV) light, transition metals, and reducing agents such as beta-mercaptoethanol or dithiothreitol. As a result, identification of Pr-SNOs are often performed by the biotin switch method, which blocks existing free thiols, then selectively reduces Pr-SNOs to the corresponding thiols using ascorbate while concurrently labeling the newly generated thiols with a biotin-containing dithiopyridine derivative (3). The biotin moiety allows selective enrichment and identification of proteins that previously contained S-nitrosocysteine residues. As an extension of this method, the protocol described here uses affinity enrichment of peptides rather than proteins, taking advantage of liquid chromatography tandem mass spectrometry (LC-MS/MS) to identify both the site of S-nitrosylation as well as the protein (Fig. 1). The detection of Pr-SNO is described for cell culture (Subheading 3.1); however, the method can be used for the identification of Pr-SNO from tissues (5,7). Importantly, rapid isolation and processing of starting material should always be performed under reduced lighting conditions in buffers containing metal chelators and in the absence of reducing compounds. This method enables the identification of S-nitrosocysteine-containing peptides starting from 1 mg of protein that contains between 0.1–.5 nmol Pr-SNO/mg of protein. These levels can be generated by exposing cells to NO donor compounds such as the family of NONOates (6,8,9). In contrast, for the identification of endogenous Pr-SNO, between 10–50 mg of initial protein is recommended (see Note 7). Also of critical importance is the inclusion of appropriate negative controls to confirm that the sites of S-nitrosylation were derived from authentic S-nitrosocysteine residues. For example, authentic Pr-SNO sites should not be identified from control samples that have been exposed to UV light (Subheading 3.2, step 2) or from control samples in which ascorbate has been omitted (Subheading 3.2, step 7). Although not discussed in detail, the suppression of nitric oxide production by pharmacological NOS enzyme inhibitors, such as L-NAME, or by genetic NOS knockout mice models can be used as negative controls. As an

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Fig. 1. Scheme of the biotin switch method, modified for the site-specific identification of Pr-SNO residues by peptide affinity capture and tandem mass spectrometry. Depicted are multiple cysteine oxidation states that can occur in proteins: RS-H, reduced thiol; RS-SR, disulfide; RS-NO; S-nitrosated; RS-SG, S-glutathiolated; RS-OH, sulfenated; RS-CR’, alkylated.

optional step (Subheading 3.3), a Western blotting protocol for immunodetection of biotinylated proteins is provided, which can be used to optimize the biotin switch assay before performing protein digestion, peptide capture, and LC-MS/MS analysis (Subheadings 3.4 and 3.5). 3.1. Cell Culture and Lysis 1. Cells are maintained in appropriate growth medium containing serum (e.g., DMEM containing 10% FBS). As the speed and efficiency of sample processing is critical for preservation of Pr-SNO, 2–3 treatment conditions are recommended. For example, a typical experiment consists of four cell culture flasks (175 cm2) that are grown to 90–95% confluence and are either untreated (endogenous Pr-SNO) or treated with an NO donor (supplemented Pr-SNO), in duplicate. 2. To prepare cultures for treatment, aspirate serum-containing media and wash the cultures with warm serum-free media. Replace the media with either serum-free media alone or a serum-free media that has been freshly supplemented with 100 µM CysNO (see Note 2) and incubate cultures at 37°C for 20 min in the dark.

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3. Transfer cultures to reduced lighting conditions and aspirate the treatment media. Wash with D-PBS-MC (1 × 10 mL) and then incubate with TE solution (5 mL) to detach the cells from the flask. After the cells have detached, inactivate TE solution by addition of ice-cold media containing 0.1% FBS (5 mL). Transfer cell suspensions from a single flask (10 mL) to one 15 mL centrifuge tube. Wash the culture flasks with ice-cold D-PBS-MC (1 × 5 mL) and combine with the cell suspensions. Pellet the cells by centrifugation (200 × g, 6 min, 4°C), then wash cell pellets with ice-cold D-PBS-MC (3 × 10 mL). 4. Add ice-cold lysis buffer (1 mL) to the cell pellets and triturate with a P200 pipette followed by gentle vortexing. Rotate crude cell lysates at 4°C in the dark for 30 min, followed by centrifugation (14,000 × g, 10 min, 4°C). Recover supernatant and rapidly determine the protein concentration using the Bradford reagent and bovine serum albumin as a standard. If necessary, dilute cell lysates to a final concentration of 0.8–1 mg/mL with lysis buffer (see Note 8).

3.2. Biotin Switch Assay: Derivitization of Pr-SNO 1. Unless otherwise stated, perform all steps on ice and either protected from direct sunlight or in the dark. 2. If a UV transilluminator or UV photolysis instrument is available, a negative control sample can be generated as UV light effectively reduces the S-NO bond (7, 10) (λmax = 330–340 nm). For example, to use a UV transilluminator to generate Pr-SNO depleted samples, divide equal aliquots (2 × 1 mL) of cell lysates into Petri dishes. For negative control samples that are to be depleted, leave the dishes uncovered. For the aliquots where Pr-SNO is to be preserved, wrap the Petri dishes in aluminum foil. Place both sets of dishes in an ice trough and expose to UV light for 60 min by placing an inverted UV transilluminator over the dishes (7). 3. Transfer lysates (1 mL) to 15 mL centrifuge tubes, washing the Petri dishes with an equal volume of lysis buffer to maximize protein recovery. 4. Precipitate proteins with 2 volumes (4 mL) of acetone (–20°C) and incubate at −20°C for 10 min, followed by centrifugation (3,000 × g, 10 min, 4°C). Discard the supernatant. 5. Resuspend the protein pellet in blocking buffer (1 mL) and incubate at 50°C for 30 min with frequent mixing. 6. Precipitate blocked proteins with 4 volumes (4 mL) of acetone (see step 4). Discard supernatant and solubilize pellet in resuspension buffer. Repeat acetone precipitation, then wash pellet with acetone (3 × 4 mL). 7. Resuspend pellets in labeling buffer (1 mL) and aliquot (2 × 0.5 mL) in 1.5 mL centrifuge tubes. To one aliquot, add 5 uL of sodium ascorbate solution (see Notes 9 and 10). Incubate at room temperature with gentle rotation. 8. Precipitate biotinylated proteins with 2 volumes (1 mL) acetone (see step 4). Discard supernatant, then wash pellet with acetone (3 × 1 mL). Resuspend protein in bicarbonate buffer (0.5 mL) (see Note 11). Samples no longer need to be protected from light. Estimate protein concentration using either the Bradford reagent or BCA assay.

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3.3. SDS-PAGE and Western Blotting for Biotinylated Proteins 1. Prepare 1X MOPS running buffer (1 L) and assemble XCell Surelock Mini-cell system using a NuPAGE 4–12% Bis-Tris gel. 2. If necessary dilute samples to 1 mg/mL. Remove an aliquot of each sample (30 ul) and add 6X nonreducing sample buffer (6 µL). 3. Load 5 uL of SeeBlue molecular weight marker, followed by 30 uL of each sample per lane. 4. Electrophorese samples for at 150 V for 5 min, then at 200 V for 55 min at room temperature. 5. Transfer electrophoresed proteins to PVDF membrane using a Mini Trans-blot apparatus filled with 1X Transfer buffer. Using an ice cooler tank insert, transfer at 100 V for 1 h at 4°C. 6. After transfer, incubate PVDF membrane in membrane blocking buffer (25 mL) for at least 1 h at room temperature or overnight at 4°C under rotation. 7. Incubate PVDF membrane in primary antibody solution (10 mL) for 1 h at room temperature under rotation. Meanwhile, prepare 1X TBS-T wash buffer (0.5 L). 8. Discard primary antibody solution and wash PVDF membrane with TBS-T wash buffer (25 mL; 2 × 5 min, 2 × 10 min). 9. Incubate PVDF membrane with secondary antibody solution (10 mL) for 1 h at room temperature under rotation. Protect membrane from light. 10. Discard secondary antibody solution and wash PVDF membrane with TBS-T wash buffer (25 mL; 2 × 5 min, 1 × 10 min). Perform a final wash with TBS (25 mL; 1 × 10 min). 11. Scan PVDF membrane on the Odyssey Infrared Imaging system (see Note 12). A representative anti-biotin Western blot from CysNO-treated lysates that were processed by the biotin switch is shown (Fig. 2). Omission of ascorbate served as a negative control (Fig. 2, lanes 1, 3, 5).

3.4. Protein Digestion and Peptide Capture 1. Rinse Microcon YM-10 filters with 100% methanol (0.2 mL), and centrifuge (5,000 × g, RT). Wash with water (0.2 mL) and repeat centrifugation. 2. Immediately transfer biotinylated proteins (from Subheading 3.2, step 8) to prerinsed Microcon YM-10. Digest proteins by addition of trypsin (1:100 enzyme:protein) for 16 h at 37°C. 3. Centrifuge (5,000 × g, RT) until entire volume (~0.45 mL) has passed through the filter. Discard retentate and recover the filtrate containing a mixture of biotinylated and non-biotinylated peptides. 4. Remove 0.1 mL of streptavidin-agarose suspension from the stock (1:1 as supplied by Pierce) per mg of initial protein that was digested (across all samples). Wash streptavidin-agarose beads with 10 volumes of 0.1 M ammonium bicarbonate. Centrifuge (500 × g), discard supernatant, and resuspend beads in 0.1 M ammonium bicarbonate, preparing a homogenous 1:1 beads:bicarbonate suspension. Add 0.1 mL of streptavidin-agarose suspension per mg of initial protein that was digested (single sample) to each peptide mixture and incubate for 1 h at RT with gentle rotation.

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Fig. 2. Biotin switch Western blot for biotinylated proteins. Brain lysates were untreated (control) (1,2) or treated with 20 µM CysNO (3–6), then processed by the biotin switch (Subheading 3.2– 3.3). All samples were incubated in labeling buffer containing biotin-HPDP. The detection of Pr-SNO is observed in both ascorbate-treated (4) and copper(II)/ascorbate-treated (6) samples (see Note 9). The absence of signal from the copper(II)/ascorbate-treated control sample (2) is due to the low levels of endogenous Pr-SNO (< 0.01 nmol/mg protein), which are below the limit of detection for this approach (~1 nmol/mg protein). Importantly, the omission of ascorbate from aliquots of 2, 4, and 6 displayed little to no signal (1, 3, and 5).

5. Centrifuge (500 × g, 1 min) and carefully aspirate the supernatant. Resuspend the beads in bicarbonate wash buffer (0.5 mL) and centrifuge (500 × g, 1 min). Repeat washing an additional 4 times, followed by 5 washes with water (0.5 mL). 6. Add 70% formic acid (100 µL) to the dry, washed streptavidin-agarose beads and incubate for 30 min at RT with gentle rotation to elute the biotinylated peptides. Centrifuge samples (1,000 × g, 5 min) and transfer the supernatant to clean microcentrifuge tubes. Repeat centrifugation and supernatant transfer to ensure agarose beads are removed from the sample. 7. Concentrate eluted peptides in vacuo to near dryness and resuspend in 0.1% formic acid (20 µL). If desired, peptides can be stored at −80°C for several months. 8. Transfer an aliquot of resuspended peptide mixture (10 µL) to an autosampler injection vial.

3.5. Peptide Identification by LC-MS/MS Liquid chromatography coupled to electrospray ionization (ESI) tandem mass spectrometry (MS/MS) affords significantly improved sensitivity compared to matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) or threedimensional ion trap instruments. In particular, two-dimensional ion traps, such as the Thermo LTQ, are ideal for the identification of low abundance post-translational

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modifications from complex mixtures. This is due to its low femtomole limit of detection achieved by improved ion capacity, shorter duty cycle, and data-dependent MS2 acquisition, as well as its ability to site-specifically map post-translational modifications via collision-induced dissociation (CID) peptide sequencing. Yet this improved sensitivity, which typically results in the acquisition of 12,000–15,000 MS/MS spectra per sample, can be a double-edged sword. While the number of high quality, matched spectra is increased, the number of poor quality spectra resulting from the detection of low abundance peptides or simply noise is also increased. Therefore, this protocol employs a multistep analysis that uses both automated computational algorithms (SEQUEST and Scaffold) as well as manual inspection of MS/MS spectra to reduce false-positive peptide assignments (see Subheading 3.5.2). 3.5.1. LC-ESI/MS/MS Analysis Liquid separation of peptides is achieved by reverse phase C18 using a Thermo Surveyor autosampler and pump, while ESI-MS/MS is performed by a Thermo LTQ ion trap mass spectrometer. Biotinylated peptides (7 µL) are subjected to reverse phase LC separation on a 0.1 × 110 mm fused silica capillary column fitted with a 60 mm frit (precolumn) packed with Jupiter C18 (5 µm particle size) using a flow rate of 0.7 µL/min. The mobile phase consisted of 0.1% formic acid in either HPLC grade water (A) or acetonitrile (B). Peptides are initially washed onto the column with 100% A for 10 min, followed 100 to 98% A for 5 min. Peptides are eluted with linear gradients of increasing solvent B from 2 to 25% B for 30 min, followed by 25% to 90% B for 15 min. As peptides are eluted from the capillary LC column they are subjected electrospray ionization (capillary voltage = 2 kV). Full scan (MS) spectra are acquired in positive ion mode from 400–2000 m/z, followed by data-dependent acquisition of MS/MS spectra on the four most intense ions from the MS scan. 3.5.2. Database Searching and Validation of Peptide Assignments MS/MS spectra are extracted from the raw files using SEQUEST. SEQUEST output files are generated by searching raw spectra against theoretical MS/MS spectra generated from the in silico trypsinolysis of protein databases, such as the NCBI nr or IPI sequence databases, with a peptide mass tolerance of 2.5 amu and a fragment ion tolerance of 1.0 amu (see Note 6). Cysteine modification by methyl methanethiosulfonate (+46 atomic mass units) and by biotin-HPDP (+428 atomic mass units) are specified as variable modifications. To compare SEQUEST sequence-to-spectrum assignments across several MS/MS experiments, SEQUEST output files are loaded into Scaffold. Only peptide assignments containing a biotin-HPDP-modified cysteine (+428 amu) are considered (see Note 13). Peptides assignments are first filtered by the following

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SEQUEST threshold scores: XC ≥ 2.5 for doubly charged and ≥ 3.5 for triply charged ions; RSp ≤ 5; preliminary score (Sp) ≥ 350. Assignments that satisfy these criteria and are identified in at least 2 out of 3 independent experiments are manually reviewed (11). A peptide assignment with below threshold scores and/ or marginal spectrum quality is accepted if it exhibits a pattern of fragmentation and relative ion peak intensity that is similar to an accepted peptide assignment present in another replicate. Shown in Fig. 3 are representative MS/MS spectra that contain cysteine-modified biotin-HPDP, which were either accepted (A) or rejected (B). Finally, accepted peptide assignments are compared between controls (e.g. –ascorbate or +photolysis) and experimental treatments. Only peptides assignments unique to the experimental group are considered as authentic Pr-SNO peptide identifications. 4. Notes 1. Unless stated otherwise, “water” should be of > 18 MΩ-cm resistance and used for preparing all solutions. 2. After thawing the aliquot, the concentration is determined spectrophotometrically. First, blank a quartz cuvette against 999 uL of water, then add 1 uL of the stock solution, mix by inversion and read absorbance at 335 nm. Calculate the concentration of the stock solution by the following formula: [CysNO] = (Abs335 nm × 1000)/900 M−1cm−1. After determining the stock concentration store on ice and use within 30 min. 3. A moderate amount of MMTS is included in the lysis buffer to limit potential loss of Pr-SNO and/or artifactual transnitrosation chemistries that may occur during homogenization. However, if isolating tissue or cellular lysates for the purpose of post-lysis NO donor treatment, MMTS should be omitted. 4. Gel electrophoresis is described using NuPAGE Bis-Tris gradient mini gels, however, any standard SDS-PAGE system can be employed as long as the buffers (e.g., running and loading buffer) do not contain reducing agents. These gradient gels were used for their ability to efficiently resolve non-reduced protein samples as well as for their consistent transfer characteristics. Fig. 3. (continued) This fragmentation pattern is consistent with the presence of glutamine residues (Q) in the sequence; therefore this assignment would be accepted. (B) Identified as the doubly-charged sequence TLEIEAC428R with an Xc of 2.59. Although this spectrum had a reasonable series of continuous y-ions, several of the most intense peaks (arrows) could not be ascribed to singly or doubly-charged fragment ions or to fragment ions containing neutral losses of either water. Without additional evidence, this assignment would be rejected.2+ indicates a doubly-charged y- or b-ion fragment. * designates fragments that have lost ammonia, while @ designates fragments that have lost water.

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Fig. 3. Representative MS/MS spectra of Cys-biotin-HPDP labeled peptides. Both spectra passed the initial Sequest threshold scoring filter and were subjected to manual inspection (Subheading 3.5.2). (A) Identified as the doubly-charged sequence, VIQC428FAETGQVQK, with an Xc = 4.45. This spectrum contains well-matched peaks containing several y- and b-ions in series with most major peaks matched. Diagnostic pairs of b-ion fragments (b5/ b5*-b12/b12*) are observed; corresponding to the expected fragment (b6, m/z = 1090.3) and the fragment resulting from a loss of ammonia (b6*, m/z = 1073.3).

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5. Visualization of anti-biotin reactivity has been successfully performed with anti-mouse IgG conjugated to hRP; however, antimouse IgG conjugated to Alexa Fluor 680 allows near-infrared detection via the Odyssey Imaging system. For Western applications, this system provides linear range over 4–5 orders of magnitude as well as superior reproducibility compared to chemiluminescent detection. 6. SEQUEST evaluates peptide assignments primarily by cross correlation scores (Xc). This threshold scoring system allows defined cutoffs to be set for “correct” and “incorrect” peptides. However, the appropriate threshold will likely vary between different experimental conditions, such as instrument type, sample complexity, and protein database size. Also, SEQUEST scores do not inherently provide the probability that a peptide assignment is correct or incorrect. Currently, there are several statistical methods that can address these issues. For example, SEQUEST database searches can be performed against in silco digests of “reversed” databases, which contain the protein sequences in the normal orientation and an inverted orientation, where the protein sequences are listed from the C to N terminus. Peptide assignments to the reversed sequences are considered incorrect assignments and can be used to calculate the false positive (FP) or error rate for a particular experiment ( (12, 13) ). Appropriate SEQUEST threshold scores are chosen to obtain a desired FP rate. Usually ≤ 5% peptide FP rate is satisfactory for two-dimensional ion traps. As a complementary method for evaluating the quality of peptide assignments, the PeptideProphet and ProteinProphet algorithms utilize Bayesian statistics to determine the probability that an individual peptide assignment is correct (14,15). By assuming that correct and incorrect peptide assignments follow normal distributions, these algorithms in combination with manual inspection, provide an effective strategy for validating MS/MS peptide assignments generated from database search algorithms such as SEQUEST and Mascot. These algorithms are available as open source packages, implemented in the Trans-Proteomic Pipeline (http://tools.proteomecenter.org) or utilized within commercial software such as Scaffold (Proteome Software, Portland, OR). 7. In various rat tissues, endogenous Pr-SNOs are present between 1–10 pmol per mg of total protein (16). Similarly, in a cell culture model of murine macrophages (RAW 264.7), LPS-induced iNOS induction generated approximately 25 pmol of Pr-SNO per mg of total protein (17). For the current approach, between 0.1–0.5 nmol total Pr-SNO isolated from cells or tissue is the lowest recommended amount of modification to perform the analysis. These estimates suggest a starting protein amount of between 10 and 50 mg of protein per sample is appropriate. Given a single experimental

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9.

10.

11.

12.

13.

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condition could be split into as many as 4 samples (±photolysis and ±ascorbate), obtaining this amount of starting material from cellular cultures is challenging. Currently, for cell culture experiments this usually requires samples to be processed by the biotin switch (Subheading 3.2) and then pooled from several experiments before proceeding to mass spectrometry analysis (Subheading 3.4). As technological advancements in MS sensitivity, mass accuracy, and sample processing are made these experiments will likely become more manageable. Although protein yield from a nearly confluent 175 cm2 cell culture flask will depend on the cell line used, a reasonable expectation is 2–5 mg. As described in the method, the yield from a single 175 cm2 flask was 2 mg (primary human aortic smooth muscle cells). Recent studies have suggested that not all S-NO bonds are equally susceptible to ascorbate reduction (18,19). Therefore, it is important to note that Pr-SNOs identified by this method likely reflect the subset, which are most susceptible to ascorbate reduction. As an alternative approach, our lab has observed significantly increased efficiency of Pr-SNO reduction using copper(II)/ascorbate (100 µ M/50 µ M) compared to ascorbate alone (5 mM) (Fig. 2, Lane 3 vs 5). Thus far we have not observed any nonspecificity due to Cu/Asc treatment by Western blotting (Lane 1 vs 2) or by LC-MS/MS (data not shown). Several studies have suggested that high concentrations of ascorbate (>10 mM) can generate nonspecific biotin-HPDP labeling through the reduction of disulfide bonds. This direct reaction is unlikely as even the one electron reduction of cystine by the dehydroascorbate radical is thermodynamically unfavorable (10). While direct mechanistic evidence for the ascorbate-dependent nonspecificity has not been demonstrated in the context of the biotin switch, it is likely that inadvertent exposure to UV light underlies this observation (10), highlighting the importance of protecting samples from sunlight exposure. After bicarbonate resuspension, ensure that large clumps of the protein pellet are disrupted. It is normal if the solution appears cloudy; the protein will be digested efficiently by overnight trypsin incubation. The default instrument settings as specified by the “membrane” preset are usually sufficient. Nonetheless, increased sensitivity and lower background has been observed if the membrane is first dehydrated in methanol, and then allowed to dry before scanning. This may require a lower intensity setting compared to the default method. The specificity of peptide identification for biotin-HPDP labeled cysteines should be ≥90%. If this was not achieved, consider increasing the volume or number of washes before eluting the biotinylated peptides from the

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streptavidin-agarose beads. Alternatively, manual inspection may reveal that these assignments are low quality spectra. In this case, the SEQUEST threshold filters should be adjusted and/or searching against a reverse database is recommended to ensure false-positive rates are ≤5% (see Note 6). References 1. Choi, Y. B., Tenneti, L., Le, D. A., Ortiz, J., Bai, G., Chen, H. S., and Lipton, S. A. (2000) Molecular basis of NMDA receptor-coupled ion channel modulation by S-nitrosylation. Nat. Neurosci. 3, 15–21. 2. Mannick, J. B., Hausladen, A., Liu, L., Hess, D. T., Zeng, M., Miao, Q. X., Kane, L. S., Gow, A. J., and Stamler, J. S. (1999) Fas-induced caspase denitrosylation. Science 284, 651–654. 3. Jaffrey, S. R. and Snyder, S. H. (2001) The biotin switch method for the detection of S-nitrosylated proteins. Sci. STKE PL1. 4. Marshall, H. E. and Stamler, J. S. (2001) Inhibition of NF-kappa B by S-nitrosylation. Biochemistry 40, 1688–1693. 5. Hao, G., Derakhshan, B., Shi, L., Campagne, F., and Gross, S. S. (2006) SNOSID, a proteomic method for identification of cysteine S-nitrosylation sites in complex protein mixtures. Proc. Natl. Acad. Sci. U. S. A. 103, 1012–1017. 6. Greco, T. M., Hodara, R., Parastatidis, I., Heijnen, H. F., Dennehy, M. K., Liebler, D. C., and Ischiropoulos, H. (2006) Identification of S-nitrosylation motifs by sitespecific mapping of the S-nitrosocysteine proteome in human vascular smooth muscle cells. Proc. Natl. Acad. Sci. U. S. A. 103, 7420–7425. 7. Derakhshan, B., Wille, P. C., and Gross, S. S. (2007) Unbiased identification of cysteine S-nitrosylation sites on proteins. Nat. Protoc. 2, 1685–1691. 8. Keefer, L. K., Nims, R. W., Davies, K. M., and Wink, D. A. (1996) “NONOates” (1-substituted diazen-1-ium-1,2-diolates) as nitric oxide donors: convenient nitric oxide dosage forms. Methods Enzymol. 268, 281–293. 9. Zhang, Y. and Hogg, N. (2005) S-Nitrosothiols: cellular formation and transport. Free Radic. Biol. Med. 38, 831–838. 10. Forrester, M. T., Foster, M. W., and Stamler, J. S. (2007) Assessment and application of the biotin switch technique for examining protein S-nitrosylation under conditions of pharmacologically induced oxidative stress. J. Biol. Chem. 282, 13977–13983. 11. Tabb, D. L., Friedman, D. B., and Ham, A. J. (2006) Verification of automated peptide identifications from proteomic tandem mass spectra. Nat. Protoc. 1, 2213–2222. 12. Cargile, B. J., Bundy, J. L., and Stephenson, J. L., Jr. (2004) Potential for false positive identifications from large databases through tandem mass spectrometry. J. Proteome Res. 3, 1082–1085. 13. Peng, J., Elias, J. E., Thoreen, C. C., Licklider, L. J., and Gygi, S. P. (2003) Evaluation of multidimensional chromatography coupled with tandem mass spectrometry (LC/LC-MS/MS) for large-scale protein analysis: the yeast proteome. J. Proteome Res. 2, 43–50.

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14. Keller, A., Nesvizhskii, A. I., Kolker, E., and Aebersold, R. (2002) Empirical statistical model to estimate the accuracy of peptide identifications made by MS/MS and database search. Anal. Chem. 74, 5383–5392. 15. Nesvizhskii, A. I., Keller, A., Kolker, E. and Aebersold, R. (2003) A statistical model for identifying proteins by tandem mass spectrometry. Anal. Chem. 75, 4646–58. 16. Bryan, N. S., Rassaf, T., Maloney, R. E., Rodriguez, C. M., Saijo, F., Rodriguez, J. R. and Feelisch, M. (2004) Cellular targets and mechanisms of nitros(yl)ation: an insight into their nature and kinetics in vivo. Proc. Natl. Acad. Sci. U. S. A. 101, 4308–13. 17. Zhang, Y. and Hogg, N. (2004) Formation and stability of S-nitrosothiols in RAW 264.7 cells. Am. J. Physiol. Lung Cell. Mol. Physiol. 287, L467–74. 18. Zhang, Y., Keszler, A., Broniowska, K. A. and Hogg, N. (2005) Characterization and application of the biotin-switch assay for the identification of S-nitrosated proteins. Free Radic. Biol. Med. 38, 874–81. 19. Weichsel, A., Brailey, J. L. and Montfort, W. R. (2007) Buried S-nitrosocysteine revealed in crystal structures of human thioredoxin. Biochemistry. 46, 1219–27.

156 Detection of Nitrotyrosine-Containing Proteins Xianquan Zhan and Dominic M. Desiderio 1. Introduction Tyrosine nitration in a protein results from the peroxynitrite anion (ONOO−, generated from the chemical reaction of nitric oxide and superoxide anion) in a cell and from oxidative stress, and might be an important molecular event in the physiological and pathological processes of the human pituitary. The pituitary is main component of multiple hypothalamic–pituitary–target organ axis systems. Studies have indicated that nitric oxide (NO) (1) and nitric oxide synthase (NOS) participate in those axis systems (2–4), including luteinizing hormone (LH) (4–7), follicle-stimulating hormone (FSH) (5), prolactin (PRL) (8), growth hormone (GH) (9–11), and adrenocorticotropin (ACTH) (12) axis systems. We recently discovered several nitrotyrosine-containing proteins in human pituitary postmortem and nonfunctional adenoma tissues (1,13,14), and this chapter describes different methods for detecting nitrotyrosine containing proteins, using pituitary tissue as an example. The detection and identification of an in vivo nitrotyrosine-containing protein is very challenging. Commercial antinitrotyrosine polyclonal/monoclonal antibodies, combined with different technologies such as enzymelinked immunosorbent assay (ELISA) (15,16), one/two-dimensional sodium dodecyl sulfate polyacrylamide gel electrophoresis (1-D/2-D SDS-PAGE)based Western blot assay (13,14,17,18), and immunoprecipitation (14), are effective methods to measure a nitrotyrsoine level, and to isolate, enrich, and detect an endogenous nitroprotein. ELISA has been extensively used to analyze redox production (15,16,19). ELISA is a classic method to measure the content of nitrotyrosine, and the ELISA assay kit is commercially available (Upstate Catalog No. 17–136). 1-D/2-D SDS-PAGE can separate and preferentially enrich nitroproteins based on the unique properties (molecular weight MW and electric point pI) of a protein. Western blot assay not only detects the From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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tyrosine-containing protein, but also determines the relative level of nitrotyrosine. Modern tandem mass spectrometry (1,13,14) methods can determine the amino acid sequence of proteolytic fragment of a protein to locate the nitrotyrosine sites in a protein sequence to clarify the function and role of endogenous tyrosine nitration. 2. Materials 2.1. Tissue Homogenization 1. Liquid nitrogen, freezer (−80°C), polytron homogenizer (Model P710/35; Brinkmann Instruments, Westbury, NY), and Fisher sonicator (Model FS30H; Fisher, Pittsburgh, PA). 2. Sodium chloride (0.9%) solution in deionized destilled water (see Note 1). 3. Homogenizing buffer (10 ml): 2 M acetic acid and 0.1% (v/v) mercaptoethanol. Store at 4°C. 4. Bicinchoninic acid (BCA) protein assay reagent kit (Pierce, Rockford, IL).

2.2. Enzyme-Linked immunosobent assay (ELISA) 1. Slide-A-Lyzer dialysis cassette 2K MWCO, 0.2–0.5 ml (Pierce Product number 66205). 2. Phosphate-buffered saline (PBS) (200 mM; 20 ×): 4.6 g NaH2PO4 (or 5.3 g NaH2PO4•H2O), 23 g Na2HPO4, 175 g NaCl, and 0.2 g NaN3 are dissolved in c.750 ml deionized destilled water. Adjust pH to 7.22. Add deionized destilled water to 1 liter. Store at room temperature. Dilute 50 ml of 20 × PBS with 950 ml deionized destilled water for use. 3. Bovine serum albumin (BSA), Fraction V (Sigma). 4. Tetranitromethane (Sigma Product Number T25003) (see Note 2). 5. 50 mM Tris-HCl, pH 8.0, 50 ml: 302.75 mg Tris-base is dissolved in 40 ml deionized destilled water, adjust pH value to 8.0 with hydrolytic chloride, and add deionized destilled water to 50 ml. 6. Mouse antinitrotyrosine monoclonal antibody, clone 1A6 (100 µg/100 µl Catalog number 05-233, Upstate Biotechnology, Lake Placid, NY). Store at −20°C. 7. Rabbit antimouse IgG conjugated with alkaline phosphatase. Store at 4°C. 8. Alkaline phosphatase substrate pNPP (Sigma Fast™ P-Nitrophenyl phosphate Tablet set: pNPP 1.0 mg/ml and Tris buffer 0.2 M) (Sigma). Store at 4°C. 9. Costar® EIA/RIA plate with 96-well flat bottom (Product number 122 06049; Corning Incorporated, Corning, NY).

2.3. 1-D Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (1-D SDS-PAGE) 1. Protein extraction buffer: 60 mM Tris-HCl pH 6.8, 2% SDS, 10 mM dithiothreitol, 100 µM butylated hydroxytoluene (BHT) (see Note 3), and 2 mM EDTA. 2. Resolving gel stock buffer: 1.5 M Tris-HCl, pH 8.8. Store at 4°C.

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Stacking gel stock buffer: 0.5 M Tris-HCl, pH 6.8. Store at 4°C. 30% Acrylamide/ 0.8% bisacrylamide solution (see Note 4). 10% SDS: 10 g SDS is dissolved in 100 ml deionized destilled water. N,N,N′,N′-tetramethyl-ethylenediamine (TEMED, Bio-Rad, Hercules, CA). Ammonium persulfate (10%): 100 mg of ammonium persulfate is dissolved in 1 ml deionized destilled water prior to use (see Note 5). 50% isobutanol in water: 50 ml isobutanol mix with 50 ml deionized destilled water. Store at room temperature. Mix well prior to use. Electrophoresis running buffer (10×): 250 mM Tris, 1.92 M glycine, and 1% (w/v) SDS. Store at room temperature. Sample loading buffer: 30% (v/v) glycerol, 2% SDS, 62.5 mM Tris pH 6.8, 50 mM DTT, 5 mM EDTA, 0.02% NaN3. Store in 200 µl aliquots in −20°C. Prestained molecular weight markers: All Blue Precision Plus ProteinTM Standards (Catalog No. 161-0373, Bio-Rad, Hercules, CA). Store in 20 µl aliquots at −20°C.

2.4. 2-D Gel Electrophoresis (2DGE) 1. Protein extracting buffer (1 ml): 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 100 mM DTT, 0.5% pharmalyte, and trace bromophenol blue (see Notes 6 and 7). 2. Rehydration buffer (1 ml): 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 60 mM DTT, 0.5% Pharmacia IPG buffer, and trace bromophenol blue (see Note 7). 3. Resolving-gel buffer stock (4×): 1.5 M Tris-HCl, pH 8.8. Store at 4°C. 4. Reducing equilibration buffer (50 ml): 375 mM Tris-HCl pH 8.8, 6 M urea, 2% (w/v) SDS, 20% (v/v) glycerol, 2% DTT, and trace bromophenol blue (see Note 8). 5. Alkylation equilibration buffer (50 ml): 375 mM Tris-HCl pH 8.8, 6 M urea, 2% (w/v) SDS, 20% (v/v) glycerol, 2.5% w/v iodoacetamide, and trace bromophenol blue (see Note 8). 6. 40% acrylamide/bisacrylamide stock solution (29:1): 40% w/v acrylamide, 1.38% w/v N, N′-methylenebisacrylamide (see Note 4) (Catalog No. 161-0146, Bio-Rad). 7. 10% Ammonium persulfate (3 ml) is made just prior to use (see Note 5). 8. SDS electrophoresis buffer (1 ×; 25 L): 25 mM Tris, 192 mM glycine, and 0.1% SDS. Store at room temperature. (see Note 9). 9. 1% Agarose sealing solution (100 ml) is made by heating until the agarose is completely dissolved and kept at 80°C prior to use. (see Note 10).

2.5. Western Blotting for Nitrotyrosine Based on 1-D SDS-PAGE 1. Transfer buffer (1 L): 25 mM Tris, 192 mM glycine, and 10% (v/v) methanol. 2. Polyvinylidene fluoride (PVDF) membrane: Immobilon-P transfer membrane (Catalog No. IPVH 081 00; Millipore, Bedford, MA) (see Note 11). 3. Mini Trans-blot filter paper (7.5 × 10 cm, Catalog No. 1703932; Bio-Rad). 4. Tris-buffered saline (TBS) stock solution (10×): 250 mM Tris-HCl pH 7.4, 1.37 M NaCl, 27 mM KCl. Dilute 100 ml with 900 ml deionized water for use. 5. Blocking solution: 1% (w/v) fraction V bovine serum albumin in TBS.

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6. Primary antibody: mouse antinitrotyrosine monoclonal antibody, clone 1A6 (100 µg/100 µl Catalog number 05-233, Upstate Biotechnology, Lake Placid, NY). Dilute in 1:2000 with 3% BSA in TBS solution (0.5 µg Ab/mL) for use (see Note 12). 7. Secondary antibody: Sheep antimouse IgG conjugated with horse radish peroxidase (Catalog No. NXA931, Amersham). Dilute 1:7500 in 5% nonfat dry milk (NFDM) in TBS solution for use. 8. Enhanced chemiluminescent (ECL) reagents (Catalog No. RPN2106; Armersham Amersham Biosciences, Piscataway, NJ) and Bio-Max ML film (Kodak, Rochester, NY).

2.6. Western Blotting for Nitrotyrosine Based on Large-Format 2DGE 1. Gel equilibration buffer (1 L): 25 mM Tris and 192 mM glycine. 2. PVDF membrane equilibration buffer (1 L): 25 mM Tris, 192 mM glycine, and 20% (v/v) methanol. 3. Polyvinylidene fluoride (PVDF) membrane: Hybond-P 20 × 20 cm (Product No. RPN2020F; Amersham), or Immobilon-P transfer membrane (IPVH 20200; Millipore, Bedford, MA). 4. NovaBlot Electrode Filter Paper (Product No. 80-1105-19; Amersham). 5. Anode transfer buffer R (1 L): 3.63% (w/v) Tris, 20% (v/v) methanol, pH 10.4. 6. Anode transfer buffer S (1 L): 0.303% (w/v) Tris, 20% (v/v) methanol, pH 10.4. 7. Cathode transfer buffer T (1 L): 0.52% (w/v) 6-amino-n-hexanoic acid, 20% (v/v) methanol, pH 7.6). 8. Phosphate-buffered saline (PBS) (200 mM; 20 ×): 4.6 g NaH2PO4 (or 5.3 g NaH2PO4•H2O), 23 g Na2HPO4, 175 g NaCl, and 0.2 g NaN3 are dissolved in c.750 ml deionized destilled water, adjust pH to 7.22, add deionized destilled water to 1 liter. Store at room temperature. 9. PBS: 10 mM PBS. 50 ml of 20 × PBS is diluted with 950 ml deionized water. 10. PBST: 10 mM PBS, 0.2% (v/v) Tween-20, and 0.01% sodium azide. 11. 0.3% BSA/PBST: 0.3% w/v BSA, 10 mM PBS, 0.2% v/v Tween-20, and 0.01% w/v sodium azide. 12. Primary antibody: Rabbit anti-human nitrotyrosine antibody (Sigma, N0409, St. Louis, MO). Dilute (1:1000=v:v) in a 0.3% BSA/PBST solution (1 µg Ab/mL) for use. 13. Secondary antibody: goat antirabbit alkaline phosphase-conjugated IgG (Product No. 31340; Pierce, Rockford, IL). Dilute (1:5000 = v:v) in a 0.3% BSA/PBST solution for use. 14. Development reagent: 5-bromo-4-chloro-3-indolyl phosphate/nitro blue tetrazolium (BCIP/NBT) (1-Step™ NBT/BCIP; Pierce, Rockford, IL).

2.7. Immunoprecipitation of Nitrotyrosine-Containing Protein 1. M-PER® mammalian protein extraction reagent (Pierce). 2. ImmunoPure® Immobilized Protein G plus (Pierce). Store at 4°C (see Note 13).

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3. Binding/washing buffer: 140 mM NaCl, 8 mM sodium phosphate, 2 mM potassium phosphate, and 10 mM KCl, pH 7.4. 4. Rabbit Anti-nitrotyrosine polyclonal antibody (Chemicon® International, Inc.), or mouse anti-nitrotyrosine monoclonal antibody (Chemicon® International, Inc.). 5. Disuccinimidyl suberate (DSS), 2 mg/tube (Pierce) (see Note 14). 6. ImmunoPure® IgG Elution Buffer (Pierce), pH 2.8 contains primary amine. 7. Handee™ Spin Cup Columns (Pierce Product No. 69700). 8. Handee™ Microcentrifuge Tubes (Pierce Product No. 69715). 9. The pH-neutralized solution: 1 M Tris, pH 9.5.

2.8. Tandem Mass Spectrometry Characterization of Nitrotyrosine-Containing Protein 1. Sequencing grade modified trypsin (Catalog No. V511A, 5 × 20 µg aliquots; Promega). Store at −20°C (see Note 15) 2. Trypsin resuspension buffer (Promega Catalog No. V542A): 50 mM acetic acid. 3. 30 mM potassium ferricyanide (100 ml) is stored at 4°C. 4. 100 mM sodium thiosulfate (100 ml) is stored at 4°C. 5. 100 mM ammonium bicarbonate (100 ml) is stored at 4°C. 6. 50 mM ammonium bicarbonate (100 ml) is stored at 4°C. 7. 1% trifluoroacetic acid (TFA) (100 ml) is stored at 4°C. 8. 0.1% TFA (1 ml) is made prior to use. 9. 50% acetonitrile/0.1% TFA (1 ml) is made prior to use. 10. ZipTipC18 microcolumn (Millipore ZTC18S096, Bedford, MA). 11. 85% v/v acetonitrile/0.1% v/v TFA (1 ml) is made prior to use. 12. 2% actonitrile/0.5% acetic acid (1 ml) is made prior to use.

3. Methods Tyrosine nitration in a protein is a relatively stable redox product that is produced due to oxidative/nitrative stress associated with physiological and pathology processes. The chemical structure of nitrotyrosine is a nitro (NO2) group added to position 3 of the phenolic ring of a tyrosine residue. Commercially available anti-nitrotyrosine monoclonal and polyclonal antibodies enable one to determine the level of nitrotyrosine by ELISA (amount of nitrotyrosine per unit weight tissue), to detect the profile of nitrotyrosine-containing proteins by Western blotting based on 2DGE (1,13) (see Fig. 1) or 1-D SDSPAGE (17,18), and to isolate and enrich nitroproteins by immunoprecipitation (14) and 2DGE Western analysis (1,13). For those isolated and enriched nitroproteins, tandem mass spectrometry is used to obtain each amino acid sequence and to identify each nitrotyrsoine-containing protein and nitrationsites (1,13,14) (see Fig. 2).

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Fig. 1. Two-dimensional Western blot analysis of anti-3-nitrotyrosine proteins in a human pituitary (70 µg protein per 2-D gel). (A) Silver-stained image on a 2D gel before transfer of proteins to a PVDF membrane. (B) Silver-stained image on a 2D gel after transfer of proteins to a PVDF membrane. (C) Western blot image of anti-3-nitrotyrosine proteins (anti-3-nitrotyrosine antibodies + secondary antibody). (D) Negative control of Western blot to show the cross-reaction of the secondary antibody (only the secondary antibody; no anti-3nitrotyrosine antibody). (Reproduced from ref. 1 with permission from Elsevier Science).

Fig. 2. SEQUEST (top-right) and de novo (bottom) analysis of an MS2 spectrum of the precursor ion ( (M+2H)2+, at m/z = 686, RT = 52.3 min, and scan number 2180) for a nitrotyrosyl peptide (Tyr-237) 228GQC#KDALEI*YK238 that contained 11 amino acids, and that derived from synaptosomal-associated protein. (Reproduced from ref. 1 with permission from Elsevier Science).

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3.1. Preparation of Samples 1. The human normal (control) pituitary tissue that is obtained from autopsy or the pituitary adenoma tissue that is obtained from neurosurgery is frozen immediately in liquid nitrogen, stored at − 80°C until processed. 2. Weigh the wet tissue: weigh an empty clean 15-ml tube (WT1), transfer the frozen tissue quickly and carefully, weigh the tube with tissue (WT2). The tissue weight = WT2-WT1. 3. Add 2 ml of 0.9% sodium chloride, and lightly shake to remove the blood on the surface of tissue. Repeat 3 times (see Note 16). 4. Add 10 ml homogenization buffer for c.0.5–0.6 g tissue, and homogenize fully (1 min × 10) with a Polytron homogenizer (13 000 rpm) in 4°C cold room. The homogenate is sonicated with a Fisher sonicator (Power setting = 5) for 20 s. 5. 1 ml of homogenate is centrifuged (10,000 g, 10 min). The supernatant is used for ELISA analysis to determine the nitrotyrosine concentration (see step 1 in Subheading 3.2). 6. The remaining homogenate is lyophilized in 1 ml aliquots, and stored at − 80°C. 7. The total protein content in the lyophilized pituitary is measured with a bicinchoninic acid (BCA) protein assay reagent kit. (see Note 17). (a) 280 µg of lyophilized pituitary sample is added into 264 µl solution that contains 8 M urea and 4% CHAPS. Stand for 2 h, sonicate for 5 min, rotate for 1 h, sonicate for 5 min, rotate for 1 h, centrifuge at 12,000 rpm for 20 min. The supernatant is used for the BCA analysis. (b) Preparation of BSA standard solution: Dilute BSA standard (2 mg/ml) for the following dilution series: 2000, 1500, 1000, 750, 500, 250, 125, 25, 0 µg/ml. (c) BCA working solution: 50 part of reagent A + 1 part of reagent B (50:1), mix fully prior to use. (d) 0.1 ml sample or standard dilution is mixed with 2 ml BCA working solution (1:20). Incubate at 37°C for 30 min. Cool to room temperature (c.10 min). Measure the O.D. value (A562 nm) on the spectrophotometer. (e) Use linear regression to make the standard line (BSA concentration vs. A562 nm), and obtain a regression equation that use A562 nm to estimate the protein concentration. The regression equation is used to calculate the protein concentration of a lyophilized pituitary sample.

3.2. ELISA for Tissue Nitrotyrosine Level 1. The supernatant of 1 ml homogenate of pituitary tissue is dialyzed with a Slide-ALyzer dialysis cassette in 1000 ml PBS for 120 min. Change fresh PBS every 40 min. 2. Preparation of 20 µM standard nitrated BSA (50 ml): 66 mg BSA is dissolved in 50 ml of 50 mM Tris-HCl pH 8.0. Add 599 µl tetranitromethane into the above BSA solution, react for 5 min at room temperature. For removal of salt and tetranitromethane, the nitrated BSA solution is dialyzed with a Slide-A-Lyzer dialysis cassette in 1000 ml PBS for 120 min; change fresh PBS every 40 min. The dialyzed nitrated BSA is stored at − 80°C.

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3. Make coating reagent: dilute 20 µM nitrated BSA with PBS in 1:20 dilution. 4. Add 100 µl coating reagent per well into Costar® EIA/RIA plate with 96-well flat bottom. Incubate at 37°C for 1 h. Wash with 300 µl PBS/well for 3 times. 5. During coating, prepare the blocking solution: 1% Chicken ovalbumin in PBS. 6. Add 300 µl blocking solution per well, and incubate at 37°C for 1 h (Note: this step can be incubated longer). Wash with 300 µl PBS/well for 3 times. 7. During blocking, prepare 11 standard dilution series (0.4 dilution) of 20 µM (20000 pmol/ml) nitrated BSA: 20,000, 8,000, 3,200, 1,280, 512, 205, 82, 33, 13, 5, 2 pmol/ml, and each concentration 125 µl. Prepare 6 sample dilution series (0.2 dilution) of pituitary samples, and each concentration 125 µl. Meanwhile, prepare primary antibody: dilute mouse anti-nitrotyrosine monoclonal antibody in 1:2,000 dilution with 0.2% chicken ovalbumin in PBS. 8. Mix 125 µl of diluted primary antibody and 125 µl standard nitrated-BSA serial sample (or sample dilution series); incubate at 37°C for 1 h. 9. Add 100 µl of mixture above per well (duplicates), and shake at room temperature for 1 h. Wash with 300 µl PBS/well for 3 times. 10. Dilute secondary antibody (anti-mouse IgG-alkaline phasphatase) in 1:2000 dilution with 1% chicken ovalbumin in PBS. Add 100 µl diluted secondary antibody/ well, shake at room temperature for 1 h. Wash with 300 µl PBS for 3 times. 11. Add alkaline phosphatase substrate pNPP solution (100 µl/well). Stand at room temperature until maximum absorbance reaches 0.6–1.0. Measure the absorbance at 405 nm and 655 nm. 12. Make the standard curve (log value of nitrated BSA concentration vs. O.D.) with the SigmaPlot software, and obtain a curve equation. Use sample O.D. value to calculate the nitrotyrosine concentration of pituitary sample (see Note 18). Convert the nitrotyrosine concentration to the nitrotyrosine amount per gram pituitary tissue.

3.3. 1-D SDS-PAGE and Western Blot for Nitrotyrosine Immunoactivity 1. Protein extraction for SDS-PAGE: An amount of lyophilized pituitary that contains 100 µg protein is dissolved in 100 µl protein extraction buffer, stand for 1 h, sonicate 20 s, rotate 1 h, sonicate 20 s. Centrifuge for 20 min at 13,000 × g. The supernatant is the extracted protein. 2. Bio-Rad Mini PROTEAN® Electrophoresis system is used; easily adaptable to other formats. The glass plates for gel is cleaned with the detergent; rinsed extensively with tap water, ethanol, distilled water, and ethanol; dried in a clean plastic rack for use. Install the long and short glass plates in the gel cassette (avoid the bottom leaking). 3. Preparation of a 1.5 mm thick, 12% resolving-gel: 4 ml of 30% acrylamide/0.8% bisacrylamide, 3.34 ml deionized water, 2.5 ml of 1.5 M Tris-HCl pH 8.8, and 100 µl of 10% SDS are mixed in a clean 50-ml tube. Add 50 µl of 10% ammonium persulfate and mix quickly, add 5 µl of TEMED, and mix quickly and fully. Pour the resolving gel solution into the assembled gel cassette (avoid generating

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bubbles), leave 1.0–1.5 cm space for a stacking gel. Overlay the resolving gel with 1 ml of 50% isobutanol-water solution. Stand (room temperature) and avoid moving the gel cassette. The gel will be polymerized in 30–40 min. Remove the isobutanol and rinse with distilled water 3 times, and absorb the redundant water with KimWipe paper. Preparation of a 4% stacking gel: 0.8 ml of 30% acrylamide/0.8% Bis, 3.6 ml deionized water, 1.5 ml 0.5 M Tris-HCl pH 6.8, and 60 µl 10% SDS are mixed in a clean 50-ml tube. Add 30 µl of 10% ammonium persulfate and mix quickly. Add 6 µl TEMED and mix quickly and fully. Pour the stacking gel solution onto the polymerized resolving gel up to the top-end of short glass plate. Insert quickly 1.5-mm thick and 10-well comb into the resolving gel. The stacking gel will be polymerized in 30–40 min. Preparation of electrophoresis running buffer: dilute 100 ml of 10 × running buffer with 900 ml distilled water in a 1000-ml measuring cylinder. Assemble the prepared gel cassette into the Bio-Rad Mini PROTEAN® electrophoresis cell. Add the running buffer into the upper and lower chambers of the gel unit. Let the electrophoresis running buffer cover the top-end of prepared gel. Remove gently and carefully the comb, and the running buffer will automatically fill in each well. Mix 10 µl of the extracted protein and 20 µl of sample loading buffer in a 0.5-ml tube, and boil in 100°C water. Cool to room temperature. Load 30 µl of the prepared sample in a well. Meanwhile, load 5 µl of prestained molecular weight markers into another well. Complete the assembly of the gel unit by covering the lid and connect to the power supply. The gel is run at constant 80 V for 10 min, then at constant 120 V. Stop electrophoresis when the dye front is 0.5 cm from the bottom of glass plate. After electrophoresis, the separated protein is visualized with development reagent, such as Gel-code blue (see step 9) and silver stain (see step 21 in Subheading 3.4), or is transferred to PVDF membrane for Western blot analysis (see step 10 and forward). Disassemble the mini-gel unit. Remove the gel carefully, and avoid breaking the gel. Put gel in the clean flat tank, wash with distilled water (shake, 5 min). Replace with 100 ml Pierce Gel-Code blue reagent and shake gently for 1–2 h. Destain with distilled water until the gel band is clearly visible and the gel background is very low. Disconnect the power supply and disassemble the mini-gel unit. Remove the gel, cut a small corner for labeling, and gently shake in transfer buffer for 10 min. Preparation of PVDF membrane: cut a piece of PVDF membrane (6 × 9 cm) and put it in the 100% methanol and shake for 10 min. Replace methanol with distilled water to check quickly its hydrophilic side (water moves slowly on the hydrophilic side and fast on the hydrophobic side). Use pencil to write the date for label on the margin of the hydrophilic side. Replace water with transfer buffer and shake for at least 10 min. Wet two Mini Tans-blot Filter papers (6 × 9 cm) and two clean fiber pads in the transfer buffer.

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13. Assemble the transfer cassette: Put the black-side of the clean plastic transfer cassette down and open the cassette. Lay on one wet fiber pad onto the blackside. Lay on one sheet of wet filter paper on the wet fiber pad, and remove any bubbles. Put the gel on the filter paper and remove any bubbles. Put the PVDF membrane (the hydrophilic side down) on the gel and remove any bubbles. Put one sheet of wet filter paper on the PVDF and remove bubbles. Put one wet fiber pad on the filter paper. Close the transfer cassette. 14. The assembled transfer cassette is placed into the transfer tank with sufficient transfer buffer and a magnetic stir bar. The black side of the transfer cassette is toward the black side (cathode, -). Put into a flat box of cooling ice. The lid is put on the transfer tank and connected to the power supply. The transfer tank is covered with ice. Transfer is carried out at 320 mA constant for 2 h or at 300 V constant for 1 h. 15. After transfer, disconnect the power supply, disassemble the transfer cassette (Position the gel up and PVDF down), and use a pencil to draw a line around the gel on the hydrophilic side of PVDF for labeling. Remove the gel. Place the PVDF membrane into a block solution. 16. The PVDF membrane (protein-side up) is blocked in 1% BSA-TBS solution (gently shake for 1 h) (see Note 19). After block, the PVDF is washed two times with deionized water. 17. Add the primary antibody solution 30 ml (15 µl mouse anti-nitrotyrosine monoclonal antibody is diluted in 30 ml of 3% BSA in TBS solution), and incubate for 1 h at room temperature. Pour out the primary antibody solution. Wash with 100 ml of 0.2% Tween-20 in TBS for 10 min for 6 times. 18. Add the secondary antibody solution (6.7 µl antimouse IgG conjugated with horse radish peroxidase diluted in 50 ml of 5% nonfat dry milk in TBS), and incubate at room temperature for 1 h with gentle shaking. Pour out the secondary antibody solution. Wash with 100 ml of 0.2% Tween-20 in TBS for 10 min for 3 times. Wash with 100 ml of TBS for 10 min for 3 times. Rinse with deionized water for 4 times (see Note 20). 19. ECL visualization of the blotted PVDF membrane: Absorb the redundant water on the PVDF membrane between two sheets of filter paper. Place PVDF membrane on a clean glass plate (protein-side up) and cover with pre-prepared ECL reagent (4 ml ECL reagent A is mixed fully with 4 ml ECL reagent B prior to use). Incubate for 3–5 min. Dry the PVDF membrane between two sheets of filter paper. The PVDF membrane is placed between two transparent plastic sheets, and fixed in an X-ray film cassette. The remaining following steps are carried out in a dark room under the safe red light conditions. 20. Put a sheet of film on the transparent plastic sheet that contains the PVDF membrane and close the X-ray film cassette for a suitable exposure time, typically a few minutes. If the detected protein is very low, then expose for several hours to overnight. If the detected protein is very high, then expose for 10 s to 1 min. Remove the film and place it into an automatic image instrument to obtain the Western blotting image.

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3.4. 2DGE and Western Blot for Nitrotyrosine Immunoactivity 1. Protein extraction for 2DGE: Weigh sufficient lyophilized pituitary sample to mix with 250 µl of the protein extracting buffer. The mixture is vortexted for 5 min, sonicated for 5 min, and rotated for 50 min. Add 110 µl of rehydration buffer. The mixture is sonicated for 5 min, rotate for 50 min, vortexed for 5 min, and centrifuged for 20 min at 15,000 g. The supernatant is the “protein sample solution” (20). 2. Rehydration of IPG strip (see Note 21): add 350 µl of the “protein sample solution” into a slot of the rehydration tray. Remove the plastic cover from the Dry IPG strip (do not touch gel-side). Place slowly and carefully strip gel-side-down on the “protein sample solution” in the rehydration tray; and distribute the “protein sample solution” evenly along the whole strip length under the IPG strip; avoid bubbles. Overlay strip with 3–4 ml of mineral oil. Rehydrate the strip overnight (c.18 h; room temperature). 3. The first-dimension, isoelectric focusing (IEF) is performed on an Amersham Multiphor II Electrophoresis system (see steps 4–10). 4. Assemble the Amersham Multiphor II instrument, connect the red bridging cable, and set the Multitemp water circulator to 20°C. 5. Add 6–7 ml of mineral oil to the center of the cooling plate. Position the Drystrip tray onto the mineral oil on the cooling plate (the red wire towards the rear); distribute mineral oil evenly between the Drystrip tray and the cooling plate (no bubbles). Insert red electrode wire (anode, +) into the rear receptacle; connect black electrode to the pin in the front. 6. Add 12 ml of mineral oil into the Drystrip tray. Place immobiline strip aligner (plastic sheet with grooves up) on top of the oil in the Drystip tray; distribute evenly the mineral oil between plastic sheet and the Drystrip tray (avoid bubbles; avoid getting mineral oil on top of plastic sheet); and line up grooves with the pattern on the cooling plate. 7. Take the rehydrated IPG strip out of the rehydration tray, rinse fully with deionized distilled water (see Note 22), drain. Place the IPG strip (gel-side-up; pointed end (acidic/anode) towards the rear) into a groove in the strip aligner. An immobiline strip aligner can analyze up to 12 IPG strips at a time. 8. Cut two electrode paper strips (length = 110 mm), and moisten each electrode strip with 0.5 ml of distilled water. Blot the excess water on the electrode paper strips with paper tissue to make them slightly damp. Place the two moistened electrode paper strips on the gel surface of those aligned IPG strips (one across the cathode and one across the anode edges) (see Note 23). Position the electrodes onto the electrode paper strips (Red electrode goes to the rear right, black to the front left). 9. Pour 70–80 ml of mineral oil to completely cover the IPG strips. Place the lid onto the Multiphor II instrument, and connect to Amersham EPS3500XL power supply. The IEF parameters are: Step 1: 100 V, 2 mA, 5 W, 1 min gradient; Step 2: 100 V, 2 mA, 5 W, 2 h set; Step 3: 500 V, 2 mA, 5 W, 1 min gradient; Step 4: 3,500 V, 2 mA, 5 W, 1.5 h gradient; and Step 5: 3,500 V, 2 mA, 5 W, 8 h set. The total time is 11 h 32 min, Vh c.31,000.

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10. After IEF, disassemble the unit, remove IPG strips, gently blot off mineral oil, wrap loosely in plastic wrap, and store at −80°C (see Note 24). 11. The second-dimension, SDS-PAGE is performed on a Bio-Rad PROTEAN plus® Dodeca™ cell vertical electrophoresis system (see steps 12–19). 12. Cast 12% PAGE resolving gel with a Bio-Rad PROTEN-plus Multicasting chamber: Assemble the PROTEIN-plus Multicasting chamber system. For 12 gels, mix 180 ml of 40% w/v acrylamide/bisacrylamide stock solution (29:1), 150 ml of 1.5 M Tris-HCl pH 8.8, 270 ml deionized distilled water; de-gas with vacuum pump for 10 min. Add 3 ml of 10% ammonium persulfate, and 150 µl TEMED to the mixture solution (mix gently and avoid any bubbles). Gently pour the solution into holding chamber, and connect the holding chamber to the inlet port on multicasting chamber. Place a gel comb in the first gel cassette. Elevate the holding chamber above the level of multi-casting chamber to fill the gel solution up to the level of the comb. Remove the comb, and overlay the gels with deionized distilled water immediately. Allow the gels to polymerize for >1 h (see Note 25). 13. Connect Dodeca cell tank to carboy that holds 25 L electrophoresis buffer to allow electrophoresis buffer to flow into the Dodeca buffer tank. Set the circulating water bath temperature to 15°C, and turn on the circulator at least 1 h prior to the actual run. 14. Take out the frozen focused IPG strips, stand at room temperature for 3 min, and put them (one in each slot) in the equilibrium tray (gel-side-up). Add c.4 ml of reducing equilibrium buffer, and rock gently for 10 min. Replace immediately the reducing equilibrium buffer with c.4 ml of alkylation equilibrium buffer for each slot, and rock gently for 10 min. 15. During equilibrium, discard the Multi-casting chamber. Remove the prepared gel plate, and rinse the top surface of gel with deionized distilled water three times. Absorb excess water on the top surface of gel with a KimWipe paper. Position the prepared gel plate on the gel-stander. 16. Remove the equilibrated IPG strip, rinse once in the electrophoresis buffer in a 20-cm length of cylinder; remove the redundant electrophoresis buffer on the surface of IPG strip. Position the IPG strip over the top of resolving gel, gel-side facing front, plastic back contacting the glass plate, the pointed-end of IPG strip to the left and the square end to the right. Load quickly ca. 3 ml of 1% agarose solution (c.80°C) onto the top of the resolving gel, push quickly the IPG strip into the un-polymerized agarose solution, and align the up-side of IPG strip onto the top of the shorter glass plate. Polymerize in c.10 min. 17. Use two hands to insert the prepared gel plate hinge side down (PROTEIN plus hinged spacer plate) near the bottom of the tank; the IPG stip-end of gel is positioned next to the cathode (black electrode card; -) so that the sample migrates toward the anode (red electrode card; +); and make sure that the gasket is flared out toward the electrode. Adjust the level of electrophoresis buffer to the middle of the top spacer (see Note 26). 18. Place the lid on the tank, and connect the pump tubing to the top of the lid. Set the Buffer Recirculation Pump to 100 and turn it on. Connect the Dodeca Cell to

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the PowerPac 200 power supply. Electrophoresis is run under the conditions of constant voltage 200 V for 370 min. After the electrophoresis, disassemble the Dodeca cell, take out the PROTEAN plus hinged spacer plate on the benchtop, short plate facing upward and the hinge to the left. Insert the gel releaser between the short and the long plate at the top right corner. Pull the gel releaser up, lift the short plate until the gel cassette is opened completely (180°). Separate gently the gel from the plate, and prevent the gel from tearing (see Note 27). The separated protein on the 2DGE gel can be visualized with development reagent, such as silver stain (see step 21) and Gel-code blue (see step 9 in Subheading 3.3), or transfer to PVDF membrane for Western blot analysis (see step 22 and forward). After electrophoresis, the gel is taken out, followed by silver staining: (1) Fix gel in 250 ml of (50% v/v methanol and 5% v/v acetic acid) (20 min); wash in 250 ml of 50% v/v methanol (10 min); wash in deionized water (10 min). (2) Sensitize the gel in 250 ml of 0.02% w/v sodium thiosulfate (1 min); wash with deionized water (1 min, 2 times). (3) Silver reaction (20 min) in 250 ml of the solution (0.1% w/v silver nitrate plus 200 µl 37% v/v formaldehyde prior to use); wash with deionized water (1 min, 2 times). (4) Develop in 250 ml of 3%w/v sodium carbonate with 100 µl 37% v/v formaldehyde (add prior to use) until the desired intensity of staining occurs (usually c.3 min). (5) Stop the development within 250 ml of 5% v/v acetic acid (10 min); wash the gel in deionized water (5 min). (6) Store the gel in 250 ml of 8.8% glycerol. After electrophoresis, remove the 2D gel and orientate the 2D gel by removing the left-upper corner (acidic end), and soak the 2D gel in the gel equilibration buffer for at least 10 min. Preparation of PVDF membrane: Cut a sheet of PVDF membrane to match the gel size (20 × 17.2 cm), and place into 100% methanol for 10 min, wash with destilled water for 5 min, and check its hydrophilic side (see Note 28). Use pencil to write the date on the margin of hydrophilic side of PVDF membrane. Equilibrate in the PVDF membrane equilibration buffer for at least 10 min (see Note 29). The transfer is performed on an Amersham Multiphor II semi-dry electrotransfer system. Assemble the electroblotting cassette: (1) saturate anode electrode plate with deionized destilled water, and remove excess water with paper. Put it onto the buffer tank. (2) Immerse 6 filter papers in anode transfer buffer R, and place them carefully onto the anode plate. (3) Soak 3 filter papers in anode transfer buffer S, and place them carefully on the top of the first 6 filter papers. (4) Wet PVDF membrane in anode transfer buffer S for 30 s, and place PVDF (hydrophilic side up) onto the layer of second 3 sheets of filter papers. (5) Put the 2D gel onto the PVDF membrane. (6) Immerse 9 filtering papers in cathode transfer buffer T, and place them carefully on the top of the gel. (7) Saturate the cathode electrode plate with deionized distilled water, and remove excess water with a filter paper, place it on the 9 filtering papers. (8) Connect all units of the transfer system (see Note 30).

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25. Close the lid and connect to the power supply (Amersham EPS 3501XL). Electrotransfer is performed at a constant current of 0.8 mA/cm2 for 1 h 40 min. After transfer, remove the top filtering papers and draw a line around the 2D gel on the margin of PVDF membrane. 26. PVDF membrane (protein side up) is blocked in 100 ml of 0.3% BSA/PBST at room temperature for 60 min with gentle shaking. After blocking, the PVDF membrane is rinsed twice with water. 27. Add 100 ml of diluted primary antibody (100 µl rabbit anti-nitrotyrosine anbibody is diluted with 100 ml of 0.3% BSA/PBST). Incubate for 1 h at room temperature with gentle shaking. Pour out the primary antibody solution. Wash with 200 ml PBST with shaking (15 min; 4 ×). Rinse twice with deionized distilled water. 28. Add 100 ml of diluted secondary antibody (20 µl goat antirabbit alkaline phosphase-conjugated IgG is diluted in 100 ml of 0.3% BSA/PBST). Incubate at room temperature for 1 h with gentle shaking. Pour out the secondary antibody solution. Washing with 200 ml of PBST with shaking (15 min; 3 ×). Wash with 200 ml of PBS with shaking (15 min; 3 ×). Rinse 4 times with water (see Note 20). 29. The blot proteins are visualized by adding 1-Step™ NBT/BCIP substrate to cover the PVDF membrane (protein-side up) with gentle shaking until the desired color is obtained. Wash in deionized destilled water for 10 min. Dry the PVDF in the air or between two sheet of filter papers. Store between two leaves of transparent plastic sheet. 30. Scan the visualized PVDF membrane and the corresponding silver-stained 2D gel into a digitized image. Import the digitized image into the Bio-Rad PDQuest 2D gel image analysis software to generate the 2D gel spots and the PVDF blot spots. Match the immunopositive Western blotting spot to the corresponding silverstained 2D gel spot (see Figure 1) (see Note 31).

3.5. Immunoprecipitation of Nitrotyrosine-Containing Proteins 1. Protein extraction for immunoprecipitation: A portion (e.g., c.62 mg wet weight tumor tissue) of pituitary tissue in 1.5-ml Eppendof tube is rinsed 3 times with binding/washing buffer to remove blood from the tissue surface. Add 600 µl of Pierce M-PER® mammalian protein extraction buffer that is compatible with immunoprecipitation (10:1 = buffer:tissue), votex 5 min, homogenize 5 min, sonicate 20 s, rotate 2 h, sonicate 20 s, and centrifuge at 13,000 g for 30 min. The supernatant is the extracted protein sample, which is transferred to a new tube. The protein content is measured with the Bio-Rad Bradford Protein Assay reagent. 2. Immunoprecipitation of nitrotyrosine-containing proteins is carried out with a Pierce Seize X mammalian immunoprecipitation kit. Equilibrate Immobilized Protein G, Anti-nitrotyrosine antibody, and binding/washing buffer to room temperature. 3. Gently swirl the bottle of ImmunoPure Immobilized Protein G beads to resuspend fully the beads, and add 400 µl of beads into a 0.5-ml Handee spin-cup column that is placed inside a Handee microcentrifuge tube. Centrifuge at 3,000 g for 1 min. Discard the flow-through. Wash the beads with 400 µl of binding/washing buffer

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(invert tube 10 times, centrifuge 3,000 g, 1 min) for 2 times. Put the spin-cub into a new microcentrifuge tube. Add 300 µl binding/washing buffer and 100 µl (= 100 µg) anti-nitrotyrosine antibody, and invert quickly the spin-cup to mix the antibody and beads. Incubate at room temperature for 1 h with gently rotating to allow antibody to bind the protein G. Centrifuge at 3,000 g for 1 min and discard the flow-through. Wash with 500 µl binding/washing buffer (invert 10 times, centrifuge at 3,000 g for 1 min) for 3 times. Put the spin-cup into a new microcentrifuge tube. Dissolve 2 mg disuccinimidyl suberate (DSS) in 80 µl dimethyl sulfoxide (DMSO). Dilute 25 µl DSS with 400 µl binding/washing buffer, and add 425 µl diluted DSS solution to the washed beads with immobilized protein G antibodies. Invert the tube quickly and incubate for 30 min with gently rotating to allow the antibodies to crosslink with the immobilized protein G (see Note 14). Centrifuge at 3,000 g for 1 min, and discard the flow-through. Wash the beads with crosslinked protein G-antibodies with 500 µl of elution buffer (pH 2.8) (invert 10 times, centrifuge at 3,000 g for 1 min) for 5 times. Put the spincup into a new microcentrifuge tube. Wash the antibody-protein G-beads with 500 µl binding/washing buffeer (invert tube 10 times, centrifuge 3,000 g for 1 min) for 4 times. Dilute the extracted pituitary protein samples with binding/washing buffer (at least 1:1 dilution). Add 500 µl of diluted sample to the prepared antibody-protein G-beads, mix quickly by inverting the tube. Incubate overnight (gently rocking, 19 h, 4°C) to bind nitrotyrosine-containing proteins with anti-nitrotyrosine antibodies. Centrifuge at 3,000 g for 1 min, and discard the flow-through. Wash the beads with nitroprotein with 400 µl binding/washing buffer (invert tube 10 times, centrifuge 3,000 g for 1 min) for 3 times to remove any nonbound proteins. Elute the bound nitroproteins with 200 µl elution buffer (pH 2.8) that contains a primary amine (gently mix well, centrifuge 3,000 g, 1 min) for 3 times. The eluates that contain nitroproteins are collected, add 10 µl of the pH-neutralized solution per 200 µl eluate. If necessary, the eluate may be dried partially to elevate the concentration of nitroprotein.

3.6. Tandem Mass Spectrometry Characterization of Nitrotyrosine-Containg Proteins 1. Trypsin digestion of immunoprecipitated nitroprotein: The immunoprecipitated protein is digested with trypsin (see Note 32). The procedure (14) is: (a) To a vial of sequencing grade-modified trypsin (20 µg, Promega, Madison, WI), add 5µl trypsin resuspension buffer that contains 50 mM acetic acid (pH 2.8) and 95 µl of 200 mM NH4HCO3 (pH 8.2), and mix fully. (b) Add 20 µl of the eluate that contains nitroproteins into a 0.5-ml siliconized tube (see Note 33). (c) Add 1µl of solution that contains 1 M Tris (pH 9.5), 100 mM dithiothreitol, and 100 mM iodoacetamide.

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(d) Add 25 µl trypsin solution and 54 µl ddH2O, mix fully to form the enzyme digestion reaction system (100 µl, final concentration of NH4HCO3 = 50 mM, pH 8.1). Incubate overnight (37°C, 23 h). 2. Trypsin digestion of in-gel nitroprotein: Excise the protein spot that corresponded to the nitrotyrosine immunopositive spot from the 2-D gels, and digest with trypsin (see Note 32). The procedure (20) is: (a) Cut the silver-stained gel spots and place into a 1.5 mL siliconized tube (see Note 33). Wash 6 times with 500 µl of deionized water. Transfer the gel into another 1.5-mL siliconized tube, and mince it into several pieces (c.0.5–1 mm3) with a pipette tip. (b) Pior to use, prepare the destaining solution by mixing 30 mM potassium ferricyanide with 100 mM sodium thiosulfate (1:1 v/v). Add 20 µl of fresh destaining solution to destain the gel pieces until the brownish color disappears (c.1–2 min). Remove the destaining solution, and wash 5–6 times the gel pieces with 20 µl water until the yellow color disappears. (c) Incubate the gel pieces for 20 min in 20 µl of 200 mM ammonium bicarbonate. Discard the buffer, and wash once the gel pieces with 20 µl water. (d) Dehydrate the gel pieces with 30 µl acetonitrile for 2 times until the gel pieces turn an opaque white. Dry in a vacuum centrifuge for 30 min at 30°C. (e) Add 100 µl of trypsin resuspension buffer to a vial of lyophilized trypsin powder (20 µg, 833 pmol) (Promega V5111), and mix fully. Dilute the trypsin solution (20 µl, 200 ng/µl) to 16 ng/µl with 230 µl of 50 mM ammonium bicarbonate. (f) Add 20–30 µl (0.32–0.48 µg) of the diluted trypsin solution to each dried gel piece. The gel piece should be covered by the digestion solution. Incubate (37°C, c.18–20 h), cool (4°C, 30 min). (g) Gently centrifuge for 10 s, sonicate in a water bath (30°C) for 5–6 min, and centrifuge at 12,000 g for 2 min. Transfer carefully the supernatant into a 0.5-ml siliconized tube. (h) Add 10–15 µl of 50 mM ammonium bicarbonate to the gel pieces to repeatedly extract (2 ×) tryptic peptides (step 7). Combine all supernatants (the tryptic peptide mixture). 3. The tryptic peptide mixture from a 2-D gel spot or 20–40 µl tryptic peptide mixture from immunoprecipitated proteins is purified with a ZipTipC18 beads microcolumn (Millipore ZTC18S096, Bedford, MA). Set up the pipette to the 10-µl scale. Wash 5 times the beads with 10 µl acetonitrile, and 5 times with 10 µl of 50% acetonitrile. Equilibrate 5 times with 10 µl of 0.1% TFA. Bind the tryptic peptides by pipetting the sample up and down 15 times. Wash twice with 10 µl of 0.1% TFA. 4. Elute the purified tryptic peptide mixture with 6 µl of 85% v/v acetonitrile/0.1% v/v TFA by pipetting up and down for 10 times. The eluate is air-dried. Prior to MS analysis, redissolve the dried tryptic peptide mixture with 6 µl of 2% actonitrile/0.5% acetic acid. 5. The tandem mass spectrometry analysis of the purified tryptic peptide mixture is carried out with a capillary liquid chromatography (LC)- electrospray

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ionization (ESI)-quadropole ion-trap (Q-IT) mass spectrometer system (LCQDeca; ThermoFinnigan, San Jose, CA). 6. Set the LCQDeca instrument parameters as follows: ESI voltage 2.0 kV, capillary probe temperature 110°C, and electron multiplier −900 V. Use a solution (1 pmol/ul) of synthetic des-arg bradykinin to determine the instrument sensitivity and mass accuracy. In the MS mode, the (M + 2H)2+ ion of des-arg bradykinin at m/z 452.7 should have a signal intensity of 1 × 107 (arbitrary units) at a flow-rate of 0.5 µl/min. In the MS/MS mode, the precursor ion at m/z 452.7 should produce m/z 404.2, 710.4, and 807.4. Acquire the MS/MS data in the “triple-play” datadependent scan mode that includes four scan events: a full-range MS scan, and three MS/MS scans that depend on the “triple-play” data scan mode. 7. Inject manually 6 µl tryptic peptide mixture into the LC system. Separate the peptide mixture on a capillary column: New Objective PicoFrit 360 µm (OD), 75 µm (ID), 15 µm tip pores (ID) packing an 8 cm length Magic C18AQ material material (5 µm beads, 200 Å pores) (Michrom Bioresources, Auburn, CA). Elute (35 µl/ min flowrate) peptides with the following five gradients (Solution A = water + 0.1% v/v formic acid; Solution B = 90% v/v acetonitrile + 0.1% v/v formic acid): (1) 100% solution A and 0% solution B (5 min); (2) linear gradient up to 35% solution A and 65% solution B within 30 min; (3) maintain 35% solution A and 65% solution B (15 min); (4) linear gradient down to 100% solution A and 0% solution B within 5 min; and (5) maintain 100% solution A and 0% solution B. The peptides elute directly into the ESI source. All MS/MS data are acquired and managed with the ThermoFinnigan Xcalibur software. 8. Identification of a nitrotyrosine-containing protein. Use amino acid sequence to identify the protein by searching the SWISS-PROT and NCBInr databases with the SEQUEST software that is a part of the LCQDeca software package. Mass modifications of +45 kDa (+ NO2–H) at Tyr, of +57 kDa (+NH2COCH2–H) at Cys, and of +18 kDa at Met are considered in the search. Each positive search result – nitration of a Tyr residue – is confirmed with a manual check of the original LC, MS, and MS/MS data to determine each nitration site. During the analysis of those nitrated proteins, the following experimental criteria are applied: K or R at the C-terminus; K, R, or D (1) preceding the N-terminus; 0 or 1 missed trypsin cleavage sites; singly charged product b- and y-ions, and Homo sapiens. De novo sequencing is used to independently and accurately determine each amino acid sequence. The amino acid sequence from de novo sequencing is used to search the human SWISS-PROT protein database with the SIB BLAST search engine (http:// us.expasy.org/tools/blast/). An example (Fig. 2) demonstrates the identification of a nitrotyrosine-containing protein.

4. Notes 1. The water that is used to make all buffers/solutions and to wash gels should be deionized distilled (dd) water that has a resistivity of 18.2 MΩ−cm. We often use Millipore water.

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2. Tetranitromethane is an oxidizing reagent; very toxic; irritating to the eyes, respiratory system, and skin with very serious irreversible effects. Experimental gloves should be worn, and one should operate in the hood. 3. Butylated hydroxytoluene (BHT) is an effective antioxidant to protect a sample from oxidation. 4. Acrylamide and bisacrylamide in the monomeric form are neurotoxins. Avoid inhaling and exposing to skin. Dispose ecologically of the remains by polymerizing the remains with an excess of ammonium persulfate (21,22). 5. Use fresh ammonium persulfate solution. Fresh ammonium persulfate “crackles” when water is added. It it does not, replace it with fresh stock. 6. High temperature will decompose urea. Work at < to 30°C. 7. For “protein extraction buffer” and “rehydration buffer”, generally a stock solution (7 M urea, 2 M thiourea, 4% (w/v) CHAPS, and trace bromophenol blue) is made, and stored in 1 ml aliquots at −80°C. Prior to use, the protein extracting buffer is made by adding 15.4 mg DTT (final concentration = 100 mM) and 20 µl pharmalyte (final concentration = 0.5%) to 1 ml of rehydration stock solution, and mix well. The rehydration buffer is made by adding 9.2 mg DTT (final concentration = 60 mM) and 5 µl Pharmacia IPG buffer (final concentration = 0.5%) to 1 ml of rehydration stock solution, and mix well. Thus, DTT, pharmalyte, and IPG buffer will be fresh. 8. For “reducing equilibration buffer” and “alkylation equilibration buffer,” generally an SDS equilibration stock buffer (375 mM Tris-HCl pH 8.8, 6 M urea, 2% (w/v) SDS, 20% (v/v) glycerol, and trace bromophenol blue) is made, and stored in aliquots of 50 ml at −80°C. Prior to use, the reducing equilibration buffer is made by adding 1 g DTT (final concentration = 2% w/v) to 50 ml of SDS equilibration stock buffer, and mixed well. The alkylation equilibration buffer is made by adding 1.25 g iodoacetamide (final concentration = 2.5% w/v) to 50 ml of SDS equibration stock buffer, and mix well. Thus, DTT and iodoacetamide will be fresh. 9. SDS is a strong detergent that irriates the respiratory system. Avoid inhaling the SDS powder when SDS is weigh. 10. The Agarose solution should not be boiled over. If the solution turns light yellow, it should be replaced with a fresh agarose solution. 11. There are commonly two type of Immobilon-P transfer membrane: 0.45 µm and 0.2 µm. The 0.45-µm PVDF membrane is often used to transfer the protein, whereas the 0.2-µm PVDF membrane is often used to transfer small molecular weight proteins or peptides. If a 0.2-µm PVDF membrane is used to transfer protein, then there will be more resistance that decreases the transfer efficiency. 12. The anti-nitrotyrosine antibody is currently a commercial product that can be obtained from Upstate Biotechnology, International Chemicomn, and

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16. 17.

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Sigma, etc. A polycolonal or monocolonal antinitrotyrosine antibody can be obtained. Pierce Immobilized protein G plus cross-linked 6% beaded agarose supplied as a 50% slurry (for example, 400 µl of settled gel is equivalent to 200 µl of 50% slurry) that contains 0.02% sodium azide. Binding capacity is > 20 mg human IgG per ml of settled gel. Disuccinimidyl suberate (DSS) is a very strong crosslinker, whose activity is much stronger than DMP, another commonly used crosslinker. DSS is water-insoluble and should be first dissolved in an organic solvent such as DMSO or DMF, and added to the aqueous reaction mixture. DSS is moisture-sensitive. Store at 4–8°C in a desiccator. To avoid moisture condensation onto the product, the vial that contains DSS must be equilibrated to room temperature before opening. Prepare DSS solution just prior to use because DSS is readily hydrolyzed and becomes non-reactive. Overcrosslinking proteins with biological activity (e.g., enzymes, antibodies, etc.) could result in loss of activity possibly because of a conformational changes of the molecule. Activity loss also may occur when DSS modifies lysine groups that are involved in binding substrate or antigen. Adjusting the molar ratios of reagent to the target may overcome activity loss. Alternatively, use a cross-linker that targets a different functional group. Hydrolysis of DSS is a competing reaction and increases with increasing pH. Hydrolysis occurs more readily in dilute protein or peptide solutions. In concentrated protein solutions, the acylation reaction is favored. Promega’s Sequencing Grade Modified Trypsin is porcine trypsin modified by reductive methylation. Its resistance to proteolytic digestion is two times greater activity than unmodified trypsin. Sequencing Grade modified trypsin is further treated with TPCK followed by affinity purification to yield a highly active and stable molecule. Modified trypsin has maximal activity at pH 7–9, and is reversibly inactivated at pH 4. It is resistant to mild denaturing conditions: 0.1% SDS, 1 M urea, or 10% acetonitrile. Modified trypsin retains 48% activity in 2 M guanidine HCl. It is important to wash the blood from the tissue surface. Care should be taken with small tissue samples because of loss of tissue. In order to obtain consistent results among different samples when comparing gels, a lyophyilized pituitary is redissolved in a solution of 8 M urea and 4% CHAPS for BCA protein concentration analysis because BCA reagent is not compatible with thiourea, DTT, IPG buffer, and bromophenol blue. For different experiment, each time use the protein concentration of a fixed sample to verify the protein concentration of other samples. The standard curve is an inverted “S”-like curve. The sample points that correspond to the linear section of the inverted “S”-like curve are used to determine the nitrotyrosine concentration of sample.

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19. PVDF membrane with protein can be kept in blocking solution for 2–3 days at 4°C with gently shaking. 20. After blocking, incubation with primary antibody, and incubation with secondary antibody, wash fully because salt and Tween-20 could result in particles on the visualized PVDF that would increase the background. 21. The size of the commercially Amersham immobilized pH gradient dry strip is 0.5-mm thick and 3-mm wide with different length (7, 11, 13, 18, and 24 cm). The pH range is 3–10, 4–7, 6–9, 4–5, etc., with a linear pH gradient or non-linear pH gradient. The pH range of the IPG buffer should be consistent with the pH range of the IPG strip; for example, IPG buffer pH 3–10 NL matches the IPG stip pH 3–10 NL, IPG buffer pH 3–10 matches the IPG strip pH 3–10 (22). 22. It is necessary for a rehydrated IPG strip to be rinsed fully with ddH2O to remove excess mineral oil and undissolved components. 23. The 110-cm long electrode paper strips not only conduct but also remove the salt from samples. The latter significantly improves the IEF effect because tissue samples all contain salts that will affect the IEF. If the salt is too high, during the IEF, the electrode paper strips may be replaced once with a new electrode strip. 24. After IEF, use only water and liquinox detergent (hot water works nicely to remove mineral oil) to clean electrodes, Dry strip tray, and strip aligner. Use paper towel to clean the cooling plate. 25. The cast gels can be used within a week when they are covered with moist filter paper and stored at 4°C. Moreover, for our 2-D gel studies, the reproducibility of a single-concentration gel is better than a gradient gel (23,24). 26. If the gel plate is completely immersed in buffer, then there will be a significant current leak that negatively affects the electrophoresis results. If the buffer level is not high enough to cover the entire gel area, then the end of the gel will show no separation. 27. After electophoresis, use only water to wash the Dodeca tank, gaskets, electrode cards, and lid. For long-term storage, flush all parts thoroughly with water to completely remove any residual buffer. 28. A PVDF membrane has two different sides: hydrophobic and hydrophilic. The water moves slowly on the hydrophilic side, and fast on the hydrophobic side. The protein should be transferred to the hydrophilic side of PVDF. 29. A PVDF membrane must always be kept wet; if it dries, then re-wet in methanol and water. 30. More attention should be paid to avoid any air bubbles in steps 1 to 7) because they severely affect the semi-dry electrotransfer. 31. Our studies (23,24) have demonstrated the high levels of reproducibility and resolution of 2DGE. For the first dimension IEF, a high reproducibility

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can be obtained with commercially available IPGstrips. For the seconddimensional SDS-PAGE, different types of gel system are available: vertical versus horizontal gel systems; constant-percentage versus gradient gels. Compared to the horizontal Multiphor gel system (pre-cast gradient SDS gel = 180 × 245 × 0.5 mm; analyzes a gel at a time), the vertical Dodeca gel system (single-concentration gel = 190 × 205 × 1 mm; analyze up to 12 gels at a time) demonstrates a better spatial and quantitative reproducibility (23), and a wider linear dynamic range to measure the change in the protein abundance in a complex proteome (24). Moreover, for betweengel comparison, the same experiment conditions (including the loading amount of protein, sample-dissolving solution, IEF, IPGstrip equilibration solution, SDS-PAGE, and gel image process) should be used to conduct two-dimensional gel electrophoresis experiments and image analysis for each experiment (25). 32. In these experiments, especially for digestion with trypsin, gloves and a head cap are always used to avoid any keratin from skin and hair contamination of the protein sample (20). 33. All tubes and pipette tips that contact protein sample or peptides should be siliconized (or low-retention) to avoid any loss of protein or peptide. Acknowledgments The authors acknowledge financial support from NIH (NS 42843 to DMD; RR 16679 to DMD).

References 1. Zhan, X. and Desiderio, D. M. (2004) The human pituitary nitroproteome: detection of nitrotyrosyl-proteins with two-dimensional Western blotting, and amino acid sequence determination with mass spectrometry. Biochem. Biophys. Res. Commun. 325, 1180–1186. 2. Lloyd, R. V., Jin, L., Qian, X., Zhang, S., and Scheithauer, B. W. (1995) Nitric oxide synthase in the human pituitary gland. Am. J. Pathol. 146, 86–94. 3. Ueta, Y., Levy, A., Powell, M. P., Lightaman, S. L., Kinoshita, Y., Yokota, A., Shibuya, I., and Yamashita, H. (1998) Neuronal nitric oxide synthase gene expression in human pituitary tumours: a possible association with somatotroph adenomas and growth hormone-releasing hormone gene expression. Clin. Endocrinol. (Oxf.) 49, 29–38. 4. Ceccatelli, S., Hulting, A. L., Zhang, X., Gustafsson, L., Villar, M., and Hokfelt, T. (1993) Nitric oxide synthase in the rat anterior pituitary gland and the role of nitric oxide in regulation of LH secretion. Proc.Natl. Acad. Sci. USA 90, 11292–11296. 5. McCann, S. M., Karanth, S., Mastronardi, C.A., Dees, W. L., Childs, G., Miller, B., Sower, S., and Yu, W. H. (2001) Control of gonadotropin secretion by folliclestimulating hormone-releasing factor, luteinizing hormonereleasing hormone, and leptin. Arch. Med. Res. 32, 476–485.

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6. McCann, S. M., Haens, G., Mastronardi, C., Walczewska, A., Karanth, S., Rettori, V., and Yu, W. H. (2003) The role of nitric oxide (NO) in control of LHRH release that mediates gonadotropin release and sexual behavior. Curr. Pharm. Des. 9, 381–390. 7. Pinilla, L., Gonzalez, L. C., Tena-Sempere, M., Bellido, C., and Aguilar, E. (2001) EVects of systemic blockade of nitric oxide synthases on pulsatile LH, prolactin, and GH secretion in adult male rats. Horm. Res. 55, 229–235. 8. Duvilanski, B. H., Zambruno, C., Seilicovich, A., Pisera, D., Lasaga, M., Diaz, M. C., Belova, N., Rettori, V., and McCann, S. M. (1995) Role of nitric oxide in control of prolactin release by the adenohypophysis. Proc.Natl. Acad. Sci. USA 92, 170–174. 9. Bocca, L., Valenti, S., Cuttica, C.M., Spaziante, R., Giordano, G., and Giusti, M. (2000) Nitric oxide biphasically modulates GH secretion in cultured cells of GHsecreting human pituitary adenomas. Minerva Endocrinol. 25, 55–59. 10. Cuttica, C.M., Giusti, M., Bocca, L., Sessarego, P., De Martini, D., Valenti, S., Spaziante, R., and Giordano, G. (1997) Nitric oxide modulates in vivo and in vitro growth hormone release in acromegaly. Neuroendocrinology 6, 426–431. 11. Pinilla, L., Tena-Sempere, M., and Aguilar, E. (1999) Nitric oxide stimulates growth hormone secretion in vitro through a calcium- and cyclic guanosine monophosphate-independent mechanism. Horm. Res. 51, 242–247. 12. Riedel, W. (2002) Role of nitric oxide in the control of the hypothalamic – pituitary – adrenocortical axis. Z. Rheumatol. 59, 36–42. 13. Zhan, X. and Desiderio, D. M. (2007) Linear ion-trap mass spectrometric characterization of human pituitary nitrotyrosine-containing proteins. International J. Mass Spectrom. 259, 96–104. 14. Zhan, X. and Desiderio, D. M. (2006) Nitroproteins from a human pituitary adenoma tissue discovered with a nitrotyrosine affinity column and tandem mass spectrometry. Analytical Biochem. 354, 279–289. 15. Khan, J., Brennan, D. M., Bradley, N., Gao, B., Bruckdorfer, R., and Jacobs, M. (1998) 3-nitrotyrosine in the proteins of human plasma determined by an ELISA method. Biochem. J. 330, 795–801. 16. Torreilles, J. and Romestand, B. (2001) In vitro production of peroxynitrite by haemocytes from marine bivalves: C-ELISA determination of 3-nitrotyrosine level in plasma proteins from mytilus galloprovincialis and crassostrea gigas. BMC Immunology 2, 1. 17. Aulak, K. S., Miyagi, M., Yan, L., West, K. A., Massillon, D., Crabb, J. W., and Stuehr, D. J. (2001) Proteomic method identifies protein nitrated in vivo during inflammatory challenge. Proc.Natl. Acad. Sci. USA. 98, 12056–12061. 18. Miyagi, M., Sakaguch, H., Darrow, R. M., Yan, L., West, K. A., Aulak, K. S., Stuehr, D. J., Hollyfield, J. G., Organisciak, D. T., and Crabb, J. W. (2002) Evidence that light modulates protein nitration in rat retina. Mol. Cell. Proteomics 1, 293–303. 19. Gu, X., Meer, S. G., Miyagi, M., Rayborn, M. E., Hollyfield, J. G., Crabb, J. W., and Salomon, R. G. (2003) Carboxyethylpyrrole Protein Adducts and Autoantibodies, Biomarkers for Age-related Macular Degeneration. J. Biol. Chem. 278, 42027–42035.

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20. Zhan, X. and Desiderio, D. M. (2003) A refernce map of a human pituitary adenoma proteome. Proteomics 3, 699–713. 21. Westermeier, R. (1997) Electrophoresis in Practice: A guide to Methods and Applications of DNA and Protein Separations, 2nd ed. VCH Wiley Company Press, Weinheim, Federal Republic of Germany. 22. Zhan, X. (2005). Two-Dimensional Electrophoresis, in Experimental Protocols for Medical Molecular Biology in Chinese and English (Zheng, W., ed.), Peking Union Medical College Press, Beijing, China, pp. 93–108. 23. Zhan, X. and Desiderio, D. M. (2003) Differences in the spatial and quantitative reproducibility between two second-dimensional gel electrophoresis. Electrophoresis 24, 1834–1846. 24. Zhan, X. and Desiderio, D. M. (2003) Spot volume vs. amount of protein loaded onto a gel. A detailed, statistical comparison of two gel electrophoresis systems. Electrophoresis 24, 1818–1833. 25. Zhan, X. and Desiderio, D. M. (2003) Heterogeneity analysis of the human pituitary proteome. Clin. Chem. 49, 1740–1751.

157 Mass Spectrometric Determination of Protein Ubiquitination Carol E. Parker, Maria Warren Hines, Viorel Mocanu, Susanna F. Greer, and Christoph H. Borchers 1. Introduction Cellular homeostasis requires a delicate balance of protein synthesis and degradation, a balance which is maintained by the actions of regulatory proteins. In turn, proteins which are no longer required must be degraded in a rapid and selective manner. The selective degradation of proteins involved in diverse regulatory mechanisms such as signal transduction (1), transcriptional regulation (2), cell cycle regulation (3), and stress response (4) have been linked to the covalent attachment of ubiquitin to proteins. Ubiquitin is a 76-residue polypeptide, which attaches via its carboxy-terminus to lysine ε amino groups of the target protein. Ubiquitin-protein conjugates are short lived, primarily because of proteolysis by the 26S proteosome or, in some cases, dissociation of the complex with removal of the ubiquitin by ubiquitin isopeptidases (5). Other roles for monoubiquitination have recently been uncovered, including acting as a signal for protein trafficking, cell division, targeting proteins to subnuclear structures, endocytosis, signal transduction, and kinase activation (6–11). Both the dynamic nature of these important regulatory proteins, and the low protein levels in vivo, make analysis of protein ubiquitination inherently difficult. The use of mass spectrometry to identify sites of ubiquitination within target proteins in vivo will allow greater understanding of the ways in which ubiquitination alters protein function. Therefore, the development of broadly applicable identification approaches is critical. The importance of ubiquitination and the role of mass spectrometry in the study of ubiquitination has recently been reviewed by Kirkpatrick, Denison, and Gygi (12,13), Xu and Peng (14), and Drews, Zong, and Ping (15).

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2. Materials 2.1. MALDI Analysis of Intact Proteins 1. Bruker Reflex III or a Bruker Ultraflex matrix-assisted laser desorption (MALDIMS, (Bruker Daltonics, Billerica, MA). 2. α-cyano 2-hydroxycinnamic acid (Aldrich; St. Louis, MO). 3. Ethanol. 4. Water, HPLC-grade. 5. Formic acid.

2.2. LC/MS/MS 1. 2. 3. 4. 5. 6.

Waters/Micromass Q-TOF (Waters/Micromass Corp., Milford, MA). PepMap C1815 cm × 75 µm id capillary column (Dionex; Sunnyvale, CA). Trapping column 5 mm × 800 Å id C18 P3 (Dionex). Water, HPLC-grade (Pierce). Acetonitrile, HPLC-grade (Pierce). Formic acid.

2.3. Affinity Purification 1. 2. 3. 4. 5. 6.

Anti-HA antibody beads (Sigma, St. Louis, MO). Anti-FLAG antibody beads (Sigma). Ammonium bicarbonate. Ethanol. Water (Pierce). Formic acid (Fisher).

2.4. In-Solution Digestion 1. 2. 3. 4. 5. 6.

Water, deionized or HPLC-grade (Pierce). Trypsin, sequencing-grade (Promega; Madison, WI). GluC (Sigma). Ammonium bicarbonate. Low-retention Eppendorf tubes (Axygen; Union City, CA). Thermomixer (Eppendorf; Hamburg, Germany).

2.5. Lyophilization and Reconstitution 1. Freeze dryer (Labconco; Kansas City, MO). 2. Water (Pierce).

3. Methods 3.1. Evidence from Gels. Most direct evidence for ubiquitination comes from gel electrophoresis (Fig. 1), where a series of higher molecular weight bands are observed above the molecular weight of the protein, or from Western blot analysis using antiubiquitin antibodies (Fig. 2).

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Fig. 1. Ubiquitination evidence from Gels. PAGE gel showing a series of bands above the molecular weight of the non-ubiquitinated protein. (Collaborator: W. C. Patterson).

Fig. 2. Ubiquitination evidence from Western blot analyses. PAGE gel showing a series of bands above the molecular weight of the non-ubiquitinated protein: Western blot analysis of CIITA, showing ubiquitination.

3.2. Mass Spectrometric Evidence for Protein Ubiquitination While ubiquitination is clearly important for protein degradation, most mass spectrometric studies on ubiquitination have focused on protein phosphorylation, rather than direct mass spectrometric studies of protein ubiquitination. There are three types of mass spectrometric evidence for ubiquitination: the first is ubiquitination of the intact protein, the second is co-electrophoretic migration of the target protein and the attached ubiquitin and, and the third is the mass shift of a ubiquitinated peptide relative to the nonubiquitinated peptide. 3.2.1. Evidence from MS of the Intact Protein Direct mass spectrometric evidence of intact ubiquitinated HSP70 is shown in Fig. 3. This spectrum was obtained by direct MALDI-MS analysis of ubiquitinated HSP70, affinity-bound to anti-HSP70 beads.

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Fig. 3. Ubiquitination evidence from MALDI of intact proteins. MALDI-MS of HSP70 with a series of ubiquitin attached moieties.

3.2.1.1. DIRECT MALDI ANALYSIS OF PROTEINS BOUND TO AFFINITY BEADS (SEE NOTE 1) 1. Use antibodies covalently bound to the affinity beads (see Note 2). 2. Bind the antibody according to the bead manufacturers’ instructions (see Notes 3 and 4). 3. Rinse beads 3 times with 2–3 bead volumes of 100 mM ammonium bicarbonate (see Note 5). 3.2.1.2. SPOTTING THE MALDI TARGET 1. Prepare the MALDI matrix solution – a saturated solution of recrystallized (see Note 6) α-cyano 2-hydroxycinnamic acid in 45:45:10 ethanol:water:formic acid. (see Note 7). 2. Pipet 0.5 µL of settled beads onto the MALDI target, followed by 0.5 µL of MALDI matrix solution.

3.2.2. Evidence from MS and MS/MS of Peptides The second type of mass spectrometric evidence for ubiquitination comes from in-gel digestion and protein identification studies. In-gel digestion of the higher molecular weight bands followed by protein identification by MALDI-MS

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Fig. 4. Peptides from the target protein CIITA and peptides from ubiquitin from a gel band at mw ∼200 KDa.

or LC/MS/MS on a gel can sometimes provide evidence for ubiquitination – evidence comes from peptides rather than proteins. In the example shown in Fig. 4, peptides from the target protein CIITA and peptides from ubiquitin were found in a gel band of approximately 200 Kda, clearly higher than the unmodified 150 kDa CIITA or the 8.7 kDa unbound modified monoubiquitin (see Subheading 3.1.2.4). 3.2.3. MS and MS/MS Spectra of Ubiquitinated Peptides The third type of mass spectrometric evidence comes from the modified peptide itself, either from a shift in peptide molecular weight, or from MS/MS data. Ubiquitin is a 76-amino acid protein. An E3 ligase attaches ubiquitin to the ε amino group of a lysine residue in the target protein. This covalent linkage is formed at C-terminal glycine residue of the ubiquitin, with loss of the elements of water. Cleavage with trypsin or gluC (see Note 8) leaves characteristic “tails” on the modified lysine. These “tails” cause a shift in the molecular weight of the peptide, which can be used to distinguish these modified peptides from the unmodified peptides (Fig. 5). The actual site of ubiquitination can then be determined by MS/MS sequencing.

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Fig. 5. Schematic showing the expected peptide molecular weight shift caused by ubiquitination.

These peptide mass shifts can be used to find and sequence ubiquitinated peptides. Mascot, for example, allows the user to create a modification of a particular mass, which can then be used to search the peptide data from a given digest. Due to their transient nature and low natural abundances, ubiquitinated peptides are difficult to detect. A study by Gururaja et al (16) on Hela cell lysates, used a 6 × his-tagged ubiquitin, IMAC purification, digested by Lys-C and trypsin, and protein identification by 2D-LC/MS/MS using strong cation exchange and C18 reversed phase media. Ubiquitination was confirmed by anti-his tag and anti-ubiquitin Western blotting on the undigested lysates. A total of 244 proteins were found, which the authors categorized into functional groups, but determination of the exact ubiquitination sites was not the focus of this study. Very few ubiquitinated peptides were found by mass spectrometry from a single preparation until the work by the Gygi group in 2003 (17). As in the studies described above, a 6 × his-tagged ubiquitin was also used, and ubiquitinated proteins from yeast were purified by IMAC. The ubiquitin-enriched fraction (0.2 mg out of each original 100 mg of yeast cell lysate) was then digested with trypsin, fractionated into 80 fractions on a SCX column, and the 80 fractions were analyzed by capillary LC/MS/MS. Peptide MS/MS data was searched using Sequest software, allowing for a ubiquitin-modified lysine with a mass shift of +114 Da. In this manner, 110 ubiquitination sites were determined. In another study using 6 × his-myc-Ub, also from the Gygi group, 211 proteins were identified. Ubiqui-

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tination sites were identified on 15 of these proteins (18). A detailed description of this method has been reported by Peng and Cheng (19). Using a similar approach, Kirkpatrick et al (20) also studied the “ubiquitinome” of human cells, this time HEK293 cells. As in the above studies, 6 × his-tagged ubiquitin was used, followed by Ni-IMAC purification. Lysates from as many as 80 plates were pooled, and the eluted proteins were digested with trypsin, followed by LC/MS/MS. Twenty-two ubiquitinated proteins were identified, along with 19 additional proteins non-specifically-bound to the IMAC beads. An attempt was made to find the ubiquitinated peptides, but only branched peptides from polyubiquitin were identified in this study – by the presence of the GG modification on lysine. No consistent fragmentation (i.e., no loss of 114 Da) was observed in these GGK-modified peptides. Confirmatory evidence of the ubiquitination at these specific sites was the lack of tryptic cleavage at the modified K. An alternative, but still “shotgun” approach is to use an anti-ubiquitin antibody to accomplish the enrichment. This approach was used by Figeys’ group, and the digestion of 30 bands resulted in the identification of 70 proteins (21). Several GGK-containing “signature” peptides were located by their mass shifts, using a Mascot database search. An interesting and useful improvement on the antiubiquitin affinity purification approach was the use of the anti-polyubiquitin antibodies which had been developed in 1994 by Fujimuro et al (22). The use of one of these antibodies (FK2) instead of a conventional anti-ubiquitin antibody has been shown to work even in denaturing conditions and improves the selectivity of ubiquitinated protein enrichment by avoiding the capture of free ubiquitin (23,24). 3.2.3.1. MS/MS FRAGMENTATION OF UBIQUITINATED PEPTIDES

When the work described in this book chapter was initiated (2004), we could only find two reports of ubiquitination site determination which showed detailed MS/MS spectra of ubiquitinated peptides. These were on specificallytargeted proteins of interest – the first was the work of Laub et al (25), who used gluC for proteolytic digestion of the protein, and the second was the work of Dohlman’s group (26), who used trypsin. Both of these papers showed MS/MS spectra of the ubiquitinated peptides, and we decided to use these peptides as the starting point for a detailed study of the fragmentation of ubiquitinated peptides with the goal of finding specific diagnostic fragment ions to aid in detecting low levels of ubiquitinated peptides at lower levels in biological materials (27). When a ubiquitinated protein is enzymatically digested, a portion of the ubiquitin side chain remains attached to the modified lysine (Fig. 6). A ubiquitinated peptide therefore has 2 N-termini – one from the original peptide and one from the ubiquitin side chain. Thus, it is possible to have two series of b ions and y

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Fig. 6. Schematic showing fragmentation of ubiquitinated peptides.

ions. For the sake of clarity, we have chosen to refer to those b and y ions involving the ubiquitin side chain as b and y ions. Obviously, diagnostic ions for the modification must come from fragmentation of this side chain. Fragment ions involving any part of the “normal” peptide will vary in mass according to the peptide being modified and will therefore not be of general diagnostic use. 3.2.3.2. EXAMINATION OF THE LITERATURE TRYPTIC PEPTIDE SPECTRA

MS/MS spectra of GG-tagged tryptic peptides are shown in both the paper from Dohmann’s group, and that of the Gygi group. Examination of these spectra show that b and y ions are produced by dissociation of the peptide from the target protein, but there do not appear to be any b ions from the GG side chain, or the GGK portion of the peptide. 3.2.3.3. EXAMINATION OF THE LITERATURE GLUC PEPTIDE SPECTRA

The MS/MS spectrum of the gluC ubiquitinated peptide from rXL-calmodulin which was shown in the Laub et al paper (25), reveals b and y ions from the calmodulin portion of the peptide (Fig. 7). Interestingly, it also shows doublycharged (b7-H2O) 2+, (b14-H2O)2+, and (b12-H2O)2+ ions which contain both of the N-termini (one from the calmodulin peptide and one from the ubiquitin side chain) of the original branched peptide. These two ions (athough of low relative abundance) can be seen in the MS/MS spectrum of the ubiquitinated peptide from natural BT-calmodulin. Careful examination of the both spectra

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Fig. 7. Calmodulin ubiquitinated peptide structure after cleavage with gluC. Circled peptide fragments were not noted in the original publication by Laub et al (25), but appear to have the appropriate masses.

reveals ions which might be from the ubiquitin side chain, and therefore of possible diagnostic utility. The y14 ion includes ions from both the side chain and the C-terminus of the calmodulin-portion of the peptide, so it cannot be used as a ubiquitination marker ion. However, there are also ions from the side chain, b4 at m/z 478.4 (obs) and, possibly, b2, at m/z ∼191.2 (obs), which appear to be diagnostic ions of ubiquitination, after digestion with gluC. 3.2.3.4. PREPARATION OF MODEL UBIQUITINATED PEPTIDES

In order to further study the fragmentation of ubiquitinated peptides, we had a model peptide synthesized. A peptide was synthesized with the structure shown in Fig. 7. These modified peptides should show a characteristic mass shift from their unmodified analogues: in the case of trypsin, a mass shift of 114.0428 Da from the GG “tag” left on the modified lysine. In the case of gluC, a mass shift of 1302.7883 Da, a much larger mass shift resulting from the much longer “tail” (STLHLVLRLRGG-) on the modified lysine, is expected. Unfortunately, the model peptide was synthesized with an amide at the C-terminus, and an acetyl group at the N-terminus, so it is not an exact model of the peptide found by Laub et al (25). Fortunately, the acetyl group was in the “calmodulin” portion of the peptide, so we expected that diagnostic fragments from the ubiquitin side chain would still be present in the MS/MS spectrum. ESI-MS/MS spectra of the model peptides were obtained by LC/MS/MS analysis using a Waters/Micromass Q-tof API US, equipped with a Waters capLC system. An aliquot of the sample was injected first onto a Dionex trapping column, which was then switched so that it was connected on-line to a 75 µm Dionex Pep-map analytical column and the ESI source. The original synthetic peptide, the model gluC peptide, was dissolved in water, and injected without further purification. It proved to be a mixture of acetylated and methylated forms, whose MS/MS spectra could be analyzed separately after separation by LC/MS/MS. The actual methylation sites can be deduced from the MS/MS spectrum: one is on the S in the calmodulin part of the peptide, the other is on one of the first two residues (S or T) of the ubiquitin tail.

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The model ubiquitinated peptide was prepared from the synthetic peptide by means of an in-solution tryptic digest (see Subheading 3.1.2.5.3), using 2 µg of trypsin and 1 µg of the original peptide in 100 µL of 100 mM ammonium bicarbonate. As stated earlier, an aliquot of the digest was injected into the LC/ MS/MS system, and went first onto a Dionex trapping column, which was then switched so that the column was connected on-line to a 75 µm i.d. Dionex Pepmap analytical column and the ESI source. Two main products were formed: a peptide with the expected “GG” tail on the ubiquitinated lysine, and a second peptide with an “LRGG” tail, resulting from a missed cleavage. Even though this was an in-solution tryptic digest with a large amount of trypsin, a significant amount of a peptide was produced with a missed cleavage site on the side chain. Although this missed cleavage was unexpected, it is not unreasonable, since this cleave site is close to the branch point so cleavage at this site is likely to be sterically hindered. MS/MS spectra were obtained for both of these products. The formation of this peptide is of significant analytical interest as it provides a second characteristic molecular weight shift for ubiquitinated peptides after tryptic digestion. 3.2.3.5. FRAGMENTATION OF MODEL GLUC UBIQUITINATED PEPTIDE

Because of the various possible dimethylated isoforms, the first model gluC peptide examined was the dimethylated version. The resulting spectrum is shown in Fig. 8. Both b and y fragment ions are found from the “normal” part of the peptide. Most interestingly, several fragments are found which only involve the side chain. As were observed in the literature spectrum, three characteristic ions were detected from the “ubiquitin” side chain: these are b2, b3, and b4. These ions, in their un-methylated forms, (m/z 189.088, 302.172, 439.231, 555.315, and 651.383) should thus be diagnostic ions for peptides with ubiquitin side chains that have been cleaved with gluC. 3.2.3.6. FRAGMENTATION OF MODEL TRYPTIC PEPTIDES

Cleavage of the synthetic model peptide with trypsin resulted in a GG-tagged peptide whose MS/MS spectrum is shown in Fig. 9. Unfortunately, as in the literature spectra, no characteristic fragment ions could be found which were diagnostic of the critical GGK portion of the molecule. The MS/MS spectrum of the model tryptic peptide which had the LRGG tag (resulting from a missed cleavage) was more analytically useful (Fig. 10). Diagnostic ions of the ubiquitin tag were found. These include b2, b4, and the internal fragment ion (LRGGK-28). There is also a LRGGKD ion fragment ion, but since this includes the “D” from the calmodulin peptide, it cannot be used as a diagnostic ion for modification by ubiquitin. The assignment of these peptide fragments is confirmed by the MS/MS spectrum of the S-methylated calmodulin

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Fig. 8. MS/MS Fragmentation of synthetic model calmodulin gluC peptide, dimethyl form. (Reproduced with permission from ref. 27, copyright John Wiley & Sons, Limited).

peptide is shown in Fig. 11. The same diagnostic ions b2, b4, (LRGGK-28) ions are observed in this spectrum. 3.2.4. Enrichment of Samples for Specific Ubiquitin-Modified Proteins and/or Proteins That Interact with Them As described above (Subheading 3.2.3), the work of Gururaja (16), Kirkpatrick (20), and Peng (17) depended upon enriching the sample in ubiquitinated peptides. Their goals were to find as many ubiquitination sites as possible in any protein, so their approach was to use a 6 × his-tagged ubiquitin, and enrich the sample in ubiquitinated proteins through the use of IMAC or 6 × his –myc-Ub (18). Other researchers have used antiubiquitin (21) or antipolyubiquitin (23,24) for “shotgun” enrichment. To find ubiquitination sites in a particular target protein requires a different approach. One method is the use of a GST-tagged substrate, and anti-GSH affinity beads. This method was used by the Marshall group for an FT-MS study of polyubiquitinated GST-Ubc5, ubiquitinated in vitro (28). FT-MS identified fifteen ubiquitination sites in GST-Ubc5, and four sites in ubiquitin, although large quantities of material were used (e.g., 700 µg GST-Ubc5).

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Fig. 9. Fragmentation of model “GG-” ubiquitinated tryptic peptide. Reproduced with permission from ref. 27, copyright John Wiley & Sons Limited.

If an antibody is available against the target protein, then it can be used to affinity purify the target protein from the cell lysate. If an antibody against the protein of interest is not available, an affinity tag can be incorporated into the target protein sequence. In our case, we used a FLAG tag added to the aminoterminus of the CIITA target protein so that it could be immunoprecipitated with anti-FLAG antibody beads. As a second strategy for increasing the proportion of ubiquinated vs. nonubiquitinated protein, a plasmid was used which coded for a modified ubiquitin which had all of the lysines modified to arginines and a HA tag on the C-terminus. This construct was designed to prevent the formation of polyubiquitin chains and thus to inhibit degradation of the target protein (29). The HA tag allows a second affinity purification step, either before or after proteolysis, this time on anti-HA beads. To avoid proteolysis of the ubiquitin (and loss of the HA tag), LysC was used for the initial digestion of the protein, instead of trypsin. The affinity-bound protein can then be digested overnight with trypsin or with gluC (see Note 8) while still attached to the beads, using the protocols described in the next subheading. Alternatively, the protein can be eluted from the affinity beads with 1:1:8 ethanol:formic acid:water, lyophilized and digested in solution.

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Fig. 10. Fragmentation of model “LRGG-” ubiquitinated tryptic peptide. (Reproduced with permission from ref. 27, copyright John Wiley & Sons Limited).

3.2.5. Elution and Enzymatic Digestion Procedures 3.2.5.1. ELUTION OF PROTEINS FROM BEADS 1. Place ∼50–100 µL of beads in an Eppendorf tube. 2. Wash beads 3 × with 200 µL of 100 mM ammonium bicarbonate, and discard wash solutions. 3. Add 100 µL 1:1:8 Ethanol:formic acid:water. 4. Vortex, let settle. 5. Remove and save eluates. 6. Repeat extraction 2 more times. 7. Save and combine eluates. 8. Lyophilize. 9. Store at −80°C. 3.2.5.2. ON-BEAD DIGESTION TRYPTIC PROCEDURE (30) (SEE NOTE 9) 1. Place ∼50–100 µL of antibody beads (with the attached affinity-bound protein) in an Eppendorf tube. 2. Wash 3 × with 200 µL of 100 mM ammonium bicarbonate. 3. Add 100 µL of 100 mM ammonium bicarbonate.

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Fig. 11. Fragmentation of model “LRGG-” ubiquitinated tryptic peptide, methylated form. (Reproduced with permission from ref. 27, copyright John Wiley & Sons Limited).

4. Reconstitute Promega trypsin in 20 µL Promega resuspension buffer (0.015 M acetic acid) (Promega trypsin comes in aliquots of 20 µg per vial). 5. Add 2 µL trypsin solution to each sample (see Note 10). 6. Vortex/spin down at 200 min) are preferred for separating complex mixtures of peptides in order to reduce suppression effects and to reduce the number of coeluting peptides, since selection of the precursor ions is done based on ion abundances. For an unknown modification site in a protein containing many potential sites, where the peptide molecular weight is therefore not known, automatic data-dependent triggering of MS/MS data collection (called “survey scan” mode in the Micromass MassLynx software) is the only feasible automatic scanning option. The resulting MS/MS spectra are then analyzed by commercially-available software packages (such as Mascot or SEQUEST) which can be programmed (see Notes 15 and 16) to consider a lysine modified with a GG or, as we have learned from the above experiments on our model peptides, LRGG, for a tryptic digest, or with STLHLVLRLRGG for a gluC digest). Ideally, the ubiquitination will be found from this automated search routine (see Notes 17 and 18). Fig. 12 shows an example of a database search of MALDI-MS data from a tryptic digest of a ubiquitinated protein. Here, a ubiquitinated peptide has been identified. Although promising, these results simply mean that there is a peak in the mass spectrum which has the mass of an expected tryptic peptide where a lysine has been modified with a GG tag. Since a peak at this mass could

Fig. 12. Mascot database search results from a tryptic digest of ubiquitinated CIITA, showing a “hit” for a potential ubiquitinated peptide. This potential ubiquitination site has not yet been confirmed by MS/MS data.

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have come from another peptide in the mixture, this ubiquitination site cannot be confirmed without MS/MS sequence data of this peptide. If MS/MS data has been acquired and searched, as is the case when LC/MS/ MS has been used, the identity of the peptide can be confirmed and site of ubiquitination found from the database search results (see Note 19). This was the method used to find the ubiquitination sites in the Peng, et al. paper (17). Unfortunately, this approach often fails where lower amounts of biological material are available. Very low levels of the modified peptide mean that there may not be sufficient intensity of the modified peptide to trigger this automatic datadependent MS/MS sequencing. In this case, another much more time-consuming option is to examine the MS spectra to search for a peptide shifted by the masses corresponding these possible modifications. This can be done manually by creating a list of expected “normal” peptide masses, calculating the modified masses, and examining the MS spectra obtained during the LC/MS/MS run. Obviously, at 1 s per scan, thousands of spectra are obtained throughout the course of an extended LC/MS/ MS run. Current software systems allow the combination of groups of spectra, and these groups of combined spectra can be examined. Most current software packages also allow the deconvolution of the spectra to singly-charged species, which reduces the complexity of the manual data analysis, since multiple charge states of the possible peptides are deconvoluted to singly-charged species. A semi-automated approach to this task is to combine all of the spectra, perform a deconvolution on the MS data to generate a pseudo-singly charged spectrum (see Note 20), and then submitting this data to the database search software for searching as an MS data file. As above, the MS data can then be searched for modified peptides (see Note 21). If a possible modified peptide is identified in this manner, it is useful to examine the original MS data to see if the calculated +2 and +3 charge states for this peptide co-chromatograph. Now that the masses are known, it is possible to perform a different type of MS scanning, where the precursor ion is specified – in Micromass software, it is called “include only” MS/MS. In this scan mode, MS/MS is only performed on the preselected precursor ions. In a previous study (31), we have found that this can lead to an increase in sensitivity of a factor of 50–100 for these ions. Although this scanning mode is dependent on intensity-based triggering, and thus has the same sensitivity limitations as the “survey scan” mode, knowing these characteristic fragment ions also allows the possibility of “precursor ion scanning”. Here, when a preset characteristic ion is detected, the data system switches to the MS mode, detects the precursor, and collects the MS/MS data. Similar approaches are already commonly used, for example, in order to find peptides containing acetylated lysine from the acetylated lysine immonium

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ion (32) or to find phosphotyrosine-containing peptides from its characteristic immonium ion (33,34). The identification of the specific and characteristic fragment ions which we have described above, provides a powerful new approach for finding ubiquitinated peptides. Searching the MS/MS chromatograms (see Note 21) for these characteristic ions (which should cochromatograph because they are fragment ions from the same peptide) should allow one to find peptides containing ubiquitin side chains. Since we first suggested the use of extended ubiquitin tags to improve the detection of ubiquitinated peptides in 2005 (35), two other groups have used the LRGG tag and have demonstrated the improvement resulting from inclusion of this second mass shift. Jeon, et al. (36) used GG and LRGG mass shifts resulting from GG and LRGG tags to search LC/MS/MS data in a mouse heart digest. The use of the GG tag resulted in the identification of 27 ubiquitination sites on 21 proteins. The use of the LRGG tag resulted in the identification of an additional 6 sites. These two tags were also used to identify the polyubiquitination sites on ubiquitin itself (in this case, Lys 48 and Lys 63). One important observation by this group is that the ubiquitinated proteins from mouse heart were insoluble in detergent-free buffer – CNBr cleavage resulted in smaller more soluble proteins, while preserving the ubiquitination. Recently the Figey group used both GG and LRGG tags to increase the detection of ubiquitinated peptides from a human MCF7 breast cancer cell culture (37). This group found a total of 96 ubiquitination sites. Of these sites, 53% were found using internal GGK residue, an additional 8% from the internal LRGGK residue, and 39% from C-terminal GGK peptides (which were cleaved by trypsin at the modified residue). These C-terminal peptides (with the standard tryptic cleavage after K) would, of course, have been detected by the database searches, since at least the 1 missed cleavage expected from blocked tryptic would have been allowed. However, the authors noted that no C-terminal LRGG peptides were detected, only C-terminal GGK peptides. This provides support for the hypothesis that the missed cleavage of the LR-GGK bond results from steric hindrance of the trypsin. A “two-stage” affinity purification strategy was used by the Figeys group for the study of the ubiquitination of proteins co-purifying with valosin-containing protein (VCP), a ubiquitin-dependent chaperone (38). In this study, first immobilized anti-VSP and then anti-ubiquitin beads were used. Using this strategy, along with a high-pressure device to improve tryptic digestion efficiencies, 27 ubiquitination sites were mapped on 21 proteins, and 58 additional “probably-ubiquitinated” proteins were also identified by LC/MS/MS, using the 114 or 383-Da mass shift associated with the GG- or LRGG-modified lysine residues.

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3.2.6. Determination of Ubiquitin Branched Structure Polyubiquitinated chains can be thought of as multiple ubiquitinations of ubiquitin. Tryptic cleavage of these linked ubiquitins results in GGK and LRGGK-tagged peptides from ubiquitin. A detailed strategy for determining this branching structure has been reported by Kaiser and Wohlschlegel (39), who used the GGK tagged peptides. They reported that they preferred to perform LC/MS/MS or LC/LC/MS/MS on digests of gel-separated proteins because if a mixture of proteins was present, it could not be determined which protein these polyubiquitin fragments came from. Also, without separation of the ubiquitinated peptides from the protein (prior to digestion of the polyubiquitin tail), the “average” branch structure would be determined if there were more than one ubiquitin site on each protein. This, or course, would be difficult to accomplish because enzymes that would separate the ubiquitination sites on the protein would also cleave the polyubiquitin chains. 3.2.7. Quantitation of Ubiquitination Several recent papers have reported different methods of quantifying the ubiquitination at various sites in the protein, as well as quantitation of the various polyubiquitin isoforms. Cripps et al (40) used targeted detection of ubiquitinated peptides from a targeted protein, and also performed quantitation determination of the ubiquitinated peptides. Relative quantitation using selected reaction monitoring (see Note 22) was performed on GGK-modified tryptic peptides from Tau protein, immunoprecipitated from Alzheimers brain tissue, using unmodified Tau peptides as internal standards. From this MRM data, the authors determined the relative ubiquitination levels of the 3 sites found in this protein. Although the possible formation of the LRGG tag waas not considered in this studym, or any of the quantitation studies described below, it probably would not significantly affect the results. Gygi and coworkers have recently published two ubiquitination studies with absolute quantitation (AQUA (42); see Note 23), the first on the epidermal growth factor receptor (EGFR) (41), and the second on cyclin B1 (43). In the EGFR study, recombinant EGFR receptor was purified, and LC/MS/MS was performed. Six ubiquitination sites (GGK) were identified, and quantitative determination of the “average” branch structure of the polyubiquitin chain at each site was performed by comparison of the selected reaction monitoring peak areas of each peptide with those from stable-isotope labeled peptides (the “ub-AQUA” method (42)). This “ub-AQUA” method was also used by the Gygi group to determine the branch structure of ubiquitinated cyclin B1 (43). The absolute amounts of each of the 10 possible polyubiquitin cleavage products (7 branched and 3 unbranched)

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were determined by comparison with the amounts of 10 synthetic isotopicallylabeled analogues. (For example, the heavy leucine in one of the reference peptide contained six 13C’s and one 15N). As in the study by Kaiser and Wohlshlegel described above (39), ubiquitinated cyclin B1 was first separated by gel electrophoresis, and the quantitation was performed on the extract containing the in-gel digested peptides. Calculations were performed to determine the amount of ubiquitin in long chains vs. short chains. To determine the relative quantitation of the branch structures in a set of 5 ubiquitin-conjugating enzymes and ubiquitin ligases, the Gygi group used a different strategy, utilizing a set of mutant ubiquitins, each having all but one of the 7 lysines replaced by alanines (44). The relative amounts of each type of linkage was based on spectral counting (45) of the resulting GGK-containing peptides, i.e. the number of spectra observed in a data-dependent LC/MS/MS acquisition, rather than the abundance of the spectra. 3.2.8. Emerging Strategies In addition to new quantitative studies, there have been several interesting studies designed to improve detection of ubiquitinated peptides which is still extremely challenging when the amount of biological material is limited. As mentioned above (see Subheading 3.1.2.3.1), ubiquitinated peptides differ from “standard” peptides in that they have 2 N-termini. Recently, Cotter and his group have developed a new method which uses this feature (46,47). This method is based on a derivatization procedure, similar to the chemistry behind Edman sequencing (48). This reagent attaches an SPITC (4-sulfophenyl isothiocyanate) moiety to the N-termini – to both N-termini for ubiquitinated peptides. Under MS/MS conditions, ubiquitinated peptides can then be distinguished from nonubiquitinated peptides by the loss of two SPITC groups forming a set of “signature” ions resulting from complete or partial loss of one or two of these tags (“normal” peptides can only have one loss of SPITC) (Fig. 13). This derivatization procedure, first developed by Keough et al (49), produces a y series from a derivatized peptide, while reducing the b series (ions which contain the N-terminus). The Cotter group first modified this reagent for more efficient use in aqueous media (50), and then used it to derivatize tryptic peptides prior to MALDI-MS and MALDI-MS/MS. They used this method for determination of ubiquitination sites in a synthetic peptide containing a GGK tag, and to tetraubiquitin (46). In a subsequent paper (47), the Cotter group used this procedure to find the ubiquitination sites on His-tagged CHIP (C-terminal HSP70-interacting protein), which was affinity purified using Cobalt beads, and IMAC. Three peptides were identified as doubly-tagged based on their MS/ MS spectra, which produced “signature” ions from the SPITC moieties and

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Fig. 13. Fragmentation of an SPITC-labeled ubiquitinated peptide. (Reprinted from ref. 46, with permission)

sequence ions which were used to identify the site of ubiquitination. In a recent report, the Perales group (51) used the SPITC approach, and reported improved quality of the resulting MS/MS sequencing, which is another advantage of this approach. A different method for N-terminal derivatization was recently suggested by the Gebler group (52). Like the SPITC method, this is a new look at an older derivatization technique for mass spectrometry (53). This method relies on derivatization of the N-terminal peptides with tris(2,4,6-trimethoxyphenyl)

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phosphonium acetic acid N-hydroxysuccinimide ester (TMPP-Ac-OSu), which introduces a fixed charge on the N-terminus and simplifies MALDI-.MS/MS sequencing. This method also has potential for improving the detection of “double N-terminal” ubiquitinated peptides. These N-terminal labeling approaches should be very useful for identifying ubiquitinated peptides from a mixture, especially if the label could be combined with an affinity capture moiety such as biotin. This approach is proving extremely useful for the identification of crosslinked peptides (54), and it would seem that a similar strategy could be developed for the labeling and recovery of ubiquitinated peptides. 4. Notes 1. We call this “direct” analysis because the affinity beads are placed directly on the target, in contrast to methods were the affinity-bound proteins are eluted first, and the eluate is spotted on the target. 2. If binding has to be done through protein A, or the antibody is dissolved in ascites, the antibody can be crosslinked to the beads (55) before the target protein is affinity-bound. 3. Since only a small number of beads will be placed on the MALDI target, it is important to get much protein as possible on each bead, so use a small amount (∼20 µL) of settled affinity bead slurry. 4. To avoid releasing parts of the antibody into the solution, try to avoid reducing agents such as β-mercaptoethanol or dithiothreitol (DTT) in the binding buffer. 5. Most salts will be removed during the wash steps. Other components not compatible with mass spectrometric analysis (such as glycerol or detergents) should be avoided or minimized. Although Zwittergent is thought to be compatible with mass spectrometry, it seems to make agarose beads turn “gummy,” so it should not be used. 6. To recrystallize α-cyano 2-hydroxycinnamic acid, make a saturated solution of α-cyano 2-hydroxycinnamic acid in boiling methanol. Pour off the solution and discard it. Add more methanol, and again make a saturated solution in boiling methanol. This time, pour off the methanol and save it. Evaporate the methanol to dryness in a hood, while protecting the solution and the crystals from light with aluminum foil. Store in the dark or in a vial wrapped with aluminum foil. 7. The matrix solvent must contain both organic and acid so that it dissociates the affinity bound protein from the antibody on the MALDI target. 8. Also known as Staphloccus aureus V-8 protease. 9. The antibody used should be covalently attached to the beads, and DTT should not be used (or used in a very low concentration (56)) in the purification step. If the disulfide bridges in the antibody are reduced, the

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10. 11. 12. 13.

14.

15.

16.

17.

18. 19.

20. 21. 22.

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antibody can be enzymatically digested along with the attached protein. This will lead to a high background of IgG peptides along with the peptides from the target protein, and will make finding the modified peptide more difficult. For proteins affinity-bound to antibody beads, much higher ratios of enzyme (e.g., 5:1) to substrate should be used than for proteins in solution (30). Be sure to add enough digestion buffer so that the beads can “slosh around” in the Eppendorf tube and don’t dry out during the overnight digestion. If you do not know the protein concentration, use ∼2 µL of a 1 µg/µL solution. If the sample to be digested is already dissolved in water or a buffer such as PBS, add enough ammonium bicarbonate to the solution to ensure that the pH will be ∼7–8. The specificity of gluC depends on buffer used and pH of the solution. Cleavage can be either C-terminal to glutamic acid (in ammonium acetate at pH 4.0) or (ammonium bicarbonate at pH 7.8), or to both glutamic and aspartic acid (in PBS, pH 7.8). For a solution digest, you can add a second buffer to the original buffer in order to adjust the pH, but be sure to take both enzymes into account when calculating the expected of the peptide molecular weights. This requires a site license for Mascot, although the company says that if there is sufficient interest, they will add new modifications to their on-line website (www.matrixscience.com). Programming in these modification means, in effect, telling the software that there are three additional types of lysines, with new masses: 242.1374 for GGK, 511.6294 for LRGGK, and 1430.8824 for STLHLVLRLRGGK. In searching using Mascot, first search with no variable modifications, then select the target protein and search in the error tolerant mode, specifying the appropriate ubiquitination modifications you have previously entered. Be sure to specify at least two missed cleavages. Cleavage is not expected to occur at the modified lysine. The database search can also be “forced” to consider a specific target program. In MASCOT, this is done by adding the accession number from the appropriate database as the first line of the peak list being searched. (e.g., accession=XXXXX, followed by a blank line). For Micromass MassLynx software, this is MaxEnt 3, under “Tools”. If multiple levels of MS/MS spectra are produced, as in the Micromass MassLynx software, all of the MS/MS functions must be searched. Multiple reaction monitoring (or selected reaction monitoring) is a technique performed on a tandem mass spectrometer. In this technique, a both the precursor and the product ion (formed in a collision cell) are selected, which makes it a very selective technique for quantitation.

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23. This is a confusing term, partly because the AQUA acronym is used by two groups (both from Harvard) for different things, automated quantitation (57) and absolute quantitation (42). AQUA as used now in mass spectrometry (42) is a new name for a mass spectrometric technique dating back at least to the early 1970s (58, 59) where a stable-labeled form of the target analyte is introduced into the sample to act as an internal standard. This isotopically labeled molecule has properties nearly identical to the target analyte, but has a different mass (because of the isotopic label). This allows it to be distinguished from the native form by mass spectrometric analysis. Acknowledgments This study was funded by a gift from an anonymous donor to support research in proteomics and cystic fibrosis, and grants from the Cystic Fibrosis Foundation (CFFTI STUTTS01U0) and from NIH (ES11997, 5U54HD035041-07, and P30 CA 16086-25).

References 1. Hochstrasser, M. (1996) Ubiquitin-dependent protein degradation. Ann. Rev. Genet. 30, 405–439. 2. Levinger, L. and Varshavsky, A. (1982) Selective arrangement of ubiquitinated and D1 protein-containing nucleosomes within the Drosophila genome. Cell 28, 375–385. 3. Glotzer, M., Murray, A. W., and Kirschner, M. W. (1991) Cyclin is degraded by the ubiquitin pathway. Nature 349, 132–138. 4. Sasaki, A., Inagaki-Ohara, K., Yoshida, T., Yamanaka, A., Sasaki, M., Yasukawa, H., Koromilas, A., and Yoshimura, A. (2003) The N-terminal truncated isoform of SOCS3 translated from an alternative initiation AUG codon under stress conditions is stable due to the lack of a major ubiquitination site, Lys-6. J. Biol. Chem. 278, 2432–2436. 5. Hershko, A., and Ciechanover, A. (1992) The ubiquitin system for protein degradation. Ann. Rev. Biochem. 61. 6. Pickart, C. M. (2001) Ubiquitin enters the new millennium. Mol. Cell 8, 499–504. 7. Hicke, L. (2001) A new ticket for entry into budding vesicles—ubiquitin. Cell 106, 527–530. 8. Johnson, E. S. (2002) Ubiquitin branches out. Nature Cell Biol. 4, E295–E298. 9. Sun, Z. W. and Allis, C. D. (2002) Ubiquitination of histone H2B regulates H3 methylation and gene silencing in yeast. Nature 418, 104–108. 10. Conaway, R. C., Brower, C. S., and Conaway, J. W. (2002) Emerging roles of ubiquitin in transcription regulation. Science 296, 1254–1258. 11. Hochstrasser, M. (2002) Evolution and function of ubiquitin-like proteinconjugations ystems. Nature Cell Biol. 2, E153–E157. 12. Kirkpatrick, D. S., Denison, C., and Gygi, S. P. (2005) Weighing in on ubiquitin: the expanding role of mass-spectrometry-based proteomics. Nature Cell Biol. 7, 750–757.

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13. Denison, C., Kirkpatrick, D. S., and Gygi, S. P. (2005) Proteomic insights into ubiquitin and ubiquitin-like proteins. Curr. Opin. Chem. Biol. 9, 69–75. 14. Xu, P. and Peng, J. (2006) Dissecting the ubiquitin pathway by mass spectrometry. Biochim. Biophys. Acta, Proteins Proteomics 1764, 1940–1947. 15. Drews, O., Zong, C., and Ping, P. (2007) Exploring proteasome complexes by proteomic approaches. Proteomics 7, 1047–1058. 16. Gururaja, T., Li, W., Noble, W. S., Payan, D. G., and Anderson, D. C. (2003) Multiple functional categories of proteins identified in an in vitro cellular ubiquitin affinity extract using shotgun peptide sequencing. J. Proteome Res. 2, 394–404. 17. Peng, J., Schwartz, D. R., Elias, J. E., Thoreen, C. C., Cheng, D., Marsischky, G., Roelofs, J., Finley, D., and Gygi, S. P. (2003) A proteomics approach to understanding protein ubiquitination. Nature Biotechnol. 21, 921–926. 18. Hitchcock, A. L., Auld, K., Gygi, S. P., and Silver, P. A. (2003) A subset of membrane-associated proteins is ubiquitinated in response to mutations in the endoplasmic reticulum degradation machinery. Proc. Natl. Acad. Sci. U.S.A. 100, 12735–12740. 19. Peng, J., and Cheng, D. (2005) Proteomic analysis of ubiquitin conjugates in yeast. Methods Enzymol. 399 (Ubiquitin and Protein Degradation, Part B), 367–381. 20. Kirkpatrick, D. S., Weldon, S. F., Tsaprailis, G., Liebler, D. C., and Gandolfi, A. J. (2005) Proteomic identification of ubiquitinated proteins from human cells expressing His-tagged ubiquitin. Proteomics 5, 2104–2111. 21. Vasilescu, J., Smith, J. C., Ethier, M., and Figeys, D. (2005) Proteomic Analysis of Ubiquitinated Proteins from Human MCF-7 Breast Cancer Cells by Immunoaffinity Purification and Mass Spectrometry. J. Proteome Res. 4, 2192–2200. 22. Fujimuro, M., Sawada, H., and Yokosawa, H. (1994) Production and characterization of monoclonal antibodies specific to multi-ubiquitin chains of polyubiquitinated proteins. FEBS Lett. 349, 173–80. 23. Hatakeyama, S., Matsumoto, M., and Nakayama, K. I. (2005) Mapping of ubiquitination sites on target proteins. Methods Enzymol. 399, 277–286. 24. Matsumoto, M., Hatakeyama, S., Oyamada, K., Oda, Y., Nishimura, T., and Nakayama, K. I. (2005) Large-scale analysis of the human ubiquitin-related proteome. Proteomics 5, 4145–4151. 25. Laub, M., Steppuhn, J. A., Bluggel, M., Immler, D., Meyer, H. E., and Jennissen, H. P. (1998) Modulation of calmodulin function by ubiquitin-calmodulin ligase and identification of the responsible ubiquitylation site in vertebrate calmodulin. Eur. J. Biochem. 255, 422–431. 26. Marotti, L. A., Jr., Newitt, R., Wang, Y., Aebersold, R., and Dohlman, H. G. (2002) Direct Identification of a G Protein Ubiquitination Site by Mass Spectrometry. Biochemistry 41, 5067–5074. 27. Warren, M. R. E., Parker, C. E., Mocanu, V., Klapper, D. G., and Borchers, C. H. (2005) Electrospray ionization tandem mass spectrometry of model peptides reveals diagnostic fragment ions for protein ubiquitination. Rapid Communic. Mass Spectrom. 19, 429–437.

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28. Cooper, H. J., Heath, J. K., Jaffray, E., Hay, R. T., Lam, T. T., and Marshall, A. G. (2004) Identification of sites of ubiquitination in proteins: A Fourier transform ion cyclotron resonance mass spectrometry approach. Analytical Chemistry 76, 6982–6988. 29. Greer, S. F., Zika, E., Conti, B., Zhu, X.-S., and Ting, J. P.-Y. (2003) Enhancement of CIITA transcriptional function by ubiquitin. Nature Immunol. 4, 1074–1082. 30. Parker, C. E. and Tomer, K. B. (2000). In Methods in Molecular Biology (Chapman, J. R., Ed.), Vol. 146 (Mass Spectrometry of Proteins and Peptides), pp. 185–201, Humana Press, Totowa, NJ. 31. Kast, J., Parker, C. E., van der Drift, K., Dial, J. M., Milgram, S. L., Wilm, M., Howell, M., and Borchers, C. H. (2003) Matrix-assisted laser desorption/ionization directed nano-electrospray ionization tandem mass spectrometric analysis for protein identification. Rapid Comm. Mass Spectrom. 17, 1825–1834. 32. Borchers, C., Parker, C. E., Deterding, L. J., and Tomer, K. B. (1999) A preliminary comparison of precursor scans and LC/MS/MS on a hybrid quadrupole time-offlight mass spectrometer. J. Chromatogr. A 854, 119–130. 33. Steen, H., Kuester, B., Fernandez, M., Pandey, A., and Mann, M. (2001) Detection of tyrosine phosphorylated peptides by precursor ion scanning quadrupole TOF mass spectrometry in positive ion mode. Anal. Chem. 73, 1440–1448. 34. Steen, H., Kuster, B., Fernandez, M., Pandey, A., and Mann, M. (2002) Tyrosine phosphorylation mapping of the epidermal growth factor receptor signaling pathway. J. Biol. Chem. 277, 1031–1039. 35. Warren, M. R. E., Parker, C. E., Mocanu, V., Klapper, D. G., and Borchers, C. H. (2004) In 52nd Annual Conference on Mass Spectrometry and Allied Topics, Nashville, TN. 36. Jeon, H. B., Choi, E. S., Yoon, J. H., Hwang, J. H., Chang, J. W., Lee, E. K., Choi, H. W., Park, Z.-Y., and Yoo, Y. J. (2007) A proteomics approach to identify the ubiquitinated proteins in mouse heart. Biochem. Biophys. Res. Commun. 357, 731–736. 37. Denis, N. J., Vasilescu, J., Lambert, J.-P., Smith, J. C., and Figeys, D. (2007) Tryptic digestion of ubiquitin standards reveals an improved strategy for identifying ubiquitinated proteins by mass spectrometry. Proteomics 7, 868–874. 38. Vasilescu, J., Zweitzig Daniel, R., Denis Nicholas, J., Smith Jeffrey, C., Ethier, M., Haines Dale, S., and Figeys, D. (2007) The proteomic reactor facilitates the analysis of affinity-purified proteins by mass spectrometry: application for identifying ubiquitinated proteins in human cells. J Proteome Res. 6, 298–305. 39. Kaiser, P., and Wohlschlegel, J. A. (2005) Identification of ubiquitination sites and determination of ubiquitin-chain architectures by mass spectrometry. Methods Enzymol. 399 (Ubiquitin and Protein Degradation, Part B), 266–277. 40. Cripps, D., Thomas, S. N., Jeng, Y., Yang, F., Davies, P., and Yang, A. J. (2006) Alzheimer Disease-specific Conformation of Hyperphosphorylated Paired Helical Filament-Tau Is Polyubiquitinated through Lys-48, Lys-11, and Lys-6 Ubiquitin Conjugation. J. Biol. Chem. 281, 10825–10838. 41. Huang, F., Kirkpatrick, D., Jiang, X., Gygi, S., and Sorkin, A. (2006) Differential regulation of EGF receptor internalization and degradation by multiubiquitination within the kinase domain. Molecular Cell 21, 737–748.

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42. Kirkpatrick, D. S., Gerber, S. A., and Gygi, S. P. (2005) The absolute quantification strategy: a general procedure for the quantification of proteins and posttranslationalmodif ications. Methods (San Diego, CA) 35, 265–273. 43. Kirkpatrick, D. S., Hathaway, N. A., Hanna, J., Elsasser, S., Rush, J., Finley, D., King, R. W., and Gygi, S. P. (2006) Quantitative analysis of in vitro ubiquitinated cyclin B1 reveals complex chain topology. Nature Cell Biol. 8, 700–710. 44. Kim, H. T., Kim, K. P., Lledias, F., Kisselev, A. F., Scaglione, K. M., Skowyra, D., Gygi, S. P., and Goldberg, A. L. (2007) Certain pairs of ubiquitin-conjugating enzymes (E2s) and ubiquitin-protein ligases (E3s) synthesize nondegradable forked ubiquitin chains containing all possible isopeptide linkages. J. Biol. Chem. 282, 17375–17386. 45. Liu, H., Sadygov, R. G., and Yates, J. R. I. (2004) A model for random sampling and estimation of relative protein abundance in shotgun proteomics. Anal. Chem. 76, 4193–4201. 46. Wang, D., and Cotter, R. J. (2005) Approach for determining protein ubiquitination sites by MALDI-TOF mass spectrometry. Anal. Chem. 77, 1458–1466. 47. Wang, D., Xu, W., McGrath, S. C., Patterson, C., Neckers, L., and Cotter, R. J. (2005) Direct identification of ubiquitination sites on ubiquitin-conjugated CHIP using MALDI mass spectrometry. J. Proteome Res. 4, 1554–1560. 48. Edman, P. (1950) Preparation of phenyl thiohydantoins from some natural amino acids. Acta Chem. Scand. 4, 277–282. 49. Keough, T., Youngquist, R. S., and Lacey, M. P. (2003) Sulfonic acid derivatives for peptide sequencing by MALDI MS. Anal. Chem. 75, 156A–165A. 50. Wang, D., Kalb, S. R., and Cotter, R. J. (2004) Improved procedures for N-terminal sulfonation of peptides for matrix-assisted laser desorption/ionization post-source decay peptide sequencing. Rapid Ccommun. Mmass Spectrom. 18, 96–102. 51. Leon, I. R., Neves-Ferreira, A. G. C., Valente, R. H., Mota, E. M., Lenzi, H. L., and Perales, J. (2007) Improved protein identification efficiency by mass spectrometry using N-terminal chemical derivatization of peptides from Angiostrongylus costaricensis, a nematode with unknown genome. J. Mass Spectrom. 42, 781–792. 52. Chen, W., Lee, P. J., Shion, H., Ellor, N., and Gebler, J. C. (2007) Improving de novo sequencing of peptides using a charged tag and C-terminal digestion. Anal. Chem. 79, 1583–1590. 53. Huang, Z.-H., Wu, J., Roth, K. D. W., Yang, Y., Gage, D. A., and Watson, J. T. (1997) A picomole-scale method for charge derivatization of peptides for sequence analysis by mass spectrometry. Anal. Chem. 69, 137–144. 54. Sinz, A. (2006) Chemical cross-linking and mass spectrometry to map threedimensional protein structures and protein-protein interactions. Mass Spectrom. Rev. 25, 663–682. 55. Peter, J. F., and Tomer, K. B. (2001) A general strategy for epitope mapping by direct MALDI-TOF mass spectrometry using secondary antibodies and crosslinking. Anal. Chem. 73, 4012–4019. 56. Hall, M. C., Torres, M. P., Schroeder, G. K., and Borchers, C. H. (2003) Mnd2 and Swm1 Are Core Subunits of the Saccharomyces cerevisiae Anaphase-promoting Complex. J. Biol. Chem. 278, 16698–16705.

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158 Detection of Sumoylated Proteins Ok-Kyong Park-Sarge and Kevin D. Sarge 1. Introduction Small ubiquitin–related modifier (SUMO) was discovered as a modifier of mammalian proteins in 1997 (1–2). SUMO has since been demonstrated to be a modifier of many important proteins, giving this modification a vital role in modulating a large number of important cellular processes (3–5). SUMO proteins are very similar to ubiquitin structurally, but sumoylation does not promote degradation of proteins and instead regulates key functional properties of target proteins. These properties include subcellular localization, protein partnering, and transactivation functions of transcription factors, among others (3–5). Protein sumoylation plays a particularly vital role in regulating many important processes occurring in the nucleus, and although sumoylation can be found on proteins that exist in a number of cellular compartments, most of the sumoylation characterized to date occurs on nuclear proteins (3–5). Indeed, proteins of the SUMO conjugation machinery have been found to be localized to nuclear pore complexes, in addition to other locations in the nucleus. SUMO proteins are covalently attached to lysine residues of proteins, which are generally found within the consensus motif ΨKXE where Ψ is a hydrophobic amino acid and X is any residue. Like ubiquitination, the covalent attachment of SUMO to other proteins involves a series of enzymatic steps (see Fig. 1), but the proteins involved are distinct from those in the ubiquitin conjugation pathway. First, the SUMO proteins have to undergo proteolytic processing near their C-terminal end to form the mature proteins, a step which is performed by SUMO proteases (Ulps). These proteases are dual-functional, as they are also responsible for cleaving SUMO groups from substrate proteins by cleaving the isopeptide bonds by which they are joined (3–5). The mature processed SUMO protein is covalently attached via a thioester bond to the Uba2 subunit of the heterodimeric SUMO E1 activating enzyme in an ATP-dependent reaction (6–9). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. The SUMO conjugation pathway. After they are translated, SUMO proteins must first be processed by a SUMO protease such as Ulp1, which removes four C-terminal residues so that the mature form ends with a glycine. These SUMO proteases are also responsible for removing SUMO groups from proteins. This mature form is then activated in an ATPdependent manner by forming a thioester bond with a cysteine residue in the SAE2 subunit of the heterodimeric E1 activating enzyme. Following this activation step, the SUMO moiety is transferred to the E2 conjugating enzyme ubc9. In the final step SUMO is transferred in a ligation reaction from ubc9 to substrate proteins, forming an isopeptide bond between the terminal glycine on SUMO and the ε-amino group of a lysine in the target protein. The efficiency of sumoylation of some proteins is enhanced by SUMO ligase E3 proteins, via their ability to bind both ubc9 and the target protein, thereby increasing the kinetics of the SUMO transfer.

The SUMO moiety is transferred from the E1 to ubc9, the SUMO E2 enzyme, which then binds to the ΨKXE consensus sequence in target proteins and forms an isopeptide bond between the ε-amino group of the lysine within this sequence and the carboxyl group of the C-terminal glycine of the SUMO polypeptide (10–13). SUMO E3 proteins have been identified that enhance the efficiency of SUMO attachment by interacting with both ubc9 (the E2 enzyme) and the substrate, thereby acting as bridging factors (3–5). Vertebrate cells contain three SUMO paralogs. SUMO-2 and SUMO-3 are very similar to each other in sequence, and have approximately 50% sequence identity with SUMO-1, which is the best characterized of the three vertebrate SUMO proteins. In this chapter we describe two different experimental approaches for determining whether a specific protein is sumoylated. One method employs immunoprecipitation of the protein of interest, either endogenous or transfected epitope-tagged protein, followed by Western blot with SUMO antibodies. The sond method involves incubating the protein, either as a 35S-labeled in vitro translation product or purified recombinant protein, with a reconstituted in vitro sumoylation enzymatic reaction, followed by SDS-PAGE and autoradiography or Western blot, respectively, to look for the appearance of higher molecular

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weight bands indicative of sumoylation. By comparing wild type protein constructs with those containing non-sumoylatable arginine substitutions of candidate target lysine residues, these protocols can also allow identification of those lysine residues where SUMO attachment actually occurs in a given protein. This information then provides the critical reagents for testing the functional consequences of blocking sumoylation of that particular protein. To illustrate the types of data that can be obtained using these methodologies we present figures showing the results of immunoprecipitation and in vitro sumoylation analyses of a transcription factor we study called HSF2. 2. Materials 2.1. Detection of Sumoylated Proteins by Immunoprecipitation Analysis 1. 2. 3. 4. 5. 6. 7.

Cells expressing the protein of interest. Phosphate Buffered Saline (PBS). Lysis solution: 0.15 M Tris-HCl (pH 6.7), 5% SDS, & 30% glycerol. N-ethylmaleimide. Complete protease inhibitor (Roche; Indianapolis, IN). Protein G-sepharose or protein A-sepharose. Primary antibody capable of immunoprecipitating protein of interest (or against epitope tag if analyzing a transfected tagged protein), species-matched nonspecific IgG, and antibodies against SUMO-1, SUMO-2, or SUMO-3 (Invitrogen). 8. SDS loading buffer 9. Polyacrylamide gel electrophoresis equipment and SDS-PAGE solutions. 10. Reagents for immunoblotting and detection (nonfat dried milk for blocking, and ECL or other detection system).

2.2. Detection of Sumoylated Proteins by In Vitro Sumoylation Analysis 1. Plasmid containing open reading frame of protein of interest oriented to be expressed from a T7 promoter in the vector. 2. pGEX-SUMO-1, pQE30-SUMO-1, pGEX-Ubc9, and pGEX-SAE2/SAE1 bicistronic expression construct, or purified SUMO, ubc9, SAE1/SAE2 available commercially (e.g., LAE Biotech International). 3. Ampicillin and LB media. 4. Isopropyl-β-D-thio-galactopyranoside (IPTG). 5. Phenylmethylsulfonyl fluoride (PMSF). 6. Glutathione-agarose and Ni-NTA-agarose. 7. In vitro translation kit (e.g., Promega TNT T7 Quick for PCR DNA kit). 8. Polyacrylamide gel electrophoresis equipment and SDS-PAGE solutions. 9. Whatman paper, X-ray film, and cassettes for detection of 35S-labeled proteins in sumoylation reaction.

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3. Methods 3.1. Detection of Sumoylated Proteins by Immunoprecipitation Analysis In this protocol, proteins to be tested for sumoylation are immunoprecipitated using lysis buffers designed to block the action of desumoylating enzymes (see Note 1) and then subjected to Western blot analysis using anti-SUMO antibodies to look for the appearance of a band with a size consistent with a sumoylated form of the protein. Although the theoretical molecular weight of the SUMO proteins is approximately 11 kDa, the size increase for each SUMO added on SDS-PAGE is typically in the range of 15–17 kDa. In the case of a protein with multiple sumoylation sites, or where SUMO chains form on a lysine target site (see Note 2), multiples of this size increase are expected, sometimes yielding very large shifts in mobility. Multiple bands representing different sumoylation states of the protein are also possible. This approach can be used to analyze endogenous proteins, or transfected proteins containing an epitope tag (FLAG, myc, etc.) (see Note 3). The transfection approach can also be used to determine the lysine residue(s) to which the SUMO group is attached, by comparing the sumoylation of wildtype constructs to ones in which candidate lysines have been changed to non-sumoylatable arginines. Two different lysis conditions are described, one using SDS to inhibit desumoylase enzymes and the other containing N-ethylmaleimide, a chemical inhibitor of these enzymes. 1. Tissue culture cells are grown in media appropriate for that cell line or primary cell type, and typically at least 1 × 106 cells are needed for each immunoprecipitation. 2. For harvesting place the plate of cells on ice, remove media by aspiration and add 1 ml of ice-cold PBS to the plate. Scrape cells off plates with a cell scraper, transfer to a 1.5 ml microcentrifuge tube. 3. Collect cells by centrifugation at 13,000 ×g for 30 sonds at room temperature, and remove supernatant by aspiration. 4. Lyse cells in 150 µl of lysis solution which is then diluted 1:10 in PBS/0.5% NP40 plus complete protease inhibitor (Roche) and centrifuged (16,000 g, 10 min, 4°C) to remove cellular debris. Sonication can be done prior to the centrifugation step if the lysate is highly viscous. Alternatively, the cells can be lysed in any of the standard immunoprecipitation lysis buffers (e.g., RIPA, etc.) known to extract the protein of interest, providing the lysis solution is supplemented with 20 mM N-ethylmaleimide (desumoylase inhbitor, freshly dissolved). 5. While the cell lysate is being centrifuged prepare 30 µl of 50% protein G-Sepharose (or protein A-sepharose if that is preferential for the particular antibody) as per manufacturer’s instructions. After protein G-Sepharose has been prepared add the above cell lysate to the protein G-Sepharose and rotate at 4°C for a half hour to preclear the lysate. 6. Pellet the protein G-Sepharose by centrifugation (16,000 g, 20 s, 4°C) and transfer the supernatant to a new tube. At this point take 30 µl of cell lysate, place in a separate tube with 10 µl 4 × SDS load buffer and label “input.”

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7. Divide the remainder of each treatment lysate into 2 equal amounts in separate 1.5 ml centrifuge tubes. To one of the tubes add a sufficient amount of primary antibody and to the other tube add a species-matched nonspecific IgG. Place samples on rotator at 4°C for 1 hr after which add 20 µl of PBS washed protein G-Sepharose and rotate at 4°C for 3 h. 8. Collect beads by centrifugation (16,000 g, 10 sonds, 4°C) and discard supernatant. 9. Wash the beads 4 times with PBS/0.5% NP40 plus complete protease inhibitor, or other lysis buffer if another was chosen, collecting the beads by centrifugation after each wash (16,000 g, 10 s, 4°C). Add 30 µl 2 × SDS-PAGE loading buffer to beads after removing supernatant from final wash. 10. Separate immunoprecipitated proteins by SDS-polyacrylamide gel electrophoresis. 11. Transfer proteins to nitrocellulose or nylon membrane and subject to Western blotting using antibodies against SUMO-1, SUMO-2, or SUMO-3 (commercially available). This methodology can also be used to examine the sumoylation state of an epitope-tagged version of the protein of interest being expressed in transfected cells (see Note 3). The results of immunoprecipitation analysis to examine sumoylation of the HSF2 protein is shown in Fig. 2.

3.2. In vitro Sumoylation Assay In this protocol the protein of interest is in vitro translated (typically with S-methionine incorporation) and then incubated in a reaction containing the SUMO E1 and E2 enzymes and SUMO-1, followed by SDS-PAGE and autoradiography to determine whether a lower mobility band appears that would be consistent with a sumoylated form of the target protein. Because of the high concentrations of SUMO E1 and E2 enzymes used in this in vitro sumoylation reaction, the need for SUMO E3 proteins is diminished and thus sumoylation can be detected without their addition. Performing the in vitro sumoylation reaction using mutants of the protein in which candidate sumoylation site lysine 35

Fig. 2. Detection of sumoylation by immunoprecipitation/SUMO-1 Western blot. HSF2 protein was immunoprecipitated from extracts of HeLa cells followed by Western blot using anti-SUMO-1 antibodies. The positions of molecular weight standards are indicated on the left side of the panel. (Reprinted with permission from ref. 15).

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residues are changed to nonsumoylatable arginines can be used to determine the site(s) at which SUMO attachment is occurring (e.g., see Fig. 4). The expression and purification of the recombinant proteins required for the in vitro sumoylation assay is described in Subheading 3.2.1, and the protocol for performing assay itself is described in Subheading 3.2.3. 3.2.1. Expression If Recombinant SUMO, ubc9 (SUMO E2), and SAE1/SAE2 (SUMO E1) Heterodimer The SUMO proteins are expressed and purified as fusion proteins with GST or 6xHis at the N-terminal end of the SUMO proteins, and these affinity tags do not need to be removed prior to using these SUMO proteins for the sumoylation reaction. The size difference of GST-SUMO vs. 6xHis-SUMO can also provide a useful control for the sumoylation reaction as it yields a predictable size difference between the sizes of the sumoylation products (e.g., see Fig. 3). The SUMO E1 is a heterodimer of SAE1/SAE2, and is active when expressed in E. coli from a bicistronic contruct of GST-SAE2 and untagged SAE1; the two proteins complex and can be purified using glutathione-agarose affinity chromatography (14). The SUMO E2 can be expressed which appears to be more active, at least in our hands when the GST tag is removed by thrombin cleavage. The following is the general protocol for expressing and purifying these recombinant proteins needed for the in vitro sumoylation assay. Once purified these recombinant proteins can be stored at −80°C for extended periods of time. 1. Transform expression construct into DH10B E. coli cells using standard molecular biology methods. 2. Plate cells on LB plates containing ampicillin and incubate overnight at 37°C. 3. Select single colonies and grow overnight at 37°C in LB media containing ampicillin. 4. Inoculate individual liters of LB containing ampicillin with aliquots (5 mL) of overnight culture and grow cells to an O.D.600 nm0.6–0.8. 5. Induce the cells with IPTG (1 mM) for 3 hs. 6. After 3 hr induction harvest cells by centrifugation (4,000 g; 5 min; 4°C) and resuspend pellet in PBS (4°C) then repellet cells. At this point the cell pellet can be extracted or stored at −80°C. 7. To extract protein from the cells they are resuspended in PBS (4°C) with PMSF (phenylmethylsulfonyl fluoride) to a final concentration of 1 mM. 8. Pass cells through a French press at 10,000 to 14,000 psi, and repeat to ensure complete lysis. 9. Centrifuge the cell lysate (30,000 g;1 h; 4°C) and retain the supernatant. 10. Purify proteins using standard nondenaturing affinity chromatography techniques suitable for their fusion tags (glutathione agarose affinity chromatography for GST-fusion proteins and Ni-NTA-agarose affinity chromatography for 6 × Hisfusionprote ins).

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Fig. 3. Analysis of SUMO-1 modification by reconstituted in vitro sumoylation reaction. (A) In vitro translated 35S-labeled HSF2 protein was incubated with HeLa cytosol (as a source of E1), Ubc9, SUMO-1, SUMO-2 or with various combinations of each of these, and then subjected to SDS-PAGE followed by autoradiography. The positions of unmodified and SUMO-modified HSF2 are indicated to the right of the panel. (B) In vitro translated 35S-labeled HSF2 protein was subjected to the in vitro SUMO-1 modification assay using either 6xHis-SUMO-1 or GSTSUMO-1 as the SUMO-1 substrate for the reaction. (Reprinted with permission from ref. 15).

11. Check purified proteins by Coomassie blue staining of a SDS-PAGE gel (GSTSUMO1 = 38 kDa, GST-Ubc9 = 44 kDa, and 6His-SUMO1 = 14 kDa). 12. For the in vitro sumoylation assay the GST-Ubc9 needs to be thrombin cleaved to remove the GST-tag. This can be done following standard protocols. Check thrombin cleavage of GST-Ubc9 by Coomassie staining of a SDS-PAGE gel (Ubc9 = 18 kDa).

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Fig. 4. In vitro translated 35S-labeled wildtype HSF2 protein and the HSF2 SUMO-1 consensus site mutants K82R, K139R, and K151R were used as substrates in in vitro SUMO-1 modification reactions. The positions of unmodified and SUMO-modified HSF2 proteins are indicated to the right of the panel. (Reprinted with permission from ref. 15).

3.2.2. In Vitro Sumoylation of Rabbit Reticulocyte System Translated Proteins The in vitro sumoylation assay uses 35S-methionine radiolabelled protein as the substrate. We generate radiolabelled protein using the TNT T7 Quick for PCR DNA kit (Promega, Madison WI) following the manufacturers instructions. Described below is the SUMO modification procedure to be utilized with radiolabelled translated proteins. 1. Translate target protein fresh before each sumoylation assay following the manufacturer’s instructions. Place freshly translated protein on ice until step 3. 2. For each sumoylation reaction prepare a mixture containing the following (on ice): 1 µl 10× sumoylation buffer, 0.4 units creatine phosphokinase, 10 mM creatine phosphate, 0.6 units inorganic pyrophosphatase, 100 ng purified SAE1/SAE2 heterodimer (E1), 400 ng purified ubc9 (E2), 1 µg purified 6 × His-SUMO-1, and H2O to 10 µl. 10× sumoylation buffer contains 500 mM Tris-HCl (pH 7.6), 500 mM KCl, 50 mM MgCl2, 10 mM DTT, and 10 mM ATP. As a negative control, make another mix identical to the one above except that it lacks SUMO protein. If desired a third mix can be made in which GST-SUMO-1 is added instead of 6 × His-SUMO. 3. Set up the reactions (no SUMO control, +6 × His-SUMO, and +GST-SUMO) by adding 2 µl in vitro translated protein to the 10 µl reaction mixes, and then incubate at 37°C for 1 hr. 4. Terminate the reaction by adding 12 µl 2 × SDS-PAGE load buffer. Store at −20°C until gel electrophoresis. 5. Boil the samples for 5 min and separate on SDS-PAGE gel. After electrophoresis the gel is fixed for 10 min in SDS-PAGE fixing solution (50% methanol/10% acetic

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acid), then dried on Whatman paper, and finally placed on x-ray film.35S emissions are low energy and so it is advisable not to leave plastic wrap between the dried gel and the X-ray film if possible, as this can increase required exposure times. 6. One important experiment to do to give confidence in the in vitro sumoylation assay is to do a reconstitution test where each of the components required for sumoylation (E1, ubc9, SUMO-1) are individually left out of the reaction and compared to a reaction where all components are present (Fig. 3A). As shown in this figure, such an experiment can also reveal the relative efficiency of sumoylation of your target protein by different SUMO proteins (e.g., SUMO-1 vs. SUMO-2). Another experiment which increases confidence that your protein is indeed being sumoylated vs. being targeted by some other modification is to compare the effect of using of 6 × His-SUMO vs. GST-SUMO as the donor SUMO, because this gives a predictable size shift between the sumoylated reactions (Fig. 3B). 7. To identify the sumoylation site in the protein of interest site directed mutagenesis is done to change candidate lysine (changed to arginine) in the sumoylation consensus sequence (ΨKXE) to non-sumoylatable arginine residues utilizing protocols such as the Quickchange Site Directed Mutagenesis Kit (Stratagene). The extent of sumoylation of the wild type vs. lysine-to-arginine mutant can then be compared using either the transfection/immunoprecipitation approach or the in vitro sumoylation assay. Results of such an experiment using the in vitro sumoylation assay are shown in Fig. 4.

4. Notes 1. SUMO-modified proteins are highly susceptible to SUMO proteases. The SDS in the lysis buffer described in this protocol inactivates the SUMO proteases allowing for easier detection of sumoylated proteins. However, a common complication with the SDS-lysis method is that the cell lysates tend to be very viscous and sticky due to genomic DNA in the lysate. This problem is remedied by brief sonication which shears the DNA and makes the samples easier to manipulate. SUMO proteases can also be inhibited by the addition isopeptidase inhibitor N-ethylmaleimide (20 mM) to standard lysis buffers such as NP-40 lysis buffer (50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 1% NP-40) if another lysis buffer besides the SDS-lysis is more desirable. 2. The size of putative sumoylated forms of a protein on SDS-PAGE will depend on how many different SUMO attachment sites the protein has. In addition, SUMO-2 and SUMO-3 have been reported to form polymeric chains reminiscent of ubiquitin, which could result in large increases in size for a sumoylated protein on SDS-PAGE compared to the non-sumoylated form (14). Thus, it is possible to observe bands that are multiples of the approximate 15–17 kDa size of each SUMO unit, as well as multiple bands representing different sumoylation states.

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3. Investigating sumoylation may also be done using cells transfected with fusion-tagged plasmid constructs of the protein thought to be sumoylated with immunoprecipitation utilizing fusion tag antibodies which are available from commercial sources (e.g., GFP, FLAG, Myc, and 6 × His tags). In these types of experiments it is advisable to co-transfect the cells with a SUMO expression construct (often this is epitope-tagged) to ensure that sufficient SUMO protein is present in the transfected cells to allow for efficient sumoylation of the transfected target protein being tested. Acknowledgments We are very grateful to Mike Matunis (Johns Hopkins), Ron Hay (University of St. Andrews), and Chris Lima (Sloan Kettering Institute) for providing constructs and reagents.

References 1. Mahajan, R., Delphin, C., Guan, T., Gerace, L., and Melchior, F. (1997) A small ubiquitin-related polypeptide involved in targeting RanGAP1 to nuclear pore complex protein RanBP2. Cell 88, 97–107. 2. Matunis, M. J., Wu, J., and Blobel, G. (1998) SUMO-1 modification and its role in targeting the Ran GTPase-activating protein, RanGAP1, to the nuclear pore complex. J Cell Biol 140, 499–509. 3. Mukhopadhyay, D. and Dasso, M. (2007) Modification in reverse: the SUMO proteases. Trends Biochem. Sci. 32, 286–295. 4. Bossis, G. and Melchior, F. (2006) SUMO: regulating the regulator. Cell Div. Jun 29, 1–13. 5. Kerscher, O., Felberbaum, R., and Hochstrasser, M. (2006) Modification of Proteins by Ubiquitin and Ubiquitin-Like Proteins. Annu. Rev. Cell Dev. Biol. 22, 159–180. 6. Johnson, E.S., Schwienhorst, I., Dohmen, R.J., and Blobel, G. (1997) The ubiquitin-like protein Smt3p is activated for conjugation to other proteins by an Aos1p/ Uba2p heterodimer. EMBO J. 16, 5509–5519. 7. Desterro, J.M., Rodriguez, M.S., Kemp, G.D., and Hay, R.T. (1999) Identification of the enzyme required for activation of the small ubiquitin-like protein SUMO-1. J. Biol. Chem. 274, 10618–10624. 8. Gong, L., Li, B., Millas, S., and Yeh, E.T. (1999) Molecular cloning and characterization of human AOS1 and UBA2, components of the sentrin-activating enzyme complex. FEBS Lett. 448, 185–189. 9. Okuma, T., Honda, R., Ichikawa, G., Tsumagari, N., and Yasuda, H. (1999) In vitro SUMO-1 modification requires two enzymatic steps, E1 and E2. Biochem. Biophys. Res. Commun. 254, 693–698. 10. Desterro, J.M., Thomson, J., and Hay, R.T. (1997) Ubch9 conjugates SUMO but not ubiquitin. FEBS Lett 417, 297–300. 11. Johnson, E.S. and Blobel, G. (1997) Ubc9p is the conjugating enzyme for the ubiquitin-like protein Smt3p. J. Biol. Chem. 272, 26799–26802.

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12. Rodriguez, M.S., Dargemont, C., and Hay, R.T. (2001) SUMO-1 conjugation in vivo requires both a consensus modification motif and nuclear targeting. J. Biol. Chem. 276, 12654–12659. 13. Sampson, D.A., Wang, M., and Matunis, M.J. (2001) The small ubiquitin-like modifier-1 (SUMO-1) consensus sequence mediates Ubc9 binding and is essential for SUMO-1 modification. J. Biol. Chem. 276, 21664–21669. 14. Tatham, M.H., Jaffray, E., Vaughan, O.A., Desterro, J.M., Botting, C.H., Naismith, J.H., and Hay, R.T. (2001) Polymeric chains of SUMO-2 and SUMO-3 are conjugated to protein substrates by SAE1/SAE2 and Ubc9. J. Biol. Chem. 276, 35368–35374. 15. Goodson, M.L., Hong, Y., Rogers, R., Matunis, M.J., Park-Sarge, O.K., and Sarge, K.D. (2001) Sumo-1 modification regulates the DNA binding activity of heat shock transcription factor 2, a promyelocytic leukemia nuclear body associated transcription factor. J. Biol. Chem. 276, 18513–18518.

159 Efficient Enrichment of Intact Phosphorylated Proteins by Modified Immobilized Metal-Affinity Chromatography Anna Dubrovska 1. Introduction The lack of an efficient technique to enrich for phosphorylated proteins is limiting of phosphoproteomes studies. Enrichment with antiphospho serine, antiphospho threonine, or anti-phospho tyrosine antibodies depends on the affinity and specificity of antibodies that limit comprehensiveness of the analysis (1–3). Metabolic labeling of cells with inorganic (32P) phosphate, followed by an analysis of the whole proteome, requires efficient separation to exclude comigration with nonphosphorylated proteins (4,5). Chemical modifications of phosphorylated residues in the peptides, followed by MS, allow identification of phosphorylation sites (1,5–8). However, these techniques do not provide information about full length proteins, e.g., molecular mass of intact proteins. Immobilized metal-affinity chromatography (IMAC) has been successfully used for enrichment of phosphopeptides (1,5,9,10), but the efficiency of purification of phosphoproteins has been low, indicating significant losses of phosphoproteins (5,9,10). Another important limitation of available IMAC methods is that bound proteins are eluted into a solution which is incompatible with further separation by 2-DE. A modified IMAC technique developed in our laboratory makes possible efficient and comprehensive analysis of phosphoproteins, and is compatible with 2-DE (11).

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2. Materials 2.1. Cell Culture and Lysis 1. Clonetics® Normal Human Mammary Epithelial Cell Systems (Lonza Walkersville, Inc.). 2. Cell lysis buffer: 1% (v/v) Triton X100, 150 mM NaCl, 10 mM Tris-HCl, pH 8.0, protease inhibitors (Roche Diagnostics; complete inhibitor cocktail tablets EDTAfree) (see Note 1). Store at 4°C. Add protease inhibitors just prior to use. 3. Transforming growth factor-β1 (TGFβ1, Peprotech) is reconstituted with sterile 4 mM HCl containing 1 mg/mL BSA to a final concentration 50 µg/mL and stored in single use aliquots at −80°C, and then added to tissue culture dishes at the concentration5 ng/mL. 4. Radioactive orthophosphate ( (33P) or (32P) orthophosphate) (GE Healthcare Uppsala, Sweden).

2.2. Preparation of Fe-NTA Agarose 1. NTA agarose (Qiagen, VWR International AB). 2. 6 M guanidine-HCl in 0.2 M acetic acid. Store at room temperature. 3. Water solution of 2% (w/v) sodium dodecyl sulfate (SDS). Store at room temperature. 4. 25%, 50%, 75% and 100% (v/v) ethanol-water solutions. 5. Aqueous solution of 100 mM ethylenediamine tetraacetic acid (EDTA), pH 8.0 6. Solution of 0.1 M FeCl3 in 0.1 M acetic acid. Prepare just prior to use. 7. Aqueous solution of 0.1 M acetic acid. 8. Rotor shaker. 9. Empty econo-pac chromatography columns (Bio-Rad, Hercules, CA).

2.3. Phosphoprotein Enrichment Procedure 1. Washing buffer: 5 mM Tris-HCl, pH 6.3, 150 mM NaCl. Store at 4°C. 2. Elution buffer: 100 mM dithiotriethol (DTT), 2% (w/v) SDS, 50 mM Tris-HCl, pH 6.8. Prepare just prior to use. 3. Methanol 100%. 4. Glacial acetic acid. 5. Aqueous solution of 25% (v/v) methanol, 10% (v/v) acetic acid. 6. Quick Start Bradford Protein Assay Kit (Bio-Rad, Hercules, CA).

2.4. 2-DE Electrophoresis 1. Rehydration solution: 8 M urea, 2.5% (w/v) CHAPS, 0.28% (w/v) DTT, 0.5% (v/v) IPG buffer, pH 3–10 (GE Healthcare), 0.002% bromophenol blue. Store at −80°C. Add DTT just prior to use. 2. DTT or iodoacetamide-containing equilibration buffer: 2% SDS, 50 mM Tris-HCl pH 8.8, 6 M urea, 30% (v/v) glycerol, 0.002% bromophenol blue. Store in 10 ml aliquots at −80°C. Add DTT to 1% or iodoacetamide to 2.5% just prior to use.

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3. SDS electrophoresis buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS, pH 8.3. Store at room temperature. 4. Agarose sealing solution: 0.5% agarose, 0.002% bromophenol blue in SDS electrophoresis buffer. 5. 4× resolving gel buffer: 1.5 M TrisHCl, pH 8.8. Filter solution through a 0.45 µm filter. Store at 4°C. 6. 10% ammonium persulfate. 7. 10% SDS. 8. Acrylamide/bis acrylamide solution (37.5:1; T = 40%, C = 2.6%). 9. N,N,N,N’- Tetramethyl-ethylenediamine (TEMED) (GE Heralthcare, Uppsala, Sweden). 10. Water-saturated isobutanol. Shake equal volumes of water and isobutanol in a glass bottle and allow to separate. Use a top layer. Store at room temperature. 11. Prestained molecular weight markers (New England BioLabs, Ipswich, MA). 12. IPGDry strips with immobilized pH gradient, pH range 3–10, 18 cm, linear (GE Healthcare, Uppsala, Sweden). 13. Ettan IPGphor first-dimension isoelectric focusing system (GE Healthcare, Uppsala, Sweden). 14. Ettan DALTsix Electrophoresis System (GE Healthcare, Uppsala, Sweden). 15. ImagePlatinum software (GE Healthcare, Uppsala, Sweden).

2.5. Silver Staining Procedure 1. Fixing solution: 20% (v/v) methanol, 10% (v/v) acetic acid. 2. Sensitizing solution 30% (v/v) ethanol, 0.2% (w/v) sodium thiosulphate (Na2S2O3), 6.8% (w/v) sodium acetate (CH3COONa). 3. Silver solution: 0.25% silver nitrate, AgNO3. 4. Developing solution: 2.5% (w/v) sodium carbonate (Na2CO3), 0.015% (v/v) formaldehyde. Add formaldehyde immediately before use. 5. Gel drying solution: 30% (v/v) methanol, 5% (v/v) glycerol. 6. Cellophane sheets. 7. Rocking platform. 8. Fuji ImageReader software (Fuji). 9. FujiX2000 PhosphorImager (Fuji). 10. AIDA 2D Densitometry software (Bio Imaging).

2.6. Western Blotting for Phosphorylated Proteins 1. 2. 3. 4. 5. 6.

Criterion precast gel system (Bio-Rad, Hercules, CA). Criterion TrisHCl Gels, 12.5% (Bio-Rad, Hercules, CA). Criterion electrophoresis cell (Bio-Rad, Hercules, CA). Criterion blotter cell (Bio-Rad, Hercules, CA). Hybond P membrane (Amersham Biosciences, Piscataway, NJ). Tris-buffered saline with Tween (TBS-T): make 10× stock solution with 1.37 M NaCl, 27 mM KCl, 250 mM Tris-HCl, pH 7.4, 1% Tween-20. Dilute 100 mL with 900 mL water to use.

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7. Blocking solution and antibody dilution buffer: 5% (w/v) BSA fraction V in TBS-T. 8. Primary antibody: antiphospho tyrosine; antiphospho threonine (Santa Cruz, CA); anti-phospho serine (Sigma-Aldrich). 9. Secondary antibody: Antimouse IgG conjugated to horse radish peroxidase (GE Healthcare). 10. Chemiluminescence Luminol Reagent (Santa Cruz, CA). 11. Kodak BioMax MR film (Sigma-Aldrich).

3. Methods The described Fe-IMAC procedure permits significant enrichment of phosphoproteins, and allows generation of good quality 2-D gels. Combined with the simplicity of the procedure described here, it provides a tool for the efficient and comprehensive analysis of phosphoproteins. This technique may be applied to various types of biological material from cultured cells to tissues. 3.1. Preparation of Fe-NTA Agarose NTA or Ni-NTA agaroses are commercially available from Qiagen and can be used for preparation of Fe-NTA resin using the following procedure: 1. Place the column on a stand above the capacious beaker for flow-through collection. Add 50% NTA agarose slurry in the column on the basis of 1 mL the beads per 1.5 mL of total cell lysate. 2. Wash the column with 2 volumes of 6 M guanidine-HCl in 0.2 M acetic acid. 3. Wash the column with 5 volumes of ddH2O. 4. Wash the column with 3 volumes of 2% SDS. 5. Wash the column with 1 volume of 25% ethanol. 6. Wash the column with 1 volume of 50% ethanol. 7. Wash the column with 1 volume of 75% ethanol. 8. Wash the column with 5 volumes of 100% ethanol. 9. Wash the column with 1 volume of 75% ethanol. 10. Wash the column with 1 volume of 50% ethanol. 11. Wash the column with 1 volume of 25% ethanol. 12. Wash the column with 1 volume of ddH2O. 13. Wash the column with 5 volumes of 100 mM EDTA, pH 8.0. 14. Wash the column with 10 volumes of ddH2O. 15. Wash the column with 5 volumes of 0.1 M acetic acid. 16. Wash the column with 5 volumes 0.1 M FeCl3 in 0.1 M acetic acid. 17. Close the column tip and add 5 volumes 0.1 M FeCl3 in 0.1 M acetic acid. 18. Close the upper side of column and incubate the NTA beads with 0.1 M FeCl3 solution on rotor shaker for 16 h at room temperature. 19. Wash the column with 5 volumes of 0.1 M acetic acid. Beads can be stored in 0.1 M acetic acid at 4°C for 2 weeks.

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20. Wash the column with 5 volumes of lysis buffer (1% Triton X100, 150 mM NaCl, 10 mM TrisHCl, pH 8.0, protease inhibitors). Washed Fe-NTA beads can be used for phosphoprotein enrichments procedure.

3.2. Metabolic Labeling To monitor phosphorylated proteins, we performed metabolic labeling of MCF10A human breast epithelial cells with radioactive (33P) orthophosphate or (32P) orthophosphate. (33P) isotope has lower energy, as compared to (32P) isotope, which may be of importance for safety considerations (see Note 2). 1. The MCF10A cells were growing to confluence in mammary epithelial growth medium (MEGM) in 150 mm dishes. 2. The cells were metabolically labeled with serum free MEGM medium supplemented with 0.25 mCi/mL (32P) inorganic phosphoate for 16 h to ensure an efficient incorporation of radioactive isotope into the proteins. 3. The cell were treated or not with 5 ng/ml TGFβ1 for 3 h to initiate TGFβ1dependent protein phosphorylation. 4. The cells were washed 3 times with cold washing buffer (see Note 3). 5. Proteins were extracted from cells using lysis buffer (3 mL of lysis buffer for 15 cm dish). To provide complete protein extraction cell were incubated with lysis buffer on ice for 30 min with agitation and the material was scraped into the appropriate labeled tubes.

3.3. Phosphoprotein Enrichments Procedure for the Cell Culture It is important to pick up a small amount of each chromatography fraction (30–60 mkl) for the next analysis. Make a note about total volume of each fraction taken for analysis as it might be helpful for estimation of phosphoprotein binding. 1. Cell extracts were clarified by centrifugation at 13000 g for 20 min at 4°C, and were incubated with washed Fe-IMAC beads for 12 h at 4°C on rotor shaker. We applied 3 mL of lysate with average concentration of total proteins 0.85 mg/ml for 2 mL of Fe-IMAC beads directly into the chromatography column (see Note 4). 2. After incubation with cell extract, Fe-IMAC beads were washed six times with washing buffer. Washing procedure was performed at room temperature. The bound proteins were eluted by incubation with a buffer containing 100 mM dithiotriethol (DTT), 2% SDS, 50 mM TrisHCl, pH 6.8 for 5 min at 95°C (see Note 5). 3. Eluted proteins were precipitated by adding of methanol to 25% and acetic acid to 10% directly to the samples followed by incubation on ice for 30 min (see Note 6). 4. Precipitated proteins were centrifuged at 13000 g for 30 min at 4°C. 5. The pellet was washed twice with 25% (v/v) methanol, 10% (v/v) acetic acid solution followed by centrifugation at 13000 g for 30 min at 4°C (1 mL of precipitation solution per 1.5 mL tube).

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6. The result white protein-containing pellet was dried in vacuum centrifuge to remove traces of a precipitation solution (see Note 7). 7. The dried protein pellet was dissolved in rehydration solution. 8. Complete solubilization of proteins required 5–12 h vortexing at room temperature; prolonged drying in a vacuum centrifuge increased the time of solubilization.

3.4. Phosphoprotein Enrichments from Tissue The Fe-IMAC technique described here is also suitable for enrichment of phosphoproteins from tissues. 1. Frozen tissue (0.5- 1 cm3) was placed in Petry dish in 3 mL of ice-cold lysis buffer and cut into small pieces (1 mm3) with scalpel. 2. Tissue pieces were transferred in 50 mL Falcon tube and sonicated on ice by performing 10 cycles at 30 s/cycle with 1 min interval between the cycles to cool down the tube content. 3. Tissue extract was clarified by centrifugation at 13000 g for 20 min at 4°C, and protein concentration was measured. 3 mL of the tissue lysate with protein concentration 0.85 mg/mL was incubated with 2 mL of Fe-IMAC beads for 12 h at 4°C on rotor shaker. 4. All the next stages of the procedure are the same as described above for the cell culture. Using this described technique we observed 81% recovery of phosphoproteins extracted from chicken liver, as evaluated by immunoblotting with antithreonine antibodies (Fig. 2E). An example result for 2-DE of phosphoproteins enriched from chicken liver is shown in Fig. 2F and 2G.

3.5. Evaluation the Efficiency of Phosphoprotein Enrichment To evaluate the efficiency of phosphoprotein enrichment by Fe-IMAC, we performed 1-DE and 2-DE, followed by monitoring of phosphorylated proteins by Western blotting or authoradiography. 3.5.1. Silver Staining Procedure 1. To estimate the efficiency of phosphoprotein binding to and elution from Fe-IMAC beads all fraction of Fe-NTA based chromatography were taken for 1-DE (input, flow-through, washes, elution and combined elution fraction). Combined elution fraction is the resulted protein solution obtained after dissolving the washed protein pellet in rehydration solution. 2. Equal volume of each chromatography fraction (50 mkl) was taken for SDS-PAGE separation using Criterion electrophoresis system. 3. For silver staining the gel was placed in a tray containing fixing solution and agitated on a shaker for at least 2 h. 4. The fixing solution was drained from the tray and the gel was washed ddH2O for 10 min. 5. The sensitizing solution was added and gel was agitated in sensitizing solution for 30 min.

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6. The sensitizing solution was drained from the tray and the gel was washed with ddH2O 4 times for 15 min. 7. The gel was incubated in silver solution for 30 min. 8. After 30 min, the silver nitrate solution was drained into an appropriate waste beaker and gel was washed with ddH2O two times, 1 min each time. 9. The developing solution was added to the gel, and agitated until tiny yellow or brown precipitate appears. Then developer was poured off, add fresh developer was added. The developing step was continued until desired intensity of protein staining is achieved. 10. Adding the fixing solution terminated the staining. 11. The gel was then agitating in drying for 20 min. 12. The soaked gel was placed between the wet cellophane sheets and dried on the gel drier at 65°C for 1 h (see Note 8).

3.5.2. Estimation a Chromatography Efficiency Using 1-DE Gels 1. To monitor efficiency of phosphoprotein binding FE-NTA chromatography fractions were separated by 1-DE and the gels were exposed in a FujiX2000 PhosphorImager. Examples of the silver stained gel and autoradiography are shown in Fig. 1A and 1B. 2. Quantification of radioactivity associated with proteins separated by SDS-PAGE was performed using Fuji ImageReader software, and was normalized to volumes of fraction using the formula Appf = D/V, where Appf is an amount of phosphoproteins in each fraction, D is densitometric intensity of this fraction and V is a fraction volume. 3. AIDA 2D Densitometry module (Bio Imaging) was used to evaluate the relative amount of total proteins in each chromatography fraction. Data obtained by densitometry of silver stainined gel and autoradiography were used to calculate an yield of phosphoproteins (Ypp), concentration factor (Cpp) and and level of phosphoproteins in the elution fraction (Epp) as shown in Fig. 1C.

3.5.3. Monitoring the Phosphoprotein Binding by Western Blotting for Phosphorylated Proteins 1. Criterion electrophoresis system was used for protein SDS-PAGE separation and blotting. 2. Once the transfer was completed, the nitrocellulose membrane was washed with 1× TBS-T with agitation 3 times for 15 min each wash. 3. After washing the membrane was blocked with 5% BSA in TBS-T for 1 h at room temperature on a rocking platform. 4. The blocking buffer was discarded and the membrane was incubated with primary antibody for antiphospho tyrosine (dilution 1:500), anti-phospho threonine (dilution 1:500), antiphospho serine (dilution 1:1000), anti-Smad2 and antiphospho Smad2 proteins. Anti-Smad2 and antiphospho Smad2 antibody were prepared in our institute and used with dilution 1:500. All antibodies were diluted in 5% BSA in TBS-T. The membrane was incubated with primary antibody solution overnight at 4°C on a rocking platform (see Note 9).

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Dubrovska C

A

Cpp = (A/B) / (C/D)=2.3 Ypp=(D+F)/Bx100%=70-90% Epp=D/Cx100%=70-90%

relative amount of proteins

3500

B

3000 total proteins phosphoproteins

2500 2000 1500 1000 500

elu tio elu n 1 tio n 2

washings

flo

w

in p th ut ro ug h

0

FeNTA chromatography fractions

D

G

E

F

Fig. 1. Fe-IMAC allows efficient enrichment of phosphoproteins (A, B). Aliquots of the whole cell lysate (input; 1/50 of the input), flow-through (fl-thr; 1/50 of the fraction), washes, elution fractions from Fe-IMAC beads (elution; 1/50 of the elution fraction), and elution sample prepared for 2-DE (comb. elution fraction; 1/50 of the sample) were subjected to SDS-PAGE, as indicated. Gels were dried and exposed in a phosphorimager. An image of 32P-labeled proteins (A) and an image of silver-stained gel (B) are shown.

Efficient Enrichment of Intact Phosphorylated Proteins

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5. The membrane was washed 3 times with TBS-T for 15 min each wash. 6. The membrane was incubated with secondary antibody 30 min at room temperature on the rocking platform. The secondary antibody was freshly prepared prior to use with dilution 1:10000 in 5%BSA in TBS-T. 7. The membrane was washed 3 times with TBS-T for 15 min each wash. 8. The membrane was then incubated with Chemiluminescence Luminol Reagent (Santa Cruz, CA), and proteins were visualized by a suitable exposure with Kodak BioMax MR film. An example result is shown in Fig. 1D, 1E, 1F, and 1G.

3.5.4. Separation of Eluted Fraction with 2-DE 1. Prepared samples (eluted proteins solubilized in rehydration solution from step 3.3) were subjected to isoelectric focusing using IPGDry strips with immobilized pH gradient, pH range 3–10, 18 cm, linear (GE Healthcare, Uppsala, Sweden) according to the following protocol: rehydration 10 h; 50 V, 3 h; 1000 V, 1 h; 8000 V, 10 h or to 50,000 Vh. 2. 2-D SDS–PAGE was performed in 12% polyacrylamide gels. The Ettan DALTsix gel caster was used to cast the gels 5–13 h prior to 2-D electrophoresis. Gels were stored at room temperature prior to use. 3. The 2-D gels were run during the day (about 7 h) at constant power 5W per gel through the sealing agarose and 60 W in total through the acrylamide gel. 4. After the electrophoresis, gels were fixed in 10% acetic acid and 20% methanols for 10–12 h. Proteins were detected by silver staining, as described above. 5. To monitor efficiency of phosphoprotein enrichment the gels were exposed in a FujiX2000 PhosphorImager for 18 h.

Fig. 1. (continued) Data obtained by densitometry of silver stainined gel and autoradiography were used to calculate an yield of phosphoproteins (Ypp), concentration factor (Cpp) and level of phosphoproteins in the elution fraction (Epp) as shown in panel (C), where A, C, and E is densitometrical intensity of the input fraction, elution fraction 1 and elution fraction 2 correspondingly for silver stained gel; B, D, and F is densitometrical intensity of the input fraction, elution fraction 1 and elution fraction 2 correspondingly for autoradiography. To monitor phosphorylated proteins, the gels were subjected to immunoblotting with antibodies specific to phosphotyrosine (D), to phosphoserine (E), or to phosphothreonine (F). To monitor TGFb-induced phosphorylation of Smad2, MCF10A cells were treated or not with TGFb1 (5 ng/mL), as indicated, and phosphoproteins were enriched with Fe-IMAC procedure (G). Collection of fractions and gel electrophoresis were performed as described in panels A–E. Immunoblotting was performed with antibodies specific to phosphorylated Smad2 (phosphorylated Smad2) or to total Smad2 (Smad2), as indicated. Lanes corresponding to the whole cell extract (input), flow-through (fl-thr), washing fractions (washes), and elution fractions (comb. elution fraction) are indicated. Combined elution fractions represent samples prepared for 2-DE (precipitated and solubilized elution fraction). Longer exposure with anti-Smad2 antibodies indicates presence of Smad2 in elution fractions. (Reproduced with permission from ref. 11. Copyright Wiley-VCH Verlag GmbH & Co. KgaA.)

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3.5.5. Estimation a Chromatography Efficiency Using 2-DE Gels 1. We performed 2-DE with eluted fractions, measured quantity-reflecting values of the silver-stained protein spots (quantitating volumes, as defined by the ImagePlatinum software, GE Healthcare), and matched them with the radioactively labeled spots, which were detected in a phosphorimager. 2. Then, we compared the quantitating volume of proteins in silver-stained spots which corresponded to radioactively labeled proteins, to the total quantitating volume of silver-stained proteins in 2-D gels. We found that up to 90% of proteins of elution fractions, which were separated by 2-DE, were phosphorylated (Fig. 2A and 2B). 3. The elution procedure allows the generation of good quality 2-D gel. In 2-D gels of proteins prepared with Fe-IMAC technique, we detected in average 500 33P- or 32 P-labeled proteins (Fig. 2B) and in average 700 silver-stained spots (Fig. 2A). For comparison, we identified in average only 240 32P-labeled proteins in 2-D gels of whole cell lysate from MCF10A cells (Fig. 2D). Silver staining revealed in average 1400 proteins spots in the same gel (Fig. 2C). 4. To make sure that Fe-IMAC technique described here is compartible with massspectrometry, we randomly selected and identified more than 50 proteins from 2-D gels, which are shown in Fig. 2, using MALDI-TOF MS (Table 1 shows identification of six proteins, as example). Searches of PubMed and prediction of phosphorylation sites with NetPhos showed that the identified proteins were reported as phosphorylated in cells and/or have a number of well-defined phosphorylation sites (Table 1). Together with 33P- and 32P-metabolic labeling and immunoblottings with anti-phospho amino acids antibodies (Figs. 1 and 2), this confirms efficient enrichment of phosphoproteins with Fe-IMAC.

4. Notes 1. It is important to use EDTA-free inhibitor cocktail. Presence of EDTA in lysis buffer may prevent phosphoprotein binding. 2. Be sure that you have a permission to work with radioactive material, otherwise contact your lab coordinator. 3. Do not use 1×PBS (phosphate buffered saline) for cell washing. Nonorganic phosphates may interfere with phosphoprotein binding by Fe-NTA resin and cause significant losses of phosphoproteins. 4. Clarified cell lysate has been incubated with Fe-NTA beads directly into the chromatography column. Be sure that column tip is closed prior to add cell lysate. 5. The tubes content must change color from dark to white otherwise extend incubation time to 10 min to make sure that all bound phosphoproteins were eluted. In average 0.3–0.4 mg of proteins were eluted when 2.5 mg of cellular proteins were taken as the input. The composition of the elution buffer was optimized for the most efficient recovery of proteins, and to

Efficient Enrichment of Intact Phosphorylated Proteins

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be compatible with further 2-DE separation. Elution with EDTA, phosphate salts, or with a buffer containing only SDS were found to be incompatible with 2-DE. 6. Shake the sample immediately after adding of methanol and acetic acid. The flakes of precipitated proteins should appear after incubation the samples on ice. Acetone or ethanol precipitation resulted in formation of

Fig. 2. Fe-IMAC protocol improves resolution of phosphorylated proteins in 2-DE. Images of 2-D gels obtained with samples of MCF10A cells prepared according to the Fe-IMAC protocol (A,B), prepared from the whole MCF10A cell extract (C,D), and prepared from chicken liver (F,G) are shown. Radioactive orthophosphate labeled proteins of MCF10A cells were detected after exposure gels in a phosphorimager, and images are shown in panels B and D. Images of the same gels after silver staining are shown in panels A and C. Arrows show migration positions of proteins listed in Table 1, as examples of identified proteins. Annotation of proteins (spot numbers) is as in Table 1.

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Fig. 2. (continued) Fe-IMAC-based enrichment of phosphoproteins from chicken liver was monitored by 1-DE, followed by immunoblotting with antiphosphothreonine antibodies (E). Images of silverstained 2-D gels obtained with Fe-IMAC sample (F), and with whole liver extract (G) are shown. 2-DE was performed as described by Kanamoto et al. (13). Directions of isoelectrofocusing (pH) and SDS-PAGE are indicated (directions in panel A are also for gels in panels A–D). (Reproduced with permission from ref. 11. Copyright Wiley-VCH Verlag GmbH & Co. KgaA.)

NudE Anaphase -promoting complex (APC, subunit 8) GTP-bind ing protein alpha q subunit HSP 70 (mortalin) Cytohesin1 B2-1 Grb14

b)

1.0e 1 000 1.0e 1 000

Probability

AAH30634.1 1.0e 1 000 1bc9 1.0E 1 000 AAH53559.1 1.0e 1 000

AAC50363.1 1.0e 1 000

NP_110435 T51168

Accession number b)

b)

2.43 1.43 1.19

2.43

2.43 2.43

22 43 18

27

14 21

Z-score b) Seq. cover.,% b)

5.8 5.5 6.5

5.7

5.6 6.7

pI

6.0 6.3 9.0

5.6

26d) 75 21 46d)

5.2 6.6

pI

74 23 61d)

42d)

38 69

Mr

Theoretical values

42 63

Mr

Experim ental values

[17] [18] [19]

[16]

[14] [15]

Reference c)

a) Annotation of protein spots is as indicated in Fig. 2. b) Name of proteins, accession number, probability, Z-score, sequence coverage, and theoretical pI and M r values were obtained searching NCBI database w ith ProFound search engine. Experimental values of pI and M r were calculated from m igration positi on of proteins in 2-D gels. c) References to papers describing identified proteins as phosphoproteins. d) Lower molecular masses of identified GTP-binding protein alpha q subunit and Grb14 may be due to the limited proteolysis of proteins.

4 5 6

3

1 2

Spot a) Name of protein

Table 1

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precipitates which were not compatible with 2-DE. Dialysis of elution fraction did not eliminate completely the compounds interfering with 2-DE, or caused significant losses of phosphoproteins. 7. Be sure that no solution left in the tube prior to dissolve the pellet with rehydration solution. 8. Any bubbles trapped between the gel and the cellophane must be removed. 9. We monitored the phosphorylation of Smad2, which is a component of transforming growth factor-b (TGFb) signaling pathway, and which is phosphorylated at two C-terminal serine residues upon its activation (12). You can use anti-phospho- antibodies to any other proteins, which you can induce to be phosphorylated instead. It will provide you a good marker to check the phosphoprotein distribution between the chromatography fractions. Acknowledgments The author would like to thank Serhiy Souchelnytskyi for his advices and encouragement and J. Ericsson, V. Lukiyanchuk, C.-H. Heldin, N. Bhaskaran for comments, U. Hellman for discussions and access to a mass spectrometer. This work was supported in part by grants from the Swedish Cancer Society, the Swedish Research Council, the Hiroshima University, and Merck KGaA to Serhiy Souchelnytskyi.

References 1. Figeys, D. (2002) Proteomics approaches in drug discovery. Anal. Chem. 74, 412–419. 2. Stancato, L. F. and Petricoin, E. F. III (2001) Fingerprinting of signal transduction pathways using a combination of anti-phosphotyrosine immunoprecipitations and two-dimensional polyacrylamide gel electrophoresis. Electrophoresis 22, 2120–2124. 3. Imam-Sghiouar, N., Laude-Lemaire, I., Labas, V., and Pflieger, D. et al. (2002) Subproteomics analysis of phosphorylated proteins: application to the study of B-lymphoblasts from a patient with Scott syndrome. Proteomics 2, 828–838. 4. Stasyk, T., Dubrovska, A., Lomnytska, M., Yakymovych, I. et al. (2005) Phosphoproteome profiling of transforming growth factor (TGF)-beta signaling: abrogation of TGFbeta1-dependent phosphorylation of transcription factor-II-I (TFII-I) enhances cooperation of TFII-I and Smad3 in transcription. Mol. Biol. Cell. 16, 4765–4780. 5. Adam, G. C., Sorensen, E. J., and Cravatt, B. F. (2002) Mapping enzyme active sites in complex proteomes. Mol. Cell. Proteomics, 1, 781–790. 6. Zhou, H., Watts, J. D., and Aebersold, R. (2001) Quantitative proteome analysis by solid-phase isotope tagging and mass spectrometry. Nature Biotech. 19, 375–378. 7. Oda, Y., Nagasu, T., and Chait, B. T. (2001) Enrichment analysis of phosphorylated proteins as a tool for probing the phosphoproteome. Nature Biotech. 19, 379–382. 8. Knight, Z. A., Schilling, B., Row, R. H., and Kenski, D. M. et al. (2003) Phosphospecific proteolysis for mapping sites of protein phosphorylation. Nature Biotech. 21, 1047–1054.

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9. Ficarro, S. B., McCleland, M. L., Stukenberg, P. T., and Burke, D. J. et al. (2002) Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nature Biotech. 20, 301–305. 10. Phosphoprotein Purification Handbook, Vol. 12, Qiagen, Crawley 2002, pp. 7–19. 11. Dubrovska, A. and Souchelnytskyi, S. (2005) Efficient enrichment of intact phosphorylated proteins by modified immobilized metal-affinity chromatography. Proteomics 5, 4678–4683. 12. Souchelnytskyi, S., Moustakas, A., and Heldin, C.-H. (2002) TGF-beta signaling from a three-dimensional perspective: insight into selection of partners. Trends Cell Biol. 12, 304–307. 13. Kanamoto, T., Hellman, U., Heldin, C.-H., Souchelnytskyi, S. (2002) Functional proteomics of transforming growth factor-beta1-stimulated Mv1Lu epithelial cells: Rad51 as a target of TGFbeta1-dependent regulation of DNA repair. EMBO J. 21, 1219–1230. 14. Yan, X., Li, F., Liang, Y., and Shen, Y. et al. (2003) Human Nudel and NudE as regulators of cytoplasmic dynein in poleward protein transport along the mitotic spindle. Mol. Cell. Biol. 23, 1239–1250. 15. Vodermaier, H. C. (2004) APC/C and SCF: controlling each other and the cell cycle. Curr. Biol. 14, R787–R796. 16. Matousek, P., Durchankova, D., Svandova, I., Novotny, J., and Svoboda, P. (2005) Agonist-induced tyrosine phosphorylation of Gq/G11 alpha requires the intact structure of membrane domains. Biochem. Biophys. Res. Comm. 328, 526–532. 17. Mizukoshi, E., Suzuki, M., Misono, T., and Loupatov, A. et al. (2001) Fibroblast growth factor-1 interacts with the glucose-regulated protein GRP75/mortalin. Biochem. Biophys. Res. Commun. 280, 1203–1209. 18. Dierks, H., Kolanus, J., and Kolanus, W. (2001) Actin cytoskeletal association of cytohesin-1 is regulated by specific phosphorylation of its carboxyl-terminal polybasic domain. J. Biol. Chem. 276, 37472–37481. 19. Shen, T. L. and Guan, J. L. (2004) Grb7 in intracellular signaling and its role in cell regulation. Front. Biosci. 9, 192–200.

160 Analyzing Protein Phosphorylation John Colyer 1. Introduction Protein phosphorylation is a ubiquitous modification used by eukaryotic cells to alter the function of enzymes, ion channels, and other proteins in response to extracellular stimuli, or mechanical or metabolic change within the cell. In many instances, phosphorylation results in a change in the catalytic activity of the phosphoprotein, which influences one particular aspect of cellular physiology, thereby allowing the cell to respond to the initiating stimulus. A number of different residues within a protein can be modified by phosphorylation. Serine, threonine, and tyrosine residues can be phosphorylated on the side chain hydroxyl group (o-phosphoamino acids), whereas others become phosphorylated on nitrogen atoms (N-phosphoamino acids, lysine, histidine, and arginine). The former group are involved in dynamic “regulatory” functions and have been studied extensively (1), whereas the latter group may perform both structural/catalytic roles and signaling functions, the study of which has occurred more recently (2). The disparity in our understanding of the role of o- and N-phosphoamino acids is in part a consequence of the acid lability of N-phosphoamino acids, which leads to their destruction during the analysis of many phosphorylation experiments. In terms of the process of studying an individual phosphoprotein, a number of key issues can be identified. First, one must demonstrate that phosphorylation of the protein takes place; then define the number of sites within the primary sequence that can be phosphorylated and by which protein kinase; identify the individual residue(s) phosphorylated; the functional implication of phosphorylation of each site; and describe the use of each site of phosphorylation in vivo. This chapter aims to describe the conduct of an experiment performed to identify a protein as a phosphoprotein. In the case of oligomeric enzymes, it will identify the subunit(s) phosphorylated by a particular kinase. The determination of From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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the stoichiometry of phosphorylation is also described, which provides the first information concerning the number of phosphorylation sites within a polypeptide. These procedures are most straightforward if one has access to the purified protein kinase of interest and the protein substrate. In the case of the kinase, this can be served in many instances by a number of commercial sources, but the approach may be limited by the availability of sufficient pure protein substrate. If this is the case, phosphorylation of a particular target as part of a complex mixture of proteins (e.g., whole-cell extract) can be performed. Under these conditions, identification of the protein of interest will require exploitation of a unique electrophoretic property of the protein (3) or require purification of the protein by immunoprecipitation or other comparable affinity-interaction means prior to electrophoresis. The identification of the protein as a phosphoprotein can thereby be achieved, although analysis of the stoichiometry of phosphorylation in this way is inadvisable. In each case, the experimental procedure has a common design: an in vitro phosphorylation reaction is followed by separation of the phosphoproteins by SDS-PAGE and subsequent identification by autoradiography. The incorporation of labeled phosphate can be determined by excising the phosphoprotein band from the dried gel, scintillation counting this gel piece, and converting 32P cpm into molar terms from the knowledge of the specific activity of the initial ATP stock, and the amount of protein substrate analyzed. 2. Materials 1. Purified and partially purified multifunctional protein kinases can be obtained from several commercial sources. The availability of a number of enzymes is illustrated in Table 1. The list is not exhaustive, and inclusion in the table does not constitute endorsement of the product: Table 1 Commercial Source of Multifunctional Protein Kinases Kinase

Source

Protein kinase A Protein kinase C Calmodulin kinase II Casein kinase II CDC2 kinase src kinase

a,c,d,e a,b,c,d c a,b,d d d

a

Boehringer Mannheim. Calbiochem-Novabiochem. c Sigma Chemical Co. d TCS Biologicals Ltd. b

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2. Phosphorylation buffer for the catalytic subunit of protein kinase A (c-PKA): 50 mM Histidine-KOH, pH 7.0, 5 mM MgSO4, 5 mM NaF, 100 nM c-pKA, 100 µM ATP. a. Histidine-KOH, pH 7.0 is prepared as a concentrated stock, 200 mM stored at 4°C for 1 mo, or −20°C for >12 mo. (Warm to 30°C for 30 min to dissolve histidine following storage at −20°C.) b. MgSO4, EGTA, NaF: all prepared as 100-mM stock, stable at 4°C >12 mo. c. ATP, nonradioactive: 20- or 100-mM stock (pH corrected to 7.0 with KOH) stable at −20°C for >12 mo. Aliquot to avoid repeat freeze-thaw cycles. d. c-pKA, Mr 39,000 (4), sources Table 1: stable at −70°C for ~12 mo. Avoid dilute solutions—enzyme tends to aggregate and inactivate. 3. γ-32P-ATP, ICN Pharmaceuticals: dispense into small aliquots and store −20°C. Avoid freeze to thaw cycles. T1/2 ~14 d; discard 1 mo after reference date. 4. SDS-PAGE sample buffer, (double-strength): 125 mM Tris-HCl, pH 6.8, 20% glycerol, 2% SDS, 10% 2-mercaptoethanol, 0.01% bromophenol blue, stable at −20°C >12 mo. Aliquot and avoid freeze-thaw cycles. 5. Filter paper: Whatman 3MM. 6. X-ray film, X-ray cassettes, intensifying screens, developer and fixative: X-Ograph Ltd.; developer and fixative are stored at room temperature, are reusable, and are stable for approx 2 wk. 7. Scintillation fluid: Emulsifier-safe, Packard.

3. Method 3.1. Phosphorylation Reaction Using Purified Protein Substrates 1. In a designated radioactive area with appropriate acrylic screening prepare a stock of radioactive ATP. The addition of 50 µCi [γ-32P]-ATP to a 0.5-mL solution of 1 mM ATP will produce a suitable experimental ATP stock of 220 cpm/pmol (see Note 1). Warm to 37°C. 2. Incubate the purified protein of interest (0.1–1.0 mg/mL) in the phosphorylation assay medium lacking ATP. Allow the sample to warm to 37°C for 2 min (see Note 2). A number of control samples should be set up in parallel. One control should contain target protein, but no exogenous kinase, and another, kinase but no target protein. 3. Start the phosphorylation reaction by the addition of ATP, containing [γ-32P]-ATP (as defined in step 1, Subheading 3.). Cap the tube and vortex briefly. Follow the phosphorylation as a function of time by removing aliquots of the reaction at specific points in time, every 20 s for the first minute, and then at 60-s intervals for the next 4 min. 4. Terminate the reaction by mixing the sample with an equal volume of doublestrength SDS-PAGE sample buffer at room temperature. Dispense the phosphorylation sample into an Eppendorf tube containing an equal volume of double-strength sample buffer, cap the tube, and vortex briefly. Store these samples at room temperature, behind appropriate screens, until all samples have been collected.

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5. Incubate all samples for 30 min at 37°C. Perform SDS-PAGE in a gel of suitable acrylamide composition, loading a minimum of 5 µg pure target protein/lane (details of electrophoresis in Chapter 21). Allow the dye front to migrate off the bottom of the gel (depositing most of the radioactivity into the electrode buffer), stain with Coomassie brilliant blue, and destain the gel (see Chapter 21). 6. Mount the gel on filter paper and cover with clingfilm. The filter paper should be wet with unused destain solution prior to contact with the SDS-PAGE gel. Lower the gel onto the wet filter paper slowly, and flatten to remove air bubbles. Cover with clingfilm, and dry using a vacuum-assisted gel drier for 2 h at 90°C. 7. Once dry, an X-ray film should be placed in contact with the gel for a protracted period to image the location of phosphoproteins. This procedure must be performed in the dark, although light emitted by dark room safety lamps is permitted. An X-ray film is first exposed to a conditioning flash of light from a flash gun. Hold a single piece of X-ray film and the flash gun 75 cm apart. Set the flash gun to the minimum power output, and discharge a single flash directly onto the film (see Note 3). Place the film on top of a clean intensifying screen within an X-ray cassette. Take the dried SDS-PAGE gel, still sandwiched between filter paper and clingfilm, and place it gel side down onto the X-ray film. Do not allow the gel to move once in contact with the film, and use adhesive tape to secure the contact. With a permanent marker pen, draw distinctive markings from the filter paper backing of the gel onto the X-ray film to facilitate orientation of film and gel once autoradiography is complete (see Note 4). Close the X-ray cassette, label the cassette with experimental details, including the current date and time, and store at −70°C. 8. After 16 h of exposure, the autoradiograph can be developed. Remove the cassette from the −70°C freezer, and allow at least 30 min for it to thaw. Once in the dark room, with safety light illumination only, the cassette should be opened, and SDSPAGE gel removed from the X-ray film. The film should be placed in 2 L developer and agitated for 4 min at room temperature, in the dark. Using plastic forceps, the film is removed from developer, rinsed in water, and then agitated for a further 90 s in 2 L of fixative (room temperature, dark). At the end of 90 s, the autoradiograph is no longer light-sensitive, and normal lighting can be resumed. Wash the autoradiograph extensively, for 10 min in a constant flow of water, and allow to air-dry. 9. Identification of phosphorylated polypeptides can be performed by superimposition of autoradiograph on the SDS-PAGE gel (see Note 5). A uniform, almost transparent background should be achieved, with phosphorylated proteins identified as black bands on the autoradiograph of variable intensities (depending on the level of phosphorylation), but regular width and shape (see Note 6). Exposure times can be altered in the light of results obtained, and repeated autoradiographs of various durations performed on a single gel (see Note 7).

3.2. Phosphorylation of Components in Complex Protein Mixtures 1. The phosphorylation experiment is performed largely as detailed in Subheading 3.1. with the following modifications. A protein concentration of 1–5 mg/mL is recommended supplemented with 100 nM purified c-pKA and 10 µM adenosine

Analyzing Protein Phosphorylation

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cyclic 3,5-monophosphate, and with 1 µM microcystin-LR for additional Ser/Thr phosphatase control. 2. Perform a phosphorylation time-course experiment and process as described in Subheading 3.1. If the identity of a particular protein cannot be gauged from a peculiar electrophoretic feature (e.g., dissociation of oligomer to monomer on boiling; 3), then an affinity purification step must be introduced prior to electrophoresis. 3. In this case, phosphorylation will be terminated by placing the sample on ice. Immunoprecipitation of the protein of interest should be performed, taking care to solubilize membrane proteins effectively if they are of interest. Immunoprecipitates should be processed as described in Subheading 3.1., step 5 onward (see Note 8).

3.3. Determination of Phosphorylation Stoichiometry 1. The specific activity of the ATP (cpm/pmol) needs to be determined empirically. At the time of the phosphorylation experiment, dilute a sample of the experimental ATP (1 mM containing 100 µCi/mL [γ-32P]-ATP, as described in Subheading 3.1., step 1) to 1 µM by serial dilution in water. Dispense triplicate 10-µL aliquots of the 1-µM ATP (10 pmol) into separate scintillation vials, and add 4.6 mL scintillation fluid to each. Cap and label 10 pmol ATP (see Note 9). 2. Perform steps 1–9 of Subheading 3.1. To excise the phosphorylated protein bands from the dry SDS-PAGE gel, identify the location by superimposition of gel and autoradiograph. With a marker pen, outline the autoradiographic limits of the phosphoprotein on the gel. Overlay again to confirm the accuracy of demarkation. Mark similar-sized areas of gel that do not contain phosphoproteins to determine background [32P]. Excise these gel pieces with scissors, remove the clingfilm, and place the acrylamide piece and filter paper support in a scintillation vial. Add 4.6 mL scintillation fluid, and cap the vial. 3. Scintillation count each vial for 5 min or longer, using a program defined for 32P radionucleotides. Minimal quenching of 32P occurs under these conditions. 4. To calculate the pmol phosphate incorporated/µg protein, subtract the background radioactivity (cpm) from experimental data (cpm) to obtain phosphate incorporation into the protein sample (in units of cpm). Convert this to pmol incorporation/ µg using the formula: Phosphorylation (pmol/µg protein) = [protein phosphorylation (cpm)/SA of ATP (cpm/pmol) × µg protein]

(1)

With a knowledge of the molecular weight of the polypeptide, a molar phosphorylation stoichiometry can be calculated from these data using the formula: Phosphorylation (mol/mol protein) = [phosphorylation (pmol/µg protein) × molecular weight/106]

(2)

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5. Experimental conditions that result in maximal protein phosphorylation will have to be optimized. Parameters worth considering include alteration of the pH of the reaction, extension of the time-course of phosphorylation (up to several hours), addition of extra protein kinase during the phosphorylation process, and addition of extra ATP throughout the time-course.

4. Notes 1. The specific activity of ATP must be tailored to the experiment intended. Phosphorylation and autoradiography of proteins require ≥200 cpm/pmol, studies that require analysis beyond this point (e.g., phosphoamino acid identification) require ~2000 cpm/pmol, while phosphorylation of peptides requires ~ 20–50 cpm/pmol. 2. The stability of proteins at low concentration is sometimes an issue; inclusion of an irrelevant protein, but not a phosphoprotein (e.g., bovine serum albumin) at 1 mg/mL (final) is recommended. The phosphorylation example used to illustrate this method (c-pKA) displays catalytic activity in the absence of signaling molecules. In other instances this is not so. Therefore, relevant activators should be included as dictated by the kinase (e.g., Ca2+, calmodulin, acidic phospholipids, and so forth). 3. Film developed at this stage will exhibit very slight discoloration compared to unexposed film. 4. Phosphorescent labels (Sigma-Techware) can be used to label an SDSPAGE gel prior to autoradiography. It will also facilitate superimposition of gel and autoradiograph. These can also highlight the position of mol-wt markers, an image of which will be captured on the X-ray film. 5. Protein kinases invariably autophosphorylate. This can be identified clearly in control samples lacking phosphorylation target (Subheading 3.1., step 2). 6. Autoradiographs sometimes have a high background signal. Uniform black coloration over the whole film, extending beyond the area exposed to the gel is indicative of illumination of the X-ray film. Discoloration of part of the film is indicative of light entering the X-ray cassette. Examine the cassette carefully, particularly the corners that are prone to damage by rough handling. 7. Phosphoimage technology represents an alternative to autoradiography, it has the advantage of collecting the image quickly. Exposure times are reduced by an order of magnitude. However, in my experience, this benefit is at the expense of the quality of the image, which is granular. 8. The time required for immunoprecipitation or similar procedure should be kept to a minimum to limit the dephosphorylation of proteins by endogenous phosphatase enzymes. A cocktail of phosphatase inhibitors should be included for the same reason.

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9. The hydrolysis of ATP to ADP and Pi occurs at a low rate in the absence of any enzyme. The extent of hydrolysis of [γ-32P]-ATP can affect the determination of phosphorylation stoichiometry, since it will result in the overestimation of the specific activity of ATP if a correction is not made. Quantification of the purity of ATP is quoted in the product specification from suppliers. Only fresh [γ-32P]-ATP should be used in these procedures or the degree of hydrolysis confirmed by thin-layer chromatography (6). References 1. Krebs, E. G. (1994) The growth of research on protein phosphorylation. Trends Biochem. Sci. 19, 439. 2. Swanson, R. V., Alex, L. A., and Simon, M. I. (1994) Histidine and aspartate phosphorylation: two component systems and the limits of homology. Trends Biochem. Sci. 19, 485–490. 3. Drago, G. A. and Colyer, J. (1994) Discrimination between two sites of phosphorylation on adjacent amino acids by phosphorylation site-specific antibodies to phospholamban. J. Biol. Chem. 269, 25,073–25, 077. 4. Peters, K. A., Demaille, J. G., and Fischer, E. H. (1977) Adenosine 3':5'-monophosphate dependent protein kinase from bovine heart. Characterisation of the catalytic subunit. Biochemistry 16, 5691–5697. 5. Otto, J. J. and Lee, S. W. (1993) Immunoprecipitation methods. Meth. Cell Biol. 37, 119–127. 6. Bochner, B. R. and Ames, B. N. (1982) Complete analysis of cellular nucleotides by two dimensional thin layer chromatography. J. Biol. Chem. 257, 9759–9769.

161 Mass Spectrometric Analysis of Protein Phosphorylation Stefan Gander, Alessio Cremonesi, Johana Chicher, Suzette Moes, and Paul Jenö 1. Introduction Phosphorylation is one the most frequently occurring posttranslational modifications in proteins, playing an essential role in transferring signals from the outside to the inside of a cell and in regulating many diverse cellular processes such as growth, metabolism, proliferation, motility and differentiation. It is estimated that up to one-third of all proteins in a typical mammalian cell are phosphorylated (1). Phosphorylation is carried out by a vast group of protein kinases which are thought to constitute 3% of the entire eukaryotic genome (1–3). To decipher the recognition signal of protein kinases and protein phosphatases acting on a given molecular target, and to understand how the activity of the target protein is regulated by phosphorylation, it is important to define the sites and the extent of phosphorylation at each specific site. Earlier techniques of phosphoprotein analysis involves in vivo or in vitro labeling with [32P]-phosphate (4–8). The radiolabelled protein is subsequently digested with a suitable protease and radiolabelled peptides are separated either by high-performance liquid chromatography (HPLC) (7,8) or by two-dimensional phosphopeptide mapping (4–8). The site of phosphorylation is then determined by solid-phase Edman sequencing (6). Identification of phosphorylation sites by Edman degradation of [32P]-labelled proteins has been almost completely replaced by mass spectrometric techniques mainly because of speed, versatility and the availability of powerful computational tools (9–12). MS-based techniques have been steadily improved in the past so that the global analysis of protein phosphorylation, its dynamics and the elucidation of substrates phosphorylated

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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by protein kinases have become possible (13,14). These studies have provided remarkable insights into signaling cascades and their organization. The basic elements of mass spectrometric phosphoprotein analysis consist of proteolytic digestion of the protein(s) of interest followed by measuring the masses of the resulting peptides. Individual phosphopeptides can be detected by a variety of techniques such as MALDI-mass mapping and phosphatase treatment (15), neutral loss scanning (16), and precursor ion scanning (17). The phosphorylation site in a given phosphopeptide can be identified by subjecting the peptide to fragmentation in the collision cell of the mass spectrometer. From the mass difference between the fragment ions, the sequence can be “read” and the site of phosphorylation can be identified when the mass difference is either 167, 181, or 243 Da, corresponding to the residue masses of P-Ser, P-Thr, or P-Tyr. Powerful search programs such as SEQUEST and MASCOT (18,19) exist to identify both, the phosphoprotein and the site at which the phosphate resides. Phosphorylation at Ser, Thr, and Tyr residues may modulate the activity of a given protein. Thus, the determination of the extent to which a protein becomes phosphorylated is important for the understanding of the dynamics of how a protein responds to a particular stimulus. Recently, mass-spectrometry based techniques for relative quantification of protein phosphorylation have been developed, using either stable-isotope tagged amine-reactive labels (iTRAQ) (20) or stable isotope labeling by amino acids in cell culture (SILAC) (21). Alternatively, chemical (22) or enzymatic approaches (23) can be used to quantitate relative changes in phosphorylation. One major obstacle in phosphoprotein analysis is the low relative level of phosphorylation, in particular tyrosine phosphorylation. To access phosphorylation events, enrichment strategies have been developed that allow selective isolation of serine/threonine phosphorylation by immobilized metal affinity chromatography (IMAC) (24) or tyrosine phosphorylation by immunoprecipitation with antiphosphotyrosine antibodies (25). In this chapter, we will describe the basic steps of mass spectrometric phosphopeptide mapping, namely enzymatic digestion of phosphoproteins, reversephase high-performance liquid chromatography of peptides with mass spectral analysis (LC/MS), and determination of the site of phosphorylation by tandem mass spectrometry (MS/MS). 2. Materials 2.1. Equipment 1. HPLC system: For LC/MS coupling, a Rheos 2200 pump (Flux Instruments, Basel, Switzerland) was operated at a flow of 150 ml/min. A precolumn split was used to reduce the flow to approximately 500 nl/min.

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2. Mass Spectrometers: LTQ Orbitrap mass spectrometer (Thermo Electron, San Jose, CA) equipped with a Finnigan Nanospray Probe (Thermo Electron, San Jose, CA) and Bruker Reflex III MALDI-TOF instrument (Bruker Daltonics GmbH, Switzerland).

2.2. Materials and Reagents 1. Fused silica capillaries (100 µm i.d., 280 µm o.d.) and PEEK sleeves (300 µm i.d.) (LC Packings, Dionex Switzerland AG, Olten, Switzerland). F120 PEEK Fingertight fittings (Upchurch Scientific, ERCATECH AG, Berne, Switzerland). 2. C18 capillary columns for LC/MS: 300SB C-18 columns, 0.3 × 50 mm (Agilent Technologies, Basel, Switzerland). C18 ZipTips, P10 size, were from Millipore (Bedford, MA). PHOS-Select Iron affinity Gel was purchased from SIGMAALDRICH (Buchs, Switzerland). 3. Chemicals: TFA (Applied Biosystems, Rotkreuz, Switzerland) and acetonitrile (Lab-Scan, MACHEREY-NAGEL AG, Oensingen, Switzerland). DTT and ten times concentrated dephosphorylation buffer (0.5 M Tris-HCl, pH 8.5, 1 mM EDTA) (Roche Diagnostics, Mannheim, Germany). Iodoacetamide (Fluka Chemie AG, Buchs, Switzerland). 4. Enzymes: Proteases were obtained from the following suppliers: modified trypsin: (Promega, Madison, WI), endoproteinase LysC (Achromobacter protease) (Wako Pure Chemical Industries, Osaka, Japan), endoproteinase GluC, sequencing grade, and calf intestinal alkaline phosphatase (1000 U/ml) (Roche Diagnostics, Mannheim, Germany). 5. Isotopically labeled H218O, >95% pure (CAMPRO Scientific GmbH, Berlin, Germany). α-cyano-4-hydroxycinnamic acid for MALDI-TOF (Sigma-Aldrich, Buchs, Switzerland).

3. Methods 3.1. Enzymatic Digestions of Phosphoproteins To efficiently trace sites of phosphorylation in proteins, it is important to obtain an essentially complete proteolysis of the protein of interest (see Note 1). This is best achieved if the protein is fully denatured by reduction and alkylation with DTT and iodoacetamide. For reduction, the protein is dissolved in 10 µl 100 mM Tris-HCl, pH 8.0, 8 M urea (freshly prepared), 1.5 µl 75 mM DTT dissolved in water) is added and the protein is reduced for 1–2 h at 37°C. Alkylation is done by adding 1 µl of 625 mM iodoacetamide and incubating the protein for 15 min at room temperature. To obtain an as complete a digest as possible, digestion with a combination of proteases is preferable. After reduction and alkylation the protein is first digested with the endoproteinase LysC. The residual iodoacetamide and urea (approximately 6 M) do not appreciably inhibit the enzymatic activity. The enzyme to substrate ratio is kept between 1:50 and 1:20 (w/w). Incubation with endoproteinase LysC is carried out at 37°C for 1 h. The reaction is stopped by the addition of 10% TFA

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to a final concentration of 0.5% TFA. For subsequent trypsin digestion, the urea concentration has to be lowered to 2 M by dilution with 100 mM Tris-HCl, pH 8.0, whereas endoproteinase GluC is severely inhibited even by 2 M urea. When using this enzyme, it is best to remove urea completely. This can be achieved by reversephase chromatography. Digestion with trypsin or endoproteinase GluC is carried out at an enzyme to substrate ratio of 1:50 to 1:20 (w/w) for 18 h at 37°C. 3.2. Phosphorylation Analyzed by Liquid ChromatographyMass Spectrometry To find phosphopeptides in protein digests, reverse-phase chromatography in MS-compatible solvents such as formic acid or acetic acid is directly coupled to the mass spectrometer. The eluting peptides are measured in a data-dependent mode. First, full-range mass spectra are analyzed in real time, and then ions are collected for fragmentation to generate sequence-specific ions for data bank searching. For phosphopeptide detection we rely in most cases on neutral loss scanning. The neutral loss is the result of a gas-phase b-elimination reaction of phosphoric acid (−98 Da) from phosphoserine and phosphothreonine. Peptides phosphorylated on serines and threonines can be found by searching the LC/MS data for those peptides that had lost either 49, 32.6, or 24.5 Da for doubly, triply, or quadruply charged peptides. P-Tyr containing peptides do not tend to lose phosphoric acid and escape detection by neutral loss scanning. Individual MS/ MS spectra are then compared against the known protein sequence using TurboSequest software (26). Fig. 1A shows the LC/MS analysis of a tryptic digest of the yeast ribosomal protein RPL24A (systematic yeast gene annotation is YGL031C). In particular, we were interested in the peptide GDSKIFR surrounding Ser21 that had been reported to be phosphorylated in vivo (21). The phosphorylated doubly charged peptide eluted at 30.49 min from the reverse phase column (Fig. 1B). Searching the LC/MS data for the neutral loss of phosphoric acid for doubly charged peptides (49 Da) revealed a strong candidate peptide whose elution coincides with the predicted phospho-GDS*KIFR peptide (Fig. 1C). Fragmentation of the phosphopeptide was triggered at scan 5208 (30.43 min). The primary fragmentation product is an intense ion (402.67 Da) 49 Da less than the precursor ion (Fig. 2A). While the doubly charged ion of 451.7 Da matches exactly the singly phosphorylated peptide GDSKIFR, positive identification by data bank searching with the TurboSEQUEST software was not possible due to the few fragment ions present in the spectrum. This is typical for phosphopeptides that tend to predominantly lose phosphoric acid via b-elimination and therefore suppress the formation of informative fragment ions. Therefore, the 402.67 ion that underwent neutral loss of phosphoric acid was brought to fragmentation in the subsequent scan. This resulted in a richer fragmentation spectrum from which the phosphopeptide and

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Fig. 1. (A) Ion chromatogram of an endoproteinase LysC/tryptic digest of yeast ribosomal proteins eluting from a capillary column into the mass spectrometer. (B) Selected ion tracing for the 451.71 Da ion that corresponds to the singly phosphorylated GDS*KIFR from ribosomal protein RPL24A (YGL031C). (C) Neutral loss scanning experiment showing the ion tracing for doubly charged peptides that underwent loss of phosphoric acid (49 Da). The asterisk indicates phosphorylation, and D marks the peptide that had lost phosphoric acid during collisional activation.

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Fig. 2. Fragmentation spectrum of the doubly charged GDS*KIFR phosphopeptide. (A) The 451.71 Da precursor ion caused a prominent loss of phosphoric acid (402.67 Da) with little fragmentation of the peptide backbone. (B) Abundant peptide fragmentation occurs upon further fragmentation of the 402.67 Da neutral loss fragment ion (MS/MS/MS). The asterisk indicates phosphorylation, while D marks the serine that had lost phosphoric acid during collisional activation.

the site of phosphorylation could be unambiguously identified (Fig. 2B). The “phosphate-specific” MS scan takes advantage of the neutral loss of phosphoric acid from phosphopeptides in a fully automated fashion (see notes). Once all the data have been collected, data bank search is initiated with the following settings: variable modification of Ser, Thr, and Tyr set to 80 Da (the mass of a phosphate) and 57 Da static modification for Cys (alkylation by iodoacetamide). 3.3. Stoichiometry of Phosphorylation Stoichiometries of phosphorylation by mass spectrometry can be quantified by dividing the sum of the peak areas of all intensities of a given phosphopeptide (all charge states must be included) by the sum of all intensities of the phosphorylated and its nonphosphorylated forms. However, this often results in underestimating the phosphorylation stoichiometriy, as phosphopeptides tend to ionize with lower efficiency than their corresponding unphosphorylated peptides (27). Quantitation in site-specific phosphorylation has been achieved by the use of stable isotope labeling (28) and most recently, by iTRAQ (29). For the quantification of relative changes in the extent of phosphorylation that occur upon drug treatment of yeast cells we rely on a simple method that involves the labeling of individual pools of differentially phosphorylated proteins by enzymatic digestion in isotopically labeled water, affinity selection of phosphopeptides followed by MALDI time-of-flight mass spectrometry (23). The method consists of three steps: (i) enzymatic digestion of a protein in H216O or isotopically enriched H218O to label individual pools of differentially phosphorylated proteins; (ii) affinity selection of phosphopeptides from combined digests by immobilized metal-affinity

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chromatography; and (iii) dephosphorylation with alkaline phosphatase to allow for quantitation of changes of phosphorylation by MALDI-TOF. We have successfully used 18O labeling to trace relative changes in phosphorylation of the yeast protein kinase Npr1 (systematic yeast gene name YNL183C). Upon treatment of yeast cells with rapamycin or following nitrogen starvation, Npr1 becomes activated by dephosphorylation to support sorting of the general amino acid permease Gap1 to the plasma membrane (30). We mapped the global changes of phosphorylation in Npr1 following rapamycin treatment. Particularly sensitive phosphorylation sites to rapamycin treatment are found on Ser353 and Ser356. The 1227.4 Da MALDI-TOF signal corresponds to the doubly phosphorylated peptide SQHSSIGDLKR (comprising residues 353–363) from untreated cells while the 1231.4 Da signal derives from the same phosphopeptide of Npr1 that had been obtained from rapamycin-treated yeast cells (Fig. 3A). The same Npr1 phosphopeptide was isolated from a genomically deleted Sit4 mutant. In this case, no decrease in the extent of phosphorylation of

Fig. 3. (A) MALDI-TOF spectra of the doubly phosphorylated peptide (S*QHS*SIGDLKR) encompassing residues 353–363 of the yeast protein kinase Npr1 (YNL183C) that had been isolated from untreated (1227.4 Da) or from rapamycin-treated (1231.4 Da) cells. The 1231.4 Da signal indicates an approximately four-fold dephosphorylation. (B) The same phosphopeptide was obtained from a mutant in which the sit4 phosphatase responsible for Npr1 dephosphorylation was deleted. The 1231.5 Da signal indicates that no dephosphorylation occurred at these sites. (C) Coomassie-blue stained SDS gel of GST-Npr1 from wildtype cells (lane 1 and 2) or sit4 mutant cells, either untreated (−) or treated (+) with 200 nM rapamycin for 15 min. The arrows indicate phosphorylated (upper) and dephosphorylated (lower) GST-Npr1. (Modified from ref. 23.)

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the SQHSSIGDLKR phosphopeptide was detectable due to the lack of the Sit4 phosphatase responsible for dephosphorylation of Npr1 (Fig. 3B). 3.4. Procedure for 18O Labeling of Peptides 1. Dry 20 µl of reduced and carboxamidomethylated phosphoprotein (in 100 mM Tris-HCl, pH 8.0, 150 mM NaCl, 0.2 µg/µl protein concentration) in a SpeedVac. 2. Dissolve one pool of phosphoprotein in 20 µl H216O (for example, phosphoprotein derived from non-treated or wild type cells) and the other pool (for example, phosphoprotein derived from treated or mutant cells) in 20 µl isotopically enriched H218O. 3. Digest the two protein pools separately with trypsin at an enzyme/substrate ratio of 1:25 at 37°C for 2 h. During digestion in H218O, two atoms of 18O will be incorporated into every newly generated peptide C-terminus. 4. Stop the digestion by adding TFA to a final concentration of 0.1%. Mix the digest from normal and from H218O water and desalt them by adsorbing the peptides onto a C18 ZipTip. The peptides are washed with 0.1% TFA and desorbed with 5 µl 80% acetonitrile containing 0.1%TFA. 5. Select phosphopeptides by adding approximately 2 ml of a 50% suspension of PHOS-Select IMAC beads (in 0.1% acetic acid/30% acetonitrile) to the desalted peptide pool. Incubate the peptide-IMAC suspension in an Eppendorf Thermoshaker (set to 1,200 rpm) at room temperature for 1 h. 6. Wash the beads three times with 25 ml 0.1% acetic acid/30% acetonitrile. 7. Release the bound phosphopeptides with 10 ml 100 mM Tris-HCl, pH 8.0 containing 0.5 units of alkaline phosphatase by shaking the suspension at 37°C for 15 min. 8. For MALDI-TOF analysis desalt the peptides with ZipTips as above, except that peptide elution is done with 80% acetonitrile, 0.1% TFA, containing 1 mg/ml a-cyano-4-hydroxycinnamic acid.

4. Notes 1. For routine phosphopeptide mapping, we usually digest the proteins first with the endoproteinase LysC followed by a second digestion with trypsin. This digestion regime generates peptides with at least two positive charges on each peptide for efficient ionization. The C-terminal lysine or arginine favours fragmentation in the collision cell, which leads to a high sequence coverage, a prerequisite for unambiguous localization of the site of phosphorylation. Also, since the endoproteinase LysC is active in 6 M urea, highly insoluble phosphoproteins can be efficiently digested. Subsequent trypsin digestion also is possible after lowering the urea concentration to 2 M. 2. One major obstacle in phosphopeptide analysis is that they are often too hydrophilic to bind to reverse-phase supports used for LC/MS separation.

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4.

5.

6.

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A simple solution to this problem has been suggested by combining graphite powder for retaining phosphopeptides that are otherwise lost in the flow through (31). In cases where phosphorylation is suspected but could not be proven due to poor fragmentation, we dephosphorylate the digest with alkaline phosphatase and repeat the analysis. If the candidate peptide is phosphorylated, dephosphorylation leads to a reduction of its mass by 80 Da (or multiples thereof in the case of multiple phosphorylation). Enzymatic dephosphorylation with calf-intestinal alkaline phosphatase (CIP) can be directly carried out in the digestion mix, since CIP is completely resistant to proteolysis by enzymes such as trypsin, endoproteinase LysC or -GluC, although urea tends to inactivate the enzyme. For protein digests in the 1–500 picomole range, 1 U of CIP was found to be sufficient to lead to complete dephosphorylation of phosphopeptides. For phosphopeptide identification of LC/MS/MS data we rely on the SEQUEST software (Thermo Finnigan, San Jose, CA). The routine search settings are as follows: Delta CN for peptides is 0.1; Xcorr versus charge state is set to 1.50 for singly, 2.0 for doubly, and 2.5 for triply charged peptides, respectively; minimal protein probability is 0.01. In principle, phosphorylation stoichiometries can be quantified by comparing the integrated peak intensities of a phosphopeptide and its nonphosphorylated counterpart. In doing so, it is assumed that both forms of the peptide ionize with equal efficiencies. This estimation of stoichiometries is in many cases valid for singly and doubly phosphorylated peptides. However, with increasing clustering of phosphorylation sites we have found that extensively phosphorylated peptides tend to become poorly ionized. In such cases, we prefer H218O-labelling and phosphatase treatment. This allows estimation of relative changes in the extent of phosphorylation because an isotopically labelled pair of peptides with equal ionization prope ties is generated. Also, for MALDI-TOF experiments, neutral loss of phosphate does not occur, while the relative extent of phosphorylation is preserved. Absolute quantitation of the extent of phosphorylation is possible with the recently introduced AQUA technology (32). The search parameters for the identification vary, of course, with the software used for protein identification. With the SEQUEST protein/peptide identification, we set the search parameters variable for Ser/Thr/Tyr phosphorylation (+80 Da) and variable Ser/Thr modification to −18 Da for those peptides that underwent loss of phosphoric acid and that had been fragmented in an MS3 scan.

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References 1. Hubbard, M.J., and Cohen, P. (1993) On target with a new mechanism for the regulation of protein phosphorylation. Trends Biochem. Sci. 18, 172–177. 2. Hunter, T. (1991) Protein kinase classification. Methods Enzymol. 200, 3–37. 3. Cohen, P. (1992) Signal integration at the level of protein kinases, protein phosphatases and their substrates. Trends Biochem. Sci. 17, 408–413. 4. Boyle, W.J., van der Geer, P., and Hunter, T. (1991) Phosphopeptide mapping and phosphoamino acid analysis by two-dimensional separation on thin-layer cellulose plates. Methods Enzymol. 201, 110–152. 5. Luo, K., Hurley, T.R., and Sefton, B.M. (1991) Cyanogen bromide cleavage and proteolytic peptide mapping of proteins immobilized to membranes. Methods Enzymol. 201, 149–152. 6. Wettenhall, R.E., Aebersold, R.H., and Hood, L.E. (1991) Solid-phase sequencing of 32P-labeled phosphopeptides at picomole and subpicomole levels. Methods Enzymol. 201, 186–199. 7. Kuiper, G.G.J.M., and Brinkmann, A.O. (1995) Phosphotryptic peptide analysis of the human androgen receptor: detection of a hormone-induced phosphopeptide. Biochemistry 34, 1851–1857. 8. Winz, R., Hess, D., Aebersold, R., and Brownsey, R.W. (1994) Unique structural features and differential phosphorylation of the 280-kDa component (isozyme) of rat liver acetyl-CoA carboxylase. J. Biol. Chem. 269, 14438–14445. 9. Payne, M.D., Rossomando, A.J., Martino, P., Erickson, A., K., Her, J.-H., Shabanowitz, J., Hunt, D.F., Weber, M.J., and Sturgill, T.W. (1991) Identification of the regulatory phosphorylation sites in pp42/mitogen-activated protein kinase (MAP kinase). EMBO J. 10, 885–892. 10. Resing, K.A., Johnson, R.S., and Walsh, K.A. (1995) Mass spectrometric analysis of 21 phosphorylation sites in the internal repeat of rat profilaggrin, precursor of an intermediate filament associated protein. Biochemistry 34, 9477–9487. 11. Verma, R., Annan, R.S., Huddleston, M.J., Carr, S.A., Reynard, G., and Deshaies, R.J. (1997) Phosphorylation of Sic1p by G1 Cdk required for its degradation and entry into S phase. Science 278, 455–460. 12. Kalo, M.S., and Pasquale, E.B. (1999) Multiple in vivo tyrosine phosphorylation sites in EphB receptors. Biochemistry 38, 14396–14408. 13. Li, X., Gerber, S.A., Rudner, A.D., Beausoleil, S.A., Haas, W., Villén, J., Elias, J.E., and Gygi, S.P. (2007) Large-scale phosphorylation analysis of alpha-factor-arrested Saccharomyces cerevisiae. J Proteome Res. 6, 1190–11977. 14. Chi, A., Huttenhower, C., Geer, L.Y., Coon, J.J., Syka, J.E, Bai, D.L., Shabanowitz, J., Burke, D.J., Troyanskaya, O.G., and Hunt, D.F. (2007) Analysis of phosphorylation sites on proteins from Saccharomyces cerevisiae by electron transfer dissociation (ETD) mass spectrometry. Proc. Natl. Acad. Sci. USA 104, 2193–2198. 15. Larsen, M.R., Sørensen, G.L., Fey, S.J., Larsen, P.M., and Roepstorff, P. (2001) Phospho-proteomics: evaluation of the use of enzymatic de-phosphorylation and differential mass spectrometric peptide mass mapping for site specific phosphorylation assignment in proteins separated by gel electrophoresis. Proteomics 1, 223–238.

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16. Schlosser, A., Pipkorn, R., Bossemeyer, D., and Lehmann, W.D. (2001) Analysis of protein phosphorylation by a combination of elastase digestion and neutral loss tandem mass spectrometry. Anal. Chem. 73, 170–176. 17. Tholey, A., Reed, J., and Lehmann, W.D. (1999) Electrospray tandem mass spectrometric studies of phosphopeptides and phosphopeptide analogues. J. Mass Spectrom. 34, 117–123. 18. Lu, B., Ruse, C., Xu, T., Park, S.K., and Yates, J. 3rd (2007) Automatic validation of phosphopeptide identifications from tandem mass spectra. Anal. Chem. 79, 1301–1310. 19. Perkins, D.N., Pappin, D.J., Creasy, D.M., and Cottrell, J.S. (1999) Probabilitybased protein identification by searching sequence databases using mass spectrometry data. Electrophoresis 20, 3551–3567. 20. Zhang, Y., Wolf-Yadlin, A., Ross, P.L., Pappin, D.J., Rush, J., Lauffenburger, D.A., and White, F.M. (2005) Time-resolved mass spectrometry of tyrosine phosphorylation sites in the epidermal growth factor receptor signaling network reveals dynamic modules. Mol. Cell. Proteomics 4, 1240–1250. 21. Gruhler, A., Olsen, J.V., Mohammed, S., Mortensen, P., Faergeman, N.J., Mann, M., and Jensen, O.N. (2005) Quantitative phosphoproteomics applied to the yeast pheromone signaling pathway. Mol. Cell. Proteomics 4, 310–327. 22. Tao, W.A., Wollscheid, B., O’Brien, R., Eng, J.K., Li, X.J., Bodenmiller, B., Watts, J.D., Hood, L., and Aebersold, R. (2005) Quantitative phosphoproteome analysis using a dendrimer conjugation chemistry and tandem mass spectrometry. Nat. Methods 2, 591–598. 23. Bonenfant, D., Schmelzle, T., Jacinto, E., Crespo, J.L., Mini, T., Hall, M.N., and Jenoe, P. (2003) Quantitation of changes in protein phosphorylation: a simple method based on stable isotope labeling and mass spectrometry. Proc. Natl. Acad. Sci. USA 100, 880–885. 24. Ficarro, S.B., McCleland, M.L., Stukenberg, P.T., Burke, D.J., Ross, M.M., Shabanowitz, J., Hunt, D.F., and White, F.M. (2002) Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nat. Biotechnol. 20, 301–305. 25. Blagoev, B., Ong, S.E., Kratchmarova, I., and Mann, M. (2004) Temporal analysis of phosphotyrosine-dependent signaling networks by quantitative proteomics. Nat. Biotechnol. 22, 1139–1145. 26. Gatlin, C.L., Eng, J.K., Cross, S.T., Detter, J.C., and Yates, III, J.R. (2000) Automated identification of amino acid sequence variations in proteins by HPLC/microspray tandem mass spectrometry. Anal. Chem. 72, 757–763. 27. Radimerski, T., Mini, T., Schneider, U., Wettenhall, R.E., Thomas, G., and Jenoe, P. (2000) Identification of insulin-induced sites of ribosomal protein S6 phosphorylation in Drosophila melanogaster. Biochemistry 39, 5766–5774. 28. Oda, Y., Huang, K., Cross, F.R., Cowburn, D., and Chait, B.T. (1999) Accurate quantitation of protein expression and site-specific phosphorylation. Proc Natl Acad Sci USA. 96, 6591–6596. 29. Bantscheff, M., Eberhard, D., Abraham, Y., Bastuck, S., Boesche, M., Hobson, S., Mathieson, T., Perrin, J., Raida, M., Rau, C., Reader, V., Sweetman, G., Bauer, A., Bouwmeester, T., Hopf, C., Kruse, U., Neubauer, G., Ramsden, N., Rick, J., Kuster, B.,

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and Drewes, G. (2007) Quantitative chemical proteomics reveals mechanisms of action of clinical ABL kinase inhibitors. Nat Biotechnol., Epub ahead of print. 30. Beck, T., Schmidt, A., and Hall, M.N. (1999) Starvation induces vacuolar targeting and degradation of the tryptophan permease in yeast. J. Cell Biol. 146, 1227–1238. 31. Larsen, M.R., Graham, M.E., Robinson, P.J., and Roepstorff, P. (2004) Improved detection of hydrophilic phosphopeptides using graphite powder microcolumns and mass spectrometry: evidence for in vivo doubly phosphorylated dynamin I and dynamin III. Mol. Cell. Proteomics 3, 456–465. 32. Gerber, S.A., Rush, J., Stemman, O., Kirschner, M.W., and Gygi, S.P. (2003) Absolute quantification of proteins and phosphoproteins from cell lysates by tandem MS. Proc. Natl. Acad. Sci. USA 100, 6940–6945.

162 Protein Microarrays for Phosphorylation Studies Birgit Kersten and Tanja Feilner 1. Introduction Posttranslational modification of proteins by phosphorylation is the most abundant type of cellular regulation which affects essentially every intracellular process of eukaryotes. Phosphorylation of a protein can cause changes in its structure, stability, enzymatic activity, the ability to interact with other molecules, or its subcellular localization. Protein kinases are catalyzing the reversible phosphorylation of protein substrates at serine, threonine or tyrosine residues. In Arabidopsis, e.g., serin/threonine kinases represent about 4% of the proteome (1), but their biological function is not well understood. Therefore, high-throughput proteomics methods (2,3) for global analysis of protein kinase function are needed to identify downstream substrates. Various techniques for determining consensus phosphorylation site sequences, including peptide libraries (4) and peptide arrays (5,6) led to the identification of kinase substrates. Furthermore, a solid-phase phosphorylation screening of λ phage c-DNA expression libraries (7,8) as well as different protein-protein-interaction screening methods, such as overlay methods (9–11) and the yeast two hybrid system (12–14) have been applied in this respect. More recently protein kinases were engineered to accept unnatural adenosine triphosphate (cyclopentyl ATP) analogues and have been used to identify specific substrates (15–17). Furthermore, preliminary studies demonstrated the suitability of protein microarrays to study phosphorylation by kinases (18,19). To elucidate the sites of phosphorylation, antibodies against phosphorylated protein epitopes (6) may be used for detection on protein microarrays. Furthermore, peptide arrays (5,6) or MSbased methods (20,21) may be applied in this respect. We here describe a protein microarray-based method for the phosphorylation screening of proteins and the identification of potential kinase substrates in a high-throughput manner. We successfully used this screening tool to From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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identify novel targets for the barley casein kinase 2α (CK2α) (22) as well as for different Arabidopsis thaliana mitogen activated kinases (MAP kinases) (23). Our method utilizes plant protein microarrays which have been generated as described in detail previously (22, 24). Then microarrays have to be incubated with the soluble and active kinase in the presence of radioactive [γ33-P]ATP. Radioactive signals detected by phosphor imager or X-ray film were then evaluated for the selection of potential substrates. We verified the potential substrates by on blot-phosphorylation in vitro. As a whole, our approach allows to shortlist candidate substrates of plant protein kinases for further analysis. Follow-up in vivo experiments are essential to evaluate their physiological relevance (13,25,26). 2. Materials 2.1. Purification of Recombinant Kinases under Native Conditions 1. Medium for culturing cells: 2YT or LB containing 100 µg/ml ampicillin, 15 µg/ml kanamycin and 2% glucose. 2. Isopropyl-β-D-thiogalactopyranoside, IPTG. 3. Native lysis buffer: 300 mM NaCl, 50 mM NaH2PO4, 10 mM imidazole, pH 8.0. 4. Native wash buffer: 300 mM NaCl, 50 mM NaH2PO4, 20 mM imidazole, pH 8.0. 5. Native elution buffer: 300 mM NaCl, 50 mM NaH2PO4, 250 mM imidazole, pH 8.0. 6. Lysozyme. 7. PMSF. 8. Ultrasonic homogeniser (Branson Ultrasonic, Danbury, CT). 9. 1 ml polypropylen columns (Qiagen, Hilden, Germany). 10. Bradford reagent (Bio-Rad Labratories GmbH, Munich).

2.2. Kinase Assay on Protein Microarrays 1. TBS+Tween (TBST): 10 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.1% (v/v) Tween. 2. Blocking solution: 2% bovine serum albumin, BSA (Sigma, St. Louis, MO) in TBST. 3. [γ33-P]ATP, 250 µCi/ml (Amersham Pharmacia Biotech Europe GmbH, Freiburg, Germany). 4. CK2α buffer: 25 mM Tris–HCl pH 8.5, 10 mM MgCl2, 1 mM DTT. 5. FAST™-slides (Whatman Schleicher & Schuell, Dassel, Germany). 6. Cover slide (Carl Roth GmbH, Karlsruhe, Germany). 7. X-ray cassette (Hypercassette, Amersham Pharmacia, Freiburg, Germany). 8. X-ray film (Kodak, Stuttgart, Germany). 9. Imaging screen (BAS-SR 2025, Fujifilm, Japan). 10. Phosphor imager (BAS-Reader-5000, Fujifilm, Japan).

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3. Methods 3.1. Purification of Recombinant Kinases under Native Conditions For the phosphorylation screening the kinase has to be soluble, active and pure, which means that no other kinase should be present. To achieve this a possibility would be a large scale production and purification of recombinant His-tagged protein kinases from cDNA expression libraries, e.g. a barley expression library constructed in the E. coli vector pQE30NST (accession number: AF074376; 22) or an Arabidopsis expression library in vector pQE30NASTattB (accession number: AY386205; 23) (see also 24). In order to express and purify protein kinases from these libraries we apply the following protocol: For expression: 1. 50 ml falcon tubes were filled with 10 ml of medium for culturing cells. 2. Cultures were inoculated with bacteria from 384-well plates, which were stored at −80°C. 3. After overnight growth at 37°C with vigorous shaking the cultures were transferred into 300 ml Erlenmeyer flasks and 90 ml of prewarmed 2YT medium supplemented with 100 µg/ml of ampicillin, and 15 µg/ml of kanamycin were added. The incubation was continued until an OD of 0.7 was reached. 4. To induce protein expression, IPTG was added to a final concentration of 1 mM, and incubation was continued for 4 h at 37°C. 5. Cells were transferred into 50 ml falcon tubes (two for each culture), harvested by centrifugation at 1,900 × g for 10 min and stored immediately at −80°C.

For native protein purification (all of the following steps were performed at 0−4°C): 1. The frozen pellets were thawed on ice. 2. Cells were lysed in 0.5 ml of native lysis buffer supplemented with 0.25 mg/ml of lysozyme and 0.1 mM of PMSF. 3. Lysates were pooled and DNA was sheared with an ultrasonic homogeniser for 3 × 1 min at 50% power on ice. 4. Lysates were transferred into 1.5 ml Eppendorf-tubes and centrifugated at 20,000 × g, 4°C. 5. Supernatants were transferred into fresh Eppendorf-tubes. 6. Then 250 µl of NiNTA–agarose were added to bind the His-tagged proteins by shaking for 1 h at 300 rpm at 4°C. 7. The mixtures were transferred to 1 ml polypropylen columns. 8. The columns were washed with 10 bed volumes native wash buffer. 9. Proteins were eluted with 4 elution steps using 0.5 ml of native elution buffer each. 10. Proteins were mixed with glycerol (20% (v/v) end concentration) and stored at −80°C (see Note 1).

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Fig. 1. 15% SDS-Page of two native purified kinases and calmodulin (Coomassie-stained). Nine samples of each purification step were separated: 1: pellet (insoluble fraction after centrifugation); 2: supernatant (soluble fraction after centrifugation); 3: first column flowthrough; 4+5: first and second washing fraction; 6–9: eluates. M, molecular weight marker. Approximate size of the protein markers are shown at the left in kDa.

We would recommend collecting an aliquot from each centrifugation-, lysis-, wash- and elution step in order to control the efficiency of the purification steps using a polyacrylamide gel, e.g. 15% (see Fig. 1). Protein concentration was determined by Bradford assay (27). 3.2. Kinase Assay on Protein Microarrays We recommend checking the activity of the kinase prior to microarray experiments (see Note 2) using a known substrate as a positive control (see Note 3). Furthermore, prior to microarray experiments, one has to exclude that the kinase will phosphorylate 3′- or 5′-tags of the recombinant proteins, such as His-tags (see Note 4). We intended to perform a qualitative signal evaluation after kinase assay. Therefore we have spotted proteins and controls (see Note 3) in quadruplicates on FAST™-slides (see Note 5) using a 10 × 10 spotting pattern (see Note 6) as demonstrated in Fig. 2 (22). Spotting pattern for further quantitative evaluation has been described recently (23). For kinase assay: 1. Microarrays, which were spotted the day before and stored at 4°C in a closed slide holder, were washed for 1 h in TBST with vigorous shaking at room temperature (see Note 7). 2. Washed microarrays were blocked for 1 h at room temperature with 2% BSA/TBST. 3. Blocked microarrays were placed on Whatman paper soaked with TBST to avoid drying of the micorarrays during the subsequent kinase reaction.

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4. Microarrays were incubated with 250 µl kinase solution containing 13 µg/ml of CK2α and 25 µCi/ml of radioactive labelled [γ33-P]ATP in CK2α buffer for 1 h at room temperature (see Note 1). This incubation was performed underneath a cover slide. 5. Six wash steps of 30 min each were performed in TBST. 6. Finally, the microarrays were dried using a microtiter plate centrifuge (e.g., Eppendorf AG, Hamburg, Germany: 5810R) or by manual fanning and transferred into an X-ray cassette. 7. Signal detection was performed by means of an X-ray film. The film was laid on the microarrays and the cassette was stored for a suitable time at −80°C (see Note 8). Afterwards, the film was developed in a dark chamber. Alternatively, the microarrays were exposed to an imaging screen, which was screened afterwards by a phosphor imager for signal detection (see Note 8).

Fig. 2 (left) shows a typical result of a phosphorylation screening experiment with CK2α after exposure of the microarrays to X-ray film. Quadruples of several proteins gave distinct signals on the film. In order to determine the immobilisation of the recombinant His-tagged proteins on the microarrays we screened them with an anti-RGS-His6 antibody as described in detail previously (24). Nearly all of the spotted proteins were detectable (~95%) as shown in Fig. 2 (right). 3.3. Selection of Potential Kinase Targets For qualitative signal evaluation, the results of two independent experiments with different preparations of the kinase have been used. The criteria for the selection of the potential kinase targets were: 1. A protein was considered as positive in one experiment if all 4 spots of the corresponding quadruplicate gave a distinct signal on the X-ray film or in the phosphor imager (see Fig. 2). 2. A protein was regarded as potential target protein if it attained positive results in both experiments.

Thus, for example 21 potential CK2α target proteins were identified out of nearly 800 different barley proteins (22). The selection of potential targets using a threshold-based quantification system after phosphorylation on microarrays has been described in high throughput studies in Arabidopsis thaliana (23) as well as in yeast (28). 3.4. Verification of Potential Phosphorylation Targets As this microarray-based phosphorylation method is a screening tool to identify potential phosphorylation targets in vitro, we would recommend a verification of the identified targets by other methods. For such verifications various in vitro or in vivo approaches may be applied. A few of them have already been mentioned in the introduction.

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We performed an on blot-phosphorylation assay to verify potential substrates: 1. Proteins were separated using an SDS–PAGE, e.g., 15%, followed by blotting the proteins on PVDF membranes. After blotting, gels were Coomassie stained. 2. Phosphorylation reaction was carried out on the blot membrane using reaction conditions as described for the microarray-based assay in an appropriate volume.

Fig. 3 shows a typical example of such an on blot-phosphorylation experiment. Fourty-eight proteins, which were identified in both microarray experiments as potential substrates using a MAP kinase were verified with this method. Nearly all of them were confirmed as shown in Fig. 3. We used the same positive controls as in the microarray experiments: different concentrations of myelin basic protein (MBP; artificial substrate of MAP kinases). For negative controls proteins were taken, which were tested in the microarray-based kinase assay and were not identified as potential substrates (see Fig. 3). Other in vitro verification methods are phosphorylation assays performed in solution with subsequent detection of the phosphorylated proteins by SDS electrophoresis and autoradiography (8,23). Such assay has been used by Fukunaga and Hunter to verify candidate kinase substrates identified by phosphorylation screening of an expression phage cDNA library (8). 4. Notes 1. We recommend to store kinases at −80°C by using glycerol (end concentration 20%) or to lyophilise the kinases in their according buffer and to store them afterwards at −80°C. The lyophilised kinases are only being dissolved in ddH2O shortly before use. 2. Prior to performing micorarray-based kinase experiments we recommend to test the activity of the prepared kinase by in gel assays (29) or kinase assays in solution (8) using a known substrate of the respective kinase (see

Fig. 2. Kinase- and immunoassay with 768 different barley proteins, immobilized on FAST™-slides (384 proteins in each field). Left: X-ray film image with the results of a representative CK2α-assay. Right: equal microarrays screened with an anti-RGS-His6 antibody. The map below shows the positions of the 16 controls. The purified proteins and the controls were spotted in quadruplicates. Controls: 1, 4, 13, 16: HMG(1) (GenBank Acc.: CK124404), 0.29 mg/ml; 2: HMG(2) (GenBank Acc.: CK122788), 0.76 mg/ml; 3: HMG(3) (GenBank Acc.: CK124847), 0.28 mg/ml; 5: CK2α (GenBank Acc.: CK123377), 0.22 mg/ml; 6: other library kinase (GenBank Acc.: CK125548), 0.21 mg/ml; 7: human gapdh (His-tagged recombinant human glyceraldehyde-3-phosphate dehydrogenase), 0.36 mg/ml; 8: BSA, 20 pmol/µl in PBS; 9: native elution buffer; 10: denaturing elution buffer; 11: mouse anti-RGS-His6 antibody, diluted 1:10 in PBS; 12: rabbit-anti-mouse IgG-Cy3 conjugate, diluted 1:25 in PBS; 14: H2O; 15: PBS. (Reprinted with permission from ref. 22. Copyright 2004 Elsevier Science).

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Fig. 3. Verification of kinase substrates. 15% SDS-PAGE gels of potential kinase substrates (upper panel) and autoradiograms of these proteins after blotting and kinase reaction (lower panel). Gels were Coomassie-stained after electrophoresis and blotting. MBP1 – MBP5: positive controls, MBP in various dilution steps (MBP1: 2,000 ng/µl; MBP2: 1,000 ng/µl; MBP3: 500 ng/µl; MBP4: 250 ng/µl; MBP5: 125 ng/µl). 1 – 48: potential kinase substrates identified in the microarray experiments. C1-C3: negative controls; these are proteins, which were not identified as potential substrates using microarrays. RM: rainbow marker; M: molecular weight marker. Approximate size of the protein markers are shown at the left in kDa.

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4.

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Note 3). The optimal conditions for kinase activation (e.g., optimal buffer conditions, kinase concentration and incubation time of the kinase) must be determined experimentally for each individual kinase, if they are not known from the literature. The use of a positive control is recommendable (e.g., a known substrate of the kinase). In studies with barley CK2α (22) we used as positive controls different barley proteins, which share a strong homology with different plant HMG (high mobility group) proteins. These proteins are well known targets of the CK2α (30,31). In another study we analyzed different Arabidopsis MAP kinases (23). In this case a known artificial substrate, the myelin basic protein (MBP), was used as a positive control. Recombinant proteins often contain 3’- or 5’- tags which are encoded by the cloning vector besides the coding sequence of the respective protein. Phosphorylation of these tags may result in false-positive results and should therefore be excluded prior to microarray experiments. One possibility is to perform test assays in solution with the active kinase using some recombinant proteins which are known to be no substrates of the respective kinase and which were expressed the same manner (same tags) as the recombinant proteins used for the microarray experiments. These selected proteins should yield negative results in the test. With respect to surface coatings we tried to phosphorylate immobilized denatured proteins on FAST™-slides (coated with a nitrocellulose-derived polymer) and epoxy slides. We detected phosphorylation only on FAST™slides although His-tagged proteins were detectable with anti-RGS-His6 antibody on both surfaces. In other studies, nanowells (19), BSA-NHS (BSA-N-hydroxysuccinimide) slides (18), or SAM2 (streptavidin-coated membrane) slides (32) have been used as surfaces in microarray-based kinase assays. We reviewed different micorarray surfaces for different microarray applications including phosphorylation studies (33). The maximum spot density, at these individual spots can still be differentiated in a phosphorylation assay, is highly dependent on the resolution of the phosphor imager, which scanned the imaging plates. For the determination of this density we recommend the use of commercially available protein kinase (e.g., PKA from New England Biolabs GmbH). Our experience is, when using a scanner with a resolution of 10 µm (e.g., BioImager FLA 8,000, Fujifilm, Japan) in combination with spotting patterns up to 11 × 11 (spot distance: 410 µm), that we received sufficient resolution to evaluate the radioactive signals. Applications of higher spotting pattern (14 × 14-pattern with a spot distance of 321 µm; 15 × 15-pattern with a spot distance of 300 µm) were not suitable for radioactive detection, as signals of intense spots were interfering with neighbouring spot signals. This lead

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to faulty analysis of the signal intensity of neighbouring proteins as well as faulty background corrections. 7. This washing step in TBST was performed to remove urea from the microarrays, as we recognised that the urea reduces the activity of the kinase in the microarray assay. 8. For the detection of radioactive signals with X-ray film, the film was laid onto the microarrays, which have been covered with cling film. The exposition time is dependent on the signal intensity and can vary between 30 min to a few days. The sensitivity of the X-ray films can cause problems, as the blackening of the films is only in a very limited area linear and proportional to the signal intensity. For the detection with a phosphor imager the imaging plates are laid onto the covered microarrays and are then scanned. Detection by phosphor imager is 10–100 × more sensitive than by X-ray films and therefore it is possible to get faster results and/or detect poor signals respectively. Furthermore, the system has a higher dynamic range compared to the X-ray film. Strong signals as well as poor signals are detected within one exposition and the linear signal-to-intensity range has a dimension over 5 (100,000:1). This shows that the phosphor imager is of better use for quantification of radioactive signals than X-ray films. Acknowledgments We grateful thank Armin Kramer for providing Fig. 1 and Jasmin Bastian for reading the manuscript.

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23. Feilner, T., Hultschig, C., Lee J., Meyer S., Immink, R.G.H., Koenig A., Possling A., Seitz, H., Beveridge, A., Scheel, D., Cahill, D.J., Lehrach, H., Kreutzberger, J., and Kersten, B. (2005) High throughput identification of potential Arabidopsis Mitogen-activated protein kinases substrates. Mol. Cell. Proteomics 4, 1558–1568. 24. Kersten, B. and Feilner, T. (2007) Generation of plant protein microarrays and investigation of antigen-antibody interactions. Methods Mol. Biol. 355, 365–378. 25. Liu, Y. and Zhang, S. (2004) Phosphorylation of 1-aminocyclopropane-1-carboxylic acid synthase by MPK6, a stress-responsive mitogen-activated protein kinase, induces ethylene biosynthesis in Arabidopsis. Plant Cell 16, 3386–3399. 26. Riera, M., Figueras, M., Lopez, C., Goday, A., and Pages, M. (2004) Protein kinase CK2 modulates developmental functions of the abscisic acid responsive protein Rab17 from maize. Proc. Natl. Acad. Sci. U S A 101, 9879–9884. 27. 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, 248–254. 28. Ptacek, J., Devgan G., Michaud G., Zhu H., Zhu X., Fasolo J., Guo H., Jona G., Breitkreutz A., Sopko R., McCartney R.R., Schmidt M.C., Rachidi N., Lee S.J., Mah A.S., Meng L, Stark M.J., Stern D.F., De Virgilio C., Tyers M., Andrews B., Gerstein M., Schweitzer B., Predki P.F., and Snyder M. (2005) Global analysis of protein phosphorylation in yeast. Nature 438, 79–84. 29. Usami, S., Banno, H., Ito, Y., Nishihama, R., and Machida, Y. (1995) Cutting activates a 46-kilodalton protein kinase in plants. Proc. Natl. Acad. Sci. U S A 92, 8660–8664. 30. Grasser, K. D., Maier, U. G., and Feix, G. (1989) A nuclear casein type II kinase from maize endosperm phosphorylating HMG proteins. Biochem. Biophys. Res. Commun. 162, 456–463. 31. Stemmer, C., Schwander, A., Bauw, G., Fojan, P., and Grasser, K. D. (2002) Protein kinase CK2 differentially phosphorylates maize chromosomal high mobility group B (HMGB) proteins modulating their stability and DNA interactions. J. Biol. Chem. 277, 1092–1098. 32. Boutell, J. M., Hart, D. J., Godber, B. L., Kozlowski, R. Z., and Blackburn, J. M. (2004) Functional protein microarrays for parallel characterisation of p53 mutants. Proteomics 4, 1950–1958. 33. Feilner, T., Kreutzberger, J., Niemann, B., Kramer, A., Possling, A., Seitz, H., and Kersten, B. (2004) Proteomic studies using microarrays. Curr. Proteomics 1, 283–295.

163 Two-Dimensional Phosphopeptide Mapping Hikaru Nagahara, Robert R. Latek, Sergei A. Ezhevsky, and Steven F. Dowdy 1. Introduction A major mechanism that cells use to regulate protein function is by phosphorylation and/or dephosphorylation of serine, theonine and tyrosine residues. Phosphopeptide mapping of these phosphorylated residues allows investigation into the positive and negative regulatory roles these sites may play in vivo. In addition, phosphopeptide mapping can uncover the specific phosphorylated residue and hence, kinase recognition site, thus helping in the identification of the relevant kinase(s) and/or phosphatase(s). Two-Dimensional (2D) Phosphopeptide mapping can utilize in vivo and in vitro 32P labeled proteins (1–6). Briefly, 32P labeled proteins are purified by SDS-PAGE, transferred to a nitrocellulose filter and digested by proteases or chemicals. The phosphopeptides are then separated by electrophoresis on thin layer cellulose (TLC) plate in the first dimension followed by thin layer chomatography in an organic buffer in the sond dimension. The TLC plate is then exposed to autoradiographic (ARG) film or phosphor-imager screen and the position of the 32P containing peptides are thus identified. Specific phosphopeptides can then be excised from the TLC plate and analyzed further by amino acid hydrolysis to identify the specific phosphorylated residue(s) and/or by manual amino-terminal sequencing to obtain the position of the phosphorylated residue(s) relative to the protease cleavage site (3). In addition, mixing in vivo with in vitro 32P-labeled proteins can yield confirmation of the specific phosphorylated residue and the relevant kinase.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2. Materials 2.1. Equipment 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Multiphor II horizontal electrophoresis apparatus (Pharmacia). Power pack capable of 1000 V constant. Refrigerated circulating water bath. Thin layer chomatography (TLC) chamber, ~30 L × 10 W × 28H cm, and internal stand. Speedy-vac or lyophilizier. Shaking water bath. SDS-PAGE apparatus. Semi-Dry blotting apparatus (Owl). Small fan. Rotating wheel or apparatus.

2.2. Reagents 1. Phosphate-free tissue culture medium. 2. Phosphate-free dialyzed fetal bovine serum (FBS). Alternatively, dialyze 100 ml FBS against 4 L dialysis buffer for 12 h and repeat two more times using 10,000 MWCO dialysis tubing. Dialysis buffer: 32 g NaCl, 0.8 g KCl, 12 g Tris in 4 L, pH to 7.4 with HCl. 3. 32PO42− orthophosphate, 3–5 mCi per tissue culture dish. 4. Protein extraction buffer (ELB): 20 mM HEPES (pH 7.2), 250 mM NaCl, 1 mM EDTA, 1 mM DTT, 0.1–0.5% NP-40 or Triton X-100, 1 µg/ml Leupeptin, 50 µg/ ml PMSF, 1 µg/ml Aprotitin, and containing the following phosphatase inhibitors: 0.5 mM NaP2O7, 0.1 mM NaVO4, 5.0 mM NaF. 5. Rabbit anti-Mouse IgG. 6. Killed S. aureus cells (Zyzorbin). 7. Protein A agarose. 8. 2× Sample Buffer: 100 mM Tris-HCl (6.8), 200 mM DTT, 4% SDS,0.2% bromophenol blue, 20% glycerol. 9. Protein Transfer buffer: 20% methanol, 0.037% SDS, 50 mM Tris, 40 mM glycine. 10. 50 mM N H4HCO3, made fresh each usage from powder. 11. 0.5% (w/v) Polyvinyl pyrolidone-360 in 100 mM acetic acid. 12. Sequencing grade trypsin (Boehinger Mannheim), resuspend 100 µg in 1.5 ml fresh 50 mM NH4HCO3, store 10 µg/150 µl aliquots at −20°C. 13. Performic acid: Mix 1 vol hydrogen peroxide with 9 vol formic acid. Incubate for 1.5 h on ice. 14. Scintillation fluid. 15. Thin-layer chomatography cellulose plastic-backed plates, 20 × 20 cm (Bakerflex/VWR). 16. pH 1.9 Electrophorese Running Buffer: 50 ml 88% formic acid, 56 ml acetic acid and 1894 ml H2O. Do not adjust pH.

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17. Electrophoresis color marker: 5 mg/ml DNP-lysine, 1 mg/ml xylene cyanol FF in 50 µl n-butanol, 25 µl pyridine, 25 µl acetic acid, 1.9 ml H2O. 18. TLC chamber buffer: 75 ml n-butanol, 50 ml pyridine, 15 ml glacial acetic acid, 60 ml H 2O.

3. Methods 3.1. 32P Orthophosphate Labeling 1. For in vivo 32P orthophosphate labeling of cellular proteins, preplate approximately 1 × 106 cells in a 10 cm dish. Rinse adherent cells thee times with 5 ml phosphatefree media. Suspension cells can be rinsed and collected by centrifugation at ~1800 rpm for 5 at RT or 30°C, aspirate the media and repeat as above. Add 3–5 mCi of 32P ortho phosphate in 3.5 ml of phosphate-free media containing 10% dialyzed serum to the 10 cm dish and incubate cells at 37°C for 4–6 h (see Note 1). 2. Aspirate the 32P containing media with a plastic pipette, transfer supernatant waste into a 50 ml conical disposable tube. Rinse the cells twice with 10 ml PBS(-) and combine with 32P media waste in 50 ml tube and dispose of properly. 3. Add 1 ml ice cold extraction buffer (ELB; 7) and place dish on a flat bed of ice, behind a shield. Tilt dish slightly every 30 s for 3’–5’ to continually cover the cells. Collect cellular lysate and debris by tilting dish approximately 30° using a P-1000 pipetman tip and transfer to 1.5 ml eppendorf tube and mix. Alternatively, adherent cells can be released by trypsin/EDTA addition, collected and washed twice in media containing serum to inactivate the trypsin. After the final centrifugation, add 1 ml ELB, mix by using a P-1000 tip and transfer to an eppendorf tube. Place tube on ice for 20’–25’ with occasional mild inverting (see Note 2). 5. Spin out insoluble particular matter from the cellular lysate in microfuge at 12 K, 4°C, for 10’. Transfer supernatant to new eppendorf tube and preclear lysate by the addition of 50 µl killed S. aureus cells, cap tube and place on rotating wheel at 4°C for 30’–60’. 6. To remove S. aureus cells, centrifuge lysate at 12 K, 4°C for 10 . Transfer lysate to fresh eppendorf tube, making sure to leave the last 50 µl or so of lysate behind with the pellet. The presence of contaminating S. aureus cells in this portion of the sample will reduce the amount of immunocomplexes recovered if included.

3.2. Immunoprecipitation and Transfer of 32P-Labeled Protein 1. Add primary antibody to the precleared lysate supernatant, usually 100–200 µl in the case of hybridoma supernatants and 3–5 µl of commercially purified antibodies. Add 30 µl protein A agarose beads, cap the tube, and place on a wheel at 4°C for 2–4 h to overnight. If primary mouse antibody isotype is IgG1 or unknown, add 1 µl Rabbit anti-Mouse IgG to allow indirect binding of the primary antibodies to the protein A agarose (7). 2. After incubation with primary antibody, perform a “1 g spin” by placing the tube on ice for 15’, aspirate the supernatant to just above the protein A agarose bed level.

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Stop aspirating at the top of the agarose beads, avoiding drying the beads. This supernatant waste is still highly radioactive. Wash the agarose beads by addition of 1 ml ice cold ELB, cap and invert tube several times and centrifuge at 12 K, 4°, 30 s. Aspirate supernatant and repeat two to thee more times. Again, avoid drying the beads. After the final 30 s spin, aspirate supernatant off of the protein A beads until just dry, add 30 µl of 2× Sample buffer. Boil sample for 5 min, centrifuge 12 K for 10 s, cool tube on ice and load immunocomplexes onto a SDS-PAGE (8). After running the SDS-PAGE, separate the glass plates and trim down the gel with a razor blade by removing the stacking gel and any excess on the sides or bottom. Measure the trimmed gel size and cut six sheets of 3 MM Whatman filter paper and one sheet of nitrocellulose (NC) filter to the same size. Soak two sheets of cut 3 MM filter paper in transfer buffer and place on semi-dry transfer unit. This can be done by filling a tupperware-like container with transfer buffer and dipping the 3 MM paper into it. Place one soaked 3 MM sheet on the back of gel and rub slightly to adhere to gel. Then invert the glass plate with the gel-3 MM still stuck to it and peel gel-3 MM away from the glass with the use of a razor blade. Place it 3 MM side down onto the soaked 3 MM sheets on the semi-dry unit. Soak the cut NC filter with transfer buffer and place it on top of gel followed by thee presoaked 3 MM sheets on top of filter. Gently squeegee out bubbles and excess buffer with the back of your little finger or by rolling a small pipette over the stack. It is important to squeegee out the bubbles, but avoid excess squeegeeing that would result in drying the stack. Mop up the excess buffer on the sides of the stack present on the transfer unit. Place the top on the transfer unit. In this configuration, the bottom plate is the cathode (negative) and the top the anode (positive). Transfer the 32P-labeled proteins to the NC filter at 10 Volts constant for 1.5 to 2 h. The starting current will vary from ~180 to 300 mAmp depending on the surface area of the gel and will drop to around 80 to 140 mAmp by the end of the transfer.

3.3. Trypsinization of Protein on Nitrocellulose Filter 1. Following transfer of 32P-labeled proteins to NC filter, open the transfer unit and using a pair of filter forceps place the NC filter, protein side down onto saran-wrap and cover. Expose the saran-wrap covered NC filter to ARG-film for approximately 1 h to 2 h with protein side towards the film. The length of the exposure will vary with the abundance of 32P content and protein, and with the efficiencies of recovery from immunoprecipitation and transfer. Radioactive or luminescent markers are needed on the filter to determine the orientation, position and alignment of the NC filter with respect to the ARG film. Using the markers, line up your filter on top of its ARG. This is best done on a light box. Use a razor blade to excise the slice of NC filter that corresponds to the 32P-labeled protein band. 2. Place the NC filter slice into an eppendorf tube, add ~200 µl of 0.5% PVP-360 in 100 mM acetic acid, cap tube and incubate in a shaking water bath at 37°C for 30 min ( see Note 3).

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3. Wash the filter slice five times with 1 ml H2O and then twice with 1 ml fresh 50 mM NH4HCO3. 4. Add 10 µg of trypsin in 150 µl of 50 mM NH4HCO3 to the NC filter slice. Incubate in a 37°C shaking water bath over-night. Following this incubation, spike the digestion of the NC filter with another 10 µg of trypsin in 150 µl of 50 mM NH4HCO3 and incubate at 37°C for an additional 4 h. 5. Vortex the tube containing the NC filter/trypsin sample for 1 min, centrifuge at 12 K for 30 s. Transfer the supernatant to a new eppendorf tube. Wash the NC filter slice by addition of 300 µl H2O, vortex 1 min, centrifuge at 12 K for 30 s, and combine the supernatants. 6. Freeze the trypsinized 32P-labeled peptide sample on dry ice and then completely dry in a speedy-vac (without heat). This generally takes about 4 h to complete. Prepare the performic acid for the oxidation step (step 8). 7. After the sample has been completely dried, add 50 µl of ice cold performic acid and place on ice for 1 h. Stop the reaction by addition of 400 µl H2O to the sample, followed by freezing on dry ice and then drying in a speedy-vac. 8. Resuspend the sample in 8–10 µl of H2O. Determine the level of radioactivity by counting 0.5 µl of the sample on a scintillation counter. Usually a total of ≥2000 cpm is sufficient for 2D phosphopeptide mapping.

3.4. Phosphopeptide Separation: First and Second Dimensions 1. Mark the origin on the thin-layer cellulose (TLC) plate by lightly touching a pencil to the TLC plate at the position indicated in Fig. 1. Apply multiple 0.5 µl aliquots of the trypsinized 32P-labeled peptides to the origin to achieve ≥2000 cpm. Dry the TLC plate thoroughly between each aliquot application by use of a gentle fan. Pay particular attention to adding each subsequent aliquot to the same small area at the origin. Add 0.5 µl of the color marker 3 cm from the top edge of the TLC plate (Fig. 1) and then dry the TLC plate under a fan for an additional 30–60 min. 2. Prepare the Multiphor II apparatus for electrophoresis. Place the Multiphor II in a cold room, connect the cooling plate to the cooling circulator bath hoses and precool to 5°C. Prepare and chill the pH 1.9 running buffer and add to Multiphor II buffer tanks. Insert the electrode paddles into innermost chambers, and attach the wire connections. We have used the IEF electrodes in direct contact with the cellulose plate; however, using the paddles provided and wicking buffer onto the plate yields the best results (see Figs. 1 and 2). Place the cooling plate into the Multiphor apparatus. Add 1 L of prechilled pH 1.9 running buffer to each chamber of the Multiphor II. (These instructions are provided with the Multiphor II unit.) 3. Place the loaded TLC plate on top of the cooling plate. To dampen the TLC plate with buffer, first cut a 21 × 21 cm piece of 3 MM Whattman paper and make an approximately 1 cm hole at the origin by puncturing the 3 MM paper with a pencil. Soak the cut 3 MM paper in pH 1.9 running buffer, blotting it between two sheets of dry 3 MM paper, and then placing it over the loaded TLC plate sitting on top of the cooling plate. Slowly pipette running buffer onto the 3 MM paper until the entire cellulose plate is damp beneath, avoid excessive puddling. Remove the paper

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Fig. 1. The location of the origin, anode, cathode and color dye marker dye relative to each other on the 20 × 20 cm TLC plate are depicted (see Subheading 3.4, steps 1 and 2.

and wick a single piece of 13 × 21 cm buffer-soaked 3 MM filter paper from the buffer chamber onto the 2 cm outer edge on both sides of the plate (see Fig. 2). Be sure to fold the paper neatly over the edge of the cooling plate and to make sure that it is evenly contacting the TLC plate. Place the glass cover over the TLC plate touching/resting on the 2 cm overhang of the 3 MM paper wicks. Attach the Multiphor II cover. 4. Electrophorese the peptides on the loaded TLC plate at 1000 V constant for 28–30 min. The run time may be increased up to 38 min. If further separation of 32 P-labeled peptides in this dimension is required, the run time may be increased up to 38 min. 5. Following the first dimension separation by electrophoresis, remove the TLC plate from the Multiphor II apparatus and dry for 1 h with a fan. 6. Place the TLC plate on a stand in a thin-layer chomatography chamber preequilibrated for 48 h in chomatography buffer. The TLC buffer should cover approximately 1 cm of the bottom of the TLC plate when placed on the stand.

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Fig. 2. A cross stional view of the Multiphor II apparatus. Loaded TLC plate, 3 MM Whatman filter paper wicks, and glass cover plate are depicted in the running position (see Subheading 3.4, steps 2 and 3).

Leave the TLC plate in the chamber until the solvent line diffuses to the dye position at the top of the TLC plate, 3 cm below the top of the plate. This usually takes 7–8 h ( see Note 4). 7. Remove the TLC plate from the chamber and dry for >1 h. Expose the TLC plate to ARG-film or a phosphor-imager screen overnight and develop. If the signal is too weak, expose for 5 to 7 days. The use of a phosphor-imager greatly diminishes the length of time required to obtain a 2D phosphopeptide map (see Fig. 3).

4. Notes 1. We routinely label 1 × 106; however, depending on the abundance of the specific protein of interest and on the number of phosphorylation sites this number may vary from 1 × 105 to 1 × 107 cells. In addition, adherent and non adherent cells may both be labeled in suspension. The cells can be labeled with 32P orthophosphate in a 15 or 50 ml disposable conical tube or T-flask. Please note that the activity of kinase(s)/phosphatase(s) present in adherent cells may be altered when labeling in suspension. Dish size is not important as long as enough media is added to cover the bottom of the dish/flask. Attempt to achieve a final 32P orthophosphate concentration of 1.0 to 1.3 mCi/ml of media. 2. We use ELB to lyse cells and generate cellular extracts; however, any extraction buffer containing Triton X-100, SDS, DOC, NP-40 or similar detergent that will lyse cells will suffice. If a strong background is observed following the SDS-PAGE, transferring the immobilized protein A agarose

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Fig. 3. 2-D phosphopeptide map of the retinoblastoma tumor suppressor gene product (pRb) labeled with 32P phosphate in vivo. pRb contains thirteen cyclin-dependent kinase (cdk) phosphorylation sites, hence the complexity of the phosphopeptide map. Note the presence of several levels of 32P intensity associated with specific peptides. This can arise by a number of mechanisms, including in vivo site preferences and/or accessibility of the nitrocellulose immobilized 32P-labeled protein to trypsin. The origin, first and sond dimension runs are as indicated (see Subheading 3.4, step 7).

immunocomplexes to a new eppendorf tube prior to the final wash and centrifugation step can result in a reduced background with minimal loss of specific signal. 3. When running multiple lanes of the same 32P-labeled protein immunocomplexes on SDS-PAGE treat each lane/NC filter slice separately. The trypsinized peptides from as many as five NC filter slices can be combined together, then frozen on dry ice and dried. 4. Seal the top of the TLC chamber with vacuum grease and minimize the amount of time the lid is off of the chamber. Pre-equilibrate chamber for >48 h prior to use. We routinely change the TLC chamber buffer every 8 weeks. Poor separation in the sond dimension is usually indicative of buffer alterations due to evaporation and/or hydration. 5. The procedures described in this chapter can be stopped at the following steps: (1) when the immunocomplexes are in 2× sample buffer following the immunoprecipitation; (2) after lyophilization following trypsinization of the NC filter slices; and (3) after drying the TLC plate following the first dimension eletrophoresis. Acknowledgments We thank Fung Chen for protocols and Jeff Settleman for the “1 g spin.” S.F.D. is an Investigator of the Howard Hughes Medical Institute.

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References 1. Kamp, M.P. and Sefton, B.M. (1989) Acid and base hydrolysis of phosphoproteins bound to immmobilon facilitates analysis of phosphoamino acid in gel-fractionated proteins. Anal. Biochem. 176, 22–27. 2. Lees, J.A., Buchkovich, K.J., Marshak, D.R., Anderson, C.W., and Harlow, E. (1991) The retinoblastoma protein is phophorylated on multiple sites by human cdc2. EMBO 13, 4279–4290. 3. Luo, K., Hurley, T.R., and Sefton, B.M. (1991) Cynogen bromide cleage and proteolytic peptide mapping of proteins immobilized to membranes. Methods Enzymol. 201, 149–152. 4. Desai, D., Gu, Y., and Morgan, D.O. (1992) Activation of human cyclin-dependent kinase in vitro. Mol. Biol. Cell. 3, 571–582. 5. Hardie, D.G. (1993) Protein Phosphorylation: A Practical Approach. IRL Press, Oxford, UK. 6. van der Geer, P., Luo, K., Sefton, B.M., and Hunter, T. (1993) Phosphopeptide mapping and phosphoamino acid analysis on cellulose thin-layer plates. Protein Phosphorylation - A Practical Approach. Oxford University Press, NY. 7. Harlow, E. and Lane, D. (1988) Antibodies: A Laboratory Manual. Cold Spring Harbor Press, NY. 8. Judd, R.C. (1994) Electrophoresis of peptides. Methods Mol. Biol. 32, 49–57.

164 Identification of Proteins Modified by Protein (D-Aspartyl/L-Isoaspartyl) Carboxyl Methyltransferase Darin J. Weber and Philip N. McFadden 1. Introduction The several classes of S-adenosylmethionine-dependent protein methyltransferases are distinguishable by the type of amino acid they modify in a substrate protein. The protein carboxyl methyltransferases constitute the subclass of enzymes that incorporate a methyl group into a methyl ester linkage with the carboxyl groups of proteins. Of these, protein (d-aspartyl/l-isoaspartyl) carboxyl methyltransferase, EC 2.1.1.77 (PCM) specifically methyl esterifies aspartyl residues that through age-dependent alterations are in either the d-aspartyl or the l-isoaspartyl configuration (1,2). There are two major reasons for wishing to know the identity of protein substrates for PCM. First, the proteins that are methylated by PCM in the living cell, most of which have not yet been identified, are facets in the age-dependent metabolism of cells. Second, the fact that PCM can methylate many proteins in vitro, including products of overexpression systems, can be taken as evidence of spontaneous damage that has occurred in these proteins since the time of their translation. The biggest hurdle in identification of substrates for PCM arises from the extreme base-lability of the incorporated methyl esters, which typically hydrolyze in a few hours or less at neutral pH. Thus, many standard biochemical techniques for separating and characterizing proteins are not usefully applied to the identification of these methylated proteins. In particular, the electrophoresis of proteins by the most commonly employed techniques of sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) results in a complete loss of methyl esters incorporated by PCM, owing to the alkaline pH of the buffers employed. Consequently, a series of systems employing polyacrylamide gel From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Schematic of acidic discontinuous gel system. The system employs a pH 1.4 stacking gel on top of a pH 2.4 resolving gel with chloride as the leading ion and acetate as the trailing ion to stack the proteins tightly. The presence of the anionic detergent SDS allows the separation of proteins on the basis of molecular weight. The low pH preserves labile protein methyl esters, and so allows the identification of age-altered substrates of PCM.

electrophoresis at acidic pH have been utilized in efforts to identify the substrates of PCM. A pH 2.4 SDS system (3) using a continuous sodium phosphate buffering system has received the most attention (1,3–14). The main drawback of this system is that it produces broad electrophoretic bands. Acidic discontinuous gel systems using cationic detergents, (15), have proven useful in certain situations (16–21) and can be recommended if the cationic detergent is compatible with other procedures that might be utilized by the investigator (e.g., immunoblotting, protein sequencing). Recently, we have developed an electrophoresis system that employs SDS and an acidic discontinuous buffering system (Fig. 1). This procedure results in sharp electrophoretic bands and would be a good choice for investigators wishing to adhere to SDS as the anionic detergent. This system is described below, and examples of its ability to resolve proteins are provided. 2. Materials 2.1. Equipment 1. Slab gel electrophoresis unit: We have used the mini-gel electrophoresis units from Idea Scientific (Minneapolis, MN) (10 × 10 × 0.1 cm) with great success, as well as the Sturdier large-format gel system from Hoeffer Scientific Instruments (San Francisco, CA) (16 × 18 × 0.15 cm). 2. Electrophoresis power supply, constant current.

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3. X-ray film and photo darkroom. 4. Scintillation counter.

2.2. Reagents 1. 40% (w/v) Acrylamide stock solution containing 37:1 ratio of acrylamide to N,N′ methylene-bis-acrylamide (Bio-Rad, Richmond, CA). 2. Resolving gel buffer: 0.1 M NaH2PO4 (Sigma, St. Louis, MO), 2.0 % SDS (United States Biochemical, Cleveland, OH [USB], ultrapure), 6 M urea (ultrapure) pH 2.4, with HCl. 3. Modified Clark and Lubs buffer (C & L buffer): Add 25.0 mL of 0.2 M NaCl to 26 mL of 0.2 M HCl, and bring to a final volume of 0.1 L. The buffer pH should be ~1.4 (22) (see Note 1). 4. Stacking gel buffer (2X): 2.0% (w/v) SDS, 6.0 M urea and C & L buffer such that the C & L buffer makes up 66% (v/v) of the total volume with the remaining 34% consisting of water and other buffer components. Buffer will be 0.033 M in NaCl. Readjust pH to 1.4 with HCl. 5. Sample solubilization buffer (2X): 2.0% (w/v) SDS, 6.0 M urea, 10% glycerol ([USB], ultrapure), 0.01% pyronin Y dye (Sigma), and C & L 33% (v/v) of the total volume, with the remaining 67% consisting of water and other buffer components. Buffer will be 0.0165 M in NaCl. Readjust pH to 1.4 with HCl. 6. Electrode buffer (1X): 0.03 M NaH2PO4, 0.1% SDS, 0.2 M acetate, pH 2.4 with HCl. 7. Gel polymerization catalysts: 0.06% FeSO4, 1.0% H2O2, 1.0% ascorbic acid, prepared fresh in separate containers. 8. Colloidal Coomassie G-250 protein stain stock solution: 125 g ammonium sulfate, 25 mL 86% phosphoric acid, 1.25 g Coomassie brilliant blue G-250 (Sigma), deionized water to 1.0 L. The dye will precipitate, so shake well immediately before use. Stable indefinitely at room temperature (11). 9. Destain solution: 10% (v/v) acetic acid. 10. Fluorography solution: 1.0 M sodium salicylate brought to pH 6.0 with acetic acid (23). 11. X-ray film: Kodak X-Omat AR or equivalent.

2.3. Gel Recipes 2.3.1. 12% Acrylamide Resolving Gel, pH 2.4 The following volumes are sufficient to prepare one 7.5 × 10.5 × 0.15 cm slab gel: 2X Resolving gel buffer 40% Acrylamide (37:1) 0.06% FeSO4 1.0% Ascorbic acid 0.3% H2O2

3.75 mL 2.25 mL 0.06 mL 0.06 mL 0.06 mL

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2.3.2. 4.0% Stacking Gel, pH 1.4 The following volumes are sufficient to prepare one 3.0 × 10.5 × 0.15 cm stacking gel: 2X Stacking gel buffer 40% Acrylamide (37:1) 0.06% FeSO4 1.0% Ascorbic acid 0.3% H2O2

3.75 mL 0.75 mL 0.06 mL 0.06 mL 0.06 mL

3. Methods 3.1. Sample Solubilization 1. An equal volume of 2X solubilization buffer is added and the samples are heated in a 95°C heating block for no more than 30 s (see Note 2). 2. To separate samples under reducing conditions, it is critical to add any reducing agent before addition of the solubilization buffer. Up to 10 µg/gel protein band can be resolved with this system; total sample loaded in a single well should not exceed about 100 µg.

3.2. Gel Preparation 3.2.1. Resolving Gel 1. Add all the components listed under Subheading 2.3.1, except the 0.3% H2O2, together in a small Erlenmeyer side-arm flask. 2. Degas the solution for at least 5.0 min using an in-house vacuum. 3. Assemble together gel plates and spacers that have been scrupulously cleaned and dried. 4. Using a pen, make a mark 3.0 cm from the top of the gel plates to denote the space left for the stacking gel. 5. To the degassed gel solution, add the H2O2 catalyst. Gently mix solution by pipeting the solution in and out several times. Avoid introducing air bubbles into the solution. 6. Quickly pipet the acrylamide solution between the glass gel plates to the mark denoting 3.0 cm from the top of the gel plates. 7. Carefully overlay the acrylamide solution with about a 2.0-mm layer of watersaturated butanol using a Pasteur pipet, so that the interface will be flat on polymerization. 8. Allow the gel to polymerize at room temperature until a distinct gel–butanol interface is visible. 9. After polymerization is complete, pour off the overlay, and gently rinse the top of the gel with deionized water. Invert the gel on paper towels to blot away any remaining water between the gel plates.

3.2.2. Stacking Gel 1. Add all the components listed under Subheading 2.3.2, except the 0.3% H2O2, together in a small Erlenmeyer sidearm flask.

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2. Degas the solution for at least 5.0 min using in-house vacuum. 3. Soak the well-forming combs in the H2O2 catalyst solution while preparing the stacking gel. 4. To the degassed gel solution, add the H2O2 catalyst. Gently mix solution by pipeting the solution in and out several times. Avoid introducing air bubbles into the solution. 5. Quickly pipet the acrylamide solution between the glass gel plates to the very top of the gel plates. 6. Remove the combs from the H2O2 catalyst solution, shake off some of the excess solution, and insert the comb in between the gel plates by angling the comb with one hand and guiding the comb between the plates with the other hand. 7. Ensure the comb is level relative to the top of the resolving gel and that no bubbles are trapped under the comb. 8. After the stacking gel has completely polymerized, carefully remove the comb. Remove any unpolymerized acrylamide from each well by rinsing with deionized water (see Note 3).

3.3. Electrophoresis 1. Assemble the gel in the electrophoresis unit. 2. Add sufficient electrode buffer to cover the electrodes in both the upper and lower reservoirs. 3. Remove any air bubbles trapped under the bottom edge of the plates with a bent 25-gage needle and syringe containing electrode buffer. 4. Rinse each well with electrode buffer immediately before adding samples. 5. Load all wells with 40–60 µL of protein samples; load 1X solubilization buffer into any empty wells. 6. Run the gels at 15 mA, constant current, at room temperature for 4–6 h, or until proteins of interest have been adequately resolved (see Note 4). 7. Fix the gel in 12% (v/v) TCA with gentle shaking for 30 min. 8. Pour off fixative, and mix 1 part methanol with 4 parts of colloidal Coomassie G-250 stock solution. Slowly shake gel with staining solution for ~12 h (see Note 5). 9. Pour off staining solution. Any nonspecific background staining of the gel can be removed by soaking the gel in 10% (v/v) acetic acid. Little destaining of protein bands occurs even after prolonged times in 10% acetic acid.

3.4. Detection of Radioactively Methylated Proteins Several protocols exist for radioactively methylating proteins with PCM (2,16). Following electrophoresis, gel bands containing radioactively labeled proteins can be detected by fluorography or scintillation counting of gel slices. 3.4.1. Fluorography of Gels 1. If gels have been stained, they are destained using 10% methanol/7% acetic acid to decolorize the gel bands. 2. Expose the gel to fluorography solution for 30 min at room temperature with gentle shaking.

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Fig. 2. Comparison of discontinuous acid gel with continuous acid gel. The following experiment tested for the presence of age-altered proteins in a commercial preparation of collagenase. The collagenase preparation (Sigma, type IV) was methylated with purified rabbit erythrocyte PCM and 3H-AdoMet. Aliquots from the same methylation reaction were then resolved on (top) 12% discontinuous acid gel, described in text, or (bottom) 12% continuous acid gel system prepared according to the method of Fairbanks and Auruch (3). Lane 1: Rainbow molwt markers (Amersham); Lane 2: 18 µg of methylated collagenase were loaded on each gel. Following electrophoresis and staining, 0.5-cm gel slices were treated with base to detect radioactivity as described under Subheading 3. Both gel systems are capable of preventing the loss of methyl esters from protein samples, but the discontinuous system provides much higher resolution of individual polypeptide bands.

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3. The gels are then placed on a piece of filter paper and dried under vacuum without heat for 3 h. 4. In a dark room, X-ray film (Kodak X-Omat AR) is preflashed twice at a distance of 15 cm with a camera flash unit fitted with white filter paper (3M) to act as a diffuser. 5. The gel is placed in direct contact with the film and taped in place. For future alignment, puncture holes in an asymmetric pattern in a noncrucial area of the gelfilm sandwich. A 25-gage needle is useful for this purpose. 6. After sealing in a film cassette, the cassette is wrapped with aluminum foil, and exposure takes place at −70°C for several weeks. 7. After exposure, remove the cassette from −70°C, and allow to warm to room temperature. Develop film in darkroom. 8. An example of this technique is shown in Fig. 2.

3.4.2. Scintillation Counting Radioactive Methanol Evolved by Base Hydrolysis of Protein Methyl Esters in Gel Slices 1. Stained gels are soaked in 10% acetic acid containing 3% (v/v) glycerol for 1.0 h, placed on a piece of filter paper, and dried under vacuum without heat for 3.0 h. The glycerol keeps the gel from cracking and keeps it pliable for the steps described below. 2. Using a ruler and fine-tip marking pen, a grid is drawn directly on the surface of the dried gel. 3. A sharp scalpel is then used to cut out uniform slices precisely from each gel lane. Alternatively, selected bands can be individually excised from the gel. 4. 4.0 mL of scintillation fluid are added to each 20-mL scintillation vial. A glass 1-dram vial is then placed inside the scintillation vial, carefully avoiding spilling scintillation fluid into the 1-dram vial. 5. Each dried gel slice is then placed in the inner dram vial of a scintillation vial. 6. After all the gel slices have been placed in a separate inner 1-dram vial, 0.3 mL of 0.2 M sodium hydroxide is added to each inner vial. The scintillation vial is then immediately tightly capped and allowed to sit undisturbed for at least 3 h. Several hours are required for any volatile methanol that has formed by methyl ester hydrolysis to equilibrate and partition into the organic scintillation fluid, where it can then be detected by the scintillation counter. 7. Controls for measuring the efficiency of equilibration are performed by using a 14C methanol standard, which is added to an inner 1-dram vial containing a nonradioactive gel slice and base hydrolysis solution. This is then placed inside a scintillation vial and allowed to equilibrate along with the other samples. An equal aliquot of the 14C methanol standard is mixed directly with the scintillation fluid. 8. Figures 3 and 4 show examples of gels that have been sliced and counted in a scintillation counter under the conditions just described.

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Fig. 3. Coomassie staining and autoradiography of complex protein mixtures by acidic discontinuous SDS gel electrophoresis. The following experiment was performed to measure the varieties of age-altered proteins in a cell cytoplasm. (Left Panel) Coomasie G-250 stained acidic discontinuous gel. (Right Panel) Autoradiogram of same gel. Lane 1: Cytoplasmic proteins, following incubation cells with 3H-S-adenosyl-methione (AdoMet). Lane 2: Cytoplasmic proteins, following incubation of PC12 cytoplasm with 3H-AdoMet. Lane 3: Cytoplasmic proteins, following incubation of intact PC12 cells with 3H-AdoMet and purified rabbit PCM. Lane 4: Cytoplasmic proteins, following incubation of lysed PC12 cells with 3H-AdoMet and purified rabbit PCM. Lane 5: Positive control; 20 µg of ovalbumin methylated with 3H-AdoMet and purified rabbit PCM. Cells, lysates, and subfraction incubated with IU PCM, 300 pmol Adomet in final volume of 50 µL 0.2 M citrate, pH 6.0, 20 min at 37°C.

4. Notes 1. The buffering system employed in the stacking gel and sample solubilization buffer is based on a modification of the C & L buffering system. NaCl is employed rather than KC1 of the original system, because K+ ions cause SDS to precipitate out of solution. 2. On occasion, the solubilization buffer will contain precipitates. These can be brought back into solution by briefly heating the buffer at 37°C. The solubilization buffer is stable for at least 2 wk at room temperature; by storing the buffer in small aliqouts at −20°C, it is stable indefinitely. 3. Gels can be stored for up to two weeks at 4°C by wrapping them in damp paper towels and sealing tightly with plastic wrap. 4. Since the dye front is a poor indicator of protein migration, use of prestained mol-wt markers, such as the colored Rainbow markers from Amersham, allows the progress of protein separation to be monitored by simply identifying the colored bands, which are coded according to molecular weight. 5. It is essential that all fixing and staining steps occur at acidic pH. The colloidal Coomassie G-250 procedure described under Subheading 3. has

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Fig. 4. Coomassie staining and radioactive methyl ester determination in gel slices of electrophoresed proteins from diseased human brain tissue. Extracts prepared from homogenates of Alzheimer’s diseased brain (obtained from the Department of Pathology, Oregon Health Sciences University) were methylated in vitro with purified rabbit erythrocyte PCM and 3H-AdoMet. Methylated proteins were then separated on 12% discontinuous, acidic gels, and radioactivity in each gel slice was quantified with scintillation counting as described under Subheading 3. Top: Distribution of methyl acceptor proteins in tissue protein that was insoluble in the nonionic detergent Triton X-100. Middle: Distribution of methyl acceptor proteins in tissue proteins that was soluble in an aqueous homogenization buffer. Bottom: Distribution of methyl acceptor proteins in crude homogenates of Alzheimer’s diseased brains. Incubation conditions are similar to those described in Fig. 3.

several advantages: it is acidic, simple to perform, and has higher sensitivity than other dye-based staining methods, including those using Coomassie R-250. Additionally, nonspecific background staining is very low, so only minimal destaining is necessary to visualize protein bands.

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6. Avoid using higher glycerol concentrations or prolonged incubation of the gel in this solution. Otherwise, the gel will be sticky after drying and contract sharply away from the paper backing on cutting. References 1. Aswad, D. W. and Deight, E. A. (1983) Endogenous substrates for protein carboxyl methyltransferase in cytosolic fractions of bovine brain. J. Neurochem. 31, 1702–1709. 2. Lou, L. L. and Clarke, S. (1987) Enzymatic methylation of band 3 anion transporter in intact human erythrocytes. Biochemistry 26, 52–59. 3. Fairbanks, G. and Avruch J. (1973) Four gel systems for electrophoretic fractionation of membrane proteins using ionic detergents. J. Supramol. Struct. 1, 66–75. 4. Barber, J. R. and Clarke, S. (1984) Inhibition of protein carboxyl methylation by S-adenosyl-l-homocysteine in intact erythrocytes. J. Biol. Chem. 259(11), 7115–7122. 5. Bower, V. E. and Bates, R. G. (1955) pH Values of the Clark and Lubs buffer solutions at 25°C. J. Res. Natl. Bureau Stand. 55(4), 197–200. 6. Gingras, D., Menard, P., and Beliveau, R. (1991) Protein carboxyl methylation in kidney brush-border membranes. Biochim. Biophys. Acta. 1066, 261–267. 7. Johnson, B. A., Najbauer, J., and Aswad, D. W. (1993) Accumulation of substrates for protein l-isoaspartyl methyltransferase in adenosine dialdehyde-treated PC12 cells. J. Biol. Chem. 268(9), 6174–6181. 8. Johnson, B. A., Freitag, N. E., and Aswad, D. W. (1985) Protein carboxyl methyltransferase selectively modifies an atypical form of calmodulin. J. Biol. Chem. 260(20), 10,913–10,916. 9. Lowenson, J. D. and Clarke, S. (1995) Recognition of isomerized and racemized aspartyl residues in peptides by the protein L-isoaspartate (d-aspartate) O-methyltransferase, in Deamidation and Isoaspartate Formation in Peptides and Proteins. (Aswad, D. W., ed.), CRC, Boca Raton, pp. 47–64. 10. McFadden, P. N., Horwitz, J., and Clarke, S. (1983) Protein carboxyl methytransferase from cow eye lens. Biochem. Biophys. Res. Comm. 113(2), 418–424. 11. Neuhoff, V, Stamm, R., Pardowitz, I., Arold, N., Ehrhardt, W., and Taube, D. (1988) Essential problems in quantification of proteins following colloidal staining with Coomassie brilliant blue dyes in polyacrylamide gels, and their solutions. Electrophoresis 9, 255–262. 12. O’Conner, C. M. and Clarke, S. (1985) Analysis of erythrocyte protein methyl esters by two-dimensional gel electrophoresis under acidic separating conditions. Analyt. Biochem. 148, 79–86. 13. O’Conner, C. M. and Clarke, S. (1984) Carboxyl methylation of cytosolic proteins in intact human erythrocytes. J. Biol. Chem. 259(4), 2570–2578. 14. Sellinger, O. Z. and Wolfson, M. F. (1991) Carboxyl methylation affects the proteolysis of myelin basic protein by staphylococcus aureus V8 proteinase. Biochim. Biophys. Acta. 1080, 110–118. 15. MacFarlane, D. E. (1984) Inhibitors of cyclic nucleotides phosphodiesterases inhibit protein carboxyl methylation in intact blood platelets. J. Biol. Chem. 259(2), 1357–1362.

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16. Aswad, D. W. (1995) Methods for analysis of deamidation and isoaspartate formation in peptides, in Deamidation and Isoaspartate Formation in Peptides and Proteins (Aswad, D. W., ed.), CRC, Boca Raton, pp. 7–30. 17. Freitag, C. and Clarke, S. (1981) Reversible methylation of cytoskeletal and membrane proteins in intact human erythrocytes. J. Biol. Chem. 256(12), 6102–6108. 18. Gingras, D., Boivin, D., and Beliveau, R. (1994) Asymmetrical distribution of L-isoaspartyl protein carboxyl methyltransferases in the plasma membranes of rat kidney cortex. Biochem. J. 297, 145–150. 19. O’Conner, C. M., Aswad, D. W., and Clarke, S. (1984) Mammalian brain and erythrocyte carboxyl methyltranserases are similar enzymes that recognize both d-aspartyl and l-iso-aspartyl residues in structurally altered protein substrates. Proc. Natl. Acad. Sci. USA 81, 7757–7761. 20. O’Conner, C. M. and Clarke, S. (1983) Methylation of erythrocyte membrane proteins at extracellular and intracellular d-aspartyl sites in vitro. J. Biol. Chem. 258(13), 8485–8492. 21. Ohta, K., Seo, N., Yoshida, T., Hiraga, K., and Tuboi, S. (1987) Tubulin and high molecular weight microtubule-associated proteins as endogenous substrates for protein carboxyl methyltransferase in brain. Biochemie 69, 1227–1234. 22. Barber, J. R. and Clarke, S. (1983) Membrane protein carboxyl methylation increase with human erythrocyte age. J. Biol. Chem. 258(2), 1189–1196. 23. Chamberlain, J. P. (1979) Fluorographic detection of radioactivity in polyacrylamide gels with the water soluble fluor, sodium salicylate. Analyt. Biochem. 98, 132.

165 Analysis of Tyrosine-O-Sulfation Jens R. Bundgaard, Jette W. Sen, Anders H. Johnsen, and Jens F. Rehfeld 1. Introduction Tyrosine O-sulfation was first described about 50 years ago as a posttranslational modification of fibrinogen (1). In the following thirty years it was considered to be a rare modification affecting only a few proteins and peptides. However, in the beginning of the 1980s tyrosine (Tyr) sulfation was shown to be a common modification and since then an increasing number of proteins have been identified as sulfated. The target proteins belong to the classes of secretory, plasma membrane, and lysosomal proteins, which reflects the intracellular localization of the enzymes catalyzing Tyr sulfation, the tyrosylprotein sulfotransferases (TPSTs). TPSTs are type II integral membrane glycoproteins that reside in the trans Golgi network and use adenosine 3′-phosphate 5′-phosphosulfate (PAPS) as a co-substrate and sulfate donor. In higher organisms, 2 enzymes have been identified and cloned and they appear to be ubiquitously expressed in tissues (2–4). Moreover, homologous genes are present in lower eukaryotes as well as in the nematode Caenorhabditis elegans, fruit fly, and zebrafish. Both TPST-1 and TPST-2 knock-out mice models have been established and both display distinct phenotypes. The TPST-1 knock out mice are slightly smaller than wild-type mice and the female produces smaller litters (5). The TPST-2 knockout mouse has delayed growth but reaches normal weight at 10 wk. Strikingly, the TPST-2 knock-out male is infertile (6). The biological function of Tyr sulfation is modulation of protein-protein interactions. For instance, Tyr sulfation is necessary for binding of the peptide hormone cholecystokinin (CCK) to the CCK-1 receptor (7); it is critical for interactions between p-selectin and PSGL-1 (P-Selectin Glycoprotein Ligand-l) (8); between hirudin and thrombin (9); and it modulates the ability of HIV −1 virus to enter cells via the chemokin receptor CCR5 and CD4 (10). In addition, sulfation has been reported to modulate From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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proteolytic processing (11) and secretion rates of secretory proteins (12). From these functions, it follows that it is not only important to detect Tyr sulfation, but also to identify the sulfation site to establish effects on the activity on the protein. Attempts have been made to determine a consensus site for tyrosine sulfation and generate algorithms that predict sulfation sites (13–15), one of these is the “sulfinator,” which is available at www.expasy.org. However, it is known that the sulfinator at best gives an indication of possible sulfation sites, and it has been shown that nontypical structures, which are not identified by the sulfinator, can be sulfated (16,17). Traditionally, Tyr sulfation has been analyzed by incorporation of radio labelled sulfate into target cells followed by purification of the target protein. Subsequently, the protein is degraded enzymatically or by alkaline hydrolysis followed by thinlayer electrophoresis to demonstrate the presence of radioactively labeled tyrosine. These techniques have been described in detail previously (18). The aim of this chapter is to present alternative analytical methods of Tyr sulfation other than radioisotope incorporation before analysis. Thus, sulfation of protein or peptide fragments can be demonstrated by a combination of chromatography and specific assays. This has several advantages. First, depending on the assay used, the analysis can be quantitative. Second, problems with incorporation of isotope and the high degree of incorporation of radiolabeled sulfate into carbohydrate moieties that might interfere with detection can be avoided. In addition, it allows two ways of identification of sulfated tyrosines depending on purity of the protein. One strategy demands purified protein using mass spectrometry or chromatography, whereas another uses Tyr sulfate sensitive antisera to distinguish between sulfated and nonsulfated forms in a crude extract. We provide protocols for establishment of sulfation specific antisera and give examples of chromatographic systems that are useful for analysis of Tyr sulfation. Establishment of a specific immunoassay is costly and time-consuming, and initiation of such a process demands an interest exceeding the mere confirmation of the modification. If such an assay is desirable, a number of considerations have to be taken into account. The size of an antibody binding site is complementary to that of a peptide ligand (or epitope) of 4–6 amino acid residues (19). Within such an epitope the exact structure of each residue profoundly influences the binding of the antibody (20). Moreover, the protein or peptide structures surrounding the particular epitope also modulate binding affinity considerably (21). Accordingly, it is not surprising that attempts to raise antibodies that specifically bind a single O-sulfated Tyr residue as such— irrespective of neighboring structures—have failed (Hakanson, R. and Huttner, W. B. personal communications, and our own results). However, recently a promising monoclonal antibody was reported, which was raised against tyrosine sulfated P-selectin glycoprotein ligand-1. It has tyrosine sulfate specificity and show a wide tolerance to sequence variance around the sulfated tyrosine.

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Nevertheless, it does not bind sulfotyrosine alone and is thus not entirely sequence independent (22). The discrepancy with the success to raise antibodies against the structurally similar phosphorylated Tyr residues (23) is at present unexplained. In accordance with the aforementioned consideration and with prevailing immunochemical theories, antibodies that recognize sulfation of a given Tyr residue consequently have to be raised against the entire epitope containing the sulfated tyrosine, i.e., a peptide sequence of approx 5 or 6 amino acid residues. It has, however, proven difficult to raise high-titer and high-affinity antibodies or antisera against peptides of less than 10 residues (24). Therefore, a peptide of approx 10 residues containing the desired epitope in an otherwise expedient sequence should be designed, synthesized, and used for immunization. Hence, measurement of Tyr sulfation by immunochemical methods requires precise knowledge of the sequence and structure surrounding the O-sulfated residue in a peptide or protein. It is necessary to consider different strategies in the design of appropriate radioimmunoassays (RIA) (see Note 1). Sulfation of a protein or peptide introduces a highly acidic group that alters its local physical and chemical properties such as conformation, hydrophilicity, and pK-value. This can be used to separate sulfated and nonsulfated forms, by various chromatographic techniques, e.g., ion-exchange chromatography and reverse phase high-performance liquid chromatography (RP-HPLC) at near neutral pH under which the conditions for ionization of the sulfate group induces the maximum difference between the two forms (see Note 2). Also, removal of the sulfate group by digestion with arylsulfatase combined with chromatographic analysis of the peptide before and after the digestion can reveal the presence of Tyr sulfate. Often, identification of sulfated Tyr within a protein requires digestion of the protein into smaller peptides followed by separation of the fragments and isolation of relevant peptide. A number of proteinases may be used, for example trypsin or the endoproteinases Lys-C, Glu-C, and Asp- N, the choice being partly dependent on the individual protein. The details for these techniques are beyond the scope of the present chapter, but may be found in other volumes of this series. These analytical methods can profit from the availability of specific antisera or be carried out using pure proteins. Having a purified protein offers alternative ways of analysis of Tyr sulfation. The sulfate ester bond is generally considered as labile and particularly susceptible to acidic hydrolysis. Despite of this, the tyrosine sulfate can generally withstand pH in the range of 1–3, often used during many standard analyses such as reversed phase chromatography and mass spectrometry, as long as the temperature is kept at room temperature or below and the exposure time is limited (25). However, sulfated Tyr can not be identified by protein sequence analysis because of the highly acidic environment and elevated temperatures during the analysis. Instead, nonderivatized Tyr will result. Likewise, the standard acidic

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hydrolysis used for amino acid analysis is not feasible; instead, demonstration of the presence of intact tyrosin sulfate in a peptide was originally obtained by basic hydrolysis before amino acid analysis (26). This technique has a limited sensitivity because of an inevitable background. Alternatively, nondestructive digestion can be obtained by aminopeptidase M or a combination of aminopeptidase M and prolidase followed by amino acid analysis (27). However, these techniques are not used much anymore. Mass spectrometry (MS) is now the preferred tool of analysis of many post-translational modifications. MS in combination with protein sequence analysis can disclose the presence of a sulfated Tyr. Thus, if the identified sequence contains Tyr and the measured mass is 80 Da higher than the calculated value, it is a good indication that the Tyr is sulfated. However, phosphorylation also results in an 80 Da increase of the molecular mass (see Note 3; Fig. 1). It should also be noted that the sulfate group is lost during ionization in both matrix assisted laser desorption/ionisation (MALDI) and to some degree in electrospray ionization (ESI), resulting in a false negative result. Using MALDI time-of-flight (TOF) MS, the loss of sulfate can be turned into a diagnostic advantage because the loss is often complete in the positive mode (31) and limited in negative mode. Hence, comparison of spectra obtained in positive and negative linear mode of the same sample will show a vast difference in the yield of the sulfated and nonsulfated species (see Note 4), whereas the picture will be more complex in the reflector mode (see Note 5; Fig. 2). In experiments using hybrid instruments of the quadrupole-TOF type it is possible to see fractions of the sulfated species in positive mode even though the desulfated species is generally dominating. The fact that tyrosine sulfate is unstable even at standard collision energy in a quadrupole-TOF instrument, can be used in identifying new sulfated targets by using neutral loss scan experiment detecting the loss of 79.957 (32). Though tyrosine sulfate and tyrosine phosphate has almost identical mass, it is possible to distinguish between the two as the tyrosine sulfate is generally much more unstable than the tyrosine phosphate, which is not nearly as prone to neutral loss as the tyrosine sulfate. Furthermore when performing MS/MS experiments the tyrosine phosphate gives rise to a stable immonium ion of 216.043 whereas the immonium ion of tyrosine sulfate is not detected. This can be used to further confirm the identification of a phosphate or sulfate. Another option is to investigate tyrosine sulfation by a mass spectrometer capable of electron capture dissociation (ECD). This is a fragmentation method that, to a higher degree, maintains the sulfate during peptide backbone fragmentation but requires specialized and expensive instrumentation (33,34). Recently, a new method for stoichiometric identification of tyrosine sulfate in proteins using tandem mass spectrometry was described (35). The method utilize sulfosuccinimidyl acetate to block unsulfated tyrosines, followed by analysis under conditions that hyrolyze tyrosine sulfate, leaving unmodified

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Fig. 1. Mass spectra obtained in the positive and negative mode in a mass spectrometer equipped with delayed extraction (Bruker Daltonik Biflex). As model peptides were used nonsulfated and sulfated shark gastrin-8 (Asp-Tyr(SO3)-Thr-Gly-Trp-Met-Asp-Phe-NH2) with the theoretical monoisotopic molecular masses 1032.40 and 1112.36 Da, respectively. In the positive mode the measured m/z values represent the protonated molecular ion (+1.01) and to a certain degree the Na+ and K+ adducts, whereas the deprotonated form (−1.01) is recorded in the negative mode. In this example the intact sulfated peptide is only observed in the negative mode together with a small fraction of the desulfated form. See also, however, Note 3. y-axis: number of collected ions.

tyrosines as indicators of a previously sulfated site. This method is generally applicable and sensitive, but like other present methods it is not well suited for general screening of sulfated proteins. 2. Materials 2.1. Generation of Tyr Sulfation Sensitive Radioimmunoassays 2.1.1. Preparation of immunogen (common for all Three Coupling Methods) (see Note 6) 1. 5 mg sulfated peptide. 2. N, N-dimethylformamide. 3. 25 mg bovine serum albumin.

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Fig. 2. Mass spectrum of sulfated shark gastrin-8 (see Fig. 1) obtained in the negative reflected mode using α-cyano-4-hydroxycinnamic acid as matrix. The peak at m/z 1111.29 represents the intact peptide (theoretical value 1112.36 −1.01) whereas the peak at 1040.88 represents the in-flight desulfated peptide. The difference between the two is considerably less than the expected 80 (see Note 5). Note that the resolution is higher in the reflected mode than in the linear mode (Fig. 1).

4. 0.05 M sodium phosphate buffer, pH 7.5. For carbodiimide-coupling: 5. 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide HCl. For glutaraldehyde-coupling: 6. 500 g/L g lutaraldehyde. 7. One Sephadex G-10 column with a fraction collector. For maleimidobenzoyl-succinimede ester-coupling: 8. 25 mg m-maleimidobenzoyl-N-hydroxysuccinimide ester (MBS). 9. One Sephadex G-25 column with a fraction collector.

2.1.2. Immunization 1. 2. 3. 4.

6–8 rabbits. 8.5 g/L saline buffer. Freund’s complete adjuvant. Freund’s incomplete adjuvant.

2.1.3. Preparation of Tracer (see Note 7; Fig. 3) 1. Synthetic peptide 2. HPLC apparatus with reverse-phase column (Aquapore C-8 column, RP-300, 220 × 4.6 mm, 7- µm bead size) and fraction collector.

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Fig. 3. The structure of immunogens used for production of antibodies directed against tyrosyl O-sulfated peptides (upper part of the figure), and the structure of corresponding radioactive tracers (lower part of the figure).

3. Ethanol or acetonitrile (HPLC grade). 4. 1.0 mL/L trifluoroacetic acid (TFA; HPLC grade in water). 5. 0.02 M Barbital buffer, pH 8.4. Iodination using chloramin T:

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I125 (Amersham IMS 30, specific activity 16.85 mCi/µg of iodine). 0.05M phosphate buffer, pH 7.5. Disodium sulfite (Na2SO3). Chloramine T. Iodination using Bolton Hunter reagent: 10. Bolton and Hunter reagent for protein iodination (Amersham). 11. 0.2 M sodium borate buffer, pH 8.5: mixture of 0.2 M Na2B4O7 and 0.2 M H3BO3 with pH adjusted. 12. 0.1 M sodium borate buffer with 0.2 M glycine, pH 8.5 (prepared from the buffer above). 6. 7. 8. 9.

2.1.4. RIA Procedure 1. 2. 3. 4. 5.

Synthetic peptide for standard (see Note 8). Disposable plastic tubes. 0.02 M barbital buffer, pH 8.4, containing 1 g/L bovine serum albumin. Activated charcoal. Blood plasma: this is a mixture of buffer and outdated human plasma from Blood Banks in 0.02 M sodium phosphate, pH 7.4. The optimal concentration normally varies between 10–50% plasma. 6. γ-scintillation counter.

2.2. Analysis of Tyr Sulfation Using FPLC Based Ion-Exchange Chromatography (see Note 9) 1. An FPLC system equipped with an ion-exchange column, e.g., an 5/5HR MonoQ anion-exchange column (Pharmacia) and a fraction collector. 2. Buffer A: 50 mM Tris-HCl, pH 8.2 added 10% (v/v) acetonitrile. 3. Buffer B: 50 mM Tris-HCl, pH 8.2 added 10% (v/v) acetonitrile and 1 M NaCl.

Both buffers are filtered through a 0.45 µm filter, degassed and stored at 4°C where it is stable for months. 2.3. Analysis of Tyr Sulfation Using Reverse Phase HPLC 1. An HPLC instrument with gradient formation capability and equipped with a suitable RP (reversed phase) column (C8 or C18). 2. Solvents: Stock solution of 0.5 M ammonium acetate in water, filtered through 0.45 µm filter (can be kept for several months at 4°C). Solvents A and B are prepared by addition of 20 mL/L of H2O and acetonitrile, respectively (both HPLC grade), to form 10 M ammonium acetate.

2.4. Analysis of Tyr Sulfation Using Mass Spectrometry 1. Instrument: A mass spectrometer for MALDI-TOF MS. 2. Matrix solution: Prepare a 20 mg/mL solution of α-cyano-4-hydroxycinnamic acid in 30% (v/v) acetonitrile/0.1% (v/v) TFA in an Eppendorf tube and centrifuge; use the supernatant as matrix solution. Alternatively 2,5-dihydroxybenzoic acid (DHB)

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20 mg/mL in 50% Acetonitrile/1% phosphoric acid can be used (see Note 10). For larger peptides 3,5-dimethoxy-4-hydroxy-trans-cinnamic acid (sinapinic acid) can be used. Premade matrix solutions (“MALDI-grade”) are also commercially available (Hewlett-Packard). 3. For neutral loss scanning on quadrupole-TOF instrument Instrument: A hybrid instrument of quadrupole time-TOF type. For complex samples connected to a HPLC instrument equipped with a reversed phased column for separation of peptides. Desalted sample dissolved in 30–50% Acetonitrile 0.2% formic acid for direct infusion, or sample dissolved in aqueous solution for injection on reversed phase column. 4. Solution A: 95% Acetonitrile/0.2% formic acid, solution B: 5% Acetonitrile/ 0.2% formic acid.

2.5. Arylsulfatase Treatment 1. Arylsulfatase type VIII (Sigma, 20–40 U/mg). 2. Acetate buffer: 0.2 M sodium acetate, pH 5.0. 3. 0.2% Sodium chloride.

3. Methods 3.1. Generation of Tyr Sulfation Sensitive Radioimmunoassays 3.1.1. Preparation of the Immunogen 3.1.1.1. CARBODIIMIDE COUPLING 1. Dissolve 5 mg peptide hapten in 1 mL of N,N-dimethylformamide. 2. Dissolve 25 mg bovine serum albumin in 2.5 mL of 0.05 M sodium phosphate, pH 7.5. 3. Conjugate by the addition of 125 mg of 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide HCl to give molar ratios between the peptide, albumin and ethylcarbodiimide of 1:0.1:40. Mix the reagents and incubate for 20 hours at 20°C. 4. Divide the mixture into six portions and store at −20°C until immunization (stable up to one year). 3.1.1.2. GLUTARALDEHYDE COUPLING 1. Dissolve 2.5 mg of sulfated peptide hapten together with 7.5 mg bovine serum albumin in 5 mL of 0.05 M sodium phosphate, pH 7.5. 2. Conjugate by dropwise addition of 100 µL of 500 g/L glutaraldehyde. 3. Mix the solution and incubate for 4 h at 20°C. 4. Apply the mixture to a calibrated Sephadex G-10 column and elute at 20°C with 0.05 M sodium phosphate, pH 7.5, in fractions of 1 mL. 5. The void volume fractions containing the conjugate are then pooled, divided into 6 portions, and stored at −20°C until immunization (stable for months). 3.1.1.3. MALEIMIDOBENZOYL-SUCCINIMIDE ESTER COUPLING 1. Dissolve 5 mg sulfated peptide hapten together with 25 mg of bovine serum albumin (BSA) in 10 mL 0.05 M sodium phosphate buffer (pH 7.5).

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2. Dissolve 25 mg of m-maleimidobenzoyl-N-hydroxysuccinimide ester (MBS) in 1 mL N, N-dimethylformamide. 3. Conjugate by dropwise addition of the MBS-solution to the sulfated peptide/BSA solution. 4. Stir at room temperature for 30 min. 5. Free MBS is separated from the activated albumin by Sephadex G-25 gel filtration (eluted at 20°C with 0.05 M sodium phosphate, pH 7.5, in fractions of 1 mL). 6. Add the sulfated and cysteinylated peptide to the MBS-activated bovine serum albumin and stir the mixture stirred for 3 h at room temperature and a pH of 7.5.

3.1.2. Immunizations 1. The first portion of the antigen is suspended in 8.5 g/L saline to a volume of 5 mL and carefully emulsified with an equal volume of Freund’s complete adjuvant. We recommend emulsification by pushing the saline-adjuvant forwards and backwards between two connected 10-mL syringes until a drop of the emulsion remains coherent and condensed on a larger water surface. Randomly bred rabbits are used for immunizations in series of 6–8 rabbits each. 2. Two subcutaneous injections of the mixture are given over the hips in amounts corresponding to 40 µg hapten per animal. 3. Five or more booster injections using Freund’s incomplete adjuvant (prepared as described in 1) are then administered simultaneously in all rabbits at 8-wk intervals, using one-half of the initial dose of antigen per immunization. 4. The rabbits are bled from an ear vein 10 d after immunization. Sera from the bleedings are separated and stored at −20°C. 5. The antiserum obtained is evaluated (see Note 11).

3.1.3. Preparation of Tracer for Radioimmunoassay (see Note 7) 3.1.3.1. IODINATION USING CHLORAMINE T 1. Dissolve 500 µg peptide in 1 mL 0.05 M phosphate buffer, pH 7.5. Aliquot in 10 µL portions and store at −20°C. 2. Dilute 2 mCi I125 (~20 µL) with 85 µL 0.05 M phosphate buffer, pH 7.5 (final activity about 200 µCi/10 µL). 3. Dissolve 5 mg chloramin T in 10 mL 0.05 M phosphate buffer pH 7.5. 4. Dissolve 2 mg disodium sulfite in 10 mL 0.05 M phosphate buffer pH 7.5. 5. Mix 10 µL peptide, 10 µL I125 and 10 µL chloramin T gently. After 30 s the reaction is arrested with 25 µL disodium sulfite. Shake gently. If more tracer is prepared, do iodination in batches and pool the preparations. 6. Purify the tracer by HPLC. Fractions with maximal counts of reactivity is diluted with 0.02 M barbital buffer, pH 8.4 to a final activity of 106 cpm/mL. Store the tracer in 500 µL aliquots at −20°C. 3.1.3.2. IODINATION USING THE BOLTON HUNTER REAGENT 1. Evaporate the water from the I125 Bolton Hunter reagent under a slow flow of nitrogen. 2. Dissolve 5 µg of peptide in 10 µL water and add 10 µL 0.2 M sodium borate buffer.

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3. Add the peptide solution to the Bolton Hunter reagent and incubate the mixture at 4°C for about 20 h. 4. Add 500 µL 0.1 M borate buffer with 0.2 M glycine. Continue directly with HPLC purification of the tracer as described above, or store at 4°C.

3.1.4. RIA Procedure 1. At room temperature, prepare the following mixtures of total volumes of 2.4 mL (RIA is carried out as an equilibrium system at pH 8.4, using 0.02 M barbital buffer, pH 8.4, containing 1 g/L bovine serum albumin) in disposable plastic tubes: 2.0 mL of antiserum dilution, 250 µL of tracer solution (giving 1000 cpm, corresponding to 0.5 fmol of freshly prepared 125I-labeled tracer-peptide), and 150 µL of standard solution or sample. All samples should be assayed in duplicate. 2. Incubate at 4°C for 2–5 days to reach equilibrium. 3. Antibody-bound (B) and free (F) tracers are separated by the addition of 0.5 mL of a suspension of 20 mg of activated charcoal and diluted blood plasma to each tube. The tubes are centrifuged for 10 min at 2000 g. 4. Count the activities of the supernatant (B) and sedimented charcoal (F) in automatic γ-scintillation counters for 5 min. 5. Calculate the binding percentage as B − [(B + F) × D] / (B + F) − [(B + F) × D] × 100, where the “damage” (D) is defined as B × 100 / (B + F) in the absence of antiserum. The damage is usually 2–3%. The peptide concentration is determined from a standard curve based on known standards.

3.2. Analysis of Tyr Sulfation Using FPLC Based Ion-Exchange Chromatography 1. Equilibrate the column to start conditions and inject sample. 2. Elute with an appropriate gradient at the flow of 1 mL/min and collect fractions if necessary. 3. The eluting peptide is monitored by the UV signal recording or any appropriate specific assay. An example of an elution profile examined by radioimmunoassay is given in Fig. 4.

3.3. Analysis of Tyr Sulfation Using Reverse Phase HPLC 1. Equilibrate column to start conditions and inject sample. 2. Elute with an appropriate gradient. 3. Record UV signal at 214 nm and—if further analyses are wanted—collect fractions automatically at regular intervals or collect peak manually. An example of an elution profile is given in Fig. 5.

3.4. Analysis of Tyr Sulfation Using Mass Spectrometry 3.4.1. MALDI-TOF 1. Samples should be in a final concentration of 0.05–5 pmol/µL, sensitivity depends highly on the nature of the peptide (see Note 12).

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Fig. 4. Anion exchange chromatography using a FPLC system of the main processing forms of human gastrin heterologously expressed in a hamster β-cell line, HIT. Panel A shows carboxyamidated gastrin forms of 17 and 34 residues and demonstrates the separation of sulfated and nonsulfated forms. Panel B shows the immediate precursor of carboxyamidated gastrin, with the free C-terminus extended with a Gly. Fractions were eluted using a linear gradient of 20–55% buffer B over 60 min at a flow of 1 ml/min. Fractions were analyzed using specific radioimmunoassays. (Elution positions can be verified using synthetic peptides or by analysis of elution positions following arylsulfatase treatment.)

2. Mix 0.5 µL of sample with 0.5 µL MALDI matrix solution. For manual preparation, this can be done in a microcentrifuge tube, or many samples can be prepared consecutively at different positions in a trough made from a cutthrough polypropylene tube. 3. Apply immediately 0.5 µL of the mixture to the sample target (volume may depend on the target size) and let sample dry. 4. Insert sample(s) into the mass spectrometer and record the mass spectrum according to the instrument manual.

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Fig. 5. HPLC separation of the model peptides, non-sulfated and sulfated shark gastrin-8 (see Fig. 1). The use of 0.1% trifluoroacetic acid (TFA) and 10 mM ammonium acetate (NH4Ac) as buffering component are compared. In both cases 0.5%/min. gradients from H2O to acetonitrile were employed, starting from 13% acetonitrile in the TFA system and from 8% in the NH4Ac system (indicated by the straight lines). Although the two forms are indeed separated in the TFA system (by 1% acetonitrile) the separation is much more pronounced in the NH4Ac system (by 2.5% acetonitrile). For bigger peptides, where sulfation influences the chromatographic behavior relatively less, use of the NH4Ac system may be crucial for the separation of the non-sulfated and sulfated form.

3.4.2. Neutral Loss Experiment on Quadrupole-TOF: Analyses of complex samples: 1. The LC part of the system is programmed to run a gradient appropriate to the sample, e.g., 5–40% B over 120 min. (see Note 13). 2. The instrument is programmed to do neutral loss scanning for loss of 79.9568 Da and with alternating slightly higher and slightly lower collision energy than standard settings during the run of the gradient (e.g., on a Q-TOF2 instrument (Waters) collision energies 13 and 8 as opposed to a standard setting of 10), see Fig. 6. 3. Peptides detected by neutral loss scanning are sequenced by MS/MS (see Note 14). 4. Tyrosine sulfated peptides are confirmed by the MS/MS data that should not contain the tyrosine phosphate immonium ion of 216.043 Da.

3.4.3. Direct Infusion of Simple Samples 1. For direct infusion, e.g., by nanospray without reversed phase separation the sample has to be clean from salts and only contain a limited number of peptides.

Fig. 6. Mass spectra of tyrosine sulfated drosulfokinin showing the increased neutral loss as the collision energy (CE) is increased in intervals of 2 from 7 to 13. The spectra were obtained on a Q-TOF2 using nanospray ionization. Sulfated drosulfokinin is seen at 755.3 Da and desulfated drosulfokinin at 715.3 Da. Both ions are doubly charged and the neutral loss is therefore observed as 40 Da.

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2. The experiment is run as described above but without the LC part. Alternatively collision energy can be changed manually and mass shift for the peptide suspected of being tyrosine sulfated monitored.

3.5. Arylsulfatase Treatment (see Note 15) 1. Prepare an enzyme solution of arylsulfatase VIII of 2 mg/mL in 0.2% sodium chloride. 2. Prepare the lyophilized sample by resuspension in 400 µL acetate buffer. 3. Add 100 µL enzyme solution to the sample preparation. The final concentration of arylsulfatase is 0.4 mg/mL (equivalent to 0.55 U/mL). 4. Incubate for 3 h at 37°C. 5. Terminate the reaction by boiling for 10 min.

4. Notes 1. The strategy depends on several factors: (1) The size of the protein or peptide; (2) the position of the sulfated Tyr within the protein; (3) the biology of the sulfated protein or peptide (a large membrane protein; a circulating hormone; or a small neurotransmitter peptide, etc.); and (4) the concentration range in which to measure. If the sulfated Tyr is positioned at the N- or the C-terminus, production of antibodies against a synthetic analogue of the terminally sulfated decapeptide is straightforward. If, however, the sulfated Tyr is located in the middle of a protein or a long polypeptide chain, antibodies should be raised against a fragment containing the sulfated tyrosine, preferably a fragment that can be released from the protein by appropriate proteolytic cleavage. Finally, if the sulfated protein is available in substantial quantities for large immunization series, “shotgun” immunization with the entire protein and subsequent identification of epitope specificity can be attempted. 2. The presence of Met in the peptide presents a possible pitfall, because oxidation of Met to the sulfoxide (which may occur spontaneously in the presence of atmospheric oxygen) increases the hydrophilicity considerably, thus causing the peptide to elute significantly earlier than the reduced form in both a RP-HPLC and in an anion exchange chromatography system. 3. Both sulfation and phosphorylation add 80 Da to the molecular mass of the peptide (the exact monoisotopic values being 79.957 and 79.966, respectively). However, it is possible to distinguish the two by several criteria. Thus, under the standard conditions used for Edman sequencing peptides phosphorylated at Ser, Thr or Tyr give no signal (although for different reasons, (28,29)) resulting in a blank cycle, whereas sulfated Tyr is hydrolyzed and seen as Tyr in almost normal yield. Furthermore, in contrast to Tyr-sulfated peptides (Fig. 1), phosphorylated peptides are recorded as

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the intact species in MALDI-TOF linear mode (30). In the reflector mode a typical triplet is observed from phosphorylated peptides consisting of the intact molecular ion accompanied by a major and a much less abundant fragment because of the loss of H3PO4 (98 Da) and HPO3 (80 Da), respectively (30). This is in contrast to the singular loss of 80 Da from a Tyr-sulfated peptide (see Note 5). The relative yields of the sulfated and nonsulfated form depend on the composition of the peptide. Small acidic peptides like the model peptide give almost clean peaks representing the sulfated and nonsulfated form in the negative and positive (linear) mode, respectively (Fig. 1), whereas larger and more neutral peptides give a more mixed picture. A MALDI-TOF mass spectrum obtained in the negative mode using reflector will show the correct molecular ion and an additional peak at a distance less than 80 Da below the value for the molecular ion (Fig. 2). This is caused by rupture of the sulfate ester bond in the linear flight tube after the peptide was accelerated but before the peptide reaches the reflector, and the calibration for the reflector requires ions with full accelerating energy. The decrease in molecular mass caused by sulfate loss observed in the reflected mode is instrument dependent, but once established it can also be used as an indicator of a sulfated peptide. The tyrosine 0-sulfated peptide, synthetic or purified has to be available in mg amounts. After dissolution in an appropriate buffer, the peptide is coupled to a protein carrier in amounts that ensure a coupling ratio of approx 5–10 molecule haptens per carrier molecule. There are several useful carrier candidates. We recommend bovine serum albumin (BSA, approx 5 mg/mg hapten decapeptide). BSA is easily available, inexpensive and in our experience as effective as any other protein carrier. When peptide hapten has to be synthesized, we recommend synthesis of an analogue of the genuine hapten equipped with a N- or C-terminal cysteinyl residue. Which terminus or end depends on the position of the sulfated tyrosyl residue (Fig. 3). The peptide hapten is then coupled to a free amino group in the carrier protein through the terminal cysteine using maleimidobenzoic acid N-hydroxysuccinimide ester as coupling reagent (36). Again, depending on the position of the sulfated tyrosyl residue, conventional coupling using either ethylcarbodiimide or glutaraldehyde may also be used (36). With directional N- or C-terminal carrier coupling (Fig. 3), it is important to design a tracer that corroborates and further advances the antibody specificity achieved. Because 125I is the preferred RIA-isotope, and because iodine easiest is coupled to tyrosyl- residues, we recommend synthesis of a tyrosylated hapten decapeptide in which the additional tyrosyl-residue for

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iodination is placed in the same position as cysteine in the hapten analogue, i.e. at the terminus opposite to the epitopic part of the hapten containing the 0-sulfated tyrosine (Fig. 3). In this way, iodinated tyrosine will not interfere with the antibody binding. There is no risk for iodination of the O-sulfated tyrosine-residue. The sulfonate group blocks incorporation of iodine. Natural occurrence of an additional unsulfated tyrosyl in the short hapten sequence has not been experienced so far; but in such situation a new labeling strategy has to be delineated- based on the exact positions of the additional and the sulfated tyrosyl residue. The tyrosine extended peptide analogue (Fig. 3) (4 nmoles) can be monoiodinated using the mild chloramine-T method, previously described (37). If mild oxidation damages the peptide (containing for instance methionyl residues), it is possible to use the nonoxidative Bolton-Hunter iodination (38). Subsequent purification on reversed-phase HPLC ensures a high specific radioactivity of the tracer. To evaluate the chromatographic separation of labeled and nonlabeled peptides, 1 ml of the monoiodinated peak fraction is mixed with 10 pmol of the relevant tyrosine-extended peptide and reapplied to the HPLC column as described. Both the radioactivity of the labeled peptides and the immunoreactivity are measured. Using the assumption that iodinated and unlabeled peptides are measured with identical affinity the specific radioactivity of the tracers can be determined by self-displacement (39). The essence of the described procedure is to ensure monoiodination without oxidative damages to the peptide. High specific radioactivity is then achieved by efficient chromatographic purification. 8. As standard substance for RIA-measurements using antibodies specific for sequence containing a sulfated tyrosyl-residue, it is straightforward to use a peptide corresponding to the hapten decapeptide. If, however, this decapeptide in some way is partly or completely hidden within the structure or conformation of the genuine tyrosyl sulfated protein, or if the antibody requires that epitope has a free N- or C-terminus (Fig. 3), it may be necessary to release the hidden epitope by cleavage with proteases that expose the epitope for antibody binding (40). If the natural occurring tyrosyl sulfated protein or polypeptide is recognized without cleavage, the protein may as well be used as standard- if available in sufficiently pure form. 9. In this example the peptide analyzed are acidic (the sequence of gastrin-17 is: pQGPWLEEEEEAYGWMDF-NH2) for which reason an anion exchange column is used. Because Tyr sulfation predominantly occurs in acidic regions of proteins and peptides, similar conditions this will often be appropriate. However, it is also possible to use cation exchange chromatography to separate peptide forms or alternatively, if the net charges of the fragments are neutral, the pH of the buffers may be adjusted.

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10. α-cyano-4-hydroxycinnamic acid gives a more homogeneous matrix than DHB and generally also a better signal. However, DHB is a cool matrix compared to α-cyano-4-hydroxycinnamic acid, and hence gives a reduced loss of sulfate in general and especially in reflector mode. 11. Four characteristics of sera from the immunized animals have to be examined (1) The titer is defined as the antiserum dilution that binds 33% of the 0.5-fmol tracer at equilibrium. (2) Affinity is expressed by the “effective” equilibrium constant (K0eff), determined as the slope of the curve at zero peptide concentration in a Scatchard plot (41). (3) Specificity is determined in percentage as the molar ratio of the concentrations of the sulfated standard peptide, the unsulfated peptide and other related peptides that produce a 50% inhibition of the binding of the tracer. (4) Homogeneity of the antibodies with respect to binding kinetics is expressed by the Sips index (42). An index of 1 indicates homogeneity of both the tracer and the antiserum in the binding, otherwise seen only for monoclonal antibodies (21). The ability of the peptides to displace tracers from the antisera may be tested in peptide concentrations of 0, 3, 10, 30, 100, 1000, 10 000, and 100 000 pmol/L. 12. Samples in HPLC buffers are preferable (0.1% TFA in 30% acetonitrile is ideal for MALDITOF). Salt and buffer concentrations should be kept as low as possible. Volatile buffers may be removed in a vacuum centrifuge and the dried sample redissolved in 0.1% TFA/30% acetonitrile. Most detergents, especially SDS, will completely abolish the signal. 13. The gradient should be chosen to give a good separation of peptides. The tyrosine sulfate containing peptides are increasingly suppressed as more peptides are eluted simultaneously. This results in higher risk of missing a tyrosine sulfated peptide in the neutral loss scanning procedure. 14. Note that the sulfate group will be completely lost during MS/MS and the spectrum will show the fragmentation pattern of the desulfated peptide, whereas the precursor ion is that of the sulfated peptide. This constitutes a problem to the general search engines, which are dependent on intact fragment ions of modified peptides to do correct annotation. Therefore manually annotation has to be applied or the parent ion mass must be altered before search. 15. It is our experience that the arylsulfatase treatment is tricky and sometimes very inefficient. We therefore recommend that experiments with negative outcome are repeated and, whenever possible, the inclusion of a positive control peptide with the experiments. References 1. Bettelheim, F. R. (1954) Tyrosine-O-sulfate in A peptide from fibrinogen. J. Am. Chem. Soc. 76, 2838–2839.

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2. Ouyang, Y., Lane, W. S., and Moore, K. L. (1998) Tyrosylprotein sulfotransferase: purificationand molecular cloning of an enzyme that catalyzes tyrosine O-sulfation, a common posttranslational modification of eukaryotic proteins. Proc. Natl Acad Sci. U S A 95, 2896–2901. 3. Beisswanger, R., Corbeil, D., Vannier, C., Thiele, C., Dohrmann, U., Kellner, R. et al. (1998) Existence of distinct tyrosylprotein sulfotransferase genes: molecular characterization of tyrosylprotein sulfotransferase-2. Proc. Natl Acad. Sci. USA 95, 11134–11139. 4. Ouyang, Y. B. and Moore, K. L. (1998) Molecular cloning and expression of human and mouse tyrosylprotein sulfotransferase-2 and a tyrosylprotein sulfotransferase homologue in Caenorhabditis elegans. J. Biol. Chem. 273, 24770–24774. 5. Ouyang, Y. B., Crawley, J. T., Aston, C. E., and Moore, K. L. (2002) Reduced body weight and increased postimplantation fetal death in tyrosylprotein sulfotransferase1-deficient mice. J. Biol. Chem. 277, 23781–23787. 6. Borghei, A., Ouyang, Y. B., Westmuckett, A. D., Marcello, M. R., Landel, C. P., Evans, J. P., and Moore, K. L. (2006) Targeted disruption of tyrosylprotein sulfotransferase-2, an enzyme that catalyses post-translational protein tyrosine O-sulfation, causes male infertility. J. Biol. Chem. 281, 9423–9431. 7. Mutt, V. (1980) Cholecystokinin: isolation, structure and functions, in Gastrointestinal Hormones. (Glass, G. B. J., ed.). Raven Press, New York; pp. 169–221. 8. Pouyani, T. and Seed, B. (1995) Psgl-1 recognition of p-selectin is controlled by a tyrosine sulfation consensus at the psgl-1 amino-terminus. Cell 83, 333–343. 9. Stone, S. R., Betz, A., Parry, M. A., Jackman, M. P., and Hofsteenge, J. (1993) Molecular basis for the inhibition of thrombin by hirudin. Adv. Exp. Med. Biol. 340, 35–49. 10. Farzan, M., Mirzabekov, T., Kolchinsky, P., Wyatt, R., Cayabyab, M., Gerard, N. P. et al. (1999) Tyrosine sulfation of the amino terminus of CCR5 facilitates HIV-1 entry. Cell 96, 667–676. 11. Bundgaard, J. R., Vuust, J., and Rehfeld, J. F. (1995) Tyrosine O-sulfation promotes proteolytic processing of progastrin. EMBO J 14, 3073–3079. 12. Friederich, E., Fritz, H. J., and Huttner, W. B. (1988) Inhibition of tyrosine sulfation in the trans-Golgi retards the transport of a constitutively secreted protein to the cell surface. J. Cell Biol. 107, 1655–1667. 13. Monigatti, F., Gasteiger, E., Bairoch, A., and Jung, E. (2002) The Sulfinator: predicting tyrosine sulfation sites in protein sequences. Bioinformatics 18, 769–770. 14. Rosenquist, G. L. and Nicholas, H. B. Jr. (1993) Analysis of sequence requirements for protein tyrosine sulfation. Protein Sci. 2, 215–222. 15. Nicholas, H. B. Jr., Chan, S. S., and Rosenquist, G. L. (1999) Reevaluation of the determinants of tyrosine sulfation. Endocrine 11, 285–292. 16. Chen, T. and Shaw, C. (2003) Cloning of the (Thr6)-phyllokinin precursor from Phyllomedusa sauvagei skin confirms a non-consensus tyrosine O-sulfation motif. Peptides 24, 1123–1130. 17. Bundgaard, J. R., Vuust, J., and Rehfeld, J. F. (1997) New consensus features for tyrosine O-sulfation determined by mutational analysis. J. Biol. Chem. 272, 21700–21705.

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18. Huttner, W. B. (1984) Determination and occurrence of tyrosine O-sulfate in proteins. Methods Enzymol. 107, 200–223. 19. Schechter, I. (1970) Mapping of the combining sites of antibodies specific for polyL-alanine determinants. Nature 228, 639–641. 20. Rehfeld, J. F. and Johnsen, A. H. (1994) Residue-specific immunochemical sequence prediction. J. Immunol. Methods 171, 139–142. 21. Rehfeld, J. F., Bardram, L., Cantor, P., Hilsted, L., and Johnsen, A. H. (1989) The unique specificity of antibodies in modern radioimmunochemistry–an essay on assays. Scand J. Clin. Lab. Invest. Suppl. 194, 41–44. 22. Hoffhines, A. J., Damoc, E., Bridges, K. G., Leary, J. A., and Moore, K. L. (2006) Detection and Purification of Tyrosine-sulfated Proteins Using a Novel Antisulfotyrosine Monoclonal Antibody. J. Biol. Chem. 281, 37877–37887. 23. Ross, A. H., Baltimore, D., and Eisen, H. N. (1981) Phosphotyrosine-containing proteins isolated by affinity chromatography with antibodies to a synthetic hapten. Nature 294, 654–656. 24. Rehfeld, J. F. (1998) Accurate measurement of cholecystokinin in plasma. Clin. Chem. 44, 991–1001. 25. Balsved, D., Bundgaard, J. R., and Sen, J. W. (2007) Stability of Tyrosine Sulfate in Acidic Solutions. Anal. Biochem. 363, 70–76. 26. Gregory, H., Hardy, P. M., Jones, D. S., Kenner, G. W., and Sheppard, R. C. (1964) The antral hormone gastrin. Structure of gastrin. Nature 204, 931–933. 27. Johnsen, A. H. (1991) Nondestructive amino acid analysis at the picomole level of prolinecontaining peptides using aminopeptidase M and prolidase: application to peptides containing tyrosine sulfate. Anal. Biochem. 197, 182–186. 28. Meyer, H.E., Hoffmann-Posorske, E., Donella-Deana, A., and Korte, H. (1991) Sequence analysis of phosphotyrosine-containing peptides. Methods Enzymol. 201, 206–224. 29. Meyer, H.E., Hoffmann-Posorske, E., and Heilmeyer, L.M. Jr. (1991) Determination and location of phosphotyrosine in proteins and peptides by conversion to S-ethylcysteine. Methods Enzymol. 201, 169–185. 30. Annan, R.S. and Carr, S.A. (1996) Phosphopeptide analysis by matrix-assisted laser desorption time-of-flight mass spectrometry. Anal. Chem. 68, 3413–3421. 31. Talbo, G. and Roepstorff, P. (1993) Determination of sulfated peptides via prompt fragmentation by UV matrix-assisted laser desorption/ionisation mass spectrometry. Rapid. Commun. Mass. Spectrom. 7, 201–204. 32. Salek, M., Costagliola, S., and Lehmann, W. D. (2004) Protein tyrosine-O-sulfation analysis by exhaustive product ion scanning with minimum collision offset in a NanoESI Q-TOF tandem mass spectrometer. Anal. Chem. 76, 5136–5142. 33. Haselmann, K. F., Budnik, B. A., Olsen, J. V., Nielsen, M. L., Reis, C. A., Clausen H. et al. (2001) Advantages of external accumulation for electron capture dissociation in Fourier transform mass spectrometry. Anal. Chem. 73, 2998–3005. 34. Monigatti F., Hekking B., and Steen H. (2006) Protein sulfation analysis-A primer. Biochim Biophys Acta. 1764, 1904–13. 35. Yu, Y., Hoffhines, A. J., Moore, K. L., and Leary, J. A (2007) Determination of the sites of tyrosine O-sulfation in peptides and proteins. Nat. Methods 4, 583–588.

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36. Harlow, E. and Lane, D. (1988) Antibodies: A laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, pp 82–83. 37. Stadil, F. and Rehfeld, J. F. (1972) Preparation of 125I-gastrin for radioimmunoanalysis. Scand. J. Clin. Lab. Invest. 30, 361–369. 38. Bolton, A. E. and Hunter, W. M. (1973) The labelling of proteins to high specific radioactivity by conjugation to a 125I-containing acylating agent. Biochem. J, 133: 529–539. 39. Morris, B. J. (1976) Specific radioactivity of radioimmunoassay tracer determined by self-displacement. Clin. Chim. Acta 73, 213–216. 40. Bardram, L. and Rehfeld, J. F. (1988) Processing-independent radioimmunoanalysis. Anal. Biochem. 175, 537–543. 41. Ekins, R. and Newman, B. (1970) Theoretical aspects of saturation analysis. Acta Endocrinol. 147, 11–36. 42. Sips, R. (1948) On the structure of a catalyst surface. J. Chem. Phys. 16, 490–495.

166 Analysis of Protein Palmitoylation by Metabolic Radiolabeling Methods Katherine H. Pedone, Leah S. Bernstein, Maurine E. Linder, and John R. Hepler 1. Introduction Many eukaryotic signaling proteins are modified by the covalent attachment of long-chain lipids. These highly hydrophobic molecules bind target proteins and facilitate interaction with cellular membranes, lipid molecules and other proteins. Generally, there are three classes of lipids that modify target proteins: myristate, isoprenoids and palmitate (for reviews, see refs. 1,2). Myristate, a 14-carbon saturated fatty acid, binds proteins at N-terminal glycine residues via an amide linkage. This usually occurs cotranslationally after removal of the initiating methionine, although it also can occur after proteolytic cleavage and exposure of an internal glycine. Long-chain isoprenoid lipids, including farnesyl and geranylgeranyl groups, modify proteins posttranslationally and attach via a thioether linkage to C-terminal cysteine residues. Palmitate is a 16-carbon saturated fatty acid that modifies proteins posttranslationally through thioester incorporation at cysteines. Palmitate also can modify proteins at additional sites and by alternative mechanisms, as in proteins palmitoylated at serine and threonine residues via oxyester linkages and by amide-linked N-terminal cysteines and glycines. In addition, other fatty acid species form thioester linkages with proteins. For these reasons, the general term for protein lipid modification by thioester attachment is thioacylation, or S-acylation, and the term palmitoylation refers specifically to protein modification by palmitate. Proteins that undergo palmitoylation by thioester attachment at cysteine residues, or S-palmitoylation, represent the majority of protein targets of palmitoylation. This chapter will focus on methods for identifying protein substrates for thioester incorporation of palmitate.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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A wide range of proteins become palmitoylated, including intracellular signaling proteins, transmembrane enzymes, secreted morphogens and cell surface receptors. Although the precise role of palmitoylation has proven difficult to discern, the modification has well-described pleiotropic effects on protein function. The common feature shared by all palmitoylated proteins is the capacity for interaction with cellular membranes. The extended hydrocarbon chain of the palmitate group acts as a stable membrane attachment point. This is particularly important for peripheral membrane proteins that otherwise are inherently soluble. The palmitate groups have high affinity for the membrane microdomain structures called lipid rafts in particular, and modification by palmitate is thought to target proteins to these membrane regions for selective protein interactions and signaling purposes. Preventing protein palmitoylation by point mutation or pharmacologic inhibition can exclude proteins from lipid rafts, impair biological activity and has effects on protein-protein interactions, protein stability and subcellular trafficking. For comprehensive reviews of protein palmitoylation, see refs. 3–5. Protein palmitoylation is an enzyme-regulated process that differs from other types of lipid modifications by its reversible nature. Both myristoylation and prenylation are considered stable modifications for the lifetime of the protein, but some palmitoylated proteins exhibit measurable half-lives for the lipid modification. Some intracellular signaling proteins can undergo multiple cycles of depalmitoylation followed by reloading with fresh palmitate. Guanine nucleotide-regulated proteins like Ras and heterotrimeric Gα protein subunits are examples of palmitoylated proteins that dynamically incorporate palmitate during signaling events. The rate of palmitate turnover on these proteins is enhanced dramatically by pharmacologic stimulation of cell surface receptors or direct protein activation (reviewed in ref. 6). For this reason, reversible attachment of palmitate to signaling proteins is thought to be a regulatory mechanism for controlling biological activity of the targeted proteins. Two classes of enzymes that regulate this process have been identified: protein acyltransferases (PATs) catalyze the transfer of palmitate from an intracellular donor such as palmitoyl-coenzyme A to a protein substrate, and protein thioesterases (PTEs) control removal of the lipid. For detailed information on these enzymes and the molecular mechanisms that underlie protein palmitoylation, see ref. 7. The protocols presented in this chapter outline how to assess if a protein is palmitoylated in living cells. The general approach is to incubate cells expressing the protein of interest with [3H]palmitate, isolate the protein from the cells by immunoprecipitation, and use SDS-PAGE and fluorography to determine whether it incorporated radioactive palmitate. In some cases, palmitate may be incorporated into serine or threonine residues through an oxyester linkage, thus it is necessary to verify that a labeled protein is a result of authentic, thioester-linked lipid. This

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is accomplished by treatment of the gel containing the labeled proteins with neutral hydroxylamine, which hydrolyzes only thioester-linked lipids. This method also is useful for analysis of proteins that have an N-terminal glycine to discern [3H]palmitate incorporation from incorporated [3H]myristate, metabolically converted from the [3H]palmitate. Identity of the attached fatty acid is confirmed by cleaving thioester-linked fatty acids with base. The fatty acids are then extracted from the hydrolysate and analyzed by HPLC or TLC. The methods described here for palmitoylation analysis by metabolic radiolabeling and fluorography have many advantages. The general experimental techniques are basic and commonly employed in labs utilizing cell culture systems. The specific methods are straightforward and require standard laboratory equipment and few reagents. The primary limitation of this approach is that it only provides a qualitative measure of palmitoylation. The stoichiometry of palmitate incorporation in a protein substrate cannot be determined using these methods. Other methods are available for quantitative analysis of protein palmitoylation. A newly developed technique called fatty acyl exchange labeling is a sensitive and quantitative approach to evaluating protein palmitoylation. In this method, palmitate molecules are cleaved from the modified protein by treatment with neutral hydroxylamine, and the resulting free sulfhydryl groups are coordinated with radiolabeled, thiol-specific reagents like [3H]N-ethylmaleimide (NEM) (8). This approach also can be adapted for use with non-radioactive (e.g. biotin-conjugated) reagents to bind available cysteine residues. Other methods for analyzing protein palmitoylation include mass spectrometry and in vitro palmitoylation assays with purified and partially purified PATs. These methods are described in more detail with good discussion on advantages and limitations of the techniques, particularly for fatty acyl exchange labeling, in reference (8). Once it is established that a protein is palmitoylated, the site(s) for palmitate attachment can be determined by mutation of cysteines to alanine or serine and the mutant protein assayed for loss of palmitoylation. There is no known consensus sequence for palmitate incorporation, so every cysteine on the surface of a protein should be considered a candidate site for palmitoylation. For a review of some shared sequence motifs in palmitoylated proteins, see reference (6). Proteins that incorporate palmitate at more than one site may contain a primary cysteine, where palmitoylation is required prior to incorporation at other sites within the protein. For this reason, it is important to design both individual and multi-site cysteine mutants to properly map the sites and/or sequence requirements for palmitate incorporation. For our studies, we use the PCR-based QuikChange® Site-Directed Mutagenesis Kit from Stratagene for creating individual and combination cysteine-to-serine point mutations in our proteins of interest (see Fig. 1). It is important to note that site-directed mutagenesis is an effective way to establish the dependence of the modification on a specific amino acid

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Fig. 1. Palmitoylation of G16α at N-terminal cysteines determined by metabolic radiolabeling analysis combined with site-directed mutagenesis. Top, upper panel: Immunoblot analysis of G16α-EE wild type (WT) and C9,10S protein immunoprecipitated (anti-EE antibody) from membrane (M) and cytosol (C) fractions of transfected HEK293 cells. Immunoblot analysis was performed with a Gq-family-specific antibody (Z811) kindly provided by Dr. Paul Sternweis (UT Southwestern Medical Center, Dallas, TX). Top, lower panel: Fluorography of G16α-EE WT and C9,10S protein immunoprecipitated from fractionated cells as described above. Exposure time was 56 days at −80°C. Bottom: Sequence alignments of the first 41 amino acids of G16α WT and C9,10S. Cysteine residues for putative palmitoylation and selected cysteine-to-serine mutations are indicated in bold type. Cysteine-to-serine mutations were generated using QuikChange® Site-Directed Mutagenesis kit from Stratagene (cat. no. 200518). (Reproduced from ref. 14 with permission of American Society for Biochemistry and Molecular Biology).

residue. However, definitive proof of site-specific modification requires isolation and identification of the acylated peptide by mass spectrometry. To successfully perform the techniques described in this chapter, several general considerations should be made. First, specific antibodies are required in order to immunoprecipitate the expressed protein. Since it often is difficult to produce antibodies against proteins directly, epitope-tagged versions of proteins can be engineered. In this chapter, we describe palmitoylation analysis of a G protein, G16α, that contains an internal glutamate-glutamate (EE) epitope tag. Other convenient epitopes can be used instead. Second, sufficient quantities of protein need to be expressed in order to produce a [3H]-labeled signal. Sf9 insect cells have provided an excellent expression system for palmitoylation studies, particularly for heterotrimeric G protein alpha subunits (Gα) (9). The primary advantage of insect cells is that infection with recombinant baculovirus produces relatively high levels of heterologous protein expression. Because they are eukaryotic cells, Sf9 contain the cellular enzyme machinery to modify expressed proteins. Detailed protocols for radiolabeling of Sf9 cells and for analysis of lipid modifications can be found in ref. 10.

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Although Sf9 cells have proven to be a good model system for palmitoylation studies of Gα proteins, some mammalian proteins may be processed or regulated inappropriately in an insect expression system. Subtle processing differences can have significant effects on palmitate incorporation into the protein. While it is desirable to analyze palmitoylation of any protein in a native system, often proteins are difficult to detect in the endogenous cells or tissues. In this chapter, we describe methods for analyzing palmitoylation of recombinant proteins heterologously expressed in a mammalian cell line. These methods have been used regularly by our labs to evaluate palmitoylation of various G protein subunits and protein Regulators of G Protein Signaling (RGS proteins) (9,11–14). In this chapter, our example protein is G16α, a G protein alpha subunit from the Gq family of heterotrimeric G proteins. We recently showed that G16α is posttranslationally modified by palmitate and that at least two of three cysteine residues in the N-terminus of the protein are essential for palmitoylation (Fig. 1) (14). We used HEK293 cells for our studies, although COS-7, CHO, or other readily transfectable adherent cell line may be substituted (see Note 1). Most of the methods described here for mammalian cells are the same or similar to those for Sf9 cells (10). The differences between the two expression systems are the methods for introduction of the construct into the cells and cell lysis. Since many laboratories are equipped for mammalian cell culture, these protocols are likely to be straightforward to adopt. 2. Materials 2.1. Measuring Palmitate Incorporation into Recombinant EE-Tagged G16a Protein in Mammalian Cell Culture Systems 2.1.1. Transfection and Metabolic Labeling of EE-Tagged G16a in Mammalian Cells 1. 2. 3. 4. 5.

HEK293 cells (American Type Culture Collection). Transfection-quality G16α-EE plasmid DNA (see Note 2). Lipofectamine™ 2000 Transfection Reagent (Invitrogen). Opti-MEM® I reduced-serum media (Invitrogen). Complete HEK293 medium: Dulbecco’s Modified Eagle’s Medium (DMEM; Invitrogen), 10% fetal bovine serum (Atlanta Biologicals), penicillin/streptomycin (CellGro). 6. [35S]methionine wash medium: 95% methionine-free DMEM (Invitrogen, cat. no. 21013-024), 5% DMEM, 2.5% dialyzed fetal bovine serum (Invitrogen). 7. [35S]methionine labeling medium: 95% methionine-free DMEM, 5% DMEM, 2.5% dialyzed fetal bovine serum, 1 mM sodium pyruvate (Invitrogen), 100 µCi/ mL L-[35S]methionine (>1000Ci/mmol) (Amersham Biosciences). Aliquot the [35S]methionine and store at −80°C. Make media fresh before use.

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8. [3H]palmitate labeling medium: 1 mCi/mL [3H]palmitate ([9,10-3H]-Palmitic Acid (31 Ci/mmol), (PerkinElmer Life and Analytical Sciences), 1% DMSO, 2.5% fetal bovine serum, 1 mM sodium pyruvate, 1 mM non-essential amino acids (Invitrogen). Make fresh before use (see Note 3). 9. Nitrogen gas.

2.1.2. Subcellular Fractionation 1. Potter-Elvehjem tissue grinders with polytetrafluoroethylene (PTFE)-coated pestle, 2 mL capacity, 0.1–0.15 mm clearance (Wheaton Potter-Elvehjem Tissue Grinder, Thermo Fisher Scientific). 2. Ice-cold phosphate-buffered saline (PBS). 3. Hypotonic lysis buffer: 50 mM HEPES, pH 8.0, 1 mM EDTA, 1 mM DTT, protease inhibitors (2 µg/mL aprotinin, 1 µg/mL leupeptin, 100 µM phenylmethylsulfonyl fluoride (PMSF) ). Add DTT and protease inhibitors just before use. 4. 2X isotonic buffer: 50 mM HEPES, pH8.0, 10 mM MgCl2, 300 mM NaCl, 1 mM EDTA, 500 mM sucrose, protease inhibitors (4 µg/mL aprotinin, 2 µg/mL leupeptin, 200 µM PMSF). Add protease inhibitors just before use. 5. 1.5 mL capacity microfuge tubes for high-speed centrifugation (Microfuge Tube Polyallomer, Beckman). 6. 2 mL capacity microfuge tubes. 7. 1X Immunoprecipitation (IP) buffer: 50 mM HEPES, pH8.0, 5 mM MgCl2, 150 mM NaCl, 1 mM EDTA, 250 mM sucrose, protease inhibitors (2 µg/mL aprotinin, 1 µg/mL leupeptin, 100 µM PMSF). Add protease inhibitors just before use. 8. IP buffer with 4% DβM: 4% n-dodecyl-β-D-maltoside (DβM; Calbiochem) in 1X IP buffer. DβM is used at a final concentration of 2% for extraction of membrane proteins.

2.1.3. Immunoprecipitation of labeled protein, SDS-PAGE and fluorography 1. 2. 3. 4.

Anti-GLU-GLU (EE) monoclonal antibody (Covance). Protein A Sepharose (nProtein A Sepharose® 4 Fast Flow, GE Healthcare). Bovine serum albumin (Sigma). Coomassie blue stain: 30% isopropanol, 10% acetic acid, 0.25% Coomassie brilliant blue R-250. 5. Destain solution: 30% methanol, 10% acetic acid. 6. En3Hance autoradiography enhancer solution (En3Hance™, PerkinElmer Life and Analytical Sciences, cat. no. 6NE9701) (see Note 4). 7. 2X Laemmli sample buffer: 100 mM Tris-HCl pH6.8, 0.5% SDS, 20% glycerol, 0.5% β-mercaptoethanol, 0.004% bromophenol blue. Prepare stocks of 2X Laemmli sample buffer with and without 20 mM DTT.

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2.2. Hydroxylamine Sensitivity by Thin Layer Chromatography 1. Hydroxylamine solution: 0.5 M hydroxylamine hydrochloride (Sigma-Aldrich), pH7.0, in 50% isopropanol. Use concentrated NaOH to pH the solution. 2. 0.5 M Tris-HCl, pH7.0, in 50% isopropanol.

2.3. Identification of Fatty Acids on Labeled Protein 1. 13 × 100 mm glass tubes with Teflon-lined caps (Corning, cat. no. 9826-13). 2. HPLC grade methanol, chloroform, acetonitrile, acetic acid. 3. [3H]-fatty acid standards: [3H]palmitate, [3H]myristate, [3H]stearate (Perkin Elmer Life and Analytical Sciences). 4. Nitrogen gas. 5. NaOH. 6. 6N HCl, high grade (Pierce Chemicals). 7. Palmitic acid (Sigma). 8. Trifluoroacetic acid. 9. C:18 reversed-phase thin-layer chromatography plates (Whatman). 10. En3Hance™ Spray Surface Autoradiography Enhancer (Perkin Elmer Life and Analytical Sciences).

3. Methods 3.1. Measuring palmitate incorporation into recombinant G16a-EE protein in mammalian cells 3.1.1. Transfection and metabolic labeling of G16a-EE protein in mammalian cells This protocol calls for metabolically labeling parallel dishes of transfected cells: one with [3H]palmitate, the counterpart with [35S]methionine. The purpose of labeling cells with [35S]methionine is to control for expression and recovery of the recombinant protein of interest. Alternatively, immunoblot analysis of the protein of interest immunoprecipitated from unlabeled cells may be substituted for radiolabeling with [35S]methionine (Fig. 1). 1. Plate 2–10 cm2 dishes of HEK293 cells so that they will be 85–90% confluent the next day. 2. Transfect HEK293 cells with G16α-EE DNA construct using Lipofectamine™ 2000 Transfection Reagent according to manufacturer’s instructions (see Note 5). Incubate 5 h, 37°C, 5% CO2/95% air. 3. Replace transfection media with complete media, and allow cells to recover in 37°C incubator overnight (see Note 6). 4. Approximately 36 h after beginning the transfection, label one plate with [35S] methionine: wash once in [35S]methionine wash media, then add labeling media. Incubate overnight in the 37°C incubator.

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5. The following day, label the second plate with [3H]palmitate: wash cells once with pre-warmed DMEM, then incubate for 1–2 h in [3H]palmitate labeling medium (see Note 7). 6. Harvest each plate 48 h post-transfection: using a clean razor blade or rubber policeman, scrape cells into separate conical tubes on ice. 7. Collect cells by centrifugation at 500 g, 4°C for 4 min. 8. Aspirate media into a receptacle for radioactive liquids. Resuspend cells in 5 mL ice-cold PBS to wash away residual media. 9. Collect cells by centrifugation at 500 g, 4°C for 4 min.

3.1.2. Subcellular Fractionation (see Note 8) 1. Aspirate PBS, and resuspend each cell pellet in 0.5 mL hypotonic lysis buffer. Transfer cells to separate Potter-Elvehjem tissue grinders on ice. 2. Homogenize cells with 30–50 strokes, and transfer lysates to separate 1.5 mL microfuge tubes on ice (see Note 9). 3. Add 0.5 mL 2X isotonic buffer to each tube to restore normal osmotic pressure. 4. Centrifuge lysates at 600 g, 4°C for 10 min to collect nuclei, unbroken cells, and plasma membrane sheets. 5. Transfer the supernatants to Beckman polyallomer microfuge tubes. Spin in TLA100.3 rotor with adaptors, 100,000 g, 4°C for 30 min. 6. Remove supernatants from the high-speed spin (cytosol) to clean 2 mL microfuge tubes. Add 1 mL IP buffer with 4% DβM to each for a final detergent concentration of 2% DβM. 7. Resuspend pellets (membranes) in 0.5 mL IP buffer (without detergent) in the polyallomer tubes, and transfer to Potter-Elvehjem tissue grinders on ice. Lightly homogenize the membrane pellets (15 strokes). 8. Add 0.5 mL IP buffer with 4% DβM to each membrane sample for a final detergent concentration of 2% DβM. 9. Solublize membranes by incubating at 4°C for 3 h on an end-over-end rocker (see Note 10). Incubate cytosol samples similarly for the same duration. 10. Pellet the insoluble material from membrane samples by high-speed centrifugation at 100,000 g, 4°C for 30 min. 11. Remove the supernatants (membrane extract) to clean microfuge tubes (see Note 11).

3.1.3. Immunoprecipitation of Labeled Protein, SDS-PAGE, and Fluorography (see Note 12) 1. Preblock membrane extracts and cytosol samples in 0.7 mg/mL BSA solution and pre-absorb with 50 µL Protein A Sepharose. Incubate with end-over-end rotation, 4°C from 30 min to 2 h. 2. Add 6–7 µL anti-EE antibody to each sample of membrane extracts and cytosol. Incubate at 4°C at least 3 h, or overnight if convenient, rotating end-over-end. 3. Collect sepharose beads and immunoprecipitated protein complexes by centrifugation: 200 g, 4°C for 1 min. 4. Aspirate supernatant, and resuspend sepharose beads in 1 mL IP buffer containing 0.2% DβM.

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5. Repeat spin: 200 g, 4°C for 1 min. Wash sepharose beads two more times in the same fashion. 6. Suspend the washed beads in 60 µL 2X Laemmli sample buffer without DTT (see Note 13). 7. Boil samples 1 min, then spin 1 min in microcentrifuge at maximal speed. Load the supernatant samples on SDS-PAGE gels and run (see Note 14). 8. Stain gels with Coomassie blue stain, rocking, 30 min. 9. Incubate in several changes of destain solution, rocking, until proteins are visualized on the gel (see Note 15). 10. Soak gels in En3Hance™ autoradiography enhancer solution, 60 min, with gentle rocking. 11. Wash gels thoroughly in a large volume of cold water for 30 min with gentle rocking. 12. Dry down the gels to completion on a vacuum dryer, 60°C. 13. Expose the gels to film in a cassette. Store the cassette at −80°C for at least 3 days to visualize the [35S]-labeled bands. Replace with new film and expose for 1–4 weeks to visualize [3H]-labeled bands (see Note 16).

3.2. Hydroxylamine Sensitivity of [3H]palmitate-Labeled Proteins (see Note 17; Fig. 2) 1. Transfect cells, label and fractionate cells as in steps 3.1.1 and 3.1.2 (see Note 18). 2. Immunoprecipitate EE-tagged protein from cells as outlined in 3.1.3, steps 1–6 above. 3. Boil samples 1 min, then spin 1 min in microcentrifuge at maximal speed. Divide each sample in half, and run duplicate gels of each. 4. Fix gels by incubating in destain solution for 30 min, rocking.

Fig. 2. Confirmation of thioester linkage of [3H]palmitate by hydroxylamine cleavage. Fluorography of the radiolabeled G16α N-terminus with and without neutral hydroxylamine treatment. A peptide representing the N-terminal domain of G16α (amino acids 1–41) was expressed in HEK293 cells as a FLAG-tagged GFP fusion protein. The fusion protein was immunoprecipitated (Anti-FLAG M2 Affinity Gel, Sigma, cat. no. A2220) from transfected membranes of HEK293 cells metabolically radiolabeled with either [35S]methionine or [3H] palmitate. Immunoprecipitated proteins were treated with 0.5 M Tris, pH7.0 (left panel) or 0.5 M hydroxylamine, pH7.0 (NH2OH, right panel) and prepared for fluorographic analysis. Exposure time was 2 days at −80°C for [35S]-signals, 5 days for [3H]-signals. (Reproduced from ref. 14 with permission of American Society for Biochemistry and Molecular Biology.)

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5. Soak one gel in fresh 0.5 M hydroxylamine solution (pH7.0) for 12–18 hours (overnight). As a control, soak the other gel in 0.5 M Tris (pH7.0) in 50% isopropanol. Replace the solutions at least once after approximately one h. The gels will shrink as they are dehydrated by the isopropanol. 6. Wash each gel 4 times in 50% isopropanol over a period of 48 h. 7. Soak gels in Coomassie stain, then destain solutions as above to visualize precipitated proteins. The gels will rehydrate and return to original size during this step. 8. Soak gels in En3Hance™ autoradiography enhancer solution for 60 min with gentle rocking. 9. Dry down the gels on a vacuum dryer, 60°C, and expose to film at −80°C.

3.3. Identification of Fatty Acids on Labeled Protein 3.3.1. Base Hydrolysis and Extraction of Thioester-Linked Fatty Acids 1. Transfect cells, label, and fractionate cells as in steps 3.1.1 and 3.1.2. 2. Immunoprecipitate protein from cells as outlined in Subheading 3.1.3, steps 1–6. 3. Boil samples 1 min, then pellet for 1 min in microcentrifuge at maximal speed. Divide each sample in half and run duplicate SDS-PAGE gels. 4. Stain gels with Coomassie blue stain, rocking, 30 min. 5. Incubate in several changes of destain solution until proteins are visualized on the gel. 6. Excise the immunoprecipitated protein bands from membranes and cytosol with a razor blade or scalpel, keeping gel slices equal in size. Also excise three gel slices from unused portions of the gel to run with standards. 7. Place gel slices in air-tight, screw-cap 13 × 100 mm glass tubes. Wash gel slices overnight with five changes of 50% methanol. 8. Remove methanol with Pasteur pipet. To the blank gel slices, add [3H]myristate, [3H]stearate, or [3H]palmitate standards, 25,000–50,000 dpm per sample. Dry gel slices under a nitrogen stream. 9. To hydrolyze thioester-linked lipids, resuspend gel slices in 0.7 mL 1.5 M NaOH. Flush tubes with nitrogen and cap. Incubate 3 h, 37°C, mixing periodically. 10. Acidify the solutions to pH1-2 by adding 6N (high-grade) HCl. This typically requires 100–200 µL of acid. 11. To extract the released fatty acids, add 3.75 volumes of chloroform:methanol (1:2 v/v) in a fume hood. This results in chloroform:methanol:aqueous 1.25:2.5:1 v/v/v. Incubate 10 min, room temperature with occasional mixing. 12. To separate the fatty acids from water-soluble components and gel slices, add 1.25 volumes (of original aqueous volume) of chloroform and 1.25 volumes of water. 13. Vortex tubes, then centrifuge at 1,000 rpm at room temperature. This will separate the mixture into an upper aqueous phase and a lower organic phase, with the gel slice in between. 14. Remove the bottom phase containing fatty acids by bubbling a Pasteur pipette through the aqueous phase to the bottom of the tube. Transfer to new screw-cap tubes. 15. To extract remaining fatty acids, add the same amount of chloroform as in step 12. Incubate 5 min, room temperature with occasional mixing. Centrifuge 1,000 rpm and remove bottom phase as before. Combine with first extracts.

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16. Dry down extracts under nitrogen. 17. Add 0.5 mL chloroform and 40 µg cold palmitic acid as a carrier. Flush tubes with nitrogen and store at −20°C.

3.3.2. Thin-Layer Chromatographic Analysis of Hydrolyzed Lipids 1. Count a small volume of each sample (50 µL) in scintillation vials to determine how much sample to load. Let the chloroform dry before counting or it will quench. 2. Spot equivalent amounts of each sample (25,000–50,000 dpm) on a reverse-phase C:18 TLC plate. Also spot equivalent standards. 3. Resolve the extracts using 90% acetonitrile/10% acetic acid in a chromatography tank. This should take 20–30 min. 4. After the solvent has migrated to the top, remove plates and allow them to dry thoroughly in the hood. 5. Spray plates with En3Hance™ spray. Let plates dry slightly and expose to film at −80°C for 5–10 h. Re-expose for 24–48 h if signal is undetectable.

4. Notes 1. The protocols for heterologous protein expression, cell lysis and fractionation (Subheadings 3.1.1 and 3.1.2) are described here specifically for mammalian cells. The remainder of the methods are performed the same independent of eukaryotic cell type used. 2. We use Midi- or Maxi-prep kits from Qiagen to produce DNA for transfections. Follow the manufacturer’s protocols. Our G16α-EE DNA plasmid came from UMR cDNA Resource Center (www.cdna.org; cat. no. GNA1500000). Many other DNA constructs for mediators of G protein signaling (GPCRs, G proteins, RGS proteins) also are available from UMR in epitope-tagged and untagged forms. 3. [3H]palmitate is supplied as a solution in ethanol and cannot be added directly to media. First, measure the appropriate amount into a glass tube and evaporate the ethanol under a nitrogen stream. When the palmitate is dry, it will appear as a light film on the bottom of the tube. Resuspend the palmitate first in DMSO (for final concentration of 1% in palmitate labeling media; volume will depend on final volume of labeling media), then add the fetal bovine serum (final concentration 2.5%). Add the remaining reagents to the dissolved [3H]palmitate. 4. We have successfully used a homemade version of fluorographic reagent: 1 M sodium salicylate in 15% methanol. Store in a dark bottle. 5. Detailed instructions for optimizing transfection conditions with Lipofectamine™ 2000 Transfection Reagent are provided by the manufacturer. The amount of DNA and lipofectamine used per dish can vary depending on the protein to be expressed.

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6. The optimal time to harvest transfected cells for peak protein expression may vary depending on the protein. In this protocol, cells are collected 48 h post-transfection. For shorter post-transfection incubations, labeling should be started at the appropriate time so that [35S]methionine labeling media is incubated overnight and [3H]palmitate media is incubated for 1–2 h. 7. Some proteins undergo turnover of the palmitate modification. For example, the half-life for palmitate on H-Ras has been measured as approximately 2.4 h; however, the half-life decreases to 90 min for ectopically expressed, activated H-Ras, suggesting that G protein activation is linked to palmitate turnover (15). The propensity for palmitate turnover should be considered when establishing a time period for metabolic labeling with [3H]palmitate. A 1–2 h labeling period with high concentrations of [3H]palmitate as suggested in this protocol is a good starting point. G protein alpha subunit palmitoylation has been detected with labeling periods as short as 15 min (16). 8. All solutions should be kept cold (4°C), and all samples should be kept on ice during the following steps in order to minimize protein degradation. 9. If using the same Potter-Elvehjem tissue grinder for both [35S]- and [3H]labeled cells during an experiment, pass the [3H]-labeled samples first to avoid accidental transfer of the higher energy [35S]-signal into the [3H]labeled samples. Wash out the tissue grinder thoroughly with ethanol to remove trace [3H]-residue. Alternatively, cells may be lysed by another convenient method (e.g. repeated cycles of snap-freezing and quick-thawing). 10. Other buffers may be used to solublize proteins successfully. We found this HEPES-based buffer formulation with 2% DβM detergent to work well for extraction and immunoprecipitation of our Gα proteins with the anti-EE antibody. If a different antibody is used, the buffer may need to be adjusted. Also, a shorter incubation period for membrane extraction may be sufficient for other proteins. Generally, a 1-h extraction period is sufficient to release proteins from membranes prepared at 1 mg/mL membrane protein concentration. 11. At this step, the membrane extracts and cytosol samples may be snap-frozen in liquid nitrogen and stored at −80°C until later use, as limited by the half-life of the [35S]-signal. Frozen samples should be thawed quickly and kept on ice. 12. Immunoprecipitation protocols vary widely. For a review and guide to optimization of immunoprecipitation using different types of antibodies and solid support matrices, see reference (17). Other protocols may be easily substituted. 13. The thioester linkage for S-palmitoylation is vulnerable to reducing agents such as DTT. For this reason, we exclude DTT from our Laemmli sample buffer for [3H]palmitate-labeled samples. This does not affect migration

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15. 16.

17.

18.

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of our G proteins on an SDS-polyacrylamide gel, but the same may not be true for other proteins. 20 mM DTT may be included in [3H]palmitatelabeled samples if needed, but the lability of the thioester linkage should be considered. Laemmli sample buffer containing 20 mM DTT should be used for all other protein samples prepared for SDS-PAGE. Avoid loading sepharose beads onto the SDS-PAGE gel. Thioester-linked fatty acid also is susceptible to prolonged heating. We have found that excluding DTT from the sample buffer and heating for 1 min is sufficient to obtain good resolution of Gα proteins on gels without a substantial loss of radioactive palmitate on the protein. The gel can be left in destain solution overnight if desired. Do not use plastic wrap to cover the gel during exposure as it will block the [3H]-signal. Storage of the cassette at −80°C is essential for the En3Hance™ fluorography reagent to work. Do not use a screen. The [35S]-labeled samples may be visualized within a few days. The [3H]-labeled bands may be detected after one week exposure, but it can take one month or longer, depending on the amount of protein on the gel. Exposure time must be determined empirically. If the membrane-bound protein of interest incorporates [3H]palmitate, further analysis should be done to confirm authentic thioester palmitate attachment. Treatment of the gel containing the [3H]-labeled sample with neutral hydroxylamine should result in the reduction or disappearance of the labeled band. As a control, a duplicate gel is soaked in Tris buffer at the same pH. The [3H]-labeled sample should be visible in this gel. Finally, [35S]methionine-labeled proteins should be run in parallel to control for the loss of protein from the hydroxylamine- and Tris-treated gels during processing. A hydroxylamine-insensitive label can be analyzed as a potential amide-linkage by treating the gels with acid instead. Amide-linked acids are insensitive to hydroxylamine treatment, but are sensitive to acid hydrolysis. The number of cells required for these experiments may vary depending on protein expression and stoichiometry of palmitoylation. Adjust the size and/or number of dishes transfected based on the autoradiograph in Subheading 3.1.

References 1. Casey, P. J. (1995) Protein lipidation in cell signaling. Science 268, 221–225. 2. Resh, M. D. (2006) Trafficking and signaling by fatty-acylated and prenylated proteins. Nat. Chem. Biol. 2, 584–590. 3. Dunphy, J. T. and Linder, M. E. (1998) Signalling functions of protein palmitoylation. Biochim. Biophys. Acta 1436, 245–261.

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4. Resh, M. D. (2006) Palmitoylation of Ligands, Receptors, and Intracellular Signaling Molecules. Sci. STKE 359, re14. 5. Greaves, J. and Chamberlain, L. H. (2007) Palmitoylation-dependent protein sorting. J. Cell Biol. 176, 249–254. 6. Smotrys, J. E. and Linder, M. E. (2004) Palmitoylation of Intracellular Signaling Proteins: Regulation and Function. Ann. Rev. Biochem. 73, 559–587. 7. Linder, M. E. and Deschenes, R. J. (2007) Palmitoylation: policing protein stability and traffic. Nat. Rev. Mol. Cell Biol. 8, 74–84. 8. Drisdel, R. C., Alexander, J. K., Sayeed, A., and Green, W. N. (2006) Assays of protein palmitoylation. Methods 40, 127–134. 9. Linder, M. E., Middleton, P., Hepler, J. R., Taussig, R., Gilman, A. G., and Mumby, S. M. (1993) Lipid modifications of G proteins: α subunits are palmitoylated. Proc. Natl. Acad. Sci. U.S.A. 90, 3675–3679. 10. Linder, M. E., Kleuss, C., and Mumby, S. M. (1995) Palmitoylation of G-protein α subunits. Methods Enzymol. 250, 314–330. 11. Hepler, J. R., Biddlecome, G. H., Kleuss, C., Camp, L. A., Hofmann, S. L., Ross, E. M., and Gilman, A. G. (1996) Functional importance of the amino terminus of Gqα. J. Biol. Chem. 271, 496–504. 12. Rose, J. J., Taylor, J. B., Shi, J., Cockett, M. I., Jones, P. G., and Hepler, J. R. (2000) RGS7 is palmitoylated and exists as biochemically distinct forms. J. Neurochem. 75, 2103–2112. 13. Srinivasa, S. P., Bernstein, L. S., Blumer, K. J., and Linder, M. E. (1998) Plasma membrane localization is required for RGS4 function in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. U.S.A. 95, 5584–5589. 14. Pedone, K. H. and Hepler, J. R. (2007) The Importance of N-terminal Polycysteine and Polybasic Sequences for G14α and G16α Palmitoylation, Plasma Membrane Localization and Signaling Function. J. Biol. Chem. 282, 25199–25212. 15. Baker, T. L., Zheng, H., Walker, J., Coloff, J. L., and Buss, J. E. (2003) Distinct Rates of Palmitate Turnover on Membrane-bound Cellular and Oncogenic H-Ras. J. Biol. Chem. 278, 19292–19300. 16. Mumby, S. M., Kleuss, C., and Gilman, A. G. (1994) Receptor regulation of G-protein palmitoylation. Proc. Natl. Acad. Sci. U.S.A. 91, 2800–2804. 17. Harlow, E. and Lane, D. (1988) Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Plainview, NY.

167 Incorporation of Radiolabeled Prenyl Alcohols and Their Analogs into Mammalian Cell Proteins A Useful Tool for Studying Protein Prenylation Alberto Corsini, Christopher C. Farnsworth, Paul McGeady, Michael H. Gelb, and John A. Glomset 1. Introduction Prenylated proteins comprise a diverse family of proteins that are posttranslationally modified by either a farnesyl group or one or more geranylgeranyl groups (1–3). Recent studies suggest that members of this family are involved in a number of cellular processes, including cell signaling (4–6), differentiation (7–9), proliferation (10–12), cytoskeletal dynamics (13–15), and endocytic and exocytic transport (4,16,17). The authors’ studies have focused on the role of prenylated proteins in the cell cycle (18). Exposure of cultured cells to competitive inhibitors (statins) of 3-hydroxy-3-methylglutaryl Coenzyme A (HMGCoA) reductase not only blocks the biosynthesis of mevalonic acid (MVA), the biosynthetic precursor of both farnesyl and geranylgeranyl groups, but pleiotropically inhibits DNA replication and cell-cycle progression (10,18–20). Both phenomena can be prevented by the addition of exogenous MVA (10,18,19). The authors have observed that all-trans-geranylgeraniol (GGOH) and, in a few cases, all-trans-farnesol (FOH) can prevent the statin-induced inhibition of DNA synthesis (21). In an effort to understand the biochemical basis of these effects, the authors have developed methods for the labeling and two-dimensional gel analysis of prenylated proteins that should be widely applicable. Because of the relative diversity of prenylated proteins, it is important to use analytical methods that differentiate between them. A useful approach discussed here is to selectively label farnesylated or geranylgeranylated proteins using [3H] labeled FOH or GGOH (22–24), followed by one-dimensional From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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SDS-PAGE or high-resolution twodimensional gel electrophoresis (2DE) of the labeled proteins. Since the enzymes that transfer prenyl groups to proteins utilize the corresponding prenyl alcohol pyrophosphate (FPP or GGPP) as substrate, these prenols are thought to undergo two phosphorylation steps prior to their subsequent utilization (2,3). The discovery of a GGOH kinase and a geranylgeranyl phosphate kinase in eubacteria (25), together with the ability of rat liver microsomal and peroxisomal fractions to form FPP, provide additional evidence that these prenol pools serve as a source of lipid precursor for protein prenylation. When proteins are labeled in this way, and are subsequently analyzed by one-dimensional SDS-PAGE, it is possible to distinguish several major bands of radioactivity that correspond to two apparently distinct subsets of proteins: those that incorporate [3H]-FOH and those that incorporate [3H]GGOH. But when the labeled proteins are analyzed by high-resolution 2DE, the number of radioactive proteins that are observed is at least fivefold greater, and three subsets of prenylated proteins can be identified: one subset of proteins that incorporates only farnesol, a second that incorporates only geranylgeraniol, and a third that can incorporate either prenol. In this chapter, the advantages of using labeled prenols to dissect the differential effects of FOH and GGOH on cellular function will be presented in the context of studies of the role of prenylated proteins in cell-cycle progression. The ability of mammalian cells to incorporate natural and synthetic prenol analogs (Fig. 1) into specific proteins also will be discussed. In Subheading 4, some of the advantages and limitations of the methods will be discussed. 2. Materials 2.1. Prenols and Analogs: Synthesis and Labeling 1. All-trans-FOH, d20 0.89 g/mL; mevalonic acid lactone; geraniol, d20 0.89 g/mL (Sigma, St. Louis, MO). 2. Tetrahydrofarnesol was a gift from Hoffman la Roche (Basel, Switzerland). A racemic form can be made from farnesyl acetone following the procedure for hexahydro-GGOH (see Subheading 3.1.1.). 3. All-trans-GGOH, d20 0.89 g/mL; all-trans-GGOH [1-3H], 50–60 Ci/mmol; alltrans-FOH [1-3H], 15–20 Ci/mmol; geraniol [1-3H], 15–20 Ci/mmol (American Radiolabeled Chemicals, St. Louis, MO). 4. Mevalonolactone RS-[5-3H(N)], 35.00 Ci/mmol (NEN, Boston, MA). 5. Charcoal; LiAlH4 powder; MnO2; NaBH4 powder; triethyl phosphonoacetate; phosphonoacetone; sodium ethoxide (Aldrich, Milwaukee, WI). 6. [3H]-NaBH4 solid, 70 Ci/mmol (Amersham, Arlington Heights, IL). 7. Silica gel 60 (F254) (Merck, Gibbstown, NJ). 8. Aquamix liquid scintillation solution (ICN Radiochemicals, Irvine, CA).

Fig. 1. Structure of the natural and synthetic prenol analogs used in these labeling studies.

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2.2. Cell Culture Reagents 1. Dulbecco’s modified Eagle media (DMEM) (glucose 4.5 g/L); penicillin (10,000 U/mL)–streptomycin (10 mg/mL); trypsin (0.25% w/v)–1 mM ethylenediaminetetraacetate (EDTA); nonessential amino acids (NEAA) solution 10 mM; phosphate-buffered saline (PBS) without Ca2+ and Mg2+; fetal calf serum (FCS) (Gibco, Grand Island, NY). 2. Disposable culture Petri dishes (100 × 10 mm and 35 × 10 mm) (Corning Glass Works, Corning, NY). 3. Filters, 0.22 µm (Millipore, Bedford, MA). 4. Plasma-derived, bovine serum (PDS) (Irvine Scientific, Santa Ana, CA). 5. Aquasol scintillation cocktail; thymidine [methyl-3H], 2 Ci/mmol] (NEN). 6. Trichloroacetic acid (TCA) (J. T. Baker, Phillipsburg, NJ). 7. Phenylmethylsulfonyl fluoride (PMSF), aprotinin, leupeptin, and pepstatin A (Sigma). 8. Simvastatin in its lactone form (gift from Merck, Sharp and Dohme; Rahway, NJ) is dissolved in 0.1 M NaOH (60°C, 3 h), to give the active form. Adjust the pH to 7.4 and the concentration to 50 mM, then sterilize by filtration. 9. Swiss 3T3-albino mouse cell line (3T3) were from American Type Culture Collection (ATCC), Rockville, MD. 10. Human skin fibroblasts (HSF) are grown from explants of skin biopsies obtained from healthy individuals. The cells are used between the fifth and fifteenth passages.

2.3. One-Dimensional and Two-Dimensional Gel Electrophoresis 1. SDS, TEMED, ammonium persulfate, mol wt protein standards, glycine, Bradford protein assay kit, heavyweight filter paper, and Bio Gel P6-DG (Bio-Rad, Hercules, CA). 2. Duracryl™ preblended acrylamide solution (30% T, 0.65% C, used for both onedimensional and 2DE gels) and all other 2DE gel reagents, were obtained from ESA, Chelmsford, MA. 3. Ethylmaleimide, N-[ethyl-1,2-3H] ([3H]-NEM), 57 Ci/mmol (NEN). 4. N-ethylmaleimide (Aldrich). 5. Soybean trypsin inhibitor and SDS-7 mol wt standards used with 2DE gels (Sigma). 6. Amplify and Hyperfilm (Amersham). 7. Reacti-Vials™ (Pierce, Rockford, IL). 8. DNaseI and RNaseA (Worthington Biochem, Freehold, NJ).

3. Methods 3.1. Synthesis and Labeling of Prenols and Analogs 3.1.1. Synthesis of Racemic 6,7,10,11,14,15 Hexahydrogeranylgeraniol 1. This synthesis is outlined in Fig. 2. Weigh 10.4 g of farnesyl acetone and 500 mg 10% Pd on charcoal, dissolve in 100 mL methanol and hydrogenate at 50 psi in a Parr

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Fig. 2. Schematic for the synthesis of racemic 6,7,10,11,14,15 hexahydro-geranylgeraniol from farnesyl acetone.

hydrogenator for 2 d. Analysis by TLC (7:3 hexanes/diethyl ether) and GC-MS should indicate that the material is fully converted to the hexahydrofarnesyl acetone. 2. Filter the reaction mixture over filter paper, wash with ethanol, rotary-evaporate to dryness, and take up in a small volume of ethanol, then filter over Fluorosil to remove the last traces of catalyst. Concentrate the resultant oil to dryness with a rotary evaporator (yield 8.64 g, 81% recovery). 3. Dissolve 1.56 g hexahydrofarnesyl acetone in 10 g dry dimethylformamide in a round-bottom flask submerged in ice water and equipped with a dropping funnel containing 1.37 mL triethyl phosphonoacetone. Flush the entire apparatus with argon (Ar). Add triethyl phosphonoacetone over the course of 45 min, followed by 2.2 mL 21% (w/w) sodium ethoxide in ethanol, which is added over the course of 1 h.

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4. Remove the mixture from the ice water bath and continue to stir for 48 h under Ar. Analysis by TLC (7:3 hexanes/diethyl ether) should show that the majority of the starting material (Rf 0.5) has reacted (final product Rf 0.6). 5. Transfer the reaction mixture to a separatory funnel with hexane, and wash 2× with a NaCl-saturated solution. Dry the organic layer with MgS04, pass through filter paper, and evaporate the solvent to dryness by rotary evaporation (yield 1.41 g, 72% recovery). 6. Dissolve 0.75 g of the product from the previous step in 10 mL anhydrous diethyl ether in a round-bottom flask equipped with a stir bar. Cool the apparatus by stirring in an ice-water bath. 7. Add 267 mg of LiAlH4, and stir the reaction mixture overnight under Ar. The following day, add 25 mL saturated NH4Cl and stir the reaction mixture overnight under Ar. 8. Transfer the reaction mixture to a separatory funnel with diethyl ether and wash with water and saturated NaCl solution. Dry the organic phase with MgSO4, filter through filter paper and concentrate to dryness (yield 440 mg, 51% recovery). 9. Purify the material by TLC (7:3 hexanes–diethyl ether, Rf 0.25), or use in the crude state if proceeding on to make radiolabeled material.

3.1.2. Synthesis of cis-Isomers 1. Form the cis-isomers of the allylic alcohols from the respective trans-alcohols after oxidation to the aldehyde (see Subheading 3.1.3.). Allow the mixture of cis/ trans isomers to come to equilibrium (approx 36 cis:65 trans) by standing at room temperature for several days. 2. Purify the cis-isomer by TLC (7:3 hexanes/diethyl ether).

3.1.3. Tritium Labeling of Prenols 1. The method for radiolabeling the prenols is outlined in Fig. 3. First, oxidize the alcohol to the aldehyde with an excess of MnO2. Dissolve 44.7 mg of hexahydrogeranylgeraniol in 2 mL benzene, and add 517 mg of MnO2. Mix the reaction by tissue culture rotator overnight. 2. The next day, allow the reaction mixture to settle, remove the liquid phase, and filter over glass wool to remove any remaining MnO2. Add diethyl ether to the original reaction vessel, and repeat the process. Combine the two solutions, and concentrate to dryness under a stream of nitrogen. 3. Chromatograph the material by TLC (7:3 hexanes/diethyl ether, Rf 0.35). Two overlapping bands are present corresponding to the cis- and trans-double bond isomer, the upper band being the cis-isomer and the lower band being the trans. 4. Remove the lower one-third of the overlapping bands, elute with diethyl ether, dry with MgSO4, and concentrate to dryness (yield 20%). If not used immediately, this material should be stored at −70°C to prevent isomerization of the trans-aldehyde to the equilibrium mixture (~30% cis). 5. Elute the remaining material, and rechromatograph if desired. 6. Reduce the aldehyde to the alcohol using [3H]NaBH4. Dissolve 2 mg of the aldehyde in 1 mL absolute ethanol, and add 5 µL 14 N NH4OH. Dissolve the [3H]NaBH4

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Fig. 3. Schematic for the tritium labeling of naturally occurring prenols.

in ethanol at a concentration of 100 mCi/mL. Add 200 µL of the [3H]NaBH4 to the aldehyde solution, shake the solution, and then leave it vented in the hood for at least 4 h. 7. Concentrate the material to dryness in a gentle stream of Ar, take up in diethyl ether, and place over a column of silica gel overlaid with MgSO4. Purify the material by TLC (7:3 hexanes/diethyl ether) to remove unreacted starting material and the small amount of the cis-isomer that is generated during the reaction. Elute the labeled alcohols from the silica using diethyl ether. Dry using a stream of Ar. 8. Dissolve the product in ethanol, and bring to a final concentration of 2.1 mM, 2.6 mM, 6.5 mM, or 0.4 mM for labeled 2 cis-GGOH, hexahydrogeranylgeraniol, geraniol, and tetrahydrofarnesol, respectively.

3.2. Cell Culture Experiments 1. Grow HSF and 3T3 cells in monolayers, and maintain in 100-mm Petri dishes at 37°C in a humidified atmosphere of 95% air, 5% CO2 in DMEM, pH 7.4, supplemented with 10% FCS v/v, 1% (v/v) NEAA, penicillin (100 U/mL), and streptomycin (0.1 mg/mL). 2. Dissociate confluent stock cultures with 0.05% trypsin-0.02% EDTA, and seed HSF cells (3 × 105 cells/35-mm Petri dish) or 3T3 cells (2 × 105 cells/35-mm Petri dish) in a medium containing 0.4% FCS or 1% PDS, respectively, to stop cell replication. 3. HSF cells become quiescent within 3 d and the experiments can begin on d 4. For 3T3 cells, change the medium on d 2 and 4; cells become quiescent within 5 d, and the experiments can begin on d 6.

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4. At this time, stimulate the cells by replacing the medium with one containing 10% FCS, in the presence or absence of the tested compounds, and continue the incubation as needed at 37°C. Simvastatin is used at a final concentration of 40 µM, when required. 5. Dissolve unlabeled prenols in absolute ethanol and prepare stock solutions as follows: a. 2 mM all-trans-GGOH (the density for this and all other prenols used in these studies is d20 0.89 g/mL). Add 30 µL of the prenol to 49 mL ethanol and store in 1-mL aliquots at −20°C. On the day of the experiment, dilute an aliquot with ethanol (e.g., 300 µL of GGOH to 900 µL of ethanol) to obtain a working solution of 0.5 mM. b. 4 mM all-trans-FOH. Add 20 µL of the prenol to 20 mL of ethanol and store in 1-mL aliquots at −20°C. On the day of the experiment, dilute this 1:3 with ethanol to make 1 mM working stock solution. c. 4 mM geraniol. Add 20 µL of the prenol to 28.83 mL ethanol and store in 1-mL aliquots at −20°C. On the day of the experiment, dilute this 1:3 with ethanol to make 1 mM working stock solution. 6. Prenols are light-sensitive, so experiments should be performed under dim light, and incubation should be done with Petri dishes covered with foil. Be careful not to exceed 1% (v/v) ethanol in the culture medium. 7. For estimation of DNA synthesis after mitogenic stimulation, incubate cells for 22–24 h at 37°C then change the medium to one containing 10% FCS and 2 µCi/ mL [3H]-thymidine and incubate the cells for another 2 h at 37°C (19). Remove the medium and wash the monolayers once with PBS at room temperature, then add 2 mL of fresh, ice-cold 5% TCA (w/v), and keep the cells at 4°C on ice for at least 10 min. Remove TCA and wash once with 5% TCA, and dissolve the monolayers in 1 mL of 1 N NaOH for 15 min. Transfer 500 µL of the cell lysates to a liquid scintillation vial, add 150 µL glacial acetic acid and 5 mL Aquasol, and measure incorporated radioactivity in a liquid scintillation counter (Table 1; Fig. 4).

3.3. Cell Labeling and Prenylated Protein Analysis 3.3.1. Labeling of Proteins with [3H] Farnesol or [3H] Geranylgeraniol and One-Dimensional SDS-PAGE Analysis 1. Dry 9 mCi [3H]-MVA, 500 µCi[3H]-GGOH, or 1 mCi [3H]-FOH, and resuspend in 50 µL ethanol, to bring to the concentration of 4.2 mM, 166 µM, or 1.1 mM, respectively. 2. Incubate the cells (3 × 35-mm Petri dishes/sample) (see Notes 2–6) for 20 h at 37°C, with the appropriate labeled isoprenoids. Incubation periods as short as 5 h are sufficient to label most proteins, making it possible to perform time course experiments. 3. To harvest the cells, place the Petri dishes on ice, remove the medium, and wash 3× with ice-cold PBS containing 1 mM PMSF. Scrape the cells into 1.5 mL PBS/PMSF per Petri dish, collect the resuspended cells, and centrifuge for 5 min at 200g.

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Table 1 Mevalonate, Prenols, and Synthetic Analogs Vary in Their Ability to Prevent the Inhibitory Effect of Simvastatin on DNA Synthesisin Cultured Cellsas Measured by Incorporation of [3H]-Thymidine Cell type

HSF

3T3

Treatment Conditions:

(Reported as % control) 30 100 85 29 37 29 34 N.A.

30 90 96 30 33 30 46 36

Simvastatin (S) 40 µM S+ mevalonate 100 µM S+ all-trans-GGOH 5 µM S+ 2-cis-GGOH 5 µM S+ all-trans-FOH 10 µM S+ geraniol 5 mM S+ 6,7,10,11,14,15-hexahydro-GGOH 5 µM S+ tetrahydro FOH 5 µM

Experimental conditions: see Subheading 3.2. N.A., not assayed. The mean value of control (100%) without inhibitor was 104 × 103 ± 3 × 103 dpm/plate and 187 × 103 ± 10 × 103 dpm/plate for HSF and 3T3 cells, respectively.

4. Aspirate the PBS, add 150 µL PBS/PMSF to the cell pellet, and sonicate in a bath sonicator to disrupt the cell pellet. Transfer the resuspended pellet to a 1.5-mL Eppendorf tube. 5. Add 1.3 mL of cold (−20°C) acetone, mix, sonicate, and allow to stand on ice for 15 min. Centrifuge for 5 min (13,000 g at 4°C) to sediment the delipidated proteins. Re-extract the protein pellet twice with 1 mL cold acetone. 6. Add 1 mL chloroform:methanol (2:1), mix, sonicate, and incubate for 30 min at 37°C. Centrifuge for 5 min (13,000 g, 4°C), and collect the lipid extracts. 7. Remove the residual organic solvent by evaporation, and solubilize the delipidated proteins overnight at room temperature in 100 µL 3% SDS, 62.5 mM TrisHCl, pH 6.8. 8. Determine the protein content in 10 µL of sample, according to Lowry (27). 9. Transfer 10 µL of the sample to a liquid scintillation vial, add 5 mL Aquamix, and measure the incorporated radioactivity. 10. Add 80 µL of sample application buffer to the remainder, and analyze an aliquot (15–80 µg of protein) by one-dimensional SDS-PAGE, according to Laemmli (28), using a 12% gel. 11. After electrophoresis, wash the gel with destain (methanol–MilliQ water–acetic acid, 25:65:10) for 10 min, then stain the gel (0.1% Coomassie R250 in methanol– MilliQ–acetic acid, 40:50:10) for 20 min, and destain overnight. 12. Treat the gel with Amplify for 30 min in preparation for fluorography. Wash the gel twice with water, dry, and expose to Kodak XOMAT-AR film at −70°C (Figs. 5 and 6).

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Fig. 4. The ability of simvastatin-treated HSF cells to synthesize DNA was used as a measure of their ability to traverse the cell cycle. HSF cells were seeded at a density of 3 × 105/35-mm dish in medium supplemented with 0.4% FCS, and the cultures were incubated for 72 h. Quiescent cells were incubated for 24 h in fresh medium containing 10% FCS, 40 µM simvastatin, and the indicated concentrations of unlabeled GGOH or [3H]-GGOH (15 Ci/mmol). Control cells received only 10% FCS. (Left panel) the labeled cell lysates from each treatment group were analyzed by SDS-PAGE (see Subheading 3.3., step 1), the gel was fluorographed, and the resulting films analyzed by densitometry. The optical density reported for each treatment group represents the sum of all protein bands between 20 and 30 kDa observed in the corresponding gel lane (see insert). An equal amount of cell lysate (15 µg cell protein/lane) was applied to each lane. In the experiment shown, 9 × 103, 12 × 103, 42 × 103, and 142 × 103 cpm/lane were analyzed for the 0.5, 1, 2.5, and 5 µM [3H]-GGOH samples, for lanes left to right respectively. (Right panel) In a parallel experiment, DNA synthesis was determined for each treatment group. Cells were incubated as above, but with a corresponding concentration of unlabeled GGOH. After 22 h, [3H]-thymidine (2 µCi/mL) was added to treated and control samples, and the incubation was continued for another 2 h. Nuclear DNA was analyzed for incorporation of [3H]-thymidine (see Subheading 3.2., step 7). The incorporation of [3H]-thymidine for each treatment group is reported as a percentage of the control. The mean value of control (100%) was 114 × 103 ± 6 × 103 dpm/plate.

3.3.2. High-Resolution Two-Dimensional Gel Electrophoresis of Labeled Proteins 1. In 2DE analysis, proteins are first separated on the basis of pI, using isoelectric focusing (IEF), followed by separation on the basis of mol mass. Therefore, differentiation of closely related proteins with similar mol masses is facile (29,30). Two different IEF methods have been developed for use in high-resolution

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Fig. 5. Metabolic labeling of prenylated proteins in Swiss 3T3 cells after various treatments. Cells were seeded at a density of 2 × 105/dish in medium containing 1% PDS and the cultures incubated for 5 d. Quiescent cells were incubated for 20 h in fresh medium containing 10% FCS, 50 µM [3H]-MVA (35 Ci/mmol), 2.5 µM [3H]-FOH (15 Ci/mmol), or 1 µM [3H] GGOH (60 Ci/mmol), in the presence or absence of 40 µM simvastatin. Cell pellets were delipidated, and equal amounts of cell extracts (80 µg cell protein/lane) were separated by 12% SDS-PAGE and fluorographed. In the experiment shown, 8 × 108, 2.7 × 107, 6 × 107 cpm/lane were analyzed for cells labeled with [3H]-MVA, [3H]-FOH or [3H]-GGOH, respectively.

2DE. The one described here uses carrier ampholytes (31–33) to obtain the appropriate pH gradient in which sample proteins are focused. This system is available from Genomics Solutions, Chelmsford, MA. The second IEF method uses precast gel strips containing an immobilized pH gradient made by crosslinking ampholytes into a gel matrix (29,34). This system is available from Amersham Pharmacia, Piscataway, NJ (see Notes 7–10 for additional comments). 2. Prior to analysis by 2DE, cells are labeled as described in Subheading 3.3.1.

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Fig. 6. Metabolic labeling of prenylated proteins in Swiss 3T3 cells after various treatments. Experimental conditions as in Fig. 5. Quiescent cells were incubated for 20 h in medium containing 10% FCS, 40 µM simvastatin, and one of the following: lane 1, 5 µM [3H]-all transGGOH (20 Ci/mmol); lane 2, 5 µM [3H]-cis, trans-GGOH (17.5 Ci/mmol); lane 3, 5 µM [3H]-6,7,10,11,14,15 hexahydrogeranylgeraniol (17.5 Ci/mmol); lane 4, 5 µM [3H]-tetrahydrofarnesol (17.5 Ci/mmol); and lane 5, 5 µM [3H]-geraniol (17.5 Ci/mmol). Cell pellets were delipidated, and equal amounts of cell extract (60 µg cell protein/lane) were analyzed by SDSPAGE on 12% gels, and gels were fluorographed. In the experiment shown, 334 × 103, 197 × 103, 247 × 103, 271 × 103, 285 × 103 cpm/lane were analyzed for lanes 1–5, respectively.

3. On ice, wash each 100-mm dish once with 10 mL PBS, then twice with a solution containing 10 mM Tris buffer, pH 7.5, 0.1 mM PMSF, and 1.0 µg/mL each of aprotinin, leupeptin, and pepstatin A. Drain for 45 s, and remove residual buffer. 4. Add 240 µL of boiling sample buffer (28 mM Tris-HCl, and 22 mM Tris base, 0.3% SDS, 200 mM DTT, pH 8.0), and collect cells with a cell scraper. Transfer lysate to a 1.5-mL microcentrifuge tube, boil for 5 min, and cool on ice. This and subsequent steps are a modification of methods previously described (32,33). 5. Add 30 µL (or one-tenth vol) of protein precipitation buffer (24 mM Tris base, 476 mM Tris-HCl, 50 mM MgCl2, 1 mg/mL DNaseI, 0.25 mg/mL RNaseA, pH 8.0) to each cell lysate sample, and incubate on ice for 8 min. Subsequent steps are performed at room temperature, unless otherwise indicated.

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6. Add ~6000 cpm of the internal standard, [3H]NEM-labeled soybean trypsin inhibitor (preparation described in Subheading 3.3.3.), to each sample (see Note 10). 7. Add 4 vol of acetone at room temperature to each lysate, and let stand for 20 min. Centrifuge the protein precipitate for 10 min at 16,000 g. 8. Remove the acetone phase, resuspend the precipitate in 800 µL fresh acetone, using a bath sonicator, and centrifuge as above for 5 min. Repeat this wash step once. 9. Let the pellet air dry for 3 min (avoid overdrying), before adding 30 µL of a 1:4 mixture of the SDS buffer (from Subheading 3.2.2., step 4) and urea solution (9.9 M urea; 4% Triton X-100; 2.2% ampholytes, pH 3–10; 100 mM DTT). Warm the sample to 37°C, and bath-sonicate to solubilize the protein (avoid overheating the sample). Centrifuge for 5 min at 16,000 g. Set aside 1 µL for scintillation counting and 1 µL for Bradford protein assay. 10. Apply the sample (26 µL) onto a 1 × 180 mm-tube gel (4.1% T, 0.35% C; ampholyte pH range of 3.0–10.0). Focus for 17.5 h at 1000 V, then for 0.5 h at 2000 V. 11. Extrude the gel from the tube into equilibration buffer (300 mM Tris base, 75 mM Tris-HCl, 3% SDS, 50 mM DTT, and 0.01% bromophenol blue), and incubate it for 2 min before overlaying onto the second dimension SDS-PAGE gel (12.5% T, 0.27% C). 12. Perform SDS-PAGE on large format gels (220 × 240 × 1 mm) for 6 h, using the constant power mode (16 W/gel) in a prechilled tank equipped with Peltier cooling (Genomics Solutions). When five gels are used in this mode, the running buffer temperature will increase 14°C over the course of 6 h when started at 0°C. 13. Fix gels for 1 h in methanol:Milli Q:acetic acid (50:40:10), then stain overnight with a solution containing 0.001% Coomassie R250 in methanol:Milli Q:acetic acid (25:65:10). 14. Immerse each stained gel in 200 mL of Amplify, and incubate for 20 min, with rocking. Without rinsing, dry the treated gel onto heavyweight blotting paper and expose to preflashed Hyperfilm MP for 7–14 d at −70°C. Exposures of >10 wk may be needed to visualize low abundance labeled proteins (Fig. 7A,B).

3.3.3. Preparation of the Two-Dimensional Gel Internal Standard, [3H]NEM-Labeled Soybean Trypsin Inhibitor 1. Transfer 100 µL of a freshly prepared solution of 22 mM N-ethylmaleimide (NEM) in acetonitrile to a 0.5-mL Reacti-Vial, and add 250 µL [3H]NEM (1 µCi/µL in pentane; 56 Ci/mmol). Mix, and let stand for 2 h uncapped in a fume hood to evaporate the pentane. The final volume will be 110 µL, and the new specific activity will be 95.4 mCi/mmol. 2. Prepare 1 mg/mL solution of soybean trypsin inhibitor (STI) in 50 mM Tris, pH 8.5, and reduce the protein by adding DTT (2 mM, final concentration). Incubate overnight at 4°C. 3. Remove excess DTT by applying 300 µL of the reduced STI (in six 50-µL aliquots) to six SW rotor equibbed, spin columns, and centrifuge for 4 min at 1000 g in a preequilibrated countertop centrifuge. Prepare spin columns by placing 1 mL

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bed-volume of Bio Gel P6-DG (previously rehydrated in 50 mM Tris HCl, pH 7.5) into a 1-mL plastic tuberculinsyringe equipped with a plug of glass wool, followed by centrifugation for 2 min at 1000 g. (In addition to removing DTT, this step also permits buffer exchange to provide the proper pH required for the NEM reaction step which follows.) 4. Pool the six column eluates (250–300 µL total volume), and add 10–12 µL of the [3H]NEM solution ( 24 µCi; see Subheading 3.3.3., step 1). Incubate for 6 h on ice. 5. Separate the [3H]NEM-labeled STI from the unreacted [3H]NEM using five or six fresh, preequibbelated spin columns (see Subheading 3.3.3., step 3). Pool the column eluates containing the purified [3H]NEM-labeled STI (1 µCi/nmol protein), divide into 30-µL aliquots, and store at −70°C. To make up a working stock solution, dilute the eluate 1:39 with buffer (6000 cpm/5 µL), and use as an internal standard for 2DE gel fluorography (see Subheading 3.3.2., step 6; see also Note 10).

4. Notes 1. Since either all-trans-GGOH or MVA, but not all-trans-FOH, can completely prevent the inhibitory effect of simvastatin on DNA synthesis in cultured HSF and 3T3 cells (Table 1), it may be worthwhile to determine whether this effect is related to the cells’ ability to incorporate labeled isoprenoid derivatives into specific proteins, or into other isoprenoid metabolites. 2. To maximize the incorporation of [3H]-MVA into cellular proteins, it is necessary to block endogenous MVA synthesis with a statin such as simvastatin (35). One-dimensional SDS-PAGE reveals that whole-cell homogenates incorporated [3H]-MVA into at least 10 major bands, with molecular masses ranging from 21 to 72 kDa (Fig. 5). Intense bands of radioactivity are seen in the 21–28 kDa range, corresponding to Ras and Ras-related small GTPases (2,3). In contrast, only a few protein bands are visible when simvastatin is omitted from the incubation. 3. It also is necessary to block endogenous MVA synthesis with simvastatin, for efficient incorporation of [3H]-FOH into specific cellular proteins. Fig. 7. Two-dimensional gel fluorographs of [3H]-prenol-labeled cell-lysates. HSF cells were labeled with either 5 µm [3H]-FOH or 5 µm [3H]-GGOH (see Subheading 3.3.1.), lysed, delipidated and analyzed by 2DE and fluorography (see Subheading 3.3.2.). (A) Whole-cell lysate from [3H]-FOH labeled cells that contained 385 µg protein and 170 × 103 cpm. Film was exposed for 23 wk (B) Whole-cell lysate from [3H]-GGOH-labeled cells that contained 475 µg protein and 390 × 103 cpm. Film was exposed for 10 wk. The major proteins that incorporated both FOH and GGOH are circled. At least nine additional clusters of minor proteins display a similar ability to incorporate either prenol, but have not been circled (see also Note 7). The position of the internal standard, [3H]-NEM-labeled soybean trypsin inhibitor (6250 cpm per gel; see Subheading 3.3.3.), is indicated by a square.

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However, this is not the case for [3H]-GGOH labeling, which proceeds equally well in the presence or absence of simvastatin (Fig. 5). In investigations of the cellular uptake of MVA and its derivatives, the authors typically found uptake of less than 0.1% of the added [3H]-MVA, 2% of the added [3H]-FOH, and more than 10–15% of the added [3H]GGOH. The efficient uptake of GGOH by cells permits rapid labeling of geranylgeranylated proteins. Protein bands from one-dimensional SDSPAGE are detectable after labeling the cells for only 1–2 h, and subsequent gels require only a few days of film exposure. Both [3H]-GGOH labeling of proteins and the GGOH-mediated prevention of simvastatin-induced inhibition of DNA synthesis show parallel doseresponse sensitivities to a similar range of GGOH concentrations (up to 2.5 µM; Fig. 4). The labeling approach described here can also be utilized for investigating natural or synthetic isoprenoid analogs (Fig. 6). When cells were treated with [3H]-2-cis-GGOH (precursor of dolichols), proteins are only weakly labeled, compared to all trans-GGOH. Under parallel conditions, the addition of unlabeled 2-cis-GGOH failed to prevent the inhibitory effect of simvastatin on DNA synthesis (Table 1). When prenylated proteins are selectively labeled with [3H]-FOH or [3H]GGOH, and analyzed by one-dimensional SDS-PAGE, two subsets of proteins can be identified: a subset of 17 protein bands that incorporate [3H]-FOH, and a separate subset of 12 protein bands that incorporate [3H]-GGOH (see Fig. 5). However, when similarly labeled proteins are analyzed by 2DE, which allows much greater resolution, a third subset of proteins is identified. Using this approach, 82 proteins are visualized that incorporate only [3H]-FOH, 34 proteins that incorporate only [3H]GGOH, and 25 proteins that can incorporate either prenol (compare Fig. 7A,B), although with varying efficiencies. The presence of unlabeled GGOH in the media during [3H]-FOH labeling experiments or unlabeled FOH during [3H]-GGOH labeling experiments, does not affect the protein composition of these subsets. As mentioned above (see Subheading 3.3.2., step 1), there are primarily two different methods of IEF available for use in 2DE: one that uses soluble carrier ampholytes (31,32) and one that uses immobilized ampholytes in Immobiline™ gel-strips, discussed in Note 9 below (29,30,34). The analyses presented here were performed using the first method, which was, until recently, the only large format system commercially available. This system is especially convenient for small analytical samples, but can also be used reproducibly for large-sample analyses (33). When 2DE is coupled with immunoanalysis, this system permits the detection of some proteins that cannot be visualized on blots from Immobiline gels

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(M. Aepfelbacker, personal communication; C. Farnsworth, unpublished results). 9. When large samples (>500 µg protein; e.g., cell lysates) are analyzed, the Immobiline system is currently preferable, because of advances in gel technology and sample-handling techniques. These gels are now commercially available in an extended format (0.5 × 3 × 180 mm) and come as dehydrated gel strips bonded to a rigid plastic support. This permits the gel to be rehydrated directly with dilute sample lysates. Additionally, samples containing up to 5 mg protein can be analyzed in this way (36) without the need for concentration by techniques such as protein precipitation. The gels can accommodate a relatively large sample volume (upper limit of 350 µL vs 26 µL for the carrier ampholyte system). Another advantage of immobilized ampholyte pH gradients is that they allow the use of very narrow pH gradients (e.g., pH 5.5–6.5). This permits the separation of charge-isoforms that differ by only pH 0.05 units (29). Lastly, when 2DE is coupled with immunoanalysis and overlay assays, detailed maps of multimember protein families can be constructed (37). 10. In control experiments using lysates of unlabeled cells, recovery of the labeled internal standard, [3H]-NEM STI (see Subheading 3.3.3.), following complete sample workup (see Subheading 3.3.2., steps 6–9) was 91 ± 4%, n = 3. Acknowledgment Alberto Corsini (1993–1999) was visiting scientist under the terms of the US (National Heart, Lung, and Blood Institute)–Italy bilateral agreement in the cardiovascular area.

References 1. Zhang, F. L. and Casey, P. J. (1996) Protein Prenylation: Molecular mechanisms and functional consequences. Annu. Rev. Biochem. 65, 241–269. 2. Glomset, J. A., Gelb, M. H., and Farnsworth, C. C. (1990) Prenyl proteins in eukariotic cells: a new type of membrane anchor. Trends Biochem. Sci. 15, 139–142. 3. Maltese, W. A. (1990) Posttranslational modification of proteins by isoprenoids in mammalian cells. FASEB J. 4, 3319–3328. 4. Glomset, J. A. and Farnsworth, C. C. (1994) Role of protein modification reactions in programming interactions between ras-related GTPases and cell membranes. Annu. Rev. Cell Biol. 10, 181–205. 5. Casey, P. J., Moomaw, J. F., Zhang, F. L., Higgins, J. B., and Thissen, J. A. (1994) Prenylation and G protein signaling, in Recent Progress in Hormone Research, Academic, New York. 6. Inglese, J., Koch, W. J., Touhara, K., and Lefkowitz, R. J. (1995) Gβγ interactions with pH domains and Ras-MPK signaling pathways. Trends Biochem. Sci. 20, 151–156.

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7. Marshall, M. S. (1995) Ras target proteins in eukaryotic cells. FASEB J. 9, 1311–1318. 8. Kato, K., Cox, A. D., Hisaka, M. M., Graham, S. M., Buss, J. E., and Der, C. J. (1992) Isoprenoid addition to Ras protein is the critical modification for its membrane association and transforming activity. Proc. Natl. Acad. Sci. USA 89, 6403–6407. 9. Boguski, M. S. and McCormick, F. (1993) Proteins regulating Ras and its relatives. Nature 365, 643–654. 10. Jakobisiak, M., Bruno, S., Skiersky, J. S., and Darzynkiewicz, Z. (1991) Cell cycle-specific effects of lovastatin. Proc. Natl. Acad. Sci. USA 88, 3628–3632. 11. Taylor, S. J. and Shalloway, D. (1996) Cell cycle-dependent activation of Ras. Curr. Biol. 6, 1621–1627. 12. Olson, M. F., Ashworth, A., and Hall, A. (1995) An essential role for Rho, Rac, and Cdc42 GTPases in cell cycle progression through G1. Science 269, 1270–1272. 13. Fenton, R. G., Kung, H., Longo, D. L., and Smith, M. R. (1992) Regulation of intracellular actin polymerization by prenylated cellular proteins. J. Cell Biol. 117, 347–356. 14. Pittler, S. J., Fliester, S. J., Fisher, P. L., Keller, R. K., and Rapp, L. M. In vivo requirement of protein prenylation for maintenance of retinal cytoarchitecture and photoreceptor structure. J. Cell Biol. 130, 431–439. 15. Tapon, N. and Hall, A. (1997) Rho, Rac, and Cdc42 GTPases regulate the organization of the actin cytoskeleton. Curr. Opin. Cell Biol. 9, 86–92. 16. Zerial, M. and Stenmark, H. (1993) Rab GTPases in vesicular transport. Curr. Opin. Cell Biol. 5, 613–620. 17. Novick, P. and Brennwald, P. (1993) Friends and family: the role of the Rab GTPases in vesicular traffic. Cell 75, 597–601. 18. Raiteri, M., Arnaboldi, L., McGeady, P., Gelb, M. H., Verri, D., Tagliabue, C., Quarato, P., et al. (1997) Pharmacological control of the mevalonate pathway: effect on smooth muscle cell proliferation. J. Pharmacol. Exp. Ther. 281, 1144–1153. 19. Habenicht, A. J. R., Glomset, J. A., and Ross, R. (1980) Relation of cholesterol and mevalonic acid to the cell cycle in smooth muscle and Swiss 3T3 cells stimulated to divide by platelet-derived growth factor. J. Biol. Chem. 255, 5134–5140. 20. Goldstein, J. L. and Brown, M. S. (1990) Regulation of the mevalonate pathway. Nature 343, 425–430. 21. Corsini, A., Mazzotti, M., Raiteri, M., Soma, M. R., Gabbiani, G., Fumagalli, R., and Paoletti, R. (1993) Relationship between mevalonate pathway and arterial myocyte proliferation: in vitro studies with inhibitor of HMG-CoA reductase. Atherosclerosis 101, 117–125. 22. Crick, D. C., Waechter, C. J., and Andres, D. A. (1994) Utilization of geranylgeraniol for protein isoprenylation in C6 glial cells. Biochem. Biophys. Res. Commun. 205, 955–961. 23. Crick, D. C., Andres, D. A., and Waechter, C. J. (1995) Farnesol is utilized for protein isoprenylation and the biosynthesis of cholesterol in mammalian cells. Biochem. Biophys. Res. Commun. 211, 590–599.

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24. Danesi, R., McLellan, C. A., and Myers, C. E. (1995) Specific labeling of isoprenylated proteins: application to study inhibitors of the posttranslational farnesylation and geranylgeranylation. Biochem. Biophys. Res. Commun. 206, 637–643. 25. Shin-ichi, O., Watanabe, M., and Nishino, T. (1996) Identification and characterization of geranylgeraniol kinase and geranylgeranyl phosphate kinase from the Archebacterium Sulfolobus acidocaldarius. J. Biochem. 119, 541–547. 26. Westfall, D., Aboushadi, N., Shackelford, J. E., and Krisans, S. (1997) Metabolism of farnesol: phosphorylation of farnesol by rat liver microsomial and peroxisomal fractions. Biochem. Biophys. Res. Commun. 230, 562–568. 27. Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) Protein measurement with the folin phenol reagent. J. Biol. Chem. 193, 265–275. 28. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 29. Görg, A., Postel, W., and Günther, S. (1988) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 9, 531–546. 30. Jungblutt, P., Thiede, B., Simny-Arndt, U., Müller, E.-C., Wittmann-Liebold, B., and Otto, A. (1997) Resolution power of two-dimensional electrophoresis and identification of proteins from gels. Electrophoresis 17, 839–847. 31. O’Farrell, P. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–4021. 32. Patton, W. F., Pluskal, M. G., Skea, W. M., Buecker, J. L., Lopez, M. F., Zimmermann, R., Lelanger, L. M., and Hatch, P. D. (1990) Development of a dedicated two-dimensional gel electrophoresis system that provides optimal pattern reproducibility and polypeptide resolution. BioTech. 8, 518–529. 33. Lopez, M. F. and Patton, W. F. (1990) Reproducibility of polypeptide spot positions in two-dimensional gels run using carrier ampholytes in the isolelectric focusing dimension. Electrophoresis 18, 338–343. 34. Görg, A., Günther, B., Obermaier, C., Posch, A., and Weiss, W. (1995) Twodimensional polyacrylamide gel electrophoresis with immobilized pH gradients in the first dimension (IPG-Dalt): the state of the art and the controversy of vertical versus horizontal systems. Electrophoresis 16, 1079–1086. 35. Schmidt, R. A., Schneider, C. J., and Glomset, J. A. (1984) Evidence for posttranslational incorporation of a product of mavalonic acid into Swiss 3T3 cell proteins. J. Biol. Chem. 259, 10,175–10,180. 36. Sanchez, J.-C., Rouge, V., Pisteur, M., Ravier, F., Tonella, L., Moosmayer, M., Wilkins, M. R., and Hochstrasser, D. F. (1997) Improved and simplified in-gel sample application using reswelling of dry immobilized pH gradients. Electrophoresis 18, 324–327. 37. Huber, L. A., Ullrich, O., Takai, Y., Lütcke, A., Dupree, P., Olkkonen, et al. (1994) Mapping of Ras-related GTP-binding proteins by GTP overlay following twodimensional gel electrophoresis. Proc. Natl. Acad. Sci. USA 91, 7874–7878.

168 The Metabolic Labeling and Analysis of Isoprenylated Proteins Douglas A. Andres, Dean C. Crick, H. Peter Spielmann, and Charles J. Waechter 1. Introduction 1.1. Background The posttranslational modification of proteins by the covalent attachment of farnesyl and geranylgeranyl groups to cysteine residues at or near the carboxyl(C)-terminus via a thioether bond is now well established in mammalian cells (1–6). Most isoprenylated proteins are thought to serve as regulators of cell signaling and membrane trafficking. Farnesylation and geranylgeranylation of the cysteinyl residues has been shown to promote both protein–protein and protein–membrane interactions (3,7,8). Isoprenylation, and in some cases the subsequent palmitoylation, provide a mechanism for the membrane association of polypeptides that lack a transmembrane domain, and appear to be prerequisite for their in vivo activity (3,9,10). Three distinct protein prenyltransferases catalyzing these modifications have been identified (1,2,4–6). Two geranylgeranyltransferases (GGTases) have been characterized and are known to modify distinct protein substrates. The CaaX GGTase (also known as GGTase-1) geranylgeranylates proteins that end in a CaaL(F) sequence, where C is cysteine, A is usually an aliphatic amino acid, and the C-terminal amino acyl group is leucine (L) or phenylalanine (F). Rab GGTase (also known as GGTase-2) catalyzes the attachment of two geranylgeranyl groups to paired C-terminal cysteines in most members of the Rab family of GTP-binding proteins (11). These proteins terminate in a Cys-Cys, Cys-X-Cys, or Cys-Cys-X-X motif where X is a small hydrophobic amino acid. Another set of regulatory proteins is modified by protein farnesyltransferase (FTase). All known farnesylated proteins terminate in a tetrapeptide CaaX box, wherein From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. Proposed “salvage” pathway for the utilization of F-OH and GG-OH for isoprenoid biosynthesis. Evidence for the presence of microsomal enzymes catalyzing the conversion of F-P-P and GG-P-P to the free isoprenols has been reported by Bansal and Vaidya (13).

C is cysteine, A is an aliphatic amino acid, and X has been shown to be a COOHterminal methionine, serine, glutamine, cysteine, or alanine. Isoprenylated proteins are commonly studied by metabolic labeling of cultured cells by incubation with [3H]mevalonate which is enzymatically converted to [3H]farnesyl pyrophosphate (F-P-P) and [3H]geranylgeranyl pyrophosphate (GG-P-P) prior to being incorporated into protein (ref. 12; Fig. 1). Cellular proteins can then be analyzed by gel electrophoresis and autoradiography. However, [3H]mevalonate labeling does not distinguish between farnesylated and geranylgeranylated proteins directly. Other methods for the identification of the isoprenyl group attached to protein that make use of [3H]mevalonolactone labeling have been described. These methods generally require the isolation of large amounts of labeled protein, extensive proteolysis, a number of column chromatographic steps, and cleavage of the thioether bond with Raney nickel or methyl iodide, followed by gas chromatography or HPLC and mass spectrometry to identify the volatile cleavage products (see ref. 4 and papers cited therein). This chapter describes the use of either the natural free [3H]farnesol (F-OH) and [3H]geranylgeraniol (GG-OH) or the unnatural and non-radioactive anilinogeraniol (AGOH) in the selective metabolic labeling of farnesylated and geranylgeranylated proteins in cultured mammalian cells. In addition, the methods use materials and equipment readily available in most laboratories. We have recently shown that mammalian cells can utilize the free isoprenols, GG-OH and F-OH, for the isoprenylation of cellular proteins (14,15). When C6

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glioma cells were incubated with [3H]F-OH, radioactivity was also incorporated into cholesterol (14). The observation that the incorporation of label into sterol was blocked by squalestatin 1 (SQ), a potent inhibitor of squalene synthetase (16–20), suggested that F-OH, and probably GG-OH, are utilized for isoprenoid biosynthesis after being converted to the corresponding activated allylic pyrophosphates, F-P-P and GG-P-P (Fig. 1). Preliminary studies have suggested the presence of enzyme systems in mammals and lower organisms that are capable of phosphorylating F-OH and GG-OH (21–23). More recently microsomal fractions from N. tabacum have been shown to contain CTP-mediated kinases that catalyze the conversion of F-OH and GG-OH to the respective allylic pyrophosphate intermediates by two successive monophosphorylation reactions (24). Further work is certainly warranted on the isolation and characterization of these enzymes. The early developments in understanding the mechanism and physiological significance of the salvage pathway for the utilization of F-OH and GG-OH have been reviewed (25). A drawback to all metabolic labeling approaches to examine protein isoprenylation is the inherently low sensitivity of autoradiographic detection of [3H]labeled proteins and the limitation that incorporation of radiolabeled prenyl groups does not provide a convenient method for the isolation of the modified proteins. To address these limitations, incorporation of unnatural F-P-P analogues has been utilized to examine the prenylation status of cellular proteins. These methods provide a facile approach to both the detection and isolation of modified proteins (26–28). These methods include a tagging-via-substrate (TAS) approach using in vivo incorporation of 8-azido-farnesol (27) in conjunction with a biotinylated phosphine capture reagent to isolate modified proteins, which are then identified by mass spectroscopy. However, the TAS technology requires access to specialized equipment and thus does not lend itself to the routine detection of prenylated proteins. Here, we describe a more convenient method to simplify the detection of unnatural F-P-P modified proteins, making use of analogue-selective antibodies. In particular, recent studies have shown that the unnatural F-P-P analogue anilinogeranyl diphosphate (AG-P-P) is transferred by FTase to protein substrates with kinetics similar to F-P-P, that AG-OH is effectively utilized in a wide range of cells for the in vivo labeling of farnesylated proteins, and does not resemble any known natural protein modification (27,29,30). More importantly, anilinogeranyl modified proteins can be readily detected with anti-anilinogeranyl antibodies (28). Thus, the unnatural FPP analog and corresponding antibodies provide a simple and rapid method for monitoring FTase activity in cells and detection of cellular proteins modified by AG-OH. 1.2. Experimental Strategy Utilizing free F-OH or GG-OH as the isotopic precursors of F-P-P and GGP-P, or AG-OH as an unnatural alternative FTase substrate, has several experimental advantages over metabolic labeling of isoprenylated proteins with

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mevalonate. First, isoprenoid alcohols are more hydrophobic and rapidly enter cultured mammalian cells. They are efficiently utilized in a range of mammalian cell lines, and obviate the need to include HMG-CoA reductase inhibitors to lower endogenous pools of mevalonate. The experimental strategy is illustrated in Fig. 1. A key advantage of the strategy is that F-OH, AGOH and GG-OH are selectively incorporated into distinct subsets of isoprenylated proteins, providing a simple and convenient approach to specifically label farnesylated or geranylgeranylated proteins (Figs. 2 and 8).

Fig. 2. SDS-PAGE analysis of proteins labeled by incubating C6 glioma cells and CV-1 cells with [3H]mevalonate, [3H]F-OH, or [3H]GG-OH. The details of the metabolic labeling procedure and SDS-PAGE analysis are described in Subheadings 3.1.–3.3. For these analyses proteins were metabolically labeled by incubating the indicated cultured cells with [3H]F-OH in the presence of lovastatin (5 µg/mL) because it increased the amount of radioactivity incorporated into protein during long-term incubations, presumably by reducing the size of the endogenous pool of F-P-P. The gel patterns reveal that distinctly different sets of proteins are labeled by each precursor in C6 and CV-1 cells. Consistent with selective labeling by [3H]F-OH and [3H]GG-OH, the labeling pattern for [3H]mevalonate, which can be converted to 3H-labeled F-P-P and GG-P-P, appears to be a composite of the patterns seen with the individual [3H]isoprenols. SMG = proteins in the size range (19–27 kDa) of small GTP-binding proteins. (Figure reprinted with permission from ref. 14).

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Fig. 3. Experimental scheme for the rapid analysis of metabolically labeled isoprenylated cysteine residues labeled by various isotopic precursors. The experimental details of this procedure are described in Subheading 3.4.

Following metabolic labeling using [3H]F-OH, [3H]GG-OH, or [3H]mevalonolactone, the metabolically labeled proteins are exhaustively digested with Pronase E to liberate the specific isoprenyl-cysteine residues (Fig. 3). The identity of the isoprenylated cysteine residue can then be readily identified by normal or reverse-phase thin layer chromatography (TLC). Figs. 4 and 5 show representative TLC analysis of isoprenyl-cysteines released from metabolically labeled cellular proteins and from recombinant proteins that were isoprenylated in vitro. Alternatively, once the non-radioactive AG-OH is incorporated in cellular proteins, total cell lysates can be directly subjected to immunoblotting with anti-AG antibodies to detect AG-modified endogenous proteins (Fig. 8). The anilinogeranyl moiety is structurally unrelated to any other known cellular components or protein modifications making it an excellent epitope for selective antibody recognition (Troutman Bioconj Chem 2005REF). The AG-P-P prodrug anilinogeraniol (AG-OH) is incorporated into cellular protein in the presence of the endogenous F-P-P pool without the requirement for blocking endogenous FPP formation and is remarkably non-toxic (28). Thus, AG-OH and the corresponding antibodies provide a simple, rapid, and nonradioactive method for monitoring FTase activity and detection of cellular proteins modified by AG-OH. The isoprenol labeling and Pronase E methods may also be applied to the analysis of individual metabolically labeled proteins. These methods provide a simple and convenient approach for the identification of the isoprenyl group found on a specific protein. Two experimental approaches are available. In the first, separate cell cultures are incubated with [3H]mevalonate, [3H]F-OH, or [3H]GG-OH the metabolically labeled protein of interest is then purified and subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis. The specificity of the F-OH and GG-OH incorporation allows the identity of the prenyl group to be directly assessed (Fig. 6). In the second, the cells are metabolically labeled with [3H]mevalonolactone and the isolated protein of interest is subjected to Pronase E treatment followed

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Fig. 4. Chromatographic analysis of isoprenyl-cysteine residues metabolically labeled by incubating C6 glioma cells with, [3H]F-OH (upper panel), [3H]GG-OH (middle panel) or [3H]mevalonate (lower panel). From the traces illustrated, it can be seen that F-Cys and GG-Cys are both metabolically labeled when C6 cells are incubated with [3H]mevalonate, but only F-Cys or GG-Cys are labeled when C6 cells are incubated with [3H]F-OH or [3H] GG-OH, respectively. Virtually identical results were obtained by chromatographic analysis of the Pronase E digests of CV-1 proteins metabolically labeled by each isotopic precursor. Radiolabeled products are also observed at the origin of the TLC which could be incompletely digested isoprenylated peptides, or in the case of [3H]mevalonate and [3H]GG-OH, possibly mono- or digeranylgeranylated Cys-Cys or Cys-X-Cys sequences reprinted with permission (14) (see Fig. 5).

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Fig. 5. Chromatographic analyses of isoprenyl-cysteine residues liberated by Pronase E digestion of recombinant protein substrates enzymatically labeled in vitro by [3H]F-P-P or [3H]GG-P-P. The recombinant proteins were labeled by incubation with recombinant FTase (upper panel), GGTase I (upper panel) or GGTase II (lower panel), essentially under the conditions described previously (18).

by TLC analysis of the butanol-soluble products. In this case analysis the chromatogram reveals the nature of the isoprenyl group (Fig. 7). The experimental methods for these analytical procedures are presented in detail in this chapter.

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Fig. 6. Specific metabolic labeling of RDJ2-transfected HEK cells. (A) Monolayers of human embryonic kidney HEK cells transfected with pRDJ2 (an expression plasmid that contained the RDJ2 cDNA under the control of a CMV promoter) were radiolabeled with [3H]mevalonate and the expressed RDJ2 immunoprecipitated from detergent-solubilized cell extracts using a RDJ2-specific antibody. A portion of the resulting immunoprecipitate as well as a portion of the cell extract were subjected to SDS-PAGE. The gel was treated with Amplify, dried, and exposed to film for 14 d. (B) Immunoprecipitated RDJ2 after metabolic labeling of transfected HEK cells with [3H]F-OH (lane 1) or [3H]GG-OH (lane 2) was subjected to SDSPAGE analysis and fluorography. The remaining immunoprecipitated protein fractions isolated from HEK cells after metabolic labeling with [3H]F-OH (lane 3) or [3H]GG-OH (lane 4) were immunoblotted using anti-RDJ2 IgG and subjected to chemiluminescence detection.

Fig. 7. Chromatographic analysis of butanol-soluble products released by Pronase E digestion of RDJ2 protein metabolically labeled by incubation with [3H]mevalonolactone. Immunoprecipitated RDJ2, isolated from [3H]mevalonolactone-labeled HEK cells, was subjected to Pronase E digestion. The labeled products were extracted with 1-butanol, and analyzed by reverse-phase chromatography using C18 reverse-phase TLC plates, and developed in acetonitrile–water–acetic acid (75:25:1). Radioactive zones were located with a Bioscan Imaging System 200-IBM. The arrows indicate the position of authentic F-Cys and GG-Cys.

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Fig. 8. Incorporation and detection of anilinogeranyl modified proteins by immunoblotting. In vitro prenylation reactions were performed using recombinant farnesyltransferase and AGPP in the absence (lane 1) or presence (lane 2) of H-Ras. The reactions were resolved by SDS-PAGE, transferred to nitrocellulose, and subjected to immunoblotting with anti-AG polyclonal antibody (28, 33). In vivo prenylation reactions (lanes 3 and 4) were performed on HEK-293 cells preincubated with 30 µM lovostatin and 10 µM GGOH for 24 h. The media was removed and replenished with media containing either 3 µM FTI-277 (lane 3) or no inhibitor (lane 4) for 1 h. AGOH (100 µM) was then added directly to the cultures and cells incubated for an additional 2 h. Cells were harvested and total cell lysates (50 µg) resolved by SDS-PAGE and anilinogeranyl modified endogenous proteins detected by immunoblotting using anti-AG polyclonal antibody.

2. Materials 2.1. Metabolic Radiolabeling of Mammalian Cells in Culture with [3H]Farnesol, [3H]Geranylgeraniol, and [3H]Mevalonolactone 1. ω,t,t-[1-3H]Farnesol (20 Ci/mmol, American Radiolabeled Chemicals, Inc., St. Louis, MO). 2. ω,t,t,t-[3H]Geranylgeraniol (60 Ci/mmol, American Radiolabeled Chemicals, Inc., St. Louis, MO). 3. [3H]Mevalonolactone (60 Ci/mmol, American Radiolabeled Chemicals, Inc., St. Louis, MO). 4. Appropriate cell culture media, plastic ware, and cell incubator. 5. 95% Ethanol. 6. Serum Supreme (SS), an inexpensive fetal bovine serum (FBS) substitute obtained from BioWhittaker, has been successfully used with C6 glioma, Chinese hamster ovary (CHO) clone UT-2 and green monkey kidney (CV-1) cells in this laboratory. 7. Bath sonicator.

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2.2. Delipidation of Labeled Proteins 1. 2. 3. 4. 5. 6. 7.

Phosphate-buffered saline (PBS). PBS containing 2 mM EDTA. Methanol. Chloroform–methanol (2:1, v/v). 12-mL Disposable conical screw-capped glass centrifuge tubes. Benchtop centrifuge. Probe sonicator.

2.3. SDS-PAGE Analysis of Metabolically Labeled Proteins 1. 2. 3. 4. 5.

SDS-PAGE apparatus. Western blot transfer apparatus. Nitrocellulose membrane (Schleicher and Schuell, Protran BA83). 2% SDS, 5 mM 2-mercaptoethanol. Ponceau S: 0.2% Ponceau S, 3% trichloracetic acid (TCA), 3% sulfosalicylic acid (Sigma, S 3147). 6. Fluorographic reagent (Amplify, Amersham Corp.). 7. Sheets of stiff plastic (such as previously exposed X-ray film).

2.4. Pronase E Digestion and Chromatographic Analysis of Radiolabeled Proteins 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

12. 13. 14. 15. 16. 17. 18.

N-[2-hydroxyethyl]piperazine-N¢-[2′ -ethanesulfonic acid] Hepes, pH 7.4. Calcium acetate. Bath sonicator. Pronase E (Sigma, St. Louis, MO). 37°C water bath. n-Butanol saturated with water. Bench centrifuge. Oxygen-free nitrogen gas. Chloroform:methanol:H2O (10:10:3, by vol). Farnesyl-cysteine (F-Cys) was synthesized as described by Kamiya et al. (31). Geranylgeranyl-cysteine (GG-Cys) was synthesized as described by Kamiya et al. (31) except that isopropanol is substituted for methanol in the reaction solvent to improve the yield of the synthetic reaction. Silica Gel G 60 TLC plates (Sigma, St. Louis, MO). Chloroform:methanol:7 N ammonia hydroxide (45:50:5). Si*C18 reverse-phase plates (J. T. Baker Inc., Phillipsburg, NJ). Acetonitrile:H2O:acetic acid (75:25:1, by vol). Conical glass tubes. Anisaldehyde spray reagent (32) (see Note 1). Ninhydrin spray reagent (see Note 2).

2.5. Immunoprecipitation of Specific Radiolabeled Protein 1. Protein-specific immunoprecipitating antibody. 2. Protein A-Sepharose (Pharmacia).

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Wash buffer A: 20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.2% Nonidet P-40 (NP-40). Wash buffer B: 20 mM Tris-HCl, pH 7.5, 500 mM NaCl, 0.2% NP-40. Wash buffer C: 20 mM Tris-HCl, pH 7.5, 150 mM NaCl. n-Butanol saturated with water. Tabletop ultracentrifuge (Beckman).

2.6. Immunodetection of AG-Modified Cellular Proteins 1. Anilinogeraniol (30 µM, AG-OH) (Contact Drs. H. P. Spielmann and D. A. Andres, Univ. Kentucky, Lexington, KY) (28,33,34). 2. Appropriate cell culture media, plastic ware, and cell incubator. 3. Cell lysis buffer: 20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Nonidet P-40 (NP-40), 1 mM phenylmethyl+sulfonyl fluoride (PMSF) (add fresh PMSF solution and lysis is carried out at 4°C). Bradford reagent protein concentration kit (Pierce). 4. AG-specific antibody (Contact Drs. H. P. Spielmann and D. A. Andres, Univ. Kentucky, Lexington, KY) (28,33). 5. SDS-PAGE apparatus. 6. Western blot transfer apparatus. 7. Nitrocellulose membrane (Schleicher and Schuell, Protran BA83). 8. Blocking buffer PCT: 1 × phosphate buffered saline (PBS), 0.1 % Tween 20, 1% casein 9. Shaker platform. 10. Wash buffer PT: 1× PBS and 0.1% Tween 20. 11. Wash buffer PTE: 1× PBS and 0.3% Tween 20. 12. horseradish peroxidase-conjugated secondary antibody (Zymed). 13. Enhanced chemiluminescent detection reagent (Pierce).

3. Methods 3.1. Procedure for the Selective Metabolic Labeling of Farnesylated and Geranylgeranylated Proteins Procedures are described for the metabolic labeling of mammalian cells grown to near-confluence in Falcon 3001 tissue culture dishes. These protocols can be scaled up or down as appropriate. The incorporation of [3H]F-OH or [3H]GG-OH into protein was linear with respect to time and the concentration of [3H]isoprenol in tissue culture dishes ranging from 10 to 35 mm in diameter under the conditions described here (14,15,25) (see Note 3). 1. For metabolic labeling experiments with either [3H]F-OH or [3H]GG-OH, a disposable conical glass centrifuge tube (screw capped) and Teflon-lined cap are flame sterilized. The labeled isoprenol dissolved in ethanol is added to the tube and the ethanol is evaporated under a sterile stream of air. 2. An appropriate volume of sterile SS is added to yield a final concentration of 0.5–1 mCi/mL. The labeled isoprenols are dispersed in the SS by sonication in a Branson bath sonicator for 10 min. After sonication an aliquot is taken for liquid scintillation counting to verify that the 3H-labeled isoprenol has been quantitatively dispersed in SS.

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3. The growth medium is removed from the cultured cells by aspiration and 500 µL of labeling medium, consisting of an appropriate medium and SS (final concentration = 3–5%) containing [3H]GG-OH or [3H]F-OH, is added. 4. Cell cultures are typically incubated at 37°C under 5% CO2 for 6–24 h. Actual culture media and incubation conditions will vary depending on the specific cell type being studied.

3.2. Recovery of Metabolically Labeled Proteins from Adherent Cell Lines 1. The labeling medium is removed by gentle aspiration. 2. The metabolically labeled cells are washed with 1–2 mL of ice-cold PBS, to remove unincorporated isotopic precursor. 3. One milliliter of PBS containing 2 mM EDTA is added, and cells are incubated for 5 min at room temperature. The washed cells are gently scraped from the dish with a disposable cell scraper, and transferred to a 12-mL conical glass centrifuge tube. 4. The metabolically labeled cells are sedimented by centrifugation (500g, 5 min), and the PBS–EDTA is removed by aspiration (avoid disrupting the cell pellet). 5. The cells are resuspended in PBS (1–2 mL) and the PBS is removed by aspiration after the cells are sedimented by centrifugation. 6. Two milliliters of methanol (CH3OH) is added to the cell pellet, and the pellet is disrupted by sonication using a probe sonicator. 7. The suspension is sedimented by centrifugation (1500 g for 5 min). 8. The CH3OH extract is carefully removed, to avoid disturbing the partially delipidated pellet, and transferred to a glass conical tube (see Note 4). 9. The protein pellet is reextraced twice with 2 mL of CHCl3–CH3OH (2:1), and the extracts are pooled with the CH3OH extract (see Note 4). 10. Residual organic solvent is removed from the delipidated protein pellet by evaporation under a stream of nitrogen. The delipidated protein fractions are then subjected to Pronase E digestion for analysis of isoprenyl-cysteine analysis by TLC (see Subheading 3.4), or dissolved in 2% SDS, 5 mM 2-mercaptoethanol for SDS-PAGE analysis.

3.3. SDS-PAGE Analysis of Metabolically Labeled Proteins To examine the molecular weight and number of proteins metabolically labeled by incubation with [3H]F-OH or [3H]GG-OH, the delipidated protein fractions can be analyzed by SDS-PAGE. Because these experiments rely on the detection of low-energy 3H-labeled compounds, two procedures are described for the use of fluorography to increase the sensitivity of detection (see Note 5 before proceeding). 1. The delipidated protein fractions are solubilized in 2% SDS, 5 mM β-mercaptoethanol. An aliquot is used to determine the amount of labeled precursor incorporated into protein. 2. The radiolabeled polypeptides (20–60 µg of protein) were analyzed by SDS-PAGE using an appropriate percentage polyacrylamide resolving gel (4–20%) for the proteins of interest.

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3. Following SDS-PAGE, gels can be analyzed using two distinct methods. In the first, the gel is directly soaked in the fluorographic reagent, Amplify (Amersham), according to the manufacturer’s protocol, dried, and exposed to X-ray film as described in step 5. In a second approach, proteins were electrophoretically transferred to nitrocellulose filters and stained with Ponceau S to determine the efficiency of transfer. The nitrocellulose filters are destained by brief washing in distilled water and allowed to air-dry. (See Note 5 for a discussion of the merits of each method before continuing.) 4. The filters were then dipped briefly in the fluorographic reagent, Amplify (Amersham), placed on a sheet of plastic backing, and dried for 1 h at 50°C. It is important that a thin and even film of Amplify reagent remain on the filter and that it be placed protein side up to dry. 5. Fluorograms were produced by exposing preflashed X-ray film to the nitrocellulose filter, or dried SDS-PAGE gel, for 5–30 d at −80°C.

3.4. Methods for the Identification of Cysteine-Linked Isoprenyl Group These simple methods are inexpensive, rapid, and allow the identification of the isoprenyl-cysteine residue(s) from isoprenylated protein(s). Examples of this method for the identification of isoprenyl-cysteine groups from metabolically labeled cells and recombinant proteins labeled in vitro are shown in Figs. 4 and 5. As expected, [3H]F-Cys and [3H]GG-Cys were liberated from RAS(CVLS) and RAS(CVLL), respectively. A radioactive peak is also seen at the origin in the analysis of the Pronase digest of radiolabeled RAS(CVLL) (Fig. 5, middle panel). This radiolabeled product(s) is probably incompletely digested [3H]geranylgeranylated peptides. Rab 1A terminates in two cysteine residues, both of which are isoprenylated (35). Fig. 5 (lower panel) indicates that Pronase E is incapable of cleaving between the two cysteine residues. 1. To liberate the labeled isoprenyl-cysteine residues for analysis, the delipidated protein fractions (50–100 µg) are incubated with 2 mg of Pronase E; 50 mM HEPES, pH 7.4; and 2 mM calcium acetate in a total volume of 0.1 mL at 37°C for approx 16 h. The experimental scheme for this analysis is illustrated in Fig. 3 (see Note 6). 2. Proteolysis is terminated by the addition of 1 mL of n-butanol saturated with H2O and mixing vigorously. 3. Centrifuge the mixture for 5 min in a benchtop centrifuge at 1500g. Two phases will form, and the upper phase should be clear (see Note 7). 4. Transfer the upper phase, containing n-butanol, to a separate conical glass centrifuge tube. 5. Add 1 mL of H2O to the butanol extract and mix vigorously. Centrifuge at 1500g for 5 min, to effect a phase separation. Remove the lower aqueous phase with a Pasteur pipet (see Note 8). Evaporate the n-butanol under a stream of nitrogen at 30–40°C (see Note 9).

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6. The labeled isoprenyl-cysteines are dissolved in 250 µL of CHCl3:CH3OH:H2O (10:10:3, by vol) by mixing vigorously, and an aliquot (10 µL) is taken to determine the amount of radioactivity. 7. The radiolabeled products are analyzed chromatographically on a normal-phase system, using silica gel G 60 TLC plates developed in CHCl3:CH3OH:7 N NH4OH (45:50:5, by vol) or by reverse-phase chromatography using silica gel Si*C18 reverse-phase plates developed with acetonitrile–H2O–acetic acid (75:25:1, by vol) (see Note 10). 8. The desired developing solvent mixture is added to the chromatography tank to a depth of 0.5–1.0 cm and allowed to equilibrate. 9. Dry entire sample under nitrogen stream. Redissolve in chloroform–methanol– water (10:10:3, by vol). 10. Apply an aliquot (approx 10,000 dpm containing 10–12 µg of authentic F-Cys and GG-Cys) to the origin on the TLC plate, using a fine glass capillary or a Hamilton syringe. The addition of the unlabeled standards improves the resolution of the metabolically labeled isoprenylated cysteines. 11. Allow the sample to dry (this can be facilitated with a stream of warm air). Complete application of the labeled isoprenyl-cysteine extract to the plate in 5-µL aliquots, allowing time for the spot to dry between applications. 12. Place the plate(s) in the preequilibrated chromatography tank, and after the solvent has reached the top of each plate (1–2 h) remove and allow to air-dry in a fume hood. 13. When the plates are thoroughly dried, the radioactive zones are located with a Bioscan Imaging Scanner System 200-IBM or autoradiography (see Note 11). 14. Standard compounds are localized by exposure of the plate to the anisaldehyde spray reagent (32) or a ninhydrin spray reagent (see Note 12).

3.5. Application of These Methods to the Analysis of Individual Proteins To illustrate the utility of this approach for individual isoprenylated proteins, these methods are applied to the analysis of a recently isolated farnesylated protein, RDJ2 (rat DnaJ homologue 2). The cDNA clone of this DnaJ-related protein was recently identified (36). The predicted amino acid sequence is found to terminate with the tetrapeptide Cys-Ala-His-Gln, which conforms to the consensus sequence for recognition by protein farnesyltransferase, and was shown to undergo farnesylation in vivo. To perform this analysis, a means of specifically identifying the protein of interest must be available. In this example, a protein-specific immunoprecipitating antibody was used to isolate the protein from isotopically radiolabeled mammalian cells. However, other experimental approaches are available. (See Note 13 for a discussion of these stategies.) 1. Mammalian cells are grown to near confluence and metabolically labeled with either [3H]mevalonate, [3H]F-OH, or [3H]GG-OH as described in Subheading 3.1.

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2. 3.

4. 5.

6.

7. 8.

9.

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If a recombinant protein is to be analyzed, the mammalian cells should be transfected either stably or transiently with the expression vector prior to labeling (37). The metabolically labeled cells are washed with 1–2 mL of ice-cold PBS to remove unincorporated isotopic precursor. One milliliter of PBS containing 2 mM EDTA is added, and the cells are incubated for 5 min at room temperature. The washed cells are gently scraped from the dish with a disposable cell scraper and transferred to a 12-mL conical glass centrifuge tube. The metabolically labeled cells are sedimented by centrifugation (500 g, 5 min), and the PBS–EDTA is removed by aspiration (avoid disrupting the cell pellet). The cells are resuspended in lysis buffer (2 mL), disrupted by passage through a 20-gauge needle (3–4×), and centrifuged for 15 min at 100,000 g in a Beckman table top ultracentrifuge. For immunoprecipitation of recombinant RDJ2, 20 µg of rabbit anti–RDJ2 antibody was added to the cleared supernatant and the mixture was incubated for 12 h at 4°C with gentle rocking (see Note 14). Immune complexes were then precipitated by addition of 100 µL of a 50% slurry of protein A-Sepharose for 2 h at 4°C with gentle rocking. Protein A beads were collected by centrifugation (1 min in a tabletop microfuge at 10,000 rpm). The pellet was washed 3× by resuspending in 1 mL of wash buffer A, 1 mL of wash buffer B, and 1 mL of wash buffer C, sequentially. The protein is dissolved in 2% SDS, 5 mM β-mercaptoethanol for SDS-PAGE analysis (see Subheading 3.3. and Fig. 6). Alternatively, the protein A beads are subjected to pronase E digestion for analysis of isoprenyl-cysteine analysis by TLC (see Subheading 3.4, Fig. 7, and Note 15).

3.6. Immunodetection of AG-labeled Cellular Proteins A recent advance in the detection of cellular isoprenylated proteins was been the generation of the unnatural F-P-P analogue AG-P-P and anti-AG antibodies (28). These reagents have been utilized to examine the prenylation status of both general cellular proteins (Fig. 8) and specific prenylated proteins (33). The combined use of an unnatural FPP analog and specific antibody provides a simple, inexpensive, and rapid method for monitoring both FTase activity in cells and detection of cellular prenylated proteins using non-radioactive methods (Fig. 8). 1. Mammalian cells are grown to near confluence and metabolically labeled with AG-OH. This can either be performed on cells treated with lovastatin (30 µM) to reduce the intracellular F-P-P/GG-P-P pool and 100 mM AG-OH for 24 h, or cells with no lovastatin block can be incubated with 30–100 µM AG-OH from 6–72 h (28, 33) (see Note 16). If a recombinant protein is to be analyzed, the mammalian cells should be transfected either stably or transiently with the expression vector prior to labeling (37). 2. The AG-OH labeled cells are washed with 1–2 mL of ice-cold PBS to remove unincorporated AG-OH, the PBS removed by aspiration, and the cells resuspended

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in cell lysis buffer (300–500 µl for a 10-cm dish). The cells are disrupted by brief sonication, and protein concentration determined using Bradford reagent. 3. The AG-modified polypeptides (50–100 µg of total cell protein) can be analyzed by SDS-PAGE using an appropriate percentage polyacrylamide resolving gel (4–20%) for the proteins of interest and electrophoretically transferred to nitrocellulose membrane. 4. The membranes are incubated in PCT for 1 h at room temperature, after which the following sequential steps are performed at room temperature: incubate with polyAG-Ab (1; 5,000 dilution) in PCT for 1 h; 3 × 15 min washes with shaking in PT buffer; incubation for 1 h in PCT with goat-anti-rabbit horseradish peroxidaseconjugated secondary antibody (dilution of 1: 20,000); 2 × 10 min washes with PT; 1 × 10 min wash with PTT; and a final 2 × 10 min wash with PT. 5. The membranes are then subjected to enhanced chemiluminescence detection by being placed in 10 mL of detection buffer (Pierce Supersignal) for 1 min and exposed to x-ray film.

4. Notes 1. Anisaldehyde spray reagent contains 10 mL of anisaldehyde, 180 mL of 95% ethanol, and 10 mL of conc. sulfuric acid, added in that order. 2. Ninhydrin spray reagent contains 0.2% ninhydrin in 95% ethanol. 3. All mammalian cells tested (CHO, C6 glioma cells and green monkey kidney [CV-1] cells) utilized F-OH and GG-OH for protein isoprenylation except murine B cells before or after activation by lipopolysaccharide (LPS). Thus, it is possible that the “salvage” pathway for F-OH and GG-OH (Fig. 1) may not be ubiquitous in mammalian cells. 4. The pooled organic extracts can be used for analysis of lipid products metabolically labeled by [3H]F-OH (see ref. 14). The organic solvent is evaporated under a stream of nitrogen, and the lipid residue redissolved in CHCl3:CH3OH (2:1) containing 20 µg each of authentic cholesterol and squalene. An aliquot is taken to determine the amount of radioactivity incorporated into the lipid extracts. The lipid products are analyzed on Merck silica gel G 60 TLC plates (Sigma) by developing with hexane– diethyl ether–acetic acid (65:35:1) or chloroform. Radioactive zones were located with a Bioscan Imaging Scanner System 200-IBM. Standard compounds are located with iodine vapor or anisaldehyde spray reagent (32). 5. Western transfer is preferred because transferring labeled proteins to nitrocellulose membrane appears to give a gain of 2–10-fold in sensitivity. One suspects that the polyacrylamide gel acts to quench the signal from radiolabeled protein. The transfer step serves to collect proteins in a single plane, and eliminates this problem. However, care should be taken with the intrepretation of these experiments. It is possible that some radiolabeled proteins, particularly those of either small (100 kDa) molecular mass, may be inefficiently transferred. The properties of the protein, percentage of acrylamide, transfer buffer components, and transfer time will each influence the transfer efficiency. Direct analysis of the gel will be less sensitive, requiring more labeled protein and longer exposure times, but material will not be lost during transfer. Pronase E will completely dissolve over a period of 30 min at 37°C, and these protease preparations contain sufficient esterase activity to hydrolyze the carboxymethyl esters at the C-termini of isoprenylated proteins (38). The lower aqueous phase remains cloudy, and contains precipitated proteins and peptides. The addition of H2O will significantly reduce the volume of the n-butanol phase at this step (reducing the time required for the following steps). The addition of an equal volume of n-hexane to the n-butanol will speed this evaporation, by forming an azeotrope. The addition of the hexane will cause the solution to become cloudy and biphasic. The upper (butanol– hexane phase) evaporates quite rapidly, and the subsequent addition of 1 mL of 100% ethanol speeds the evaporation of the lower aqueous phase. The resolution of GG-Cys and F-Cys is better in the reverse–phase chromatography system. If the normal phase system is used, the plates should be activated by heating in a 100°C oven for at least 1 h. At least 5000 dpm are required for good detection using a Bioscan Imaging System, but analysis will be better (and faster) with 10,000 dpm or more. The entire sample from the digestion of 100 µg of labeled protein can usually be loaded on the TLC plates without any significant loss of chromatographic resolution. Spray until plate is moist, and heat in 100°C oven until spots appear. The plates are scanned prior to spraying to avoid the reagent vapors, and because the spray reagents quench the detection of radioactivity. A consideration before beginning these experiments is the abundance of the protein of interest. The analysis of a very low abundance protein will require a large number of radiolabeled tissue culture cells. Therefore, the overproduction of the protein using a mammalian expression vector may present a distinct experimental advantage. This approach also allows the cDNA to be modified to contain a unique epitope or affinity sequence at the N-terminus. In this way, proteins for which specific antibodies are not available may be studied. Optimal immunoprecipitation conditions must be established for each protein-antibodyc omplex. Pronase E digestion can be carried out directly on the protein A beads without further processing. Follow directions given in Subheading 3.4, scaling up the volume of the proteolysis reaction mixture to provide sufficient

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liquid to amply cover the protein A beads. Follow steps 2–15 as directed. Initially, proteins should be labeled with mevalonate and both free isoprenols, to ensure correct interpretation of the results. Although a variety of proteins have been tested (see Figs. 2, 4, and 7) using [3H]isoprenol labeling, the list of individual proteins is quite limited. It will be necessary to analyze a wide variety of defined isoprenylated proteins to further establish the reliability and limitations of this method. 16. Optimal AG-OH labeling conditions must be established for each cell culture system. Thus, pilot studies varying both the concentration (within the limits of 10–100 µM to avoid potential toxicity) and time of AG-OH labeling, in the presence or absence of lovastatin, should be performed (28). This will be particularly important if examining a specific cellular protein (see Note 13). Finally, it must be remembered that AGPP is a FTase substrate and that this approach can not be used to analyze geranylgeranylated proteins. Acknowledgments The methods described in this chapter were developed with support from NIH grants NINDS04103 and HL072936 (D.A.A.) and GM36065 (C.J.W.).

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via farnesyl pyrophosphate formed by successive monophosphorylation reactions. Proc. Natl. Acad. Sci. U.S.A. 96, 13080–13085. Crick, D. C., Andres, D. A., and Waechter, C. J. (1997) Novel salvage pathway utilizing farnesol and geranylgeraniol for protein isoprenylation. Biochem. Biophys. Res. Commun. 237, 483–487. Gibbs, B. S., Zahn, T. J., Mu, Y., Sebolt-Leopold, J. S., and Gibbs, R. A. (1999) Novel farnesol and geranylgeraniol analogues: A potential new class of anticancer agents directed against protein prenylation. J. Med. Chem. 42, 3800–3808. Kho, Y., Kim, S. C., Jiang, C., Barma, D., Kwon, S. W., Cheng, J., Jaunbergs, J., Weinbaum, C., Tamanoi, F., Falck, J., and Zhao, Y. (2004) A tagging-via-substrate technology for detection and proteomics of farnesylated proteins. Proceed. Natl. Acad. Sci. U.S.A. 101, 12479–12484. Troutman, J. M., Roberts, M. J., Andres, D. A., and Spielmann, H. P. (2005) Tools to analyze protein farnesylation in cells. Bioconjugate Chem. 16, 1209–1217. Chehade, K. A., Andres, D. A., Morimoto, H., and Spielmann, H. P. (2000) Design and synthesis of a transferable farnesyl pyrophosphate analogue to Ras by protein farnesyltransferase. J. Organ. Chem. 65, 3027–3033. Chehade, K. A., Kiegiel, K., Isaacs, R. J., Pickett, J. S., Bowers, K. E., Fierke, C. A., Andres, D. A., and Spielmann, H. P. (2002) Photoaffinity analogues of farnesyl pyrophosphate transferable by protein farnesyl transferase. J. Am. Chem. Soc. 124, 8206–8219. Kamiya, Y., Sakurai, A., Tamura, S., and Takahashi, N. (1978) Structure of rhodotorucine A, a novel lipopeptide, inducing mating tube formation in Rhodosporidium toruloides. Biochem. Biophys. Res. Commun. 83, 1077–1083. Dunphy, P. J., Kerr, J. D., Pennock, J. F., Whittle, K. J., and Feeney, J. (1967) The plurality of long chain isoprenoid alcohols (polyprenols) from natural sources. Biochim. Biophys. Acta 136, 136–147. Dechat, T., Shimi, T., Adam, S. A., Rusinol, A. E., Andres, D. A., Spielmann, H. P., Sinensky, M. S., and Goldman, R. D. (2007) Alterations in mitosis and cell cycle progression caused by a mutant lamin A known to accelerate human aging. Proc. Natl. Acad. Sci. U.S.A. 104, 4955–4960. Roberts, M. J., Troutman, J. M., Chehade, K. A., Cha, H. C., Kao, J. P., Huang, X., Zhan, C. G., Peterson, Y. K., Subramanian, T., Kamalakkannan, S., Andres, D. A., and Spielmann, H. P. (2006) Hydrophilic Anilinogeranyl Diphosphate Prenyl Analogues Are Ras Function Inhibitors. Biochemistry 45, 15862–15872. Seabra, M. C., Brown, M. S., Slaughter, C. A., Sudhof, T. C., and Goldstein, J. L. (1992) Purification of component A of Rab geranylgeranyl transferase: possible identity with the choroideremia gene product. Cell 70, 1049–1057. Andres, D. A., Shao, H., Crick, D. C., and Finlin, B. S. (1997) Expression cloning of a novel farnesylated protein, RDJ2, encoding a DnaJ protein homologue. Arch. Biochem. Biophys. 346, 113–124. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, New York. Stimmel, J. B., Deschenes, R. J., Volker, C., Stock, J., and Clarke, S. (1990) Evidence for an S-farnesylcysteine methyl ester at the carboxyl terminus of the Saccharomyces cerevisiae RAS2 protein. Biochemistry 29, 9651–9659.

169 Antibody Production Robert Burns 1. Introduction Antibodies are proteins that are produced by the immune systems of animals in response to foreign substances. The immune system has the ability to recognize material as nonself or foreign and mount a response to them. Substances that elicit this are known as immunogens or antigens, and one of the outcomes of the immune response is the production of antibodies that will recognize and bind to the eliciting substance. The immune response results in the degradation and processing of the antigen by cells called macrophages and its presentation in fragments to B lymphocytes that are found in the lymphatic tissues of the animal. On presentation of the antigen fragments the B lymphocytes mature and “learn” to produce antibody molecules that have a specific affinity to the antigen fragment that they have been presented with. This process gives rise to populations of B lymphocyte clones each of which produces antibody molecules to different locations known as epitopes on the target antigen (1). As individual clones produce the antibody molecules and there are many of them, the resulting antibody mix in the blood is known as polyclonal and the fluid derived from the clotted blood is known as polyclonal antiserum. Deliberate exposure to antigens producing antibodies in animals is known as immunization, and repeated exposure leads to a condition known as hyperimmunity, in which a significantly large proportion of the circulating antibodies in the animals blood will be directed toward the antigen of interest. When exposure to the antigen ceases a stable but quiescent populations of B lymphocytes known as memory cells remain that will respond to the presence of the antigen by dividing to increase their number and the level of circulating antibody in the blood.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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The primary exposure to an antigen gives rise to a pentameric form of antibody known as immunoglobulin M (IgM). Subsequent exposure to the antigen causes maturation of the cells and a shift in the antibody type to the more stable dimeric immunoglobulin G (IgG). This shift in antibody type happens as affinity maturation of the B lymphocyte clones occurs. Some antigens, particularly highly glycosylated proteins, do not elicit the production of memory B lymphocytes and also may not give rise to IgG antibodies even after repeated immunization. These are known as anamnestic antigens, and as no memory B lymphocytes are produced, the immune system always interprets exposure to them as a primary one and mounts an IgM response. It is possible to produce IgG to anamnestic antigens by treating them to remove the carbohydrate moieties prior to immunization, but in some cases this may lead to antibodies that do not recognize the native protein structure. Antibodies have two qualities known as avidity and affinity. Affinity is a measure of how well the binding site on the antibody molecule fits the epitope, and antibodies, which have high affinity also, have high specificity to the target epitope. Avidity is a measure of how strong the interaction between the epitope and the antibody molecule is. For research purposes, antibodies to the antigen of interest should be expressed at fairly high levels in the sera of donor animals, should be of high affinity, and should predominantly be composed of IgG molecules, which are stable and easily purified. This can be achieved for most antigens providing they are not too highly glycosylated, have a molecular weight >5000, and are not toxic or induce immuneparesis in the donor animal. The ability of species to respond to antigens is variable and it may be necessary to investigate which animal would be the most suitable prior to embarking on a large scale immunization project. Monoclonal antibodies are used very successfully in many areas of research and can either replace, complement, or be used in conjunction with immunoglobulins obtained from donor serum. Monoclonal antibodies are produced by hybridoma cell lines and can be grown in tissue culture in the laboratory. Hybridomas are recombinant cell lines produced from the fusion of B cells derived from the lymphatic tissue of donor animals and myeloma cell lines that imparts immortality to the cells (2,3). As each hybridoma is descended from a single B cell clone the antibody expressed by it is of a single specificity and immunoglobulin type, and is thus termed a monoclonal antibody. Each monoclonal antibody is monospecific and will recognize only one epitope on the antigen to which it has been raised. This may lead to practical problems if the epitope is not highly conserved on the native protein or where conformational changes may occur in the because of to shifts in pH or other environmental factors. Monoclonal antibodies are highly specific and will rarely if ever produce cross-reactions with other proteins. Polyclonal antibodies may cross-react with other closely related proteins where there are shared epitopes.

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Polyclonal antisera contain a heterogeneic mixture of antibody molecules, many of which will recognize different epitopes on the protein, and their binding is much less likely to be affected by poorly conserved epitopes or changes in the protein shape. Antisera are derived from individual animal bleeds and because of this are subject to batch variation. Individual animals can have very different immune responses to the same antigen, and individual bleeds from the same animal may vary in antibody content quite markedly. Monoclonal antibodies are produced from highly cloned cell lines that are stable and reliably produce a defined antibody product. Polyclonal antibodies are generally less specific than monoclonals, which can be a disadvantage, as cross-reactivity may occur with nontarget proteins. Monoclonal antibodies can be too specific, as they will recognize only a single epitope that may vary on the protein of interest. Both antibody types have their place in the research laboratory, and a careful evaluation of the required use should be undertaken before deciding which would be most applicable. In recent years the use of non-animal-based systems have been reported and in particular the use of phage display technology for antibody production appears to be promising. Phage display libraries expressing single chain antibodies can be screened using the antigen to isolate antibodies of interest which are then expressed using vector systems such as M13 (4). These systems allow the generation of antibodies without the use of animal donors and also allow the generation of antibodies to “difficult” antigens. There are limitations in that the antibodies produced are actually antibody fragments and there may be restrictions on how they may be used in assays due to molecule size and stability. Antibodies derived by phage display technology may be transfected into cell lines such as Chinese Hamster Ovary (CHO) if large quantities of product are required. Fully human antibodies can be generated using this methodology without the use of human donor tissue and its attendant ethical concerns. 1.1. Donor Animals Most polyclonal antibodies for research purposes are produced in domestic rabbits unless very large quantities are required, and then sheep, goats, donkeys, and even horses are used. Antibodies can also be produced in chickens; the antibodies are conveniently produced in the eggs. Rats and mice can also be used to produce antiserum but yield much smaller quantities of antibody owing to their relatively small size. Immunized mice can be used to produce 5–10 mL of polyclonal ascitic fluid by induction of ascites. Ascites is induced by the introduction of a nonsecretory myeloma cell line into the peritoneal cavity after priming with Pristane. Polyclonal antibodies are secreted into the peritoneum from the blood plasma of the animal and can achieve levels of 2–5 mg/mL. Fluid isaspirated

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from the peritoneum of the mouse when ascites has developed, indicated by bloating of the abdomen. The UK Home Office and other regulatory authorities now regard induction of ascites in mice as a moderate/severe procedure and this method is not normally used unless no alternative methods are available. 1.2. Adjuvants Adjuvants are substances that increase the immune response to antigen by an animal. They may be simple chemicals such as alum, which adsorbs and aggregates proteins, increasing their effective molecular weight, or they may be specific immunestimulators such as derivatives of bacterial cell walls (5). Saponins such as Quil-A derived from the tree Quillaja saponaria may also be used to increase the effective immunogenicity of the antigen. Ideally an adjuvant should not induce too great an antibody response to itself to ensure that antibodies generated are specific to the antigen of interest. The most popular adjuvant, which has been used very successfully for many years, is Freund’s adjuvant. It has two formulations, complete and incomplete, which are used for the primary and subsequent immunizations, respectively. Freund’s incomplete formulation is a mixture of 85% paraffin oil and 15% mannide monooleate. The complete formulation additionally contains 1 mg/mL of heat-killed Mycobacterium tuberculosis. In recent years the use of Freund’s adjuvant has declined owing to animal welfare concerns and also the risk that it poses to workers, as it can cause localized soft tissue damage following accidental needlestick injuries. Injection preparations that contain Freund’s adjuvant are also difficult to work with, as the resultant emulsion can be too thick to administer easily and it interacts with plastic syringes, preventing easy depression of the plunger. Several adjuvant formulations are available that contain cell wall derivatives of bacteria without the intact organisms; they are much easier to administer and will achieve a similar immunostimulatory effect to Freund’s without its attendant problems. 1.3. Legislation Most countries have legislation governing the use of animals for all experimental work, and antibody production is no exception. Although the methods used are usually mild in terms of severity they must be undertaken with appropriate documentation and always by trained, authorized staff. The legislation in terms of animal housing, immunization procedures, bleeding regimens, and choice of adjuvant vary widely according to local legislation, and it is imperative that advice be taken from the appropriate authorities prior to undertaking any antibody production work.

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1.4. Antigens The antigen chosen for an immunization program should be as close in structure and chemical identity to the target protein as possible. An exception to this is when synthetic peptides are produced to mimic parts of the native protein, an approach that is invaluable when the native antigen may be toxic to animals or nonimmunogenic. The antigen should be soluble, stable at dilutions of approx 1 mg/mL, and capable of being administered in a liquid of close to physiological pH (6.5–7.5). The antigen should also be in as pure a form as practically possible to avoid the generation of antibodies to contaminating materials. Many proteins are highly immunogenic in donor animals, particularly when the antigen is derived from a different species (see Note 1). Raising antibodies in the same species from which the antigen is derived from can be extremely difficult but can be overcome by conjugating the antigen to a carrier protein from another species prior to immunization. Carrier proteins such as hemoglobin, thyroglobulin, and keyole limpet hemocyanin (KLH) are commonly used. Animals immunized with the conjugated form of the antigen will produce antibodies to both the protein of interest and the carrier protein. Apart from a lower specific antibody titer in the serum there should be no interference from the carrier protein antibodies as it will not be present in the final testing system. 1.5. Test Protocol Ideally antiserum should be tested using the procedure in which it is to be used, as antibodies may perform well using one assay but not with another. This, however, is not always practical and so a number of tests can be carried out on antiserum to test its suitability for final use. The affinity of the antiserum to the antigen can be assayed by plate-trapped double-antibody sandwich enzyme-linked immunosorbent assay (DAS ELISA) (6). The antigen is bound to a microtiter plate and then challenged with dilutions of the test antiserum. A secondary antispecies antibody enzyme conjugate is then added to the plate and will bind to any antibody molecules present. A chromogenic enzyme substrate is then added and the degree of color development indicates the quantity of antibody bound by the antigen. Ouchterlony double-immunodiffusion can be used to observe the ability of the antiserum to produce immune complexes with the antigen in a semisolid gel matrix. This method can also be used to test cross-reactivity of the antiserum to other proteins closely related to the antigen. Radioimmunoassay and other related techniques can also be used to test the avidity of the antiserum to the antigen and will also give a measure of antibody titer.

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In all the above tests preimmune antisera should be included to ensure that results obtained are a true reflection of antibodies produced by immunization and not due to nonspecific interactions. 2. Materials 1. At least two rabbits should be used for each polyclonal antibody production project, as they may have different immune responses to the antigen. They should be purchased from a reputable source and be parasite and disease free. New Zealand whites are often used but any of the domestic breeds can be used (see Note 2). 2. The antigen should be in a buffer of pH 6.5–7.5 and be free of toxic additives (sodium azide is often added as a preservative). A concentration of 1 mg/mL is desirable but anything above 100 µg/mL is acceptable. 3. Complete and incomplete Freund’s adjuvant or any of the propriety preparations containing purified bacterial cell wall components such as Titermax and RIBI.

3. Method 1. Prior to a course of immunizations a test bleed should be taken from the rabbits to provide a source of preimmune antiserum for each animal (see step 5 below). It is usual to take only 2–3 mL for the test bleeds, which will yield 1–1.5 mL of serum. The blood should be allowed to clot at 4°C for 12 h and the serum gently aspirated from the tube. 2. The antigen should be mixed with the appropriate adjuvant according to the manufacturer’s instructions to achieve a final volume of 0.5 mL/injection containing 50–500 µg of antigen (see Note 3). If Freund’s adjuvant is to be used, the first injection only should contain the complete formulation and the incomplete one should be used for all subsequent immunizations. 3. The rabbit should be gently restrained and the antigen–adjuvant mixture injected into the thigh muscle. Alternate legs should be used for each injection (see Note 4). 4. The immunization should be repeated 14 d after the primary one and a test bleed taken 28 d after that. 5. If the antiserum shows that the desired immune status and antibody quality has been achieved then donor bleeds can be taken. The volume of blood collected and frequency of bleeding depends on animal welfare legislation and must be adhered to. Each bleed should be assayed individually, and once the antibody titer has started to fall a further immunization can be given, followed by donor bleeds, or the animal can be terminally anesthetized and exsanguinated by cardiac puncture or by severing the carotid artery. 6. Antiserum collected from rabbits can be stored for extended periods of time at 4°C but the addition of 0.02% sodium azide is recommended to prevent adventitious bacterial growth. Antiserum quite commonly has functional antibodies even after years of refrigerated storage, but storage at −20°C is recommended for long-term preservation. 7. Antiserum will often yield in excess of 5 mg/mL of antibody and can be purified to give the immunoglobulin fraction. This can be achieved with ammonium sulfate

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precipitation or by affinity chromatography using either the immobilized antigen or protein A or G. The antibody fraction can then be adjusted to 1 mg/mL, which is an ideal concentration both for its stability and for many practical applications. Purified antibody should have 0.02% sodium azide or some other preservative added to prevent the growth of adventitious organisms. Sodium azide can interfere with enzyme reactions and with photometric measurement, and this should be taken into account with regard to the final assay. Purified antibodies diluted to 1 mg/mL can be stored for long periods at 4°C with no loss of activity but for extended storage −20°C is recommended.

4. Notes 1. Some antigens will consistently fail to induce an antibody response in certain animals, and other species should be investigated as potential donors. In very rare occasions antigens will not elicit an immune response in a range of species and the nature of the antigen will then have to be investigated with a view to modifying it to increase its immunogenicity. 2. Female rabbits are less aggressive, and although smaller and yielding smaller quantities of antiserum are preferable to male rabbits for antiserum production. Many biomedical facilities use communal floor pens for donor animals and female rabbits adapt better to this form of housing. 3. The use of excessive amounts of antigen in immunizations should be avoided, as this can lead to a poor immunological response. Overloading the immune system can lead to selective deletion of the B cell clones of interest and a reduction in the specific antibody titer. High doses of antigen in the secondary and subsequent immunizations can cause anaphylactic shock and death of the donor animal. 4. It has been reported that increased stress levels in animals can depress the immune response, and appropriate measures should be taken to ensure that immunizations and bleeds are performed with the minimum of stress to the animals. General husbandry in terms of housing, noise levels, and other environmental factors should also be examined to ensure that animals for polyclonal production are maintained under suitable conditions. References 1. Roitt, I., Brostoff, J., and Male, D. (1996) Immunology, C. V. Mosby, St. Louis, pp. 1.7–1.8. 2. Kennet, R. H., Denis, K. A., Tung, A. S., and Klinman, N. R. (1978) Hybrid plasmacytoma production: fusions with adult spleen cells, monoclonal spleen fragments, neonatalspleen cells and human spleen cells. 3. Kohler, G. and Milstein, C. (1975) Continuous cultures of fused cells secreting antibody of predefined specificity. Nature 256, 495–497.

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4. Hust, M. and Dubel, S. (2005) Methods in Molecular Biology, Vol. 295: Immunochemical Protocols, Third Edition, pages 71–95. Humana Press Inc. Totowa, NJ. 5. Harlow, E. and Lane, D. (1988) Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp. 96–97. 6. Kemeny, D. M. and Chandler, S. (1988) ELISA and Other Solid Phase Immunoassays, John Wiley & Sons, pp. 1–29.

170 Production of Antibodies Using Proteins in Gel Bands Sally Ann Amero, Tharappel C. James, and Sarah C. R. Elgin 1. Introduction A number of methods for preparing proteins as antigens have been described (1). These include solubilization of protein samples in buffered solutions (ref. 2 and see Chapter 169), solubilization of nitrocellulose filters to which proteins have been adsorbed (ref. 3 and see Chapter 171), and emulsification of protein bands in polyacrylamide gels for direct injections (4–8). The latter technique can be used to immunize mice or rabbits for production of antisera or to immunize mice for production of monoclonal antibodies (9–11). This approach is particularly advantageous when protein purification by other means is not practical, as in the case of proteins insoluble without detergent. A further advantage of this method is an enhancement of the immune response, since polyacrylamide helps to retain the antigen in the animal and so acts as an adjuvant (7). The use of the protein directly in the gel band (without elution) is also helpful when only small amounts of protein are available. For instance, in this laboratory, we routinely immunize mice with 5–10 µg total protein using this method; we have not determined the lower limit of total protein that can be used to immunize rabbits. Since polyacrylamide is also highly immunogenic, however, it is necessary in some cases to affinity-purify the desired antibodies from the resulting antiserum or to produce hybridomas that can be screened selectively for the production of specific antibodies, to obtain the desired reagent. 2. Materials 1. Gel electrophoresis apparatus; acid-urea polyacrylamide gel or SDS-polyacrylamide gel. 2. Staining solution: 0.1% Coomassie brilliant blue-R (Sigma, St. Louis, MO, B-7920) in 50% (v/v) methanol/10% (v/v) acetic acid. From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Destaining solution: 5%-(v/v) methanol/7% (v/v) acetic acid. 2% (v/v) glutaraldehyde (Sigma G-6257). Transilluminator. Sharp razor blades. Conical plastic centrifuge tubes and ethanol. Lyophilizer and dry ice. Plastic, disposable syringes (3- and 1-mL). 18-gage needles. Spatula and weighing paper. Freund’s complete and Freund’s incomplete adjuvants (Gibco Laboratories, Grand Island, NY). 13. Phosphate-buffered saline solution (PBS): 50 mM sodium phosphate, pH 7.25/150 mM sodium chloride. 14. Microemulsifying needle, 18-g. 15. Female Balb-c mice, 7–8 wk old, or New Zealand white rabbits.

3. Method 1. Following electrophoresis (see Note 1), the gel is stained by gentle agitation in several volumes of staining solution for 30 min. The gel is partially destained by gentle agitation in several changes of destaining solution for 30–45 min. Proteins in the gel are then cross-linked by immersing the gel with gentle shaking in 2% glutaraldehyde for 45–60 min (12). This step minimizes loss of proteins during subsequent destaining steps and enhances the immunological response by polymerizing the proteins. The gel is then completely destained, usually overnight (see Note 2). 2. The gel is viewed on a transilluminator, and the bands of interest are cut out with a razor blade. The gel pieces are pushed to the bottom of a conical plastic centrifuge tube with a spatula and pulverized. The samples in the tubes are frozen in dry ice and lyophilized. 3. To prepare the dried polyacrylamide pieces for injection, a small portion of the dried material is lifted out of the tube with a spatula and placed on a small square of weighing paper. In dry climates it is useful to first wipe the outside of the tube with ethanol to reduce static electricity. The material is then gently tapped into the top of a 3-mL syringe to which is attached the microemulsifying needle (Fig. 1A). Keeping the syringe horizontal, 200 µL of PBS solution is carefully introduced to the barrel of the syringe, and the plunger is inserted. Next, 200 µL of Freund’s adjuvant is drawn into a 1-mL tuberculin syringe and transferred into the needle end of a second 3-mL syringe (Fig. 1B). This syringe is then attached to the free end of the microemulsifying needle. The two plungers are pushed alternatively to mix the components of the two syringes (Fig. 1C). These will form an emulsion within 15 min; it is generally extremely difficult to mix the material any further. 4. This mixture is injected intraperitoneally or subcutaneously into a female Balb-c mouse, or subcutaneously into the back of the neck of a rabbit (see refs. 13 and 14). Since the emulsion is very viscous, it is best to use 18–g needles and to anesthesize the animals. For mice, subsequent injections are administered after 2 wk and after

Production of Antibodies Using Proteins in Gel Bands Fig. 1. Preparation of emulsion for immunizations. To prepare proteins in gel bands for injections, an emulsion of Freund’s adjuvant and dried polyacrylamide pieces is prepared. (A) Dried polyacrylamide is resuspended in 200 µL of PBS solution in the barrel of a 3-mL syringe to which is attached a microemulsifying needle. (B) Freund’s adjuvant is transferred into the barrel of a second 3-mL syringe. (C) An emulsion is formed by mixing the contents of the two syringes through the microemulsifying needle.

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3 more wk. If monoclonal antibodies are desired, the animals are sacrificed 3–4 d later, and the spleen cells are fused with myeloma cells (ref. 13). The immunization schedule for rabbits calls for subsequent injections after 1 mo or when serum titers start to diminish. Antiserum is obtained from either tail bleeds or eye bleeds from the immunized mice, or from ear bleeds from immunized rabbits. The antibodies are assayed by any of the standard techniques (see Chapters 204 and 205).

4. Notes 1. We have produced antisera to protein bands in acetic acid–urea gels (see Chapter 27), Triton–acetic acid–urea gels (15, 16) (see Chapter 28), or SDS–polyacrylamide gels (see Chapter 21). In our experience, antibodies produced to proteins in one denaturing gel system will crossreact to those same proteins fractionated in another denaturing gel system and will usually crossreact with the native protein. We have consistently obtained antibodies from animals immunized by these procedures. 2. It is extremely important that all glutaraldehyde be removed from the gel during the destaining washes, since any residual glutaraldehyde will be toxic to the animal. Residual glutaraldehyde can easily be detected by smell. It is equally important to remove all acetic acid during lyophilization. Monoacrylamide is also toxic, whereas polyacrylamide is not. We do observe, however, that approx 50 mm2 of polyacrylamide per injection is the maximum that a mouse can tolerate. 3. Freund’s complete adjuvant is used for the initial immunization; Freund’s incomplete adjuvant is used for all subsequent injections. The mycobacteria in complete adjuvant enhance the immune response by provoking a local inflammation. Additional doses of mycobacteria may be toxic. 4. High-titer antibodies have been produced from proteins in polyacrylamide gel by injecting the gel/protein mixture into the lumen of a perforated plastic golf ball implanted subcutaneously in rabbits (17). This approach places less stress on the animal, as complete adjuvants need not be used, and bleeding is eliminated. The technique has also been used in rats. References 1. Chase, M. W. (1967) Production of antiserum, in Methods in Immunology and Immunochemistry, vol. I (Williams, C. A. and Chase, M. W., eds.), Academic, New York, pp. 197–200. 2. Maurer, P. H. and Callahan, H. J. (1980) Proteins and polypeptides as antigens, in Methods in Enzymology, vol. 70 (Van Vunakis, H. and Langne, J. J., eds.), Academic, New York, pp. 49–70. 3. Knudson, K. A. (1985) Proteins transferred to nitrocellulose for use as immunogens. Anal. Biochem. 147, 285–288.

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4. Tjian, R., Stinchcomb, D., and Losick, R. (1974) Antibody directed against Bacillus subtilis σ factor purified by sodium dodecyl sulfate slab gel electrophoresis. J. Biol. Chem. 250, 8824–8828. 5. Elgin, S. C. R., Silver, L. M., and Wu, C. E. C. (1977) The in situ distribution of drosophila non-histone chromosomal proteins, in The Molecular Biology of he Mammalian Genetic Apparatus, vol. 1 (Ts’o, P.O.P., ed.), North Holland, New York, pp. 127–141. 6. Silver, L. M. and Elgin, S. C. R (1978) Immunological analysis of protein distributions in Drosophila polytene chromosomes, in The Cell Nucleus (Busch, H., ed.), Academic, New York, pp. 215–262. 7. Bugler, B., Caizergucs-Ferrer, M., Bouche, G., Bourbon, H., and Alamric, F. (1982) Detection and localization of a class of proteins immunologically related to a 100kDa nucleolar protein. Eur. J. Biochem. 128, 475–480. 8. Wood, D. M. and Dunbar, B. S. (1981) Direct detection of two-cross-reactive antigens between porcine and rabbit zonae pellucidae by radioimmunoassay and immunoelectrophoresis. J. Exp. Zool. 217, 423–433. 9. Howard, C. D., Abmayr, S. M., Shinefeld, L. A., Sato, V. L., and Elgin, S. C. I. (1981) Monoclonal antibodies against a specific nonhistone chromosomal protein of Drosophila associated with active genes. J. Cell Biol. 88, 219–225. 10. Tracy, R. P., Katzmann, J. A., Kimlinger, T. K., Hurst, G. A., and Young, D. S. (1983) Development of monoclonal antibodies to proteins separated by two dimensional gel electrophoresis. J. Immunol. Meth. 65, 97–107. 11. James, T. C. and Elgin, S. C. R. (1986) Identification of a nonhistone chromosomal protein associated with heterochromatin in Drosophila melanogaster and its gene. Mol. Cell Biol. 6, 3862–3872. 12. Reichli, M. (1980) Use of glutaraldehyde as a coupling agent for proteins and peptides, in Methods in Enzymology, vol. 70 (Van Vunakis, H. and Langone, J. J., eds.), Academic, New York, pp. 159–165. 13. Campbell, A. M. (1984) Monoclonal Antibody Technology. Elsevier, New York. 14. Silver, L. M. and Elgin, S. C. R. (1977) Distribution patterns of three subfractions of Drosophila nonhistone chromosomal proteins: Possible correlations with gene activity. Cell 11, 971–983. 15. Alfagame, C. R., Zweidler, A., Mahowald, A., and Cohen, L. H. (1974) Histones of Drosophila embryos. J. Biol. Chem. 249, 3729–3736. 16. Cohen, L. H., Newrock, K. M., and Zweidler, A. (1975) Stage-specific switches in histone synthesis during embryogenesis of the sea urchin. Science 190, 994–997. 17. Ried, J. L., Everad, J. D., Diani, J., Loescher, W. H., and Walker-Simmons, M. K. (1992) Production of polyclonal antibodies in rabbits is simplified using perforated plastic golf balls. BioTechniques 12, 661–666.

171 Raising Highly Specific Polyclonal Antibodies Using Biocompatible Support-Bound Antigens Monique Diano and André Le Bivic 1. Introduction Highly specific antibodies directed against minor proteins, present in small amounts in biological fluids, or against insoluble cytoplasmic or membraneous proteins, are often difficult to obtain. The main reasons for this are the small amounts of protein available after the various classical purification processes and the low purity of the proteins. In general, a crude or partially purified extract is electrophoresed on an SDSpolyacrylamide (SDS-PAGE) gel; then the protein band is lightly stained and cut out. In the simplest method, the acrylamide gel band is reduced to a pulp, mixed with Freund’s adjuvant, and injected. Unfortunately, this technique is not always successful. Its failure can probably be attributed to factors such as the difficulty of disaggregating the acrylamide, the difficulty with which the protein diffuses from the gel, the presence of SDS in large quantities resulting in extensive tissue and cell damage, and finally, the toxicity of the acrylamide. An alternative technique is to extract and concentrate the proteins from the gel by electroelution but this can lead to loss of material and low amounts of purified protein. Another technique is to transfer the separated protein from an SDS-PAGE gel to nitrocellulose. The protein-bearing nitrocellulose can be solubilized with dimethyl sulfoxide (DMSO), mixed with Freund’s adjuvant, and injected into a rabbit. However, although rabbits readily tolerate DMSO, mice do not, thus making this method unsuitable for raising monoclonal antibodies. To obtain highly specific antibodies the monoclonal approach has been considered as the best technique starting from a crude or partially purified immunogen. However, experiments have regularly demonstrated that the use of highly heterogeneous material for immunization never results in the isolation of

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clones producing antibodies directed against all the components of the mixture. Moreover, the restricted specificity of a monoclonal antibody that usually binds to a single epitope of the antigenic molecule is not always an advantage. For example, if the epitope is altered or modified (i.e., by fixative, Lowicryl embedding, or detergent), the binding of the monoclonal antibody might be compromised, or even abolished. Because conventional polyclonal antisera are complex mixtures of a considerable number of clonal products, they are capable of binding to multiple antigenic determinants. Thus, the binding of polyclonal antisera is usually not altered by slight denaturation, structural changes, or microheterogeneity, making them suitable for a wide range of applications. However, to be effective, a polyclonal antiserum must be of the highest specificity and free of irrelevant antibodies directed against contaminating proteins, copurified with the protein of interest and/or the proteins of the bacterial cell wall present in the Freund’s adjuvant. In some cases, the background generated by such irrelevant antibodies severely limits the use of polyclonal antibodies. A simple technique for raising highly specific polyclonal antisera against minor or insoluble proteins would be of considerable value. Here, we describe a method for producing polyclonal antibodies, which avoids both prolonged purification of antigenic proteins (with possible proteolytic degradation) and the addition of Freund’s adjuvant and DMSO. Two-dimensional gel electrophoresis leads to the purification of the chosen protein in one single, short step. The resolution of this technique results in a very pure antigen, and consequently, in a very high specificity of the antibody obtained. It is a simple, rapid, and reproducible technique. Two-dimensional (2D) electrophoresis with ampholines for the isoelectric focusing (IEF) is still considered by most as time consuming and technically demanding. The introduction, however, of precast horizontal Ampholines or Immobilines IEF gels has greatly reduced this objection. Moreover, the development of 2D gel databases and the possibility to link it to DNA databases (Swiss-2D PAGE database) (1) further increases the value of using 2D electrophoresis to purify proteins of interest. Our technique was tested using two kinds of supports, nitrocellulose and polyvinylidene fluoride (PVDF), for the transfer of the protein. Both supports were tested for biocompatibility in rats and rabbits and were readily tolerated by the animals. Because PVDF is now used routinely for the analysis of proteins and peptides by mass spectrometry, it was of primary importance to test this as a prerequisite. Even if 2D electrophoresis is a time-consuming technique it actually allows one to isolate several antigens in the same experiment, which is very important for applications such as maps of protein expression between wild-type and mutant cells or organisms or identification of proteins belonging to an isolated functional complex.

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A polyclonal antibody, which by nature cannot be monospecific, can, if its titer is very high, behave like a monospecific antibody in comparison with the low titers of irrelevant antibodies in the same serum. Thus, this method is faster and performs better than other polyclonal antibody techniques while retaining all the advantages of polyclonal antibodies. 2. Materials 1. For 2D gels, materials are those described by O’Farrell (2,3) and Laemmli (4). It should be noted that for IEF, acrylamide and bis-acrylamide must be of the highest level of purity, and urea must be ultrapure (enzyme grade) and stirred with Servolit MB-1 from Serva (0.5 g of Servolit for 30 g of urea dissolved in 50 mL of deionized water) as recommended by Görg (5). 2. Ampholines with an appropriate pH range, for example, 5–8 or 3–9. 3. Transfer membranes: 0.45-µm BA 85 nitrocellulose membrane filters (Schleicher and Schull GmBH, Kassel, Germany); 0.22-µm membranes can be used for low molecular weight antigenic proteins. 4. Transfer buffer: 20% methanol, 150 mM glycine, and 20 mM Tris, pH 8.3. 5. Phosphate-buffered saline (PBS), sterilized by passage through a 0.22-µm filter. 6. Ponceau red: 0.2% in 3% trichloroacetic acid. 7. Small scissors. 8. Sterile blood-collecting tubes, with 0.1 M sodium citrate, pH 6, at a final concentration of 3.2%. 9. Ultrasonication apparatus, with 100 W minimum output. We used a IOO-W ultrasonic disintegrator with a titanium exponential microprobe with a tip diameter of 3 mm (1/8 in.). The nominal frequency of the system is 20 kc/s, and the amplitude used is 12 p.

3. Methods This is an immunization method in which nitrocellulose-bound protein purified by 2D electrophoresis is employed and in which neither DMSO nor Freund’s adjuvant is used, in contrast to the method described by Knudsen (6). It is equally applicable for soluble and membrane proteins. 3.1. Purification of Antigen In brief, subcellular fractionation of the tissue is carried out to obtain the fraction containing the protein of interest. This is then subjected to separation in the first dimension by IEF with Ampholines according to O’Farrell’s technique or by covalently bound Immobilines gradients. Immobilines allow a better resolution by generating narrow pH gradients (1:2000 and if there is a strong specific signal without any background. We have also immunized mice with nitrocellulose-bound protein by intraperitoneal injection of the powder. This is convenient when time and material are very limited, since very little protein is needed to induce a response (3–5× less than for a rabbit) and since the time lag for the response is shorter (the second immunization was 2 wk after the first, and the third

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immunization, 10 d after the second). Mice have a high tolerance for the nitrocellulose powder. Unfortunately, the small amount of serum available is a limiting factor. This technique for immunizing mice can, of course, be used for the preparation of monoclonal antibodies. For people who need to raise antibodies against Drosophila melanogaster proteins, rats are better hosts than rabbits (mice too with the above restrictions). Nitrocellulose is injected on the top of the shoulder of the rats. The foot pad is also adequate as a site of priming injection for small quantities. Blood is collected from the tail. Utilization of serum. The proper dilutions are determined. We routinely use 1:4000 for blots, 1:300–1:200 for immunofluorescence, and 1:50 for immunogold staining. Serum continues to recognize epitopes on tissue proteins after Lowicryl embedding. Labeling is highly specific, and gives a sharp image of in situ protein localization. There is no need to purify the serum. IgG purified from serum by whatever means usually gives poorer results than those obtained with diluted serum. Purification procedures often give rise to aggregates and denaturation, always increase the background, and result in loss of specific staining. Bacterial antigenic protein. When antibodies are used in screening cDNA libraries in which the host is Escherichia coli, the antibodies produced against bacterial components of Freund’s adjuvant may also recognize some E. coli components. An advantage of our technique is that it avoids the risk of producing antibodies against such extraneous components. Is nitrocellulose antigenic? Some workers have been unable to achieve good results by immunization with nitrocellulose-bound protein. They reproducibly obtain antibodies directed against nitrocellulose. We found out that it is the result from injecting powdered nitrocellulose in Freund’s adjuvant; using adjuvant actually increases the production of low-affinity IgM that binds nonspecifically to nitrocellulose. We have never observed this effect in our experiments when the technique described here was followed strictly. The same observations applied to PVDF membranes. The purification step by 2D electrophoresis implies the use of denaturing conditions (SDS), and thus is not appropriate for obtaining antibodies directed against native structures. For that purpose, the protein should be transferred onto nitrocellulose after purification by classical nondenaturing methods and gel electrophoresis under nondenaturing conditions. However, it should be pointed out that, following the method of Dunn (10), it is possible partially to renature proteins with modifications of the composition of the transfer buffer. Second dimension electrophoresis can be carried out with a first electrophoresis under native conditions, followed by a second electrophoresis

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under denaturing conditions, that is, with SDS. Because the resolution provided by a gradient is better, it should always be used in the 2D electrophoresis. Agarose may also be used as an electrophoresis support. 14. If only a limited amount of protein is available, and/or if the antigen is weakly immunogenic, another procedure may be used. The first immunization is given as a single injection, of up to 0.8 mL, into the popliteal Iymphatic ganglion (10), using a 22-gauge needle, that is, the finest that can be used to inject nitrocellulose powder. In this case, the antigen is delivered immediately into the immune system. If necessary, both ganglions can receive an injection. The small size of the ganglions limits the injected volume. The eventual excess of antigen solution is then injected into the back of the rabbit, as described in Methods. The boosters are given in the classic manner, that is, in several subcutaneous injections into the rabbit’s back. If the amount of protein available is even more limited, a guinea pig may be immunized (first immunization in the Iymphatic ganglion, and boosters, as usual). 15. The advantage of getting a high titer for the antibody of interest is that the amount of irrelevant antibodies is, by comparison, very low and, consequently, does not generate any background. Another advantage of using a crude serum with a high antibody titer is that this serum may be used without further purification to screen a cDNA expression library (12). 16. The time required for transfer and the intensity used are dependent on the molecular weight and the nature of the protein to be transferred (hydrophilic or hydrophobic). During transfer, the electrophoresis tank may be cooled with tap water. We, however, prefer semidry transfer, which is time and buffer saving and cooling is not necessary. References 1. Pennington, S. (1994) 2-D protein gel electrophoresis: an old method with future potential. Trends Cell Biol. 4, 439–441. 2. O’Farrel, P. H. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–4021. 3. O’Farrel, P. Z., Goodman, H. M., and O’Farrel, P. H. (1977) High resolution twodimensional electrophoresis of basic as well as acidic proteins. Cell 12, 1133–1142. 4. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature (Lond.) 227, 680–685. 5. Görg, A. (1998) A laboratory manual. http://www.weihenstephan.de/blm/deg. 6. Knudsen, K. A. (1985) Proteins transferred to nitrocellulose for use as immunogens. Analyt. Biochem. 147, 285–288. 7. Burnette, W. N. (1981) Electrophoretic transfer of proteins from sodium dodecylsulfate polyacrylamide gels to unmodified nitrocellulose and radiographic detection with antibody and radioiodinated protein A. Analyt. Biochem. 112, 195–203.

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8. Vaitukaitis, J., Robbins, J. B., Nieschlag, E., and Ross, G. T. (1971) A method for producing specific antisera with small doses of immunogen. J. Clin. Endocrinol. 33, 98–988. 9. Dunn, M. J. and Patel, K. (1988) 2-D PAGE using flat bed IEF in the first dimension, in Methods in Molecular Biology, Vol. 3: New Protein Techniques (Walker, J. M., ed.), Humana Press, Totowa, NJ, pp. 217–232. 10. Dunn, S. D. (1986) Effects of the modification of transfer buffer composition and the renaturation of proteins in gels on the recognition of proteins on Western blots by monoclonal antibodies. Analyt. Biochem. 157, 144–153. 11. Sigel, M. B., Sinha, Y. N., and Vanderlaan, W. P. (1983) Production of antibodies by inoculation into lymph nodes. Meth. Enzymol. 93, 3–12. 12. Preziosi, L., Michel, G. P. F., and Baratti, J. (1990) Characterisation of sucrose hydrolizing enzymes of Zymomonas mobilis. Arch. Microbiol. 153, 181–186.

172 Production of Antisera Using Peptide Conjugates Thomas E. Adrian 1. Introduction Because an immunogen requires both an antigenic site and a T-cell receptor binding site, a minimum size is necessary (1). Natural immunogens have a molecular weight >5000. Synthetic fragments of proteins may be able to bind to the surface of B cells, but do not stimulate an immune response. Such molecules are known as haptens. A hapten is an incomplete immunogen but can be made immunogenic by coupling to a suitable carrier molecule. A variety of different cross-linking agents are utilized for coupling of peptides to carrier proteins; examples of each type are covered in this chapter. In the case of larger fragments (approximately 40 amino acids), it may be possible to stimulate an immune response by presenting the peptide together with a carrier such as polyvinylpyrrolidone without the need for conjugation. This method, which is also described in this chapter, has proven to be useful for the author for a number of different peptides. Unfortunately, however, the success of this method compared with the responses to conjugated peptides is largely a matter of trial and error, and peptide fragments of this size are expensive to produce. 2. Materials 2.1. Synthetic Peptides as Haptens There is considerable advantage to be gained by using synthetic peptides as haptens. First, it is possible to raise region-specific antibodies directed perhaps to one end, or the active site of a protein. Second, it is possible to insert particular amino acids with specific side chains for coupling. For example, a peptide can be synthesized with an extra cysteine residue at one end of the molecule to enable coupling using sulfo-SMCC (2). Alternatively, a tyrosine residue can be inserted that will enable specific coupling through the bis-diazotized benzidine reaction (3). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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This latter approach is valuable because the same synthetic peptide can then serve as a radioligand for metabolic studies, for receptor binding, or for a radioimmunoassay, with the assurance that the antibodies raised will not be directed toward the tyrosine residue that will be iodinated (3). 2.2. Protein Carriers Factors governing the choice of the carrier include immunogenicity, solubility, and availability of functional groups for cross-linking. Although substances such as mucopolysaccharides, poly-L-lysine, and polyvinylpyrrolidone have been used as carriers, proteins are more widely used. Common protein carriers include serum albumin, ovalbumin, hemocyanin, and thyroglobulin. To find the very best immunogen it would be ideal to prepare conjugates with several different carriers with a range of hapten to carrier coupling ratios (see Note 1). The cost and time involved will usually make this impractical, however, and it is therefore necessary to carefully select the carrier most suitable for a particular antigen. In the classical hapten carrier system T lymphocytes recognize processed carrier determinates and cooperate with B cells which produce hapten-specific antibody response. Note that the amounts of carrier and peptide for conjugation are given in molar terms in the following conjugation protocols. This is necessary because of the wide variation in the molecular weights of potential carriers and neuropeptides to be coupled. The carrier protein represented by 100 nmol is approx 7 mg of bovine albumin, 4.5 mg of ovalbumin, 15 mg of γ-globulin, and 70 mg of thyroglobulin. 2.2.1. Bovine Serum Albumin Because of its wide availability, high solubility, and relatively high number of coupling sites albumin is a popular choice as a carrier for weakly antigenic compounds. Albumin has a molecular weight of 67,000 and has 59 lysine residues providing primary amines useful for conjugation. A drawback in the use of albumin for conjugation is that it is frequently used in immunoassays as a non-specific blocking protein. Anti-albumin antibodies induced by its use as a carrier protein can result in false positive results in such systems. 2.2.2. Ovalbumin Ovalbumin (egg albumin) also has wide availability, as it is the primary protein constituent of egg white. This protein is smaller than serum albumin with a molecular weight of 45,000, but contains 20 lysine residues, 14 aspartic acid, and 33 glutamic acid residues for conjugation (4). Ovalbumin exists as a single polypeptide chain with an isoelectric point of 4.6. Half of its 400 residues are hydrophobic. Caution should be exercised in handling of albumin, as it is denatured at temperatures above 56°C or even by vigorous shaking.

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2.2.3. Hemocyanin Keyhole limpit hemocyanin (KLH) is a copper-containing protein that belongs to a family of nonheme proteins found in arthropods and mollusca. KLH exists in five different aggregate states at neutral pH that will dissociate into subunits above pH 9.0 (5). KLH is a valuable carrier protein because of its large molecular mass (approx 1 × 106 to 1 × 107) and numerous lysine groups for coupling. This property of dissociation at high pH can be utilized because it increases the availability of angiogenic sites, and this can produce improved antigenic responses (5). The disadvantage of using KLH as a carrier protein is its poor water solubility. Although this makes the protein difficult to handle, it does not impair its immunogenicity. 2.2.4. Thyroglobulin Thyroglobulin is another large molecular weight protein with a limited solubility. The advantage of thyroglobulin as a carrier comes from its large content of tyrosine residues which can be used for conjugation using the diazo reaction. The molecular weight of thyroglobulin is 670,000. 2.3. Coupling Agents Chemical coupling agents or cross-linkers are used to conjugate small peptide haptens to large protein carriers. The most commonly used cross-linking agents have functional groups that couple to amino acid side chains of peptides (see Table 1). A host of different cross-linkers are commercially available, each with different characteristics regarding chemical groups that they are reactive toward, pH of coupling, solubility, and ability to be cleaved. The most comprehensive listing of coupling reagents is found in the Pierce catalog (Pierce, Rockford, IL; http://www.piercenet.com). This user friendly website now includes a table of all the available cross-linking agents (called: “Crosslinkers at a Glance”) and a crosslinker selection guide to identify suitable coupling agents for specific linkage sites. Several things need to be taken into consideration when selecting a bifunctional coupling reagent. First is the selection of functional groups; this can be used to produce a specific type of conjugate. For example, if the only primary amine available is in the amino-(N)-terminal end of a peptide, then a coupling agent can be selected to specifically couple in this position, leaving the carboxyl (C)-terminal end of the peptide free and available as an antigenic site. Of course, a synthetic peptide can be engineered to contain an amino acid with a specific side chain available for coupling. This is usually placed at one end of the peptide, again making the other end available as an antigenic site. Second is the length of the cross bridge; the presence of a spacer arm may make the hapten more available and therefore produce a better immune response. Third is whether the cross-linking groups are the same (homobifunctional) or different

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(heterobifunctional). Once again this can alter the specificity of the coupling reaction. The last consideration is whether the coupling reaction is chemical or photochemical. For a good antigenic response it is necessary to maintain the native structure of the protein complex, and this can be achieved only using mild buffer conditions and near neutral pH. The reactive groups that can be targeted using cross-linkers include primary amines, sulfhydryls, carbonyl, and carboxylic acids (see Note 2). It is difficult to predict the proximity of protein– peptide interactions. The use of bifunctional reagents with spacer arms can prevent steric hindrance and make the hapten more available for producing a good immune response. 2.3.1. Carbodiimide Carbodiimide condenses any free carboxyl group (nonamidated C-terminal aspartate or glutamate residue) or primary amino group (N-terminal or lysyl residue), to form a peptide bond (CO–NH). The most commonly used water-soluble carbodiimide is 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide hydrochloride (CDI or EDC). This coupling agent is very efficient and easy to use and usually couples at several alternative points on a peptide, giving rise to a variety of antigenic responses (6). Because of the unpredictable nature of the antibody responses, using this bifunctional agent the process has been termed “shotgun immunization.” If it is necessary to raise antisera to a particular region of the peptide then this is not the method of choice. Furthermore, because the peptide bond linking the hapten to the carrier cannot rotate and holds the hapten physically close, steric hindrance is considerable. Carbodiimide reacts with available carboxyl groups to form an active O-acylurea intermediate, which is unstable in aqueous solution, making it ineffective in two-step conjugation procedures. This intermediate then reacts with the primary amine to form an amide derivative. Failure to immediately react with an amine results with hydrolysis of the intermediate. Furthermore, hydrolysis of CDI itself is a competing reaction during the coupling and is dependent on temperature 4-morphylinoethansulfonic acid (MES) can be used as an effective carbodiimide reaction buffer in place of water. Phosphate buffers reduce the efficiency of the CDI reaction, although this can be overcome by increasing the amount of CDI used to compensate for the reduction and efficiency. Loss of efficiency of the CDI reaction is even greater with Tris, glycine, and acetate buffers, and therefore use of these these should be avoided. 2.3.2. Glutaraldehyde Glutaraldehyde links primary amino groups (either the N-terminal or lysyl residues) on both the peptide hapten and the carrier. This linkage allows free

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rotation of the hapten, which reduces possible steric hindrance which may otherwise block access to the immune system by the large carrier molecule. 2.3.3. Sulfo-Succinimidyl 4-(N-maleimidomethyl) Cyclohexane-1-Carboxylate (Sulfo-SMCC) A peptide with a free sulfhydryl group, such as a synthetic peptide with a terminal cysteine residue, provides a highly specific conjugation site for reacting with sulfo-SMCC. This cross-linker contains a maleimide group that reacts with free sulfhydryl groups, along with an N-hydroxysuccinimidyl ester group that reacts with primary amines (2). All peptide molecules coupled using this chemistry display the same basic antigenic conformation. They have a known and predictable orientation leaving the molecule free to interact with the immune system. This method can preserve the major epitopes on a peptide while enhancing the immune response. The water solubility of sulfo-SMCC, along with its enhanced maleimide stability, makes it a favorite for hapten carrier conjugation. 2.3.4. Bis-diazotized Benzidine Bis-diazonium salts bridge tyrosyl or hystidyl residues between the hapten and carrier. Overnight treatment of benzidine at 4°C with nitrous acid (hydrochloric acid and sodium nitrite) results in the two amino groups being diazotized. These two diazonium groups allow coupling at both ends of the molecule. Although limited in coupling points, diazotized benzidine provides a spacer arm holding the hapten away from the carrier and usually results in an excellent antigenic response. 2.4. Adjuvants The adjuvant is important for inducing an inflammatory response. The author has had continuous success over two decades using Freund’s adjuvant. Various synthetic adjuvants are available such as AdjuPrime (Pierce) and Ribi Adjuvant System (Ribi Immunochemicals) (7). The author has had very limited success with the latter adjuvant, when used to immunize with several different small peptides. In contrast, responses with Freund’s adjuvant run in parallel have always been good. The conjugate is injected in the form of an emulsion made with Freund’s adjuvant, which is a mixture of one part of detergent (Arlacel A, Sigma Chemicals) with four parts of n-hexadecane. This permits slow release of the coupled hapten into the circulation and may serve to protect labile antigens from degradation. Freund’s adjuvant alone (“incomplete”) causes an inflammatory response that stimulates antibody formation, and when made “complete” by addition of 1 mg/mL of heat-killed Mycobacterium butyricum, this response is further enhanced. It is convenient to purchase complete and incomplete Freund’s adjuvant ready mixed (Sigma Chemicals or Calbiochem).

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When preparing the emulsion, care should be taken to ensure that the oil remains in the continuous phase. Injection of aliquots of the aqueous conjugate solution into the oil via a fine bore needle, followed by repeated aspiration and ejection of the crude emulsion, will produce the required result. An alternative for making the emulsion is to use a homogenizer, although the generator should be retained specifically for this purpose to avoid subsequent peptide contamination. A simple test for the success of the preparation is to add a drop of the emulsion to the surface of water in a tube. The emulsion should stay in a single droplet without dispersing; confirming that it is immiscible and thus oil-phasec ontinuous. 2.5. Choice of Animal for Immunization Several factors need to be considered when choosing animal species for an immunization program, including cost, ease of handling and the volumes of antisera required (see Note 3). Small animals (such as rats and mice) have low blood volumes and present difficulties with bleeding. Large animals such as sheep or goats are expensive to house particularly over long periods. Rabbits or guinea pigs provide a near optimal solution, as they are relatively cheap to house and bleeding an ear vein or cardiac puncture in guinea pigs can provide between 10 and 30 mL of plasma from each bleed. For production of monoclonal antibodies immunization of mice is required. 3. Methods 3.1. Carbodiimide Procedure 1. Dissolve the peptide to be coupled (400 nmol) and protein carrier (100 nmol) in a small volume of water (7.5 (6). Although the chloroformates are highly reactive toward protein amine groups, they also hydrolyze rapidly in aqueous solutions. Therefore, they are normally added in large excess to the protein. A typical example of this procedure is given below in which vitamin B12 is coupled to BSA: 1. Suspend 84 mg of vitamin B12 in 250 µL of dry dioxan with 6 µL of pyridine and 8 µL of diphosgene. The vitamin immediately changes color from red to purple (see Note 4).

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Leave the mixture for 15 min to react. Evaporate unreacted materials. Resuspend the chloroformates in 250 µL of dry solvent. Add approx 1, 8, 25, and 50 mg of the B12–chloroformate to 2 mL aliquots of BSA (5 mg/mL in 0.1 M bicarbonate) and leave for 4 h to react. 6. Dialyze the B12–BSA conjugates for 2 d against PBS to remove uncoupled B12 and unwanted reaction products.

The pink solutions can then be measured by scanning spectrophotometry (Fig. 3), and the amount of vitamin coupled can be measured by the increase in absorbance at 280 nm, or more accurately by measuring the absorbance at 360 nm that was only due to B12. In this case, the four conjugates were found to contain 0.1, 1.1, 2.8, and 5.2 mg of B12, which represented approx 0.5, 7, 22, and 30 molecules of B12 bound to each BSA molecule. The covalent bonding of the vitamin groups can be confirmed by using the conjugates in competitive ELISA assays. 3.1.2.3. UTILIZING A COMMERCIAL LINKER CONTAINING A HYDROXYL-SPECIFIC ISOCYANATE GROUP

There is one commercial linker, PMPI, which can be used to couple hydroxyl residues to protein sulfhydryl groups (7). One end of the linker consists of an isocyanate group, which is normally first coupled at equimolar concentrations to the SM hydroxyl residue in dry organic solvents. The other end is a maleimide residue, which couples to free protein sulfhydryl groups in aqueous buffers between pH 6.5 and 7.5. 3.1.3. Small Molecule Amine Residues SMs containing an amine residue can be coupled to proteins by simply reversing the above carbodiimide reaction. NHS is added to the protein, and it is then treated with a large excess of the water-soluble carbodiimide (EDC) to generate NHS activated carboxylic acid residues in the proteins. The NHS maintains the protein solution at around pH4.5 hence preventing the protein crosslinking to its own amine groups as they are protonated. This procedure is discussed in much greater practical detail in Subheading 3.2.1. The SM (dissolved in 0.1 M bicarbonate) is then added and left to couple for 2–16 h. After extensive dialysis, conjugation can be confirmed by spectrophotometry or non-denaturing electrophoresis. Here, however, conjugated proteins would migrate more slowly due to their loss of carboxylic acid residues. If the SM is insoluble in water, then it can be coupled to a commercial crosslinker, which contains an NHS-activated ester at one end. This is done by simply adding the SM to the linker at an equimolar ratio for 24 h in dry DMF or DMSO. This type of linkage is also required when the protein (antibody

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or enzyme) loses its activity after it is treated with EDC. I have found this to happen with AP. This type of crosslinker can also be used to create a spacer arm between the SM and the protein, but as they are more commonly used to couple antibodies and enzymes together, their practical use is described below in Subheadings 3.2.2 and 3.2.3. 3.2. The Formation of Protein-Protein Conjugates Protein–protein conjugates can be made by most of the procedures already discussed. However, there is one critical difference. Proteins have to be coupled in aqueous solutions in order to prevent their denaturization, precipitation, and loss of functional activity. In practical terms, this means that the organic solvents often needed to dissolve the carbodiimides and commercial linkers should never exceed 10% of the final reaction volumes. 3.2.1. Antibody–KLH Conjugates Antibody–KLH conjugates can be used to increase the murine–murine immune response to an antibody above and beyond that obtained solely by changing the mouse strains (8). The simplest way to prepare such conjugates is to use the water-soluble carbodiimide EDC at a large molar excess to allow for the rapid hydrolysis of both the EDC and its reactive intermediates. It is now absolutely essential to include NHS in reaction mixtures prior to the addition of the carbodiimide (3). I normally treat the antibody with the carbodiimide for 15 min to activate its carboxylic acid groups and then add the activated antibody to the KLH to enable it to bind to the KLH amine groups as described below. 1. Dialyze 1 mg (0.5–3 mL) of monoclonal antibody against distilled water for at least 4 h at 0°C. This is done to remove all preservatives (e.g., azide), and buffers containing carboxylic acid and amine groups. 2. Prepare one vial of mcKLH (20 mg in 5 mL) by dialysis against 0.1 M bicarbonate for the same length of time. 3. Add 5 mg of NHS to the antibody followed by 2 mg EDC. Leave for 15 min to activate carboxylic acid groups. 4. Add 800 µL (approx 3 mg) of the KLH solution to the above and leave this antibody/ KLH mixture to conjugate overnight. This should result in a roughly 1:1 complex given the molecular weights of IgG and KLH as 165 and 480 K, respectively. 5. Dialyze the conjugate against 0.9% saline for 24 h.

Conjugation can be confirmed by electrophoresis in SDS in 5% polyacrylamide gels (Fig. 4). The complexes (b and c) are at a much higher molecular weight than the KLH (a) and the monoclonal antibody (d and e; note: probably two antibodies!) starting components. Given the low intensity of staining of b and c, most of the complexes do not even penetrate the gel.

1722 Fig. 4. The conjugation of Antibody to KLH. Lanes 1, 2, and 3 contain 15 ug mcKLH, 20 ug of antibody-KLH conjugates and 5 ug of unconjugated antibody respectively.

Thompson 1

a

2

3 b c

d e

3.2.2. Antibody–Enzyme Conjugates It is possible to conjugate antibodies to enzymes exactly as is given for KLH in Subheading 3.2.1. However, the carbodiimide-derivatized antibody can lose much of its activity. The complexes can contain several crosslinked molecules of each component! The only thing you can control is the ratio of each component to another compound. Most antibody–enzyme conjugates are therefore produced in more precise conditions using well-defined crosslinkers. The most popular consist of an NHS-ester linked to a maleimide group via a spacer arm (9). The spacer can be a benzene ring [m-maleimidobenzoyl-NHS ester (MBS)], cyclohexane [succinimidyl 4-(n-maleimido-methyl)-cyclohexane-1-carboxylate (SMCC)], or various lengths of nonaromatic carbon chain (EMCS). The selection of an appropriate spacer is very important. Rigid cyclic and aromatic spacers are themselves highly immunogeneic (9). Nonrigid carbon spacers have very little immunogenicity (9), and their flexibility can greatly enhance the sensitivity of competitive enzyme-linked immunosorbent assay (ELISAs) (10) when they are used to link SM to AP or horseradish peroxidase. NHS-esters link to primary amines at pH 7–9, whereas the maleimide groups react with sulfhydryl groups at pH 6.5–7.5. An antibody–AP conjugation procedure is given below as an example. 1. Dissolve 1–2 mg of the IgG in 1 mL 50 mM phosphate buffer, pH 7.0, containing 10 mM EDTA (see Note 5). 2. Reduce specifically at the hinge region by the addition of 2-mercaptoethylamine (6 mg in 100 µL buffer) for 90 min at 37°C. 3. Separate the reduced antibody from the reducing agent on a P10 desalting column using the same phosphate buffer containing EDTA. If 1-mL fractions are eluted,

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4.

5. 6. 7. 8.

9.

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the reduced antibody normally elutes in the fourth and fifth tubes and is stable for several hours in the presence of EDTA. Dissolve one vial of AP (10,000 units, approx 4 mg) in 1 mL distilled water and prepare for coupling by dialyzing against 50 mM phosphate, pH 7.5, to remove amine-containing buffers and preservatives (see Note 6). Dissolve 1 mg of the crosslinker (EMCS) in 100 µL DMSO and add immediately to the dialyzed AP (see Note 7). Leave 30 min (no longer!) for the amine groups of the AP to couple to the NHSester end of the linker. Remove excess unreacted linker on a P10 column. Add the AP-EMCS fraction immediately to the reduced antibody and leave the final mixture for 2–16 h for the reduced antibody to react with the maleimide end of the EMCS–AP complex. Dialyze to remove the EDTA from the antibody–EMCS–AP conjugate.

The antibody–EMCS–AP conjugate is checked by SDS electrophoresis in an 8% polyacrylamide gel (Fig. 5). Very little unreacted antibody and AP can be seen in lane 4. 3.2.3. Antibody–Antibody Conjugates Antibody–antibody conjugates are prepared in exactly the same way as for anti-body–AP conjugates, which have just been described. Equal amounts of each antibody are used. The only problem to be considered is that one of the antibodies has to be linked via its amine groups. This can inactivate an antibody when the antibody has a susceptible amine residue in its binding region. However, a highly susceptible antibody is always reduced and linked via its hinge sulfhydryl residue to the maleimide group. It is unlikely that both antibodies would be susceptible to a loss in activity when they are linked through their amine residues. Some workers prefer to use the thiolating groups SATA (11), SPDP (12), and Trauts’ reagent [iminothilane (13)] to introduce sulfhydryl groups into antibodies and enzymes rather than use a reducing agent. All these agents react spontaneously with protein amine residues in aqueous solutions. Two reagents, SATA and SPDP, produce protected sulfhydryl residues, which can be released when required for crosslinking; the third directly incorporates a free SH group, but this can be unstable under certain conditions (14). 4. Notes 1. This is by far the most commonly used coupling procedure. Always use SM and carbodiimides in dry organic solvents at 1:1 molar ratios to activate the carboxyls. If the SM is only soluble in water, a large excess of EDC (around 50X) is required, and the stabilizing NHS has to be added before you add the carbodiimide.

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Fig. 5. The conjugation of a monoclonal antibody to AP using EMCS. Lane 1 contains AP, lane 2 contains AP treated with EMCS, lane 3 contains ed antibody and lane 4 contains the antibody-AP conjugates cross-linked by the EMCS.

2. The NHS-activated SM is always added in a large molar excess (20–50fold) to the protein, and the extent of coupling required is discovered by trial and error. A pH > 7.0 is required to enable the activated ester to couple to the protein amine groups, as they have to be de-protonated to be able to

Small-Molecule–Protein Conjugation Procedures

3.

4.

5.

6.

7.

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react. At 0.1 M, sodium bicarbonate has a pH of 8.3 and is a convenient and simple coupling solution. It is very easy to couple an SM to a protein residue via a spacer arm using this technique. The NHS-activated SM in dry organic solvent is added to an equimolar solution of 6-aminocaproic acid (also in dry solvent) and left overnight. A peptide bond is formed with the amine residue on the caproic acid just as it would be with proteins, but here it is still in dry organic solvent. The SM acid group is now coupled via a five-carbon chain to another carboxylic acid. SM-COOH + 2HN-(CH2)5-COOH in the presence of DCC and NHS results in SM-CONH-(CH2)5-COOH. Another cycle of NHS and carbodiimide is added, and the SM–(CH2)5-CO–NHS-activated ester group is then coupled to the protein via the spacer. Alternatively, as this linker is formed in dry organic solutions, several cycles can be performed, and thus any length linker can be made, prior to the SM being coupled to the protein (10). Although this is a quick and reliable method of derivatizing hydroxyl residues, diphosgene dissociates into phosgene gas, which is very dangerous. Handle very carefully in a fume hood! It is very important that EDTA be present in the solutions used to reduce the antibody and in the P10 elution of the reduced antibody or it will reoxidize. It is also essential that the NHS-ester amine coupling to the enzyme be carried out at pH 7.5. If a higher pH is used (e.g., the more normal 0.1 M bicarbonate buffer, pH 8.3 for 1–2 h), the maleimide group at the other end of the linker rapidly hydrolyzes. It is then impossible to bind the maleimide end of the linker to the reduced antibody in the second stage. The crosslinker maleimide group only has a half-life of 30 min at pH > 7.5. A large excess of linker at pH 7.5 and a short reaction time (20–30 min) followed by a quick separation (of unreacted linker) in a P10 column is used to reduce this problem. Lower temperatures can also be used to reduce the rate of maleimide hydrolysis. Maleimide hydrolysis while the first protein/enzyme is coupling via its amine groups is the major problem associated with maleimide-spacer-NHS-ester crosslinkers. A five-carbon straight chain linker was used in this example (EMCS), but three other linkers [MBS, SMCC, and N-(γ-maleimidobutyryloxy) succinimide ester (GMBS)] have successfully been used in an identical procedure. More expensive sulfo-NHS derivatives of the above crosslinkers can be used if necessary. Their use is promoted because they are soluble in water. There are usually no problems using the normal linkers predissolved in a small amount of DMSO. However, some proteins and enzymes are extremely sensitive to the presence of organic solvents, and

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the sulpho-crosslinker derivatives could then be directly substituted into the conjugation procedure. References 1. Carraway, K. L. and Koshland, D. E. (1972) Carbodiimide modification of proteins. Methods Enzymol. 25, 616–623. 2. Rebek, J. and Feitler, D. (1973) An improved method for the study of reaction intermediates. The mechanism of peptide synthesis mediated by carbodiimides. J. Am. Chem. Soc. 95, 4052–4053. 3. Grabarek, Z. and Gergely, J. (1990) Zero length crosslinking procedure with the use of active esters. Anal. Biochem. 185, 131–135. 4. Thompson, S., Wong, E., Cantwell, B. M. J., and Turner, G. A. (1990) Serum alpha1-protease inhibitor with abnormal properties in ovarian cancer. Clin. Chim. Acta 193, 13–26. 5. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the bacteriophage T4. Nature [Lond.] 227, 680–685. 6. Thompson, S., Spoors, J. A., Fawcett, M.-C., and Self, C. H. (1994) Photocleavable nitrobenyl-protein conjugates, Biochem. Biophys. Res. Comm. 201, 1213–1219. 7. Annunziato, M. E., Patel, U. S., Ranade, M., and Palumbo, P. S. (1993) p-Maleimido-phenyl isocyanate: A novel heterobifunctional linker for hydroxyl to thiol coupling. Bioconjugate Chem. 4, 212–218. 8. Maruyama, M., Sperlagh, M., Zaloudik, J., et al. (2002) Immunization procedures for antiidiotypic antibody production in mice and rats. J. Immunol. Methods 264, 121–133. 9. Peeters, J. M., Hazendonk, T. G., Beuvery, E. C., and Tesser, G. I. (1989) Comparison of four bifunctional reagents for coupling peptides to proteins and the effect of the three moieties on the immunogenicity of the conjugates. J. Immunol. Methods 120, 133–143. 10. Bieniarz, C., Husain, M., Barnes, G., King, C. A., and Welch, C. J. (1996) Extended length heterobifunctional coupling agents for protein conjugations. Bioconjugate Chem. 7, 88–95. 11. Julian, R., Duncan, S., Weston, P. D., and Wrigglesworth, R. (1983) A new reagent which may be used to introduce sulphydryl groups into proteins, and its use in the preparation of conjugates for immunoassay. Anal. Biochem. 132, 68–73. 12. Carlsson, J., Drevin, H., and Axen, R. (1978) Protein thiolation and reversible protein-proteinc onjugation. Biochem. J. 173, 723–737. 13. Jue, R., Lambert, J. M. Pierce, L. R., and Traut, R. R. (1978) Addition of sulphydryl groups to Escherichia coli ribosomes by protein modification with 2-iminothiolane. Biochemistry 17, 5399–5406. 14. Singh, R., Kats, L., Blatter, W. A., and Lambert, J. M. (1996) Formation of N-substituted 2-iminothiolanes when amino groups in proteins and peptides are modified by 2-iminothiolane. Anal. Biochem. 236, 114–125.

174 The Chloramine T Method for Radiolabeling Protein Graham S. Bailey 1. Introduction Many different substances can be labeled by radioiodination. Such labeled molecules are of major importance in a variety of investigations, e.g., studies of intermediary metabolism, determinations of agonist and antagonist binding to receptors, quantitative measurements of physiologically active molecules in tissues and biological fluids, and so forth. In most of those studies, it is necessary to measure very low concentrations of the particular substance, and that in turn, implies that it is essential to produce a radioactively labeled tracer molecule of high specific radioactivity. Such tracers, particularly in the case of proteins, can often be conveniently produced by radioiodination. Two γ-emitting radioisotopes of iodine are widely available, 125I and 131I. As γ-emitters, they can be counted directly in a well-type crystal scintillation counter (commonly referred to as a γ counter) without the need for sample preparation in direct contrast to β-emitting radionuclides, such as 3H and 14C. Furthermore, the count rate produced by 1 g atom of 125I is approx 75 and 35,000 times greater than that produced by 1 g atom of 3H and 14C, respectively. In theory, the use of 131 I would result in a further sevenfold increase in specific radioactivity. However, the isotopic abundance of commercially available 131I rarely exceeds 20% owing to contaminants of 127I, and its half-life is only 8 d. In contrast, the isotopic abundance of 125I on receipt in the laboratory is normally at least 90% and its half-life is 60 d. Also, the counting efficiency of a typical well-type crystal scintillation counter for 125I is approximately twice that for 131I. Thus, in most circumstances, 125I is the radionuclide of choice for radioiodination. Several different methods of radioiodination of proteins have been developed (1,2). They differ, among other respects, in the nature of the oxidizing agent for converting 125I− into the reactive species 125I2 or 125I+. In the main, those reactive species substitute into tyrosine residues of the protein, but substitution into From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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other residues, such as histidine, cysteine, and tryptophan, can occur in certain circumstances. It is important that the reaction conditions employed should lead on average to the incorporation of one radioactive iodine atom/molecule of protein. Greater incorporation can adversely affect the biological activity and antigenicity of the labeled protein. The chloramine T method, developed by Hunter and Greenwood (3), is probably the most widely used of all techniques of protein radioiodination, and is used extensively for the labeling of antibodies. It is a very simple method in which the radioactive iodide is oxidized by chloramine T in aqueous solution. The oxidation is stopped after a brief period of time by addition of excess reductant. Unfortunately, some proteins are denatured under the relatively strong oxidizing conditions, so other methods of radioiodination that employ more gentle conditions have been devised, e.g., the lactoperoxidase method (see Chapter 175). 2. Materials Na 125I: 37 MBq (1 mCi) concentration 3.7 GBq/mL (100 mCi/mL). Buffer A: 0.5 M sodium phosphate buffer, pH 7.4 (see Note 1). Buffer B: 0.05 M sodium phosphate buffer, pH 7.4. Buffer C: 0.01 M sodium phosphate buffer containing 1 M sodium chloride, 0.1% bovine serum albumin, and 1% potassium iodide, final pH 7.4. 5. Chloramine T solution: A 2 mg/mL solution in buffer B is made just prior to use (see Note 2). 6. Reductant: A 1 mg/mL solution of sodium metabisulfite in buffer C is made just prior to use. 7. Protein to be iodinated: A 0.5–2.5 mg/mL solution is made in buffer B.

1. 2. 3. 4.

3. Method 1. Into a small plastic test tube (1 × 5.5 cm) are added successively the protein to be iodinated (10 µL), radioactive iodide (5 µL), buffer A (50 µL), and chloramine T solution (25 µL) (see Notes 3 and 4). 2. After mixing by gentle shaking, the solution is allowed to stand for 30 s to allow radioiodination to take place (see Note 5). 3. Sodium metabisulfite solution (500 µL) is added to stop the radioiodination, and the resultant solution is mixed. It is then ready for purification as described in Chapter 178 and Note 6.

4. Notes 1. The pH optimum for the iodination of tyrosine residues of a protein by this method is about pH 7.4. Lower yields of iodinated protein are obtained at pH values below about 6.5 and above about 8.5. Indeed, above pH 8.5 the iodination of histidine residues appears to be favored. 2. If the protein is seriously damaged by the use of 50 µg of chloramine T, it may be worthwhile repeating the radioiodination using much less oxidant

The Chloramine T Method for Radiolabeling Protein

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4.

5.

6.

1729

(10 µg or less). Obviously, the minimum amount of chloramine T that can be used will depend, among other factors, on the nature and amount of protein to be iodinated. The total volume of the reaction should be as low as practically possible to achieve a rapid and efficient incorporation of the radioactive iodine into the protein. It is normal to carry out the method at room temperature. However, if the protein is especially labile, it may be beneficial to run the procedure at a lower temperature and for a longer period of time. Because of the small volumes of reactants that are employed, it is essential to ensure adequate mixing at the outset of the reaction. Inadequate mixing is one of the most common reasons for a poor yield of radioiodinated protein by this procedure. It is possible to carry out this type of reaction using an insoluble derivative of the sodium salt of N-chloro-benzene sulfonamide as the oxidant. The insoluble oxidant is available commercially (Iodo-Beads, Pierce, Rockford, IL). It offers a number of advantages over the employment of soluble chloramine T. It produces a lower risk of oxidative damage to the protein, and the reaction is stopped simply by removing the beads from the reaction mixture, thus avoiding any damage caused by the reductant.

References 1. Bolton, A. E. (1985) Radioiodination Techniques, 2nd ed. Amersham International, Amersham, Bucks, UK. 2. Bailey, G. S. (1990) In vitro labeling of proteins, in Radioisotopes in Biology (Slater, R. J., ed.), IRL, Oxford, UK, pp. 191–205. 3. Hunter, W. M. and Greenwood, F. C. (1962) Preparation of iodine-131 labeled human growth hormone of high specific activity, Nature 194, 495,496.

175 The Lactoperoxidase Method for Radiolabeling Protein Graham S. Bailey 1. Introduction This method, introduced by Marchalonis (1), employs lactoperoxidase in the presence of a trace of hydrogen peroxide to oxidize the radioactive iodide 125 − I to produce the reactive species 125I2 or 125I+. These reactive species substitute mainly into tyrosine residues of the protein, although substitution into other amino acid residues can occur under certain conditions. The oxidation can be stopped by simple dilution. Although the technique should result in less chance of denaturation of susceptible proteins than the chloramine T method, it is more technically demanding and is subject to a more marked variation in optimum reaction conditions. 2. Materials 1. Na 125I: 37 MBq (1 mCi) concentration 3.7 GBq/mL (100 mCi/mL). 2. Lactoperoxidase: available from various commercial sources. A stock solution of 10 mg/mL in 0.1 M sodium acetate buffer, pH 5.6, can be made and stored at −20°C in small aliquots. A working solution of 20 µg/mL is made by dilution in buffer just prior to use. 3. Buffer A: 0.1 M sodium acetate buffer, pH 5.6 (see Note 1). 4. Buffer B: 0.05 M sodium phosphate buffer containing 0.1% sodium azide, final pH 7.4. 5. Buffer C: 0.05 M sodium phosphate buffer containing 1 M sodium chloride 0.1% bovine serum albumin and 1% potassium iodide, final pH 7.4. 6. Hydrogen peroxide: A solution of 10 µg/mL is made by dilution just prior to use. 7. Protein to be iodinated: A 0.5–2.5 mg/mL solution is made in buffer A.

It is essential that none of the solutions except buffer B contain sodium azide as antibacterial agent, since it inhibits lactoperoxidase.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Method 1. Into a small plastic test tube (1 × 5.5 cm) are added, in turn the protein to be iodinated (5 µL), radioactive iodide (5 µL), lactoperoxidase solution (5 µL), and buffer A (45 µL). 2. The reaction is started by the addition of the hydrogen peroxide solution (10 µL) with mixing (see Note 2). 3. The reaction is stopped after 20 min (see Note 3) by the addition of buffer B (0.5 mL) with mixing. 4. After 5 min, buffer C (0.5 mL) is added with mixing. The solution is then ready for purification as described in Chapter 178 (see Note 4).

4. Notes 1. The exact nature of buffer A will depend on the properties of the protein to be radioiodinated. Proteins differ markedly in their pH optima for radioiodination by this method (2). Obviously the pH to be used will also depend on the stability of the protein, and the optimum pH can be established by trial and error. 2. Other reaction conditions, such as amount of lactoperoxidase, amount and frequency of addition of hydrogen peroxide, and so forth, also markedly affect the yield and quality of the radioiodinated protein. Optimum conditions can be found by trial and error. 3. The longer the time of the incubation, the greater the risk of potential damage to the protein by the radioactive iodide. Thus, it is best to keep the time of exposure of the protein to the radioactive iodide as short as possible, but commensurate with a good yield of radioactive product. 4. Some of the lactoperoxidase itself may become radioiodinated, which may result in difficulties in purification if the enzyme is of a similar size to the protein being labeled. Thus, it is best to keep the ratio of the amount of protein being labeled to the amount of lactoperoxidase used as high as possible. References 1. Marchalonis, J. J. (1969) An enzymic method for trace iodination of immunoglobulins and other proteins. Biochem. J. 113, 299–305. 2. Morrison, M. and Bayse, G. S. (1970) Catalysis of iodination by lactoperoxidase. Biochemistry 9, 2995–3000.

176 The Bolton and Hunter Method for Radiolabeling Protein Graham S. Bailey 1. Introduction This is an indirect method in which an acylating reagent (N-succinimidyl-3 [4-hydroxyphenyl]propionate, the Bolton and Hunter reagent), commercially available in a radioiodinated form, is covalently coupled to the protein to be labeled (1). The [125I] Bolton and Hunter reagent reacts mostly with the sidechain amino groups of lysine residues to produce the labeled protein. It is the method of choice for radiolabeling proteins that lack tyrosine and histidine residues, or where reaction at those residues affects biological activity. It is particularly suitable for proteins that are sensitive to the oxidative procedures employed in other methods. 2. Materials 1. [125I] Bolton and Hunter reagent: 37 MBq (1 mCi), concentration 185 MBq/mL (5 mCi/mL) ( see Note 1). 2. Buffer A: 0.1 M sodium borate buffer, pH 8.5. 3. Buffer B: 0.2 M glycine in 0.1 M sodium borate buffer, pH 8.5. 4. Buffer C: 0.05 M sodium phosphate buffer containing 0.25% gelatin. 5. Protein to be iodinated: A 0.5–2.5 mg/mL solution is made in buffer A.

It is essential that none of the solutions contain sodium azide or substances with free thiol or amino groups (apart from the protein to be labeled), since the Bolton and Hunter reagent will react with those compounds. 3. Method 1. The [125I] Bolton and Hunter reagent (0.2 mL) is added to a small glass test tube (1 × 5.5 cm) and is evaporated to dryness under a stream of dry nitrogen. 2. All reactants are cooled in iced water (see Note 2). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. The protein to be iodinated (10 µL) is added, and the tube is gently shaken for 15 min ( see Note 2). 4. Buffer B (0.5 mL) is added (see Note 3). The solution is mixed and allowed to stand for 5 min. 5. Buffer C (0.5 mL) is added with mixing (see Note 4). The resultant solution is then ready for purification.

4. Notes 1. [125I] Bolton and Hunter reagent is available from Amersham International (Little Chalfort, UK) and Dupont NEN (Stevenage, UK). The Amersham product is supplied in anhydrous benzene containing 0.2% dimethylformamide. Aliquots can be easily withdrawn from the vial. However, the Dupont NEN is supplied in dry benzene alone, and dry dimethylformamide (about 0.5% of the sample volume) must be added to the vial with gentle shaking to facilitate the removal of aliquots. 2. [125I] Bolton and Hunter reagent is readily hydrolyzed in aqueous solution. Under the described conditions, its half-life is about 10 min. 3. Buffer B stops the reaction by providing an excess of amino groups (0.2 M glycine) for conjugation with the [125I] Bolton and Hunter reagent. Thus, the carrier protein (0.25% gelatin) in buffer C does not become labeled. 4. This method of radioiodination has been used extensively, and various modifications of the described procedure, including time and temperature of the reaction, have been reported (2). For example, it is possible first to acylate the protein with the Bolton and Hunter reagent, and then carry out radioactive labeling of the conjugate using the chloramine T method. However, in general, this procedure does not seem to offer any advantage over the method described here. Also, the time and temperature of the reaction can be altered to achieve optimal labeling. References 1. Bolton, A. E. and Hunter, W. M. (1973) The labeling of protiens to high specific radioactivities by conjugation to a 125I-containing acylating agent. Biochem. J. 133, 529–538. 2. Langone, J. J. (1980) Radioiodination by use of the Bolton-Hunter and related reagents, in Methods in Enzymology, vol. 70 (Van Vunakis, H. and Langone, J. J., eds.), Academic, New York, pp. 221–247.

177 Preparation of 125I-Labeled Peptides and Proteins with High Specific Activity Using IODO-GEN J. Michael Conlon 1. Introduction The reagent IODO-GEN (1,3,4, 6-tetrachloro-3α, 6α-diphenylglycoluril) was first introduced by Fraker and Speck in 1978 (1) and rapidly found widespread use for the radioiodination of both peptides and proteins (2). Although the usual application of the reagent in the laboratory is for introduction of an atom of 125I, IODO-GEN has been used successfully for the preparation of proteins, particularly monoclonal antibodies, labeled with 131I124 and 123I for use in immunoscintigraphy and positron emission tomography (3–5). This chapter addresses only the use of IODO-GEN to radiolabel pure peptides/proteins in aqueous solution, but iodination of proteins on the cell surface of a wide range of intact eukaryotic and prokaryotic cells (6,7) and on subcellular organelles (8) and plasma lipoproteins (9) has been accomplished using the reagent. The advantages of IODO-GEN over alternative reagents previously used for radiolabeling such as chloramine T, lactoperoxidase/hydrogen peroxide, and 125I-labeled Bolton–Hunter reagent (N-succinimidyl-3 [4-hydroxyphenyl] propionate) are that the reaction is rapid, technically simple to perform, and gives reproducibly high levels of incorporation of radioactivity with minimal oxidative damage to the protein. The structure of IODO-GEN is shown in Fig. 1. The reagent is virtually insoluble in water and so a solution in dichloromethane or chloroform is used to prepare a thin film of material on the walls of the reaction vessel. Addition of an aqueous solution of Na+ 125I generates the reactive intermediate (I+/I3+) that participates in electrophilic attack primarily at tyrosine but also at histidine residues in the peptide/protein. Reaction is terminated simply by transferring the contents of the reaction vessel to a clean tube, thereby obviating the need to add a reducing agent. In comparison to other radio-iodination methods, particularly the use of chloramine T and lactoperoxidase, oxidative damage to sensitive From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. The structure of IODOGEN (1,3,4,6-tetrachloro-3α, 6α-diphenylglycoluril).

residues in the peptide/protein (particularly methionine and tryptophan) is much less using IODO-GEN (10). To minimize oxidative damage even further and to limit production of diiodotyrosyl derivatives, the strategy of trace labeling is recommended. This involves radioiodination of only approx 10% of the molecules and necessitates the separation, as completely as possible, of the radiolabeled peptide from the unreacted starting material in order to obtain a tracer of sufficiently high specific activity to be of use in radioimmunoassay, radioreceptor assay, or autoradiography. Reversed-phase high-performance liquid chromatography (RP-HPLC) combines rapidity and ease of operation with optimum separation of labeled and unlabeled peptide (reviewed in (11)). The availability of wide-pore C3 and C4 columns generally permits good recoveries of labeled proteins with molecular mass (Mr) > 10,000. Techniques such as selective adsorption to diatomaceous materials (e.g., talc or microfine silica) and gel permeation chromatography give tracers of low specific activity, and ion-exchange chromatography is time consuming and results in a sample dilution that may be unacceptable. Recent applications of IODO-GEN in the field of protein and peptide radioiodination involve the use of the reagent in the preparation of N-succinimidyl 4-guanidinomethyl-3-[131I]iodobenzoate (12), N-succinimidyl 3-[131I]iodo4-phosphonomethylbenzoate (13), and N-succinimidyl-5-[131I]iodopyridine3-carboxylate (14) as superior conjugation alternatives to the Bolton–Hunter reagent, particularly for labelling monoclonal antibodies. 2. Materials 2.1. Apparatus No specialized equipment is required to carry out the iodination, but the reaction should be carried out in an efficient fume hood. Reaction takes place in a 1.5-mL natural-colored polypropylene microcentrifuge (Eppendorf tube) (see Note 1), immersed in an ice bath. A nitrogen or argon cylinder is required

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for removal of solvent. Liquids are dispensed with 10- and 100-µL pipets (e.g., Gilson Pipetman) that, because of inevitable contamination by radioactivity, should be dedicated to the reaction and stored in a designated area. For HPLC, a system capable of generating reproducible linear two solvent gradients is required. Again, because of contamination by radioactivity, an injector, for example, Rheodyne Model 7125 with 1-mL sample loop; a 1-mL leak-free injection syringe, for example, Hamilton Gastight (cat. no. 1001); a 25 × 0.46 cm analytical reversed-phase column (see Note 2 on column selection); and a fraction collector capable of collecting a minimum of 70 samples, for example, Frac 100 (Pharmacia, Uppsala, Sweden) should be dedicated to the radioiodination procedure. A UV detection system and chart recorder/integrator are not necessary unless it is important to demonstrate that the radiolabeled component has been completely separated from the starting material. 2.2. Chemicals 2.2.1. Iodination Reagents 1. IODO-GEN (Pierce, Rockford, IL): The reagent should be stored in a desiccator in the freezer and the bottle protected from light. 2. Dichloromethane (stabilized, HPLC grade): Redistillation is not required. 3. Carrier-free Na+ 125I− in 0.1 M NaOH (3.7 GBq/mL; 100 mCi/mL, Amersham Pharmacia Biotech, Piscataway, NJ). 4. 0.2 M Disodium hydrogen phosphate–sodium dihydrogen phosphate buffer, pH 7.5 (see Note 3).

2.2.2. Chromatography 1. Solvent A: Add 1 mL of trifluoroacetic acid (Pierce HPLC/Spectro grade) to 1000 mL of water (see Note 4). 2. Solvent B: Mix 700 mL of acetonitrile (Fisher optima grade) with 300 mL of water and add 1 mL of trifluoroacetic acid. The solvents should be degassed, preferably by sparging with helium for 1 min, but passage through a filter is unnecessary.

3. Method 3.1. Radioiodination 1. Dissolve 1.5 mg of IODO-GEN in 2 mL of dichloromethane. 2. Pipet 20 µL of the solution into a polypropylene tube and remove the solvent in a gentle stream of nitrogen or argon at room temperature. The aim is to produce a film of IODO-GEN on the wall of the tube. If the reagent has formed a visible clump, the tube should be discarded and a new tube prepared. According to the manufacturer’s instructions (Pierce), the tubes can be stored in a vacuum desiccator for up to 2 mo, but it is recommended that a tube is prepared freshly for each reaction. The tube is set in an ice bath for 10 min prior to the iodination (see Note 5).

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3. Dissolve 10 nmol of the peptide (see Note 6) in 0.2 M sodium phosphate buffer, pH 7.5 (100 µL) and pipet the solution into the chilled IODO-GEN-coated tube. 4. Add the Na+ 125I− solution (5 µL; 0.5 mCi or 10 µL; 1 mCi depending on the quantity of tracer required). 5. Allow the reaction to proceed for between 1 and 20 min (see Note 7). The contents of the tube should be gently agitated by periodically tapping the side of the tube with a gloved finger. 6. Reaction is stopped by aspirating the contents of the tube into a solution of 0.1% (v/v) trifluoroacetic acid–water (500 µL) contained in a second polypropylene tube. The reaction vessel may be washed with a further 200 µL of trifluoroacetic acid–water (500 µL) contained in a second polypropylene tube. The reaction vessel may be washed with a further 200 µL of trifluoroacetic acid–water and the washings combined with the reaction mixture.

3.2. Chromatography 1. Prior to carrying out the iodination, the column is equilibrated with 100% solvent A at a flow rate of 1.5 mL/min for at least 30 min. In the case of more hydrophobic peptides/proteins, the column is equilibrated with starting solvent containing up to 30% solvent B (21% acetonitrile). 2. The instrument is programmed to increase the proportion of solvent B from starting conditions to 70% (49 % acetonitrile) over 60 min using a linear gradient (see Note 8). 3. The reaction mixture is injected onto the column (see Note 9). The fraction collector, programmed to collect 1-min fractions, is started and the linear elution gradient is begun. A total of 60 fractions are collected. 4. At the end of the chromatography, the column is irrigated with 100% solvent B for 30 min. The column may be stored in this solvent. 5. The radioactivity in aliquots (2 mL) of each fraction is counted in a gamma scintillation counter (see Note 10 for optimum storage conditions of tracer).

The results of a typical radioiodination are illustrated in Fig. 2. The reaction mixture comprised 10 nmol of rat galanin and 0.5 mCi of Na+ 125I− and the reaction time was 2 min. Unreacted free iodide was eluted at the void volume of the column (fractions 5 and 6). The fraction denoted by the bar (tube 51) was of high specific activity (approx 74 TBq/mmol; 2000 Ci/mmol). The earlier eluting minor peak of radioactivity (tube 49) probably represented the diiodotyrosyl derivative. Before use in radioimmunoassay or radioreceptor studies, the quality of the tracer is assessed by incubating an aliquot (approx 20,000 cpm) with an excess of an antiserum raised against galanin (1 : 1000 dilution) in 0.1 M sodium phosphate buffer, pH 7.4 (final volume 300 µL) for 16 h at 4°C. Free and bound radioactivity are separated by addition of 100 µL of a 10 mg/mL solution of bovine γ-globulin (Sigma, St. Louis, MO) and 1 mL of 20% (w/v) solution of polyethylene glycol 6000 (Sigma, approx Mr 8000) in water followed by centrifugation (1600g for 30 min at 4°C). Under these conditions, >90% of the radioactivity is bound to antibody.

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Fig. 2. RP-HPLC on a (0.46 × 25 cm) Vydac 218TP54 (C18) column of the reaction mixture following incubation of 10 nmol of rat galanin with 0.5 mCi of Na 125I in an IODOGENcoated tube for 2 min at 0°C. Fractions (1 min) were collected and the fraction denoted by the bar contained tracer of high specific activity (74 TBq/mmol). The dashed line shows the concentration of acetonitrile in the eluting solvent.

4. Notes 1. A systematic comparison of the properties of different IODO-GEN-coated surfaces has concluded that soda-lime glass produced the most rapid rate of oxidation of Na 125I (15). However, irreversible binding of most peptides to the walls of polypropylene tubes is much less than to glass and so clear polypropylene Eppendorf tubes are routinely used in the author’s laboratory. The use of borosilicate glass tubes (15) and polysytrene tubes is not recommended. Glass test tubes (12 mm × 75 mm) precoated with IODO-GEN (50 µg) are available from Pierce. 2. The choice of column is dictated by the nature of the radiolabeled peptide to be purified. For relatively small (Mr < 3000) peptides, good resolution and recoveries are generally obtained with (0.46 × 25 cm) narrow pore (80 Å), 5-µm particle size octadecylsilane (C18) columns such as Supelcosil LC-18-DB (Supelco, Bellefonte, PA), Ultrasphere ODS (Beckman, Duarte, CA), or Spheri-5 RP-18 (Brownlee/Perkin Elmer, Foster City, CA). For purification of radiolabeled tracers of higher molecular mass (Mr > 3000), the use of columns containing wide-pore (300 Å) 5-µm particle size C18 packing materials is recommended. Suitable columns include

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5.

6.

7.

8.

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Vydac 218TP54 (Separations Group, Hesperia, CA), Spherisorb widepore C18 (Waters Corporation, U. K.), Waters Delta-Pak C18 and Ultrapore C18 (Beckman). For purification of tracers of molecular mass > 10,000, such as the pituitary glycoprotein hormones, sharper peaks and better recoveries of radioactivity may be obtained using wide-pore silica with propyl and butyl carbon loading. Suitable columns include Beckman Ultrapore C3 and Vydac 214TP54 C4 columns. The reaction is not markedly pH sensitive and high incorporations of 125I may be achieved in the pH range 6–9 (16). Suitable water can be obtained using a Milli-Q purification system (Millipore) supplied with water that has been partially purified by single distillation or by treatment with a deionization resin. A low reaction temperature is important in minimizing damage to the radiolabeled peptide/protein. For example, the use of IODO-GEN at room temperature to prepare 125I-labeled human growth hormone resulted in the production of tracer containing a significant amount of the hormone in an aggregated form whereas only the radio-labeled monomer was produced when the reaction was carried out at 0°C (17). As many peptides are relatively insoluble in buffers of neutral pH, it is recommended that the peptide first be dissolved in a minimum volume (approx 5 µL) of 0.1% (v/v) trifluoroacetic acid/water and the volume made up to 100 µL with 0.2 M sodium phosphate buffer, pH 7.5. The optimum reaction time must be determined for each peptide and protein, but some general guidelines can be given. For small peptides (90%) and can be used for most immunochemical procedures including conjugation with radioisotopes, enzymes, and so on, where pure IgG is required.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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2. Materials 1. 2. 3. 4. 5.

DEAE Sepharose CL-6B. 0.07 M sodium phosphate buffer, pH 6.3. 1 M NaCl. Sodium azide. Chromatography column (see Note 1).

3. Methods 1. Dialyze the serum (preferably ammonium sulfate fractionated; see Chapter 179) against 0.07 M sodium phosphate buffer, pH 6.3, exhaustively (at least two changes over a 24-h period) at a ratio of at least 1 vol of sample to 100 vol of buffer. 2. Apply the sample to the column, and wash the ion exchanger with 2 column volumes of sodium phosphate buffer. Collect the wash, which will contain IgG, and monitor the absorbance of the eluate at 280 nm (A280). Stop collecting fractions when the A280 falls to baseline. 3. Regenerate the column by passing through 2–3 column volumes of phosphate buffer containing 1 M NaCl. 4. Wash thoroughly in phosphate buffer (2–3 column volumes), and store in buffer containing 0.1% NaN3. 5. Pool the fractions from step 2 and measure the A280 (see Note 2).

4. Notes 1. The column size will vary according to the user’s requirements or the amount of antibody required. Matrix binding capacities are given by manufacturers and should be used as a guide. ) of human IgG is 13.6 (i.e., a 1 mg/mL 2. The extinction coefficient (E1% 280 solution has an A280 of 1.36). This can be used as an approximate value for IgGs from other sources. Reference 1. Johnstone, A. and Thorpe, R. (1996) Immunochemistry in Practice, 3rd ed. Blackwell Scientific, Oxford, UK.

182 Purification of IgG Using Ion-Exchange HPLC Carl Dolman, Mark Page, and Robin Thorpe 1. Introduction Conventional ion-exchange chromatography separates molecules by adsorbing proteins onto the ion-exchange resins that are then selectively eluted by slowly increasing the ionic strength (this disrupts ionic interactions between the protein and column matrix competitively) or by altering the pH (the reactive groups on the proteins lose their charge). Anion-exchange groups (such as diethylaminoethyl; [DEAE]) covalently linked to a support matrix (such as Sepharose) can be used to purify IgG in which the pH of the mobile-phase buffer is raised above the pI or IgG, thus allowing most of the antibodies to bind to the DEAE matrix. Compare this method with that described in Chapter 181 in which the IgG passes through the column. The procedure can be carried out using a laboratory-prepared column that is washed and eluted under gravity (1); however, high-performance liquid chromatography (HPLC) provides improved reproducibility (because the sophisticated pumps and accurate timers), speed (because of the small high-capacity columns), and increased resolution (because of the fine resins and control systems). 2. Materials 1. Anion-exchanger (e.g., Mono-Q HR 5/5 or HR 10/10, Pharmacia, Uppsala, Sweden. 2. Buffer A: 0.02 M triethanolamine, pH 7.7. 3. Buffer B: Buffer A containing 1 M NaCl. 4. 2 M NaOH. 5. Sodium azide.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Methods 1. Prepare serum by ammonium sulfate precipitation (45% saturation; see Chapter 179). Redissolve the precipitate in 0.02 M triethanolamine buffer, pH 7.7, and dialyze overnight against this buffer at 4°C. Filter the sample (see Note 1) before use (0.2 µm). 2. Assemble the HPLC system according to the manufacturer’s instructions for use with the Mono-Q ion-exchange column. 3. Equilibrate the column with 0.2 M triethanolamine buffer, pH 7.7 (buffer A). Run a blank gradient from 0 to 100% buffer B (buffer A + 1 M NaCl). Use a flow rate of 4–6 mL/min for this and subsequent steps. 4. Load the sample depending on column size. Refer to the manufacturer’s instruction for loading capacities of the columns. 5. Equilibrate the column with buffer A for at least 10 min. 6. Set the sensitivity in the UV monitor control unit, and zero the baseline. 7. Apply a salt gradient from 0 to 28% buffer B for about 30 min (see Note 2). Follow with 100% 1 M NaCl for 15 min to purge the column of remaining proteins. 8. Wash the Mono-Q ion-exchange with at least 3 column volumes each of 2 M NaOH followed by 2 M NaCl. 9. Store the Mono-Q ion-exchange column in distilled water containing 0.02% NaN3.

4. Notes 1. It is essential that the sample and all buffers be filtered using 0.2-µm filters. All buffers must be degassed by vacuum pressure. 2. Using Mono-Q, IgG elutes between 10% and 25% buffer B, usually approx 15%. When IgG elutes at 25% (dependent on pI), then it will tend to coelute with albumin, which elutes at approx 27%. When this occurs, alternative purification methods should be employed. 3. Other anion-exchange media can be substituted for Mono-Q, for example, Anagel TSK DEAE (Anachem, Luton, UK). Reference 1. Johnstone, A. and Thorpe, R. (1996) Immunochemistry in Practice, 3rd edition, Blackwell Scientific, Oxford, UK.

183 Purification of IgG by Precipitation with Polyethylene Glycol (PEG) Mark Page and Robin Thorpe 1. Introduction PEG precipitation works well for IgM, but is less efficient for IgG; salt precipitation methods are usually recommended for IgG. PEG precipitation may be preferred in multistep purifications that use ion-exchange columns, because the ionic strength of the Ig is not altered. Furthermore, it is a very mild procedure that usually results in little denaturation of antibody. This procedure is applicable to both polyclonal antisera and most MAb containing fluids. 2. Materials 1. PEG solution: 20% (w/v) PEG 6000 in PBS. 2. PBS: 0.14 M NaCl, 2.7 mM KCl, 1.5 mM KH2PO4, 8.1 mM Na2HPO4.

3. Methods 1. Cool a 20% w/v PEG 6000 solution to 4°C. 2. Prepare serum/ascitic fluid, and so forth, for fractionation by centrifugation at 10,000gav for 20–30 min at 4°C. Discard the pellet. Cool to 4°C. 3. Slowly stir the antibody containing fluid, and add an equal volume of 20% PEG dropwise (see Note 1). Continue stirring for 20–30 min. 4. Centrifuge at 2000–4000gav for 30 min at 4°C. Discard the supernatant (see Note 2), and drain the pellet. Resuspend in PBS or other buffer as described for ammonium sulfate precipitation (see Chapter 179).

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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4. Notes 1. Although the procedure works fairly well for most antibodies, it may produce a fairly heavy contamination with non-IgG proteins. If this is the case, reduce the concentration of PEG in 2% steps until the desired purification is achieved. Therefore, carry out a pilot-scale experiment before fractionating all of the sample. 2. PEG precipitation does not work for some antibodies. If the procedure is to be used for a valuable antibody for the first time, keep the supernatant in case precipitation has been inefficient.

184 Purification of IgG Using Protein A or Protein G Mark Page and Robin Thorpe 1. Introduction Some strains of Staphylococcus aureus synthesize protein A, a group-specific ligand that binds to the Fc region of IgG from many species (1,2). Protein A does not bind all subclasses of IgG, e.g., human IgG3, mouse IgG3, sheep IgG1, and some subclasses bind only weakly, e.g., mouse IgG1. For some species, IgG does not bind to protein A at all, e.g., rat, chicken, goat, and some MAbs show abnormal affinity for the protein. These properties make the use of protein A for IgG purification limited in certain cases, although it can be used to an advantage in separating IgG subclasses from mouse serum (3). Protein G (derived from groups C and G Streptococci) also binds to IgG Fc with some differences in species specificity from protein A. Protein G binds to IgG of most species, including rat and goat, and recognizes most subclasses (including human IgG3 and mouse IgG1), but has a lower binding capacity. Protein G also has a high affinity for albumin, although recombinant DNA forms now exist in which the albuminbinding site has been spliced out, and are therefore very useful for affinity chromatography. Other streptococcal immunoglobulin binding proteins are protein H (binds IgG Fc), protein B, which binds IgA and protein Arp, which binds IgG & IgA. These are not generally available for immunochemical use. Another bacterial IgG-binding protein (protein L) has been identified (4). Derived from Peptostreptococcus magnus, it binds to some κ (but not λ) chains. Furthermore, protein L binds to only some light-chain subtypes, although immunoglobulins from many species are recognized (5,6). Finally, hybrid molecules produced by recombinant DNA procedures, comprising the appropriate regions of IgG-binding proteins (e.g., protein L/G, protein L/A) also have considerable scope in immunochemical techniques. These proteins are therefore very useful in the purification of IgG by affinity chromatography. Columns are commercially available (MabTrap G II, Pharmacia, From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Uppsala, Sweden) or can be prepared in the laboratory. The product of this method is of high purity and is useful for most immunochemical procedures including affinity chromatography and conjugation with radioisotopes, enzymes, biotin, and so forth. 2. Materials 1. 2. 3. 4. 5.

PBS: 0.14 M NaCl, 2.7 mM KCl, 1.5 mM KH2 PO4, 8.1 mM Na2HPO4. Sodium azide. Dissociating buffer: 0.1 M glycine-HCl, pH 3.5. Adjust the pH with 2 M HCl. 1 M Tris. Binding buffer: Optimal binding performance occurs using a buffer system between pH 7.5 and 8.0. Suggested buffers include 0.1 M Tris-HCl, 0.15 M NaCl, pH 7.5; 0.05 M sodium borate, 0.15 M NaCl, pH 8.0; and 0.1 M sodium phosphate, 0.15 M NaCl, pH 7.5. 6. IgG preparation: serum ascitic fluid, or hybridoma culture supernatant. 7. Protein A, G column.

3. Methods Refer to Chapter 186 for CNBr activation of Sepharose and coupling of protein A, G. 1. 2. 3. 4.

5. 6. 7.

8.

9.

Wash the column with an appropriate binding buffer. Pre-elute the column with dissociating buffer, 0.1 M glycine-HCl, pH 3.5. Equilibrate the column with binding buffer. Prepare IgG sample: If the preparation is serum, plasma, or ascitic fluid, dilute it at least 1:1 in binding buffer and filter through 0.45-µm filter. Salt-fractionated preparations (see Chapter 179) do not require dilution, but the protein concentration should be adjusted to approx 1–5 mg/mL. Hybridoma culture supernatants do not require dilution. Apply sample to column at no more than 10 mg IgG/2-mL column. Wash the column with binding buffer until the absorbance at 280 nm is 95% of the F(ab′)2 is reduced. Following alkylation and overnight incubation, the reaction mixture typically elutes from HPLC as a triplet, containing a mixture of alkylated FabЭ and Fab′(mal), which elute in a similar postion to Fab′(SH), bispecific F(ab′)2 product, which elutes similarly to the parent F(ab′)2, and a smaller amount of bispecific F(ab′)3, which elutes similarly to IgG.

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4. Notes 1. Two SH groups may also be produced by the reduction of the heavy/light chain disulfide bond. However, under the conditions used, this bond is not fully reduced, and any SH groups that are produced are less likely to be available for conjugation (1,2). This procedure relies on one maleimidated hinge SH-group remaining free for conjugation after the intramolecular cross-linking of adjacent SH-groups with o-PDM. It follows that the Fab′ species chosen to be maleimidated must be derived from IgG with an odd number of hinge region disulfide bonds. Of the mouse IgG subclasses, IgG1 and IgG2a (three bonds), and IgG3 (one bond) qualify, whereas IgG2b (four bonds) does not. F(ab′)2 derived from rabbit Ig (one bond) and rat IgG1 (three bonds)can also be employed. However, rat IgG2a and IgG2c both have two, and rat IgG2b has four and so cannot be used as the maleimidated partner. 2. If F(ab′)3 derivatives are required, the number of SH-groups at the hinge of the unmaleimidated partner should be at least two and preferably three, because this determines the number of Fab′(mal) arms that can be conjugated. 3. We have found that a few antibodies give consistently low yields of BsAb when used as the maleimidated partner. If large quantities of a derivative are required, it is worthwhile performing small scale pilot preparations to determine which maleimidated partner gives the optimal yield. 4. It is very important to avoid contamination of the Fab′-A(SH) with 2-ME. In order to minimize the risk, stop collecting when the recorder has returned two-thirds of the way back to the baseline. In order to ensure that Fab′-B(SH) is not contaminated with 2-ME left over from the first run, make sure that it is not loaded until the chart recorder has returned to the baseline. 5. Sometimes the mixture becomes slightly cloudy. 6. To avoid contamination with o-PDM/DMF, which elutes as a second large peak, stop collecting when the chart recorder has returned halfway to baseline. 7. To avoid loss of product, slightly over concentrate, then wash the cell with a small volume of chilled buffer. 8. It is very important to add excess iodoacetamide at this stage; otherwise the BsAb derivative can precipitate. 9. To minimise contamination with endotoxin, buffers and reagents are made up using commercially available endotoxin-free water, and glass vessels washed with 0.2 M NaOH and thoroughly rinsed with endotoxin-free water before use. In addition, where possible chromatography columns are cleaned up regularly by running through with 0.2 M NaOH for 30 min

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followed by extensive washing with running buffer. However, please check the manufacturers’ instructions to ensure that this is not detrimental to the gel matrix being used. As both the preparation of the parent F(ab′)2 and the BsAb are multistage processes involving incubation with enzyme and several column chromatography and concentration steps, it is still possible that the product will be contaminated with LPS. To assay for endotoxin, we use EndosafeR –PTS cartridges (Charles River Laboratories, Charleston, SC) and collect the ‘clean product’ in an endotoxin-free disposable plastic container. To remove any endotoxin we use EndoTrapR Red endotoxin removal columns (Lonza, Walkersville, MD); these have a high binding capacity for endotoxin, and can be regenerated and re-used. To avoid re-contamination, minimise manipulations of the “clean” BsAband use endotoxin-free disposable plasticware. References 1. Glennie, M. J., McBride, H. M., Worth, A. T., and Stevenson, G. T. (1987) Preparation and performance of bispecific F(ab′γ)2 antibody containing thioether-linked Fab′γ fragments. J. Immunol. 139, 2367–2375. 2. Glennie, M. J., Tutt, A. L., and Greenman, J. (1993) Preparation of multispecific F(ab′)2 and F(ab′)3 antibody derivatives, in Tumour Immunobiology, A Practical Approach (Gallagher, G., Rees, R. C., and Reynolds, C. W., eds.), Oxford University Press, Oxford, UK, pp. 225–244.

194 Antigen Measurement Using ELISA William Jordan 1. Introduction The initial description of the enzyme-linked immunosorbent assay (ELISA) almost 30 yr ago (1) marked a technological advance that has had an immense impact in both clinical diagnostic and basic scientific applications. This assay represents a simple and sensitive technique for specific, quantitative detection of molecules to which an antibody is available. Although there are a huge number of variations based on the original ELISA principle, this chapter focuses on perhaps the two most useful and routinely performed: (1) the indirect sandwich ELISA—providing high sensitivity and specificity and (2) the basic direct ELISA—useful when only one antibody to the sample antigen is available. 1.1. The Direct ELISA During the indirect sandwich ELISA (Fig. 1A), an antibody specific for the substance to be measured is first coated onto a high-capacity protein binding microtiter plate. Any vacant binding sites on the plate are then blocked with the use of an irrelevant protein such as fetal calf serum (FCS) or Bovine serum albumin (BSA). The samples, standards, and controls are then incubated on the plate, binding to the capture antibody. The bound sample can be detected using a secondary antibody recognizing a different epitope on the sample molecule, thus creating the “sandwich.” The detection antibody is commonly directly conjugated to biotin, allowing an amplification procedure to be carried out with the use of streptavidin bound to the enzyme horseradish peroxidase (HRP). As streptavidin is a tetrameric protein, binding four biotin molecules, the threshold of detection is greatly enhanced. The addition of a suitable substrate such as 3,3′,5,5′-tetramethylbenzidine (TMB) allows a colormetric reaction to occur in the presence of the HRP that can be read on a specrophotometer with the

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Fig. 1. The indirect sandwich ELISA.

resulting optical density (OD) relating directly to the amount of antigen present within the sample. 1.2. The Indirect ELISA In some cases, however, only one specific antibody may be available, and in such a small quantity that directly conjugating it to biotin would be impractical. In this situation a direct ELISA should be used. During the direct ELISA

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the sample itself is coated directly onto the microtiter plate and is then detected using the specific antibody. If this antibody is biotinylated, then the procedure can then proceed as in the indirect ELISA; if not, then a secondary biotinylated antibody directed against the species of the detecting antibody itself can be used. Figure 1B demonstrates this technique. To set up a reliable and durable ELISA it is essential to first optimize a number of the parameters mentioned previously. The level of optimization will, of course, depend on exactly what is required from the assay. In some cases a simple “yes-or-no” answer is desired and a simple standard procedure may be sufficient. If, however, high sensitivity is the aim with accurate quantatation of the molecule in question, then carefully setting up the optimal conditions in advance will pay dividends and save a great deal of time in the long term. 2. Materials 1. Antibodies (see Note 1): For the indirect ELISA, antibody pairs can often be bought commercially and consist of a capture antibody and a biotinylated detection antibody. Both antibodies are specific for the molecule in question, with the detection antibody being directly biotinylated, and able to recognize the sample molecule when it is bound to the capture antibody on the plate. For the direct ELISA, one specific detection antibody is required, preferably biotinylated, but if not a secondary biotinylated antibody specific for the detection antibody is also needed. Aliquot and freeze at −20°C or lower in small, usable quantities with a carrier protein at such as 10% FCS or 1% BSA. 2. Blocking buffer: Phosphate-buffered saline (PBS), pH 7.4, supplemented with either 1% fatty acid free BSA or filtered 10% FCS. If measuring in tissue culture samples, substitute the FCS with protein representing that within the culture conditions, that is, 10% FCS; in effect the sample will have been preabsorbed with this during culture and thus any non-specific binding to the blocking buffer is avoided. 3. Carbonate coating buffer: 8.41 g Na2HCO3 in 1 L freshly made PBS. Dissolve and adjust to desired pH with HCl or NaOH. Store for no more than 1 mo at 4°C (see Note 2). 4. High-capacity protein binding 96-well microtiter plates: There are a large number of suitable makes including Maxisorp (Nunc), Immunoware (Pierce), Immunlon II (Dynatech), and Costar (see Note 3). 5. PBS–Tween–10% FCS: Add 0.05 mL of Tween 20 to 90 mL of PBS, pH 7.4, and 10 mL of filtered FCS. Make up fresh as required. 6. Plate sealers or cling film wrap. 7. Plate washing apparatus: Adequate washing is a vital element of achieving a successful ELISA. Although a number of automatic plate washers are available, they are expensive and the use of a wash bottle with good pressure is perfectly suitable, although a little more time consuming. 8. Samples/standards: Standards of known amounts are required for positive controls, and for estimation of levels within samples (via comparison to a titration of known

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amounts). All standards should be diluted in medium as near as, or identical to that of the sample solution. Standards are best obtained from a reliable commercial source having been mass calibrated and should be frozen in small, concentrated aliquots at −20°C or lower. Repeated freeze–thaw cycles must be avoided. Spectrophotometer: Any suitable microplate reader able to measure absorbance at the appropriate wavelength. Stop solution: 0.5 M H2SO4. Streptavidin HRP (see Note 4): Use in accordance to manufacturer’s instructions. Sources of streptavidin HRP include Sigma, Biosource, Pierce, and Zymed. Substrate: One-step TMB (Zymed). Although many other substrates are available for HRP, TMB has high sensitivity with a quick development time. OD can be monitored at 650 nm while the color develops, then at 450 nm when the reaction is stopped with H2SO4. TMB may also be obtained in a lyophilized state and made up fresh with hydrogen peroxidase. Washing buffer: Add 0.5 mL of Tween 20 to 1 L of PBS, pH 7.4. Make up fresh as required.

3. Methods 3.1. Basic Sandwich ELISA Protocol 1. Dilute the capture antibody to 1 µg/mL is coating buffer, pH 9.5. Add 100 µL to each well of a high-capacity protein binding 96-well microtiter plate. 2. Seal the plate to avoid evaporation and incubate overnight (12–18 h) at 2–8°C. 3. Wash plate: Discard unbound antibody by inverting and flicking the plate over a sink. Fill each well with washing buffer and leave for at least 10 s before discarding once more. Ensure all liquid has been removed between each wash by repeatedly tapping the plate onto clean paper towels. Repeat 3×. 4. Add 200 µL of blocking buffer to each well. Seal plate and incubate for at least 2 h at room temperature. 5. Discard blocking buffer. Wash plate 3× as described in step 3. 6. Prepare a titration series of known standards (e.g., in 1.5-mL Eppendorf tubes) diluted in a matrix representing that of the samples (e.g., culture medium or human serum). Include a negative control such as culture medium only. Transfer samples and antigen standards to the ELISA plate in duplicate at 100 µL/well. Seal plate and incubate at room temperature for at least 2 h, or overnight at 4°C for increased sensitivity. 7. Wash plate 3× as described in step 3. 8. Dilute biotinylated detection antibody to 1 µg/mL in PBS, pH 7.4. Add 50 µL/well. Incubate at room temperature for 2 h. If problems with nonspecific binding of the biotinylated antibody to the plate occur, dilute the antibody in PBS–Tween–10% FCS rather than just PBS. 9. Wash plate 3× as described in step 3. 10. Dilute streptavidin HRP according to manufacturer’s instructions. Add 100 µL/ well. Incubate at room temperature for 30 min. Again, if problems with nonspecific binding of the biotinylated antibody to the plate occur, dilute the antibody in PBS-Tween-10% FCS rather than just PBS.

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11. Wash plate 4× as described in step 3. 12. Add 100 µL/well of “one step” TMB. Allow color to develop for 10–60 min (20 min is usually sufficient). OD may be monitored at this stage at 650 nm as the color develops. 13. Add 100 µL of 0.5 M H2SO4 to each well to stop the reaction. Read OD at 450 nm. 14. Estimate amount of antigen within samples by comparing ODs to those of known standards.

3.2. Direct ELISA Protocol 1. Make serial dilutions of samples from neat to 1 : 16 in coating buffer, pH 9.5. Add 100 µL to each well of a high-capacity protein binding microtiter plate. Also set up wells with antigen standards and a negative control diluted in the same coating buffer. 2. Seal the plate with an acetate plate sealer or cling film wrap to avoid evaporation and incubate overnight (12–18 h) at 2–8°C. 3. Wash plate: Discard unbound antibody by inverting and flicking the plate over a sink. Fill each well with washing buffer leave for at least 10 s and discard once more. Ensure all liquid has been removed between each wash by repeatedly tapping the plate onto clean paper towels. Repeat 3×. 4. Add 200 µL of blocking buffer to each well. Seal plate and incubate for at least 2 h at room temperature. 5. Discard blocking buffer. Wash plate 3× as described in step 3. 6. Dilute detection antibody to 2 µg/mL in PBS. Add 50 µL/well. Incubate at room temperature for 2 h. If the antibody is biotinylated proceed to step 9. 7. Wash plate 3× as described in step 3. 8. Add secondary antibody (100 µL, 2 µg/mL) specific for the detection antibody (i.e., if the detection antibody is mouse, use biotinylated rabbit anti-mouse Ig). 9. Wash plate 3× as described in step 3. 10. Dilute streptavidin HRP according to the manufacturer’s instructions. Add 100 µL/ well. Incubate at room temperature for 30 min. 11. Wash plate 4× as described in step 3. 12. Add 100 µL/well of “one-step” TMB. Allow color to develop for 10–60 min (20 min is usually sufficient). OD may be monitored at this stage at 650 nm, as the color develops. 13. Add 100 µL of 0.5 M H2SO4 to each well to stop the reaction. Read OD at 450 nm. 14. Estimate amount of antigen within samples by comparing ODs to those of known standards of sample.

3.3. Optimization of ELISA At this stage the ELISA may be more than adequate, although optimization of the following parameters is usually required: 1. pH of carbonate coating buffer (pH 7.0–pH 10.0). Also try PBS at pH 7.4. 2. Concentration of capture antibody (0.5–10.0 µg/mL).

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3. Concentration of secondary antibody (indirect ELISA, 0.5–4.0 µg/mL). 4. Concentration of biotinylated detection antibody (0.1–2.0 µg/mL). 5. Concentration of streptavidin–HRP conjugate (usually between 1 : 1000 and 1 : 10,000).

4. Notes 1. The quality of the antibodies used is perhaps the most important aspect in seting up a good ELISA. Antibodies need to have a high affinity for the sample to be measured, reducing the chance of being “washed off” during the assay. The indirect ELISA relies on two specific antibodies and thus the increased specificity increases sensitivity of the assay by reducing background. 2. The pH of the coating buffer can have a great effect on the amount of antibody that will bind to the plate and thus to the ELISA as a whole. Basically, a higher pH will result in more antibody binding but may have a detrimental effect on its immunoreactivity. Thus a pH must be found that is suitable for the antibody in question, and this can vary dramatically. When beginning optimization of the assay, test a range of carbonate buffers from pH 7.0 to 10.0 as well as PBS, pH 7.4. We usually find a carbonate buffer pH of 9.5 gives good results. In some cases we have found commercially available coating antibodies recommended by the manufacturer to be adsorbed onto the plate at pH 7.4 to be far more effective at higher pH values, improving the sensitivity by up to 10-fold. This can allow the coating concentration to be vastly reduced, creating an extremely cost-effective assay. 3. There can be a significant difference between the protein binding capability of different makes, and even batches of microtiter plates. Some appear to be extremely good for binding antibodies, whereas others more useful for other proteins. The only real way to choose a suitable plate is by trial and error, or using a recommended make known to bind the protein you intend to coat. 4. Although the HRP–TMB system represents a good, reliable, and sensitive combination, HRP has a number of alternative substrates that can be used (2). These include o-phenylene diamine (OPD) or 2,2′-azino-di(3-ethylbenzthiazoline-6-sulfonic acid (ABTS). There are also a number of options for the enzyme used other than HRP, such as alkaline phosphatase (AP) which can be used in combination with the substrate p-nitrophenyl phosphate (p-NPP) or phenolphthalein monophosphate (PMP). The choice of enzyme–substrate system depends on a number of factors including price, sensitivity, and whether a filter is available for the substrate specific wavelength to be measured.

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5. The use of biotinylated secondary antibodies (or detection antibody in the indirect sandwich ELISA) in conjunction with enzyme-conjugated streptavidin (or avidin, extravidin, etc.) both increase sensitivity and save time in that a further step is eliminated from the assay and therefore another step of optimization is eliminated. 6. If the major aim of the ELISA is to obtain quantitation of substances present in extremely low concentrations a number of adaptations to the technique can be used. Such techniques often use AP enzyme systems which can be utilized, for example, to lock into a circular redox cycle producing an end product such as red formazan which is greatly amplified in comparison to standard amplification methods (3). Chemiluminescent amplified ELISA principles have also been shown to give very high sensitivity (4) and can be optimized to measure as little as 1 zeptomol (about 350 molecules!) of AP (5). Although extremely sensitive, such techniques are time consuming to set up and optimize, and are far more expensive than the simple colormetric ELISAs described in this chapter. 7. In some cases, molecules present in a sample are masked by the solution that they are in. This problem can sometimes be solved by diluting the samples in PBS–Tween–10% FCS. If this is performed, remember to make similar adjustments to the solution used for the standards. Possible interference molecules within samples such as soluble receptors for the antigen can also cause a problem. Commercially available matched antibody pairs for molecules should have been pretested and guaranteed against being affected by such problems. References 1. Engvall, J. R. and Perlamann, P. (1971) Enzyme-linked immunosorbent assay (ELISA). Quantitative assay of immunoglobulin G. Immunochemistry 8, 871–874. 2. Roberts, I. M., Solomon, S. E., Brusco, O. A., Goldberg, W., and Bernstein, J. J. (1991) A comparison of the sensitivity and specificity of enzyme immunoassays and time-resolved fluoromimmunoassay. J. Immunol. Meth. 143, 49–56. 3. Self, C. H. (1985) Enzyme amplification—a general method applied to provide an immunoassisted assay for placental alkaline phosphatase. J. Immunol. Meth. 76, 389–393. 4. Bronstein, I., Voyta, J. C., Thorpe, G. H., Kricka, L. J., and Armstrong, G. (1989) Chemiluminescent assay of alkaline phosphatase applied in an ultrasensative enzyme immunoassay of thyrotropin. Clin. Chem. 35, 1441–1446. 5. Cook, D. B. and Self, C. H. (1993) Determination of one-thousandth of an attomole (1 zeptomole) of alkaline phosphatase: application in an immunoassay of proinsulin. Clin. Chem. 39, 965–971.

195 Enhanced Chemiluminescence Immunoassay Richard A. W. Stott 1. Introduction Chemiluminescence results from reactions with a very high energy yield, which produce a potentially fluorescent product molecule; reaction energy passed to the product may result in an excited state and subsequent production of a single photon of light. The light yield is usually low, but can approach one photon per molecule in bioluminescent reactions catalyzed by dedicated enzymes. A wide variety of immunoassay systems that use chemiluminescence or bioluminescence have been developed with the aim of detecting low concentrations of biologically active molecules. Even when emission efficiencies are 9) or low (90%).

also work for antibodies. The most convenient strategy is probably the use of a stretch of histidines, the so-called his-tag, a series of 5 to 6 histidines that bind to affinity media such as NTA-agarose or Talon resin, when metal ions (nickel or cobalt) are bound. His-tagged proteins bind with milli-molar affinity to the column and are gently eluted with 150–300 mM imidazole (Fig. 4). Another such tag is the StrepII-tag (14), which shows affinity to streptavidin- or streptactin-sepharose (streptactin is a genetically engineered streptavidin with higher affinity to the Strep-tag) (Fig. 5). Commercial antibodies to both tags are available, so the tags can be used for detection purposes as well. Other detection tags like the V5-tag (GKPIPNPLLGLDST), which is derived from a small epitope present on the P/V proteins of the paramyxovirus, SV5, or the myc-tag (EQKLISEEDL), which correspond to residues 408–439 of the human p62 c-myc protein are also frequently used, because specific high-affinity monoclonal antibodies are commercially available. The FLAG-tag (DYKDDDDK) is of special interest since a monoclonal antibody (termed M1) exists that only binds to the tag when the N-terminus is free and not involved in a peptide bond. This has been used to monitor cleavage of signal sequences during E. coli antibody expression (15).

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2. Materials 2.1. Bacterial Cell Culture 1. 2xYT Medium: • 2xYT (Yeast Extract Tryptone) Media; Difco. • Dissolve 155 g 2xYT powder in 5 L deionized water. • After total dissolution, autoclave at 121°C for 20 min. • Store at room temperature. 2. Glucose stock (40%): • D(+)-glucose m onohydrate, ( MW = 198 g/mol). • Dissolve 44 g glucose in 100 mL deionized water. • After total dissolution, sterile filtration using 0.2 µm filter. • Store at room temperature. 3. CAM (Chloramphenicol) stock (34 mg/ml in ethanol): • Dissolve 3.4 g chloramphenicol in 100 mL ethanol (p.a.). • After total dissolution, sterile filtration using 0.2 µm filter. • Store at −20°C. 4. IPTG stock (1M in ddH2O): • Dissolve 4.2 g IPTG in 100 ml deionized water. • After total dissolution, sterile filtration using 0.2 µm filter. • Store at −20°C. 5. Preculture medium: mix 974 ml 2xYT medium with 25 ml glucose stock solution and 1 ml CAM-stock solution. 6. Expression-culture medium: mix 974 ml 2x YT-medium with 2.5 ml glucose stock solution and 1 ml CAM-stock solution. 7. Sterile filter: Acrodisc 13 mm syringe filters, 0.2 µm (Pall).

2.2. Cell Lysis 1. Cell Lysis Buffer: Take 120 ml BugBuster (Merck), add 240 mg Lysozyme (Roche), 1250 U Benzonase (Merck; 250 U/µl = 5 A/µl), and 5 PIT (Complete Protease Inhibitor) tablets, EDTA free (Roche #1873580) (see Note 1). Always prepare fresh solution. 2. BSS buffer: 200 mM boric acide, 160 mM NaCl, pH 8.0. Dissolve 12.37 g boric acid add 9.35g NaCl in 1000ml deionized water. Adjust to pH to 8.0 with NaOH. After total dissolution, sterile filtration using 0.2 µm filter. Store at room temperature. 3. PeriPrep lysis buffer: 200 mM boric acide, 160 mM NaCl, 2 mM EDTA, protease Inhibitor, pH 8.0. Dissolve 0.75g EDTA: Titriplex®III, (VWR, #108421) and 40 PIT (Complete Protease Inhibitor) tablets, EDTA free, (Roche), in 1000 ml BSS buffer (see Note 1). Adjust to pH to 8.0 with NaOH. Always prepare fresh solution. 4. Avidin stock solution 10mg/ml (for Strep-Tag purifications) (see Note 2). Dissolve 10 mg avidin in 10ml deionized water. Store at −20°C. 5. Seriflip filter device (STERIFLIP, Milipore).

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2.3. Affinity Chromatography 2.3.1. IMAC Affinity Chromatography 1. IMAC-RB (running buffer): 20 mM NaH2PO4, 500 mM NaCl, 10 mM imidazol, pH 7.4. Dissolve 2.4 g NaH2PO4, 29.2 g NaCl, 0.68g imidazole in 1000ml deionized water. Adjust to pH to 7.4. After total dissolution, sterile filtration using 0.2 µm filter. Store at room temperature. 2. IMAC-WB (washing buffer): 20 mM NaH2PO4, 500 mM NaCl, 20 mM imidazol, pH 7.4. Dissolve 2.4 g NaH2PO4, 29.2 g NaCl, 1.36 g imidazol in 1000 ml deionized water. Adjust to pH to 7.4. After total dissolution, sterile filtration using 0.2 µm filter. Store at room temperature. 3. IMAC-EB (elution buffer): 20 mM NaH2PO4, 500 mM NaCl, 250 mM Imidazole, pH 7.4. Dissolve 2.4 g NaH2PO4, 29.2 g NaCl, 17 g imidazole in 1000 ml deionized water. Adjust to pH to 7.4. After total dissolution, sterile filtration using 0.2 µm filter. Store at room temperature (see Note 3). 5. Ni-NTA agarose (Qiagen) (see Note 4). 4. Gravity flow plastic columns: Poly Prep (BioRad).

2.3.2. Strep Tag Affinity Chromatography 1. Strep-RB (running buffer): 100 mM Tris-HCl, 750 mM NaCl, 1 mM EDTA, pH 8.0. Dissolve 12.1 g Tris, 43.8 NaCl, 0.37g EDTA in 1000 ml deionized water. Adjust to pH to 8.0. After total dissolution, sterile filtration using 0.2 µm filter. Store at room temperature. 2. Strep-EB (elution buffer): Strep-RB + 5 mM D-Desthiobiotin. 100 mM Tris-HCl, 750 mM NaCl, 1 mM EDTA, 5 mM D-desthiobiotin, pH 8.0. Dissolve 12.1 g Tris, 43.8g NaCl, 0.37 g EDTA, 1.1 g D-D-desthiobiotin in 1000 ml deionized water. Adjust to pH to 8.0. After total dissolution, sterile filtration using 0.2 µm filter. Store at 4°C and not longer than 1 week (see Note 5). 3. Strep-RegB (regeneration buffer): 5 mM HABA, 100 mM Tris-HCl, pH 8.0. Dissolve 1.21 g HABA (HABA-ImmunoPure Pierce/Perbio) and 12.1 g Tris in 1000ml deionized water. Adjust to pH to 8.0 with HCl. After total dissolution, sterile filtration using 0.2 µm filter. Store at 4°C. 4. Streptactin sepharose (IBA-GMBH). 5. Gravity flow plastic columns: Poly prep (BioRad).

2.3.3. Size Exclusion Chromatography 1. PBS: 0.136 M NaCl, 2.68 mM KCl, 8.1 mM Na2HPO4, 1.46 mM KH2PO4, pH 7.4. Dissolve 0.2g KCl, 8g NaCl, 1.44g Na2HPO4 and 0.24g KH2PO4 in 1000ml deionized water. Adjust to pH to 7.4. After total dissolution, sterile filtration using 0.2 µm filter. Store at room temperature. 2. 3xPBS: 0.401 M NaCl, 8.04 mM KCl, 24.3 mM Na2HPO4, 4.38 mM KH2PO4, pH 7.4. Dissolve 0.6 g KCl, 24 g NaCl, 4.32 g Na2HPO4, and 0.72g KH2PO4 in 1000 ml deionized water. Adjust to pH to 7.4. After total dissolution, sterile filtration using 0.2 µm filter. Store at room temperature.

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3. PD10 columns: GE (Amersham Pharmacia Biotech).

2.3.4. Sterile Filtration, Concentration Determination, SDS-PAGE 1. Sterile filter: Acrodisc 13mm syringe filters, 0.2 µm (Pall). 2. UV-compatible cuvette: UVette (VWR international). 3. SDS-sample buffer 5 x concentrated: 125 mM Tris-HCl pH6,8 10% glycerol, 0,5% SDS, 0,05% (w/v) bromophenol blue, 5% beta-mercaptoethanol. 4. SDS running buffer: 25 mM Tris, 192 mM glycine, 0,1% SDS. 5. SDS-PAGE system: Criterion 15% Tris. 26W (Bio-Rad). 6. Gel staining solution: Gelcode Blue stain reagent (Pierce).

3. Methods 3.1. Strep-Tag Expression and Purification 3.1.1. Preculture 1. Fill 20 ml medium 1 (2xYT + 1% glucose + CAM) into a 100 ml Erlenmeyer flask. 2. Inoculate culture with a single colony with a sterile inoculating loop. Freshly transformed TG1F− cells are recommended (see Note 6). 3. Incubate at 30°C, O/N (18 h +/− 2h), 250 rpm.

3.1.2. Expression Culture 1. Fill 750 ml medium 2 (2xYT + 0.1% glucose + CAM) into a 2 L baffled Erlenmeyer flask. 2. Inoculate with 1/300 volume (2.5 ml) of preculture. 3. Incubate at 30°C, 3 h, 180 rpm. 4. Induce with 560 µL IPTG (1M) stock solution (0.75 mM final concentration). 5. Express Fab at 30°C, 160 rpm, O/N (20 h +/−2 h post induction time) (see Note 7).

3.1.3. Cell Harvest and Periplasmic Prep (PP) 1. Harvest cells by centrifugation at 5000 g, 30 min, 4°C. 2. Decant supernatant and discard after autoclaving (see Note 8). 3. Resuspend the bacterial pellet with 30–35 ml cold (75 mM). 4. Ni-NTA shows good results in our hands, but some authors prefer other IMAC resins (e.g., Talon-resin or Zn/Cu ions). 5. Desthiobiotin takes a long time to dissolve (gentle warming helps). Low concentrations of desthiobiotin will not interfere with many assays. It also shows no significant UV-absorption or interference in protein concentration assays. The final product is often also very pure (>90% judged on SDS-page). So it is often possible omit buffer exchange and use the eluted product directly in many applications. 6. For optimal expression performance it could be helpful to screen several clones. About 80% of all clones from the HuCAL® library will express in a range of +/−20%. In some case it is possible to find clones which show much higher expression rates (see Fig. 2). 7. These conditions show the best results for most Fab fragments. Nevertheless it could be helpful to optimise the expression conditions for optimal results. In some cases, lower temperatures (25°C) and lower IPTG concentrations (0.2 mM) will give better results. 8. In some cases (depending on the individual antibody and on the expression conditions, e.g. long expression times, or media that contains glycerol), significant amounts of the Fab can be found in the expression media. This can be checked with an anti-Fab-ELISA.

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9. Filtration can be omitted but this may cause clogging or contamination of the columns, in particular if the columns are used several times. 10. Streptactin columns can be used more than 10 times over more than 6 month without significant loss of binding activity. 11. For purification of bivalent Fab or large Fab-fusions it is not recommended to use a periplasmic preparation, because the release in the lysis buffer is often limited and the purification yields drop down. 12. The use of 3xPBS is recommended in purifications of bivalent Fab fragments, because these constructs sometimes tend to aggregate in PBS (depending on the individual antibody and the final concentration). This aggregation can be avoided by using a buffer with higher ionic strength like 3xPBS. 13. After thawing 3xPBS samples, a precipitate can often been seen at the bottom of the well or tube. This is salt from the PBS. It will disappear after thawing and gentle shaking. References 1. Knappik, A., Ge, L., Honegger, A., Pack, P., Fischer, M., Wellnhofer, G., Hoess, A., Wolle, J., Pluckthun, A., and Virnekas B. (2000) Fully synthetic human combinatorial antibody libraries (HuCAL) based on modular consensus frameworks and CDRs randomized with trinucleotides. J. Mol. Biol. 296, 57–86. 2. Soderlind, E., Strandberg, L., Jirholt, P., Kobayashi, N., Alexeiva, V., Aberg, A. M., Nilsson, A., Jansson, B., Ohlin, M., Wingren, C., Danielsson, L., Carlsson, R., and Borrebaeck, C. A. (2000) Recombining germline-derived CDR sequences for creating diverse single-framework antibody libraries. Nat. Biotechnol. 18, 852–856. 3. Hoet, R. M., Cohen, E. H., Kent, R. B., Rookey, K., Schoonbroodt, S., Hogan, S., Rem, L., Frans, N., Daukandt, M., Pieters, H., van Hegelsom, R., Neer, N. C., Nastri, H. G., Rondon, I. J., Leeds, J. A., Hufton, S .E., Huang, L., Kashin, I., Devlin, M., Kuang, G., Steukers, M., Viswanathan, M., Nixon, A .E., Sexton, D. J., Hoogenboom, H. R., and Ladner, R. C. (2005) Generation of high-affinity human antibodies by combining donor-derived and synthetic complementaritydetermining-regiondi versity. Nat. Biotechnol. 23, 344–348. 4. Hoogenboom, H. R., de Bruine, A. P., Hufton, S. E., Hoet, R. M., Arends, J. W., and Roovers, R. C. (1998), Antibody phage display technology and its applications. Immunotechnology 4, 1–20. 5. Kretzschmar, T. and von Ruden, T. (2002) Antibody discovery: phage display. Curr. Opin. Biotechnol. 13, 598–602. 6. Krebs, B., Rauchenberger, R., Reiffert, S., Rothe, C., Tesar, M., Thomassen, E., Cao, M., Dreier, T., Fischer, D., Hoss, A., Inge, L., Knappik, A., Marget, M., Pack, P., Meng, X. Q., Schier, R., Sohlemann, P., Winter, J., Wolle, J., and Kretzschmar, T. (2001) High-throughput generation and engineering of recombinant human antibodies. J Immunol. Methods 254, 67–84.

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7. Rauchenberger, R., Borges, E., Thomassen-Wolf, E., Rom, E., Adar, R., Yaniv, Y., Malka, M., Chumakov, I., Kotzer, S., Resnitzky, D., Knappik, A., Reiffert, S., Prassler, J., Jury, K., Waldherr, D., Bauer, S., Kretzschmar, T., Yayon, A., and Rothe, C. (2003). Human combinatorial Fab library yielding specific and functional antibodies against the human fibroblast growth factor receptor 3. J. Biol. Chem. 278, 38194–38205. 8. Jarutat, T., Nickels, C., Frisch, C., Stellmacher, F., Hofig, K.P., Knappik, A., and Merz, H. (2007) Selection of vimentin-specific antibodies from the HuCAL phage display library by subtractive panning on formalin-fixed, paraffin-embedded tissue. Biol. Chem. 388, 651–658. 9. Bird, R. E., Hardman, K. D., Jacobson, J. W., Johnson, S., Kaufman, B.M., Lee, S. M., Lee, T., Pope, S. H., Riordan, G. S., and Whitlow, M. (1988) Singlechain antigen-binding proteins. Science 242, 423–426. 10. Hudson, P. J. and Kortt, A. A. (1999). High avidity scFv multimers; diabodies and triabodies. J. Immunol. Methods 231, 177–189. 11. He, M. and Khan, F. (2005) Ribosome display: next-generation display technologies for production of antibodies in vitro. Expert Rev. Proteomics 2, 421–430. 12. Plückthun, A. and Pack, P. (1997) New protein engineering approaches to multivalent and bispecific antibody fragments. Immunotechnology 3, 83–105. 13. Rheinnecker, M., Hardt, C., Ilag, L. L., Kufer, P., Gruber, R., Hoess, A., Lupas, A., Rottenberger, C., Pluckthun, A., and Pack P. (1996) Multivalent antibody fragments with high functional affinity for a tumor-associated carbohydrate antigen. J. Immunol. 157, 2989–2997. 14. Schmidt, T. G. and Skerra, A. (2007) The Strep-tag system for one-step purification and high-affinity detection or capturing of proteins. Nature Protocols 2, 1528–1535. 15. Knappik, A. and Pluckthun, A. (1994). An improved affinity tag based on the FLAG peptide for the detection and purification of recombinant antibody fragments. Biotechniques 17, 754–761.

204 Screening Hybridoma Culture Supernatants Using Solid-Phase Radiobinding Assay Mark Page and Robin Thorpe 1. Introduction A large number of hybridoma culture supernatants (up to 200) need to be screened for antibodies at one time. The assay must be reliable so that it can accurately identify positive lines, and it must be relatively quick so that the positive lines, which are 75–100% confluent, can be fed and expanded as soon as possible after the assay results are known. Solid-phase binding assays are appropriate for this purpose and are commonly used for detection of antibodies directed against soluble antigens (1). The method involves immobilizing the antigen of choice onto a solid phase by electrostatic interaction between the protein and plastic support. Hybridoma supernatants are added to the solid phase (usually a 96-well format) in which positive antibodies bind to the antigen. Detection of the bound antibodies is then achieved by addition of an antimouse immunoglobulin labeled with radioactivity (usually 125I) and the radioactivity counted in a γ counter. 2. Materials 1. Antigen: 0.5–5 µg/mL in phosphate-buffered saline (PBS) with 0.02% sodium azide. 2. PBS: 0.14 M NaCl, 2.7 mM KCl, 1.50 mM KH2PO4, 8.1 mM Na2HPO4. 3. Blocking buffer: PBS containing either 5% v/v pig serum, 5% w/v dried milk powder, or 3% w/v hemoglobin (see Note 1). 4. Hybridoma culture supernatants. 5. Negative control: irrelevant supernatant or culture medium. 6. Positive control: serum from immunized mouse from which spleen for hybridoma production was derived. 7. Antimouse/rat immunoglobulin, 125I-labeled (100 µCi/5 µg protein). From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Method 1. Pipet 50 µL of antigen solution into each well of a 96-well microtiter plate and incubate at 4°C overnight. Such plates can be stored at 4°C for several weeks. Seal plates with cling film to prevent the plates from drying out (see Note 2). 2. Remove antigen solution from wells by either a Pasteur pipet (cover the tip with a small piece of plastic tubing to prevent scratching the antigen-coated wells) or by shaking the contents from the plate in one quick movement (see Note 3). 3. Wash the plate by filling the wells with PBS and rapidly discarding the contents. Repeat this twice more, tap the plate dry, then fill the wells with blocking buffer, and incubate at room temperature for 30 min to 1 h. Wash three more times with PBS. 4. Pipet 100 µL of neat hybridoma supernatant into the wells, and incubate for approx 3 h at room temperature. Include the negative and positive controls. 5. Wash with blocking buffer three times. 6. Dilute the 125I-labeled antimouse immunoglobulin in blocking buffer to give 1 × 106 cpm/mL. Add 100% µL to each well, and incubate for 1 h at room temperature. 7. Wash with blocking buffer. 8. Cut out individual wells using scissors or a hot nichrome wire plate cutter, and determine the radioactivity bound using a γ counter. A positive result would have counts that are four to five times greater than the background. Normally, the background counts would be around 100–200 cpm with a positive value of at least 1000 cpm ( see Note 4).

4. Notes 1. Filter the milk and hemoglobin solutions coarsely through an absorbent cloth (e.g., Kimnet, Kimberley Clark) to remove lumps of undissolved powder. 2. Antigens (diluted in distilled water) can be dried onto the plates by incubation in a warm room (37°C) without sealing. This method is usually preferred if the antigen is a peptide improving binding to the plastic; however, the background signal may be increased. 3. The antigen solution may be reused several times to coat additional plates, since only a small proportion of the protein adheres to the plastic. 4. Counting radioactivity in a 96-well format can also be performed by using plates designed for use in scintillation counters. The assay is performed as described, except that in the final step, a scintillation fluid (designed to scintillate with 125I, such as Microscint 20, Packard Instrument Co.) is added to each well, and the scintillation counted in a purpose-built machine (e.g., Topcount, Packard Instrument Co.). Reference 1. Johnstone, A. and Thorpe, R. (1996) Immunochemistry in Practice, 3rd ed. Blackwell Scientific, Oxford, UK.

205 Screening Hybridoma Culture Supernatants Using ELISA Mark Page and Robin Thorpe 1. Introduction Enzyme-linked immunosorbent assay (ELISA) is a widely used method for the detection of antibody and is appropriate for use for screening hybridoma supernatants (1). As with radiobinding assays, it is a solid-phase binding assay that is quick, reliable, and accurate. The method is often preferred to radioactive assays, since the handling and disposal of radioisotopes is avoided and the enzyme conjugates are more stable than radioiodinated proteins. However, most of the substrates for the enzyme reactions are carcinogenic or toxic and, hence, require handling with care. Enzymes are selected that show simple kinetics, and can be assayed by a simple procedure (normally spectrophotometric). Cheapness, availability, and stability of substrate are also important considerations. For these reasons, the most commonly used enzymes are alkaline phosphatase, β-d-galactosidase, and horseradish peroxidase. 2. Materials 1. PBS-Tween: Phosphate-buffered saline (PBS) (0.14 M NaCl, 2.7 mM KCl, 1.5 mM KH2PO4, 8.1 mM Na2HPO4) containing 0.05% Tween-20. 2. Antigen: 1–5 µg/mL in PBS. 3. Blocking buffers: PBS-Tween containing either 5% v/v pig serum or 5% w/v dried milk powder (see Note 1). 4. Hybridoma culture supernatants. 5. Alkaline phosphatase conjugated antimouse immunoglobulin. 6. Carbonate buffer: 0.05 M sodium carbonate, pH 9.6. 7. p-Nitrophenyl phosphate, disodium hexahydrate: 1 mg/mL in carbonate buffer containing 0.5 mM magnesium chloride. 8. 1 M sodium hydroxide.

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Method 1. Pipet 50 µL of antigen solution into each well of a 96-well plate, and incubate overnight at 4°C. Such plates can be stored for 4°C for several weeks. Seal plates with cling film to prevent the plates from drying out (see Note 2). 2. Remove antigen solution from wells by either a Pasteur pipet (cover the tip with a small piece of plastic tubing to prevent scratching the antigen-coated wells) or by shaking the contents from the plate in one quick movement (see Note 3). 3. Wash the plate by filling the wells with PBS and rapidly discarding the contents. Repeat this twice more, then fill the wells with blocking buffer, and incubate at room temperature for 30 min to 1 h. Wash three more times with PBS. 4. Pipet 100 µL of neat hybridoma supernatant into the wells, and incubate for approx 3 h at room temperature. Include the negative and positive controls (see Note 4). 5. Wash with blocking buffer three times. 6. Prepare 1:1000 dilution of alkaline phosphatase-conjugated antimouse immunoglobulin in PBS-Tween, and add 200 µL of this to each well. Cover, and incubate at room temperature for 2 h. 7. Shake-off conjugate into sink and wash three times with PBS. 8. Add 200 µL p-nitrophenyl phosphate solution to each well, and incubate at room temperature for 20–30 min. 9. Add 50 µL sodium hydroxide to each well, mix, and read the absorbance of each well at 405 nm (see Note 5). Typical background readings for absorbance should be less than 0.2 with positive readings usually three or more standard deviations above the average background value.

4. Notes 1. Filter the milk solution coarsely through an absorbent cloth (e.g., Kimnet, Kimberley Clark) to remove lumps of undissolved powder. 2. Antigens (diluted in distilled water) can be dried onto the plates by incubation in a warm room (37°C) without sealing. This method is usually preferred if the antigen is a peptide improving binding to the plastic; however, the background signal may be increased. 3. The antigen solution may be reused several times to coat additional plates, since only a small proportion of the protein adheres to the plastic. 4. All assays should be carried out in duplicate or triplicate. 5. Purpose-built plate readers provide a very rapid and convenient means of determining the absorbance. Reference 1. Johnstone, A. and Thorpe, R. (1996) Immunochemistry in Practice, 3rd ed. Blackwell Science, Oxford, UK.

206 Growth and Purification of Murine Monoclonal Antibodies Mark Page and Robin Thorpe 1. Introduction Once a hybridoma line has been selected and cloned, it can be expanded and seed stocks cryopreserved for future use. Relatively large amounts of purified MAb may also be required. There are a variety of procedures for this that ensure the establishment of a stable cell line secreting high levels of specific immunoglobulin. High concentrations of antibody can be generated by growing the line in the peritoneal cavity of mice/rats of the same strain as the tumor cell line donor and spleen cell donor. Antibody is secreted into the ascitic fluid formed within the cavity at a concentration up to 10 mg/mL. However, the ascites will contain immunoglobulins derived from the recipient animal that can be removed by affinity chromatography if desired. Several in vitro culture methods using hollow fibres or dialysis tubing (1) have been developed and are commercially available; this avoids the use of recipient mice/rats and contamination by host immunoglobulins, although contamination with culture medium-derived proteins may be a problem. 2. Materials 1. RPMI-1640/DMEM medium. 2. Fetal bovine serum (FBS). 3. Phosphate-buffered saline (PBS): 0.14 M NaCl, 2.7 mM KCl, 1.50 mM KH2PO4, 8.1 mM Na2HPO4. 4. HT medium: 1 mL 50X HT supplement in 50 mL 10% FBS/RPMI. 5. HT supplement (Gibco, Paisley, UK).

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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3. Method 3.1. Growth of Hybridomas 1. After cloning, grow hybridomas in 1 mL HT medium in separate wells of a 24-well plate. Check growth every day and feed with 0.5–1 mL HT medium if supernatant turns yellow (see Note 1). 2. When cells are 75–100% confluent (see Note 2), expand into two further wells with fresh HT medium. When these are confluent, transfer into a 25-cm3 flask and feed with approx 10 mL medium (see Note 3). Subsequently, cell lines should be weaned off HT medium by increasing the dilution of HT medium stepwise with normal medium lacking the HT supplement, eventually culturing the cells in normal medium. Do this by reducing the amount of HT medium by 25–50% at each feed for expansion of the hybridomas. For example, reduce the amount of HT from 100 to 75 to 50 to 25 to 0% replacing with normal medium. 3. The FBS may be reduced to 5% or lower if hybridoma growth is strong.

3.2. Purification of MAb MAb purification can be carried out by a number of methods as described in Chapters 179–190 and 207. 4. Notes 1. Phenol red is included in the medium as a visual indicator of pH. A yellow supernatant indicates acid conditions as a result of dissolved CO2 (carbonic acid) derived from active cell growth (respiration). Normally, a yellow supernatant will indicate that the cells should be split (passaged) because the active cell growth will have exhausted the medium of nutrients and hence will not sustain cell viability. 2. Cells are confluent when the bottom of the flask/well is completely covered. Usually, the cells will require feeding or expanding at this stage (see Note 1). 3. To prevent inadvertant loss of precious cell lines, it may be prudent to freeze cloned lines before expansion into flasks. Reference 1. Pannell, R. and Milstein, C. (1992) An oscillating bubble chamber for laboratory scale production of MAb as an alternative to ascitic tumors. J. Immunol. Meth. 146, 43–48.

207 Affinity P urification Techniques for Monoclonal Antibodies Alexander Schwarz 1. Introduction Monoclonal antibodies have many applications in biotechnology such as immuno-affinity chromatography, immunodiagnostics, immunotherapy, drug targeting, biosensors, and many others. For all these purposes homogeneous antibody preparation are needed. Affinity chromatography, which relies on the specific interaction between an immobilized ligand and a particular molecule sought to be purified, is a well-known technique to purify proteins from solution. Three different affinity techniques for the one-step purification of antibodies—protein A, thiophilic adsorption, and immobilized metal affinity chromatography—are described in this chapter. 1.1. Protein A Chromatography Protein A is a cell wall component of Staphylococcus aureas. Protein A consists of a single polypeptide chain in the form of a cylinder, which contains five highly homologous antibody binding domains. The binding site for protein A is located on the Fc portion of antibodies of the immunoglobulin G (IgG) class (1). Binding occurs through an induced hydrophobic fit and is promoted by addition of salts such as sodium citrate or sodium sulfate. At the center of the Fc binding site as well as on protein A reside histidine residues. At alkaline pH, these residues are uncharged and hydrophobic, strengthening the interaction between protein A and the antibody. As the pH is shifted to acidic values, these residues become charged and repel each other. Differences in the pH-dependent elution properties (Table 1) are seen between antibodies from different classes as well as different species due to minor differences in the binding sites. These

From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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Schwarz Table 1 Affinity of Protein A for IgG for Different Species and Subclasses IgG species/subclass

Affinity

Binding pH

Elution pH

Human IgG1 Human IgG2 Human IgG3 Human IgG4

High High Moderate High

7.5 7.5 8 7.5

3 3 4–5 3

Mouse IgG1 Mouse IgG2a Mouse IgG2b Mouse IgG3

Low Moderate High Moderate

8.5 8 7.5 8

5–6 4–5 3 4–5

Rat IgG1 Rat IgG2a Rat IgG2b

Low None–low Low

8.5

5–6

8.5

5–6

differences can be successfully exploited in the separation of contaminating bovine IgG from mouse IgG, as shown in Subheading 1.3. The major attraction in using protein A is its simplicity. The supernatant is adsorbed onto a protein A gel, the gel is washed, and the antibody is eluted at an acidic pH. The antibody recovered has a purity of >90%, often with full recovery of biological activity. The method described in Subheading 3.1. provides a more detailed description of the purification for high-affinity antibodies. The affinity between protein A and the antibody is due mainly to hydrophobic interactions at the binding sites (1). The interaction can be strengthened by the inclusion of higher concentrations of chaotropic salts such as sodium citrate or sodium sulfate. Although the addition of chaotropic salts is unnecessary in the case of high-affinity antibodies, it allows weakly binding antibodies such as mouse IgG1 to strongly interact with protein A. This purification scheme is outlined in Subheading 3.2. The different affinities resulting from minor differences in the Fc region can also be utilized for the separation of subclasses or the separation of different species in hybridoma supernatant using a pH gradient. However, this is possible only if the difference in affinity is large enough like in the case of the separation of mouse IgG2b from bovine IgG. If fetal calf serum supplemented growth media is used, bovine IgG will contaminate the monoclonal antibody if the method described in Subheading 3.1. or 3.2. is used. An improved protocol utilizing the different affinities is described in Subheading 3.3. 1.2. Thiophilic Adsorption Chromatography The term thiophilic adsorption chromatography was coined for gels, which contain low molecular weight sulfur-containing ligands such as divinyl sulfone

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structures or mercapto-heterocyclic structures (2,3). The precise binding mechanism of proteins to these gels is not well understood. However, thiophilic chromatography can be regarded as a variation of hydrophobic interaction chromatography inasmuch as chaotropic salts need to be added to facilitate binding of antibodies to the gel. Therefore, the interaction between the ligands and the antibody is likely to be mediated through accessible aromatic groups on the surface of the antibody. The major advantage of these thiophilic gels over protein A gels is that they bind all antibodies with sufficiently high capacity and very little discrimination between subclasses or species. Furthermore, elution conditions are much milder, which might be beneficial if the harsh elution conditions required for strongly binding antibodies to protein A lead to their denaturation. Thiophilic gels also purify chicken IgY (4). A general method is provided in Subheading 3.4. The major disadvantage of thiophilic gels is the requirement to add high concentrations of chaotropic salts to the adsorption buffer to facilitate binding. This requirement is not a problem for small-scale purifications at the bench, but it is unattractive at larger scale. Recently, a major improvement of thiophilic chromatography came about with the utilization of very high ligand concentrations (9). By immobilizing mercaptoheterocyclics such as 4-mercaptoethyl pyridine at very high concentrations, the need to add chaotropic salts such as sodium sulfate was abolished. All antibodies investigated are adsorbed at neutral pH, independent of the salt concentration. At acidic pH values, the ligand becomes charged and through charge–charge repulsion, the antibody is eluted. This variation of thiophilic adsorption chromatography is highly reminiscent of protein A chromatography in its simplicity, but with the added advantage that it is nonspecific for antibodies. Purities and recoveries obtained range from 90% for antibodies from protein-free cell culture supernatant to approx 75% from ascitic fluid. The method is outlined in Subheading 3.5. 1.3. Immobilized Metal Affinity Chromatography Immobilized metal affinity chromatography (IMAC) is a general term for a variety of different immobilization chemistries and metals utilized (10). The most commonly used gel for IMAC is nickel-loaded iminodiacetic acid (Ni-IDA) gel, used, for example, in kits for the separation of His-tail modified proteins. Under slightly alkaline conditions, the interaction between the immobilized nickel and proteins is strongest with accessible histidine residues. Ni-IDA binds to the Fc portion of the antibody (11), and similar to thiophilic adsorption chromatography, it binds all antibodies without discrimination between subclasses or species (11,12). Depending on the growth medium used, some contamination is apparent, namely transferrin and traces of albumin. The purity of monoclonal antibodies

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purified out of ascitic fluid is generally lower than out of hybridoma cell culture supernatant. As described for protein A chromatography, a shift in pH to acidic values leads to the generation of charges on the histidine residues and consequent elution of the protein bound. Alternatively, competitive elution with either imidazole or EDTA yields antibody in good purity. However, the elution with EDTA delivers the antibody with the metal still bound to the protein, while imidazole adsorbs at 280 nm, interfering with UV detection. A general protocol is outlined. 2. Materials 2.1. General Purification of Human IgG, Humanized IgG, and Mouse IgG2a and Mouse IgG2b Using Protein A 1. 2. 3. 4.

Buffer A: 50 mM Tris-HCl, pH 7.5. Buffer B: 50 mM Tris-HCl, 500 mM NaCl, pH 8.0. Buffer C: 100 mM acetate buffer, 50 mM NaCl, pH 3.0. Buffer D: 10 mM NaOH.

2.2. General Purification of Mouse IgG1, Rat IgG1, and Rat IgG2b Using Protein A 1. Buffer A: 500 mM sodium citrate, ph 8–8.4. 2. Buffer B: 100 mM sodium acetate, 50 mM NaCl, pH 4.0. 3. Buffer C: 10 mM NaOH.

2.3. General Purification of IgG with Removal of Bovine IgG Using Protein A 1. 2. 3. 4.

Buffer A: 50 mM Tris-HCl, pH 7.5. Buffer B: 50 mM citrate, pH 5.0. Buffer C: 50 mM citrate, 50 mM NaCl, pH 3.0. Buffer D: 10 mM NaOH.

2.4. Thiophilic Purification of IgG 1. Buffer A: 50 mM Tris-HCl, 500 mM sodium sulfate, pH 7.5. 2. Buffer B: 50 mM sodium acetate, pH 5.0. 3. Buffer C: 10 mM NaOH.

2.5. General Purification of IgG Using MEP-HyperCell 1. Buffer A: 50 mM Tris-HCl, pH 7.5. 2. Buffer B: 100 mM acetate buffer, 50 mM NaCl, pH 3.5. 3. Buffer D: 10 mM NaOH.

2.6. Immobilized Metal Affinity Purification of IgG 1. Buffer A: 50 mM sodium phosphate, 500 mM sodium chloride, pH 8.0. 2. Buffer B: 50 mM sodium acetate, 50 mM sodium chloride, pH 4.5. 3. Buffer C: 200 mM sodium phosphate, 200 mM sodium chloride, pH 8.0.

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4. Buffer D: 50 mM Tris-HCl, 500 mM sodium chloride, 50 mM EDTA, pH 8.0. 5. Buffer E: Buffer B, 50 mM nickel chloride.

3. Method 3.1. General Purification of Human IgG, Humanized IgG, and Mouse IgG2a and Mouse IgG2b Using Protein A (see Note 1) This protocol is designed for the purification of high-affinity IgG monoclonal antibodies from hybridoma cell culture supernatant and ascitic fluid. For lower affinity monoclonals, refer to Subheading 3.2. or the other purification methods described below. For cell culture supernatants that contain fetal calf serum and the separation of contaminating bovine IgG from the monoclonal antibody, please see Subheading 3.3. 1. 2. 3. 4. 5. 6. 7. 8.

Bring all materials to room temperature. Equilibrate a 10-mL protein A gel with 5 column volumes of buffer A. Load supernatant onto protein A–gel column at 5 mL/min. Load up to 20 mg of antibody/mL of gel. Wash with 10 column volumes of buffer B. Elute antibody with 5 column volumes of buffer C. Reequilibrate column with 2 column volumes of buffer A. Wash with 5 column volumes of buffer E followed by 10 column volumes of buffer A.

The column is ready for the next chromatography run. 3.2. General Purification of Mouse IgG1, Rat IgG1, and Rat IgG2b Using Protein A (see Note 2) For cell culture supernatants that contain fetal calf serum and the separation of contaminating bovine IgG from the monoclonal antibody, please see Subheading 3.3. 1. Bring all materials to room temperature. 2. Dilute supernatant 1:1 with 1000 mM sodium citrate solution. Filter the resulting solution through a 0.2-µm filter (important). 3. Equilibrate a 10-mL protein A gel with 5 column volumes of buffer A. 4. Load supernatant onto the protein A–gel column at 5 mL/min. 5. Load up to 12 mg of antibody/mL of gel. 6. Wash with buffer A until the UV baseline is reached. 7. Elute antibody with 10 column volumes of buffer B. 8. Reequilibrate column with 2 column volumes of buffer A. 9. Wash with 5 column volumes of buffer C followed by 10 column volumes of buffer A.

The column is ready for the next chromatography run.

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3.3. General Purification of IgG with Removal of Bovine IgG Using Protein A (see Note 3) This method is designed for the purification of human, humanized, and mouse IgG2a and IgG2b monoclonal antibodies from hybridoma cell culture supernatant containing fetal calf serum. It does not work well with weakly binding antibodies such as mouse IgG1. 1. 2. 3. 4. 5. 6. 7. 8.

Bring all materials to room temperature. Equilibrate a 10-mL protein A gel with 5 column volumes of buffer A. Load supernatant onto the protein A–gel column at 5 mL/min. Load up to 20 mg of antibody/mL of gel. Wash with 5 column volumes of buffer A. Wash with 15 column volumes of buffer B. Elute antibody with 10 column volumes of buffer C. Wash with 5 column volumes of buffer E followed by 10 column volumes of buffer A.

The column is ready for the next chromatography run. 3.4. Thiophilic Purification of IgG (see Note 4) This protocol is designed for the purification of all IgG from hybridoma cell culture supernatant and ascitic fluids. For cell culture supernatants that contain more than 5% fetal calf serum, the monoclonal antibody will be contaminated with bovine IgG. 1. 2. 3. 4. 5. 6. 7. 8. 9.

Bring all materials to room temperature. Dilute supernatant 1:1 with 1000 mM sodium sulfate solution. Filter the resulting solution through a 0.2-µm filter (important). Equilibrate thiophilic gel with 5 column volumes of buffer A. Load supernatant onto the thiophilic gel column at 5 mL/min. Load up to 10 mg of antibody/mL of gel. Wash with buffer A until the UV baseline is reached. Elute antibody with 10 column volumes of buffer B. Wash with 5 column volumes of buffer C followed by 10 column volumes of buffer A.

The column is ready for the next chromatography run. 3.5. General Purification of IgG Using MEP-HyperCell (see Note 5) This protocol is designed for the purification of all IgG from hybridoma cell culture supernatant and ascitic fluids. For cell culture supernatants that contain more than 5% fetal calf serum, the purity is relatively low at approx 50–65%.

Affinity Purification Techniques for Monoclonal Antibodies 1. 2. 3. 4. 5. 6. 7. 8.

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Bring all materials to room temperature. Equilibrate a 10-mL MEP-HyperCell gel with 5 column volumes of buffer A. Load supernatant onto the column at 5 mL/min. Load up to 20 mg of antibody/mL of gel. Wash with 10 column volumes of buffer A. Elute antibody with 8 column volumes of buffer B. Reequilibrate the column with 2 column volumes of buffer A. Wash with 8 column volumes of buffer E followed by 10 column volumes of buffer A.

The column is ready for the next chromatography run. 3.6. Immobilized Metal Affinity Purification of IgG (see Note 6) This protocol is designed for the purification of all IgG from hybridoma cell culture supernatant and ascitic fluids. For cell culture supernatants that contain more than 5% fetal calf serum, the monoclonal antibody will be contaminated with bovine IgG. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Bring all materials to room temperature. Dilute three parts of supernatant with one part of buffer C. Equilibrate a 10-mL IMAC gel with 5 column volumes of buffer B. Load the column with buffer E until the column is completely colored. Wash with 5 column volumes of buffer B. Equilibrate with 5 column volumes of buffer A. Load supernatant onto the IMAC gel column at 5 mL/min. Load up to 10 mg of antibody/mL of gel. Wash with buffer A for 15 column volumes until the UV baseline is reached. Elute antibody with 10 column volumes of buffer B. Strip column with 5 column volumes of buffer D, followed by 5 column volumes of buffer B.

The column is ready for the next chromatography run. 3.7. Concluding Remarks All the protocols provided in the yield antibodies in good yield and good purity. If not further specified in the accompanying notes, the protocols are usable with virtually all commercially available gels. Difficulties in using a protein A gel can be overcome by using a different affinity column such as a thiophilic gel or an IMAC gel. In most protocols provided, elution is accomplished at acidic pH values. Therefore, further purification can be accomplished by adsorbing the eluate onto a cation-exchange gel. Most of the impurities will flow through and, when using gradient elution, the antibody is generally the first protein to elute from the column.

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4. Notes 1. This general protocol works well with high-affinity antibodies and is independent of the particular protein A gel used. The purity of the antibody recovered can be increased by first dialyzing the supernatant or the ascitic fluid against buffer A. Sometimes, phenol red is difficult to remove and the washing step has to be increased accordingly until no red color can be detected on the gel. 2. No pH adjustments have to be made for the sodium citrate solution as the pH will be between 8.0 and 8.5. It is very important to filter the resulting solution after mixing of the supernatant and the sodium citrate solution. In the case of ascitic fluid, a precipitate is clearly visible, while in the case of hybridoma cell culture supernatant precipitating material might not be visible. The purity is somewhat lower than in the case of high-affinity antibodies, but it is very acceptable in the 85–90% range. 3. The molarity of the citrate buffer used in the intermediate wash is slightly dependent on the protein A gel used. It is possible to wash the protein A gel from Pharmacia with 100 mM citrate buffer, pH 5.0, while the higher molarity would start to elute antibody from a protein A gel from BioSepra. A good starting point is the 50 mM citrate buffer outlined above, and if no loss of the desired monoclonal antibody is detected in the wash fraction, higher buffer concentrations can be employed. 4. There are only a few thiophilic gels commercially available. Of these, the thiophilic gel from E. Merck is by far the best. The most important precaution in using thiophilic gels is the need to filter the supernatant after adjustment to 500 mM sodium sulfate. Other antibody molecules like IgA (5), bispecific antibodies (6), and single-chain antibody fragments have been purified using thiophilic gels (7,8). 5. All antibodies, independent of species and subclass, can be purified with this gel. No separation of subclasses can be achieved, however. It is highly likely that similarly to the thiophilic gels described above, other antibody classes such as IgA, IgE, IgM, and chicken IgY can be purified using MEPHyperCell. The MEP-HyperCell resin is available from BioSepra Inc., a unit of Ciphergen, Inc. 6. Generally, all chelating and amine-containing reagents such as citrate and Tris buffers should be avoided in the feedstocks and buffers used, as these buffers will strip the column of the immobilized metal. If possible, using a gradient to elute the proteins results in better purity of the antibody sought. Three different metals can be used to purify antibody, namely copper, nickel, and cobalt. A good starting metal of any purification using IMAC is nickel. Other metals that work in the purification of antibodies are cobalt and copper. The use of zinc does not yield any antibody. The addition of a

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His-tag allows the specific adsorption and elution of single-chain antibodies and antibody fragments (13,14). References 1. Diesenhofer, J. (1981) Crystallographic refinement and atomic models of a human Fc fragment and its complex with fragment B of protein A from Staphylococcus aureus at 2.9 and 2.8 Å resolution. Biochemistry 20, 2361–2370. 2. Porath, J., Maisano, F., and Belew, M. (1985) Thiophilic adsorption—a new method for protein fractionation. FEBS Lett. 185, 306–310. 3. Oscarsson, S. and Porath, J. (1990) Protein chromatography with pyridine- and alkylthioester based Agarose adsorbents. J. Chromatogr. 499, 235–247. 4. Hansen, P., Scoble, J. A., Hanson, B., and Hoogenraad, N. J. (1998) Isolation and purification of immunoglobulins from chicken eggs using thiophilic interaction chromatography. J. Immunol. Meth. 215, 1–7. 5. Leibl, H., Tomasits, R., and Mannhalter, J. W. (1995) Isolation of human serum IgA using thiophilic adsorption chromatography. Protein Exp. Purif. 6, 408–410. 6. Kreutz, F. T., Wishart, D. S., and Suresh, M. R. (1998) Efficient bispecific monoclonal antibody purification using gradient thiophilic affinity chromatography. J. Chromatogr. B Biomed. Sci. Appl. 714, 161–170. 7. Schulze, R. A., Kontermann, R. E., Queitsch, I., Dubel, S., and Bautz, E. K. (1994) Thiophilic adsorption chromatography of recombinant single-chain antibody fragments. Analyt. Biochem. 220, 212–214. 8. Fiedler, M. and Skerra, A. (1999) Use of thiophilic adsorption chromatography for the one-step purification of a bacterially produced antibody F(ab) fragment without the need for an affinity tag. Protein Exp. Purif. 17, 421–427. 9. Burton, S. C. and Harding, D. R. (1998) Hydrophobic charge induction chromatography: salt independent protein adsorption and facile elution with aqueous buffers. J. Chromatogr. A 814, 71–81. 10. Porath, J. and Olin, B. (1983) Immobilized metal ion affinity adsorption and immobilized metal ion affinity chromatography of biomaterials. Biochemistry 22, 1621–1630. 11. Hale, J. E. and Beidler, D. E. (1994) Purification of humanized murine and murine monoclonal antibodies using immobilized metal-affinity chromatography. Analyt. Biochem. 222, 29–33. 12. Al-Mashikhi, S. A., Li-Chan, E., and Nakai, S. (1998) Separation of immunoglobulins and lactoferrin from cheese whey by chelating chromatography. J. Dairy Sci. 71, 1747–1755. 13. Laroche-Traineau, J., Clofent-Sanchez, G., and Santarelli, X. (2000) Three-step purification of bacterially expressed human single-chain Fv antibodies for clinical applications. J. Chromatogr. B Biomed. Sci. Appl. 737, 107–117. 14. Kipriyanov, S. M., Moldenhauer, G., and Little, M. (1997) High level production of soluble single chain antibodies in small-scale Escherichia coli cultures. J. Immunol. Meth. 200, 69–77.

208 A Rapid Method for Generating Large Numbers of High-Affinity Monoclonal Antibodies from a Single Mouse Nguyen Thi Man and Glenn E. Morris 1. Introduction Since the first description by Kohler and Milstein (1), many variations on this method for the production of monoclonal antibodies (MAb) have appeared and recent comprehensive books on the subject have included both recombinant and conventional methods (see Further Reading). The variation we describe here, however, includes a number of refinements that enable rapid (6–10 wk) production from a single spleen of 10–30 cloned and established hybridoma lines producing antibodies of high-affinity. We initially applied this method to recombinant fusion proteins containing fragments of the muscular dystrophy protein dystrophin (2–5), and dystrophin-related proteins (6,7), to hepatitis B surface antigen (8) and to the enzyme creatine kinase (CK) (9), and have used the MAb thus produced for immunodiagnosis, epitope mapping, and studies of protein structure and function (5–12). Epitopes shared with other proteins are common, so availability of several MAb against different epitopes on a protein can be important in ensuring the desired specificity in immunolocalization and Western blotting studies (6). More recently, we have applied this same technique for nuclear proteins, such as emerin (16 MAbs; 13), lamin A (5 MAbs; 14), SMN (22 MAbs; 15), gemins (35 MAbs; 16) and the transcription factor, Six5 (18 MAbs; 17), as well as other human proteins as diverse as huntingtin (19 MAbs; 18) and dopamine receptors (8 MAbs; 19). In the standard Kohler and Milstein method, Balb/c mice are immunized with the antigen over a period of 2–3 mo, and spleen cells are then fused with mouse myelomas using polyethylene glycol (PEG) to immortalize the B-lymphocytes secreting specific antibodies. The hybrid cells, or hybridomas, are then selected From: The Protein Protocols Handbook, Third Edition Edited by: J.M. Walker © Humana Press, a Part of Springer Science + Business Media, LLC 2009

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using medium containing hypoxanthine, aminopterin, and thymidine (HAT medium) (20). All unfused myelomas are killed by HAT medium, and unfused spleen cells gradually disappear in culture; only hybridomas survive. Use of hyperimmune mice is critical for success and we have found this much easier to achieve using recombinant proteins as immunogens, rather than conjugated peptides. Fusion proteins with a bacterial component, such as pET vectors with a thioredoxin fusion (Novagen, Inc), have been especially useful. We have rarely found it necessary to take steps to ensure correct folding of the protein immunogen, since some MAbs that recognize both folded and unfolded protein were usually produced when appropriately screened. An exception that did require special steps in both immunogen preparation and hybridoma screening to obtain MAbs against native protein is described below. Fig.1A illustrates both a weak response to immunogen (little binding at 1:1000 dilution), making MAb production difficult, and a typical strong response (strong binding at 1:1000 dilution) that will make hybridoma production very much easier. The importance of appropriate screening of both mouse test bleeds and hybridoma supernatants cannot be emphasized too often. If the desired antibody is not represented in the mouse serum, there is little chance of its emerging after hybridoma fusions.

Fig. 1. Effect of aggregation of CK on the immune response in Balb/c mice. Test sera were taken from three mice immunized with untreated CK (A): circles, squares, and triangles represent individual mice) or with glutaraldehyde-aggregated CK (B), and serial dilutions were used as primary antibody in either direct ELISA for denatured CK (dotted lines) or sandwich ELISA for native CK (solid lines). (Reproduced with permission from ref. 9.)

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Special features of the detailed method to be described here include: 1. 2. 3. 4.

A culture medium for rapid hybridoma growth without feeder layers; Screening early to select high-affinity antibodies; Cloning without delay to encourage hybridoma growth and survival; and The use of round-bottomed microwell plates to enable rapid monitoring of colony growth with the naked eye.

In typical fusion experiments, cells are distributed over ten microwell plates (960 wells) and 200–700 wells show colony growth. Of these hybridomas, up to 50% (100–300) produce antibodies that show some antigen binding, though many may be too weak to be useful. After further screening for desired affinity and specificities, we normally select 25–30 as a convenient number to clone. Table 1 illustrates the results obtained from four fusions by this method. Only occasionally, in our experience, are two or more MAb produced that are indistinguishable from each other when their epitope specificities are later examined in detail. In the early stages after fusion, a particularly good cell-culture system is needed to promote rapid hybridoma growth in HAT medium. Horse serum, selected for high cloning efficiency, is a much better growth promoter than most batches of fetal calf serum, and HAT medium is prepared using myelomaconditioned medium and human endothelial culture supernatant, a hybridoma growth promoter. Under these conditions, colonies are visible with the naked eye 7–10 d after fusion, and screening should start immediately. Even unimmunized mouse spleens will produce large numbers of colonies, so the use of hyperimmunized mice, with high serum titers in the screening assay, is essential for generating a high proportion of positive colonies. The screening assays are also of critical importance, because they determine the characteristics of the MAb produced. Both these points are illustrated in Fig. 1. CK, a fairly typical

Table 1 Examples of Colony and MAb Yields Using This Method Antigen 108–kDa rod fragment of dystrophin in fusion protein (2) 59–kDa rod fragment of dystrophin in fusion protein (4) 55–kDa C--terminal fragment of dystrophin in fusion protein (5)

Wells with growth

Wells binding to antigen strongly

Wells with desired Final no. affinity//specificity of MAb

587

136

27

16

669

147

30

25

700+

42

20

17

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“globular” enzyme, is denatured and inactivated when attached directly to ELISA plastic, but it can be captured onto ELISA plates in its native form by using Ig from a polyclonal antiserum (9). When mice were immunized with untreated CK, antibody titers against denatured CK were high, whereas titers of antibody against native CK were insufficient to produce a plateau in the capture ELISA, even at high serum concentrations (Fig. 1A). Seven fusions from such mice (over 1000 colonies screened) produced only two, low-affinity MAb specific for native CK. In contrast, when we immunized with CK aggregated by glutaraldehyde treatment (Fig. 1B), titers were higher for native CK antibodies than for those against denatured CK. In a fusion performed with one of these mice, only 2% of the fusion wells were positive in the direct ELISA (denatured CK), but 13% recognized only native CK in the capture ELISA (9). This illustrates the importance of having both high-titer mice and a suitable screen. Finally, although the method is rapid and efficient, it is also labor-intensive, especially during screening and cloning. When embarking on the method, one should not be daunted by the prospect of having over 70 culture plates in the incubator simultaneously by the time that the later cloning stages of a single experiment are reached. 2. Materials 1. One or two vertical laminar flow sterile hoods (e.g., Machaire, Bolton, UK). 2. 37°C CO2 incubator (e.g., LEEC, Nottingham, UK; see Note 1). 3. Transtar 96 (Corning Inc, Lowell, MA) for plating fusions and removing hybridoma supernatants. 4. Round-bottomed 96-well tissue-culture microwell plates (Invitrogen, Carlsbad, CA, cat. no. 163320) (see Note 2). 5. Sterile tissue-culture flasks 25 mL (Invitrogen), 75 mL (Invitrogen), and 24-well plates (Invitrogen), 1.8-mL cryogenic vials (AlphaLabs, Eastleigh, UK), and sterile 25-mL universal containers (Invitrogen). 6. Freund’s adjuvant, complete (Sigma [St. Louis, MO] no. F-5881) stored at 4°C. 7. Freund’s adjuvant, incomplete (Sigma) stored at 4°C. 8. Pristane (2,6,10,14 tetramethyl pentadecane; (Sigma) stored at room temperature. 9. 500X HT: 6.8 mg/mL hypoxanthine (Sigma) and 1.95 mg/mL thymidine (Sigma) in water (see Note 3). Filter to sterilize. Store in 5-mL aliquots in sterile plastic universals at −20°C (stable for a few years). 10. 1000X Aminopterin (Sigma): 4.4 mg are dissolved in 25 mL of water (see Note 3). Since the chemical is toxic and teratogenic, it must be weighed with appropriate containment and operator protection. Filter to sterilize. Store in 2-mL aliquots at −20°C, wrapped in aluminum foil to protect from light (stable for years). 11. 50% polyethylene glycol (PEG), mol wt 1500 (Merck, Darmstadt, Germany), is made up by weighing 10 g of PEG in a glass universal and autoclaving. Although the solution is still hot (about 60°C), add 10 mL of DMEM/25 mM HEPES

High-Affinity Monoclonal Antibodies from a Single Mouse

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14.

15. 16. 17. 18.

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prewarmed in a 37°C water bath. Mix well, and store in 5-mL aliquots at room temperature in the dark; stable at least for 6 mo. The pH should be slightly alkaline (as judged by phenol red) when used for fusion. DMEM/Glutamax I/25 mM HEPES. (Invitrogen). This is used as a “physiological saline,” not for growing cells. DMEM/Glutamax I/20% HS is prepared by adding to each 80 mL of 1X DMEMGlutamax I (Invitrogen): 1 mL penicillin-streptomycin (Invitrogen, cat. no. 15070063), 1 mL nonessential amino acids (MEM) (Invitrogen, cat. no. 11140), and 20 mL selected horse serum (HS). For routine growth and maintenance of both myelomas and established hybridomas, serum is reduced to 10%. Cloning medium: DMEM/20% HS supplemented with 1X HT and 5% human endothelial culture supernatant (HECS) (No longer available from Costar-Corning; see Note 16). HAT medium: Myeloma-conditioned DMEM-Glutamax I/20% HS with 1X HAT and 5% HECS. Medium for feeding the fusion: DMEM-Glutamax I/20% HS with 5% HECS and 5X HT. Medium for freezing down cells: 90% HS/10% dimethyl sulfoxide (DMSO) (Sigma). Trypan blue (0.1%) (Sigma) in PBS.

3. Methods 3.1. Immunization of Balb/c Mice 1. Purify the protein antigen. With purified antigen, screening is faster and easier, and the yield of MAb is higher, but proteins of 50% purity or less on SDS-PAGE may still give good results. 2. Each of three Balb/c mice (6–8 wk old) is given an ip injection of 50–100 µg of antigen in 0.1 mL PBS emulsified with an equal volume of Freund’s complete adjuvant by sucking up and down many times into a disposable sterile plastic 2-mL syringe with a sterile 21-gage 1–1/2 in. no. 2 needle (see Note 4). 3. Four weeks later, an ip boost of 50–100 µg antigen in Freund’s incomplete adjuvant is given. 4. Ten days later, blood (c. 0.1 mL) is taken from the tail vein, allowed to clot, and centrifuged at 10,000 rpm for 5 min. The serum is tested by the method to be used for screening hybridomas. If the titers are high in this assay (see Fig. 1), the mice should be left to rest for an additional month before fusion. 5. The best mouse is boosted with 50–100 µg of antigen in PBS, ip, and/or iv in the tail vein, 4, 3, and 2 d before fusion.

3.2. Selection of Horse Serum 1. Most suppliers will provide test samples of different serum batches and keep larger amounts on reserve. All sera are heat-inactivated at 56°C for 30 min before use, though this is not known to be essential. Thaw serum in 37°C water bath and swirl to mix before incubating in a 56°C water bath for 30 min with occasional swirling.

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We have never yet found a batch of fetal bovine serum that performs as well as horse serum in these tests (see also Subheading 3.8). 2. Prepare cells of any hybridoma or myeloma line at 1000 cells/mL in DMEM and dilute to 40 cells/mL, 20 cells/mL, 10 cells/mL in DMEM/20% HS of different batches (cloning of established cell lines does not require feeder layers or HT and HECS). For each test serum, plate one 96-well round-bottomed microplate with four rows at four cells in 0.1 mL/well, four rows at two cells in 0.1 mL/well, and four rows at one cell in 0.1 mL/well. 3. From the fifth day onward, note the number of colonies at each cell density for each test serum, as well as the size of the clones. After 2 wk, choose the batch of serum with the highest number and the largest clones. Serum may be stored at −70°C for at least 1–2 yr without loss of activity.

3.3. Growth and Maintenance of Myelomas SP2/O (21) myeloma lines grow very fast and do not synthesize immunoglobulins They should be kept in logarithmic growth in DMEM/5% HS for at least a wk before fusion by diluting them to 5 × 104/mL when the cell density reaches 4 or 5 × 105/mL. If thawing myelomas from liquid nitrogen, it is advisable to begin at least 1–2 wk before fusion (see Note 5). We have never found it necessary to treat lines with azaguanine to maintain their aminopterin sensitivity, which can be checked by plating in HAT medium at 100 cells/well. 3.4. Fusion with Spleen Cells 1. All dissection instruments are sterilized by dry heat (160°C for 5 h), autoclaving (120°C for 30 min) or dipping in 70% ethanol, and flaming in a Bunsen burner. Two separate fusions are done with each spleen as a precaution against accidents. 2. Two days before fusion, set up 2 × 75-mL flasks with 40 mL each DMEM 20% HS and 1 × 105 myeloma cells/mL (Sp2/O). 3. On the day of fusion, place in a 37°C water bath 100 mL of DMEM/25 mM HEPES, 10 mL of DMEM/20% HS, and 1 bottle of PEG1500. 4. Count the myeloma cells in a hemocytometer after mixing an aliquot with an equal volume of 0.1% Trypan blue in PBS; cell density should be 4–6 × 105 with viability 100% and no evidence of contamination. 5. Transfer the myeloma cells to four universals and centrifuge for 7 min at 300 g (MSE 4 L). Remove the supernatants with a 10-mL pipet and retain them for plating the fusion later (myeloma-conditioned medium). Resuspend each pellet in 5 mL of DMEM/HEPES, and combine in two universals (2 × 10 mL). 6. The immunized mouse is killed by cervical dislocation outside the culture room and completely immersed in 70% ethanol (15–30 s). Subsequent procedures are performed in sterile hoods. The mouse is pinned onto a “sterile” surface (polystyrene covered with aluminum foil and swabbed with 70% ethanol), and a small incision in the abdominal skin is made with scissors. The peritoneum is then exposed by tearing and washed with 70% ethanol before opening it and pinning it

High-Affinity Monoclonal Antibodies from a Single Mouse

7.

8.

9.

10.

11.

12.

13.

14. 15.

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aside. The spleen is lifted out, removing the attached pancreas with scissors, and placed in a sterile Petri dish (see Note 6). After removing as much surrounding tissue as possible, the spleen is placed in 10 mL of DMEM/25 mM HEPES and held at one end with forceps while making deep longitudinal cuts with a sterile, curved scalpel to release the lymphocytes and red blood cells. This process is completed by scraping with a Pasteur pipet sealed at the end until only connective tissue is left (2–3 min). The spleen cell suspension is transferred to a 25-mL universal, with pipeting up and down five to six times to complete the dispersal. Large lumps are allowed to settle for 1 min before transferring the supernatant to a second universal and centrifuging for 7 min at 300 g. Resuspend the pellet in 5 mL of DMEM/25 mM HEPES, and keep at room temperature. Usually about 1 × 108 cells/spleen are obtained (see Note 7). Add 2.5 mL spleen cell suspension to each 10 mL of myeloma cell suspension, and mix gently. Centrifuge at 300 g for 7 min at 20°C. Remove supernatants with a pipet, and resuspend pellets in 10 mL DMEM/25 mM HEPES using a pipet. Centrifuge at 300 g for 7 min. Remove the supernatant completely from one pellet (use a Pasteur pipet for the last traces), and then loosen the pellet by tapping the universal gently (see Note 8). Remove the DMEM/25 mM HEPES and the 50% PEG from the water bath just before use. Take 1 mL of PEG in a pipet, and add dropwise to the cell pellet over a period of 1 min, mixing between each drop by shaking gently in the hand. Continue to shake gently for another minute. Add 10 mL DMEM/25 mM HEPES dropwise with gentle mixing, 1 mL during the first minute, 2 mL during the second, and 3.5 mL during the third and fourth minutes. Centrifuge at 300 g for 7 min. (This procedure can be repeated with the second cell pellet, while the first is spinning.) Remove supernatant, and resuspend pellet gently in 5 mL of DMEM/20% HS. Place both 5-mL cell suspensions in their universals in the CO2 incubator for 1–3 h. During this time, take the c. 80 mL of myeloma-conditioned medium and add 4.5 mL of HECS, 90 µL of aminopterin, and 180 µL of hypoxanthine/thymidine solutions. Filter 2 × 40 mL through 0.22-µm filters (47 mm; Millipore [Bedford, MA]) to resterilize, and remove any remaining myeloma cells. To each 40 mL, add the 5-mL fusion mixture from the CO2 incubator and distribute in 96-well microtiter plates (4 plates/fusion; 100 µL/well) using a Transtar 96 or a plugged Pasteur pipet (3 drops/well) (see Note 9). Put the plates from each fusion in a separate lunch box in the CO2 incubator and leave for 3 d. On d 4, add to each well 80 µL of DMEM/20% HS supplemented with HECS and 5X HT. Replace in the CO2 incubator as quickly as possible. By d 10–14, a high proportion of the wells should have a clear, white, central colony of cells, easily visible with the naked eye. If you want to select for highaffinity MAb (and you would be well advised to do so, unless you have some very special objectives), do not delay screening. If you are using a sensitive screening method, such as ELISA, immunofluorescence, or Western blotting, you will detect

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high-affinity antibodies from even the very small colonies. Do not wait for the medium to turn yellow, or your cells may start to die (if you are using a less sensitive screen, such as an inhibition assay, you may need to wait longer). A high proportion of the 768 wells (50–100%) should have colony growth by 14 d. If it is