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2D PAGE: Sample Preparation and Fractionation

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METHODS IN MOLECULAR BIOLOGY TM 

2D PAGE: Sample 

Preparation 

and Fractionation 

Volume 2 

Edited by 

Anton Posch 

Bio-Rad Laboratories GmbH, Munich, Germany

Editor 

Anton Posch 

Bio-Rad Laboratories GmbH 

Munich, Germany 

Series Editor 

John M. Walker 

School of Life Sciences 

University of Hertfordshire 

Hatfield, Herts., UK 

ISBN: 978-1-60327-209-4 e-ISBN: 978-1-60327-210-0 

ISSN: 1064-3745 

Library of Congress Control Number: 2007943017 

©2008 Humana Press, a part of Springer Science+Business Media, LLC 

All rights reserved. This work may not be translated or copied in whole or in part without the written 

permission of the publisher (Humana Press, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 USA), 

except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form 

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The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are 

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or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the 

material contained herein. 

Cover illustration: Figure 3, Chapter 14, “The Terminator: A Device for High Throughput Extraction of 

Plant Material,” by B. M. van den Berg. 

Printed on acid-free paper 

987654321 

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Preface 

This book, split into two volumes, presents a broad coverage of the principles 

and recent developments of sample preparation and fractionation tools in 

Expression Proteomics in general and for two-dimensional electrophoresis 

(2-DE) in particular. 2-DE, with its unique capacity to resolve thousands of 

proteins in a single run, is still a fundamental research tool for nearly all 

protein-related scientific projects. The methods described here in detail are not 

limited to 2-DE and can also be applied to other protein separation techniques. 

Because each biological sample is unique, a suited sample preparation 

strategy has to consider the type of sample as well as the type of biological 

question being addressed. The complex nature of proteins often requires a 

multitude of sample preparation options. In addition, sample preparation is 

not only a prerequisite for a successful and reproducible Proteomics experiment, 

but also the key factor to meaningful data evaluation. Interestingly, not 

much attention was paid to this area during Proteomics methodology development 

and therefore this book is intended to explain in depth how proteins 

from various sources can be properly isolated and prepared for reproducible 

Proteome analysis. 

The application of fractionation and enrichment strategies has become a 

major part of sample preparation. The number of possible different proteins in 

a cell or tissue sample is believed to be in the several hundreds of thousands, 

spanning concentration ranges from the level of a single molecule to micromolar 

amounts, and no single analytical method developed today is capable of 

resolving and detecting such a diverse sample. Sample fractionation reduces the 

overall complexity of the sample, and enriches low abundance proteins relative 

to the original sample. Proteins that may originally have been undetectable are 

thus rendered amenable to analysis by 2-DE and a broad variety of gel-free 

mass spectrometry-based technologies. 

This book is for students of Biochemistry, Biomedicine, Biology, and 

Genomics and will be an invaluable source for the experienced, practicing 

scientist, too. 

Anton Posch 

v

Contents 

Preface ................................................................ v 

Contributors ........................................................... xi 

1. Application of Fluorescence Dye Saturation Labeling 

for Differential Proteome Analysis of 1,000 Microdissected 

Cells from Pancreatic Ductal Adenocarcinoma Precursor 

Lesions ......................................................... 

Barbara Sitek, Bence Sipos, Günter Klöppel, Wolff Schmiegel, 

Stephan A. Hahn, Helmut E. Meyer, and Kai Stühler 

1 

2. Albumin and Immunoglobulin Depletion of Human Plasma ........ 

Rosalind E. Jenkins, Neil R. Kitteringham, 

Carrie Greenough, and B. Kevin Park 

15 

3. Multi-Component Immunoaffinity Subtraction 

and Reversed-Phase Chromatography of Human Serum .......... 

James Martosella and Nina Zolotarjova 

27 

4. Immunoaffinity Fractionation of Plasma Proteins by Chicken IgY 

Antibodies...................................................... 

Lei Huang and Xiangming Fang 

41 

5. Proteomics of Cerebrospinal Fluid: 

Methods for Sample Processing ................................. 

John E. Hale, Valentina Gelfanova, Jin-Sam You, 

Michael D. Knierman, and Robert A. Dean 

53 

6. Sample Preparation of Bronchoalveolar Lavage Fluid ............... 

Baptiste Leroy, Paul Falmagne, and Ruddy Wattiez 

67 

7. Preparation of Nasal Secretions for Proteome Analysis .............. 

Begona Casado, Paolo Iadarola, and Lewis K. Pannell 

77 

8. Preparation of Urine Samples for Proteomic Analysis ............... 

Rembert Pieper 

89 

9. Isolation of Cytoplasmatic Proteins from Cultured Cells for 

Two-Dimensional Gel Electrophoresis ........................... 101 

Ying Wang, Jen-Fu Chiu, and Qing-Yu He 

vii

viii Contents 

10. Sample Preparation of Culture Medium from Madin-Darby 

Canine Kidney Cells ............................................ 113 

Daniel Ambort, Daniel Lottaz, and Erwin Sterchi 

11. Sample Preparation for Mass Spectrometry Analysis 

of Formalin-Fixed Paraffin-Embedded Tissue: Proteomic 

Analysis of Formalin-Fixed Tissue ............................... 131 

Nicolas A. Stewart and Timothy D. Veenstra 

12. Metalloproteomics in the Molecular Study of Cell Physiology 

and Disease .................................................... 139 

Hermann-Josef Thierse, Stefanie Helm, 

and Patrick Pankert 

13. Protein Extraction from Green Plant Tissue ......................... 149 

Ragnar Flengsrud 

14. The Terminator: A Device for High-Throughput Extraction of 

Plant Material .................................................. 153 

B. M. van den Berg 

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

Etienne H. Meyer and A. Harvey Millar 

16. Isolation and Preparation of Chloroplasts from Arabidopsis 

thaliana Plants .................................................. 171 

Sybille E. Kubis, Kathryn S. Lilley, and Paul Jarvis 

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

Elisabeth Jamet, Georges Boudart, Gisèle Borderies, 

Stephane Charmont, Claude Lafitte, Michel Rossignol, 

Herve Canut, and Rafael Pont-Lezica 

18. Isolation and Fractionation of the Endoplasmic Reticulum from 

Castor Bean (Ricinus communis) Endosperm for Proteomic 

Analyses........................................................203 

William J. Simon, Daniel J. Maltman 

and Antoni R. Slabas 

19. Cell Wall Fractionation for Yeast and Fungal Proteomics ........... 217 

Aida Pitarch, César Nombela, and Concha Gil 

20. Collection of Proteins Secreted from Yeast Protoplasts in Active 

Cell Wall Regeneration ......................................... 241 


21. Sample Preparation Procedure for Cellular Fungi ................... 265 

Alois Harder

Contents ix 

22. Isolation and Enrichment of Secreted Proteins from Filamentous 

Fungi ...........................................................275 

Martha L. Medina and Wilson A. Francisco 

23. Isolation and Solubilization of Cellular Membrane Proteins from 

Bacteria ........................................................ 287 

Kheir Zuobi-Hasona and L. Jeannine Brady 

24. Isolation and Solubilization of Gram-Positive Bacterial Cell 

Wall-Associated Proteins........................................295 

Jason N. Cole, Steven P. Djordjevic, and Mark J. Walker 

25. Cell Fractionation of Parasitic Protozoa ............................ 313 

Wanderley de Souza, José Andrés Morgado-Diaz, 

and Narcisa L. Cunha-e-Silva 

Index..................................................................333

Contributors 

Daniel Ambort • University of Berne, Berne, Switzerland 

Leroy Baptiste • University of Mons-Hainaut, Mons, Belgium 

Gisèle Borderies • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, 

Castanet-Tolosan, France 

Georges Boudart • UMR 5546 CNRS-Université Paul Sabatier-Toulouse 

III, Castanet-Tolosan, France 

L. Jeannine Brady • University of Florida, Gainesville, Florida 

Herve Canut • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, 


Begona Casado • Swiss Federal Institute of Technology, Zurich, Switzerland 

Stephane Charmont • Novartis Pharma AG, Basel, Switzerland 

Jen-Fu Chiu • The University of Hong Kong, Hong Kong, China 

Jason N. Cole • University of Wollongong, Wollongong, Australia 

Narcisa L. Cunha-e-Silva • Universidade Federal do Rio de Janeiro, 

Rio de Janeiro, Brazil 

Wanderley de Souza • Universidade Federal do Rio de Janeiro, Rio de 

Janeiro, Brazil 

Robert A. Dean • Lilly Research Laboratories, Indianapolis, Indiana 

Steven P. Djordjevic • Elizabeth Macarthur Agricultural Institute, 

Menangle, Australia 

Paul Falmagne • University of Mons-Hainaut, Mons, Belgium 

Xiangming Fang • GenWay Biotech, Inc., San Diego, California 

Ragnar Flengsrud • Norwegian University of Life Sciences, Ås, Norway 

Wilson A. Francisco • Arizona State University, Tempe, Arizona 

Valentina Gelfanova • Lilly Research Laboratories, Greenfield, Indiana 

Concha Gil • Complutense University of Madrid, Madrid, Spain 

Carrie Greenough • University of Liverpool, Liverpool, Great Britain 

Stephan A. Hahn • Ruhr-University Bochum, Bochum, Germany 

John E. Hale • Lilly Research Laboratories, Greenfield, Indiana 

Alois Harder • Toplab GmbH, Martinsried, Germany 

Qing-Yu He • The University of Hong Kong, Hong Kong, China 

Stefanie Helm • University of Heidelberg, Mannheim, Germany 

Lei Huang • GenWay Biotech Inc., San Diego, California 

xi

xii Contributors 

Paolo Iadarola • University of Pavia, Pavia, Italy 

Elisabeth Jamet • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, 


Paul Jarvis • University of Leicester, Leicester, Great Britain 

Rosalind E. Jenkins • University of Liverpool, Liverpool, Great Britain 

Neil R. Kitteringham • University of Liverpool, Liverpool, Great Britain 

Günter Klöppel • Christian Albrechts University, Kiel, Germany 

Michael D. Knierman • Lilly Research Laboratories, Greenfield, Indiana 

Sybille E. Kubis • University of Leicester, Leicester, Great Britain 

Claude Lafitte • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, 


Baptiste Leroy • University of Mons-Hainaut, Mons, Belgium 

Kathryn S. Lilley • University of Cambridge, Cambridge, Great Britain 

Daniel Lottaz • University of Berne, Berne, Switzerland 

Daniel J. Maltman • University of Durham, Durham, Great Britain 

James Martosella • Agilent Technologies Inc., Wilmington, Delaware 

Martha L. Medina • Arizona State University, Tempe, Arizona 

Helmut E. Meyer • Ruhr-University Bochum, Bochum, Germany 

Etienne H. Meyer • The University of Western Australia, Perth, Australia 

A. Harvey Millar • The University of Western Australia, Perth, Australia 

José Andrés Morgado-Diaz • Instituto Nacional de Câncer, Rio de 

Janeiro, Brazil 

César Nombela • Complutense University of Madrid, Madrid, Spain 

Patrick Pankert • University of Heidelberg, Mannheim, Germany 

Lewis K. Pannell • University of South Alabama, Mobile, Alabama 

B. Kevin Park • University of Liverpool, Liverpool, Great Britain 

Falmagne Paul • University of Mons-Hainaut, Mons, Belgium 

Rembert Pieper • The Institute for Genomic Research, Rockville, Maryland 

Aida Pitarch • Complutense University of Madrid, Madrid, Spain 

Rafael Pont-Lezica • UMR 5546 CNRS-Universitè Paul Sabatier-Toulouse 


Michel Rossignol • UMR 5546 CNRS-Universitè Paul Sabatier-Toulouse 


Wolff Schmiegel • Ruhr-University Bochum, Bochum, Germany 

William J. Simon • University of Durham, Durham, Great Britain 

Bence Sipos • Christian Albrechts University, Kiel, Germany 

Barbara Sitek • Ruhr-University Bochum, Bochum, Germany 

Antoni R. Slabas • University of Durham, Durham, Great Britain 

Erwin Sterchi • University of Berne, Berne, Switzerland 

Nicolas A. Stewart • National Cancer Institute at Frederick, Frederick, 

Maryland

Contributors xiii 

Kai Stühler • Ruhr-University Bochum, Bochum, Germany 

Hermann-Josef Thierse • University of Heidelberg, Mannheim, Germany 

B. M. van den Berg • Elexa, Enkhuizen, The Netherlands 

Timothy D. Veenstra • National Cancer Institute at Frederick, Frederick, 

Maryland 

Mark J. Walker • University of Wollongong, Wollongong, Australia 

Ying Wang • The University of Hong Kong, Hong Kong, China 

Ruddy Wattiez • University of Mons-Hainaut, Mons, Belgium 

Jin-Sam You • Indiana Centers for Applied Protein Sciences, Indianapolis, 

Indiana 

Nina Zolotarjova • Agilent Technologies Inc., Wilmington, Delaware 

Kheir Zuobi-Hasona • University of Florida, Gainesville, Florida

1 

Application of Fluorescence Dye Saturation Labeling 

for Differential Proteome Analysis of 1,000 

Microdissected Cells from Pancreatic Ductal 

Adenocarcinoma Precursor Lesions 

Barbara Sitek, Bence Sipos, Günter Klöppel, Wolff Schmiegel, 

Stephan A. Hahn, Helmut E. Meyer, and Kai Stühler 

Summary 

The identification of molecular changes underlying clinical pathogenic processes is 

often hampered by significant cellular diversity of the tissue. Pathogenic aberrant cells 

are surrounded by cells originating e.g., from stroma, the vascular system or other neighbouring 

cell types, which lead to under-representation of interesting cells when analysing 

whole tissue specimen. Therefore, selection of relevant cell types for detailed analysis 

is an absolute prerequisite for in depth elucidation of underlying biological processes. 

Microdissection offers the advantage to select for a biologically relevant cell type which 

is often in low abundance. Here, we present a proteomics approach allowing us to analyse 

1,000 microdissected cells stemming from pancreatic carcinoma precursor lesions applying 

fluorescence dye saturation labeling. 

Key Words: Difference gel electrophoresis; DIGE; fluorescence dye saturation 

labeling; pancreatic ductal adenocarcinoma; panin, tumor marker; two-dimensional gel 

electrophoresis. 

1. Introduction 

To identify new candidate molecular markers for pancreatic ductal adenocarcinoma 

we established a proteomics approach analysing microdissected 

cells from precursor lesions, the so called pancreatic intraepithelial neoplasia 

From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 

Edited by: A. Posch © Humana Press, Totowa, NJ 

1

2 Sitek et al. 

(PanIN) (1). Due to a limited amount of proteins available from microdissection 

we developed a procedure which included fluorescence dye saturation labeling 

in combination with high resolution two-dimensional gel electrophoresis 

(2-DE) (2). With this procedure we were able to analyse proteins extracted 

from 1,000 microdissected cells with a high resolution of up to 2,500 protein 

spots. For differential proteome analysis we analysed microdissected cells 

from 9 patients. We compared the protein expression of the different PanIN 

grades (PanIN 1A/B, PanIN 2, PanIN 3) and carcinoma cells related to normal 

epithelial cells and found 86 significantly regulated spots (p < 0.05, ratio >1.6). 

Using protein lysates from pancreatic carcinoma tissue as a reference proteome 

we were able to successfully identify the proteins after tryptic in-gel digestion. 

2. Materials 

2.1. Microdissection and Sample Preparation 

1. Microscope: BH2 (Olympus, Wetzla, Germany). 

2. Cryostat: Cryotom SME (Shandon). 

3. Ultrasonic bath (VWR). 

4. Hand homogenisator. 

5. Injection needle: 0.65 mm × 25 mm (Braun, Melsungen, Germany). 

6. Lysis buffer: 2 M thiourea, 7 M urea, 4% 3-[(3-cholamidopropyl) 

dimethylammino]-1-propane sulfonate (CHAPS), 30 mM Tris-HCl, pH 8.0. 

7. Hematoxylin stain: 25% (v/v) hematoxylin according to Mayer (Merck). 

8. Eosin stain: 1.7% (w/v) eosin (Merck), diluted in 96% ethanol. 

2.2. Cysteine-Specific Protein Labeling Using CyDye DIGE 

Fluor Saturation Dyes 

1. Dye stock solution: CyDye DIGE Fluor saturation dyes solid compounds are 

reconstituted in dimethylformamide (DMF) giving a concentration of 2 mM (50 

μL DMF to 100 nM of dye) for analytical gels and 20 mM (20 μL DMF to 400 

nM of dye) for preparative gels. It is stable at –20°C for 3 mo. 

2. 50 mM NaOH solution (for pH adjustment). 

3. 2 mM TCEP (triscarboxethylphosphine) solution for analytical gels and 20 mM 

TCEP solution for preparative gels. 

4. Image software: ImageQuantTM (GE Healthcare Bioscience). 

5. Differential image analysis software: Decyder v5.02 (GE Healthcare Bioscience). 

6. Fluorescence Scanner: Typhoon 9400 (GE Healthcare Bioscience). 

2.3. Isoelectric Focusing 

1. Seperation gel buffer: 3.5% (w/v) acrylamide, 0.3% (w/v) piperazindiacrylamide, 

4% (v/v) carrier ampholines mixture (pH 2–11), 9.0 M urea, 5% glycerol, 0.06% 

(v/v) TEMED.

Application of Fluorescence Dye Saturation Labeling 3 

2. Cap gel buffer: 12.3% (w/v) acrylamide, 0.13% (w/v) piperazindiacrylamide, 4% 

(v/v) carrier ampholyte mixture, 9.0 M urea, 5% glycerol, 0.06% (v/v) TEMED 

3. 1.2% (w/v) APS (Ammoniumpersulfat). 

4. Anodic buffer: 3 M urea, 7.3% (w/v) phosphoric acid. 

5. Cathodic buffer: 9 M urea, 5% (w/v) glycerol, 0.75 M ethylendiamine. 

6. Sephadex solution: 270 mg Sephadex suspension (20 g Sephadex was swollen in 

500 mL water, resuspended into 1Lof25%glycerol solution and filtered), plus 

233 mg urea, plus 98 mg thiourea, and 25 μL ampholine mixture, pH 2–11 and 

25 μL DTT (1.08 g/5 mL). 

7. Protection solution: 30% urea (w/v), 5% glycerol (w/v), 2% carrier ampholytes, 

pH 2–4. 

8. Equilibration solution: 125 mM Tris-base, 40% glycerol, 65 mM DTT, 3% SDS. 

2.4. Two-dimensional Polyacrylamid Gel Electrophoresis (2D-PAGE) 

1. Glass plates compatible with fluorescence imaging (25 × 30 × 0.4 cm). 

2. Plastic spacer (30 ×1×0.15 cm). 

3. Apparatus for vertical SDS-PAGE. (System VA Sarstedt, Nümbrecht, Germany) 

4. Gel carrier grooves (self made). 

5. Gel solution: 570 mM Tris-base and 180 mM Tris-HCL, 0.06% TEMED, 0.2% 

SDS, 15% acrylamide, 0.2% bisacrylamide. 

6. Running buffer: 0.2 M Tris, 1.92 mM glycine, 0.1% (w/v) SDS. 

7. 40% (w/w) APS (Ammoniumpersulfat). 

8. Protection solution: 285 mM Tris-base and 90 mM Tris-HCL, 0.1% SDS. 

9. Agarose solution: 1% (w/w) agarose (dissolved in running buffer), 0.001% (w/w) 

bromphenol blue. 

3. Methods 

The number of cells available by manual microdissection is rather 

limited. When analysing precursor lesions of the pancreatic adenocarcinoma 

only 1,000–5,000 cells can be provided in a reasonable time window (3–4 h). 

Due to the scarce protein amount (1,000 cells are equivalent to approx 2 μg 

protein) applying classical 2-DE techniques in combination with silver staining 

(loading amount 100 μg) or difference gel electrophoresis (DIGE) minimal 

labeling (50 μg) is not feasible. Therefore, DIGE saturation labeling which 

has a 50-fold higher sensitivity for protein detection must be applied when 

analysing such scarce sample amount cells (3). 

DIGE saturation labeling is based on covalent attachment of all protein 

cysteine residues prior to 2-DE (4). In contrast to DIGE minimal labeling 

where only one lysine residue of approx 3% of a protein species is labeled by 

a fluorescence dye (Cy2, Cy3 or Cy5) (5), DIGE saturation dyes leads to a 

complete labeling of all proteins (3).


Moreover, in contrast to DIGE minimal labeling multiplexing within one gel 

for direct differential analysis is not feasible when using saturation labeling. 

Effects of fluorescence resonance energy transfer (FRET), by which the fluorescence 

emission of a CyDye can be enhanced or quenched, respectively, have to 

be considered (6). Therefore, only two saturation dyes are available, whereof 

one CyDye (mostly Cy5) is taken for differential analysis whereas the other 

CyDye (mostly Cy3) is taken as an internal standard for controlling system 

variation that ultimately provides a more accurate quantification of relative 

protein abundance. 

Furthermore, preparative protein amounts (300–500 μg) for protein identification 

can not be provided by microdissection. Therefore, a reference proteome 

stemming from a comparable source (e.g., whole tissue or cell culture lysate) 

must be determined. This reference proteome should match the proteome of 

the microdissected samples to a high degree (>90%) and thus facilitates protein 

identification in subsequent in-gel digestion using mass spectrometry (1). 

However, before commencing DIGE analysis one must optimally determine 

the labeling conditions i.e., the proportion of fluorescent dye to microdissected 

cells. It has been shown (own observations) that due to inherent differences 

in the cysteine content of a given proteome, coupled with variable sample 

conditions, one has to independently develop an optimisation procedure for each 

sample. This allows a high performance and high quality proteomic analysis. 

3.1. Optimised Manual Microdissection for the Proteome Analysis 

of Pancreatic Adenocarcinoma Cells 

1. Froze tissue resections containing pancreatic carcinoma on liquid nitrogen and 

store at –80°C (see Note 1). For histological classification of normal ducts and 

different PanIN stages prepare 5 μm frozen sections of peritumoral pancreatic 

parenchyma using a cryostat and stain with hematoxylin and eosin. 

2. For microdissection in subsequent tissue slides prepare 10-μm frozen section from 

tissue blocks containing the required grades and only stain with haematoxylin 

(Fig. 1) It has been shown that eosin interferes with 2-DE leading to reduced 

protein recovery (Fig. 2). 

3. Under microscopic observation harvest the required number of cells using a sterile 

injection needle. 

4. Lyse the microdissected cells in 100 μL lysis buffer (4°C), sonicate (6 × 10 sec 

pulses on ice) after each collection step and finally centrifuge (12,000g for 5 min) 

the lysate and store the supernatant at –80°C. 

3.2. Determination of the Minimal Number of Microdissected Cells 

1. Because microdissection is very time consuming and the number of available 

sample material is limited it is necessary to find the minimal number of cells


Fig. 1. Manual microdissection. Subsequent histological classification the 

hematoxylin stained (10 μm serial sections) PanIN lesions (A) were microdissected 

under a microscope using a sterile injection needle (B). 

Fig. 2. Compatibility of H&E staining with 2-DE. Influence of H&E staining 

was investigated applying 7000 microdissected cells, each stained with H&E (A) or 

hematoxylin only (B). After microdissection and labeling with Cy3 the samples were 

separated by 2-DE and scanned using fluorescence scanner. Due to the weak protein 

recovery shown in (A) it is obvious that eosin interferes with proteome analysis and 

that hematoxylin staining is compatible with 2-DE analysis.


Fig. 3. Determination of the minimal number of microdissected cells required for 

2DE. Aliquots with 1000 and 7000 cells were labeled with Cy3 and analysed using 

2-DE under the same conditions. The spots in the gels were detected using DIA mode 

of DeCyder software. In the gel with 1000 cells 2500 spots (A) and in the gel with 

7000 cells 2600 spots (B) could be detected. 

sufficient for a differential proteome analysis. Therefore, a pool of approx 30,000 

microdissected cells is required allowing to determine the optimal number of 

spots by a dilution experiment. 

2. Prepare four aliquots containing 1,000, 2,500, 5,000, and 7,000 cells in the volume 

of 100 μL lysis buffer, respectively (see Note 2). 

3. For protein labeling use the standard protocol according to user manual. Briefly, 

add 4 nM TCEP to each sample and incubate for 1hat37°C. Label the samples 

with 8 nM saturation dyes for 30 min at 37°C. For stopping the labeling reaction 

add 10 μL DTT and 10 μL Ampholytes, pH 2–4, before 2-DE. 

4. Subsequent to 2-DE scan the gels as described elsewhere (see Note 3) and detect 

the number of spots using DIA module of DeCyder software (see Note 4). The 

optimal number of cells considered for subsequent analyses results from highest 

number of protein spots per analysed number of cells (Fig. 3). 

3.3. Optimisation of Fluorescence Dye Labeling 

1. Having established the optimal number of cells for comprehensive proteome 

coverage, one must subsequently empirically optimise the labeling conditions. 

This avoids any effects of over- and under-labeling. Therefore, a so called


Fig. 4. Effects of different DIGE saturation labeling conditions detected by 

same/same experiment. For the optimal application of DIGE saturation labeling the 

labeling conditions have to be optimized. (A) If too much dye amount has been applied 

over-labeling leads to a horizontal shift detected in CyDye-dependent manner (arrows). 

(B) Under-labeling of proteins with CyDye results in vertical streaking. (C) The optimal 

label condition is reached if an exact overlay of both fluorescence images can be 

detected. 

“same/same” experiment titrating different dye amounts is performed. Thus, 

prepare six aliquots containing the optimal number of cells (see Chapter 3.2) from 

the pool of 30,000 cells. 

2. Label three of them with 2, 4, and 8 nM of Cy3 and another three with 2, 4, and 

8nM of Cy5. 

3. Mix samples with equal dye amount and perform 2-DE for same/same experiment. 

4. After gel scanning analyse the overlay images in order to find the optimal ratio 

of protein and dye showing an accurate overlay of Cy3 and Cy5 images using 

ImageQuant TM . As shown in Fig. 4, applying too much dye results in horizontal 

shifts could occur due to additional labeling (e.g., -amino group of lysines), 

whereas an insufficient amount of dye causes vertical streaks due to inadequate 

protein labeling. 

3.4. Reference Proteome for Internal Standardization 

and Protein Identification 

1. 2-DE analysis of 1,000 microdissected cells (∼2 μg) does not deliver sufficient 

sample material for protein identification using mass spectrometry. Therefore, 

a reference proteome from a closely related source has to be defined allowing 

protein identification by protein spot assignment. Additionally, this reference 

proteome may be considered as an internal standard (labeled with Cy3) which 

reduces consumption of microdissected samples. 

2. It is therefore imperative that a suitable reference proteome is identified i.e. 

one which not only demonstrates a high correlation to microdissected cells, but 

also provides an adequate protein concentration for subsequent identification.


Therefore freeze several tissues containing pancreatic carcinoma on liquid 

nitrogen directly after resection and store at –80°C. 

3. For homogenization use 100 mg of the tissue on liquid nitrogen in 148 μL lysis 

buffer using a hand homogeniser and sonicate the sample 6 times for 10 sec 

on ice. 

4. To remove insoluble debris centrifuge (12,000g for 5 min) the homogenisate. 

5. Label 3 μg of each lysate with 2 nM Cy3 according to the standard labeling 

protocol according to user manual. 

6. Label aliquots of 1,000 microdissected cells with 4 nM Cy5. 

7. Pair-wise mix the labeled tissue lysates with labeled cells and process by 2-DE. 

8. After scanning, analyze the images using DIA module of DeCyder (Fig. 5) and 

calculate a matching rate between microdissected cells and carcinoma tissue (see 

Note 5). 

9. Optimize the labeling conditions for the tissue lysate showing the highest matching 

rate with the microdissected cells also by same/same experiment (see Chapter 3.3). 

Fig. 5. Protein spot pattern of carcinoma tissue (reference proteome) and microdissected 

cells after 2-DE. For protein identification and internal standardization a 

reference proteome with high degree in protein spot matching is necessary. For the 

proteome of a pancreatic tissue lysate a high matching rate (>90%) has been determined. 

In section A all detected protein spots of carcinoma tissue and microdissected 

cells, respectively, can be matched (letters are shown for better assignment). Whereas 

in section B beside a number of assignable protein spots (numbers) a protein spot 

unique for microdissected cells (arrows) was revealed.


3.5. Two-Dimensional Gel Electrophoresis 

1. These instructions are modified from the 2-DE technique as described by Klose 

and Kobalz (2). This technique is based on isoelectric focusing employing carrier 

ampholyte tube gels. It can be easily adapted to the DIGE technique and other 

formats, including analytical as well as preparative gels. In spite of the fact 

that most of the instruments were constructed in-house, equivalent equipment 

is commercially available. For the second dimension the Desaga VA300 gel 

system is applied. 

2. Two days before running isoelectric focusing (IEF) tube gels (Ø 1.5 mm, 20 cm) 

are prepared. Add 45 μL of APS solution to 2 mL of separations gel solution 

and fill tube to first mark (20 cm) using a syringe. Now, add 14 μL APS solution 

to 0.7 mL cap gel solution and cast cap gel to second mark (20.5 cm) behind 

the separation gel. To prevent urea crystallisation place an air cushion under the 

cap gel to third mark (21 cm). 

3. Before starting IEF apply 2 mm sephadex solution to prevent protein precipitation 

onto the separation gel. Then load the sample (dye-labeled) and overlay 

the sample with protection solution (approx 5 mm) to prevent direct contact of 

the acidic cathodic buffer (see Note 6). 

4. Fill anodic (bottom) and cathodic buffer (top) into the IEF chambers. Ensure 

that no air bubbles hamper IEF (see Note 6). Start isoelectric focusing applying 

a step-wise voltage program (100V for 1 h, 200 V/1 h, 400 V/17.5 h, 650 V/1 

h, 1,000 V/30 min, 1,500 V/10 min, 2,000 V/5 min). 

5. While the 21.5 h IEF is running, gels for the second dimension should be 

prepared. Clean the glass plates thrice (gel side) using a lint-free paper towel— 

for the first wash use doubled distilled water followed by 100% ethanol and 

finally 70% ethanol. To ensure correct gel dimension two 1.5 mm plastic spacers 

are placed between two plates sealed by silicon. 

6. Add 288 μL APS solution into 144 mL gel buffer, cast the gel and overlay with 

water-saturated isobutanol. 

7. After polymerization (45 min) remove isobutanol and wash the surface with a 

protection solution. To protect gel drying place 2-DE protection solution onto 

the gel and store the gels at 4°C. 

8. After IEF extrude the gel by means of inserting a nylon fiber (see Note 7) into 

the gel groove of the IEF gel carrier and incubate with equilibration solution for 

15 min to load proteins with SDS. 

9. Wash the gel three times with running buffer before applying to the second 

dimension. 

10. For the transfer of the IEF gel into SDS-PAGE gel hold the groove with gel 

in contact with the edge of the glass plate and slide the gel between the glass 

plates using a wire suitably formed. 

11. Overlay the IEF gels with agarose solution, add the running buffer to the upper 

and lower (15°C) chambers and start the electrophoresis. For the entrance of the 

proteins into the SDS-PAGE gel apply low current (75 mA) for 15min. When the 

proteins have entered increase the current to 200 mA for approximately 5–7 h.


3.6. Differential Proteome Analysis of 1000 Microdissected Cells 

from Different PanIN Stages 

1. This instruction comprised all steps obtained during optimisation procedure as 

described above. 

2. Harvest 1000 cells by manual microdissection and incubate the cells in 100 μL 

lysis buffer. 

3. For protein labeling according to the optimised protocol reduce the proteins with 

2 nM TCEP and label the proteins of the microdissected cells with 4 nM Cy5. 

4. For the preparation of the internal standard label 3 μg protein from a PDAC 

tissue sample lysate with 2 nM Cy3. 

5. Add 10 μL DTT to stop labeling reaction and add 10 μL Ampholytes, pH 2–4. 

Fig. 6. Representative images for each investigated progression step of PDAC. The 

lysates from 1000 microdissected cells were labeled with Cy5 mixed with internal 

standard and processed by 2-DE. Subsequent image acquisition the spot patterns were 

analyzed using DeCyder software and 2,000–2,500 protein spots have been detected 

for the different PanIN grades. Protein spot 2574 has been detected as differentially 

expressed in PanIN 2, PanIN 3 and Carcinoma (see Fig.7)


6. Mix the sample and analyse the mixture using 2-DE. 

7. After 2-DE, leave the gels between the glass plates and acquire the images 

using the Typhoon 9400 scanner (see Note 3). Therefore, choose excitation 

wavelengths and emission filters specific for each of the CyDyes according to 

the Typhoon user guide. 

8. Before image analysis with DeCyder software crop the images with 

ImageQuant TM software (Fig. 6). 

9. For intra-gel spot detection and quantification use the Differential In-gel Analysis 

(DIA) mode of the DeCyder software. Set the estimated number of spots to 

3000. Apply an exclusion filter to remove spots with a slope greater than 1.6 

(see Note 4). 

10. After spot matching between the different gels using the Biological Variation 

Analysis mode (BVA) consider only protein spots with an expression changes 

of factor > 1.6 and p-value (Student’s t-test) < 0.05 as significantly regulated 

(Fig. 7). 

Fig. 7. Stage-dependent regulation of protein spot 2574. For each patient and PanIN 

stage the protein spot intensity is shown. The depicted protein spot 2574 (see Fig. 6) 

shows a significant up-regulation (p < 0.05) in the carcinoma stage by a factor of 3.2. 

In the PanIN stages 2 and 3 a significant down-regulation by a factor of -2.7 and -1.8 

has been determined.


3.7. Micropreparation of Significantly Regulated Protein Spots 

1. After differential analysis protein spots are identified using mass spectrometry 

(MALDI-MS, ESI-MS) subsequent tryptic in-gel digestion. Therefore, 100-fold 

protein amount of the internal standard (reference proteome) must be labeled and 

applied to 2-DE. A preparative label kit with 400 nM Cy3 is available. 

2. For preparative gels label 400 μg of tissue lysate with 260 nM of Cy3 (130 nM 

TCEP) (see Note 8). 

3. Directly after gel scanning isolate the protein spots of interest manually (see 

Note 9). Assign the positions of the spots using a printout of the gel placed 

underneath the glass plate. 

4. Put the isolated protein spots into sample cups (glass) and store them at –80°C. 

5. For protein identification different mass spectrometric (MS) techniques can 

be applied subsequent to enzymatic in-gel digestion. Matrix assisted laser 

desorption/ionization MS (MALDI-MS) allows a fast and sensitive MS analysis 

performing peptide mass fingerprinting (PMF) (7). In cases where more 

sequence information is necessary (e.g. posttranslational modification) liquid 

chromatography-coupled electrospray ionisation MS (LC-ESI-MS) with its high 

performance for peptide fragment mass fingerprinting (PFF) is preferable (8). 

4. Notes 

1. To protect samples against tissue disruption (freezer burn) after resection the 

tissue is placed in tin foil at first and then stored in a cryotube. 

2. Usually, depending of tube’s diameter not more than 50 μL can be applied. 

Therefore, we increased the volume by blowing up the glass tube so that a sample 

volume of 100–200 μL can be applied. 

3. Before scanning the glass plates have to be cleaned thoroughly. First of all use 

ethanol for removing of acrylamide or silicone and then clean the glass plates 

with water. 

4. To avoid the detection of dust particles and artifacts as protein spots in the gel, 

an exclusion filter concerning slope, area, peak height, or volume can be applied. 

For one or more of these characteristics a value that distinguishes dust particles 

and spots has to be found. 

5. For the determination of the matching rate between two images which derived 

from the same gel analyze the gel using DIA mode of DeCyder software. After a 

spot detection an average ratio of 2.0 could be set in order to find spots occurring 

in both images (Average ratio < 2.0). 

6. Air bubbles should be avoided by applying the solution slowly under the surface 

(1 mm) of the solution which has been applied before. For Sephadex application 

the small volume of separation gel buffer generated during polymerisation is 

sufficient. 

7. To prevent destruction of the IEF gel by the nylon fiber (i) the thermoplastic nylon 

fiber should be fitted to the tube inner diameter by melting one end into the tube 

and (ii) the gel should be extruded using the cap gel (acrylamide concentration


12.3%) as a cushion or (iii) another possibility is to polymerise a high concentrated 

acrylamide solution (15%) above the extruding side and use this gel piece as a 

cushion. 

8. To avoid a high sample volume for the preparative labeling TCEP and Cy3 should 

have a concentration of 20 mM respectively instead of 2 mM (see Chapter 2.2). 

The amount of TCEP and Cy3 for labeling of preparative protein lysate has to be 

calculated according to labeling conditions for analytical gels. 

9. Before scanning, mark the glass plate from the picking gel with fluorescence 

stickers The stickers are necessary for matching between the gel (the proteins are 

not visible) and the gel image. After scanning print the gel image in the original 

size. Put the print under the glass plates, align the position of the stickers on the 

glass plate with the image. 

Acknowledgments 

The authors would like to thank Kathy Pfeiffer, Conny Bieling, Sabine 

Burkert and Birgit Streletzki for excellent technical assistance and Jon Barbour 

for critical reading of the manuscript. This work was supported by the grant 

from the Deutsche Krebshilfe (B.S., J.L., S.A.H and K.S., 70-2988-Schm3), 

Bundesministerium für Bildung und Forschung (NGFN, FZ 031U119) and the 

Nordrhein Westfalen Ministerium für Wissenschaft und Forschung. 

References 

1. Sitek, B., Lüttges, J., Marcus, K., Klöppel, G., Schmiegel, W., Meyer, H. E., Hahn, S. 

A. and Stühler, K. (2005) Application of fluorescence difference gel electrophoresis 

saturation labeling for the analysis of microdissected precursor lesions of pancreatic 

ductal adenocarcinoma. J. Proteomics, 5, 2665–2679. 

2. Klose, J. and Kobalz, U. (1995) Two-dimensional electrophoresis of proteins: an 

updated protocol and implications for a functional analysis of the genome. J. 

Electrophoresis 6, 1034–1059. 

3. Kondo, T., Seike, M., Mori, Y., Fujii,K., Yamada, T., and Hirohashi, S. (2003) 

Application of sensitive fluorescent dyes in linkage of laser microdissection and 

two-dimensional gel J. Proteomics 3, 1758–1766. 

4. Sitek, B., Scheibe, B., Jung, K, Schramm ,A., and Stühler, K. (2006) Difference gel 

electrophoresis (dige): the next generation of two-dimensional gel electrophoresis 

for clinical research, in Proteomics in Drug Research (Hamacher et al., eds), Wiley- 

VCH, Weinberg, pp 33–55. 

5. Unlü, M., Morgan,M. E., and Minden,J. S. (1997) Difference gel electrophoresis: 

a single gel method for detecting changes in protein extracts. Electrophoresis 18, 

2071–2077. 

6. Gruber, H. J., Hahn, C. D., Kada, G., Riener, C. K., Harms, G. S., Ahrer, W., 

Dax, T. G., and Knaus, H.-G. (2000) Anomalous fluorescence enhancement of Cy3


and cy3.5 versus anomalous fluorescence loss of Cy5 and Cy7 upon covalent linking 

to IgG and noncovalent binding to avidin. Bioconjug. Chem. 11, 696–704. 

7. Stühler, K., and Meyer, H. E. (2004) MALDI: more than peptide mass fingerprints. 

Curr Opin Mol Ther. 3, 239–248. 

8. Marcus, K., Moebius, J., and Meyer, H. E. (2003) Differential analysis of phosphorylated 

proteins in resting and thrombin-stimulated human platelets. Anal Bioanal 

Chem. 7, 973–993.

2 

Albumin and Immunoglobulin Depletion 

of Human Plasma 

Rosalind E. Jenkins, Neil R. Kitteringham, Carrie Greenough, 

and B. Kevin Park 

Summary 

Plasma and serum have been the focus of intense study in recent years in the expectation 

that they will provide important biomarkers of health and disease, without the need for 

invasive procedures. This aim has been hindered by the fact that a few highly abundant 

proteins dominate the protein profile, masking the lower abundance proteins and limiting 

our ability to analyse the entire plasma proteome. This chapter details a simple and 

effective method for removal of two of the most dominant proteins in plasma, albumin 

and immunoglobulin. 

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

proteome. 


The dynamic range of proteins in human plasma is greater than ten orders 

of magnitude, from albumin present at around 30 mg/mL to cytokines such 

as interleukin 6 present at basal levels of 1–3 pg/mL (1). This extraordinarily 

wide range of protein concentrations within a single compartment makes 

detailed analysis of the lower abundance proteins technically demanding, yet it 

is exactly these low level species that are likely to provide insight into disease 

states, into the effects of therapeutic intervention, and to yield specific and 

sensitive biomarkers. Many methods have been developed to deplete serum and 



15

16 Jenkins et al. 

plasma of the high abundance, housekeeping proteins, including alcohol precipitation, 

ultracentrifugation, salting in/salting out (2,3), and molecular weight 

fractionation (4,5). However, removal of proteins by affinity capture is the 

most commonly used approach because of the highly selective nature of the 

depletion. 

Anti-human albumin monoclonal antibody immobilized onto a solid support 

such as sepharose forms the basis of many immunoaffinity columns for the 

removal of the most abundant protein in human plasma. Clearly the affinity 

and specificity of the antibody determine the efficiency and discrimination of 

protein removal. However, the support on which the antibody is immobilized 

will also impact on the process, not least in terms of the binding capacity of 

the matrix. Affinity matrices based on bacterial protein A, protein G, or protein 

L, or molecularly engineered versions of these, are most commonly used to 

isolate immunoglobulins (6–11). Sophisticated multicomponent immunoaffinity 

matrices that are designed to deplete 10–15 of the most abundant plasma 

proteins are available (12) and have been used with great success, but they do 

have several drawbacks: they are expensive; they have a relatively low binding 

capacity so that several aliquots of plasma must be processed individually 

and then pooled to generate sufficient material for further analysis; and the 

depletion of the target proteins is frequently far from complete. The method 

described in this chapter, although limited to the depletion of only two of the 

highest abundance proteins, is simple, rapid, and accessible and provides a 

significant improvement in coverage of the plasma proteome obtainable by 2D 

gel electrophoresis. 

2. Materials 

2.1. Blood Sampling 

1. Heparinized tubes. 

2. Refrigerated centrifuge (up to 450g). 

3. 0.5-mL Eppendorf tubes. 

2.2. Albumin Depletion 

1. 2-mL cartridge containing POROS® beads coated with monoclonal goat antihuman 

serum albumin (HSA) (Applied Biosystems). 

2. Loading and washing buffer: phosphate buffered saline (PBS): 137 mM NaCl, 1 

mM KH2PO4, 5.6 mM Na2HPO4, 2.7 mM KCl, pH 7.4. 

3. Elution buffer: 12 mM HCl. 

4. HSA column wash buffer: 1 M NaCl. 

5. High performance liquid chromatography system capable of flow rates from 0.5–3 

mL/min (for automated depletions).

Albumin and Immunoglobulin Depletion of Human Plasma 17 

6. 2-mL and 10-mL plastic syringes (for manual depletions). 

7. Blunt ended syringe needle (for manual depletions). 


9. Spectrophotometer able to read the absorbance at 280 nm (for manual depletions). 

2.3. Immunoglobulin Depletion 

1. 0.2 mL cartridge containing POROS® beads coated with recombinant protein G 

(Applied Biosystems). 

2. Loading and washing buffer: phosphate buffered saline (PBS). 

3. Elution buffer: 12 mM HCl. 

4. Protein G column wash buffer: 1 M NaCl 10% acetic acid. 

5. High performance liquid chromatography system capable of flow rates from 0.5–3 

mL/min (for automated depletions). 

6. 2-mL and 10-mL plastic syringes (for manual depletions). 

7. Blunt ended syringe needle (for manual depletions). 


9. Spectrophotometer able to read the absorbance at 280 nm (for manual depletions). 

2.4. One-Dimensional (1D) gel Electrophoresis 

1. Standard 1D gel electrophoresis apparatus. 

2. Laemmli sample buffer (1×): 3% SDS, 0.1 M Tris-HCl, 15% glycerol, 0.2% 

bromophenol blue, pH 7.6. 

2.5. TCA Precipitation of Proteins 

1. 20% trichloroacetic acid (TCA) and ice-cold acetone. 

2. Resuspension buffer: 5% SDS, 1.15% DTT. 

3. Refrigerated centrifuge (up to 14,000g). 

2.6. 2D Gel Electrophoresis 

1. Standard 2D gel electrophoresis apparatus. 

2. IPG rehydration buffer: 9 M urea, 2% (w/v) CHAPS, bromophenol blue (trace), 

2% IPG buffer (Pharmacia), 0.28% DTT 

3. Methods 

3.1. Collection of Samples 

1. Collect the blood into heparinized tubes and sediment the red blood cells as soon 

after acquisition as possible by centrifugation at 450g for 10 min (see Notes 1, 

2, and 3).


2. Recover the supernatant, divide it into small aliquots and store them at –80°C. 

Each aliquot should be thawed and used only once. 

3. Once thawed, centrifuge the aliquot briefly to remove any precipitate that may 

have formed at low temperatures and that may block the affinity cartridges. 

3.2. Albumin Depletion 

1. Samples are diluted to 6 mg/mL in PBS and 600 μL (equivalent to 3.6 mg protein 

or approx 60 μL plasma) are applied to the column (see Notes 4, 5 and 6). 

2. The POROS column is mounted in a cartridge holder supplied by the manufacturer 

with ports for insertion of standard HPLC fittings, if performing the following 

steps robotically, or for a needle port adapter at the top and a short piece of PEEK 

tubing at the base to help direct the flow-through when performing the steps 

manually. The blunt-ended needle attached to a 10-mL syringe is inserted firmly 

into the needle port adapter for manual application of the sample and buffers 

to the column, which should be applied at a flow rate of one drop per second 

throughout for manual chromatography. The flow rates for depletions performed 

on an HPLC system are given at the appropriate points in the text. 

3. The column is equilibrated by applying 10 column volumes (20 mL) of PBS at a 

flow rate of 2.4 mL/min. 

4. The diluted plasma sample is applied to the column at a flow rate of 1.2 mL/min 

and the flow-through (containing plasma proteins minus albumin) is collected as 

500-μL fractions into 1.5-mL eppendorf tubes. 

5. A further 10 column volumes (20 mL) PBS are applied to the column to ensure that 

all non-specifically bound proteins are flushed through, with continued collection 

of fractions. Collection of a total of ten fractions is generally sufficient to capture 

all of the nonbound plasma proteins. 

6. Albumin is eluted from the column by the application of 5 column volumes (10 

mL) of 12 mM HCl at a flow rate of 2.4 mL/min, with the first five 1-mL fractions 

collected into 1.5-mL eppendorf tubes. 

7. The column is cleaned by applying 10 column volumes (20 mL) of HSA column 

wash buffer (1 M NaCl) at a flow rate of 2.4 mL/min (see Note7). 

8. If further samples are to be processed, the column may be equilibrated with PBS 

as in point 3. If the column is to be stored for 1–2 d, flushing through with 10 

column volumes (20 mL) PBS and storage at 4°C will be sufficient. If it is to be 

stored for a longer period, the PBS should contain 0.05% sodium azide. 

3.3. Assessment of Fractions Containing Albumin-Depleted Plasma 

Proteins by 1D Gel Electrophoresis (see Note 8) 

1. Aliquots of 10 μL of the flow-through fractions are mixed with 4× Laemmli 

sample buffer, boiled and loaded onto SDS-PAGE (sodium dodecyl sulphate 

polyacrylamide gel electrophoresis) minigels. 

2. Similarly, 3–6 μL of the eluted HSA fractions, and 2 μL of the dilute undepleted 

plasma sample, are prepared for gel analysis.


Fig. 1. Depletion of albumin from human plasma. Coomassie blue (A) and silver (B) 

stained 1D gel of undepleted plasma (P), aliquots of the flow-through fractions 1–10 

and eluted HSA (E1). The undepleted plasma appears to contain a relatively complex 

mixture of proteins that is dominated by the albumin band at 66 kDa. The depleted 

fractions should contain the same range of proteins but the albumin band should be 

completely absent. In contrast, the fraction representing eluted albumin should contain 

no other protein bands, even at the level of sensitivity of silver stain. The multiple 

bands visible below the major HSA band were determined by mass spectrometry to 

be breakdown products, presumably because of the acid conditions under which the 

albumin was eluted from the column. (C) UV trace of albumin depletion performed by 

HPLC showing that fractions 3–7 contain the protein flow-through from the anti-HSA 

column whereas fractions 17 and 18 contain the eluted HSA (E1) itself. 

3. The samples are loaded onto 10% SDS-PAGE minigels and the gels stained with 

colloidal Coomassie Blue (Fig. 1A) or silver stain (Fig. 1B) after electrophoresis. 

3.4. Assessment of Fractions Containing Albumin-Depleted Plasma 

Proteins by UV Absorbance 

1. Once the depletion method has been optimized, the flow-through fractions may 

be assessed by spectrophotometry. If samples have been depleted using an 

HPLC system, a real-time UV trace of the chromatographic separation is usually 

available. Fig. 1 is an example of such a trace.


2. If performing the depletions manually, the fractions may be assessed using an 

off-line spectrophotometer. Aliquots of 100 μL of each fraction are placed in 

mini-cuvets or wells of a 96-well plate, and the absorbance at 280 nm recorded. 

3. All flow-through fractions producing an absorbance value are pooled for the next 

step in the protocol, the depletion of immunoglobulin (see Note 9). 

3.5. Immunoglobulin Depletion 

1. The 0.2-mL protein G POROS column is supplied with a smaller cartridge holder 

and should be mounted as described earlier. The blunt-ended needle attached to a 

2-mL syringe is used for manual application of sample and buffers to the column. 

As for the HSA depletion column, all reagents are applied to the column at a 

flow rate of 1 drop per second when the chromatography is performed manually. 

Flow rates for depletion using an HPLC system are noted at appropriate points 

in the text. 

2. The column is equilibrated by applying 10 column volumes (2 mL) of PBS at a 

flow rate of 1mL/min. 

3. The albumin-depleted plasma sample cannot be applied as a single aliquot but 

should be split into two aliquots of 600 μL (see Note 10), each being loaded 

at a flow rate of 0.5 mL/min. The flow-through (containing plasma proteins 

minus albumin and immunoglobulin) is collected as 500-μL fractions into 1.5-mL 

eppendorf tubes. 

4. A further 10 column volumes (2 mL) PBS are applied to the column at a flow rate 

of 1 mL/min to ensure that all nonspecifically bound proteins are flushed through, 

with continued collection of fractions. Collection of a total of six fractions is 

generally sufficient to capture all the nonbound plasma proteins. 

5. Immunoglobulin is eluted from the column by the application of 10 column 

volumes (2 mL) of 12 mM HCl at a flow rate of 1 mL/min, with two 1-mL 

fractions being collected into 1.5-mL eppendorf tubes. 

6. After the first aliquot has been depleted of immunoglobulin, the column is reequilibrated 

with PBS as in point 2, and the second aliquot processed as in 

steps 3–5. 

7. After the second aliquot has been processed, 10 column volumes (2 mL) of protein 

G column wash buffer (1 M NaCl containing 10% acetic acid) are applied to the 

column at a flow rate of 1 mL/min (see Note 11). As for the anti-HSA column, 

the protein G column may be stored for 1–2 days with PBS but for longer periods, 

the PBS should contain 0.05% sodium azide. 

3.6. Assessment of Fractions Containing Albuminand 

Immunoglobulin-Depleted Plasma Proteins by 1D Gel 

Electrophoresis (see Note 8) 

1. Aliquots of 10 μL of the flow-through fractions are mixed with 4× Laemmli 

sample buffer, boiled and loaded onto SDS-PAGE minigels.


Fig. 2. Depletion of immunoglobulin from albumin-depleted human plasma. 

Coomassie blue (A) and silver (B) stained 1D gel of undepleted plasma (P), 

HSA-depleted plasma (H), aliquots of the flow-through fractions 1–6 and eluted 

immunoglobulin (E2). The depletion of the immunoglobulin bands is more difficult to 

discern than the depletion of the very abundant albumin band shown in figure 1, but 

the eluted protein should be clearly seen as two bands representing the heavy and light 

chains of the immunoglobulin (Ig H and Ig L). (C) UV trace of immunoglobulin depletion 

performed by HPLC showing that fractions 1 and 2 contain the protein flow-through 

from the protein G column whereas fraction 7 contains the eluted immunoglobulin. 

2. Similarly, 3–6 μL of the eluted immunoglobulin fractions, 2 μL of the dilute 

undepleted plasma, and 10 μL of the albumin-depleted plasma are prepared for 

gel analysis. 

3. The samples are electrophoresed on 10% SDS-PAGE minigels and the gels stained 

with colloidal Coomassie Blue or silver stain (Fig. 2). 

3.7. Assessment of Fractions Containing Albumin- and 

Immunoglobulin-Depleted Plasma Proteins by UV Absorbance 

1. Once the depletion method has been optimized, the flow-through fractions may 

be assessed by simple spectrophotometry by measuring their absorbance in-line


at 214 nm using the UV detector on the HPLC system, or off-line at 280 nm in 

a cuvet or microplate format. 

2. All flow-through fractions producing an absorbance value are pooled before 

processing for 2D gel electrophoresis or other proteomic analysis (see Note 12). 

3.8. TCA Precipitation of Depleted Protein Samples 

for 2D Gel Electrophoresis 

1. The depleted samples should be maintained at 4 ° C or on ice throughout this 

procedure. 

2. A solution of 20% TCA in water is prepared just before use and placed on ice to 

chill (see Note 13). 

3. 2 mL 20% TCA are added to 2 mL of the dilute depleted plasma, and the sample 

is mixed gently and incubated on ice for 30 min. 

4. The sample is then divided equally between three 1.5-mL eppendorf tubes and 

centrifuged at 14,000g and 4 ° C for 10 min to pellet the proteins. 

Fig. 3. 2D gel images of undepleted (A) and depleted (B) plasma. The plasma 

proteins were focussed on nonlinear pH 3–10 IPG strips before 2nd dimension 

separation on 12% SDS PAGE gels. The gels were stained with colloidal Coomassie 

blue. Image analysis of the gels shows that the albumin and immunoglobulin heavy 

chain are 98% and 80% depleted, respectively. The resolution of the protein spots is 

improved significantly by depletion, particularly for proteins migrating in the top half 

of the gel close to the migration position of albumin. Indeed, peptide mass fingerprinting 

of the visible features reveals that hemopexin comigrates with albumin, yet it 

is undetectable on gels of the undepleted sample. There is an overall increase in the 

number of detectable spots of approx 50% when the gels are stained with Coomassie 

blue, indicating that the detection of lower abundance proteins has been improved.


5. The supernatant is discarded, and the pellets washed with 2×1mLofice-cold 

acetone to remove all traces of the TCA. 

6. The pellets are air-dried briefly (approx 5 min) before resuspension in 12 μL 

resuspension buffer (see Note 14). 

7. The sample is heated at 95 ° C for 10 min to aid solubilisation followed by the 

addition of 350 μL IPG-strip rehydration buffer. 

8. The sample is clarified by a high speed centrifugation step before conventional 

2D electrophoresis (Fig. 3). 

4. Notes 

1. To make meaningful comparisons between plasma samples, it is vital that the 

collection and storage is consistent. Plasma is rich in proteases and inappropriate 

or overlong storage can lead to significant changes in the protein profile. 

Whatever procedure is chosen for sample preparation, it must be strictly 

adhered to. 

2. Serum and plasma are almost identical in terms of the protein profile observed 

on 2D gels. However, some of the proteins involved in the clotting process, 

such as fibrinogen, are removed when the clotted red blood cells are separated 

from the serum by centrifugation. There is also a slightly increased tendency 

for lysis of the red blood cells when preparing serum. It is probably easier and 

more consistent to prepare plasma. 

3. The anti-HSA column may be used to deplete albumin from the plasma or serum 

of various animal species, but lower levels of total protein must be loaded onto 

the column. We have depleted albumin from mouse and rat serum after loading 

1–2 mg total protein. The protein G column works as effectively for animal 

immunoglobulins as for human. 

4. Most human plasma samples have a protein concentration of 60–70 mg/mL with 

approximately half of that comprised of albumin. The binding capacity of a 2 

mL anti-HSA POROS cartridge is 1.8–2 mg albumin so the samples will require 

a dilution step before loading onto the column. 

5. Both the anti-HSA and the protein G POROS columns may be used for the 

depletion of multiple samples without loss of effectiveness, as long as they are 

cleaned and stored correctly. The smaller protein G column does have a tendency 

to deteriorate before the larger anti-HSA column, but we have successfully 

processed at least 50 plasma samples through one pair of columns. 

6. Manual depletions are just as effective as those performed on HPLC systems, 

but they are rather tedious. A cheap but effective alternative is to use a syringe 

pump to apply samples and buffers. 

7. The anti-HSA column should be cleaned with wash buffer (1 M NaCl) after 

every depletion to prevent residual protein build-up that would lead to decreased 

efficiency of albumin removal and increased pressure during the chromatography 

steps.


8. Measuring the absorbance at 214/280 nm provides a measure of the total amount 

of protein in each fraction, but it does not indicate how much of the target 

protein has been removed (efficiency of depletion), nor whether other proteins 

have been depleted as well (specificity of depletion). A visual assessment of 

these factors can be rapidly achieved by 1D gel electrophoresis. However, once 

the protocol has been optimized, it should be necessary to perform this sort of 

quality control only if the UV trace looks anomalous, or when a new POROS 

cartridge is being employed. 

9. At this stage, it is usual to have a total volume of approx 1.5 mL of albumindepleted 

plasma containing roughly 900 μg of protein. Because the amount of 

protein loaded onto the column was approx 3.6 mg, the reduction in total protein 

content is equivalent to 75% suggesting that the depletion of albumin is complete. 

10. Immunoglobulin comprises 8–26% of the total protein in plasma and is 

therefore present at concentrations of 5–18 mg/mL. The sample has already 

been diluted at least 1:10 during the albumin depletion, so the concentration is 

now approx 0.5–1.8 mg/mL. The binding capacity of the protein G cartridge is 

up to 1.8 mg immunoglobulin, so the maximum volume of the pooled fractions 

containing HSA-depleted plasma that could be loaded is 1 mL. These fractions 

may be stored at 4 ° C for short periods of time before the immunoglobulin 

depletion, but should be processed within 8h. 

11. The protein G column should be cleaned after 2 immunoglobulin depletions 

to keep it functioning at the highest efficiency. After the first depletion, the 

column is simply re-equilibrated with PBS as in Section 2.3.5 point 1, but after 

the second, it should be exposed to wash buffer (1 M NaCl/10% acetic acid). 

12. At this stage, it is usual to have a total volume of approx 2 mL of depleted plasma 

containing roughly 500 μg protein. This is equivalent to a further 11% reduction 

in protein content of the plasma, which is approximately what would be expected 

following removal of the immunoglobulin. However, this is too dilute for most 

proteomic analyses so a method to concentrate the proteins must be employed. 

13. There are several reagents for precipitating proteins from dilute samples, 

including acetone, methanol and acetonitrile, but the method that seems to 

work most effectively for the samples described here is TCA precipitation. It 

is essential that the TCA is prepared immediately before use: even storage for 

2–3 h reduces the efficiency of precipitation. The sample may be precipitated 

with TCA for longer than 30 min, but there is a risk of protein degradation 

or modification on prolonged exposure. Alternative methods for protein 

concentration include molecular weight cut-off filters, for which a significant 

loss of total protein is a factor, or the use of reversed phase matrices to capture 

the proteins, for which the elution buffer must be very carefully selected for its 

compatibility with the 1st dimension separation. 

14. The volume of resuspension buffer (5% SDS, 1.15% DTT) to be added must 

be calculated carefully to ensure that after mixing with IPG-strip rehydration 

buffer, the level of SDS is within tolerable levels for the 1st dimension


separation, i.e., less than 0.25%. When the sample described here is mixed 

with 350-μL IPG-strip rehydration buffer, the final concentration of SDS in the 

buffer is 0.17%, well within the tolerance of the 1st dimension separation. 


This work was supported by the Wellcome Trust. Thanks to Rod Watson of 

Applied Biosystems for assistance with the establishment of protocols. 

References 

1. Anderson, N. L., and Anderson, N. G. (2002) The human plasma proteome: history, 

character, and diagnostic prospects. Mol. Cell Proteomics 1, 845–67. 

2. Cohn, E. J. (1941) The properties and functions of the plasma proteins with a 

consideration of the methods for their separation and purification Chem. Rev. 28, 

395. 

3. Simoni, R. D., Hill, R. L., and Vaughan, M. (2002) The beginning of protein 

physical chemistry. Determinations of protein molecular weights. The work of 

Edwin Joseph Cohn. J. Biol. Chem. 277, 19e. 

4. Georgiou, H. M., Rice, G. E., and Baker, M. S. (2001) Proteomic analysis of 

human plasma: Failure of centrifugal ultrafiltration to remove albumin and other 

high molecular weight proteins Proteomics 1, 1503–06. 

5. Tirumalai, R. S., Chan, K. C., Prieto, D. A., Issaq, H. J., Conrads, T. P., and 

Veenstra, T. D. (2003) Characterization of the low molecular weight human serum 

proteome Mol. Cell Proteomics 13, 13. 

6. Akerstrom, B., Brodin, T., Reis, K., and Bjorck, L. (1985) Protein G: a powerful 

tool for binding and detection of monoclonal and polyclonal antibodies J. Immunol. 

135, 2589–92. 

7. Forsgren, A., and Sjoquist, J. (1966) “Protein A” from S. aureus. I. Pseudo-immune 

reaction with human gamma-globulin J. Immunol.97, 822–7. 

8. Housden, N. G., Harrison, S., Roberts, S. E., Beckingham, J. A., Graille, M., Stura, 

E., and Gore, M. G. (2003) Immunoglobulin-binding domains: Protein L from 

Peptostreptococcus magnus Biochem. Soc. Trans. 31, 716–8. 

9. Kabir, S. (2002) Immunoglobulin purification by affinity chromatography using 

protein A mimetic ligands prepared by combinatorial chemical synthesis Immunol. 

Invest. 31, 263–78. 

10. Svensson, H. G., Hoogenboom, H. R., and Sjobring, U. (1998) Protein LA, a novel 

hybrid protein with unique single-chain Fv antibody- and Fab-binding properties 

Eur. J. Biochem. 258, 890–6. 

11. Wilchek, M., Miron, T., and Kohn, J. (1984) Affinity chromatography Methods 

Enzymol. 104, 3–55. 

12. Pieper, R., Su, Q., Gatlin, C. L., Huang, S. T., Anderson, N. L., and 

Steiner, S. (2003) Multi-component immunoaffinity subtraction chromatography: 

An innovative step towards a comprehensive survey of the human plasma proteome 

Proteomics 3, 422–32.

3 

Multi-Component Immunoaffinity Subtraction 

and Reversed-Phase Chromatography of Human Serum 

James Martosella and Nina Zolotarjova 

Summary 

Serum analysis represents an extreme challenge because of the dynamic range of 

the proteins of interest, and the high structural complexity of the constituent proteins. 

High-abundant proteins such as albumin, IgG, transferrin, haptoglobin, IgA and alpha1anti-trypsin 

represent up to 85% of the total protein mass in serum (Fig. 1). These 

major protein constituents interfere with identification and characterization of important 

moderate- and low-abundant proteins by limiting the dynamic range of mass spectral 

and electrophoretic analysis. During protein isolation, separation, and analysis, these six 

proteins often mask the detection of the more important low-abundant proteins that are 

of high interest as biomarkers of disease or drug targets. In one- and two-dimensional 

gel electrophoresis (1DGE and 2DGE) for example, the spots or bands because of these 

six highly abundant proteins, as well as their fragments, often overlap or completely 

mask large regions of the gel, making detection of the myriad low-abundant proteins very 

difficult, if not impossible. Moreover, proteomic analysis methods commonly include an 

electrophoretic or chromatographic separation step which, of course, has a finite mass 

loading tolerance. The presence of a large quantity of high-abundant proteins limits the 

mass load of targeted proteins that can be initially sampled by these separation methods 

and thus requires the need for multidimensional separation techniques to reduce sample 

complexity. 

Herein we describe immunoaffinity depletion combined with reversed-phase separation 

modes to reduce the sample complexity of human serum. We selectively immunodepleted 

six of the most abundant proteins from human serum, then employed gradient elution 

reversed-phase (RP) HPLC to fractionate the remaining serum proteins. The workflow 

shown in (Fig. 2) was optimized to process immunodepleted flow-through serum samples 

directly to a RP column with minimal sample handling. The RP operational conditions 

permitted robust and repeatable separations and have been optimized specifically for 

immunodepleted serum samples. 



27

28 Martosella and Zolotarjova 

Key Words: affinity chromatography; human plasma; human serum; HPLC; immunodepletion; 

prefractionation; proteomics; reversed-phase chromatography. 

Immunoglobulin G 

16.6% 

Albumin 

54.3% 

Alpha-1-antitrypsin 3.8% 

Immunoglobulin A 3.4% 

Transferrin 3.3% 

Haptoglobin 2.9% 

Other 15% 

Fig. 1. Composition of proteins in human serum. The protein composition is 

schematically depicted based on mass abundance in normal human serum. The six 

high-abundant proteins removed by the immunoaffinity column comprise approx 85% 

of the total protein mass in human serum. 

Human Serum or Plasma 

Immunodepletion of Six 

High-abundant Proteins 

High Temp. RP-HPLC 

Multiple RP HPLC 

Fraction Collection 

2D-PAGE 

LC/MS/MS 

Sample Denaturation 

3-Step Multi-Segment Gradient 

Fig. 2. Multidimensional chromatographic workflow.

Multi-Component Immunoaffinity Subtraction and Reversed-Phase 29 


Interest in proteomic analysis of human serum has been greatly elevated during 

the past several years as liquid chromatography (LC) and mass spectrometry (MS) 

methodologies have evolved sufficiently to investigate this challenging sample. 

The popularization of multidimensional LC methods, and the ever-improving 

sensitivity and performance of multi-stage MS instruments, combined with highspeed 

database searching, is permitting complex protein samples to undergo 

analysis by identification of constituent tryptic peptide fragments. Proteomic 

analysis of human serum represents an extreme challenge because of the dynamic 

rangeoftheproteinsofinterest.Serum(plasma)containsmanyproteins,estimated 

to include almost 500,000 molecular species and has more than 10 orders of 

magnitude concentration range, from mg/mL to pg/mL, and possibly less (1). 

This wide range of analytical target molecules is currently outside the realm of 

the dynamic range of available technologies for proteomic analyses. A means to 

address the complexity of these samples is the application of multidimensional 

separation techniques (2,3), for example, by multidimensional LC fractionation. 

WedescribeanimmunoaffinityandRPLCcolumnapproach,whichreducesthe 

complexity of serum or plasma. Samples are first immunodepleted of the six most 

abundant proteins using an immunoaffinity LC method. This separation delivers 

a flow-through fraction containing low-abundant proteins, whereas the bound 

high-abundant proteins are left behind. Second, the immunodepleted samples are 

separated under a set of optimized reversed-phase (RP) conditions using a highrecovery 

macroporous column material. The conditions and protocol have been 

designed specifically for immunodepleted human serum or plasma samples. The 

combination of sample simplification by immunoaffinity depletion, combined 

with robust and high recovery RP-HPLC fractionation, yields samples permitting 

higher quality protein identifications (4). The approach presented here enables 

an expanded dynamic range for the detection of low-abundant proteins in the 

complex proteomic samples and thereby assists in the search for novel biomarkers 

of disease states and intervention strategies. 

2. Materials 

2.1. Immunoaffinity Depletion of High-Abundant Proteins 

2.1.1. Serum Collection 

1. Becton Dickinson Vacutainer tubes (VWR International) with SST gel and BD 

clot activator. 

2.1.2. Immunoaffinity Chromatography 

1. An 1100 liquid chromatograph (Agilent Technologies, Wilmington, DE.) 

consisting of a binary gradient pumping system, a thermostatted autosampler,


a variable wavelength absorbance detector, a temperature controlled column 

compartment, and a thermostatted automated analytical scale fraction collector 

(see Note 1). 

2. Agilent Chemstation Software version B.01.03. 

3. Multiple Affinity Removal System column, 4.6 mm id × 100 mm, Immunodepletion 

Buffer A and Buffer B (Agilent Technologies, Wilmington, DE). 

4. 0.22-μm spin filters. 

2.2. Immunodepleted Serum and Bound Fraction Processing 

1. BCA Assay Kit (Pierce, Rockford, IL). 

2. Tris-glycine precast gels (4–20% acrylamide, 10 wells, 1 mm), sample preparation 

(loading) and running buffers, Mark12 unstained standards (Invitrogen, 

Carlsbad, CA). 

3. 4-mL spin concentrators with 5 kDa molecular weight cutoffs (Agilent 

Technologies, Wilmington, DE). 

4. Standard equipment for two-dimensional electrophoresis (Bio-Rad, Hercules, CA). 

5. Rehydration buffer: 8 M urea, 2% CHAPS, 2% ampholytes and 20 mM dithiothreitol. 

6. 11 cm immobilized pH gradient (IPG), pH 3–10 nonlinear strips (Bio-Rad, 

Hercules, CA). 

7. 8–16% precast Tris-glycine gels for second dimension separation (Bio-Rad). 

8. Gel Code Blue – Coomassie stain (Pierce, Rockford, IL). 

2.3. Reversed-phase Separation of Low-abundant Proteins 

1. An 1100 liquid chromatograph (Agilent Technologies, Wilmington, DE.) 

consisting of a binary gradient pumping system equipped with a 900 μL capillary 

injector loop, a thermostated autosampler, a variable wavelength absorbance 

detector, a temperature controlled column compartment, and a thermostated 

automated analytical scale fraction collector (see Note 1). 

2. 4.6 mm ID × 50 mm macroporous high recovery reversed-phase C18 column (mRP- 

C18)(AgilentTechnologies,Wilmington,DE).Particlecompositionisasilica-based 

macroparticulate material with a particle size of 5.0 μm. The column hardware is 

made of PEEK composition and the frits are 2.0 μm PEEK encapsulated. 

3. Methods 

3.1. Immunoaffinity Depletion of High-Abundant Proteins 

3.1.1. Serum Collection 

1. Collect serum samples into a Becton Dickinson Vacutainer tubes (VWR International) 

with SST gel and BD clot activator. After clot formation, centrifuge 

sample at 1000g for 15 min. Remove serum and store aliquotes at –80°C. Total 

time for serum processing is less than 60 min.


3.1.2. Immunoaffinity Chromatography 

The immunoaffinity column depletion technology offers rapid and simultaneous 

removal of six high-abundant proteins in 28 min per sample (Fig. 3). 

The column is based on the rabbit polyclonal antibodies to six major 

serum proteins—human serum albumin (HSA), transferrin, alpha1-anti-trypsin, 

haptoglobin, immunoglobulin A (IgA), and immunoglobulin G (IgG) that were 

affinity purified on corresponding protein antigen columns (4). The resulting 

affinity-purified antibodies were covalently coupled to porous beads via their 

Fc region and cross-linked. Spatially controlled cross-linking of the antibodies 

resulted in preferential orientation of the antibody binding sites away from the 

solid-phase surface, supporting maximum binding capacity of targeted proteins. 

Through the series of column loading, washing, collection, and reequilibration 

steps, multiple serum samples can be processed and depleted of 

targeted high-abundant proteins. After each pass of serum through the column, 

flow-through fractions containing low-abundant proteins can be pooled and 

concentrated for downstream postaffinity processing. The methods outlined 

below demonstrate a typical workflow for immunodepleting a given quantity 

of human serum samples. Similar procedures have been employed successfully 

for cerebrospinal fluid, amniotic fluid and for urine analysis. 

This methodology is robust, scalable, and easily automated for multiple 

samples processing, as well as compatible with downstream 1D and 2D-SDS- 

PAGE, LC, LC/MS, and/or enzymatic or chemical fragmentation methods. 

1. Purge LC system with Buffer A and Buffer B at a flow rate of 1.0 mL/min for 

10 min. without column (see Note 2). 

Serum 

Flow-through 

Fraction 

Injection Elution 

Bound Fraction 

Re-equilibration 

Fig. 3. Chromatogram for the affinity removal of high-abundant proteins from 

human serum. 35 μL of serum was diluted 5× and injected on a 4.6 mm id × 100 

mm immunoaffinity column (0.50 mL/min) and a flow-through peak (3–5.0 min) was 

collected for reversed-phase HPLC fractionation. The column was washed with Buffer 

A and the targeted high-abundant proteins were eluted with Buffer B.


Table 1 

LC timetable 

Time (mins) %B Flow rate Max. pressure 

1 0.00 000 0.500 120 

2 10.00 000 0.500 120 

3 10.01 10000 1.000 120 

4 17.00 10000 1.000 120 

5 17.01 000 1.000 120 

6 28.00 000 1.000 120 

2. Run two method blanks by injecting 200 μL of Buffer A without the column 

according to the LC timetable (Table 1) (see Note 3). Table 1 here 

3. Attach a 4.6 × 100 mm immunoaffinity column and equilibrate in Buffer A for 4 

min at a flow rate of 1 mL/min. 

4. Dilute human serum five times with Buffer A (for example 35 μL of human 

serum with 140 μL of Buffer A (see Note 4). 

5. Remove sample particulates with a 0.22 μm spin filter and centrifuge sample for 

1.0 min at 16,000g. 

6. Inject 175 μL of the diluted serum at a flow rate of 0.5 mL/min and run LC 

timetable. 

7. Collect the flow-through fraction (appears between 3.0–5.0 min; see Fig. 3 for the 

chromatogram) into 1.5-mL microcentrifuge tubes at 4°C. Store collected fraction 

at –20°C if not analyzed immediately. 

8. Wash column with buffer A (see LC timetable) and elute the bound fraction with 

Buffer B at a flow rate of 1.0 mL/min for 7.0 min. 

9. Regenerate the column by equilibrating it with Buffer A for 11.0 min at a flow 

rate of 1.0 mL/min for a total run cycle of 28.0 min. (see Note 5). 

3.2. Immunodepleted Serum and Bound Fraction Processing 

For analyses of immunodepleted serum before RP separation or without 

further RP fractionation proceed with the recommendations below. For direct 

processing of the flow-through for RP HPLC fractionation of the immunodepleted 

serum see Section 3.3. 

1. If lyophylization of the immunodepleted serum is desired, buffer exchange to 

a volatile buffer (for example, ammonium bicarbonate) because of high salt 

concentration in Buffer A. 

2. Measure protein concentration in serum, flow-through and bound fractions using 

BCA protein assay. For 1D-SDS-PAGE analysis of the flow-through fraction 

(immunodepleted serum) or bound fraction, mix sample aliquots (5–10 μg of 

protein) with the equal volume of the loading buffer, boil sample for 3 min. and


200.0 

116.3 

97.4 

66.3 

55.4 

36.5 

31.0 

21.5 

14.4 

6.0 

3.5 

1 2 3 4 5 

Fig. 4. 1D SDS gel electrophoresis of human serum protein fractions from an 

immunoaffinity column. An equal amount (9 μg) of crude serum (Lane 2), flow-through 

(Lane 3) and bound fractions (Lane 4) were separated on 4–20% SDS-PAGE gel under 

nonreducing conditions. Lanes 1 and 5 are the molecular weight standards (Mark12) 

from Invitrogen. The proteins were stained with Coomassie Blue dye. Based on the 

protein assay of the flow-through fraction, 85% of total protein was removed from the 

crude serum. 

load on the gel. Fig. 4 shows 1D gel electrophoresis data for crude serum, flowthrough 

and bound fractions from an immunoaffinity column. Equal amounts 

of protein (9 μg) were loaded in each lane. Results show that high-abundant 

proteins in serum (Lane 2) are clearly removed and are not visible in the flowthrough 

fraction (Lane 3). Also, low-abundant proteins that were not visible in 

the serum before depletion (Lane 2) became visible in the flow-through fraction 

after removal of the high-abundant proteins. 

3. For IEF, 2D-SDS-PAGE, and MS analysis, it is necessary to buffer 

exchange/desalt and concentrate fractions to an appropriate buffer. Use 4 mL 

spin concentrators with 5 kDa molecular weight cutoffs. Centrifuge samples at 

7,500g for 20 min at 4°C. Buffer exchange into 20 mM Tris-HCl, pH 7.4, by 

3 rounds of buffer addition, with 20 min centrifugation for each round. Aliquot 

the concentrated samples and store at –70°C until the analysis. Analyze protein 

concentration using a BCA protein assay. 

4. Prepare 2D electrophoresis samples by mixing 250 μg of proteins with 185 

μL of rehydration buffer containing 8 M urea, 2% CHAPS, 2% ampholytes, 

pH 3–10, and 20 mM dithiothreitol. Apply samples on 11-cm immobilized 

pH gradient (IPG), pH 3–10 nonlinear strips and process them according to


the manufacturer’s instructions. Perform the second dimension separation on 

8–16% precast Tris-glycine gels. Visualize proteins by Coomassie Blue staining. 

Fig. 5 shows the protein pattern of human serum before (Panel A) and after 

immunodepletion (Panel B). Circles indicate the areas where the targeted highabundant 

proteins reside. The depletion of high-abundant proteins unmasks the 

low-abundant proteins because of the substantial removal of protein mass from 

the sample. More than 85% of total protein was depleted after a single pass 

of serum through the immunoaffinity column. This enabled a large increase in 

low-abundant protein mass loading onto the gel (up to 10 times). As a result, lowabundant 

protein fractions become enriched and more easily detectable on the gel, 

making the protein spots more amenable to quantitation and MS identification. 

The immunoaffinity column is highly specific for the removal of highabundant 

proteins from serum (5). A small number of nontargeted proteins are 

bound to the column. None of the nonspecific proteins are bound quantitatively 

to the immunoaffinity column and represent only a small percentage of the 

total flow-through. 

Anti-trypsin IgA 

Transferrin 

IgG Heavy 

Albumin 

Chain MW 

(kDa) 

200 

116 

97 

66 

55 

Haptoglobin 

A, Crude Serum B, Serum after immunodepletion 

Ig Light 

Chain 

37 

31 

21 

14 

6 

pH 3 –10 

MW 

(kDa) 

200 

116 

97 

66 

55 

Fig. 5. 2D gel electrophoresis of human serum before and after removal of highabundant 

proteins. Panel A. Human serum before depletion. The targeted high abundant 

proteins are circled. Panel B. Human serum after depletion of the six targeted high 

abundant proteins. The positions of the removed proteins are circled. 250 μg of total 

protein loaded on each gel. Molecular weight standards–Mark12 (Invitrogen). Proteins 

were visualized by staining with Coomassie Blue. 

37 

31 

21 

14 

6


3.3. Reversed-phase Separation of Low-Abundant Proteins 

3.3.1. Reversed-Phase Chromatographic Conditions 

1. Eluent A: 0.1% TFA in water, Eluent B: 0.08% TFA in acetonitrile. 

2. Autosampler and fraction collection temperature: 4°C. 

3. Column temperature: 80°C. If column oven cannot reach or maintain 80°C, a 

lower temperature can be used, however, protein recovery and chromatographic 

resolution may be increasingly compromised with decreasing temperature. 

4. UV absorbance: 280 nm (preferred) or 210 nm. 

5. Sample flow rate: 0.75 mL/min. 

6. Prepare LC gradient elution according to LC Timetable in Table 2. 

7. Prepare flow-through sample directly for Reversed-Phase chromatographic 

separation (see Note 6). Allow sample to equilibrate at room temperature for at 

least 30 min. 

8. Inject up to 900 μL of the denatured (6M urea/1.0% acetic acid) flow-through 

proteins from immunodepletion. If it is desirable to process the entire flowthrough 

sample volume from each immunodepletion run (for example 1.5 mL 

of denaturated flow-through) or load multiple flow-throughs, an injector loading 

program is needed. This can also be useful for utilizing the columns full loading 

capacity when flow-throughs from multiple immunodepletions have been pooled 

(see Note 7). Fig. 6 is representative of a RP elution profile of immunodepleted 

human serum obtained when using the optimized multi-segmented elution conditions 

presented in Table 2. The RP separation can be fractionated and the collected 

fractions processed for downstream electrophoretic or LC/MS analyses (6). 

9. Collect fractions at 1.0 min time intervals from 0–54 min. At the recommended 

flow of 0.75 mL/min., the fraction volumes will be 0.75 mL (see Note 8). 

10. Dry each fraction in a centrifugal vacuum concentrator. To avoid protein degradation 

do not dry overnight. If sample processing is not immediate, store dried 

fractions at –80°C. 

11. For SDS-PAGE analysis of the separation efficiency (an example of the results 

produced is shown in Fig. 7), dry fractions in the vacuum concentrator and 

Table 2 

LC timetable 

Time (mins) %B 

1 0.00 3.00 

2 6.00 30.0 

3 39.0 55.0 

4 49.0 100.0 

5 53.0 100.0 

6 58.00 3.00 

7 postrun re-equilibration 68.00 3.00


Absorbance @ 280 nm (mAu) 

mAU 

35 

30 

25 

20 

15 

10 

5 

0 

0 5 10 15 20 25 

Time (min.) 

30 35 40 45 

Fig. 6. Representative RP-HPLC elution profile (absorbance at 280 nm) for human 

serum depleted of high-abundant proteins. An aqueous TFA and ACN (TFA) gradient 

was used at 80°C at a flowrate of 0.75 mL/min on a 4.6 mm id × 50mm mRP-C18 

column. The sample comprised a total of 270 μg protein in 6 M urea/1% AcOH. 

then dissolve each with electrophoresis sample preparation (loading) buffer as 

recommended below (see Note 9). 

12. For 2D PAGE analysis of the RP fractions, we suggest combining the fractions in 

a manner suitable to accommodate specific workflows and goals. Fraction pooling 

will be required to achieve appropriate protein loads for 2D PAGE analysis. To 

process the RP fractions, dry in a vacuum concentrator and dilute with the IEF 

buffer according to the manufacturers protocol. 

13. For consistent and repeatable reversed-phase separation consult Note 10. 

4. Notes 

1. Conventional autosamplers generally do not provide optimum sampling because 

of conditions which can lead to extra column band broadening and mixing, 

thereby reducing resolution. 

2. The immunoaffinity column requires a proprietary two buffer system (Buffer 

A and Buffer B) for operation. The two buffers provide the means to separate 

min


Fractions 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 

kDa 

200.0 

116.3 

97.4 

66.3 

55.4 

36.5 

31.0 

21.5 

14.4 

Fractions 27 28 29 30 31 32 33 34 35 36 

kDa 

200.0 

116.3 

97.4 

66.3 

55.4 

36.5 

31.0 

21.5 

14.4 

Fig. 7. SDS-PAGE analysis of RP-HPLC fractionated immunodepleted human 

serum from an mRP-C18 column (4.6 mm id × 50 mm). A depleted human serum 

sample was injected onto the column and eluted by a multi-segment gradient. Fifty-four 

fractions were collected (29 showing the majority of protein elution) for analysis by 

4–20% SDS-PAGE. 

low-abundant proteins from high-abundant proteins and regenerate the column, 

all in about 20–30 min per injection. Buffers A and B are optimized to minimize 

co-adsorption of nontargeted proteins to the column packing, and to ensure 

reproducibility of column performance and long column lifetime. Buffer A is 

a salt-containing neutral buffer (pH 7.4) used for loading, washing, and reequilibrating 

the column. Buffer B is a low pH urea buffer used for eluting 

the bound high-abundant proteins from the column. Serum samples are injected 

onto the column and the high-abundant proteins are simultaneously removed as 

low-abundant proteins pass through in the flow-through fraction. After collecting 

the low-abundant proteins and washing the column, the bound proteins are 

eluted with Buffer B and the column is re-equilibrated with Buffer A. Do not 

expose immunoaffinity column to solvents other than Buffers A and B. Organic 

solvents (acetonitrile, alcohols, etc.), strong oxidizers, acids, reducing agents, 

and other protein denaturing agents will cause irreversible column damage.


Under optimized operating conditions the column stable and robust for at least 

200 injections of serum samples. 

3. Ensure proper sample loop size in autosampler. 

4. Consult column certificate of analysis to verify column capacity. Concentrations 

of some high-abundant proteins can vary widely depending on the sample 

source. Proteins such as alpha1-antitrypsin, haptoglobin, IgG rise several folds 

in response to stress, infection, inflammation, or tissue necrosis and are known 

as acute phase reactants. Users need to adjust column loading volume accordingly. 

It is not recommended to load crude serum onto the column. Addition 

of protease inhibitors cocktail (Complete, Roche, IN) in buffer A for sample 

dilution helps prevent protein degradation. 

5. When not in use, store the column, with the end-caps sealed, in a refrigerator at 

2–8°C to minimize losses in column capacity. Do not freeze the column. 

6. Flow-through sample preparation: The amount of flow-through sample volume 

from a 4.6 mm id × 100 mm immunodepletion column is approx 1.0–1.5 mL. 

Immunodepleted serum fractions can either be pooled together or processed 

individually. However, by either method, they must first be denatured under 

acidic conditions. To denature before RP separation, 480 mg solid urea and 13 

μL acetic acid (AcOH) are added for every 1.0 mL of flow-through for a final 

sample concentration of approx 6 M urea/1.0% AcOH. Calculation is based on 

an immunodepletion separation of 35 μL human serum (diluted 5×), in which 

BCA protein analysis gave a flow-through protein concentration of 0.38 mg/mL. 

It is recommended to measure the actual flow-through protein concentration 

for each serum sample lot processed and adjust the 6 M urea and 1.0% AcOH 

concentrations accordingly. 

7. For loading sample volumes greater than 900 μL, an isocratic loading method 

is required. To load under isocratic conditions set 97% Eluent A (0.1% TFA in 

water) and Eluent B (0.08% TFA in acetonitrile) at 3.0% for a minimum run time 

of 3.0 min (maintain all other chromatographic conditions from Section 3.3). 

Perform desired amount of column injections without overloading the column. 

The maximum loading capacity for a 4.6 mm id × 50 mm mRP-C18 column 

is approx 400 μg of immunodepleted serum. After multiple sample injections 

have been loaded, proceed with the elution gradient in Table 2. In Chemstation, 

this process can be automated and configured under Sequence and Sequence 

Table. 

8. Depending on the user’s capacity for sampling handling and/or processing objectives, 

some workflows may require the need to collect more or a lesser amount 

of fractions and may therefore need to vary the time-based collection. However, 

fraction collection greater than 2.0 min time intervals will require collection 

tubes larger than 2.0 mL and typically require tray or instrument adjustments for 

many automated fraction collectors. Fractions collected from 1 to 6 min and 37 

to 54 min are not shown in Fig. 7. The majority of protein elution as determined 

by Commassie blue staining occurs from 7 to 36 min. If a comprehensive MS 

analysis is the goal, we recommend fraction collection throughout the entire run.


9. For fractions #1–#12 and #37–#54, dissolve each fraction in 30 μL of 

electrophoresis sample preparation buffer and heat for 3 min at 70°C. If the 

entire protein load on the column was 400 μg or less, load the entire 30 μL 

of prepared sample directly onto the gel. If protein fractions were pooled from 

several chromatographic runs and thus exceeded 400 μg of total column load, 

remove 15 μL of prepared sample and dilute with 15 μL deionized water (30 μL 

total) and load onto the 1D SDS-PAGE. 

For fractions #13–#36, dissolve each fraction in 75 μL of sample preparation 

buffer and heat for 3 min at 70°C. If the entire protein load on the column was 

400 μg or less, directly load 15 μL (without water dilution) onto SDS-PAGE. 

If protein fractions were pooled from several chromatographic runs and thus 

exceeded 400 μg of total column load, remove 7 μl of prepared sample, dilute 

with 7 μL deionized water (14 μL total) and load onto the gel. 

10. Column runs from the same depleted serum sample, during the same run 

progression, should be very repeatable with identical peak shapes and intensities. 

If variances are occurring the column may need replacing. Column life varies 

with use and conditions, but should typically last for over 75 injections. It is 

recommended to complete each RP separation with a minimum postrun time 

of 10.0 min to ensure that the column has re-equilibrated. Periodically perform 

blank injections to evaluate baseline stabilization. If peak ghosting, which is a 

characteristic of protein carryover, is present, perform a run with 100% Eluent 

B for 4 min, maintain elevated temperature, and repeat the blank injection. If 

ghosting is still present, yet minimized, repeat the run of 100% Eluent B. When 

not in use store column at room temperature in 25–75% (v/v) water-methanol 

with the column ends capped. 

References 

1. Anderson, N. L., Anderson, N. G., (2002) The human plasma proteome: history, 

character, and diagnostic prospects. Mol Cell Proteomics, 1, 845–67. 

2. Duan, X., Yarmush, D. M., Berthiaume, F., Jayaraman, A., and Yarmush, M. L. 

(2004) A moose serum two-dimensional gel map: application to profiling boon 

injury and infection. Electrophoresis, 25, 3055–65. 

3. Fujii, K., Nakano, T., Kawamura, T., Usui, F., Bando, Y., Wang, R., and 

Nishimura, T. (2004) Multidimensional protein profiling technology and its application 

to human plasma proteome. J.Proteome Res., 3, 712–18. 

4. Zolotarjova, N., Martosella, J., Nicol, G., Bailey, J., Boyes, B.E., and Barrett, 

W.C. (2005) Differences among techniques for high-abundant protein depletion. 

Proteomics, 5, 3304–13. 

5. Harlow, E. and Lane, D., (1988) Antibodies, A Laboratory Manual. Cold Springs 

Harbor Laboratory: New York, pp. 726. 

6. Martosella, J., Zolotarjova, N., Liu, H., Nicol, G., and Boyes, B.E., (2005) Reversedphase 

high-performance liquid chromatographic prefractionation of immunodepleted 

human serum proteins to enhance mass spectrometry: identification of lowerabundant 

proteins. J. Proteome Res. 4, 1522–1537.

4 

Immunoaffinity Fractionation of Plasma Proteins 

by Chicken IgY Antibodies 

Lei Huang and Xiangming Fang 

Summary 

Separation of complex mixtures having a wide dynamic range of protein concentration, 

such as plasma or serum, presents a significant challenge for proteomic analysis. 

Immunoaffinity fractionation is one of the most effective methods used during sample 

preparation to improve the ability to detect low-abundant proteins (LAP), enhancing 

biomarker discovery. Avian IgY (Immunoglobulin Yolk) antibodies have unique and 

advantageous features, which include strong avidity, high specificity, low nonspecific 

binding, and accumulative production. Polyclonal IgY antibodies covalently coupled to 

microbeads are particularly effective in specifically removing high-abundant proteins 

(HAP) from plasma, serum, CSF, urine, and other body fluid or cellular sources. IgY-12 

is a composition of IgY microbeads designed for one-step removal of the 12 most 

abundant proteins in human serum or plasma: albumin, IgG, transferrin, fibrinogen, 1antitrypsin, 

IgA, IgM, 2-macroglobulin, haptoglobin, apolipoproteins A-I and A-II, and 

orosomucoid (1-acid glycoprotein). Removal of the 12 HAPs enables improved resolution 

and dynamic range for one-dimensional gel electrophoresis (1DGE), two-dimensional gel 

electrophoresis (2DGE), and liquid chromatography/mass spectrometry (LC/MS). 

Key Words: High abundance proteins; IgY antibodies; immunoaffinity fractionation; 

low abundant proteins; protein depletion; protein separation; proteomics; sample preparation. 


IgY antibody is immunoglobulin gamma isolated from egg yolks (so 

called IgY) of certain avian and reptilian vertebrates such as birds, reptiles, 

and amphibians (1–3). Chicken IgY antibodies have been developed and 



41

42 Huang and Fang 

successfully applied for various types of immunoassays (4–8). An outstanding 

advantage of chicken IgY antibodies is that they are secreted by hens into egg 

yolk, resulting in a high-yielding and easy to access reservoir of antibodies 

(9). Compared to drawing blood, collecting eggs is noninvasive, continuous, 

convenient, and scalable. One egg contains about 100 mg of total IgY. A laying 

hen can actively produce eggs for 2 years at an average of 20 eggs per 

month. Distinct from mammalian IgG antibodies in molecular structure and 

biochemical features, IgY antibodies have been shown to have several advantages 

over IgG, particularly for their high avidity and less cross-reactivity 

to human proteins (10–12). This is because of the avian affinity maturation 

mechanism of gene conversion and the great evolutionary distance between 

chicken and mammals. When mammalian protein antigens are used to immunize 

chickens, more immunogenic epitopes are presented to the host, resulting in IgY 

antibodies with high affinity and broader recognition spectrum. In addition, the 

IgY Fc region does not bind human proteins such as complement, rheumatoid 

factor, Fc receptor, etc, thus significantly increasing IgY’s specificity of capture. 

Another unique feature of IgY antibodies is that they have a broader antigenbinding 

host range. This is also the result of greater evolutionary distance 

between chickens and mammals, and the sequence similarity among mammals. 

IgY antibodies raised against these high abundance proteins using human 

antigens also recognize the orthologous proteins from other mammalian species 

such as nonhuman primates, rat, mouse, pig, goat, cow, and dog. 

IgY microbeads are produced by covalently coupling IgY antibodies to 

60-μm polymeric beads via the oligosaccharides located on their Fc region. This 

orientated conjugation allows maximal capture of target proteins. Compared 

to other affinity reagents, including IgG microbead products, IgY microbeads 

have been shown to have distinct features and advantages (13,14). IgY-12 is a 

mixture of 12 types of IgY microbeads designed to collectively remove albumin, 

IgG, 1-antitrypsin, IgA, IgM, transferrin, haptoglobin, 1-acid glycoprotein 

(orosomucoid), 2-Macroglobulin, HDL (mainly apolipoproteins A-I and A- 

II), and fibrinogen from complex human body fluids such as serum, plasma, 

and cerebral spinal fluid (CSF) in a single step. IgY-12 spin column (0.6-mL 

bed size) can process 15–20 μL human plasma per loading, yielding 100–160 

μg of proteins partitioned of HAP. Larger samples can be processed by IgY- 

12 liquid chromatography (LC) columns. A 2-mL LC column can partition 

40–50 μL human plasma per injection and a 10-mL LC column allows a 

single loading of 200–250 μL. Through a simple procedure of sample loading, 

washing, eluting, and regenerating, approx 90–95% of total protein mass from 

human serum or plasma is removed. The LAP in the flow-through fractions 

can be further studied by 2D polyacrylamide gel electrophoresis (PAGE) or 

LC/MS. The regenerated beads can be reused many times with minimal protein

Immunoaffinity Fractionation of Plasma Proteins by Chicken 43 

carry-over (15). The IgY microbeads can also be used in 96-well filter plate and 

other formats for high-throughput partitioning of human serum/plasma samples 

or other body fluids. 

2. Materials 

2.1. IgY-12 High Capacity Spin Column Kit 

1. Prepacked IgY-12 spin column, containing 1.2-mL IgY microbeads slurry 

(Beckman Coulter, Fullerton, CA). Store at 2–8°C. Do not freeze. 

2. Dilution buffer (Tris Buffered Saline, TBS): 10 mM Tris-HCl, pH 7.4, 150 mM 

NaCl. For sample dilution, washing and equilibrating column, and rinsing pipet 

tips during resin transfer. Store at room temperature. 

3. Stripping buffer: 0.1M Glycine-HCl, pH 2.5. For stripping off bound proteins 

from column. Store at room temperature. 

4. Neutralization buffer: 1M Tris-HCl, pH 8.0. For neutralizing column and eluted 

proteins. Store at room temperature. 

5. 2-mL collection tubes. For collecting flow-through, washing, and eluted fractions. 

6. Empty spin columns with end caps. 

2.2. IgY-12 High Capacity LC2 or LC10 Column Kit 

1. Prepacked IgY-12 LC column, 2-mL or 10-mL packed bed (Beckman Coulter, 

Fullerton, CA). Store at 2–8°C. Do not freeze. 

2. Dilution, stripping, and neutralization buffers are same as for spin column. 

3. Spin filters. For sample clean up before loading column to remove sample particulates 

and extend column life. 

2.3. IgY-12 Microbeads for 96-Well Filter Plates 

1. IgY-12 microbeads, 50% slurry (GenWay Biotech, San Diego, CA). Store at 

2–8°C. Do not freeze. 

2. Dilution, stripping, and neutralization buffers are same as for spin column. 

3. 96-well filter plate, 400 μL, UHMW PE 25 μM, Long drip. (Innovative Microplate, 

Billerica, MA). Store at room temperature. 

4. Collection plate, Nunc 96-DeepWell Plates, 1.2-mL, Polypropylene (NUNC, 

Rochester, NY). 

2.4. SDS Gel Electrophoresis 

1. Tris-HCl SDS Gel, precast, 4–20% linear gradient (Rio-Rad, Hercules, CA). Store 

at 2–8°C. 

2. 5× SDS Sample Buffer: 10% (w/v) SDS, 20 mM dithiothreitol (DTT) or 25% (w/v) 

-mercaptoethanol (BME) (omitted under nonreducing condition), 20% (w/v)


glycerol, 0.2M Tris-HCl, pH 6.8, 0.05% (w/v) bromophenol blue. Store at room 

temperature. 

3. Tris/glycine/SDS electrophoresis buffer: 25 mM Tris-base, 200 mM glycine, 0.1% 

(w/v) SDS. 

4. Coomassie Blue Staining Solution: 0.25% (w/v) Coomassie Brilliant Blue R250, 

40% (v/v) methanol, 10% (v/v) acetic acid. Store at room temperature. 

5. Destain Soluton: 40% (v/v) methanol, 10% acetic acid. Store at room temperature. 

3. Methods 

3.1. Spin Column (see Note 1) 

3.1.1. Immunocapture of 12 Abundant Serum/Plasma Proteins 

1. Dilute 15–20 μL serum or plasma sample in dilution buffer to obtain a final 

volume of 600 μL. 

2. Snap off the tip from the column and place the column in a 2-mL collection tube. 

3. Centrifuge the column for 30 sec at 400g in a microcentrifuge to obtain dried 

beads. 

4. Place the end cap to the column. Immediately add 0.5 mL diluted sample to the 

dried beads in the column. Seal the column with the top snap cap. 

5. Mix the beads and the sample completely by inverting and shaking the column, 

place it on an end-to-end rotator and incubate at room temperature for 15 min. 

6. Invert the column. Remove the end cap and place the column in a 2-mL collection 

tube. Centrifuge for 30 sec at 400g. Collect flow-through (IgY-12-depleted) 

sample for further analysis (see Note 2). 

3.1.2. Washing of Column 

1. Wash beads with 0.5 mL of dilution buffer, a total of three times. To obtain 

maximum yields of flow-through samples, the fraction from the first washing can 

be collected and combine with the flow-through sample from Section 3.1.1 step 

6 for further analysis. 

2. For each wash, always first insert the end cap, and then add 0.5 mL of dilution 

buffer and seal the column with top snap cap. Mix the beads and buffer completely 

by inverting and shaking the column, remove the end cap while inverting the 

column and place it in a 2-mL collection tube. Centrifuge for 30 sec at 400g and 

save the flow-through for further analysis. 

3.1.3. Stripping of Bound Proteins 

1. Strip off bound proteins from beads using stripping buffer, a total of three to four 

times. For each elution, place the end cap to the column first after centrifugation, 

then add 0.5-mL stripping buffer and seal the column with top snap cap. Mix the 

beads and buffer completely (see Note 3) by inverting and shaking the column, 

incubate at room temperature for 3 min, remove the end cap while holding the


column upside down and place it in a 2-mL collection tubes. Centrifuge for 30 

sec at 400g and collect the eluate. It is crucial for column stability to immediately 

neutralize the beads (see Section 3.1.4.). 

2. Pool eluted samples (total 1.5–2.0 mL) and neutralize with 150–200 μL of neutralization 

buffer. Samples can be stored at -80°C if not analyzed immediately. 

3.1.4. Regeneration of IgY-12 Microbeads 

1. To regenerate IgY-12 microbeads after stripping bound serum proteins as 

described above, immediately neutralize beads with 0.6 mL of 1:10 diluted neutralization 

buffer. Mix beads and buffer completely by inverting and shaking the 

column. Incubate at room temperature for 5 min. Spin down beads in the column 

for 30 sec at 400g. 

2. Resuspend beads in 0.5-mL of dilution buffer. Beads are ready for next cycle or 

storage at 4°C. For storage of regenerated beads, add sodium azide (NaN3)to 0.02%. (w/v) in dilution buffer. 

3.2. LC Column (see Note 4) 

3.2.1. Protocol for 6.4 × 63 mm (2 mL) Column 

1. Set up the three Buffers as the only mobile phases. 

2. Purge lines with three Buffers at a flow rate of 1.0 mL/min for 10 min. 

3. Set up LC timetable (see Table 1 for details) and run two method blanks by 

injecting 125 μL of dilution buffer without a column. 

4. Attach column and equilibrate it in dilution buffer for 10 min at a flow rate of 

1.0 mL/min at room temperature. 

5. Dilute human serum five times (for example: 50 μL human serum with 200 μL 

of dilution buffer). 

6. Remove particulates with a 0.45-μm spin filter; 1 min at 9,000g. 

7. Inject 250 μL of the diluted and filtered plasma sample (Column capacity: 40–50 

μL of neat human serum/plasma per injection), start the method at a flow rate 

of 0.1 mL/min for 10 min, wash the column at a flow rate of 0.2 mL/min 

for 7 min, then change the flow rate to 1.0 mL/min to continue the wash for 

5 min, collect flow-through fraction and store collected fractions at –80°C if not 

analyzed immediately. 

8. Elute bound proteins from the column with stripping buffer at a flow rate of 

1.0 mL/min for 142 min, and neutralize the column with neutralizing buffer at 

a flow rate of 1.0 mL/min for 6 min. 

9. Regenerate column by equilibrating it with dilution buffer for an additional 

6 min at a flow rate of 1.0 mL/min. 

10. Store column after equilibrating with dilution buffer containing 0.02% (w/v) 

sodium azide (NaN3) at 2–8°C in a refrigerator. 

11. A standard chromatograph is illustrated in Fig. 1.


Table 1 

LC Method for a 6.4 × 63 mm column 

Cycle Time 

(min) 

Dilution 

buffer 

Stripping 

buffer 

Neutralization 

buffer 

Flow rate 

(mL/min) 

Max 

pressure 

(psi) 

Injection 

Wash 0 100 0 0 0.1 100 

Wash 10.01 100 0 0 0.2 100 

Wash 17.01 100 0 0 1.0 100 

Elution 22.01 0 100 0 1.0 100 

Neutralization 36.01 0 0 100 1.0 100 

Re-equilibrium 42.01 100 0 0 1.0 100 

Stop 48.00 

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

3.2.2. Protocol for 12.7 × 79.0 mm (10 mL) Column 

1. Set up the three buffers as the only mobile phases. 

2. Purge lines with three buffers at a flow rate of 1.0 mL/min for 10 min. 

3. Set up LC timetable (see Table 2 for details) and run two method blanks by 

injecting 1.25 mL of dilution buffer without a column. 

Fig. 1. Chromatography of immunoaffinity separation of human plasma using IgY- 

12 high capacity LC2 column. Fifty microliters human plasma was fractionated on the 

column.


4. Attach column and equilibrate it in dilution buffer for 10 min at a flow rate of 

2.0 mL/min at room temperature. 

5. Dilute human serum/plasma five times (for example: 250 μL human plasma with 

1.0 mL of dilution buffer). 

6. Remove particulates with a 0.45 μm spin filter; 1 min at 9,000g. 

7. Inject 1.5 mL of the diluted and filtered plasma sample (column capacity: 

200–250 μL of neat human serum/plasma), start the method at a flow rate of 

0.5 mL/min for 30 min, wash the column at a flow rate of 2.0 mL/min for 

5 min, collect flow-through fraction and store collected fractions at -80°C if not 

analyzed immediately. 

8. Elute bound proteins from the column with stripping buffer at a flow rate of 

2.0 mL/min for 15 min, and neutralize the column with neutralizing buffer at a 

flow rate of 2.0 mL/min for 10 min. 

9. Regenerate column by equilibrating it with dilution buffer for an additional 

10 min at a flow rate of 2.0 mL/min. 

10. Store column after equilibrating with dilution buffer containing 0.02% (w/v) 

sodium azide (NaN 3) at 2–8°C in a refrigerator. 

11. A standard chromatograph is illustrated in Fig. 2 

3.3. 96-Well Spin Filter Plate 

1. For each well, dilute 2–3 μL human serum or plasma in dilution buffer to obtain 

final volume of 100 μL. 

2. Aliquot 200 μL per well IgY-12 microbeads slurry into 96-well filter plate. Spin 

plate at 190g for 1 min in Eppendorf bench top centrifuge (see Note 5) with plate 

adapter to remove buffer. 

3. Add diluted sample to each well, mix with pipet tip. Incubate at room temperature 

on shaker for 15 min. 

Table 2 

LC method for a 12.7 × 79.0 mm column 

Cycle Time 

(min) 

Dilution 

buffer 

Stripping 

buffer 

Neutralization 

buffer 

Flow rate 

(mL/min) 

Max 

pressure 

(psi) 

Injection 

Wash 0 100 0 0 0.5 100 

Wash 30.01 100 0 0 2.0 100 

Elution 35.01 0 100 0 2.0 100 

Neutralization 50.01 0 0 100 2.0 100 

Re-equilibrium 60.01 100 0 0 2.0 100 

Stop 70.00 

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


Fig. 2. Chromatography of immunoaffinity separation of human plasma using IgY- 

12 LC10 column. Two hundred fifty microliters human plasma was fractionated on the 

column. 

4. Spin plate at 190g for 1 min. Collect flow-through fraction in collection plate, 

about 100 μL. 

5. Add 100 μL dilution buffer to each well. Gently shake the plate and spin at 190g 

for 1 min. Collect flow-through faction into the same collection plate from step 4. 

6. Wash beads with 100 μL dilution buffer, a total of 3 times. For each wash, add 

100 μL dilution buffer to each well, gently shake the plate and spin at 190g for 

1 min. Collect flow-through faction into collection plate for future analysis. 

7. Add 100 μL stripping buffer to each well. Gently shake the plate and incubate at 

room temperature on shaker for 2 min. Spin the plate at 190g for 1 min. Repeat 

for three to four times. Collect and combine flow-through factions into collection 

plate for future analysis. 

8. Immediately add 100 μL neutralization buffer to each well. Gently shake the plate 

and incubate at room temperature on shaker for 5 min. 

9. Spin the plate at 190g for 1 min. Add 100 μL dilution buffer to each well. Beads 

are ready for next cycle or storage at 4°C. For storage of regenerated beads, add 

sodium azide (NaN 3) to 0.02%. (w/v) in dilution buffer. 

3.4. Evaluation of Fractionation Efficiency by SDS-PAGE (see Note 6) 

1. Take a small aliquot of samples from neat plasma, flow-through and eluted 

fraction. Mix with 5× SDS sample buffer. Load approx 25–30 μL (see Note 7)


Fig. 3. SDS-PAGE analysis of neat plasma, flow-through and eluted fractions using 

IgY-12 high capacity spin column (1.0 mL slurry). Twenty microliters of human plasma 

was partitioned on the column. Five cycles were repeated. Fifteen microliters of 1:70 

dilution of neat plasma, 15 μL of flow-through fraction, and 15 μL of eluted fraction 

were loaded to 4–20% SDS gradient gel. Coomassie blue staining. M: molecular weight 

marker, S: Neat plasma. F1-F5: Flow-through fractions from cycle 1 to 5, E1-E5: 

Eluted/bound fractions from cycle 1 to 5. 

of each sample on 4–20% SDS gel. The gel is run in Tris/Glycine/SDS 

electrophoresis buffer at 200 volts for 35 min. 

2. Remove gel from the gel cassette. Rinse the gel with deionized water. Stain gel 

in Coomassie Blue Staining solution for 30 min to 1honshaker. 

3. Rinse the gel with deionized water. Place gel in destaining solution and shake 

until the bands emerge clearly from the background. Replace destaining solution 

with deionized water. 

4. A successful fractionation will result in distinct banding patterns on the gel as shown 

in Fig. 3. The major protein bands of albumin, transferrin, IgG, and Apo-A1 that 

disappeared in the flow-through fraction will be shown in the eluted fraction. 

4. Notes 

1. Before loading plasma sample to the new IgY-12 Spin Column, perform the full 

procedure with buffers only for one or two cycles. The purpose is to remove any 

residual uncoupled IgY antibodies in the column. 

2. The flow-through fraction is now greatly diluted. The samples can be concentrated 

to desired concentration and volume for downstream analysis using molecular


weight cutoff centrifugal concentrators, such as Vivaspin (Sartorius, Goettingen, 

Germany). An alternative method is TCA/Acetone precipitation. 

3. The beads tend to be packed tighter in acidic solution after centrifugation. 

Make sure beads are resuspended well for effective stripping. For more efficient 

stripping, 0.25 M Glycine-HCl, pH 2.5 can be used. In this case, 1 to 4 dilution 

of neutralization buffer should be used to regenerate beads and 250 μL of neutralization 

buffer should be added per 1 mL of eluted samples. 

4. The LC protocols are optimized for Beckman System Gold HPLC, Pump Module 

1 Type: 118, Detector Model: 166. If using other HPLC systems, some adjustment 

may be required. 

5. 96-well filter plate format can be adapted to automated liquid handling system. 

Some adjustments in procedure may be required. 

6. SDS-PAGE is a simple way to evaluate the efficiency of sample fractionation by 

IgY microbeads. The major plasma proteins, such as albumin, transferrin, IgG, 

and Apo-A1, can be easily visualized by Coomassie blue staining of SDS gel. 

The different protein banding patterns of neat plasma, flow-through fraction and 

eluted fraction represent the protein composition in each sample. Under reducing 

condition (with DTT or BME in SDS sample buffer), the heavy and light chains 

of IgG are separated and migrate at different speed; while under nonreducing 

conditions (no DTT or BME in SDS sample buffer), the heavy and light chains of 

IgG are linked by disulphide bounds and migrate on gel as single band at higher 

molecular weight. 

7. The total protein mass in plasma is about 60–80 mg/mL. Approximately 90% of 

proteins are captured by IgY-12 microbeads. The recovery rate is about 85–90%. 

The protein concentration in flow-through fraction is very low. To see protein 

banding pattern in SDS gel, load maximal volume of sample that a well of the gel 

can hold for flow-through and eluted/bound fractions, usually 25–30 μL(including 

5× Sample Buffer). As a control, load 15–20 μL of diluted neat plasma, usually 

at 1 to 70–80 dilutions. After flow-through fraction is concentrated and protein 

concentration is measured, an equal amount of protein (5–10 μG) from each 

fraction and neat plasma can be loaded for comparison on SDS gel. Fractionation 

efficiency can be assessed. 

8. IgY microbeads can also be used to partition plasma/serum from other species, 

such as nonhuman primates, mouse, rat, cow, dog, etc. To ensure maximal 

separation efficiency, use 50% of human sample loading for other species. 

9. IgY microbeads can be recycled for at least 100 times under proper conditions. 

It is important to neutralize the beads immediately after stripping. 


The author would like to thank Dr. Wei-Wei Zhang and Mr. Robert Gans 

for critical reading and editing of the manuscript. The IgY-12 microbeads 

products were developed and manufactured by GenWay Biotech and marketed 

by Beckman Coulter.


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and Zhang, W.-W. (2003) Polyclonal gene-specific IgY antibodies for proteomics 

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divide and conquer the proteome. Drug Discovery Today: Technology 1, 141–48 

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needs of proteomic sample preparation and analysis. Proteomics 5, 3314–28.

5 

Proteomics of Cerebrospinal Fluid: 

Methods for Sample Processing 

John E. Hale, Valentina Gelfanova, Jin-Sam You, Michael D. Knierman, 

and Robert A. Dean 

Summary 

Cerebrospinal fluid (CSF) provides an important source of potential biomarkers for 

brain disorders and therapeutic drug development. Applications of proteomic technology 

to the identification and quantification of proteins in CSF are increasing rapidly. Key to 

obtaining reproducible and reliable data about protein levels in CSF are standardization 

of methods for sample collection, storage, and subsequent sample processing. Methods 

are described here for all steps of sample processing for a number of different proteomic 

approaches. 

Key Words: Cerebrospinal fluid; mass spectrometry; proteomics; silver staining; 

two-dimensional gel electrophoresis. 


Cerebrospinal fluid is the interstitial fluid that bathes the ventricles of the 

brain. CSF is produced at a rate of approx 500 mL/d (1) and participates in 

maintenance of hydrodynamic pressure, transportation of nutrients, and removal 

of metabolites from the brain (2). Because of its proximity to the different 

regions of the brain, CSF has long been considered an important source for 

biomarkers of diseases of the brain. CSF is physically separated from plasma 

by the blood-brain barrier (bbb). This prevents the free flow of large molecules 

(such as proteins) from one space to the other. Smaller molecules may diffuse 

more freely however penetration of the bbb is dependent on the physical 



53

54 Hale et al. 

properties of the individual molecule. The proteins of CSF have been studied 

by two-dimensional (2-D) gel electrophoresis for some time (3). Many of 

the proteins seen in CSF are abundant serum proteins, which has led to the 

mistaken impression that CSF is simply an ultrafiltrate of serum. There are 

many differences between the protein composition of CSF and serum however. 

Thus CSF is not a simple ultrafiltrate of serum (1). Several proteins have been 

identified that are present at much higher concentration in CSF than in serum 

(4). Proteins produced and secreted in the brain will be prevented from rapidly 

diffusing into the serum by the same blood brain barrier that limits diffusion 

of molecules into the brain. The use of CSF for biomarker discovery and 

measurement is of obvious importance. However, differential levels of proteins 

may arise for many different reasons. Of major concern is blood contamination, 

which may occur during sample collection. The protein concentration of blood 

is 200–400 times higher than that for CSF so a very small percentage of blood 

can have a very dramatic effect on the protein profile of CSF of some proteins 

may be altered by differential handling of CSF samples and care should be 

taken to ensure that samples are collected and stored in as similar a fashion as 

possible. The success of any proteomic analysis of CSF is largely dependent 

on the quality of the sample analyzed. Although the samples are not collected 

by the analytical chemist directly, an understanding of the clinical methods 

used for collection and the criteria for acceptance of samples is important in 

downstream interpretation of the results. This section is intended as a guide 

to aid the analytical chemist in understanding the issues involved in sample 

collection and in assessing the suitability of individual samples for subsequent 

analysis. 

Lumbar puncture (LP) is routinely performed to collect cerebral spinal fluid 

(CSF) for confirmation of suspected meningeal infection and subarachnoid 

hemorrhage. CSF is also frequently examined to detect malignancies, neurodegenerative 

processes, and other pathology involving the central nervous system 

(CNS). LP also provides an opportunity for direct measurement of intracranial 

pressure (5,6). Although gross assessment of CSF provides clues about the 

presence of disease, physicians rely on a broad spectrum of laboratory 

techniques to evaluate patient specimens. The wealth of data available from 

standardized chemical, cytological, and microbiological tests on CSF from 

healthy individuals and patients with specific diseases provides the background 

against which clinical laboratories and physicians compare data for individual 

patients. This knowledge base allows physicians to narrow down the diagnostic 

possibilities suggested by a patient’s complaint and presentation (5,7). 

In clinical research, LP has increasingly been paired with varied analytical 

technologies to better characterize normal and disease biology and enhance 

diagnostic accuracy (8,9). The procedure also is used to characterize the central

Proteomics of Cerebrospinal Fluid 55 

disposition, pharmacodynamic and clinical responses produced by established 

and candidate therapeutics. In drug development, analyses of CSF are used 

to determine if drugs penetrate the CNS and alter biochemical pathways 

and cellular responses. This approach aims to more thoroughly define the 

mechanism of action and the optimal dose for drugs designed to act on 

the CNS (10). Collection of CSF for clinical research generally employs LP 

techniques similar to those used in routine clinical practice. In attempting to 

identify central pharmacodynamic effects, LP is occasionally performed before 

and following multidose administration of drug designed to achieve circulating 

steady state concentrations (11). Placement of an indwelling catheter 

for continuous sampling of CSF from the thecal sac has also been reported in 

clinical research (12). This approach provides an opportunity to characterize 

diurnal changes in CSF. Continuous CSF sampling also creates an opportunity 

to evaluate acute pharmacokinetic and pharmacodynamic responses with 

various pharmacological interventions (10). 

Lumbar puncture for CSF collection in research is generally safe (13). 

Nevertheless, LP is an invasive procedure. Whether done for diagnostic or 

research purposes, the procedure carries a number of inherent risks. The most 

common complication from LP is postdural puncture headache. The headache is 

typically frontotemporal and may be accompanied by neck stiffness, dizziness, 

and nausea. The headache may result from added tension on anchoring structures 

of the brain because of removal of CSF (14). Leakage of CSF at the dural 

puncture site may be an important factor. The latter explanation is supported 

by a decreased incidence of headache when LP is performed using a small 

gauge spinal needle shown to reduce CSF leakage in an in vitro cadaveric dural 

model (15). Maintaining the patient in a prone, slightly head down position 

can help resolve the headache and associated symptoms. Persistent headache 

can be treated by intravenous administration of caffeine or epidural injection of 

autologous whole blood (blood patch) at the LP site (14). Other more serious, 

but rare risks include hemorrhage, infection, and herniation of the brain. As a 

result, LP is contraindicated in individuals with a bleeding diathesis, thrombocytopenia 

(platelet count


or L3–L4 interspaces through which the spinal needle will pass (5,16). If the 

impact of the position on the planned analyses is unknown, it may be useful 

to record or predetermine the position in the research protocol. Strict attention 

to maintenance of sterile conditions is required. Administration of a local 

anesthetic reduces the risk of pain and tends to reduce unwanted movement of 

the patient during the course of the procedure. 

The care taken to avoid risk to a patient also helps ensure access to a high 

quality CSF specimen. A traumatic LP increases the likelihood of blood in the 

CSF specimen. Excessive blood contamination is particularly problematic as 

it can distort chemical measurements and cell counts performed on CSF. If 

sufficiently severe, this issue can compromise clinical interpretation or research 

conclusions. The possibility of a traumatic tap producing a blood contaminated 

specimen can sometimes occur even with careful preparation of the patient and 

close attention to procedural technique. Accordingly, CSF is typically collected 

in a series of sterile tubes that are submitted for laboratory analyses in a fashion 

that reduces the potential for such confounds. In diagnostic settings, the first 

tube is typically used for chemical and immunological analyses, a second for 

microbiological examination, and a third for total cell count and differential 

cell counts (17). Despite such precautions, clinical laboratories may refuse 

to perform selected analyses on grossly bloody CSF specimens. Similarly, 

research laboratories should pay close attention to this common, undesired 

source of preanalytical variability and define criteria that render a CSF specimen 

unacceptable for the intended use. Once collected, the CSF must be processed, 

preserved, and transported in a timely manner so as not to render a good 

specimen unacceptable. Preservation of specimen constituents consistent with 

the state at the time of collection requires attention to the potential impact of the 

collection device and vials, materials used for processing time and temperature. 

It is important to assure that “biology in the tube” does not confound attempts 

to characterize the biological state under investigation through degradation of 

chemical and cellular elements. 

Although we realize we cannot anticipate all possible proteomic methodologies 

that may be applied to CSF, the purpose of this review is to provide 

basic methodology for the collection, storage, qualification, and processing of 

CSF for proteomic applications. 

2. Materials 

2.1. Sample Simplification 

1. Montage equilibration buffer, wash buffer and columns are provide with the 

Montage Albumin Deplete Kit (Millipore). 

2. Protein G Sepharose (Amersham Biosciences).


2.2. Gel Electrophoresis 

1. Ammonium carbonate solution: 1M ammonium carbonate adjusted to pH 11 with 

30 % ammonium hydroxide. 

2. Reduction-alkylation cocktail: 97.5% ACN, 2% iodoethanol, and 0.5% 

triethylphosphine. 

3. Rehydration sample buffer: 11 cm IPG strips, (pH 3–10 nonlinear) and 8–16% 

linear gradient Protean II gels were from Bio-Rad, Hercules, CA, USA. 

4. IPG reduction buffer: 0.1M Tris-HC1, pH 6.8, 6M urea, 2M thiourea, 1% SDS, 

6.4 mM dithiothreitol (DTT), 30% glycerol, and a few grains of bromophenol 

blue. 

5. IPG alkylation buffer: 0.1M Tris-HC1, pH 6.8, 6M urea, 2M thiourea, 1% SDS, 

24 mM iodoacetamide , 30% glycerol, and a few grains of bromophenol blue. 

2.3. Silver Stain 

1. Fixer 1: 50% MeOH, 10% Acetic acid. 

2. Fixer 2: 5% MeOH, 10% acetic acid. 

3. DTT solution: 5 mg/mL dithiothreitol. 

4. Silver solution: 0.1% AgNO 3 

5. Developer: 30 g sodium carbonate in 1LH 2O + 500 μL 37% formaldehyde. 

6. Stopping solution: 2.3M citric acid. 

2.4. In-Gel Digestion 

1. Farmers reagent: 30 mM potassium ferricyanide (solution 1) and 100 mM sodium 

thiosulphate (solution 2). The working reagent is 1 part solution 1 and 1 part 

solution 2 mixed together just before use. 

2. Gel reduction-alkylation reagent: 50% 0.1M ammonium carbonate solution, 

48.75% acetonitrile (by volume), 1% iodoethanol, and 0.25% triethylphosphine 

(final pH 10). 

3. Gel trypsin solution: 20 ng/μL analytical grade (Promega,Madison, WI). One vial 

containing 20 μg was reconstituted with 1 mL 50 mM sodium bicarbonate, pH 8. 

4. Extraction solution: 100 mM NH 4HCO 3, pH 8.5. 

5. Destain reagent: 50% acetonitrile, 25 mM NH 4HCO 3, pH 8.0. 

6. DTT solution 2: 50 mM DTT in 100 mM NH 4HCO 3, pH 8.0. 

7. Iodoacetamide solution: 100 mM iodoacetamide in 100 mM NH 4HCO 3, pH 8.0. 

2.5. Processing CSF for LC/MS/MS Analysis 

1. Chicken lysozyme (Sigma, St Louis, MO) is dissolved at 1 mg/mL in H2O and 

stored in single use aliquots at –80°C. Working solutions are prepared by diluting 

at 12.5 μg/mL in urea solution before use. 

2. Urea solution: 8M urea in 100 mM ammonium carbonate, pH 11.0. 

3. LC/MS reduction-alkylation reagent: 97.5 % acetonitrile, 2% iodoethanol, 0.5% 

triethylphosphine (see Note 1).


4. LC/MS trypsin solution: TPCK treated bovine pancreatic trypsin (Worthington, 

Lakewood, NJ) is dissolved at 1 mg/mL in H 2O and stored in single use aliquots 

at –80°C. Working solutions are prepared by diluting to 5 μg/mL in 100 mM 

ammonium bicarbonate pH 8.0 before use. 

2.6. MALDI-TOF Mass Spectrometry 

1. O-methylisourea solution: 1M solution O-methylisourea-hemisulfate (Arcos) in 

100 mM sodium carbonate buffer adjusted to pH 10 with 5.0N NaOH (see Note 2). 

2. Matrix solution: Recrystallized -cyano-4-hydroxycinnamic acid (see Note 3) is 

reconstituted with 200 μL 50 % acetonitrile, 0.05 % TFA to make saturated matrix 

solution. 

2.7. LC-MS/MS Mass Spectrometry. 

1. The C-18 reversed phase column was a Zorbax SB300 1× 50 mm (Agilent). 

2. Solvent A: 0.1% formic acid (Aldrich) in water (Burdick and Jackson HPLC 

grade). 

3. Solvent B: 50% acetonitrile, 0.1% formic acid (Aldrich) in water (Burdick and 

Jackson HPLC grade). 

4. Solvent C: 80% acetonitrile, 0.1% formic acid (Aldrich) in water (Burdick and 

Jackson HPLC grade). 

3. Methods 

3.1. Simplification of CSF by Removal of Abundant Proteins 

Similar to serum, albumin and immunoglobulin comprise more than 50 % of 

the protein concentration of CSF. To visualize proteins of lower abundance, it 

is desirable to reduce the levels of these proteins before separation or analysis. 

One procedure for the reduction of these proteins is described here. 

1. Resuspend Protein G Sepharose beads in Montage equilibration buffer, centrifuge 

at 500g for 2 min, and discard supernatant. Repeat Montage equilibration buffer 

addition and centrifuging once. Resuspend beads pellet in equal volume of 

Montage equilibration buffer. 

2. Dilute aliquots (50 μg protein) of CSF with Montage equilibration buffer to a 

volume of 300 μL. 

3. To the CSF samples, add 20μL Protein G Sepharose bead suspension and agitate 

slowly for 1hatRT. 

4. While samples are incubating with Protein G Sepharose, rehydrate Montage 

columns via manufacturer specifications: add 400 μL of Montage equilibration 

buffer to column insert and centrifuge at 500g for 2 min. Discard eluate from the 

collection tube. 

5. Pellet Protein G Sepharose beads at 500g for 2 min.


6. Carefully remove (without disturbing beads) and transfer 280 μL of the effluent to 

a rehydrated Montage column. Centrifuge column at 500g for 2 min. Reapply the 

eluate to the column and centrifuge again. Add two consecutive 100-μL washes 

of Montage wash buffer over the column via 500g centrifugation for 2 min (final 

volume approx 500 μL). Discard the column insert. 

7. Speed vacuum samples to approx 30–50 μL. Monitor volumes during speed 

vacuuming to prevent evaporation to dryness. 

3.2. Processing CSF for 2-Dimensional Gel Electrophoresis 

Separation of proteins by 2-dimensional gel electrophoresis may be adversely 

affected by the presence of salts in the sample. Additionally, smearing of bands 

has been noted in 2D gel electrophoresis that is attributable to incomplete 

reduction of disulfides in proteins (18). We have addressed both of these issues 

in the procedure that follows. 

1. Dialyze CSF against deionized water using 3-kDa MWCO dialysis tubing. 

2. To reduce and alkylate the protein samples, place 50 μg of CSF protein (approx 

50 μL) in a tube. Add 5 μL of ammonium carbonate solution followed by 50 μL 

of reduction-alkylation cocktail ( see Note 1). Cap the tube and incubate for 60 

min at 37°C. 

3. The sample is then uncapped and evaporated in a speedvacuum with medium heat 

for 2 h. After the sample is dried, reconstitute it in Bio-Rad rehydration sample 

buffer. 

4. Samples are isoelectrically focused using 11 cm IPG strips, (pH 3–10, nonlinear). 

Rehydrate the IPG strips overnight at room temperature. The IPG strips are run 

at 8,000 V for 60,000 Vh using an IEF cell. 

5. The second dimension separation is performed in 8–16% linear gradient Protean 

II gels at 120 V for 2 h. 

6. Stain the gels with silver (Section 3.3) and scan with a Bio- Rad FX-Imager at 

50-μm resolution. 

3.2.1. Alternative 2-Dimensional Gel Electrophoresis Procedure 

This procedure incorporates the alkylation of cysteines into the prepping of 

the IPG strips for the second dimension. 

1. After dialysis against water, dissolve samples in approx 0.4 mL of rehydration 

solution and apply to immobilized pH gradient strips. The rehydration of the IPG 

strips is performed overnight at room temperature. Run the rehydrated IPG strips 

at 8,000 V for 60,000 Vh using an IEF cell. 

2. Following focusing, reduce the proteins on the strips by immersing the strips in 

IPG reduction buffer for 15 min at room temperature. 

3. Alkylate the proteins by immersing the strips in IPG alkylation buffer for 15 min 

at room temperature.


4. The IPG strips are then overlayed on SDS PAGE gels (8–16 % linear gradient 

Protean II gels) and separated at 120 V for 2 h. 

3.3. Mass Spectrometry Compatible Silver Stain 

Some silver staining procedures use reagents, such as glutaraldehyde, which 

will interfere with proteomic analysis of proteins (19). Glutaraldehyde will react 

with lysines in proteins and cross-link them. This interferes with trypsin, which 

is the typical enzyme used for in-gel digestion procedures. Formaldehyde is 

capable of inducing the reduction of silver while not cross linking proteins. The 

following procedure has been used for staining of gels before in-gel digestion 

and mass spectral analysis. 

1. Following electrophoresis soak the gels in fixer 1 for 30 min (see Note 4). Then 

for an additional 30 min in fixer 2. Agitate on a rocker platform gently. 

2. Rinse gels with Milli-Q H 2O with gentle agitation for 20 min to 1 h (200 mL at 

a time). 

3. Soak gels in a DTT solution for 30 min (volume sufficient to cover gel i.e., 

100 mL) with gentle rocking. 

4. Pour the DTT solution off and add 100 mL silver solution to cover gel which is 

soaked 30 min with gentle rocking. 

5. Pour off the silver solution and rinse the gel once with 100 mL H 2O (rapidly). 

Rinse the gel twice with 100 mL developer for ∼30 sec. Then soak in 100 mL 

developer until proteins reach the desired intensity (see Note 5). 

6. Stop the development by adding 5 mL stopping solution per 100 mL developer 

and agitating for 10 min. 

7. Rinse the gel with several changes of distilled H 2O over 30 min. Fig. 1 shows an 

image of a silver stained 2-D gel of albumin depleted CSF. Fig. 2 lists the spots 

that were identified after digestion and LC/MS/MS analysis. 

3.4. In-Gel Digestion Procedure 

3.4.1. Standard Protocol 

Following gel separation and staining, a common procedure for identification 

of proteins is excising the gel band or spot and digestion of the protein in the gel 

with a proteolytic enzyme (most commonly trypsin). The following procedure 

may be used with dye or silver stained gels. 

1. Place Coomassie blue or Sypro ruby-stained gel pieces in Eppendorf centrifuge 

tubes or in the wells of a 96-well plate. 

2. Silver stained gel pieces are first de-stained by immersing them in Farmers reagent 

for 5–10 min followed by 3 washes with Milli-Q water (see Note 6).


Fig. 1. Albumin was removed from a pooled CSF sample using the procedure in 

Section 3.1 without the protein G step. Proteins were separated by two-dimensional 

electrophoresis. The gel was silver stained. 

3. Immerse the gel pieces in gel reduction-alkylation reagent sufficient to completely 

cover them (usually ∼100 μL (see Note 1). Cap the tubes, or seal the plate, and 

incubate the samples at 37°C for 60 min. 

4. After incubation, the reagent is drawn off and discarded. Dehydrate the gel pieces 

with 100 μL of pure acetonitrile for approx 5 min. 

5. Decant the excess acetonitrile, and dry the gel pieces on a speed vacuum for at 

least 15min with medium heat. 

6. Rehydrate the gel pieces with gel trypsin solution (usually 5–10 μL,) and incubate 

overnight at 37°C (see Note 7). 

7. For LC/MS/MS analysis, extract peptides with extraction solution for 60 min at 

37°C and desalt with a μC-18 Ziptip before injection onto the mass spectrometer. 

For MALDI analysis, extract peptides as described in Section 3.6. 

3.4.2. Alternative Procedure for Reduction and Alkylation of Gel Pieces 

For proteins separated in gels without prior reduction and alkylation this 

procedure can be substituted for the reduction/alkylation step described previously.


Fig. 2. Spots were cut out and digested with trypsin (see 3.4). LC-MS/MS was 

performed with the procedure listed in Section 3.7 and proteins identified searching 

the non-redundant database with SEQUEST. 

1. Crush excised Coomassie blue or Sypro stained gel pieces and soak in destain 

reagent for at least 5 min. Decant destain and add fresh destain reagent for another 

5 min. Repeat at least 3 times total. 

2. Silver stained gel pieces are destained as described previously. 

3. Reduce protein in the gel with 50 μL of DTT solution 2, for 30 min at 56°C. 

4. Remove the DTT solution and replace with 50 μL of iodoacetamide solution, for 

30 min at 45°C. 

5. Remove the iodoacetamide solution, wash the gel pieces with extraction solution, 

and then with acetonitrile and dry on a speedvacuum. 

6. Rehydrate the gel pieces with gel trypsin solution (usually 5–10 μL,) and incubate 

overnight at 37°C (see Note 7). 

7. For LC/MS/MS analysis, extract peptides with extraction solution for 60 min at 

37°C and desalt with a μC-18 Ziptip before injection onto the mass spectrum. For 

MALDI analysis, extract peptides as described in Section 3.6.


3.5. Processing CSF for LC/MS/MS Analysis 

More frequently, gel electrophoresis procedures are being eliminated and 

protein mixtures are being analyzed by chromatographic separation of proteolytic 

digests of the mixtures. This procedure has been used for processing of 

unfractionated cerebrospinal fluid before LC/MS/MS analysis. 

1. Prepare the albumin and immunoglobulin depleted CSF as in Section 3.1. 

2. Add 40 μL of chicken lysozyme solution (see Note 8). 

3. Add 100 μL of LC/MS reduction/alkylation reagent (see Note 1). Close the tubes 

and incubate for 1hat37°C. 

4. Speedvacuum the solutions to dryness, at least 3 h with medium heat. 

5. Redissolve the pellet in 200 μL of LC/MS trypsin solution to produce a 1.6M 

urea solution and an enzyme: substrate ratio of 1:50 (w/w). 

6. Incubate at 37°C overnight. 

7. Typically, 100 μL of this digest is injected onto the mass spectrometer. 

3.6. MALDI-TOF Mass Spectrometry 

Peptides from in-gel digests of proteins may be identified from their peptide 

fingerprint. MALDI-TOF mass spectrometry can provide that fingerprint which 

consists of the singly protonated mass of each tryptic peptide in a given protein. 

These fingerprints may be used to search protein databases using software 

written for this purpose. Conversion of lysines into homoarginines increases 

the sensitivity of detection of lysine containing tryptic peptides (20–22). This 

procedure has been used for 1 and 2 D gel separated proteins. 

1. Following in-gel trypsin digestion (Section 3.4), extract peptides with Omethylisourea 

solution for 60 min at 37°C to convert lysines to homoarginine 

2. Desalt samples with a μC-18 Ziptip before MALDI mass spectral analysis. 

3. Spot the desalted peptides onto a MALDI target (1 μL) and add an equal volume 

of saturated matrix solution. 

4. After the spots dry, MALDI-TOF spectra are obtained on a Voyager DE-Pro mass 

spectrometer (see Note 9). 

5. Search peptide masses using the program Knexus. The mass of the homoarginine 

residue (170.23) and the S-ethanol-cysteine residue (147.03) are added to the 

list of user defined modifications. For peptides obtained from in-gel digests, the 

oxidized methionine modification is also selected. 

3.7. LC-MS/MS Mass Spectrometry 

For unfractionated protein digests or for gel digests that contain multiple 

proteins, LC-MS/MS analysis can separate and fragment peptides providing


spectra that can be used to search databases for peptide identification. The 

following procedure is capable of separating peptides from digests of unfractionated 

CSF. 

1. Inject tryptic digests (100–200 L from Section 3.5) onto a C-18 reversed 

phase column at a flow rate of 50 L/min on a Surveyor HPLC system 

(ThermoFinnigan). 

2. The gradient conditions are 90% solvent A, 10% solvent B to 5% solvent A, 95% 

solvent B over 120 min, followed by a 0.1 min ramp to 100% solvent C, and held 

at 100% solvent C for 5 min, followed by a 0.1 min ramp to 90 % solvent A, 

10% solvent B, and held for 17 min. Divert the effluent to waste for the first 

5 min to keep the source clean. The total column effluent is connected to the 

electrospray interface of an LTQ ion trap mass spectrometer (ThermoFinnigan). 

The source is operated in positive ion mode with 4.8 kV electrospray potential, a 

sheath gas flow of 20 arbitrary units, and a capillary temperature of 225°C. The 

source lenses are set by maximizing the ion current for the 2+ charge state of 

angiotensin. 

3. Collect data in the triple play mode with the following parameters: centroid parent 

scan set to 1 microscan and 50 ms maximum injection time, profile zoom scan 

set to 3 microscans and 500 ms maximum injection time, and a centroid MS/MS 

scan set to 2 microscans and 2000 ms maximum injection time. Set dynamic 

exclusion settings to a repeat count of one, exclusion list duration of two minutes, 

and rejection widths of –0.75 m/z and +2.0 m/z. 

4. Carry collisional activation out at a relative collision energy of 35% and an 

exclusion width of 3 m/z. 

5. Search MS/MS spectra against a nonredundant protein database with 

SEQUEST (23). 

4. Notes 

1. The reduction alkylation solution should be prepared just before use. Triethylphosphine 

is pyrophoric and should be handled in a fume hood in accordance with the 

material safety data sheet. 

2. If the pH is not properly adjusted to 10.0 the reaction may not proceed to 

completion. 

3. -cyano-4-hydroxycinnamic acid (Sigma, St Louis, MO) is recrystalized with 

the following procedure. Place 50 mg -cyano-4-hydroxy-cinnamic acid in a 50 

mL tube and add 10 mL of DI water. Add 30 % ammonium hydroxide until 

completely dissolved. Centrifuge and pour into a new 50-mL tube. In a fume 

hood add neat trifluoroacetic acid until pH is below 3 to precipitate the -cyano- 

4-hydroxycinnamic acid (check with indicator paper), centrifuge, and decant the 

supernatant. Wash the pellet 3× with 0.1 % TFA. Dissolve the final pellet into 

enough 50% acetonitrile 0.1% TFA to just dissolve it, no more than 40 mL. 

Aliquot 200 μL into 1.0 mL tubes, speed vaccuum to dryness and store at –20°C.


4. To minimize background staining, clean glassware with which the gel will come 

into contact with 50% HNO 3. Wear gloves when handling the gel as fingerprints 

will produce background stain. 

5. Development should be executed with continuous agitation. If a precipitate should 

begin to form discard the developer and replace it with fresh developer. 

6. The gel pieces should be clear. If yellow color remains continue washes until it 

is gone. 

7. The volume of trypsin solution added should be just enough to rehydrate the gel. 

There should not be excess liquid immersing the gel. 

8. Lysozyme is added as an internal standard to monitor the efficiency of 

reduction/alkylation and digestion. 

9. The mass spectrometer was calibrated with des-Arg-bradykinin, angiotensin I, 

glu1-fibrinopeptide,and ACTH [1–17, 18–39, and 7–38], insulin (bovine), thioredoxin 

(Escherichia coli), and apomyoglobin (horse) (Sigma, St. Louis, MO). 

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sclerosis treated with interferon beta. Clin. Neurol. Neurosurg. 108:124–28, 

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(1992) Serial cerebrospinal fluid corticotropin-releasing hormone concentrations 

in healthy and depressed humans. J. Clin. Endocrinol. Metab. 74:1325–30 

13. Jhee S. S., Zarotsky V. (2003) Safety and tolerability of serial cerebrospinal 

fluid (CSF) collections during pharmacokinetic/pharmacodynamic studies: 5 years 

experience. Clin. Res. Reg. Affairs 20(3):357–63. 

14. Raskin N. H. Headache. In: Harrison’s Principles of Internal medicine, 16th 

ed. Kasper D. L., Fauci A. S., Longo D. L., Brauwald E., Hauser S. L., and 

Jameson J. L. eds. McGraw Hill, New York, pp 85–94 

15. Angle P. J., Kronberg J, E., Thompson D. E., et al. (2003) Dural tissue trauma 

and cerebrospinal fluid leak after epidural needle puncture: effect of needle design, 

angle, and bevel orientation. Anesthesiology 99(6):1376–82. 

16. Sandoval M., Shestak W., Sturmann K., and Hsu C. (2004) Optimal patient 

position for lumbar puncture, measured by ultrasonography. Emergency Radiology 

10(4):179–81. 

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2nd ed. American Society of Clinical Pathologists Press, Chicago, IL. 

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proteins in preparation of two-dimensional map analysis: why, when, and how. 


19. Shevchenko, A., Wilm, M., Vorm, O., Mann, M.(1996) Mass spectrometric 

sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68, 850–58. 

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of tryptic peptide detection by MALDI-TOF mass spectrometry is achieved 

by conversion of lysine to homoarginine. Anal. Biochem. 287, 110–17. 

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laser desorption/ionization mass spectrometric analysis of tryptic hydrolysates of 

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Anal. Chem. 71, 4981–88.

6 

Sample Preparation of Bronchoalveolar Lavage Fluid 

Baptiste Leroy, Paul Falmagne, and Ruddy Wattiez 

Summary 

Respiratory diseases are important health problem throughout the world. The 

bronchoalveolar lavage (BAL) fluid obtained by fiber-optic bronchoscopy is a biofluid 

reflecting the expression of secreted pulmonary proteins and the products of activated 

cells. The characterization of the BALF proteome provides an opportunity to establish 

diagnostic and get prognostic indicators of airway diseases. The main part of the chapter 

is devoted to the description of the most effective and reliable method of BALF sample 

preparation and processing for proteomic studies. Principal BALF proteome characteristics 

and the difficulties to work with this fluid are also introduced. Finally, the best 

conditions for high resolution and high reliability 2-dimensional electrophoresis (DE) of 

BALF samples are given. 

Key Words: BALF sampling; bronchoalveolar lavage fluid; epithelial lining fluid; 

lung; mass spectrometry; proteome. 


The respiratory tract, and in particular the alveoli, are covered by a thin film 

called epithelium lining fluid (ELF). This fluid plays important functions of 

protection against external aggressions and preservation of the gas-exchange 

capability of the airways. ELF contains several categories of cells (mainly 

of the immune system) and a wide variety of soluble components (lipids, 

nucleic acids, and proteins/peptides) (1). Diverse techniques exist to sample 

this particular fluid, but the most commonly used remains the bronchoalveolar 

lavage (BAL) during fiber-optic bronchoscopy. This minimally invasive and 



67

68 Leroy et al. 

reproducible procedure consists of liquid instillation through a fibrobronchoscope 

and recovery of the bronchoalveolar lavage fluid (BALF) by gentle 

aspiration (2,3). This biofluid allows the monitoring of the expression of 

secreted pulmonary proteins and the products of activated cells. 

In the last decade, the number of BALF focusing studies has grown exponentially. 

BALF proteome study has been initiated by some groups that have 

demonstrated that investigations of BALF proteome and the discovery of 

disease associated proteins should contribute to a better knowledge of the 

lung at the molecular level and to the study of the lung disorders at the 

clinical level (4–8). The most abundant proteins are plasma proteins that 

derive by diffusion across the blood barrier. A comparative analysis of serum 

and BALF proteomes revealed the presence of proteins specifically produced 

in the airways (9). These proteins are, therefore, good candidates for lung 

specific disease biomarkers. The proteins produced locally are very heterogeneous 

and can be classified according to their function: proteins involved 

in defense mechanisms, tissue remodelling, lipid metabolism, inflammatory 

processes, cell growth, and oxidant-antioxidant systems. Differential-display 

proteomics studies showed that nonphysiological conditions cause significant 

modifications of the BALF proteome; these modifications can be used to 

better understand pathogenesis mechanisms or to reveal diseases. Significant 

efforts have been devoted to large scale identification of BALF proteins 

or their degradation products and to the evaluation of potential markers 

for lung diseases, especially, for fibrosing interstitial lung diseases such as 

sarcoidosis and idiopathic pulmonary fibrosis (4,6,8,10,11). A majority of 

BALF proteomic studies use the two dimensional gel electrophoresis (2-DE) 

approach associated with mass spectrometry technologies. The technological 

progress encountered recently also allow scientists to reach the objective 

of quantitative analysis that is of crucial importance in differential-display 

proteomic. Reproducible two-dimensional BALF proteome patterns can be 

obtained using immobilized pH gradients (IPG) in the first dimension (Fig. 1) 

(12). After staining, the two dimensional patterns are compared to reveal 

up- or downregulated proteins. High throughput identification of proteins by 

mass spectrometry techniques also permits the creation of reference gels of 

the BALF proteome (9). These gels are now available on the World Wide 

Web (http://W3.umh.ac.be/biochim/proteomic.htm). The complexity of BALF 

proteome has been demonstrated by the great diversity of proteins present in 

this fluid. 

Although BALF proteome analysis is now largely facilitated by previous 

works, BALF sample preparation remains a critical step (13). Indeed, majors 

problems associated with BALF proteome analysis are ascribable to the low

Sample Preparation of Bronchoalveolar Lavage Fluid 69 

100 kDa 

MW 

5 kDa 

3.5 

α1-antitrypsin 

SP-A 

CC16 

Albumin 

Immunoglobulin light chain, κλ 

Transthyretin 

pI 10 

IgG heavy chain, γ 

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

protein content and the high salt (because of the phosphate-buffered saline used 

for the lavage procedure) and lipid concentrations of this sample. Moreover, as 

for other human fluid, the wide dynamic range of protein concentration with the 

occurrence of some major proteins gives rise to difficulties in the study of lowabundance 

proteins. In this context, several efforts have been made to optimize 

BALF sample preparation and to identify less abundant proteins (13). Recently, 

alternative or supplement methods to the 2-DE approach have been developed 

such as multidimensional liquid chromatography (multi-LC): proteins from 

BALF are separated by chromatography (ionic and/or reverse phase) coupled 

“on line” with mass spectrometry analysis (multi LC-MS) (10,14). This new 

promising approach allows the identification and the quantification of small, 

minor, and hydrophobic proteins present in the BALF samples. Nevertheless, 

at this moment, LC–MS technology dedicated to BALF proteome study lacks 

quantitative data. 

In this chapter, we describe BALF sample preparation for 2-DE, the most 

currently used proteomic tool. Clearly, this sample preparation dedicated to 

2-DE is also adequate to LC-MS approach. Finally, the conditions used in our 

laboratory to achieve 2-DE separation of BALF proteins are also detailed.


2. Materials 

1. Sterile saline solution: 0.9% (w/v) NaCl, filtered through a 0.2-μm syringe filter 

unit (Minisart, Sartorius, Germany). 

2. Ultrapure water (double-distilled, deionized, >18) is used for all reagent 

preparations. 

3. Protease inhibitor cocktail tablets (Complete mini, Roche, Germany). 

4. Dialysis membrane: Spectra/Por membrane (MWCO : 3.5 kDa) (Spectrum). 

5. Dialysis buffer: 50 mM NH4HCO3. 6. Speed Vac. 

7. Immobilized pH gradient 3–10 NL 180mm (Pharmacia-Amersham). 

8. Rehydration solution: 7 M urea, 2 M thiourea, 4% CHAPS (w/v), 2% ampholytes 

3–10 (v/v), 65 mM DTE, and a trace of bromophenol blue. Prepare fresh each 

time. 

9. Sample buffer: 7 M urea, 2 M thiourea, 4% CHAPS (w/v), 2% ampholytes 

3–10 (v/v), 65 mM DTE and trace amounts of bromophenol blue. Prepare fresh 

each time. 

10. Equilibration buffer: 50 mM Tris-HCl, pH 6.8, 6 M urea, 30% glycerol (w/v), 

2% SDS (w/v). Prepare fresh each time. 

11. DryStrip Reswilling tray (Amersham biosciences or Bio-Rad). 

12. Multiphor II (Amersham biosciences) or Protean IEF Cell (Bio-Rad) device. 

13. Standard vertical electrophoresis units for SDS-PAGE. 

14. Programmable power supply able to deliver >3000 V. 

15. Thermostatic circulator (Multitemp II, Amersham biosciences). 

3. Methods 

3.1. BALF Sampling 

1. Under medical controle, place a flexible fiberoptic bronchoscope through an 

endotracheal tube wedged into a subsegmental bronchus of an anesthetized patient 

(see Note 1 and 2). 

2. Instill, through the bronchoscope, 20 mL of sterile saline solution warmed to 

37°C. 

3. Collect the fluid by gentle aspiration and dispose in sterile siliconated bottles on 

ice (see Note 3). All subsequent manipulation of the samples should be realized 

on ice. 

4. Repeat procedure 1–3 four times and combine harvested fluids except the first 

sample (see Note 4). 

5. To elimine cells from the sample, centrifuge the combined fluids at 800g for 5min 

at 4°C (see Note 5, 6, 7). 

6. To avoid sample degradation, add an adequat amount (1 tablet for 60 mL of BALF 

sample) of protease inhibitor cocktail tablets (Complete mini, Roche, Germany) 

(see Note 8). 

7. Store BALF at –80°C until use (see Note 9).


3.2. BALF Preparation for 2-DE 

Numerous BALF sample preparations have been used by different authors. 

Among these, precipitation (using trichloracetic acid or two-step combination 

of precipitant) is one of the most commonly used methodologies for protein 

analysis. However, limitations are observed with such a procedure because of a 

nonquantitative protein resolubilization after the precipitation step (13). Finally, 

the most efficient method of preparing a BALF sample before 2-DE analysis 

uses ultra filtration and dialysis-lyophilization (see Note 10). 

1. Rehydrate dialysis membranes (cut-off value 3.5 kDa) by soaking them in 

ultrapure water heated to about 90°C for 5 min or following manufacturer’s 

instructions. Length of the membrane must be adapted to the sample volume. 

Alternatively or if small volumes need to be dialyzed, ready to use dialysis 

devices are available commercially (Slide-A-Lyser 3.5K, Pierce). 

2. Rinse extensively the rehydrated membranes with ultrapure water. The 

membranes may not run dry. 

3. Close the dialysis tubes on one side and improve sealing with ultrapure water. 

4. Fill the membrane with BALF sample and close the second side of the dialysis 

tube. Check seal. 

5. Immerse the membrane tube in dialysis buffer (see Note 11) and place on a 

magnetic stirrer at 4°C for 3 h. 

6. Replace dialysis buffer twice. 

7. Harvest dialyzed BALF sample into an appropriate container. 

8. Reduce BALF sample volume to 20 μL in the Speed Vac (see Note 12). 

10. Suspend the sample in a minimum volume of sample buffer dedicated to 2-DE 

analysis or LC-MS method. 

11. Centrifuge at 18,000g for 15 min at 4°C to remove any unsolubilized material. 

12. Sample is now ready for protein assay and 2-DE or LC-MS analysis 

(see Note 13). 

Proteomic analysis of BALF is often hampered by the predominance of 

several highly abundant proteins including albumins and immunoglobulins. 

Depletion of these proteins is necessary before proteome analysis for detection 

of minor proteins. See other chapters in this book. 

3.3. BALF 2-DE analysis 

Here we propose the optimum conditions for obtaining high resolution and 

reliable 2-DE of BALF samples (see Note 14). Use of different pH gradients in 

IEF or reticulations in SDS-PAGE should be needed in narrower study of BALF 

proteins. Indeed, the high number of protein spots in the pH range 4.5–6.7 

are increased or decreased in different lung pathologies such as sarcoidosis 

or fibrosis (6,8,9). In this context, the use of narrow range IPG strips for


IEF improves the resolution of the separation and increases the probability of 

detecting less abundant proteins. 

1. Place the IPG strips in an adequate DryStrip Reswilling tray. 

2. Cover the IPG strips first with 500 μL of rehydration buffer and second with lowviscosity 

paraffin oil, and let the strips rehydrate overnight at room temperature. 

3. Remove the rehydrated IPG gels from the grooves, rinse them with ultrapure 

water, and place them, gel-side down, on water saturated filter paper. Filter 

papers are first blotted to remove excess water. 

4. Place rehydrated IPG strip (pH 3–10, 18 cm) in the Multiphor (Amersham 

Biosciences) or Protean IEF Cell (Bio-Rad) device set to 20°C. The correct 

settings of strips and cup loading can be achieved by following the manufacturer’s 

instructions. 

5. Apply 100 μg of protein/strip in cups at the anodic side of the IPG strip (see 

Note 15, 16). 

6. Increase voltage linearly from 300–5,000 V during the first 3 h and stabilize the 

voltage at 5,000 V for 20 h (see Note 17). 

7. After electrofocusing, place the IPG strips individually in capped glass tubes 

and equilibrate them for 20 min at room temperature in 10 mL of equilibration 

buffer containing 2% DTE (w/v) under gentle agitation. 

8. Replace buffer by 10 mL of fresh equilibration buffer containing 2% iodoacetamide 

(w/v) and incubate for 20 min at room temperature under gentle 

agitation. 

9. Run the second dimension on a 9–16% polyacrylamide linear gradient gel 

(18 × 20 × 1.5 cm) at 40 mA/gel constant current and 10°C. 

10. The gels are then ready to be stained using standard silver staining (see Note 18). 

4. Notes 

1. Human bronchoalveolar lavage is an invasive method that must be carried 

out under the informed consent of the concerned subjects and approved by a 

competent ethics committee. 

2. Human bronchoalveolar lavage requires topical lidocaine anaesthesia and 

endotracheal tube manipulation and must thus be performed by a competent 

physician in an adequate environment. 

3. Typically, the mean recovery of BALF is 55 % of the instilled volume. 

4. The first sample is separated from the others to avoid bronchial contamination. 

5. With this procedure, typical protein concentration of BALF ranges from 0.05 to 

1.20 mg/mL. 

6. During the BAL process, the cellular elements can secrete a variety of components. 

Therefore, the cells should be removed immediately to provide optimal 

proteome stability. 

7. After centrifugation, the phenotype of cells (macrophages, lymphocytes) can 

be analyzed. The normal cellular pattern of BALF contains mainly alveolar


macrophages, a small percentage of lymphocytes and less than 2% of 

polymorphs. This cellular pattern changes in many lung pathologies. 

8. Protease inhibitors are necessary but peptide inhibitors, e.g., high concentration 

of aprotinin, may interfere with mass spectrometry analysis. 

9. Samples should be divided into small aliquots before storage to avoid repeated 

freezing and thawing of samples. 

10. Desalting by dialysis and ultra membrane centrifugation are very effective 

techniques for salt removal, leading to minimal sample loss compared to precipitation 

or filtration methods. However, during membrane ultracentrifugation, 

protein adsorption onto the membrane surface is a problem that can be, in part, 

circumvented by repeated washing steps with sample solution after centrifugation 

(13). 

11. Bath dialysis volume must be 100× sample volume. 

12. Speed Vac centrifugation is an easier method that generates less protein loss 

than freeze-drying (13). 

13. If electrophoresis is not to be run at this time, store the lyophilized BALF at 

–80°C until used. Protein concentration of BAL fluid must be determined using 

a suited protein assay. 

14. IEF running conditions depend on the pH gradient and the length of the IPG gel 

strip used. Conditions presented here assume the use of nonlinear wide-range 

immobilized pH gradient (3–10) 18 cm long IPG strips (optimized for body 

fluids) and the Multiphor II electrophoresis system of Amersham Biosciences. 

We also recommend the second dimension to be run on linear gradient polyacrylamide 

gels (9–16%) for best resolution. 

15. Different methods can be used to apply BALF sample on IPG strips such as 

the sample cup method or during the strip rehydration process. Classically, to 

increase the loading capacity, and enhance the resolution of 2-DE, the entire 

IPG gel can be used for sample application, with the proteins entering the gel 

during rehydration. Nevertheless, the best BALF 2-DE quality is obtained using 

the cup loading approach. A recent new type of in-gel sample application named 

“paper bridge sample application” has been optimized for the BALF proteome 

analysis. This procedure allows increasing the loading capacity of BALF sample 

without loss of the 2-DE resolution (8). 

16. After IEF but before equilibration, strips may be stored at –80°C until the second 

dimension. 

17. The quantity may be varied according to the sensitivity of the staining method. 

18. Quantification of proteins is a major problem of the 2-DE approach, especially 

after silver staining. However, a new fluorescent protein labeling protocol (2D- 

DIGE) before electrophoretic separation has been developed. (See other chapters 

in this book). 


R. Wattiez is Research Associate of the Belgian FNRS. The authors thanks 

Catherine S’Heeren for her technical assitance.


References 

1. Wattiez, R. and Falmagne, P. (2005) Proteomics of bronchoalveolar lavage fluid. 

J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 815, 169–78. 

2. Baughman, R. P. and Drent, M. (2001) Role of bronchoalveolar lavage in interstitial 

lung disease. Clin. in Chest Med. 22, 331–41. 

3. Reynolds, H.Y. (2000) Use of bronchoalveolar lavage in humans—past necessity 

and future imperative. Lung 25, 271–93. 

4. Wattiez, R., Hermans, C., Bernard, A., Lesur, O., and Falmagne, P. (1999) 

Human bronchoalveolar lavage fluid: two-dimensional gel electrophoresis, amino 

acid microsequencing and identification of major proteins. Electrophoresis 20, 

1634–45. 

5. Noel-Georis, I., Bernard, A., Falmagne, P., and Wattiez, R. (2002) Database of 

bronchoalveolar lavage fluid proteins. J. Chromatogr. B Analyt. Technol. Biomed. 

Life Sci. 771, 221–36. 

6. Magi, B., Bini, L., Perari, M.G., Fossi, A., Sanchez, J.C., Hochstrasser, D., 

Paesano, S., Raggiaschi, R., Santucci, A., Pallini, V., and Rottoli, P. (2002) 

Bronchoalveolar lavage fluid protein composition in patients with sarcoidosis 

and idiopathic pulmonary fibrosis: a two-dimensional electrophoretic study. 


7. Lindahl, M., Stahlbom, B., and Tagesson, C. (1999) Newly identified proteins in 

human nasal and bronchoalveolar lavage fluids: potential biomedical and clinical 

applications. Electrophoresis 20, 3670–6. 

8. Sabounchi-Schutt, F., Astrom, J., Hellman, U., Eklund, A., and Grunewald, J. 

(2003) Changes in bronchoalveolar lavage fluid proteins in sarcoidosis: a 

proteomics approach. Eur. Respir. J. 21, 414–20. 

9. Noel-Georis, I., Bernard, A., Falmagne, P. and Wattiez, R. (2001) Proteomics as 

the tool to search for lung disease markers in bronchoalveolar lavage. Dis. Markers 

17, 271–8. 

10. Kriegova, E., Melle, C., Kolek, V., Hutyrova, B., Mrazek, F., Bleul, A., du 

Bois, R. M., von Eggeling, F. and Petrek, M. (2006) Protein Profiles of 

Bronchoalveolar Lavage Fluid from Patients with Pulmonary Sarcoidosis. Am. J. 

Resp. Crit. Care Med. 26, 1145–54. 

11. Lenz, A. G., Meyer, B., Costabel, U., and Maier, K. (1993) Bronchoalveolar lavage 

fluid proteins in human lung disease: analysis by two-dimensional electrophoresis. 


12. Lenz, A. G., Meyer, B., Weber, H., and Maier, K. (1990) Two-dimensional 

electrophoresis of dog bronchoalveolar lavage fluid proteins. Electrophoresis 11, 

510–3. 

13. Plymoth, A., Lofdahl, C. G., Ekberg-Jansson, A., Dahlback, M., Lindberg, H., 

Fehniger, T. E., and Marko-Varga, G. (2003) Human bronchoalveolar lavage: 

biofluid analysis with special emphasis on sample preparation. Proteomics 

3, 962–72. 

14. Wu, J., Kobayashi, M., Sousa, E., Lieu, W., Cai, J., Goldman, S. J., Dorner, A. J., 

Projan, S. J., Kavuru, M. S., Qiu, Y., and Thomassen, M. J. (2005) Differential


proteomic analysis of bronchoalveolar lavage fluid in asthmatics following 

segmental antigen challenge. Mol. Cell. Proteomics 4, 1251–64. 

15. Chromy, B., Gonzales, A., Perkins, J., Choi, M., Corzett, M., Chang, B. C., 

Corzett, C. H., and McCutchen-Maloney, S. L. (2004) Proteomic analysis of 

human serum by two-dimensional differential gel electrophoresis after depletion 

of high-abundant proteins. J. Proteome Res. 6, 4–8.

7 

Preparation of Nasal Secretions for Proteome Analysis 

Begona Casado, Paolo Iadarola, and Lewis K. Pannell 

Summary 

The determination of protein patterns in nasal secretions of healthy subjects can help in 

the early diagnosis of diseases such as acute sinusitis. The comparison of nasal lavage fluid 

collected from subjects with acute sinusitis before and after pharmacological treatment 

gives information about the drug effects on glandular secretions. Nasal secretions were 

stimulated with 1× NS (0.9% Normal Saline) and 24× NS in healthy subjects and in 

sinusitis subjects before and after pharmacological treatment. The nasal lavage fluid 

(NLF) proteins are precipitated with a solution of “acid-ethanol.” Using this solution, the 

high molecular weight proteins precipitate and separate from the low molecular weight 

proteins. The proteins are digested and the peptides are separated using a capillary liquid 

chromatographic system. Eluted peptides are analyzed on ESI-Q-TOF mass spectrometry 

instrument. 

Key Words: CapLC-ESI-Q-ToF; liquid-liquid extraction; Nasal secretions; pharmacological 

treatment; proteomics; sample preparation; sinusitis. 


Nasal secretions (NS) are a barrier against pathogenic (e.g., bacteria and 

viruses) and nonpathogenic (e.g., fine particles) antigens that are present in 

the air and are breathed in through the nose. NS serve to humidify, heat 

or cool, and clean inhaled air and contain proteins of the innate immune 

system. These proteins are from plasma, glandular mucous and serous cells (1,2) 

and their release is started during allergen exposure, rhinovirus, adenovirus, 

influenza, bacterial rhinosinusitis, cystic fibrosis, and occupational exposure. 

The hyperresponsiveness is a typical characteristic of inflamed mucosa and 

airways (3–5) although different molecular mechanisms may be involved in 

allergic, infectious, and nonallergic disorders. The release of nasal secretions, 



77

78 Casado et al. 

is generally induced by spraying a solution of normal (S) and hypertonic 

saline (HTS). The latter, in humans, is an airway irritant of nasal mucosa by 

stimulating nociceptive nerves and glandular secretion. For example, substance 

P is released in the mucosa after a neural depolarization with local axon 

responses caused by hypertonic saline stimulation (6,7). HTS nasal provocations 

have been employed also to understand the different neural response between 

acute sinusitis, acute rhinitis, and the nonallergic rhinitis present in subjects 

with chronic fatigue syndrome (CFS) (8,9). In the last 4 years, the interest of 

the scientific community on nasal mucosa and nasal secretions has increased. 

The collection of nasal lavage, in fact, is a simple way for obtaining samples 

from the upper airways and may be performed using noninvasive procedures. 

For example, the agent responsible for the Creutzfeldt-Jakob disease can be 

identified “in vivo” in nasal mucosa (10), and an anthrax vaccine based on 

the use of an anthrax protective antigen (PA) protein carried by liposomeprotamine-DNA 

(LPD) is nasally dosed in mice (11). 

The complete knowledge of the nasal secretion constituents has not been 

achieved yet. It is apparent that the identification of specific mucous protein 

profiles may help elucidate the different mechanisms involved in host defense. 

To date the determination of mucus protein profiles can be achieved using 

a proteomic procedure. Different proteomic approaches have been used so 

far to analyze nasal lavage fluid (NLF). Lindahl et al. used two dimensional 

electrophoresis (2-DE) with matrix-assisted laser desorption/ionization 

(MALDI) mass spectrometry to analyze either the proteome of NLFs 

from subjects exposed to methyltetrahydrophthalic anhydride (MHHPA) 

or dimethylbenzylamine (DMBA), and that from healthy nonsmokers and 

smokers (12–18). Another approach has been described by Casado et al. who 

applied liquid chromatography (LC) with electrospray ionization (ESI) mass 

spectrometry to analyze NLFs of subjects affected by acute sinusitis before 

and after pharmacological treatment, and for the comparison of NLFs of 

normal subjects before and after nasal provocation (19,20). Kristiansson et al. 

have used the same procedure for the analysis of HHPA-(hexahydrophthalic 

anhydride) adducted albumin tryptic peptides in nasal lavage fluid as 

biomarkers of exposure (21,22). These two complementary approaches 

provided new information on proteins involved in host protection and defence 

against microorganisms and occupational exposure. 

2. Materials 

2.1. Pharmacological Treatment 

1. Antibiotic: amoxicillin-clavulanic acid (Augmentin, GlaxoSmithKline, Research 

Triangle Park, NC, USA).

Preparation of Nasal Secretions for Proteome Analysis 79 

2. Steroid: fluticasone propionate nasal spray (Flonase, GlaxoSmithKline, Research 

Triangle Park, NC, USA). 

3. 2 adrenergic agonist vasoconstrictor: oxymetazoline nasal spray (Super G, 

Landover, MD, USA). 

4. United States Pharmacopea normal saline: saline nasal spray (0.9% NaCL) (Abbott 

Laboratories, IL, USA). 

2.2. Nasal Provocation 

1. United States Pharmacopea normal saline: saline nasal spray (0.9% NaCL) (Abbott 

Laboratories, IL, USA). 

2. Hypertonic saline 21.6% 

3. Beconase AQ pump aspirator spray device (23,24) (Glaxo-Wellcome, Triangle 

Park, NC, USA). 

4. Nasal secretion collection: 5-ounce Dixie wax-paper cup (James River Corp., 

Norwalk, CT, USA) or polypropylene beakers (Fisher Scientific, Fair Lawn, 

NJ, USA). 

2.3. Nasal Lavage Fluid Preparation for Liquid Chromatography 

and Mass Spectrometry Analysis 

1. Protein assay on 96-well micro plates using MRX Microplate Reader Instrument 

(Dynex Technologies, Chantilly, VA, USA). 

2. Bovine albumin as standard protein for total protein assay (Sigma, MO, USA). 

3. Acid-ethanol solution: 50% 0.2N acetic acid, 50% ethanol, 0.02% sodium bisulfite 

(25) stored at 4°C. Ethanol and acetic acid obtained from Fisher Scientific 

(Fair Lawn, NJ, USA), and sodium bisulfite from Mallinckrodt Laboratory 

Chemicals (Phillipsburg, NJ, USA). 

4. Protein digestion: sequencing grade modified trypsin (Promega, Madison, WI, 

USA). 

5. Digestion buffer: 0.1M ammonium bicarbonate (pH 7.8) (Sigma, St. Louis, 

MO, USA). 

2.4. Liquid Chromatography and Mass Spectrometry Analysis 

of Nasal Lavage Fluid 

1. Desalt and concentration: 35 × 0.32 mm BioBasic C18 precolumn ( Thermo 

Hypersil-Keystone, Bellefonte, PA, USA). 

2. Peptide separation: Reverse-phase Zorbax C18 column (100 mm × 150 μm id) 

(Micro-Tech Scientific, Sunnyvale, CA, USA). 

3. Solvent A: HPLC grade H2O with 0.2% formic acid (Fisher Scientific, 

Fair Lawn, NJ, USA). 

4. Solvent B: HPLC grade acetonitrile with 0.2% formic acid (Fisher Scientific, 

Fair Lawn, NJ, USA).


5. Capillary LC instrument (Waters Inc, Milford, MA, USA). 

6. Electrospray-Quadrupole-Time of Flight mass spectrometer (Waters Inc, Milford, 

MA, USA). 

3. Methods 

3.1. Nasal Provocation 

1. Subjects’ nasal cavities need to be pre-washed with 24 sprays (100 μL each 

nostril) of sterile normal saline (1 × NS, 0.9% NaCl) using a Beconase AQ 

pump aspirator spray device. 

2. Subjects gently blow out through their noses, and the lavage fluid from both 

nostrils into a 5-ounce Dixie wax-paper cup. This material is discarded. 

3. 10 min later step 2 is repeated using 12 sprays of 1 × NS, and the lavage fluid 

discarded. Now the nasal provocation is performed. 

4. 100 μL of 0.9% normal saline is administered separately into each nostril. 

5. After 5 min, NLF is collected using 12 sprays of 0.9% NS (see Note 1). 

6. The NLF must gently blow out into a cup. NLF from left and right nostrils are 

mixed together (first series) (see Note 2). 

7. Immediately after this collection, the same subject is sprayed with hypertonic 

saline (HTS) (21.6% NaCl, 24 times the tonicity of NS (24 × NS). The pH of 

HTS (freshly prepared solution with double distilled deionized water) is 6.07 

(see Note 3). 

8. After 5 min, 12 puffs of NS are sprayed into each nostril. 

9. The NLF must gently blow out into a cup. NLFs from left and right nostrils are 

mixed together (second series). 

10. After pharmacological treatment on day 6, the first and second series for day 6 

are collected repeating the procedure indicated in steps 1–9. 

11. Lavage fluids are gently shaken to disperse mucous globules and pipetted into 

Eppendorf tubes. Samples are frozen at –20°C until analysis is performed. 

3.2. Nasal Lavage Fluid Preparation 

3.2.1. Total Protein Assay 

1. Measure the total protein concentration in each sample using modified Lowry’s 

method (see other chapters in this book). 

2. Place standard human albumin or real samples (10 μL) in triplicate in polystyrene 

microtiter plates, and add assay reagents. 

3. Measure the optical densities (650 nm) with a microtiter-plate reader. 

4. Interpolate the protein concentrations in the samples from the regression analysis 

of the standard curve (protein concentration in normal NLF: 1st series 878 μg/mL 

and 2nd series 1,700 μg/mL; protein concentration in sinusitis NLF: day 1 1st 

series 1,321 μg/mL and 2nd series 1,512 μg/mL, and day 6 1st series 638 μg/mL 

and 2nd series 725 μg/mL).


3.2.2. Acid-Ethanol Precipitation 

1. 30 μL of nasal lavage fluid are mixed with an equal volume of 50% ethanol, 50% 

0.2 N acetic acid, 0.02% sodium bisulfite. 

2. Leave the protein to precipitate at –20°C overnight (see Note 4). Samples are 

stable in acid-ethanol solution for long time if stored at –20°C. 

3. Centrifuge the mixture for 30 min at 4°C. The supernatant containing endogenous 

peptides, lipids and sugars is discarded. 

3.2.3. Protein Digestion 

1. Prepare a fresh solution of 0.1 M ammonium bicarbonate pH 7.8. 

2. Dissolve the protein pellet in 10 μL of 0.1 M ammonium bicarbonate pH 7.8 

vortex until the protein pellet is dissolved. 

3. Dissolve 25 μg of trypsin in 25 μL of ammonium bicarbonate buffer (see Note 5). 

4. Trypsin is added to the samples in trypsin:protein ratio of 1:20 (w/w). 

5. Incubate the solution at 37°C overnight. 

6. Inactivate the trypsin by adding 1 μL of 0.1% formic acid. 

3.3. Analyis of Nasal Lavage Fluid Preparations by Liquid 

Chromatography Coupled to Mass Spectrometry 

Electrospray has the advantage of ionizing macromolecules in a liquid. The 

ions observed are formed by addition of proton (hydrogen ion) to give the [M+H] 

ion in which M = analyte molecule, H = hydrogen ion. For large macromolecules 

(such as peptides) there will often be a distribution of many charge states. 

1. Same amounts of tryptic peptides are injected into a capillary liquid chromatography 

(CapLC) system after testing the LC system (see Note 6). 

2. Peptide mixture are concentrated and desalted on a BioBasic C18 precolumn 

applying an isocratic procedure (95% water in 0.2% formic acid (FA)) with a 

flow rate of 20 μL/min for 10 min (see Note 7). 

Table 1 

m/z, charge state, mean and SD of elution time, and CV of five chosen tryptic 

peptides from albumin from NLF. 

m/z Charge state Time (min) (X+±) CV (%) 

682.38 +3 47±0.9 1.8 

812.41 +2 61±0.9 1.5 

693.82 +2 43±1.2 2.8 

671.83 +2 47±1.3 2.8 

575.32 +2 34±0.9 2.6


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

Fig. 2. The histograms showing the proteins tabulated according biological function 

and origin, found in sinusitis NLFs pre- (Day1) and post- (Day6) pharmacological 

treatment.


3. The peptides are separated on a Zorbax C18 column (100 mm x 150 μm I.D.) 

using a gradient from 95% water in 0.2% FA to 95% acetonitrile in 0.2% FA 

over 100 min. The flow is set at 10 μL/min. A splitter is used to carry 1 μL/min 

on the analytical column (see Note 8). 

4. Separated peptides are analyzed using Electrospray-Quadrupole-Time of Flight 

mass spectrometer. The samples are run in duplicate. The reproducibility of elution 

times is determined by comparison with the retention time of five tryptic peptides 

of endogenous albumin (Table 1). Fig. 1 shows a total ion chromatogram (TIC) 

profile of nasal lavage fluid (see Note 9). 

Table 2 

Keratins from normal NLF are reported. We detect type I and type II reflecting 

the spectrum of cutaneous, transitional, and type I and II form heterodimers in 

intermediate fibers respiratory mucosal cells. Sinusitis had a more limited 

spectrum with k1, k5, k6a, k6f, k10, and k13 

Keratin Epithelium Location 

k1–k2 and k9–k10 epidermis anterior nares 

k6, k4, k16 terminally differentiated 

squamous cells 

anterior nares 

k1–k10 hairs nasal vestibule 

k25 inner root sheaths 

k6, k16 outer root sheath’s inner layer 

k5–k14 outer layer and sebaceous 

glands 

k8–k18, k7, k19 transitional cuboidal 

epithelium and 

pseudoatratified respiratory 

epithelium 

nasopharynx 

k5–k14 basal cells (progenitors of 

respiratory epithelium 

from k5–k14 to k1–k10 differentiation of 

pseudostratified respiratory 

epithelial suprabasal cells 

k6, k16 or (k17) wet stratified squamous 

epithelial lining epithelial 

oral and esophageal 

invaginations of submucosal 

glands and ducts 

mucoseae 

k5–k14 myoepithelial cells surround 

submucosal glands


3.4. Data Interpretation 

1. The protein identification can be performed using MASCOT MS/MS ion search 

software (see Note 10). 

2. If the spectra are manually sequenced, the new peptide sequence can be matched 

to a protein using peptide match program in the protein identification resource 

(PIR, www.pir.georgetown.edu). 

3. PIR BLAST similarity search is used to search unknown protein query sequences. 

4. Protein sequence alignments are constructed using the CLUSTAW program 

in PIR. 

5. The identified proteins are compared on the basis of their origin and function 

(26). In Fig. 2, the proteins identified before and after pharmacological treatment 

were grouped according the biological function and origin. All the inflammatory 

proteins were identified only on Day 1 and not on Day 6. We can hypothesize that 

the treatment was successful and blocked the influx of inflammatory cells (e.g., 

IL-16 and IL-17E), the generation of their mediators (e.g., TGF- 2 receptor), 

vascular permeability, and glandular hypersecretion. On Day 1, keratins associated 

with respiratory epithelium and the squamous metaplasia present in sinusitis were 

detected. Different actin protein (actin , 1, and 2) reflect the hyperplasia 

of the respiratory epithelium. Keratin proteome in normal NLF reflected the 

anticipated normal type of basal, pseudostratified, respiratory, glandular, and 

stratified nonkeratinized and keratinized squamous epithelium that is present in 

the nose. A large number of keratins have been detected in normal NLF then in 

sinusitis NLF. Keratin profile in sinusitis NLF was consistent with desquamation 

of terminally differentiated cells and the presence of squamous metaplasia. The 

results demonstrate the changes that are taking place in respiratory epithelial cells 

during inflammation (Table 2). 

4. Notes 

1. The subjects pressed their left nostrils closed, and then spritzed 12 sprays of 

1× NS into their right nostrils. 

2. Because the amount of nasal secretions blown out from left and right nostrils 

are different, it is important to mix the two samples together. Because both 

nostrils are not always simultaneously closed or open, this situation may cause 

a different pattern of proteins. It is very important to mix the samples as, 

mixing the specimens from the two nostrils, the sample homogeneity is strongly 

improved. Big globules of mucus must be dissolved to liberate the proteins in 

the mucous net. 

3. Hypertonic saline stimulates the glandular secretions, local mucosal substance 

P release and pain. Because it is a provocation, the presence of a physician is 

recommended. 

4. Acid-ethanol solution precipitates high molecular weight proteins. Supernatant 

contains peptides that can be used for following determinations. Although


proteins precipitate in few hours, it is recommended to carry out the procedure 

overnight. 

5. If the trypsin stock is not entirely used, we suggest resuspending it in acetic acid 

to prevent autodigestion of trypsin. 

6. Before injecting the NLF samples, it is important to test the LC system. one 

microliter of nine peptide mixtures (neurotensin, angiotensin I, angiotensin 2, 

Glu-fibrinopetide B, somatostatin, bradykynin, bombesin, enkephalin, and 

substance P; 1 pmol of each peptide) is injected on LC system. Resolution and 

sensitivity can be checked. 

7. During peptide concentration and desalting, small (3–4 amino acids) and highly 

hydrophilic peptides are washed off from the precolumn. Those losses can 

prevent damage of the column. 

8. It is important to look carefully at the samples before injecting them on the 

CapLC to check if samples are contaminated with mucous globules. This material 

in fact can block the capillary causing the damage to the system. For LC- 

ESI-MS/MS formic acid replaces trifluoroacetic acid (TFA) in the LC mobile 

phase because an efficient ionization is prevented by the strong ion pairing 

characteristics of TFA. 

9. Duplicate and triplicate runs are necessary to examine the reproducibility of the 

elution. Two options are available for checking consistency of the chromatographic 

system. First, it is possible to spike the sample with one or more standard 

peptides. Second, it is possible to choose tryptic peptides from an endogenous 

protein present in the sample. You must know before analyzing your sample 

which abundant protein is present in the sample and if it is easily cleaved by 

the enzyme you are using. Those peptides need to be consistently present in 

your runs. We choose the second option and we looked at five peptides from 

albumin. The mean, standard deviation of retention time, and CV % of the five 

peptides are calculated to see how reproducible the experiments are. 

10. The program can be found in the web (www.matrixscience.com). There is a 

disadvantage in using MASCOT in the web. Only the first 300 peptides can be 

searched on the database. Using the nonrestricted MASCOT more information 

can be retrieved from the raw data. The following general search parameters 

were used: monoisotopic molecular masses, enzyme trypsin, peptide tolerance 

of ± 0.4 Da and MS/MS tolerance of ± 0.3 Da. The search is restricted to Homo 

sapiens species to make the search easier. 

References 

1. Baraniuk, J. N. (2000) Immunology and Allergy Clinics of North America 

(Lasley, M. and V. Altman, L. C. eds.), Saunders, Philadelphia, pp. 245–64. 

2. Baraniuk, J. N., Staevska, M. (2004) Current Therapy in Allergy Immunology and 

Rheumatology (Lichtenstein, L. M., Busse, W. W. and Geha, R. S., eds.), Mosby, 

Philadelphia, pp. 17–24.


3. Meyer, P., Andersson, M., Persson, C.G., and Greiff, L. (2003) Steroid-sensitive 

indices of airway inflammation in children with seasonal allergic rhinitis. Pediatr. 

Allergy Immunol. 14, 60–65. 

4. Dahl, R., and Mygind, N. (1998) Mechanisms of airflow limitation in the nose and 

lungs. Clin. Exp. Allergy 28, 17–25. 

5. Svensson, C., Andersson, M., Greiff, L., and Persson, C.G. (1998) Nasal mucosal 

end organ hyperresponsiveness. Am. J. Rhinol. 12, 37–43. 

6. Baraniuk, J. N., Ali, M., Yuta, A., Fang, S-Y., and Naranch, K. (1999) Hypertonic 

saline nasal provocation stimulates nociceptive nerves, substance P release, and 

glandular mucus exocytosis in normal humans. Am. J. Respir. Crit. Care Med. 

160, 655–62. 

7. Sanico, A. M., Philip, G., Lai, G. K., and Togias, A. (1999) Hyperosmolar saline 

induces reflex nasal secretions, evincing neural hyperresponsiveness in allergic 

rhinitis. J. Appl. Physiol. 86, 1202–10. 

8. Baraniuk, J. N., Clauw, D. J., and Gaumond, E. (1998) Rhinitis symptoms in 

chronic fatigue syndrome. Ann. Allergy Asthma Immunol. 81, 359–65. 

9. Fukuda, K., Straus, S. E., Hickei, I., Sharpe, M. C., Dobbins, J. C., and Komaroff, A. 

(1994) The chronic fatigue syndrome: a comprehensive approach to its definition 

and study. Ann. Intern. Med. 121, 953–9. 

10. Tabaton, M., Monaco, S., Cordone, M. P., Colucci, M., Giaccone, G., Tagliavini, F., 

and Zanusso, G. (2004) Prion deposition in olfactory biopsy of sporadic 

Creutzfeldt-Jakob disease. Ann Neurol. 55, 294–6. 

11. Sloat, B. R., and Cui, Z. (2005) Strong mucosal and systemic immunities 

induced by nasal immunization with anthrax protective antigen protein incorporated 

in liposome-protamine-dna particles. Pharm Res. Dec 6; [Epub ahead 

of print] 

12. Lindahl, M., Stahlbom, B., and Tagesson, C. (1995) Two-dimensional gel 

electrophoresis of nasal and bronchoalveolar lavage fluids after occupational 

exposure. Electrophoresis 16, 1199–1204. 

13. Lindahl, M., Stahlbom, B., Svartz, J., and Tagesson, C. (1998) Protein patterns of 

human nasal and bronchoalveolar lavage fluids analyzed with two-dimensional gel 

electrophoresis. Electrophoresis 19, 3222–29. 

14. Lindahl, M., Stahlbom, B., and Tagesson, C. (1999) Newly identified proteins in 

human nasal and bronchoalveolar lavage fluids: potential biomedical and clinical 

applications. Electrophoresis 20, 3670–76. 

15. Lindahl, M., Svartz, J., and Tagesson, C. (1999) Demonstration of different forms 

of the anti-inflammatory proteins lipocortin-1 and Clara cell protein-16 in human 

nasal and bronchoalveolar lavage fluids. Electrophoresis 20, 881–90. 

16. Ghafouri, B., Stahlbom, B., Tagesson, C., and Lindahl, M. (2002) Newly identified 

proteins in human nasal lavage fluid from non-smokers and smokers using twodimensional 

gel electrophoresis and peptide mass fingerprinting. Proteomics 2, 

112–20. 

17. Lindahl, M., Stahlbom, B., and Tagesson, C. (2001) Identification of a new 

potential airway irritation marker, palate lung nasal epithelial clone protein, in


human nasal lavage fluid with two-dimensional electrophoresis and matrix-assisted 

laser desorption/ionization-time of flight. Electrophoresis 22, 1795–1800. 

18. Lindahl, M., Irander, K., Tagesson, C., and Stahlbom, B. (2004) Nasal lavage fluid 

and proteomics as means to identify the effects of the irritating epoxy chemical 

dimethylbenzylamine. Biomarkers 9, 56–70. 

19. Casado, B., Pannell, L. K., Viglio, S., Iadarola, P. et al. (2004) Analysis of 

the sinusitis nasal lavage fluid proteome using capillary liquid chromatography 

interfaced to electrospray ionization-quadrupole time of flight- tandem mass 

spectrometry. Electrophoresis 25, 1386–93. 

20. Casado, B., Pannell, L. K., Iadarola, P., Baraniuk, J.N. (2005) Identification of 

human nasal mucous proteins using proteomics. Proteomics 5, 2949–59. 

21. Kristiansson, M. H., Lindh, C. H., and Jonsson, B. A. (2003) Determination of 

hexahydrophthalic anhydride adducts to human serum albumin. Biomarkers 8, 

343–59 

22. Kristiansson, M. H., Lindh, C. H., and Jonsson, B. A. (2004) Correlations between 

air levels of hexahydrophthalic anhydride (HHPA) and HHPA-adducted albumin 

tryptic peptides in nasal lavage fluid from experimentally exposed volunteers. 

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Allergy Immunol. 103, 202–8. 

25. Baraniuk, J. N., Okayama, M., Lundgren, J. D., Mullol, M. et al. (1990) Vasoactive 

intestinal peptide in human nasal mucosa. J. Clin. Invest. 86, 825–31. 

26. Wu, H. C. H., Huang, H., Yeh, Lai-Su, L., Barker, C. W. (2003) Protein family 

classification and functional annotation. Comput. Biol. Chem. 27, 37–47.

8 

Preparation of Urine Samples for Proteomic Analysis 

Rembert Pieper 

Summary 

Reproducible procedures for the preparation of protein samples isolated from human 

urine are essential for meaningful proteomic analyses. Key applications are the discovery 

of novel proteins or their modifications in the human urine as well as protein biomarker 

discovery for diseases and drug treatments. The methodology presented here features 

experimental steps aimed at limiting protein losses because of organic solvent precipitation, 

effective separation of proteins from other compounds in the human urine 

and molecular weight-based enrichment of proteins in two distinct fractions. Urinary 

proteins are separated from cellular debris in the urine via centrifugation, concentrated 

with 5-kDa-cutoff membrane concentration devices and separated via size exclusion 

chromatography into fractions with a higher and a lower molecular weight than 30 kDa, 

respectively. A successive optional affinity removal step for highly abundant plasma 

proteins is described. Finally, buffer exchange steps useful for specific downstream 

proteomic analysis experiments of urinary proteins are presented, such as 2-dimensional 

gel electrophoresis, differential protein or peptide labeling and digestion with trypsin for 

LC-MS/MS analysis. 

Key Words: Biomarker discovery; gel electrophoresis; human urine; multidimensional 

liquid chromatography; proteomic sample preparation; urinary proteome. 


Human urine plays a central role in clinical diagnostics. The human urinary 

proteome has been investigated particularly in the context of renal and bladder 

malfunction and cancer (1–6). Under normal physiological conditions, small 

protein amounts are excreted with the urinary fluid (0.5–5 mg per voiding), 

because the kidney restricts passage of plasma proteins, particularly in the 



89

90 Pieper 

M r range above 40 kDa during the filtration process in the glomeruli. More 

protein is lost in diseases, particularly those affecting the kidney, leading to 

proteinurea (7). Marshall and Williams as well as Anderson et al. pioneered the 

research in the characterization of the human urinary proteome in the eighties 

and nineties including sample preparation methods to isolate proteins from other 

matter in the urinary fluid (2,5,8–13). The most frequently used urine sample 

preparation methods for proteomic analysis are based on selective protein 

precipitation (12–17) or ultrafiltration and molecular weight-based enrichment 

steps (3,11,15,16,18). Urinary sample preparation should be performed at 

4°C and in the presence of protease inhibitors to avoid protein degradation. 

Cellular debris in urinary fluid should be removed before protein enrichment 

to avoid contamination with cellular proteins. Urine concentration is required 

to effectively separate proteins in size exclusion chromatography (SEC) experiments 

into protein fractions of distinct M r ranges. Immunoaffinity subtraction 

(IAS) permits the selective removal of highly abundant plasma proteins in 

urine concentrates and enrichment of lower abundance urinary proteins (3,19). 

Concentrated or lyophilized urinary protein samples are eventually prepared in 

buffers compatible with a variety of proteomic analysis techniques. 

2. Materials 

2.1. Urinary Protein Concentrate Preparation 

1. 250-mL conical bottom polypropylene centrifugation tubes (Fisher Scientific). 

2. CompleteTM protease inhibitor cocktail tablets (Roche, Indianapolis, IN). 

3. Centricon ® Plus-80 (5,000 NMWL) centrifugal filter devices (Millipore, 

Billerica, MA). 

4. Amicon ® Ultra-4 (5,000 NMWL) centrifugal filter devices (Millipore, 


5. Swinging bucket rotor with 250-mL conical tube adaptors and centrifuge for 

velocities up to 4,000g (Beckman-Coulter, Fullerton, CA). 

6. Buffer A: 100 mM sodium phosphate, pH 7.0, 150 mM NaCl, 0.02% sodium 

azide. 

7. BCA assay reagents (Pierce Chemicals, Rockford, IL). 

2.2. Size Exclusion Chromatography of Urinary Protein Concentrates 

1. HiLoad 16/60 Superdex 75 prep grade column (GE Healthcare, Piscataway, NJ). 

2. Liquid chromatography system (FPLC) with fraction collector adjustable to 4°C. 

3. Centricon ® Plus-20 (5,000 NMWL) centrifugal filter devices (Millipore, 


4. Broad range gel filtration standard (Bio-Rad, Hercules, CA) (see Note 1). 

5. Buffer B: 25 mM ammonium bicarbonate, 1 mM benzamidine, 1 mM Na-EDTA.

Preparation of Urine Samples for Proteomic Analysis 91 

2.3. Immunoaffinity Subtraction of Proteins 

1. Multiple Affinity Removal Spin Cartridge for the Depletion of High Abundance 

Proteins in Human Serum (Agilent Technologies) or Vivapure Anti-HSA/IgG 

Removal Kit (Sartorius AG) or Affinity Depletion Cartridge for Removal of HSA 

and Immunoglobulins from Human Serum (Applied Biosystems). 

2. Elution buffer: 0.5% glycine, 0.25% CHAPS, 150 mM NaCl, 2M urea, pH 2.5. 

3. Ultrafree ® -CL 0.45-μm centrifugal filter devices, (Millipore, Billerica, MA) 

2.4. Final Sample Preparation for Proteomic Analysis 

1. IPG rehydration solution: 8M urea, 2M thiourea, 4% CHAPS, 18 mM DTT 

and 0.5% Bio-Lyte ® pH 3–10 carrier ampholytes (Bio-Lyte ® is from Bio-Rad, 

Hercules, CA). 

2. Freeze-dry/lyophilization unit (vacuum pump, evacuable centrifuge, cold trap). 

3. Methods 

An overview of sample preparation and fractionation steps for urinary 

proteins is provided in Fig. 1. The schematic also shows downstream applications 

for urinary proteome analysis. 

3.1. Urinary Protein Concentrate Preparation 

1. The urine sample is collected, e.g., from a patient in a clinical laboratory, and 

transferred into a 250-mL tube with a conical bottom. It is cooled on ice and 

two CompleteTM protease inhibitor cocktail tablets are added to minimize protein 

degradation. The sample tube is centrifuged at 3,000g for 60 min at 4°C (see 

Note 2). In order not to disturb the precipitate, the supernatant is pipeted carefully 

into a new polypropylene tube. It can be frozen and stored for days at –80°C 

or processed immediately. The precipitate containing cellular debris and other 

insoluble matter is discarded. 

2. The supernatant is transferred to a Centricon ® Plus-80 device and spun at 3,000g 

at 4°C, until the urine sample volume is reduced to approx 4–5 mL (see Note 3). 

This sample is collected and transferred into an Amicon ® Ultra-4 centrifugal filter 

device. It is concentrated to 1 mL by spinning at 4,000g at 4ºC, rediluted with 

buffer A to 4 mL and reconcentrated to approx 500 μL. 

3. The protein concentrate is transferred to a 1.5-mL microtube and usually has 

a brownish color. It can be frozen and stored for days at -80°C or processed 

immediately. 

4. The urinary protein concentrate is spun at 10,000g for 15 min at 4°C. The 

supernatant of the centrifugation step is recovered and the pellet discarded. The 

protein amount is measured and the sample is ready to be subjected to the size 

exclusion chromatography experiment.

92 Pieper 

Fig. 1. Overview of urinary protein sample preparation procedures. 1. Removal of 

precipitates via centrifugation at 3,000g; 2. Concentration of urine in Centricon ® Plus- 

80 centrifugal filter devices; further concentration in Amicon ® Ultra-4 centrifugal filter 

devices; 3. Size exclusion chromatography (Superdex 75) generating two fraction pools 

(SEC ≤30 kDa fraction and >30 kDa fraction); 4. Reconcentration of SEC samples; 

5. Immunoaffinity subtraction generating the IAS >30 kDa fraction; 6. Final sample 

concentration in Amicon ® Ultra-4 centrifugal filter devices. Downstream applications 

for proteomic analysis: 2-DE gel electrophoresis; digestion with trypsin (followed 

by LC-MS/MS analysis); LC separation of urinary proteins; covalent (isotope-coded) 

labeling of urinary proteins for differential quantitation using MS methods. 

5. For protein quantitation using the BCA assay, 1- or 2-μL aliquots are transferred 

to a microtiter plate, incubated for 10 min with 100 μL BCA solution at 37ºC and 

measured in a spectrophotometer at =A 562. In parallel, a BCA assay standard 

curve with concentrations of 0.25 to 2 mg/mL bovine serum albumin is generated 

to calculate the total protein amount in the urinary sample. 

3.2. Size Exclusion Chromatography of Urinary Protein Concentrate 

1. The 16/60 Superdex 75 column is equilibrated in buffer A using an FPLC system 

at 4ºC in a cold cabinet (see Note 4). Once a stable baseline is observed monitoring 

UV light absorption at =A280, the initial experiment pertains to the molecular


weight (M r) column calibration. This experiment should be repeated on separate 

days to ascertain reproducibility. 

2. 100 μL (approx 3.5 mg protein) of the Bio-Rad gel filtration standard (M r range 

from 670 to 1.4 kDa) are loaded into the sample loop. The flow rate for the LC 

separation is 0.5 mL/min and fractions are eluted in volumes of approx 4–5 mL. 

As shown in the chromatogram of Fig. 2A, resolved LC peaks appear for the gel 

filtration standard proteins with the exception of the two high M r proteins (670 

and 150 kDa), which elute as a double peak. 

3. Using an X/Y scatter diagram with the M r units in logarithmic scale, the graphic 

display of M r values and elution volumes should yield a nearly linear fit and 

enable the determination of the elution volume corresponding to the M r of 30 

Fig. 2. Size exclusion chromatography of urinary protein concentrates on a Superdex 

75 column. Chromatogram A: Bio-Rad gel filtration standard with thyroglobulin (670 

kDa) and Ig G (150 kDa) in double peak 1, ovalbumin (45 kDa) in peak 2, myoglobin 

(18 kDa) in peak 3 and vitamin B12 (1.4 kDa) in peak 4. B and C: two urinary protein 

concentrates, fractions 3–7: SEC >30 kDa sample pool; fractions 8–14: SEC ≤ 30 kDa 

sample pool. The UV 280 traces were monitored. This Figure has been reproduced with 

permission 3 .

94 Pieper 

kDa (see Note 5). The fraction number corresponding to this elution volume is 

determined. 

4. After the protein concentration measurement of urinary samples using the BCA 

assay, the sample volume equivalent to an amount of 4 mg protein is determined. 

If the 500 μL urinary concentrate contains more than 4 mg protein, the appropriate 

sample volume is aliquoted and re-diluted to 500 μL with buffer A, while freezing 

the remaining concentrate. If there is less than 4 mg protein, all of the urinary 

protein concentrate is applied to the LC experiment. This sample is kept on ice 

before application to the SEC experiment. 

5. Using the same LC column, LC method, and fraction collector settings, urinary 

protein concentrates are loaded onto the 16/60 Superdex 75 SEC column and 

fractionated. As shown in the chromatograms of Fig. 2B and C for two different 

urinary protein samples, A 280 elution traces may vary from sample to sample. 

6. The fractions should be placed on ice after collection and combined into two 

fraction pools: (1) the fraction pool with proteins corresponding to a M r higher 

than 30 kDa (SEC >30 kDa) and (2) the fraction pool with proteins corresponding 

toaM r equal to and lower than 30 kDa (SEC ≤30 kDa). The 30 kDa M r fraction 

itself is added to the latter fraction pool. No fractions are collected in the baseline 

area (A 280 = 0), usually for fractions collected before the LC peak for the 670/150 

kDa gel filtration standards and after elution of the LC peak for the 1.4 kDa 

standard (peaks 1 and 4 in Fig. 2A, respectively). 

7. The two urinary protein sample pools are concentrated in Centricon ® Plus-20 

units to approx 1 mL. If fraction pool volumes are larger than 20 mL, concentrate 

in a stepwise process in the same Centricon ® tubes. Protein amounts in the SEC 

>30 kDa and SEC ≤30 kDa urinary protein concentrates are measured using 

the BCA assay as described under Section 8.3.1. step 5 and are frozen at –80°C. 

3.3. Immunoaffinity Subtraction of the SEC >30 kDa Fraction 

1. An optional fractionation step is the enrichment of less abundant urinary proteins 

depleting highly abundant plasma proteins via immunoaffinity subtraction (IAS). 

A detailed protocol for plasma protein depletion is provided in a different chapter 

in this book. 

2. Removal of highly abundant plasma proteins such as albumin and immunoglobulins 

(Ig) is desirable to increase the dynamic range for protein detection and 

quantitation in downstream proteomic analyses. Most abundant plasma proteins 

have Mr values higher than 30 kDa and are present in the SEC >30 kDa urinary 

fraction. This is the fraction to be subjected to IAS (see Note 6). 

3. Different commercial products are available to deplete abundant plasma proteins 

from serum or urine and may be available in batch and/or as sealed cartridges 

(see Note 7). 

4. This short protocol describes depletion with IAS resin in batch form. Approx 

0.5–1 mL resin (available in batch or removed from a cartridge) is suspended 

in 1 mL buffer A and placed in a approx 5-mL microtube. The SEC >30 kDa


fraction (500 μL) is added to the suspended IAS resin and incubated for 15 min 

at room temperature. During the incubation step, the suspension is occasionally 

agitated by gently pipeting. 

5. The suspension is transferred to an Ultrafree-CL 0.45-μm filter device and 

centrifuged at 1,000g for 1–2 min. The filtrate (bottom tube) is collected and 

placed in an Amicon ® Ultra-4 centrifugal filter device. The resin is resuspended 

gently in 1.5 mL buffer A, briefly equilibrated, spun again at 1,000g for 1–2 min 

and the filtrate added to the Amicon ® Ultra-4 tube. This step is repeated and 

the three combined filtrates are concentrated to approx 250 μL by spinning the 

Fig. 3. Protein spot profiles in 2-DE gels in a IAS >30 kDa fraction (top gel) and 

a SEC ≤ 30 kDa fraction (bottom gel). Samples with approx 200 μg protein were 

loaded in each IEF tube gel. In the first dimension, proteins were focused in the pI 

range between 4 and 7 applying 25,000 Volt-hours (Vh). The tube gel was stacked 

on an 8–15 %T second-dimension slab gel and proteins were resolved in the M r 

range between 8 and 200 kDa over 1,300 Vh. Protein spots in the gels were stained 

with Coomassie Brilliant Blue G. Spot trains denoted in the gels are: (1) albumin*; 

(2) (-1-acid glycoprotein*; (3) Ig light chains*; (4) prostaglandin H2 D-isomerase; 

(5) (-1-microglobulin. *These proteins were of very high abundance in the SEC >30 

kDa fraction and mostly removed via IAS. Spot trains for Ig light chains (3) and 

prostaglandin H2 D-isomerase (4) overlap in the bottom gel.

96 Pieper 

Amicon ® Ultra-4 device at 4,000g at 4°C. This concentrate is the IAS >30 kDa 

(flow-through) fraction. 

6. To recycle the resin, it is resuspended in 2 mL elution buffer in the Ultrafree-CL 

device at room temperature, equilibrated for 2–3 min and spun at 1,000g for 

1–2 min. The resin re-suspension and elution step is repeated. The eluates are 

discarded and the resin is immediately neutralized with 4 mL buffer A and spun 

at 1,000g for 1–2 min. The resin is resuspended in a small volume of buffer A 

(0.5–1 mL) and stored in suspension at 4°C. The resin can be reused for IAS 

using another urinary protein SEC >30 kDa fraction (see Note 8). 

3.4. Final Urinary Protein Sample Preparation 

for Proteomic Analysis 

1. The SEC ≤30 kDa fraction and the SEC >30 kDa fraction, if not processed via 

IAS, were stored at –80°C. After thawing, they are transferred to Amicon ® Ultra-4 

filter devices and concentrated to approx 250 μL at 4°C spinning at 4,000g, as 

described for the IAS >30 kDa fraction in Section 3.3. step 5. 

2. Final protein amounts are determined using the BCA assay as described in 

Section 3.1. step 5. 

3. A 40-fold buffer exchange in the Amicon ® Ultra-4 follows rediluting and reconcentrating. 

For 250 μL, the total exchange buffer volume is therefore 10 mL, which 

is added stepwise to the sample in the Amicon ® Ultra-4 tube (4 mL capacity). 

4. For the preparation of 2-DE gel samples and for further LC separation steps, 

buffer B is used as the exchange buffer. The 2-DE gels in Fig. 3 display the spot 

profiles for two urinary protein fractions. To prepare urinary protein samples for 

trypsin digestion, 25 mM ammonium bicarbonate is used as the exchange buffer. 

These protein samples are lyophilized for 24 h and the protein amounts of the 

freeze-dried samples are noted. 

5. For the preparation of samples in which proteins are to be covalently labeled, e.g., 

with isotope-coded amine-reactive tags for differential MS analysis, the exchange 

buffer is 50 mM HEPES, pH 7.8 and the volume is reduced to obtain a final 

protein concentration of approx 5 mg/mL. 

4. Notes 

1. The gel filtration standard contains thyroglobulin (670 kDa), bovine -globulin 

(150 kDa), chicken ovalbumin (45 kDa), equine myoglobin (18 kDa), and vitamin 

B12 (1.4 kDa) and, once reconstituted in 500 μL water, has a protein concentration 

of 36 mg/mL. 

2. Under ideal circumstances, human urine specimens collected in clinical laboratories 

should be cooled on ice and supplemented with protease inhibitor tablets


(Complete TM ) immediately. Centrifugation at 3,000g to remove cellular debris 

should also occur in the clinical laboratory right after sample cooling. The resulting 

supernatant can be frozen at –80°C and shipped on ice. 

3. The 200–250 mL urinary fluid is added stepwise to the same Centricon ® Plus-80 

device (capacity of 80 mL). If precipitation occurs during the concentration step, 

the collected and concentrated sample is aliquoted into three 1.5-mL microtubes 

and spun for 15 min at 10,000g. 

4. It is ideal to equilibrate and run the Superdex 75 column at 4ºC. If a cold cabinet 

or a cooling system is not available, the LC experiments (gel filtration standard, 

urinary protein samples) can also be performed at room temperature. At room 

temperature, 1 mM benzamidine and 1 mM EDTA should be added to buffer A. 

It is important to equilibrate the reservoir with buffer A and the chromatography 

column to the same temperature for LC separation of the samples (either at 4ºC 

or at room temperature). 

5. A less ideal alternative is to approximate the 30 kDa elution volume as the 

fraction located equidistantly between the 45 kDa (ovalbumin) and the 18 kDa 

(myoglobin) LC peaks. 

6. Detailed protocols for the immunoaffinity subtraction (IAS) technology are 

described in a separate chapter in this book. In the human urine, several blood 

plasma proteins are frequently observed as highly abundant proteins (3). These 

proteins are human albumin (approx 67 kDa), IgG (approx 150 kDa) and -1-acid 

glycoprotein (approx 42 kDa). However, their concentrations are known to vary 

dramatically in human urine, sample- and donor-dependent fashion. Other proteins 

frequently of high abundance in human urine are prostaglandin H2 d-isomerase 

(approx 25 kDa) and -1-microglobulin (approx 31 kDa). 

7. Three simple-to-use IAS cartridges for the removal of plasma proteins are 

available (see Materials). Albumin and IgG are subtracted by all three 

immunoaffinity removal cartridges. For additional plasma protein subtraction 

specificities, further information should be obtained from the cartridge manufacturer. 

In a proteomic experiment comparing several urinary protein samples, only 

one of the IAS resin products should be used. The methods for the use of an IAS 

LC cartridge are different. Commercially available IAS LC resin cartridges (with 

a volume greater than 0.5 mL) are typically sufficient for high-abundance plasma 

protein removal from a SEC >30 kDa fraction with up to 3 mg total urinary protein. 

8. The described protocol works particularly well with protein A- or protein 

G-derivatized resins in which proteins A/G are cross-linked to plasma proteinspecific 

polyclonal antibodies via dimethylpimelimidate. In particular, this 

pertains to the elution and resin recycling steps which allow for effective elution 

of affinity-bound proteins, retain the cross-linkage and maintain the binding 

functions of the immobilized antibodies. Whether the elution buffer is compatible 

with the IAS resins of all depletion cartridges (based on cross-linkage chemistry 

between resin and antibodies) should be verified with the manufacturers. An 

alternative elution buffer provided by the manufacturer may be used for the 

recycling of the resin.

98 Pieper 

References 

1. Celis, J. E., Wolf, H., and Ostergaard, M. (2000) Bladder squamous cell carcinoma 

biomarkers derived from proteomics. Electrophoresis 21, 2115–21 

2. Edwards, J. J., Anderson, N. G., Tollaksen, S. L., von Eschenbach, A. C., 

and Guevara, J., Jr. (1982) Proteins of human urine. II. Identification by twodimensional 

electrophoresis of a new candidate marker for prostatic cancer. Clin 

Chem 28, 160–63 

3. Pieper, R., Gatlin, C. L., McGrath, A. M., et al. (2004) Characterization of the 

human urinary proteome: a method for high-resolution display of urinary proteins 

on two-dimensional electrophoresis gels with a yield of nearly 1400 distinct protein 

spots. Proteomics 4, 1159–74 

4. Saito, M., Kimoto, M., Araki, T., et al. (2005) Proteome analysis of gelatin-bound 

urinary proteins from patients with bladder cancers. Eur Urol 48, 865–71 

5. Williams, K. M., Williams, J., and Marshall, T. (1998) Analysis of Bence Jones 

proteinuria by high resolution two-dimensional electrophoresis. Electrophoresis 

19, 1828–35 

6. Decramer, S., Wittke, S., Mischak, H., et al. (2006) Predicting the clinical outcome 

of congenital unilateral ureteropelvic junction obstruction in newborn by urinary 

proteome analysis. Nat Med 12, 398–400 

7. Waller, K. V., Ward, K. M., Mahan, J. D., and Wismatt, D. K. (1989) Current 

concepts in proteinuria. Clin Chem 35, 755–765 

8. Anderson, N. G., Anderson, N. L., and Tollaksen, S. L. (1979) Proteins of human 

urine. I. Concentration and analysis by two-dimensional electrophoresis. Clin Chem 

25, 1199–1210 

9. Edwards, J. J., Tollaksen, S. L., and Anderson, N. G. (1982) Proteins of human 

urine. III. Identification and two-dimensional electrophoretic map positions of some 

major urinary proteins. Clin Chem 28, 941–48 

10. Marshall, R. J., Turner, R., Yu, H., and Cooper, E. H. (1984) Cluster analysis of 

chromatographic profiles of urine proteins. J Chromatogr 297, 235–44 

11. Marshall, T., and Williams, K. M. (1993) Centriprep ultrafiltration for fractionation 

of serum and urinary proteins before electrophoresis. Clin Chem 39, 1558 

12. Marshall, T., and Williams, K. (1996) Two-dimensional electrophoresis of human 

urinary proteins following concentration by dye precipitation. Electrophoresis 17, 

1265–72 

13. Marshall, T., and Williams, K. M. (1997) Two-dimensional electrophoresis of 

human urine and cerebrospinal fluid following protein concentration by dye precipitation. 

Biochem Soc Trans 25, S657 

14. Thongboonkerd, V., Chutipongtanate, S., and Kanlaya, R. (2006) Systematic evaluation 

of sample preparation methods for gel-based human urinary proteomics: 

quantity, quality, and variability. J Proteome Res 5, 183–91 

15. Thongboonkerd, V., McLeish, K. R., Arthur, J. M., and Klein, J. B. (2002) 

Proteomic analysis of normal human urinary proteins isolated by acetone precipitation 

or ultracentrifugation. Kidney Int 62, 1461–69


16. Tantipaiboonwong, P., Sinchaikul, S., Sriyam, S., Phutrakul, S., and Chen, S. T. 

(2005) Different techniques for urinary protein analysis of normal and lung cancer 

patients. Proteomics 5, 1140–49 

17. Sun, W., Li, F., Wu, S., et al. (2005) Human urine proteome analysis by three 

separation approaches. Proteomics 5, 4994–5001 

18. Oh, J., Pyo, J. H., Jo, E. H., (2004) Establishment of a near-standard twodimensional 

human urine proteomic map. Proteomics 4, 3485–97 

19. Pieper, R., Su, Q., Gatlin, C. L., Huang, S. T., Anderson, N. L., and 

Steiner, S. (2003) Multi-component immunoaffinity subtraction chromatography: 

an innovative step towards a comprehensive survey of the human plasma proteome. 

Proteomics 3, 422–32

9 

Isolation of Cytoplasmatic Proteins from Cultured Cells 

for Two-Dimensional Gel Electrophoresis 

Ying Wang, Jen-Fu Chiu, and Qing-Yu He 

Summary 

Cytoplasma is the cell interior place between the cellular membrane and the nucleus, 

where various intracellular activities take place, including energy production, reactive 

oxygen species (ROS) detoxification, heme synthesis, nitrogen and lipid metabolism, 

phosphorylation in signal transduction, and cytoskeletal meshwork construction. The rich 

cytoplasmatic proteins carrying out these intracellular functions are interesting targets 

for biochemical and molecular biological studies. The relatively recent discipline of 

proteomics offers a chance to globally analyze the changes in cytoplasmic proteins corresponding 

to drug treatments or disease conditions, and thus provide target candidates for 

further biological validation in drug development and biomarker discovery. Isolation of 

cytoplasmic proteins from cells is a necessary step for high resolution protein separation 

by two-dimensional gel electrophoresis (2DE) and specific proteomic analysis. 

Key Words: Cytoplasmatic proteins; protein isolation; proteomics; sample preparation; 

subcellular fractionation; two-dimensional gel electrophoresis. 


The cytoplasm is crowded and highly ordered with transport vesicles, 

mitochondria, chloroplasts, and other organelles. The endocytosis and exocytosis 

in cytoplasm provide paths between the cell interior and the surrounding 

medium, allowing for the uptake of extracellular components and the secretion 

of proteins and other components produced within the cell. The cytoplasm is 

the place for energy metabolism (1), reactive oxygen species (ROS) detoxification 

(2), heme synthesis (3), nitrogen and lipid metabolism (4,5), mixed 



101

102 Wang et al. 

phospholipids biosynthesis (6), cytoskeleton rearrangements (7), and various 

protein-protein interactions (8). Mapping the altered expression in cytoplasmic 

proteins by proteomic analysis under given conditions can provide a network 

of responses to intracellular and extracellular signals (9). 

Proteomics is a research technique that can identify, characterize, and 

quantitate proteins expressed in cells, tissues, or organisms under given conditions 

such as chemotherapeutic drug challenge (10,11). The altered proteins 

identified by proteomic analysis can be further characterized as potential drug 

targets; the global analysis of the protein alterations can result in valuable 

information for understanding the drug action mechanisms. By comparing the 

cytoplasmic protein profiles of HONE1 cells treated by gold(III) porphyrin 

1a to untreated control, we identified a number of differentially expressed 

proteins by peptide-mass-finger printing (PMF) (12). The identification of the 

altered proteins provided valuable clues to illustrate the underlying drug action 

mechanisms. 

2. Materials 

2.1. Cell Culture and Wash Buffer 

1. RPMI 1640 Medium or Dulbecco’s Modified Eagle’s Medium (DMEM) plus 

10% FBS, supplemented with 2 mM/L l-glutamine, 100 units/mL penicillin, and 

100 μg/mL streptomycin. 

2. Solution of trypsin (0.25%) and ethylenediamine tetraacetic acid (EDTA) (1 mM). 

3. Cell washing buffer for 2DE: 10 mM Tris-HCl, pH 7.0, 250mM sucrose. 

4. Teflon cell scrapers. 

5. Tissue grinder. 

2.2. Buffers and Reagents for Cytoplasmic Protein Precipitation 

1. Extraction buffer: 10 mM Tris-HCl, pH 7.6, 10 mM KCl, 5 mM MgCl2. 2. Nuclei isolation buffer: 10 mM Tris-HCl, pH 7.6, 10 mM KCl, 5 mM MgCl2, 0.35M sucrose. 

3. Radioimmunoprecipitation assay buffer (RIPA): 10 mM Tris-HCl, pH 7.5, 150 

mM NaCl, 1% NP 40 (w/v), 0.1% SDS (w/v), 0.5% sodium deoxycholate (w/v) 

(see Note 1),1mMDTT. 4. Protease inhibitors (see Note 2): 200 mM stock solution of phenylmethanesulfonyl 

fluoride (PMFS) in isopropanol (store at room temperature); 1 mg/mL leupeptin 

in water (store frozen in aliquots), 1 mg/mL aprotinin in water (store frozen in 

aliquots), 1 mg/mL pepstatin in methanol (store frozen in aliquots). 

5. Phosphatase inhibitors (see Note 3): activated 200 mM stock solution of sodium 

vanadate in water and 200 mM sodium fluoride stock solution, store at room 

temperature.

Isolation of Cytoplasmatic Proteins from Cultured Cells 103 

6. Methylene blue solution: 1.4% (w/v) methylene blue in 95% ethanol and filtered 

through 0.45 μm filter paper. 

7. 10% trichloroacetic acid (TCA). 

2.3. Buffers for Western Blot Analysis 

1. Transfer buffer: 25 mM Tris base, 192 mM glycine, 0.05% SDS (w/v), 20% 

methanol. 

2. TBS-T: 20 mM Tris-HCl, pH 7.6, 0.15M NaCl, 0.1% (w/v) Tween 20. 

3. Stripping buffer: 50 mM glycine, 1 % SDS (w/v), pH 2.0. 

4. ECL reagents (GE Healthcare). 

2.4. Buffers and Reagents for Two-Dimensional Gel 

Electrophoresis (2DE) 

1. 2DE clean up kit (GE Healthcare). 

2. Rehydration solution: 8M urea, 2% CHAPS (stored frozen in aliquots). Add 

0.002% bromophenol blue, 0.005% IPG buffer, and 2.8 mg/mL DTT freshly 

before use. 

3. SDS equilibration buffer: 6M urea, 50 mM Tris-HCl, pH 8.8, 30% glycerol, 2% 

SDS. 

4. 1.5M Tris-HCl (pH 8.8): dissolve 181.7 g Tris base into 750 mL ddH 2O, add 12N 

HCl to adjust to pH 8.8 and then add ddH 2O to final volume of 1 L, after that, 

filtered by 0.45 μm filter paper. 

5. Thirty percent acrylamide gel stock solution: 30% acrylamide (w/v), 0.8% N’N’methylenebiasacrylamide 

(w/v), and filtered by 0.45 μm filter paper. 

6. SDS running buffer: 25 mM Tris base, 0.192 M glycine, 0.1% SDS (w/v). 

7. Agarose sealing solution: 0.5% agarose (normal or low-melting point), 0.002% 

bromophenol blue in SDS running buffer. 

8. Bromophenol blue stock solution: 1% (w/v) bromophenol blue powder, 50 mM 

Tris-Base, filtered through 0.45 μm filter paper. 

9. Silicon oil (e.g., DryStrip ® Cover Fluid from GE Healthcare). 

2.5. Silver Staining Solutions 

1. Fixation solution: 40% ethanol, 10% acetic acid. 

2. Incubation solution: 30% ethanol, 4.1% sodium acetate (anhydrous) (w/v), 0.2% 

sodium thiosulfate (anhydrous) (w/v). 

3. Silver nitrate solution: 0.1% silver nitrate (w/v), 0.02% formaldehyde (v/v). 

4. Development solution: 2.5% sodium-carbonate (w/v), 0.01% formaldehyde. 

5. Stop solution: 1.46% sodium-EDTA·2H2O (w/v). 

6. Preservation solution: 4.0% glycerol (w/v), 30% ethanol.


2.6. Coomassie Brilliant Blue Staining Solutions 

1. Fixation solution: 40% methanol (v/v) and 5% phosphoric acid (v/v). 

2. Coommasie blue G-250 staining solution: 0.08% Coomassie brilliant blue G-250 

(w/v) in 12% trichloroacetic acid, pH < 1.0. 

3. Coomassie blue R-250 staining solution: 0.1% Coomassie brilliant blue R-250 

(w/v), in 50% methanol (v/v), and 10 % acetic acid (v/v). 

4. Destaining solution: 15% methanol (v/v), 10% acetic acid (v/v), 7% acetic acid 

(optional) (stored up to 1 month at room temperature). 

3. Methods 

3.1. Cell Harvest (see Note 4) 

1. When the attached cells reach about 80% confluence, discard the media and wash 

two times with ice cold washing buffer (see Note 5), keep 1 mL washing buffer 

in the dish. 

2. Harvest cells by prechilled cell scrapper, then transfer 1 mL of cell suspension 

to a clean 2.0-mL Eppendorf tube, spin down at 3,000g for 5 min (4°C), and 

wash two times with washing buffer, 1 mL each time. Cell pellet should be lysed 

immediately for extraction of cytoplasmatic proteins. 

3. When suspended cells reach about 80% confluence, spin down at 3,000g for 5 

min (4°C), and wash two times with ice cold washing buffer, 10 mL for each 

plate, and spin down to get cell pellet. Cell pellet should be lysed immediately 

for extraction of cytoplasmatic proteins. 

3.2. Precipitation of Cytoplasmatic Proteins (see Note 6) 

1. Re-suspend about 1×107 cells in 1 mL extraction buffer with protease inhibitors 

(0.2 mM PMSF, 5 μg/mL leupeptin, 2 μg/mL aprotinin, and 2 μg/mL pepstatin). 

Incubate the cells on ice for 10 min, and lysis by addition of Triton X-100 to the 

final concentration of about 0.3% (w/v). 

2. Homogenize cells in an ice-cold tissue grinder. 30–50 passes with the grinder are 

recommended; however, efficient homogenization may depend on the cell type. 

To check the efficiency of homogenization, pipet 2–3 μL of the homogenized 

suspension onto a cover slide, stained with methylene blue solution and observe 

under a microscope. Nuclei appear as dark blue and cytoplasm as light purple 

blue under white field lens. If about 80% of the nuclei are not surrounded by 

cytoplasm, proceed to step 3. Otherwise, perform 10–20 additional passes using 

the tissue grinder. Excessive homogenization should also be avoided, as it can 

cause damage to the heavy membranes, e.g., mitochondrial membrane, which 

triggers release of mitochondrial components (see Note 7). 

3. Slowly add 0.5 volume of nuclei isolation buffer to the bottom of extraction buffer, 

then remove nuclei by centrifugation at 500–700g for 10 min (swinging-bucket 

rotor) (4°C).


4. Further centrifuge the supernatant from nuclei isolation at 10,000g for 30 min 

(fixed-angle rotor) (4°C). Save the supernatant as cytoplasmatic fraction, and the 

pellet as heavy membrane fraction containing mitochondria. 

5. Add 0.25 volumes of 10% TCA to cytoplasmatic fraction to precipitate cytoplasmatic 

proteins; incubate on ice for 10 min. Spin down at 5,000g for 5 min 

(4°C). Remove supernatant, leaving protein pellet intact. Wash pellet with 200 

μL prechilled acetone. Spin down at 5,000g for 5 min (4°C) (see Note 8). Wash 

twice with acetone again. Air dry the pellet, and add rehydration solution for 2DE 

analysis (see Note 9). Or add RIPA assay buffer for Western blot analysis to test 

the purity of the isolated subcellular fraction. 

3.3. Western Blot Analysis to Detect the Purity of the Sample 

1. Determine the purity of the isolated subcellular fractions by Western blot 

analysis against specific protein markers (Table 1). Separate the samples by onedimensional 

SDS PAGE. 

Table 1 

Marker proteins of individual subcellular fractions. 

Subcellular 

localization 

Marker protein Database 

accession 

number 

References 

Mitochondria Cox II gi 142786 (13) 

Cox IV gi 142789 (14, 15) 

Cytochrome oxidase 1 gi 551699 (16) 

VDAC gi 340201 (14, 15, 17) 

MnSOD gi 34707 (18, 19) 

HSP60 gi 77702086 (12) 

mtHSP70 gi 292160 (20) 

Cytochrome P450 gi 5733409 (21) 

Cytoplasma Tubulin gi 4507729 (22) 

GAPDH gi 7669492 (16, 23) 

-actin gi 4501885 (12) 

LDH gi 9257228 (20) 

Calpain gi 791040 (24) 

Cytokeratin gi 1419564 (25) 

Vimentin gi 62414289 (26) 

Nuclei Histone gi 3649600 (18, 19) 

c-Jun gi 20986521 (27) 

c-Fos gi 29904 (28, 29)


2. Before transfer, soak PVDF membrane in 100% methanol for 1 min, then soak 

with transfer buffer. 

3. After SDS PAGE, transfer the gel to the membrane electrophoretically, using 

transfer buffer. Assemble the filter paper, SDS gel and membrane into a blotting 

sandwich as shown in Fig. 1 (see Note 10). The current for transfer should be 0.8 

times the area of the membrane in mA. 

4. After transfer, incubate the membrane in 5% nonfat milk in TBS-T buffer (4°C 

overnight or room temperature for 2 h). 

5. Wash the membrane three times with TBS-T buffer (10 min each, room temperature), 

followed by incubation with primary antibody, in 1% nonfat milk (4°C 

overnight or room temperature for 2 h). 

6. Wash the membrane three times by TBS-T buffer (10 min each, room temperature), 

then incubate with secondary antibody in 1% nonfat milk (4°C overnight 

or room temperature, 45 min to 1 h). 

7. Discard the secondary antibody and wash the membrane three times by TBS-T 

buffer (10 min each, room temperature). Develop the membrane using the ECL 

reagents. 

8. Once a satisfactory exposure for the results has been obtained, the membrane 

is stripped with stripping buffer and then reprobed with another antibody. Wash 

the membrane twice with TBS-T buffer (10 min each, room temperature), then 

incubate the membrane in stripping buffer for 20 min (room temperature), wash 

twice with TBS-T again (10 min each, room temperature), then go to steps 

4–7. Fig. 2 shows an example of Western blot analysis of specific protein 

markers of cytoplasmic and heavy membrane fraction from cisplatin treated 

HONE1 cells. 

Fig. 1. Western Blot assembly.


Fig. 2. Western blot analysis of cytoplasmatic fraction obtained from HONE1 cells 

treated with cisplatin for 6, 12, and 24 h. 

3.4. Two-Dimensional Electrophoresis 

1. Add 2.8 mg DTT, 5 μL carrier ampholytes or IPG buffer of the respective 

pH gradient, and 2 μL bromophenol blue to 1 mL rehydration stock solution 

(without DTT and IPG buffer) immediately before adding the protein samples. 

2. Add protein samples to the above mentioned rehydration solution (volume 

recommendations are given in Table 2), vortex and spin down the samples. The 

recommended protein loading is given in Table 3. 

3. IPG-strip sample loading by in-gel-rehydration. Pipet the sample solution into 

an IPG-strip holding device. Remove the protective cover from the IPG strip, 

position the IPG strip with the gel side down. To help coat the entire IPG strip, 

gently lift and lower the strip and slide it back and forth along the surface of the 

solution. Be careful not to trap bubbles under the IPG strip. Overlay the strips by 

Immobiline DryStrip® Cover Fluid (GE Healthcare) to ensure that rehydrated 

Table 2 

Rehydration solution volume per Immobiline DryStrip 

IPG Strip Length (cm) Total volume per strip (μL) 

7 125 

13 250 

18 340


Table 3 

Protein loads for silver and Coomassie blue staining. 

IPG Strip Length (cm) pH-range 

Recommended protein load (μg) 

Silver stain Coomassie stain 

7 4–7 25–50 50–100 

3–10, 3–10 NL 50–75 50–150 

13 3–10, 3–10 NL 50–100 100–200 

18 4–7 200–400 400–1000 

3–10, 3–10 NL 100–400 200–1000 

Immobiline DryStrip gels do not dry out during electrophoresis. Finally, place 

the cover on the strip holder. 

4. IEF is performed according to a step-wise voltage increase procedure: 

rehydration at 30 V for 10–16 h, followed by 500 V and 1000 V for 1 h 

each, and 5,000–8,000 V for about 10 h with a total of 64,000 V hours 

(56,000 V hours is acceptable for 7-cm strips). After IEF, the strips can be 

subjected to second dimension immediately or can be kept at –70 ° C for several 

weeks. 

5. Prepare the 1.5-mm thick polyacrylamide gels. Seal the gel surface by 1-butanol, 

allow the gel to be polymerized for at least 30 min at room temperature or 

polymerize the gel overnight (see Notes 11 and 12). 

6. The strips after IEF are subjected to two-step equilibration in equilibration 

buffers with 1% DTT (w/v) for the first step, and 2.5% (w/v) iodoacetamide for 

the second step. Wash strips by SDS running buffer for three times before load 

onto the acrylamide gel (see Note 13). 

7. Electrophoresis conditions: set buffer circulation temperature to 10ºC and start 

the run at 15 mA per gel. After 30 min increase current to 30 mA per gel. 

8. After SDS page, visualize proteins with silver staining or Coomassie brilliant 

blue staining. For silver staining, fix the gels in fixation solution overnight, 

and then change to incubation solution for 30 min. After washing three times 

in water for 10 min each, stain the gels in silver nitrate solution for 40 

min. Perform development for 15 min in development solution. Stop staining 

by stop solution and then wash the stained gels three times in water for 

5 min each. 

9. Alternatively, visualize the gels by Coomassie brilliant blue stain. Fix the gels 

in fixation solution for Coomassie brilliant blue stain overnight, followed by 

Coommasie blue G-250 or R-250 stain for more than 12 h. Destain by destaining 

solution until background is acceptable. 

10. Acquire images by a suitable Image Scanner and preserve gels in preservation 

solution for further analysis. Fig. 3 shows 2DE pattern of cytoplasmatic proteins 

compared with the pattern of a whole cell protein extract.


Fig. 3. 2DE pattern of cytoplasmatic proteins compared with the pattern of whole 

cell protein. One hundred μg of cytoplasmatic protein or whole cell protein from human 

nasopharyngeal carcinoma HONE1 cells was separated on 13-cm IPG-strips, pH 3–10, 

followed by SDS-PAGE (12% SDS gel), and visualized by silver staining. 

4. Notes 

1. Sodium deoxycholate is an ionic detergent to extract proteins. Prepare 10 % 

stock solution in water, protect the solution from light. Do not add sodiumdeoxycholate 

when preparing lysates for kinase assays. Ionic detergents can 

denature enzymes, causing them to lose activity. 

2. Commercially available protease inhibitor cocktails can be used instead, for 

example, protease inhibitor cocktail (Catalog number P8340) from Sigma- 

Aldrich, protease inhibitor cocktail set I (Catalog number 539131) from 

Calbiochem. 

3. Do not add phosphatase inhibitors when preparing lysates for phosphatase assays. 

4. Wash cells intensively to avoid contamination of serum proteins in the culture 

medium. 

5. Do not wash the cells with PBS in the last washing step, because PBS contains 

150 mM sodium chloride, which will interfere with isoelectric focusing. Use 

instead 250 mM sucrose, 10 mM Tris-HCl, pH 7.5. 

6. Always perform the isolation of cytoplasmic proteins on ice. Chill all buffers 

before use. Perform the isolation steps on ice to ensure the purity of subcellular 

fractions, because activities of most cellular proteases are inhibited at low 

temperature. 

7. Under ideal homogenization conditions, particulate organelles such as nuclei, 

mitochondria, lysosomes, and peroxisomes remain intact. Golgi complexes, 

plasma membranes, and reticular organelles will fragment and can, at least to 

some extent, vesiculate. It is not possible to outline a general protocol suitable 

for the production of a reasonable homogenate for all kinds of cultured cells,


because different conditions are required for different cell types and experimental 

purposes. 

8. Long centrifuge at high speed will cause difficulties in dissolving protein pellet 

in rehydration solution or RIPA assay buffer. 

9. 2DE clean-up kit can be used to improve the quality of 2DE results by removing 

interfering contaminants. 

10. Bubbles between the SDS gel and the membrane will lead to inefficient transfer. 

11. It is critical that all the glass plates for 2DE have been cleaned and rinsed 

extensively with distilled water. Clean the glass plates with distilled water 

followed by 95% ethanol to remove the acid and air-dry prior use. 

12. Prepare 12.5% second dimensional SDS gels for standard separations; prepare 

8–10% gels to better separate large molecular weight proteins and 15% gels 

for low molecular weight proteins. For the equilibration step, add DTT and 

iodoacetamide to equilibration buffer just prior use. 


This investigation was partially supported by grants from Hong Kong 

University funding (No. 200511159099 to Q.Y.H.) and the Area of Excellence 

Scheme of the Hong Kong University Grants Committee. 

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10 

Sample Preparation of Culture Medium from 

Madin-Darby Canine Kidney Cells 

Daniel Ambort, Daniel Lottaz, and Erwin Sterchi 

Summary 

A reproducible, standardized and simple sample preparation methodology is the key 

to successful two-dimensional gel electrophoresis (2-DE). This chapter describes step-bystep 

the sample preparation of culture medium from Madin-Darby canine kidney (MDCK) 

cells. Tips and tricks are given to circumvent possible pitfalls. 

Key Words: Bicinchoninic acid (BCA) assay; culture medium (CM); isoelectric 

focusing (IEF); Madin-Darby canine kidney (MDCK); rehydration loading; 

two-dimensional gel electrophoresis (2-DE) ultracentrifugation; ultrafiltration . 


Two-dimensional gel electrophoresis (2-DE), introduced by O’Farrell and 

Klose in 1975 (1,2), enabled separation of complex protein mixtures into 

individual protein species according to their net charge (pI) in the first 

dimension by isoelectric focusing (IEF) and in the second dimension according 

to their molecular mass (Mr) by sodium dodecyl sulfate-polyacrylamide gel 

electrophoresis (SDS-PAGE) (3). In the conventional approach, IEF was 

performed in carrier ampholyte-generated pH gradients, which moved towards 

the cathode on prolonged focusing time. This “cathodic drift” phenomenon 

was thereafter remedied by nonequilibrium pH gradient gel electrophoresis 

(NEPHGE) (4) and finally eliminated with the invention of fixed immobilized 

pH gradients (IPG) (5–8). The development of microanalytical techniques, 

namely Edman sequencing (9–11) and mass spectrometry (12–14) enabled 



113

114 Ambort et al. 

identification of proteins at amounts available from a single 2-D gel. 2-DE 

advanced to the core technology of proteome analysis (7,8,15) and was brought 

from art to craft in an industrial standard. 

Appropriate sample treatment is the key to good results. The ideal sample 

solubilization procedure should result in the disruption of all noncovalently 

bound protein complexes and aggregates into a solution of individual 

polypeptides (15). Denaturation and reduction of proteins is achieved in the 

standard lysis buffer (O’Farrell 1975) (1) which is composed of 8–9 M urea, 

2–4% CHAPS, 1% DTT or DTE and 0.8–2% carrier ampholytes. Hydrophobic 

proteins are better dissolved in 2 M thiourea and 7 M urea instead of 9 M urea 

(16). Optimized procedures for different sample types do exist (17). However, 

a general “Prepares them all” procedure is not available (18). Another highpriority 

issue is the removal and inactivation of all interfering substances: 

nucleic acids, lipids, salts, small ionic compounds, polysaccharides, proteases, 

and insoluble particles. Sample preparation for 2-DE is a very cumbersome and 

time-consuming task that is subject to trial and error. 

Because of the complex biological architecture of eukaryotic cells into 

organelles and large cellular structures, fractionation techniques are applied 

before comprehensively studying the subproteomes. In such reductionist 

approaches classic biochemical techniques, namely centrifugation and affinitymediated 

isolation using antibodies against molecular tags, are applied to enrich 

for subcellular fractions as reviewed by Yates 3rd (19). Beside analysis of 

cytosolic, organelle-specific and transmembrane proteins several investigations 

were aimed at identifying those proteins secreted by various cell types into the 

extracellular milieu or medium (20–24). 

This chapter describes the sample preparation of culture medium from 

Madin-Darby canine kidney (MDCK) cells for 2-DE (Fig. 1). The methodology 

is subsectioned into four parts with basic introductory information on each topic: 

cell culture (see Subheading 3.1.), ultracentrifugation and ultrafiltration (see 

Subheading 3.2.), protein quantitation by BCA assay (see Subheading 3.3.) 

and finally, rehydration loading and isoelectric focusing (see Subheading 3.4.). 

2. Materials 

2.1. Cell Culture 

2.1.1. Equipment 

1. BD Falcon TM bulk packaged serological pipets (10 mL) (BD Biosciences, Franklin 

Lakes, NY, USA) 

2. BD Falcon TM individually wrapped serological pipets (25 mL) (BD Biosciences, 

Franklin Lakes, NY, USA)

Sample Preparation of Culture Medium 115 

Fig. 1. Two-dimensional gel electrophoresis of Madin-Darby canine kidney cell 

culture supernatant (80 μg protein). First dimension: isoelectric focusing in an immobilized 

pH gradient (IPG) pH 3–10 nonlinear in a 24-cm-long gel strip. Second dimension: 

SDS-PAGE in a 12.5% gel. Silver stained. 

3. BD Falcon TM standard cell culture dish, standard tissue-culture treated 

(100 × 20 mm) (BD Biosciences, Franklin Lakes, NY, USA) 

4. CELLSTAR ® PP-test tubes (50 mL, sterile) (Greiner Bio-One Inc., Longwood, 

FL, USA) 

5. Erlenmeyer flasks (250 mL) 

6. Laminar air flow cabinet (Brouwer AG, Luzern, Switzerland) 

7. Neubauer improved counting chamber (Assistent, Sondheim, Germany) 

8. NUAIRE TM US autoflow CO 2 water-jacketed incubator NU-4750 (Vitaris AG, 

Baar, Switzerland) 

9. Water bath 

2.1.2. Solutions and Reagents 

1. Dulbecco’s Phosphate Buffered Saline (D-PBS) (1X) (500 mL) (GIBCO Invitrogen 

corporation, Grand Island, NY, USA) 

2. Foetal bovine serum (FBS) (E. U. approved South American origin, virus and 

mycoplasma tested) (500 mL) (GIBCO Invitrogen corporation, Grand Island, NY, 

USA) 

3. Minimum essential medium (MEM) (1X) (with Earle´s salts, without l-glutamine) 

(500 mL) (GIBCO Invitrogen corporation, Grand Island, NY, USA) 

4. Penicillin-streptomycin-glutamine (100X) (100 mL) (GIBCO Invitrogen corporation, 

Grand Island, NY, USA) 

5. Trypsin-EDTA (1X) (100 mL) (GIBCO Invitrogen corporation, Grand Island, 

NY, USA)


6. Culture medium: 1X MEM (see Note 1), 5% (v/v) FBS, 100 U/mL penicillin, 

100 μg/mL streptomycin and 292 μg/mL l-glutamine. To 500 mL of MEM (one 

bottle) aseptically add 25 mL of FBS (see Note 2) and 5 mL of 100X penicillinstreptomycin-glutamine 

stock solution. Store at 4°C. Before use warm up to 37°C 

in a water bath. 

7. Serum-free medium: 1X MEM (see Note 1), 100 units/mL penicillin, 100 μg/mL 

streptomycin, and 292 μg/mL l-glutamine. To 500 mL of MEM (one bottle) 

aseptically add 5 mL of 100X penicillin-streptomycin-glutamine stock solution. 

Store at 4°C. Before use warm up to 37°C in a water bath. 

2.2. Ultracentrifugation and Ultrafiltration 


1. Centricon ® Plus-70 centrifugal filter devices (Millipore corporation, Billerica, 

MA, USA) 

2. Centrifuge filter system (50 mL, 0.2 μm) (Costarcorporation, Cambridge, MA, 

USA) 

3. Eppendorf centrifuge 5415R (Eppendorf AG, Hamburg, Germany) 

4. Eppendorf tubes (1.5 mL, 2 mL) 

5. KONTRON CENTRIKON TFT 70.38 fixed-angle rotor (KONTRON Instruments 

AG, Zürich, Switzerland) 

6. KONTRON CENTRIKON T-2060 ultracentrifuge (KONTRON Instruments 


7. KONTRON CENTRIKON ultracentrifuge tubes (32.5 mL) (KONTRON Instruments 


8. Mettler AC 100 analytical balance (Mettler Instrumente AG, Greifensee, Zürich, 

Switzerland) 

9. Sorvall RT6000D centrifuge (Kendro Laboratory Products AG, Zürich, 


10. Sorvall H1000B swinging bucket rotor (Kendro Laboratory Products AG, Zürich, 


11. Water bath 


1. Ethylenedinitrilo tetraacetic acid disodium salt dihydrate (Na 2-EDTA 2H 2O, 

Titriplex ® III) (GR for analysis) 

2. Phenylmethylsulfonyl fluoride (PMSF) (Sigma, St. Louis, MO, USA) 

3. 2-Propanol (GR for analysis) 

4. Sodium hydroxide (NaOH) (pellets GR for analysis) 

5. Tris(hydroxymethyl)aminomethane (Tris) (GR for analysis buffer substance)


6. 0.5 M EDTA pH 8.0 stock solution: To make 100 mL of stock solution, dissolve 

2 g of NaOH pellets in 80 mL of ddH 2O. Add 18.6 g of Na 2-EDTA-2H 2O under 

constant stirring at RT (see Note 3). Titrate solution to pH 8 with 5 N NaOH 

(liquid). Adjust to a final volume of 100 mL with ddH 2O and check pH again. 

Filter solution with a 0.2 μm bottle top filter. This solution can be stored at RT. 

7. 0.1 M PMSF stock solution: To prepare 25 mL, dissolve 0.44 g of PMSF in 25 

mL of 2-propanol (isopropanol). Warm up to 37°C in a water bath (see Note 4). 

Portion solution into 2 mL aliquots in 2 mL Eppendorf tubes and store at –20°C. 

8. 0.2 M Tris pH 10.5 stock solution: To make 0.5 L, dissolve 12.12 g of Tris in 

0.5 L of ddH 2O(see Note 5). Filter solution with a 0.2 μm bottle top filter. This 

solution can be stored at RT. 

9. Sample solubilization buffer I: 20 mM Tris-HCl pH 9.0, 1 mM EDTA, 1 mM 

PMSF. To prepare 150 mL, dilute 15 mL of 0.2 M Tris pH 10.5 stock solution to 

a final volume of 150 mL in ddH 2O. Add 0.3 mL of 0.5 M EDTA pH 8.0 stock 

solution and 1.5 mL of 0.1 M PMSF stock solution under constant stirring at RT 

(see Note 6). Filter solution with a 0.2 μm bottle top filter. This solution should 

be prepared freshly just before use! 

2.3. Protein Quantitation by BCA Assay 


1. Beaker (25 mL) 

2. CELLSTAR ® micro-plate (TC, sterile) (Greiner Bio-One Inc., Longwood, FL, 

USA) 

3. Incubator 

4. Vortex mixer 

5. Vmax ® microplate reader (Molecular Devices Corporation, Sunnyvale, CA, USA) 


1. Albumin Standard Ampules (2 mg/mL, 10 x 1 mL) (Pierce, Rockford, IL, USA) 

2. BCATM Protein Assay Kit (Pierce, Rockford, IL, USA) 

3. BCATM Protein Assay Reagent A (500 mL) (Pierce, Rockford, IL, USA) 

4. BCATM Protein Assay Reagent B (25 mL) (Pierce, Rockford, IL, USA) 

2.4. Rehydration Loading and Isoelectric Focusing 


1. Beaker (50 mL) 

2. Centrifuge filter system (50 mL, 0.2 μm) (Costarcorporation, Cambridge, MA, 

USA) 

3. Eppendorf centrifuge 5415R (Eppendorf AG, Hamburg, Germany) 

4. EPS 3501 XL power supply (Amersham Biosciences, Uppsala, Sweden)


5. IEF electrode strips (Amersham Biosciences, Uppsala, Sweden) 

6. Immobiline TM DryStrip Kit (Amersham Biosciences, Uppsala, Sweden) 

7. Immobiline TM DryStrip Reswelling Tray (7–24 cm) (Amersham Biosciences, 

Uppsala, Sweden) 

8. Vortex mixer 

9. Multiphor TM II horizontal electrophoresis apparatus (Amersham Biosciences, 

Uppsala, Sweden) 

10. Multitemp TM III thermostatic circulator (Amersham Biosciences, Uppsala, 

Sweden) 

11. Multi-purpose rotator (Scientific Industries Inc., Queens Village, NY, USA) 

12. Parafilm (50 × 15 m) (American National Can Company, Chicago, IL, USA) 

13. Petri dishes 

14. Tweezers 


1. Bromophenol blue (BPB) (Bio-Rad Laboratories, Richmond, CA, USA) 

2. 3-[(3-cholamidopropyl)-dimethylammonio]-1-propane sulfonate (CHAPS) 

(ultrapure) (USB corporation, Cleveland, OH, USA) 

3. 1,4-dithioerythritol (DTE) (for biochemistry) (Merck, Darmstadt, Germany) 

4. ImmobilineTM DryStrip pH 3–10 NL (IPG) (240 × 3 × 0.5 mm) (Amersham 

Biosciences, Uppsala, Sweden) 

5. Mixed bed ion exchanger resin 

6. Paraffin (Merck, Darmstadt, Germany) 

7. PharmalyteTM 3–10 (for IEF) (Amersham Biosciences, Uppsala, Sweden) 

8. Thiourea (puriss. p. a. ACS; ≥99% [RT]) (Fluka, Buchs, Switzerland) 

9. ZOOM ® urea (Invitrogen life technologies, Carlsbad, CA, USA) 

10. DTE aliquots: 65 mM in 1.5 mL of sample solubilization buffer II (working 

solution). Portion 0.015 g (15 mg) of DTE in 1.5 mL Eppendorf tube and store 

at 4°C until use (see Note 7). 

11. Sample solubilization buffer II (stock solution): 7 M urea, 2 M thiourea, 4% 

(w/v) CHAPS. To prepare 25 mL, dissolve 10.5 g of urea, 3.8 g of thiourea in 

10 mL of ddH2O under constant stirring at RT. Fill up to a final volume of 30 

mL with ddH2O(see Note 8).Add5gofmixed bed ion exchanger resin and 

stir for 10 min. Remove beads by filtration through a 0.2 μm bottle top filter. 

Dissolve 1gofCHAPS, add a trace of BPB and portion solution into 1.5 mL 

aliquots in 1.5 mL Eppendorf tubes. Store at –20°C until use. 

12. Sample solubilization buffer II (working solution): 7 M urea, 2 M thiourea, 4% 

(w/v) CHAPS, 1% (w/v) (65 mM) DTE, 2% (v/v) (0.8% (w/v)) Pharmalyte 

3–10. To make up 1.5 mL, add 1.5 mL of sample solubilization buffer II 

(stock solution) to one DTE aliquot in 1.5-mL Eppendorf tube. Add 30 μL of 

Pharmalyte 3–10 and incubate for 15 minutes at RT with occasional vortexing 

until DTE is completely dissolved. This solution is prepared just before use (see 

Note 9).


3. Methods 


Mammalian renal tubular epithelium consists of at least seven different 

segments, complicating biochemical investigation of this heterogeneous tissue. 

Cultured monolayers of dog kidney (Madin-Darby canine kidney (MDCK)) 

cells display many typical features of renal tubular epithelia, such as brush 

border membrane, tight junctions and adherent junctions. MDCK strain II (26) 

was used to prepare samples of cell culture supernatants for 2-DE. High-quality 

protein samples for 2-DE are only obtained from serum-free media. Serumderived 

proteins heavily contaminate cell-derived secreted proteins in culture 

media (see Note 10). Hence it is essential to wash the cells thoroughly (twice 

in PBS or serum-free medium) when changing from complete culture media to 

serum-free conditions. 

1. Harvest confluent MDCK cells by trypsinization. To five confluent 100-mm cell 

culture dishes add 1.5 mL of prewarmed (37°C) Trypsin-EDTA solution per dish 

and incubate at 37°C in a humidified incubator in an atmosphere of 5% CO 2 until 

cells detach (see Note 11). 

2. Resuspend trypsinized cells in culture medium. To each dish add 6.5–7 mL of 

prewarmed (37°C) culture medium and pool resuspended cells from the five 

dishes into one 50-mL Greiner tube. Fill up to a total volume of 40 mL with 

culture medium (see Note 12). 

3. Seed 1.15 × 10 6 cells onto 100-mm cell culture dishes. Adjust final volume with 

prewarmed (37°C) culture medium to 9 mL per dish. Prepare a total of 18 dishes 

per condition (see Note 13). Incubate the cultures for about three days at 37°C 

in a humidified incubator in an atmosphere of 5% CO 2 until cells are confluent 


4. Aspirate medium and wash cells twice in 4 mL of prewarmed (37°C) serum-free 

medium per dish (see Note 15). Add 4 mL of serum-free medium to each dish 

and incubate for 22 h at 37°C in a humidified incubator in an atmosphere of 

5% CO 2. 

5. Harvest cell culture supernatants. Pool media from 18 dishes into one 250-mL 

Erlenmeyer flask to give a final volume of 70–72 mL (see Note 13). Immediately 

proceed to ultracentrifugation and ultrafiltration (see Subheading 3.2.). 

3.2. Ultracentrifugation and Ultrafiltration 

Beside body fluids, such as human plasma, urine, and cerebrospinal fluid, 

cell culture supernatants are among the most difficult samples to be prepared 

for 2-D PAGE. Culture media contain the complete set of interfering substances 

that are incompatible with the first dimensional isoelectric focusing: insoluble 

particles (dead cells, cell debris), nucleic acids (from dead cells), lipids 

(membranes and exosomes (22)), salts (from medium see Note 1), small ionic


compounds (amino acids from medium), phenolic compounds (Phenol red 

from medium) and proteases (secreted during cultivation). Therefore these 

contaminants are removed in a two-step purification strategy: 1) insoluble 

particles, nucleic acids and lipids by high-speed centrifugation; 2) salts, small 

ionic and phenolic compounds by ultrafiltration. Proteases are inhibited by 

addition of inhibitors and basic pH conditions. Alternative methods such as 

TCA/acetone precipitation (27) and dialysis must not be applied. Unspecific 

salt precipitation and loss of proteins are associated with these methods (see 

Note 16). 

1. Add 140 μL of 0.5 M EDTA pH 8.0 and 700 μL of 0.1 M PMSF to 70 mL of 

culture medium in 250 mL Erlenmeyer flask and put on ice (see Note 17). 

2. Transfer 4 × 17.5 mL of medium to four prechilled KONTRON CENTRIKON 

ultracentrifuge tubes. Adjust precise volumes with ddH 2O on an analytical 

balance and put tubes on ice. 

3. Place precooled (4°C) KONTRON CENTRIKON TFT 70.38 fixed-angle rotor 

into KONTRON CENTRIKON T-2060 ultracentrifuge. Place tubes into the rotor 

in appropriate positions and close the lid. Ultracentrifuge for 60 min at 31,200 

rpm (100,000g) at 4°C. 

4. Carefully remove tubes from ultracentrifuge and put on ice. 

5. Rinse Centricon ® Plus-70 components consisting of cap, concentrate/retentate 

cup, sample filter cup and filtrate collection cup with ddH 2O to remove dust 

particles (see Note 18). 

6. Place sample filter cup into filtrate collection cup and leave on ice. 

7. Pool the medium by inverting tubes into the same sample filter cup and close. 

Then place assembled Centricon ® Plus-70 centrifugal filter device into precooled 

(4°C) Sorvall H1000B swinging bucket rotor fixed in a Sorvall RT6000D 

centrifuge. Centrifuge for 60 min at 3,200 rpm (2,190g) at 4°C. 

8. Discard the flow-through in the filtrate collection cup. Add 70 mL of prechilled 

sample solubilization buffer I (20 mM Tris pH 9.0, 1 mM EDTA, 1 mM PMSF) 

into sample filter cup (see Note 19). Centrifuge for 60 min at 3,200 rpm (2,190g) 

at 4°C. 

9. Repeat step 8 twice. 

10. After the last washing step place concentrate/retentate cup upside down onto the 

filtrate collection cup. Invert assembly and place back into centrifuge. Recover 

sample concentrate for 5 min at 2,200 rpm (1,000g) at 4°C. 

11. Determine volume with a pipet. Typical final concentrate volumes are between 

250 μL and 350 μL. 

12. Transfer protein concentrate to a 1.5 mL Eppendorf tube and spin for 5 min 

at 13,200 rpm (16,100g) at 4°C to remove precipitates (see Note 20). Put 

samples on ice and proceed to protein quantitation (see Subheading 3.3.) or store 

at –20°C until use.


3.3. Protein Quantitation by BCA Assay 

Quantitative determination of protein solubilized in modified lysis buffer 

(16) (7 M urea, 2 M thiourea, 4% CHAPS, 65 mM DTE and 2% Pharmalyte 

3–10) is not possible. The Bradford assay (29) cannot be used for two reasons: 

1) Coomassie Brilliant Blue G-250 binds to detergents (CHAPS) and carrier 

ampholytes (Pharmalyte 3–10); 2) Coomassie Brilliant Blue G-250 may not 

bind to protein at all under basic pH conditions (urea). The second problem 

may be remedied by acidification of sample with 0.1 N HCl before quantitation 

(30). Standard Lowry (31), Biuret (32), and BCA (bicinchoninic acid) (33) 

assays based on the reduction of Cu 2+ to Cu + for development of color interfere 

with thiol reducing agents (DTE) and thiourea. Thiourea forms complexes with 

copper ions. TCA/acetone precipitation (27) must not be applied in combination 

with any of these techniques. Protein may be lost on precipitation leading 

to underestimation of solubilized protein. The best solution to all problems 

mentioned above is quantitation of protein before solubilization in lysis buffer. 

In this section the BCA assay from Pierce is used to accurately quantitate 

protein concentration in sample solubilization buffer I (20 mM Tris pH 9.0, 

1mM EDTA, 1 mM PMSF). Although the BCA assay interferes with chelating 

agents (EDTA) concentrations below 10 mM are tolerated (34). 

1. Prepare a set of albumin (BSA) standards in 1.5-mL Eppendorf tubes (see 

Table 1 for details). Use ddH 2O as diluent. Gently vortex tubes. There will be 

sufficient volume for two replications of each diluted standard. 

2. Prepare protein samples in 1.5-mL Eppendorf tubes. Dilute 5 μL of each protein 

concentrate to a final volume of 100 μL in ddH 2O (1:20 dilution). Gently vortex 

tubes. While performing BCA assay put undiluted protein concentrates on ice 

(see Subheading 3.2. step 12). 

Table 1 

Preparation of diluted albumin (BSA) standards 

Vial Volume of diluent a Volume and source of BSA Final BSA concentration 

A 150 μL 150 μL of stock b 1000 μg/mL 

B 25 μL 75 μL of A 750 μg/mL 

C 50 μL 50 μL of A 500 μg/mL 

D 75 μL 25 μL of A 250 μg/mL 

E 88 μL 12.5 μL of A 125 μg/mL 

F 98 μL 2.5 μL of A 25 μg/mL 

a Use ddH2O to prepare albumin (BSA) standards. 

b The concentration of albumin standard stock solution is 2 mg/mL.


3. Pipet 25 μL of each standard or unknown sample replicate into a microplate 

well. Standards are applied in duplicates, protein samples in triplicates. Use 

ddH 2O for blanks. 

4. Prepare BCA working reagent by mixing 50 parts of BCA TM Reagent A with 1 

part of BCA TM Reagent B in a 25 mL beaker. Mix thoroughly (see Note 21). 

For each standard, unknown sample or blank 200 μL of BCA working reagent 

is required. Include two extra replicates in your calculation. 

5. Add 200 μL of BCA working reagent to each well. Gently shake microplate by 

hand for a few seconds. 

6. Incubate microplate for 30 min at 37°C in an incubator. 

7. Cool plate to RT and measure absorbance at 550 nm on a microplate reader (see 

Note 22). 

8. Subtract the average 550 nm absorbance measurement of the blank replicates 

from the 550 nm absorbance measurements of all other individual standards and 

unknown sample replicates. 

9. Prepare a standard curve by plotting the average blank-corrected 550 nm 

absorbance measurement (A 550, y-axis) for each albumin standard versus its 

amount (in μg, x-axis) in increasing order (see Note 23). 

10. Use the standard curve to determine the protein amount of each unknown 

sample (Fig. 2). The protein concentration in unknown sample is calculated as 

follows: (protein amount of unknown in μg)/(volume of diluted sample in μL) × 

(dilution factor)= protein concentration in mg/mL (see Note 24). Typical protein 

concentrations are between 5 mg/mL and 7 mg/mL. 

11. Portion undiluted protein concentrates into appropriate aliquots (80 μg for 

analytical load) in 1.5 mL Eppendorf tubes and store at –20°C until use (see 

Subheading 3.4.1). 

3.4. Rehydration Loading and Isoelectric Focusing 

Traditionally, protein samples prepared in standard lysis buffer (O’Farrell 

1975) (1) were loaded onto the basic end (cathode) of an isoelectric focusing 

(IEF) tube gel. Before sample loading the gel rods were prerun to establish a 

carrier ampholyte-derived pH gradient. On development of fixed pH gradients 

samples were applied with rubber frames or sample cups to rehydrated immobilized 

pH gradient (IPG) strips at the acidic or basic end (5–8) or simultaneously 

at both ends (35). The problem with cup-loading sample application 

techniques is that proteins may precipitate during the sample entry phase, which 

leads to horizontal streaking at the sample application point. This problem is 

remedied by in-gel sample rehydration where protein solubilized in lysis buffer 

is directly diluted with the rehydration solution used for IPG strip reswelling 

(36,37). Unfortunately, some proteins that are soluble in lysis buffer may be 

lost on dilution into rehydration solution because of lower concentrations of 

chaotropic agents and detergents. For simplicity protein sample preparation


Fig. 2. Typical color response curve for albumin (BSA) standards using the BCA 

assay. Each point represents the mean of two replications. 

and rehydration can be done all-in-one in modified lysis buffer (16) (7 M 

urea, 2 M thiourea, 4% CHAPS, 65 mM DTE and 2% Pharmalyte 3–10). This 

strategy works very well in combination with the Multiphor II horizontal flatbed 

isoelectric focusing system with final maximum voltage limited to 3,500 V 

for steady-state IEF (38). Higher voltage settings may become problematic, 

because zwitterionic detergent (CHAPS), reducing agent (DTE) and carrier 

ampholytes (Pharmalyte 3–10) heavily contribute to the current in the 

strip. 

1. Thaw protein samples on ice (see Subheading 3.3.11). Dilute each aliquot of 

protein solution (80 μg for analytical load) to a final volume of 450 μL in sample 

solubilization buffer II (see Note 25). 

2. Gently vortex tubes and solubilize protein for 60 min on a rotary shaker at RT. 

3. Centrifuge for 30 min at 13,200 rpm (16,100g) at 22°C in a tabletop centrifuge 

to remove insoluble particles. 

4. Slide the protective lid completely off the Immobiline TM DryStrip Reswelling 

Tray and level the tray by turning the leveling feet until the bubble in the spirit 

level is centered. 

5. Remove IPG strips (240 mm long, 3 mm wide ready-made Immobiline TM 

DryStrips pH 3–10 NL cast on GelBond PAGfilm) from the freezer and warm 

up to RT.


6. Evenly apply the entire sample-containing solution into the groove of the 

reswelling tray (see Note 26). 

7. Peel off the protective cover sheet from the ready-made IPG strip starting at the 

acidic (+) end and grip the strip with tweezers at the overlapping basic plastic 

end (see Note 27). 

8. Slowly lower the IPG strip (gel side down) onto the solution with the 

acidic (+) end oriented towards the number labels of the reswelling tray (see 

Note 26). 

9. Cover the strip with 3 mL of paraffin oil (see Note 28). Repeat steps 6–9 for 

each sample. 

10. Slide the lid onto the reswelling tray and rehydrate the IPG strips overnight 

at RT. 

11. To remove rehydrated IPG strips sequentially from the reswelling tray, open 

the lid, slide the tip of tweezers along the sloped end of the slot and into the 

slight depression under the IPG strip. Grab the acidic (+) end of the strip with 

tweezers and lift the strip out of the tray. While still holding the strip, rinse it 

briefly with ddH 2O. Place it on a piece of damp filter paper at one edge to drain 

off excess liquid. Repeat procedure for each strip. 

12. Set the temperature on the MultiTemp TM III thermostatic circulator to 20°C. 

13. Place the ceramic cooling plate in Multiphor II unit and make sure the surface 

is level. 

14. Starting at the top of the plate near the cooling tubes pipet 1 mL of paraffin oil 

in a straight line onto the middle of the plate. Use the grid of the cooling plate 

as a guide. Then pipet 1 mL of paraffin oil on each side of the paraffin oil line 

(a total of 3 mL) onto the bottom of the plate. The additional 1 mL of paraffin 

oil on each side is evenly spread on the cooling plate to form a triangle which 

begins with the base line at the bottom and extends to the middle of the paraffin 

oil line (see Note 29). 

15. Slowly lower the Immobiline TM DryStrip tray onto the bottom of the paraffin 

oil triangle with the red (anodic, positively charged) electrode connection of the 

tray positioned at the top of the plate near the cooling tubes. 

16. Connect the red and black electrode leads on the tray to the Multiphor II unit. 

17. Pour 10 mL of paraffin oil into the tray at the bottom of the cooling plate. 

18. Slowly lower the Immobiline TM DryStrip aligner, 12 grooves side up, onto the 

bottom of the paraffin oil layer next to the black electrode. 

19. Transfer the rehydrated IPG strips with tweezers to adjacent grooves of the 

aligner in the tray. Place the strips gel side up with the acidic (+) end at the top 

of the tray near the red electrode. 

20. Cut one IEF electrode strip into two pieces each to a length of 110 mm. Moisten 

the two IEF electrode strips with deionized water (see Note 30). 

21. Place the damp electrode strips across the acidic and basic ends of the aligned 

IPG strips.


22. Align each electrode over an electrode strip, ensuring the marked side corresponds 

to the side of the tray giving electric contact (see Note 31). When the 

electrodes are properly aligned, press them down to contact the electrode strips. 

23. Pour 100 mL of paraffin oil into the tray to cover the IPG and electrode strips. 

24. Close the lid of the Multiphor II unit. Connect the leads on the lid to the EPS 

3501 XL power supply and start IEF according to programmed parameters (see 

Note 32). 

25. After IEF remove electrodes and IEF electrode strips. 

26. Grip each IPG strip with tweezers at the overlapping basic plastic end, carefully 

remove it from the tray and rinse it with ddH 2O. 

27. Place each IPG strip into a petri dish with the plastic side of the strip facing the 

inner wall of the petri dish and cover. Seal it with a piece of parafilm and store 

at –20° C until use. 

28. IPG strip equilibration, SDS-PAGE and postseparation visualization techniques 

applied following sample preparation and IEF are not topic of this chapter. 

Useful tips and tricks concerning these methods can be found in the Amersham 

2-D electrophoresis handbook (40). As an example the final 2-D map of Madin- 

Darby canine kidney (MDCK) cell culture supernatant is shown in Fig. 1. 

4. Notes 

1. Composition of Minimum Essential Medium (MEM) (25): 264 mg/mL CaCl2 2H2O, 400 mg/mL KCl, 200 mg/mL MgSO4 7H2O, 6800 mg/mL NaCl, 

2200 mg/mL NaHCO3, 158 mg/mL NaH2PO4 2H2O, 1000 mg/mL 

d-glucose, 10 mg/mL Phenol red, 126 mg/mL l-arginine HCl, 24 mg/mL 

l-cystine, 42 mg/mL l-histidine HCl H2O, 52 mg/mL l-isoleucine, 52 

mg/mL l-leucine, 73 mg/mL l-lysine HCl, 15 mg/mL l-methionine, 32 mg/mL 

l-phenylalanine, 48 mg/mL l-threonine, 10 mg/mL l-tryptophan, 36 mg/mL 

l-tyrosine, 46 mg/mL l-valine, 1 mg/mL d-Ca pantothenate, 1 mg/mL choline 

chloride, 1 mg/mL folic acid, 2 mg/mL i-inositol, 1 mg/mL niacinamide, 1 

mg/mL pyridoxine HCl, 0.1 mg/mL riboflavin, and 1 mg/mL thiamine HCl. 

2. Before use fetal bovine serum (FBS) is heat-inactivated! Incubate one bottle 

(500 mL) of FBS for 30 min at 56°C in a water bath. Portion solution into 

25 mL aliquots and store at –20°C. 

3. The EDTA may not completely dissolve in 0.5 M stock solution below pH 8.0. 

Therefore titration of EDTA stock solution with a few drops of 5 N NaOH 

(liquid) is necessary. The solution is ready when the pale white color turns into 

a crystal clear solution. 

4. PMSF is very toxic! Protect your eyes and skin. PMSF is not very stable in 

water and has a half-life of about 30 min. Hence the solution is prepared in 

isopropanol. PMSF is difficult to dissolve at RT; therefore the solution is warmed 

up to 37°C. 

5. The 0.2 M Tris stock solution has a pH of 10.5–10.6. Do not titrate with HCl! 

The chloride ions extremely contribute to the current (heat production) during


isoelectric focusing (IEF). Any heat produced during IEF will cause protein 

precipitation and produce horizontal streaks in the final 2-D gel. 

6. The sample solubilization buffer I has a pH of 9.0–9.1. Do not titrate with HCl! 

The Tris serves as a positively charged ion that helps in solubilization of proteins 

and not to maintain constant pH (see Note 19). This solution should be prepared 

shortly before use because of the poor stability of PMSF in water (see Note 4). 

The 0.1 M PMSF stock solution should be warmed up to 37°C in a water bath. 

Otherwise the PMSF may precipitate! 

7. DTE is not very stable in solution. Hence it is stored as solid in small aliquots 

at 4°C. 

8. The final volume of 30 mL compensates for the dead volume of the magnetic 

stir bar in a 50-mL beaker and equals to a total volume of 25 mL. Add urea 

and thiourea in small portions with the help of a spatula under constant stirring 

at RT. Urea and thiourea will cool down the solution and decrease solubility! 

Therefore in between additions wait for several minutes until each small portion 

is fully dissolved. Do not heat urea-containing solutions above 37°C to avoid 

carbamylation of proteins! 

9. The sample solubilization buffer II (working solution) should be prepared 

freshly. Never reuse remaining buffer, better discard it! 

10. It is essential to wash the cells thoroughly (twice in PBS or serum-free medium) 

to remove serum proteins. Culture media heavily contaminated with fetal bovine 

serum (FBS) resemble human plasma! 

11. Before trypsinization confluent cells may be thoroughly washed (twice in 2–3 mL 

of PBS or serum-free medium) (see Note 10). Prewarm PBS and Trypsin-EDTA 

solution to 37°C in a water bath! 

12. To each dish treated with 1.5 mL of Trypsin-EDTA solution 6.5–7 mL of culture 

medium is added to give a total volume of 40 mL. Prewarm culture medium to 

37°C in a water bath! 

13. In total 18 dishes per condition are prepared. The final volume of culture medium 

used per dish is 9 mL. Once MDCK cells reach confluence serum-free medium is 

added. The final volume of serum-free medium used per dish is 4 mL. This gives 

a total volume of 70–72 mL per condition and corresponds to the appropriate 

processing volume for the Centricon ® Plus-70 centrifugal filter devices (see 

Note 18). 

14. The doubling time of MDCK strain II cells is one day. Confluence is reached 

after 3–4 days. 

15. Alternatively, use PBS instead of serum-free medium if costs are a major 

concern. 

16. TCA/acetone (27) may precipitate calcium-phosphate and small amino acids 

from the culture media. Very high contaminant concentrations may be achieved 

that extremely interfere with isoelectric focusing. Dialysis must not be used! 

Very high solute volumes are needed to remove salts and unspecific protein 

binding to the dialysis membrane may occur.


17. The 0.1 M PMSF stock solution is warmed up to 37°C in a water bath. PMSF 

is added before putting medium on ice to avoid precipitation. EDTA inhibits 

metalloproteases by chelation of free metal ions. PMSF inhibits serine proteases 

and some cysteine proteases. Inhibitor cocktails (for example Complete Mini, 

EDTA-free from Roche) must not be used! These cocktails contain protein- and 

peptide-based inhibitors that can reach very high concentrations on ultrafiltration 

(up to 300X) and hence abundantly mask the secreted proteins present in the 

medium. 

18. Usage guidelines for Centricon ® Plus-70 centrifugal filter devices are given in 

the user guide (28). The filter material is made of a polyethersulfone Biomax 

membrane with a 5 kDa cut-off. 

19. The pH of 9.0 from Tris serves two functions: 1) it maximizes protein extraction 

at basic pH conditions (almost any protein is in deprotonated state); 2) minimizes 

protease activity. The Tris itself serves as a positively charged ion that helps in 

solubilization of proteins (see Note 6). Very basic proteins may be lost! 

20. On concentration protein precipitation may occur! 

21. When Reagent B is first added to Reagent A, turbidity is observed that quickly 

disappears on mixing to yield a clear, green color. 

22. Alternatively, wavelengths from 540–590 nm may be used with this 

method (34). 

23. Amount of albumin standards (see Table 1) used: F, 0.625 μg; E, 3.125 μg; D, 

6.25 μg; C, 12.5 μg; B, 18.75 μg, and A, 25 μg. The standard amount is referred 

to a volume of 25 μL. Average blank-corrected 550 nm absorbance values above 

0.8 must not be used for standard curve preparation! 

24. For example: (protein amount of unknown is 7 μg)/(volume of diluted sample 

is 25 μL) × (dilution factor is 20) = 5.6 mg/mL. 

25. A final volume of 450 μL is recommended by the supplier (Amersham 

Biosciences) for rehydration of one 24 cm Immobiline TM DryStrip. For first trial 

prepare a duplicate! The upper limit is twelve samples per run. 

26. Avoid trapping of air bubbles. 

27. Do not wear gloves during removal of protective cover sheet! The rubber material 

tends to stick to the “naked” gel and hence will damage it. 

28. Overlaying of IPG strips with paraffin oil reduces risk of urea crystallization 

during rehydration. 

29. In this case the paraffin oil evenly distributes the heat produced during IEF 

between the tray and the cooling plate. 

30. Do not use ddH 2O or tap water! The former leads to very low conductivity 

between electrode and IPG strip, the latter to very high. 

31. Each electrode has a side marked red or black. 

32. Program for 24 cm IPG pH 3–10 NL strips using the EPS 3501 XL power 

supply (adapted from Hoving (40)): Phase 1, 300 V, 1 Vh (0.006 h); Phase 2, 

300 V, 900 Vh (3 h); Phase 3, 3500 V, 9500 Vh (5 h), and Phase 4, 3500 V, 

52500 Vh (15 h). Current and power are set non-limiting (2 mA, 5 W). Phases 

1–4 are programmed in the gradient mode. The voltage will be ramping up to


the maximum set in the phase, starting from zero in the first phase and in phases 

to follow from the end point of the phase before. Therefore in phases 1 and 3 

voltage is linearly increased and in phases 2 and 4 held constant. The current 

check option must be switched off! 


The authors wish to acknowledge and thank Ursula Luginbühl for excellent 

technical assistance. This work was funded by the Swiss National Science 

Foundation (SNSF) (grant 3100A0-100772 to E.E.S.) and the European Science 

Foundation (ESF) Integrated Approaches for Functional Genomics (grant 0341 

to D. A.). 

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Centricon ® Plus-70 centrifugal filter devices from Millipore)


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11 

Sample Preparation for Mass Spectrometry Analysis 

of Formalin-Fixed Paraffin-Embedded Tissue 

Proteomic Analysis of Formalin-Fixed Tissue 

Nicolas A. Stewart and Timothy D. Veenstra 

Summary 

One of the great hopes in biomedical research is that proteomic technology can be used 

to identify novel biomarkers for diseases such as cancer. The challenge to discovering 

biomarkers starts with sample collection and continues right through data acquisition and 

bioinformatic analysis. Because the ultimate goal is to find indicators of human disease it 

is ideal to be able to study clinical samples. Unfortunately clinical samples such as serum, 

plasma, urine, and especially tissue biopsies are precious and are often difficult to obtain 

in sufficient quantities or numbers to conduct proteomic discovery studies. There exists, 

however, a vast archive of pathologically characterized clinical samples in the form of 

formalin fixed paraffin embedded tissue blocks. This chapter describes methods that have 

been developed to allow the proteins from these tissue samples to be excised in a form 

that is amenable for proteomic analysis by mass spectrometry. 

Key Words: Cancer biomarkers; formalin-fixed paraffin embedded tissue; mass 

spectrometry; proteomics; tissue microdissection. 


One of the greatest determinants on the survival rate from any cancer is 

the stage at which it is detected. The survival rate from cancers detected at an 

early stage is generally higher but decreases as the stage of detection increases. 

Therefore a major impetus in proteomics research is to identify biomarkers 

of early stage diseases. Many of these biomarker discovery efforts are aimed 

towards the use of biofluids such as serum and plasma. These sample types, 



131

132 Stewart and Veenstra 

however, are inherently difficult to analyze and are often too precious to be 

given over to proteomic-type discovery studies that are continuing to evolve in 

both efficacy and success. Another complication is that although biomarkers 

that originate from the site of a tumor may have a high local concentration, 

by the time they become diluted within the circulatory system, their effective 

concentration in the acquired clinical sample may be vanishingly small. Ideally, 

it would be beneficial to identify a biomarker at the tumor level and determine 

if it could be translated to a marker detectable in serum or plasma. Tissue 

or tumor biopsies therefore, are ideal sample candidates for the discovery of 

potential biomarkers. However, the feasibility of their use for such studies is 

hampered because of the fact that they are relatively difficult to obtain and 

require careful storage and handling. 

Formalin-fixed, paraffin-embedded (FFPE) tissues, on the other hand, 

represent a vast resource for retrospective protein biomarker investigation. 

Formalin fixation and paraffin embedding of tissue is practiced in pathology 

labs worldwide as a standard processing method by which tissues can be 

stored and catalogued as stable entries. Unfortunately, the storage stability 

of these tissues partially arises from the high degree of covalently crosslinked 

proteins. Currently, immunohistochemistry (IHC) is the only published 

technology capable of providing protein information from these samples (1,2). 

In addition, because IHC requires a priori knowledge of individual proteins 

being analyzed, it is not a discovery-based technology. 

As with tissue biopsies, FFPE sections are heterogeneous in that they 

contain many different types of cells. To acquire a homogeneous population 

of cells, laser-capture microdissection (LCM) is used. This technique has the 

ability to directly isolate a user-defined population of cells from their tissue 

microenvironment (3). Although LCM of fresh tissue is widely practiced, 

outside of the interest in conducting tissue microdissection of FFPE tissues for 

mRNA extraction, microdissection of FFPE tissues is not widely practiced for 

extraction and analysis of soluble protein (4). 

Mass spectrometry (MS) is arguably the driving technology in discoverydriven 

proteomics. Dramatic technical improvements in MS instrumentation 

combined with the rapid growth of genomic and proteomic databases have 

enabled development of approaches to identify and quantify large numbers of 

proteins from complex samples such as serum and tissues (5). Combining LCM 

of FFPE tissues and MS has the potential for generating protein biomarker data 

necessary for discovering proteins that are key determinants or indicators of 

diseases such as cancer. 

A common misnomer is that MS identifies proteins in proteomic studies. 

Actually in its present form, MS is best suited to the identification of peptides 

that are produced from the enzymatic digestion of proteins. It is through

Sample Preparation for Mass Spectrometry Analysis 133 

the analysis of peptide surrogates that proteins are identified. Because of 

their size, peptides are very amenable to tandem MS (MS/MS) methods that 

produce partial amino acid sequence information that leads to their identification. 

Besides the advances made in MS instrumentation, its coupling with 

nanoflow reversed-phase liquid chromatography (nanoRPLC) has made the 

analysis of complex peptide mixtures possible. To compare the tryptic peptide 

abundances obtained from two different FFPE tissues, stable-isotope labeling 

using trypsin-mediated 16 O/ 18 O incorporation may also be employed. 

2. Materials 

2.1. Tissue Processing 

1. Mayer’s hematoxylin stain. 

2. Eosin stain. 

3. Graded ethanol solutions. 

4. SubX organic solvent (Surgipath Medical Industries, Richmond, IL). 

2.2. Protein Extraction 

1. 50% glycerol. 

2. LCM instrument. 

3. Low-binding microcentrifuge tubes. 

4. The Liquid Tissue-MS protein prep kit (Expression Pathology, Inc., 

Gaithersburg, MD). 

2.3. Trypsin Digest 

1. Porcine sequencing grade modified trypsin (Promega, Madison, WI). 

2. Dithiothreitol (DTT, Sigma, St. Louis, MO). 

3. Trifluoroacetic acid (TFA) (≥ 98.0% pure). 

4. Incubator for the digest at 37°C in microcentrifuge tubes. 

5. High performance liquid chromatography (HPLC)-grade acetonitrile (ACN) 

(EMD Chemicals Inc., Gibbstown, NJ). 

6. C-18 reverse phase microcolumns (e.g. ZipTip®, Millipore, Billerica, MA). 

7. Conditioning, loading and eluting buffers for reverse phase desalting microcolumns: 

ACN, 0.1% TFA in ddH2O, and 60% ACN, 0.1% TFA solution, respectively. 

2.4. LC-MS/MS analysis of tryptic peptides 

1. HPLC system capable of NanoRPLC (e.g., Agilent 1100 capillary LC system, 

Agilent Technologies, Palo Alto, CA). 

2. Fused silica capillary column: 75 μm inner diameter × 360 outer diameter × 10 

cm long, slurry packed with C-18 silica-bonded stationary phase; 3 μm, 300 Å 

pore size.


3. Linear ion trap MS coupled to the NanoRPLC (e.g., LTQ, Thermo Electron, San 

Jose, CA). 

4. High resolution MS coupled to the NanoRPLC for 18 O isotope labeling analysis 

(e.g., LTQ-FTICR, Thermo Electron, San Jose, CA). 

5. Trifluroroacetic acid is diluted to 0.1% (v/v) and is used to resolubilize tryptic 

peptides before LC-MS/MS. 

6. A solution of 0.1 % (v/v) formic acid (FA) in ddH 2O (NANOPure Diamond water 

system, Barnstead International, Dubuque, IA) (Mobile Phase A) and a solution 

of 0.1% FA in HPLC-grade ACN (Mobile Phase B) are prepared for the gradient 

used in the reverse phase liquid chromatographic separation of tryptic peptides. 

2.5. Mass Spectrometry Analysis and Bioinformatic Analysis 

1. Accompanying software for the collection of the raw MS/MS data generated. 

2. Software to search raw data files to a database (e.g., SEQUEST). 

3. Methods 

3.1. Tissue Processing 

1. For tissue microdissection, 10 μm thick tissue sections are cut from FFPE whole 

mount tissue blocks and placed on coated slides. 

2. The section is then heated for 60 min at 58°C. To remove paraffin, treat the 

section with SubX organic solvent (Surgipath Medical Industries, Richmond, IL) 

twice for 5 min. 

3. The tissue is then rehydrated through multiple, graded ethanol solutions and 

distilled water. Tissue is then counterstained with Mayer’s hematoxylin, and 

dehydrated through graded ethanol solutions, and air-dried. 

3.2. Laser-capture Microdissection 

1. Rehydrate the tissue with 50% glycerol in water for 5 min before laser capture 

microdissection (LCM). 

2. Place the slide containing the tissue upside-down below the objective lens and 

locate cellular regions with specific histological features of interest (see Note 1). 

3. Capture cells with the LCM instrument utilizing an excimer laser (MPB 

Technologies PSX-100) operating at the following conditions: 248 nm 

wavelength, 2.5 ns pulse, Emax= 5 mJ, repetition rate = 0.1–100 Hz (see Note 2). 

4. Captured cells within the selected area are then transferred into a 1.5 mL 

low-binding microcentrifuge receiving tube. Approx 100,000–200,000 cells are 

required for subsequent proteomic analysis (see Note 3). 

3.3. Protein Extraction and Trypsin digest 

1. Cells collected by microdissection for nano-reversed-phase liquid 

chromatography-tandem mass spectrometry (RPLC-MS/MS) analysis were


processed using proprietary reagents according to the manufacturer’s recommendations 

(Liquid Tissue, Expression Pathology Inc., Gaithersburg, MD). 

2. The material for nanoRPLC-MS/MS analysis was suspended in 20 μL of Liquid 

Tissue reaction buffer, and incubated for 90 min at 95°C, followed by cooling 

on ice for 3 min. 

3. DTT is added to a final concentration of 10 mM. 

4. Heat samples for 5 min at 95°C, and let cool to room temperature. 

5. Trypsin (15–18 units) is added and the sample is incubated at 37°C for 18 h. 

6. The resulting proteome digestates may be stored at –20°C until analysis. 

7. Tryptic peptides generated from the comparative FFPE samples are desalted 

using C-18 microcolumns. Microcolumns are conditioned by aspirating and 

dispensing with conditioning buffer (10 μL, 3×), followed by loading buffer 

(10 μL 3×). 

8. Peptide samples are re-solubilized in loading buffer (10 μL) and washed with 

loading buffer (10 μL 3×). 

9. Peptides are eluted from the microcolumn with the elution buffer (10 μL) into 

microcentrifuge tubes, and lyophilized. 

10. Peptide samples are re-solubilized in loading buffer (10 μL) and transferred to 

vials for autosampler. 

3.4. Trypsin-mediated 18O-Labeling 1. For isotope labeling, the protein extracts from equivalent numbers of cells 

16 18 microdissected from FFPE tissues are reconstituted separately in H2 O and H2 O 

each containing 20% methanol (v/v). 

2. Sequencing grade trypsin is resuspended in the appropriately labeled water (i.e. 

16 18 H2 OorH2 O), and added to the related sample at an enzyme to protein ratio 

of 1:20. 

3. Incubate at 37°C for 16 h. After this time, an additional equivalent aliquot of the 

trypsin is added and the samples incubated at 37°C for an additional 6h(see 

Note 7). 

4. TFA is added to a final concentration of 0.4% (v/v) to stop the reaction (see Note 8). 

5. The differentially labeled proteome samples are pooled and lyophilized to dryness. 

6. Samples are resolubilized in loading buffer (10 μL) and transferred to vials for 

autosampler. 

3.5. Nanoflow RPLC-MS/MS Analysis 

1. Nanoflow RPLC is performed on a capillary LC system coupled online to a ion 

trap MS or an MS capable of high resolution (i.e., 100,000) for quantitation of 

16 18 O/ O-labeled samples (see Note 9). 

2. Wash column for 30 min with 98% mobile phase A at a flow rate of 0.5 μL/min 

prior to sample injection. 

3. Inject sample (typically 1–6 μL) onto the column.


4. Elute peptides using a linear gradient of 2–40% mobile phase B in 110 min, then 

to 98% mobile phase B in an additional 30 min, all at a constant flow rate of 0.25 

μL/min. 

3.6. Mass Spectrometry Analysis and Bioinformatic Analysis 

1. The MS is operated in a data dependent MS/MS mode in which each full MS scan is 

followed by five MS/MS scans of the five most abundant peptide molecular ions. 

2. Subject peptides to collision-induced dissociation (CID) using a normalized 

collision energy of 35% (see Note 10). The heated capillary temperature and 

electrospray voltage of the mass spectrometer are typically set at 160°C and 

1.5 kV, respectively. 

3. Although data collection may be dictated based on the available mass to charge 

(m/z) range of the instrument, it is typically collected over a broad range of 

400–2,000. 

4. Because the starting amount of protein obtained from the LCM cells is low, 

multidimensional fractionation prior to MS analysis is inefficient owing to sample 

handling losses associated with chromatography. 

5. To maximize the number of peptides identified, segmented precursor selection 

scan ranges (i.e., gas phase fractionation in the m/z dimension, GPFm/z) are 

used. The following overlapping m/z intervals may be used as a guideline: 

m/z 400–605, 595–805, 795–1005, 995–1,205, 1,195–1,405, 1,395–1,605, 

1,595–1,805, 1,795–2,000, 400–805, and 795–1,200. Data for the 16O/ 18O-labeled experiments are acquired using FTICR detection in centroid mode for the full MS 

scan (m/z 400–2000) at 100,000 resolution followed by MS/MS of the top five 

most abundant molecular ions detected. 

6. The tandem mass spectra are searched against the UniProt human 

proteomic database from the European Bioinformatics Institute (http:// 

www.ebi.ac.uk/integr8) using a combination of database and searching algorithm 

software (e.g. SEQUEST). Peptides are searched using fully tryptic cleavage 

constraints and a dynamic 4.008 amu modification on the C-terminus for when 

18O isotope labeling analysis. When using SEQUEST, a legitimately identified 

peptide should have cross correlation (Xcorr) scores of 1.9 for [M+H] 1+ , 2.2 for 

[M+2H] 2+ and 3.5 for [M+3H] 3+ and a minimum delta correlation score (Cn) of 0.08 

4. Notes 

1. Microdissection is best performed using software-directed laser pulses to strike at 

a constant velocity and rate throughput over the previously defined and mapped 

cellular regions. 

2. Target slides are optically transparent quartz coated with an energy transfer 

coating with the exact dimensions of a standard histology glass slide. The slide 

stage is a computer-controlled, XY translation stage. A 1/8 beam-splitter is


used to split the laser to an energy meter with the remaining beam traveling 

to an ultraviolet reflective mirror and directed down (-Z) to a ×10 microscope 

objective (LMU-10X-UVR, OFR). The objective focuses the laser onto the slide 

and the LCM process is observable using a confocally aligned CCD camera. 

3. Because many FFPE tissues have been in storage for years, if not decades, an 

obvious concern when conducting proteomic investigations of these samples is 

the effect of long-term storage. Presently there are not enough studies on this 

topic to make any solid conclusions, however, no obvious detrimental effect has 

been observed in those studies that have examined FFPE tissues using MS (8,9). 

One of the studies showed that formalin-fixation and storage does not result in 

an inordinate degree of oxidation to methionyl and cysteinyl residues nor were 

any adverse effects on the tryptic digestion efficiency observed (8). 

4. Unfortunately the maximum number of cells that can typically be microdissected 

from FFPE tissues is on the order of 200,000. Although this number will 

vary depending on the tissue and its heterogeneity, it typically does not yield 

enough protein to employ multidimensional fractionation before MS analysis. 

The protein yield for a typical experiment is on the order of 10–20 μg, whereas 

typically 200 μg is required for a multidimensional fractionation consisting of 

strong cation exchange prior to RPLC to be employed. 

5. There are experimental and data analysis issues related to quantitative proteomics 

whether it is done using O 16 /O 18 labeling or subtractively (i.e., when the number 

of peptides identified from one proteome sample is compared to that identified 

in another). The trypsin-mediated incorporation of O 18 is not always absolutely 

complete because of the incomplete exchange at the peptide’s carboxy-terminus. 

Although no absolute reason for this effect has been established, it may be 

because of the low abundance of the peptide within the complex mixture. Manual 

analysis of the MS spectrum should always be conducted when a potential 

interesting abundance change is suggested. When using a subtractive proteomics 

approach, the precision level is such that abundance changes below three-fold 

are not considered reliable. 

6. A constant issue when dealing with the identification of large numbers of 

peptides from complex mixtures is the false positive rate. It is impossible to 

orthogonally validate all of the peptides/proteins identified in these studies, 

therefore acceptable filtering criteria are necessary when evaluating correct 

identifications. SEQUEST is a commonly used software program used for identifying 

peptides based on tandem MS spectra. The filtering criteria based on X corr 

andC n scores are provided as a guideline only. In bioinformatic analyses, they 

have been shown to provide a false positive rate of approx 1.5% (5). 

7. Because the proteins are extracted from the FFPE tissues as tryptic peptides, the 

role of trypsin during this step is to enzymatically exchange the C-terminal carboxyl 

oxygen atoms with the appropriate oxygen (i.e., 16 Oor 18 O) isotope (6). 

8. An alternative means to stop the trypsin-mediated incorporation of 18 O is to boil 

the sample at the end of the digest. This is acceptable, but the sample should


not be boiled after addition of acid as deamidation of asparagine or glutamine 

residues may occur. 

9. An MS capable of high resolution (e.g., FTICR-MS) is necessary for accurately 

quantifying the different isotopically-labeled counterparts from the two proteome 

samples to be compared. 

10. To minimize the selection of peptides that have already been subjected to CID, 

dynamic exclusion is used. While this is a user defined value, 90 s. is typical. 


This project has been funded in whole or in part with federal funds from the 

National Cancer Institute, National Institutes of Health, under Contract N01- 

CO-12400. The content of this publication does not necessarily reflect the views 

or policies of the Department of Health and Human Services, nor does mention 

of trade names, commercial products, or organization imply endorsement by 

the United States Government. 

References 

1. MacIntyre, N. (2001) Unmasking antigens for immunohistochemistry. Br. J. Biomed. 

Sci. 58, 190–96. 

2. Shi, S. R., Cote, R. J. and Taylor, C. R. (2001) Antigen retrieval techniques: current 

perspectives. J. Histochem. Cytochem. 49, 931–37. 

3. Emmert-Buck, M. R., Bonner, R. F., Smith, P. D., et al. (1996) Laser capture 

microdissection. Science 274, 998–1001. 

4. Gillespie, J. W., Best, C. J., Bichsel, V. E., et al. (2002) Evaluation of non-formalin 

tissue fixation for molecular profiling. Am. J. Pathol. 160, 449–57. 

5. Peng, J. and Gygi, S. P. (2001) Proteomics: the move to mixtures. J. Mass Spectrom. 

36, 1083–91. 

6. Yao, X., Freas, A., Ramirez, J., Demirev, P. A., and Fenselau, C. (2001) Proteolytic 

18O labeling for comparative proteomics: model studies with two serotypes of 

adenovirus. Anal. Chem. 73, 2836–42. 

7. Yates, J. R. III, Eng, J. K., McCormack, A. L., and Schieltz, D. (1995) Method to 

correlate tandem mass spectra of modified peptides to amino acid sequences in the 

protein database. Anal. Chem. 67, 1426–36. 

8. Hood, B. L., Darfler, M. M., Guiel, T. G., et al. (2005) Proteomic analysis of 

formalin-fixed prostate cancer tissue. Mol. Cell. Proteomics 4, 1741–53. 

9. Crockett, D. K., Lin, Z., Vaughn, C. P., Lim, M. S., Elenitoba-Johnson, K. S. 

(2005) Identification of proteins from formalin-fixed paraffin-embedded cells by 

LC-MS/MS. Lab. Invest. 85, 1405–15.

12 

Metalloproteomics in the Molecular Study of Cell 

Physiology and Disease 

Hermann-Josef Thierse, Stefanie Helm, and Patrick Pankert 

Summary 

Physical and chemical stresses as well as metal-related diseases can disrupt the 

normal trafficking of metal ions. Moreover, homeostatic imbalance of such metal ions 

may modulate essential cellular functions (including signal transduction pathways), may 

catalyze oxidative damage, and may affect the folding of nascent proteins. Here we 

describe a new qualitative subproteomic method for the detection, isolation, and identification 

of metal-interacting proteins. Combining both classical immobilized metal ion 

affinity chromatography (IMAC) and modern proteomic techniques (e.g., two dimensional 

gel electrophoresis [2-DE]), metal-specific proteins have been successfully isolated and 

identified to define a metalloproteome. These metal-specific proteomes may give new 

insights into metal-related pathophysiological processes, such as the allergic reaction to 

nickel, which represents the most common form of human contact hypersensitivity. 

Key Words: 2-DE; 2-dimensional gel electrophoresis; disease proteomics; IMAC; 

immobilized metal ion affinity chromatography; metal affinity; metalloproteome; nickel; 

nickel allergy; nitrilotriacetic acid. 


Because immobilized metal ion affinity chromatography (IMAC) was first 

developed for metal-specific protein isolation by Porath et al. (1,2), a large 

number of IMAC protocols have been developed. All IMAC protocols share 

the same principal that proteins with exposed histidine and cysteine side chains 

have a distinct affinity for certain metals, like Ni2+ ,Co2+ ,Zn2+ ,Cu2+ ,Fe3+ , 

or Mn2+ (3,4). Depending on the chosen metal-chelating group and metal ion 

more or fewer coordination sites are accessible for potential protein binding. 



139

140 Thierse et al. 

In general, binding affinity of proteins to metal ions is dependent on the 

amino acid composition of a given protein, with histidine showing the highest 

metal affinity followed by tryptophan, phenylalanine, tyrosine, and cysteine 

(5). However, the affinity between a metal ion and an amino acid is also highly 

dependent on the specific metal ion itself, with Fe 3+ , for example, having the 

highest affinity to carboxyl- and phosphate-groups. Besides the amino acid 

sequence and the distribution of a certain amino acid, surface characteristics 

and protein folding (3-dimensional structure) are additional important parameters 

for binding affinity. In an optimal column or bead-based performance 

the selected metal is strongly held by the matrix bound metal-chelating group, 

e.g., nitrilotriacetic acid (NTA) or iminodiacetic acid (IDA), still leaving metal 

coordination sites available to interact with the metal-specific protein ligand. 

Tetradentate NTA, for example, was found to bind Ni 2+ with three carboxyl 

groups and one nitrogen (6), leaving two other ligand positions accessible to 

Ni 2+ -specific proteins or recombinant 6His-tagged proteins (7). Interestingly, 

when compared to the dissociation constant (K d) of most antibody bindings (K d 

from10 −5 M to 10 −12 M), 6His-tagged protein to Ni 2+ K d has been shown to be 

relatively high, 10 −13 M at pH 8 (8,9). Moreover, Ni-NTA itself has been shown 

to be stable over a pH range of 2.5–13 and withstands extreme conditions like 

2% SDS and 100% ethanol. 

Today, IMAC is not only commonly used for purification of the mentioned 

recombinant 6His-tagged proteins or the evaluation of protein folding and 

endotoxin removal from protein preparations, but also advantageous in the 

enrichment of phosphopeptides (10). Reversible protein phosphorylation (e.g., 

at Ser, Thr, and Tyr residues) is one of the most important post-translational 

modifications, controlling signal transduction pathways that direct cellular 

activation, differentiation, and proliferation, as well as apoptosis (11). Because 

phosphopeptides are acidic and show strong binding characteristics to some 

metal ions (Fe 3+ ,Ga 3+ , and Al 3+ ) different IMAC protocols have been introduced 

for isolation of phosphopeptides (10,12). Among the most recently 

described phosphopeptide IMAC methods are titanium dioxide (TiO 2) phosphopeptide 

enrichment (13,14), and metal oxide affinity chromatography (MOAC) 

where the affinity of the phosphate group for Al(OH) 3 is exploited (15). For 

detailed information on IMAC phosphopeptide enrichment see Corthals et al., 

2005 and Ueda et al., 2003 (5,10). 

Affecting up to 15% of the women in industrialized countries, human 

nickel (Ni) allergy represents one of the major metal-related diseases in the 

human population. However, according to several studies transition metal Ni 

is two-faced. Nickel is considered as a beneficial physiological agent used for 

essential functions as an ultra trace element. Conversely, Nickel can act as a 

pathological agent by interacting directly with DNA or DNA-binding proteins

Metalloproteomics in the Molecular Study of Cell Physiology 141 

or metal-specific T cells, thus causing cellular toxicity or a metal-specific 

allergic contact dermatitis (ACD). To elucidate such physiological and diseaserelated 

molecular processes, a new qualitative subproteomic method, combining 

IMAC and 2-dimensional gel electrophoresis (2-DE), has been developed to 

enrich and identify Ni 2+ -interacting proteins from different human cell types 

(e.g., human antigen presenting cells, keratinocytes). After affinity binding of 

metal-interacting proteins to Ni-NTA beads, followed by stringent washing 

steps and elution with imidazole, proteins are applied to modern 2-DE using 

immobilized pH-gradients (16–19). In a typical proteomic workflow, proteins 

are identified by mass spectrometry to define the metalloproteome. Complementary 

methods such as the use of recombinant proteins and metal detection 

by atomic absorption spectrometry allow confirmation of direct metal-protein 

interactions (20). Thus, with a clear differential pattern to Cu-binding proteins 

in human liver cells, Ni-NTA affinity enrichment of proteins from human 

B-cell lysates resulted in the subproteomic identification of so far unknown 

Ni-interacting proteins (Fig. 1) (21–23). Quite unexpectedly, 16 of these Niinteracting 

proteins were found to belong to the group of stress-inducible 

heatshock proteins or chaperonins, including HSP-60, HSP-70, BIP, and the 

high-molecular heterooligomeric complex of TriC/CCT (21). Thus Ni 2+ ,in 

addition to the induced formation of T cell epitopes recognizable by the 

A B 

Protein solution 

cell lysate 

0.1% Triton X-100 

Ni-NTA-beads 

1h incubation, 4°C 

Washing steps 

500 mM NaCl (high salt) 

137 mM NaCl (low salt) 

Elution of Ni-interacting 

proteins 

250 mM imidazole 

2-DE, staining, spot 

picking, trypsinization 

MW 

pH 4 pH 7 

Fig. 1. Ni-affinity enriched proteins were isolated from human antigen presenting 

cells (2* 10 7 cells) as demonstrated in the subproteomic workflow (A) and analyzed 

by 2-DE (B, silver staining), and subsequently identified by mass spectrometry (for 

details of mass spectrometric protein identification see (21)).


acquired immune system, intimately interacts with essential constituents of the 

innate defense system, thereby linking both arms of the immune system. In 

summary, the development and combined usage of new metal-specific and 

proteomic techniques gives new insights into metal-related metabolic pathways 

and frequent metal-specific disorders, like human nickel ACD, leading to 

improved strategies in diagnosis and therapy of such environment-induced 

disease (21,24,25). 

2. Materials 

2.1. Cell Culture and Lysis 

1. RPMI 1640 medium for human antigen presenting cells containing 10% fetal calf 

serum, 2 mM l-glutamine, 1 mM sodium pyruvate, nonessential amino acids and 

25 mM HEPES buffer (all from Gibco BRL Life Technologies, Paisley, UK). 

For primary keratinocytes Keratinocyte Basal Medium 2 was supplemented with 

SupplementPack/Keratinocyte Growth Medium 2 (KGM) (Promocell, Heidelberg, 

Germany). 

2. A solution of trypsin (0.1%) and ethylenediamine tetraacetic acid (EDTA) (0.02%) 

(Biochrom, Berlin, Germany). 

3. Phosphate buffered saline (D-PBS) (Invitrogen, Karlsruhe, Germany). 

4. Fetal calf serum (FCS) (Biochrom, Berlin, Germany). 

5. Lysis buffer (0.1% Triton): 137 mM NaCl, 20 mM Tris-HCl, 10% (v/v) glycerol, 

0.1% (v/v) Triton X-100, pH 8.2. Add protease inhibitor cocktail tablets, 

“complete mini, EDTA free” (Roche, Mannheim, Germany) before use. Store in 

aliquots at –20°C (see Note 1). 

2.2. Isolation of Nickel-Binding Proteins 

1. Nickel-nitrilotriacetic acid (Ni-NTA) Magnetic Agarose Beads 5% suspension, 

with binding capacity: 300 μg/mL (Qiagen, Hilden, Germany). 

2. High salt lysis buffer: 500 mM NaCl, 20 mM Tris-HCl, pH 8.2, 10% (v/v) 

Glycerol, 0.1% Triton X-100. 

3. Low salt lysis buffer: 137 mM NaCl, 20 mM Tris-HCl, pH 8.2, 10% (v/v) 

Glycerol, 0.1% Triton X-100. 

4. Imidazole solution: 250 mM imidazole (Sigma, Taufkirchen, Germany) in distilled 

water. 

5. Magnetic device for 1.5 mL Eppendorf tubes (Qiagen, Hilden, Germany). 

6. Standard 2-D electrophoresis equipment. 

2.3. Silver Staining of Metal-Affinity Enriched Proteins Compatible 

for Mass Spectrometry (MS) 

Several protocols of 2-dimensional gel electrophoresis (2-DE) are useful 

for detection of IMAC separated proteins, without negatively affecting the


principal method described here (16,17,21). Nevertheless, after separating 

affinity-enriched proteins by 2-DE a protein staining method compatible for 

mass spectrometric analysis has to be applied. Following protein gel scanning, 

e.g., with the LabScan Image Scanner (GE Healthcare, München, Germany) or 

the new laser scanner FLA 5100 (FUJIFILM Life Science, Düsseldorf), spot 

picking can be performed by hand or automatically using the PROTEINEER 

spII system (Bruker Daltonics, Bremen, Germany). Subsequent MALDI mass 

spectra can be recorded e.g., with a Ultraflex MALDI-TOF spectrometer 

(Bruker Daltronics, Bremen, Germany) equipped with a 337 nm nitrogen laser 

(for details see (21)). 

1. Thiosulfate solution: 0.02% (w/v) sodium thiosulfate pentahydrate (Merck, 

Darmstadt, Germany). 

2. Silver staining solution: 0.2% (w/v) silver nitrate (Merck, Darmstadt, Germany), 

0.02% formaldehyde solution (34%) (J.T. Baker, Deventer, NL). 

3. Developer: 3% (w/v) sodium carbonate (J.T. Baker, Deventer, NL), 0.05% 

formaldehyde solution (37%) (J.T. Baker, Deventer, NL), 0.0005% thiosulfate 

solution. 

4. Stopping solution (suitable for mass spectrometry) (see Note 2): 50% methanol 

(Merck, Darmstadt, Germany), 12% acetic acid (glacial) (Merck, Darmstadt, 

Germany). 

5. Stopping solution (not suitable for mass spectrometry): 0.5% glycine (Roth, 

Karlsruhe, Germany). 

6. Ethanol. 

3. Methods 

3.1. Preparing Samples for Isolation of Nickel-Binding Proteins 

1. Culture of immune cells in RPMI medium and/or primary keratinocytes in KGM 

in 10 mm tissue culture dish until passage 3 or earlier. 

2. When e.g., keratinocytes are confluent, wash with PBS and incubate with 3 mL 

of trypsin/EDTA for 5 min at 37°C. Rinse cells with a pasteur pipet and transfer 

into a tube supplied with 6 mL 10% FCS in PBS. Rinse the dish with additional 

3 mL of 10% FCS and transfer it into the tube. 

3. Wash cells 3× with cold PBS and count the cells. 

4. Resuspend cells in lysis buffer (2* 107 /mL), incubate at 4°C for 1 h with gentle 

shaking. 

5. Clarify the lysate by centrifugation (20,000g, 10 min, 4°C). (see Note 3) 

6. The supernatant can be stored in aliquots at –20°C or –80°C. 

3.2. Isolation of Nickel-Binding Proteins 

1. Resuspend the Ni-NTA Magnetic Agarose Beads by vortexing or pipeting. 

2. Incubate 150 μL Ni-NTA Beads with 1 mL lysate (recommended ratio) by rotation 

for2hat4°Cina1.5-mL Eppendorf tube (see Note 4).


3. Insert the tube into a magnetic device for 1.5-mL Eppendorf tubes and remove 

supernatant (see Note 5). 

4. Wash pellet 2× with high salt lysis buffer, once with low salt lysis buffer. The 

pellet has to be resuspended thoroughly during each wash. Discard supernatants 

(see Note 6). 

5. For elution add the imidazole solution to the pellet and incubate for 10 min at 

room temperature. Use a volume equal to the original volume of Ni-NTA beads. 

6. Insert the tube into a magnetic device for 1.5-mL Eppendorf tubes and carefully 

transfer the eluate into a fresh tube. Store aliquots at –80°C. 

7. For first-dimension isoelectric focussing sample (e.g., 25 μg) can be applied by 

including it in the rehydration solution of the IPG strip, followed by standard 

protocols of 2-D electrophoresis. 

3.3. Silver Staining of Metal-Affinity Enriched Proteins Compatible 

for Mass Spectrometry (MS) 

After 2-DE of metal-affinity enriched proteins spots have to be stained with 

staining protocols adapted to mass spectrometric analysis (for details see (21)). 

Silver staining has to be performed in glass dishes and see also Notes 7–11. 

1. Fix the gel in 40% ethanol/10% (v/v) acetic acid for at least 1 h. 

2. Wash 3× in 30% (v/v) ethanol for 20 min. 

3. Reduce the gel in thiosulfate solution for 1 min. 

4. Wash 3× in distilled water for 20 sec. 

5. Stain the gel in staining solution for 20 min. 

6. Wash 3× in distilled water for 20 sec. 

7. Develop in developer for 3–5 min. 

8. Wash 3x in distilled water for 30 sec. 

9. Stop the reaction in stopping solution for 5 min. 

10. Wash 2× in distilled water for 30 sec and an additional time for 15 min. 

11. Gel can be stored in 1% (v/v) acetic acid or shrink-wrap. An example result is 

demonstrated in Fig. 1. 

Alternatively, gels (e.g., loaded with 100 μg protein concentration) may 

be stained by MS-compatible Coomassie staining, e.g., according to Jungblut 

et al. (26). 

4. Notes 

1. Freshly prepared lysis buffer can be stored at 4°C for several months under 

sterile conditions. 

2. The color of the gel deepens after a while. 

3. At this point a determination of the protein concentration is often useful. 

4. Incubation of the cell-lysate with Ni-NTA Magnatic Agarose Beads can be 

extended to overnight incubation.


5. The supernatant contains all proteins that do not bind to Nickel and maybe 

additional Nickel-binding Proteins. Therefore, store the supernatant at –20°C or 

–80°C just in case. 

6. Supernatants of the washing steps can be stored at –20°C or –80°C just in case. 

7. All steps should be performed at room temperature with gentle shaking. 

8. Because the silver stain is very sensitive, nitrile gloves are recommended. Latex 

gloves may leave fingerprints on the gel. 

9. The amount of fluid per gel depends on the size of the gel and the staining 

dishes. Approximately 250 mL for 20 × 20 cm gels is sufficient. 

10. The gel becomes less flexible through the staining procedure and tears very 

easily. To transport the gel safely onto the scanning device, very careful handling 

is required. 

11. All solutions, containing methanol or silver nitrate have to be disposed of 

according to the directions given by the local authorities. 


We thank Doris Wild and Stefanie Eikelmeier for excellent technical assistance, 

and Dr. Ian Haidl, Depts. of Pediatrics, Microbiology and Immunology, 

Halifax, Canada, for very careful reading of the manuscript. This work 

was supported in part by the Landesstiftung Baden-Wüerttemberg, Germany, 

Forschungsprogramm “Allergologie” by grant P-LS-AL/26 (to HJT), and the 

European Union, as part of the project Novel Testing Strategies for In Vitro 

Assessment of Allergens (Sens-it-iv), LSHB-CT-2005 – 018681, (www.sensit-iv.eu). 

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3. Mondal, K. and Gupta, M. N. (2006) The affinity concept in bioseparation: evolving 

paradigms and expanding range of applications. Biomol. Eng. 23, 59–76. 

4. Sun, X., Chiu, J. F., and He, Q. Y. (2005) Application of immobilized metal 

affinity chromatography in proteomics. Expert Rev. Proteomics. 2, 649–57. 

5. Ueda, E. K., Gout, P. W., and Morganti, L. (2003) Current and prospective applications 

of metal ion-protein binding. J. Chromatogr. A. 988, 1–23. 

6. Hochuli, E., Dobeli, H., and Schacher, A. (1987) New metal chelate adsorbent 

selective for proteins and peptides containing neighbouring histidine residues. J. 

Chromatogr. 411, 177–84. 

7. Terpe, K. (2003) Overview of tag protein fusions: from molecular and biochemical 

fundamentals to commercial systems. Appl. Microbiol. Biotechnol. 60, 523–33.


8. Harlow, E., and Lane, D. (1988) Antibodies: A Laboratory Manual, Cold Spring 

Harbor Laboratory, Cold Spring Harbor, NY. 

9. Schmitt, J., Hess, H., and Stunnenberg, H. G. (1993) Affinity purification of 

histidine-tagged proteins. Mol. Biol. Rep. 18, 223–30. 

10. Corthals, G. L., Aebersold, R., and Goodlett, D. R. (2005) Identification of 

phosphorylation sites using microimmobilized metal affinity chromatography. 

Methods Enzymol. 405, 66–81. 

11. Bollen, M. and Beullens, M. (2002) Signaling by protein phosphatases in the 

nucleus. Trends Cell Biol. 12, 138–45. 

12. Zhou, W., Merrick, B. A., Khaledi, M. G., and Tomer, K. B. (2000) Detection and 

sequencing of phosphopeptides affinity bound to immobilized metal ion beads by 

matrix-assisted laser desorption/ionization mass spectrometry. J. Am. Soc. Mass 

Spectrom. 11, 273–82. 

13. Hata, K., Morisaka, H., Hara, K., et al. (2006) Two-dimensional HPLC on-line 

analysis of phosphopeptides using titania and monolithic columns. Anal. Biochem. 

350, 292–7. 

14. Larsen, M. R., Thingholm, T. E., Jensen, O. N., Roepstorff, P., and Jorgensen, 

T. J. (2005) Highly selective enrichment of phosphorylated peptides from peptide 

mixtures using titanium dioxide microcolumns. Mol. Cell Proteomics. 4, 873–86. 

15. Wolschin, F., Wienkoop, S., and Weckwerth, W. (2005) Enrichment of phosphorylated 

proteins and peptides from complex mixtures using metal oxide/hydroxide 

affinity chromatography (MOAC). Proteomics. 5, 4389–97. 

16. Gorg, A., Obermaier, C., Boguth, G., et al. (2000) The current state of twodimensional 

electrophoresis with immobilized pH gradients. Electrophoresis. 21, 

1037–53. 

17. Gorg, A., Weiss, W., and Dunn, M. J. (2004) Current two-dimensional 

electrophoresis technology for proteomics. Proteomics. 4, 3665–85. 

18. Klose, J. (1975) Protein mapping by combined isoelectric focusing and 

electrophoresis of mouse tissues. A novel approach to testing for induced point 

mutations in mammals. Humangenetik. 26, 231–43. 

19. O’Farrell, P. H. (1975) High resolution two-dimensional electrophoresis of 

proteins. J. Biol. Chem. 250, 4007–21. 

20. Thierse, H. J., Moulon, C., Allespach, Y., et al. (2004) Metal-protein complexmediated 

transport and delivery of Ni2+ to TCR/MHC contact sites in nickelspecific 

human T cell activation. J. Immunol. 172, 1926–34. 

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proteins in human B cells. Proteomics. 5, 3614–22. 

22. She, Y. M., Narindrasorasak, S., Yang, S., Spitale, N., Roberts, E. A., and 

Sarkar, B. (2003) Identification of metal-binding proteins in human hepatoma lines 

by immobilized metal affinity chromatography and mass spectrometry. Mol. Cell. 

Proteomics. 2, 1306–18. 

23. Smith, S. D., She, Y. M., Roberts, E. A., and Sarkar, B. (2004) Using immobilized 

metal affinity chromatography, two-dimensional electrophoresis and mass 

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proteins by immobilized metal affinity chromatography and two-dimensional 

electrophoresis. Electrophoresis. 14, 638–43.

13 

Protein Extraction from Green Plant Tissue 

Ragnar Flengsrud 

Summary 

A method for preparation of protein from green plant tissue for two-dimensional 

electrophoresis is described. The method is demonstrated on barley leaves, potato leaves 

and spruce needles and appears to overcome the obstacles inherent in green plants to 

proteomic analysis. The yield and the representation of proteins are discussed. 

Key Words: Barley leaves; green plant tissue; potato leaves; protein extraction; 

spruce needles; two-dimensional electrophoresis. 


Preparation of proteins for two-dimensional (2-D) electrophoresis is an 

important, and sometimes crucial, part of this central proteomics technique, a 

fact that may be overlooked or underestimated. 

Plants and especially green plant tissues constitute considerable challenges 

here, because of low protein concentration and the presence of deleterious 

compounds in the cell. 

Plant proteases may contribute to the challenge because remarkable stability 

has been reported relevant to temperature (1–3), pH(2,3) and even urea or 

guanidine hydrochloride (2,4). 

During the work with green plant tissues several principles were applied 

that seem to overcome these problems, resulting in good, reproducible 2-D 

separation of green tissue proteins from three different species (5). These 

principles are: (a) Cell disruption and homogenization in liquid N2 and insoluble 

polyvinylpyrrolidone; (b) an extraction buffer including thiourea and SDS with 

a pH lower than 5.5; (c) protein precipitation in 90% (v/v) acetone at –20°C; 



149

150 Flengsrud 

(d) dialysis against a buffer containing 9.5M urea, nonionic detergent and 

lysine. The sample preparation method described in this chapter is well suited 

for barley leaves (Fig. 1), potato leaves and spruce needles. Electropherograms, 

both in the isoelectric focusing (Fig. 1) and the nonequilibrium pH gradient 

mode, is presented. There is no indication of protease activity. Total recovery 

of protein is 6.7–16.5% (5). Yet, the possibility exists that all or most proteins 

are represented in the extract. 

A study on the effect of different extraction solutions on the solubilization 

of endosperm proteins showed that the same proteins were extracted, but to 

different degree. SDS/urea was one of the extraction solutions with best overall 

results in this work. Even with 22% protein recovery, the extract was shown 

to contain hordein proteins (6). The sample preparation method was utilized in 

a stress diagnosis study on spruce needles (7,8). This study used both soluble 

and immobilized pH gradients and about 1,500 spots were detected by image 

analysis. Here, the changes in needle protein pattern were studied by image 

analysis of 300–350 spots. Protease activity was found to be negligible in this 

study. 

Fig. 1. Two-dimensional electrophoresis in the isoelectric focusing mode of proteins 

extracted from leaves of a mutant line (H354-33-7-5). of the barley variety cv. Carlsberg 

II. The loading was 33 μg protein and silver staining was used for detection.

Protein Extraction from Green Plant Tissue 151 

2. Materials 

1. Extraction buffer: 50 mM pyridine, 10 mM thiourea, 1% (w/v) SDS, adjusted to 

pH to 5.0 by HCl. The purity of pyridine (see Note 1), the pH-range (see Note 2) 

and the use of SDS (see Note 3) is important. 

2. Lysis solution: 9.5M urea, 2% (v/v) nonionic detergent (Igepal CA-630), 1.6% 

(v/v) ampholytes pH 5–7, 0.4% ampholytes pH 3–10, 2.5% (w/v) dithiothreitol 

(DTT). 

3. Modified lysis solution for dialysis: 2% (w/v) lysine substitutes the ampholytes. 

3. Methods 

1. Cut the leaves into 0.5 to 1.0 cm parts and needles into 2–4 mm parts. Typically, 

start with 0.4 g green tissue and grind it twice in a mortar with liquid N2 to give 

a fine powder. The storage of plant material before homogenization should be 

considered (see Note 4). 

2. Add an amount of insoluble polyvinylpyrrolidone (Polyclar AT) twice the weight 

of plant material (see Note 5) and mix. 

3. Add extraction buffer (9.5 mL) to the mixture in the mortar, stir for a few minutes 

and centrifuge at +5°C for 40min. at 8,000g. 

4. Transfer the supernatant to a thick-walled glass centrifuge-tube, add ice cold 

acetone (see Note 6) to give a final concentration of 90% (v/v) and mix well. 

Allow proteins to precipitate at –20°C for 2 h. The yield of the protein depends 

on the concentration of acetone (see Note 7). 

5. Collect proteins by centrifugation for 20 min at +5°C and 5,000g. Discard the 

supernatant and wash the precipitate once with ice cold acetone and centrifuge as 

above. 

6. Carefully dry the resulting precipitate in a stream of N2, add 400 μL of the lysis 

solution and mix well. 

7. Dialyse the mixture of precipitate and lysis solution overnight against 25 mL of 

the modified lysis solution. 

8. Centrifuge the dialysed sample at 8,000g for 10 min. Add the appropriate 

ampholytes to the clear supernatant to give a final 2% (v/v) concentration and 

store at –20°C in suitable aliquots. The suitability depend on the detection method 

to be used following the 2-D electrophoresis (see Note 8). 

4. Notes 

1. The pyridine in the extraction buffer should be distilled over ninhydrin and 

stored under nitrogen. Alternatively, at least HPLC-grade is used and stored under 

nitrogen. 

2. The study of thiourea as phenoloxidase inhibitor concluded that the pH in the 

extraction buffer should not be above 5.5 (9). 

3. Extraction with and without SDS in the extraction buffer showed that SDS was 

necessary for the solubilisation of membrane-bound proteins (5).

152 Flengsrud 

4. Storage of barley leaves at –20°C or –80°C up to 3 mo and at +5°C for 1 wk, 

before homogenization does not seem to affect the protein pattern. 

5. The original study (5) used twice the amount (g/g) of insoluble polyvinylpyrrolidone 

(Polyclar AT) to plant tissue for its homogenization. A later work (7) 

used routinely equal amounts and observed that less polyvinylpyrrolidone resulted 

in poor electropherograms. 

6. Acetone is kept at –20°C before its use in protein precipitation. 

7. A final concentration of 90% acetone at –20°C for 1 h will totally precipitate the 

proteins (10). 

8. The following aliquots of the extract are suitable for 2-D electrophoresis: for 

silver staining, 40–60 μL, corresponding to 30–46 μg protein; for Coomassie Blue 

staining, 150–260 μL. The aliquots should not be refrozen. 

References 

1. Fahmy, A. S., Ali, A. A., and Mohammed, S. A. (2004) Characterization of a 

cysteine protease from wheat Triticum aestivum (cv. Giza 164). Bioresour. Technol. 

91, 297–304. 

2. Kaneda, M., Yonezawa, H., and Uchikoba, T. (1995) Improved isolation, stability 

and substrate specificity of cucumisin, a plant serine endopeptidase. Biotechnol. 

Appl. Biochem. 22, 215–22. 

3. Patel, B. K. and Jagannadham, M. V. (2003) A high cysteine containing thiol 

proteinase from the latex of Ervatamia heyneana: purification and comparison with 

ervatamin B and C from Ervatamia coronaria. J. Agric. Food Chem. 51, 6326–34. 

4. Uchikoba, T., Niidome, T., Sata, I., and Kaneda, M. (1993) Protease D from the 

sarcocarp of honeydew melon fruit. Phytochemistry 33, 1005–08. 

5. Flengsrud, R. and Kobro, G. (1989) A method for two-dimensional electrophoresis 

of proteins from green plant tissues. Anal. Biochem. 177, 33–6. 

6. Flengsrud, R. (1993) Separation of acidic endosperm proteins by two-dimensional 


7. Davidsen, N. B. (1995) Two-dimensional electrophoresis of acidic proteins isolated 

from ozone-stressed Norway spruce needles (Picea abies L. Karst): Separation 

method and image prosessing. Electrophoresis 16, 1305–11. 

8. Davidsen, N. B. (1996) Improved two-dimensional electrophoretic separation of 

acidic proteins extracted from Norway spruce needles by using immobilized pH 


9. Van Driessche, E., Beeckmans, S., Dejaegere. R., and Kanarek, L. (1984) Thiourea: 

the antioxidant of choice for the purification of proteins from phenol-rich plant 

tissues. Anal. Biochem. 141, 184–88. 

10. Neuhoff, V. (1973) Micromethods in Molecular Biology. Springer-Verlag 

(Kleinzeller, A., Springer, G.F., and Wittmann, H.G., eds.) 14, p133.

14 

The Terminator: A Device for High-Throughput 

Extraction of Plant Material 

B. M. van den Berg 

Summary 

The Terminator is a device for cost-efficient high-throughput homogenization of plant 

material and sample preparation. Protein and DNA samples can easily be prepared from 

large numbers of crude material for further analysis such as protein electrophoresis or 

polymerase chain reaction (PCR) followed by DNA electrophoresis. The functioning of the 

device is based on vibration of 96 stainless steel pegs in wells of a standard 96-well micro 

plate. Using the Terminator all types of plant tissue, including seeds, can be homogenized 

in standard micro plates in 3 min. 

Key Words: DNA extraction; electrophoresis; high throughput analysis; plant 

material; protein extraction; seeds. 


High-throughput homogenization and sample preparation is often a timelimiting 

step in large-scale analysis of biological material. This certainly holds 

for the analysis of seed or other plant tissue in several agricultural businesses— 

for example the seed business (1)—where many hundreds or even thousands 

of individual samples are daily analyzed on a routine basis. 

In the seed business, gel isoelectric focusing of protein (but also other 

electrophoretic techniques) is widely used to determine the genetic purity of 

seed lots or the percentage of inbreds that may occur in hybrid seed lots. With 

the advance of isoelectric focusing in the 1980s, the need for high-throughput 

sample preparation techniques and equipment became important (1). During the 



153

154 van den Berg 

years 1988 and 1989 a device was developed—later named the Terminator— 

that is perfectly suited to address the needs of high-throughput sample preparation 

for isoelectric focusing of protein (2). In more recent years, the Terminator 

has proven to be an efficient tool also in high-throughput extraction of 

DNA from seed and other plant tissue (1,3). In 2004, the Terminator was 

improved. The device was restyled, another power supply was added, and the 

design of the stainless steel pegs was improved to increase the efficiency of 

homogenization. Here, the functioning of the Terminator is presented in detail. 

2. Materials 

All chemicals were purchased from Sigma (St. Louis, OH, USA), unless 

otherwise stated. 

2.1. Sample Preparation 

1. The Terminator used is the restyled and improved Terminator as produced and 

sold by Elexa (Enkhuizen, the Netherlands). This device consists of three parts: the 

Terminator base plate (Fig. 1A), the Terminator head consisting of a vibromotor 

and an aluminum plate with 96 stainless steel pegs (Fig. 1B), and a variable power 

supply (Fig. 1C). The base plate is a4cmthick stainless steel plate equipped 

with a micro plate holder consisting of an aluminum base, plastic holders, and 

micro plate clips. Standard 96-wells micro plates can be placed on the micro 

plate holder and fixed with the special clips. The heavy weight of the steel plate 

and the rubber feet assure that the Terminator remains at a fixed position during 

operation. 

Fig. 1. Image showing the three parts of the Terminator. A, the Terminator base 

plate; B, the Terminator head consisting of the vibrating motor and the 96-peg plate; 

C, the variable AC power supply (0–220 V).

A Device for High-Throughput Extraction of Plant Material 155 

The Terminator head consists of a special design vibromotor that operates at a 

vibration frequency of 50 Hz, and an aluminum plate attached to it which bears 

96 stainless steel pegs that fit perfectly in the wells of a standard micro plate. 

The variable power supply is a standard AC 220 V transformer that can deliver 

0–220 V (see Notes 1–3). 

2. 96-well flat-bottom micro plates (see Note 4) from Costar (Cambridge, USA). 

3. Seeds from Seminis Vegetable Seeds (Oxnard, CA, USA) and Syngenta (Nerac, 

France). 

4. Sunflower and corn seed protein extraction buffer for isoelectric focusing (IEF): 

10 mM Tris-HCl, pH 7.0, 2% (v/v) ampholytes of same pH range as that of the 

gel. Make fresh just before use. 

5. Tomato seed ADH (alcohol dehydrogenase) extraction buffer: 2% ampholytes 

(v/v) pH range 3–10, 0.25% (w/v) dithiothreitol. Make fresh just before use. 

6. Ampholytes: SinuLytes 3-7 and 3-10 from Sinus (Heidelberg, Germany). Store 

at 4°C (see Note 5). 

7. DNA seed extraction buffer (XB): 0.2M Tris-HCl pH 8.0, 0.5% (w/v) sodium 

dodecyl sulphate (SDS), 0.3M NaCl, 25 mM ethylenediamine tetraacetic acid 

(EDTA). Store at room temperature. 

8. Protein precipitation (PP) buffer: 2.5M potassium acetate pH 6.5. Add 245.0 g 

potassium acetate to 800 mL water. Stir until fully dissolved. Adjust the pH to 

6.5 with acetic acid. Bring the final volume to 1 L. Store at room temperature. 

9. Tris-EDTA (TE) buffer: dissolve 1.21 g Tris and 37.2 mg EDTA in 1 L water. 

Store at 4°C. 

2.2. Isoelectric Focusing (IEF) 

These instructions assume knowledge of making thin horizontal isoelectric 

focusing gels between glass plates and the use of equipment for horizontal 

isoelectric focusing (4–7). 

1. 16% (w/v) glycerol. Store at 4°C. 

2. 30% acrylamide/bis-acrylamide solution (29:1). Store at 4°C. 

3. SinuLytes 3-7 and 3-10 (Sinus, Heidelberg, Germany). Store at 4°C. 

4. 10% (w/v) ammonium persulfate. Store at 4°C, but no longer than 1 wk. 

5. Electrode paper: Whatman #17 

6. Electrode solution: 2% (v/v) ampholytes with pH-range identical to that of 

the gel. 

7. 96-well sample applicator strips from Elexa (Enkhuizen, The Netherlands). 

8. Gel backing: GelGrip from Sinus (Heidelberg, Germany). 

9. Coomassie gel staining solution: 0.2% (w/v) coomassie brilliant blue (CBB), 

50% (v/v) water, 40% (v/v) ethanol, 10% (v/v) acetic acid. Add 0.1 g coomassie 

brilliant blue to 20 mL ethanol. Stir until fully dissolved. Then add 25 mL water 

and 5 mL acetic acid. Prepare fresh before use. 

10. CBB destaining solution: 50% (v/v) water, 40% (v/v) ethanol, 10% (v/v) acetic 

acid. Store at room temperature.


11. ADH staining solution: 0.1M Tris-HCl, pH 7.5, 5% (v/v) ethanol, 0.2% 

-nicotinamide adenine dinucleotide (NAD), 0.2% (w/v) 1-(4,5-dimethylthiazol- 

2-yl)-3,5-diphenylformazan (MTT), 0.05% (w/v) phenazine methosulfate (PMS). 

Prepare fresh. 

12. 20% (w/v) trichloroacetic acid (TCA). Prepare fresh. 

2.3. Inter Simple Sequence Repeat PCR (ISSR-PCR) 

ISSR-PCR is a technique based on amplification of DNA between two simple 

sequence repeat (SSR; head-to-tail tandem arrays of short DNA repeat motifs) 

regions, that uses 5 ′ or 3 ′ anchored SSR PCR primers (8). 

1. Anchored primer: 5’-DVDTCTCTCTCTCTCTC (D = A,G,T ;V = A,C,G) from 

Invitrogen (Carlsbad, CA, USA). 

2. PCR mix: 9.06 μL water, 1.50 μL PCR buffer (10x), 0.60 μL 50 mM magnesium 

chloride, 0.6 μL dNTP mixture (2.5 mM each), 0.12 μL 25 μM primer, 0.12 

μL DNA polymerase, 3 μL pepper DNA solution. The PCR buffer (10×) and 

the magnesium chloride (MgCl 2) come at the appropriate concentration with the 

DNA polymerase. The dNTP’s and primer must be diluted to the appropriate 

concentration with water. 

3. DNA polymerase from Invitrogen (Carlsbad, CA, USA). 

4. PCR reaction plates from Invitrogen (Carlsbad, CA, USA). 

2.4. DNA Electrophoresis 

These instructions assume knowledge of making thin horizontal gels between 

glass plates and the use of equipment for horizontal electrophoresis (3). 

1. Gel buffer (2×): 120 mM Tris-formic acid, pH 9.0. 

2. 10% (w/v) ammonium persulfate. Store at 4ºC, but no longer than 1 wk. 

3. 20% (w/v) glycerol. Store at 4ºC. 

4. 30% acrylamide/bis-acrylmide solution (29:1). Store at 4ºC. 

5. Gel backing: GelGrip from Sinus (Heidelberg, Germany). 

6. Sample buffer for DNA electrophoresis (SB): 2.5% gel buffer (2×), 0.02% 

Bromophenol Blue. 

7. In-gel well template and spacers (0.2 mm thick) from Elexa (Enkhuizen, The 

Netherlands). 

8. Electrode paper: Whatman #17. 

9. Electrode solution: 10% (w/v) Tris-Base, 1.73% (w/v) boric Acid, 0.02% 

bromophenol blue. 

10. Fixing solution: 2% (v/v) nitric acid. 

11. Staining solution: 0.2% (w/v) silver nitrate. 

12. Developer solution: 0.05% (v/v) formaldehyde, 3% (w/v) sodium carbonate. 

Prepare fresh just before use. 

13. Stop solution: 5% (v/v) acetic acid.


3. Methods 

3.1. Sample Preparation 

The Terminator homogenizes tissue in the wells of the micro plate by the 

vibration in all directions of the pegs in the wells. Fig. 2A illustrates a diagrammatic 

view of how plant tissue is squeezed and homogenized between the 

vibrating pegs and the wall and bottom of the micro plate wells. Optimal 

voltage for operation of the Terminator lies between 90 and 130 V depending 

on the type of tissue to be homogenized. For locations with 110 net Voltage, a 

pretransformer that gives 220V output can best be used. 

1. Small parts of the corn and sunflower seeds are punched using a home-made 

device to get samples that fit in the wells of a 96-well plate (see Note 7). 

Alternatively, parts can be cut using a scalpel. To cut the seed parts easier, the 

seeds may be soaked in water overnight. The seed parts are put in the wells of 

a 96-well micro plate, 200 μL seed extraction buffer is added, and the tissue is 

disrupted and fully homogenized using the Terminator. After 3 min operation of 

the Terminator the homogenates are centrifuged in a micro plate centrifuge and 

the supernatant samples are used for IEF. 

2. Tomato seeds are put in the wells of a 96-well micro plate, 200 μL extraction buffer 

is added, and the seeds are disrupted and fully homogenized using the Terminator. 

Fig. 2. Composite image showing a close view of the pegs of the Terminator in 

micro plate wells. A drawing of a peg in a well of a micro plate illustrating how tissue 

is squeezed between the pegs and the walls and bottom of the plate. B and C show a 

photograph at close range of the pegs in the micro plate.


After 3 min operation of the Terminator the homogenates are centrifuged in a 

micro plate centrifuge and the supernatant samples are used for IEF. 

3. Pepper seeds are put in the wells of a 96-well micro plate, 200 μL DNA 

extraction buffer is added, and the seeds are disrupted and fully homogenized 

using the Terminator. After 3 min operation of the Terminator the homogenates 

are centrifuged in a micro plate centrifuge and 90 μL of each of the individual 

supernatants are brought (taking care not to disturb the pellets) in another micro 

plate. 90 μL PP buffer is added to the supernatants and the micro plate is mixed 

using an orbital shaker for 3 min at moderate speed. Then the plate is centrifuged 

at 4°C, 1,300g, for 10 min with moderate deceleration. 

Of the resulting supernatants 90 μL is brought to a new micro plate and 90 

μL ice-cold isopropanol (–20°C) is added to the supernatants. The micro plate 

is briefly shaken using the orbital shaker at medium speed and then centrifuged 

at 4°C, 1,300g for 10 min with moderate deceleration. The resulting supernatant 

is discarded and the pellet dried at 65°C for 20 min. To the pellet 100 μL 

TE-buffer is added, and the resulting DNA solution is stored at 4ºC. For long 

term storage –20°C is recommended. 

3.2. Isoelectric Focusing (IEF) 

1. For IEF of corn and sunflower seed extracts, gels of size 260 × 188 mm, and 

thickness of 0.2 mm are made fresh by combining the following solutions: 9.5 

mL 16% glycerol, 2.0 mL acrylamide/bis acrylamide solution, 1.0 mL SinuLytes 

3-7, 12 μL TEMED, 35 μL 10% ammonium persulfate. After swirling the 

solution the gel was poured. 

2. The gel is divided in two fields using electrode paper wicks on the long sides 

and in the middle of the gel. In this way 96 samples can be run with a running 

distance of 5 cm. 

3. Prefocusing is carried out at settings 600 Volts, 60 mA, 12 Watts for 75 Volthours 

4. The corn and sunflower seed extracts (8 μL per sample) are applied to the 

horizontal IEF gel in the range 3–7 using the 96-well applicator strip. 

5. Focusing is carried out in two runs. Run 1 with 200 Volts, 60 mA, 12 Watts 

for 50 Volthours and run 2 at settings 1000 Volts, 60 mA, 12 Watts for 1,000 

Volthours. 

6. After focusing, proteins are fixed in TCA solution for 10 min, stained with CBB 

solution for 10 min, and then the gel is destained several times for 10 min until 

the background is clear. 

7. For ADH IEF of tomato seed extracts gels of size 260 × 188 mm, and thickness 

of 0.2 mm are made fresh by combining the following solutions: 9.5 mL 16% 

glycerol, 2.0 mL acrylamide/bis acrylamide , 1.0 mL SinuLytes 3-10, 12 μL 

TEMED, 35 μL l10% ammonium persulfate. After swirling the solution the gel 

was poured.





9. Prefocusing is carried out at settings 600 Volts, 60 mA, 12 Watts for 75 Volthours 

10. The tomato seed extracts (8 μl per sample) are applied to the horizontal IEF gel 

using the 96-well applicator strip. 

11. Focusing is carried out in two runs. Run 1 with 200 Volts, 60 mA, 12 Watts 

for 50 Volthours and run 2 at settings 1000 Volts, 60 mA, 12 Watts for 700 

Volthours. 

12. The ADH staining solution is prepared just before the end of the focusing. 

Tris-HCl buffer is brought to 37°C. 5 mL ethanol, 0.05 g NAD, 0.05 g MTT 

and 0.01 g PMS are added, and dissolved by mixing. The gel is stained at 37°C 

until the bands can be clearly visualized (5–10 min). After removal of the stain, 

the gel is destained 10 min in 2% (v/v) acetic acid and then rinsed 10 min with 

water. 

3.3. Inter-SSR PCR 

1. The PCR mix is prepared just before PCR by adding the reaction mixture components 

together, including 3 μL of the DNA solution gained by extraction under 

Section 3.2. 

2. The PCR mix is added to the PCR plates and PCR is carried out with the following 

program steps: an initial 5 min step at 94ºC, the 40 cycles of 0.3 min at 94°C, 

0.45 min at 55°C, and 2 min at 72°C, which is followed by a final step of 5 min 

at 72°C. 

3.4. DNA Electrophoresis 

1. For horizontal electrophoresis of pepper ISSR PCR fragments, gels of size 260 × 

188 mm and thickness of 0.2 mm are used. The following gel solution is used to 

pour the gels: 6.25 mL gel buffer, 9.5 mL 20% glycerol, 2.08 mL acrylamide/bis 

acrylamide 29: 1 solution, 0.02 mL TEMED, 0.14 mL 10% ammonium persulfate. 

The gel is poured and used the same day or stored at 4°C (for maximally 5 d). 

2. To the PCR mix 4 μL of SB is added and of these samples, 4 μL is pipetted into 

the in-gel wells of the gel. 




4. Electrophoresis is carried out at 15°C at power settings of 600 V, 40 mA and 

24 W for 75 min 

5. After electrophoresis the gel is incubated in fix solution for at least 3 min and then 

rinsed with water for 30 s. There after the gel is incubated in staining solution for 

20 min, washed in water for 1 min, and then the DNA fragments are visualized 

by adding developer solution. The developer solution is refreshed after 1 min. 

The time for development is 3–5 min depending on the amount of DNA present


in the gel. Staining reaction is stopped by discarding the developer solution and 

adding stop solution. After 5 min in the stop solution the gel is washed with water 

for at least 30 min and dried on the air. 

4. Notes 

1. Before operation of the Terminator, the head with the 96 pegs must be placed. 

It must be assured to place the head carefully, by lowering the Terminator head 

slowly above the Terminator base plate. Then it is slowly lowered to assure that 

the pegs fit into the wells of the micro plate. After operation, the head must be 

removed carefully to avoid sample going from one well to another. 

2. The Voltage to be applied for operation of the Terminator must be determined 

empirically. The knob of the variable power supply must be turned slowly to 

increase the Voltage until the head starts clearly vibrating and macerating the 

tissue. This generates quite some noise by the pegs that hit the bottom of the 

plate. If the tissue is disrupted—this may take only a few seconds—the noise 

decreases, and homogenization can continue at a constant voltage somewhere 

between 90 and 130 V. The time needed for complete homogenization may also 

be determined empirically but is no more than 3 min. 

3. Cleaning and maintenance of the Terminator is very simple. After operation, the 

Terminator head can be cleaned by spraying the pegs with water using a siphon 

and then the pegs can be dried on the air. The head can be placed with the 

pegs on a tissue. When kept clean, the Terminator needs no maintenance. Several 

Terminators operate now for more than 10 yr in several labs without maintenance. 

For thorough cleaning and decontamination the 96-peg plate can be detached 

from the vibrator head by turning the vibrator head counter clockwise. 

4. Micro plates from Costar are used. However, as standard 96-well flat-bottom 

micro plates are produced worldwide using the same format, the Terminator is 

compatible with 96-well flat-bottom micro plates of all major suppliers. 

5. SinuLytes are used as ampholytes in IEF gels. Alternatively, other sources of 

ampholytes are available. But SinuLytes are superior in our hands because of 

a low molecular weight (average molecular weight is between 400 and 700 

Dalton) of the many amphoteric compounds, which means fast fixing, staining, 

and destaining of gels. In addition, high buffer capacity and solubility at pI, even 

conductivity along the gel and linear pH results in superior performance. 

6. If one views the operation of the Terminator, one may easily suspect (at first sight) 

that cross-contamination (sample of one well contaminates sample of another well) 

is likely. However, extensive experiments to study possible cross-contamination 

were carried out. One may also view Fig. 3B. The single-banded pattern does not 

contain a trace of other bands that are present in other lanes. Further, for making 

the image of Fig. 4, pepper seeds were placed in such a way in the micro plates 

that each variety lacking the intense band (see Fig. 4) is surrounded by seeds 

of a variety that has the intense band. In this way cross-contamination would 

become easily visible from the resulting DNA banding pattern. This shows that


Fig. 3. Images of isoelectric focusing gels showing analysis of homogenates prepared 

using the Terminator. A CBB stained banding pattern of corn seed extract focused 

in the pH range 3–7. The image shows genetic variability of the corn seed storage 

proteins. B CBB stained banding pattern of sunflower seed extract focused in the pH 

range 3–7. The image shows genetic variability of the sunflower seed storage proteins 

C ADH stained banding pattern of tomato seed extract in the pH range 3–10. The 

image shows the two known variants of the dimeric enzyme alcohol dehydrogenase 

from tomato seeds. 

Fig. 4. Banding pattern resulting from ultra-thin layer electrophoresis of PCR 

samples using pepper seed DNA as template. One anchored primer is used for PCR. 

Pepper varieties were taken that differ in presence of an intense PCR band. Prior to 

homogenization using the Terminator, the pepper seeds were put in the micro plate 

in such a way that each seed having the band was surrounded by a seed lacking the 

band. The variable band is easily visible in the rectangle on the left in the image. An 

exploded view is made in the right at the bottom. The white square indicates the area 

of the gel that was eluted for further PCR (see Note 6).


no cross-contamination occurred. To exclude even traces of cross-contamination, 

the gel area indicated with a white square in Fig. 4 was eluted and the sample was 

used for PCR. DNA electrophoresis showed no trace of DNA at that particular 

place in the gel. 

7. Several types of cork borers are commercially available to cut round parts from 

large seeds such as corn, sunflower, and squash. Also leaf samples can best be 

taken using a cork borer or a puncher that cuts small leaf discs. It works best to 

cut discs of similar size as the bottom of the micro plate wells. First put the discs 

in the wells and then add the extraction fluid. In this way the leaf tissue can be 

optimally homogenized. 

References 

1. van den Berg, B.M. (1998) Isoelectric focusing in the vegetable seed industry. 


2. van den Berg, B.M. and Tamboer, J.H.A. (1992) The terminator, an apparatus for 

simultaneous homogenization of 96 small seeds individually. Electrophoresis 13, 

9, 10. 

3. van den Berg, B.M. (1997) Horizontal ultrathin-layer multi-zonal electrophoresis of 

DNA: an efficient tool in analysis of PCR fragments. Electrophoresis, 18, 2861–64. 

4. van den Berg, B.M., Burg, H.C.J., Tamboer, J.H.A., and Grapendaal, B. (1992) 

Equipment for rapid homogenization of high numbers of plant tissue for 

electrophoretic analysis of proteins. Electrophoresis 13, 76–81. 

5. van den Berg, B.M. and Gabillard, D. (1994) Isoelectric focusing in immobilized pH 

gradient of melon (Cucumis melo L.) seed protein: methodical and genetic aspects 

and application in breeding. Electrophoresis 15, 1541–51. 

6. van den Berg, B.M. (1990) Inbred testing of tomato (Lycopersicon esculentum) F1 

varieties by ultrathin-layer isoelectric focusing of seed protein. Electrophoresis 11, 

824–29. 

7. van den Berg, B.M. (1991) A rapid and economical method for hybrid purity testing 

of tomato (Lycopersicon esculentum L.). F1 hybrids using ultrathin-layer isoelectric 

focusing of alcohol dehydrogenase variants from seeds. Electrophoresis 12, 64–9. 

8. Zietkiewicz, E., Rafalski, A. and Labuda, D. (1994) Genome fingerprinting by 

simple sequence repeat (SSR)-anchored polymerase chain reaction amplification. 

Genomics 20, 176–83.

15 

Isolation of Mitochondria from Plant Cell Culture 

Etienne H. Meyer and A. Harvey Millar 

Summary 

Mitochondria carry out a variety of important processes in plants. Their major role 

is the synthesis of ATP through the coupling of a membrane potential to the transfer of 

electrons from NADH to O 2 via the electron transport chain. The NADH is generated by 

the oxidation of organic acids via the tricarboxylic acid cycle. However, mitochondria also 

perform many important secondary functions such as synthesis of nucleotides, amino acids, 

lipids, and vitamins. Mitochondria contain their own genome and undertake transcription 

and translation by some unique mechanisms; they actively import proteins and metabolites 

from the cytosol, are involved in programmed cell death processes in plants, and respond 

to cellular stress conditions. To understand the extent and mechanisms of mitochondrial 

functions in plants and the way in which their functions are perceived by the nucleus 

requires detailed information about the protein components of these organelles. Isolation 

of mitochondria to identify their proteomes and the changes in these proteomes during 

development and environmental stresses is growing area of research. In this chapter we 

provide a useful method for the isolation of mitochondria from plant cell culture using 

a gentle method of cell disruption based on protoplasts isolation that provides relatively 

high mitochondrial yields. 

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

isolation. 


Plant mitochondria play an important role in plant metabolism. They provide 

energy in the form of ATP, but also they provide a large number of metabolic 

precursors in the form of tricarboxylic acids to the rest of the cell for nitrogen 

assimilation and biosynthesis of amino acids. They play key roles in plant 



163

164 Meyer and Millar 

development, fertility, and susceptibility to disease. Between 1,000 and 1,500 

proteins are expected to be found in this cellular compartment and currently 

many hundreds of these proteins have been identified by proteomics. Studies of 

the plant mitochondria proteome require isolation methods that avoid rupture 

of mitochondrial membranes. These methods should be able to eliminate 

most of the cellular contaminant released after disruption of the plant cell. 

Several extensive methodology reviews (1–3) and more specific methodology 

papers (4–6) are already available on plant mitochondrial purification. All these 

methods are based on a cell disruption by grinding. Here we describe a gentle 

method based on protoplasts isolation. Protoplasts are plant cells after removal 

of the cell wall. This cell wall is made of fibrils of cellulose embedded in a 

matrix of several other kinds of polymers such as pectin and lignin. This rigid 

structure can be digested using two enzymatic activities, cellulase will digest the 

cellulose and pectolyase will break the intercellular pectin bonds. The resulting 

protoplasts are highly fragile and can easily be broken by filtration or homogenization. 

Protoplasts can be isolated from virtually any plant tissue. Plant organs 

give low yields of protoplasts, whereas cell cultures are an excellent starting 

material for protoplast isolation. The method we describe here was optimized 

for the purification of mitochondria from plant cell culture. 

2. Materials 

1. Plant cell material for protoplasts: Depending on the growth rate of the plant 

culture, 5–7 day old cell culture should be used. This culture should be sterile to 

avoid fungal and bacterial contamination. 

2. Enzyme buffer: 0.4M mannitol, 0.7 g/L MES-KOH pH 5.7. Just before use 0.4% 

(w/v) of cellulase and 0.05% (w/v) of pectolyase are added (see Note 1). This 

buffer has to be prepared just before use. 

3. Disruption buffer: 0.4M sucrose, 3 mM EDTA, 50 mM Tris-HCl, pH 7.5, 0.1% 

BSA, 2 mM DTT (Dithiothreitol) which is added just prior the disruption. The 

osmoticum (sucrose) maintains the mitochondria structure and prevents physical 

swelling and rupture of membranes, the buffer (Tris) prevents acidification from 

the contents of ruptured vacuoles, the EDTA inhibits the function of phospholipases 

and various proteases, the BSA will remove free fatty acids and the 

reductant (DTT) prevents damage from oxidants present in the tissue or produced 

on homogenization. This media can be freshly prepared, stored overnight at 4°C 

or frozen at –20°C and stored for many weeks (see Note 2). 

4. Wash buffer: 0.3 M sucrose, 1 mM EDTA, 10 mM MOPS-KOH, pH 7.2. This 

buffer is then used for resuspension of organelle pellets, as the base media for 

Percoll gradients and for washing purified organelle pellets. This media can be 

freshly prepared, stored overnight at 4°C or frozen at –20°C and stored for many 

weeks (see Note 2).

Isolation of Mitochondria from Plant Cell Culture 165 

5. Percoll gradient solutions: Gradients of Percoll (Pharmacia, Uppsala, Sweden) 

(see Note 3) are prepared in the wash buffer on the day of use. This is aided by 

making a 2× wash buffer and adding Percoll and distilled water to make 1× wash 

buffer with the appropriate percentage of Percoll required. The discontinuous step 

gradient was cast using a simple inverted syringe bodies (see Note 4). 

3. Methods 

3.1. Protoplasts Isolation 

The cell culture (5–7 day-old cells) is filtered through two layers of muslin. 

The cells are resuspended in the enzyme buffer (a maximum of 500 g of cells 

per L of buffer) and incubated for 3hinthedark under low agitation (45 

rpm) at 25°C. Then the protoplasts are washed twice to remove all traces of 

the digestion enzymes. The suspension is centrifuged (for 10 min at 800g) and 

the pelleted protoplasts are resuspended in enzyme buffer without enzyme (see 

Note 5). After the second wash, the protoplast pellet is resuspended in cold 

disruption buffer. 

The following steps should be done either in a cold room or on ice using 

4ºC cooled glassware and centrifuge tubes (see Note 2). 

3.2. Protoplasts Disruption 

The protoplasts can be disrupted by filtration through Nylon meshes. The 

suspension is filtrated successively through three different Nylon meshes 

(100 μm, 75 μm, and 30 μm holes). Alternatively, the protoplasts can be broken 

by homogenization in a Dounce homogeniser (or potter) (see Note 6). 

We recommend checking the digestion as well as the disruption by optical 

microscopy (see Note 7). 

3.3. Differential Centrifugation to Obtain a Crude Organelle Pellet 

1. Transfer filtered homogenate into 50, 250, or 500 mL centrifuge tubes, depending 

on the volume of the preparation, and centrifuge in a precooled rotor for 5 min 

at ∼3,000g in a fixed angle rotor in a preparative centrifuge at 4°C. 

2. Decant supernatant gently into another set of centrifuge tubes taking care not 

to transfer the pellet material which contains plastids, nuclei and cell debris. 

Centrifuge supernatant for 15 min at ∼18,000g and the resulting high speed 

supernatant is discarded. The tan, yellow, or green coloured pellet in each tube 

contains an unwashed crude organelle pellet. 

3. Resuspend the pellet in 2–10 mL of wash medium with the aid of a clean, soft 

bristle paint brush.


Fig. 1. Percoll gradient purification of plant mitochondria. (A) Three-step Percoll 

gradient for the purification of Arabidopsis mitochondria made from 5 mL of 50% 

Percoll, 25 mL of 25% Percoll, and 5 mL of 18% Percoll solution from bottom to top. 

This gradient was centrifuged at 40,000g in a fixed angle rotor for 45 min. Amyloplast 

envelopes are concentrated in the 18 to 25% interphase (“a”), the fraction containing 

mitochondria in the 25–40% interphase (“m”). (B) One-step Percoll gradient made 

with 35 mL of 28% Percoll. The fraction containing mitochondria from the three-step 

Percoll gradient was loaded on this gradient which was then centrifuged at 40,000g in 

a fixed angle rotor for 45 min. Mitochondria (“m”) are present on top of the gradient 

whereas peroxisomes and other contaminants (“p”) are located in the bottom part of 

the gradient.


3.4. Density Gradient Purification of Mitochondria 

The crude mitochondrial preparation described above is contaminated by 

thylakoid or amyloplast membranes, peroxisomes and endoplasmic reticulum. 

Further purification is carried out using Percoll (Pharmacia, Uppsala, Sweden) 

density gradients. 

1. Layer washed mitochondria, from up to 80 g of etiolated plant tissue or up to 40 

g green plant tissue, over 35 mL of a 18–25–40% step gradient (from bottom to 

top:5mLof50%, 25 mL of 25%, 5 mL of 18% Percoll in wash buffer) in a 50 

mL centrifuge tube (see Note 2). 

2. Centrifuge at ∼40,000g for 45 min in a fixed angle rotor of a preparative centrifuge 

without braking on the deceleration. 

3. After centrifugation, mitochondria form a white-brown band in the bottom part 

of the gradient (Fig. 1). Aspirate the mitochondria with a Pasteur pipet avoiding 

collection of the yellow or green plastid fractions. Dilute suspension with at least 

four volumes of standard wash medium and centrifuge at ∼18,000g for 15 min 

in 50 mL tubes. 

4. The resultant loose pellets is resuspended in a small amount of wash medium and 

loaded on top of a 28% Percoll continuous gradient (35 mL of 28% Percoll in 

wash buffer in a 50-mL centrifuge tube). 

5. Centrifuged at ∼40,000g for 45 min in an angle rotor of a preparative centrifuge 

without braking on the deceleration. 

6. After centrifugation the mitochondria form a white band in the top part of the 

gradient whereas contaminants such as peroxisomes are located in the bottom part 

of the gradient (Fig. 1). Aspirate the mitochondria with a Pasteur pipet and dilute 

them with at least four volumes of wash buffer and centrifuge again at ∼18,000g 

for 15 min in 50-mL tubes. 

7. Remove the supernatant and resuspend the pellet in wash buffer. Centrifuge again 

at ∼18,000g for 15 min. Resuspend the mitochondrial pellet in wash medium at 

a concentration of 5–20 mg mitochondrial protein/mL. This can be determined 

using a Bradford or Lowry assay. 

8. In our hands 100 g of cell culture yields 95 g of protoplasts and 15 mg of purified 

mitochondria. 

9. Once isolated by density gradient purification, plant mitochondria can be kept 

on ice for 5–6 h without significant losses in membrane integrity and respiratory 

function. Longer-term storage of mitochondria can be achieved by rapid-freezing 

of mitochondrial samples in liquid N 2. Frozen samples can be then kept at –80°C. 

4. Notes 

1. At these concentrations of enzymes, more than 95% of the cells will be converted 

in protoplasts after 3 h. Increasing the concentrations may result in a faster


digestion but also a lot of protoplasts could break due to a longer incubation in 

the buffer. 

2. Preparation of mitochondria should be undertaken as quickly as is possible and 

without samples warming above 4°C or storage for extended periods between 

centrifugation runs. So we highly recommend using chilled buffers. The time 

between homogenization and preparation of the washed crude pellet is the most 

critical for ensuring integrity and high yield. 

3. The colloidal silica sol, Percoll, allows the formation of iso-osmotic gradients 

and through isopycnic centrifugation facilitates a range of methods for the density 

purification of mitochondria. The most common method is the sigmoidal, selfgenerating 

gradient obtained by centrifugation of a Percoll solution in a fixedangle 

rotor. The density gradient is formed during centrifugation at >10,000g 

due to the sedimentation of the poly-dispersed colloid (average particle size 29 

nm diameter, average density = 2.2 g/mL). The concentration of Percoll in 

the starting solution and the time of centrifugation can be varied to optimise a 

particular separation 

4. Step gradients of Percoll are often used as these aids the concentration of 

mitochondria fractions on a gradient at an interface between Percoll concentrations. 

Step gradients can easily be formed by setting up a series of inverted 20-mL 

syringes (fitted with 19-gauge needles) strapped to a flat block of wood, clamped 

to a retort stand over a rack at an angle of 45° containing the centrifuge tubes 

(Fig. 2). The needles are lowered to touch the bevels against the inside, lower 

edge of the tubes. The step gradient solutions are then added (from bottom to 

top) to the empty inverted syringe bodies and each allowed to drain through in 

turn before the addition of the next step solution. 

Fig. 2. Making density gradient. Home-made apparatus for discontinuous gradients, 

see explanation in Note 4


5. The protoplasts are very fragile. Thus the agitation should be very gentle (45 rpm) 

on an orbital shaker. Also, the resuspension of the pelleted protoplasts should be 

as soft as possible to avoid disruption of protoplasts. We recommend resuspending 

the pellet by slowly swirling the tube. 

6. These methods of disruption are dependant on the size of the cells. Small cells 

(diameter below 10 μm) will not be disrupted. Mesh with smaller holes should 

then be used. The disruption of large cells will release a lot of membrane fragment 

which will block the holes of the 10-μm mesh. Then the filtration through the 

10-μm mesh will be replaced by a second filtration through the 30-μm mesh. 

7. The digestion has to be checked by optical microscopy after three hours. A drop 

(approx 50 μL) is largely sufficient. If a lot of nondigested cells remain (nonround 

cells), incubate the suspension a longer time in the enzyme medium. The 

disruption should also be checked to ensure that all the protoplasts are broken. 

Unbroken protoplasts will be pelleted during the low-speed centrifugation and 

lost. If some unbroken protoplasts remain, repeat the filtration or homogenisation 

step. 

References 

1. Neuburger, M. (1985) Higher-Plant Cell Respiration, vol. 18 (Douce, R., Day, DA, 

ed.), pp. 7–24, Springer-Verlag, Berlin. 

2. Douce, R. (1985) Mitochondria in higher plants: Structure, function and biogenesis, 

American Society of Plant Physiologists, Academic Press, Orlando, Florida. 

3. Millar, A. H., Liddell, A., and Leaver, C. J. (2001) Isolation and subfractionation 

of mitochondria from plants. Meth. Cell Biol., 65, 53–74. 

4. Neuburger, M., Journet, E. P., Bligny, R., Carde, J. P., and Douce, R. (1982) 

Purification of plant-mitochondria by isopycnic centrifugation in density gradients 

of percoll. Arch. Biochem. Biophys. 217, 312–23. 

5. Leaver, C. J., Hack, E., and Forde, B. G. (1983) Protein-synthesis by isolated 

plant-mitochondria. Meth. Enzymol. 97, 476–84. 

6. Day, D. A., Neuburger, M., and Douce, R. (1985) Biochemical-characterization of 

chlorophyll-free mitochondria from pea leaves. Aust. J. Plant Physiol. 12, 219–228.

16 

Isolation and Preparation of Chloroplasts 

from Arabidopsis thaliana Plants 

Sybille E. Kubis, Kathryn S. Lilley, and Paul Jarvis 

Summary 

A major area of research in the postgenomic era has been the proteomic analysis 

of various subcellular and suborganellar compartments. The success of these studies is 

to a large extent dependent upon efficient protocols for the preparation of highly pure 

organelles or suborganellar components. Here we describe a simple, rapid, and low-cost 

method for isolating a high yield of Arabidopsis chloroplasts. The method can readily 

be applied to wild-type plants and different mutants, and at different developmental 

stages ranging from 10-day-old seedlings to rosette leaves from older plants. The isolated 

chloroplast fraction is highly pure, with immunologically undetectable contamination from 

other cellular organelles. Chloroplasts isolated using the method described here have been 

successfully used for proteomic analysis, as well as in studies on chloroplast protein import 

and other aspects of chloroplast biology. 

Key Words: Arabidopsis thaliana; chloroplast isolation; chloroplast proteomics; 

organelle isolation; Percoll gradient; plastids; polytron homogenizer. 


Chloroplasts belong to a diverse group of organelles called plastids (1,2). 

Plastids are ubiquitous in plants and algae, and perform numerous essential 

functions including important steps in the biosynthesis of amino acids, 

lipids, nucleotides, hormones, vitamins, and secondary metabolites, as well 

as oxygenic photosynthesis (2,3). In the latter process, energy from sunlight 

is converted into usable chemical bond energy, and the associated redox 

reactions lead to the generation of oxygen from water. Chloroplasts are therefore 



171

172 Kubis et al. 

important sites for the production of organic matter and oxygen, and so provide 

the fuels essential for all higher forms of life (4). 

Completion of the genome sequencing projects for Arabidopsis, rice 

and other species, and the development of efficient methods for routine 

protein identification by mass spectrometry, have enabled numerous largescale 

proteomic studies. Because of the dynamic range limitations associated 

with analyses on highly complex mixtures (i.e., the tendency of abundant 

proteins to mask the presence of less abundant proteins), these studies have 

tended to focus on isolated subcellular components. In plants, chloroplasts have 

received considerable attention in this regard (5–8). The uses of such proteomic 

studies are several-fold: they can confirm the expression and structure of genes 

predicted based on genome sequence analysis in silico; they can provide information 

on subcellular and suborganellar protein localization; and they can even 

be used to estimate the relative abundance of different proteins. Information of 

this nature is particularly important, because it has been estimated that up to 

50% of the proteins encoded by the >26,000 genes in the Arabidopsis genome 

are of unknown function (9). Proteomics is one of the tools being used to 

address this deficiency. 

Most proteins targeted to chloroplasts possess a cleavable, amino-terminal 

targeting signal called a transit peptide (10,11). Using computer programs (e.g., 

TargetP) to detect the presence of a transit peptide, it has been estimated that 

∼4,000 proteins are targeted to chloroplasts in Arabidopsis (9). Unfortunately, 

these in silico methods are not 100% reliable (12), and so the only truly 

dependable method for the determination of protein localization is laboratory 

experimentation. However, there are presently less than 700 entries in a database 

of experimentally determined chloroplast proteins (13), clearly indicating a 

need for further studies. Furthermore, several recent reports have indicated 

that protein targeting to chloroplasts is not as simple as was once thought. 

In a large-scale study of the Arabidopsis chloroplast proteome, only ∼60% 

of the proteins identified were predicted to have a transit peptide (6,14). Of 

the remainder, many appeared to have a signal peptide for ER translocation, 

or no cleavable targeting sequence at all. Intriguingly, direct evidence for 

a protein transport pathway to chloroplasts through the ER and Golgi has 

now been presented (15), and the targeting of proteins lacking a cleavable 

peptide has been described in some detail (16,17). These data demonstrate that 

transit peptide prediction in silico cannot provide a complete picture of the 

chloroplast proteome. The existence of dual-targeted proteins (e.g., proteins 

targeted to mitochondria or the ER as well as chloroplasts) adds an additional 

level of complexity (18,19), further emphasizing the need for experimental 

determination of protein localization.

Isolation and Preparation of Chloroplasts 173 

A number of chloroplast proteomic studies have already been described. In 

addition to the whole chloroplast study mentioned earlier (14), several reports 

have described the proteomes of individual suborganellar compartments of the 

chloroplast: e.g., the thylakoid lumen (20,21), the envelope membrane (22,23), 

the stroma (24), and the lipid-containing structures called plastoglobuli (25). 

In addition, an analysis of plastids in dark-grown plants, called etioplasts, 

revealed a proteome consistent with what one would expect of plastids in 

heterotrophic tissue, along with some novel functions (26). As well as studies 

that simply catalogue the proteins present in a particular subcellular or suborganellar 

compartment, comparative proteomics has been employed with considerable 

success. For example, the chloroplasts from mesophyll cells and bundlesheath 

cells of maize, a C4 plant, were recently compared (27). The data 

not only revealed differential accumulation of carbon metabolism enzymes 

consistent with the C4 photosynthetic mechanism, but also shed light on how 

other plastidic functions are distributed between the two cell types. In another 

example, chloroplasts isolated from Arabidopsis mutants lacking different 

protein import receptor isoforms were compared with wild-type chloroplasts 

(28,29). Different groups of chloroplast proteins were found to be selectively 

deficient in different receptor mutants, indicating that the different receptor 

isoforms likely possess a degree of preprotein recognition specificity (10,11). 

These various studies have demonstrated the utility and value of chloroplast 

proteome analysis, and it is anticipated that proteomics will continue to form 

an essential component of chloroplast research in the future. The chloroplast 

isolation procedure described in technical detail here, and previously (30), has 

been successfully used in proteomic studies (28,29), as well as in other research 

on chloroplast biology (31–33). 

2. Materials 

2.1. Growth of Arabidopsis Seedlings 

1. Seeds, stored in a 1.5-mL microfuge tubes (with a small hole in the lid to allow 

evaporation of any residual moisture) at room temperature. 

2. 70% (v/v) ethanol containing 0.05% (v/v) Triton X-100 (Sigma-Aldrich Ltd., 

Poole, UK), in a Duran bottle at room temperature. For 200 mL, mix 140 mL 

100% ethanol, 60 mL sterile deionized water, and 100 μL Triton X-100. 

3. 100% ethanol, in Duran bottle at room temperature. 

4. Laminar flow hood (e.g., Model P5HB, Bassaire Ltd., Southampton, UK). 

5. Circular filter papers, 9 cm in diameter (Whatman, Banbury, UK; or Fisher 

Scientific, Loughborough, UK). 

6. Industrial methylated spirit (IMS). 

7. Orbital shaker (e.g., Model S01, Stuart Scientific, Stone, UK).


8. Petri dishes, 9 cm in diameter (e.g., Bibby Sterilin Ltd., Stone, UK). 

9. Murashige and Skoog (MS) medium: MS salt and vitamin mixture (Sigma), 

0.5% (w/v) sucrose, and 0.6% (w/v) agar. Sterilize the medium in an autoclave 

(15 min, 121°C, 15 psi), cool to 50°C in a water bath, and then pour into Petri 

plates to a depth of ∼3–4 mm in a laminar flow hood (400 mL medium is 

sufficient for ∼15–20 plates). Allow the plates to dry for 1hinthehood before 

replacing the lids. Prepoured plates can be stored at 4°C, up-side down, sealed 

in a plastic bag for up to 1 month. 

10. Leukopor tape (Beiersdorf AG, Hamburg, Germany) or equivalent (e.g., 

Micropore tape, 3M, Bracknell, UK). 

11. Refrigerator or cold-room (4°C). 

12. Plant tissue culture chamber (e.g., Model CU-36L5, Percival Scientific Inc., 

Perry, Iowa) set at 20°C, providing 100–120 μmol/m 2 /s white light with a longday 

cycle (16-h-light/8-h-dark). 

2.2. Chloroplast Isolation 

These materials are sufficient for one isolation on a single plant sample. 

If multiple samples (e.g., different genotypes) are to be analyzed, additional 

materials will be required (i.e., in points 1, 3, 6, 7, and 9 below). 

1. For one isolation procedure, 25–40 Petri plates of 10-day-old plants, each plate 

containing ∼150–200 seedlings as shown in Fig. 1B (see Notes 1, 2). 

2. Two ice buckets, containing ice. 

3. Two 1-L beakers, one 50-mL beaker, measuring cylinders, and one funnel. 

4. Polytron; e.g., Kinematica Model PT10-35 (Kinematica AG, Littau, 

Switzerland), with a large rotor (PTA 20 S) and speed set to 4 on scale of 11 

(see Note 3). 

5. Cold-room (4°C). 

6. Miracloth (Calbiochem Ltd., Nottingham, UK); two squares of about 15 × 15 cm. 

7. Two 30-mL Nalgene tubes and one 250-mL Nalgene tube (Fisher). 

8. Percoll (Amersham Biosciences, Little Chalfont, UK); an opened bottle can be 

stored at 4°C for several months. 

9. Continuous Percoll gradient. Before use, make up 26 mL of gradient mixture as 

follows: 13 mL Percoll, 13 mL 2× chloroplast isolation buffer (see below), and 5 

mg glutathione (roughly, the tip of a small spatula). Mix the components together 

in a 30-mL Nalgene tube, ensuring that the glutathione is completely dissolved. 

Precentrifuge in a fixed angle rotor at 43,000g max for 30 min (brake off) at 4°C; this 

is equivalent to 19,000 rpm in an SS-34 rotor in a Sorvall RC6 centrifuge (Kendro, 

Asheville, North Carolina), with acceleration set to 7 and deceleration set to 2. 

Gradients can be prepared the day before, and then stored overnight at 4°C. 

10. Chloroplast isolation buffer (CIB): 0.3M sorbitol, 5 mM MgCl 2,5mM EGTA, 

5mM EDTA, 20 mM HEPES/KOH pH 8.0, 10 mM NaHCO 3. This is prepared 

as a 2× CIB stock (see Notes 4, 5). The final pH of the solution should be 8.0.


Fig. 1. Different steps of the chloroplast isolation procedure. (A) An MS agar plate 

carrying Arabidopsis seeds sown at an appropriate density (∼150 seeds per plate) for 

use in chloroplast isolation after growth for 10–14 d. (B) A plate similar to that shown 

in A, after the plants have been allowed to grow for 14 d in a tissue culture cabinet. 

(C, D) The Arabidopsis seedlings are harvested from the plates by hand, taking care 

not carry over any of the agar medium. (E) The Arabidopsis tissue is transferred to a 

50-mL beaker containing cold chloroplast isolation medium, and then disrupted using 

five consecutive rounds of homogenization using a polytron blender. (F) Following 

filtration through Miracloth, the homogenate is loaded onto a preformed continuous 

Percoll gradient. (G) After centrifugation of the loaded Percoll gradient, two green 

bands are apparent: the upper band contains broken material, whereas the lower band 

contains intact chloroplasts.


11. HEPES-MgSO 4-sorbitol (HMS) buffer: 50 mM HEPES/NaOH pH 8.0, 3 mM 

MgSO 4, 0.3M sorbitol (see Note 6). The final pH of the solution should be 8.0. 

2.3. Establishing Yield and Intactness of Chloroplasts 

1. Hemocytometer with a 0.1 mm deep counting chamber and a ruling pattern of 1/400 

mm 2 (e.g., Improved Neubauer BS748, Hawksley Technology, Lancing, UK). 

2. Cover glass (e.g., 22 × 22 mm, No.1, Chance Propper Ltd., Warley, UK). 

3. Phase-contrast microscope (e.g., Carl Zeiss AG, Oberkochen, Germany). 

4. HMS buffer (see Section 2.2.). 

5. Tissue paper (e.g., Kimcare, Kimberly-Clarke Europe Ltd., Reigate, UK). 

2.4. Preparation for Proteomics 

1. Ice bucket containing ice, with lid. 

2. Lysis buffer (see Table 1). Recommended detergent components: ASB- 

14 (amidosulfobetaine-14) (Calbiochem Ltd.); CHAPS (3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate) 

(Sigma-Aldrich Ltd.); SB3-10 

Table 1 

Lysis buffers 

Buffer Component Concentration 

1. Standard buffer CHAPS 4% (w/v) 

Urea 8M 

Tris-HCl, pH 168–169 10–30 mM 

Magnesium acetate 5 mM 

2. Thiourea buffer CHAPS 4% (w/v) 

Urea 7M 

Thiourea 2M 

Tris-HCl, pH 8.0–9.0 10–30 mM 


3. ASB-14 buffer a ASB-14 2% (w/v) 

Urea 7M 

Thiourea 2M 

Tris-HCl, pH 8.0–9.0 10-30 mM 


4. SDS buffer b SDS 2% (w/v) 

Tris-HCl, pH 8.0–9.0 10-30 mM 


a ASB-14 can be substituted with NP40, SB3-10 or various other sulfobetaine-derived detergents. 

b SDS-containing solutions must be diluted to a final concentration of 0.2% or less before 

successful isoelectric focusing can take place.


(3-(decyldimethylammonio)propanesulfonate inner salt) (Sigma-Aldrich Ltd.); 

Nonidet P40 (NP40) substitute ([octylphenoxy]polyethoxyethanol) (USB Corp., 

Cleveland, Ohio). 

3. Hand-held, plastic pestles for use with 1.5-mL microfuge tubes (e.g., pellet pestle, 

blue polypropylene, Sigma-Aldrich Ltd.). 

4. Bench-top microcentrifuge (e.g., Eppendorf 5415D, Eppendorf UK Ltd., 

Cambridge, UK). 

5. Protein concentration estimation kit (e.g., DC Protein Assay, Bio-Rad Laboratories 

Ltd., Hemel Hempstead, UK; or PlusOne 2-D Quant, Amersham Biosciences). 

6. Plastic cuvets, 1 mL (e.g., Sarstedt Ltd., Leicester, UK). 

7. Spectrophotometer (e.g., Spectronic; Thermo Electron Corp., Waltham, 

Massachusetts). 

3. Methods 

3.1. Growth of Arabidopsis Seedlings 

1. Transfer the appropriate amount of seeds (e.g., for 40 Petri plates each carrying 

∼150–200 seeds, an amount equivalent to ∼240–320 μL will be required) into a 

sterile 1.5-mL microfuge tube, and add 1 mL of 70% (v/v) ethanol, 0.05% (v/v) 

Triton X-100 (see Note 7). 

2. Shake the tube by hand to ensure that all seeds are suspended in the solution, and 

then place the tube, oriented horizontally, onto an orbital shaker. Shake at 250 

rpm for 5 min. 

3. Allow the seeds to settle at bottom of tube, remove the supernatant with a pipet, 

and then add 1 mL of 100% ethanol. Shake the tube by hand first of all, and then 

on the orbital shaker at 250 rpm for 10 min. 

4. Meanwhile, switch on the laminar flow hood and sterilize the interior surfaces 

with IMS. Take an appropriate number of round filter papers (at least one per 

seed sample), and fold them in half to create a crease (this will facilitate seed 

sowing later). In the hood, soak (and sterilize) the filter paper(s) with IMS. Allow 

the filter paper(s) to dry. 

5. Using a cut 1-mL Gilson pipet tip (lacking ∼5 mm from the fine end, to increase 

the aperture size), pipet the seeds onto the sterilized filter paper(s) and leave to 

dry. This takes about 15 min. 

6. Sow seeds onto Petri plates containing MS medium. For 10- to 14-day-old 

plants, an appropriate density is ∼150–200 seeds/plate, as shown in Fig. 1A. (see 

Notes 7, 8). 

7. Seal each plate with Leukopor tape. 

8. Incubate plates upside down (to prevent condensation accumulating on the surface 

of the agar) at 4°C for at least 2d(upto4dispossible) to break seed dormancy 

and synchronize germination. 

9. Grow plants for 10–14 d in a plant tissue culture chamber. Plants grown for 14 d 

are shown in Fig. 1B.


3.2. Isolation of Chloroplasts 

The isolation procedure should be started as early as possible in the morning. 

During the isolation procedure, plant material should kept at 4°C. The first 

steps (stages 2–4 below) can be carried out on the bench in the laboratory, but 

the isolation itself should be carried out at 4°C in the cold-room. The method 

below describes an isolation on a single plant sample. If multiple samples (e.g., 

different genotypes) are to be used, additional materials will be required (see 

Section 2.2). 

1. Preparation, on the day before the isolation: Place a 200-mL aliquot of 2× CIB, a 

50-mL aliquot of 2× CIB, a 50-mL aliquot of HMS buffer, and 200 mL deionized 

H 2O into the refrigerator or cold-room; in the morning, the frozen solutions 

should have thawed. Place all rotors in the cold-room to precool overnight. 

2. Preparation, on the day of the isolation: Prepare CIB by adding 200 mL chilled 

sterile deionized H 2O to 200 mL thawed 2× CIB; mix well and keep on ice. Put 

100 mL of the CIB into a 1-L plastic beaker and keep on ice. Place a second 

1-L plastic beaker on ice, with funnel containing two layers of Miracloth. The 

250-mL and 30-mL Nalgene tubes should also be placed on ice to precool. 

3. Prepare a continuous Percoll gradient as described in Section 2.2 (point 9), and 

keep the tube on ice after precentrifugation (see Note 9). 

4. Take plates out of plant tissue culture chamber and remove the Leukopor tape. 

Remove the seedlings from the medium by gently scraping them off with a 

gloved hand (see Fig. 1C, D), avoiding carry over of medium because this 

interferes with the isolation, and place them into the 100 mL CIB in the 1-L 

beaker on ice. 

5. During homogenization, a total of 100 mL CIB is used per sample; this is used 

in five, consecutive rounds of homogenization, each one using 20 mL CIB (see 

Note 10). 

6. Place 20 mL fresh CIB into the 50-mL plastic beaker, and then transfer the 

seedlings into the beaker. 

7. Place the plant material under the rotor of polytron, and homogenize for 1-2 s (see 

Fig. 1E). The optimal conditions for the homogenization have to be established 

empirically (see Note 11). 

8. Filter the homogenate through two layers of Miracloth into the 1-L beaker on 

ice. Gently squeeze the Miracloth around the plant material. 

9. Place a second 20-mL aliquot of fresh CIB into 50-mL beaker, and return the 

plant material to the beaker. 

10. Repeat points 7–9 until all five 20-mL aliquots of CIB have been used, and 

five rounds of homogenization and filtration have been completed. The plant 

material will gradually become disrupted during the procedure. 

11. Transfer the pooled, filtered homogenate into the 250-mL Nalgene tube on ice, 

and centrifuge at 1,000g max for 5 min (brake on) at 4°C; this is equivalent 

to 3,000 rpm in an SLA-1500 rotor in a Sorvall RC6 centrifuge, with both 

acceleration and deceleration set to 7).


12. Pour off most of the supernatant, and resuspend the pellet in the residual ∼500 

μL supernatant by rotating the tube on ice; do not resuspend by pipeting. 

13. Transfer the resuspended homogenate onto the top of the preformed Percoll 

gradient, using a cut 1-mL Gilson pipet tip (lacking ∼5 mm from the fine end, 

to increase the aperture size). Pipet very slowly so as not to disturb the gradient 

(see Fig. 1F). 

14. To separate the intact chloroplasts from broken chloroplasts and other debris, 

centrifuge in a swing-out rotor at 7,800g max for 10 min (brake off) at 4°C; this 

is equivalent to 7,000 rpm in an HB-6 rotor in a Sorvall RC6 centrifuge, with 

acceleration set to 7 and deceleration set to 2. 

15. After centrifugation, remove the tube carefully and place it on ice. The lower 

green band in the gradient contains intact chloroplasts, whereas the upper band 

contains broken chloroplasts (see Fig. 1G). Broken chloroplasts are removed 

and discarded by pipeting, and then the intact chloroplasts are recovered using 

a 1-mL Gilson pipet tip (cut at the end), and transferred into a precooled 30 

mL Nalgene tube. The volume of recovered intact chloroplasts can range from 

2mLto6mL. 

16. Add 25 mL HMS buffer to the chloroplasts and invert the tube carefully 2–3 

times to wash off the Percoll. 

17. Centrifuge the chloroplasts in a swing-out rotor at 1,000g max for 5 min (brake 

on) at 4°C; this is equivalent to 2,000 rpm in an HB-6 rotor in a Sorvall RC6 

centrifuge, with both acceleration and deceleration set to 7. 

18. Gently pour off the supernatant, and then resuspend the chloroplasts in 

∼150–400 μL fresh HMS buffer by rotating the tube on ice; do not resuspend 

by pipeting. 

3.3. Establishing the Yield, Intactness and Purity of the Isolated 

Chloroplasts 

If necessary, the yield of chloroplasts and their intactness can be assessed as 

follows. Alternatively, for some applications it may be appropriate to proceed 

directly to downstream procedures (e.g., Section 3.4). 

1. Add 5 μL of isolated chloroplasts to 495 μL of HMS buffer in a 1.5-mL microfuge 

tube, and then mix gently by flicking the tube to obtain a 1:100 dilution. 

2. Pipet ∼60–80 μL of the diluted suspension onto the counting chamber of the 

hemocytometer, and place a cover glass on top. 

3. Drain the excess suspension with tissue paper. 

4. Count the number of chloroplasts in 10 different 1/400 mm 2 squares (e.g., those 

on each diagonal line), using a phase contrast microscope with a 16× objective. 

The number of chloroplasts per square should average between 10 and 20. If too 

few or too many chloroplasts are present, adjust the dilution factor (point 1 above) 

accordingly and repeat the procedure. Intact chloroplasts appear round and bright 

green (see Fig. 2A), and under phase-contrast are surrounded by a bright halo of 

light.


5. The concentration (number of chloroplasts per mL) is calculated as follows: n (the 

average number of chloroplasts per 1/400 mm 2 square) × 25 (the total number 

of squares in the grid) × 100 (the dilution factor employed) × 10 4 (scaling factor 

needed to express the data per mL, because the volume above the 25 squares is 

only 0.1 μL). 

6. Calculate the actual yield of chloroplasts by multiplying the concentration (number 

of chloroplasts per mL) by the volume of chloroplast suspension used in Section 

3.2, point 18. 

Samples prepared using the methodology described here are mostly intact, 

and exhibit minimal contamination from other cellular compartments. To illustrate 

these points, we analyzed typical chloroplast preparations by phasecontrast 

light microscopy (see Fig. 2A,B), by transmission electron microscopy 

(see Fig. 2C), and by immunoblotting using high-titre antibodies against components 

of various cellular organelles (see Fig. 3). Microscopic analysis did not 

reveal any evidence of significant contamination from other cellular organelles. 

Fig. 2. Light and electron micrographs of isolated chloroplasts. (A, B) Chloroplasts 

isolated from 14-day-old Arabidopsis seedlings were analysed by phase-contrast 

light microscopy, at both low (A) and high (B) magnification, and the majority were 

adjudged to be intact. Size bars indicate 100 μm. (C) The integrity of the chloroplasts 

was confirmed by transmission electron microscopy, as described previously 

(30). No evidence of significant contamination of the chloroplast preparation with other 

organelles was observed using either method. Size bar indicates 2 μm.


Immunologically, the chloroplasts contained undetectable contamination from 

all organelles tested:


3.4. Preparation of the Chloroplasts for Proteomics 

1. Chloroplast lysis and solubilization: Resuspend the intact chloroplast sample in 

a small volume of the appropriate lysis buffer (see Table 1). To ensure optimal 

lysis, a plastic hand-held pestle should be employed in conjunction with a 1.5-mL 

microfuge tube. Solubilization should be allowed to proceed on ice for 30 min, 

with frequent grinding of the chloroplast material using the pestle (see Note 12). 

After this period, centrifuge at 1,400gmax (∼3,900 rpm in an Eppendorf 5415D 

microfuge using the standard 24-place rotor) for 10 min at 4°C, and then remove 

and retain the supernatant. 

2. Protein Precipitation: It is often necessary to precipitate proteins derived from 

plant material lysis, to ensure the removal of phytophenols, lipids and other 

cellular components that may interfere with downstream processing steps. There 

are several commercial kits available for this purpose; e.g., PlusOne 2-D Clean-Up 

Fig. 4. Analysis of chloroplast protein samples by CyDyeDIGE 2D-PAGE. Chloroplasts 

were isolated from 10-day-old Arabidopsis seedlings using the described protocol, 

and then analysed using CyDyeDIGE technology as described previously (28,29). An 

image of a typical pH 3–10 nonlinear 2D-DIGE gel is shown (the pH range is indicated 

at the top). Two different chloroplast protein samples (one from wild-type Arabidopsis, 

and another from a mutant) were labeled with minimal CyDyeDIGE fluors, and then 

analysed simultaneously. The images from the wild-type sample (Cy5) and the mutant 

sample (Cy3) have been overlaid. If the Cy5 signal were colored red and the Cy3 signal 

were colored green, spots corresponding to proteins spots corresponding to proteins 

deficient in the mutant would appear red, whereas proteins enriched in the mutant would 

appear green. The large (LSU) and small (SSU) subunits of Rubisco are indicated.


kit (Amersham Biosciences), or PerfectFOCUS (Genotech, St. Louis, Missouri). 

Alternatively, refer to other chapters in this book. 

3. Protein estimation: The protein concentration of the sample(s) in lysis buffer 

must be determined before proceeding with any proteomics methodologies. There 

are several commercial kits available for this purpose, but it is imperative that 

the method used is compatible with samples containing detergents; e.g., RC/DC 

Protein Assay (Bio-Rad Laboratories Ltd.), or PlusOne 2-D Quant (Amersham 

Biosciences). 

4. Once these various preparative procedures have been completed, the solubilized 

chloroplast protein samples can be subjected to a variety of different methodologies 

for proteome analysis, such as difference gel electrophoresis (DIGE) (see 

Fig. 4). 

4. Notes 

1. For 10-day-old plants, this is roughly equivalent to ∼6,000–8,000 individuals 

or ∼20 g tissue. For older plants, few individuals are needed, and the density of 

plants per plate should be reduced. 

2. It is likely that more plates will be needed when working with sick, mutant 

plants. 

3. The method works just as well using a Kinematica PT20 polytron with a small 

rotor (13 mm diameter) at ∼40% maximum speed, as described previously (30). 

4. Prepare 2× CIB and store in 200-mL aliquots in 400 mL Duran bottles at –20°C. 

Before use, thaw overnight at 4°C, then make CIB by adding 200 mL sterile 

deionized H2O and mixing well. 

5. Small amounts of 2× CIB are needed for preparation of Percoll gradients, so 

aliquots of 2× CIB in 50-mL tubes are recommended for storage at –20°C. The 

buffer can be thawed and frozen several times (up to 5 times). Mix well after 

thawing. 

6. Prepare 50-mL aliquots and store at –20°C. The buffer can be thawed and frozen 

several times (up to 5 times). Mix well after thawing. 

7. Do not wear gloves when handling seeds. The seeds will stick to the gloves 

because of static electricity. Instead, sterilize hands with IMS and avoid touching 

the seeds directly. 

8. When working with certain, particularly sick mutants, it may be beneficial to 

use MS medium supplemented with 100 mM (∼3%) sucrose. 

9. When using newly thawed aliquots of 2× CIB, mix well before use to obtain a 

homogeneous solution. 

10. Sometimes, yields can be improved by increasing the volume of CIB and 

increasing the number of rounds of homogenization; this may be particularly 

advantageous when using large amounts of tissue. 

11. When using small rotors (e.g., PTA10S with the Kinematica PT10-35, or the 

13-mm rotor with the Kinematica PT20) good homogenization is achieved by 

moving the small beaker up and down 8–10 times quickly (taking ∼3–4sin


total), in each one of the five rounds of homogenization. When working with 

chlorotic or sick mutants, homogenization times should be reduced according to 

the severity of the mutant phenotype. For example, when isolating chloroplasts 

from the ppi1 mutant (28), homogenization time was reduced to ∼1–2 s for each 

of the five rounds of homogenization. When using larger rotors (e.g., PTA20S 

with the Kinematica PT10-35), the up and down movement of the beaker is not 

possible because of the strong suction generated. 

12. Buffers containing ASB-14 will solidify at 4°C. Therefore, lysis and solubilization 

using such buffers should be carried out at room temperature, and the 

lysis buffer should include a protease inhibitor cocktail (e.g., Complete Protease 

Inhibitor Cocktail Tablets, Roche Diagnostics Ltd., Lewes, UK). 


We thank Ramesh Patel for technical assistance, and Natalie Allcock 

and Stefan Hyman (Electron Microscope Laboratory, Faculty of Medicine 

and Biological Sciences, University of Leicester) for transmission electron 

microscopy. We are grateful to Sabina Kovacheva and Ramesh Patel for 

their helpful comments on the manuscript. We thank Marc Boutry (PMA2 

H + -ATPase), Karl-Josef Dietz (PrxII F), Richard Trelease (catalase), Peter 

Shaw (histone H3), Henrik Scheller (PSI-D), and Kenton Ko (SSU) for generously 

providing antibodies. This work was supported by the Biotechnology 

and Biological Sciences Research Council (BBSRC) Genomic Arabidopsis 

Resource Network (GARNet), by the Royal Society Rosenheim Research 

Fellowship (to P.J.), and by BBSRC Grants 91/P12928 and BBS/B/03629 

(to P.J.). 

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17 

Isolation of Plant Cell Wall Proteins 

Elisabeth Jamet, Georges Boudart, Gisèle Borderies, 

Stephane Charmont, Claude Lafitte, Michel Rossignol, Herve Canut, 

and Rafael Pont-Lezica 

Summary 

The quality of a proteomic analysis of a cell compartment strongly depends on the 

reliability of the isolation procedure for the cell compartment of interest. Plant cell walls 

possess specific drawbacks: (1) the lack of a surrounding membrane may result in the loss 

of cell wall proteins (CWP) during the isolation procedure; (2) polysaccharide networks 

of cellulose, hemicelluloses, and pectins form potential traps for contaminants such as 

intracellular proteins; (3) the presence of proteins interacting in many different ways 

with the polysaccharide matrix require different procedures to elute them from the cell 

wall. Three categories of CWP are distinguished: labile proteins that have little or no 

interactions with cell wall components, weakly bound proteins extractable with salts, and 

strongly bound proteins. Two alternative protocols are decribed for cell wall proteomics: 

(1) nondestructive techniques allowing the extraction of labile or weakly bound CWP 

without damaging the plasma membrane; (2) destructive techniques to isolate cell walls 

from which weakly or strongly bound CWP can be extracted. These protocols give very 

low levels of contamination by intracellular proteins. Their application should lead to 

a realistic view of the cell wall proteome at least for labile and weakly bound CWP 

extractable by salts. 

Key Words: Arabidopsis thaliana; bioinformatics; cell fractionation; cell wall; cell 

wall protein; plant; proteomics. 


Plant cell wall proteins (CWP) present specific complexities in addition to the 

difficulties usually encountered in proteome analysis, such as protein separation 

and detection of scarce proteins (1). They are embedded in an insoluble 



187

188 Jamet et al. 

polysaccharide matrix and interact with other cell wall components, making 

their extraction challenging. Current models of cell wall structure describe the 

arrangement of their components into two structurally independent and interacting 

networks, embedded in a pectin matrix (2,3). Cellulose microfibrils and 

hemicelluloses constitute the first network; the second one is formed by structural 

proteins. Three types of CWP can be distinguished, according to their 

interactions with cell wall components (4). CWP can have little or no interactions 

with cell wall components and thus move freely in the extracellular 

space. Such proteins can be found in liquid culture media of cell suspensions 

and seedlings or can be extracted with low ionic strength buffers. We call this 

fraction “labile proteins,” most of them have acidic pI ranging from 2 to 6 

(Fig. 1A). Alternatively, CWP might be weakly bound to the matrix by Van 

der Waals interactions, hydrogen bonds, and hydrophobic or ionic interactions. 

Such proteins may be extracted by salts and most of them have basic pI ranging 

from8to11(Fig. 1B) so that they are positively charged at the acidic pH of cell 

walls. Even though most of the cell wall polysaccharides are neutral, negatively 

Fig. 1. pIs of labile and weakly-bound CWP. pIs of CWP identified in 

several proteomic studies (4) were calculated (www.iut-arles.up.univ-mrs.fr/w3bb/d_ 

abim/compo-p.html) after removal of their predicted signal peptides (http:// 

psort.nibb.ac.jp/form.html). Three groups of proteins were considered: (A) labile 

proteins; (B) salt-extracted proteins, i.e. proteins extracted with salt solutions or 

chelating agent; (C) all proteins. Reprinted from (4), Copyright (2005), with permission 

from Elsevier.

Isolation of Plant Cell Wall Proteins 189 

charged pectins contain polygalacturonic acid that provides negative charges 

for interactions with basic proteins. Such interactions would be modulated by 

pH, degree of pectin esterification, Ca 2+ concentration, and by the mobility 

and diffusion coefficients of these macromolecules (3,5). Finally, CWP can 

be strongly bound to cell wall components so that they are still resistant to 

salt-extraction. As examples, extensins are cross-linked by covalent links (6,7) 

and peroxidases can have a high affinity for Ca 2+ -pectate (8). 

The available techniques described in this chapter allow the extraction of 

labile and weakly bound CWP. Because labile proteins can be lost during 

the preparation of cell walls, they must be extracted from tissues by nondestructive 

techniques such as vacuum infiltration (9), or recovered from liquid 

culture media from cell suspension cultures or seedlings (10,11). Weakly bound 

CWP can be extracted with salts or chelating agents from living cells with 

nondestructive techniques (9,10) or from purified cell walls with destructive 

techniques. At present, there is no efficient procedure to release CWP strongly 

bound to the extracellular matrix. Structural proteins, for instance extensins or 

PRP, can be cross-linked via di-isodityrosine bonds (6,12). Purified cell walls 

appear as the most suitable material to isolate such proteins. However, until 

now, extensins have only been eluted with salts before their insolubilization 

from cell suspension cultures (13). 

2. Materials 

A major problem in proteomics is the occurrence of keratins that can contaminate 

materials and working solutions. The presence of keratins can prevent the 

identification of proteins of interest by mass spectrometry. It is necessary to pay 

attention to all possible sources of contamination at all steps of the following 

protocols. Powder-free gloves should be permanently worn and washed with 

soap before their first use. Chemicals should be reserved for proteomic studies 

and should not be manipulated with spatula. Buffers should be filtered on 0.22 

μm pore size filters. Glass plates for electrophoresis should be cleaned with 

alcohol before use. 

2.1. Extraction of Labile or Weakly Bound CWP by Nondestructive 

Techniques 

2.1.1. CWP extraction and Analysis from Liquid Culture Medium 

of Seedlings 

1. Murashige and Skoog (MS) culture medium: Murashige and Skoog (14) liquid 

medium (Sigma Chemical, St Louis, MO, USA) is supplemented with 10 g/L 

sucrose and adjusted to pH 5.8 with KOH. 

2. PVPP (Sigma, St. Louis, MO, USA) is treated with acid to increase polymerization 

and to remove metal ions and contaminants. One g PVPP in 10 mL 10%


HCl is boiled for 10 min, filtered through a G4 filter, and rinsed until neutral 

pH is reached. The residue is dehydrated with acetone and grinded in a mortar 

to obtain a fine powder (15). 

3. Low binding 12 kDa cutoff Spectra/Por ® cellulose ester (CE) dialysis bags 

(Merck Eurolab Poly Labo, Strasbourg, France). 

4. Centriprep ® device (MWCO: 10 kDa) (Millipore Corporation, Bedford, MA, 

USA). 

5. Bradford protein assay (Coomassie ® Protein assay Reagent Kit, Pierce, Rockford, 

IL, USA) (16). 

6. Immobilized pH gradient (IPG) buffers and 13 cm-strips pH 4 –7 or 6–11 (GE 

Healthcare Europe GmbH, Orsay, France). 

7. 2-DE (2-dimensional electrophoresis) sample buffer: 7M urea, 2M thiourea, 

4% (w/v) CHAPS, 65 mM DTE, 0.5% (v/v) IPG buffer (pH 4-7 or 6-11), 

bromophenol blue trace. 

2.1.2. CWP Extraction and Analysis from Cell Suspension Cultures 

1. Gamborg liquid medium: Gamborg B5 medium supplemented with 20 g/L 

sucrose, 2.5 μM naphthalene acetic acid (Sigma Chemical, St Louis, MO, USA) 

and adjusted to pH 5.7 with KOH (17). 

2. Cell washing buffer: 50 mM sodium acetate buffer pH 6.5, 10 mM DTT, 1 mM 

PMSF, 1% ethanol, 50% glycerol. 

3. C1 protein extraction buffers: 0.15M NaCl in cell washing buffer. 

4. C2 protein extraction buffer: 1M NaCl in cell washing buffer. 

5. C3 protein extraction buffer: 0.2M CaCl2 dihydrate in cell washing buffer. 

6. C4 protein extraction buffer: 2M LiCl in cell washing buffer. 

7. C5 protein extraction buffer: 50 mM 1,2-cyclohexanediamine tetracetic acid 

(CDTA) in cell washing buffer. 

8. Low binding 2 kDa cutoff Spectra/Por ® CE dialysis bags (Merck Eurolab Poly 

Labo, Strasbourg, France). 

9. One mL Hi-Trap SP Sepharose cation exchanger (GE Healthcare Europe GmbH, 

Orsay, France). 

10. Hi-Trap SP equilibration buffer: 10 mM MES-KOH, pH 5.2. 

11. Hi-Trap SP elution buffer: 10 mM MES-KOH, pH 5.2, 2M NaCl. 

12. Econo-Pac ® 10DG desalting column (Bio-Rad, Hercules, CA,USA). 

13. Desalting column equilibration buffer: 50 mM ammonium formate. 

14. Bradford protein assay (Coomassie ® Protein assay Reagent Kit, Pierce, 

Rockford, IL, USA) (16). 

15. 1-DE (1-dimensional electrophoresis) sample buffer: 62 mM Tris-HCl, pH 6.8, 

2% SDS, 10% glycerol, 5% mercapto-ethanol. 

16. Resuspending solution: 1M thiourea, 10 mM DTT, 1% (v/v) protease inhibitor 

cocktail for plant (Sigma, St. Louis, MO, USA) in UHQ water (see Note 1). 

Prepare as required. 

17. Immobilized pH gradient (IPG) buffers and 13-cm strips pH 4–7 or 6–11 (GE 

Healthcare Europe GmbH, Orsay, France).


18. 2-DE sample buffer: 7M urea, 2M thiourea, 4% (w/v) CHAPS, 65 mM DTE, 

0.5% (v/v) IPG buffer (pH 4–7 or 6–11), bromophenol blue trace. 

2.1.3. Extraction and Analysis of CWP from Rosette Leaves 

2.1.3.1. Extraction of Proteins 

1. Recovering solution: 0.3M mannitol, 66 mM DTT, 330 mM thiourea, 3.3% (v/v) 

protease inhibitor cocktail for plant (Sigma, St. Louis, MO, USA) (see Note 1). 

Prepare as required. 

2. R1 protein extraction buffer: 1M NaCl, in recovering solution. Adjust pH to 6.9 

with 0.5N NaOH. Prepare as required 

3. R2 protein extraction buffer: 0.2M CaCl2 dihydrate, in recovering solution. Adjust 

pH to 6.9 with 0.5N NaOH. Prepare as required. 

4. R3 protein extraction buffer: 2M LiCl, 0.3 in recovering solution. Adjust pH to 

6.9 with 1N NaOH. Prepare as required. 

5. R4 protein extraction buffer: 50 mM CDTA, in recovering solution. Adjust pH to 

6.9 with 5N NaOH. Prepare as required. 

6. Malate dehydrogenase (MDH) assay mixture: 50 mM Tris-HCl, pH 7.8 (2.15 

mL), 50 mM MgCl2 hexa-hydrate (300 μL), 150 mM DTT (100 μL), 10 mM 

NADP (150 μL), 30 mM malic acid (300 μL). Store DTT and NADP solutions in 

single use aliquots at –20°C. Store malic acid solution at –20°C no longer than 

one month. Prepare the MDH assay mixture as required 

2.1.3.2. Analysis of Labile CWP 


Labo, Strasbourg, France). Store at –20°C. 

2. Resuspending solution: 1M thiourea, 10 mM DTT, protease inhibitor cocktail (1% 

v/v) in UHQ water. Prepare as required. 

3. Immobilized pH gradient (IPG) buffers and 7-cm strips pH 4–7 (GE Healthcare 

Europe GmbH, Orsay, France). 

4. 2 DE-sample buffer: 7M urea, 2M thiourea, 4% (w/v) CHAPS, 65 mM DTE, 


2.1.3.3. Analysis of Weakly Bound CWP 


Labo, Strasbourg, France). Store at –20°C. 

2. One mL Hi-Trap SP Sepharose cation exchanger (GE Healthcare Europe GmbH, 

Orsay, France). 

3. Hi-Trap SP equilibration buffer: 10 mM MES-KOH, pH 5.2. 

4. Hi-trap SP elution buffer: 10 mM MES-KOH, pH 5.2, 2M NaCl. 

5. Econo-Pac ® 10DG desalting column (Bio-Rad, Hercules, CA, USA). 


7. 1-DE sample buffer: 62 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, 5% 

mercapto-ethanol.


8. Resuspending solution: 1M thiourea, 10 mM DTT, 1% (v/v) protease inhibitor 

cocktail in UHQ water. Prepare as required. 

9. Immobilized pH gradient (IPG) buffers and 7-cm strips pH 4-7 (GE Healthcare 

Europe GmbH, Orsay, France). 

10. 2 DE-sample buffer: 7M urea, 2M thiourea, 4% (w/v) CHAPS, 65 mM DTE, 


2.2. Extraction of Weakly Bound CWP by Destructive Techniques 

2.2.1. Cell wall preparation 

1. MS solid medium: Murashige and Skoog (14) liquid medium (Sigma Chemical, 

St Louis, MO, USA) is supplemented with 20 g/L sucrose and 12 g/L agar, and 

adjusted to pH 5.8 with KOH. 

2. PVPP (Sigma, St. Louis, MO, USA) is treated with acid to increase polymerization 

and to remove metal ions and contaminants. One g PVPP in 10 mL 10% HCl 

is boiled for 10 min, filtered through a G4 filter, and rinsed until neutral pH is 

reached. The residue is dehydrated with acetone and grinded in a mortar to obtain 

a fine powder (15). 

3. Nylon nets (1.5-mm pore size and 25-μm pore size). 

4. Waring blender with a 2-L flask (SEB Moulinex, Ecully, France). 

5. Grinding buffer: 10 mM acetate buffer, pH 4.6, 0.4M sucrose, 0.2 % (v/v) protease 

inhibitor cocktail for plant (Sigma, St. Louis, MO, USA) (see Note 1). 

6. Cell wall purification buffers: 5 mM acetate buffer, pH 4.6, 0.6M or 1M sucrose, 

0.2% (v/v) protease inhibitor cocktail. 

7. Cell wall washing buffer: 5 mM acetate buffer, pH 4.6. 

2.2.2. Extraction and Separation of Proteins 

1. H1 protein extraction buffer: 5 mM acetate buffer, pH 4.6, 0.2M CaCl 2, 0.1% 

protease inhibitor cocktail for plant (Sigma, St. Louis, MO, USA). 

2. H2 protein extraction buffer: 5 mM acetate buffer, pH 4.6, 2M LiCl, 0.1% protease 

inhibitor cocktail. 

3. Econo-Pac ® 10DG desalting column (Bio-Rad, Hercules, CA,USA). 


5. Bradford protein assay (Coomassie ® Protein assay Reagent Kit, Pierce, Rockford, 

IL, USA) (16). 

6. 1-DE sample buffer: 62 mM Tris pH 6.8 (HCl), 2% SDS, 10% glycerol, 5% 

mercapto-ethanol. 

3. Methods 

The choice of a protocol to extract CWP for proteomic analysis is dependent 

on the plant material and of the type of proteins to be released from cell walls. 

Working on living cells is probably the best solution to avoid intracellular


contamination. This is possible for cell suspension cultures or seedlings grown 

in liquid medium as well as for any plant organ that can be infiltrated under 

vacuum with extraction buffers. Both labile and weakly bound CWP can be 

released. When this is not possible, it is necessary to purify cell walls. The main 

problem is to avoid intracellular contaminants that will stick nonspecifically to 

cell walls. Only weakly bound CWP can be extracted from purified cell walls 

because labile CWP are lost during cell wall preparation. 

Another important point is the choice of the extraction solution. For example, 

a solution of 0.3M mannitol infiltrated in living tissues such as leaves can 

solubilize a few CWP expected to be located only in intercellular spaces. Indeed, 

identified proteins are acidic, suggesting no ionic interactions with negatively 

charged cell wall components (9). NaCl is usually used for extraction of proteins 

retained by ionic interactions in the cell wall. LiCl can extract hydroxyprolinerich 

glycoproteins from intact cells in Chlamydomonas reinhardii (18). Calcium 

chloride is probably the most efficient salt to extract CWP (9,19). The ability 

of acidic and neutral carbohydrates to strongly chelate calcium (20,21) might 

explain, through a competition mechanism, that proteins or glycoproteins 

weakly bound to cell wall polysaccharides can be selectively solubilized by 

CaCl 2. CDTA, a chelating agent, solubilizes Ca 2+ -pectate. It releases a small 

number of proteins having domains of interaction with polysaccharides, notably 

proteins showing homology to lectins. This suggests an interaction of these 

proteins with polysaccharides associated to pectins (9). 

3.1. Extraction of Labile or Weakly Bound CWP by Nondestructive 

techniques 

3.1.1. Liquid Culture Medium of Seedlings 

1. Soak A. thaliana ecotype Columbia seeds (see Note 2) in tap water for 2 h, then 

sterilize in diluted bleach (2.4% w/v) for 45 min, and rinse several times with 

deionized water. 

2. Sterilized seeds (100 mg) are germinated and grown in MS liquid culture medium 

in ten 1-L flasks on a rotary shaker (90 rpm) at 26°C in the dark (22). Each flask 

contains 130 mL of culture medium. After 14 d, etiolated seedlings are harvested 

and the culture medium is filtered through nylon net (60 μm) to remove cell 

debris. 

3. Collect 900 mL of culture medium (see Note 3). Mix with 9 g PVPP. Shake the 

mixture at 4°C for at least 30 min, filter and centrifuge to pellet the insoluble 

residue. Dialyze against 10 L distilled water during 10–12 h at 4°C using a dialysis 

bag (MWCO: 12 kDa) with three changes. Reduce the volume of the sample by 

repeated centrifugations (3,500g for 15 min at 4°C) through a Centriprep ® device 

(MWCO: 10 kDa) to about 1 mL. 

4. Quantify proteins using the Bradford protein assay.


5. Dilute 250 μg proteins extracted from culture medium with 250 μL 2-DE sample 

buffer. Proteins are separated by 2-DE using 13 cm- IPG gel strips pH 4–7 or 

6–11 for the first dimension. 

3.1.2. Cell Suspension Cultures 

1. A cell suspension culture of A. thaliana ecotype Columbia is grown on Gamborg 

liquid medium. From this culture, 50 mL (25 g) is transferred every 2 wk to 250 

mL fresh medium in 1-L Erlenmeyer flask and shaken at 70 rpm in an orbital 

shaker, under continuous light (30 μE.m −2 .s −1 ) at 22°C. 

2. Wash cells of 7-day-old A. thaliana suspension culture with water and pellet 

them by centrifugation at 200g. Plasmolyze by successive immersion in 25 % 

glycerol, 50 % glycerol for 10 min each, and finally wash in 50 % cold glycerol. 

All subsequent extractions are performed at 0°C except otherwise stated. 

3. Before protein extraction, wash cells with cell washing buffer to remove contaminant 

proteins coming from broken cells that nonspecifically stick to cell walls. 

4. Extract proteins by washings of the plasmolyzed cells under gentle stirring (30 

min) in the proportion of 25 mL of pelleted cells per 50 mL of solution. First 

extraction is performed with C1 buffer. 

5. Wash cells with the same extraction buffer, then with 50 % glycerol before 

centrifugation at 200g for 5 min. 

6. Extract weakly bound CWP with C2, C3, C4, or C5 buffer in the same way (see 

Note 4). 

7. Exhaustively dialyze protein extracts at 4°C against 20 L H 2O using a dialysis bag 

(MWCO: 2 kDa). Measure the protein content of each extract using the Bradford 

protein assay. 

8. Dialyze against Hi-Trap SP equilibration buffer and apply to a Hi-Trap SP 

Sepharose column equilibrated with Hi-Trap SP equilibration buffer at 1 mL/min. 

Elute the retained basic proteins with Hi-Trap SP elution buffer at 1 mL/min. 

Desalt the basic proteins on an Econo-Pac ® 10DG desalting column equilibrated 

with desalting column equilibration buffer. Freeze-dry the eluate. Resuspend the 

dry residue in 40 μL 1-DE sample buffer and separate proteins by 1-DE on a 

10–17% gradient polyacrylamide gel (16.5 × 13.5 × 0.15 cm). 

9. Freeze-dry the acidic and neutral proteins from the Hi-Trap SP Sepharose column 

effluent. Solubilize the dry residue with a minimal volume of resuspending 

solution and desalt on an Econo-Pac ® 10DG desalting column. Freeze-dry the 

proteins, dissolve in 2-DE sample buffer and perform a 2-DE using a 7 cm-IPG 

gel strip pH 4–7 for the first dimension. 

3.1.3. Rosette Leaves 

3.1.3.1. Extraction of Proteins 

1. Sterilize A. thaliana ecotype Columbia seeds by soaking in diluted bleach (2.6% 

w/v) for 5 min and rinse several times with deionized water. Sow the seeds on


humid compost in 10 × 10 cm pots and cover the pots with a plastic film. Remove 

the plastic film after 48 h and transfer the cultures in a growth chamber at 70% 

humidity, with a photoperiod of 9 h light at 110 μE.m −2 s −1 at 22°C, and 15 h 

dark at 20°C. Plants should be moderately watered with a nutrient solution once 

a week. 

2. Remove carefully 4- to 5-week-old plants (Fig. 2A) from the pots and wash 

compost off with deionized water. Cotyledons and yellowish leaves should be 

systematically removed from plants. Process whole plants for vacuum-infiltration 

as follows. Make a small noose with a piece of string and pass the root through 

the noose. Tighten the noose around the collar then twist the root around the 

string and wrap in parafilm. In a large beaker, immerse completely the rosettes 

first in distilled water for a few seconds in recovering solution. Put the beaker 

with the immersed rosettes in a dessicator connected to a vacuum pump (Fig. 2B). 

Vacuum-infiltrate the rosettes for 2 min after starting the pump. Reintroduce 

carefully air in the dessicator after vacuum breakage (Fig. 2C). Transfer the 

infiltrated plants to a centrifuge tube, with the collar at about 1 cm at the edge 

of the tube (Fig. 2D). Paste the lower part of the root outside of the tube with 

adhesive tape. Introduce at the bottom of the centrifuge tube 300 μL of recovering 

solution. Centrifuge infiltrated plants in swinging buckets at 200g for 17 min at 

Fig. 2. Vacuum-infiltration of rosette leaves. Four steps of the procedure are illustrated: 

(A) 4–5 wk-old plants; (B) vacuum-infiltration of immersed rosettes in a dessicator 

connected to a vacuum pump; (C) rosette leaves after vacuum-infiltration, note 

the darker part of a leaf after successful infiltration (black arrow); (D) infiltrated plant 

transferred to a centrifuge tube, note the drop of solution containing the protease 

inhibitor cocktail at the bottom of the tube (black arrow).


20°C (see Note 5). Collect the apoplastic washing fluids with a micropipet and 

estimate the volume. Vacuum-infiltration and centrifugation should be repeated 

twice. 

3. Assay the apoplastic fluids for malate dehydrogenase (MDH) activity to detect 

cytoplasmic contaminations. Measure MDH activity at room temperature in 3 mL 

MDH assay mixture and one-twentieth of the volume of the recovered apoplastic 

fluids. Reduction of NADP is followed at = 340 nm. Pool only those apoplastic 

washing fluids with no detectable MDH activity. 

4. Vacuum-infiltrate rosettes with R1, R2, R3 or R4 buffer. Check for MDH activity 

on the recovered apoplastic fluids as described above. Discard any apoplastic 

washing fluids with MDH activity. Pool the remaining apoplastic washing fluids 

free of MDH activity. 

3.1.3.2. Analysis of Labile CWP by 2-DE 

1. Exhaustively dialyze the apoplastic washing fluids from rosettes infiltrated with 

the recovering solution at 4°C against deonized water in low binding 2 kDa 

cutoff Spectra/Por ® CE dialysis bags. Freeze-dry the dialysates. Resuspend the 

dry residues in 3 mL of resuspending solution and desalt on an Econo-Pac ® 10DG 

desalting column equilibrated with desalting column equilibration buffer for the 

complete removal of mannitol. Freeze-dry the eluate. 

2. Solubilize the dry residue in 2-DE sample buffer and separate proteins by 2-DE 

using a 7 cm-IPG gel strip pH 4–7 for the first dimension. 

3.1.3.3. Analysis of Weakly Bound CWP 

1. Exhaustively dialyze the apoplastic washing fluids from rosettes infiltrated with 

R1, R2, R3, or R4 buffer against Hi-Trap SP equilibration buffer as described 

above. Apply to a Hi-Trap SP Sepharose column equilibrated with Hi-Trap SP 

equilibration buffer at 1 mL/min. Elute the retained basic proteins with Hi-Trap 

SP elution buffer at 1 mL/min. Desalt the basic proteins on an Econo-Pac ® 10DG 

desalting column equilibrated with desalting column equilibration buffer. Freezedry 

the eluate. Resuspend the dry residue in 40 μL 1-DE sample buffer and 

separate proteins by 1-DE on a 10–17% gradient SDS-polyacrylamide gel (16.5 

× 13.5 × 0.15 cm). 

2. Freeze-dry the acidic and neutral proteins in the Hi-Trap SP Sepharose column 

effluent. Solubilize the dry residue with a minimal volume of resuspending 

solution and desalt on an Econo-Pac ® 10DG desalting column equilibrated with 

desalting column equilibration buffer. Freeze-dry the proteins and perform a 2-DE. 

3.2. Extraction of Weakly Bound CWP by Destructive Techniques 

3.2.1. Cell Wall Preparation 

1. Soak A. thaliana ecotype Columbia seeds (see Note 2) in tap water for 2 h, 

then sterilize in diluted bleach (2.4 %) for 45 min, and rinse several times with 

deionized water. Sow the seeds (150 mg) in a Magenta box (6 × 6 cm) containing


50 mL of solid MS medium. Grow seedlings at 23°C in the dark for 11 d (see 

Note 6). 

2. Harvest hypocotyls (around 2 cm high) of an average of 20 Magenta boxes as 

follows. First, remove carefully the solid MS medium carrying the seedlings from 

each box. Then, cut hypocotyls below cotyledons and above root with a pair of 

scissors. Wash the 1-cm-long hypocotyls with distilled water onto a nylon net 

(1.5-mm pore size) to remove all the cut cotyledons and seed coats that stick 

to hypocotyls (see Note 7). Transfer the hypocotyls into 500 mL of grinding 

buffer and add PVPP (1g/10g fresh weight of hypocotyls) to complex phenolic 

compounds. 

3. Grind the mixture in cold room using a Waring blender at full speed for 15 min. 

4. Separate cell walls from soluble cytoplasmic fluid by centrifugation of the 

homogenate for 15 min at 1,000g and 4°C. Further purify the pellet by two 

successive centrifugations in 500 mL of cell wall purification buffers, 0.6M and 

1M sucrose respectively. 

5. Wash the residue with 3 L of cell wall washing buffer on a nylon net (25-μm pore 

size) to eliminate all soluble compounds. Grind the resulting cell wall fraction in 

liquid nitrogen in a mortar with a pestle before lyophilization. Starting with 16 g 

fresh weight of hypocotyls, this procedure usually results in 1.3 g dry powder. 

3.2.2. Extraction of Proteins 

1. Typically, 0.65 g of lyophilized cell walls is used for one experiment. Extract 

proteins by successive salt solutions in this order: two extractions with 6 mL H1 

buffer, followed by two extractions with 6 mL H2 buffer. Resuspend cell walls 

by vortexing for 5-10 min at room temperature, and then centrifuge for 15 min 

at 4,000g and 4°C. Supernatants from the same extracting buffer are pooled. 

2. Desalt supernatants using Econo-Pac ® 10DG desalting columns equilibrated with 

desalting column equilibration buffer. Lyophilize the extracts and resuspend in 

H2O2. 3. Quantify proteins using the Bradford protein assay. 

4. Add 1-DE sample buffer. Separate proteins by 1-DE on a 12.5% SDSpolyacrylamide 

gel. 

3.3. Analysis of CWP 

3.3.1. Specific Constraints for Separation by Electrophoresis and 

Protein Identification by Mass Spectrometry 

The separation of CWP by classical two-dimensional gel electrophoresis 

(2-DE) is difficult. Because most CWP are basic glycoproteins (Fig. 1C), they 

are poorly resolved by this technique (22). They are better separated by 1-DE. 

However, protocols including chromatographic steps to separate proteins before 

1-DE are available (24,25). In this chapter, a method able to separate acidic


and basic CWP is proposed for a better resolution of these two types of CWP 

in 2-DE or 1-DE respectively. 

Frequently, in 1-DE, proteins are not well separated from one 

another, and a protein sample can contain a mixture of proteins. 

However, the peptide mass mapping technology using high resolution 

(< 20 ppm) MALDI-TOF mass spectrometry (MS) permits the identification 

of several proteins from a mixture. Search engines such as MS- 

FIT from Protein Prospector (http://prospector.ucsf.edu/ucsfhtml4.0/msfit.htm) 

or MASCOT (http://www.matrixscience.com/search_form_select.html) allow 

multiple searches. In case of difficulties, proteins can also be identified by 

peptide sequencing using LC (liquid chromatography)-MS/MS systems (9). 

3.3.2. Use of Bioinformatics for the Evaluation of the Efficiency 

of an Extraction Protocol 

The reliability of protein profiling for a compartment like the cell wall, 

strongly depends on the quality of the preparation. Unfortunately, the classical 

methods to check the purity of a particular fraction are not conclusive 

for proteomic studies, because the sensibility of the analysis by mass 

spectrometry is 10–1,000 times more sensitive than enzymatic or immunological 

tests using specific markers. Our experience in the field has shown 

that the most efficient way to evaluate the quality of a cell wall preparation 

is (1) to identify all the proteins extracted from the cell wall by 

mass spectrometry, and (2) to perform extensive bioinformatic analysis to 

determine if the identified proteins contain a signal peptide, and no retention 

signals for other cell compartments. Comparison of the results obtained 

with different programs is necessary to ensure a reliable prediction: PSORT 

allows predicting any subcellular localization (http://psort.ims.u-tokyo.ac.jp/ 

form.html); TargetP looks for the presence of signal peptides for protein 

secretion or of transit peptides for mitochondrion or chloroplast targeting 

(http://www.cbs.dtu.dk/services/TargetP/); Aramemnon compares the results of 

several programs predicting the presence of signal peptides and transmembrane 

domains (http://aramemnon.botanik.uni-koeln.de/). It is then possible to 

conclude about the quality of the cell wall preparation by calculating the ratio 

of predicted secreted proteins to intracellular ones. 

4. Notes 

1. Protease inhibitor cocktail for plant is required to prevent proteolysis during 

the extraction procedure. Proteolysis induces the production of smaller broken 

proteins that can be spread over 1-D or 2-D polyacrylamide gels. Thus, proteolysis 

can prevent the identification of both broken proteins and other proteins of interest


by mass spectrometry. Moreover, the occurrence of these polypeptides is a great 

problem for quantitative and comparative proteomics. 

2. Seeds germinate in culture media that are favourable to development of bacteria 

or fungi. Because of the high amount of seeds (150 mg) introduced in a culture 

flask or in a Magenta box, the possible contamination events are multiplied. So, 

seeds should be carefully sterilized, and the healthy state of plants should be very 

good during their production in greenhouses. 

3. Culture media should be processed immediately after recovery. Otherwise, proteolysis 

may occur even if they are stored frozen. 

4. No more than two successive extractions with salt solutions should be performed. 

Otherwise, cells are damaged and intracellular contaminants are released in the 

culture medium and can stick non-specifically to cell walls (10). 

5. Be careful setting minimal acceleration to avoid seriously damaging the vacuuminfiltrated 

plants during the centrifugation step. Imperatively centrifuge in 

swinging buckets to get undamaged plants during spinning. 

6. All seedlings should grow at about the same rate to reach the same size after 11 

d. If germination is not homogeneous, place the boxes at 6°C during 2dtoallow 

all seeds to start germination without growth. Then, all boxes can be put at 23°C 

for 11 d. 

7. Cotyledons should be carefully removed. They contain few protein species but 

each of them in a huge amount. Because of their density, cotyledons co-sediment 

with cell walls. As a consequence, few cotyledons induce a significant contamination 

during extraction of cell wall proteins, especially by storage proteins. 

This contamination prevents the identification of proteins of interest by mass 

spectrometry. 


The authors are grateful to the Université Paul Sabatier (Toulouse III, France) 

and the CNRS for support. 

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1125–1135.

18 

Isolation and Fractionation of the Endoplasmic 

Reticulum from Castor Bean (Ricinus communis) 

Endosperm for Proteomic Analyses 

William J. Simon, Daniel J. Maltman, and Antoni R. Slabas 

Summary 

This chapter describes the preparation and isolation of highly purified endoplasmic 

reticulum (ER) from the endosperm of developing and germinating castor bean (Ricinus 

communis) seeds to provide a purified organelle fraction for differential proteomic 

analyses. The method uses a two-step ultracentrifugation protocol first described by 

Coughlan (1) and uses sucrose density gradients and a sucrose flotation step to yield 

purified ER devoid of other contaminating endomembrane material. Using a combination 

of one dimensional (1D) and two dimensional (2D) gel electrophoresis the complexity 

and reproducibility of the protein profile of the purified organelle is evaluated prior to 

detailed proteomic analyses using mass spectrometry based techniques. 

Key Words: Castor bean; electrophoresis; endoplasmic reticulum; proteomics; 

Ricinus communis. 


The endoplasmic reticulum (ER) is a specialized endomembrane system 

within all cells responsible for a number of important biological processes 

including, protein folding, protein sorting, and secretion (2), and protein Nglycosylation 

(3). In the seeds of higher plants the ER is the processing site for 

the synthesis of storage proteins (4) and is also the site for fatty acid modification 

(5–7), triacyglycerol (TAG) biosynthesis (8), complex lipid biosynthesis 

and the primary site for membrane biogenesis. 



203

204 Simon et al. 

Understanding the processes of TAG biosynthesis and complex lipid partitioning, 

particularly in the major commercially grown oil producing plant 

species, is an important step towards maximizing both oil quality and yield 

from these crops. This is particularly so if advances in molecular biology are 

to be used to rationally design new crops using transgenic technologies. 

Although the plant ER plays a central role in metabolism there have been 

a limited number of studies on the organelle due to the difficulties associated 

with its isolation and purification. 

In order to overcome this we have taken advantage of the liquid properties 

of castor bean endosperm to prepare preparative amounts of ER using modifications 

of the method first described by Beevers (9). 

The ER of developing seeds is the major site of synthesis of both lipid and 

storage proteins – such biosynthetic capacity is lost during germination. In order 

to elucidate the changes in the ER during seed development and germination we 

have prepared material from both to allow for a detailed differential proteomic 

analysis of the two tissues (10). 

2. Materials 

2.1. Tissue Homogenization and ER Purification 

1. 5% Hypochlorite solution in water (see Note 1) for seed surface sterilization. 

2. Homogenization medium: 500 mM sucrose, 10 mM KCl, 1 mM EDTA, 1 mM 

MgCl2 2 mM dithiothreitol (DTT), 0.1 mM phenylmethyl-sulfonyl fluoride 

(PMSF), 150 mM Tricine-KOH pH 7.5. Keep at 4°C on ice. 

3. Sucrose solutions for gradient formation: prepare fresh in sterile water 50 mL of 

each of the following, 20%, 30%, 40%, and 60% sucrose each containing 1 mM 

EDTA and 0.1 mM PMSF. Keep at 4°C on ice. 

4. 10% glycerol solution for –80°C storage of purified ER fractions. 

5. The following Beckman centrifuges, rotors and centrifuge tubes: 

Avanti 30 centrifuge, L-70, and Optima TLX ultra-centrifuges, F0850, SW28, 

SW41, and TL100.4 rotors, 326823 and 344061 ultra-clear tubes and 343776 

micro tubes. 

2.2. Protein Estimations 

1. Protein standard: Bovine serum albumin (BSA) 1.0 mg/mL in water. 

2. Protein assay kit (modified Bradford) from BIO-RAD. 

2.3. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis 

of Purified ER 

The SDS-PAGE method described is a modification of that originally 

described by Laemmli (11). All reagents are from BIO-RAD.

Isolation and Fractionation of the Endoplasmic Reticulum 205 

1. SDS-PAGE resolving gel (10%): 3.33 mL Acrylamide-Bis solution (37.5:1), 2.5 

mL 1.5M Tris-HCl pH 8.8, 100 μL of 10% SDS, 100 μL of 10% ammonium 

persulphate, 10 μL of TEMED in a final volume of 10 mL water. 

2. SDS-PAGE stacking gel (4%): 1.3 mL Acrylamide-Bis solution (37.5:1), 2.5 mL 

0.5M Tris-HCl pH 6.8, 100 μL of 10% SDS, 100 μL of 10% ammonium persulphate, 

10 μL of TEMED (added last once all other constituents are mixed) in a 

final volume of 10 mL water. 

3. Water saturated isobutanol solution: Mix together equal volumes of water and 

isobutanol in a glass bottle and shake vigorously to mix. Allow to settle and use 

the top layer. 

4. Gel running buffer: 25 mM Tris-HCl pH 8.3, 250 mM glycine, 0.1% SDS (see 

Note 2). 

5. Sample loading buffer: 50 mM Tris-HCl pH 6.8, 100mM DTT, 2% SDS, 0.1% 

bromophenol blue, 10% glycerol (see Note 3). 

6. Dalton Mark VII molecular weight markers (Sigma) made up in sample loading 

buffer as the manufacturers instructions. 

2.4. Mass Spectrometer Compatible Silver Staining 

Prepare all reagents fresh immediately before use. 

1. Fixing solution: 10% acetic acid, 40% methanol. 

2. Sensitizing solution: 75 mL methanol, 10 mL sodium-thiosulphate (5 %), 17 g 

sodium-acetate in 250 mL water. 

3. Washing solution: water. 

4. Silver solution: 0.25% silver nitrate in water. 

5. Developing solution: 6.25 g sodium carbonate, 100 μL formaldehyde in 250mL 

water. 

6. Stop solution: 3.65 g ethylenediaminetetracetic acid (EDTA) in 250 mL water. 

2.5. Mini 2 Dimensional Gels (2D gels) 

1. The method described in this text uses the Multiphor II electrophoresis unit 

combined with the Immobiline Dry Strip (IPG) kit and the Immobiline Dry 

Strip Reswelling Tray from GE Healthcare for the first dimension isoelectric 

focusing (IEF) step of 2D electrophoresis. The method should be adaptable to 

IEF kit from any manufacturer that uses IPG strips, however focusing parameters 

may need to optimized. For a useful guide to 2D electrophoresis see (12). 

2. Immobiline Dry Strip 7 cm ready made IPG strips pH 3–10 from GE Health care. 

3. Lysis-rehydration buffer: 9M urea, 2M thiourea, 4% CHAPS (3-[(3 

cholamidopropyl)dimethylammonio]-1-propanesulfonate), 1% DTT, 2% carrier 

ampholytes pH 3–10 (GE Healthcare). Buffer is made up without DTT and 

carrier ampholytes, aliquoted and stored at –80°C. Once thawed DTT and carrier 

ampholytes are added immediately before use.


4. Equilibration buffer 1: 50 mM Tris-HCl pH 8.8, 6M urea, 30% glycerol, 10% 

SDS, 1% DTT. Prepare 10mL. 

5. Equilibration buffer 2: 50 mM Tris-HCl pH 8.8, 6M urea, 30% glycerol, 10% 

SDS, 4.8% iodoacetamide, 0.01% bromophenol blue. Prepare 10 mL. 

6. Low melting point (LMP) agarose solution: 5% made up in SDS-PAGE running 

buffer. Prepare 20 mL for each mini 2D gels being run by heating the solution to 

100°C immediately before use. 

7. SDS-PAGE gel reagents as described in Section 2.3. 

8. Mass spectrometer compatible silver staining reagents as described in Section 2.4. 

3. Methods 

3.1. Isolation of ER Membranes from Germinating and Developing 

Castor Bean Endosperm 

1. In order to make a proteomic comparison between the ER from germinating and 

developing castor bean, material must be prepared from the seeds at both growth 

stages. For the germinating sample mature seeds are first surface sterilized for 

1.0 min in 5% hypochlorite solution and then washed overnight in running tap 

water. The soaked seeds are then sown in moist vermiculite and germinated in the 

dark at 30°C for 3–4 d before dissecting out the endosperm. For the developing 

material seedpods are harvested from castor plants 25 d after flowering (see 

Note 4) and individual seeds are removed for processing. 

2. Take 40–50 seeds (see Note 5) from each of the germinating or developing 

samples and carefully dissect out the endosperm and remove the embryo and 

cotyledons. Place the endosperm halves into a large glass Petri dish containing 

10 mL ice-cold homogenization buffer and manually chop them for 10 min 

on ice using two single-sided razor blades (see Note 6). Carry out all further 

procedures at 4°C unless otherwise stated. 

3. Filter the crude homogenate through a 100-μm nylon mesh to remove large debris 

and transfer the filtrate to 50-mL ultra-clear centrifuge tubes and centrifuge in a 

Avanti 30 bench-top centrifuge using a F0850 rotor at 1,000g at 4°C for 15 min. 

4. During this centrifugation step prepare in two Beckman Ultraclear 25 × 89-mm 

centrifuge tubes, discontinuous sucrose density gradients consisting of 7 mL of 

20% sucrose carefully layered on top of 13 mL of 30% sucrose. Both sucrose 

solutions contain 1 mM EDTA and 0.1 mM PMSF. 

5. With a glass rod carefully remove and discard the fat pad (see Note 7) which 

has formed on top of the supernatant during centrifugation of the homogenate 

and divide the supernatant into two halves each of which is carefully layered 

onto the top of the prepared sucrose gradients (step 4 above). 

6. Transfer the tubes to the buckets of the Beckman SW28 rotor and centrifuge at 

100,000g for2hat2°CinaBeckman L70 ultra-centrifuge. 

7. Following centrifugation mount the tubes vertically in a laboratory clamp 

positioned under a lamp and the membranes should be clearly visible as a 

distinct band at the 20-30% sucrose interface (Fig. 1). Without disturbing the


Fig. 1. Following centrifugation at 100,000g for 2 h the centrifuge tube is carefully 

removed from the rotor without disturbing the sucrose gradient and mounted in a 

laboratory clamp positioned under a lamp. This enables the membrane layer to be 

clearly seen (marked by the arrow) at the 20–30% sucrose interface (∼1.12 g/cm 3 

density). As shown a hypodermic needle is used to pierce the centrifuge tube wall just 

below the membrane layer and the membranes are removed using a syringe. 

sucrose gradient carefully pierce the wall of the tube with a hypodermic needle 

immediately below the interface and using a syringe remove the membrane layer 

(Fig. 1). 

8. Pool the membrane fractions from both tubes and mix with an equal volume of 

ice cold 60% sucrose containing 1 mM EDTA and 0.1 mM PMSF. 

9. Pipet 4.0 mL of diluted membrane fraction into the bottom of Beckman Ultraclear 

14 × 89 mm centrifuge tubes and carefully overlay with 3 mL of 40% sucrose 

solution, 3 mL of 30% sucrose solution and 2 mL of 20% sucrose all containing 

1mM EDTA and 0.1 mM PMSF.


10. Transfer the tubes to the buckets of the Beckman SW41 rotor and centrifuge 

at 250,000g for 22 h at 2°C in a Beckman L70 ultra-centrifuge. During this 

centrifugation step the ER fraction will float through the dense sucrose layers 

to resolve as a highly purified fraction at the 20–30% sucrose interface (∼1.12 

g/cm 3 density. 

11. Carefully remove the purified ER from each tube as described in step 7 and 

pool together. Dilute with an equal volume of ice-cold water and aliquot into 

Beckman thick walled 500-μL polycarbonate centrifuge tubes. 

12. Transfer the tubes to the Beckman TL100.4 rotor and pellet the membranes 

by centrifugation in a Beckman Optima TLX ultracentrifuge at 250,000g for 

45 min. 

Pour off the supernatant and re-suspend the membrane pellet in a minimum 

volume of 10% (v/v) glycerol aliquot into tubes suitable for storage, snap freeze 

in liquid nitrogen and store at –80°C (see Note 8). 

3.2. Protein Estimation – Modified Bradford 

1. Using the 1.0 mg/mL BSA stock solution (Section 2.2.1) prepare a calibration 

curve as outlined in Table 1. Vortex solutions and incubate at room temperature 

for 15 min. 

2. In duplicate mix 2 μL of sample with 798 μL of water and 200 μL of Bradford 

reagent. Vortex solutions and incubate at room temperature for 15 min. 

3. Using a spectrophotometer measure the absorbance of the standards and samples 

at 595 nm. 

Plot the calibration curve and estimate the protein concentration of the 

samples from this curve. 

3.3. (SDS-PAGE) Analysis of Purified ER from Developing 

and Germinating Castor Bean Endosperm 

1. For SDS-PAGE analyses we use the mini Protean gel kit from BIO-RAD 

although any gel format may be used including commercially available 

precast gels. 

Table 1 

Bradford assay calibration curve standard dilutions. 

BSA Series (μg) 0 1 2 5 10 15 20 40 

1mg/mLBSA (μL) - 1 2 5 10 15 20 40 

H 2O (μL) 800 799 798 795 790 785 780 760 

Bradford reagent (μL) 200 200 200 200 200 200 200 200


2. Assemble the gel kit with spacers for 0.75 mm gels and prepare 10 mL of 

resolving gel solution as outlined in Section 2.3.1. Using a Pasteur pipet carefully 

pipet the gel solution between the plates avoiding air bubbles. Leave sufficient 

space between the top of the gel solution and the top of the glass plates to allow 

for the gel comb and approximately 1 cm of stacker gel. 

3. Overlay the gel solution with water-saturated isobutanol and allow the gel to 

polymerize at room temperature which should take around 30 min. 

4. Pour off the isobutanol and rinse the top of the gel thoroughly with several 

washes of water. 

5. Prepare 10 mL of stacking gel solution as described in Section 2.3.2 and using a 

Pasteur pipet carefully overlay this gel solution onto the top of the polymerized 

resolving gel. Insert the gel comb avoiding the formation of air bubbles and 

allow the stacking gel to polymerize, which should take around 30 min at room 

temperature. 

6. While the stacking gel is polymerizing prepare 800 mL of 1× gel running buffer 

as described in Section 2.3.4. 

7. Once the stacking gel has set carefully remove the gel comb and rinse out the 

wells with running buffer. Assemble the gel kit and fill the upper and lower 

buffer compartments with running buffer. 

8. For SDS-PAGE analyses of purified ER fractions using mass spectrometer 

compatible silver stain dissolve an aliquot of the purified preparation equivalent 

to between 10 and 20 μg of total protein in SDS-PAGE sample loading buffer 

(see Note 9), centrifuge for 5 min in a bench-top microfuge at maximum speed 

and load directly into the washed wells of the gel. 

9. Include at least one lane of SDS-VII molecular weight markers as described in 

Section 2.3.6. on every gel. 

10. Connect the electrophoresis tank to the power supply and run the electrophoresis 

at 100 volts until the bromophenol blue dye front passes through the stacker 

gel into the resolving gel. At this point increase the voltage to 200 volts and 

continue until the dye front reaches the end of the gel. 

11. Switch off the power supply, disassemble the gel kit and carefully transfer the 

gel to a clean polythene container for staining. 

3.4. Mass Spectrometer Compatible Silver Staining 

1. Mass spectrometric methods for proteomic analyses are highly sensitive 

techniques capable of detecting and identifying subfemtomole amounts of protein 

in a sample. For this reason contamination of samples particularly with human 

keratin can be a major problem during analyses. To avoid this as much as possible 

use freshly prepared staining reagents, handle gels as little as possible and use 

clean containers for all staining procedures. 

2. Following electrophoresis transfer the gel to a clean staining tray and fix the 

proteins in the gel by incubating the gel in 100 mL of fixing (Section 2.4.1) 

solution (see Note 10) for 15 min with gentle shaking (see Note 11) at room 

temperature. Repeat this step with a second 100 mL of fixing solution.


3. Discard the fixing solution and add 100 mL of sensitizing solution (Section 2.4.2). 

Incubate the gel for 30 min at room temperature with gentle shaking. 

4. Discard the sensitizing solution and wash the gel thoroughly with 3 × 5 min 

washes with fresh water. 

5. Pour off the last water wash, add 100 mL of silver solution (Section 2.4.4) and 

incubate the gel for 20 min at room temperature with gentle shaking. 

6. Discard the silver solution and wash the gel thoroughly with 3 × 1 min washes 

with fresh water. 

7. Pour off the last water wash, add 100 mL of developing solution (Section 2.4.5) 

and incubate the gel for 4 minutes at room temperature with gentle shaking. 

Monitor the intensity of the staining during this incubation period and if necessary 

lengthen or shorten the developing time to achieve bands of reasonable intensity 

without staining the gel background. 

8. Discard the developing solution, add 100 mL of stopping solution (Section 2.4.6) 

and incubate the gel for 10 min at room temperature with gentle shaking. 

9. Discard the stopping solution and wash the gel thoroughly with 3 × 5 min washes 

with fresh water. This staining procedure should result in discrete protein bands 

stained from brown to black, depending on protein concentration, against a clear 

gel background. A representative 1D-SDS PAGE gel showing three independent 

kDa 

66 

45 

36 

29 

24 

20 

14 

S 

Developing ER Germinating ER 

1 

2 

3 

Fig. 2. Protein estimations were made on each preparation using the modified 

Bradford procedure as described in Section 3.2. Protein samples (10 μg) were 

mixed with SDS-PAGE sample buffer vortexed at room temperature for 10 min and 

microfuged at maximum speed before loading into wells. Electrophoresis was carried 

out as described in Section 18.3.3. Lane S contains molecular weight markers. The 

protein profiles are highly reproducible between preparations and clear differences are 

seen between the developing and germinating ER samples on these gels. 

1 

2 

3


castor bean ER preparations from both developing and germinating endosperm 

stained using this protocol is shown in Fig. 2. 

3.5. Mini 2D Gel Protein Profiling of Purified ER from Developing 

and Germinating Castor Bean Endosperm 

1. For differential proteomic analyses of the developing and germinating ER a 

number of independent biological samples need to be prepared. This is to allow 

for the true biological differences between the two tissue types to be compared 

and to enable the elimination of technical artifacts such as gel to gel variability, 

sample solubilization and staining artifacts. The use of mini 2D gels allows 

the rapid and detailed evaluation of the reproducibility and complexity of the 

protein profiles of the purified ER fraction from independent preparations before 

beginning large-scale proteomic profiling experiments. Fig. 3 shows the mini 

2D gel profiles from three independent biological preparations of germinating 

and developing ER prepared using the protocols outlined in this text. 

2. To an aliquot of purified ER material equivalent to 50 μg of total protein from 

each preparation add 4 volumes of ice-cold absolute acetone and incubate on 

ice for 60 min. Pellet the proteins from this 80% acetone precipitation step by 

centrifugation for 10 min in a microfuge at maximum speed. 

3. Allow the pellet to air dry but do not over-dry. 

4. Add 125 μL (see Note 12) of 2D lysis-rehydration buffer (Section 2.5.1) to 

the pellet and gently disperse it using a pipetman fitted with a 100 μL pipet 

tip. Sonicate the sample for 10 min in a sonicating water bath and incubate on 

a vortex shaker at 30°C for 60 min. Centrifuge for 10 min in a microfuge at 

maximum speed. 

5. Using the reswelling tray rehydrate an IPG strip with the entire solubilized 

sample for a minimum of 6 h but ideally overnight (see Note 13). 

6. Assemble the Multiphor kit for isolelectric focusing. 

7. Using forceps remove the re-hydrated IPG strip from the reswelling tray, rinse 

with water and gently blot dry. 

8. Place the rehydrated IPG strip in the plastic insert of the IEF tray, position the 

electrodes and connect them to the power supply. 

9. Fill the IEF tray with mineral oil so that the gel strip is completely covered. 

10. Fit the lid to the Multiphor unit and begin isoelectric focusing (IEF). 

11. IEF is typically carried out in three phases or voltage steps (see Note 14). For 

7-cm IPG strips with a sample loading of 50 μg of purified ER fraction these 

steps are outlined in Table 2. 

12. While the IEF gel is running prepare an SDS-PAGE mini gel for the second 

dimension. For this gel no stacking gel is required and the gel should be prepared 

with 1-mm spacers to allow for the IPG strip to be positioned on the gel surface. 

13. Prepare 10 mL of resolving gel solution as described in Section 2.3.1. Using a 

Pasteur pipet carefully pipette the gel solution between the gel plates avoiding 

air bubbles until the solution is about 0.8 cm from the top of the lower glass 

plate.


Germinating Developing 

pH 3.0 pH 10 S pH 3.0 pH 10 S 

Fig. 3. ER was independently purified from developing and germinating castor bean 

in three separate experiments using the protocols described in the text. Molecular 

weight standards were loaded adjacent to the basic end of the IEF strip and are marked 

S. An aliquot equivalent to 50 μgoftotal protein from each preparation was evaluated 

on mini 2D gels (pH 3.0–10) and stained with mass spectrometer compatible silver 

stain. As can be clearly seen the independent preparations are reproducible and there 

are significant differences in the proteomic profiles between the two tissues. 

14. Overlay the gel solution with water-saturated isobutanol and allow the gel to 

polymerize at room temperature until the IEF is completed. 

15. Following IEF disconnect the gel kit from the power supply and disassemble 

the unit (see Note 15). 

16. Place the focused IPG strip into equilibration buffer 1 (Section 2.5.4) and 

incubate at room temperature for 15 min with gentle shaking. 

17. Transfer strip to equilibration buffer 2 (Section 2.5.5) and incubate at room 

temperature for 15 min with gentle shaking.


Table 2 

Isoelectric focusing parameters a 

step volts mA Watts Time (h) kVh 

1 200 2 5 0.01 

2 3500 2 5 1.30 2.8 

3 3500 2 5 1.00 2.2 

a Step 1 is a low voltage step which minimizes sample aggregation. Voltage is then gradually 

increased during step 2 to the final focusing voltage and held at this voltage during step 3 until 

focusing is complete. 

18. During this second equilibration step pour of the isobutanol solution from the 

polymerized second dimension gel, wash the gel surface extensively with water 

and blot dry. 

19. Prepare low melting point agarose solution as described in Section 2.5.6. 

20. Remove the gel strip from the equilibration solution and place it resting on 

its edge on a piece of moistened filter paper to allow any excess equilibration 

solution to drain away. 

21. Using forceps position the strip between the glass plates of the second dimension 

gel with its backing strip against one of the glass plates and gently push the 

IPG strip down until the entire lower edge is in contact with the SDS-PAGE 

gel surface. Ensure there are no air bubbles trapped between the IPG strip and 

the gel. 

22. If required SDS-VII molecular weight markers can be spotted onto a small piece 

of filter paper and positioned in contact with the SDS-PAGE gel surface at one 

end of the IPG strip. 

23. Overlay the IPG strip and with low melting point agarose solution that has 

cooled to between 40–50°C. 

24. Assemble the gel kit, connect it to the power supply and begin running the 2nd 

dimension electrophoresis at 60 volts for 10 min. This will allow the proteins 

to migrate from the IPG strip into the SDS-PAGE gel. Increase the voltage to 

150 volts and continue the electrophoresis until the bromophenol dye front has 

reached the end of the gel. 

25. Following electrophoresis stain the gels using the mass spectrometer compatible 

silver protocol described in Section 3.4. 

4. Notes 

1. Unless otherwise stated “Water” in this text refers to high purity water with a 

resistivity of 18 M-cm. 

2. SDS-PAGE running buffer can be made up as a 10× stock solution, stored at 

room temperature and diluted as required for use.


3. SDS-PAGE sample loading buffer can be conveniently made up for use as a 5× 

stock solution, divided into aliquots and stored at –20°C. 

4. In order to ensure that developing seeds are collected at the correct developmental 

stage plants are monitored daily during growth and flowers are tagged 

as they emerge. The seedpods are then allowed to develop and are harvested 

after 25 d. 

5. The protocol described here has been optimised to yield the maximum amount of 

purified ER devoid of contamination with other plant endomembrane material. 

Our attempts to scale up this procedure using higher seed numbers as starting 

material have only resulted in contaminated preparations. We recommend that 

for scale up use the protocol as described but increase the number of preparations. 

6. Carefully chopping the endosperm with single sided razor blades allows for 

successful tissue homogenisation without significant membrane disruption. 

7. During the low speed centrifugation step of the crude homogenate to remove 

cell debris a fat pad will form on top of the supernatant in the centrifuge tube. 

It is important that this fat pad is carefully removed without disruption before 

the supernatant containing the membrane fraction is loaded onto the sucrose 

gradient. 

8. The typical yield of purified ER from a Ricinus germinating or developing 

preparation is approx 1.0 mg. We usually re-suspend the final pellet of purified 

ER in 500 μL of 10% (v/v) glycerol and divide into 5× 100-μL aliquots for 

storage at –80°C. 

9. Samples for SDS-PAGE analyses are best prepared by adding a volume of 5 x 

sample buffer to an aliquot of sample equivalent to 10–20 μg of total protein 

and then diluting to 5 times the volume to give a sample ready for loading in 

1× sample buffer. 

10. For mass spectrometric compatible silver staining use 100mL of each solution 

for each mini gel being processed. 

11. Gels can be left in fixing solution overnight or longer if necessary until staining 

is required. During staining procedures gels are shaken gently on a gel rocker 

platform or on an orbital shaker set at low speed. 

12. The volume of 2D lysis-rehydration buffer required to solubilize a sample is 

dependent on the dimensions of the IPG strip being re-hydrated. For mini gels 

using 7-cm IPG strips the volume is 125 μL, for 11-cm IPG strips 200 μL, for 

13-cm IPG strips 250 μL and for 18-cm large format gels 350 μL. 

13. Using the IEF reswelling tray up to 12 IPG can be rehydrated with samples at 

the same time. 

14. The voltage and duration of IEF is dependent on the pH and length of the IPG 

strip being used and on the total amount of protein loaded onto the gel. For 

guidelines on the parameters used with different IPG strips see the technical 

data sheets supplied with the IPG strips by the manufacturers. 

15. Following IEF gel strips can be equilibrated immediately ready for the SDS- 

PAGE second dimension step or they can be stored at –20°C and the equilibration 

and second dimension done at a later date.


References 

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of plant endoplasmic reticulum – identification of protein disulphide-isomerase as 

the major reticuloplasmin. Eur. J. Biochem 235, 215–24. 

2. Vitale, A. and Denecke, J. (1999) The endoplasmic reticulum – gateway of the 

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19 

Cell Wall Fractionation for Yeast and Fungal Proteomics 


Summary 

The cell wall is an external envelope shared by yeasts and filamentous fungi that 

defines the interface between the microorganism and its environment. It is an extremely 

complex structure consisting of an elastic framework of microfibrillar polysaccharides 

(glucans and chitin) that surrounds the plasma membrane and to which a wide array of 

different proteins, often heavily glycosylated, are anchored in various ways. Intriguingly, 

these cell wall proteins (CWPs) play a key role in morphogenesis, adhesion, pathogenicity, 

antigenicity, and as a promising target for antifungal drug design. However, the CWPs are 

difficult to analyze because of their low abundance, low solubility, hydrophobic nature, 

extensive glycosylation, covalent attachment to the wall polysaccharide skeleton, and 

high heterogeneity. We describe a typical procedure of cell wall fractionation to isolate 

and solubilize different CWP species from yeasts and filamentous fungi according to 

the type of linkages that they establish with other wall components and under suitable 

conditions for following reproducible proteomic analyses. CWPs retained noncovalently or 

by disulfide bonds are first extracted from isolated yeast or fungal cell walls by detergents 

and reducing agents. Subsequently, CWPs covalently linked to or closely entrapped within 

the internal glucan-chitin network are sequentially released either by mild alkali conditions 

or by enzymatic treatments first with glucanases and then with chitinases. This strategy 

is a powerful tool not only for obtaining an overview of the sophisticated cell wall 

proteome of yeasts and filamentous fungi, but also for characterizing mechanisms of 

incorporation, assembly and retention of CWPs into this intricate cellular compartment 

and their interactions with structural wall polysaccharides. 

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

proteins; proteomics; yeast. 



217

218 Pitarch et al. 

Fig. 1. Schematic representations of the cell wall from S. cerevisiae and C. albicans, 

and molecular modules of CWPs solubilized by the present procedure of cell wall 

fractionation. The cell wall of S. cerevisiae and C. albicans basically consists of -1,3 

and -1,6-glucans, chitin, mannoproteins and proteins. -1,3-glucan and chitin form 

an elastic microfibrillar polysaccharide skeleton surrounding the plasma membrane to 

which mannoproteins are attached through -1,6-glucan, alkali-sensitive bonds, and/or 

other hitherto uncharacterized linkages. Cell wall proteins (CWPs; mannoproteins and

Cell Wall Fractionation for Yeast and Fungal Proteomics 219 


The cell wall is an intricate structure common to yeasts and filamentous fungi 

that surrounds the plasma membrane and is strategically placed at the interface 

between the cell and its environment, including the host (1–3). This external 

envelope, accountable for 20–30% of the cell dry weight, is essential for the 

survival of the microorganism. In fact, it is involved in many vital functions, 

such as physical protection, osmotic stability, selective permeability barrier, 

immobilized enzyme support, cell-cell interactions (e.g., cell recognition and 

adhesion) and morphogenesis, to name but a few. In pathogen fungi, this 

cellular compartment further takes an active part in virulence, pathogenicity, 

antigenicity, immunomodulation of the immune response, and adhesion to host 

substrates (2). Most importantly, its essential nature and its fungal specificity 

(given its absence in mammalian cells) interestingly make the cell wall an 

attractive target site (see Note 1) to design antifungal drugs with selective 

toxicity for human pathogen fungi, such as Candida albicans or Aspergillus 

fumigatus, among others (4–6). 

Although the cell wall structure and organization have been investigated 

most extensively in the prototype yeast Saccharomyces cerevisiae, a similar 

molecular model is also applicable for other ascomycetes, and in particular 

for C. albicans, a dimorphic fungus capable of growing either in yeast form 

or as hyphae (see Note 2) (1,7,8). The cell wall of S. cerevisiae is mainly 

composedofglucans(with-1,3and-1,6linkages),chitin(N-acetylglucosamine 

polymers), and proteins (often highly O- and/or N-mannosylated) interconnected 

by covalent and/or non-covalent bonds, leading to an elevated complexity (see 

Fig. 1). -1,3-glucan, the major component of the cell-wall (electron-transparent) 

inner layer, forms an elastic three-dimensional microfibrillar framework, encircling 

the cell, to which other wall constituents are covalently anchored. Chitin 

is often cross-linked to the -1,3-glucan microfibrillar backbone on its inner 

side (close to the plasma membrane) and, to a lesser extent, to short sidechains 

of -1,6-glucan. This N-acetylglucosamine polymer presents low levels 

(see Note 2), except in the budding neck ring, in the primary septum, in and 

around the bud scars, or under stress conditions. Both -1,3-glucan and chitin 

(structural wall polysaccharides) provide mechanical strength and elasticity 

◭ 

Fig. 1. proteins) can also be loosely associated, either by non-covalent bonds or 

through disulfide bridges, with other covalently linked CWPs. See Introduction for 

further information. The callouts depict details for potential mechanisms of CWP 

retention into the cell wall on the basis of the procedure of cell wall fractionation 

described in this chapter to isolate and solubilize different CWP species from yeasts 

and filamentous fungi.


to the cell wall. -1,6-glucan, a flexible minor wall component, interconnects 

certain cell wall proteins (CWPs), the so-called glycosyl phosphatidylinositol 

(GPI)-CWPs, with -1,3-glucan (∼90% of GPI-CWPs) or chitin (∼10% 

of GPI-CWPs) through a phosphodiester bridge in their GPI remnant (see 

Note 3). The CWPs are mostly located on the outside of this -1,3-glucanchitin 

network (i.e., at the cell-wall electron-dense outer layer) and, in minor 

amounts, throughout the cell wall, determining its porosity. These CWPs can be: 

1. Loosely associated, either noncovalently or through disulfide bonds, with other 

cell wall components. This group of CWPs comprises (1) soluble precursor 

forms of covalently linked CWPs, (2) proteins related to the biosynthesis and 

modulation of wall constituents, such as -1,3-glucosyltransferase (Bgl2p), exoglucanase 

(Exg1p) and chitinase (Cts1p), and (3) noncanonical proteins 

“classically” considered to be confined to the intracellular compartment because 

they lack the conventional secretory signal sequence (2,9–11). Nevertheless, the 

mechanism by which these nonconventional proteins are targeted to the cell 

surface remains enigmatic (11–13). This array of loosely associated proteins is 

commonly found at the cell surface but also, in a smaller ratio, in the cell-wall 

inner layers. These CWPs can be extracted using detergents and reducing agents 

(see Fig. 1). 

2. Covalently linked to -1,3-glucan: 

a. Directly via an alkali-labile linkage (speculatively through a O-linked sidechain), 

such as PIR-CWPs (CWPs with internal repeats). The PIR-CWPs are 

highly O-mannosylated proteins with one or more internal repeat regions, a 

N-terminal signal peptide, a Kex2 proteolytic processing site, and a C-terminal 

sequence with four cysteine residues at highly conserved positions (1,14–17). 

They are normally located in the cell-wall inner layer (18). Other CWPs belong 

to this category are also present in the yeast cell walls (see the following and 

Fig. 2). This type of CWPs can be solubilized under mild alkali conditions or 

by enzymatic treatment with -1,3-glucanases but not with -1,6-glucanases 

(see Figs. 1–3). 

b. Indirectly by a -1,6-glucan moiety through their GPI remnant, such as GPI- 

CWPs. The GPI-CWPs are highly O-glycosylated proteins with an N-terminal 

signal peptide, a C-terminal GPI anchor addition signal, and serine- and 

threonine-rich regions (see Note 3) (1,7,17,19). These CWPs are predominantly 

placed in the cell-wall outer layer. This group of CWPs can be released 

either by enzymatic treatment with -1,3 or -1,6-glucanases or by using 

hydrofluoric acid (HF)-pyridine, which cleavages the phosphodiester bond in 

the GPI remnant (20) (GPI-CWP 1 in Figs. 1–3). 

3. Covalently anchored to chitin by a -1,6-glucan moiety via their GPI remnant, 

such as some GPI-CWPs (21,22). This type of CWP-polysaccharide complex is 

largely found in the lateral walls or under stress conditions. These CWPs can be


Fig. 2. Schematic representations of the different known types of covalently linked 

CWPs in S. cerevisiae and C. albicans, and the most commonly used methods to 

solubilize them from isolated cell walls. These diagrams are based on information from 

references (1,14,18-21,23). See Introduction for further details. PIR-CWPs, a small 

number of hybrid GPI-CWPs (e.g., Cwp1p (23); GPI-CWPs 2 and 3) at acidic pHs, and 

other CWPs (25,32) are directly attached to -1,3-glucan through an alkali-sensitive 

linkage (ASL). GPIr denotes GPI remnant, and ASL alkali-sensitive linkage. 

extracted either by enzymatic treatment with chitinases or -1,6-glucanases or by 

using HF-pyridine (GPI-CWP 4 in Figs. 1–3). 

However, it is unsurprising that other types of linkages, hitherto uncharacterized, 

among CWPs and structural wall components are also present in the 

yeast cell wall (see Fig. 1). Be that as it may, there is no doubt that the molecular 

model of the yeast cell wall is even more sophisticated than that outlined above. 

This is because certain CWPs can simultaneously be retained into the -1,3glucan/chitin 

skeleton in various ways. For instance, the Cwp1p, a S. cerevisiae 

GPI-CWP, is double-anchored to the -1,3-glucan framework both through an 

alkali-sensitive linkage and by its -1,6-glucan moiety, playing an important 

role in stress response (23). Hence, two additional GPI-CWP-polysaccharide 

complexes are defined (GPI-CWPs 2 and 3 in Figs. 2 and 3). 

Taking into account the special architecture and nature of the cell wall, the 

isolation and solubilization of CWPs from this complex cellular compartment 

is not therefore an evident and easy affair. Indeed, CWPs from yeasts


Fig. 3. Venn diagram summarizing the most commonly used methods to extract the 

different known types of CWPs in S. cerevisiae and C. albicans. This chart is based 

on data from references (1,10,14,18–21,23). The GPI-CWP numbers refer to those 

indicated in Fig. 2. 

and filamentous fungi are tricky enough to resolve by two-dimensional 

electrophoresis (2-DE) gels, because of their low abundance, low solubility, 

hydrophobicity, high heterogeneity, extensive glycosylation (especially, Oand/or 

N-mannosylation), and covalent attachment to the wall polysaccharide 

(-1,3-glucan/chitin) skeleton (11,24). These problems can, at least to some 

degree, be solved by sequential solubilization of CWPs on the basis of the type 

of attachments that they establish to other cell wall components (11,17,25). 

This procedure implies breakage of the covalent linkages between CWPs and 

wall polysaccharides (see Chapter 20). Intriguingly, cell wall fractionation is 

an appropriate paradigm system to: 

1. Reduce the intricacy of the cell wall. 

2. Enrich samples for CWPs and thus increase the detection of low-abundance 

species by removing the most abundant soluble gene products. 

3. Enhance the solubility of large, low abundance, and/or hydrophobic CWPs. 

4. Define (map and identify) the proteins that make up the cell wall (the cell wall 

proteome), and elaborate a comprehensive and integrated view of the complex 

CWP composition. The CWP resolution can be increased using the cell wall 

fractionation procedure described here, because the heterogeneous population of 

protein species present in the yeast and fungal cell envelope can be distributed 

over several 2-DE gels (11). 

5. Characterize mechanisms of incorporation, assembly and retention of CWPs into 

the cell wall. 

6. Elucidate the CWP interactions with cell wall polysaccharides.


7. Study protein-protein interactions or regulatory networks exclusive to the cell 

wall. 

8. Monitor abnormal protein expression localized to this external envelope. 

9. Discover novel diagnostic/prognostic markers, antifungal targets and/or therapeutic 

candidates for human mycoses. 

This chapter will integrate a typical procedure of cell wall fractionation to 

extract different CWP species from isolated cell walls of yeasts or filamentous 

fungi according to their interactions with other wall components. The resulting 

selectively enriched CWP fractions can then directly be (1) analyzed by 2- 

DE (see Note 4) and mass spectrometry (MS) (11,17) or (2) digested with 

trypsin, followed by liquid chromatography (LC) in tandem with MS analyses 

(25) to circumvent some of the difficulties associated with in-gel digestion of 

the heavily glycosylated CWPs (12). The purity and quality of these enriched 

fractions of CWPs should be screened by using bona fide markers both of the 

cell wall and of intracellular compartments before carrying out any interpretation 

of the results. However, the unambiguous evidence for their cell wall 

location will only be established after (1) their in situ immunolocalization (i.e., 

immunoelectron microscopy or immunofluorescence studies) and/or (2) the use 

of tagged fusion proteins (e.g., c-myc-tag or green fluorescence protein (GFP) 

fusion proteins) based on the fusion of the protein in question to modified 

versions of extracellular enzymes that rely on a detectable phenotype. 

2. Materials 

Growth media, solutions and buffers should be sterilized by autoclaving 

before use when working under sterile conditions. Their labile components 

should be sterilized separately using a 0.22-μm filter (Millipore, Bedford, MA) 

and then added to the other autoclaved ingredients. All solutions and buffers 

should be prepared with ultrapure water, as provided by Nanopure or Milli-Q 

18 M/cm resistivity systems (Millipore), and precooled when the procedure 

is carried out at 4°C. 

2.1. Cell Wall Isolation from Yeasts and Filamentous Fungi 

1. Yeast-Peptone-D-glucose (YPD) plates: 1% (w/v) yeast extract (Difco Laboratories, 

Detroit, MI), 2% (w/v) peptone (Difco), 2% (w/v) d-glucose, 2% (w/v) 

agar (Difco). 

2. YPD medium: 1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) d-glucose. 

3. Bead mill homogenizer (model MSK; Braun Biotech International GmbH, 

Melsungen, Germany). 

4. Liquid carbon dioxide (CO2).


5. 0.40- to 0.60-mm chilled, acid-washed glass beads (Sartorius, Goettingen, 

Germany) (see Note 5). 

6. PMSF stock solution: 0.1 M in isopropanol. Dissolve 174 mg of phenylmethylsulfonyl 

fluoride (PMSF; Fluka, Chelmsford, MA) in a final volume of 10 mL 

isopropanol, and store at –20°C (see Note 6). PMSF should be handled with 

caution because it is highly toxic. Weigh this hazardous compound in a fume 

hood, and wear gloves, goggles and a mask. 

7. Lysis buffer: 10 mM Tris-HCl, pH 7.4, 1 mM PMSF. 

8. YPD-chloramphenicol plates: 1% (w/v) yeast extract, 2% (w/v) peptone, 2% 

(w/v) d-glucose, 2% (w/v) agar (Difco), 10-μg/mL chloramphenicol. When the 

1-L autoclaved YPD and agar solution cools to ∼65°C, add 1 mL of 10× 

chloramphenicol solution. 

9. Chloramphenicol solution (10X): Dissolve 10 mg of chloramphenicol in a final 

volume of 1 mL ethanol, and sterilize using a 0.2-μm filter. 

10. Wash solution A: 1 mM PMSF. 

11. Wash solution B: 5% (w/v) NaCl, 1 mM PMSF. 

12. Wash solution C: 2% (w/v) NaCl, 1 mM PMSF. 

13. Wash solution D: 1% (w/v) NaCl, 1 mM PMSF. 

2.2. Protein Solubilization from Isolated Yeast and Fungal Cell Walls 

2.2.1. By Detergents and Reducing Agents 

1. Wash buffer: 50 mM Tris-HCl, pH 8.0, 1 mM PMSF. 

2. Extraction buffer: 50 mM Tris-HCl, pH 8.0, 0.1 M EDTA, 2% (w/v) SDS, 10 mM 

DTT (dithiothreitol; see Note 7). 

2.2.2. Under Mild Alkali Conditions 

1. Wash solution: 1 mM PMSF. 

2. Wash buffer: 0.1 M sodium acetate, pH 5.5, 1 mM PMSF. 

3. Extraction solution: 30 mM NaOH, 1 mM PMSF. 

4. Stop solution: acetic acid. 

2.2.3. By -1,3-Glucanase Treatment 


2. Wash buffer: 50 mM Tris-HCl, pH 7.5, 1 mM PMSF. 

3. Extraction buffer: 1,500 U Quantazyme ylgTM (Quantum Biotechnologies Inc, 

Montreal, Canada; see Note 8) per gram of wet weight of cell walls, in 2 mL of 

a solution containing 50 mM Tris-HCl, pH 7.5, 10 mM DTT, 1 mM PMSF (see 

Note 9). 

4. Stop solution: 10% SDS.


2.2.4. By Exochitinase Treatment 


2. Wash buffer: 50 mM sodium phosphate buffer, pH 6.3. 

3. Extraction buffer: 0.3 U exochitinase (Sigma, St. Louis, MO) per gram of wet 

weight of cell walls, in 2 mL of a solution containing 50 mM sodium phosphate 

buffer, pH 6.3 (see Note 10). 

4. Stop solution: 10% SDS. 

2.3. Protein Precipitation. 

1. 100% Trichloroacetic acid (TCA) solution: Dissolve 100 g of TCA in sufficient 

water to yield a final volume of 100 mL (see Note 11). TCA should be handled 

with caution, because it is extremely caustic. Protect eyes and avoid contact with 

skin when working with TCA solutions. 

2. Acetone precooled to –20°C. 

3. Neutralizing solution: 0.1 N NaOH. 

3. Methods 

The protocols described below outline a typical procedure of cell wall 

fractionation to isolate and solubilize different CWP species from yeasts and 

filamentous fungi on the basis of the type of linkages that they establish 

with other wall components. This method involves (1) cell homogenization 

by physical disruption techniques, (2) isolation of cell walls by differential 

centrifugation, (3) sequential solubilization of CWPs from isolated cell walls 

using different chemical agents (detergents, reducing agents, and alkalis) and 

enzymes (glucanases and chitinases), and (4) precipitation of the resulting 

selectively enriched CWP fractions under suitable conditions for subsequent 

proteomic analyses. A flowchart of the strategy presented here is shown in 

Fig. 4. This is based on earlier methods described by Kapteyn et al. (20,21) 

and Mrsa et al. (26), and on recent modifications reported by Pitarch et al. (11) 

that have proved effective at properly extracting CWPs from S. cerevisiae and 

C. albicans. Nevertheless, adaptation of growth and extraction conditions may 

be required for other species of yeasts and filamentous fungi. 

3.1. Cell Wall Isolation from Yeasts and Filamentous Fungi (see 

Note 12) 

Although cell disintegration can be accomplished using a wide variety of 

techniques, mechanical breakage of cells using glass beads is certainly one of 

the most quick and reliable procedures to disrupt the cell walls and plasma 

membranes of yeasts and filamentous fungi (see Note 13). After cell disruption,


Fig. 4. Flowchart of a typical procedure of cell wall fractionation to isolate and 

solubilize different CWP species from yeasts and filamentous fungi according to the 

type of attachments that they establish to other cell wall components. 

cell walls can be separated from other cytosolic and membranous components 

by differential centrifugation of the cell homogenate at relatively low speed 

(see Note 14).


1. Grow yeast or fungal cells on an YPD plate (stock maintenance medium) at 

30°C for 2 days. Use a single colony to inoculate 50 mL of YPD (or selective) 

medium in a 250-mL flask (see Note 15), and grow overnight at 30ºC with 

vigorous rotary shaking (200 rpm). 

2. Use this 50-mL preculture to inoculate four 2-L flasks containing 500 mL fresh 

YPD medium each, and grow at 30°C in a shaking incubator (200 rpm) until 

the culture reaches early-log phase growth (OD 600nm = 0.5–1; see Note 16). 

3. Harvest the cells by centrifugation at 4,500g for 5 min (see Note 17) and discard 

the supernatant. 

4. Resuspend the cell pellet in 200 mL ice-cold water, and centrifuge 5 min at 

4,500g. Decant the supernatant. 

5. Resuspend the cell pellet in 200 mL ice-cold lysis buffer, and centrifuge 5 min 

at 4,500g. Discard the supernatant. 

6. Resuspend the cells in 3 volumes of ice-cold lysis buffer, and add 3–4 volumes 

of 0.5-mm acid-washed glass beads (see Note 18). Transfer the cell suspension 

to an appropriate sized shaking flask (see Note 19). 

7. Grind the suspension at maximum speed for 30–60 s using a bead mill homogenizer 

and cooling with liquid CO 2 (for example, in a CO 2-refrigerated MSK 

homogenizer), and then place the shaking flask on ice for 1–2 min (see Note 20). 

Repeat this step until complete cell breakage. Monitor the degree of cell breakage 

with a phase-contrast microscope and by plating on YPD-chloramphenicol plates 


8. Enable the glass beads to settle out and collect the supernatant carefully 

(see Note 22). Wash the glass beads with ice-cold lysis buffer and collect the 

washings until they are clear (see Note 23). Pool the supernatant and all the 

washings. 

9. Centrifuge the pooled supernatant and washings (cell homogenate) at 1,000g for 

10 min at 4°C. Discard the supernatant carefully (see Note 24). 

10. Resuspend the isolated cell walls in 200 mL ice-cold wash solution A. Centrifuge 

10 min at 1,000g and at 4°C. Carefully decant the supernatant. Repeat this step 

two more times. 

11. Resuspend the cell walls in 200 mL ice-cold wash solution B. Centrifuge 10 min 

at 1,000g and at 4°C. Carefully decant the supernatant. Repeat this step four 

more times (see Note 25). 

12. Resuspend the walls in 200 mL ice-cold wash solution C. Centrifuge 10 min 

at 1,000g and at 4°C. Carefully remove the supernatant. Repeat this step four 

more times. 

13. Resuspend the pellet in 200 mL ice-cold wash solution D. Centrifuge 10 min at 

1,000g and at 4°C. Carefully decant the supernatant. Repeat this step four more 

times (see Note 26). 

14. Resuspend the walls in 200 mL ice-cold wash solution A. Centrifuge 10 min at 

1,000g and at 4°C. Carefully discard the supernatant. Repeat this step two more 

times using preweighed centrifuge bottles. 

15. Weigh the wet wall pellet (see Note 27).


3.2. Protein Solubilization from Isolated Yeast and Fungal Cell Walls 

CWPs can then be solubilized sequentially from isolated cell walls by a wide 

variety of reagents (e.g., detergents, reducing agents, alkalis, and hydrolytic 

enzymes, among others) in connection with their attachments to other wall 

components (see Fig. 4). All procedures are performed at 4°C with prechilled 

solutions, reagents and apparatus (see Note 28) unless otherwise indicated. 

3.2.1. By Detergents and Reducing Agents 

Detergents, such as sodium dodecyl sulfate (SDS) or n-octylglucoside, can 

be used to extract CWPs that are associated noncovalently with other wall 

components. The use of reducing agents, such as dithiothreitol (DTT) or mercaptoethanol 

(ME), (1) enables the solubilization of CWPs that are loosely 

associated, either by disulfide bridges or through non-covalent bonds, with other 

covalently linked CWPs, (2) results in an increase in the cell wall porosity, and (3) 

facilitates the subsequent action of wall degrading enzymes (see Figs. 1 and 4). 

1. Resuspend the purified cell walls (prepared as described in Subheading 3.1) in 

200 mL ice-cold wash buffer. Centrifuge 10 min at 1,000g and carefully decant 

the supernatant (see Note 24). 

2. Add 5 mL of extraction buffer for each wet gram of cell walls and resuspend. Boil 

the cell wall suspension at 100°C for 10 min, and centrifuge 10 min at 1,000g. 

3. Collect the supernatant and store the pellet. 

4. Precipitate the supernatant (containing SDS/DTT-extractable CWPs) and store 

at –80°C (see Note 29 and Fig. 4). 

5. Repeat step 2 with the stored pellet and carefully discard the supernatant (see 

Note 30). 

6. Weigh the wet wall pellet (containing SDS/DTT-resistant cell walls) and divide 

it equally between two tubes (see Fig. 4 and Notes 27 and 31). Store them for 

further extractions (see Subheadings 3.2.2, 3.2.3 and 3.2.4). 

3.2.2. Under Mild Alkali Conditions 

Treatment of SDS/DTT-resistant cell walls under mild alkali conditions 

(using 30 mM NaOH) results in the extraction of CWPs linked to -1,3-glucan 

through an alkali-sensitive linkage (of unknown nature) by the -elimination 

process (1, 14–17). PIR-CWPs, some GPI-CWPs, and other CWPs belong this 

group (see Figs. 1–4). 

1. Resuspend one tube of the purified SDS/DTT-resistant cell walls (prepared as 

described in Subheading 3.2.1) in 200 mL ice-cold wash solution. Centrifuge 

10 min at 1,000g and carefully decant the supernatant (see Note 24). Repeat this 

step two more times. 

2. Resuspend the walls in 200 mL ice-cold wash buffer. Centrifuge 10 min at 1,000g 

and carefully discard the supernatant. Repeat this step five to seven more times.


3. Add 4 mL of ice-cold extraction solution for each wet gram of cell walls and 

resuspend. Incubate the cell wall suspension at 4°C for 17 h with gentle shaking 


4. Stop the chemical reaction by adding neutralizing amounts of acetic acid (see 

Note 33). 

5. Centrifuge 10 min at 1,000g and collect the supernatant (containing alkalisensitive 

CWPs). Precipitate or dialyze the clear supernatant (see Note 33) and 

store at –80°C. 

3.2.3. By -1,3-Glucanase Treatment 

-1,3-glucanases (commercially available) can be used to extract CWPs 

covalently anchored to -1,3-glucan. This group of CWPs released from 

SDS/DTT-resistant cell walls by these wall hydrolytic enzymes contains (1) 

GPI-CWPs, which are indirectly attached to -1,3-glucan, via -1,6-glucan, 

through a phosphodiester bridge and can alternatively be solubilized either by - 

1,6-glucanases (see Note 34) or by using HF-pyridine, (2) alkali-sensitive CWPs 

(including PIR-CWPs and some GPI-CWPs, among others), which are anchored 

to -1,3-glucan through uncharacterized linkages (perhaps by a O-linked sidechain) 

and can also be released under mild alkali conditions (using 30 mM 

NaOH; see Subheading 3.2.2), and (3) other CWPs linked to -1,3-glucan 

through other types of hitherto unidentified bridges (see Figs. 1–4). 

1. Resuspend the other tube of the purified SDS/DTT-resistant cell walls (prepared 

as described in Subheading 3.2.1) in 200 mL ice-cold wash solution. Centrifuge 




and carefully remove the supernatant. Repeat this step five to seven more times. 

3. Add 2 mL of extraction buffer for each wet gram of cell walls and resuspend (see 

Note 34). Incubate the cell wall suspension at 37°C for 17 h with gentle shaking. 

4. Stop the enzymatic reaction by adding the stop solution at a final concentration 

of 0.4% (w/v) and heating at 100°C for 3–5 min. 

5. Centrifuge 10 min at 1,000g. 

6. Collect the supernatant (containing -1,3-glucanase-extractable CWPs) and store 

the pellet (containing SDS/DTT and -1,3-glucanase-resistant cell walls) for 

further extractions (see Subheading 3.2.4). 

7. Precipitate the clear supernatant and store at –80°C. 

3.2.4. By Exochitinase Treatment 

Enzymatic treatment of the SDS/DTT- and -1,3-glucanase-resistant cell 

walls with exochitinases leads to the solubilization of CWPs covalently 

anchored to chitin. These comprise (1) a small subgroup of GPI-CWPs, which


are indirectly linked to chitin, via -1,6-glucan, through their GPI remnant and 

can also be extracted either by -1,6-glucanases or by using HF-pyridine, and 

(2) other CWPs attached to chitin through other types of hitherto uncharacterized 

linkages (see Figs. 1–4). 

1. Resuspend the SDS/DTT and -1,3-glucanase-resistant cell walls (prepared as 

described in Subheading 3.2.3) in 200 mL ice-cold wash solution. Centrifuge 




and carefully discard the supernatant. Repeat this step five to seven more times. 

3. Add 2 mL of extraction buffer for each wet gram of cell walls and resuspend. 

Incubate the cell wall suspension at 37°C for 17 h with gentle shaking. 

4. Stop the enzymatic reaction by adding the stop solution at a final concentration 

of 0.4% (w/v) and heating for 3–5 min at 100°C. 

5. Centrifuge 10 min at 1,000g and collect the supernatant (containing 

-1,3-glucanase-resistant and exochitinase extractable CWPs). 

6. Precipitate the clear supernatant and store at –80°C. 

3.3. Protein Precipitation. 

The different selectively enriched CWP fractions obtained in the 

Subheading 3.2 should be concentrated and desalted before carrying out further 

proteomic analyses. These can be (1) dialyzed against a volatile buffer and 

dried, or (2) precipitated with TCA as described below (see Note 35), among 

other methods. 

1. Add 1/9th the total volume of the protein sample of an ice-cold 100% TCA 

solution to a final concentration of 10%. 

2. Mix thoroughly and incubate on ice for 30 min. 

3. Centrifuge the suspension at 10,000g for 15 min, and discard the supernatant (see 

Note 36). 

4. Wash protein pellet twice with cold acetone to remove residual TCA. 

5. Air dry for 30 min. 

6. Add neutralizing amounts of 0.1N NaOH (see Note 37) and store at –80°C. 

4. Notes 

1. Supporting this enterprise, -1,3-glucan is targeted by a new antifungal drug 

class in recent clinical use, i.e., echinocandins (including caspofungin and 

micafungin), which blocks the biosynthesis of this cell wall polysaccharide (4–6). 

2. It must be borne in mind that unlike S. cerevisiae, many filamentous fungi 

contain further -glucans and a high chitin content in their cell walls (1). 

3. The GPI-proteins are translocated into the endoplasmic reticulum (ER), where 

(1) the N-terminal signal peptide (secretion signal necessary to enter the classical


ER-Golgi secretory pathway) is cleaved, (2) the C-terminal GPI anchor addition 

signal is replaced with a GPI anchor, and (3) O- and/or N-linked core glycosylation 

takes place. Remarkably, further glycosylation also then occurs in the 

Golgi apparatus. These GPI-anchored proteins are directed through the secretory 

pathway to the outer side of the plasma membrane, where they are attached 

through their C-terminal GPI anchors (see Fig. 5). Intriguingly, some of them are 

released from the plasma membrane by cleavage of their GPI anchors, resulting 

in GPI anchor remnants (a truncated, lipidless GPI anchors) (1,19,27,28). These 

proteins are then covalently incorporated into the cell wall, by the attachment 

of their GPI remnants to -1,6-glucan (19). 

4. It is convenient to use gradient gels that enable protein separation up to at 

least 500 kDa, because the extensive glycosylation (especially, O- and/or Nmannosylation) 

of a large proportion of CWPs results in extremely high apparent 

molecular masses on SDS-PAGE or 2-DE gels (7). 

5. The size of the glass beads is crucial to achieve an efficient cell disruption. 

Optimal bead size for spores is 0.1 mm and for yeast and mycelia 0.5 mm. 

6. The PMSF can also be solubilized in ethanol, methanol and 1,2-propanediol. It 

is unstable in aqueous solution. PMSF is added to reduce possible proteolytic 

processes. It inhibits serine proteases (e.g., trypsin, chymotrypsin, and thrombin) 

and thiolproteases (e.g., papain). 

7. 40 mM -mercaptoethanol may be substituted for 10 mM DTT. mercaptoethanol 

or DTT should be added just before use. 

8. Quantazyme ylg TM is a recombinant yeast -1,3-glucanase purified from E. coli 

that is completely free of protease, endo- and exonuclease activity. It is highly 

stable in solution for months at 4°C, maintaining its whole activity. 

9. The use of reducing agents facilitates the ability of Quantazyme ylg TM to degrade 

the cell wall -1,3-glucan. 40 mM -mercaptoethanol or 10 mM cysteine may 

be substituted for 10 mM DTT. 

10. Exochitinase is a cell wall lytic enzyme isolated from Serratia marcescens that 

catalyzes the progressive degradation of chitin, starting at its nonreducing end. 

This preparation contains phosphate buffer salts and shows an optimum pH 

of 6.0. 

11. Because TCA is very hygroscopic, the whole content of a newly opened TCA 

bottle should be used to prepare the TCA stock solution. 

12. Perform all procedures from this subheading until isolated cell walls are obtained 

under sterile conditions. Use sterile centrifuge bottles. 

13. Mechanical cell breakage using a bead mill homogenizer is considered the 

technique of choice for disrupting cells with cell walls, especially spores, yeasts 

and fungi, but it also works successfully with algae, bacteria, plant, and animal 

tissue culture in suspension. Cell disruption takes place by the crushing action 

of the glass beads, which are vigorously agitated by shaking or stirring, after 

crashing with the cells. The Braun MSK cell homogenizer combines two types 

of motions in the shaking flask: rotation and tumbling. For small-scale preparations, 

a Fast-Prep cell breaker (Q-Biogene, Carlsbad, CA) can be used in lieu


Fig. 5. Putative model of the incorporation of GPI-anchored proteins into the cell 

wall. This is based on information from references (19,27,28). Their GPI anchor is 

processed at the plasma membrane leading to a GPI remnant (a truncated, lipidless GPI 

anchor). This is cross-linked to -1,6-glucan when these GPI proteins are covalently 

incorporated into the cell wall. See Note 3 for further details.


of the Braun MSK cell homogenizer. Cell breakage of filamentous fungi can 

alternatively be achieved by (1) grinding freeze-dried mycelium in a mortar and 

pestle, (2) grinding mycelium in liquid nitrogen, or (3) homogenizing thick cell 

pastes. When handling liquid nitrogen, use insulated gloves, avoid any potential 

contact because of the risk of frostbite, and never utilize glass containers because 

they may break. 

14. Differential centrifugation in sucrose density gradients may alternatively be used 

to isolate the cell wall fraction. 

15. Liquid cultures should be grown in a flask that is at least 4–5 times larger than 

the culture volume. 

16. It is important that the yeast culture is in early-log phase growth 

(∼ OD 600nm= 0.5–1) because it is easier to disrupt their cell walls with the bead 

mill homogenizer than those close to or in stationary phase growth. It must be 

borne in mind that the composition of the cell wall changes with the growth 

stage and culture conditions (growth temperature, external pH, oxygen levels, 

and composition of the growth medium (1,29)), among others. Hence, it may be 

necessary to adjust the cell density according to the specific objectives of the 

experiment. 

17. Overall, yeasts are successfully harvested by centrifugation, whereas vacuumassisted 

filtration, rather than centrifugation, is often preferred for harvesting 

filamentous fungi. 

18. The wet weight (in grams) of the cell pellet is nearly equal to the packed cell 

volume (in milliliters). Add about 3 mL of ice-cold lysis buffer for each wet 

gram of cell pellet and resuspend. 

19. The shaking flasks can be made of glass or stainless steel. The latter are better at 

transferring heat. It is essential to exclude all air from the shaking flask before 

the cell breakage to avoid foaming and denaturation of proteins. 

20. It is extremely important that the temperature of the cell suspension remains at 

4°C during the cell disruption to prevent heat inactivation and denaturation of 

proteins. Use liquid CO 2 to cool the protein sample during cell breakage. 

21. This procedure is carried out until complete cell breakage, which will vary with 

yeast/fungal strain and growth stage. The degree of cell breakage should be 

examined: 

a. Before proceeding: by observing cell lysis under a phase-contrast microscope. 

b. After proceeding: by plating an aliquot of the cell suspension after and 

before cell disruption on YPD-chloramphenicol plates and growing at 30°C. 

The ratio of CFUs after to before cell breakage is then calculated to 

estimate the efficiency of cell disruption and, therefore, potential intracellular 

contamination in the subsequent steps. Failure of cells to grow on 

YPD-chloramphenicol plates should be evidenced. 

22. The supernatant (cell homogenate) and following washings (residual cell 

homogenate) can be collected (1) by decanting after allowing the glass beads to 

settle out by gravity, or (2) by straining through a perforated tube or “strainer”


(see Fig. 6). To perform the last method, transfer the cell homogenate containing 

the glass beads to a plastic tube and make holes of a diameter less than 0.4–0.6 

mm in its bottom with a flamed needle to obtain a “strainer.” After straining 

the cell homogenate off, collect it and wash the glass beads with ice-cold lysis 

buffer. Collect the washing, and wash again until the collected washings are 

clear. Pool the cell homogenate and all the washings. The washing step is critical 

Fig. 6. Proposed procedure for removing the glass beads from the cell homogenate. 

In the method described here, the cell homogenate is strained through a perforated tube 

with holes of a diameter less than 0.4–0.6 mm in its bottom, whereas the glass beads 

(with a diameter of 0.4–0.6 mm) are retained in the perforated tube or “strainer” (see 

Note 22). Successive washings with ice-cold lysis buffer should then be performed to 

increase the recovery of cell homogenate, and thus of cell wall material.


to avoid any loss of material while removing the glass beads from the cell 

homogenate. 

23. To reuse the glass beads, rinse them by soaking in concentrated nitric acid for 

1 h, and then flush through with water. Dry them in a baking oven, cool, and 

store at 4°C (see Fig. 6). 

24. It is important to decant the supernatant carefully, because the cell-wall pellet 

is less compact than the preceding cell pellets. In general, care should be taken 

during the following steps of this procedure (especially during decanting of the 

washings) to prevent any loss of cell walls, and therefore of CWPs. Be that as 

it may, this potential loss of protein material does not interestingly result in a 

preferential deficiency of certain CWP species or families. 

25. The purpose of extensively washing the isolated cell walls with solutions 

of decreasing concentrations of NaCl is to remove potential extracellular, 

membranous and/or cytosolic protein contaminants that can be adhered to them 

through nonspecific ionic interactions (see Fig. 7). The number of washing steps 

will therefore rely on the objectives of the experiment. 

26. A further washing step with 0.1 M Na 2CO 3 overnight at 4°C under gentle shaking 

may reduce potential cytoplasmic contamination that is found into microsomes, 

because this treatment allows them to be opened and washed (30,31). 

27. The wet weight (in grams) of cell walls in the pellet can be calculated by taking 

the weight of the centrifuge bottle (which has been previously weighed) away 

from the total weight (i.e., the weight of wall pellet plus centrifuge bottle). 

28. Centrifugation must be performed in refrigerated centrifuges at 4°C, with 

prechilled rotors, to avoid undesirable proteolytic activity. 

29. This fraction is not a pure preparation of cell wall, but rather is enriched in CWPs 

(loosely associated with other wall components) and may potentially contain a 

small amount of membrane proteins. 

30. This extra step is important to remove any remaining SDS/DTT-extractable 

CWPs and potential membranous components from the isolated cell walls, which 

will be used in subsequent CWP extraction steps (see Fig. 4). 

31. This pellet, containing SDS/DTT-resistant cell walls, is divided equally between 

two tubes to independently extract alkali-sensitive CWPs and -1,3-glucanaseextractable 

CWPs in the following steps (see Fig. 4). 

32. It is convenient to place the cell wall suspension into a container with ice in a 

cool room at 4°C (see Fig. 4). 

33. The chemical reaction must be stopped by acid neutralization. This can be 

performed: 

a. by adding neutralizing amounts of acetic acid to the wall suspension. The 

clear supernatant containing alkali-sensitive CWPs should then be (i) precipitated 

by adding nine volumes of ice-cold methanol (18) or (ii) dialyzed 

against water or 20 mM bis-Tris, pH 6.0 (32). 

b. by subsequent protein precipitation of the clear supernatant containing alkalisensitive 

CWPs with TCA (see Subheading 3.3).


Fig. 7. Basic principle of the procedure used for washing the isolated cell walls 

of yeasts and filamentous fungi. Potential extracellular, membranous and/or cytosolic 

protein contaminants that may nonspecifically associate with the isolated cell walls 

through ionic interactions can be removed under relatively more stringent conditions. 

Extensive washings of the isolated cell walls with high ionic strength (e.g., 5% NaCl) 

can disrupt the nonspecific ionic interactions between the isolated cell walls and these 

putative contaminants.


34. Endo--1,6-glucanase isolated from Trichoderma harzianum (33) can be used 

to release GPI-CWPs (0.8 U/g wet weight of cell walls in 100 mM sodium 

acetate pH 5.5) (18,20). Subsequently, -1,6-glucanase-digested cell walls can 

be treated with Quantazyme or ice-cold 30 mM NaOH to extract (1) the - 

1,6-glucanase-resistant PIR-CWPs, (2) the -1,6-glucanase-resistant GPI-CWPs 

(GPI-CWPs linked to the -1,3-glucan through a alkali-sensitive linkage), and 

(3) hybrid GPI-CWPs, such as Cwp1p (see Fig. 2) (23). 

35. The protein concentration should be higher than 100 μg/mL before precipitating 

with TCA. If the amount of protein precipitated is less than 1 nmole, the protein 

sample should be concentrated and desalted by other ways (e.g., by ultrafiltration 

or forced dialysis), because the pellet is imperceptible. 

36. The supernatant should be stored in case the protein did not precipitate. 

37. Protein preparation can then be resuspended in a small volume of buffer suitable 

for the subsequent analytical procedures (e.g. sample buffer for SDS-PAGE or 

2-DE). 


We thank the Merck, Sharp & Dohme (MSD) Special Chair in Genomics 

and Proteomics, European Community (STREP LSHB-CT-2004-511952), 

Comunidad de Madrid (S-SAL-0246/2006) and Comisión Interministerial de 

Ciencia y Tecnología (BIO-2003-00030 and BIO-2006-01989) for financial 

support of our laboratory. 

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20 

Collection of Proteins Secreted from Yeast Protoplasts 

in Active Cell Wall Regeneration 


Summary 

The yeast cell wall is a dynamic and complex matrix of polysaccharides (glucans, 

mannans, and chitin), proteins and minor amounts of lipids that isolate the yeast from 

the extracellular medium, protecting it against osmotic and physical injuries. Removal of 

this essential structure for cell integrity and viability by controlled enzymatic digestion 

in an iso-osmotic medium brings about protoplast formation. When yeast protoplasts 

are incubated in an osmotically stabilized liquid nutrient medium, cell wall precursors 

are secreted into the culture medium to de novo synthesize the cell wall. During the 

early stages of the regeneration process of protoplast walls, many wall protein precursors 

(presumably structural proteins along with remodeling and cross-linking enzymes) are 

shed into the extracellular medium but not covalently incorporated into the nascent cell 

wall, intriguingly enabling their easy isolation and solubilization. We have developed a 

method to collect proteins secreted from yeast protoplasts in active cell wall regeneration 

under conditions that are suitable for subsequent proteomic analyses. This procedure 

circumvents some of the troubles intrinsically related to other extraction protocols of cell 

wall proteins, such as chemical or enzymatic modifications, and poor quality in protein 

resolution and identification because of linkages to glucan/chitin residues. It further offers 

a valuable model system to understand how the de novo cell wall biosynthesis occurs in 

the yeast cell or how the yeast cell wall participates in morphogenesis. 

Key Words: Candida albicans; cell wall; protoplasts; regeneration; Saccharomyces 

cerevisiae; yeast. 


Yeasts are unicellular eukaryotic organisms that, unlike mammalian cells, 

are surrounded by an elastic and highly dynamic structure, namely the cell wall. 



241


This external envelope, basically composed of -1,3 and -1,6-glucans, chitin, 

mannoproteins, and proteins (1,2), is crucial for preserving the morphology 

and osmotic integrity of the yeast cell and, therefore, essential for cell viability 

(see chapter on Cell Wall Fractionation for Yeast and Fungal Proteomics for 

further details). The yeast cell wall can be completely eliminated by controlled 

enzymatic digestion in an iso-osmotic medium, resulting in the formation of 

protoplasts (see Fig. 1). Yeast protoplasts are uniquely enveloped by the plasma 

membrane, which displays typical invaginations and dictates their distinctive 

spherical shape on the basis of physical laws. When yeast protoplasts are cultivated 

in osmotically stabilized nutrient media, these are intriguingly able to 

synthesize a new cell wall and revert to normal cells, which are capable of proliferating 

and inducing proper morphogenesis (see Note 1) (3–6). Both processes, 

cell wall regeneration and protoplast reversion, are intrinsically associated with 

yeast cell survival (see Note 2) (5). 

Chitin is the first cell wall component to be deposited on the surface of 

regenerating protoplasts (7–9). The microfibrils of nascent chitin, distributed 

irregularly over the plasma membrane at the early stages of reversion, undergo 

a gradual rise in density during the subsequent stages of cell wall regeneration, 

leading to the formation of a regular fibrillar mesh. This de novo chitin skeleton, 

located around the plasma membrane of reverting protoplasts, is soon overlaid 

with -1,3-glucan. Remarkably, although cell wall proteins (i.e., mannoproteins 

and proteins) begin to be synthesized early in the course of protoplast reversion 

to normal cells, their covalent incorporation into this de novo glucan-chitin 

framework is delayed until an adequate amount of -1,3-glucan molecules is 

assembled on the nascent polysaccharide lattice. In fact, cell wall proteins are 

the last components to be effectively bound to the regenerating wall. For this 

reason, during the initial stages of the reversion process, cell wall proteins 

(biologically intended for being anchored to the wall polysaccharide network) 

are not covalently retained into the nascent wall of regenerating protoplasts and, 

consequently, are shed into the extracellular medium. A schematic representation 

of the dynamics of protoplast formation and de novo wall construction 

in reverting protoplasts is shown in Fig. 1. 

The use of regenerating protoplasts and, in particular the collection of 

proteins secreted from yeast protoplasts in active cell wall regeneration (i.e., 

during the first stages of the regeneration process of protoplast walls) into the 

culture medium, are a good and simple model system to: 

1. Easily isolate and solubilize proteins of the complex yeast cell wall, because these 

are freely released from actively reverting protoplasts into the culture medium. 

2. Reduce the complexity of this subcellular compartment (see Chapter 19 for 

details on intricate structure and molecular organization of the yeast cell wall) and

Collection of Proteins Secreted from Regenerating Yeast Protoplasts 243 

Fig. 1. Dynamics of protoplast formation, cell wall regeneration and protoplast 

reversion to normal yeast cells. Elimination of the yeast cell wall by controlled 

enzymatic digestion in an iso-osmotic medium leads to the protoplast formation (see 

Note 14). Yeast protoplasts have typical intrinsic features, such as (1) spherical shape, 

(2) invaginations of the plasma membrane in many areas, and (3) ability to synthesize 

new cell walls and revert to normal growing cells in iso-osmotic regenerating conditions 

(see Note 1). Regenerating protoplasts in an osmotically stabilized nutrient medium


facilitate further related analytical procedures, exploiting the fact that the yeast 

cell is deprived of its cell wall and stimulated to resynthesize it step by step. 

3. Bypass some of the problems inherently associated with chemical and/or 

enzymatic treatments to solubilize cell wall proteins (see Chapter 19), such as: 

a. Potential protein modifications. These are avoided with the method described 

here because cell wall proteins are collected from the growth medium and 

directly analyzed in ensuing proteomic studies without the use of chemical 

agents or enzymes that may in some measure modify them. 

b. The extraction of proteins bearing glucan and/or chitin side-chain 

residues, which hinder their subsequent resolution by two-dimensional gel 

electrophoresis (2-DE) and/or identification by mass spectrometry (MS) (10, 

11). This is circumvented using the present approach, because this does not 

break the covalent linkages between proteins and structural wall polysaccharides 

(glucan/chitin). 

4. Characterize cell wall protein precursors before their incorporation into the cell 

wall (i.e., those gene expression products involved in the de novo wall biosynthesis). 

These mainly include structural proteins, as well as remodelling and 

cross-linking enzymes (10,12–14)). 

5. Study the de novo generation and, therefore, all steps of the biosynthesis of the 

cell wall of yeasts, because this stratagem yields information not only about 

the composition of cell wall precursors but also about their interactions (at the 

different stages of protoplast reversion to normal cells) (7–9,12). 

6. Elucidate the role of the yeast wall in cell morphogenesis, given that cell wall 

regeneration by reverting protoplasts is a de novo process of morphogenesis 

(3–5,15). 

◭ 

Fig. 1. (Continued) first form a fibrillar chitin network, whereupon -1,3-glucan 

molecules are promptly deposited. Mannoproteins and proteins are the last components 

to be assembled on the nascent fibrillar mesh, because their covalent incorporation 

into the regenerating cell wall only occurs after the establishment of a structural 

glucan-chitin matrix around the plasma membrane of reverting protoplasts. Consequently, 

during the early stages of the regeneration process of protoplast walls, cell 

wall protein precursors are not retained into the nascent wall but are secreted into the 

culture medium. It follows that deposition of the fibrillar component (polysaccharide 

framework in the inner wall layer) and amorphous component (protein precursors in 

the outer wall layer) into the de novo cell wall is therefore desynchronized (5). The 

filasomes, located in the reverting site of the protoplasts, appear to be involved in 

the transport of cell wall components from the cytoplasm to the plasma membrane 

in association with secretory vesicles. Interestingly, actin plays a key role in (1) the 

initiation of cell wall regeneration, (2) the correct deposition of cell wall precursors, 

and (3) preservation of the proper shape of the regenerating protoplasts (see Note 2) 

(15,23).


7. Discover new diagnostic/prognostic markers and/or therapeutic targets for human 

mycoses (14). 

8. Increase the productivity and facilitate downstream processing of useful (recombinant 

or nonrecombinant) proteins extracellularly expressed in yeasts (16), 

because the use of regenerating protoplasts eliminates the physical barrier of the 

cell wall, which may limit their excretion. However, it is worth mentioning that 

this stratagem cannot be exploited for long-term production processes because of 

(1) the extreme fragility of yeast protoplasts, and (2) complete cell wall regeneration 

around the active protoplasts during long-term culture periods (see Note 3). 

In this chapter, we describe a method for obtaining proteins secreted into 

the culture medium during the early stages of the regeneration process of 

protoplast walls under suitable conditions for subsequent proteomic analyses 

(10,13,14,17,18). We also provide a rapid and straightforward protocol for 

checking the degree of purity of the cell wall proteins excreted from yeast protoplasts 

in active cell wall regeneration. Proteins involved in cell wall construction 

(such as -1,3-glucosyltransferases, exoglucanases, glycosyl phosphatidylinositol 

(GPI)-proteins, and proteins with internal repeats [PIR], among others), 

heat shock proteins, glycolytic enzymes and other proteins have successfully 

been identified using the procedure presented here in combination with 2- 

DE and MS (10,13,14,17). Given the nature of the protein sample, (multidimensional) 

liquid chromatography techniques in tandem with MS could also 

alternatively be applied. Although other methods to isolate and solubilize cell 

wall proteins from yeast species have been reported in the Chapter 19, the 

choice between these or the one outlined here, which each have advantages 

and disadvantages, will depend on the specific application. 

2. Materials 

All solutions and buffers should be prepared with ultrapure water (doubledistilled, 

deionized water with a resistivity >18M/cm), and prechilled when 

working at 4°C. Growth media, solutions and buffers should be sterilized 

by autoclaving before use. Their labile components should be filter-sterilized 

separately and added to the other ingredients after autoclaving. 

2.1. Collection of Proteins Secreted from Yeast Protoplasts in Active 

Cell Wall Regeneration 

2.1.1. Preparation of Yeast Protoplasts 

1. Yeast-Peptone-D-glucose (YPD) plates: 1% (w/v) yeast extract (Difco Laboratories, 

Detroit, MI), 2% (w/v) peptone (Difco), 2% (w/v) d-glucose, 2% (w/v) 

agar (Difco).


2. YPD medium: 1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) d-glucose. 

3. Pretreatment buffer: 10 mM Tris-HCl, pH 9.0, 5 mM EDTA, 1% (v/v) 

-mercaptoethanol. 

4. 1 M sorbitol solution: Dissolve 182.17 g of sorbitol in sufficient water to yield a 

final volume of 1 L. 

5. Glusulase ® (Du Pont; NEN Life Science Products, Boston, MA). 

2.1.2. Active Cell Wall Regeneration of Yeast Protoplasts 

1. Complete minimal (CM) medium: 0.17% (w/v) yeast nitrogen base (YNB) without 

amino acids or ammonium sulfate (Difco), 0.5% (w/v) (NH4) 2SO4, 2% (w/v) 

d-glucose, 0.13% (w/v) dropout powder (see Note 4). 

2. Dropout powder: 2.5 g adenine, 1.2 g l-arginine, 6.0 g l-aspartic acid, 6.0 g lglutamic 

acid, 1.2 g l-histidine, 3.6 g l-leucine, 1.8 g l-lysine, 1.2 g l-methionine, 

3.0 g l-phenylalanine, 22.5 g l-serine, 12.0 g l-threonine, 2.4 g l-tryptophan, 

1.8 g l-tyrosine, 9.0 g l-valine, and 1.2 g uracil. 

3. Lee medium (19): 0.02% (w/v) MgSO4, 0.25% (w/v) K2HPO4, 0.5% (w/v) NaCl, 

0.5% (w/v) (NH4) 2SO4, 0.05% (w/v) l-alanine, 0.13% (w/v) l-leucine, 0.1% 

(w/v) l-lysine, 0.01% (w/v) l-methionine, 0.007% (w/v) l-ornitine, 0.05% (w/v) 

l-phenylalanine, 0.05% (w/v) l-proline, 0.05% (w/v) l-threonine (see Note 5). After 

autoclaving, add 25 mL of 50% glucose solution and 2 mL of 0.1% biotin solution. 

4. 50% glucose solution: Dissolve 12.5 g of d-glucose in a final volume of 25 mL 

water and sterilize using a 0.22-μm filter (Millipore, Bedford, MA). 

5. 0.1% biotin solution: Dissolve 2 mg of biotin in a final volume of 2 mL water 

and sterilize using a 0.22-μm filter. 

2.1.3. Recovery of Proteins Secreted from Regenerating Protoplasts 

1. Phenylmethylsulfonyl fluoride (PMSF) stock solution: 0.1 M PMSF in 

isopropanol. Dissolve 174 mg of PMSF (Fluka, Chelmsford, MA) in a final 

volume of 10 mL isopropanol, and store at –20 ºC (see Note 6). PMSF should be 

handled with caution, because it is highly toxic. Weigh this hazardous chemical 

in a fume hood, and wear gloves, goggles and a mask. 

2. Antipain stock solution: Dissolve 5 mg of antipain (Sigma, St. Louis, MO) in a 

final volume of 1 mL water, and store at –20 ºC (see Note 7). 

3. Leupetin stock solution: Dissolve 5 mg of leupeptin (Sigma) in a final volume of 

1 mL water, and store at –20 ºC (see Note 8). 

4. Pepstatin stock solution: Dissolve 2.5 mg of pepstatin (Sigma) in a final volume 

of 1 mL methanol, and store at –20 ºC (see Note 9). 

5. Protease inhibitor mix: 0.1 mM PMSF, 2 μg/mL antipain, 2 μg/mL leupeptin, and 

1 μg/mL pepstatin (i.e., for 450 mL of protein solution, add 450 μL of PMSF stock 

solution and 180 μL each of antipain, leupeptin and pepstatin stock solutions). 

Mix just before use. 

6. Filtration unit (Millipore). 

7. 0.22-μm pore-size nitrocellulose filter (Millipore).


2.1.4. Concentration of Proteins Secreted from Regenerating 

Protoplasts 

1. Ultrafiltration apparatus (300-mL stirred ultrafiltration cell system; Amicon; 

Beverly, MA). 

2. YM-10 Diaflo ® ultrafiltration membrane (pre-equilibrated 10,000-Da pore-size 

ultrafilter; Amicon). 

3. Magnetic stirrer. 

4. Oxygen-free nitrogen with pressure regulator (see Note 10). 

5. Membrane wash solution: 1–2 M NaCl. 

6. Membrane store solution: 10% ethanol. 

7. Lyophilizer. 

2.2. Assay of Cell Wall Protein Purity: Alkaline Phosphatase Assay 

1. Lysis buffer: 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM dithiothreitol (DTT), 

0.5 mM PMSF and 5 μg/mL each of antipain, leupeptin and pepstatin (see Notes 6–9). 

2. NaCl-glycine buffer (alkaline buffer): 0.1 M NaCl, 0.1 M glycine, pH 9.7 (adjusted 

with 1 N NaOH). 

3. Stock p-nitrophenol (PNP) solution: 8 mM PNP (Sigma) in NaCl-glycine buffer. 

4. Substrate solution: 20 mM disodium p-nitrophenyl phosphate (PNPP; Sigma) in 

NaCl-glycine buffer. 

5. Stop solution: 0.05 M NaOH. 

6. Spectrophotometer. 

3. Methods 

3.1. Collection of Proteins Secreted from Yeast Protoplasts in Active 

Cell Wall Regeneration 

The flowchart in Fig. 2 summarizes the different steps in the collection of 

proteins excreted into the osmotically stabilized liquid nutrient medium during 

the first stages of the regeneration process of protoplast walls. These involve 

(1) the complete elimination of the yeast cell wall by controlled enzymatic 

digestion (i.e., protoplast formation), (2) active regeneration of the cell wall 

and ensuing secretion of wall protein precursors into the growth medium, 

(3) recovery of the culture medium with secreted proteins by centrifugation 

and filtration, and (4) concentration of secreted proteins by ultrafiltration and 

lyophylization. This method to isolate and solubilize protein precursors of 

the yeast cell walls is taken from the published protocols in Saccharomyces 

cerevisiae and Candida albicans by Pardo et al. (10) and Pitarch et al. (17), 

respectively, and is based on earlier methodology developed by Elorza et al. 

(20). Although this procedure has been used successfully and reproducibly on


Fig. 2. Flowchart of basic steps in the collection of proteins secreted from yeast 

protoplasts in active cell wall regeneration into the osmotically stabilized liquid nutrient 

medium.


these yeast species for subsequent proteomic analyses (10,13,14,17), it may be 

necessary to adjust it for other yeast species or even for filamentous fungi. 

3.1.1. Preparation of Yeast Protoplasts (see Note 11) 

1. Grow yeast cells on a YPD plate (stock maintenance medium) at 28–30 ºC for 

2 d. Inoculate 50 mL of YPD medium in a 250-mL flask (see Note 12) with a 

single colony from the YPD plate, and grow overnight at 28–30 ºC in a shaking 

incubator (200 rpm). 

2. Use this 50-mL preculture to inoculate two 2-L flasks containing 500 mL 

fresh YPD medium each, and grow at 28–30 ºC with vigorous rotary shaking 

(200 rpm) until the culture reaches mid-log phase growth (OD 600nm= 4;see 

Note 13). 

3. Harvest the yeast cells by centrifugation at 4,500g for 5 min and discard the 

supernatant. 

4. Resuspend the cell pellet in 150 mL water, and centrifuge 5 min at 4,500g. 

Decant the supernatant. 

5. Gently resuspend the yeast cells in 100 mL of pretreatment buffer to a density 

of 1–2 × 10 9 cells/mL, and incubate at 28 ºC with gentle rotary shaking (80 

rpm) for 30 min (see Note 14). Centrifuge 10 min at 600g, and discard the 

supernatant. 

6. Gently resuspend the cell pellet in 150 mL of a 1 M sorbitol solution (see 

Note 15). Centrifuge 10 min at 600g to harvest cells, and discard the supernatant. 

7. Gently resuspend the cell pellet in a 1 M sorbitol solution to a density of 5 × 

10 8 cells/mL, and add 30 μL/mL ice-cold Glusulase ® (see Note 16). 

8. Incubate cells with very gentle shaking (80 rpm; see Note 17) at 28 ºC until 

more than 90–95% of them are protoplasts (∼45 min to 1 h). Monitor the degree 

of protoplast formation with a phase-contrast microscope (see Note 18). 

9. Harvest protoplasts by very gentle centrifugation at 600g for 15 min. Decant the 

supernatant carefully (see Note 19). 

10. Gently wash the protoplast pellet with 150 mL of a 1 M sorbitol solution (see 

Note 20). Centrifuge 15 min at 600g and decant the supernatant carefully. 

11. Repeat this step two more times to eliminate any trace of Glusulase ® (see Note 21). 

12. Remove a small aliquot of the protoplast preparation for use in Subheading 3.2 

to evaluate enzymatic activity of alkaline phosphatase (see Note 22). 

3.1.2. Active Cell Wall Regeneration of Yeast Protoplasts (see Notes 11 

and 17) 

1. Gently resuspend the protoplasts in the regenerating buffer supplemented with 

1 M sorbitol to a density of 3×108cells/mL (see Notes 20 and 23). 

2. Incubate protoplasts with gentle rotary shaking (80 rpm) at 28 ºC for 2htoinduce 

their active cell wall regeneration.


3.1.3. Recovery of Proteins Secreted from Regenerating Protoplasts 

(see Notes 17 and 24) 

1. Centrifuge the regenerating protoplasts at 600g for 15–20 min at 4 ºC, and collect 

the supernatant carefully (see Note 19 and 25). 

2. Add the protease inhibitor mix. 

3. Filter the supernatant through a 0.22-μm pore-size nitrocellulose filter without 

any external device in ice at 4 ºC (see Note 26). 

4. Remove a small aliquot of cell-free culture filtrate to further monitor the cell lysis 

by quantitative determination of alkaline phosphatase (see Subheading 3.2). 

3.1.4. Concentration of Proteins Secreted from Regenerating 

Protoplasts (see Note 24) 

1. Wash an YM-10 Diaflo ® ultrafiltration membrane by floating it skin (glossy) side 

down in a beaker of water for 1h(see Note 27). Change water at least three 

times. 

2. Concentrate the cell-free culture filtrate by ultrafiltration using the washed YM-10 

ultrafilter (see Fig. 3A and Notes 10 and 28). 

3. Dilute with 300 mL water, and concentrate again. Repeat this step three more 

times to eliminate any trace of sorbitol (see Note 29). 

4. Remove the ultraconcentrated protein solution (molecular weight fraction above 

10,000 Da) from the ultrafiltration cell (see Fig. 3B and Note 30). 

5. Quick-freeze the ultraconcentrated protein solution at –80 ºC (see Note 31), and 

concentrate it by lyophilization (see Note 32). 

6. Resuspend the freeze-dried protein sample in a small volume of water (see 

Note 33), and quantify the protein content by standard protein determination 

methods. 

3.2. Assay of Cell Wall Protein Purity: Alkaline Phosphatase Assay 

(see Note 34) 

The method given below outlines an easy and prompt screening procedure 

to check the efficiency, purity, and quality of the collection of proteins secreted 

from yeast protoplasts in active cell wall regeneration before performing further 

proteomic analyses and establishing any interpretation of the results. This is 

based on measurement of enzymatic activity of alkaline phosphatase, which is 

exclusive to the cytosol in yeasts and thus exploited as a marker of cytosolic 

contamination. This intracellular enzyme catalyzes the cleavage of phosphate 

ester bonds from many compounds (including the chromogenic phosphatase 

substrate used in this protocol, p-nitrophenylphosphate; PNPP) under alkaline 

conditions and an optimum temperature of 37 ºC. If there is protoplast lysis 

during sample preparation and, therefore, cytosolic contamination (containing 

alkaline phosphatase, among other intracellular proteins) into the sample of


Fig. 3. Schematic representation of a stirred ultrafiltration cell on a magnetic stirrer. 

Pressurized stirred ultrafiltration cells can efficiently be used to simultaneously concentrate 

and desalt samples of dilute proteins (from initial volumes of 3 mL–2 L to final 

concentrate volumes of 50 μL–60 mL). In the procedure described here, the cell-free 

culture filtrate is forced through an YM-10 Diaflo ® ultrafilter, located on a polymer 

grid at the bottom of the cell (see Note 28). Protein sample is separated into two groups 

according to molecular weight and size (see Note 27). The ultraconcentrate (molecular 

weight fraction above 10,000 Da) is subjected to lyophilization or centrifugal microconcentration 

(see Note 32) before carrying out further proteomic analyses. The protein 

content of the ultrafiltrate (molecular weight fraction below 10,000 Da) can also be 

quantified by standard protein determination methods to evaluate the final protein 

recovery.


wall protein precursors, then alkaline phosphatase will hydrolyze PNPP (which 

is colorless), releasing p-nitrophenol (PNP, which is yellow-colored at alkaline 

pH but colorless at acidic pH values) and inorganic phosphate (see Fig. 4). 

Accordingly, only protein solutions with intracellular contamination will turn 

yellow following the addition of PNPP. The enzyme activity of alkaline 

phosphatase can consequently be detected and quantified by spectrophotometry. 

Fig. 4. Biochemical principle of the alkaline phosphatase assay. The pnitrophenylphosphate 

(PNPP) is a chromogenic substrate for several phosphatases, such 

as acid phosphatases and alkaline phosphatases. Alkaline phosphatase hydrolyzes this 

artificial substrate, which is colorless, at alkaline pH values (pH = 9.7) and 37°C to form 

p-nitrophenol (PNP), which is yellow-colored in basic solutions and can be measured 

at 410 nm on a spectrophotometer. The intensity of the yellow color correlates with the 

amount of PNP issuing from the hydrolysis of PNPP catalyzed by phosphatase alkaline, 

according to Beer’s law. In contrast, acid phosphatase hydrolyzes PNPP under acidic 

conditions (pH = 4.8) and 37°C leading to PNP, which is colorless at acidic pH values.


The present method is extremely sensitive, because very small amounts of 

alkaline phosphatase generate sufficient PNP after an adequate incubation 

period, bringing a measurable color about. This protocol is modified from that 

of Cabib et al. (21), which was developed for rapid checking of colonies with 

lysed yeast cells (embedded into a gelled medium supplemented with PNPP at 

pH 9.7). These acquired a yellow color on and around colonies as a result of 

the release of alkaline phosphatase. 

Interestingly, the present assay is not affected by the acid phosphatases, 

which are located in the outermost and innermost cell wall layers of the yeast 

cells (1) and, therefore, detected in intact cells (see Fig. 4) . This is because 

of the fact that these extracellular enzymes are only able to hydrolyze PNPP 

under acidic conditions (optimum pH = 4.8) and the pH of the reaction buffer 

is highly alkaline (pH = 9.7). 

1. Resuspend the aliquot of protoplasts (see Subheadings 3.1.3 and Note 22)in1mL 

of ice-cold lysis buffer. Vortex for 1 min, and cool on ice for 1–2 min. Repeat this 

step until complete cell breakage (monitored with a phase-contrast microscope). 

Remove cell debris from the homogenate by centrifugation at 12,500g for 10 min 

at 4 ºC. Collect the supernatant and use it as a source of alkaline phosphatase for 

the positive control (see Note 35). 

2. Use known amounts of the reaction product PNP in NaCl-glycine buffer (10–110 

μg/mL) to obtain a standard curve. Mix thoroughly by careful vortexing. Measure 

and record the absorbance at 410 nm in the spectrophotometer. Use the same 

plastic cuvet for all samples, starting with the most dilute sample. 

3. Include controls of (1) NaCl-glycine buffer alone (buffer control), (2) cell-free 

culture filtrate alone (enzyme control), (3) substrate without cell-free culture 

filtrate (substrate control), and (4) substrate with protoplast lysate (positive 

control) as indicated in Fig. 5 to determine superfluous absorbance either 

from other compounds in buffer and cell-free culture filtrate, or from substrate 

breakdown (see Note 36). 

4. Place 500 μL of cell-free culture filtrate of products secreted from regenerating 

protoplasts in a 1.5-mL Eppendorf tube (see Subheadings 3.1.3 and Note 36), 

and add 500 μL of substrate solution (see Note 36). 

5. Vortex carefully and incubate in the dark at 37 ºC for 30 min. Protect from bright 

light. 

6. Stop the enzyme activity with 400 μL of stop solution. Mix carefully and cool 

the tubes to room temperature (25–35ºC). 

7. Measure and record the absorbance at 410 nm in the spectrophotometer. Use the 

same plastic cuvet for all tubes. 

8. Determine enzyme activities in all tubes using the standard curve (10–110 μg 

p-nitrophenol/mL). The units of enzyme (see Note 37) are calculated as: 

U.E. = 14.38 × c/t, where c is the PNP concentration in μg/mL and t is the reaction 

time in min.


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

4. Notes 

1. In all yeast species studied so far, protoplasts embedded into gelled nutrient 

media are always able to regenerate a complete cell wall and revert to normal 

reproducing cells (5). In contrast, the cell wall regeneration of reverting yeast 

protoplasts grown in liquid nutrient media is (1) incomplete in Saccharomyces 

cerevisiae and other budding yeasts, bringing about cells with aberrant walls 

(albeit with all cell wall components) incapable of dividing and inducing 

normal morphogenesis (5,12), but (2) complete in Candida albicans (4,22), 

Schizosaccharomyces pombe (23,24), Nadsonia elongata (5) and Endomycopsis 

fibuliger (5), resulting in normal growing cells. Accordingly, certain physical


factors can actively be involved in the cell wall construction and protoplast 

reversion (5). 

2. The morphology of the de novo cell wall formed on the protoplast surface follows 

that of the protoplast (i.e., spherical shape) (5). On the contrary, the shape of 

new cells originated from the walled protoplasts is at least partly determined by 

the cell wall (i.e., oval shape; see Fig. 1). 

3. The complete reversion of S. pombe, N. elongata and C. albicans protoplasts to 

normal cell walls and growing cells is attained after 12, 20, and 24 h, respectively, 

of incubation in osmotically stabilized liquid nutrient media (4,5,23). 

4. To promote optimal regeneration of protoplast cell walls, it is convenient to 

use a 10× dropout powder solution that has been autoclaved separately or filter 

sterilized, and added to the remaining ingredients after autoclaving. 

5. To avoid precipitation of MgSO 4 and K 2HPO 4, these should be autoclaved 

separately or filter sterilized, and added to the other ingredients after autoclaving. 

6. The PMSF can also be solubilized in ethanol, methanol and 1,2-propanediol; and 

is unstable in aqueous solution. PMSF inhibits serine proteases (e.g., trypsin, 

chymotrypsin, and thrombin) and thiolproteases (e.g., papain). It is added to 

reduce possible proteolytic processes (see Table 1). 

7. Antipain is also soluble in methanol and dimethylsulfoxide (DMSO). It inhibits 

papain and trypsin and, to a lesser extent, plasmin. It is added to reduce possible 

proteolytic processes (see Table 1). 

8. The inhibition specificity of leupeptin is of broad spectrum. It inhibits serine 

and thiol-proteases. It is added to reduce possible proteolytic processes (see 

Table 1). 

9. Pepstatin is insoluble in water. It inhibits acid proteases (e.g., pepsin, chymosin, 

cathepsin D and renin, among others). It is added to reduce possible proteolytic 

processes (see Table 1). 

10. It is important not to use compressed air because it can bring about (1) large 

pH changes as a result of carbon dioxide dissolution, and/or (2) oxidations in 

sensitive protein solutions. 

11. Perform all procedures from this subheading under sterile conditions. Use sterile 

centrifuge bottles. 

12. Liquid cultures should be grown in a flask that is at least 4–5 times larger than 

the culture volume. 

13. The cell density of inoculum and incubation time should be adjusted according 

to the yeast strain. It is important that the yeast culture is in mid-log phase 

growth (∼ OD 600nm= 4) because (1) yeast cells are more susceptible to wall 

lytic enzymes (Glusulase ® treatment) for complete protoplasting than those in 

stationary phase growth and (2) there is sufficient biomass accumulation. 

14. This step is critical for a successful protoplast preparation because mercaptoethanol 

pretreatment facilitates the subsequent action of wall lytic 

enzymes (Glusulase ® treatment) by (1) breaking disulfide bonds of cell 

wall proteins and (2) slightly disorganizing the cell wall. In fact, recent 

studies have demonstrated that a better protoplasting effect is attained in

Table 1 

Protease inhibitors 

Effective 

concentrations Stock solutions Soluble in Notes 

Inhibitor Specificity 

Isopropanol Ethanol 

Methanol 

17 mg/mL in 

isopropanol 

17–174 μg/mL (0.1–1 

mM) 

PMSF Serine proteases 

Thiolproteasesa Stable ∼1 month 

at –20 ºC 

1, 2-propanediol 

Water 

Methanol 

2–50 μg/mL 5 mg/mL in 

water 

Antipain Trypsinb Papainc DMSO d 

Water Stable ∼6 months 

at –20 ºC 

Stable ∼1 week at 

+4 ºC 

Methanol 

Ethanole Stable ∼1 week at 

+4 ºC 

0.5–2 μg/mL 5 mg/mL in 

water 

Leupeptin Serine proteases 

Thiolproteases 

Pepstatin Acid proteases 0.7–1 μg/mL 2.5 mg/mL in 

methanol 

a Reversible by dithiothreitol (DTT) treatment. 

b Serine protease. 

c Thiolprotease. 

d Dimethylsulfoxide. 

e Sit overnight.


-mercaptoethanol-pretreated yeast cells than in nonpretreated cells (25). 

Centrifugation for isolation of -mercaptoethanol-treated yeast cells should be 

carried out with the brake in 1.5. 

15. 1 M sorbitol is used as an osmotic stabilizer. In general, sugars and sugar alcohols 

(e.g., sorbitol, mannitol) are consistently used for yeast protoplast formation and 

reversion, whereas MgSO 4 and KCl are commonly chosen for preparation and 

regeneration of protoplasts from filamentous fungi. 

16. Viable protoplasts can be achieved by controlled autolysis of the yeast cell wall 

using snail or microbial lytic enzymes (see Table 2) (25). Glusulase ® is a 

preparation of the intestinal juice of the Roman garden snail Helix pomatia, and 

consists of a mixture of lytic enzymes (-glucuronidase, sulfatase, and cellulase). 

It has proven particularly useful for obtaining protoplasts (i.e., for completely 

digesting yeast cell walls) in nearly all yeast species (5). Sterilized tweezers 

should be used to take the lid off the Glusulase ® bottle. It is important not to do 

it with the hands. Gently mix Glusulase ® with cell suspension (without shaking 

it). 

17. Perform all procedures from this point on with very gentle shaking. It is very 

important to handle the protoplasts gently in this protocol, because yeast protoplasts 

are extremely fragile. This can prevent further intracellular contamination 

into the sample of proteins secreted from actively reverting protoplasts. 

18. In general, 90–95% of protoplasts should be routinely achieved. The incubation 

time and amount of Glusulase ® needed to attain protoplasts (i.e., complete yeast 

cell wall lysis) can vary with yeast strain and growth stage and may therefore 

be necessary to adjust the incubation time and Glusulase ® amount given in the 

protocol. The degree of protoplast formation is assessed under a phase-contrast 

microscope by: 

a. Counting spherical cells. 

b. Observing cell lysis in hypotonic solution (e.g., after the addition of water; 

see Fig. 6). The number of osmotically-resistant cells (non-protoplasted cells) 

is determined by diluting an aliquot of the cell suspension after and before 

Glusulase ® treatment in water and plating on YPD plates (and growing at 

28–30 ºC). The ratio of CFUs after to before treatment is then calculated to 

estimate the efficiency of protoplast formation. 

19. Perform all centrifugations for protoplast isolation with the brakes off. It is 

important to decant the supernatant carefully, because the protoplast pellet is 

less compact than the preceding cell pellets. 

20. Protoplast resuspension is difficult and sticky. For this reason, a small volume 

should first be used to resuspend the protoplasts by gently swirling liquid across 

the surface of the pellet. Then add more solution until reaching the correct final 

volume. 

21. It is important to wash the protoplasts several times (before regenerating protoplast 

walls) to remove sulfatases, and on the whole any enzymatic activity,

Table 2 

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

from yeast cells 

Types Source Location Composition Commercial name References 

(10, 14, 26) 

Glusulase ® - Du 

Pont; NEN Life 

Science Products, 

– -glucuronidase 

– sulfatase 

– cellulase 

Gut juice 

(digestive 

enzymes) 

Snail enzymes Helix pomatia 

(the Roman 

garden snail) 

Boston, MA, US 

Not available (25, 27) 

Achatina achatina 

(the giant African 

(20, 28) 

Zymolyase 20T ® 

– MP 

Biomedicals, 

Aurora, Ohio, US 

– Seikagaku 

Corporation, 

Tokyo, Japan 

– Miles 

Laboratories, 

Elkhart, Ill., US 

Gut juice 

– -glucuronidase 

(digestive 

– endo--glucanase 

enzymes) 

– arylsulfatase 

Culture fluid – -1,3-glucan 

laminaripentaohydrolase 

– -1,3-glucanase 

– protease 

– mannase 

– amylaseb – xylanaseb – phosphataseb snail) 

Arthrobacter 

luteus (bacterium) 

Microbial 

enzymes a 

a Very expensive for use in industrial processes. 

b Trace.


Fig. 6. Evaluation of the efficiency of protoplast formation. When an osmotically 

stabilized protoplast/cell suspension is diluted with water, the extracellular medium 

becomes hypotonic to cytosol (i.e., the free water concentration is greater outside the 

cell). This results in a passive movement of free water from extracellular medium to 

cytoplasm (an influx of free water). Consequently, protoplasts gain water, swell and 

burst, whereas nonprotoplasted cells do not undergo cytolysis but an increase in turgor 

pressure because of the presence of the cell wall (see Note 18). In isotonic solutions, 

the movement of water into and out of the cell maintains equilibrium. 

present in the Glusulase ® preparation that may modify the protein precursors 

secreted into the medium. 

22. The protoplast preparation can be lysed to release intracellular proteins and used 

as a source of alkaline phosphatase (positive control; see Subheading 3.2). 

23. Lee medium is reliably used as a regenerating buffer for Candida spp. (9,17), 

whereas complete minimal (CM) medium is preferred for Saccharomyces spp. 

(10). These regeneration media are chosen because they are rich and chemically


defined media. Hence, there are no substances in their composition that interfere 

in the subsequent proteomic analyses of the secreted proteins. YPD medium is 

a complex medium, and cannot be used because it contains proteins (from the 

yeast extract), which could result in misidentifications. 

24. Perform all procedures from this point on at 4 ºC (in a cool room at 4 ºC) 

with precooled solutions, reagents and apparatus to avoid undesirable proteolytic 

activity. 

25. It is probable that the protoplast pellet is resuspendend when there is a small 

volume of the supernatant left before being carefully collected. If this happens, 

then centrifuge once more at 600 g for 20 min, and gently collect the supernatant 

again. 

26. If there is not enough time to finish the entire protocol in one laboratory period, 

the procedure can be stopped after filling the filtration unit with the centrifuged 

medium (supernatant) and placing it in a container with ice in a cool room at 

4 ºC. Do not use any external device, because this may facilitate protoplast lysis 

and subsequent intracellular contamination in the sample of secreted proteins. 

27. The YM-10 Diaflo ® ultrafiltration membrane had a pore size of 10,000 Da 

to yield molecular weight fractions above and below 10,000 Da. It should be 

handled carefully and only at the edge. The washing step is important to remove 

preservative substances (e.g., sodium azide). 

28. Place the YM-10 membrane in an ultrafiltration cell, skin (glossy) side toward 

cell-free culture filtrate, and then fill the cell with the culture filtrate. Place it on a 

magnetic stirrer, and connect the inlet line to a regulated nitrogen pressure source 

(see Note 10 and Fig. 3A). Pressurize the cell and pressure-check following the 

supplier’s instructions. Turn on the stirrer, and adjust the stirring rate. When 

the ultrafiltration is accomplished, depressurize and continue stirring for a few 

minutes to increase protein recovery (see Fig. 3B). 

29. It is essential to remove any trace of sorbitol because this interferes in the protein 

resolution of subsequent proteomic analyses. 

30. The ultrafiltration membrane should be washed with 1–2 mL of water to elute 

proteins potentially sticking to it. Add this volume to the ultraconcentrate. To 

reuse the ultrafilter, rinse it with 1–2 M NaCl, and then flush it through with 

water. Store the ultrafilter in 10% ethanol at 4 ºC. 

31. The ultraconcentrate can also be frozen quickly into liquid nitrogen. 

32. Centrifugal microconcentration techniques can alternatively be used to reconcentrate 

the ultraconcentrated protein sample. 

33. The protein sample can be used directly in two-dimensional gel electrophoresis 

(see Fig. 7) (10,13,14,17). Alternatively, freeze-dried protein preparation can be 

resuspended in a small volume of buffer suitable for the subsequent proteomic 

analyses. 

34. This assay should be handled throughout with caution, because p-nitrophenol 

is a skin irritant. Wear gloves and a laboratory coat. Duplicates of each tube 

should be performed.


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

wall regeneration. 

35. The clarified supernatant (protoplast lysate) can be stored at –80 ºC and used in 

future assays. 

36. This assay can also be carried out on a microtiter plate, using 100 μL, rather 

500 μL, of each component. 

37. Although substantial enzyme activity is found in the absence of Mg 2+ ions, it 

is convenient to use 1.5 mM MgCl 2 or1mM MgSO 4 in the substrate solution, 

because these divalent cations stimulate alkaline phosphatase activity. 

38. One unit of enzyme (U.E.) is defined as the amount of enzyme that will produce 

1 nmol of p-nitrophenol per minute.



We thank the Merck, Sharp & Dohme (MSD) Special Chair in Genomics 

and Proteomics, European Community (STREP LSHB-CT-2004-511952), 

Comunidad de Madrid (S-SAL-0246/2006) and Comisión Interministerial de 

Ciencia y Tecnología (BIO-2003-00030 and BIO-2006-01989) for financial 

support of our laboratory. 

References 

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J. P. (1998) Cell wall and secreted proteins of Candida albicans: identification, 

function, and expression. Microbiol Mol.Biol Rev. 62, 130–80. 

2. Klis, F. M., Boorsma, A., and de Groot, P. W. (2006) Cell wall construction in 

Saccharomyces cerevisiae. Yeast 23, 185–02. 

3. Osumi, M. (1998) The ultrastructure of yeast: cell wall structure and formation. 

Micron. 29, 207–33. 

4. Nishiyama, Y., Aoki, Y., and Yamaguchi, H. (1995) Morphological aspects of 

cell wall formation during protoplast regeneration in Candida albicans. J.Electron 

Microsc. 44, 72–8. 

5. Necas, O. (1971) Cell wall synthesis in yeast protoplasts. Bacteriol.Rev. 35, 149–70. 

6. Takagi, T., Ishijima, S. A., Ochi, H., and Osumi, M., (2003) Ultrastructure and 

behavior of actin cytoskeleton during cell wall formation in the fission yeast 

Schizosaccharomyces pombe. J.Electron Microsc. 52, 161–74. 

7. Rico, H., Carrillo, C., Aguado, C., Mormeneo, S., and Sentandreu, R. (1997) Initial 

steps of wall protoplast regeneration in Candida albicans. Res.Microbiol. 148, 

593–603. 

8. Kapteyn, J. C., Dijkgraaf, G. J., Montijn, R. C., and Klis, F. M. (1995) Glucosylation 

of cell wall proteins in regenerating spheroplasts of Candida albicans. 

FEMS Microbiol Lett. 128, 271–77. 

9. Elorza, M. V., Marcilla, A., Sanjuan, R., Mormeneo, S., and Sentandreu, R. (1994) 

Incorporation of specific wall proteins during yeast and mycelial protoplast regeneration 

in Candida albicans. Arch.Microbiol. 161, 145–51. 

10. Pardo, M., Monteoliva, L., Pla, J., Sanchez, M., Gil, C., and Nombela, C. (1999) 

Two-dimensional analysis of proteins secreted by Saccharomyces cerevisiae regenerating 

protoplasts: a novel approach to study the cell wall. Yeast 15, 459–72. 

11. Pitarch, A., Sanchez, M., Nombela, C., and Gil, C. (2002) Sequential fractionation 

and two-dimensional gel analysis unravels the complexity of the dimorphic fungus 

Candida albicans cell wall proteome. Mol.Cell Proteomics 1, 967–82. 

12. Klis, F. M. (1994) Review: cell wall assembly in yeast. Yeast, 10, 851–69. 

13. Pardo, M., Ward, M., Bains, S., et al. (2000) A proteomic approach for the study 

of Saccharomyces cerevisiae cell wall biogenesis. Electrophoresis 21, 3396–3410. 

14. Pitarch, A., Jimenez, A., Nombela, C., and Gil, C. (2006) Decoding serological 

response to Candida cell wall immunome into novel diagnostic, prognostic, and


therapeutic candidates for systemic candidiasis by proteomic and bioinformatic 

analyses. Mol.Cell Proteomics 5, 79–96. 

15. Kobori, H., Yamada, N., Taki, A., and Osumi, M. (1989) Actin is associated 

with the formation of the cell wall in reverting protoplasts of the fission yeast 

Schizosaccharomyces pombe. J.Cell Sci. 94, 635–46. 

16. Tanaka, H., Kamogawa, T, Aoyagi, H., Kato, I. and Nakajima, R. (2000) Invertase 

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alginate gel beads. J. Biosci. Bioeng. 89, 498–500. 

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pombe. Fungal.Genet.Biol. 24,178–206. 

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21 

Sample Preparation Procedure for Cellular Fungi 

Alois Harder 

Summary 

A crucial step in quantitative proteomics is an artefact free and reproducible sample 

preparation protocol, which has to be adapted and optimized to nearly all types of cells. 

Here we provide a sample preparation method for quantitative proteomics of cellular fungi. 

Two different protein extraction methods were compared with focus on reproducibility, 

minimized proteolytic degradation and protein losses during the sample preparation. 

In the first preparation the cells were lysed by sonication followed by protein solubilization 

in “standard” lysis buffer. The second preparation was performed with a SDSpresolubilization 

step followed by sonication and further boiling, before diluting the 

sample with lysis buffer. We have shown that the sample preparation for cellular fungi 

is performed with maximum protein solubilization, higher reproducibility and a reduced 

proteolytic activity by including a SDS-presolubilization step in the sample preparation 

protocol. 

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

yeast. 


Fungal cells have nowadays found their way in nearly all areas of biochemistry 

and pharmacy. Famous examples are active medicaments like PerenterolTM with Saccharomyces boulardii as the active ingredient (1,2), yeast host cell 

systems like the vaccine production cycle or diverse secondary metabolite 

discovering systems like the Novobiocin synthesis to name only few (3,4). To 

push those applications or detect new fungal targets or metabolites, a solid 

basis of information in cell growth, cell cycle, gene- and protein data are 

essential. Whereas cell growth and cycle can be researched in conceivable 



265

266 Harder 

time, gene and protein data are missing for the majority of the fungi. The 

finished and currently running fungal genome sequencing projects are mainly 

concentrating on model- or pathogen strains: Saccharomyces cerevisiae, Ashbya 

gossypii, Aspergillus fumigatus, Candida parapsilosis, Candida dubliniensis, 

Pneumocystis carinii, Phytophthora infestans, and Schizosaccharomyces pombe 

have been sequenced or will be finished soon and are available for the 

community (5,6). 

For identification of fungal proteins, genome data of the organism to be 

researched has to be available. Although the fungal protein databases are 

growing, the listed open reading frames are far from their complete and correct 

functional characterization. 

However,withthesequencedfungalgenomes,theavailableproteindataandthe 

homology – search programs with a solid foundation for further proteome analysis 

where eumycotic cells are present. Fungal cells are easy to cultivate, to manipulate, 

and in most cases are non-toxic, but the preparation of a robust and artefactfree 

protein extract for a proteomics approach is a challenge. Major problems in 

fungal proteomics result from the cells strong proteolytic activity. Most of the 

fungalproteases,intensivelyresearchedintheeumycoticmodelorganismSaccharomyces 

cerevisiae, are located in the cytosol and the vacuole. Few proteases 

are located in the membrane, the endoplasmic reticulum, the mitochondria, 

and the Golgi complex (7,8). These include endoproteinases, carboxypeptidases, 

aminopeptidases, and dipeptidyl-aminopeptinase. By activation of these 

proteasesproteinmodificationswilloccurandcanruleoutquantitativeproteomics 

research (9–10,11). 

Of the many cellular proteases, the lumenal vacuolar proteases comprise the 

major source of problems for protein analysis and proteome research. Endoproteinase 

A and B (PrA and PrB), carboxypeptidases Y and S, aminopeptidase 

I (LAP IV), and the aminopeptidase yscCo (ApCo) are found soluble in the 

vacuole (12,13). Polypeptide inhibitors of these vacuolar proteases are found 

in the cytosol (e.g., inhibitor for PrB is I B; inhibitor for yscCo is yscCo F). 

By preparing the crude extract all cell compartments are broken up, meaning 

that corresponding protease inhibitors will bind to their substrate by forming 

an inactive complex—in this state proteolytic activity is inhibited. As reported 

from Jones et al. the fungal proteases PrA and PrB, can be activated out of the 

crude extract by lowering the pH to a level of 4–5. The reason for this activation 

might be the hydrolysis of the polypeptide inhibitors (7,14). Further research 

on protease deficiency mutants showed that the addition of denaturating agents 

activates proteolysis by removing the corresponding inhibitor from the protease 

molecule. Due to the exigency in proteomics research, for working in strict

Sample Preparation Procedure for Cellular Fungi 267 

denaturation conditions, the necessity of a “non standard” sample preparation 

procedure for fungal cells is obvious. 

The fungal sample preparation for proteomics research has to 

• (BL)be carried out in strong denaturating conditions for its stability during 

the 2D - electrophoretic separation 

• display robustness to guarantee reproducibility by repeating the experiments 

• solubilize low abundant protein as well as high and low molecular weight 

proteins in all pH ranges 

• effectively inactivate the strong proteolytic activity of fungal cells 

2. Materials 


1. Standard aerobic incubator with an inbuilt horizontal rotary shaker. 300 mL 

Erlenmeyer flasks with silicone plugs are used for aerobic cell cultivation. 

2. Horizontal rotary shaker (e.g., HS 260, IKA). 

3. Incubation mask (e.g., Certomat H/HK). 

4. S. cerevisiae haploid wild type strain N318C (“Deutsche Sammlung für Mikroben 

und Zellinien”, DSMZ Braunschweig, Germany). 

5. Defined synthetic YNB medium (Difco Labs, San Francisco CA, USA) is 

dissolved to a concentration of 6.7 g/L in distilled water (

268 Harder 

2.4. Scintillation Counting and Protein Assay 

1. For S35-labelled cells, a scintillation counting is performed: 4.5 mL POPOP 

solution is required for one vial; 2 × 4 cm cut filter paper is used for the counting 

(see Note 1). As a calibration standard a C12 isotope (GE Healthcare) is applied. 

2. Lowry protein assay kit for SDS containing samples. 

3. The Bradford protein assay kit (Bio-Rad) for samples dissolved in lysis buffers. 

4. BSA (Bio-Rad) is used as a protein standard for both protein assays. 

UV-spectral photometer e.g., Beckman DU-64 (see Note 2). 

3. Methods 

3.1. S. cerevisiae Cell Culture 

1. The S. cerevisiae haploid wild type strain N318C is cultivated at 30 °C in defined 

synthetic YNB medium. First a preculture is grown, in which about 105 cells are 

inoculated in 30 mL culture medium (see previous section) and grown to a cell 

density of OD 0.6 measured at =610nm (see Note 3). 

2. Then the main culture (30 mL, 300-mL Erlenmeyer flasks) is started with an 

inoculum from 10 μL out of the preculture. The cells are harvested by centrifugation 

(5,000g, 15 min) in the late midlogarithmic phase, when the culture reached 

an OD of 1.1 followed by washing in ice-cold sterilized water (see Note 4). 

3. A control smear is performed on a Sabouraud agar plate to check for cellular 

contaminations. The incubation of the agar plate was done for 48 h at 37 °C in 

sterile conditions. 

4. Harvested yeast cells are then transferred into individual 1.5 mL Eppendorf tubes 

and stored at –78 °C, if not processed immediately. 

3.2. S-35 In Vitro Labeling (optional) 

1. The S-35 in vitro labelling can be done additionally to the standard cell growth 

procedure. For the in vitro S 35 - labelling experiment the main culture is grown 

to an OD of 1.0 (=610 nm) 

2. Then 10 mL of the cell suspension is inoculated with 100 μCi and kept shaking for a 

quarter of the fungal generation time (for S. cerevisiae in YNB – medium→ 30min). 

3. After the S-35 incubation the cells are harvested by centrifugation (5 min, 5,000g) 

and the supernatant is discarded to the radioactive waste. The amount of protein, 

which will be extracted out of the obtained S-35 labeled pellet will be approx 

5 mg protein solubilized in lysis buffer. Only methionine deficient mediums are 

suited for S-35 in vitro labeling. 

3.3. Cell Disruption and Protein Solubilization 

1. The yeast cell pellet from 10 mL culture OD 1.1 (see Section 21.3.1) is resuspended 

in 200 μL hot (95 °C) SDS buffer for presolubilization (see Note 5 and 6).


2. After brief vortexing the hot suspension is sonicated 10 times for 1s (60 W, 20 

kHz). The sonication has to be performed in such way that the probe is dipping 

as deep as possible in the suspension and not touching the sample cup during 

the agitation time. A 1.5-mL cup (conic bottom) is used for maximum abrasive 

agitation. Avoid foaming of the sample during this process (see Note 7). 

3. After cell lyses the sample is boiled additionally for 5 min and then cooled down 

in an ice cold water bath to a sample temperature of maximum 20 °C (see Note 

8 and 9). 

4. Then the suspension is diluted with 500 μL lysis buffer and kept shaking for 20 

min at room temperature. 

5. After spinning down for 5 min at 10,000g, the protein concentration is measured 

from the clear supernatant (see Section 21.3.4). 

6. The clear supernatant is stored at –78 °C or subjected to 2D-electrophoresis. 

An example of four replicate gels (IPG 4-7, 250 μg protein load, S. cerevisiae, 

SDS presolubilisation) is shown in Fig. 2 compared to a “standard preparation 

protocol” without the SDS presolubilization step (Fig. 1). 

1 97kD 2 

67kD 

45kD 

29kD 

21kD 

12kD 

6kD 

3 4 

Fig. 1. Displaying four replicate 2D gels (IPG 4-7L, 250 μg protein load, fluorescence 

stain) from the yeast proteome (S. cerevisiae) performed without the SDS - 

presolubilization step (see Note 12 and 13).

270 Harder 

5 6 

97kD 

67kD 

45kD 

29kD 

21kD 

12kD 

6kD 

7 8 

Fig. 2. Displaying four replicate 2D gels (IPG 4-7L, 250 μg protein load, fluorescence 

stain) from the yeast proteome (S. cerevisiae) performed with the SDS - presolubilization 

step (see Note 14). 

7. An approximate 7 μg/μL protein concentration can be expected in that obtained 

700-μL sample volume (see Note 10) 

3.4. Scintillation Counting and Protein Assay 

1. 1. For S35 - labeled cells, the protein concentration is measured by removing 

1 μL of the lysis buffer extract to a filter. After drying the sample, the filter is 

inserted into a counting vial, which is filled with 4.5 mL POPOP - solution. The 

scintillation counter is calibrated to a C12 – standard, which was first solubilized 

in lysis buffer. The scintillation counting should be repeated three times for each 

sample. Approximately 3 × 106 cpm should be loaded onto an IPG strip 4–7, 

2.5×106counts onto the gradient 3–10. This method is working with and without 

the SDS presolubilization step (see Note 11). 

2. 20 μL of the SDS extract are used for the Lowry protein assay. 5 μL of the lysis 

buffer extract are taken for the Bradford protein assay.


4. Notes 

1. For S35 counting: after an appropriate amount of sample is pipeted onto the 

filter, the sample has to be dried for at least 30 min, due to water residues 

suppress the radio signal significantly. 

2. The determination of the protein content is optional, but an optimum and reproducible 

protein load is essential for successful protein quantification. 

3. Significantly increased proteolytic activity in fungi is observed in cells grown in 

minimal medium compared to cells grown in full medium, in cells harvested in 

their stationary phase (increases proteases activity to a level at least 100 times 

that of log phase cells), in vital cells stored or grown in nitrogen starvation and 

in the presence of peptone in the culture medium. 

4. When you are researching fungal stress responses, washing the pellet with ice 

cold water will induce a deficient cell response. In that case the cell washing 

can be performed with 15 °C water as well. 

5. The whole procedure is carried out in one single Eppendorf tube. By establishing 

the yeast sample preparation procedure main focus was given to its 

reproducibility, robustness, and maximum protein solubility with respect to the 

yeast’s peculiarities like proteolytic activity and cell wall constitution. 

6. Dried pellets (for measuring the amount of cells) are not suited for further 

proteome analysis, due to heat shock responses of the cells. Homologue pellets 

should be used for protein extraction and 2D separation. 

7. The correct sonication will provide a quantitative cell disruption and protein 

disaggregating by simultaneously cracking the fungal DNA and RNA strands, 

which otherwise can produce horizontal streaks in the 2D PAGE. The DNA and 

RNA fragments should become visible as a thin white foam after the sonication 

step. 

8. By presolubilization in hot SDS buffer fungal proteases are denatured and inactivated 

during the cell breakage. Boiling the sample in SDS will increase protein 

solubility, especially for the high molecular weight and hydrophobic proteins, 

due to the high affinity of SDS to proteins in solution. 

9. It is crucial to cool down the SDS-boiled sample below 20 °C, to avoid carbamylation 

reactions with the urea containing lysis buffer. The final SDS concentration 

should not exceed 0.25% in the extract to be applied onto the IPG strip, due to 

the interference of the SDS with the isoelectric focussing, so be sure that during 

the SDS boiling step the total volume is kept. 

10. By starting with about 1×108cells (S. cerevisiae) the obtained protein concentration 

will be approximately 7μg/μL in a 700 μL volume. 

11. The protein concentration within this preparation method can be measured with 

the Lowry Kit after the SDS-presolubilization or with a scintillation counter (by 

S-35 labeled cells) out of the final extract. Determination of the dry weight or 

cell counting after cell harvesting should be done at least one time to check 

purity and approximate amount of cultivated cells.

272 Harder 

12. A major problem in fungal protein extraction is the strong fungal proteolytic 

activity. By applying a standard preparation protocol significant degradation can 

occur (see Fig. 1) and consequently a protein quantification is useless. 

13. In many cases proteolytic activity is not as obvious as in the gels 1–4 (Fig. 1), 

especially when degradation only begins. It can show up in disappearance of 

single spots or in the decrease of the quantities of some spots, which can be 

spuriously interpreted as a cell response. Therefore the complete and irreversible 

inactivation of the mycotic proteome is essential. 

14. Note, that the obtained “crude extract” will include proteins from the cytoplasm, 

the organelles, membrane bound proteins and surface proteins. Integral plasma 

membrane proteins as well as integral cell wall proteins are not extracted with 

that recipe (see Fig. 2). 

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6. [No authors listed]. (1997) The yeast genome directory.Nature 387 (6632 Suppl) 5. 

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8. Groll, M. and Huber, R. (2005) Purification, crystallization, and X-ray analysis of 

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9. McIntyre, J., Podlaska, A., Skoneczna, A., Halas, A., and Sledziewska-Gojska, 

E. (2006) Analysis of the spontaneous mutator phenotype associated with 20S 

proteasome deficiency in S. cerevisiae. Mutat Res. 593(1–2), 153–63. 

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22 

Isolation and Enrichment of Secreted Proteins 

from Filamentous Fungi 

Martha L. Medina and Wilson A. Francisco 

Summary 

Filamentous fungi have been recognized as extraordinary producers of secreted proteins 

and are known to produce novel proteins and enzymes through dispensable metabolic 

pathways. Here, methods are described for the isolation and enrichment of samples of 

secreted proteins from cultures of filamentous fungi for analysis by gel electrophoresis 

and mass spectrometry techniques. These methods can be readily applied to the study of 

differential protein expression and secretion and metabolic pathways in filamentous fungi 

by proteomic approaches. 

Key Words: Exoproteome; extracellular proteins; protein deglycosylation; protein 

precipitation; secreted proteins; secretome; gel electrophoresis. 


Protein secretion plays an important role in filamentous fungi, particularly in 

nutrition, as secreted enzymes degrade complex biological molecules to serve 

as carbon and nitrogen sources. Filamentous fungi are known for their ability 

to secrete a broad spectrum of enzymes, the majority of which are hydrolytic, 

into the extracellular matrix (1). This ability has been widely exploited by 

the biotechnology industry for the production of enzymes for commercial 

and industrial use. Most commonly, filamentous fungi secrete proteins via a 

classical secretory pathway (2) and most, if not all, of the secreted proteins 

are glycosylated, containing modifications such as oligomannose N- and Oglycans 

(3). These attached sugars increase the stability of the secreted proteins 



275

276 Medina and Francisco 

and provide resistance to environmental influences, as well as increase their 

solubility in the culture media. Although many studies of protein secretion 

have been carried out in yeast and animal systems, studies on protein secretion 

by filamentous fungi are limited (4). Typical studies have focused on the 

identification, purification and characterization of secreted proteins, but only 

a few studies have been conducted on the global analysis of fungal extracellular 

proteomes. Proteomic studies on the analysis of secreted proteins from a 

number of fungi, including Aspergillus flavus (5,6), A. oryzae (7), Fusarium 

graminearum (8), and Pleurotus sapidus (9), have recently appeared in the 

literature. Much of the delay in the use of proteomic techniques for the study 

of fungal protein expression in general can be attributed to the fact that there is 

a lack of complete and publicly available genome sequences from filamentous 

fungi. It is expected that as the number of published fungal genomes increases, 

proteomic studies on both intracellular and extracellular proteins will follow. 

Within their high capacity to produce secreted proteins, filamentous fungi are 

able to express and secrete proteins for “dispensable” metabolic functions (10). 

These enzymes participate in pathways that are either not required for growth 

or are only required for growth under a limited range of conditions. These 

changes in protein secretion provide an excellent platform for the systematic 

study of the overall protein secretion expression (secretome or exoproteome) as 

a function of culture conditions. Proteomic analysis using two-dimensional gel 

electrophoresis (2-DE) and mass spectrometry (MS) has proven to be the most 

powerful and sensitive method for the identification of proteins in complex 

mixtures. 2-DE provides an excellent platform to assess differential secreted 

protein expression. Although the number of proteins that can be analyzed by 2- 

DE is still limited to 1,000–2,000 on one gel, the maximum number of secreted 

proteins by a filamentous fungi under a given set of conditions is well within 

this range, as a recent analysis of the genome of Aspergillus niger identified 

only about 400 putative secreted proteins from a total of about 5,100 genes (11). 

Obtaining MALDI-TOF MS data from tryptic digests of gel bands and spots 

and searching against online protein databases can identify 2-DE separated 

proteins. The success of peptide mass fingerprinting depends on the detection of 

a representative set of peptide masses derived from a protein and that the protein 

in question is known (it exists in a protein database). Alternatively, proteins can 

be identified by MS/MS with custom databases that contain sequences of all 

publicly known fungal proteins or by de-novo sequencing using this technique. 

Several technical issues need to be considered when studying the secretome 

of filamentous fungi by proteomic analysis. First, although fungi have the 

ability to secrete large amounts of protein, these proteins are highly diluted in 

the culture medium. Second, glycosylation poses a problem for visualization 

of proteins separated by gel electrophoresis, as glycosylated samples tend to

Isolation of Secreted Proteins from Filamentous Fungi 277 

smear in gels. In addition, glycosylation can protect proteins against proteolysis, 

making the identification of these proteins more difficult as peptide mass 

fingerprinting relies on efficient protease digestion before mass spectrometry 

(12). Also, the attached sugars greatly increase the size of generated peptide 

fragments, adding another layer of complexity in their identification by mass 

spectrometry. These problems can be overcome by concentrating the media, 

most commonly by lyophilization or ultrafiltration, and by removing the 

sugars with commercially available glycosidases or by chemical deglycosylation 

methods. 

Here we describe a general procedure for the isolation and enrichment of 

secreted proteins from cultures of filamentous fungi for proteomic analysis 

by gel electrophoresis and mass spectrometry, including protocols for protein 

deglycosylation. The methods described here can be easily adopted for the 

isolation and characterization of the secretome of filamentous fungi grown 

under varied conditions. 

2. Materials 

2.1. Filtration and Lyophilization 

1. Miracloth (Calbiochem, EMD Biosciences, Inc., San Diego, CA) or No. 2 

Whatman filter paper. 

2. Lyophilizer 

2.2. Centrifugation and Ultrafiltration 

1. Centricon Plus-70 Centrifugal Filter Device (Millipore, Billerica, MA). 

2. Centrifuge 

2.3. Precipitation with Trichloroacetic Acid 

1. 20% (w/v) Trichloroacetic acid (TCA) (Sigma-Aldrich, St. Louis, MO), stored 

at 4°C. 

2. 70% Ethanol, stored at –20°C. 

3. Acetone 

2.4. Precipitation with Methanol and Chloroform 

1. Methanol 

2. Chloroform 

2.5. Enzymatic Deglycosylation 

1. Ultrafree-0.5 centrifugal filter devices with Biomax-5 membranes 5,000 NMWL 

(Millipore, Billerica, MA).


2. Peptide: N-glycosidase F (PNGase F) (New England BioLabs, Inc., Ipswich, MA). 

PNGase F is provided with 10× glycoprotein denaturing buffer (0.5% SDS, 1% 

-mercaptoethanol), 10× G7 reaction buffer (50 mM sodium phosphate, pH 7.5), 

10% Nonidet P-40 (NP-40), and a solution of PNGase F (see Note 1). 

2.6. Chemical Deglycosylation (see Note 2) 

1. Reacti-Vial Reaction Vials (5 mL) (Pierce Chemical Co., Rockford, IL). 

2. Trifluoromethanesulfonic acid (TFMS) (Sigma/Aldrich, St. Louis, MO, USA) 

(see Note 3). 

3. Anisole, anhydrous (Sigma/Aldrich, St. Louis, MO), stored at 4°C. 

4. 60% (v/v) Pyridine solution, stored at –20°C. 

5. Diethyl ether, stored at –20°C. 

6. 95% Ethanol, stored at 4°C. 

2.7. SDS-PAGE 

1. SDS Sample Reducing Buffer: Mix 4.05 mL of deionized water, 1.25 mL 0.5M 

Tris-HCl, pH 6.8, 2.50 mL glycerol, 2.00 mL 10% SDS and 0.20 mL 0.5% (w/v) 

bromophenol blue. 

2. -Mercaptoethanol 

2.8. Two-Dimensional Gel Electrophoresis 

1. 2-DE Buffer: 8M urea, 2% (w/v) CHAPS, 50 mM dithiothreitol, 0.2% (w/v) 100× 

Bio-Lyte 3–10 ampholytes (Bio-Rad Laboratories, Inc., Hercules, CA), 0.001% 

(w/v) bromophenol blue. 

3. Methods 

This chapter describes general-purpose sample preparation methods for the 

isolation and enrichment of secreted proteins from cultures of filamentous 

fungi. The methods described in the following section can be divided into (1) 

isolation and concentration by filtration and lyophilization or ultrafiltration, 

(2) precipitation, and (3) deglycosylation of concentrated protein samples by 

enzymatic (PNGase F) or chemical deglycosylation using TFMS acid (13). 

The samples can then be further analyzed by gel electrophoresis and mass 

spectrometry by established techniques described in other chapters. A schematic 

representation of these protocols is shown in Fig. 1. 

3.1. Isolation and Concentration of Supernatant Broth 

1. After the studied filamentous fungus has been grown in liquid culture media for 

the desired time, the broth containing the secreted proteins is collected by filtration


Fig. 1. Simplified schematic for the isolation and enrichment of secreted proteins 

from cultures of filamentous fungi for proteomic analysis (see Methods section for 

details). 

through a Miracloth or No. 2 Whatman filter paper. Alternatively, the fungal 

mycelia can be separated from the supernatants by centrifugation at 10,000g for 

10 min. (see Note 4). 

2. The supernatants are concentrated by lyophilization (steps 3–5) or ultrafiltration 

(steps 6–8). 

3. For lyophilization, place the filtered supernatants in individual round bottom 

flasks and freeze in liquid nitrogen. (see Note 5).


4. Place the round bottom flasks on the lyophilizer until they are completely dry. 

5. Redissolve the contents of the round bottom flasks in a minimal amount of 

deionized water, and store at –20°C until further analysis. 

6. For ultrafiltration, place the filtered supernatant in a Centricon Plus-70 centrifugal 

filter device to a maximum of 70 mL. 

7. Centrifuge at 3,500g until the desired final volume is achieved. 

8. The concentrated supernatant can be washed with an appropriate buffer (e.g., 

50 mM phosphate buffer, pH 6.0) and reconcentrated to remove excess salts, 

pigments, and metabolites. (see Note 6). 

3.2. Protein Precipitation 

Protein precipitation is an efficient method for the removal of most contaminants, 

including detergents, salts, peptides, lipids, and phenolic compounds 

from protein samples. Either one of the methods described below can be used 

to precipitate the secreted proteins from the concentrated broth supernatant in 

preparation for gel electrophoresis analysis. It should be noted that no method 

will precipitate all proteins and some proteins will be difficult to resuspend 

following precipitation. 

3.2.1. Precipitation with Trichloroacetic Acid (see Note 7) 

1. Pipet 0.3 mL sample of concentrated broth supernatant into a 1.5-mL siliconized 

microcentrifuge tube and add an equal amount of cold TCA solution. 

2. The mixture is incubated for 2hat–20°C to allow the proteins to precipitate. 

After 2 h, allow samples to thaw if lightly frozen. 

3. Centrifuge for 10 min at 14,000g. 

4. The samples are decanted, and 1 mL of cold 70% ethanol is added, vortexed, and 

recentrifuged for 3 min. 

5. Step 4 is repeated 3 times. 

6. To completely dry the secreted protein pellet, add 1 mL of acetone, vortex and 

centrifuge for 1 min. The acetone is decanted, and the pellet is allowed to air dry 

for 30 min. 

7. The dried pellet can be stored at –20°C until further analysis. 

8. For electrophoresis analysis, the pellet is dissolved in SDS-PAGE or 2-DE sample 

buffer, as described below (see Subheading 3.5 and Note 8). 

3.2.2. Precipitation with Chloroform/Methanol 

1. To 100 μL of the concentrated supernatant in a siliconized microcentrifuge tube, 

add 400 μL of methanol, 100 μL of chloroform, and 300 μL of H2O, and mix 

well. 

2. Incubate at 4°C for 5 min and centrifuge at 9,000g at 4°C for 2 min.


3. The upper phase is carefully removed and discarded. Add another 300 μL of 

methanol to the rest of the lower chloroform phase and the interphase with the 

precipitated protein and mix well. 

4. Incubate at 4°C for 5 min and pellet the proteins by centrifugation at 13,000g for 

5 min at 4°C. The supernatant is removed and the protein pellet is dried under a 

stream of air. 

5. The dried pellet can be stored at –20°C until further analysis. 

6. For electrophoresis analysis, the pellet is dissolved in SDS-PAGE or 2-DE sample 

buffer (see Subheading 3.5). 

3.3. Protein Deglycosylation 

3.3.1. Enzymatic Protein Deglycosylation with PNGase F 

1. Concentrate a 500 μL sample of the concentrated supernatant broth to 50 μL 

using a Ultrafree-0.5 with a Biomax-5 membrane (5,000 NMWL) centrifugal 

filter device by centrifugation at 12,000g at 4°C. To desalt the sample, add 400 μL 

of deionized water to the concentrated sample and centrifuge again until a final 

volume of 50 μL is reached. 

2. The concentrated, desalted solution is removed from the centrifugal filter device 

and placed in a 600 μL siliconized microcentrifuge tube. 

3. Enzymatic digestion is carried out as follows and according to the manufacturer’s 

instructions (New England BioLabs, Inc.): 15 μL of denaturing buffer is added to 

the sample and boiled at 100°C for 10 min. To this sample, 3.5 μL of G7 buffer 

and 3.5 μL of NP-40 buffer are added, and the samples are digested with 1,000 U 

of PNGase F (see Note 9) for 18 hours at 37°C. 

4. Following deglycosylation, the samples are precipitated as described above (see 

Subheading 3.3). 

3.3.2. Chemical Protein Deglycosylation 

with Trifluoromethanesulfonic Acid 

1. Glycoprotein samples should be relatively free of salts, minerals and detergents. 

Samples must also be completely dried. Lyophilize protein sample in a 5-mL 

Reacti-vial. 

2. Incubate lyophilized sample with 0.3 mL of cold anisole and 0.6 mL of cold 

TFMS at 0°C in an ice-bath for 4 h under nitrogen with occasional shaking. 

3. The reaction mixture is cooled to below –20°C by placing in a dry ice-ethanol 

bath and neutralized by slowly adding 1.2 mL of cold 60% aqueous pyridine 

solution. 

4. The deglycosylated peptides are freed of reagents and low-molecular weight 

sugars by adding 2 mL of cold diethyl ether. The suspension is vortexed and 

extracted twice with cold ether.


5. The aqueous layer is lyophilized, redissolved in deionized water, and precipitated 

as described in Subheading 3.3. 

3.4. Preparation of Samples for Gel Electrophoresis Analysis (see 

Note 10) 

3.4.1. SDS-PAGE 

1. Add 50 μL of -mercaptoethanol to 950 μL SDS Reducing Buffer before use. 

2. Dissolve the precipitated protein pellet in the appropriate volume of SDS reducing 

sample buffer determined by the size of the gel and the system used. 

3. Boil sample for 4 min. 

3.4.2. Two-dimensional Gel Electrophoresis 

1. Dissolve the precipitated protein pellet in the appropriate volume of 2-DE Buffer 

determined by the size of the IPG strip and the system used for isoelectric focusing 


2. Incubate the sample for2hatroom temperature and remove any insoluble material 

by centrifugation at 10,000g for 10 min. 

4. Notes 

1. PNGase F is an amidase that cleaves between the innermost N-acetylglucosamine 

and asparagine residues of high mannose, hybrid, and complex oligosaccharides 

from N-linked glycoproteins. PNGase F is the enzyme of choice for 

removing most N-linked oligosaccharides. Although secreted proteins could 

contain O-linked oligosaccharides, a recent study done on commercial cellulose 

enzyme preparation from the filamentous fungus Trichoderma reesei demonstrated 

that PNGase F treatment was superior to other enzymatic or chemical 

deglycosylation treatements in terms of yielding peptides through MALDI-MS 

that resulted in actual protein identification when searched against a database 

(12). 

2. For chemical deglycosylation, the use of a commercially available kit is recommended. 

Two of such kits are the GlycoProfile IV, Chemical Deglycosylation 

Kit from Sigma/Aldrich (Sigma/Aldrich, St. Louis, MO, USA) and Glycofree 

Chemical Deglycosylation Kit from ProZyme, Inc. (San Leandro, CA, USA). 

3. Trifluoromethanesulfonic acid is a strong acid, highly corrosive and hygroscopic. 

Protective goggles, laboratory coat and gloves should be worn when working 

with TFMS. 

4. Typically, centrifuging is used to separate cells, in this case mycelia, from the 

supernatants. However, centrifuging does not work as well for filamentous fungi, 

as mycelia do not pack well and can float to the surface of the supernatant; 

therefore, filtering is preferred over centrifugation.


5. Culture supernatants in the round bottom flasks should be frozen by spinning 

the flask while immersing in liquid nitrogen to ensure even distribution of the 

supernatant in the flask. This will allow better lyophilization of the samples. 

6. An alternate method for desalting is dialysis. Concentrated supernatant broth 

can be dialyzed against water or any appropriate buffer at 4°C. 

7. An alternate protocol for precipitation of fungal secreted proteins by 

trichloroacetic acid has been described by Suárez et al. (2005) (14). In this 

protocol, the concentrated, dialyzed and lyophilized culture filtrate is resuspended 

in 20% TCA in acetone containing 0.2% dithiothreitol (DTT), stored at 

–20°C. The suspension is kept at –20°C overnight. The sample is centrifuged at 

16,000g at 4 C for 10 min and the resulting pellet is washed three times with 

acetone containing 0.2% DTT. The supernatants are removed by centrifugation 

and the pellet is dried overnight at room temperature. The pellet is dissolved 

in SDS-PAGE or 2-DE sample buffer before gel electrophoresis analysis (see 

Subheading 3.5). In this protocol, acetone is used to increase the solubility of 

interfering organic compounds and increase protein precipitation and DTT is 

included to prevent protein modification. 

8. TCA precipitation allows for further concentration of the proteins in the sample, 

as well as for removal of non-protein substances, salts and other agents that may 

interfere with electrophoresis separation. Care should be taken to make sure the 

protein pellet is completely dry before adding sample buffer, as any leftover 

TCA will turn the Bromophenol Blue in the sample buffer yellow. 

9. The majority of the proteins secreted by filamentous fungi can have carbohydrate 

contents reaching up to 50% of the total molecular weight of the protein (1). 

This heavy glycosylation is thought to be responsible for the tendency of these 

proteins to show “smearing” on SDS-PAGE gels (12). Enzymatic protein deglycosylation 

using PNGase F or chemical deglycosylation using TFMS acid will 

help to obtain better resolution of the protein bands on the SDS-PAGE and 2-DE 

gels. Deglycosylation can also aid in identification of proteins by MALDI-MS 

by obtaining better-resolved peaks in the mass spectra. 

10. The preparation of samples for gel electrophoresis requires prior knowledge 

of protein concentration to determine the amount of protein to be loaded. The 

protein concentration in the concentrated supernatant broths can be determined 

according to the method of Bradford (15) using the Bio-Rad Protein Assay 

Reagent (Bio-Rad Laboratories, Inc., Hercules, CA), or any other commercially 

available kit. As fungal culture broths may contain phenolic compounds, it is 

important to note that many of these compounds interfere with several of the 

most popular methods for protein determination. It is suggested that protein 

concentration is determined following protein precipitation and resolubilization 

in SDS Reducing Buffer or 2-DE sample buffer. Unfortunately, several components 

of these buffers may also cause problems in the assessment of protein 

concentration. Two commercially available protein determination kits that can 

be used with samples prepared for electrophoresis techniques are 2-D Quant


Kit (GE Healthcare Bio-Sciences Corp., Piscataway, NJ) and Advanced Protein 

Assay (Cytoskeleton, Inc., Denver, CO). 

11. Resolubilization of the precipitated protein pellet in 2-DE buffer may require 

vortexing and/or sonication. It has been recently shown that adding 20–30 μL 

of 0.2M NaOH to the TCA precipitated pellet for 2 min, before adding the 

solubilization buffer, increases the amount of soluble protein in the sample 

buffer (16). 


The techniques described here were adapted from the appropriate original 

papers. This work was supported by NSF grant MCB-0317126 to W.A.F. 

M.L.M. was supported in part by a fellowship through the Research Training 

Group in Optical Biomolecular Devices provided under NSF grant DBI- 

9602258-003 and the NSF-funded MGE@MSA Program, an Alliance for 

Graduate Education and the Professoriate, headquartered at Arizona State 

University. 

References 

1. Peberdy, J. F. (1994) Protein secretion in filamentous fungi - trying to understand 

a highly productive black-box, Trends Biotechnol. 12, 50–7. 

2. Conesa, A., Punt, P. J., van Luijk, N., and van den Hondel, C. A. A. J. J. (2001) 

The secretion pathway in filamentous fungi: A biotechnological view, Fungal. 

Genet. Biol. 33, 155–71. 

3. Archer, D. B. and Peberdy, J. F. (1997) The molecular biology of secreted enzyme 

production by fungi, Crit. Rev. Biotechnol. 17, 273–306. 

4. Wallis, G. L. F., Swift, R. J., Hemming, F. W., Trinci, A. P. J., and Peberdy, J. F. 

(1999) Glucoamylase overexpression and secretion in Aspergillus niger: analysis 

of glycosylation, BBA-Gen. Subjects 1472, 576–86. 

5. Medina, M. L., Haynes, P. A., Breci, L., and Francisco, W. A. (2005) Analysis of 

secreted proteins from Aspergillus flavus, Proteomics 5, 3153–61. 

6. Medina, M. L., Kiernan, U. A., and Francisco, W. A. (2004) Proteomic analysis of 

rutin-induced secreted proteins from Aspergillus flavus, Fungal. Genet. Biol. 41, 

327–335. 

7. Zhu, L. Y., Nguyen, C. H., Sato, T., and Takeuchi, M. (2004) Analysis of secreted 

proteins during conidial germination of Aspergillus oryzae RIB40, Biosci. Biotech. 

Bioch. 68, 2607–12. 

8. Phalip, V., Delalande, F., Carapito, C., et al. (2005) Diversity of the exoproteome 

of Fusarium graminearum grown on plant cell wall, Curr. Genet. 48, 366–79. 

9. Zorn, H., Peters, T., Nimtz, M., and Berger, R. G. (2005) The secretome of 

Pleurotus sapidus, Proteomics 5, 4832–38. 

10. Keller, N. P. and Hohn, T. M. (1997) Metabolic pathway gene clusters in 

filamentous fungi, Fungal. Genet. Biol. 21, 17–29.


11. Semova, N., Storms, R., John, T., et al. (2006) Generation, annotation, and analysis 

of an extensive Aspergillus niger EST collection, BMC Microbiol. 6:7. 

12. Fryksdale, B. G., Jedrzejewski, P. T., Wong, D. L., Gaertner, A. L., and Miller, B. S. 

(2002) Impact of deglycosylation methods on two-dimensional gel electrophoresis 

and matrix assisted laser desorption/ionization-time of flight-mass spectrometry 

for proteomic analysis, Electrophoresis 23, 2184–93. 

13. Edge, A. S. B., Faltynek, C. R., Hof, L., Reichert, L. E., and Weber, P. (1981) 

Deglycosylation of glycoproteins by trifluoromethanesulfonic acid, Anal. Biochem. 

118, 131–7. 

14. Suarez, M. B., Sanz, L., Chamorro, M. I., et al. (2005) Proteomic analysis of 

secreted proteins from Trichoderma harzianum - Identification of a fungal cell 

wall-induced aspartic protease, Fungal Genet. Biol. 42, 924–34. 

15. Bradford, M. M. (1976) Rapid and sensitive method for quantitation of microgram 

quantities of protein utilizing principle of protein-dye binding, Anal. Biochem. 72, 

248–54. 

16. Nandakumar, M. P., Shen, J., Raman, B., and Marten, M. R. (2003) Solubilization 

of trichloroacetic acid (TCA) precipitated microbial proteins via NaOH for 

two-dimensional electrophoresis, J. Proteome Res. 2, 89–93.

23 

Isolation and Solubilization of Cellular Membrane 

Proteins from Bacteria 

Kheir Zuobi-Hasona and L. Jeannine Brady 

Summary 

Membrane proteins are rarely identified in two-dimensional electrophoretic (2-DE) 

proteomics maps. This is because of low abundance, poor solubility, and inherent 

hydrophobicity. In this study, membrane preparations from the Gram-positive bacterium 

Streptococcus mutans were isolated from protoplasts and by mechanical grinding. 

Membrane proteins were extracted using a mixture of trifluroethanol and chloroform, 

solubilized using highly chaotropic buffer containing ASB-14 and Triton X-100 and 

subjected to two-dimensional gel electrophoresis. 

Key Words: Gram-positive; membrane proteins; solubilization; streptococcus 

mutans; two-dimensional gel electrophoresis. 


Membrane proteins are notably limited in proteomic analysis of bacteria 

(1–4). The primary reason is their intrinsically hydrophobic nature leading to 

poor solubility (1,5). This hydrophobicity likely causes self-aggregation during 

isoelectric focusing leading to poor resolution and streaking in 2-D gels. Many 

approaches have been reported to improve recovery of hydrophobic proteins 

including fractionation for enrichment of proteins of interest and reduction of 

sample complexity (6,7), experimenting with different detergents to extract and 

solubilize these proteins (8–10), and extraction using organic solvents (11–13). 

While extraction of proteins from eukaryotic membranes and analysis by 2- 

D electrophoresis has been technically feasible, such an approach to analyze 



287

288 Zuobi-Hasona and Brady 

bacterial membrane proteins, particularly those from Gram-positive organisms 

has proved much more challenging. In this study membrane-associated proteins 

of the Gram-positive bacterium Streptococcus mutans were prepared using two 

techniques, one by mechanical grinding (14) and the other from protoplasts (15, 

16). Both methods involve differential centrifugation to enrich for membranes 

within the preparation. The membrane associated proteins were extracted using 

a mixture of trifluroethanol and chloroform (13). This treatment produced three 

separate phases: An upper aqueous phase containing most of the proteins, an 

insoluble interphase containing few proteins and a lower chloroformic phase 

containing low concentration of proteins with low molecular weight (17). The 

extracted proteins were solubilized using highly chaotropic buffer containing 

ASB-14 and Triton X-100 and subjected to two-dimensional gel electrophoresis 

(Fig. 1). These techniques were efficient and highly reproducible for analysis 

of membrane proteins in S. mutans and suggest the potential use for other 

streptococci and Gram-positive bacteria in general. 

2. Materials 


1. UA-159, Streptococcus mutans, wild-type strain. 

2. Todd-Hewitt broth supplemented with 0.3% yeast extract (THYE). 

3. TM buffer: 50 mM maleate buffer, pH 6.0, containing 20 mM MgCl2. 4. Protease inhibitor cocktail (Sigma), reconstitute with DMSO and water, aliquot 

and store at –20°C. 

5. Alumina Powder (A-5, Sigma). 

6. Prechilled mortar and pestle (–20°C). 

7. Wash buffer: 20 mM Tris-HCl, pH 6.8, containing 1 mM MgCl2 and protease 

inhibitor cocktail at a 1:200 v/v ratio. 

2.2. Preparation of Membranes from Protoplasts 

1. Phosphate-buffered saline (PBS). 

2. Buffer A: 20% sucrose, 20 mM Tris-HCl, pH 7.0, and 10 mM MgCl2. Store 

at 4°C. 

2. Mutanolysin (INC Corp), store at -20°C. 

3. Lysozyme (Sigma), store at –20°C. 

4. Buffer B: 10 mM Tris-HCl, pH 8.1, 50 mM MgCl2 and 10 mM Glucose. Store 

at 4°C. 

5. Buffer C: 10 mM Tris-HCl, pH 8.1, 50 mM NaCl and 20 mM MgCl2. Store 

at room temperature. 

6. Buffer D: 20 mM Tris-HCl, pH 7.2, and 10 mM MgCl2

Isolation and Solubilization of Cellular Membrane Proteins 289 

2.3. Trifluroethanol/Chloroform Extraction and Analysis 

1. 50 mM ammonium bicarbonate pH 11.0, store at 4°C. 

2. Protease inhibitor cocktail (Sigma). 

3. Trifluroethanol/chloroform mixture (2:1 v/v). 

4. Ready Prep 2-D Cleanup Kit (Bio-Rad). 

5. Solubilization buffer: 7M urea, 2M thiourea, 2% Triton X-100, 0.5% ASB-14, 

50 mM dithiothreitol (DTT) and 0.2% Bio-Lytes pH 3–10. Aliquot and store 

at –70°C. 

6. RC DC protein assay kit (Bio-Rad). 

7. Standard electrophoresis equipment from Bio-Rad: PROTEAN IEF Cell, Criterion 

Cell. 

3. Methods 


1. Transfer 100 mL of overnight culture of Streptococcus mutans, strain UA159 

into 2 L of prewarmed Todd-Hewitt broth supplemented with 0.3% yeast extract. 

(THYE). 

2. Incubate at 37°C with gentle agitation, until an absorbance reading of 0.7 at 600 

nm is reached. 

3. Place the culture immediately on ice for 20 min. 

4. Harvest the cells by centrifugation at 12,000g for 10 min at 4°C. 

5. Wash the cell pellet twice in TM buffer containing protease inhibitor at 1:30 v/v 

ratio. 

6. Resuspend the cell pellet with 6 mL of the same buffer divide into three aliquots 

and store as slurries (see Note 1). 

7. Transfer each frozen slurry to a prechilled mortar. 

8. Add 3-g Alumina powder (A-5, Sigma). (see Note 2) 

9. Use a prechilled pestle to grind the cells for 20 min. 

10. Transfer the mixture into a 15 mL conical tube and centrifuge at 3,000g for 5 

min at 4°C to remove the Alumina. 

11. Centrifuge the supernatant at 16,000g for 10 min at 4°C to remove intact cells 

and cell debris. Discard the pellet. 

12. Ultracentrifuge the supernatant at 100,000g for 18 h at 4°C to pellet the 

membranes. 

13. Rinse the pellet three times with wash buffer (total volume 15 mL). 

14. Resuspend the pellet in 5 ml wash buffer and centrifuge at 45,000g for5h 

at 4°C. 

15. Remove the supernatant by pipette; it represents the cell cytoplasm and store 

at –70°C. 

16. Keep the pellet in the centrifuge tube covered with parafilm, place in a box and 

store at –70°C until used. This represents the membrane-containing fraction.


(A) 

(B) 

Fig. 1. Silver stained 2-D gel of Streptococcus mutans. (A) Membrane proteins 

obtained using the grinding method. (B) Cytoplasmic proteins. The 2-D gel profile 

of cytoplasmic proteins showed an entirely different distribution of protein spots as 

compared with the gel of extracted membrane proteins. Nanoelectrospray quadrupole 

time of flight (QTOF)-tandem mass spectrometry (MS/MS) analysis of several protein 

spots isolated from this gel (B) identified proteins predicted to be cytoplasmically 

localized, including translation elongation factor (spot #8), the 10 kDa chaperone GroES 

(spot #9), and ribosome recycling factor (spot #10). None of the protein spots identified


3.2. Preparation of Membranes from Protoplasts 

1. Transfer 100 ml of overnight culture of Streptococcus mutans, strain UA159 

into 2 L of prewarmed Todd-Hewitt broth supplemented with 0.3% yeast extract 

(THYE). 

2. Incubate at 37°C with gentle agitation, until an absorbance reading of 0.7 at 

600 nm is reached. 

3. Harvest the cells by centrifugation at 12,000g for 10 min at 4°C. 

4. Wash the pellet with phosphate-buffered saline (PBS). 

5. Re-suspend the cells with 50 mL Buffer A containing 150 μL protease inhibitor 

cocktail, 1 mg mutanolysin and 18 mg lysozyme. Swirl gently. 

6. Incubate the mixture for additional 3hat37°C, swirling gently every 10 min. 

7. Examine by light microscope under high power (see Note 3). 

8. Collect the protoplast at 12,000g for 10 min at 4°C. Store supernatant at –20°C. 

8. Wash the protoplasts with 30 mL Buffer B. 

10. Add 100 μL protease inhibitor cocktail and passage twice using French Press 

under 10,000 psi. 

11. Remove debris and unbroken protoplasts by centrifugation at 6,000g for 10 min. 

at 4°C. 

12. Ultra-centrifuge the supernatant for 30 min. at 45,000g at 4°C. 

13. Label the supernatant as cytoplasmic proteins and store at –70°C. 

14. Resuspend the pellet in 3 mL Buffer C at room temperature. 

15. Ultracentrifuge at 100,000g for 45 min at 4°C. 

16. Decant the supernatant and wash the pellet twice with 2 mL Buffer D and decant 

the wash. 

17. Store the pellet at –70°C until used. 

3.3. Trifluoroethanol/Chloroform Extraction and Analysis 

1. Re-suspend each membrane pellet (from Sections 3.1 and 3.2) in 150 μL of 

50 mM ammonium bicarbonate (pH 11), containing 15 μL protease inhibitor 

cocktail and vortex well for one minute. 

2. Add 1 mL of trifluroethanol/chloroform mixture (2:1 v/v) and vortex well for 

one minute. 

3. Maintain the mixture on ice for 1 h and vortex every 5 min for 10 s each time. 

◭ 

Fig. 1. (Continued) from membrane extracts were observed in the 2-D gel of 

cytoplasmic proteins. On the other hand QTOF/MS/MS analysis of several spots 

isolated from gel (A) revealed known membrane and surface-associated proteins, 

including Enolase (spot #1), Biotin carboxyl carrier protein (spot #2), LemA-like protein 

(spot #3), Glucose 6-phosphate isomerase (spot #4), Phosphoglycerate kinase (spot 

#5), Glyeraldehyde 3-phosphate dehydrogenase (spot #6) and 50S ribosomal protein 

L7/L12 (spot #7).


4. Centrifuge at 10,000g for 5 min. at 4°C to separate into three phases. 

5. Carefully transfer the aqueous upper phase and the chloroformic lower phase 

into separate microcentifuge tubes. Keep the middle phase in the original microcentrifuge. 

6. Dry using vacuum centrifuge for 2 h. 

7. Solubilize each residue with 200 μL solubilization buffer, vortex for 1 min; let 

it set at RT then repeat vortexing twice. 

8. Precipitate membrane proteins with the Bio-Rad clean-up kit (see Note 4). 

9. Re-suspend the resulting pellets in solubilization buffer (from step 7), vortex 

well as in step 7. 

10. Centrifuge at 13,000g for 5 min at room temperature (see Note 5). 

11. Measure the protein concentration using RC DC protein assay and adjust to 

∼250 μg/mL solubilization buffer from step 7 (see Note 6). 

12. Analyze membrane fraction by standard two-dimensional electrophoresis 

protocols and silver stain the resulting 2-D gels. 

4. Notes 

1. Passage the cells through a syringe few times. The slurries can be placed in 

weighing boats wrapped, and stored frozen at –70°C until needed to be used. 

2. Use mask to avoid inhaling the Alumina, it’s recommended to do the grinding in 

an ice pocket to avoid protein degradation. 

3. More than 90% of cell wall digestion occurs after 3hofincubation. Continuous 

gentle agitation can speed the process. 

4. This step should be done using centrifuge at 4°C. Avoid over-drying the pellets, 

as it will be difficult to resuspend, resulting in loss of certain proteins. 

5. It is necessary to clarify the protein sample from insoluble particles that might 

cause streaking in 2D gel. The supernatant can be used directly for isoelectricfocusing 

(IEF) in IPG strips. Store any remaining protein sample at –70°C for 

later analysis. 

6. RC DC Protein Assay (Bio-Rad cat # 500-0121) or Plus One 2-D Kit (Amersham) 

can be used for protein quantitation. Dilute as necessary with solubilization buffer 

to yield the desired quantity of protein (50–100 μg, when using silver stain). 

References 

1. Santoni, V., Molloy, M. P. and Rabilloud, T. (2000) Membrane proteins and 

proteomics: un amour impossible? Electrophoresis 21, 1054–70. 

2. Molloy, M. P., Herbert, B. R., Slade, M. B., Rabilloud, T., et al. (2000) Proteomics 

analysis of the Escherichia coli outer membrane. Eur. J. Biochem. 267, 2871–88. 

3. Nouwens, A. S., Cordwell, S. J. Larsen, M. R., Molloy, M. P. et al. (2000) Complementing 

genomics with proteomics: the membrane subproteome of Pseudomonas 

aeruginosa PAO1. Electrophoresis, 21, 3797–3809.


4. Wilkins, M. R., Gasteiger, E., Sanchez, J. –C., Bairoch, A. and Hochstrasser, D. F. 

(1998) Two-dimensional gel electrophoresis for proteome projects: the effect of 

protein hydrophobicity and copy number. Electrophoresis , 19, 1501–05. 

5. Pasquali, C. Fialka, I., and Huber, L. A. (1997) Preparative two-dimensional gel 

electrophoresis of membrane proteins. Electrophoresis 18, 2573–81. 

6. Molloy, M. P., Herbert, B. R., Walsh, B. J., Tyler, M. I., et al. (1998) Extraction 

of membrane proteins by differential solubilization for separation using twodimensional 

gel electrophoresis. Electrophoresis 19, 2573–81. 

7. Lehner, I., Niehot, M., and Borlak, J., (2003) An optimized method for the isolation 

and identification of membrane proteins. Electrophoresis 24, 1795–1808. 

8. Rabilloud, T., Blisnick, T., Heller, M., Luche, S., et al. (1999) Analysis of 

membrane proteins by two-dimensional electrophoresis: comparison of the proteins 

extracted from normal or Plasmodium faciparum-infected erythrocyte ghosts. 


9. Wissing, J., Heim, S., Flohe, L., Bilitewski, U., and Frank, R. (2000) Enrichment 

of hydrophobic proteins via Triton X-114 phase partitioning and hydroxyapatite 

column chromatography for mass spectrometry. Electrophoresis 21, 2589–93. 

10. Wasinger, V. C., Pollack, J. B., and Humphery-Smith, I., (2000) The proteome 

of Mycoplasma genitalium. Chaps-soluble component. Eur. J. Biochem. 267, 

1571–82. 

11. Molloy, M. P., Herbert, B., Williams, K. L., and Gooley, A. A., (1999) Extraction 

of Escherichia coli proteins with organic solvents before two-dimensional 


12. Seigneurin-Berny, D., Rolland, N., Garin, J., and Joyard, J., (1999) Technical 

Advance: Differential extraction of hydrophobic proteins from chloroplast envelope 

membranes: a subcellular specific proteomic approach to identify rare intrinsic 

membrane proteins. Plant. J. 19, 217–228. 

13. Deshusses, J. M. P., Burgess, J. A., Scherl, A., Wenger, Y., Walter, N., et al. (2003) 

Exploitation of specific properties of trifluroethanol for extraction and separation 

of membrane proteins. Proteomics, 3, 1418–24. 

14. Vadeboncoeur, C., St Martin, S., Brochu, D., and Hamilton, I. R., (1991) Effect of 

growth rate and pH on intracellular levels and activities of the components of the 

phosphoenolpyruvate: sugar phosphotransferase system in streptococcus mutans 

Ingbritt. Infect. Immun. 59, 900–6. 

15. Wessel, D. and Flugge, U. I., (1984) A method for the quantitative recovery of 

protein in dilute solution in the presence of detergents and lipids. Anal. Biochem. 

138, 141–3. 

16. Bordier, C. (1981) Phase separation of integral membrane proteins in Triton X-114 

solution. J. Biol. Chem. 256, 1604–7. 

17. Zuobi-Hasona, K., Crowley, P. J., Hasona, A., Bleiweis, A. S., and Brady, L. J. 

(2005) Solubilization of cell membrane proteins from Streptococcus mutans for 

two-dimensional gel electrophoresis. Electrophoresis 26, 1200–6.

24 

Isolation and Solubilization of Gram-Positive Bacterial 

Cell Wall-Associated Proteins 

Jason N. Cole, Steven P. Djordjevic, and Mark J. Walker 

Summary 

This chapter describes a simple, rapid and reproducible method to prepare bacterial cell 

wall extracts for two-dimensional gel electrophoresis (2DE). The extraction process uses 

mutanolysin, an N-acetylmuramidase, to gently solubilize cell wall-associated proteins 

from Gram-positive prokaryotes. The cells are first washed with buffer and resuspended in 

a solution containing mutanolysin. Following incubation at 37 °C, the sample is centrifuged 

and the supernatant containing the soluble cell wall-associated proteins is harvested. 

Following a brief precipitation step, the pellet is solubilized in sample buffer ready for 

isoelectric focusing and 2DE analysis. 

Key Words: Bacterial proteome; cell wall-associated; Gram-positive; group A streptococcus; 

mutanolysin; Streptococcus pyogenes; two-dimensional gel electrophoresis. 


Gram-positive bacteria are bounded by a thick cell wall composed of 

primarily of peptidoglycan, a large macromolecule of acetamido sugars and 

amino acids (1). The glycan chains of peptidoglycan consist of alternating 

units of N-acetylglucosamine (GlcNAc) and N-acetylmuramic acid (MurNAc) 

in -1,4 linkage (2). Short cell wall peptides are cross-linked to the glycan 

chains to form a three-dimensional molecular network. The major function of 

the cell wall envelope is to provide a rigid exoskeleton for protection against 

mechanical and osmotic lysis. The cell wall also maintains a defined cell shape 

and plays an integral role in the anchoring of proteins to the cell surface (3,4). 



295

296 Cole et al. 

Most Gram-positive cell walls are resistant to dissolution with lysozyme (5). 

Mutanolysin, a muralytic enzyme derived from Streptomyces globisporus 1829, 

cleaves the -1,4 linkage of the N-acetylmuramyl-N-acetylglucosamine in the 

glycan backbone of the peptidoglycan-polysaccharide polymer, which is highly 

conserved among bacterial species (6). Although mutanolysin has the same 

specificity as lysozyme, it has a wider range of activity among peptidoglycanpolysaccharide 

from bacterial strains resistant to lysozyme, including Streptococcus 

pyogenes (group A streptococcus), a Gram-positive pathogen responsible 

for numerous human diseases. 

In this chapter, the authors describe a convenient and efficient mutanolysin 

extraction method for the solubilization of cell wall-associated proteins from 

S. pyogenes. A detailed discussion is presented on bacterial growth conditions, 

cell wall purification, preparative SDS-PAGE analysis, sample preparation, 

isoelectric focusing and 2DE. 

2. Materials 

Unless stated otherwise, all solutions were prepared with glass distilled 

water and high purity electrophoresis grade reagents. Review the manufacturers’ 

safety data sheets (MSDS) and follow the safety precautions and general 

handling procedures for hazardous materials. 

2.1. Bacterial Culture 

1. Horse blood agar plates (BioMérieux). Store at 4 °C. 

2. THBY medium: 30 g/L Todd-Hewitt broth, 10 g/L yeast extract (Difco). 

Autoclave and store at room temperature. 

2.2. Extraction of Cell Wall-Associated Proteins 

1. TE buffer: 50 mM Tris-HCl, 1 mM EDTA, pH 8.0. Autoclave and store at 4 °C. 

2. TE-Sucrose (TES) buffer: 20% (w/v) sucrose in TE buffer (pH 8.0). Autoclave 

and store at 4 °C. 

3. Phenylmethylsulfonyl fluoride (PMSF, Sigma) dissolved at 1 mM in chilled TES 

buffer (see Note 1). PMSF is very toxic and rapidly degrades in aqueous solutions. 

Prepare immediately before use and place on ice. 

4. Mutanolysin: Resuspend 10,000 units of chromatographically purified mutanolysin 

from Streptomyces globisporus ATCC 21553 (Sigma) in 2 mL of chilled filtersterilized 

0.1 M K2HPO4 (pH 6.2) for a working solution of 5,000 units/mL. Store 

200 μL aliquots at –20 °C. 

5. Lysozyme dissolved in chilled TES buffer at 100 mg/mL. Avoid foaming by 

gently pipeting up and down. Prepare fresh each time and store on ice before use.

Isolation of Gram-Positive Bacterial Cell Wall Proteins 297 

6. Mutanolysin mix: 1 mL TES buffer, 100 μL lysozyme (100 mg/mL in TES), 

50 μL mutanolysin (5,000 U/mL in 0.1 M K 2HPO 4, pH 6.2). Prepare immediately 

before use and place on ice. 

2.3. Bicinchoninic Acid (BCA) Protein Assay 

1. Bicinchoninic Acid Protein Assay Kit (Sigma): Store Reagents A and B at room 

temperature. 

2. Bovine serum albumin (BSA, Sigma). 

3. TES buffer: 20% (w/v) sucrose in TE buffer (50 mM Tris-HCl, 1 mM EDTA, pH 

8.0). Autoclave and store at 4 °C. 

4. Microtiter plate reader capable of measuring absorbance in the 560 nm region. 

5. Microtiter plate and sealing film. 

2.4. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 

1. 0.5 M Tris-HCl (pH 6.8) and 1.5 M Tris-HCl (pH 8.8) prepared in Milli-Q ® 

water. Adjust pH with concentrated HCl. Store at 4 °C. 

2. 10% (w/v) SDS dissolved in Milli-Q ® water. Store at room temperature. 

3. 40% acrylamide/bis solution (37.5:1, Amresco). Acrylamide monomer is a 

neurotoxin and suspected carcinogen; care should be taken to avoid exposure. 

Store in dark at 4 °C. 

4. N,N,N ′ ,N ′ -Tetramethylethylenediamine (TEMED, Amresco). Store at room 

temperature. 

5. Ammonium persulfate (APS, Amresco) prepared at 10% (w/v) in Milli-Q ® water 

(see Note 2). 

6. Gel overlay solution: 70% (v/v) ethanol. Store at room temperature. 

7. 10× running buffer: 250 mM Tris-HCl, 1.92 M glycine, 1% (w/v) SDS, pH 8.3. 

Do not adjust pH with acid or base. Dilute to 1× working solution and mix 

thoroughly before use. Store at room temperature. 

8. 1 M Dithiothreitol (DTT) dissolved in 0.01 M sodium acetate (pH 5.2). Filtersterilize 

and store 1 mL aliquots at –20 °C. 

9. 5× loading buffer: 225 mM Tris-HCl (pH 6.8), 50% (v/v) glycerol, 5% (w/v) 

SDS, 0.05% (w/v) Bromophenol Blue, 250 mM DTT (see Note 3). Store at 

room temperature. Dilute sample 1:5 with loading buffer and heat at 95 °C for 

10 min. 

10. Molecular weight markers: PageRuler Protein Ladder (Fermentas). Store 

at –20 °C. 

11. Coomassie Blue staining solution: 0.2% (w/v) Coomassie Blue R250, 40% 

(v/v) methanol, 10% (v/v) acetic acid. Store at room temperature. 

12. Rapid destain solution: 40% (v/v) methanol, 10% (v/v) acetic acid. Store at room 

temperature. 

13. Final destain solution: 4% (v/v) glycerol, 10% (v/v) acetic acid. Store at room 

temperature.


2.5. Trichloroacetic Acid (TCA) Precipitation 

1. 10% (v/v) TCA (Sigma): Protect from light and store at room temperature. TCA 

is a highly corrosive acid. Use safety equipment when handling to avoid exposure. 

2. Ethanol 100% (v/v) analytical reagent stored at –20 °C. 

3. Sample solubilization solution: 8 M urea, 100 mM DTT, 4% (w/v) CHAPS, 0.8% 

(v/v) Bio-Lyte ® 3/10 ampholyte (Bio-Rad), 40 mM Tris-HCl. Make up to volume 

with Milli-Q ® water (see Note 4). Store 1 mL aliquots at –20 °C. Thaw required 

number of aliquots and discard leftover solution. 

2.6. Two-Dimensional (2D) Gel Electrophoresis 

2.6.1. Sample Preparation and Rehydration 

1. Sample solubilization solution: 8 M urea, 100 mM DTT, 4% (w/v) CHAPS, 0.8% 

(v/v) Bio-Lyte ® 3/10 ampholyte (Bio-Rad), 40 mM Tris-HCl. Make up to volume 

with Milli-Q ® water. Store 1 mL aliquots at –20 °C. Thaw required number of 

aliquots and discard leftover solution. 

2. Water-bath sonicator or equivalent. 

3. Bromophenol Blue solution 1% (w/v) in Milli-Q ® water. Pass through a 0.22-μm 

filter unit (Millipore) to remove particulate matter and store at room temperature. 

4. 11-cm ReadyStrip linear pH 4-7 IPG strips (Bio-Rad). 

5. Disposable plastic rehydration tray (Bio-Rad). 

6. Mineral oil (Bio-Rad). Store at room temperature. 

2.6.2. First Dimension Isoelectric Focusing (IEF) 

1. Electrode wicks (Bio-Rad). 

2. PROTEAN ® IEF Cell and 11-cm focusing tray (Bio-Rad). 

3. Rehydrated 11-cm ReadyStrip linear pH 4–7 IPG strip (Bio-Rad). 

4. Mineral oil (Bio-Rad). Store at room temperature. 

2.6.3. Casting 2D Gels 

1. 1.5 M Tris-HCl (pH 8.8) prepared in Milli-Q ® water. Adjust pH with concentrated 

HCl. Store at 4 °C. 

2. 10% (w/v) SDS dissolved in Milli-Q ® water. Store at room temperature. 

3. 40% acrylamide/bis solution (37.5:1, Amresco). Store in dark at 4 °C. 

4. TEMED (Amresco). Store at room temperature. 

5. 10% (w/v) APS in Milli-Q ® . Prepare fresh each time. 

6. Water-saturated n-butanol gel overlay solution: Combine equal volumes of 

n-butanol and Milli-Q ® water and shake to mix. Allow to settle and use top 

(aqueous layer) to overlay gels. Store at room temperature. 

7. Gel storage solution: 0.375 M Tris-HCl (pH 8.8), 0.1% (w/v) SDS. Store at 4 °C.


2.6.4. Second Dimension SDS-PAGE 

1. Disposable plastic equilibration tray (Bio-Rad). 

2. Equilibration buffer: 6 M urea, 2% (w/v) SDS, 375 mM Tris-HCl (pH 8.8), 20% 

(v/v) glycerol, 2.5% (v/v) acrylamide, 130 mM DTT. Store 4 mL aliquots at 

–20 °C. Thaw required number of aliquots and discard leftover solution. 

3. Molecular weight markers: PageRuler Prestained Protein Ladder (Fermentas). 

Store at –20 °C. 

4. 10× running buffer: 250 mM Tris-HCl, 1.92 M glycine, 1% (w/v) SDS, pH 8.3. 

Do not adjust pH. If the pH is not accurate remake buffer. Dilute to 2× and 1× 

working concentrations and mix thoroughly before use. Store at room temperature. 

5. Bromophenol Blue solution 1% (w/v) in Milli-Q ® water. Pass through a 0.22 μm 

filter unit (Millipore) to remove particulate matter and store at room temperature. 

6. Agarose gel overlay solution: 1% (w/v) agarose in 1× running buffer (pH 8.3). 

Do not adjust pH. Add 0.1 mL of 1% (w/v) Bromophenol Blue per 100 mL 

of overlay solution and heat in a microwave oven until completely melted (see 

Note 5). Allow to cool slightly (60 °C) before using. Store at room temperature 

after use. 

2.6.5. Protein Detection and Gel Documentation 

1. Colloidal Coomassie stain: 17% (w/v) ammonium sulfate, 3% (v/v) phosphoric 

acid, 34% (v/v) methanol, 0.1% (w/v) Coomassie G250 (see Note 6). Store at 

room temperature. 

2. Destain solution: 1% (v/v) glacial acetic acid. Store at room temperature. 

3. Methods 

For full information pertaining to chemical and electrical hazards refer to the 

manufacturers’ material safety data sheets (MSDS) and instruction manuals. 

S. pyogenes (UN2814) is a potential human pathogen and safety precautions 

must be taken to avoid exposure. 

3.1. Bacterial Culture 

1. Sixteen-streak the S. pyogenes strain onto a fresh horse blood agar plate (see 

Note 7). Seal with Parafilm ® and incubate at 37 °C overnight (approx 16 h). 

2. Inoculate a single, well-isolated hemolytic colony (see Note 8) intoa5mLsterile 

tube containing 2 mL of THBY medium. Incubate at 37 °C overnight for 16 h 

without shaking. 

3. To a 250 mL conical flask containing 98 mL of THBY, add the entire volume 

of overnight culture and incubate at 37 °C without agitation until late stationary 

phase is reached (approx 16 h) (see Note 9).


3.2. Extraction of Cell Wall-Associated Proteins 

1. Transfer the overnight culture to a sterile centrifuge tube and harvest the bacterial 

cells by centrifugation at 7,560g for 20 min at 4 °C. 

2. Carefully decant the culture supernatant and discard. Place the bacterial pellet on 

ice for 5 min. 

3. Resuspend the pellet in 5 mL of chilled TE buffer containing 1 mM PMSF (see 

Note 10) by pipetting up and down. Take care to avoid foaming and ensure no 

bacterial clumps are visible. Centrifuge at 7,560g for 20 min at 4 °C and discard 

the supernatant. 

4. Repeat step 3. Resuspend the pellet in 1.15 mL of ice-cold mutanolysin mix by 

pipeting up and down. Take care to avoid foaming and ensure no bacterial clumps 

are visible. 

5. Transfer to a sterile microcentrifuge tube and incubate for 2hat37°Cwith 

shaking (200 rpm) (see Note 11). 

6. Centrifuge at 14,000g for 5 min at room temperature in a bench-top microcentrifuge. 

7. Collect the supernatant (solubilized cell wall-associated proteins) by aspiration. 

Store 500 μL aliquots at –20 °C (see Note 12). 

8. Determine the protein concentration of the cell wall extract using the BCA protein 

assay described below. 

9. Prepare for SDS-PAGE analysis by diluting 20 μL of 5× loading buffer with 80 

μL of cell wall extract in a microcentrifuge tube. Pierce the lid and boil for 10 

min. Allow to cool to room temperature before loading (see Note 13). 

3.3. BCA Protein Assay 

1. Prepare the required amount of BCA Working Reagent by mixing 50 parts 

of Reagent A (Sigma proprietary solution containing BCA, sodium carbonate, 

sodium tartrate and sodium bicarbonate in 0.1 M NaOH, pH 11.25) with 1 part 

of Reagent B, a 4% (w/v) solution of copper(II) sulfate pentahydrate. Mix by 

vortexing until the solution is a uniform light green color (see Note 14). 

2. Prepare BSA protein standards ranging from 0.2 to 1 mg/mL in TES buffer (see 

Note 15). Include a blank containing buffer with no protein. Protein standards 

may be stored at –20 °C. 

3. Dilute the cell wall extract 1:5 in TES buffer to ensure the concentration is within 

the linear range of 0.2 to 1 mg/mL. 

4. Add 25 μL of each standard, cell wall extract and blank to a microtiter plate 

before adding 200 μL of Working Reagent (see Note 16). 

5. Seal the plate, incubate at 37 °C for 30 min and cool to room temperature (see 

Note 17). 

6. Measure the absorbance at 562 nm with a microtiter plate reader and estimate 

the protein concentration of unknown samples from the BSA standard curve (see 

Note 18).


3.4. SDS-PAGE 

This procedure is for use of the discontinuous Laemmli system (7) in a 

Mini-PROTEAN ® 3 Cell (Bio-Rad) and can be readily adapted for other gel 

formats. 

1. Thoroughly clean the glass plates with a laboratory detergent, rinse completely 

with distilled water and air dry. 

2. Prepare a 12% resolving gel by mixing 3 mL of 40% acrylamide/bis solution 

with 2.5 mL of 1.5 M Tris-HCl (pH 8.8), 4.35 mL Milli-Q ® water, 100 μL 10% 

(w/v) SDS, 15 μL TEMED and 50 μL 10% (w/v) APS (see Note 19). Pour a 0.75 

mm thick resolving gel to the required level and immediately overlay with 70% 

(v/v) ethanol to give a flat gel surface (see Note 20). Allow the gel to polymerize 

for45minto1hatroom temperature (see Note 21). The resolving gel can be 

submerged in 1.5 M Tris-HCl (pH 8.8) diluted 1:4 in distilled water and stored 

at 4 °C for up to 1 wk. 

3. Decant the ethanol and thoroughly rinse the gel surface with distilled water. 

Remove excess water between the glass plates above the resolving gel with a 

piece of filter paper, taking care to avoid the gel surface. 

4. Prepare the 4% stacking gel by mixing 0.5 mL of 40% acrylamide/bis solution 

with 2.5 mL of 0.5 M Tris-HCl (pH 6.8), 3.18 mL Milli-Q ® water, 50 μL 10 

% (w/v) SDS, 5 μL TEMED and 25 μL 10% (w/v) APS. Pour the stacking gel, 

insert the comb and allow the stacking gel to polymerize for 30–45 min at room 

temperature. 

5. Prepare the running buffer by diluting 100 mL of 10× running buffer with 900 

mL of distilled water. Mix thoroughly before use. 

6. After the stacking gel has set, gently remove the comb and thoroughly rinse the 

wells with running buffer. Place the gel into the electrophoresis module and add 

running buffer to the upper and lower chambers. Load 15 μL of sample (approx 

20 μg protein) and molecular weight markers for size comparison (see Note 22). 

7. Connect the lid of the electrophoresis unit to a suitable power supply and run at 

a constant 200 V for approximately 45 min, or until the dye front reaches the 

bottom of the gel (see Note 23). 

8. Once electrophoresis is complete, discard the running buffer and remove the 

stacking gel from the resolving gel. Place the resolving gel in a plastic tray and 

submerge in Coomassie Blue staining solution (see Note 24). Cover the tray to 

minimize exposure to vapors, microwave (600 W for 15 s) and shake at room 

temperature for1horleave overnight. Decant stain (see Note 25) and add enough 

destain solution to completely cover the gel. Microwave for 15 s and shake for 20 

min at room temperature. Repeat with fresh destain solution until the background 

is clear (see Note 26). 

9. Discard destain and submerge gel in final destain solution ready for documentation 

with the GS-800 calibrated densitometer (Bio-Rad) or equivalent (see Note 27). 

A representative SDS-PAGE gel of cell wall extracts from S. pyogenes is shown 

in Fig. 1.


Fig. 1. Coomassie Blue stained 12% SDS-PAGE reducing gel of a mutanolysin 

cell wall extract harvested from S. pyogenes strain 5448 (serotype M1) after growth 

at 37 °C to late stationary phase in THBY medium without agitation. Molecular mass 

markers are given in kilo-Daltons (kDa). 

3.5. TCA Precipitation 

1. Add 1 mL of 10% (v/v) TCA to 1 mL of cell wall extract and mix immediately 

by vortexing at maximum output (see Note 28). Incubate on ice for 20 min. 


3. Discard supernatant and wash pellet with 1 mL of ice-cold 100% (v/v) ethanol to 

remove residual TCA (see Note 29). 


5. Discard the supernatant and dry the pellet for 30–60 min at room temperature. 

Store TCA precipitated pellets at –20 °C before 2DE (see Note 30). 

3.6. 2D Gel Electrophoresis 

3.6.1. Sample Preparation and Rehydration 

1. Completely resuspend the TCA precipitated pellet in 500 μL of sample solubilization 

solution by pipeting up and down, taking care to avoid foaming (see 

Note 31).


2. Sonicate at 14 W for 30 sec and vortex for 15 sec at room temperature to aid 

solubilization. Repeat this step a further 3 times (see Note 32). 

3. Centrifuge at 14,000g for 15 min at room temperature in a bench-top microcentrifuge 


4. Transfer the supernatant to a new microcentrifuge tube and discard the pellet 


5. Calculate the volume of sample required for 200 μg protein and dispense into 

a new microcentrifuge tube. Adjust the volume to 250 μL with sample solubilization 

solution (see Note 35). 

6. Add 2 μL of Bromophenol Blue solution and mix by briefly vortexing (see 

Note 36). 

7. Pipette 250 μL of sample along the length of a channel in an 11 cm rehydration 

tray. Eliminate air bubbles using forceps to ensure even distribution of the 

sample in the IPG strip. 

8. Carefully place the IPG strip onto the sample gel side down. Eliminate air 

bubbles under the strip by gently lifting the strip from one end with forceps. 

9. Carefully overlay the strip with 2 mL of mineral oil to prevent evaporation (see 

Note 37). Attach the rehydration tray lid and rehydrate overnight (11–16 h) at 

room temperature (see Note 38). The sample will be passively absorbed by the 

strip during rehydration. 

10. Remove the IPG strip from the rehydration tray with forceps (see Note 39). 

Remove the excess oil by holding the strip vertically and gently touching the 

tip of the backing strip to blotting paper (see Note 40). 

3.6.2. First Dimension IEF 

The following IEF method assumes the use of the PROTEAN ® IEF Cell 

(Bio-Rad) (see Note 41). 

1. Equilibrate four electrode wicks per IPG strip in Milli-Q ® water (see Note 42). 

2. Remove excess water by briefly blotting the wicks on filter paper. Place two 

hydrated wicks directly on top of the cathode and anode electrode wires of the 

focusing tray. 

3. Place the IPG strip gel side down onto the wicks, with the acidic (positive) 

end of the strip at the anode (positive) electrode of the focusing tray (see 

Note 43). 

4. Overlay the IPG strip with 2 mL of mineral oil and eliminate air bubbles under 

the strip by gently lifting it from one end with forceps (see Note 44). Place the lid 

on the focusing tray and place into the IEF Cell, ensuring good contact between 

the electrodes of the focusing tray and the electrodes of the IEF Cell. 

5. Program the IEF Cell with the following focusing conditions: 100 V for 1 h; 

300 V for 1 h; 600 V for 1 h; 1,000 V for 1 h; 2,000 V for 1 h; 4,000 V for 

40,000 Volt-hours (Vh) (approx 10 h); and 100 V hold (12 h) (see Note 45). Use 

an IEF Cell temperature of 20 °C with a maximum current of 50 μA per IPG 

strip.


6. Remove the IPG strip from the IEF tray and drain the excess mineral oil with 

blotting paper as described in step 10 above. Proceed directly to the equilibration 

step (see Note 46). 

3.6.3. Casting 2D Gels 

This protocol is for the casting of homogenous polyacrylamide gels with the 

Ettan DALTsix Gel Caster (GE Biosciences). 

1. Lay the Ettan DALTsix Gel Caster on its back. Remove the faceplate and place 

the triangular rubber wedge in the V-shaped base. 

2. Place a separator sheet against the back wall of the caster (square corners against 

the rubber wedge) followed by a 1.0 mm gel casting cassette. Fill the caster by 

alternately layering separator sheets and gel cassettes, ending with a separator 

sheet. Add thicker filler sheets until the stack of cassettes is flush with the edge 

of the caster (see Note 47). 

3. Carefully place the faceplate onto the caster, clamp with six spring clips and 

tighten the faceplate screws (see Note 48). 

4. Attach the cap to the filler port on the faceplate to prevent leakage. Place the Gel 

Caster in an upright position on a level surface ready for casting. 

5. Prepare the 12.5% gel solution by mixing 156 mL of 40% acrylamide/bis with 

125 mL of 1.5 M Tris-HCl (pH 8.8), 6 mL 10% (w/v) SDS and 208 mL Milli-Q ® 

water. Mix well with a magnetic stirrer before adding 6 mL 10% (w/v) APS and 

100 μL TEMED (see Note 49). To cast a homogenous 1.0-mm-thick gel, add 

the APS followed by TEMED and slowly pour the gel solution into the filling 

channel located at the back of the caster. Continuing pouring until the level is 

approx 4 cm below the top edge of the short plate (see Note 50). 

6. Immediately overlay each gel with 4 mL of water-saturated n-butanol to exclude 

air and ensure a level surface. Cover the top of the Gel Caster with plastic wrap 

and allow gels to polymerize for 3–4 h at room temperature. 

7. Pour off the gel overlay solution and rinse the surface of each gel with Milli-Q ® 

water to remove residual n-butanol and unpolymerized acrylamide. Add 5 mL of 

gel storage solution to the top of each gel. Seal the top of the Caster with plastic 

wrap and store at 4 °C overnight (see Note 51). 

3.6.4. Second Dimension SDS-PAGE 

The following procedure is for the running of large format (26 × 20 cm) 

acrylamide gels with the vertical Ettan DALTsix Electrophoresis Unit (GE 

Biosciences) (see Note 52). 

1. Turn on the MultiTemp III heat exchanger and equilibrate to 10 °C. 

2. Place the IPG strip gel side up into an 11 cm equilibration tray and add 2 mL of 

Equilibration buffer. Rock gently for 20 min at room temperature (see Note 53).


3. During the equilibration step, rinse the glass cassettes with water to remove 

adhering acrylamide. Rinse the surface of each gel with Milli-Q ® water and 

invert to drain (see Note 54). 

4. Remove the IPG strips from the equilibration tray and wash each side with 3 

mL of 1× running buffer (see Note 55). 

5. Hold the gel cassette in a horizontal position with the short glass plate side up. 

Using forceps, place the IPG strip onto the long glass plate with the plastic 

backing against the plate. By carefully pushing against the plastic backing of 

Fig. 2. Examination of the cell wall proteome of S. pyogenes strain 5448 (serotype 

M1). The mutanolysin cell wall extract was harvested after growth at 37 °C to late 

stationary phase in THBY medium without agitation. The extracts were concentrated 

by TCA precipitation, isoelectric focused over a pH range of 4–7 and resolved with 

a 12.5% SDS-PAGE gel. The gel was stained with colloidal Coomassie and several 

landmark proteins identified by MALDI-TOF peptide mass fingerprinting analysis. 

Identified protein spots are denoted by numbered arrows, which correspond to the 

proteins in Table 1. Molecular mass markers are given in kilo-Daltons (kDa).


the strip with a spatula, slide the strip between the glass plates down to the gel 

surface. Take care not to damage the gel matrix with the spatula during this step. 

The strip should rest gently against the gel surface with the positive (acidic) end 

against the left edge of the glass plate (see Note 56). 

6. Apply 10 μL of prestained molecular weight markers to a piece of filter paper 

or electrode wick paper. Allow the markers to soak in and position on the gel 

surface next to one end of the IPG strip with forceps. 

7. Overlay the IPG strip and prestained markers with 3 mL of molten agarose gel 

overlay solution (see Note 57) and allow to set for 1 min. 

8. Insert the gels into the cassette carrier and add blank casset inserts to empty gel 

slots. Secure the gel cassets with the upper buffer chamber seal and place the 

assembly into the lower buffer chamber. 

9. Add approximately 4.0 L of 1× running buffer to the lower chamber and approximately 

0.8 L of 2× running buffer to the upper chamber. Adjust the buffer levels 

of both chambers until they are equal and between the minimum and maximum 

fill lines marked on the tank (see Note 58). 

10. Attach the lid of the Electrophoresis Unit and connect to a suitable power supply. 

Run at 2.5 W per acrylamide gel for 30 min; 40 W for 1 h; and 80 W for 4–5 h 


3.6.5. Protein Detection and Gel Documentation 

1. Carefully remove the IPG strips from the top of the gel and trim the gel bottom 


2. Submerge the gel in colloidal Coomassie stain and incubate overnight with 

gentle rocking (see Note 61). 

3. Discard the stain and rock the gel in destain solution overnight. Repeat at least 

once with fresh destain solution until the background is clear. 

4. Document gel with the GS-800 calibrated imaging densitometer (Bio-Rad) or 

equivalent. The 2DE pattern of a representative S. pyogenes cell wall extract 

is presented in Fig. 2. The mutanolysin extracts are highly enriched with cell 

wall-associated proteins, as evidenced by the MALDI-TOF mass spectrometry 

identification of several well-characterized cell wall proteins of S. pyogenes 

(Table 1). 

5. Store gel at 4 °C in a sealed plastic bag containing water (short term) or water 

with 0.005% (w/v) sodium azide for long term storage. 

4. Notes 

1. Dissolve the required amount of PMSF in a small volume of ice-cold isopropanol 

and adjust to final volume with chilled TES buffer. 

2. APS breaks down in water resulting in rapid loss of reactivity. APS solutions 

should be prepared fresh each time. 

3. Add required volume of 1 M DTT to buffer just before use.

Table 1 

Landmark cell wall-associated proteins identified by MALDI-TOF peptide mass fingerprinting analysis for S. pyogenes 

strain 5448 (Serotype M1) 

Spot Protein Function or pathway Accession no. a Molecular mass (kDa) b pI b Peptidematch c Coverage (%) d 

1 Enolase Virulence factor P69950 47.2 4.74 24 68.9 

(SEN) 

2 GAPDH Virulence factor Q5XDW3 35.8 5.34 19 57.0 

3 Manganese- Virulence factor Q8P0D4 22.5 4.87 9 57.0 

dependent 

Superoxide 

dismutase 

a Swiss-Prot/TrEMBL accession number. 

b Theoretical values obtained from Swiss-Prot/TrEMBL database. 

c Number of tryptic peptides detected by MALDI-TOF MS that could be matched to the protein. 

d Percentage of protein sequence covered by the matched peptides.


4. Bio-Lyte ® 3/10 ampholyte (Bio-Rad) is recommended for all pH ranges. 

5. Bromophenol Blue tracking dye allows monitoring of electrophoresis. 

6. Combine ammonium sulfate, methanol and phosphoric acid. Bring to volume 

with Milli-Q ® water and dissolve by vigorous stirring. Add the Coomassie 

G250 and stir vigorously with heating. Do not filter solution in order to retain 

colloidal dye particles. Discard stain after each use. 

7. To minimize contamination and ensure cell viability, streak from glycerolized 

stocks of S. pyogenes stored at –80 °C rather than plates stored at 4 °C. Use 

fresh blood agar plates so that hemolysis is clearly visible. 

8. Hold plate up to light or use a light box to ensure the colony is hemolytic. 

9. The optical density at 600 nm (OD 600) for late stationary phase cultures of 

S. pyogenes is approx 1.2. 

10. PMSF irreversibly inhibits serine proteases and some cysteine proteases which 

may be liberated during the extraction process. 

11. Seal the microcentrifuge tube lid with Parafilm ® and secure to incubator in a 

horizontal position to ensure thorough mixing. 

12. Mutanolysin cell wall extracts can be stored for at least 2 yr at –20 °C. 

13. This step is used to verify the extraction of cell wall-associated proteins before 

sample preparation for 2DE. 

14. The BCA assay is more sensitive than the Lowry assay and has less variability 

than the Bradford assay. Protein quantitation should be performed before 2DE 

as the solubilization solution reagents interfere with the BCA assay. Working 

Reagent is stable for one day at room temperature. 

15. Standards should be prepared using the same diluent as the cell wall extracts to 

compensate for buffer interference. 

16. To facilitate mixing, add each sample to the microtiter plate before adding the 

Working Reagent. 

17. A purple colored BCA-copper complex is generated in the BCA protein assay. 

The samples may also be incubated at 60 °C for 15 min or at room temperature 

(25°C)from2htoovernight. Incubation at higher temperatures will increase 

the sensitivity of the assay. 

18. An absorbance range of 540–590 nm may also be used with minimal loss of 

signal. A standard curve should be determined for each assay. 

19. Atmospheric oxygen inhibits the polymerization of acrylamide. For consistent 

results, degas the solution without the APS and TEMED under vacuum for at 

least 15 min. Add APS and TEMED and swirl the solution gently but thoroughly 

to minimize the introduction of oxygen. 

20. The 12% resolving gel has an approximate separation range of 14.4 kDa to 

120 kDa. 

21. Remove the ethanol overlay after 1htoprevent dehydration of the gel surface. 

22. Slowly load the sample to avoid a diffuse loading zone and loss of band 

sharpness. Load 15 μL of 1× loading buffer in unused wells to prevent the lateral 

spread of adjoining samples.


23. The voltage and current used for SDS-PAGE is potentially lethal. Use all 

equipment in accordance with the manufacturer’s instructions. 

24. The detection limit for Coomassie Blue staining is 0.1–0.5 μg protein. 

25. The Coomassie Blue stain can be reused several times. 

26. The addition of a piece of sponge or paper towel to a corner of the tray will 

facilitate the removal of excess Coomassie Blue stain. Rapid destain solution 

can be recycled by filtering through activated carbon (Sigma). If loss of band 

intensity occurs because of excessive destaining, restain with Coomassie and 

destain as described. 

27. The addition of glycerol to the final destain solution prevents the cracking of 

gels during drying. 

28. TCA is a hazardous substance and care should be taken to avoid exposure. The 

TCA precipitation step is used to increase protein concentration before 2DE. 

29. The pellet is usually invisible, but can sometimes be seen as a white smear. To 

avoid disturbing the pellet, carefully add the ethanol and gently tap the tube to 

mix. 

30. Avoid over-drying as this will make the pellet difficult to resolubilize. TCA 

precipitated cell wall proteins can be stored for at least 2 yr at –20 °C. 

31. Poor protein solubilization may cause horizontal or vertical streaking in 2D gels. 

Do not heat to resolubilize. Samples containing urea must not be heated above 

35 °C to avoid protein carbamylation. 

32. A water-bath sonicator is suitable for this step. Alternatively, a sonicator with a 

microtip may be used. 

33. Centrifugation removes particulate material which can block the gel pores of 

the IPG strip, resulting in poor focusing and horizontal streaking. 

34. The protocol may be stopped at this point and the sample stored at –20 °C. 

35. Load up to 1 mg protein for Coomassie stained 2D gels. Sample overload may 

cause horizontal or vertical streaking. Do not exceed a total loading volume of 

250 μL per strip. 

36. A trace amount of Bromophenol Blue is included for monitoring of IPG strip 

rehydration and subsequent electrophoresis. 

37. Evaporation will cause the urea to precipitate. 

38. The minimum rehydration time is 11 h. 

39. Thoroughly cleaned rehydration trays may be re-used. 

40. Drainage of the oil removes unabsorbed protein and minimizes horizontal 

streaking in 2D gels. The IPG strip may be wrapped in plastic wrap and stored 

at –80 °C. 

41. Always wear laboratory gloves when handling IPG strips and all apparatus/ 

solutions used in their preparation to prevent contamination from skin keratin. 

42. Electrode wicks are highly recommended because they remove salts and other 

contaminants in the sample. 

43. The majority of S. pyogenes cell wall-associated proteins have a pI between 4 

and 7 (8). The appropriate pH range for other Gram-positive prokaryotes should 

be determined empirically.


44. Covering the strip with mineral oil prevents evaporation and carbon dioxide 

absorption during focusing. 

45. The maximum permissible voltage is 8,000 V. The focusing range for 11 cm 

IPG strips is 15,000–60,000 Vh. Incomplete or excessive isoelectric focusing 

may cause horizontal or vertical streaking in 2D gels. Focusing time should be 

kept to the minimum necessary and must be determined empirically. 

46. If required, the IPG strip may be wrapped in plastic wrap and stored indefinitely 

at –80 °C without having a detrimental effect on the final 2D pattern. Thaw at 

room temperature for 10–15 min before equilibration. 

47. Firmly push the separator sheets and cassettes against the bottom of the caster 

to ensure a good seal. If a full set of gels is not required, use blank cassette 

inserts with separator sheets to occupy the extra space. 

48. Push down firmly during this step to ensure the faceplate forms a tight seal. 

49. A homogenous gel containing 12.5% total acrylamide has a separation size range 

of 14 to 100 kDa. 

50. For consistent results, degas the monomer solution (excluding the APS and 

TEMED) under vacuum for 15 min before adding the APS and TEMED. 

Degassing is not essential, but will accelerate polymerization. 

51. For longer term storage, unload the gel cassettes from the caster, wrap in paper 

towel and completely submerge in gel storage buffer. Store in a sealed container 

at 4 °C for up to 3 months. 

52. Wear gloves when preparing buffers and handling 2DE gels to prevent protein 

contamination, primarily with skin keratin. 

53. IPG strip equilibration should be undertaken before second dimension separation. 

This step enhances the solubility of focused proteins and allows the binding 

of SDS for second dimension separation. Ineffective equilibration may cause 

vertical streaking in 2D gels. Thoroughly cleaned equilibration trays may be 

re-used. 

54. Ensure the glass casset and gel surface are dry before loading the IPG strips. 

Remove excess water from the wells with blotting paper, taking care to avoid 

the gel surface. 

55. This step lubricates the strip, preventing it from adhering to the glass plates 

during loading. 

56. Two 11 cm strips or three 7 cm strips fit onto a 26 × 20 cm gel. Avoid air bubbles 

between the strip and the gel surface or between the strip backing and glass 

plate. A gap between strip and gel, or damage to the strip during application, 

may cause vertical streaking in 2D gels. 

57. Melt the agarose gel overlay solution in a microwave oven. Allow to cool slightly 

(60 °C) to prevent the decomposition of urea in the equilibration buffer. Pipet 

slowly to avoid introducing air bubbles under or behind the strip, which may 

interfere with protein resolution. Air bubbles should be removed immediately 

with a spatula. 

58. Depletion of ions in the running buffer can result in poor resolution and vertical 

streaking. To avoid this, discard the running buffer after approx 3 runs.


59. Gels can be run overnight at 1.5 W per gel. The Bromophenol Blue in the 

agarose gel overlay solution is used to monitor migration. 

60. Trim around the sides of the gel to ensure it does not adhere to the spacers and 

tear. 

61. Colloidal Coomassie can detect down to 100 ng/protein spot. 


This work was supported by the National Health and Medical Research 

Council of Australia. We wish to thank Stuart Cordwell (School of Molecular 

and Microbial Biosciences, The University of Sydney, NSW, Australia) for 

assistance with the MALDI-TOF MS analysis. 

References 

1. Ton-That, H., Marraffini, L. A., and Schneewind, O. (2004) Protein sorting to 

the cell wall envelope of Gram-positive bacteria. Biochim. Biophys. Acta 1694, 

269–78. 

2. van Heijenoort, J. (2001) Formation of the glycan chains in the synthesis of 

bacterial peptidoglycan. Glycobiology 11, 25R–36R. 

3. Sjoquist, J., Movitz, J., Johansson, I. B., and Hjelm, H. (1972) Localization of 

protein A in the bacteria. Eur. J. Biochem. 30, 190–4. 

4. Navarre, W. W., and Schneewind, O. (1999) Surface proteins of Gram-positive 

bacteria and mechanisms of their targeting to the cell wall envelope. Microbiol. 

Mol. Biol. Rev. 63, 174–229. 

5. Chassy, B. M., and Giuffrida, A. (1980) Method for the lysis of Gram-positive, 

asporogenous bacteria with lysozyme. Appl. Environ. Microbiol. 39, 153–8. 

6. Yokogawa, K., Kawata, S., Takemura, T., and Yoshimura, Y. (1975) Purification 

and properties of lytic enzymes from Streptomyces globisporus 1829. Agric. Biol. 

Chem. 39, 1533–43. 

7. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the 


8. Cole, J. N., Ramirez, R. D., Currie, B. J., Cordwell, S. J., Djordjevic, S. P., 

and Walker, M. J. (2005) Surface analyses and immune reactivities of major cell 

wall-associated proteins of group A Streptococcus. Infect. Immun. 73, 3137–46.

25 

Cell Fractionation of Parasitic Protozoa 

Wanderley de Souza, José Andrés Morgado-Diaz, 

and Narcisa L. Cunha-e-Silva 

Summary 

Cell fractionation, a methodological strategy for obtaining purified organelle preparations, 

has been applied successfully to parasitic protozoa by a number of researchers. 

These studies have provided new information of the cell biology of these parasites and 

have supported investigators to assume that some of the protozoa form the roots of the 

evolutionary tree of eukaryotic cells. The cell fractionation usually starts with disruption of 

the plasma membrane, using conditions that minimize damage to the membranes bounding 

intracellular organelles. An important requirement for successful cell fractionation is the 

evaluation of the isolation procedure that can be made by morphological and biochemical 

methods. The morphological approaches use light and electron microscopy of thin section 

of different fractions obtained, and the biochemical methods are based on the quantification 

of marker enzymes or other molecules (for instance, a special type of lipid, an 

antigen, etc.). Here we will present our experience in the isolation and characterization of 

some structures found in trypanosomatids and trichomonads. 

Key Words: Cell fractionation; electron microscopy; marker enzymes; parasitic 

protozoa; trichomonads; trypanosomatids. 


Parasitic protozoa are responsible for a large number of diseases affecting 

millions of people throughout the world. Some are caused by protozoa, which 

establish an initial binding to cells from the host and then invade them 

using several alternative mechanisms. Among these protozoa we can mention 

members of the Trypanosomatidae family, causative agents of Chagas’ disease 

and leishmaniasis, and members of the Apicomplexa group, which include 

From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractioation, Volume 2 


313

314 de Souza et al. 

agents of malaria and toxoplasmosis. Other protozoa exert their parasitic 

activity via interaction with epithelial cells from the gastrointestinal and 

urogenital cavities. Examples include the agents of giardiasis, trichomoniasis, 

and amebiasis. To exert a parasitic activity these protozoa present 

several biosynthetic and metabolic pathways, which make them different from 

mammalian cells. 

Transmission electron microscopy of thin sections has provided most of the 

available information on the structural organization of parasitic protozoa. Structures 

typically found in mammalian cells, such as the nucleus, endoplasmic 

reticulum, Golgi complex, mitochondria and lysosomes are found in most, 

but not in all protozoa. Some lack structures such as the Golgi complex 

and mitochondria. Others present special secretory organelles, such as the 

micronemes, rhoptries, and dense granules found in Apicomplexa, or metabolic 

organelles, such as the hydrogenosome, found in trichomonads. One approach 

that has been used to obtain new information on structure and composition of 

structures and organelles found in parasitic protozoa is cell fractionation. Here 

we will present our experience in the isolation and characterization of some 

structures found in trypanosomatids and trichomonads. Figs. 1 and 2 show 

general views of the structural organization of Trypanosoma cruzi and Tritrichomonas 

foetus, respectively, based on information obtained by transmission 

electron microscopy. As can be seen in the figures these protozoa present well 

Fig. 1. Longitudinal section of a Trypanosoma cruzi epimastigote depicting the 

the typical positions of the nucleus (N), kinetoplast (K), anterior flagellum (F) and 

reservosomes (R) at the posterior region of the parasite. Top inset shows a detail of 

reservosome structure and bottom inset shows Golgi complex (GC), always positioned 

next to the flagellar pocket, but that was not at the plane of the longitudinal section. 

Bars: 0.5 μm.

Cell Fractionation of Parasitic Protozoa 315 

Fig. 2. Longitudinal section of Tritrichomonas foetus showing the nucleus (N), Golgi 

complex (G), hydrogenosomes (H) and glycogen particles (asterisk) (Micrograph by 

M. Benchimol). Bar = 0.5 μm. 

elaborated cytoskeletal structures, known as the subpellicular microtubules and 

the pelta-axostyle system, which make them resistant to cell breakage. 

2. Materials 

2.1. Cell Culture and Standard Equipment 

1. Trypanosomatids: Liver Infusion Trypticase Medium (LIT) (1) supplemented with 

10% heat-inactivated bovine serum (Gibco/BRL). 

2. Trichomonads: Trypticase Yeast Maltose Medium (TYM) (2) supplemented with 

10% heat-inactivated bovine serum (Gibco/BRL). 

3. Ultrasonic apparatus (Sigma, GEX 600 Model) with standard probe (13 mm 

radiating diameter. 

4. Sorvall RC 5B centrifuge with GSA and SS-34 Rotors.


5. Ultracentrifuge with swinging-bucket (Beckman SW 50.1, SW 28 and fixed angle 

Type 65) rotors. 

6. Potter-type homogenizer with Teflon pestle. 

2.2. Electron Microscopy 

1. Glutaraldehyde grade II (Ted Pella Inc.) is stored at 25% at –20 °C. Working 

solutions of 2.5% is prepared by diluting in Cacodylate buffer 0.1M, pH 7.2 

(Sigma). 

2. Osmium tetroxide (Ted Pella Inc.) is dissolved at 2% in distillated water and 

stored in a single use aliquot at –20 °C. Working solutions is prepared by dilution 

in cacodylate buffer plus 5 mM CaCl2 and 0.8% potassium ferrocyanide. 

3. Acetone p.a: solutions of 30, 50, 70, 90, and 100%. 

4. Epoxy resin: Polybed 812 (Electron Microscopy Sciences, EMS). 

5. Aqueous solutions of uranyl acetate (Electron Microscopy Sciences, EMS) and 

lead citrate. 

2.3. Isolation and Subfractionation of the Flagellum 

of Trypanosomatids 

1. Buffer A: 25 mM Tris–HCl, pH 7.4, 0.2 mM EDTA, 5 mM MgCl2,12mM μ-mercaptoethanol, 320 mM sucrose. 

2. Sonication buffer: buffer A supplemented with 1% bovine serum albumin (BSA) 

and1mMCaCl2 and protease inhibitors cocktail (Sigma). 

3. Sucrose gradient solutions for the isolation of flagellum: 0.8, 1.65, and 1.85M 

prepared in buffer A without sucrose. 

4. 2% Triton X-100 prepared in buffer A. 

5. Sucrose gradient to isolation of the flagellar membrane: 1.3, 1.6, 1.9, and 2.2M 

prepared in buffer A without sucrose. 

6. 0.0001% trypsin (type XIII-treated, Sigma). 

7. Soybean trypsin inhibitor (Sigma). 

8. Continuous sucrose gradient ranging from 1.8 to 2.2M 

2.4. Isolation of the Reservosome 

1. TMS buffer: 20 mM Tris-HCl, pH 7.2, 2mM MgCl 2, 250 mM sucrose. 

2. Sucrose solutions: 2.3, 1.2, 1.0, and 0.8 M prepared in TMS buffer without sucrose 

(TM buffer). 

3. TMS buffer supplemented with a cocktail of protease inhibitors (Sigma). 

2.5. Isolation of the Golgi Complex 

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

inhibitors.


3. Gradient sucrose solutions: 2.3, 1.2, 1.0, and 0.8M prepared in G buffer without 

sucrose. 

4. 150 mM sodium carbonate, pH 11.5. 

2.6. Isolation and Subfractionation of the Hydrogenosomes 

1. H buffer: 10 mM Tris-HCl, pH 7.2, 0.25M sucrose and 2 mM MgCl 2. 

2. Percoll gradient solutions (Pharmacia): 53, 45, 24, and 18% prepared in G buffer. 

3. V buffer: 2% Triton X-100, 50 mM 3-(N-morpholino)propanesulfonic acid 

(MOPS))-NaOH, pH 7.0, 0.1 mM MgCl 2,1nM dithiothreitol (Sigma) and 10% 

sucrose. 

4. Proteinase K (Sigma) at a final concentration 0.5 mg/mL. 

5. Quench solution: phenylmethylsulphonyl fluoride (PMSF) (final concentration 40 

μg/mL). 

3. Methods 

3.1. Isolation of Trypanosomatid Structures 

3.1.1. Isolation and Subfractionation of the Flagellum 

of Trypanosomatids 

Trypanosomatid flagellum exhibits a huge lattice-like filamentous structure 

called paraflagellar rod (PFR), running alongside a canonical 9+2 axoneme 

(3,4). The flagellar membrane, which has been considered a special domain, 

although continuous with cell body membrane, surrounds both structures. PFR 

plays a role in protozoa movement, as was indicated by a Trypanosoma brucei 

mutant that did not assemble the structure and was immotile (5). Flagellar 

membrane has a crucial participation in the interaction of T. cruzi with invertebrate 

host and parasite differentiation (6). The knowledge concerning the role 

played by trypanosomatid flagellum in the life cycle of these parasites and 

in the pathogenesis they cause in their vertebrate hosts has advanced using 

flagellar fractions as immunogen (7) or for biochemical and ultrastructural 

studies (8). There are two successful protocols for obtaining purified flagellar 

fractions: a classical one (9), deflagellating the protozoa and isolating the 

detached flagella by sucrose gradients, and a more simple method (10) that 

extracts the membranes of the whole protozoa and depolymerizes the cell body 

cytoskeleton by high salt treatment. The advantage of the latter method is that 

it does not take long, but it has a low yield, being limited to 105 parasites at 

a time. Here we describe the classical method that, although laborious, can be 

applied to 1012 parasites in a single experiment, yielding an abundant highly 

purified flagellar fraction. 

PFR is a highly conserved structure among trypanosomatids (11), allowing 

the adoption of the PFR from Herpetomonas megaseliae, a nonpathogenic


trypanosomatid, as a model for ultrastructural and biochemical studies. The 

protocols presented below have already been published for whole flagella purification 

(9), PFR (12), and flagellar membrane isolation from both H. megaseliae 

and T. cruzi (8). 

1. Start with 10 11 –10 12 protozoa. Harvest protozoa by centrifugation and wash 

twice in ice cold buffer A. All subsequent steps should be performed in an ice 

bath (see Note 1). 

2. Washed protozoa (the pellet of the last washing step) are resuspended in 

sonication buffer: buffer A supplemented with 1% bovine serum albumin (BSA) 

and1mM CaCl 2 and protease inhibitors cocktail. Cell density is very important 

in this step. If you started with 8×10 11 protozoa, resuspend in 200 mL and 

sonicate in 50-mL aliquots (see Note 2). 

3. We use an ultrasonic apparatus adjusted to 8% of maximum amplitude output. 

Typically, 6–8 sonication cycles of 10 s on and 5 s off are sufficient to deflagellate 

most cells (see Note 3). 

4. Whole cells, deflagellated cell bodies and liberated nuclei are pelleted with a 

low speed centrifugation at 120g, 10 min (see Note 4). 

5. Supernatants are centrifuged at 6,800g, 15 min, the pellet resuspended to 15 mL 

with buffer A and divided in six aliquots. 

6. Each aliquot is equilibrated on top of 0.8M sucrose cushions and spun at 1,080g 

for 20 min (see Note 5). 

7. Upper layer of each cushion is carefully removed and pelleted all together at 

17,300g. This is the crude flagellar fraction (see Note 6). 

8. The purification is achieved by laying 1-mL aliquots of crude flagellar fraction 

on top of a 1.65M/1.85M sucrose gradient, each step with 2 mL, and centrifuging 

at 130,000g for3h. 

9. Flagella equilibrate at the interface 1.65M/1.85M that must be carefully removed 

with a long and thin Pasteur pipet. All the interfaces together are diluted with 

buffer A without sucrose and pelleted at 17,300g for 20 min. The pellet is 

resuspended in a small volume of buffer A (0.5 mL) and contains purified 

flagella. 

10. Phase contrast microscopy can be used for initial examination of the flagellar 

fraction, but the only way of evaluating purity is electron microscopy. A small 

aliquot can be fixed by adding glutaraldehyde to the final concentration of 2.5% 

in buffer A and processed for electron microscopy (Fig. 3). (see Note 7). 

11. H. megaseliae or T. cruzi flagella purified as above are submitted to detergent 

extraction by doubling sample volume with ice cold 2% Triton X-100 in buffer 

A and incubated for 15 min. on ice with intermittent agitation. 

12. Sample is laid on top of a 1.3/1.6/1.9/2.2M sucrose and centrifuged at 130,000g 

for3h. 

13. Purified flagellar membrane that equilibrates at the interface 1.3/1.6M is carefully 

removed, diluted with buffer A without sucrose, pelleted at 17,300g for 20 min, 

and resuspended in a small volume (0.05 mL) (Fig. 4).


Fig. 3. Flagellar fractions from Herpetomonas megaseliae. Shows purified whole 

flagella, with preserved axonemes and PFRs enclosed by loose-fitting membrane 

profiles. Bar = 0.2 μm; Adapted from (8). 

14. H. megaseliae or T. cruzi purified flagella are submitted to extensive membrane 

solubilization with three rounds of incubation in 2% Triton X-100 or Nonidet 

P-40 for 15 min at 4°C. After each round flagella are pelleted at 17,300g, 20 min. 

15. Demembranated flagella are resuspended in buffer A and incubated very briefly 

(30 s) at 28°C with 0.001% trypsin (type XIII, TPCK-treated, Sigma). The 

reaction is stopped by addition of a 20-fold excess of specific trypsin inhibitor 

(soybean trypsin inhibitor, Sigma). Shake in a Vortex at maximum speed for 1 

Fig. 4. Flagellar fractions from Herpetomonas megaseliae. Shows flagellar 

membranes purified from Fig. 3 preparation. Bar = 0.2 μm; Adapted from (8).


min, divide in six aliquots and apply on a continuous sucrose gradient ranging 

from 1.8 to 2.2M. 

16. Centrifuge at 130,000g for 3 h. Collect 0.8-mL aliquots from top to bottom, 

dilute with buffer A without sucrose and pellet at 17,300g for 30 min. The 

third and fourth fractions from top to bottom are highly purified PFRs, as can 

be evaluated by electron microscopy (Fig. 5) and by using antibodies against 

major PFR proteins (Fig. 6B) and alpha-tubulin (Fig. 6C) in Western blots. 

SDS-PAGE analysis of purified PFRs (Fig. 6A) allowed for the first time the 

identification of some minor proteins that constitute this complex structure. 

3.1.2. Isolation of the Reservosome 

Reservosomes are storage organelles from T. cruzi epimastigotes that concentrate 

proteases and are essential for differentiation into trypomastigote forms. 

They were considered prelysosomes but the lack of a molecular marker has 

precluded a more detailed studied of their role in T. cruzi life cycle. Their 

purification (13) opened up the possibility of defining the necessary marker. 

1. Start with 3×10 10 epimastigotes from mid log phase cultures (see Note 8). 

2. Harvest protozoa by centrifugation at 4,800g, 10 min, 4°C and wash in cold 

TMS buffer (see Note 9). 

3. Resuspend in 45 mL of TMS (about 6.5 × 10 8 parasites/mL) and divide in three 

aliquots. 

4. Sonicate each aliquot at 10% of maximum amplitude of ultrasonic apparatus, in 

about 15 cycles of 2sonand1soff. (see Note 3). 

5. Centrifuge at 2,450g, 10 min, pour the supernatant carefully into another tube 

and discard the pellet, containing non-ruptured cells, nuclei, kinetoplasts etc. 

Fig. 5. Flagellar fractions from Herpetomonas megaseliae. Shows that PFRs remain 

organized after purification from demembranated H. megaseliae purified flagella. 

Bar = 0.1 μm; Adapted from (12).


Fig. 6. (A) SDS-PAGE gel of fractions from the PFR purification gradient. Lanes 

1–4 represent fractions 3–6, in this order. Besides the major PFR proteins (arrowhead), 

fractions 3 and 4 are clearly enriched in a number of minor bands whose positions 

and estimated molecular masses are indicated on the left. Molecular mass markers are 

indicated on the right. (B and C) Western blots of different steps and gradient fractions 

from the paraflagellar rod (PFR) purification procedure probed with anti PFR major 

proteins antibody (panel B), and antialpha-tubulin monoclonal antibody (panel C). In 

both panels lanes 1–4 represent fractions 3–6, in this order, from the PFR purification 

gradient. Molecular mass markers are indicated on the left. Adapted from (12).


6. Mix the supernatant with an equal volume of 2.3M sucrose in TMS and divide 

in two Beckman SW28 centrifuge tubes (usually 12 mL in each tube) (see 

Note 10). 

7. Repeating the procedure with the other two aliquots, all six tubes of the rotor 

will be used. 

8. Overlay carefully with 10 mL of 1.2M sucrose in TMS, 9 mL of 1.0M sucrose 

and8mLof0.8M sucrose. 

9. Centrifuge at 97,000g for 150 min, without brake. 

10. Collect the interfaces 0.8M/1.0M (B1) and 1.0M/1.2M (B2), 1.2M/1.27M (B3) 

and the pellet (P), dilute with TM (TMS without sucrose) and pellet at 90,000g 

for 20 min. Reservosomes are highly purified in B1 (Fig. 7) and very enriched 

in B2. Microsomes and glycosomes are the major components of B3. Acidocalcisomes 

are enriched in the pellet. Fig 7 here 

11. A small aliquot of each fraction is fixed by adding glutaraldehyde directly 

in TMS for electron microscopy (see Note 7), another aliquot is reserved for 

marker enzyme assays (see next step), and protease inhibitors are added to a 

third aliquot for SDS-PAGE and Western blot assays. Purified fractions must 

be stored below –70°C. 

12. To evaluate the purity of the fractions marker enzymes for the organelles that 

could be contaminants of reservosome fraction should be assayed: succinate 

cytochrome c reductase for mitochondria (14), pyrophosphatase for acidocalcisomes 

(Table 1) and hexokinase for glycosomes (15). 

Fig. 7. Trypanosoma cruzi reservosomes purified fraction. Adapted from (13). 

Bar = 1 μm.


3.1.3. Isolation of the Golgi Complex 

In parasitic protozoan the Golgi complex is much less well organized, less 

prominent, and correspondingly more difficult to isolate than in mammalian 

cell. The success of the organelle isolation is dependent on choosing a good cell 

disruption method, which is dependent on the morphological characteristics of 

the parasite. Morphological studies have shown that the Golgi of trypanosomatids, 

and in particular of Trypanosoma cruzi, is made up of 4–10 stacked 

cisternae localized at the anterior region of the cell, close to the flagellar pocket 

(16). A method that preserves the Golgi with minimum damage during cell 

disruption was developed resulting in a fraction highly enriched in stacked Golgi 

cisternae and vesicles (19), as determined by electron microscopy (Fig. 8), and 

also in galactosyltransferase, O-- GlcNA transferase and acid phosphatase 

activities. Contamination with other organelles (endoplasmic reticulum, plasma 

membrane, mitochondria, and glycosomes) is minimal, as analyzed by their 

respective marker enzymes (Table 2). 

All centrifugation steps and other operations are performed at 4 °C or in an 

ice bath. 

1. Harvest cells in a GSA Rotor at 1,000g for 10 min and wash three times in G 

buffer. 

2. Resuspend the cells in a hypotonic solution (G buffer without sucrose) containing 

a cocktail of protease inhibitors and incubated in this solution for 10 min. 

3. Homogenize the cells by sonication on ice with 15 cycles of 2 s with 1 s rest 

between cycles in an ultrasonic apparatus. Monitor the cell disruption by phasecontrast 

microscopy (see Note 3). 

4. Immediately add to the homogenate a concentrated sucrose solution, prepared in 

G buffer to a final concentration of 0.25M sucrose to minimize osmotic damage. 

Table 1 

Enzyme assays in the epimastigote subcellular fractions of the purification of 

reservosome of Trypanosoma cruzi 

Succinate cytochrome c reductase a Pyrophosphatase b 

Total homogenate 0.030 0.11 

Supernatant from 2,450g 0.043 0.14 

B1 0 0 

B2 0.006 0.02 

B3 0.044 0 

B4 0.830 0.15 

a Specific activity in mM min −1 mg protein −1 . 

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


Fig. 8. General view of the Golgi fraction of Trypanosoma cruzi showing Golgi 

complex cisternae and several Golgi cisternae profiles. After (19). Bar = 0.5 μm. 

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

nuclei, and kinetoplasts. 

Table 2 

Enzyme activities in Golgi fraction and other subcellular fractions of 

Trypanosoma cruzi. a 

Assay 

Enzyme Activity bc 

Homogenate fraction PNS GF 

NADPH cyt c reductase b 025 034 0043 

Acid phosphatase b 30 45 152 

5’-nucleotidase b 0101 0157 006 

Succinate cyt c reductase b 007 004 003 

Hexokinase b 0032 00436 00034 

O-a-GlcNAc transferase c 0076 0248 2044 

Gal transferase c 128 656 11480 

PNS, post-nuclear supernatant; GF, Golgi fraction 

a The results are of a typical experiment in which each number represents the means of 

duplicate determinations. Minor modifications were observed in at least three independent experiments. 

b Specific activity expressed as μM of product released min −1 mg protein −1 . 

c Specific activity expressed as pM of [ 3 H]GlcNAc or [ 3 H]Gal transferred h −1 mg protein −1 .


6. Harvest the postnuclear supernatant (PNS), add an equal volume of 2.3M sucrose 

and homogenize for 10 min. 

7. Load 15 mL of PNS onto the bottom of Beckman SW 28 Rotor ultracentrifuge 

tubes and overlay sequentially with 7 mL each of 1.2M, 1.0M, and 0.8M sucrose. 

8. Centrifuge the gradient at 95,000 x g for 1h 30 min. The band at the interface of 

1.0/1.2M sucrose corresponds to highly enriched Golgi complex fraction. 

9. Collect the fraction by diluting with G buffer without sucrose and centrifuging at 

80,000g for 45 min (Beckman Type 65 Rotor). 

3.2. Isolation of Trichomonad Structures 

3.2.1. Isolation of the Golgi Complex 

Tritrichomonas foetus, a member of the Trichomonadidae family, possesses a 

well developed Golgi complex, consisting of tightly packed elongated cisternae 

often with dilated terminal regions, localized at the anterior portion of the 

cell and in close association with the parabasal filament (18). The following 

protocol yields two subfractions containing light (GF1) and heavy (GF2) Golgi 

membranes (17). At the ultrastructural level GF1 fraction is constituted only 

of smooth-membrane structures, particularly cisternae and secretory vesicles 

(Fig. 9A) and contains a 20-fold enrichment of galactosyltransferase activity. 

GF2 fraction is composed of profiles of stacked Golgi membranes with three or 

more tightly apposed cisternae with dilated borders and associated to vesicles 

(Fig. 9B). This latter fraction contains a 7-fold enrichment in galactosyltransferase 

activity. Minimal contamination with other organelles, such as 

endoplasmic reticulum, hydrogenosomes, and plasma membrane is observed in 

these fractions, as quantified by their respective enzymatic markers (Table 3). 


ice bath. 

1. Harvest the cells in a GSA Rotor at 1,000g for 10 min and wash three times in 

G buffer. 

2. Resuspend the cells in G buffer containing a cocktail of protease inhibitors and 

homogenize using 180 strokes of a Potter-type homogenizer. Stop the homogenization 

procedure when about 90% cell breakage is achieved as ascertained by 

phase-contrast microscopy. 

3. Centrifuge at 2,500g for 10 min to discard the pellet containing unbroken cells, 

nuclei, and hydrogenosomes. 

4. Add to the supernatant (PNS) an equal volume of 2.3M sucrose to make a 1.4M 

sucrose final concentration and homogenize for 10 min. 

5. Load 15 mL of PNS onto the bottom of SW28 ultracentrifuge tubes and overlay 

in succession with 7 mL each of 1.2M, 1.0M, and 0.8M sucrose. 

6. Centrifuge at 95,000g for 2.5 h and carefully remove two bands from the top at 

0.8/1.0 (GF1) and 1.0/1.2M (GF2).


Fig. 9. General aspect of the Golgi fractions of Tritrichomonas foetus. (A) GF1 

fraction showing several vesicles (arrowheads) and cisternae profiles (arrows). (B) GF2 

fraction showing several intact stacks of Golgi complex (arrowheads). After (17). Bars 

= 0.2 μm (A) and 0.5 μm (B). 

7. Dilute these fractions in G buffer without sucrose and collect them by centrifugation 

at 80,000g for 30 min (Beckman Type 65 Rotor). 

8. Further fractionation of these two compartments can be attempted by alkaline 

treatment to obtain the Golgi content and Golgi membrane subfraction. Blend the 

Table 3 

Distribution of Galactosyltransferase and other enzyme markers in Golgi 

fractions of Tritrichomonas foetus. a 

Fractions Total 

protein 

(mg) 

Galactosyl 

transferase 

(pmole/30 

min/mg 

protein) 

NADPH cit c 

reductase 

(μmole/min/mg 

protein) 

NADP-malic 

enzyme 

(μmole/min/mg 

protein) 

5’Nucleotidase 

Homogenate 3920 4434 0035 0012 0.93 

PNS 1040 6804 00079 0026 0.89 

GF2 20 31494 00029 0006 1.50 

GF1 10 87400 n.d. n.d. 0.66 

a Results are expressed as the means of duplicate determinations. n.d., not detectable; Number 

of experiments is given in parenthesis; PNS, postnuclear supernatant; GF2, heavy Golgi fraction; 

GF1, light Golgi fraction.


Golgi fractions (GF1 and GF2), collect by centrifugation for 30 min at 80,000g 

(Beckman Type 65 Rotor), and treat with 150 mM sodium carbonate, pH 11.5, 

for 30 min. 

9. Collect the subfractions, again as in step 7. 

3.2.2. Isolation and Subfractionation of the Hydrogenosome 

T. foetus is a facultative anaerobic protist that lacks mitochondria, but 

contains another organelle, designated as the hydrogenosome. This organelle 

appears as a spherical or slightly elongated structure with a mean diameter 

of 0.5 μm, reaching 2 μm in dividing organelles. A common characteristic of 

most hydrogenosomes is the presence of a peripheral vesicle, which has been 

suggested to be a specialized sub-compartment (20). Additional information on 

this important organelle, which seems to be the main target of drugs used in 

the chemotherapy of trichomoniasis, can be obtained using hydrogenosomes 

isolated in a way that preserves their structure. Using Percoll gradients, we 

developed a procedure (21), which yielded the purest fraction obtained for 

hydrogenosomes of T. foetus (Fig. 10A) and allowed further sub-fractionation 

to isolate the peripheral vesicle (Fig. 10B). It is enriched about 12-fold in malic 

enzyme activity and free of other contaminant organelles (Table 4). Major 

hydrogenosome proteins were identified by SDS-PAGE (Fig. 10C). 


ice bath. 

1. Harvest the cells in a GSA Rotor at 1,000g for 10 min and wash three times in 

H buffer. 

2. Resuspend the cells in 30 mL of H buffer and homogenize using about 180 strokes 

of a Potter-type homogenizer. Stop the homogenization while some unbroken 

cells are still present. 

3. Dilute the homogenate in the proportion 1:10 with H buffer and centrifuge at 

1,500g for 10 min. Discard the pellet containing unbroken cells, nuclei and costa. 

4. Centrifuge the supernatant at 4,000g for 10 min. The pellet of this centrifugation 

corresponds to an enriched hydrogenosomal fraction. 

5. Resuspend the pellet in ice-cold H buffer and layer aliquots of 1 mL on the top 

of a Percoll gradient of 53, 45, 24, and 18% in H buffer (1 mL each solution). 

6. Centrifuge the gradient using a swinging bucket rotor (Beckman SW 50.1) at 

36,000g for 30 min, without using the brake. The pure fraction containing wellpreserved 

hydrogenosomes is recovered from the pellet of the Percoll gradient. 

7. Incubate a sample of the purified hydrogenosomal fraction in V buffer on ice for 1 

h. and centrifuge the suspension at 25,000g for 20 min (Beckman Type 65 Rotor). 

The supernatant (TX-supernatant) is made up of solubilized hydrogenosome 

membranes and the pellet (TX-pellet) of hydrogenosomal matrix attached to the 

peripheral vesicle.


Fig. 10. (A) Electron micrograph showing the purified hydrogenosomes of Tritrichomonas 

foetus. The hydrogenosome matrix is homogeneous and the peripheral 

vesicles present an electron-dense content. Bar = 1 μm (B) Purified fraction of 

hydrogenosomal flat vesicles Bar = 0.5 μm (C) SDS-PAGE of the fractions obtained 

during the fractionation of T. foetus. Lane 1: whole homogenate, Lane 2: fraction before 

the Percoll gradient, Lane 3: purified hydrogenosomes. After (21). 

Table 4 

Specific activity and recovery of malic dehydrogenase in fractions of the 

purification of hydrogenosomes of Tritrichomonas foetus. a 

Fractions Total protein 

(mg) 

Specific activity (μmol 

min −1 mg protein) 

Enrichment 

(-fold) 

Yield (%) 

Whole homogenate 4500 0035 10 100 

Po fraction 580 0147 42 54 

Pellet gradient 80 0421 120 22 

a Po is the fraction before the Percoll gradient. Results are expressed as the means of duplicate 

determinations in three representative experiments.


8. Wash a sample of the TX-pellet twice in H buffer and incubate for 45 min at 

room temperature with proteinase K (final concentration 0.5 mg/mL). 

9. Quench the mixture with PMSF (final concentration 40 μg/mL) on ice, for 15 

min and centrifuge at 350,000g for 30 min (Beckman Type 65 Rotor). The pellet 

of this centrifugation corresponds to the purified peripheral vesicle. 

4. Notes 

1. Three liters of axenic culture of H. megaseliae promastigotes in Warren’s 

medium supplemented with 5% fetal calf serum are enough for obtaining 8 

×1011 protozoa, after 30–36 h at 28°C. The same cell density of T. cruzi 

epimastigotes can be obtained with 5Lofparasites cultivated for 5dat28°C 

in LIT medium supplemented with 10% fetal calf medium (1). A good rotor to 

harvest a large volume culture is GSA of Sorvall ultracentrifuges. 

2. It is not necessary to use purified BSA to prepare sonication buffer; fraction V 

grade is good enough. The composition of protease inhibitors cocktail may vary, 

it is very important that it contains an irreversible cysteine protease inhibitor, 

such as E-64, because this class of protease is the most active in trypanosomatids. 

We use protease inhibitors cocktail for general use, from Sigma. 

3. It is essential to control deflagellation by phase contrast microscopy and stop 

the procedure when the majority, but not all the protozoa are deflagellated, 

otherwise some nuclei may rupture and free chromatin aggregates the organelles, 

including free flagella. 

4. Turn off centrifuge brake because this loose pellet resuspends easily. This step 

and the next are performed with Sorvall SS-34 fixed angle rotor. The supernatant 

of the low speed centrifugation should be controlled by phase contrast 

microscopy and this step should be repeated twice to clean the supernatant from 

whole cells and deflagellated cell bodies. 

5. The sucrose cushion must be spun in a swinging bucket rotor, as Beckman 

SW50.1, which must have each tube filled up to total volume (5 mL). In this 

step as well as in the gradient, the rotor should stop without using the brake. 

6. To improve the isolation yield, the cushion can be repeated using the bottom 

layer as the starting sample of step 6. 

7. A brief protocol for sample processing for electron microscopy: as fractions are 

easy to fix, 30 min of incubation in fixative solution is enough. On the other 

hand, fractions are not easy to wash, as the usual centrifugation of few minutes 

in a microfuge may not be sufficient to pellet the sample adequately and thus 

you would be losing your sample in the washing steps. To avoid this, wash in the 

ultracentrifuge at the same speed used to obtain the fraction until before osmium 

incubation when the fraction becomes denser. If you cannot process the samples 

in a short time (next day), change the fixative solution to 2.5% glutaraldehyde 

in 0.1 M cacodylate buffer pH 7.2, as Tris buffers are not good for electron 

microscopy. After fixation, samples are washed twice in 0.1M cacodylate buffer 

and post fixed with 1% osmium tetroxide in 0.1M cacodylate buffer containing


0.8% potassium ferrocyanide and 5 mM calcium chloride for 20 min at room 

temperature in the dark (wear gloves and work in a fume hood!). Wash three 

times (or until the supernatant is clear) in 0.1M cacodylate buffer. Dehydrate in 

acetone series from 30 to 100%, 10 min incubation in each bath, repeating the 

last one twice. Infiltrate slowly in Epon (acetone + Epon 1:1 overnight, Epon 

for 6 h). Embed in Epon at 60 o Cfor2d. 

8. Epimastigotes from 5-day-old cultures are ideal for reservosome purification. 

Three liters of culture in LIT medium supplemented with 10% fetal calf serum 

(1) maintained at 28°C under gentle orbital agitation are enough to obtain 3 × 

10 10 parasites. 

9. It is very important to work fast and in an ice bath, because reservosomes start to 

degrade their contents as soon as parasites are washed out from culture medium. 

It is adequate to add protease inhibitors to TMS buffer (mainly E-64, inhibitor 

of cysteine proteases, highly concentrated in the organelle), unless you just want 

to study the properties of reservosome proteases and their inhibitors. 

10. The bottom step of this gradient contains the sample and corresponds to 1.27M 

sucrose, very close in density to the next step. So, take care when preparing 

and measuring volumes of sucrose solutions before mixing sample with 2.3M 

sucrose. 


The authors dedicate this chapter to the memory of Luiz Henrique Monteiro 

Leal, who greatly contributed to improve the methods of cell fractionation of 

parasitic protozoa, and passed away during the elaboration of this manuscript. 

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Index 

A 

Acrylamide-Bis solution, 205 

Affinity Depletion Cartridge for removal 

of HSA, 91 

Albumin and immunoglobulin-depleted 

plasma proteins 

1D gel electrophoresis assessment of 

fractions, 20–21 

UV absorbance assessment of 


Albumin (BSA) standards color response 

curve using BCA assay, 123 

Albumin-depleted plasma proteins 

1D gel electrophoresis assessment of 


UV absorbance assessment of 


Alkaline phosphatase assay, biochemical 

principle, 252, 254 

Anti-HSA POROS cartridge, 23 

Anti-human albumin monoclonal 

antibody, 15 

Arabidopsis thaliana chloroplasts, 

isolation and preparation, 

171–172 

chloroplast isolation, 174–176, 

178–181 

establishing yield and intactness of 

chloroplasts, 176 

growth of Arabidopsis seedlings, 

173–174, 177 

preparation for proteomics, 176–177, 

182–183 

B 

BCA Protein Assay Kit, 117 

Beconase AQ pump aspirator spray 

device, 79, 80 

333 

Bicinchoninic acid protein assay, 30, 

121, 297 

Biological Variation Analysis mode 

(BVA), 11 

Bovine serum albumin (BSA), 204 

Bradford protein assay, 121 

Bronchoalveolar lavage fluid (BALF) 

cellular pattern of, 72–73 

2-DE analysis, 71–72 

during fiber-optic bronchoscopy, 67 

proteome analysis for, 68 

sample preparing 

dialysis-lyophilization, 71 

ultra filtration, 71 

C 

C. albicans protoplasts in active cell wall 

regeneration, proteomics of 

proteins secreted from, 261 

Castor bean endosperm isolation and 

fractionation of ER for 

proteomic analysis, 203–214 

isolation of, 206–208 

mass spectrometer compatible silver 

staining, 205, 209–211 

2D gel protein profiling of, 205–206, 

211–213 

SDS-PAGE analysis of developing 

and germinating of, 208–209 

SDS-PAGE analysis of purified ER, 

204–205 

tissue homogenization and ER 

purification, 204 

Cell physiology and disease molecular 

study, 139 

cell culture and lysis, 142 

isolation of nickel-binding proteins, 

142, 143–144

334 Index 

preparing samples for isolation of 

nickel-binding 

proteins, 143 

silver staining of metal-affinity 

enriched proteins compatible for 

mass spectrometry (MS), 

142–144 

Cell wall incorporation of GPI-anchored 

proteins model, 232 

Cell wall proteins (CWP), 187 

analysis 

and extraction protocol by 

bioinformatics, 198 

separation and identification of 

protein, 197–198 

extraction and analysis from liquid 

culture medium of 

seedlings, 189 

Cellular fungi sample preparation 

procedure, 261–266 

cell culture, 267 

cell disruption and protein 

solubilization, 267, 268–270 

S-35 in vitro cell labeling, 267–268 

scintillation counting and protein 

assay, 268, 270 

Cerebrospinal fluid (CSF) 

LC/MS/MS analysis processing, 63 

processing for 2-Dimensional gel 

electrophoresis, 59–60 

proteomics, 53–56 

gel electrophoresis, 57 

LC-MS/MS mass spectrometry, 58 

MALDI-TOF mass 

spectrometry, 58 

processing for 2-dimensional gel 


processing for LC/MS/MS analysis, 

57–58 

silver staining and in-gel 

digestion, 57 

simplification by removal of 

abundant proteins, 58–59 

Chicken IgY antibodies, immunoaffinity 

fractionation of plasma proteins 

IgY-12 and LC2/ LC10 high capacity 

spin column kit, 43 

IgY-12 microbeads for 96-well filter 

plates, 43 

SDS gel electrophoresis, 43–44 

Chloramphenicol, 224 

Chloroplast 

isolation buffer (CIB), 174 

isolation procedure, 175 

light and electron micrographs, 180 

lysis and solubilization, 182 

protein samples by DIGE 

2D-PAGE, 182 

proteomic studies, 173 

CLUSTAW program in PIR, 84 

Collision-induced dissociation (CID), 136 

Cysteine-specific protein labeling using 

CyDye DIGE fluor saturation 

dyes, 2 

Cytoplasmatic proteins 

buffers for western blot analysis, 103 

cell culture and wash buffer, 102 

cell harvest, 104 

comparison of 2DE pattern with 

pattern of whole cell 

protein, 109 

coomassie brilliant blue staining 

solutions, 104 

isolation from cultured cells 2-D gel 

electrophoresis, 101, 107–108 

buffers and reagents for, 102–103 

precipitation of cytoplasmatic 

proteins, 104–105 

silver staining solutions, 103 

western blot analysis for detection of 

purity, 105–107 

D 

DeCyder software, 10–11 

Depleted protein samples TCA 

precipitation for 2D gel 

electrophoresis, 22–23

Index 335 

Differential in-gel analysis (DIA), 11 

DIGE saturation labeling conditions, 7 

DNA electrophoresis, 156 

DryStrip Reswilling tray, 70 

Dulbecco’s Phosphate Buffered Saline 

(D-PBS), 116 

E 

Econo-Pac ® 10DG desalting column, 190 

Electrospray quadrupole time of flight 

mass spectrometer, 80, 83 

Eosin stain, 133 

Epithelium lining fluid (ELF), 67 

S-Ethanol-cysteine residue, 63 

Etioplasts, dark-grown plants, 173 

F 

FFPE tissue, 137 

Filamentous fungi secreted proteins 

isolation and enrichment, 

274–284 

centrifugation and ultrafiltration, 277 

enzymatic and chemical 

deglycosylation, 277–278 

filtration and lyophilization, 277 

isolation and concentration of 

supernatant broth, 278–280 

precipitation with trichloroacetic acid, 

methanol and chloroform, 277 

SDS-PAGE, 278, 282, 301 

two-dimensional gel electrophoresis, 

278, 282 

Fluorescence dye labeling 

optimization, 6–7 

Fluorescence resonance energy transfer 

(FRET), 4 

Foetal bovine serum (FBS), 116 

Formalin-fixed, paraffin-embedded 

(FFPE) tissues, 132 

sample preparation for mass 

spectrometry analysis, 131 

laser-capture microdissection, 134 

LC-MS/MS analysis of tryptic 

peptides, 133–134 

mass spectrometry analysis and 

bioinformatic 

analysis, 136 

nanoflow RPLC-MS/MS analysis, 

135–136 

protein extraction and trypsin 

digest, 133–135 

tissue processing, 133, 134 

trypsin digest, 133 

trypsin-mediated 18 O-labeling, 135 

G 

Gamborg liquid medium, 190 

Gel pieces, reduction and alkylation, 

61–62 

Glycosyl phosphatidylinositol (GPI)-cell 

wall proteins, 220 

Gram-positive bacterial cell 

wall-associated proteins 

isolation and solubilization 

bacterial culture, 296, 299 

BCA protein assay, 300 

casting 2D gels, 304 

cell wall-associated proteins 

extractions, 296–297, 300 

first dimension IEF, 303–304 

protein detection and gel 

documentation, 306 

SDS-polyacrylamide gel 

electrophoresis (SDS-PAGE), 

297 

second dimension SDS-PAGE, 

304–306 

trichloroacetic acid (TCA) 

precipitation, 298, 302 

Green plant tissue protein extraction, 

149–152 

H 

Hematoxylin stained histological 

classification, 5 

Herpetomonas megaseliae, 319, 320 

HiLoad 16/60 Superdex 75 prep grade 

column, 91

336 Index 

Hi-Trap SP equilibration 

buffer, 190 

HONE1 cells, Western blot analysis of 

cytoplasmatic fraction, 107 

Human BALF proteins, 2-D gel 

electrophoresis, 69 

Human nickel allergy, 140 

Human plasma 

albumin and immunoglobulin 

depletion of, 15–19 

proteins by 1D gel electrophoresis, 

20–21 

albumin depletion, 16–17, 18 

albumin-depleted plasma proteins, 

18–20 

blood sampling, 16 

chromatography of immunoaffinity 

separation of using IgY-12 high 

capacity LC2 column, 46 

1D and 2D gel Electrophoresis, 17 

immunoglobulin depletion, 17, 20 

samples collection of, 17–18 

TCA precipitation of proteins, 17 

Human serum 

chromatogram for affinity removal of 

high-abundant proteins from, 31 

2D gel electrophoresis of, 34 

1D SDS gel electrophoresis of, 33 

high-abundant proteins, 

immunoaffinity depletion of, 

29–30 

immunodepleted serum and bund 

fraction processing, 30 

low-abundant proteins, reversed-phase 

separation of, 30, 35–36 

multi-component immunoaffinity 

subtraction and reversed-phase 

chromatography, 27–39 

proteins composition, 28 

I 

IgY-12 Microbeads regeneration, 45 

ImageQuant software, 11 

Iminodiacetic acid (IDA), 140 

Immobiline Dry Strip (IPG) kit, 205 

Immobiline Dry Strip Reswelling 

Tray , 205 

Immobiline DryStrip Kit, 118 

Immobilized metal ion affinity 

chromatography (IMAC), 139 

Immobilized pH gradient, 70 

Immunoaffinity subtraction (IAS), 94 

Immunoglobulin depletion, 20 

In-gel digestion procedure, 61–62 

Inter simple sequence repeat PCR 

(ISSR-PCR) technique, 156 

IPG strips, 71–72 

IPG-strip rehydration buffer, 22–23 

Isoelectric focusing (IEF) tube 

gels, 9 

K 

Keratin proteome in normal 

NLF, 84 

Kinematica Model PT10-35, 174 

Knexus program, 63 

L 

Labile CWP analysis, 191 

Labile /weakly bound CWP extraction by 

nondestructive techniques 

cell suspension cultures, 194 

liquid culture medium of seedlings, 

193–194 

Laemmli sample buffer, 17 

LC-MS/MS mass spectrometry, 63–64 

Liquid chromatography-coupled 

electrospray ionisation MS 

(LC-ESI-MS), 12 

Liquid Tissue -MS protein prep 

kit, 133 

Liver Infusion Trypticase Medium 

(LIT), 315 

Lowry protein assay, 80 

LTQ ion trap mass spectrometer, 64 

Lumbar puncture (LP), 54 

Lysis buffers, 176

Index 337 

M 

Madin-darby canine kidney (MDCK) 

cells sample preparation from 

culture medium, 113–128 

cell culture, 114–116, 119 

2-D gel electrophoresis of, 115 

protein quantitation by BCA assay, 

117, 121–122 

rehydration loading and isoelectric 

focusing, 117–118, 122–125 

ultracentrifugation and ultrafiltration, 

116–117, 119–120 

MALDI-TOF spectrometry, 58, 63 

MASCOT MS/MS ion search 

software, 84 

Mayer’s hematoxylin stain, 133 

Metal oxide affinity chromatography 

(MOAC), 140 

Metal-specific allergic contact dermatitis 

(ACD), 141 

Mixed bed ion exchanger resin, 118 

Multidimensional liquid chromatography 

(multi-LC), 69 

Multiple Affinity Removal Spin 

Cartridge, 91 

Multiple Affinity Removal System 

column, 30 

Murashige and Skoog (MS) 

medium, 174 

N 

Nanoflow reversed-phase liquid 

chromatography 

(nanoRPLC), 133 

Nasal lavage fluid (NLF) analysis 

data interpretation, 84 

by liquid chromatography and mass 

spectrometry, 79, 81–83 

preparation, 80–81 

acid-ethanol precipitation, 81 

protein digestion, 81 

total protein assay, 80 

total ion chromatogram (TIC) profile 

of, 82 

Nasal secretions 

nasal provocation, 79–80 

pharmacological treatment, 78–79 

preparation for proteome analysis, 

77–78 

Ni-affinity enriched proteins from human 

antigen, 141 

Nickel-nitrilotriacetic acid 

(Ni-NTA), 142 

Ni-NTA Magnetic Agarose Beads, 143 

Nitrilotriacetic acid (NTA), 140 

Nonequilibrium pH gradient gel 

electrophoresis (NEPHGE), 113 

P 

Pancreatic adenocarcinoma cells 

proteome analysis, 4–5 

Pancreatic intraepithelial neoplasia, 1–2 

PanIN, Pancreatic intraepithelial 

neoplasia, 2 

Parasitic protozoa cell fractionation, 

313–314 

cell culture and standard equipment, 

315–316 

electron microscopy, 316 

hydrogenosome isolation and 

subfractionation, 327–329 

isolation and subfractionation 

flagellum of trypanosomatids, 316 

hydrogenosomes, 317 

isolation of 

golgi complex, 316–317, 323–327 

reservosome, 316, 320–321 

trypanosomatid structures, 317–320 

Pepper seed DNA, 161–162 

Peptide mass fingerprinting (PMF), 12 

Percoll gradient solutions, 165 

Phenylmethylsulfonyl fluoride 

(PMSF), 116 

PIR BLAST software, 84 

Plant cell culture isolation of 

mitochondria, 163–169 

crude organelle pellet differential 

centrifugation of, 165–166

338 Index 

density gradient purification of 

mitochondria, 167 

materials, 164–165 

protoplasts disruption and 

isolation, 165 

Plant cell wall poteins isolation, 191–192 

extraction and analysis from cell 

suspension cultures, 190–191 

labile/weakly extraction by 

nondestructive techniques, 

189–190 

Rosette leaves extraction and analysis, 

191–192 

Plant material high-throughput extraction 

device 

DNA electrophoresis, 156, 159–160 

inter simple sequence repeat PCR 

(ISSR-PCR), 156, 159 

isoelectric focusing (IEF), 155–156, 

158–159 

sample preparation, 154–155, 157–158 

Plant mitochondria Percoll gradient 

purification, 166 

Plant proteases, 149 

Plastids organelles, 171 

Plastoglobuli lipid-containing structures, 

173 

POROS ® beads, 17 

Protean IEF Cell, 70 

Protease inhibitor cocktail tablets, 70 

Protein 

concentration estimation kit, 176 

content after, sinusitis NLFs pre- and 

post-pharmacological 

treatment, 82 

deglycosylation, 281–282 

with PNGase F, 282 

trifluoromethanesulfonic acid with, 

282–283 

Proteins 

analysis by fluorescence dye 

saturation labeling and 2-DE, 2 

cysteine-specific protein labeling 

using CyDye DIGE fluor 

saturation dyes, 2 

2D-PAGE, 3 

fluorescence dye labeling 

optimisation of, 6–7 

isoelectric focusing, 2–3 

microdissection and sample 

preparation, 2 

microdissection for proteome 

analysis of pancreatic 

adenocarcinoma cells, 4 

minimal number of microdissected 

cells determination of, 4–6 

protein spots micropreparation 

of, 12 

reference proteome for internal 

standardization and protein 

identification, 7–8 

two-dimensional gel 


extraction from green plant tissue, 

149–150, 152 

materials and methods, 151 

G POROS column, 20 

G Sepharose, 58 

identification and reference proteome 

for internal standardization, 8–9 

loads for silver and Coomassie blue 

staining, 108 

precipitation 

chloroform/methanol/trichloroacetic 

acid with, 280–281 

quantitation by BCA Assay, 117–118 

secretion from yeast protoplasts in 

active cell wall 

regeneration, 241 

alkaline phosphatase assay, 247, 

250–254 

collection of proteins, 245–249 

concentration of proteins secreted 

from regenerating protoplasts, 

247, 250

Index 339 

preparation of yeast 

protoplasts, 249 

recovery of proteins secreted from 

regenerating protoplasts, 

246, 250 

Protoplast formation efficiency of 

evaluation, 259 

R 

Recombinant His-tagged proteins, 140 

Reverse-phase C18 column, 79 

Rosette leaves 

labile CWP analysis by 2-DE, 196 

proteins extraction, 194–196 

vacuum-infiltration, 195 

S 

S. cerevisiae and C. albicans, cell wall, 

218, 221–222 

S. pyogenes strain 5448 cell wall 

proteome, 305 

SEQUEST, 64 

Silver staining procedures, 60 

Size exclusion chromatography 

(SEC), 91 

Spectra/Por ® cellulose ester, 190 

Speed Vac centrifugation, 73 

Streptococcus mutans, silver stained 2-D 

gel, 290 

T 

TCA/Acetone precipitation, 50, 120 

Terminator device, 154–155 

Triacyglycerol (TAG) biosynthesis, 203 

Trichomonad structures isolation, 325 

Trifluoromethanesulfonic acid, chemical 

protein deglycosylation with, 

281–282 

Tritrichomonas foetus, 315 

Trypanosoma cruzi, 314 

Trypsin-mediated incorporation of 

O 18 , 137 

Trypticase Yeast Maltose Medium 

(TYM), 315 

U 

Ultraflex MALDI-TOF spectrometer, 143 

Urinary protein concentrates, size 

exclusion chromatography, 

90, 93 

Urine samples preparation for proteomic 

analysis, 89 

final sample preparation for proteomic 

analysis, 91 

immunoaffinity subtraction of 

proteins, 91 

size exclusion chromatography of 

urinary protein concentrates, 90, 

92–94 

urinary protein concentrate 

preparation, 90, 91 

V 

Vivapure Anti-HSA/IgG removal kit, 91 

Vmax ® microplate reader, 117 

Voyager DE-Pro mass spectrometer, 63 

W 

Weakly bound CWP analysis, 

191–192, 196 

Weakly bound CWP extraction 

destructive techniques, 192, 

196–197 

cell wall preparation, 192, 

196–197 

protein extraction and separation, 

192, 197 

Western blot assembly, 106 

Y 

Yeast and fungal cell walls protein 

solubilization 

-1,3 glucanase treatment, 

224–225, 229 

detergents and reducing agents, 

224, 228

340 Index 

and exochitinase treatment, 225, 

229–230 

under mild alkali conditions, 224, 

228–229 

Yeast and fungal proteomics cell wall 

fractionation, 217–237 

cell wall isolation from yeasts and 

filamentous fungi, 223–228 

protein precipitation, 225, 230 

protein solubilization from isolated 

yeast and fungal cell walls, 

224–225, 228 

Yeast proteome 2-D electrophoresis, 

269–270 

Yeast-Peptone-D-glucose 

(YPD), 222

2D PAGE: Sample Preparation and Fractionation - Mycobacteriology ...

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