Clinical Proteomics: Methods and Protocols [2 ed.] 1493918710, 9781493918713

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Methods in Molecular Biology 1243

Antonia Vlahou Manousos Makridakis Editors

Clinical Proteomics Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

Clinical Proteomics Methods and Protocols Second Edition Edited by

Antonia Vlahou Biomedical Research Foundation, Academy of Athens, Athens, Greece

Manousos Makridakis Biomedical Research Foundation, Academy of Athens, Athens, Greece

Editors Antonia Vlahou Biomedical Research Foundation Academy of Athens Athens, Greece

Manousos Makridakis Biomedical Research Foundation Academy of Athens Athens, Greece

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-4939-1871-3 ISBN 978-1-4939-1872-0 (eBook) DOI 10.1007/978-1-4939-1872-0 Springer New York Heidelberg Dordrecht London Library of Congress Control Number: 2014953785 © Springer Science+Business Media New York 2015 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. Exempted from this legal reservation are brief excerpts in connection with reviews or scholarly analysis or material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Duplication of this publication or parts thereof is permitted only under the provisions of the Copyright Law of the Publisher's location, in its current version, and permission for use must always be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright Clearance Center. Violations are liable to prosecution under the respective Copyright Law. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. While the advice and information in this book are believed to be true and accurate at the date of publication, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is a brand of Springer Springer is part of Springer Science+Business Media (www.springer.com)

Preface Clinical proteomics is a rapidly evolving field, characterized by continuous technological and methodological advancements. Since the publication of the Clinical Proteomics Methods and Protocols (volume I, Humana Press, 2008), significant methodological developments have resulted in increased sensitivity in protein identification, including analysis of protein posttranslational modifications as well as more reliable protein quantification. Multiple different types of biological samples can now be analyzed comprehensively for their protein context. In addition, tools for protein functional annotation, assignment to regulatory pathways and correlation to other—omics datasets are being continuously developed and optimized. Even more, significant progress towards proteomic biomarker implementation has been achieved with mass spectrometry-based approaches now being used for biomarker validation. This new volume of Clinical Proteomics captures these latest developments in the field and brings together a multidisciplinary team of leading researchers to provide step by step their latest protocols for clinical proteomics analysis. It is expected that the presented approaches may facilitate activities and serve as a reliable guide to researchers, including clinicians, chemists, molecular biologists, bioinformaticians, and computational biologists and, in general, investigators working on biomarker development. Athens, Greece

Antonia Vlahou Manousos Makridakis

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Acknowledgments The editors gratefully acknowledge all contributing authors for their collaboration which made this project possible and brought it into fruition; the series editor Prof. John Walker whose help and guidance have been instrumental; Mr. Patrick Marton, Ms. Monica Suchy, Mr. David Casey, Mr. Jeffin Thomas Varghese and the whole production team at SPi Global for making the production of this book possible.

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

PART I

SAMPLE COLLECTION

1 Biological Sample Collection for Clinical Proteomics: Existing SOPs . . . . . . . . Vasiliki Lygirou, Manousos Makridakis, and Antonia Vlahou 2 Targeting the Proteome of Cellular Fractions: Focus on Secreted Proteins. . . . Agnieszka Latosinska, Maria Frantzi, William Mullen, Antonia Vlahou, and Manousos Makridakis 3 Preparation of Urinary Exosomes: Methodological Issues for Clinical Proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marina Pitto, Samuele Corbetta, and Francesca Raimondo 4 Sample Treatment Methods Involving Combinatorial Peptide Ligand Libraries for Improved Proteomes Analyses . . . . . . . . . . . . . . . Pier Giorgio Righetti and Egisto Boschetti 5 Glycoprotein Enrichment Method Using a Selective Magnetic Nano-Probe Platform (MNP) Functionalized with Lectins. . . . . . . . Marta Cova, Rui Oliveira-Silva, José Alexandre Ferreira, Rita Ferreira, Francisco Amado, Ana Luísa Daniel-da-Silva, and Rui Vitorino

PART II

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TECHNOLOGIES FOR DISCOVERY AND VALIDATION

6 The Latest Advancements in Proteomic Two-dimensional Gel Electrophoresis Analysis Applied to Biological Samples . . . . . . . . . . . . . . . Laura Santucci, Maurizio Bruschi, Gian Marco Ghiggeri, and Giovanni Candiano 7 2DE Maps in the Discovery of Human Autoimmune Kidney Diseases: The Case of Membranous Glomerulonephritis . . . . . . . . . . . . . . . . . . . . . . . . Maurizio Bruschi, Laura Santucci, Gian Marco Ghiggeri, and Giovanni Candiano 8 MALDI-Imaging Mass Spectrometry on Tissues . . . . . . . . . . . . . . . . . . . . . . . Veronica Mainini, Maciej Lalowski, Athanasios Gotsopoulos, Vasiliki Bitsika, Marc Baumann, and Fulvio Magni

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9 Laser Capture Microdissection of Fluorescently Labeled Amyloid Plaques from Alzheimer’s Disease Brain Tissue for Mass Spectrometric Analysis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Diana A.T. Nijholt, Christoph Stingl, and Theo M. Luider 10 Urine Sample Preparation and Fractionation for Global Proteome Profiling by LC-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . Magali Court, Jérôme Garin, and Christophe D. Masselon 11 Methods in Capillary Electrophoresis Coupled to Mass Spectrometry for the Identification of Clinical Proteomic/Peptidomic Biomarkers in Biofluids. . . . . . . . . . . . . . . . . . . . . . . . Angelique Stalmach, Holger Husi, Khedidja Mosbahi, Amaya Albalat, William Mullen, and Harald Mischak 12 Quantification of Proteins in Urine Samples Using Targeted Mass Spectrometry Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nina Khristenko and Bruno Domon

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PART III EXTRACTING BIOLOGICAL RELEVANT BIOMARKERS AND PROCEEDING WITH IMPLEMENTATION 13 Statistical Issues in the Design and Planning of Proteomic Profiling Experiments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . David A. Cairns 14 Integrating Proteomics Profiling Data Sets: A Network Perspective . . . . . . . . . Akshay Bhat, Mohammed Dakna, and Harald Mischak 15 The European Medicines Agency Experience with Biomarker Qualification . . . Efthymios Manolis, Armin Koch, Dieter Deforce, and Spiros Vamvakas Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors AMAYA ALBALAT • BHF Glasgow Cardiovascular Research Centre, Institute of Cardiovascular and Medical Sciences, College of Medical Veterinary and Life Sciences, University of Glasgow, Glasgow, UK FRANCISCO AMADO • QOPNA, School of Health Sciences, University of Aveiro, Aveiro, Portugal MARC BAUMANN • Meilahti Clinical Proteomics Core Unit, Department of Biochemistry and Developmental Biology, Institute of Biomedicine, Biomedicum Helsinki, University of Helsinki, Helsinki, Finland AKSHAY BHAT • Mosaiques diagnostics & therapeutics, Hannover, Germany VASILIKI BITSIKA • Biotechnology Division, Biomedical Research Foundation, Academy of Athens, Athens, Greece EGISTO BOSCHETTI • Independent Consultant, Paris, France MAURIZIO BRUSCHI • Laboratory on Pathophysiology of Uremia, Istituto Giannina Gaslini, Genoa, Italy DAVID A. CAIRNS • Section of Oncology and Clinical Research, Leeds Institute of Cancer and Pathology, St. James’s University Hospital, Leeds, UK GIOVANNI CANDIANO • Laboratory on Pathophysiology of Uremia, Istituto Giannina Gaslini, Genoa, Italy SAMUELE CORBETTA • Department of Health Sciences, University of Milano-Bicocca, Monza, Italy MAGALI COURT • Université Grenoble-Alpes, Grenoble, France; CEA, iRTSV, Biologie à Grande Echelle, Grenoble, France; INSERM, U1038, Grenoble, France MARTA COVA • Department of Chemistry, Mass Spectrometry Centre, QOPNA, University of Aveiro, Aveiro, Portugal MOHAMMED DAKNA • Mosaiques diagnostics & therapeutics, Hannover, Germany ANA LUÍSA DANIEL-DA-SILVA • Department of Chemistry, CICECO, Aveiro Institute of Nanotechnology, University of Aveiro, Aveiro, Portugal DIETER DEFORCE • European Medicines Agency, London, UK; Ghent University, Laboratory of Pharmaceutical Biotechnology, Ghent, Belgium BRUNO DOMON • Luxembourg Clinical Proteomics Center (LCP), CRP‐Santé, Strassen, Luxembourg JOSÉ ALEXANDRE FERREIRA • Department of Chemistry, Mass Spectrometry Centre, QOPNA, University of Aveiro, Aveiro, Portugal RITA FERREIRA • Department of Chemistry, Mass Spectrometry Centre, QOPNA, University of Aveiro, Aveiro, Portugal MARIA FRANTZI • Mosaiques diagnostics & therapeutics, Hannover, Germany JÉRÔME GARIN • Université Grenoble-Alpes, Grenoble, France; CEA, iRTSV, Biologie à Grande Echelle, Grenoble, France; INSERM, U1038, Grenoble, France GIAN MARCO GHIGGERI • Division of Nephrology, Dialysis, and Transplantation, Istituto Giannina Gaslini, Genoa, Italy

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ATHANASIOS GOTSOPOULOS • Brain and Mind Laboratory, Department of Biomedical Engineering and Computational Science (BECS), Aalto University School of Science, Helsinki, Finland HOLGER HUSI • BHF Glasgow Cardiovascular Research Centre, Institute of Cardiovascular and Medical Sciences, College of Medical Veterinary and Life Sciences, University of Glasgow, Glasgow, UK NINA KHRISTENKO • Luxembourg Clinical Proteomics Center (LCP), CRP‐Santé, Strassen, Luxembourg ARMIN KOCH • European Medicines Agency, London, UK; Medizinische Hochschule Hannover, Institut für Biometrie, Hannover, Germany MACIEJ LALOWSKI • Meilahti Clinical Proteomics Core Unit, Department of Biochemistry and Developmental Biology, Institute of Biomedicine, Biomedicum Helsinki, University of Helsinki, Helsinki, Finland; Folkhälsan Institute of Genetics, Helsinki, Finland AGNIESZKA LATOSINSKA • Biotechnology Division, Biomedical Research Foundation, Academy of Athens, Athens, Greece THEO M. LUIDER • Laboratory Neuro Oncology/Clinical & Cancer Proteomics, Department of Neurology, Erasmus MC, Rotterdam, The Netherlands VASILIKI LYGIROU • Biotechnology Division, Biomedical Research Foundation, Academy of Athens, Athens, Greece FULVIO MAGNI • Department of Experimental Medicine, University Milano-Bicocca, Monza, Italy VERONICA MAININI • Department of Experimental Medicine, University Milano-Bicocca, Monza, Italy MANOUSOS MAKRIDAKIS • Biotechnology Division, Biomedical Research Foundation, Academy of Athens, Athens, Greece EFTHYMIOS MANOLIS • European Medicines Agency, London, UK CHRISTOPHE D. MASSELON • Université Grenoble-Alpes, Grenoble, France; CEA, iRTSV, Biologie à Grande Echelle, Grenoble, France; INSERM, U1038, Grenoble, France HARALD MISCHAK • Mosaiques diagnostics & therapeutics, Hannover, Germany KHEDIDJA MOSBAHI • BHF Glasgow Cardiovascular Research Centre, Institute of Cardiovascular and Medical Sciences, College of Medical Veterinary and Life Sciences, University of Glasgow, Glasgow, UK WILLIAM MULLEN • BHF Glasgow Cardiovascular Research Centre, Institute of Cardiovascular and Medical Sciences, College of Medical Veterinary and Life Sciences, University of Glasgow, Glasgow, UK DIANA A.T. NIJHOLT • Laboratory Neuro Oncology/Clinical & Cancer Proteomics, Department of Neurology, Erasmus MC, Rotterdam, The Netherlands RUI OLIVEIRA-SILVA • Department of Chemistry, Mass Spectrometry Centre, QOPNA, University of Aveiro, Aveiro, Portugal MARINA PITTO • Department of Health Sciences, University of Milano-Bicocca, Monza, Italy FRANCESCA RAIMONDO • Department of Health Sciences, University of Milano-Bicocca, Monza, Italy PIER GIORGIO RIGHETTI • Department of Chemistry, Materials and Chemical Engineering “Giulio Natta”, Politecnico di Milano, Milano, Italy

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LAURA SANTUCCI • Laboratory on Pathophysiology of Uremia, Istituto Giannina Gaslini, Genoa, Italy ANGELIQUE STALMACH • BHF Glasgow Cardiovascular Research Centre, Institute of Cardiovascular and Medical Sciences, College of Medical Veterinary and Life Sciences, University of Glasgow, Glasgow, UK CHRISTOPH STINGL • Laboratory Neuro Oncology/Clinical & Cancer Proteomics, Department of Neurology, Erasmus MC, Rotterdam, The Netherlands SPIROS VAMVAKAS • European Medicines Agency, London, UK RUI VITORINO • Department of Chemistry, Mass Spectrometry Centre, QOPNA, University of Aveiro, Aveiro, Portugal ANTONIA VLAHOU • Biotechnology Division, Biomedical Research Foundation, Academy of Athens, Athens, Greece

Part I Sample Collection

Chapter 1 Biological Sample Collection for Clinical Proteomics: Existing SOPs Vasiliki Lygirou, Manousos Makridakis, and Antonia Vlahou Abstract Proteomic study of clinical samples aims at the better understanding of physiological and pathological conditions, as well as the discovery of diagnostic and prognostic markers for the latter. Quantitative and/ or qualitative variations of body fluid proteome may reflect health- or disease-associated events connected to the adjacent or distant body regions of the fluid production/secretion/excretion and/or systemic reactions to the presence of disease. Sample collection and preparation is a critical step in order to obtain useful and valid information in clinical proteomics analysis. In this chapter, we present the current protocols and guidelines for human body fluid collection and storage, prior to proteomic analysis. A variety of body fluids that are currently being used in proteomic analysis and have potential interest in clinical practice are presented including blood plasma and serum, urine, cerebrospinal fluid, cerumen, nasal secretions, saliva, tears, breast milk, bronchoalveolar fluid, nipple aspirate fluid, amniotic fluid, bile, cervico-vaginal fluid, and seminal plasma. With no doubt these body fluids differ in the extent of their application in clinical proteomics investigations, hence in some cases the presented SOPs are established following more extensive testing (e.g., plasma, serum, urine, CSF) than others (nasal secretions, saliva, tears, breast milk, bronchoalveolar fluid, nipple aspirate fluid, amniotic fluid, bile, cervico-vaginal fluid, and seminal plasma). However, even in these latter cases, the presented protocols were reported by at least two independent groups according to the literature. We hope they can thus serve as a reliable guide for sample collection based on our current knowledge in the field and excellent starting points for proteomics investigators. It should also be pointed that variations to these protocols exist and their further refinement in the future is foreseen following the evolution of the proteomics technologies. Key words Biological sample collection, Body fluids, Protocols, Clinical proteomics, Biomarkers

1 1.1

Commonly used body fluids Blood Plasma

1.1.1 Introduction

Blood plasma is the straw-colored liquid component of blood in which the blood cells are suspended. It amounts to about 55 % of the total blood volume and is composed of water (92 %) and proteins of normal total protein concentration of 6.0–8.3 g/dL. The main functions of plasma include maintenance of the blood

Antonia Vlahou and Manousos Makridakis (eds.), Clinical Proteomics: Methods and Protocols, Methods in Molecular Biology, vol. 1243, DOI 10.1007/978-1-4939-1872-0_1, © Springer Science+Business Media New York 2015

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pressure and pH, immunity, and transportation of electrolytes, glucose, clotting factors, hormones and other excretory products [1]. Plasma is easily obtained from blood after centrifugation with anticoagulants. Blood collection is a minimally invasive and routine procedure, thus blood specimens are of the most readily available samples from patients and participants in clinical research [2]. However, the existence of highly abundant proteins in plasma hinders an efficient proteomic analysis. Specifically, albumin accounts for approximately 50 % of the total protein mass in plasma while the most abundant proteins (including albumin, IgG, IgA, IgM, α-2 and β-lipoproteins, fibrinogen, α1-antitrypsin, α2macroglobulin, transferrin, α-1-Acid glycoprotein, hemopexin 1, α-lipoproteins, haptoglobin, prealbumin, and ceruloplasmin) are estimated to account for 99 % of total protein mass. As a result, depletion of these highly abundant proteins is often necessary prior to proteomic analysis [2, 3]. However, it must be taken into consideration that depletion methods might be responsible for the partial loss of low abundant proteins (such as cytokines) due to carrier protein interactions. Disrupting the bonds between high (carrier proteins) and low abundant proteins can be a solution to this problem [4]. Several methods have been developed for depletion of highly abundant proteins including affinity chromatography using dye ligands, protein A or immunoaffinity columns specific for abundant proteins; antibody-based microarrays (or other affinity molecules such as aptamers); or protein enrichment through the use of hexapeptide ligand library (5, reviewed in 6). The Human Proteome Organization (HUPO) initiated the HUPO Plasma Proteome Project (HPPP) in 2002 and during 2003–2005 the 55 participating laboratories prepared and distributed reference specimens of human serum and EDTA-, citrate-, and heparin-plasma for proteomics analysis. The Pilot Phase of the HPPP resulted in the generation of the human plasma proteome database (currently including 10,546 proteins). The conclusion of the comparative study was that EDTA-plasma is the preferred specimen for proteomic analysis. The HPPP project evaluated all the aspects concerning plasma and serum proteome analysis, from specimen stability to analysis methods and search engines performance [7–9]. 1.1.2 Sample Collection Protocol

The protocol described below is based on the HPPP and National Institutes of Health (http://edrn.nci.nih.gov/resources/standardoperating-procedures/standard-operating-procedures) guidelines for sample collection and handling [10–12]. 1. Arterial or venous blood is obtained from an existing arterial or venous line or via venipuncture (with a 19-gauge needle) by a trained phlebotomist under the direction of a qualified and licensed physician (see Note 1).

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2. The blood is collected in blood collection tubes with anticoagulant (Potassium EDTA) (see Note 2). 3. Within 1–2 h, the tubes are inverted (ten times) and centrifuged at ≤1,300 × g for 10 min at 2–6 °C, in a horizontal, swinging bucket centrifuge (see Notes 2 and 3). 4. After centrifugation, the supernatant is transferred immediately into new tubes (cryovials) and is stored at −80 °C or in liquid nitrogen until use. Protease inhibitor cocktail can be added before storage if desired (see Note 2). 1.1.3

Notes

1. If the plasma is icteric or exhibits signs of hemolysis, this should be recorded and, depending on the disease under investigation, may not be included in the final analysis [10, 11, 13]. 2. Fasting before the blood withdrawal, the type and concentration of anticoagulant (Potassium EDTA, Lithium Heparin or Sodium Citrate) and treatment with protease inhibitor cocktail, are some of the variables that should be kept consistent among different subjects and samples when data comparability is desired [10, 11, 13]. 3. In the case that Lithium Heparin is used as anticoagulant, the tube should be inverted ten times and then centrifuged at ≤1,300 × g for 10 min at 2–6 °C. If Sodium Citrate is used, the tube should be inverted four times and then centrifuged at 1,500 × g for 15 min at 2–6 °C.

1.2

Blood Serum

1.2.1 Introduction

Blood serum is a clear, yellow-colored liquid which is obtained by allowing the blood to clot prior to centrifugation (without anticoagulants). Coagulation makes serum qualitatively different from plasma, mainly because of the removal of a large portion of clotting factors, and results in a lower (3–4 %) protein concentration compared to plasma. However, clotting factors are not totally absent in serum and protein and peptide levels may be affected by the coagulation process and the specific or nonspecific interaction with the fibrin clots. Similar to plasma, serum is an easily obtained specimen requiring, nevertheless, application of depletion steps for the removal of high abundance proteins [11, 14, 15]. The use of blood serum as a specimen for proteomic analysis is almost as common as blood plasma and, in many studies, both specimens are examined and the results are combined or compared. Similar to plasma, the Pubmed search (October 2013) results for “blood serum” and “proteomic” are a few thousand. The protein content in serum is slightly reduced due to the lack of most of the clotting factors, allowing more low-abundant proteins to be detected compared to plasma. In some cases, serum is the preferred specimen because of the elimination of

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interference that may be caused by the anticoagulants in plasma. However, it must be taken into consideration that the peptides released during the coagulation process may result in high levels of artificial peptide signals [11]. Moreover, during coagulation, platelets, erythrocytes and leucocytes may secrete certain proteins, thus increasing ex vivo the apparent concentration of proteins in serum [14]. 1.2.2 Sample Collection Protocol

The protocol described below is based on the HPPP and National Institutes of Health (http://edrn.nci.nih.gov/resources/standardoperating-procedures/standard-operating-procedures) guidelines for sample collection and handling [10–12]. 1. Arterial or venous blood is obtained from an existing arterial or venous line or via venipuncture (with a 19-gauge needle) by a trained phlebotomist under the direction of a qualified and licensed physician. 2. The blood is collected in blood collection tubes with silica clot activator (see Note 4). The samples are allowed to rest and clot for 30 min and then centrifuged at 2–6 °C, in a horizontal, swinging bucket centrifuge, at ≤1,300 × g for 10 min. 3. After centrifugation, the supernatant is transferred immediately into new tubes (cryovials) and is stored at −80 °C until use. Protease inhibitor cocktail can be added before storage if desired (see Note 5).

1.2.3

Notes

4. The presence of a clot activator in the blood collection tube promotes rapid coagulation and is optional [10, 11]. 5. Fasting before the blood withdrawal and treatment with protease inhibitor cocktail may have an impact on the serum composition and should be kept consistent among different subjects and samples for the results of their analysis to be comparable [10, 11].

1.3

Urine

1.3.1 Introduction

Urine is the clear, amber-colored liquid product of the body that is secreted by the kidneys in the urinary bladder and then excreted through the urethra. Kidneys filter the blood plasma and dispose of the water-soluble waste via urine. Healthy human urine consists primarily of water (95 %) with solutes including urea, glucose, salts, proteins, and other compounds. The protein concentration of urine is approximately 1,000-fold lower than that of blood plasma and the relative concentration (in reference to an internal standard) of a protein or a peptide may be indicative of a condition [16, 17]. There are many reasons why urine is a favorable specimen for proteomic analysis. Urine collection is a noninvasive procedure that does not require trained personnel or special equipment and

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can be repeated as frequently as needed, without causing any discomfort to the subjects. Moreover, urine is a relatively stable specimen. As it remains in the urinary bladder for hours, almost all endogenous proteolytic activity is completed before voiding. Thus, it is not surprising that the Pubmed search (October 2013) results for “urine” and “proteomic” are exceeding 1,000. On the other hand, high variability of protein content exists depending on diet, time of collection, presence of kidney dysfunction and other diseases and these factors should be taken under consideration [17]. The Human Kidney and Urine Proteome Project (HKUPP) was initiated in 2005 and the European Kidney and Urine Proteomics (EuroKUP) COST Action in 2008, to promote proteomics research in the field of renal disease and to define novel biomarkers and therapeutic targets. Among others, HKUPP and EuroKUP together have achieved the establishment of a standard protocol for urine collection and storage [18–21]. 1.3.2 Sample Collection Protocol

The protocol described below is based on the standard protocol for urine collection and storage for proteomics analysis established by HKUPP and EuroKUP [19, 21]. 1. Mid stream of the second morning urine (7–11 a.m.) is collected by the subject in a urine container. A minimum volume of 2 mL is regularly needed (see Notes 6–8). 2. No centrifugation is generally needed (see Note 9). Furthermore, no storage on ice or additives are required in case aliquoting and freezing within 3 h from collection is possible. Urine is aliquoted in working volumes (for most proteomics methods 1–2 mL aliquots are appropriate). Samples can be stored frozen at −20 °C for years, however if −80 °C long-term storage is available it should be preferred (see Note 10).

1.3.3

Notes

6. 24 h urine can be used as well, as long as it is collected in the presence of NaN3 (sodium azide) (10 mL of a 5 % solution) as preservative in order to avoid any microbial contamination. NaN3 may also be required in case prompt freezing following collection is not possible [18, 19]. 7. First morning urine or random catch is not preferred as they tend to show increased variabilities [18, 19]. 8. Containers with coating must be avoided as they may interfere in the analysis. A urine monovette can also be used for the collection of urine [18, 19]. 9. If urine is turbid and characterized by hematuria or if cellular contents are visible, the sample may be centrifuged at ≤1,000 × g

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for 10 min. After centrifugation the supernatant is transferred to a new container and then stored at ≤−20 °C [18, 19]. 10. Repeated freeze–thaw cycles should be avoided as possible [18, 19]. 1.4 Cerebrospinal Fluid 1.4.1 Introduction

1.4.2 Sample Collection Protocol

Cerebrospinal fluid (CSF) is a clear, colorless body fluid surrounding the brain and the spinal cord of vertebrates. It provides mechanical support to the brain, circulates nutrients and chemicals, and removes waste products. CSF is produced in the choroid plexus and is reabsorbed into venous sinus blood via arachnoid granulations. The total volume of CSF is approximately 160 mL in adults and is replaced four to five times per day [22]. Its composition resembles that of blood plasma. Water, gases, and lipid-soluble compounds move freely from the blood into CSF while glucose, amino acids and cations are transported by carrier-mediated processes. The protein concentration in CSF is approximately 100-fold lower than that of plasma [22, 23]. CSF can be considered as the most suitable specimen for biomarker discovery and drug development in neurological disorders, as its composition reflects the biochemical state of the central nervous system. The CSF proteome is similar to that of plasma with protein concentrations covering a large dynamic range. Of note, it also, consists of 20 % brain-derived proteins not regularly detected in the peripheral blood [22, 23]. Currently, the Pubmed search (October 2013) results for “cerebrospinal fluid” and “proteomic” are a few hundred. Challenges for clinical proteomics applications among others include adoption of the standardized sample collection and storage protocol (described below), optimization of the study design from the clinical perspective (e.g., optimal selection of cases and controls), consideration of the blood contribution to the sample and optimization of the analysis methods to detect the low abundant proteins [22, 24]. The protocol described below is based on a recently developed consensus SOP for cerebrospinal fluid collection and biobanking [25]. 1. The subject lies on his/her side, with knees pulled up toward the chest and chin tucked downward. After the back is cleaned, the trained physician injects a local numbing medicine (anesthetic) into the lower spine. 2. An atraumatic spinal needle is inserted into the vertebral body L3–L5 and CSF is collected in polypropylene tubes (screw cap, volume 1–2 mL) without any additives. A volume of at least 12 mL of CSF is recommended to be collected if possible (first 1–2 mL for basic CSF assessment) (see Notes 11–15).

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3. The CSF collected from each subject is pooled and then centrifuged at room temperature, at 2,000 × g for 10 min. After centrifugation, the supernatant is divided into aliquots in polypropylene tubes and stored at −80 °C within 1–2 h (see Note 16). 1.4.3

Notes

11. Lumpar puncture, described above, is the most common and recommended method for CSF collection. If CSF is collected through an external ventricular drain, the protein concentration may vary [25]. 12. Matched samples of blood plasma and/or serum may be collected simultaneously (see Subheadings 1.1 and 1.2) for comparison and evaluation of protein concentration in CSF [25]. 13. The same volume of CSF (12 mL minimum) must be collected for all subjects under comparison, as the concentration of proteins exhibits a rostrocaudal gradient and as a consequence, different volumes of CSF will have different composition and be representative of different spine or brain areas [25]. 14. If the CSF sample is bloody, it should not be used for proteomic analysis as the blood proteins may affect the results [25]. 15. The subject may suffer from postlumbar puncture headache after the procedure [25]. 16. Repeated freeze–thaw cycles should be avoided [25].

2 2.1

Head and neck fluids Nasal Secretions

2.1.1 Introduction

Nasal secretions (mucus) are a gelatinous fluid produced by mucous membranes in the nasal passages. Their main purpose is to protect the respiratory system by blocking potentially pathogenic antigens, but also to humidify, heat or cool and clean the inhaled air. Nasal secretions contain proteins of the innate immune system derived from plasma, glandular mucous, and serous cells. Proteomic study of these secretions is possible by analyzing nasal lavage fluid (NLF) which is obtained through nasal provocation [26]. Proteomic studies of NLF can provide valuable information toward discovering diagnostic biomarkers for special conditions (presence of disease or chemical exposure) of the upper airways. NLF is easily collected but the Pubmed search (October 2013) results for “nasal secretions” and “proteomic” are only a couple of dozens. Further analysis, including determination of a normal nasal mucus proteomic profile may increase understanding of the role of nasal secretions to immune response [27].

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2.1.2 Sample Collection Protocol

The protocol described below is based on the detailed instructions in Methods in Molecular Biology [28] and has been used further in follow-up studies [26, 27, 29]. 1. Subjects’ nasal cavities are prewashed with 24 sprays (100 μL each nostril) of sterile Normal Saline (1 × NS, 0.9 % NaCl) using a Beconase AQ pump aspirator spray device (see Note 17). 2. Subjects then gently blow out through their noses, and the lavage fluid from both nostrils is collected into a beaker. This material is discarded. 3. After 10 min, the two first steps are repeated, but with 12 sprays each time. This material is discarded (see Note 18). (This is the phase of the nasal provocation.) 4. 100 μL of sterile normal saline is administered separately into each nostril. 5. After 5 min, NLF is collected using 12 sprays of 0.9 % sterile normal saline. (Subjects must close their left nostrils and then use 12 sprays of sterile normal saline into their right nostril and vice versa.) 6. Subjects gently blow the NFL into a cup mixing the fluid from both nostrils. Any big globules of mucus must be dissolved (First series). 7. Immediately, subjects use one spray of freshly prepared (with double-distilled deionized water) hypertonic saline (24 × NS, 21.6 % NaCl) pH 6.07 (see Note 19). 8. After 5 min, subjects use 12 sprays of sterile normal saline in each nostril. 9. Subjects gently blow the NFL into a cup mixing the fluid from both nostrils. Any big globules of mucus must be dissolved (Second series). 10. NLF is gently shaken to disperse mucous globules and pipetted into eppendorf tubes. Samples are stored at −20 °C until use [26, 30].

2.1.3

Notes

17. Pregnancy or lactation, history of smoking, immunodeficiency, cystic fibrosis, airway bacterial or viral infection, antibiotic therapy in the past week, allergy immunotherapy in the past year, intranasal medications, nonprescribed medications, active rhinitis of any type, history of allergic rhinitis or nasal polyps, sinusitis, use of antihistamines or glucocorticoids in the previous 4 weeks and any chronic disease that could interfere with airway mucosal and neuronal response should be recorded and, depending on the study, be avoided as they largely affect the NSL proteomic profile [26, 30].

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11

18. Proteins identified in the first series of NLF specimens are of plasma origin while those of the second series are secreted from glands [26, 30]. 19. The presence of a physician is recommended through provocation because hypertonic saline stimulates the glandular secretions including local mucosal substance P release and may cause pain [26, 30]. 2.2

Saliva

2.2.1 Introduction

2.2.2 Sample Collection Protocol

Saliva is a watery fluid that bathes oral mucosa and covers every structure of the oral cavity. It is produced from secretions of the three paired major salivary glands (parotid, submandibular, and sublingual), numerous minor salivary glands (labial mucous gland, palatine, von Ebner’s glands, etc.), and gingival crevicular fluid. It consists mostly of water (99.5 %) but also salt, proteins, peptides, hormones, lipids, sugar as well as epithelial cells, food debris and microorganisms. Saliva’s main purpose is the protection and maintenance of mucosa integrity in the upper alimentary tract and is involved in mastication, mineralization of teeth, control of microorganisms, taste perception, and digestion [31, 32]. Saliva is a body fluid that can be easily obtained and at large quantities through a noninvasive procedure. Study of the saliva proteome will contribute to the understanding of the oral environment physiology and, thus, will allow the identification of salivary biomarkers for local or systemic diseases. The Pubmed search (October 2013) results for “saliva” and “proteomic” are a few hundred. To date, many new potential diagnostic salivary biomarkers of oral and systemic diseases, such as gingivitis [33], chronic periodontitis [34], dental caries [35–37], Sjögren’s syndrome [38], type 2 diabetes [39, 40], type 1 diabetes [33], head and neck squamous cell carcinoma [41, 42], lung cancer [43], sclerosis and psychiatric and neurological diseases [29], have been proposed based on proteomic approaches. The protocol described below is based on the World Health Organization and International Agency for Research on Cancer guidelines for saliva proteomics [44] (see Note 20). 1. Collection steps for: Whole saliva 1. The subject drinks water (bottled) and rinses their mouth out well (without drinking the water). 2. 5 min after this oral rinse, the subject spits whole saliva into a 50-ml sterile Falcon tube, about once a minute for up to 10 min. The collection tube must be on ice during the procedure and the subject should not cough up mucus and must refrain

12

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from talking. It is recommended to drop down the head and let the saliva run naturally to the front of the mouth, hold for a while and spit into the tube. 3. The target volume for each whole saliva donation is about 5 mL. Submandibular saliva 1. 2 × 2 in. cotton gauzes are used to block the opening of each parotid duct and the openings of the sublingual gland (both sides) and the floor of the mouth is left to dry up. 2. Subject raises their tongue slightly to elevate the opening to the submandibular gland and saliva is collected through a sterilized disposable yellow tip (for pipette P200) attached to a sterilized Wolf device. 3. Collection takes place at 2 min intervals. During this time, a few grains of citric acid powder are swabbed with a moistened cotton applicator onto the lateral dorsum of the tongue to stimulate the secretion. 4. The target volume for each submandibular saliva collection is about 200 μL. Sublingual saliva 1. 2 × 2 in. cotton gauzes are used to block the opening of each parotid duct and the ductal orifices of the submandibular gland and the floor of the mouth is left to dry up. 2. Subject raises their tongue slightly to elevate the opening to the sublingual gland and saliva is collected through a sterilized disposable yellow tip (for pipette P200) attached to a sterilized Wolf device. 3. Collection takes place at 2 min intervals. During this time, a few grains of citric acid powder are swabbed with a moistened cotton applicator inferior to the tongue to stimulate the secretion. 4. The target volume for each sublingual saliva collection is about 100 μL. Parotid saliva 1. Parotid cups are used to collect parotid saliva and they may be placed bilaterally for simultaneous collection from each parotid gland. 2. During the collection, at 2 min intervals, a few grains of citric acid powder are swabbed with a moistened cotton applicator on the inner surface of the mandibular ramus to stimulate the secretion.

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3. The first 0.1 mL of parotid saliva collected are discarded to ensure that fresh parotid saliva is obtained. 4. The target volume for each parotid saliva collection is 1 mL. 2. Following collection, 0.2 μL protease inhibitor cocktail (standard stock solution) and 0.2 μL 400 mM Na3VO4 per 100 μL of saliva are added and the samples are gently inverted (see Note 21). 3. Specimens are centrifuged at 4 °C at 2,600 × g for 15 min. If incomplete separation is noted, the centrifugation time should be increased at 20 min (see Note 22). 4. The supernatant is carefully transferred to a new tube (in order not to disturb the pellet). 5. Samples are stored at −80 °C. 2.2.3

Notes

20. Subjects should have no evidence of oral pathologies or inflammatory diseases [45]. 21. Collected samples must be kept on ice all the time prior to processing [45]. 22. Centrifugation, as a clearance step, should be critically considered in saliva analysis since the hydrophobic peptides that tend to aggregate with high molecular components may be lost [45].

2.3

Tears

2.3.1 Introduction

2.3.2 Sample Collection Protocol

Tears are clear, colorless secretions that overlay the epithelium of the eye’s surface. They play an important role in the optical system as they lubricate the eye, provide nutrients and growth factors to the epithelium and serve as a barrier to the outside environment. Tears consist of water, mucins, lipids, proteins, electrolytes, and various other metabolites. Their protein content is about 6–10 mg/mL and the major proteins include lysozyme, lactoferrin, secretory immunoglobulin A, lipocalin, albumin, and lipophilin [46, 47]. Tears are collected through a noninvasive procedure and the quantitative and qualitative composition of their proteome reflects the physiology of the underling tissues, providing useful information about the health of the ocular environment and systemic conditions. The Pubmed search (October 2013) results for “tear” and “proteomic” are approaching a hundred. Tears have been used as specimen for discovery of biomarkers indicative of diseases, such as dry eye conditions [48], blepharitis [49], Sjögren’s syndrome [50], conjunctivochalasis [49] and diabetic corneal epithelium [51], and to study patients under anti-glaucoma medications [52]. The protocols described below are the most commonly used in existing publications on tears proteomics investigations [47].

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Collection using capillaries 1. Subject is seated comfortably, with their head raised and against some direct source of light or flow of air. 2. Subject’s reflex tears are collected with sterile 10 μL capillary tubes without touching the eye globe or the lid. 3. It is recommended that 60–80 μL of reflex tears are collected from healthy subjects and a minimum volume of 10 μL be obtained from every subject if possible. 4. The samples are centrifuged at 7,800 × g for 10 min at 4 °C to remove cellular debris and stored in liquid nitrogen or at −80 °C (see Note 23). Collection using strips (Schirmer’s tear test) 1. Prior to the tear collection, a local anesthetic may be applied to ensure collection of only basal and not reflex tears. 2. A Schirmer strip is placed in the lower cul-de-sac region of the subject’s eye and allowed to absorb the tear for 5 min in an open eye condition from the inferior prism without any contact with the lower lid, the cornea, or conjunctiva. 3. After the collection, Schirmer strips are stored in sterile vials at −80 °C (see Notes 23 and 24). 2.3.3

Notes

23. Tear collection using capillary tubes yields greater amount of specimen but in cases that the subjects suffer from a dry eye condition the Schirmer’s tear test is preferred, which also allows to evaluate dryness [47]. 24. The tear protein is extracted from the Schirmer strips using an elution buffer (e.g., 8 M urea buffer containing 3 % CHAPS and 25 mM DTT, pH 7.4) [47].

3 3.1

Thorax fluids Breast Milk

3.1.1 Introduction

Breast milk is the most important food for neonates as its consumption provides nutrients, immune components, anti-infective factors and metabolic enzymes, which all contribute to growth and development during early life. Breast milk is a complex mixture of proteins, lipids, lactose, oligosaccharides, and several bioactive factors. Milk proteins can be grouped into three main fractions: caseins in colloidal dispersion (such as micelles), soluble proteins (whey proteins), and proteins associated with the milk fat globule membranes (MFGMP) [53, 54]. Breast milk is an easily obtained body fluid, and the Pubmed search (October 2013) results for “breast milk” and “proteomic” are nearly a hundred. According to the American Academy of

Body Fluids-SOPs

15

Pediatrics, human milk is the preferred source of nutrients for all infants, including premature and sick newborns, with rare exceptions, as it contains proteins that have significant short- and longterm beneficial effects on the infant. A comprehensive understanding of the breast milk proteome will contribute not only to the understanding of milk biogenesis and its nutrition value for the newborn, but may also provide guidance on how to develop infant formulas. Moreover, some food allergens, derived from milk, eggs, peanuts, citrus fruits, chocolate, nuts, some cereal grains, certain meats and fish, can circulate through the mother’s body and be present in the breast milk hence into the infant’s gastrointestinal tract and body. Thus, study of the breast milk proteome and identification of these allergens may yield useful information for the mother’s diet improvement [53, 55]. 3.1.2 Sample Collection Protocol

The protocol described below is the most commonly used method for breast milk collection [53, 55, 56] (see Note 25). 1. The subject collects breast milk from one breast with a breast pump in sterile polypropylene containers, at least 2–4 h after prior nursing. 2. Sample must be stored in home freezers and/or transferred to the laboratory in dry ice where they are stored at −80 °C until use. 3. When ready for use, milk is thawed at 4 °C, then ultracentrifuged at 4 °C at 100,000 × g for 60 min so that precipitation of the casein micelles while separation of the fat layer (on the top) and de-lipidated whey supernatant (in the middle) also occur. The de-lipidated whey layer will be utilized in the subsequent proteomic analysis.

3.1.3

Notes

3.2 Bronchoalveolar Lavage Fluid 3.2.1 Introduction

25. Colostrum is collected within the first 48 h of lactation [53]. Lungs, and specifically the airways and the alveoli, are covered with a thin layer of epithelial lining fluid, which is a rich source of many different cells and soluble components of the lung playing an important role in airway integrity and pulmonary defense [44]. Specifically, bronchoalveolar lavage fluid (BALF) consists of both resident alveolar cells and recruited inflammatory cells, their secreted products and proteins from leakage across the endothelial–epithelial barrier. The BALF proteome is dominated by plasmaderived proteins and as such, presents difficulties (e.g., masking effects of the low from the high abundant plasma proteins) in its analysis [57, 58]. The protein composition of BALF reflects the state of the lung. The Pubmed search (October 2013) results for “bronchoalveolar lavage fluid” and “proteomic” are about a hundred.

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Vasiliki Lygirou et al.

3.2.2 Sample Collection Protocol

The collection protocol described below is based on the American Thoracic Society procedures [59] while the described centrifugation and storage steps are commonly followed in the existent BALF proteomics studies [60–62]. 1. Bronchoalveolar lavage is performed by a physician. 2. The procedure should be planned to be performed prior to any other bronchoscopic procedure to avoid specimen contamination. 3. The minimum amount of 2 % lidocaine is applied topically, as there may be bacteriostatic effects. Additional sedatives may also be given to the subjects in order to avoid gag reflex. 4. The subject is positioned in supine position. 5. The bronchoscope is advanced until wedged in a desired subsegmental bronchus at the desired location. The ideal site is determined by the physician after reviewing previously collected radiographs. 6. 20 mL of saline are infused with a syringe while observing the flow of saline at the distal tip of the bronchoscope. 7. Gentle suction (50–80 mmHg) is applied with a suction tube and a vacuum, maintaining wedge position. BALF specimen is collected in a sterile collection trap. 8. Infusion of saline and collection of the BALF are repeated up to five times to obtain an adequate specimen (40–70 % fluid recovery) [59]. 9. BALF samples are mildly centrifuged at 13,000 × g for 15 min to remove cellular debris. 10. The supernatant is transferred into new tubes and the samples are stored at −80 °C until use.

3.3 Nipple Aspirate Fluid 3.3.1 Introduction

Nipple aspirate fluid (NAF) is the liquid that is obtained by aspiration from a nonlactating breast. Fluid secretions remain in the nonlactating breast because of a sphincter at the base of the nipple as well as keratin plugs which may occlude the duct. NAF is therefore considered to have a composition representative of the breast epithelial cell environment. It has high protein concentration ranging from 1 to 200 mg/mL and the most abundant proteins are immunoglobulins, albumin, lactoferrin, and poly-Ig receptor [63]. The Pubmed search (October 2013) results for “nipple aspirate fluid” and “proteomic” are only about 20. Proteomic approaches have yielded valuable information about the NAF protein quantitative and qualitative composition in physiological and pathological conditions. Most studies have focused on the discovery of biomarkers for the diagnosis of cancer or to determine

Body Fluids-SOPs

17

cancer risk [64–67], or as a means of measuring the estrogenic effect of a compound on the breast [63]. 3.3.2 Sample Collection Protocol

The protocol described below is the most commonly reported NAF collection procedure [66, 68] (see Note 26). 1. Prior to NAF collection, the subject massages their breast by applying moisturizing lotion onto it. Massage of the breast is carried out from the chest wall toward the nipple for 2 min. 2. The nipple area is cleansed with an alcohol pad and then a cotton swab in order to remove keratin plugs. 3. Nipple aspiration is performed using a handheld suction device attached to a 10 mL syringe. The collection cup is placed over the nipple and the plunger is withdrawn up to a maximum of 10 mL until NAF is visualized on the nipple surface. 4. NAF samples are stored in eppendorf tubes at −80 °C for up to 6 months.

3.3.3 Notes

4 4.1

26. Nipple aspiration can be performed either by a physician or by the subject itself after training [63].

Abdomen and pelvis fluids Amniotic Fluid

4.1.1 Introduction

4.1.2 Sample Collection Protocol

Amniotic fluid is the liquid component of the amniotic sac in pregnant females. It is breathed and swallowed by the fetus and, thus, contributes to its normal lung development and the formation of urine and meconium. Amniotic fluid provides also physical support to the fetus by cushioning against blows to the mother's abdomen. It also allows for fetus easier movement that promotes muscular and skeletal development. Its composition is complex and includes fetal and maternal proteins, amino acids, carbohydrates, hormones, lipids, and electrolytes [69]. Amniotic fluid has a lower protein concentration relative to plasma and contains highly abundant proteins [70]. Although amniotic fluid is mainly examined for genetic disorders through cytogenetic analysis, its proteome study may contribute to the understanding of the physiology of pregnancy and pathogenesis of gestation. Amniotic fluid is in direct contact with multiple organs of the fetus and contains large amounts of proteins that reflect the fetus’ health and development as well as its physiological interaction with the mother. Although its collection involves an invasive procedure, it is routinely used and the Pubmed search (October 2013) results for “amniotic fluid” and “proteomic” are more than a hundred. The amniotic fluid collection procedure described below is based on the amniocentesis protocol by the Maternal Fetal Medicine,

18

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University of New Mexico [71] while the described centrifugation and storage steps are commonly followed in amniotic fluid proteomics studies [69, 72, 73]. 1. Amniotic fluid is obtained by amniocentesis performed by a physician. It is usually performed in the second trimester between 15 and 20 weeks of gestation but it can be performed at multiple periods throughout pregnancy (see Note 27). 2. Prior to amniocentesis, ultrasound examination is performed in order to determine gestational age, placental position, and amniotic fluid location. Injection of a local anesthetic at the desired puncture site is optional. 3. A 21- or 22-gauge spinal needle with stylet is percutaneously inserted in the subject’s abdomen, under ultrasound guidance, avoiding the fetus, cord, and placenta. Aspiration is performed slowly and steadily in order to avoid needle displacement. 4. Approximately 20 mL of amniotic fluid is aspirated and collected in sterile test tubes. The first 1–2 mL may be discarded to avoid maternal cell contamination [71] (see Note 28). 5. Within 1–2 h, samples are centrifuged at 13,000 × g for 10 min to isolate amniocytes and the cell-free supernatant is transferred to new tubes. 6. Samples are stored at −80 °C until use. 4.1.3

Notes

27. Amniocentesis should be avoided during the first trimester of gestation because of the high risk of pregnancy complications [71]. 28. If the collected amniotic fluid contains blood, proteomic analysis of the specimen may be impaired [71].

4.2

Bile

4.2.1 Introduction

Bile is the fluid that is constantly produced by the liver and is drained by biliary ducts into the gallbladder, where it is stored and concentrated. After meals, the gallbladder releases concentrated bile onto the duodenum, through the common bile duct, where it aids in adsorption and emulsification of fat and serves as a vehicle for excretion of various endogenous (cholesterol, steroid hormones, bilirubin, etc.) or exogenous (drugs, environmental chemicals, etc.) compounds. Bile is composed of salts, lipids, proteins, hormones, inorganic ions, and other metabolites [74, 75]. Proteomic analysis of the bile can yield valuable information about pathologies involving the biliary tract. Specifically, recent studies have focused on the discovery of biomarkers for cholangiocarcinoma and pancreas adenocarcinoma. The Pubmed search (October 2013) results for “bile” and “proteomic” are exceeding one hundred.

Body Fluids-SOPs 4.2.2 Sample Collection Protocol

19

The protocol described below is the commonly used in existing publications [74, 76]. 1. Bile is collected by a surgeon during an endoscopic retrograde cholangiopancreatography or a cholecystectomy (see Note 29). 2. Endoscopically or using a syringe the surgeon aspirates 10–30 mL of the bile. 3. Immediately after collection, the samples are transferred on ice and stored at −80 °C until use. 4. When ready to be used, the samples are centrifuged at 16,000 × g for 10 min at 4 °C to separate from cellular debris.

4.2.3

Notes

4.3 Cervico-vaginal Fluid 4.3.1 Introduction

4.3.2 Sample Collection Protocol

29. In the case of endoscopic retrograde, cholangiopancreatography bile must be collected upstream to the bile duct stenosis before the contrast medium injection [74, 76]. Cervico-vaginal fluid (CVF) is the mucosal fluid that covers the vaginal epithelium and is formed by secretions from cervical vestibular glands, plasma transudate, and endometrial and oviductal fluids. It plays an important role in innate defense and, particularly, in host protection against invading microbes and viruses. CVF contains leukocytes and a range of different molecules including inorganic salts, urea, amino acids, proteins, and a number of fatty acids that derive from commensal microorganisms [77, 78]. CVF is collected through noninvasive procedures and can be used to study the pathophysiology of the female reproductive system. The CVF proteome has primarily been studied for potential markers for pregnancy-associated conditions, such as preterm labor. It can also serve as a source for biomarkers for gynecological malignancies and peri-vaginal infections [79, 80]. The Pubmed search (October 2013) results for “cervical vaginal fluid” and “proteomic” are less than 20. The protocol described below is the most commonly used method for cervico-vaginal fluid collection [81–83] (see Notes 30–33). 1. A sterile double-tipped swab is placed into the subject’s posterior vaginal fornix for 15–20 s until saturation. 2. The swab is removed and placed directly into a polypropylene tube containing buffer solution (e.g., 50 mmol/L HEPES buffer, 150 mmol/L NaCl, 0.1 % SDS, 1 mmol/L EDTA, 1 mmol/L AEBSF protease inhibitor, pH 7.5). 3. The samples are then centrifuged to remove cell debris (for example, at 4 °C, 2,000 × g for 10 min) and the supernatant is transferred to new tubes and stored at −80 °C until use.

20 4.3.3

Vasiliki Lygirou et al. Notes

30. Alternative protocol 1: 5 mL of 0.9 % NaCl are added to the subject’s vaginal cavity using a syringe provided with a 4 cm tubing extension which allows a thorough rinsing of the mucosal surface. The washing is recovered and a second washing is repeated as before. Both washings are combined and a cocktail of protease inhibitors is added. Samples must be kept on ice during the procedure. They are centrifuged at 4 °C at 800 × g for 10 min yielding an average of 7 mL of CVF lavage supernatant which is transferred to a new tube. A second centrifugation at 4 °C at 3,220 × g for 10 min is performed to remove completely all debris and possible microorganisms. The clear, cell-free supernatants are transferred to new tubes and stored at −80 °C until use [77]. 31. Alternative protocol 2: A piece of sterile gauze (5 × 5 cm) is inserted into the vagina, by the subjects themselves, and left there for 1 h. When the gauze is removed, it is stored in 50 mL plastic conical tubes at −20 °C until use. When the gauze is thawed, 10 mL of sterile phosphate-buffered saline (PBS) are added to the tube with the gauze. Then, they are mixed by rotation for 6 h. The extract is removed from the gauze using a 20 mL syringe, which is also used to squeeze the fluid out of the gauze [78]. 32. The ideal CVF collection procedure should be selected depending on the subjects’ condition and their access to trained personnel and special equipment [78]. 33. CVF samples collected with different procedures may have different compositions [78].

4.4

Seminal Plasma

4.4.1 Introduction

Seminal plasma is the liquid part of semen. It is a mixture of secretions from several male accessory glands, including prostate, seminal vesicles, epididymis, and Cowper’s gland. The average protein concentration of seminal plasma ranges from 35 to 55 g/L [84]. It contains many proteins that are important in the capacitation of the spermatozoa, modulation of the immune responses in the uterus, formation of the tubal sperm reservoir and in both sperm–zona pellucida interaction and sperm–egg fusion [85]. Seminal plasma has relatively high protein concentration but also a large dynamic range of protein abundance, making low abundance proteins difficult to detect. Semen is usually easily collected through ejaculation (with the exception of pathological conditions that do not allow its release). The Pubmed search (October 2013) results for “seminal plasma” and “proteomic” are

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more than a hundred. Seminal plasma proteome reflects the physiological and pathological state of the testis, epididymis, and other accessory sex glands; therefore, its analysis can contribute to the establishment of biomarkers for many disorders of the male reproductive system, and particularly male infertility/low fertility and prostate cancer. There are proteins that have been indicated as potential biomarkers for infertility/low fertility [86] and prostate cancer [87] but the degree of results’ overlap between the independent studies is very low due to the different proteomics techniques used in each case [84, 85, 87]. 4.4.2 Sample Collection Protocol

The protocol described below is based on the World Health Organization laboratory manual for the examination and processing of human semen [88] while the centrifugation and storage steps are commonly followed in seminal plasma proteomic studies [87, 89]. 1. The sample should be collected after a minimum of 2 days and a maximum of 7 days of subject’s sexual abstinence. If additional samples are required, the number of days of sexual abstinence should be as consistent as possible at each visit. 2. Semen is obtained by masturbation and ejaculated into a clean, wide-mouthed container made of glass or plastic that has been confirmed to be nontoxic for spermatozoa. 3. The specimen is kept at ambient temperature (20–37 °C) while the semen liquefies, to avoid large changes in temperature that may affect the spermatozoa after ejaculation [88]. 4. Within 1 h after collection and after the semen liquefies, seminal parameters (volume, total sperm count, concentration, sperm motility, morphology, round cells, neutrophils, vitality, and red blood cells) are evaluated. 5. Immediately after semen analysis, samples are centrifuged at 9,200 × g for 20 min for complete separation of seminal plasma from sperm cells and cell debris. 6. Seminal plasma samples are stored at −20 °C until use. Relevant information for all the protocols described in this chapter is summarized in Table 1.

Atraumatic spinal needle, polypropylene tubes

Local anesthesia

–

Cerebrospinal fluid (Subheading 1.4)

Nasal secretions (Subheading 2.1)

Saliva (Subheading 2.2) Refraining from eating, drinking or oral hygiene procedures for at least 1 h

Urine container or urine monovette

Urine (Subheading 1.3) –

Collection tubes and cups, cotton gauzes

Beconase AQ pump aspirator spray device, collection cups

19-gauge needle, blood collection tubes

–

Blood serum (Subheading 1.2)

19-gauge needle, blood collection tubes

–

Necessary equipment

Blood plasma (Subheading 1.1)

Body fluid

Preparation before the collection

Table 1 Summarized information on body fluid collection protocols

No special training needed

No special training needed

Trained physician

No special training needed (Can be collected at home)

Trained phlebotomist

Trained phlebotomist

Training needed

Noninvasive

Noninvasive

Invasive

Noninvasive

Minimally invasive

Minimally invasive

Invasiveness

Whole saliva: 5 mL, Submandibular saliva: 200 μL, Sublingual saliva: 100 μL, Parotid saliva: 1 mL

Depends on the volume of the saline dilutions administered

12 mL minimum

2 mL minimum

As much as needed

As much as needed

Body fluid volume that can be collected

Yes

–

Yes

Only if sample is turbid

Yes

Yes

Centrifugation before storage

Pubmed entries October 2013

−80 °C

−20 °C

−80 °C

−20 or −80 °C if possible

−80 °C

353

20

569

1,049

4,056

−80 °C or liquid 3,396 nitrogen

Storage temperature

Endoscope, syringe, tubes

Syringe or swab or Trained physician sterile gauze, tubes

2 min breast massage

Ultrasound examination

General anesthesia

–

2–7 days sexual abstinence

Nipple aspirate fluid (Subheading 3.3)

Amniotic fluid (Subheading 4.1)

Bile (Subheading 4.2)

Cervico-vaginal fluid (Subheading 4.3)

Seminal plasma (Subheading 4.4)

Wide-mouthed container

21- or 22-gauge spinal needle with stylet, ultrasound, sterile test tubes

Handheld suction device attached to a 10 mL syringe, collection cup

Bronchoscope, syringe, suction tube, vacuum

Radiographs, local anesthesia, sedatives

Bronchoalveolar lavage fluid (Subheading 3.2)

Noninvasive

Invasive

Invasive

No special training Noninvasive needed (Can be collected at home)

Surgeon

Trained physician

As much as possible

Capillary tear collection: 10 μL, Schirmer strips: as much as possible

–

Yes

–

Yes

Ultracentrifugation

Yes

As much as possible

Yes

Depends on the buffer Yes solution volume

10–30 mL

~20 mL

As much as possible

Minimally invasive 40–70 mL

Noninvasive

Noninvasive

Trained physician Noninvasive (Can be collected at home after training)

Trained physician

No special training needed (Can be collected at home)

Breast pump

–

Breast milk (Subheading 3.1)

Trained physician

Capillary tubes or Schirmer strips

Tears (Subheading 2.3) –

−20 °C

−80 °C

−80 °C

−80 °C

−80 °C

−80 °C

−20 °C until transferred in dry ice and then −80 °C

−80 °C

157

16

158

132

21

90

89

74

24

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References 1. American Red Cross. Plasma. Available from http://www.redcrossblood.org/learn-aboutblood/blood-components/plasma 2. Omenn GS, Menon R, Adamski M, Blackwell T, Haab BB, Gao W, States DJ (2007) The human plasma and serum proteome. In: Thongboonkerd V (ed) Proteomics of human body fluids. Humana, Totowa 3. Schulte I, Tammen H, Selle H et al (2005) Peptides in body fluids and tissues as markers of disease. Expert Rev Mol Diagn 5:145–157 4. Granger J, Siddiqui J, Copeland S et al (2005) Albumin depletion of human plasma also removes low abundance proteins including the cytokines. Proteomics 5:4713–4718 5. Guerrier L, Righetti PG, Boschetti E (2008) Reduction of dynamic protein concentration range of biological extracts for the discovery of low-abundance proteins by means of hexapeptide ligand library. Nat Protoc 3:883–890 6. Bodzon-Kulakowska A, Bierczynska-Krzysik A, Dylag T et al (2007) Methods for samples preparation in proteomic research. J Chromatogr B Analyt Technol Biomed Life Sci 849:1–31 7. Omenn GS (2007) THE HUPO human plasma proteome project. Proteomics Clin Appl 1:769–779 8. Muthusamy B, Hanumanthu G, Suresh S et al (2005) Plasma Proteome Database as a resource for proteomics research. Proteomics 5:3531–3536 9. Plasma Proteome Database. Available from http://www.plasmaproteomedatabase.org/ index.html 10. Rai AJ, Gelfand CA, Haywood BC et al (2005) HUPO Plasma Proteome Project specimen collection and handling: towards the standardization of parameters for plasma proteome samples. Proteomics 5:3262–3277 11. Kim MR, Kim CW (2007) Human blood plasma preparation for two-dimensional gel electrophoresis. J Chromatogr B Analyt Technol Biomed Life Sci 849:203–210 12. Di Domenico M, Scumaci D, Grasso S et al (2013) Biomarker discovery by plasma proteomics in familial Brugada Syndrome. Front Biosci (Landmark Ed) 18:564–571 13. Tammen H (2008) Specimen collection and handling: standardization of blood sample collection. Methods Mol Biol 428:35–42 14. Issaq HJ, Xiao Z, Veenstra TD (2007) Serum and plasma proteomics. Chem Rev 107: 3601–3620 15. Adkins JN, Varnum SM, Auberry KJ et al (2002) Toward a human blood serum pro-

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26.

27.

teome: analysis by multidimensional separation coupled with mass spectrometry. Mol Cell Proteomics 1:947–955 Putnam DF (1971) Composition and concentrative properties of human urine. National Aeronautics and Space Administration, Washington Rodriguez-Suarez E, Siwy J, Zurbig P et al (2013) Urine as a source for clinical proteome analysis: from discovery to clinical application. Biochim Biophys Acta 1844(5):884–898 Mischak H, Kolch W, Aivaliotis M et al (2010) Comprehensive human urine standards for comparability and standardization in clinical proteome analysis. Proteomics Clin Appl 4: 464–478 Yamamoto T (2010) The 4th Human Kidney and Urine Proteome Project (HKUPP) workshop. 26 September 2009, Toronto, Canada. Proteomics 10:2069–2070 Yamamoto T, Langham RG, Ronco P et al (2008) Towards standard protocols and guidelines for urine proteomics: a report on the Human Kidney and Urine Proteome Project (HKUPP) symposium and workshop, 6 October 2007, Seoul, Korea and 1 November 2007, San Francisco, CA, USA. Proteomics 8:2156–2159 Zurbig P, Dihazi H, Metzger J et al (2011) Urine proteomics in kidney and urogenital diseases: moving towards clinical applications. Proteomics Clin Appl 5:256–268 van Gool AJ, Hendrickson RC (2012) The proteomic toolbox for studying cerebrospinal fluid. Expert Rev Proteomics 9:165–179 Ramström M, Bergquist J (2007) The human plasma and serum proteome. In: Thongboonkerd V (ed) Proteomics of human body fluids. Humana, Totowa Kroksveen AC, Opsahl JA, Aye TT et al (2011) Proteomics of human cerebrospinal fluid: discovery and verification of biomarker candidates in neurodegenerative diseases using quantitative proteomics. J Proteomics 74: 371–388 Teunissen CE, Petzold A, Bennett JL et al (2009) A consensus protocol for the standardization of cerebrospinal fluid collection and biobanking. Neurology 73:1914–1922 Casado B, Pannell LK, Iadarola P et al (2005) Identification of human nasal mucous proteins using proteomics. Proteomics 5:2949–2959 Casado B, Viglio S, Baraniuk JN (2007) Proteomics of sinusitis nasal lavage fluid. In: Thongboonkerd V (ed) Proteomics of human body fluids. Humana, Totowa

Body Fluids-SOPs 28. Casado B, Iadarola P, Pannell L (2008) Preparation of nasal secretions for proteome analysis. In: Posch A (ed) 2D PAGE: sample preparation and fractionation. Humana, Totowa, pp 77–87 29. Caseiro A, Ferreira R, Padrao A et al (2013) Salivary proteome and peptidome profiling in type 1 diabetes mellitus using a quantitative approach. J Proteome Res 12(4): 1700–1709 30. Casado B, Iadarola P, Pannell LK (2008) Preparation of nasal secretions for proteome analysis. Methods Mol Biol 425:77–87 31. Lamy E, Mau M (2012) Saliva proteomics as an emerging, non-invasive tool to study livestock physiology, nutrition and diseases. J Proteomics 75:4251–4258 32. Vitorino R, Guedes S, Manadas B et al (2012) Toward a standardized saliva proteome analysis methodology. J Proteomics 75:5140–5165 33. He H, Sun G, Ping F et al (2011) A new and preliminary three-dimensional perspective: proteomes of optimization between OSCC and OLK. Artif Cells Blood Substit Immobil Biotechnol 39:26–30 34. Goncalves Lda R, Soares MR, Nogueira FC et al (2010) Comparative proteomic analysis of whole saliva from chronic periodontitis patients. J Proteomics 73:1334–1341 35. Rudney JD, Staikov RK, Johnson JD (2009) Potential biomarkers of human salivary function: a modified proteomic approach. Arch Oral Biol 54:91–100 36. Vitorino R, de Morais Guedes S, Ferreira R et al (2006) Two-dimensional electrophoresis study of in vitro pellicle formation and dental caries susceptibility. Eur J Oral Sci 114: 147–153 37. Cabras T, Pisano E, Mastinu A et al (2010) Alterations of the salivary secretory peptidome profile in children affected by type 1 diabetes. Mol Cell Proteomics 9:2099–2108 38. Hu S, Wang J, Meijer J et al (2007) Salivary proteomic and genomic biomarkers for primary Sjogren’s syndrome. Arthritis Rheum 56:3588–3600 39. Ambatipudi KS, Swatkoski S, Moresco JJ et al (2012) Quantitative proteomics of parotid saliva in primary Sjogren’s syndrome. Proteomics 12:3113–3120 40. Wu ZZ, Wang JG, Zhang XL (2009) Diagnostic model of saliva protein finger print analysis of patients with gastric cancer. World J Gastroenterol 15:865–870 41. Jarai T, Maasz G, Burian A et al (2012) Mass spectrometry-based salivary proteomics for the discovery of head and neck squamous cell carcinoma. Pathol Oncol Res 18:623–628

25

42. Dowling P, Wormald R, Meleady P et al (2008) Analysis of the saliva proteome from patients with head and neck squamous cell carcinoma reveals differences in abundance levels of proteins associated with tumour progression and metastasis. J Proteomics 71:168–175 43. Xiao H, Zhang L, Zhou H et al (2012) Proteomic analysis of human saliva from lung cancer patients using two-dimensional difference gel electrophoresis and mass spectrometry. Mol Cell Proteomics 11(M111):012112 44. WHO (2007) Common minimum technical standards and protocols for biological resource centres dedicated to cancer research. International Agency for Research on Cancer, Working Group Reports. vol. 2 45. Vitorino R, Barros AS, Caseiro A et al (2012) Evaluation of different extraction procedures for salivary peptide analysis. Talanta 94:209–215 46. Ananthi S, Santhosh RS, Nila MV et al (2011) Comparative proteomics of human male and female tears by two-dimensional electrophoresis. Exp Eye Res 92:454–463 47. Saijyothi AV, Angayarkanni N, Syama C et al (2010) Two dimensional electrophoretic analysis of human tears: collection method in dry eye syndrome. Electrophoresis 31:3420–3427 48. Zhou L, Beuerman RW, Chan CM et al (2009) Identification of tear fluid biomarkers in dry eye syndrome using iTRAQ quantitative proteomics. J Proteome Res 8:4889–4905 49. Acera A, Rocha G, Vecino E et al (2008) Inflammatory markers in the tears of patients with ocular surface disease. Ophthalmic Res 40:315–321 50. Argueso P, Balaram M, Spurr-Michaud S et al (2002) Decreased levels of the goblet cell mucin MUC5AC in tears of patients with Sjogren syndrome. Invest Ophthalmol Vis Sci 43:1004–1011 51. Saghizadeh M, Brown DJ, Castellon R et al (2001) Overexpression of matrix metalloproteinase-10 and matrix metalloproteinase-3 in human diabetic corneas: a possible mechanism of basement membrane and integrin alterations. Am J Pathol 158:723–734 52. Wong TT, Zhou L, Li J et al (2011) Proteomic profiling of inflammatory signaling molecules in the tears of patients on chronic glaucoma medication. Invest Ophthalmol Vis Sci 52:7385–7391 53. Liao Y, Alvarado R, Phinney B et al (2011) Proteomic characterization of human milk whey proteins during a twelve-month lactation period. J Proteome Res 10:1746–1754 54. Conti A, Giuffrida MG, Cavaletto M (2007) Proteomics of human milk. In: Thongboonkerd V (ed) Proteomics of human body fluids. Humana, Totowa

26

Vasiliki Lygirou et al.

55. Gao X, McMahon RJ, Woo JG et al (2012) Temporal changes in milk proteomes reveal developing milk functions. J Proteome Res 11:3897–3907 56. Dallas DC, Guerrero A, Khaldi N et al (2013) Extensive in vivo human milk peptidomics reveals specific proteolysis yielding protective antimicrobial peptides. J Proteome Res 12: 2295–2304 57. Govender P, Dunn MJ, Donnelly SC (2009) Proteomics and the lung: analysis of bronchoalveolar lavage fluid. Proteomics Clin Appl 3:1044–1051 58. Foster MW, Thompson JW, Que LG et al (2013) Proteomic analysis of human bronchoalveolar lavage fluid after subsgemental exposure. J Proteome Res 12:2194–2205 59. Lee AS (2004) American Thoracic Society. Bronchoalveolar Lavage. Available from http:// www.thoracic.org/clinical/critical-care/criticalcare-procedures/bronchoalveolar-lavage.php 60. Pastor MD, Nogal A, Molina-Pinelo S et al (2013) Identification of proteomic signatures associated with lung cancer and COPD. J Proteomics 89:227–237 61. Kosanam H, Sato M, Batruch I et al (2012) Differential proteomic analysis of bronchoalveolar lavage fluid from lung transplant patients with and without chronic graft dysfunction. Clin Biochem 45:223–230 62. Cederfur C, Malmstrom J, Nihlberg K et al (2012) Glycoproteomic identification of galectin-3 and -8 ligands in bronchoalveolar lavage of mild asthmatics and healthy subjects. Biochim Biophys Acta 1820:1429–1436 63. Ruhlen RL, Sauter ER (2007) Proteomics of nipple aspirate fluid, breast cyst fluid, milk, and colostrum. Proteomics Clin Appl 1:845–852 64. Ruhlen RL, Sauter ER (2007) Proteomic analysis of breast tissue and nipple aspirate fluid for breast cancer detection. Biomark Med 1:251–260 65. Pavlou MP, Kulasingam V, Sauter ER et al (2010) Nipple aspirate fluid proteome of healthy females and patients with breast cancer. Clin Chem 56:848–855 66. Pawlik TM, Hawke DH, Liu Y et al (2006) Proteomic analysis of nipple aspirate fluid from women with early-stage breast cancer using isotope-coded affinity tags and tandem mass spectrometry reveals differential expression of vitamin D binding protein. BMC Cancer 6:68 67. Alexander H, Stegner AL, Wagner-Mann C et al (2004) Proteomic analysis to identify breast cancer biomarkers in nipple aspirate fluid. Clin Cancer Res 10:7500–7510 68. Noble J, Dua RS, Locke I et al (2007) Proteomic analysis of nipple aspirate fluid throughout the menstrual cycle in healthy pre-

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

menopausal women. Breast Cancer Res Treat 104:191–196 Cho CK, Smith CR, Diamandis EP (2010) Amniotic fluid proteome analysis from Down syndrome pregnancies for biomarker discovery. J Proteome Res 9:3574–3582 Kolialexi A, Tounta G, Mavrou A et al (2011) Proteomic analysis of amniotic fluid for the diagnosis of fetal aneuploidies. Expert Rev Proteomics 8:175–185 Amniocentesis protocol (2011) Available from http://hsc.unm.edu/som/obgyn/docs/ protocols/20.pdf Tsangaris GT, Kolialexi A, Karamessinis PM et al (2006) The normal human amniotic fluid supernatant proteome. In Vivo 20:479–490 Cho CK, Shan SJ, Winsor EJ et al (2007) Proteomics analysis of human amniotic fluid. Mol Cell Proteomics 6:1406–1415 Barbhuiya MA, Sahasrabuddhe NA, Pinto SM et al (2011) Comprehensive proteomic analysis of human bile. Proteomics 11:4443–4453 Farina A, Dumonceau JM, Lescuyer P (2009) Proteomic analysis of human bile and potential applications for cancer diagnosis. Expert Rev Proteomics 6:285–301 Farina A, Dumonceau JM, Frossard JL et al (2009) Proteomic analysis of human bile from malignant biliary stenosis induced by pancreatic cancer. J Proteome Res 8:159–169 Tang LJ, De Seta F, Odreman F et al (2007) Proteomic analysis of human cervical-vaginal fluids. J Proteome Res 6:2874–2883 Shaw JL, Smith CR, Diamandis EP (2007) Proteomic analysis of human cervico-vaginal fluid. J Proteome Res 6:2859–2865 Lockwood CJ, Senyei AE, Dische MR et al (1991) Fetal fibronectin in cervical and vaginal secretions as a predictor of preterm delivery. N Engl J Med 325:669–674 Kalinka J, Sobala W, Wasiela M et al (2005) Decreased proinflammatory cytokines in cervicovaginal fluid, as measured in midgestation, are associated with preterm delivery. Am J Reprod Immunol 54:70–76 Dasari S, Pereira L, Reddy AP et al (2007) Comprehensive proteomic analysis of human cervical-vaginal fluid. J Proteome Res 6: 1258–1268 Di Quinzio MK, Oliva K, Holdsworth SJ et al (2007) Proteomic analysis and characterisation of human cervico-vaginal fluid proteins. Aust N Z J Obstet Gynaecol 47:9–15 Pereira L, Reddy AP, Jacob T et al (2007) Identification of novel protein biomarkers of preterm birth in human cervical-vaginal fluid. J Proteome Res 6:1269–1276

Body Fluids-SOPs 84. Davalieva K, Kiprijanovska S, Noveski P et al (2012) Human seminal plasma proteome study: a search for male infertility biomarkers. Balkan J Med Genet 15:35–38 85. Milardi D, Grande G, Vincenzoni F et al (2012) Proteomic approach in the identification of fertility pattern in seminal plasma of fertile men. Fertil Steril 97:67–73 e61 86. Davalieva K, Kiprijanovska S, Noveski P et al (2012) Proteomic analysis of seminal plasma in men with different spermatogenic impairment. Andrologia 44:256–264

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87. Hassan MI, Kumar V, Kashav T et al (2007) Proteomic approach for purification of seminal plasma proteins involved in tumor proliferation. J Sep Sci 30:1979–1988 88. WHO (2010) Laboratory manual for the examination and processing of human semen, 5th edn. WHO, Geneva 89. da Silva BF, Souza GH, lo Turco EG et al (2013) Differential seminal plasma proteome according to semen retrieval in men with spinal cord injury. Fertil Steril 100: 959–969

Chapter 2 Targeting the Proteome of Cellular Fractions: Focus on Secreted Proteins Agnieszka Latosinska, Maria Frantzi, William Mullen, Antonia Vlahou, and Manousos Makridakis Abstract The high complexity of the total cellular proteome underscores the need for a more targeted investigation of particular subcellular fractions as a means to detect the changes at the level of low abundance proteins. However, this approach requires the application of an enrichment strategy. In this chapter, we present the protocols, which have been used for the analysis of secretome from cell lines, targeting the investigation of protein expression changes. Key words Subcellular proteomics, Cell lines, Secretome, Secretopeptidome, Endoplasmic reticulum, Golgi apparatus

1

Introduction The cell proteome is characterized by a wide dynamic concentration range which extends over seven orders of magnitude [1]. As a result, the low-copy number proteins, which can serve as putative biomarkers for diseases, are extremely difficult to detect [1]. To overcome this limitation, application of a pre-fractionation strategy is necessary to decrease sample complexity. Analysis of secreted proteins is particularly interesting in the context of biomarker discovery. As an example, proteins secreted by cancer cells can be detected in body fluids such as blood or urine. Moreover, investigation of secretome along with the analysis of extracellular matrix proteins may help to elucidate molecular mechanisms underlying cancer aggressiveness. The theoretical basis and summary of different methods allowing for enrichment of secreted proteins are presented in this chapter. The secretome refers to the proteins released by cells through conventional (i.e. signal peptide-dependent) and non-classical ways for secretion. The latter include vesicular (e.g. autophagy-based

Antonia Vlahou and Manousos Makridakis (eds.), Clinical Proteomics: Methods and Protocols, Methods in Molecular Biology, vol. 1243, DOI 10.1007/978-1-4939-1872-0_2, © Springer Science+Business Media New York 2015

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secretion) and non-vesicular pathways (e.g. ATP-binding cassette (ABC) transporters, which belong to transmembrane proteins and utilize the energy from ATP hydrolysis to transport cargo) [2]. In order to investigate the secretome, various in vitro (cell culture), in vivo (tissue samples), and ex vivo (animal models) model systems have been used. However, the majority of the studies have employed in vitro systems and conditioned medium (CM) as a source of secreted proteins. This is mainly attributed to a more straightforward sample collection, allowing for collection of sufficient material to support downstream analysis. In addition, experimental conditions are more easily controlled in in vitro versus in vivo or ex vivo systems. For example, in vivo secretome profiling relies on the analysis of interstitial fluid (component of interstitial space outside parenchymal cells, lymphatic and blood vessels), which is not readily accessible. Even though various strategies have been described to extract interstitial fluid, the process is demanding and may require freshly collected (and not frozen) tissue specimens [3]. The classical procedure requires culturing the cells in serumfree medium (SFM) prior to the collection of conditioned medium. Bearing in mind the broad dynamic range of protein concentration in plasma (3.5–5.0 × 1010 pg/mL to 0–5 pg/mL for serum albumin and interleukin-6, respectively), this is a critical step in order to avoid masking effect and contamination of secretome profile by the highly abundant serum proteins [4]. Nevertheless, it should be noted that in some cases serum starvation may cause a series of undesirable effects to the cell homeostasis such as metabolic stress, reduced growth or enhanced autolysis. Therefore, a fine balance should be targeted with the cells being cultured in serumsupplemented medium until a sufficient confluence is achieved (70–80 %), followed by sequential washes of the cell layer with phosphate-buffered saline (PBS) and SFM to remove the excess of serum proteins. It has been shown that these washing steps have a great impact on the purity of collected material [5]. Cell confluence and incubation time with SFM depends mainly on the cell type and has to be optimized in order to minimize cell lysis and/or cell death. Usually, SFM incubation time varies from 12 to 48 h [6–8]. However, in the case of cells that are strongly dependent on the presence of serum proteins, optimization of the minimum FBS (fetal bovine serum) concentration or adaptation of cells to gradually decreasing serum concentration may be required [9]. After SFM incubation, it is highly recommended to observe the cellular morphology, shape, assess the percentage of dead-necrotic cells (Trypan blue exclusion dye), or evaluate the contamination level from cytoplasmic proteins (e.g. via western blotting of conditioned medium for actin or tubulin) [10]. Conditioned medium is collected and centrifuged to remove dead cells and cellular debris. Due to the low concentration of secreted proteins (ng/mL scale) [11], the medium has to be concentrated prior to analysis. Manifold

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31

strategies have been described in the literature toward that end, including ultrafiltration [12], precipitation [13, 14], and dialysis followed by lyophilization [15, 16]. Depending on the downstream analysis, further desalting may also be required (e.g. gel filtration chromatography using desalting columns or ultrafiltration filter units following buffer exchange protocols). In parallel to the analysis of proteins collected from conditioned medium, the naturally occurring peptides can be also investigated. In this case, the collection of the CM (conditioned medium) is performed in the same way as for the analysis of secreted proteins. However, due to the high sample complexity, a fractionation step is required in order to deplete the high molecular weight compounds. In this case, a strategy based on the use of filter devices (e.g. molecular weight cut off (MWCO): 20 kDa) is adopted to separate larger molecules (MW > 20 kDa) from smaller compounds (MW < 20 kDa, flow through). Therefore, for the peptidomics analysis, the latter (flow through) is collected. Following this step, samples have to be purified by desalting (e.g. PD-10 columns, 5 kDa MWCO). In this way, the final fraction is enriched for molecules between 5 and 20 kDa. The extracted peptides can then be analyzed by capillary electrophoresis coupled to mass spectrometry (CE-MS [17]) or LC-MS/MS. Since the conventional approach of secretome analysis, based on collection of conditioned medium, requires serum starvation (i.e. SFM), which in some cases may result in changes of the secretome profile, application of alternative methodologies is also tested. An interesting approach relies on targeting the secretory pathway organelles (endoplasmic reticulum (ER), Golgi apparatus) and enables investigation of the “secretory cargo” (proteins “on their way” to be secreted). This approach was developed by Sarkar et al. [18] by utilizing human embryonic stem cells and mouse embryonic fibroblasts. The enrichment strategy is based on differential centrifugation. Briefly, after cell lysis, the suspension is centrifuged at 3,000 × g to remove the nuclei. The sample is then depleted of mitochondria via affinity (e.g. Sarkar et al. [18], utilized magnetic microbeads (anti-Tom22)) or centrifugation, as shown in our protocols below. This strategy enriches for the group of putatively secreted proteins, not regularly detected using standard approaches [18]. As such, analysis using the aforementioned different methodologies in combination can lead to better secretome coverage. In this chapter, we present a protocol for secreted proteome enrichment established and regularly used in our laboratory. The presented cell culture conditions refer to the bladder cancer cell line models T24 and T24M. An overview of the major steps of the analytical workflow is presented in Fig. 1. The selected strategies are characterized by high reproducibility (Fig. 2a, b) and optimum extraction yield (Fig. 2b). The efficiency of enrichment is estimated via comparison of results to the total cell extract (Table 1).

Fig. 1 Workflow of classical and non-conventional analysis of secreted proteins in cell line models. The standard approach includes analysis of secreted proteins collected from conditioned medium, whereas the non-conventional method is based on the investigation of protein components derived from secretory pathway organelles (ER/Golgi Apparatus)

Fig. 2 Evaluation of sample preparation procedures for secreted proteins. Representative gel images of total cell extract [1], ER/Golgi fraction [2], and secretome (CM) [3] are shown (a). Three biological replicates were analyzed in order to confirm the reproducibility of applied procedures. In addition, graphical representation of the extracted proteins from conditioned medium and secretory pathway organelles (endoplasmic reticulum/ Golgi apparatus) is presented (b). The secreted proteins from CM were collected from 10 million cells resulting in extraction of 92 μg of protein whereas 20 million cells were used in order to enrich for the ER/Golgi fraction yielding in 76 μg of protein. Regularly, approximately 900 μg of protein (total cell extract) from 4 million cells can be extracted

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Table 1 Evaluation of enrichment efficiency for secreted proteins extracted using classical and nonconventional (ER/Golgi) approaches

Secretome

ER/Golgi fraction

Cell extract

Average number of identifications

994

1,412

1,723

Number of unique identifications

1,613

2,048

2,036

Prediction of signal peptide

379 (24 %)

279 (14 %)

168 (8 %)

Prediction of non-classical secretion

461 (29 %)

632 (31 %)

715 (35 %)

Secreted according to UniprotKB subcellular localization

248 (15 %)

98 (5 %)

70 (3 %)

Extracellular according to Gene Ontology annotationa

332 (21 %)

181 (9 %)

149 (7 %)

The average number of proteins identified using LC-MS/MS techniques (Orbitrap Velos, 8 h LC run in all cases) from five biological replicates per preparation (two for the case of total cell extracts) is reported (5 % FDR). The number of total unique identifications (specifically: sum of unique identifications from all biological samples per preparation) is also shown. In silico analysis of the latter was performed using software for the prediction of signal peptides (SignalIP [19]) and non-classical secretion (SecretomeP [20]) a The following Gene Ontology annotations were included in the analysis: extracellular space (GO:0005615), extracellular matrix (GO:0031012), extracellular region (GO:0005576), proteinaceous extracellular matrix (GO:0005578), and extracellular vesicular exosome (GO:0070062). All annotations were reported in UniProt-GOA annotation database [21]

Classification of proteins as secreted was based on the UniprotKB, Gene Ontology as well as tools for prediction of the classical (SignalIP [19]) and non-conventional secretory pathways (SecretomeP [20]). Detailed comparison of identified proteins following high-resolution LC-MS/MS analysis of all fractions is presented in Fig. 3. Even though contamination of the secretome by intracellular proteins cannot be avoided (Fig. 3a, b), it is clearly shown that the applied procedures enable enrichment in secreted proteins (Table 1 and Fig. 3c, d) in a complementary fashion (Fig. 3e). In parallel, protocols for the peptidomic analysis of secretome via CE-MS are presented. This strategy allowed for the detection of approximately 700 peptides included in 0.9 mL of conditioned medium collected from 4 million bladder cancer cells.

2 2.1

Materials Cell Culture

1. Serum-supplemented growth medium: DMEM (Dulbecco’s modified Eagle medium), High Glucose, GlutaMAX™, Pyruvate, supplemented with 10 % (v/v) FBS and 1 % (v/v) Penicillin/Streptomycin (P/S) (see Note 1). Store at 4 °C (see Note 2). 2. Phosphate-Buffered Saline: 1× in sterile Ultra Pure Water.

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Fig. 3 Comparative analysis of identified proteins from secretome (classical and non-conventional approach) and total cell extract. (I) Comparison of all identified proteins. Venn diagrams representative of all identified proteins from conditioned medium (a) and ER/Golgi fraction (b). In both cases, total cell extract was used as reference. (II) Comparison of secreted proteins. Comparison of the proteins annotated as “secreted” according to UniprotKB in conditioned medium (c) and ER/Golgi fraction (d) compared to secreted proteins identified in total cell extract (Table 1—Secreted according to UniprotKB). Complementarity of the different methods; Overview of common and unique secreted proteins identified in the secretome, ER/Golgi fraction and total cell extract (e). Venn diagrams were prepared using Venn Diagram Plotter (Pacific Northwest National Laboratory, http://omics.pnl.gov/)

Enriching Secretome

2.2 Enrichment of Secreted Proteins

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1. Serum-free medium: DMEM, High Glucose, no Glutamine, no Phenol Red, no HEPES containing medium, supplemented with 4 mM Glutamine and 1 % (v/v) Penicillin/Streptomycin (see Note 3). Store at 4 °C (see Note 2). 2. Amicon Ultra Centrifugal Filter Units, 5 kDa MWCO (see Note 4). 3. Benchtop centrifuge.

2.3 Enrichment of Secreted Peptides

1. Centristart 1 ultrafilter tubes (20 kDa MWCO) (see Note 5). 2. PD-10 desalting Columns (see Note 6). 3. Equilibration/Elution buffer for PD-10 column: Prepare 1 L of 0.01 % v/v ammonium hydroxide solution in distilled water. The pH should be about 10–11. Usually, there is no need for pH adjustment. Store at 4 °C. 4. Lyophilizer.

2.4 Enrichment of Endoplasmic Reticulum and Golgi Apparatus: Secretory Pathway Organelles

1. Lysis buffer: 250 mM sucrose, 25 mM KCl, 5 mM MgCl2, 10 mM triethanolamine, 10 mM acetic acid, 1 mM β-glycerophosphate disodium salt, 1 mM sodium orthovanadate, pH 7.6 adjusted using triethanolamine or acetic acid. The buffer can be prepared as a 5× concentrated stock solution. Store at 4 °C. 2. Solubilization buffer: 7 M urea, 2 M thiourea, 4 % CHAPS, 100 mM DTE, 1 % ampholytes (see Note 7). Store buffer at −20 °C (see Note 8). 3. Bath sonicator (see Note 9). 4. Benchtop centrifuge.

2.5 Total Protein Extraction

1. Lysis buffer: 7 M urea, 2 M thiourea, 4 % CHAPS, 100 mM DTE, 1 % ampholytes. Store buffer at −20 °C (see Note 8). 2. Bath sonicator. 3. Benchtop centrifuge.

3 3.1

Methods Cell Culture

3.2 Collection of Proteins from Conditioned Medium

1. Cultivate cells (37 °C, 5 % CO2 in a humidified atmosphere) in DMEM (High Glucose, GlutaMAX™, Pyruvate) supplemented with 10 % FBS and 1 % Penicillin-Streptomycin until cells reach 80–90 % confluency (0.5–1 × 106 cells/mL), then apply the protocol for CM collection and/or harvest the cells (see Note 10). 1. Remove the medium and rinse the cell layer three times with 1× PBS following by additional wash with serum and phenol red free medium.

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2. Incubate cells with SFM for 24 h (37 °C, 5 % CO2 in a humidified atmosphere) (see Note 11). 3. Collect the conditioned medium and centrifuge at 2,000 × g for 10 min at room temperature (see Note 12); the supernatant should be stored at −80 °C until further processing. 4. Concentrate the conditioned medium with Amicon Ultra Filters up to 50–70 μL (4,000 × g, 12 °C) (see Note 13). 5. Estimate protein concentration, e.g. using Bradford assay. 6. Samples were subjected to LC-MS/MS analysis (see Note 14). 3.3 Enrichment of Secreted Peptides

1. Collect the conditioned medium from approximately four million cells as described in Subheading 3.2 (“Collection of proteins from conditioned media”, steps 1–3) (see Note 11). 2. Fractionate the supernatant with Centrisart filters and collect the filtrate (unconcentrated material