Clinical Applications of Capillary Electrophoresis: Methods and Protocols [2nd ed.] 978-1-4939-9212-6, 978-1-4939-9213-3

Table of contents :
Front Matter ....Pages i-xii
Front Matter ....Pages 1-1
An Overview of CE in Clinical Analysis (David S. Hage)....Pages 3-11
Front Matter ....Pages 13-13
Rapid and Sensitive Determination of Branched-Chain Amino Acids in Human Plasma by Capillary Electrophoresis with Contactless Conductivity Detection for Physiological Studies (Petr Tůma)....Pages 15-24
Monitorization of α1-Acid Glycoprotein Deglycosylation Using SU-8 Microchips Electrophoresis with LIF Detection (María del Mar Barrios-Romero, Agustín G. Crevillén, Angel Puerta, Mercedes de Frutos, José Carlos Diez-Masa)....Pages 25-39
Glycoform Analysis of Alpha1-Acid Glycoprotein by Capillary Electrophoresis Using Electrophoretic Injection (Chenhua Zhang, William Clarke, David S. Hage)....Pages 41-56
On-Line Immunoaffinity Solid-Phase Extraction Capillary Electrophoresis-Mass Spectrometry for the Analysis of Serum Transthyretin (Roger Pero-Gascon, Laura Pont, Victoria Sanz-Nebot, Fernando Benavente)....Pages 57-76
Measurement of Neutral and Sialylated IgG n-Glycome at Asn-297 by CE-LIF to Assess Hypogalactosylation in Rheumatoid Arthritis (Christian Schwedler, Véronique Blanchard)....Pages 77-93
The Control of Glucose and Lactate Levels in Nutrient Medium After Cell Incubation and in Microdialysates of Human Adipose Tissue by Capillary Electrophoresis with Contactless Conductivity Detection (Petr Tůma)....Pages 95-108
Flow-Induced Dispersion Analysis (FIDA) for Protein Quantification and Characterization (Morten E. Pedersen, Jesper Østergaard, Henrik Jensen)....Pages 109-123
Front Matter ....Pages 125-125
A Chiral Generic Strategy for Enantioseparation of Acidic and Basic Drugs Using Short End Injection Capillary Electrophoresis: Application to Design of Experiment (Hassan Y. Aboul-Enein, Ahmed M. Abdel-Megied)....Pages 127-136
Front Matter ....Pages 137-137
New Advances for Newborn Screening of Inborn Errors of Metabolism by Capillary Electrophoresis-Mass Spectrometry (CE-MS) (Meera Shanmuganathan, Philip Britz-McKibbin)....Pages 139-163
Capillary Electrophoresis-Mass Spectrometry for Metabolic Profiling of Biomass-Limited Samples (Wei Zhang, Thomas Hankemeier, Rawi Ramautar)....Pages 165-172
Front Matter ....Pages 173-173
Device Fabrication and Fluorescent Labeling of Preterm Birth Biomarkers for Microchip Electrophoresis (Anna V. Nielsen, Adam T. Woolley)....Pages 175-184
Analysis of Inflammatory Mediators in Newborn Dried Blood Spot Samples by Chip-Based Immunoaffinity Capillary Electrophoresis (Terry M. Phillips, Edward F. Wellner)....Pages 185-198
Triplet-Repeat Primed PCR and Capillary Electrophoresis for Characterizing the Fragile X Mental Retardation 1 CGG Repeat Hyperexpansions (Indhu-Shree Rajan-Babu, Samuel S. Chong)....Pages 199-210
Front Matter ....Pages 211-211
A Capillary Electrophoresis UV Detection-Based Method for Global Genomic DNA Methylation Assessment in Human Whole Blood (Angelo Zinellu, Elisabetta Sotgiu, Salvatore Sotgia, Ciriaco Carru)....Pages 213-219
Capillary Electrophoresis Analysis of Prostate-Specific Antigen (PSA) (Noemi Farina-Gomez, Diana Navarro-Calderon, Angel Puerta, Monica Gonzalez, José Carlos Diez-Masa, Mercedes de Frutos)....Pages 221-234
Prostate Protein n-Glycosylation Profiling by Means of DNA Sequencer-Assisted Fluorophore-Assisted Carbohydrate Electrophoresis (Tijl Vermassen, Nico Callewaert, Sylvie Rottey, Joris R. Delanghe)....Pages 235-250
Aptamer-Based Microchip Electrophoresis Assays for Amplification Detection of Carcinoembryonic Antigen (Shulin Zhao)....Pages 251-259
Front Matter ....Pages 261-261
Highly Sensitive SDS Capillary Gel Electrophoresis with Sample Stacking Requiring Only Nanograms of Adeno-Associated Virus Capsid Proteins (Chao-Xuan Zhang, Michael M. Meagher)....Pages 263-270
Back Matter ....Pages 271-273

Citation preview

Methods in Molecular Biology 1972

Terry M. Phillips Editor

Clinical Applications of Capillary Electrophoresis Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

Clinical Applications of Capillary Electrophoresis Methods and Protocols

Second Edition

Edited by

Terry M. Phillips Department of Pharmaceutics School of Pharmacy, Virginia Commonwealth University Richmond, VA, USA Department of Pharmaceutics, Virginia Commonwealth University, Washington, DC, USA

Editor Terry M. Phillips Department of Pharmaceutics School of Pharmacy Virginia Commonwealth University Richmond, VA, USA Department of Pharmaceutics Virginia Commonwealth University Washington, DC, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9212-6 ISBN 978-1-4939-9213-3 (eBook) https://doi.org/10.1007/978-1-4939-9213-3 Library of Congress Control Number: 2019933290 © Springer Science+Business Media, LLC, part of Springer Nature 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover illustration: This image represents detection of neuropeptides in human cerebral spinal fluid. Cover Image courtesy of Dr. Terry M. Phillips. This Humana Press imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface It is over 30 years since I purchased my first capillary electrophoresis (CE) instrument with the purpose of applying it to clinical analysis, and since that time, it has been very gratifying to see the increased use of this versatile technique in clinically relevant studies. CE is a rapid technique capable of performing complex analyses of a number of different molecular species ranging from small inorganic ions to large nucleic acid fragments and proteins. Perhaps the greatest attribute of this technique is its consumption of minute samples (from 1 μL and less). This coupled with extreme sensitivity, depending upon the type of detector, has led to CE being applied to numerous different fields ranging from analytical chemistry and pharmaceutical analysis to molecular biology and proteomics. The past two decades have shown an increase in CE applications, especially in the clinical sciences where there has been a need for the introduction of rapid, accurate technologies capable of measuring specific analytes in complex biological matrices. CE is a powerful and often rapid tool for performing such analyses, being capable of measuring a number of different molecular species. CE is becoming established as a useful tool in clinical medicine due to several factors, namely, its small sample requirement, low reagent costs, and— depending upon the detection system (LIF, electrochemical, or mass spectrometers)— high sensitivity. Additionally, many CE analyses can be rapidly performed: a strong advantage in diagnostic situations where time can be of the essence. Further, new trends in chipbased CE systems offer clinical analysis, further reduction in sample consumption, and faster analyses. These factors are essential when performing analyses on pediatric or neonatal samples. CE is now replacing many enzyme-based immunoassays because of the less likelihood of false positive and fast turnaround time. Previously published work has also indicated that CE immunoassays on fluids extracted from tissue biopsy samples can aid pathological diagnosis by adding biochemical measurements to the classical histopathology. CE and MCE have been applied to a number of different clinical fields including clinical chemistry, drug analysis and monitoring, endocrinology, hematology, bacteriology and virology, analysis of genetic disorders, pediatric and neonatal analysis, immunology and immunoassays, urology, and nephrology. The 19 papers that comprise this book are divided into seven parts starting with an overview of the application of CE to clinical analysis; this will be followed by the second part, which is dedicated to applications in clinical chemistry and small molecule analysis. Part three will give an example of application in drug analysis, and part four will give a further example of CE applied to metabolomics. Part five will give examples of application in pediatrics, and part six will deal with CE analysis on oncology. Finally part seven will give an example of CE analysis in virology. In conclusion, it is intended that this book will provide a valuable source of information on the application of CE to the many different aspects of clinical medicine. The techniques described may even stimulate new research in the clinical sciences. It is hoped that the book will become a resource not only for clinical chemists but also physicians and scientists, alike, who wish to apply these techniques to diagnosis and clinical research. The techniques outlined in this book will also be useful to biomedical researchers looking for new ways to analyze small biological samples or precious archival samples at a sophisticated level.

v

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Preface

As editor of this book, I would like to express my sincere thanks to all of the authors for their valuable contributions and the referees for their generous gift of their time and expertise in evaluating these chapters. Further, I would like to especially thank Professor John Walker, the editor-in-chief of the series, for his continuous support during the preparation of this issue. My thanks also extend to the staff members of Springer who have performed a wonderful job in producing this book. Washington, DC, USA

Terry M. Phillips

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

PART I

OVERVIEW

1 An Overview of CE in Clinical Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . David S. Hage

PART II

v xi

3

APPLICATIONS IN CLINICAL CHEMISTRY

2 Rapid and Sensitive Determination of Branched-Chain Amino Acids in Human Plasma by Capillary Electrophoresis with Contactless Conductivity Detection for Physiological Studies . . . . . . . . . . . . . . . . 15 Petr Tu˚ma 3 Monitorization of α1-Acid Glycoprotein Deglycosylation Using SU-8 Microchips Electrophoresis with LIF Detection . . . . . . . . . . . . . . . . . 25 Marı´a del Mar Barrios-Romero, Agustı´n G. Creville´n, Angel Puerta, Mercedes de Frutos, and Jose´ Carlos Diez-Masa 4 Glycoform Analysis of Alpha1-Acid Glycoprotein by Capillary Electrophoresis Using Electrophoretic Injection . . . . . . . . . . . . . . . . . 41 Chenhua Zhang, William Clarke, and David S. Hage 5 On-Line Immunoaffinity Solid-Phase Extraction Capillary Electrophoresis-Mass Spectrometry for the Analysis of Serum Transthyretin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57 Roger Pero-Gascon, Laura Pont, Victoria Sanz-Nebot, and Fernando Benavente 6 Measurement of Neutral and Sialylated IgG N-Glycome at Asn-297 by CE-LIF to Assess Hypogalactosylation in Rheumatoid Arthritis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Christian Schwedler and Ve´ronique Blanchard 7 The Control of Glucose and Lactate Levels in Nutrient Medium After Cell Incubation and in Microdialysates of Human Adipose Tissue by Capillary Electrophoresis with Contactless Conductivity Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Petr Tu˚ma 8 Flow-Induced Dispersion Analysis (FIDA) for Protein Quantification and Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Morten E. Pedersen, Jesper Østergaard, and Henrik Jensen

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Contents

PART III

APPLICATION IN DRUG ANALYSIS

9 A Chiral Generic Strategy for Enantioseparation of Acidic and Basic Drugs Using Short End Injection Capillary Electrophoresis: Application to Design of Experiment . . . . . . . . . . . . . . . . . . . . . . . 127 Hassan Y. Aboul-Enein and Ahmed M. Abdel-Megied

PART IV APPLICATION IN METABOLOMICS 10

11

New Advances for Newborn Screening of Inborn Errors of Metabolism by Capillary Electrophoresis-Mass Spectrometry (CE-MS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 Meera Shanmuganathan and Philip Britz-McKibbin Capillary Electrophoresis-Mass Spectrometry for Metabolic Profiling of Biomass-Limited Sample . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 Wei Zhang, Thomas Hankemeier, and Rawi Ramautar

PART V APPLICATIONS IN PAEDIATRICS 12

13

14

Device Fabrication and Fluorescent Labeling of Preterm Birth Biomarkers for Microchip Electrophoresis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 Anna V. Nielsen and Adam T. Woolley Analysis of Inflammatory Mediators in Newborn Dried Blood Spot Samples by Chip-Based Immunoaffinity Capillary Electrophoresis. . . . . . . . 185 Terry M. Phillips and Edward F. Wellner Triplet-Repeat Primed PCR and Capillary Electrophoresis for Characterizing the Fragile X Mental Retardation 1 CGG Repeat Hyperexpansions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199 Indhu-Shree Rajan-Babu and Samuel S. Chong

PART VI

APPLICATIONS IN ONCOLOGY

15

A Capillary Electrophoresis UV Detection-Based Method for Global Genomic DNA Methylation Assessment in Human Whole Blood . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Angelo Zinellu, Elisabetta Sotgiu, Salvatore Sotgia, and Ciriaco Carru 16 Capillary Electrophoresis Analysis of Prostate-Specific Antigen (PSA) . . . . . . . . . Noemi Farina-Gomez, Diana Navarro-Calderon, Angel Puerta, Monica Gonzalez, Jose´ Carlos Diez-Masa, and Mercedes de Frutos 17 Prostate Protein N-Glycosylation Profiling by Means of DNA Sequencer-Assisted Fluorophore-Assisted Carbohydrate Electrophoresis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tijl Vermassen, Nico Callewaert, Sylvie Rottey, and Joris R. Delanghe 18 Aptamer-Based Microchip Electrophoresis Assays for Amplification Detection of Carcinoembryonic Antigen . . . . . . . . . . . . . . . . . . . Shulin Zhao

213

221

235

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Contents

PART VII 19

ix

APPLICATION IN VIROLOGY

Highly Sensitive SDS Capillary Gel Electrophoresis with Sample Stacking Requiring Only Nanograms of Adeno-Associated Virus Capsid Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 263 Chao-Xuan Zhang and Michael M. Meagher

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

271

Contributors AHMED M. ABDEL-MEGIED  Pharmaceutical Analytical Chemistry Department, Faculty of Pharmacy and Pharmaceutical Manufacturing, Kafrelsheikh University, Kafrelsheikh City, Egypt HASSAN Y. ABOUL-ENEIN  Pharmaceutical and Medicinal Chemistry Department, Pharmaceutical and Drug Industries Research Division, National Research Center, Dokki, Cairo, Egypt FERNANDO BENAVENTE  Department of Chemical Engineering and Analytical Chemistry, Institute for Research on Nutrition and Food Safety (INSA·UB), University of Barcelona, Barcelona, Spain VE´RONIQUE BLANCHARD  Charite´—Universit€ atsmedizin Berlin, Campus Virchow Klinikum, Institut fu¨r Laboratoriumsmedizin, Klinische Chemie und Pathobiochemie, Berlin, Germany PHILIP BRITZ-MCKIBBIN  Department of Chemistry and Chemical Biology, McMaster University, Hamilton, ON, Canada NICO CALLEWAERT  Center for Medical Biotechnology, VIB, Ghent, Belgium CIRIACO CARRU  Department of Biomedical Sciences, University of Sassari, Sassari, Italy SAMUEL S. CHONG  Department of Pediatrics, Yong Loo Lin School of Medicine, National University of Singapore, Singapore, Singapore; Khoo Teck Puat—National University Children’s Medical Institute, National University Health System, Singapore, Singapore; Department of Laboratory Medicine, National University Hospital, Singapore, Singapore WILLIAM CLARKE  Johns Hopkins University School of Medicine, Baltimore, MD, USA AGUSTI´N G. CREVILLE´N  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain; Science Faculty—UNED, Madrid, Spain MERCEDES DE FRUTOS  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain JORIS R. DELANGHE  Department of Clinical Chemistry, Microbiology and Immunology, Ghent University, Ghent, Belgium MARI´A DEL MAR BARRIOS-ROMERO  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain JOSE´ CARLOS DIEZ-MASA  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain NOEMI FARINA-GOMEZ  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain MONICA GONZALEZ  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain DAVID S. HAGE  Department of Chemistry, University of Nebraska, Lincoln, NE, USA THOMAS HANKEMEIER  Biomedical Microscale Analytics, Leiden Academic Center for Drug Research, Leiden University, Leiden, The Netherlands HENRIK JENSEN  FIDA-Tech Aps, C/O University of Copenhagen, Copenhagen, Denmark; Department of Pharmacy, University of Copenhagen, Copenhagen, Denmark MICHAEL M. MEAGHER  Department of Therapeutics Production and Quality, St. Jude Children’s Research Hospital, Memphis, TN, USA DIANA NAVARRO-CALDERON  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain ANNA V. NIELSEN  Department of Chemistry and Biochemistry, Brigham Young University, Provo, UT, USA JESPER ØSTERGAARD  FIDA-Tech Aps, C/O University of Copenhagen, Copenhagen, Denmark; Department of Pharmacy, University of Copenhagen, Copenhagen, Denmark

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Contributors

MORTEN E. PEDERSEN  FIDA-Tech Aps, C/O University of Copenhagen, Copenhagen, Denmark ROGER PERO-GASCON  Department of Chemical Engineering and Analytical Chemistry, Institute for Research on Nutrition and Food Safety (INSA·UB), University of Barcelona, Barcelona, Spain TERRY M. PHILLIPS  Department of Pharmaceutics, School of Pharmacy, Virginia Commonwealth University, Richmond, VA, USA; Department of Pharmaceutics, VCU, Washington, DC, USA LAURA PONT  Department of Chemical Engineering and Analytical Chemistry, Institute for Research on Nutrition and Food Safety (INSA·UB), University of Barcelona, Barcelona, Spain ANGEL PUERTA  Institute of Organic Chemistry (IQOG-CSIC), Madrid, Spain INDHU-SHREE RAJAN-BABU  Department of Pediatrics, Yong Loo Lin School of Medicine, National University of Singapore, Singapore, Singapore; Department of Medical Genetics, The University of British Columbia, Vancouver, BC, Canada RAWI RAMAUTAR  Biomedical Microscale Analytics, Leiden Academic Center for Drug Research, Leiden University, Leiden, The Netherlands SYLVIE ROTTEY  Department of Medical Oncology, Ghent University Hospital, Ghent, Belgium; Drug Research Unit Ghent, Ghent University Hospital, Ghent, Belgium VICTORIA SANZ-NEBOT  Department of Chemical Engineering and Analytical Chemistry, Institute for Research on Nutrition and Food Safety (INSA·UB), University of Barcelona, Barcelona, Spain CHRISTIAN SCHWEDLER  Charite´—Universit€ a tsmedizin Berlin, Campus Virchow Klinikum, Institut fu¨r Laboratoriumsmedizin, Klinische Chemie und Pathobiochemie, Berlin, Germany MEERA SHANMUGANATHAN  Department of Chemistry and Chemical Biology, McMaster University, Hamilton, ON, Canada SALVATORE SOTGIA  Department of Biomedical Sciences, University of Sassari, Sassari, Italy ELISABETTA SOTGIU  Department of Biomedical Sciences, University of Sassari, Sassari, Italy PETR TU˚MA  Department of Hygiene, Third Faculty of Medicine, Charles University, Prague, Czech Republic TIJL VERMASSEN  Department of Medical Oncology, Ghent University Hospital, Ghent, Belgium; Drug Research Unit Ghent, Ghent University Hospital, Ghent, Belgium EDWARD F. WELLNER  National Institute of Bioimaging and Bioengineering, National Institutes of Health, Bethesda, MD, USA ADAM T. WOOLLEY  Department of Chemistry and Biochemistry, Brigham Young University, Provo, UT, USA CHAO-XUAN ZHANG  Department of Therapeutics Production and Quality, St. Jude Children’s Research Hospital, Memphis, TN, USA CHENHUA ZHANG  Department of Chemistry, University of Nebraska, Lincoln, NE, USA WEI ZHANG  Biomedical Microscale Analytics, Leiden Academic Center for Drug Research, Leiden University, Leiden, The Netherlands SHULIN ZHAO  Key Laboratory for the Chemistry and Molecular Engineering of Medicinal Resources, College of Chemistry and Pharmacy, Guangxi Normal University, Guilin, China ANGELO ZINELLU  Department of Biomedical Sciences, University of Sassari, Sassari, Italy

Part I Overview

Chapter 1 An Overview of CE in Clinical Analysis David S. Hage Abstract The development and general applications of capillary electrophoresis (CE) in the field of clinical chemistry are discussed. It is shown how the early development of electrophoresis was closely linked to clinical testing. The rise of gel electrophoresis in clinical chemistry is described, as well as the eventual developments that lead to the creation and the use of modern CE. The general principles of CE are reviewed and the potential advantages of this method in clinical testing are examined. Finally, an overview is presented of several areas in which CE has been developed and is currently being explored for use with clinical samples. Key words Capillary electrophoresis, Clinical chemistry, Clinical applications, History of capillary electrophoresis

1

Introduction Electrophoresis has been an important tool in clinical analysis for decades [1–3]. The primary mode of separation in this method is based on the different rates of migration of analytes in an electric field. However, there are many formats for this type of separation and a variety of schemes by which the migration rates of chemicals in a sample can be modified [4–9]. These features, plus the fact that many biological agents hold some charge, have made electrophoresis a key method in clinical analysis for the examination of amino acids, peptides, proteins, and nucleic acids, as well as many small charged solutes [1–3]. One way of using electrophoresis is to apply small amounts of a sample to a support (usually a gel or paper) and allow the components in this sample to travel in a running buffer through the support in the presence of an electric field. These approaches are known as “gel electrophoresis” or “paper electrophoresis” and they have historically been the most common types of electrophoresis found in clinical laboratories [3, 4]. It is also possible to separate the components of a sample by using a narrow capillary that is filled with a running buffer, followed by placement of this buffer and its

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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David S. Hage

contents into an electric field. This second method, known as “capillary electrophoresis” or “CE” [5–8], is the focus of this text. This chapter provides an overview of CE as related to the historical development of this technique and its use in clinical analysis. A summary of the clinical applications of CE, as discussed in more detail in later chapters, is also presented.

2

Origins of Electrophoresis in Clinical Testing It has been known for over a century that substances like proteins and enzymes will travel in an electric field [10–12]. However, the use of this phenomenon for routine chemical separations did not occur until the late 1930s, when Arne Tiselius demonstrated that electrophoresis could be utilized for the separation of serum proteins [13]. The approach used by Tiselius, as later recognized by the 1948 Nobel Prize in Chemistry, was the first practical example in which electrophoresis was used for clinical analysis. The apparatus that was used by Tiselius in these experiments consisted of a U-shaped tube into which he placed his sample and a running buffer (see Fig. 1). When an electric field was applied across this tube, the proteins in the sample began to separate based on their charge and size as they migrated toward the electrode of opposite charge. The result was a series of broad and only partially resolved bands that were then used to study the protein content in the sample [8, 9]. The method that was employed by Tiselius is now known as “moving boundary electrophoresis” because it produced a series of moving boundaries between regions that contained different mixtures of proteins [3, 9]. In modern laboratories, it is more common to use more efficient separation devices and to instead use a small amount of sample that will allow analytes to be separated into narrow bands or zones. These conditions result in a general approach that is now known as “zone electrophoresis” [5–9]. It is interesting that even though Tiselius employed an open tube system, the use of gels or other supports instead of open tubes became the main format for electrophoresis that was employed in clinical laboratories for over 50 years [1–4]. The emphasis on gel or paper electrophoresis during this time was due to the smaller sample requirements, greater ease, and better reproducibility of conducting separations by this approach as opposed to using large open tubes filled with a running buffer. The popularity of gel electrophoresis in particular was further enhanced through the development of improved supports for these separations, such as polyacrylamide gels, and the introduction of new methods based on gel electrophoresis. Two examples of these methods that are still common in clinical and biomedical laboratories are sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and isoelectric focusing (IEF) [3–5, 9, 14].

An Overview of CE in Clinical Analysis

5

Buffer

Protein A Proteins A & B Protein C Proteins B & C

Applicaon of Electric Field

Sample with a Mixture of Proteins A, B & C

Proteins A, B & C

Fig. 1 General design of the apparatus that was used by Arne Tiselius in his early work with electrophoresis and the separation of serum proteins

Interest did continue in further pursuing the use of open tube systems for electrophoresis [14–19]. For instance, Hjerten developed a system in 1967 that utilized 1–3 mm I.D. quartz capillaries to carry out electrophoretic separations in free solution [15]. This system was used for the separation of proteins, nucleic acids, and microorganisms, among other analytes, and included an online detector. Unfortunately, the relatively large diameters of the capillaries in this system required the continuous rotation of these capillaries to minimize the effects of Joule heating and convection [14]. It was demonstrated in 1974 by Virtanen that narrower 0.2 mm I.D. capillaries could be used to eliminate the need for rotation to minimize the effects of heating convection [14, 16]. However, it was not until the commercial development of small diameter silica capillaries in the late 1970s, and the subsequent work by Jorgenson and Lukacs with 75–100 μm I.D. silica capillaries in the early 1980s, that CE became a viable alternative to gel electrophoresis for the separation of clinical and biological samples [17–19].

3

Basic Principles and Advantages of Capillary Electrophoresis Figure 2 shows a typical system for CE, as might be found in a clinical laboratory [3, 6–8]. This system includes a power supply, which can often provide an applied potential up to 25–30 kV, and a computer for control of the system and for the collection of data. There are also two electrodes for applying an electric field across the

6

David S. Hage

Data acquision (& control)

Detector

Capillary (+) Electrode

(-) Electrode Outlet reservoir

Inlet reservoir or sample

High voltage power supply Fig. 2 General design of a modern capillary electrophoresis system. This particular configuration is based on the “normal polarity” mode, in which the sample is applied to a silica capillary at the side of the positive electrode

capillary and buffer containers that supply a contact between these electrodes and the solution within the capillary. Modern CE systems include an online detector, which may be based on the use of UV-vis absorbance, laser-induced fluorescence, electrochemical detection, or mass spectrometry [3, 8]. In addition, the system has some means for injecting samples onto the capillary. Typical volumes for the injected samples are in the pL-nL range and can be applied by using methods such as electrokinetic injection or hydrodynamic injection [6–8]. The capillaries in most modern CE systems have inner diameters of 20–100 μm and lengths of 20–100 cm [6–8]. The use of these narrow bore tubes allows the heat that is generated in the presence of an electric field to be quickly dissipated to the surrounding environment. The removal of this heat helps to provide much more efficient and faster separations than gel or paper electrophoresis. The absence of a gel or support in most types of CE also eliminates eddy diffusion and minimizes secondary interactions that may occur with the support. These conditions create a situation in which longitudinal diffusion is often the main source of

An Overview of CE in Clinical Analysis

7

band-broadening and in which more efficient separations are obtained as the voltage is increased and analytes spend less time in the capillary. The result is a fast separation with high efficiency and narrow peaks [6–8, 17–19]. There are many features of CE that make this method attractive as an alternative to gel or paper electrophoresis for clinical analysis. For instance, CE is faster and more efficient than these traditional methods. CE is also easier to perform as part of an automated system. The small sample size requirements of CE and its ability to be used with various detectors and detection formats are additional features that make this method appealing for clinical analysis [3, 20–25].

4

Applications of CE in Clinical Analysis Following the introduction of the first commercial CE instruments in the late 1980s, there has been a steady increase in the use of CE for various samples of clinical interest. Some early reviews of clinical applications for CE can be found in references [4, 20–23]. Figures 3 and 4 illustrate how the interest in CE for clinical analysis has grown during the last three decades. This interest is indicated by both the number of publications that have appeared on this topic (Fig. 3) and citations that have been made to these papers (Fig. 4). Most of these applications have involved the use of urine, serum, plasma, or blood. In 1990, there were only two publications that dealt with such samples and that included “capillary electrophoresis” within their titles. However, between 1990 and 2017 there were over 1380 publications dealing with these samples and that included “capillary electrophoresis” in their titles. There were also over 27,000 citations to these articles over the same period of time. In addition, work appeared on more exotic specimens such as saliva, cerebrospinal fluid, and tears, with over 80 publications and almost 1760 citations to these papers appearing between 1990 and 2017. Many applications for CE in clinical chemistry have been explored and developed, as is illustrated in Fig. 5 [1, 3, 23–25]. One set of these applications has involved the use of CE in the analysis of clinical biomarkers. These biomarkers may be serum proteins, enzymes, peptides, carbohydrates, lipids, and small organic or inorganic compounds [21, 24]. For instance, some chapters in this text will examine the use of CE to analyze glucose and lactose or serum transthyretin in clinical samples [1, 2]. The use of CE for metabolomics will also be considered in this text. In this latter field, a group of analytes that are involved in one or more metabolic pathways are monitored to determine how their composition changes in the presence or absence of a given clinical condition [24].

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Year Fig. 3 Number of publications appearing between 1990 and 2017 that included the term “capillary electrophoresis” in their titles and that involved typical clinical samples. The results in (a) are for papers that contained “urine,” “serum,” “plasma,” or “blood” in the title of the paper. The results in (b) are for papers that contained “tear(s),” “saliva,” or “cerebrospinal fluid” (CSF) in the title. The results were generated on March 23, 2018, using the Web of Science. The phrases “inductively coupled plasma” and “inductively-coupled plasma” were excluded during the search for papers that included the term “plasma” in their titles

An Overview of CE in Clinical Analysis

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Year Fig. 4 Number of citations to publications that involved the use of capillary electrophoresis and typical clinical samples. The search parameters and procedures were the same as used in Fig. 3. The results in (a) are for papers that contained “urine,” “serum,” “plasma,” or “blood” in the title of the paper. The results in (b) are for papers with titles that contained the terms “tear(s),” “saliva,” or “cerebrospinal fluid” (CSF)

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Analysis of Clinical Biomarkers

Hematology

Endocrinology

Metabolomics Applications of CE in Clinical Chemistry

Genetic Analysis

Bacteriology & Virology

Drug Analysis & Monitoring

Immunology & Immunoassays

Fig. 5 Applications for CE in the field of clinical chemistry

Several additional applications for CE in clinical chemistry will be examined in this text [20–29]. One area in which CE has been employed is hematology, or the study of diseases that are related to blood and blood-forming components [1, 27–29]. Other uses that have reported for CE are in the areas of bacteriology and virology, for the detection or characterization of bacteria or viruses, and in the field of endocrinology, for the analysis and study of hormones within the body [1, 2, 24]. Another important set of applications for CE is in the field of immunology, which deals with diseases of the immune system [1, 2, 24]. A closely related topic is that of CE-based immunoassays, in which CE is combined with the use of antibodies or antibody-related agents for the selective binding and recognition of particular analytes in a sample [26]. CE has also been utilized for the analysis of drugs in clinical samples [21, 24, 25], as might be used for therapeutic drug monitoring or for the detection of drugs of abuse [1, 2]. Finally, CE can be used as a tool for genetic analysis and the identification of inherited disorders [24]. Many of these areas will be explored in more detail in the following chapters. References 1. Rifai N, Horvath AR, Wittwer CT (eds) (2018) Tietz textbook of clinical chemistry and molecular diagnostics, 6th edn. Amsterdam, Elsevier 2. Gornall AG (1986) Applied biochemistry of clinical disorders. Lippincott, New York 3. Hage DS (2016) Chromatography and electrophoresis. In: Clarke W (ed) Contemporary practice in clinical chemistry, 3rd edn. AACC Press, Washington, DC

4. Allen RC, Griffiths JC (1991) Electrophoresis. Anal Chem 63:209R–213R 5. Jorgenson JW (1986) Electrophoresis. Anal Chem 58:743A–760A 6. Skoog DA, Holler FJ, Crouch SR (2017) Principles of instrumental analysis, 7th edn. Cengage Learning, Boston 7. Hage DS, Carr JD (2011) Analytical chemistry and quantitative analysis. Prentice Hall, Boston

An Overview of CE in Clinical Analysis 8. Cazes J (ed) (2005) Ewing’s analytical instrumentation handbook, 3rd edn. CRC Press, Boca Raton 9. Karger BL, Snyder LR, Hovath C (1973) An introduction to separation science. Wiley, New York 10. Hardy WB (1899) On the coagulation of proteid by electricity. J Physiol 26:288–304 11. Hardy WB (1905) Colloidal solution. The globulins. J Physiol 33:251–337 12. Michaelis L (1909) Elektrische uberfuhrung von fermenten. Biochem Z 16:81–86 13. Tiselius AWK (1937) A new apparatus for electrophoretic analysis of colloidal mixtures. Trans Faraday Soc 33:524–531 14. Wehr T, Zhu M (1994) Capillary electrophoresis: historical perspectives. In: Landers JP (ed) Handbook of capillary electrophoresis. CRC Press, Boca Raton 15. Hjerten S (1967) Free zone electrophoresis. Chromatogr Rev 9:122–219 16. Virtanen R (1974) Zone electrophoresis in a narrow-bore tube employing potentiometric detection. Acta Polytech Scand Chem 123:1–67 17. Jorgenson JW, Lukacs KD (1981) Zone electrophoresis in open-tubular glass capillaries. Anal Chem 53:1298–1302 18. Jorgenson JW, Lukacs KD (1981) High resolution separations based on electrophoresis and electroosmosis. J Chromatogr 218:209–216 19. Jorgenson JW, Lukacs KD (1983) Capillary zone electrophoresis. Science 222:266–272

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20. Xu Y (1995) Capillary electrophoresis. Anal Chem 65:425R–433R 21. Xu Y (1995) Capillary electrophoresis. Anal Chem 67:463R–473R 22. Landers JP (1995) Clinical capillary electrophoresis. Clin Chem 41:495–509 23. Petersen JR, Okorodudu AO, Mohammad A, Payne DA (2003) Capillary electrophoresis and its application in the clinical laboratory. Clin Chim Acta 330:1–30 24. Phillips TM, Kalish H (eds) (2013) Clinical applications of capillary electrophoresis. Human Press, Totowa, NJ 25. Bojarski J, Szymura-Oleksiak J (2003) Applications of capillary electrophoresis in clinical analysis of drugs. In: Aboul-Enein HY (ed) Separation techniques in clinical chemistry. Marcel Dekker, New York 26. Moser AC, Willicott CW, Hage DS (2014) Clinical applications of capillary electrophoresis-based immunoassays. Electrophoresis 35:937–955 27. S¸ahin A, Laleli YR, Ortancil R (1995) Haemoglobin analysis by capillary zone electrophoresis. J Chromatogr A 709:121–125 28. Mario N, Baudin B, Bruneel A, Janssens J, Vaubourdolle M (1999) Capillary zone electrophoresis for the diagnosis of congenital hemoglobinopathies. Clin Chem 45:285–288 29. Greene DN, Vaughn CP, Crews BO, Agarwal AM (2015) Advances in detection of hemoglobinopathies. Clin Chim Acta 439:50–57

Part II Applications in Clinical Chemistry

Chapter 2 Rapid and Sensitive Determination of Branched-Chain Amino Acids in Human Plasma by Capillary Electrophoresis with Contactless Conductivity Detection for Physiological Studies Petr Tu˚ma Abstract Capillary electrophoresis (CE) with contactless conductivity detection (C4D) represents a strong tool for determining amino acids in clinical samples. This chapter provides detailed instructions for CE/C4D determination of the branched-chain amino acids (BCAAs) valine, isoleucine, and leucine in human plasma, which can be readily employed in physiological studies. Baseline separation of all the BCAAs is achieved on a short separation length equal to 18 cm in optimized background electrolyte consisting of 3.2 M acetic acid dissolved in 20% v/v methanol with addition of 1.0% v/v INST-coating solution. The analysis time does not exceed 3 min and the limit of detection is 0.4 μM for all BCAAs. The pretreatment of human plasma is very simple and is based on fourfold plasma dilution by acetonitrile and subsequent filtration. Only 50 μL of plasma is used for the analysis. The high sensitivity of the CE/C4D method is achieved by injecting a large volume of sample, combined with application of negative pressure to flush the acetonitrile zone out of the capillary. Key words Branched-chain amino acids, Capillary electrophoresis, Contactless conductivity detection, Human plasma, Large-volume sample stacking, Pressure-assisted analysis, Rapid determination

1

Introduction The proteinogenic branched-chain amino acids (BCAAs) valine (Val), isoleucine (Ile), and leucine (Leu) are essential for humans and other mammals and their synthetic metabolic pathways are localized within the plastids of plants [1]. These three proteinogenic BCAAs account for 35% of the essential amino acids in muscle proteins and simultaneously 40% of the essential amino acids occurring in the human diet [2]. Moreover, BCAAs in their free form have a number of other effects on intermediary metabolism; growth and protein balance; stimulation of secretion of insulin, glucagon, growth hormone, and insulin-like growth factor 1 (IGF 1) [3].

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Supplementation with BCAAs is traditionally used in sports medicine for their anabolic effects and the accelerated muscle convalescence following physical performance. In contrast to other amino acids, BCAAs are oxidized directly in the skeletal muscle, for which they are a significant source of energy during exercise [4]. Similarly, supplementation with BCAAs is used in clinical practice for patients with muscle wasting caused by cancer or age [5]. Further, there has been growing evidence in recent years that elevated plasma BCAA levels are correlated with the risk of diabetes mellitus type 2 and that the BCAA metabolism is disturbed in insulin-resistant conditions. In fact, BCAA levels can predict the risk of development of diabetes mellitus type 2 up to 12 years prior to manifestation [6]. The medicinal requirement for the simultaneous determination of three structurally very similar substances such as Val, Ile, and Leu in large sets of human plasma samples leads to the need to develop a sensitive and rapid determination based on the use of highperformance separation techniques. Capillary electrophoresis (CE), characterized by small requirements on the amount of clinical material for analysis and its uncomplicated laboratory pretreatment prior to analysis, represents a valuable alternative to traditionally used GC [7] and HPLC [8, 9] methods. Moreover, the use of capacitively coupled contactless conductivity detection (C4D) [10–12] instruments, which are currently commercially available (www.istech.at, www.edaq.com, and Admet through www.hpst. cz), has a great advantages in the CE determination of UV-Vis non-absorbing compounds, such as BCAAs [13, 14]. This contribution describes a method for the rapid electrophoretic monitoring of BCAAs in human blood plasma performed on a short separation pathway [15]. The determination is characterized by low LODs at the submicromolar level, which is achieved by using a new type of a large-volume sample stacking [16]. The stacking technique is based on (1) the addition of acetonitrile to the plasma to decrease its conductivity and (2) the application of negative pressure to flush the acetonitrile sample zone out of the capillary.

2

Materials The aqueous solutions should be prepared using ultrapure deionized water with an electric resistance of 18 MΩ cm and all the reagents should be of analytical grade of purity. All solutions should be prepared at room temperature and stored in a refrigerator at 4
C. Prior to use in CE experiments, the solution should be filtered using syringe filters with nylon membranes and a pore size of 0.45 μm to avoid creating a capillary plug.

Rapid Electrophoretic Analysis of Branched-Chain Amino Acids

2.1 Preparation of the Separating Capillary

17

1. Using a capillary cutter, separate a piece of capillary (fused silica capillary with standard polyimide coating) 31.5 cm long from the capillary roll (see Note 1) with inner and outer diameters of 25 and 360 μm, respectively. Use a gas lighter to remove the protective polyimide layer from 5 mm long sections at both ends of the capillary and also cut the capillary ends if necessary (see Note 2). 2. Install the capillary carefully in the cassette of the electrophoretic instrument with integrated C4D and place the cassette in the electrophoretic instrument. 3. Activation of the new capillary: Flush the capillary by a pressure of 1000 mbar with the following solutions in sequence: flush with 0.1 M NaOH for 20 min, wait 10 min, flush with deionized water for 5 min, with INST-coating solution for 5 min, again with deionized water for 5 min, and finally with background electrolyte (BGE) for 30 min (see Note 3). Then apply the maximum separation voltage of +30 kV and monitor the current for approx. 10 min. The current should stabilize at a value of 4.0 μA.

2.2

Solutions for CE

1. BGE: 3.2 M acetic acid (HAc) dissolved in 20% v/v methanol (MeOH) with addition of 1.0% v/v INST-coating solution, pH 2.0; pipet 9.2 mL of concentrated HAc (99%, Mr 60, 1.05 g/mL, 17.3 M) and 10.0 mL MeOH in a 50 mL volumetric vessel and finally dilute with water to the mark, store at laboratory temperature (see Note 4). 2. Stock solutions of BCAAs and internal standard (IS): 10 mM Leu, 10 mM Ile, 10 mM Val, 10 mM tris(hydroxymethyl) aminomethane (Tris, Mr 121.14, use as IS); weigh 65.6 mg of Leu (Mr 131.18), 65.6 mg of Ile (Mr 131.18), 58.6 mg of Val (Mr 117.15), and 60.6 mg of Tris and dissolve each substance in an individual 50 mL volumetric flask and dilute to the mark with deionized water (see Note 5), store in storage bottles at 4
C. 3. 0.1 M Sodium hydroxide (NaOH) for capillary activation: weigh 400 mg of solid NaOH (Mr 40.0), dissolve in deionized water and dilute to the mark in a 100 mL volumetric flask, store in a storage bottle. 4. 0.154 M Sodium chloride (NaCl) for salinization of model samples: weigh 0.45 g of solid NaCl (Mr 58.4), dissolve it and dilute to the mark with deionized water in a 50 mL volumetric flask, store in a storage bottle; or use a commercial infusion solution (154 mM NaCl) (see Note 6). 5. 0.01 M HCl acidified acetonitrile (ACN) for plasma deproteinization: pipet 25 mL of ACN into a storage bottle, add 20 μL of hydrochloric acid (37%, Mr 36.5, 1.2 g/mL, 12.1 M).

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2.3 Laboratory Pretreatment of Human Blood Plasma Samples

The obtained clinical samples of human blood plasma, sampled according to the rules for sampling clinical materials [17], can be stored in micro-centrifuge tubes for long time periods at a temperature of 80
C. Prior to CE analysis, the samples are melted at laboratory temperature (see Note 7). Use protective gloves when working with biological materials. 1. Laboratory pretreatment of plasma samples. Pipet 50 μL of blood plasma into a 250 μL micro-centrifuge tube, add 1.0 μL of IS and shake briefly on a laboratory shaker. Then add 150 μL of acidified ACN (see Note 8), shake for 2 min and centrifuge for 2 min at an acceleration of 4000  g on a laboratory centrifuge without cooling. Collect the supernatant and filter it using a centrifugal filter device with microporous membrane and pore size of 0.45 μm, at an acceleration of 4000  g, for 2 min. Pipet 50 μL of the filtered sample into a plastic vial with insert and use it for CE analysis.

3

Methods

3.1 CE Analyses of a Model BCAA Sample

1. Preparation of a model BCAA sample at 10 μM concentration level: Pipet 1.0 μL of 10 mM stock solution of Val, 1.0 μL of 10 mM stock solution of Ile, 1.0 μL of 10 mM stock solution of Leu, and 5 μL of 10 mM stock solution of IS, add 250 μL of 0.154 M NaCl, and finally add 750 μL of acidified ACN. 2. Pipet 0.99 mL of 3.2 M HAc dissolved in 20% v/v MeOH (BGE) into three vials, add 10 μL of INST-coating solution to each one: the first serves as the input CE vial, the second as the output CE vial, and the third as the washing vial (see Note 9). Place the vials with the BGE, an empty vial for the waste and the vial containing the sample in the CE instrument. 3. Program the following individual actions on the CE instrument [18] in sequence: (a) Washing of the capillary with the BGE for 2 min (input vial: BGE; output vial: waste). (b) Hydrodynamic sample injection under a pressure of 50 mbar for 16 s into the long end of the separation capillary (input vial: sample; output vial: waste). (c) CE separation with simultaneous forcing of the ACN sample zone out of the capillary: switch on the separation voltage with a linear ramp from 0 to +30 kV over a time of 12 s, and simultaneously apply a negative pressure of 50 mbar for 16 s to force the ACN sample zone out of the capillary (input vial: BGE; output vial: BGE; see Note 10).

Rapid Electrophoretic Analysis of Branched-Chain Amino Acids

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Fig. 1 CE/C4D analysis of a model mixture of BCAAs performed in BGE consisting of 3.2 M HAc dissolved in 20% v/v MeOH with addition of 1.0% v/v INST-coating solution, pH 2.0. Sample composition: 5 μM mixture of Val, Ile, Leu; 50 μM Tris (IS), dissolved in 0.154 M NaCl and finally diluted by acidified ACN in a ratio of 1:3 (75% v/v ACN content in the sample). Capillary: inner diameter, 25 μm, total length, 31.5 cm, length to the detector, 18 cm. Hydrodynamic injection at a pressure of 50 mbar for 16 s; separation voltage, +30 kV with ramp 12 s; current, 4.1 μA; forcing the rest of the sample zone out of capillary by a pressure of 50 mbar for 16 s. Peak identification: 1 sodium, 2 Tris (IS), 3 Val, 4 Ile, 5 Leu. (a) The complete electropherogram with a large sodium zone, (b) detail of the BCAA zones

4. Carry out the CE separation for 3 min. 5. Evaluate the electropherogram obtained and determine the BCAA migration times (t) and peak areas (A) (see Notes 11, 12 and Fig. 1). 3.2 Calibration for Model BCAA Samples

1. Prepare the model calibration solutions of BCAAs at five concentration levels: 5, 10, 50, 100, and 200 μM, dissolved in the acidified ACN with final content of 75% v/v and with addition of the infusion solution (154 mM NaCl): (a) The 5 μM calibration solution: Pipet 0.5 μL volumes of the Val, Ile, and Leu 10 mM stock solutions, 5 μL of 10 mM Tris, 250 μL of 154 mM NaCl, and 750 μL of ACN into the CE vial. The exact BCAA concentrations should be calculated according the formula: c ¼ 10 mM  0.5 μL/ (3  0.5 μL + 5 μL + 250 μL + 750 μL) ¼ 5.0 μM. (b) The 10 μM calibration solution: Pipet 1.0 μL volumes of Val, Ile, and Leu 10 mM stock solutions, 5 μL of 10 mM Tris, 250 μL of 154 mM NaCl, and 750 μL of ACN into the CE vial. The exact BCAA concentrations are: c ¼ 10 mM  1 μL/ (3  1 μL + 5 μL + 250 μL + 750 μL) ¼ 9.9 μM.

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(c) The 50 μM calibration solution: Pipet 5.0 μL volumes of the Val, Ile, and Leu 10 mM stock solutions, 5 μL of 10 mM Tris, 250 μL of 154 mM NaCl, and 750 μL of ACN into the CE vial. The exact BCAA concentrations are: c ¼ 10 mM  5 μL/ (3  5 μL + 5 μL + 250 μL + 750 μL) ¼ 49.0 μM. (d) The 100 μM calibration solution: Pipet 10.0 μL volumes of the Val, Ile, and Leu 10 mM stock solutions, 5 μL of 10 mM Tris, 250 μL of 154 mM NaCl, and 750 μL of ACN into the CE vial. The exact BCAA concentrations are: c ¼ 10 mM  10 μL/ (3  10 μL + 5 μL + 250 μL + 750 μL) ¼ 96.6 μM. (e) The 200 μM calibration solution: Pipet 20.0 μL volumes of the Val, Ile, and Leu 10 mM stock solutions, 5 μL of 10 mM Tris, 250 μL of 154 mM NaCl, and 750 μL of ACN into the CE vial. The exact BCAA concentrations are: c ¼ 10 mM  20 μL/ (3  20 μL + 5 μL + 250 μL + 750 μL) ¼ 187.8 μM. 2. Analyze all the calibration solutions three times according the procedure (see Subheading 3.1, step 3 and, after every 20 analyses, replace the BGE in the vials with a fresh solution. Measure the areas under the peaks (A [mV s]) in the electropherograms and construct the calibration dependences for all the BCAAs in the form, A ¼ S·c + I, where S [mV s/μM] is the slope of the calibration plot, c [μM] is the exact BCAA concentration, and I [mV s] is the intercept of the calibration plot. 3. If necessary, the calibration dependence for model samples can be used to determine the BCAA level in blood plasma, e.g., when there is an insufficient amount of clinical material for the preparation of the calibration solutions. When recalculating the concentration to the untreated plasma, it is necessary to take into account its fourfold dilution during sample treatment (the resultant concentration in the blood plasma is four times greater compared to the treated sample). 3.3 Calibration and Determination of BCAAs in Blood Plasma Samples

1. Prepare five plasma calibrators by spiking one untreated plasma sample with stock BCAA solutions at five concentration levels: 10, 25, 50, 100, and 200 μM: (a) The 10 μM plasma calibrators: Pipet 1.0 μL volumes of the Val, Ile, and Leu 2 mM stock solutions and 1 μL of 10 mM Tris into a 250 μL micro-centrifuge tube, add 50 μL of plasma, and shake in a laboratory shaker for 2 min. Then add 150 μL of ACN and perform the deproteinization procedure according to item 1 of Subheading 2.3 and finally pipet 50 μL of the supernatant into a plastic vial with insert and use it for CE analysis. The exact BCAA concentrations in the treated plasma sample should be

Rapid Electrophoretic Analysis of Branched-Chain Amino Acids

21

calculated according to the formula: c ¼ 2 mM  1 μL/ (3  1 μL + 1 μL + 50 μL + 150 μL) ¼ 9.8 μM. (b) The 25 μM plasma calibrators: Pipet 0.5 μL volumes of the Val, Ile, and Leu 10 mM stock solutions and 1 μL of 10 mM Tris into a 250 μL micro-centrifuge tube, add 50 μL of plasma and, after a shaking, add 150 μL of ACN and perform the deproteinization. The exact BCAA concentrations are: c ¼ 10 mM  0.5 μL/ (3  0.5 μL + 1 μL + 50 μL + 150 μL) ¼ 24.7 μM. (c) The 50 μM plasma calibrators: Pipet 1.0 μL volumes of the Val, Ile, and Leu 10 mM stock solutions and 1 μL of 10 mM Tris into a 250 μL micro-centrifuge tube, add 50 μL of plasma and, after a shaking, add 150 μL of ACN and perform the deproteinization. The exact BCAA concentrations are: c ¼ 10 mM  1 μL/ (3  1 μL + 1 μL + 50 μL + 150 μL) ¼ 49.0 μM. (d) The 100 μM plasma calibrators: Pipet 2.0 μL volumes of the Val, Ile, and Leu 10 mM stock solutions and 1 μL of 10 mM Tris into a 250 μL micro-centrifuge tube, add 50 μL of plasma and, after a shaking, add 150 μL of ACN and perform the deproteinization. The exact BCAA concentrations are: c ¼ 10 mM  2 μL/ (3  2 μL + 1 μL + 50 μL + 150 μL) ¼ 96.6 μM. (e) The 200 μM plasma calibrators: Pipet 4.0 μL volumes of the Val, Ile, and Leu 10 mM stock solutions and 1 μL of 10 mM Tris into a 250 μL micro-centrifuge tube, add 50 μL of plasma and, after a shaking, add 150 μL of ACN and perform the deproteinization. The exact BCAA concentrations are: c ¼ 10 mM  4 μL/ (3  4 μL + 1 μL + 50 μL + 150 μL) ¼ 187.8 μM. 2. Analyze one blank plasma sample and its 5 spiked calibrators three times according to the procedure (see Subheading 3.1, step 3) and, after every 10–15 analyses, replace the BGE in the vials with a fresh solution. Measure the areas under the peaks (A [mV s]) in the electropherograms and construct the calibration dependences for all the BCAAs in the form, A ¼ S·c + I, where S [mV s/μM] is the slope of the calibration plot, c [μM] is the exact BCAA concentration, and I [mV s] is the intercept of the calibration plot. When recalculating the concentration in the untreated plasma, it is once again necessary to take into consideration its fourfold dilution during sample treatment. 3. Perform analysis of a set of clinical plasma samples, each in triplicate. Evaluate the positions of the individual BCAAs in the electropherograms according the migration times compared to the calibration procedure (see Fig. 2). Measure the peak areas of the BCAAs and IS. Normalize the obtained

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Fig. 2 Records of CE/C4D analysis of a single blood plasma sample for BCAA determination. (a) Treated plasma without IS addition; (b) treated plasma with IS addition, (c) plasma spiked by BCAAs at a level of 50 μM. Peak identification: (1) sodium, (2) Tris (IS), (3) Val, (4) Ile, (5) Leu. For the other experimental conditions see Fig. 1

BCAA areas according to (1) the migration time of the BCAAs in the calibrators and (2) the area of IS. Finally, compute the BCAAs using the corresponding calibration dependence. 4. The procedures of calibration and of biological sample analyses can be automated, with work in a sequential regime. Program the replacement of the BGE in the vials for the fresh solution after 20 analyses.

4

Notes 1. A capillary length of 31.5 cm is the minimum for the Agilent 7100 and HP3DCE systems capillary electrophoresis instruments (Agilent Technologies); the effective capillary length (the distance from the sample introduction end to the detection cell) is 18 cm [18]. 2. A detection window need not be created on the capillary when using C4D. 3. Capillary coating using the INST-coating solution effectively reduces the EOF in acidic BGEs [19], which is necessary for large-volume sample stacking experiments and also to achieve baseline separation of the BCAAs on a short separation pathway.

Rapid Electrophoretic Analysis of Branched-Chain Amino Acids

23

4. 10 μL addition of INST-coating solution is used only in the final electrophoretic vial to minimize the consumption of expensive additive. 5. The volumetric flasks can be inserted in an ultrasonic bath for 5 min to accelerate dissolution of the solid substances. 6. Addition of an infusion solution to the sample (finally fourfold diluted) simulates the natural composition of human plasma after its dilution. The presence of sodium ions in the sample creates the conditions for a special electrophoretic mode called transient-isotachophoresis [20]. 7. The melting of the samples can be accelerated by placing a sample containing a micro-centrifuge tube in an ultrasonic bath for 5 min. 8. Pipette liquid using a special pipette equipped with a tip with movable piston for exact addition of organic solvent. 9. Maintain the same solution level heights in the input and output vials containing the BGE [18]. 10. The nonconductive ACN sample zone, formed by the migration of ions out of the zone, presents excessive resistance for the passage of electric current. If it is not forced out of the capillary by a pressure impulse, it causes a current fluctuation or complete interruption of the separation [15]. 11. BCAAs appear as negative peaks on the C4D electropherograms. This is caused by the fact that the BCAA electrophoretic mobility is lower than that of the BGE co-ions - H3O+ ions [21]. 12. For instance, the separation of a 5 μM BCAA mixture depicted in Fig. 1 should be evaluated as: t (IS) 1.76 min, A (IS) 1.77 mV s; t (Val) 2.42 min, A (Val) 0.37 mV s; t (Ile) 2.46 min, A (Ile) 0.36 mV s; t (Leu) 2.53 min, A (Leu) 0.33 mV s.

Acknowledgements This work was supported by the Grant Agency of the Czech Republic, Grant No. 18-04902S. References 1. Singh BK, Shaner DL (1995) Biosynthesis of branched-chain amino-acids - from test-tube to field. Plant Cell 7(7):935–944 2. Murray RK, Bender DA, Botham KM, Kennelly PJ, Rodwell VW, Weil PA (2007)

Harper’s illustrated biochemistry, Chapter 28 and 33., 29th edn. McGraw-Hill Lange, China 3. Adeva MM, Calvino J, Souto G, Donapetry C (2012) Insulin resistance and the metabolism of branched-chain amino acids in humans. Amino Acids 43(1):171–181

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4. Devlin TM (1992) Biochemistry with clinical correlations. Wiley-Liss, New York 5. Tazi EM, Errihani H (2010) Treatment of cachexia in oncology. Indian J Palliat Care 16 (3):129–137 6. Wang TJ, Larson MG, Vasan RS, Cheng S, Rhee EP, McCabe E, Lewis GD, Fox CS, Jacques PF, Fernandez C, O’Donnell CJ, Carr SA, Mootha VK, Florez JC, Souza A, Melander O, Clish CB, Gerszten RE (2011) Metabolite profiles and the risk of developing diabetes. Nat Med 17(4):448–U483 7. Husˇek P, Sˇimek P, Hartvich P, Zahradnı´cˇkova´ H (2008) Fluoroalkyl chloroformates in treating amino acids for gas chromatographic analysis. J Chromatogr A 1186(1–2):391–400 ˇ akova´ P, Jirosˇova´ J, Sladka´ M 8. Kand’a´r R, Z (2009) Determination of branched chain amino acids, methionine, phenylalanine, tyrosine and alpha-keto acids in plasma and dried blood samples using HPLC with fluorescence detection. Clin Chem Lab Med 47 (5):565–572 9. Sharma G, Attri SV, Behra B, Bhisikar S, Kumar P, Tageja M, Sharda S, Singhi P, Singhi S (2014) Analysis of 26 amino acids in human plasma by HPLC using AQC as derivatizing agent and its application in metabolic laboratory. Amino Acids 46(5):1253–1263 10. Kuba´nˇ P, Hauser PC (2017) Contactless conductivity detection for analytical techniques developments from 2014 to 2016. Electrophoresis 38(1):95–114 11. Kuba´nˇ P, Hauser PC (2015) Contactless conductivity detection for analytical techniquesdevelopments from 2012 to 2014. Electrophoresis 36(1):195–211 12. Kuba´nˇ P, Hauser PC (2013) Contactless conductivity detection for analytical techniques:

developments from 2010 to 2012. Electrophoresis 34(1):55–69 13. Coufal P, Zuska J, van de Goor T, Smith V, Gasˇ B (2003) Separation of twenty underivatized essential amino acids by capillary zone electrophoresis with contactless conductivity detection. Electrophoresis 24(4):671–677 14. Tu˚ma P, Ma´lkova´ K, Samcova´ E, Sˇtulı´k K (2010) Rapid monitoring of arrays of amino acids in clinical samples using capillary electrophoresis with contactless conductivity detection. J Sep Sci 33(16):2394–2401 15. Tu˚ma P, Gojda J (2015) Rapid determination of branched chain amino acids in human blood plasma by pressure-assisted capillary electrophoresis with contactless conductivity detection. Electrophoresis 36(16):1969–1975 16. Tu˚ma P, Jacˇek M, Fejfarova´ V, Pola´k J (2016) Electrophoretic stacking for sensitive determination of antibiotic ceftazidime in human blood and microdialysates from diabetic foot. Anal Chim Acta 942:139–145 17. Guder WG, Narayanan S, Wisser H, Zawta B (2009) Diagnostic samples: from the patient to the laboratory. Wiley-VCH, Weinheim 18. Lauer HH, Rozing GP (2010) High performance capillary electrophoresis. Agilent Technologies, Germany 19. Tu˚ma P (2014) Rapid determination of globin chains in red blood cells by capillary electrophoresis using INSTCoated fused-silica capillary. J Sep Sci 37(8):1026–1032 20. Krˇiva´nkova´ L, Pantu˚cˇkova´ P, Bocˇek P (1999) Isotachophoresis in zone electrophoresis. J Chromatogr A 838(1–2):55–70 21. Kuba´nˇ P, Hauser PC (2009) Ten years of axial capacitively coupled contactless conductivity detection for CZE - a review. Electrophoresis 30(1):176–188

Chapter 3 Monitorization of α1-Acid Glycoprotein Deglycosylation Using SU-8 Microchips Electrophoresis with LIF Detection Marı´a del Mar Barrios-Romero, Agustı´n G. Creville´n, Angel Puerta, Mercedes de Frutos, and Jose´ Carlos Diez-Masa Abstract In the last few years, biopharmaceuticals—therapeutic drugs which are generally obtained by using molecular biology techniques—have become a major growing sector in pharmaceutical industry. A large part of these biopharmaceuticals are therapeutic glycoproteins. The production of these drugs and their purification process are implying the development of efficient analytical methods, which allow quick and reliable control of the manufacturing process and ensuring the regulatory compliance about the quality of these drugs. Capillary gel electrophoresis (CGE) in the presence of sodium dodecyl sulfate (SDS) is becoming a method of choice in the quality control of these biopharmaceuticals. On the other hand, CGE can be improved if analyses are carried out in microchip format. This chapter reports a detailed microchips gel electrophoresis (MGE) method to separate glycosylated and deglycosylated forms of α1-acid glycoprotein (AGP) labeled with Chromeo P540, using SU-8 microchips and laser induced fluorescence detection. Due to the analogy between AGP and some therapeutic glycoproteins, we have selected AGP as a model system to illustrate the potential of MGE in the analysis of this type of biopharmaceutical compounds. Key words Microchip gel electrophoresis, SU-8 microchips, LIF detection, α1-acid glycoprotein, AGP, Glycoprotein, Deglycosylation

1

Introduction In recent years, the biotechnological field has been the main growing sector in pharmaceutical industry. One of the main reasons for this expansion is the development of new type of products, named as biopharmaceuticals, biosimilars, and biobetters (see Note 1), which are generally obtained using molecular biology techniques [1, 2]. Many of these biotherapeutic products approved or in clinical trials are glycoproteins, in which glycosylation is recognized as a Critical Quality Attribute (see Note 1). These glycoproteins include a large number of monoclonal antibodies, recombinant cytokines, and recombinant enzymes used in replacement therapies. They are

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_3, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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used in the treatment of various diseases such as different types of cancer, some chronic inflammations, and several infections [3]. The production of biotherapeutic drugs and their purification process is highly complex. It implies the development of efficient analytical methods that allow controlling quickly and reliably the manufacturing process and ensuring also the drug quality. Many controls are carried out to ensure the biological activity of these products, as well as their stability and consistency between batches [4, 5]. A critical point in glycoprotein production is glycosylation. A small alteration throughout the production process can lead to an uncompleted or altered glycosylation and therefore to a batch without the proper biological activity or inconsistent with the previous ones. Consequently, one important assay in manufacturing biopharmaceuticals is the glycosylation control [6]. However, there is no accepted standardized approach for therapeutic glycoprotein characterization and therefore the industry must use customized approaches for controlling therapeutic glycoproteins. SDS-PAGE and capillary gel electrophoresis (CGE) are analytical techniques commonly used for purity determination by separation and quantitation of size-based protein variants and, as a consequence, these techniques are useful for glycosylation control. Whereas SDS-PAGE is a manual and semiquantitative technique, CGE allows developing fast and automatic analytical methodologies [7, 8]. Moreover, CGE can be improved if analyses are carried out in microchip format, that is, in microchip gel electrophoresis (MGE). This is due to the multiple advantages that microchips offer, such as faster analysis, high separation efficacy, low reagent consumption, high throughput analysis, and the possibility of integrating all analytical operations in a single device [9]. Commercial equipment employ glass or quartz microchips, but polymeric chips (i.e., polymethyl methacrylate, polydimethylsiloxane, or SU-8) are arising as an interesting alternative because they can be fabricated using mass-replication technologies, which are less expensive than photolithographic techniques used in preparation of glass or quartz microchips. This allows the production of low-cost disposable chips for single-use applications, minimizing cross-contamination and avoiding the use of cleaning protocols [10]. This chapter reports a detailed method to separate glycosylated and deglycosylated forms of α1-acid glycoprotein (AGP) labeled with Chromeo P540, using SU-8 microchips, and laser induced fluorescence (LIF) detection. Due to the analogy between AGP and some therapeutic glycoproteins, we have selected AGP as a model protein for this kind of biopharmaceuticals to illustrate the potential of MGE in the analysis of this type of unconventional medicaments.

Electrophoresis of AGP Deglycosylation in SU-8 Microchips

2 2.1

27

Materials Microchip Setup

1. The commercial microchips used are made of SU-8 (reference MLCE4515-5050, MicroLIQUID, Arrasate-Mondragon, Spain). They are cross-shaped with external dimensions 15
45
0.9 mm and square section channels (50
50 μm). The length of the short channels is 5 mm and the length of the long channel (separation channel) is 35 mm. At the end of the channels, there are 2 mm diameter reservoirs. The SU-8 chips with microchannel structure are supported on a Pyrex glass layer. 2. The SU-8 microchips are used in a homemade microchip holder. This holder is made of polymethyl methacrylate (PMMA) and consists of two plates, fastened by using plastic screws (Fig. 1B). The top plate (Fig. 1A) (dimensions 36
68
15 mm) has six holes for M-6 screws to affix it to the bottom plate. In the top plate, there are also four threaded holes, vertically aligned with the microchip reservoirs, which allow the connection of the reservoirs to the syringe pump. The top plate has too four integrated 0.5 mm diameter platinum electrodes for applying voltage. Silicone loaded O-rings (6
1.5 mm) (Grema, Madrid, Spain) placed between the top plate and the microchip reservoirs allow closing hermetically the device and prevent liquid leakage. The bottom plate (dimensions 86
88
4 mm) has six threaded holes which allow fastening all the elements of the holder device through M-6 polyethylene screws while clamping the microchip in place. In the center of this plate, there is an elongated hole aligned with the microchip separation channel that allows the excitation light to impinge directly on the microchip. This hole prevents the absorption of the excitation light by the PMMA of the bottom plate. This microchip holder allows the easy and automatic handling of the chip, which improves separation reproducibility.

2.2 Power Supply for Microchip Electrophoresis

Voltage is independently applied to each reservoir through four negatives HCN 7E-12 500 (Fug, Rosenheim, Germany) power supplies. They are controlled by LabView 8.2 software and the interfaces NI PXI 8331, MXI-4, and TB2705 (National Instruments, Austin, TX, USA). This electronic ensemble controls eight 3300-2290-046 high voltage reed relays (Switching Technology Gu¨nther, Nuremberg, Germany) through PC817 optocouplers (Sharp Electronics Corporation, Munich, Germany) and SN7545N Darlington transistors (Texas Instruments, Dallas, TX, USA). This system allows the computer to control the voltage applied to and the application time for each reservoir all along the electrophoretic process. The electronic system also allows connecting each reservoir to ground or to leave it floating.

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Fig. 1 Diagram (A) and picture (B) of the microchip holder. (1) Microchip. (2) Top plate. (3) Holes for screws (picture B). (4) Threaded holes for connection of the syringe pump. (5) Platinum electrodes (0.5 mm diameter). (6) Silicone O-rings. (7) Bottom plate. (8) Threaded holes. (9) Elongated hole aligned with the microchip separation channel. A coin of 20 cents of euro was placed in B as a reference for the size of the microchip holder

Electrophoresis of AGP Deglycosylation in SU-8 Microchips

2.3

Fluids Control

29

1. Fluid manipulation is performed by using an Aladdin 1000 syringe pump (WPI, Sarasota, FL, USA) by a forward (pushing) or backward (aspirating) flow rate of 2600 μL/h (see Note 2). 2. Plastic syringes with Luer-lock fittings are used with the syringe pump. The syringe is connected to the microchip holder through Teflon tubing with 0.5 mm i.d. - 1/1600 o.d. (Symta, Madrid, Spain). The tubing is connected to the syringe using a JR-070123 polypropylene female Luer 1/400 -28 male adaptor, a JR-060 polyamide 1/400 –28 union, and a JR-201580BK polypropylene nut for flanged 1/800 , 1/400 -28 connection, all of them from Vici (Schenkon, Switzerland). The other end of the tubing is connected to the chip holder using a Vici JR-55060 polyphenylene sulfide flanged 1/1600 , 1/400 -28 nut. Nitrile O-rings (5
1 mm) (Grema) were used in flanged fittings. It is particularly critical to achieve a good tightness between the Teflon tubing and the microchip holder (see Note 3).

2.4

LIF Detection

1. An AE31 epifluorescence microscope is used for fluorescence detection with a 20
objective (Motic, Xiamen, China). The microchip holder is placed on the microscope stage. 2. The fluorescence excitation source is a MLM-CDR Excelsior DPSS 532 nm laser (Spectra-Physics, Irvine, CA, USA) with 20 mW nominal power output. 3. The laser is mounted into the microscope as follows. The laser is placed on an aluminum platform with an M-460 3-axis micrometric translation linear stages (Newport, Irvine, CA, USA) which allows its alignment. The laser beam and the light path of the microscope are isolated from the ambience light by a black polyvinyl chloride flexible light shield. This flexible shield is connected to the microscope through a laboratory-designed aluminum flange. The other end of the shield is connected to the laser case through another laboratory-designed aluminum adaptor, which allows creating a light-tight enceinte for laser illumination. Aluminum pieces are anodized in black to avoid any laser light reflection problems (see Note 4). 4. A neutral density filter (optical density value 1 OD from Spectra-Physics) was used to decrease the power of the laser beam at the detection point (power 1.5 mW). It is installed in a filter-holder just at the exit of the laser beam (see Note 5). 5. A Motic MG-1 filter cube consisting of an excitation filter (λexc ¼ 535/25 nm), a dichroic mirror which reflects light below 560 nm, and an emission filter (λem ¼ 600/40 nm) are used. 6. Fluorescence emitted by the sample is collected through the microscope objective and directed to a CMOS Orca Flash 2.8

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camera (Hamamatsu, Hamamatsu City, Japan). Fluorescence signal was recorded by the camera software (HCImage Live) in a Precision T3500 computer (Dell Inc., Round Rock, TX, USA). The software Origin Pro 7.0 (OriginLab, Northampton, MA, USA) was used for signal treatment and representation. 2.5 Working Solutions

All the solutions are made using ultrapure water and analytical grade reagents and (with the exception of samples) filtered with OlimPeak nylon syringe filters (Teknokroma, Barcelona, Spain), 0.2 μm pore size and 25 mm diameter. 1. Separation buffer: 5 mM tetraborate, pH 8.5, 0.1% (w/v) SDS, 10% (w/v) dextran, 10% (v/v) EOTrol LN. Prepare a 5 mM sodium tetraborate (from now on tetraborate) buffer solution pH 8.5, and add the necessary amount of SDS to prepare a 0.1% (w/v) solution of this surfactant. Add to the previous solution the necessary amount of sieving polymer, dextran (Mw 425–575 kDa, Sigma-Aldrich, St. Louis, MO, USA), to obtain a 10% (w/v) concentration. Finally, add the coating agent EOTrol LN commercial solution (Target Discovery, Palo Alto, CA, USA) to a final concentration of 10% (v/v) (see Note 6). 2. 5 mM tetraborate/0.1% SDS: Sample solutions for electrophoretic separations are adequately diluted in 5 mM tetraborate buffer solution, pH 8.5, with 0.1% (w/v) SDS. 3. Denaturing reagent: Mix a solution containing octyl-β-D-glucopyranoside (Sigma-Aldrich) (100 mg octyl-β-D-glucopyranoside solved in 4 mL of water) and a solution containing 35 μL 2-mercaptoethanol diluted with 0.965 mL of water. 4. PNGase F solution: Add 100 μL water to a vial of 50 units of PNGase F (P7367, Sigma-Aldrich) to make a 500 units/mL solution. One Sigma unit of PNGase F activity is equal to 1 IUB milliunit. 5. 4% SDS solution: Dissolve SDS in water to 4% (w/v) final concentration. 6. Proteins used for the molecular weight standard curve: α-lactalbumin (LA, Mw ¼ 14.2 kDa), β-lactoglobulin A (LG, Mw ¼ 18.4 kDa), and bovine serum albumin (BSA, Mw ¼ 66 KDa), all from Sigma-Aldrich. Aqueous solutions were at 102 M for LA and LG and at 103 M for BSA.

2.6 Equipment and Devices for Sample Treatment

1. Concentrator: Ipwisch, UK.

MiVac

Duo,

from

Barnstead/Genevac,

2. Centrifuge: Biofuge Stratos, from Heraeus Instruments, Hanau, Germany. 3. Ultracentrifugation devices: Nanosep 10 k Omega, from Pall Life Sciences, Port Washington, NY, USA.

Electrophoresis of AGP Deglycosylation in SU-8 Microchips

2.7 AGP Mass Spectrometry Analysis

3

31

A Voyager-DE PRO equipment (Applied Biosystems, Foster, CA, USA) is employed for matrix assisted laser desorption ionization time of flight mass spectrometry (MALDI-TOF-MS).

Methods

3.1 AGP Deglycosylation Protocol

The procedure to deglycosylate AGP is performed in three steps following a published method [11] with modifications: 1. AGP denaturation. Mix 20 μL of 8 mg/mL AGP solution with 8 μL of denaturing reagent. Heat the mixture at 100  C for 10 min. 2. Rupture of chemical bonds between glycans and the peptidic chain by using PNGase F. Cool the denatured AGP solution (indicated in 1) to room temperature. After that, mix this solution with 17 μL of 25 mM ammonium bicarbonate and 8 μL of PNGase F solution (ratio 1 nmol AGP/1 unit PNGase F). Heat the mixture at 37  C for 24 h. 3. Purification of deglycosylated AGP by removing glycans from the solution. Freeze the solution indicated in 2 at 20  C for 30 min to stop the enzymatic reaction. Add 1 mL of ethanol and keep the mixture in an ice bath for 30 min. Concentrate the sample solution to 200 μL using a concentrator for 30 min. Centrifuge the mixture at 8000
g for 10 min to precipitate the deglycosylated protein. Remove the supernatant containing mainly the glycans. Finally, wash the precipitate of deglycosylated AGP as follows: resuspend the precipitate in 150 μL of ethanol and keep it in an ice bath for 30 min, centrifuge at 8000
g for 10 min, remove the supernatant, and evaporate the precipitate to dryness using the concentrator.

3.2 AGP Mass Spectrometry Analysis

MALDI-TOF-MS is employed to check the AGP deglycosylation process by the following procedure: 1. Solve the deglycosylated or the glycosylated AGP in ultrapure water to a concentration in the range 10–20 μM. Note that this solution of glycosylated AGP (obtained by using the procedure indicated in Subheading 3.1 but excluding the PNGase F treatment) is also analyzed by MALDI-TOF-MS as control. 2. Mix the AGP sample (either intact or deglycosylated) with MALDI matrix (1/5 v/v mixture) (see Note 7) and apply 1 μL on the MALDI target. Let the spot on the MALDI target to dry at room temperature. 3. Analyze the AGP sample (either glycosylated or deglycosylated) in linear positive mode using the conditions indicated in the caption of Fig. 2. The MS spectrum gives rise to a double main peak corresponding to a molecular weight slightly lower

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Fig. 2 MALDI-TOF mass spectra of glycosylated and deglycosylated AGP, both obtained in the same conditions. Sinapinic acid at 10 mg/mL in 0.3% trifluoroacetic acid (v/v)/acetonitrile (70:30, v/v) was used as matrix. Samples (10–20 μM) were mixed with the matrix at a ratio of approximately 1:5 and 1 μL of this solution was spotted onto a flat stainless-steel sample plate and dried in air. Instrument parameters: acceleration voltage, 25 kV; grid voltage, 90%; guide wire voltage, 0.01%; delay time, 350 ns in the linear positive ion mode

than 22 kDa. This mass is about 40% lower than the one of glycosylated AGP, whose molecular weight was around 35 kDa according to the MALDI-TOF-MS results. This molecular weight difference matches with the reported data in the literature [12], which gives a content of 45% of the molecular weight in N-glycans for AGP. These results confirm the efficiency of the deglycosylation protocol described in Subheading 3.1. 3.3 Protein Derivatization with Chromeo P540

In order to detect proteins with good sensitivity in microchip separations they should be derivatized to render them fluorescent. We use Chromeo P540 as derivatizing agent as follows: 1. Denaturate AGP (both glycosylated and deglycosylated). Take 20 μL of 0.01 M AGP in water and mix it with 20 μL of SDS aqueous solution and heat it at 100  C for 5 min. 2. Remove the excess of SDS from the protein solution. The solution media is replaced with 5 mM tetraborate buffer solution (pH ¼ 8.5) by using ultracentrifugation devices with 10 kDa molecular weight cutoff membrane. The procedure is as follows: apply the protein on the membrane and centrifuge it for 7 min at 14,000
g, wash it with tetraborate buffer solution two times: the first (50 μL) for 7 min at 14,000
g,

Electrophoresis of AGP Deglycosylation in SU-8 Microchips

33

and the second (50 μL) for 3 min at the same g-force. Finally, recover the protein with a micropipette (see Note 8). 3. Add the protein recovered (step 2) into a 0.1 mg Chromeo vial (see Note 9). Add 500 μL of 5 mM tetraborate buffer pH ¼ 8.5 (at this point the AGP concentration is around 10 mg/mL). Stir this mixture softly at room temperature for 1 h protected from light. 3.4 EOF Control and EOF Measurement

One of the key points in MGE is to suppress or minimize the EOF because it causes a loss of separation efficiency. Electroosmotic mobility in SU-8 microchips is 1.8
104 cm²/Vs (at pH 8.5) [10], almost one order of magnitude smaller than the one in fused silica capillaries. It has been demonstrated that EOTrol is a good coating agent for EOF control in SU-8 microchips [10]. In our case, minimum values of the electroosmotic mobility (4.1
105 cm²/Vs) are obtained using solutions with EOTrol LN concentrations 10% (v/v). The optimization of EOTrol concentration was carried out by measuring the EOF according to the following procedure [13]. 1. Prepare a 1 μM rhodamine B solution (neutral marker) in 5 mM tetraborate buffer (pH ¼ 8.5) with different concentrations of EOTrol LN. 2. Inject this solution in the microchip (frontal electrophoresis) by applying 625 V at the sample reservoir (SR) while the waste reservoir (W) is grounded (see Fig. 3). 3. Measure the rhodamine B migration time at half height of the front and calculate the value of the electroosmotic mobility.

3.5 Molecular Weight Standard Curve

1. Label LA, LG, and BSA solutions with Chromeo P540 according to Subheading 3.3. 2. Dilute these solutions until a final concentration of 50 μM LA, 50 μM LG, and 10 μM BSA in separation buffer. 3. Analyze these solutions by MGE (see Subheading 3.8) and measure the migration time of each protein (see Subheading 3.6). 4. Build the molecular weight standard curve using the protein molecular weights and their corresponding migration times as data [14].

3.6 Image Processing

Record the videos of fluorescence signal at the detection point by the camera software and process them as follows: 1. Take any video frame in which fluorescence light occurs. 2. Choose a calculation area (it has to be the same in all the experiments) and set it in the middle of the channel (see Note 10). Process the video with the software. Generate a

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Marı´a del Mar Barrios-Romero et al.

Fig. 3 Schematic representation of the microchip with reservoirs. Buffer reservoir (BR), sample reservoir (SR), sample waste reservoir (SW), and buffer waste reservoir (W). In the scheme the reservoirs are labeled according to their use

spreadsheet with the average fluorescence intensity of each frame versus the time of the frame. 3. Export these data to a data analysis software for signal treatment and representation. 3.7 Microchip Conditioning

1. In all the cases (brand new chip, at the beginning of each working day, or before each injection) rinse the chip successively with water (10 min), 0.1 M NaOH (10 min), and water (10 min) by pushing with the syringe pump (2600 μL/h) connected to the reservoir W of the chip (Fig. 3). Then, fill the four channels of the chip with separation buffer by aspirating (20 min at 2600 μL/h) from W reservoir and, at the same time, replenish the other reservoirs BR, SW, and SR with separation buffer with the aid of an automatic pipette as the buffer is flowing into the channels. In this way bubble formation in the channels is avoided. 2. Replace the buffer in the SR reservoir with the sample using an automatic pipette. After that, fill the injection channel (that one going from SR to SW) with sample by aspirating from the SW reservoir for 2 min (see Note 11).

3.8 Electrophoretic Analysis

Electrophoretic analysis of AGP is carried out in three steps—load, injection, and separation. Load step is carried out to ensure that the cross of the chip does not contain any amount of the sample that could reach there by diffusion during previous operations. Injection is performed in mode gated for 2 s (see Note 12). In the separation step the detection point is placed at 3 cm from the cross. Table 1 indicates the potential applied to each reservoir in these steps. AGP samples, labeled with Chromeo P540, are analyzed by MGE. The result of the separation of the sample of AGP deglycosylated is shown in Fig. 4A, which shows a major peak (1, at 187 s) followed by some minor peaks (3, 4, and 5), which are partially

Electrophoresis of AGP Deglycosylation in SU-8 Microchips

35

Table 1 Voltage sequences applied on each reservoir to perform loading, gated injection, and separation Reservoirs Steps

Sample (SR)

Buffer (BR)

Sample waste (SW)

Waste (W)

Load

625 V

750 V

Grounded

Grounded

Injection (2 s)

750 V

Floating

Floating

Grounded

Separation

625 V

750 V

Grounded

Grounded

overlapped. Probably, these minor peaks were generated during the deglycosylation process and they could be due to partially deglycosylated fractions of AGP. In fact, as mentioned in Subheading 3.2, in the MS spectrum (Fig. 2) minor broad peaks in the m/z range of 23,000–30,000 are also observed. According to these results, it can be considered that deglycosylation by PNGase F, although it is effective, did not render a 100% yield in these conditions. On the contrary, the electropherogram of glycosylated AGP (Fig. 4B) only presents a single broad peak (centered at migration time of 300 s) for this protein. When the sample of the deglycosylated AGP is spiked with glycosylated AGP, an electropherogram (Fig. 4c) containing two major peaks (peaks 1 and 2) at migration times of 175 and 300 s and the three minor peaks observed in the deglycosylated AGP sample are obtained. These two major peaks could be tentatively assigned to the deglycosylated and glycosylated forms of the AGP, respectively. To confirm these results a molecular weight standard curve was generated (see Subheading 3.5). The fit gives rise to a straight line, which can be described by the equation Log M W ¼ 5:92  281:12 ð1=t m Þ with good correlation coefficient (r > 0.999). Using this equation, the molecular weight corresponding to peak 1 (Fig. 4A) is 26 kDa (average of migration time for peak 1, tm  SD ¼ 187  24 s, n ¼ 3). This value of the Mw agrees relatively well to the one obtained from the MS spectra, around 22 kDa, and with the value 21.5–21.6 kDa provided by the literature [11]. Additionally, the molecular weight value for glycosylated AGP calculated using the same equation is 99 kDa (tm  SD ¼ 305  34 s, n ¼ 3). This value is higher than the one obtained by MALDI-TOF-MS, 35 kDa, and the value 41–43 kDa given in the literature [12]. It is well known that the association of SDS with the glycans is prevented in glycoproteins [15]. This is particularly critical for those glycoproteins with high content in carbohydrates (AGP with 45% (w/w) glycosylation can be considered as a heavily glycosylated protein). Glycoproteins, therefore,

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Fig. 4 Electropherograms of (A) 10 μM deglycosylated AGP, (B) 5 μM glycosylated AGP, (C) 10 μM deglycosylated AGP spiked with 5 μM glycosylated AGP. Both glycosylated and deglycosylated AGP derivatized with Chromeo P540. Separation buffer: 5 mM tetraborate buffer (pH 8.5), 0.1% (w/v) SDS, 10% (w/v) dextran, and 10% (v/v) EOTrol LN. Injection type: gated, 2 s. Voltage applied for separation: 750 V BR, 625 V SR, rest of the reservoirs were grounded. Separation channel length: 3 cm. Excitation source: 532 nm laser, power 1.5 mW at the detection point. Peak assignment: (1) deglycosylated AGP, (2) glycosylated AGP, and (3), (4), and (5) minor compounds associated to partially deglycosylated AGP

due to low binding with SDS have a small charge-to-mass ratio and therefore they migrate slower than non-glycosylated proteins in SDS-CGE. In fact, it has been reported [16] that employing a commercial system (2100 Bioanalyzer microchip gel electrophoresis instrument), which uses glass microchips and where the proteins are separated, as in our case, in a sieving matrix of an entangled linear polymer in an SDS containing buffer, AGP shows two major peaks at Mw of 98 and 116 kDa, which are considerably higher in Mw than the values of 32 kDa and 47 kDa obtained by these authors using MALDI-MS and SDS-PAGE, respectively. Relatively good migration repeatability can be obtained with the method presented here. The RSD values (n ¼ 3) for migration time for inter-day analysis (in the same chip) were 13% (tm  SD ¼ 187  24 s) for deglycosylated AGP and 12% (tm  SD ¼ 305  34 s) for glycosylated AGP. However, RSD values (n ¼ 3) obtained for peak area, 61% for deglycosylated and 55% for glycosylated APG (average peak area  SD ¼ 4000  2374 counts and 20,000  11,747 counts, respectively) were rather limited.

Electrophoresis of AGP Deglycosylation in SU-8 Microchips

4

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Notes 1. In this chapter, we follow the denomination used by the European Medicine Agency for this type of nonstandard pharmaceutical compounds. Biopharmaceuticals refer to pharmaceutical drugs produced through biotechnological processes using molecular biology methods. Biosimilars indicate biopharmaceuticals containing a slightly different version of the active pharmaceutical ingredient previously registered as reference biological medical product. Biobetters are biopharmaceuticals that have been structurally and/or functionally altered to achieve an improved or different clinical performance, compared to an approved reference product. A Critical Quality Attribute of a pharmaceutical product is a physical, chemical, biological, or microbiological property or characteristic that should be within an appropriate limit, range, or distribution to ensure the desired product quality [17]. 2. Higher flow rates could cause fluid overpressure in the chip holder, leakages from the microchip reservoirs and intercommunication among them, which can produce electrical shortcircuits when voltage is applied. 3. One of the most critical steps during microchip manipulation is to screw/unscrew the Teflon tubing to/from the microchip holder. During this step, dust from the laboratory environment can be introduced inside the microchip channels causing clogging problems. To minimize this effect, the syringe was not directly connected to the reservoir, but to a three-way plastic valve (Sebdal, Caceres, Spain) with a short Teflon tubing to minimize dead volume. In this way, it is not necessary to screw/ unscrew the connector from the microchip holder every time a different liquid is introduced into the microchip. 4. Since the commercial microscope has a 100 W mercury lamp as standard illumination source, it was provided by the manufacturer with a fluorescence attachment (lens condenser and condenser and field apertures). We remove it before installing the laser illuminator. 5. The use of higher power of the laser (3–12 mW) at the detection point can cause instability of the baseline, likely due to photo-degradation of the SU-8 material. 6. EOTrol LN is a coating agent that decreases the EOF in fused silica capillaries. The ability of EOTrol LN to control the EOF and to prevent wall protein adsorption in SU-8 channels has also been demonstrated [10]. 7. The MALDI matrix solution is a mixture of 10 mg/mL of sinapinic acid in 0.3% trifluoroacetic (TFA) (v/v)/acetonitrile (70/30, v/v).

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8. This procedure for SDS removal was done to prevent any possible interference of SDS in the labeling reaction. 9. Chromeo P540 handling. This reagent, supplied by Active Motif (Tegernheim, Germany), is a pyrilium derivative, which is used to label proteins through the amino groups. The derivatization reaction is carried out by adding a solution of the denaturated AGP on 0.1 mg of a dried aliquot of Chromeo. These aliquots are prepared by dissolving 1 mg of Chromeo, as received, in 100 μL methanol. Aliquots of 10 μL of this solution are transferred to 500 μL microcentrifuge tubes. The solvent is removed using a miVAC Duo centrifugal evaporator. The aliquots of dried Chromeo are stored at 20  C and protected from light. In aqueous solution Chromeo P540 has a violet color, and when it reacts with terminal lysine and amine groups of the proteins, the solution turns to red because the conjugated form has a wavelength shift of 54 nm. Conjugated Chromeo P540 has an absorption maximum at 533 nm and an emission maximum at 627 nm. The free dye has an absorption maximum at 587 nm, though its fluorescence quantum efficiency is below 1%. As a consequence, it is not necessary to remove the excess of dye from the sample after the labeling reaction. 10. The software allows the drawing of any geometrical figure of any size. We have selected always a circle with a diameter of 25 μm (half size of the channel). 11. This step allows removing the buffer containing sieving polymer from the injection channel (SR - SW channel) and replace it with sample, which does not contain dextran. By doing so, a rough viscosity change is created at the interface between the injection cross and the entrance of the separation channel, so that the sample is pre-concentrated by stacking. 12. Gated injections are very precise and can be run in a continuous sampling mode. Furthermore, the injected amount is controlled by the injection time so that it can be easily optimized [18].

Acknowledgments The authors acknowledge the Spanish Ministries of Science and Innovation and of Economy, Industry, and Competitiveness (grants PSS-010000-2008-6 and CTQ2013-43236-R, respectively) and CSIC (joint project 2009JP003 with the Japanese Society for Promotion of Science) for financial support. M.M.B-R thanks the CSIC for a JAE-Pre contract and A.G.C. acknowledges CSIC for JAE-Doc contract. These contracts are co-financed by the European Union under the European Social Fund (ESF).

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References 1. Walsh G (2014) Biopharmaceutical benchmarks 2014. Nat Biotech 32(10):992–1000. https://doi.org/10.1038/nbt.3040 2. Malgorzata KB (2017) Progress in biopharmaceutical development. Biotechnol Appl Biochem 65:306–322. https://doi.org/10. 1002/bab.1617 3. Jefferis R (2017) Recombinant proteins and monoclonal antibodies. Adv Biochem Eng Biotechnol. https://doi.org/10.1007/10_2017_ 32 4. Little MJ, Paquette DM, Roos PK (2006) Electrophoresis of pharmaceutical proteins: status quo. Electrophoresis 27 (12):2477–2485. https://doi.org/10.1002/ elps.200500951 5. Zhang YJ, An HJ (2017) Technologies and strategies for bioanalysis of biopharmaceuticals. Bioanalysis 9(18):1343–1347. https://doi. org/10.4155/bio-2017-4981 6. O’Flaherty R, Trbojevic-Akmacic I, Greville G, Rudd PM, Lauc G (2018) The sweet spot for biologics: recent advances in characterization of biotherapeutic glycoproteins. Expert Rev Proteomics 15(1):13–29. https://doi.org/10. 1080/14789450.2018.1404907 7. Creamer JS, Oborny NJ, Lunte SM (2014) Recent advances in the analysis of therapeutic proteins by capillary and microchip electrophoresis. Anal Methods 6(15):5427–5449. https://doi.org/10.1039/c4ay00447g 8. Szekrenyes A, Roth U, Kerekgyarto M, Szekely A, Kurucz I, Kowalewski K, Guttman A (2012) High-throughput analysis of therapeutic and diagnostic monoclonal antibodies by multicapillary SDS gel electrophoresis in conjunction with covalent fluorescent labeling. Anal Bioanal Chem 404(5):1485–1494. https://doi.org/10.1007/s00216-012-62132 9. Smith MT, Zhang S, Adams T, DiPaolo B, Dally J (2017) Establishment and validation of a microfluidic capillary gel electrophoresis platform method for purity analysis of therapeutic monoclonal antibodies. Electrophoresis 38(9–10):1353–1365. https://doi.org/10. 1002/elps.201600519 10. Barrios-Romero MM, Creville´n AG, DiezMasa JC (2013) Development of an SDS-gel electrophoresis method on SU-8 microchips for protein separation with LIF detection: application to the analysis of whey proteins.

J Sep Sci 36(15):2530–2537. https://doi. org/10.1002/jssc.201300275 11. Ongay S, Neususs C (2010) Isoform differentiation of intact AGP from human serum by capillary electrophoresis - mass spectrometry. Anal Bioanal Chem 398(2):845–855. https:// doi.org/10.1007/s00216-010-3948-5 12. Fournier T, Medjoubi-N N, Porquet D (2000) Alpha-1-acid glycoprotein. Biochim Biophys Acta 1482(1–2):157–171. https://doi.org/ 10.1016/S0167-4838(00)00153-9 13. Shakalisava Y, Poitevin M, Viovy JL, Descroix S (2009) Versatile method for electroosmotic flow measurements in microchip electrophoresis. J Chromatogr A 1216(6):1030–1033. https://doi.org/10.1016/j.chroma.2008.12. 029 14. Guttman A, Nolan J (1994) Comparison of the separation of proteins by sodium dodecylsulfate slab gel-electrophoresis and capillary sodium dodecyl-sulfate gel-electrophoresis. Anal Biochem 221(2):285–289. https://doi. org/10.1006/abio.1994.1413 15. Segrest JP, Jackson RL, Andrews EP, Marchesi VT (1971) Human erythrocyte membrane glycoprotein: a re-evaluation of molecular weight as determined by SDS polyacrylamide gel electrophoresis. Biochem Biophys Res Commun 44(2):390–395. https://doi.org/10.1016/ 0006-291x(71)90612-7 16. Engel N, Weiss VU, Wenz C, Ruefer A, Kratzmeier M, Glueck S, MarchettiDeschmann M, Allmaier G (2015) Challenges of glycoprotein analysis by microchip capillary gel electrophoresis. Electrophoresis 36 (15):1754–1758. https://doi.org/10.1002/ elps.201400510 17. International conference on harmonisation of technical requirements for registration of pharmaceuticals for human use. Pharmaceutical development Q8(R2). Current step 4 version dated August 2009. https://www.ich.org/ fileadmin/Public_Web_Site/ICH_Products/ Guidelines/Quality/Q8_R1/Step4/Q8_R2_ Guideline.pdf 18. Lacher NA, Garrison KE, Martin RS, Lunte SM (2001) Microchip capillary electrophoresis/electrochemistry. Electrophoresis 22 (12):2526–2536. https://doi.org/10.1002/ 1522-2683(200107)22:123.0.CO;2-K

Chapter 4 Glycoform Analysis of Alpha1-Acid Glycoprotein by Capillary Electrophoresis Using Electrophoretic Injection Chenhua Zhang, William Clarke, and David S. Hage Abstract Human alpha1-acid glycoprotein (AGP) is an acute phase glycoprotein that has a heterogeneous glycosylation pattern. This pattern can change in certain diseases, which has resulted in interest in using AGP glycoforms as potential biomarkers for these diseases. This report describes a method that uses capillary electrophoresis to characterize and analyze AGP glycoforms both in purified samples of AGP and in human serum. This method uses static and dynamic coatings of poly(ethylene oxide) that are applied to a silica capillary for separation of AGP glycoforms in the reversed-polarity mode of CE and in the presence of negligible electroosmotic flow. Electrophoretic injection is performed onto such capillaries by using a stationary stacking interface between the sample and running buffer. In addition, acidic precipitation and desalting are used to allow for the isolation and the analysis of AGP from only 65 μL of serum. Up to eleven AGP glycoform bands can be reproducibly separated by this method, with the difference in migration time between neighboring bands being 12- to almost 60-fold larger than the standard deviation for the migration time of any given band. A limit of detection down to about 2 nM per glycoform band can be obtained by this method for AGP in serum based on absorbance detection and without the need for further sample modification or labeling. Key words Alpha1-acid glycoprotein, Capillary electrophoresis, Capillary modification, Electrophoretic injection, Glycoforms, Sample stacking, Serum

1

Introduction Human alpha1-acid glycoprotein (AGP) is an acute phase glycoprotein that has a single chain of 183 amino acids and two disulfide bonds [1–3]. AGP is a heavily glycosylated protein. The molecular weight of AGP is between 41 and 43 kDa, with about 45% of this mass being due to glycan moieties [1]. As shown in Fig. 1, there are five N-linked glycosylation sites on AGP, which occur at Asn15, Asn38, Asn54, Asn75, and Asn85. A heterogeneous mixture of bi-, tri-, or tetra-antennary glycans can be attached to these sites [1]. AGP has a low isoelectric point (2.8 to 3.8) due to the presence of multiple sialic acids in these glycan chains [1, 2].

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Fig. 1 Examples of complex N-linked glycans that can occur on AGP and the locations of the glycosylation sites on AGP. Abbreviations: GlcNAc N-acetylglucosamine, Man mannose, Gal galactose, Fuc fucose, NeuAc neuraminic acid (or sialic acid). The nomenclature used here for the representation of glycans is based on Ref. [24]

Both the concentration and composition of AGP can change with various diseases and clinical conditions [2, 4–7]. For instance, the normal concentration of AGP in serum ranges from 0.5 to 1.0 mg/mL, or 12 to 24 μM; however, this concentration can increase by up to tenfold in some clinical situations [2]. In addition, alterations in the glycosylation of AGP have been observed. For instance, a decrease in the degree of glycan branching has been reported in patients with infection and systemic lupus erythematosus (SLE) [4]. An increase in α1–3 fucosylation has been noted in pancreatic cancer [5], and an increase in levels of AGP sialylation has been seen in ovarian cancer and lymphoma [6, 7]. These changes have made AGP glycoforms and their related glycosylation patterns of interest as potential biomarkers for these and other disease states [4–7]. Capillary electrophoresis (CE) is one approach that can be used to separate and examine the glycoforms of AGP [6, 8–11]. During the development of these CE methods, various approaches have been proposed to reduce adsorption of AGP onto the capillary wall and to modify electroomostic flow for improved precision and better resolution of the glycoform bands. These approaches have included the use of buffer additives such as putrescine and urea or the use of capillaries that contained a cross-linked coating of dimethylpolysiloxane [6, 8–10]. In this report, an alternative method was employed in which static and dynamic coatings of poly (ethylene oxide) (PEO) were applied to a silica capillary. This coating method was used because it has been shown to allow highly reproducible separations of AGP glycoforms and to provide differences in migration times between neighboring glycoform bands that are 12- to almost 60-fold larger than the standard deviations of the migration times for these bands [11]. In this report, these coated capillaries were combined with electrophoretic injection and

Analysis of AGP by CE and Electrophoretic Injection

43

UV absorbance detection to examine the major glycoform bands of human AGP. The use of absorbance detection alone for AGP tends to provide only modest detection limits in CE due to the short path length of light in the capillary and the capillary’s small loading capacity [12]. For instance, a limit of detection of 0.09–0.38 μM is obtained for the glycoform bands of AGP when absorbance detection is employed with standard hydrodynamic injection [11]. Mass spectrometry is an alternative detection method that has been used with CE, in combination with immunoaffinity extraction, to examine AGP glycoforms in serum; however, the cost of the instrumentation and level of training needed for CE-mass spectrometry has tended to limit its use in routine clinical testing [13–15]. CE with laser-induced fluorescence detection and fluorescent-labeled AGP can also be used to detect AGP glycoforms [16], but this approach adds extra steps and time for sample treatment and modifies the structure of AGP. Electrophoretic injection is a type of electrokinetic injection that can be performed with AGP when there is a negligible amount of electroosmotic flow present in this system [17]. This situation creates an essentially stationary stacking interface between the sample and running buffer during injection. This technique can allow for the sensitive detection of AGP (i.e., down to 0.05–0.2 nM) without the need for AGP derivatization and by using absorbance detection [17]. Sample pretreatment was further needed to use CE with absorbance detection and electrophoretic injection for the analysis of AGP in serum (see overall scheme provided in Fig. 2). Multiple chromatographic steps, such as ion-exchange and dye-ligand chromatography, have been used to isolate AGP from serum or plasma; unfortunately, this approach generally requires a relatively large sample volume [18, 19]. Immunoaffinity extraction with antiAGP immobilized support has been used as well but needs multiple steps for sample loading, elution, and column regeneration [20]. Acidic precipitation has been employed for the pretreatment of AGP from plasma but tends to give a low purity for AGP when compared with immunoaffinity extraction [20, 21]. The method that is described in this current report combined acidic precipitation, desalting, and electrokinetic injection for the selective enrichment of AGP from serum using sample volumes of as little as 65 μL [17].

2

Materials The water used in the following procedures was prepared with a Milli-Q water purification system (EMD Millipore Corporation); however, other types of water purification systems can also be used.

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65 μl of human serum

Acidic precipitaon

Desalng

Electrophorec injecon and CE analysis

Data analysis Fig. 2 General scheme for the sample preparation and CE analysis of glycoform bands for AGP that has been isolated from serum 2.1

Samples

1. Commercial samples of normal human AGP (
99% purity, from pooled human plasma) were obtained from SigmaAldrich. 2. Commercial samples of human serum were also obtained from Sigma-Aldrich, as were made from pooled human male AB plasma of U.S. origin (sterile-filtered). Each donor that was used to prepare this commercial sample had been tested by ELISA and found to be nonreactive for hepatitis B, hepatitis C, and human immunodeficiency virus (see Note 1). 3. The specimens from patients with SLE were de-identified and preexisting serum samples that were supplied by W. Clarke; this work has been determined to be exempt from IRB review by the Johns Hopkins School of Medicine, according to the Code of Federal Regulations 45 CFR46.101 b.

2.2

Equipment

1. A Beckman P/ACE MDQ capillary electrophoresis system was used in this study. Other brands or models of CE systems with equivalent performance can also be used. 2. Spin column desalting column (0.5 mL, 7 kDa cutoff; Thermo Fisher Scientific).

Analysis of AGP by CE and Electrophoretic Injection

2.3 Buffers and Solutions

45

1. 0.5 M Perchloric acid solution, 10 mL: Add 0.429 mL of perchloric acid (70%) into a 20 mL glass vial and add 9.571 mL water, giving a final perchloric acid concentration of 0.5 M (see Note 2). 2. 1 M Sodium hydroxide (NaOH) solution, 100 ml: Add 4.0 g of NaOH into a 150 mL beaker, add 100 mL of water and dissolve the NaOH through stirring or sonication. 3. 1 M Hydrochloric acid (HCl) solution, 100 mL: Add 8.333 mL of 37% (w/w) HCl into a 100 mL volumetric glassware and add water to the mark. 4. 0.2% (w/v) PEO (viscosity average molecular weight, 8,000,000; Sigma-Aldrich) prepared in 0.1 M HCl, 100 mL: Add 0.2 g PEO slowly (see Note 3), with stirring, into 100 mL water that is held in a container within an 80  C water bath. Cover the opening of the flask with an upside-down 100 mL beaker to avoid water evaporation. Stir the solution for 2 h until all of the PEO has dissolved. After this solution has been cooled to room temperature, add 0.833 mL of 37% of HCl to this solution. 5. 20 mM Acetate buffer, pH 4.2, containing 0.05% (w/v) PEO and 0.1% (w/v) Brij 35, 200 mL: Solvent A: 0.05% (w/v) PEO and 0.1% (w/v) Brij 35, 300 mL. Add 0.15 g PEO slowly, and with stirring, into a container holding 300 mL water within an 80  C water bath. Cover the opening of the flask with an upside-down 100 mL beaker to prevent the evaporation of water. Stir this solution for an additional 2 h until all of the PEO has dissolved. After this solution has been cooled to room temperature, add 0.3 g Brij 35 to the solution with stirring. Weigh 0.1196 g of sodium acetate trihydrate into a 200 mL beaker. Add approximately 180 mL of solvent A into this beaker and stir until all the sodium acetate salt has dissolved. Add 178 μL of glacial acetic acid to the solution while stirring. Adjust the pH of this solution to 4.2 by adding small amounts of 1 M NaOH or 1 M HCl and while monitoring the pH of solution (see Note 4). Transfer the final solution into a 200 mL volumetric glassware and add solvent A to fill this flask to the mark (see Note 5).

2.4 Spin Desalting Column

Remove the spin column’s bottom closure and loosen the cap without fully removing it. A 2.0 mL microcentrifuge tube can be used as a collection tube with its cap removed. Place the column in this collection tube and centrifuge at 6600 rpm (2000  g) for 1 min to remove any storage solution that is present in the device. Place a mark on the side of the spin column where the compacted resin is slanted upward; this is to make sure the resin is placed the same way after every wash or pre-equilibration step. Place the spin

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column into a minicentrifuge, with the mark facing outward in all subsequent centrifugation steps. Add 300 μL water (i.e., the solution to be used for desalting) to the top of the resin bed in the spin column. Centrifuge at 6600 rpm for 1 min to remove any water. Repeat the addition of water and this centrifugation process two more times, discarding the water from the collection tube. 2.5 Fused Silica Capillary

3

A fused silica capillary with a length of 72 cm should be cut from capillary tubing with an inner diameter of 50 μm and an outer diameter of 360 μm, as obtained from Polymicro Technologies (Phoenix, AZ, USA) (see Note 6). This cut can be achieved by using a cleaving stone. Remove the outer polyimide coating from the capillary in the desired region of the detection window. This can be done by placing the capillary on top of the U-shaped opening of an open-ended ¼ in wrench, with this opening being placed at least 11 cm away from the near end of the capillary. A lighter can then be used to burn off the polyimide coating from the capillary over a length of approximately 5 mm (see Note 7). This area should then be gently cleaned by using a Kimwipe soaked with ethanol. To install the capillary cartridge for the P/ACE MDQ CE system, insert the long side of the capillary into the outlet side of cartridge. Carefully pull the capillary from the inlet side until the detection window appears in the cartridge window. Select the required aperture clip (size, 100  800 μm) (see Note 8), replace from the back of the cartridge, and use an insertion tool to seat the O-ring into the front of the aperture (see Note 9). Place seal retainer clips on both sides of the capillary and snap these retainer clips into position. Place the cartridge face down against the capillary length template. Use a cleaving stone to cut and remove the marked portion of the capillary. Use the rough side of the cleaving stone to burnish the ends of the capillary until these ends are smooth (see Note 10). After this step, the final capillary will have an effective length of 50 cm and a total length of 60.2 cm. Mount the cartridge with the prepared fused silica capillary into the CE system. To clean the capillary and activate its interior silanol groups, rinse the capillary with 1 M NaOH for 30 min at a pressure of 50 psi, and clean the capillary with water for 10 min at the same pressure [11, 17].

Methods

3.1 Acid Precipitation and Desalting

1. Acid precipitation: Combine a 65 μL portion of serum and 130 μL of 0.5 M perchloric acid in a 0.6 mL microcentrifuge tube and vortex mix for 20 s. Place the resulting mixture into a minicentrifuge and allow it to spin at 6600 rpm for 10 min at room temperature. This step precipitates major proteins from serum while allowing AGP to be recovered in the supernatant [21].

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47

2. Sample desalting: Place a 130 μL portion of the supernatant from the acid precipitation step into a spin desalting column. Place the spin desalting column into a 2.0 mL microcentrifuge tube with the cap removed as a collection tube and spin the column for 2 min at 6600 rpm. This desalting step helps to improve both the loading capacity and extent of zone broadening in the final CE method through its effect on the relative resistivity (or conductivity) for the sample solution versus the running buffer, as is illustrated in Fig. 3 [17]. The use of both acid precipitation and desalting for sample preparation has also been found to result in good recoveries for AGP glycoforms from serum. For instance, recoveries between 72.3 and 80.9% have been measured when using this pretreatment method to look at AGP glycoform bands from human serum by CE (see Table 1) [17]. 3. Sample dilution: Add the filtrate from the previous step to a 20 ml glass vial that also contains 5 mL of water (see Note 11). This step is used to reduce the concentration of AGP down to around 1 μg/mL and generates a relatively large volume of a working sample solution that can be employed for multiple injections, or reanalysis as needed.

Fig. 3 Electropherograms obtained for AGP glycoforms when using human serum that has been processed by desalting or that was used with no sample pretreatment. This figure is modified with permission from Ref. [17]

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Table 1 Characteristics of an electrophoretic injection CE method for AGP glycoform analysis in serum using acid precipitation and desalting for sample pretreatment Glycoform band

Limit of detection (nM)a

Precision of migration timeb

Precision of peak areab

Recoveryc

3

2.5

0.08%

0.86%

72.3%

4

8.2

0.11%

0.94%

80.9%

5

11.4

0.12%

0.71%

80.4%

6

8.5

0.12%

0.52%

80.5%

7

4.8

0.12%

0.34%

80.3%

8

2.4

0.13%

0.86%

78.2%

9

2.1

0.12%

1.18%

73.6%

Adapted with permission from Ref. [17] a The detection limit was calculated by using a signal-to-noise ratio of three, as based on the standard deviation of the intercept and the slope from a calibration curve for this assay b The relative standard deviations listed in these columns are based on measurements made in triplicate (n ¼ 3) for normal serum that was spiked with 2.5 g/L AGP. The ranges shown represent 1 relative standard deviation c The recovery was calculated by dividing the slopes of the corresponding calibration curves that were obtained before and after sample pretreatment with normal serum that had been spiked with a known amount of AGP

3.2 Capillary Electrophoresis and Electrophoretic Injection

1. CE operating conditions: The following conditions are used for the final method that is described in this report. The capillary temperature should be maintained at 25  C during the separation, and the applied voltage for the separation should be set at 30 kV (see Note 12). AGP glycoforms can be detected by using their absorbance at 200 nm [11, 17]. 2. Preparation of static and dynamic coatings of PEO: The capillary is first cleaned by using a 5 min rinse with 1 M NaOH, followed by a 3 min rinse with water. The coating is applied by rinsing the capillary for 5 min with 1 M HCl and then rinsing for 5 min with a 0.2% (w/v) PEO solution that contains 0.1 M HCl. The capillary is then rinsed for 5 min with a running buffer that consists of pH 4.2, 20 mM acetate buffer containing 0.05% (w/v) PEO and 0.1% (w/v) Brij 35. All of these rinse steps should be performed at an applied pressure of 50 psi, and this rinsing procedure should be repeated before each CE separation. The effects of changing the buffer pH, capillary coating material or concentration of Brij 35 on the separation of AGP glycoforms are discussed under Note 13. 3. Electrophoretic injection: Add 1.2 mL of the sample to a buffer vial. Carry out electrophoretic injection by using a voltage of 5 kV that is applied for 5 min (see Note 14). This electrophoretic injection step enables selective injection of AGP from a sample that has undergone the pretreatment steps described under Subheading 3.1. The selectivity of

Analysis of AGP by CE and Electrophoretic Injection

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Fig. 4 Relationship between (a) the total peak area for AGP glycoform bands and the time used for electrophoretic injection, and (b) the width for the ninth glycoform band for AGP versus the time used for electrophoretic injection. The error bars represent a range of 1 S.D. (n ¼ 3) in (a) and a range of 1 S.D. of the mean (n ¼ 3) in (b). The plots used in this figure were adapted with permission from Ref. [17]

this approach is aided by the fact that AGP is one of few proteins that can have a negative charge at an acidic pH around 4 [22]. The use of an injection time longer than 5 min can result in a higher loading capacity and signal intensity for AGP, as illustrated in Fig. 4a; however, a lower resolution will also be observed under these conditions due to the zone broadening that occurs during injection, as shown in Fig. 4b. In contrast, the use of an injection time that is shorter than 5 min will result in smaller signal intensity with similar broadening to what is seen at an injection time of 5 min. Based on these trends, an injection time of 5 min has been determined to provide a good compromise between detection limits, overall analysis times, and peak resolution [17]. 4. Capillary cleaning and storage: After a single separation or multiple separations that are carried out as a sequence, the capillary should be rinsed with 1 M NaOH at 50 psi for 15 min and then rinsed with water at 50 psi for 10 min. The capillary should be stored in water when it is not in use (see Note 15). 3.3

Data Analysis

Most of the following procedures for data analysis are specific for the type of CE system and analysis software that were used in this study. Equivalent procedures can be employed with alternative CE systems or software. 1. An electropherogram that has been obtained by the CE system can be exported as a .cdf file for analysis of its peaks by a program such as Peakfit version 4.12.

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2. Use the “section” option to select a specific region of the electropherogram for closer analysis (i.e., a suggested range of 10–25 min for most applications of this CE method). 3. To carry out a baseline correction using Peakfit, select the “progressive line” option. This option can be used to correct for artifacts such as baseline drifting. 4. When using Peakfit for data analysis, select the options “Chromatography” and “Exponentially modified Gaussian or EMG” for peak type. Select the options “Vary widths,” “Vary shape,” and “Allow negative” for auto scan. Click on the electropherogram to add multiple simulated peaks and adjust their shape to give an approximate fit to the electropherogram. 5. When using Peakfit, select and click the button that says “Fast peak fit with numerical update” until the best fit is obtained. Next, select and click the options “Review fit” and “Numeric” to acquire the measured parameters for each individual peak or band. To obtain more parameters for the simulated peaks, select “Options” to add in additional factors such as “Chromatography analysis.” 3.4 Analysis of AGP Glycoform Bands in Serum Samples

1. The electropherograms in Fig. 5a show some typical results that were obtained by the overall sample pretreatment and CE method for normal and SLE serum samples. The migration times and peak areas of the individual glycoform bands in these electropherograms were obtained by following the data analysis procedures that were described under Subheading 3.3. 2. The total peak area for AGP in the electropherogram can be obtained in this particular method by using a summation of the peak areas for the individual glycoform bands (see Note 16). A correction of the peak areas based on their apparent mobilities, as is common in other CE methods, is not needed in this approach with negligible electroosmosis because the bias of peak area in detection due to differential mobility has already been corrected during the electrophoretic injection process [17]. The relative peak area due to each individual band can be found by dividing the separate band/peak areas by the total peak band/peak for AGP. 3. The glycoform band distribution pattern for AGP can be visualized by making a plot of the percentage peak area for each glycoform band (placed on the y-axis) versus the migration time of each individual band (on the x-axis) [17]. Examples of such plots are shown in Fig. 5b. 4. This sample preparation and CE method has been found to have a limit of detection ranging from 2.1 to 11.3 nM for the major glycoform bands of AGP in human serum [17]. These limits of detection are approximately 3000-fold lower than the

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Fig. 5 (a) Electropherograms obtained for AGP from SLE serum or pooled normal serum, in which the mobility marker (Lucifer yellow CH, LyCH) was introduced by hydrodynamic injection after sample injection. (b) Glycoform patterns obtained for AGP from SLE serum or pooled normal serum by this CE method; the error bars shown for the migration times and % peak areas represent 1 S.D. of the mean (n ¼ 3). These plots are reproduced with permission from Ref. [17]

concentration of AGP that is normally observed in human serum and allow this method to be used with only 65 μl of serum without the need for any sample labeling [17]. 5. The precision of the migration times for the AGP glycoform bands is between 0.08 and 0.13%; the precisions for the peak areas are between 0.34 and 1.18% when using an internal standard to correct the peak areas for their variations in loading capacity during the injection process (see Table 1). This reproducible separation allows the direct use of migration times to identify and distinguish between AGP glycoform bands in this method [17].

4

Notes 1. Any work with human samples such as serum that may contain blood-borne pathogens should be handled according to appropriate biosafety protocols and in BSL-2 facilities. 2. Use appropriate chemical hazard precautions and facilities when handling perchloric acid [23]. 3. This step is to avoid the formation of aggregates of PEO, which will be difficult to dissolve. 4. Be sure to calibrate the pH meter before use, as the separation of AGP glycoforms by CE can be sensitive to a small change in the pH of the running buffer [11].

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5. Do not filter this buffer solution because the PEO may not be able to pass through the filter (e.g., one using a 0.2 μm pore size membrane). 6. The protocol described in this report has been developed for these particular capillary dimensions. The use of other capillary dimensions will affect the resolution, overall analysis time, and peak broadening that are obtained in the final method, and will require further optimization of parameters such as the applied voltage and capillary pretreatment/rinsing protocol, among other factors. 7. Extra care needs to be taken when handling a capillary that contains a region without a polyimide coating because the exposed region of this capillary can be quite brittle. 8. Make sure the O-ring has been removed from the aperture clip before installing the aperture clip. A preinstalled O-ring inside the aperture clip can crack the capillary when you are installing the aperture clip. 9. Make sure the O-ring lays flat after its installation. An improper orientation of the O-ring will block the light path and create resistance when connecting the optical fiber with the aperture clip. 10. A magnifying lens can be used to inspect the end of capillary. A rough end on the capillary will result in a tailing effect for all the AGP glycoform bands in the final method and should be avoided. 11. The remaining contents of the serum sample are diluted by 115.4-fold as a result of the overall pretreatment procedure. 12. An applied voltage of 30 kV is the highest setting that is available in the CE system that was used in this work. A lower applied voltage can be used but will generate a smaller electric field, leading to reduced migration speeds and longer analysis times. The peaks or bands will also appear broader in the time domain under lower voltage conditions. 13. Several other pH conditions have been evaluated for use with this method. When using a pH range of 3.8 to 5.0 for the running buffer, it was found that the best compromise between resolution and peak broadening was obtained at pH 4.2 [11]. As is shown in Fig. 6, an increase in resolution was observed for AGP glycoform bands in going from pH 5.0 to 4.2; however, the zone broadening increased and led to a reduction in resolution when going from pH 4.2 to 3.8. The effect of changing the concentration of Brij 35 as a buffer additive has been evaluated over the range of 0 to 0.5% (w/v). It was found that the use of 0.1% Brij 35 gave the best resolution for the glycoform bands, as is illustrated in Fig. 7 [11].

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Fig. 6 Electropherograms showing the effect of pH on the separation and resolution of AGP glycoform bands by CE. This figure is reproduced with permission from Ref. [11]

Fig. 7 Average resolution obtained for AGP glycoform bands by CE as a function of the concentration of Brij 35 that was placed into the running buffer. This figure was reproduced with permission from Ref. [11]. The error bars represent a range of 1 S.D. (n ¼ 3)

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Table 2 Effect on electroosmotic flow when using various modification agents and methods to treat fused silica capillariesa

Modification agent and method

Mobility due to electroosmotic flow, μosm ( 103cm2min1V1)

Reduction in electroosmotic flow (%)

Poly(vinyl alcohol) (or PVA) – Permanent coating

7.05 (0.02)

65.8 (1.4)

PVA – Permanent plus dynamic coating

7.09 (0.23)

66.8 (1.8)

11.04 (0.27)

48.0 (2.6)

PVA – Static plus dynamic coating

9.85 (0.06)

52.3 (2.0)

Poly (ethylene oxide) (or PEO) – Static coating

1.83 (0.01)

91.1 (0.4)

0.222 (0.002)

98.9 (0.1)

Dextran – Static coating

9.76 (0.01)

52.8 (2.0)

Dextran – Static plus dynamic coating

9.89 (0.10)

51.7 (2.1)

PVA – Static coating

PEO – Static plus dynamic coating

Reproduced with permission from Ref. [11] a The mobility due to electroosmotic flow for a fused silica capillary was 20.66 (0.87)  103 cm2 min1 V1. The electroosmotic flow was measured in pH 4.2, 20 mM acetate buffer with a sample containing 1.0 g/L thiourea in water. All of the results represent the average of three trials, where the number in parentheses corresponds to a range of 1 S.D

Table 2 compares the results that were obtained when using various types of coatings for the CE capillary. It was found that the use of PEO as both a static and dynamic coating reagent resulted in the greatest reduction of electroosmotic flow and the shortest overall analysis time when compared to capillary coatings based on poly(vinyl alcohol) (or PVA) or dextran for the separation of AGP glycoforms [11]. 14. The “inject” event during the method program has a time limit of up to 99 s. For the particular CE system that was employed in this study, a “separate” event can be used instead for sample injection during the method program for applying a voltage for 5 min. The sample can be added to a 2.0 mL vial and placed into the buffer tray to use this event. Every sample vial is only used for injection once in this approach because electrophoretic injection will deplete AGP from the sample vial without depleting the buffer or solvent. This means the concentration of AGP in the original sample vial will be reduced after even a single injection step. 15. This procedure should be performed right away after a separation sequence or multiple separations to avoid the permanent adsorption of serum proteins to the inner surface of the capillary, as may occur if this capillary has been left unrinsed for a long time.

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16. The peak area obtained by this method represents the true value without bias due to differences in the mobilities of the separate AGP glycoform bands [17]. When two analytes with the same quantity are injected by this approach, the analyte peak that has the longer migration time will have a larger area than the peak for the second analyte with the shorter migration time. This bias effect is automatically corrected during electrophoretic injection because a smaller amount is injected for an analyte with a smaller apparent mobility. More details on the theory of this approach can be found in Ref. [17].

Acknowledgement This work was supported by the National Institutes of Health under grant R01 GM044931. References 1. Fournier T, Medjoubi N, Porquet D (2000) Alpha-1-acid glycoprotein. Biochim Biophys Acta 1482:157–171 2. Ceciliani F, Pocacqua V (2007) The acute phase protein α1-acid glycoprotein: a model for altered glycosylation during diseases. Curr Protein Pept Sci 8:91–108 3. Scho¨nfeld DL, Ravelli RBG, Mueller U, Skerra A (2008) The 1.8-A˚ crystal structure of α1-acid glycoprotein (orosomucoid) solved by UV RIP reveals the broad drug-binding activity of this human plasma lipocalin. J Mol Biol 384:393–405 4. Mackiewicz A, Marcinkowska-Pieta R, Ballou S, Mackiewicz S, Kushner I (1987) Microheterogeneity of alpha 1-acid glycoprotein in the detection of intercurrent infection in systemic lupus erythematosus. Arthritis Rheum 30:513–518 ˜ a M, Gime´nez E, Puerta A, Llop E, 5. Balman Figueras J, Fort E, Sanz-Nebot V, de Bolo´s C, Rizzi A, Barrabe´s S, de Frutos M, Peracaula R (2016) Increased α1-3 fucosylation of α-1-acid glycoprotein (AGP) in pancreatic cancer. J Proteome 132:144–154 6. Lacunza I, Sanz J, Diez-Masa JC, de Frutos M (2006) CZE of human alpha-1-acid glycoprotein for qualitative and quantitative comparison of samples from different pathological conditions. Electrophoresis 27:4205–4214 7. Lacunza I, Sanz J, Diez-Masa JC, de Frutos M (2007) Erratum: CZE of human alpha-1-acid glycoprotein for qualitative and quantitative

comparison of samples from different pathological conditions. Electrophoresis 28:492 8. Paca´kova´ V, Hubena´ S, Ticha´ M, Madeˇra M, Sˇtulı´k K (2001) Effects of electrolyte modification and capillary coating on separation of glycoprotein isoforms by capillary electrophoresis. Electrophoresis 22:459–463 9. Kinoshita M, Murakami E, Oda Y, Funakubo T, Kawakami D, Kakehi K, Kawasaki N, Morimoto K, Hayakawa T (2000) Comparative studies on the analysis of glycosylation heterogeneity of sialic acidcontaining glycoproteins using capillary electrophoresis. J Chromatogr A 866:261–271 10. Kakehi K, Kinoshita M, Kawakami D, Tanaka J, Sei K, Endo K, Oda Y, Iwaki M, Masuko T (2001) Capillary electrophoresis of sialic acidcontaining glycoprotein. effect of the heterogeneity of carbohydrate chains on glycoform separation using an α1-acid glycoprotein as a model. Anal Chem 73:2640–2647 11. Zhang C, Hage DS (2016) Glycoform analysis of alpha1-acid glycoprotein by capillary electrophoresis. J Chromatogr A 1475:102–109 12. Simpson SL, Quirino JP, Terabe S (2008) On-line sample preconcentration in capillary electrophoresis. J Chromatogr A 1184:504–541 ´ lvarez PJ, Neusu¨ß C, de 13. Ongay S, Martı´n-A Frutos M (2010) Statistical evaluation of CZE-UV and CZE-ESI-MS data of intact α-1-acid glycoprotein isoforms for their use as potential biomarkers in bladder cancer. Electrophoresis 31:3314–3325

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14. Ongay S, Neusu¨beta C, Vaas S, Dı´ez-Masa JC, de Frutos M (2010) Evaluation of the effect of the immunopurification-based procedures on the CZE-UV and CZE-ESI-TOF-MS determination of isoforms of intact alpha-1-acid glycoprotein from human serum. Electrophoresis 31:1796–1804 15. Marino K, Bones J, Kattla JJ, Rudd PM (2010) A systematic approach to protein glycosylation analysis: a path through the maze. Nat Chem Biol 6:713–723 16. Garrido-Medina R, Puerta A, Rivera-MonroyZ, de Frutos M, Guttman A, Diez-Masa JC (2012) Analysis of alpha-1-acid glycoprotein isoforms using CE-LIF with fluorescent thiol derivatization. Electrophoresis 33:1113–1119 17. Zhang C, Bi C, Clarke W, Hage DS (2017) Glycoform analysis of alpha1-acid glycoprotein based on capillary electrophoresis and electrophoretic injection. J Chromatogr A 1523:114–122 18. Kishino S, Miyazaki K (1997) Separation methods for glycoprotein analysis and preparation. J Chromatogr B 699:371–381 19. Kremmer T, Szo¨llo¨si E´, Boldizsa´r M, Vincze B, Luda´nyi K, Imre T, Schlosser G, Ve´key K

(2004) Liquid chromatographic and mass spectrometric analysis of human serum acid alpha-1-glycoprotein. Biomed Chromatogr 18:323–329 20. Ongay S, Lacunza I, Dı´ez-Masa JC, Sanz J, de Frutos M (2010) Development of a fast and simple immunochromatographic method to purify alpha 1-acid glycoprotein from serum for analysis of its isoforms by capillary electrophoresis. Anal Chim Acta 663:206–212 21. Stumpe M, Miller C, Morton NS, Bell G, Watson DG (2006) High-performance liquid chromatography determination of alpha1-acid glycoprotein in small volumes of plasma from neonates. J Chromatogr B 831:81–84 22. Righetti PG, Caravaggio T (1976) Isoelectric points and molecular weights of proteins. J Chromatogr A 127:1–28 23. Muse LA (1972) Safe handling of the perchloric acid in the laboratory. J Chem Educ 49: A463 24. Varki A, Cummings RD, Esko JD, Freeze HH, Stanley P, Marth JD, Bertozzi CR, Hart GW, Etzler ME (2009) Symbol nomenclature for glycan representation. Proteomics 9:5398–5399

Chapter 5 On-Line Immunoaffinity Solid-Phase Extraction Capillary Electrophoresis-Mass Spectrometry for the Analysis of Serum Transthyretin Roger Pero-Gascon, Laura Pont, Victoria Sanz-Nebot, and Fernando Benavente Abstract The analysis of low abundant proteins in biological fluids by capillary electrophoresis (CE) is particularly problematic due to the typically poor concentration limits of detection of microscale separation techniques. Another important issue is sample matrix complexity that requires an appropriate cleanup. Here, we describe an on-line immunoaffinity solid-phase extraction capillary electrophoresis-mass spectrometry (IA-SPE-CE-MS) method for the immunoextraction, preconcentration, separation, detection, and characterization of serum transthyretin (TTR). TTR is a protein biomarker related to diverse types of amyloidosis, such as familial amyloidotic polyneuropathy type I (FAP-I), which is the most common hereditary systemic amyloidosis. Key words Analyte concentrator, Capillary electrophoresis, Immunopurification, In-line, Mass spectrometry, Microcartridge, On-line, Preconcentration, Sensitivity, Solid-phase extraction, Transthyretin

1

Introduction

1.1 Enhancing the Concentration Sensitivity in Capillary Electrophoresis

Capillary electrophoresis (CE) is regarded nowadays as a very suitable technique for the highly efficient separation of charged biomolecules, including peptides, proteins, and protein complexes [1–4]. However, the low concentration sensitivity for most analytes, due to the reduced sample injection volumes (typically 1–2% of the capillary volume), is very often a limitation that hinders a more widespread application, as in many other microscale separation techniques [5–10]. To decrease the limit of detection (LOD) in CE, several concentration strategies have been proposed in conjunction with the use of more sensitive and selective detectors (e.g., fluorescence and mass spectrometry (MS)) [5–10]. Some of them are based on the application of electrophoretic preconcentration techniques [5, 6], such as sample stacking [11], transient

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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isotachophoresis [12], and dynamic pH junction [13]. Although considerable concentration sensitivity enhancement can be obtained with these techniques, their dependence to the analyte and sample matrix characteristics hinders their performance in some applications. Furthermore, in general, the maximum loadability is limited to one capillary volume and the gain in sensitivity is achieved at the expense of losing separation resolution, as a large portion of the separation capillary is used to load the sample and not for the separation [5, 6]. Within the chromatographic preconcentration techniques, on-line solid-phase extraction capillary electrophoresis (SPE-CE) is widely recognized as an excellent alternative to improve loadability and selectivity and, consequently, the LODs [7–9]. Typically, in on-line SPE-CE, the extraction microcartridge, which contains the sorbent to retain the analyte from a large volume of sample (~50–100 μL), is inserted near the inlet of the separation capillary [8]. After sample loading, the capillary is rinsed to eliminate non-retained molecules and filled with background electrolyte (BGE). Then, the analyte is eluted in a small volume of suitable solution (~25–50 nL), resulting in sample cleanup and concentration enhancement before the electrophoretic separation and detection. Nowadays, the wide variety of commercial or lab-made sorbents, as well as microcartridge designs, have broadened the applicability of on-line SPE-CE [9]. Within the microcartridge designs, particle-packed microcartridges with frits have been widely applied because they prevent bleeding of the sorbents with the smallest particle size and the largest active surface area [8]. Other authors, for ease of construction, have preferred fritless microcartridges prepared from membranes [14], fibers [15], monoliths [16, 17], molecularly imprinted polymers [18], magnetic particles [19], or larger particles than diameter of the separation capillary [20]. These microcartridges are usually mounted in parallel to the separation capillary, but they can be mounted in zigzag (i.e., staggered) configuration, in multicapillary or multichannel instruments. In this last case, it is possible to do multiple parallel analyses, while avoiding the passage of sample through the separation capillary, which could decrease the analysis performance [7, 21, 22]. 1.2 On-Line Immunoaffinity SolidPhase Extraction Capillary Electrophoresis

SPE-CE has been extensively explored using the silica or polymeric sorbents typically used in off-line SPE such as C18, C8, or ion exchange because of their versatility and commercial availability [8, 9, 20]. However, the limited selectivity of such sorbents hampers the analysis of complex samples such as biological fluids. Sorbents with improved selectivity for specific analytes have been proposed, such as immobilized metal affinity chromatography (IMAC) [23], lectin [24], aptamer [17], or antibodies (Ab) [7, 21, 22, 25, 26]. Immunoaffinity (IA) sorbents prepared by immobilization of antibodies or antibody fragments provide

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excellent sample cleanup and extraction efficiency if the immunoreactivity and orientation of the antibody and the active surface area are optimum and nonspecific adsorption is minimized [25, 26]. On-line immunoaffinity solid-phase extraction capillary electrophoresis mass spectrometry (IA-SPE-CE-MS) is a state-of-the-art hyphenated technology that combines the selectivity of immunocapture by IA-SPE with the separation efficiency of CE, and the characterization power of on-line MS detection [25–29]. However, it is a challenge to make compatible the requirements of IA-SPE (i.e., IA sorbent stability and high recovery on-line immunoextraction) with on-line MS detection. Thus, on the one hand, high ionic strength BGEs, elevated temperatures, and acidic or alkaline conditions may cause Ab denaturation or elution and, on the other hand, BGEs must be volatile enough to prevent salt build-up and ionization suppression with on-line MS detection. In addition, the analysis of proteins by IA-SPE-CE-MS is especially challenging due to the potential issues related to low extraction efficiency, poor ionization efficiency, and adsorption onto the inner capillary walls [28, 29]. In recent years, different activated supports for immunoextraction have become available at a reasonable price, with a wide range of surface properties to easily and reproducibly couple intact Ab or Ab fragments [25–29]. IA sorbents prepared by covalent immobilization are the best alternative for IA-SPE-CE because of the improved stability, which makes the microcartridge reusable. In recent years, many different magnetic beads (MB) have become commercially available with a wide range of surface chemistries ready to covalently immobilize different affinity ligands [30, 31]. The robustness, versatility, and reasonable price of the commercial MBs and the simplicity of operation are rapidly expanding their application areas. In IA-SPE-CE, MBs are becoming very much appreciated because they facilitate the packing procedures and preparation of fritless microcartridges [19, 28, 32–34], as permanent magnets or electromagnets can be used to trap or move the particles. 1.3 Transthyretin (TTR) Amyloidosis Analysis

In recent years, interest has grown in the detection, characterization, and quantification of normal and variant forms (i.e., proteoforms) of different proteins as predictive indicators of an ongoing disease state [35]. Transthyretin (TTR) is a homotetrameric protein (relative molecular mass (Mr) of the monomer (MO) ~14,000) known to misfold and aggregate as stable insoluble fibrils causing diverse types of familial amyloidotic polyneuropathies. Among them, familial amyloidotic polyneuropathy type I (FAP-I), which is related to a single amino acid substitution of valine for methionine at position 30 of the TTR sequence (Met 30), is the most common [36, 37]. Traditional methods used to analyze the proteoforms of serum TTR require an off-line sample pretreatment before separation and characterization by LC-MS [38–41]. In this chapter, we detail a protocol for constructing and using in commercial CE instruments, capillaries for the analysis of TTR

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by IA-SPE-CE-MS using MBs derivatized with a polyclonal Ab (IA-MBs). The analytical method covers immunoextraction, preconcentration, detection, and characterization of TTR proteoforms from human serum to screen for individuals with suspected TTR amyloidosis, including FAP-I. The protocol is based on the IA-SPE-CE-MS applications that we recently described for the analysis of serum TTR [28, 29]. It is intended as a clear guide with detailed information of the most critical steps to engage non-initiated users concerned about the analysis of protein biomarkers of clinical interest by IA-SPE-CE, with or without on-line MS detection.

2

Materials Prepare all solutions using LC-MS grade water and analytical grade reagents or better. Store all solutions at 4
C and allow standing at room temperature before use. Degas the BGEs and the sheath liquid in an ultrasonic bath for 10 min before use. Pass all the solutions, excepting the sheath liquid, through 0.22 μm nylon filters before CE-MS and IA-SPE-CE-MS. 1. TTR standard solutions: Prepare an aqueous 1000 μg·mL1 stock solution of TTR dissolving with the appropriate volume of water the whole amount of solid in the vial provided by the manufacturer (see Note 1). Screw the cap onto the vial and invert several times to ensure complete homogenization of the stock solution of TTR. Centrifuge at 7000  g for 1 min at 25
C to bring the solution down. Store the stock solution of TTR as 50 μL aliquots in plastic vials at 20
C and thaw before use (see Note 2). Before using an aliquot of the stock solution of TTR, remove the excipients of low Mr by passage through 10,000 Mr cutoff cellulose acetate filters (Amicon® Ultra-0.5, Millipore, Bedford, MA, USA) (see Note 3). Prepare diluted standard solutions of TTR in phosphate buffered saline (PBS) at different concentrations from the 1000 μg mL1 stock solution (see Note 4). 2. Human serum samples: Collect human venous blood samples in collection tubes with Z serum separation clot activator and allow to coagulate leaving it undisturbed at room temperature for 9 h. Keep the blood at 4
C for 12 h to improve the clot retraction. Separate the supernatant from the clot with a Pasteur pipette and centrifuge at 1200  g for 20 min at 4
C. Separate the clear serum, and store as 50 μL aliquots in plastic vials at 20
C. Thaw before use (see Note 2). 3. Anti-TTR antibody (Ab): rabbit antihuman TTR polyclonal antibody (Dako, Glostrup, Denmark) (see Note 5). Exchange the solvent of a 50 μL aliquot of commercial 2400 μg mL1 Ab solution to PBS by passage through 10,000 Mr cutoff cellulose acetate filters (see Note 3). The Ab in PBS solution is prepared

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immediately before preparing the immunoaffinity sorbent with magnetic beads (IA-MBs). 4. Magnetic beads (MBs) AffiAmino Ultrarapid Agarose™ (UAAF) of 45–165 μm diameter (Lab on a Bead, Uppsala, Sweden) (see Note 6). Pipette 50 μL of UAAF MBs and transfer to a clean 0.5 mL vial. Vortex, sediment the particles using a cube magnet, and remove the supernatant. Wash the UAAF MBs with 100 μL of PBS twice and resuspend in 50 μL of PBS. Add 50 μL of Ab in PBS solution (2400 μg mL1) to the UAAF MBs suspension. Moderately shake the mixture for 40 min at room temperature using a Vortex Genius 3 (Ika®, Staufen, Germany). Remove the supernatant by magnetic separation and wash the IA-MBs three times with 100 μL of PBS. Add 80 μL of PBS and 20 μL of ethanol (EtOH) to store the IA-MBs in PBS with 20% (v/v) EtOH at 4
C when not in use (see Note 7). 5. Separation electrolyte or BGE: 10 mM ammonium acetate (NH4Ac, pH 7.0) (see Note 8). To prepare 100 mL of BGE, dissolve and mix 77 mg NH4Ac in about 90 mL water in a glass beaker. Transfer to a 100 mL volumetric flask and make up to the final volume with water. Transfer to a 125 mL glass bottle and check the pH value. If necessary, readjust the pH value with the smallest volume of diluted NH4OH solution (e.g., 100 mM NH4OH). 6. Sheath liquid: 60:40 (v/v) propan-2-ol:water with 0.25% (v/v) HFor (see Note 9). To a 125 mL glass bottle, add 60 mL of propan-2-ol, 40 mL of water, and 250 μL of HFor. 7. Eluent: 100 mM NH4OH (pH 11.2) (see Note 10). To prepare 100 mL of eluent, add 756 μL of concentrated ammonia solution (25%) to about 90 mL water in a glass beaker. Transfer to a 100 mL volumetric flask and make up to the final volume with water. 8. Capillary activation solution: 1 M NaOH (see Note 11). Weigh 4 g of NaOH pellets in a 125 mL glass bottle. Dissolve with 100 mL of water. 9. Bare fused silica capillary: 75 μm internal diameter (id)  360 μm outer diameter (od) (at least 1 m) and 250 id  360 μm od (at least 5 cm). 10. Binocular stereomicroscope with a magnification range up to 100 (see Note 12). 11. Fused silica capillary column cutter with a rotating diamond blade (see Note 13). 12. Peristaltic pump Tygon® plastic tubing (250 μm id, orangeblue retaining stops) (see Note 14). 13. Centrifugal filters of 10,000 Mr cutoff (see Note 15). 14. Neodymium cube magnet (12 mm, N48) (Supermagnete, Gottmadingen, Germany) (see Note 16).

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Methods

3.1 Serum Sample Pretreatment

Add 8 mg of NaCl to 100 μL of human serum at 4
C. Screw the cap onto the vial and invert several times to ensure complete homogenization. Add dropwise 100 μL of 5% (v/v) phenol. Invert several times to ensure complete homogenization and keep at 4
C for 30 min. Centrifuge the mixture at 11,000  g for 10 min at 4
C and collect the supernatant. Measure the volume of the recovered supernatant with an appropriate micropipette and add the same volume of PBS to dilute 1:1 (v/v) before the analysis (see Note 17).

3.2

Some of the capillary flushes are performed manually (see Note 18). In all the other cases, work at room temperature with the capillaries installed in the CE instrument using the commercial cartridge cassette for CE-MS (see Note 19).

IA-SPE-CE-MS

3.2.1 Fused Silica Capillary Preparation

1. Cut a 72 cm total length (LT)  75 μm id  360 μm od capillary to use as separation capillary (see Note 20). Activate the separation capillary by flushing the capillary with 1 M NaOH (15 min) and water (15 min) (see Note 11). Cut the separation capillary into two pieces of 7.5 and 64.5 cm (see Note 21). Flush manually with water (see Note 18). 2. Cut a 0.9 cm LT  250 μm id  360 μm od capillary to use as microcartridge body. Flush manually with water (see Note 18).

3.2.2 Construction of the IA Microcartridge

The construction of a fritless microcartridge using IA-MBs is described in Fig. 1. All the steps must be controlled with the microscope (see Note 12). 1. Cut a 0.5 cm long piece of the Tygon® plastic tubing with a scalpel. Push carefully the microcartridge body until it is 0.25 cm inside the plastic tube. Connect to the microcartridge body a disposable 3 cm LT  75 μm id  360 μm od capillary using the plastic tube (Fig. 1a) (see Notes 14 and 22). Push carefully the disposable capillary inside the plastic connector until you do not observe dead volume between the capillaries. Connect the free end of the disposable capillary to the adapted needle (see Note 18). Flush manually with water. 2. Prepare a reusable device to connect the adapted needle with the vacuum system. Cut 1 cm from the bottom of the body of a 5 mL disposable polypropylene syringe and fit it in an appropriate plastic vacuum hose (see Note 23). Attach the adapted needle connected to the disposable capillary-microcartridge body piece to the syringe tip. Pack the microcartridge by vacuum with the IA-MB sorbent until it is completely full (Fig. 1b) (see Note 24). Disconnect the adapted needle from the vacuum and use a soft paper towel to eliminate the particles from the outside of the microcartridge.

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Fig. 1 Construction procedure of a fritless IA-SPE-CE microcartridge (or analyte concentrator) using IA-MBs

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3. Connect the outlet end of the microcartridge body to the 64.5 cm piece of separation capillary, using a 0.5 cm plastic connector (see Note 14). Pull the disposable capillary carefully out of the connection with the microcartridge body inlet and connect the 7.5 cm piece of separation capillary. 4. Check that the microcartridge is completely full of IA sorbent under the microscope and that there is no dead volume between the IA-SPE-CE capillary connections (Fig. 1c). 5. Flush manually the IA-SPE-CE capillary from the inlet with water to check the system for abnormal flow restriction. Discard the IA-SPE-CE capillary if water is not flowing. 6. Install the IA-SPE-CE capillary inlet in the cartridge cassette (Fig. 1d) (see Note 21). 7. Install the IA-SPE-CE capillary outlet in the CE-MS sheathflow interface. Be careful not to damage the separation capillary exit tip. Leave 10 cm of capillary outside the electrode tip. Screw the PEEK connector of the interface to fix capillary position. Burn with a lighter the polyimide coating of the outlet end and carefully remove the ashes with a soft paper towel soaked with ethanol (see Note 25). Unscrew the PEEK connector and fix the capillary outlet protrusion to 0.1 mm with regard to the electrode outlet (see Note 26). Set sheath liquid flow rate at 3.3 μL/min (see Note 19) and the MS parameters optimized for detection of TTR (see Note 27). 8. Flush the new IA-SPE-CE capillary with BGE (10 min) and apply the separation voltage (25 kV, normal polarity, cathode in the outlet) 15 min for equilibration (see Note 20). Discard the capillary if current is zero, much lower than expected without on-line extraction or very unstable (drifts, spikes, etc.). 3.2.3 Immunoextraction, Preconcentration, Separation, and Characterization of TTR

For the optimum immunoextraction, preconcentration, separation, detection, and characterization of TTR, is required an appropriate balance between the loading, washing and BGE solutions composition and step durations, the elution plug composition and volume, and separation voltage. 1. For conditioning, flush the IA-SPE-CE capillary with BGE for 2 min (see Note 28). 2. For sample loading, flush diluted TTR standards in PBS or pretreated serum samples for 10 min (see Note 29). 3. Eliminate non-retained molecules and equilibrate the capillary before the electrophoretic separation flushing with BGE for 2 min (see Note 30). 4. Inject the eluent at 50 mbar for 10 s (see Note 31). 5. Introduce BGE at 25 mbar for 150 s (see Note 31).

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Fig. 2 (a) CE-MS for a 50 μg·mL-1 TTR standard. (b) IA-SPE-CE-MS for a 25 μg·mL-1 TTR standard. (i) Extracted ion electropherogram (EIE), (ii) mass spectrum, and (iii) deconvoluted mass spectrum. *The mass accuracy and resolution of the mass spectrometer were not enough to differentiate between these proteoforms, which had the same or very close Mr (Table 1) (MO monomer, DI dimer)

6. Apply the separation voltage of 25 kV for 30 min using BGE (Figs. 2 and 3, see Note 32). Refresh the BGE from the vial used to apply this voltage after each analysis. 7. Flush with 100 mM NH4Ac (pH 7.0) for 2 min and water for 2 min to prevent carryover between consecutive analyses. Discard the IA-SPE-CE capillary if you observe current instability and breakdowns, deterioration of the extraction performance or, always, at the end of the day (see Note 33).

4

Notes 1. TTR standard (Mr of the tetramer ~56,000) is provided as a lyophilized powder and is difficult to weigh accurately. 2. Thaw the standard TTR or serum aliquots using a water bath. It is not recommended to freeze an aliquot previously frozen and thawed.

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Fig. 3 IA-SPE-CE-MS for pretreated human serum samples (a) healthy control, (b) FAP-I patient. (i) Extracted ion electropherogram (EIE), (ii) deconvoluted mass spectrum. *The mass accuracy and resolution of the mass spectrometer were not enough to differentiate between these proteoforms, which had the same or very close Mr (Table 1) (MO monomer)

3. Centrifuge the solution at 11,000  g for 10 min at 25
C in a controlled-temperature centrifuge and wash the residue three times for 10 min in the same way, with PBS (0.011 M sodium hydrogenphosphate, 0.0015 M potassium dihydrogenphosphate, 0.14 M sodium chloride, 0.0027 M potassium chloride, pH 7.2). Recover the final residue by inverting the upper reservoir in a new vial and spin once more at a reduced centrifugal force (300  g for 2 min). Add sufficient PBS to adjust the concentration of TTR to 1000 μg mL1 or the concentration of Ab to 2400 μg mL1. 4. PBS is a buffer solution isotonic to the human body fluids and promotes an appropriate interaction between TTR and the Ab. The 1000 μg mL1 stock solution of TTR in PBS can be stored at 4
C for one week (no stability tests have been performed). The diluted standard solutions of TTR in PBS and treated serum samples have to be immediately discarded after the analysis (see Note 2).

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5. The polyclonal Ab from this manufacturer have been widely used by different experts in the field of FAP-I for immunoprecipitation of TTR [36], because of its good performance and reasonable cost. Other monoclonal and polyclonal antibodies can be used. Although monoclonal antibodies are more specific for a target antigen, polyclonal antibodies can recognize multiple epitopes of a single antigen, leading to higher tolerance to minor changes in the antigen and higher affinity than monoclonal antibodies [21]. Antibody fragments (Fab’) can be used instead of intact antibodies to further enlarge the immunoreactive area. This IA sorbent can be prepared by covalent attachment of Fab’ antibody fragments to succinimidyl silica particles [26, 29]. The performance of this sorbent for TTR analysis by IA-SPE-CE-MS is described in detail in reference [29]. 6. UAAF MBs are functionalized with amino-reactive groups to covalently bound the Ab without a preferred orientation. This IA sorbent is appropriate to obtain reproducible results by IA-SPE-CE-MS. We have also tested several commercial Protein A MBs with different particle size and binding capacity: Protein A Ultrarapid Agarose™ (Lab on a Bead), Dynabeads® Protein A (Life Technologies, Carlsbad, CA, USA), and SiMAG-Protein A (Chemicell GmbH, Berlin, Germany). Protein A strongly interacts with the Fc region of the Ab ensuring an appropriate Ab orientation. In all three cases, good results are obtained by off-line immunoprecipitation and CE-MS analysis of serum TTR. However, as there is no covalent bond between the Ab and the MBs, the conditions are harsh enough to elute the Ab together with the antigenic protein. For IA-SPE-CE-MS, this leads to gradual elution of the Ab during the analyses, poor repeatability, and decrease of the TTR recovery [28]. 7. The IA-MBs can be kept in the storage solution at 4
C for four weeks. The storage solution contains 20% (v/v) EtOH as an antimicrobial preservative to conserve the IA-MBs. 8. A BGE with neutral pH value is used to obtain sensitive and reproducible analysis in IA-SPE-CE-MS. In CE-MS, a BGE of 1 M HAc (pH 2.3) provides slightly better LODs (~10 vs. ~25 μg mL1) for TTR analysis in electrospray positive ion mode (ESI+) [42]. However, this acidic BGE is not recommended in IA-SPE-CE-MS because of Ab denaturation and protein elution during capillary conditioning and filling with BGE before the separation. Using the neutral BGE, the mass spectrum of a 50 μg·mL1 TTR standard shows the presence of protein monomer (MO) and dimer (DI) although the DI ions can detected with very low abundance (Fig. 2a (ii)). The tetramer cannot be detected by scanning until 3200 m/z. This is probably due to tetramer disruption during detection, because of the acidic sheath liquid (see Note 9) or the vacuum pressure

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inside the mass spectrometer. The vacuum pressure in the first stages of the 6220 oa-TOF LC/MS spectrometer (Agilent Technologies) cannot be tuned by the operator. Check if this vacuum pressure can be tuned in your instrument, because it could be interesting to detect TTR as the native tetramer [43]. Otherwise, for TTR characterization, we recommend to focus on the TTR proteoforms of the MO. The five most abundant TTR proteoforms for the MO (Table 1) can be detected in TTR standards by CE-MS (Fig. 2a (iii)). The main TTR proteoform corresponds to TTR showing a mixed disulfide with the amino acid cysteine at position 10 of the sequence (N ¼ 1, TTR-Cys). As can be observed in Table 1, depending on the mass spectrometer used for the analysis, the instrument mass accuracy and resolution can be unsatisfactory to differentiate between some of the proteoforms. In these cases, the reliability of the identification could be improved running -MS and -MS/MS experiments using mass spectrometers with improved mass accuracy and resolution. 9. In order to obtain sensitive and reproducible CE-MS and IA-SPE-CE-MS analysis of TTR with a BGE of 10 mM

Table 1 Theoretical average Mr for the detected TTR proteoforms by CE-MS and IA-SPE-CE-MS in standards and serum samples

Detected monomer TTR N proteoforms

Theoretical average Mr

1 TTR-Cys

13,880.4022

✓

2 Free-TTR

13,761.2640

3 TTR-Phosphorylated or TTR-Sulfonated

IA-SPE-CE-MS Healthy control

FAP-I patient

✓

✓

✓

✓

✓

✓

✓

13,841.2439 13,841.3283

✓

✓

✓

✓

4a Mutant Free-TTR (Met30) or TTR-Dehydroxylated or TTR-Sulfinic

13,793.3301 13,793.2628

– ✓

– ✓

– ✓

✓ ✓

5 (10) C-G

13,715.1713

✓

✓

✓

✓

6 TTR-Glutathione

14,066.5696

–

✓

–

–

7 TTR-CysGly

13,937.4541

–

✓

–

–

8 TTR-CysGlu

14,009.5177

–

✓

–

–

9 Mutant TTR-Cys (Met30)

13,912.4683

–

–

–

✓

a

a

CE-MS TTR standard (50 μg·mL1)

TTR standard (25 μg·mL1)

The mass accuracy and resolution of the mass spectrometer can be unsatisfactory to differentiate between these proteoforms, which had the same or very close Mr. For a reliable identification in these cases is necessary to run -MS and -MS/MS experiments using cutting-edge mass spectrometers with the best mass accuracy and resolution

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NH4Ac (pH 7.0) in ESI+ is necessary to have a 0.25% (v/v) HFor in the sheath liquid. In CE-MS with a BGE of 1 M HAc (pH 2.3), the best results are obtained with 0.05% (v/v) HFor. 10. No pH adjustment is necessary under these optimized conditions. 11. Use 1 M NaOH solution to activate the fused silica capillary before assembling the microcartridge. Later, it will ruin the IA sorbent in IA-SPE-CE-MS. The best option is to perform the activation without installing the capillary outlet in the CE-MS sheath-flow interface to avoid contamination of the mass spectrometer. Otherwise, turn off the nebulizer gas and the ESI voltage and leave the capillary outlet tip outside the metal electrode of the interface. Before the analysis, readjust the capillary protrusion in relation to the electrode outlet in the interface. 12. Control the capillary cuts, fabrication of the IA-SPE microcartridges, capillary connection dead volumes, and polyimide removal from the capillary outlet for on-line MS detection, with the stereomicroscope at 100 magnification. 13. Prepare clean-cut capillaries to obtain a proper connection between capillaries without dead volumes and a proper separation capillary outlet tip for CE-MS. The capillary cutter with a rotating diamond blade is a better alternative for unexperienced users than the ceramic capillary cutter to achieve clean-cut 360 μm od fused silica capillaries. 14. Tightly join two pieces of clean-cut 360 μm od fused silica capillaries using 0.5 cm long connectors made from this tubing. Use nitrile gloves or a soft paper towel to avoid capillary contamination during manipulation. No adhesive sealing is necessary and the microcartridge is completely replaceable. To obtain a proper connection without dead volumes between two pieces of capillary, introduce the shortest capillary piece until it is 0.25 cm inside the plastic tube and push the longest one against the connector. Be careful not to twist or bend the capillaries during connection. The 250 μm id fused silica capillary is especially fragile because the wall is very thin. 15. In Amicon® Ultra-0.5 filters of 10,000 Mr cutoff (Millipore), the cellulose acetate membrane is in vertical position for tangential (or cross) flow filtration. This configuration prevents membrane fouling, due to solute polarization, and a physical deadstop in the filter device prevents spinning to dryness and potential sample loss. 16. Use the neodymium cube magnet for magnetic separation when preparing the IA-MBs sorbent. In this protocol, IA-MBs are retained in the microcartridge due to their large average particle size (i.e., >75 μm, which is the inner diameter

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of the separation capillary, see Subheading 2, item 4). Therefore, neither frits nor magnet are required (Fig. 1). If smaller MBs are used, we propose a design taking advantage of the magnetic properties of the sorbent. In this case, select a neodymium magnet of the appropriate size and shape to use with the commercial cartridge cassette for CE-MS. For example, a neodymium block magnet (7  6  1.2 mm, N50, Supermagnete) can be installed in the cartridge cassette using tape, specifically under the microcartridge, to prevent small MBs bleeding to the separation capillary during flushing [28]. 17. This simple off-line sample pretreatment is necessary to analyze TTR from serum samples, to prevent microcartridge saturation and capillary inner surface damage due to the presence of other high-abundance proteins, especially albumin. The TTR recoveries for this off-line sample pretreatment are around 90% (comparing the CE-MS analysis of a serum sample pretreated and desalted with 10,000 Mr cutoff filters and a 250 μg mL1 TTR standard, which is the typical concentration of TTR in serum). 18. Use a hypodermic metal needle (40 mm LT  0.8 mm od) to prepare a needle adapted to fit 360 μm od capillaries. Sand the beveled tip of the hypodermic needle with small grit sandpaper. Wash the flat-tip needle with water. Connect to the needle tip a 0.5 cm long piece of the Tygon® plastic 250 μm id tubing. Use the adapted needle in combination with a 5 mL disposable polypropylene syringe to flush manually 360 μm od capillaries (avoid syringes with rubber tips on plungers). While connecting a capillary, avoid contact between the flat-tip metal needle and the capillary to prevent capillary end erosion. Be careful not to push the capillary too much inside the plastic connector. All capillaries need to be easily pulled out later. 19. The IA-SPE-CE-MS procedures are explained to be performed on a 7100 CE system (Agilent Technologies, Waldbronn, Germany). The instrument allows flushing capillaries at 930 mbar and hydrodynamic injections at low pressures ranging from 0 to 100 mbar. The CE-MS interface is an orthogonal sheath-flow interface (Agilent Technologies) with the separation capillary outlet protruding from the electrode approximately 0.1 mm. The sheath liquid is delivered by a syringe pump KD Scientific 100 Series (Holliston, MA, USA) at the flows required in CE-MS with this interface, usually between 2 and 4 μL min1. 20. Considering the distance between the CE instrument and the mass spectrometer entrance, a 72 cm LT separation capillary allows working comfortably. The separation voltage is optimized for analysis of TTR in a reasonable time. Electrophoretic currents lower than 50 μA are recommended to prevent

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electrical arcs between the electrode tip of the CE-MS interface and the mass spectrometer entrance. 21. Considering the commercial cartridge cassette for CE-MS from Agilent Technologies, the best position near the inlet to fit the microcartridge is at 7.5 cm from the inlet of the separation capillary. The microcartridge is completely inside the cartridge cassette. Use adhesive tape to fix the separation capillary and the microcartridge positions. Turn the deuterium lamp off if the UV detection is not used. 22. Connect the microcartridge body to a short and disposable 75 μm id  360 μm od capillary rather than to the separation capillary inlet for packing by vacuum. In this way, the IA-MBs smaller than the id of the capillary (i.e., particle size 18 MΩ cm 1 at 25  C and TOC < 5 ppb was obtained from Milli-Q UF-Plus system (Millipore). 2.2

Solutions

1. 50 mM phosphate–triethanolamine buffer, pH, 2.5: 2. 1 M NaOH: Weigh 4 g sodium hydroxide pellets and dissolve in 100 mL of water (see Notes 1 and 2). 3. 0.1 M NaOH: Add 10 mL of the 1 M NaOH solution to a 100 mL volumetric flask and make to the mark with water. Store in a tightly closed container at room temperature. 4. BGE. 50-mM phosphate-triethanolamine buffer (pH 2.5) with varying concentrations of CDs.

2.3

Instrumentation

1. A Beckman P/ACE MDQ Capillary Electrophoresis System. 2. A Beckman diode array detector The detection wavelength was fixed at 220 nm. 3. A BeckmanPACE station Version 1.1 used for data acquisition. 4. Fused-silica capillaries of 50 μm ID cut to total lengths 31.2 cm with an effective separation length of 21.0 cm.

2.4 Experimental Design Software

3

DOE and optimization were performed using Minitab®17 software (State College, PA, USA). The screening stage aims to achieve rapid and acceptable separations for all compounds under investigation.

Methods

3.1 Electrophoretic Conditions

1. All separations were performed using phosphate–triethanolamine buffer with varying concentrations of CDs. 2. Activation and preconditioning step for a new capillary were performed by rinsing with 1.0 M NaOH, 0.1 M NaOH, ultrapure water, and finally BGE solution. 3. Each preconditioning step was performed at a pressure of 60 psi for 10 min at temperature 20  C. 4. After each run, the capillary was first washed with 0.1 N NaOH for 3 min. Then, it was rinsed for 2 min with phosphate buffer (pH 2.5) and equilibrated for 2 min with chiral selector solution. 5. At the end of the day, the capillary was flushed with 0.1 M NaOH for 5 min and finally with water for 10 min.

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3.2 Analysis Procedures

1. Stock solutions of 1.0 mg mL 1 of the racemic investigated drugs were prepared in the least amount of methanol and diluted with water/ BGE solution for method development studies. 2. Standard solutions of 100 μg mL 1 of each compound were obtained by appropriate dilution with BGE solution.

3.3 Short-End Injection

1. The samples were injected at the end of the capillary near to the detector; this could be performed by applying negative pressure (see Note 3).

3.4

1. FLU is an acidic drug (pKa, 2.03) while DON is a basic drug (pKa, 8.82). Protonation of the piperidinic nitrogen of DON was obtained at acidic pH values. Since ionoselective selective interaction (the dissociated form complexes with the CD) is expected to occur at low pH values where the analytes are in their charged form. So, pH ranging from 2.5 to 5.0 was selected to represent the levels of pH in the DOE.

Interpretation

2. For screening phase, it is preferable to start by HS-CDs when developing new chiral enantioseparation strategy. 3. HS-CDs have a strong electrophoretic mobility toward the positive electrode in the CE environment that may be due the negatively charged groups. In addition, HS-CDs with phosphate-TEA buffer, pH 2.5 have been selected to enhance solubility and to generate mobility by protonation of the amine buffers (see Note 4). 4. DON (which is strongly cationic at low pH) would interact with the hydrophobic cavity as well sonically with the negatively charged sulfates. At pH from 2.5 to 3, zwitterionic analytes will be positively. 5. FLU will be primarily protonated and interact with the hydrophobic cavity of the HS-CDs. 6. Optimization steps were needed except for cases where Rs exceeded 1.5 meaning baseline acceptable separation. While in case of 0 < Rs < 1.5 after screening, where the peaks are only partially separated, some optimization steps were performed to increase the resolution. 7. It was found that SBE-β-CD led to the best enantioseparation in most of the investigated drugs as shown in Figs. 2 and 3. 3.5 Modeling the Separation

1. DOE allows experiments to be performed over the whole range of the factor space thus allowing accuracy with a minimum experimental effort. 2. The most crucial parameters are CDs concentration and pH which influence markedly in the chiral separation for drugs under investigation.

A Chiral Generic Strategy for Enantioseparation of Acidic and Basic Drugs. . .

131

Fig. 2 Representative CE electropherogram of the enantiomeric separation for (a) Fluconazole and (b) Donepezil (100 μg mL 1) for each drug under the following conditions: BGE consist of HS- γ-CD (10% m/v) in 50 mM phosphate buffer (pH 2.5), voltage 25 KV, temperature 25  C, and injection pressure 50 mbar for 10 s

3. The influence of two parameters, each at three levels, on the resolution of investigated drugs was studied. Nine experiments were carried out in duplicate (32) with high, low, and intermediate values for both factors. 4. The full factorial design and their responses (Rs) for these compounds are shown in Table 1. For each factor three effects can be estimated, for the intervals between the levels [1, 0], [0, 1], and [1, 1]. 3.5.1 Hydroxy Propyl-βCD (HP-β-CD)

1. For acidic and basic compounds, almost no peaks were detected for HP-β-CD (50–100 mM) and no baseline separations were obtained. 2. It was found that screening with neutral CDs does not result in sufficient separation; it is advised to investigate with charged CDs.

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Fig. 3 Representative CE electropherogram of the enantiomeric separation for (a) Fluconazole and (b) Donepezil (100 μg mL 1) for each drug under the following conditions: BGE consist of SBE (2.5% m/v) in 50 mM phosphate buffer (pH 2.5), voltage 30 KV, temperature 25  C, and injection pressure 50 mbar for 10 s 3.5.2 Highly Sulfated Gamma-CD (HS-γ-CD)

1. A study of the main effects showed that %CD was of little effect on the resolution of DON, with higher effect on the resolution of FLU. 2. Best Rs is obtained at pH 2.5 for the two racemates as shown in Fig. 4a. 3. Contour plots clearly show that maximum Rs were obtained at low pH values. %CD has a variable effect on Rs but is generally best when kept at low values with different thresholds as shown in Fig. 4b.

3.5.3 Highly Sulfated Alpha-CD (HS-α-CD)

1. While there was no separation for FLU, at any of the studied levels of all factors, the main effect study showed that for DON, the Rs improved at low values of pH and low or intermediate values of %CD as shown in Fig. 5a. 2. Contour plots control of %CD and pH is critical in case of DON where Rs drops to very low values ( 0.05) in all sample pairs relative to QC as healthy reference, (c) a PKU case reflected by a fourfold elevation in Phe, and (d) an MSUD case characterized by elevation in branched-chain amino acids, such as Val, unlike its isobar glycine betaine (Bet). An internal standard is included in all DBS extracts, which allows for correction for differences in injection volume between samples during peak integration while correcting for changes in EOF for RMT determination (reproduced from Ref. [39] with permission from ACS)

analysis (OPLS-DA). Several metabolites from DBS extracts were found to be significantly lower among CF neonates, including ophthalmic acid (OPA), several amino acids (Tyr, Ser, Thr, Asn), and an unknown polypeptide. Structural elucidation of OPA (a non-cysteinyl glutathione analog) was confirmed by MS/MS spectral matching based on three diagnostic product ions [41]. 6. Figure 6 illustrates the top four sweat metabolites that differentiate CF infants from screen-positive yet unaffected CF carriers when using nontargeted metabolite profiling by MSI-CEMS. Box-whisker plots and their corresponding receiver operating characteristic curves (ROCs) highlight good discrimination between CF cases and non-CF controls (AUC > 0.77; q < 0.05) beyond sweat chloride, which provides new insights

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Fig. 4 Inter-laboratory method comparison of MSI-CE-MS (McMaster) and DI-MS/MS (NSO) for quantification of tyrosine (Tyr) from independent 3.2-mm DBS cut-out specimens, including CF, non-CF, and screen-negative neonates (n ¼ 154). (a) A Passing-Bablok regression similarly confirms good mutual agreement for measured Tyr concentrations when using two independent methods from matching DBS specimens with a slope close to linearity (slope ¼ 1.17). (b) A Bland-Altman % difference plot also highlights good mutual agreement between the two methods when analyzing two different DBS cut-out samples with a modest mean bias of 6.9% for Tyr with eight outliers ( p ¼ 0.05; 8 out of 154) exceeding agreement limits at a 95% confidence level (reproduced from Ref. [41] with permission from ACS)

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(A)

(B)

Fig. 5 Metabolic signatures from DBS extracts associated with CF neonates as compared to unaffected screen-positive carriers. (a) A 2D scores plot using PLS-DA together with a variables of importance in projection (VIP) scores ranking for selection of top-ranked metabolites associated with CF neonates as compared to screen-positive/likely unaffected CF (SP/Non-CF) identified by two-tiered IRT/DNA algorithm based on 70 cationic metabolites reliably measured in retrospective DBS specimens. (b) Structural elucidation of unknown ion after collision-induced dissociation (CID) of precursor ion resulting in formation of three characteristic product ions that was consistent with ophthalmic acid or OPA (reproduced from Ref. [41] with permission from ACS)

into the complex pathophysiology of CF and individual treatments responses to nutritional and pharmacological therapy. Interestingly, two exogenous/biotransformed metabolites from exposures (pilocarpic acid, MEHP) and two endogenous metabolites (Asn, Gln) were determined to be associated with higher risk for paraoxonase deficiency and nutrient loss due to defective chloride transport, respectively [40].

6 Batch corrected RPA

B

7 5 4

Pilocarpic acid p = 1.12E-06 2.7x

3 2 1

0

D

Asparagine p = 3.88E-05

5 4

7.2x

3 2 1

40 Asparagine AUC = 0.863 95% CI = 0.702 to 0.898

0 Non-CF Group

0

F

MEHP p = 2.67E-04

0.2 0.1

60 40 MEHP AUC = 0.782 95% CI = 0.666 to 0.873

0

-0.1

Glutamine p = 5.44E-04 2.2x

6 5 4 3 2

0

H

20 40 60 80 100-Specificity

100

100 80

Sensitivity

Non-CF Group

8 7

100

100

20

0.0

CF

20 40 60 80 100-Specificity

80

2.0x

Sensitivity

Batch corrected RPA

60

20

0.5

0.3

100

100

0

0.4

20 40 60 80 100-Specificity

80 Sensitivity

Batch corrected RPA

Non-CF Group

6

CF

Batch corrected RPA

Pilocarpic acid AUC = 0.863 95% CI = 0.758 to 0.935

0

-1

G

40

0 CF

E

60

20

-1

C

100 80

Sensitivity

A

60 40 Glutamine AUC = 0.769 95% CI = 0.651 to 0.863

20

1 0

0 CF

Non-CF Group

0

20 40 60 80 100-Specificity

100

Fig. 6 Boxplots with scatter plot overlays and receiver operating characteristic (ROC) curves for the four top-ranked sweat metabolites in screen-positive CF infants. Plots compare differentiating metabolites (q < 0.05) in affected (n ¼ 18) and unaffected (n ¼ 50) screen-positive CF infants based on batch-corrected relative peak areas (RPA), including pilocarpic acid, PA (a, b), asparagine, Asn (c, d), monoethylhexylphthalate, MEHP (e, f), and glutamine, Gln (g, h). ROC curves indicate the area under the curve (AUC) and their 95% confidence interval (95% CI). For comparison, the ROC curve for sweat chloride had an AUC ¼ 1.0 with a median fold-change of 7.1 and p ¼ 7.85  107 in affected CF cases relative to unaffected screen-positive infants (non-CF) (reproduced from Ref. [40] with permission from ACS)

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157

Notes 1. Alternative CE systems from other manufacturers can be coupled with other mass analyzers (e.g., ion trap, triple quadrupole, Orbitrap) using conventional coaxial sheath liquid or recent commercially available low flow/sheathless CE-MS interface designs (e.g., porous sprayer, EOF-driven nanospray). 2. A segment of the outer polyimide coating of the fused-silica capillary (7 cm) is removed using a capillary window maker (MicroSolv) to reduce sample carryover between injections, as well as potential swelling effects when in contact with organic solvent in buffer. An optional polishing of the capillary ends can also be performed while rinsing with water to ensure more consistent cut surfaces at site of injection (inlet) and emitter (ion source). 3. Full-scan data acquisition over a wide mass range was used when using TOF-MS or QTOF-MS systems when performing nontargeted metabolite profiling of DBS extracts for biomarker discovery, as well as targeted quantification of known metabolites that serve as screening biomarkers for IEMs. Unknown metabolite identification was performed via collisional-induced dissociation of isolated precursor ions at three collisional energies (10, 20, and 40 V) when using a Q-TOF system [40, 41]. 4. Two CE-MS configurations are adopted depending on specific requirements of study. In cases when high sensitivity is needed for detection of low abundance metabolites, or the acquisition of high quality MS/MS spectra from weak precursor ion signals, then a single injection format is used with CE-MS in conjunction with online sample preconcentration. Alternatively, when higher sample throughput and data fidelity is required in support of large-scale clinical or epidemiological studies, including pilot studies for biomarker discovery, then a multiplexed separation format is optimally applied for analysis based on MSI-CE-MS that relies on serial injection of seven or more samples within a single run. In the latter case, one of the samples functions as a QC (i.e., pooled healthy control or reference sample), which is injected at random positions in all runs when using MSI-CE-MS since it allows for monitoring of long-term system drift, as well as batch correction of data [40, 41]. Additionally, identification of putative biomarkers for specific IEMs is facilitated since individual cases/samples can be directly compared to a reference population within the same run [39]. 5. The continuous pressure gradient used in conjunction with voltage application during CE separations allows for faster total analysis times by eluting slower migrating metabolites

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near the EOF without compromising isobar/isomer resolution or separation efficiency. 6. When suitable matching deuterated internal standards (D-IS) are not commercially available or too expensive, then a single non-deuterated IS was used to correct for differences in injection volume in-capillary between samples/runs to improve overall technical precision, namely 4-fluoro-L-phenylalanine, F-Phe. In this case, the ion response ratio of a metabolite (i.e., peak area ratio) was normalized to F-Phe that originated from the same sample injection position. Additionally, a second compound was used as a recovery standard, 3-chloro-L-tyrosine, Cl-Tyr, which was added to all DBS or sweat specimens prior to sample processing, including dilution and/or ultrafiltration. Control charts for the relative ion response of Cl-Tyr (relative to F-Phe) are often used to monitor long-term instrumental drift, including detection of potential outliers. 7. The Metabolomics Standards Initiative recommends that a minimum of two orthogonal parameters is used to annotate a molecular feature/unknown metabolite in a biological sample, namely its m/z and relative migration time (RMT). In this case, RMT corrects for changes in EOF between runs in CE by normalizing apparent migration times to an IS (F-Phe). Additionally, the charge state of an ion (i.e., single or doubly charged protonated molecular ion), mode of detection (e.g., positive or negative mode ESI), most likely molecular formula for neutral molecule, characteristic product ions or neutral losses when performing MS/MS, as well as specific chemical or enzymatic reactivity (i.e., functional group identification) can also support the annotation and structural elucidation of unknown metabolites of clinical significance. Unambiguous (level 1) identification requires confirmation of co-migration and MS/MS spectral match of metabolite in sample with its purified reference material [40, 41]. 8. All ultrafiltration tubes for deproteinization were first prerinsed with several aliquots of deionized water and spun down at 150 g for 1 min in a centrifuge in order to remove organic additives prior to processing of reconstituted DBS extracts within an hour. 9. Prechilled 1:1 MeOH:H2O (on ice) was found to greatly enhance the rate of hemolysis of DBS specimens immediately on contact relative to the same solvent mixture at room temperature. Subsequent liquid extraction was then continued under sonication for 10 min to maximize amino acid and acylcarnitine recovery. Routine clinical laboratories typically use MeOH to reduce co-extraction of salt without specifying the importance of temperature.

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10. Ultrafiltration is the deproteinization method of choice when processing whole blood or plasma samples without oxidation artifacts, which are induced by protein denaturation (e.g., oxygenases, hemoglobin) with acid or organic solvent precipitation. 11. The solvent of the DBS extract can also be evaporated under a gentle stream of N2. 12. Sample workup of DBS extracts should be processed within about 1 h while samples are kept on ice 4
C at all times using degassed/prechilled solutions. Otherwise, DBS extracts are preferably stored dried at 80
C and then reconstituted in solution with internal standards prior to CE-MS analysis. 13. Reproducible alignment of the capillary emitter into the ion source is facilitated by the Agilent coaxial sheath liquid ESI-MS interface, where the distal end of the bare fused-silica capillary was allowed to protrude from the sprayer by about 0.1 mm in order to minimize post-capillary dilution effects. 14. The average fused-silica capillary ($10 US/m) lifespan is typically 1 week (or up to 1000 sample injections) when analyzing DBS extracts or diluted sweat specimens by MSI-CEMS under continuous operation. In most cases, capillary replacement is made evident by an unstable capillary current and/or increasing noise in the total ion electropherogram that cannot be improved with subsequent conditioning or rinsing with BGE as a result of a capillary fracture. 15. A standard mixture, a quality control sample (QC), and a blank were analyzed at the beginning of each day to equilibrate the CE-MS system before analysis of a randomized batch of DBS extracts or sweat specimens and confirm absence of background signals for metabolites being analyzed. For overnight storage, capillaries were flushed with water for 10 min and air dried for 10 min. As preventative maintenance, the electrode was daily cleaned with 50% v/v isopropanol-water and methanol to avoid sample carryover. The incorporation of a QC check comprising pooled DBS specimens or sweat from screen-negative neonates is run also to confirm system suitability, such as resolution of key isomers/isobars and detection of internal/recovery standards. The same QC is also introduced randomly within each run when using MSI-CE-MS with a serial 7-sample injection format, which also allows for monitoring of long-term instrumental drift, and applied for QC-based batch correction of data. Also, daily preventative maintenance protocols including mass tuning, or mass calibration as required [39–41]. 16. A small fraction of organic modifier is added to the aqueous background electrolyte or BGE (e.g., 15% v/v acetonitrile) in

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order to improve the solubility of hydrophobic analytes or suppress micellar formation of surface-active solutes, such as long-chain acylcarnitines [36]. The buffer is prepared by diluting concentrated formic acid in water and 15% v/v acetonitrile until an apparent pH 1.8 is obtained under positive ion mode detection (ESI+). If metabolite analysis of acidic/anionic metabolites is required (e.g., organic acids, sugar phosphates), then an alkaline BGE comprised of 50 mM ammonium bicarbonate, pH 8.5 (adjusted with 10% v/v ammonium hydroxide) is used under negative ion mode (ESI-) detection. It is important that ammonia based buffers do not exceed a pH > 9 due to irreversible ammonia hydrolysis of outer polyimide capillary coating resulting in incidental capillary fractures [45]. As a result, comprehensive profiling of cationic and anionic metabolites from the same DBS extract or sweat specimen is performed in two independent runs in MSI-CE-MS [39–41]. 17. In all cases, the current generated during voltage application in CE should be kept under 50 μA in order to minimize Joule heating effects and improve ion signal stability in ESI-MS. 18. The TOF or Q-TOF system was calibrated every morning before analysis using the Agilent tune mixture for m/z 50–1700 range. An Agilent 1260 Infinity series isocratic pump equipped a 100:1 splitter was used to deliver sheath liquid to the CE-MS interface at a flow rate of 10 μL/min. The sheath liquid for ESI+ consisted of 60% v/v methanol in water with 0.1% v/v formic acid, whereas for ESI it was 50% v/v methanol in water. Purine and HP-0921 (API-TOF Reference Mass Solution Kit, Agilent Technologies) were added to the sheath liquid as reference masses for real-time automatic mass recalibration. The nebulizer gas kept at 8 psi, while the flow rate for the drying gas was maintained at 16 L/min (300
C), and the sheath gas flow was kept at 3.5 L/min (200
C). The instrument was kept in extended dynamic range (EDR, 2 GHz) for ESI+ experiments to prevent saturation of highly abundant cationic metabolites, whereas ESIexperiments were performed in high resolution (HiRes 4 GHz) to improve detection of anionic metabolites that in general have lower sensitivity. 19. Hydrodynamic injection of a long sample plug (10% of capillary length) allows for online sample preconcentration of amino acids directly in-capillary during electromigration prior to ionization based on differences in co-ion electrolyte mobility and pH at the sample and BGE interface. Up to a 50-fold improvement in concentration sensitivity can be realized by this method without compromising separation efficiency or resolution when using conventional instrumentation [36].

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20. The second injection sequence is used to displace the original sample plug within the capillary past the electrode interface at the inlet (anode), which is required to avoid CE-induced oxidation artifacts when analyzing low micomolar levels of oxidized glutathione (GSSG) in the presence of excess reduced glutathione (GSH) derived from filtered cell lysates [46]. 21. Under these conditions, neutral metabolites (e.g., urea) co-migrate with the suppressed EOF (>20 min), whereas strongly acidic anions (e.g., chloride) migrate out of the capillary at the inlet upon voltage application. The relative migration time (RMT) of an ion can be accurately predicted in CE based on its characteristic μ0 and thermodynamic pKa as derived from its putative chemical structure [42, 44], which provides a novel strategy for identification of amino acids and acylcarnitines complementary to ESI-MS. 22. In most cases, amino acids and acylcarnitines are detected as their singly charged protonated molecular ion (M + H+) with the exception of peptides such as oxidized glutathione (GSSG), which predominately forms a divalent molecular ion (M + 2H2+) in the gas-phase. Optimization of ionization conditions can reduce the extent of in-source fragmentation that can compromise sensitivity. Amino acid salt adducts are rarely detected by CE-MS due to the efficient separation of major electrolytes (Na+, K+) in conjunction with post-capillary mixing with the sheath liquid solution effluent in the ion source. 23. Serial sample injections used in MSI-CE-MS for alternating BGE/sample plugs are introduced via hydrodynamic injections. However, electrokinetic injections may also be used for BGE spacers in order to initiate separations following sample introduction in order to improve resolution without loss in effective capillary length. Additionally, nonaqueous BGE systems may also be used to analyze water-insoluble metabolites of clinical significance, such as long-chain fatty acids, bile acids, and intact lipids [47]. Various sample injection configurations can be designed in MSI-CE-MS in order to enhance sample throughput while ensuring data quality. Customized software tools that are optimal for processing multiplexed separations based on temporal signal pattern recognition are currently under development.

Acknowledgements The author wishes to acknowledge funding support from National Science and Engineering Research Council of Canada, Cystic Fibrosis Canada, and Genome Canada.

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References 1. Levy PA (2010) An overview of newborn screening. J Dev Behav Pediat 31:622–631 2. Chace DH (2009) Mass spectrometry in newborn and metabolic screening: historical perspective and future directions. J Mass Spectrom 44:163–170 3. Sweetman L (2010) Newborn screening by tandem mass spectrometry. Clin Chem 47:1937–1938 4. Zytkovicz TH et al (2001) Tandem mass spectrometric analysis for amino, organic, and fatty acid disorders in newborn dried blood spots. Clin Chem 47:1945–1955 5. Wilcken B et al (2009) Expanded newborn screening: outcome in screened and unscreened patients at age 6 years. Pediatrics 124:e241–e248 6. American College of Medical Genetics Newborn Screening Expert Group (2006) Newborn screening: toward a uniform screening panel and system--executive summary. Pediatrics 117:S296–S307 7. Burlina AB et al (2018) Newborn screening for lysosomal storage disorders by tandem mass spectrometry in North East Italy. J Inhert Metabol Dis 41:209–219 8. Ia Marca G et al (2013) Tandem mass spectrometry, but not T-cell receptor excision circle analysis, identifies newborns with late-onset adenosine deaminase deficiency. J Allergy Clin Immunol 131:1604–1610 9. Ia Marca G et al (2013) Diagnosis of immunodeficiency caused by a purine nucleoside phosphorylase defect by using tandem mass spectrometry on dried blood spots. J Allergy Clin Immunol 131:1604–1610 10. Gurian EA et al (2006) Expanded newborn screening for biochemical disorders: the effect of a false-positive result. Pediatrics 117:915–1921 11. Tarini BA, Christakis DA, Welch HG (2006) State newborn screening in the tandem mass spectrometry era: more tests, more falsepositive results. Pediatrics 117:448–456 12. Lehotay DC et al (2011) LC-MS/MS progress in newborn screening. Clin Biochem 44:21–31 13. Minkler PE et al (2015) Quantitative acylcarnitine determination by UPLC-MS/MS—going beyond tandem MS acylcarnitine “profiles”. Mol Genet Metabol 116:231–241 14. Seo JY et al (2014) Steroid profiling for congenital adrenal hyperplasia by tandem mass spectrometry as a second-tier test reduces follow-up burdens in a tertiary care hospital: a

retrospective and prospective evaluation. J Perinat Med 42:121–127 15. Kuehnbaum NL et al (2013) New advances in separation science for metabolomics: resolving chemical diversity in a post-genomic era. Chem Rev 113:2437–2468 16. Miller JH 4th et al (2012) A quantitative method for acylcarnitines and amino acids using high resolution chromatography and tandem mass spectrometry in newborn screening dried blood spot analysis. J Chromatogr B 903:142–149 17. Roy C et al (2016) Quantitative analysis of amino acids and acylcarnitines combined with untargeted metabolomics using ultra-high performance liquid chromatography and quadrupole time-of-flight mass spectrometry. J Chromatogr B 1027:40–49 18. Farez-Vidal ME et al (2008) Multi-mutational analysis of fifteen common mutations of the glucose 6-phosphate dehydrogenase gene in the Mediterranean population. Clin Chim Acta 395:94–98 19. Suksangpleng T et al (2017) A novel system for newborn screening of thalassemia and hemoglobinopathies using capillary electrophoresis is superior than isoelectric focusing. Blood 130:2089 20. Klingenberg O et al (2017) HbA1c analysis by capillary electrophoresis – comparison with chromatography and an immunological method. Scand J Clin Lab Invest 77:458–464 21. Barbas C et al (2002) Evaluation of filter paper collection of urine samples for detection and measurement of organic acidurias by capillary electrophoresis. J Chromatogr B 780:73–82 22. Boulat O et al (2001) Separation of free amino acids in human plasma by capillary electrophoresis with laser induced fluorescence: potential for emergency diagnosis of inborn errors of metabolism. J Chromatogr B 754:217–228 23. Lochman P et al (2003) High-throughput capillary electrophoretic method for determination of total aminothiols in plasma and urine. Electrophoresis 24:1200–1207 24. Vernez L, Thormann W, Krahenbuhl S (2000) Analysis of carnitine and acylcarnitines in urine by capillary electrophoresis. J Chromatogr A 895:309–316 25. Friedecky D, Adam T, Bartak P (2002) Capillary electrophoresis for detection of inherited disorders of purine and pyrimidine metabolism: a selective approach. Electrophoresis 23:565–571

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Chapter 11 Capillary Electrophoresis-Mass Spectrometry for Metabolic Profiling of Biomass-Limited Samples Wei Zhang, Thomas Hankemeier, and Rawi Ramautar Abstract Capillary electrophoresis-mass spectrometry (CE-MS) is a strong separation technique for the highly efficient and selective analysis of polar and charged metabolites in biological samples. The CE approach is especially suited for the analysis of limited sample amounts due to its nanoliter injections from only a few microliters of material in the sample injection vial. In this protocol, a CE-MS strategy is outlined for the profiling of cationic metabolites in biomass-limited samples using a small amount of human hepatocellular carcinoma (HepG2) cells as a model system. By employing a sheathless interfacing design for coupling CE to MS, it is shown that information-rich profiles for cationic metabolites can be obtained when working with a starting amount of 10,000 HepG2 cells and even lower. Overall, the proposed CE-MS-based analytical workflow may be considered a useful tool for biomass-limited metabolomics studies. Key words Capillary electrophoresis, Mass spectrometry, Sheathless interface, Metabolic profiling, Biomass-limited samples, Cationic metabolites

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Introduction Metabolomics aims at the comprehensive profiling of small (endogenous) molecules in a living system, unraveling biological processes and helping to tackle physiological issues [1–3]. This relatively new field of –omics has therefore assisted researchers in understanding the compositions and functions of living organisms and the environment. Various analytical tools have been introduced in order to deliver metabolic profiles that cover crucial metabolites, including nuclear magnetic resonance spectroscopy (NMR) and mass spectrometry (MS)-based platforms. Over the last few years, MS-based platforms have emerged as the main technique in the field of metabolomics [4]. In spite of the advancements made on liquid chromatography column technology and methodology, the analytical selectivity and efficiency for especially highly polar and charged metabolites still remains a challenge. Capillary electrophoresis (CE) separates compounds

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_11, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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based on the difference in their size-to-charge ratios, which makes this approach highly suitable for the analysis of polar and charged metabolites. Due to the distinctive and unique separation mode of CE, it can offer complementary information about the composition of metabolites in a given biological sample. At present, CE-MS is not as widely used as other techniques in global metabolic profiling studies [5], as within the separation science community this analytical technique is still often perceived to be a technically challenging approach. In 2015, Christian Wenz et al. reported a peptide mapping study involving 13 different labs, using the same sample, method, PVA-coated capillaries but a different set of instrumentations [6]. The results showed that by employing well-described suitability tests and protocols, CE-MS is capable of delivering reproducible outcomes for peptide mapping and that analytical method transfer can be performed in an effective manner among different laboratories. Recently, Harada et al. has used CE-MS for a large-scale metabolomics study [7]. In this study, 8000 human plasma samples were analyzed over a 52-month period. A unique and broad coverage of 94 polar metabolites was obtained with a similar or better reproducibility when compared to other analytical platforms employed for large-scale targeted metabolomics studies. With the introduction of a recently developed software denoted as ROMANCE, researchers now can easily convert typical CE-MS timescales into electrophoretic mobility scales by using a reference peak with known electrophoretic mobility, enabling better peak alignment and normalized peak areas, provided that the same background electrolyte is used [8]. As such, this tool may facilitate comparative metabolic profiling studies by CE-MS. In the classical CE-MS approach, the CE effluent is significantly diluted by the sheath-liquid which is provided at a flow-rate between 2 and 10 μL/min, resulting in compromised detection sensitivities. A lot of efforts have been dedicated to abolishing or minimizing the usage of the sheath-liquid, among which the sheathless porous tip interface has yielded very encouraging results so far. Developed by Moini [9], the sheathless porous tip interface is now commercially available and has been recently applied in areas like proteomics [10] and metabolomics [11, 12]. The sheathless CE-MS approach has very strong analytical features for the highly sensitive analysis of metabolites in particular for small-volume biological samples. In order to demonstrate the usefulness of sheathless CE-MS for biomass-limited metabolomics studies, a protocol is presented here illustrating how this technique can be used for the analysis of cationic metabolites extracted from limited amounts of mammalian cell samples, with the example here being human hepatocellular carcinoma (HepG2) cells. Part of the procedures outlined here has also recently been reported in a peer-reviewed video-article [13]. Overall, it is shown

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that the proposed sheathless CE-MS-based analytical workflow can deliver a decent coverage of cationic metabolites from as little as 10,000 HepG2 cells, denoting its potential for materialrestricted metabolomics studies.

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Materials Water used for the preparation of all solutions is obtained from a Milli-Q system equipped with 0.22 μM pore-size filter. Reagents used are of analytical grade unless stated otherwise.

2.1 Solutions and Samples for Analysis

1. Background electrolyte (BGE) solution: Add 1.0 mL of acetic acid into 9.0 mL of water and sonicate the mixture for 10 min under room temperature. Store at 4  C. 2. 1 M ammonium acetate: Weigh 3.854 g ammonium acetate into a 50 mL volumetric flask and add water to the marked line. Vortex thoroughly till the solid is completely dissolved. Transfer the solution to a 50 mL falcon tube. 3. 0.1 M sodium hydroxide: Weigh 0.112 g of sodium hydroxide into a 25 mL volumetric flask and add water up to the mark. Vortex and sonicate the mixture till no solid is observed. Transfer the solution to a vial. Store the prepared solutions at 4  C prior to use. 4. Metabolite standard mixture: Weigh and dissolve 31 cationic metabolite standards separately in small tainted vials with a mixture of water and acetonitrile (95:5, containing 0.1% formic acid) to a concentration of 10 mM (unless otherwise stated). The cation mixture includes (1) spermine; (2) homoserine; (3) hypoxanthine (1 mM); (4) GSSG; (5) adenosine; (6) histidine; (7) methionine; (8) L-alanine; (9) tyrosine (1 mM); (10) threonine; (11) proline; (12) glutamine; (13) asparagine (6.67 mM); (14) serine; (15) valine; (16) glutamic acid; (17) glycine; (18) 4-hydroxyproline; (19) phenylalanine (6.67 mM); (20) cytidine; (21) lysine; (22) aspartic acid; (23) isoleucine; (24) leucine; (25) spermidine; (26) tryptophan; (27) anthranilic acid; (28) cytosine (6.67 mM); (29) tyramine (6.67 mM); (30) adenine (6.67 mM); and (31) creatine. Add 50 μL of each stock solution (10 mM; 75 μL for 6.67 mM; 500 μL for 1 mM) into a 10 mL volumetric flask and fill the flask up to the marked line with water:acetonitrile (95:5, containing 0.1% formic acid) mixture to make a standard mix of 50 μM. Shake the flask vigorously. Transfer the solution to a glass vial and store at 20  C.

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2.2 Analytical Equipment

1. The reported protocol here can only be conducted on a sheathless CE instrument, CESI 8000. The sheathless CE is hyphenated to MS via a dedicated nanospray source, and relevant information can be acquired from Refs. [13, 14]. 2. For the electrophoretic separations, fused-silica capillary cartridges (30 μm I.D.  90 cm total length) are used.

3

Methods The protocol presented here for the application of sheathless CE-MS in biomass-limited cationic profiling studies is for laboratory use only. Please comply with proper laboratory safety procedures, and wear safety goggles, lab coats, and gloves, when carrying out the experiments described below. The setup of sheathless CE-MS has been described previously and interested readers are referred to another publications [13–15].

3.1 Preparation of Extracts from the Human Hepatocellular Carcinoma (HepG2) Cell Line

1. Culture the human hepatocellular carcinoma (HepG2) cell line in Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F-12 Ham (DMEM) supplemented with 10% (v/v) fetal calf serum and 1% (v/v) of penicillin/streptomycin (see Note 1). The medium is replaced every three days. 2. The harvesting of the cells is done by detaching the adherent cells using trypsinization. Cells are stained with Trypan Blue and counted using an automated cell counting system. 3. Resuspend the cells in 37  C PBS to the concentration of 2  106/mL and distribute the mixture to clean Eppendorf tubes (1 mL per tube). Centrifuge the tubes and remove the PBS. Store the dry cell pellets at 80  C. 4. Add 2 metal beads (see Note 2) and 1 mL cold methanol:water (80:20, v/v) mixture to the cell pellet. Subject the mixture to a bullet blender for 2 min at high speed. 5. Take 50 μL of the lysed cell mixture and mix it with 450 μL cold methanol:water (80:20, v/v) mixture, so that the diluted mixture has the concentration of 10,000 cells per 50 μL (see Note 3). 6. Transfer 50 μL of the diluted mixture (step 5) to a clean Eppendorf tube for metabolite extraction. Add chloroform, methanol, water and internal standard solution to make a final mixture of 750 μL, with the ratio of methanol:water: chloroform being 2:2:1. Vortex for 1 min and centrifuge the mixture for 10 min at 16,100  g. 7. Transfer 500 μL of the supernatant after centrifuge and subject it to ultrafiltration using 5 kDa cutoff membrane filters to further remove (residual) proteins by centrifugation at

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Fig. 1 An overview of the analytical workflow used for the preparation of extracts from HepG2 cells

9000  g for 2.0 h. Collect 380 μL of the filtered aqueous phase and evaporate it to dryness in a LabConco SpeedVac. Store the dried extracts at 20  C briefly before analysis. 8. Add 50 μL of 250 mM ammonium acetate solution to reconstitute the dry material. Vortex and centrifuge for 10 min at 16,100  g prior to sheathless CE-MS analysis. This procedure is summarized in Fig. 1. 3.2 Analysis of Metabolite Standards and Biological Samples

1. Add 50 μL of the metabolite standard mixture into an empty 250 μL microvial and 50 μL of 500 mM ammonium acetate. Mix well and put the vial in the inlet sample tray, where the temperature is maintained at 10  C. 2. Between runs, precondition the capillary by flushing with water (forward, 50 psi for 2 min), 0.1 M sodium hydroxide (forward, 50 psi for 3 min), water (forward, 50 psi for 2 min), and BGE (forward, 50 psi for 3 min; reverse, 50 psi for 2 min) (see Note 4). Inject the sample hydrodynamically at 6 psi for 60 s, which corresponds to circa 42 nL or 6.6% of the total capillary volume. Then perform a BGE injection at 1 psi for 10 s. 3. Start MS data acquisition and apply a voltage of 30 kV (ramp time of 1.0 min) for cationic metabolic profiling for 22 min. After the electrophoretic separation, stop MS data acquisition, and decrease the CE voltage to 1 kV using a ramp time of 5 min. 4. Assess the recorded data by determining the migration times of the signal intensity of the analyzed metabolite standard mixture (see Note 5). 5. Use the procedures described in Subheading 3.1, steps 1–8, for cationic metabolic profiling of extracts of the human hepatocellular carcinoma (HepG2) cell line. A typical profile for cationic metabolites extracted from the HepG2 cells by sheathless CE-MS is shown in Fig. 2 (see Note 6).

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Fig. 2 Multiple extraction ion electropherograms for a selected number of metabolites obtained in an extract of circa 10,000 HepG2 cells with sheathless CE-MS in positive ion mode using a porous tip emitter. Peaks: (1) lysine; (2) s-adenosylmethionine; (3) histidine; (4) glycine; (5) creatine; (6) cytidine; (7) l-alanine; (8) serine; (9) asparagine; (10) anthranilic acid; (11) phenylalanine; (12) tyrosine; (13) hypoxanthine; (14) hydroxyproline

6. After the analyses or when not in use, rinse the capillary with water (forward, 50 psi for 10 min; reverse, 50 psi for 3 min), and put the capillary on water and rinse at 5 psi until further use (see Notes 7 and 8).

4

Notes 1. The cells are incubated in 75 cm2 tissue culture flasks at 37  C under 5% CO2 in an incubator. 2. The metal beads can come in different sizes, the ones used are about 2 mm in diameter. When dealing with smaller beads, more beads can be added to ensure thorough lysis. 3. The dilution process can be further repeated to achieve the targeted cell concentration as required for the experiment. The whole sample extraction procedure is performed on ice. 4. The sheathless capillary cartridge consists of a separation capillary and a conductive capillary. Both of them need to be thoroughly flushed before the start of measurements (see Refs. [13–15]). 5. Please check for the following test compounds of the standard mixture whether the migration times are around the indicated values, i.e., for histidine: 12.4 min; l-alanine: 13.5 min; phenylalanine: 14.7 min, and 4-hydroxyproline: 15.4 min. 6. Check the resolution for the two peaks of phenylalanine and tyrosine and determine whether their resolution factor is around 0.9.

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7. When the sheathless CE-MS method is not in use, it is important to avoid drying out the porous tip capillary. One practice is to disconnect the separation capillary from the MS instrument and to store the outlet section of the capillary in water. The practice mentioned here is to rinse the capillary with water at a low flow-rate, i.e., at 5 psi for 10 min in reverse mode and at 5 psi for 300 min in forward mode, which procedure also guarantees that the position of the porous tip emitter in front of the MS inlet will remain the same, thereby making follow-up analyses easier. 8. Concerning durability of a single porous tip capillary emitter, at this stage up to 100 samples, including both standards and HepG2 cell extracts, can be analyzed. Further improvement of the sample preparation procedure is needed to improve the long-term performance of sheathless CE-MS for metabolic profiling of extracts from HepG2 cells.

Acknowledgements Wei Zhang would like to acknowledge the Chinese Scholarship Council (CSC, No. 201507060011). Dr. Rawi Ramautar would like to acknowledge the financial support of the Veni and Vidi grant scheme of the Netherlands Organization for Scientific Research (NWO Veni 722.013.008 and Vidi 723.016.003). This work was also supported by the European Union’s Seventh Framework Programme for research, technological development and demonstration (FP7/CAM-PaC) under grant agreement no 602783. References 1. Patti GJ, Yanes O, Siuzdak G (2012) Innovation: metabolomics: the apogee of the omics trilogy. Nat Rev Mol Cell Biol 13(4):263–269. https://doi.org/10.1038/nrm3314 2. Gitto S, Schepis F, Andreone P, Villa E (2018) Study of the serum metabolomic profile in nonalcoholic fatty liver disease: research and clinical perspectives. Metabolites 8(1):E17. https://doi.org/10.3390/metabo8010017 3. Ramautar R, Berger R, van der Greef J, Hankemeier T (2013) Human metabolomics: strategies to understand biology. Curr Opin Chem Biol 17(5):841–846. https://doi.org/10. 1016/j.cbpa.2013.06.015 4. Theodoridis GA, Gika HG, Want EJ, Wilson ID (2012) Liquid chromatography-mass spectrometry based global metabolite profiling: a review. Anal Chim Acta 711:7–16. https://doi. org/10.1016/j.aca.2011.09.042 5. Kuehnbaum NL, Britz-McKibbin P (2013) New advances in separation science for

metabolomics: resolving chemical diversity in a post-genomic era. Chem Rev 113 (4):2437–2468. https://doi.org/10.1021/ cr300484s 6. Wenz C, Barbas C, Lopez-Gonzalvez A, Garcia A, Benavente F, Sanz-Nebot V, Blanc T, Freckleton G, Britz-McKibbin P, Shanmuganathan M, de l’Escaille F, Far J, Haselberg R, Huang S, Huhn C, Pattky M, Michels D, Mou S, Yang F, Neusuess C, Tromsdorf N, Baidoo EE, Keasling JD, Park SS (2015) Interlaboratory study to evaluate the robustness of capillary electrophoresis-mass spectrometry for peptide mapping. J Sep Sci 38(18):3262–3270. https://doi.org/10. 1002/jssc.201500551 7. Harada S, Hirayama A, Chan Q, Kurihara A, Fukai K, Iida M, Kato S, Sugiyama D, Kuwabara K, Takeuchi A, Akiyama M, Okamura T, Ebbels TMD, Elliott P, Tomita M, Sato A, Suzuki C, Sugimoto M,

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Soga T, Takebayashi T (2018) Reliability of plasma polar metabolite concentrations in a large-scale cohort study using capillary electrophoresis-mass spectrometry. PLoS One 13(1):e0191230. https://doi.org/10.1371/ journal.pone.0191230 8. Gonzalez-Ruiz V, Gagnebin Y, Drouin N, Codesido S, Rudaz S, Schappler J (2018) ROMANCE: a new software tool to improve data robustness and feature identification in CE-MS metabolomics. Electrophoresis 39 (9–10):1222–1232. https://doi.org/10. 1002/elps.201700427 9. Moini M (2007) Simplifying CE-MS operation. 2. Interfacing low-flow separation techniques to mass spectrometry using a porous tip. Anal Chem 79(11):4241–4246. https://doi. org/10.1021/ac0704560 10. Gahoual R, Busnel JM, Beck A, Francois YN, Leize-Wagner E (2014) Full antibody primary structure and microvariant characterization in a single injection using transient isotachophoresis and sheathless capillary electrophoresistandem mass spectrometry. Anal Chem 86 (18):9074–9081. https://doi.org/10.1021/ ac502378e 11. Gulersonmez MC, Lock S, Hankemeier T, Ramautar R (2016) Sheathless capillary electrophoresis-mass spectrometry for anionic

metabolic profiling. Electrophoresis 37 (7–8):1007–1014. https://doi.org/10.1002/ elps.201500435 12. Ramautar R, Shyti R, Schoenmaker B, de Groote L, Derks RJ, Ferrari MD, van den Maagdenberg AM, Deelder AM, Mayboroda OA (2012) Metabolic profiling of mouse cerebrospinal fluid by sheathless CE-MS. Anal Bioanal Chem 404(10):2895–2900. https:// doi.org/10.1007/s00216-012-6431-7 13. Zhang W, Gulersonmez MC, Hankemeier T, Ramautar R (2016) Sheathless capillary electrophoresis-mass spectrometry for metabolic profiling of biological samples. J Vis Exp (116). https://doi.org/10.3791/54535 14. Ramautar R (2018) Sheathless capillary electrophoresis-mass spectrometry for the profiling of charged metabolites in biological samples. Methods Mol Biol 1738:183–192. https://doi.org/10.1007/978-1-4939-76430_12 15. Zhang W, Guled F, Hankemeier T, Ramautar R (2019) Utility of sheathless capillary electrophoresis-mass spectrometry for metabolic profiling of limited sample amounts. J Chromatogr B Analyt Technol Biomed Life Sci 1105:10-14. https://doi.org/10.1016/j. jchromb.2018.12.004

Part V Applications in Paediatrics

Chapter 12 Device Fabrication and Fluorescent Labeling of Preterm Birth Biomarkers for Microchip Electrophoresis Anna V. Nielsen and Adam T. Woolley Abstract An unmet need exists for a clinical diagnostic to determine preterm birth (PTB) risk. Such an assessment is possible with high sensitivity and specificity using a panel of nine biomarkers. An integrated microfluidic analysis system for these biomarkers is being developed which includes microchip electrophoresis (μCE) separation. A t-shaped microchip device can be used to test the μCE portion of this integrated system to find appropriate separation conditions. These t-shaped microchips can be fabricated using photolithographically patterned Si templates and hot embossing. PTB biomarkers can be fluorescently labeled using an amine-reactive dye prior to μCE. The μCE conditions established using this t-shaped device should be useful in developing a complete integrated microfluidic system for PTB risk assessment. Key words Microfluidics, Point-of-care diagnostics, Rapid analysis, Photolithography, Cyclic olefin copolymer, Hot embossing, Laser induced fluorescence

1

Introduction Defined as birth before the 37th week of gestation, preterm birth (PTB) affects about 10% of infants born worldwide annually [1]. This high rate of early births has made PTB the leading cause of infant mortality and birth-related complications. The average cost to care for a premature infant in 2013 was over $54,000 compared to $4400 for a full term infant [2]. One of the primary reasons for these statistics is that no early clinical diagnostic is available to determine PTB risk. If such an assessment could be realized, it might be possible to develop treatments against the onset of early labor. A panel of nine biomarkers has been discovered which allows for PTB risk diagnosis with high sensitivity and specificity [3]. Using this biomarker panel, an integrated microfluidic PTB risk assessment is being developed [4–7], which includes microchip electrophoresis (μCE) separation of these biomarkers. The μCE portion of this integrated device can be evaluated using a t-shaped

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microchip and fluorescently labeled biomarkers. Using this system, we have separated six PTB biomarkers [7]. Here, we describe the fabrication of these microchips and the process for fluorescently labeling PTB biomarkers for μCE.

2

Materials Prepare all solutions in ultrapure water (18 MΩ-cm). All chemicals should be reagent grade unless otherwise specified.

2.1 μCE Device Fabrication

1. 400 p-type Si wafers with orientation and ~500 nm oxide surface layer. 2. S1805 photoresist and MF-26A developer (MicroChem; Westborough, MA) or another similar photoresist and developer pair. 3. 48% buffered hydrofluoric acid (HF) etching solution (see Note 1). 4. 40% potassium hydroxide (KOH) etching solution. Prepare by adding 160 g KOH pellets to 240 mL water in a plastic (not glass) container (see Note 2). 5. Isopropyl alcohol (IPA). 6. Acetone. 7. Hotplate set to 110  C. 8. 1- and 2-mm-thick Zeonor 1060R cyclic olefin copolymer (COC) sheets (Zeon Chemicals, Louisville, KY) or similar COC plates cut into 22  50 mm pieces (see Note 3). 9. Photografting solution: 4% poly(ethylene glycol) diacrylate (PEGDA; MW 575) and 1% benzoin methyl ether (BME) in a 50–50 methanol-water mixture. Prepare by combining 5 mg BME, 20 mg PEGDA, 250 mg methanol, and 250 mg water. 10. Glass microscope slides (75  50  1 mm), copper blocks (75  50  5 mm), and C-clamps (1 or 2 in.) for hot embossing, flattening, and bonding (see Note 4). 11. Oven set to 110  C or 138  C. 12. Drill press and 2 mm diameter drill bits.

2.2 Fluorescent Labeling of Biomarkers

1. Two or more of the PTB biomarkers listed in Table 1. Amino acid sequences for Peptides 1–3 are given in Table 2 (see Note 5). 2. Labeling buffer: 10 mM bicarbonate buffer, pH 10. Prepare with 42 mg NaHCO3 in 50 mL water. Adjust to pH 10 using 1 M NaOH. 3. Fluorescein 5-isothiocyanate (FITC) or a similar aminereactive fluorescent dye. Make a 10 mM solution by dissolving 0.97 mg FITC in 250 μL dimethyl sulfoxide.

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Table 1 Panel of nine PTB biomarkers, their molecular weights, fluorescent dye-to-biomarker ratios during labeling, and biomarker concentrations for labeling

PTB biomarker

Dye-to-biomarker Biomarker concentration Mass (kDa) labeling ratio for labeling (μM)

PTB Peptide 1

2.0

1:1

500

Corticotropin-releasing factor

2.7

1:1

500

PTB Peptide 2

4.2

3:2

500

PTB Peptide 3

4.2

3:2

500

Defensins

3–6

3:2

250

Tumor necrosis factor-α receptor Type 1 26

10:1

35

Lactoferrin

80

15:1

50

Thrombin-antithrombin III

95–110

20:1

30

Ferritin

470

30:1

10

Table 2 Amino acid sequences for three PTB peptides PTB Peptide

Amino acid sequence

Peptide 1

QLGLPGPPDVPDHAAYHPF

Peptide 2

NVHSAGAASRMNFRPGVLSSRQLGLPGPPDVPDHAAYHPF

Peptide 3

NVHSAGAASRM(O)NFRPGVLSSRQLGLPGPPDVPDHAAYHPF

4. Centrifugal filters: 3, 10, 30, and 50 kDa cutoff values are recommended. 5. Sample buffer: 1 mM bicarbonate, pH 9. Prepare with 4.2 mg NaHCO3 in 50 mL water. Adjust to pH 9 using 1 M NaOH. 2.3

μCE Analysis

1. Running buffer: 100 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 8. Prepare with 1.19 g HEPES in 50 mL water. Adjust to pH 8 using 1 M NaOH. 2. Instrument: A home-built μCE system composed of a 488 nm laser, inverted microscope with wavelength-appropriate filter set, photomultiplier tube detector and power supply, analogto-digital converter, and computer with data-recording software [7]. 3. Two high voltage power supplies and four Pt-wire electrodes (see Note 6).

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Methods

3.1 μCE Device Preparation 3.1.1 Silicon Template Patterning

1. Clean and dry an oxidized Si wafer with acetone and IPA on a spinner at 2000 rpm (200  g). Bake on the hot plate for 3–5 min to evaporate residual solvent. 2. Apply ~2 mL of S1805 photoresist onto the center of the Si wafer, then spin at 3000 rpm (500  g, with 500 rpm/s acceleration) for 1 min. Bake the wafer on the hotplate for 1 min (see Note 7). 3. Expose the coated wafer with a 350 W mercury lamp through a chrome-glass mask similar to the one shown in Fig. 1 for 8 s (see Note 8). Bake the wafer on the hotplate for 5 min. 4. Submerge the wafer in MF-26A developer for 60 s or until the exposed photoresist has completely dissolved. Swirl the solution every ~5 s for more uniform developing across the surface. Rinse with water to remove excess developer and dry. Bake the wafer on the hotplate for 5 min (see Note 9). 5. Submerge the wafer in HF etch, perturbing the solution every ~10 s, for 5 min or until the oxide surface is completely etched (see Note 10). Rinse thoroughly with water and dry. 6. Clean off any remaining photoresist with acetone and IPA. Cut the wafer into individual t-shaped device templates. 7. Submerge the templates in KOH etching solution at 70  C for about 25 min to etch the Si surface until there is a 15–20 μm height difference between the Si surface and the oxide pattern. Rinse with water and dry.

3.1.2 COC Device Fabrication

1. Clean and dry two glass microscope slides, a Si device template, and a piece of 1-mm-thick COC. Make a stack in the following order: copper block, glass slide, Si template, COC, glass slide, and copper block (Fig. 2a). Make sure to keep the template and COC aligned so that the t-shaped pattern will transfer into the COC. 2. Use 4–6 C-clamps to clamp the stack from Step 1 together (Fig. 2b, c). Hot emboss the piece by heating in an oven at 138  C for 25 min. 3. Using a Si template aligned with a 2-mm-thick piece of COC, mark the positions of the four reservoirs on the COC. Use a drill press to drill holes through the COC in these locations. 4. Flatten the 2 mm COC piece by making another stack similar to the one described in step 1 except without the Si template: copper, glass, COC, glass, and copper. Clamp the pieces together and heat in an oven at 110  C for 20 min. 5. Allow both stacks time to cool (~30 min), then release the clamping.

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Fig. 1 Mask design for t-shaped device templates on a Si wafer. All channels are 20 μm wide. The injection channel is 12 mm long, and the separation channel is 4 cm long, intersecting the injection channel 5 mm down from the top reservoir. Reservoirs have a 1 mm radius. Each template is designed to be 22  50 mm, which allows for 5 templates per 400 wafer. The wafer alignment circle is shown here but is not included on the printed mask

Fig. 2 Stack arrangement used for hot embossing, flattening, and bonding of COC microfluidic devices. (a) Cross section of stack. The Si template shown should be used during hot embossing but not during flattening. During bonding, the Si template and COC piece should be replaced by the embossed and flattened COC pieces. (b) Top view of the stack. (c) Photograph of a stack after clamping

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6. Create a final COC stack using copper blocks, glass slides, and the two previous pieces of COC. The COC pieces should be aligned such that the patterned reservoirs on the 1-mm-thick piece are aligned with the holes in the 2-mm-thick piece. Clamp them together and bond them by baking at 110  C for 25 min (see Note 11). 3.1.3 Photografting

1. Fill both channels and all four reservoirs in a t-shaped device with photografting solution. Place tape over or otherwise block the reservoirs to prevent evaporation. 2. Turn the device upside down and expose it to 25 mW of UV light from a mercury lamp for 12 min. 3. Remove the tape from or unblock the reservoirs and aspirate the excess photografting solution. Rinse out all reservoirs and channels three times with water (see Note 12).

3.2 Fluorescent Labeling of PTB Biomarkers

1. Dissolve each biomarker in labeling buffer to the concentration indicated in Table 1 in a microcentrifuge tube with a total volume of 20–200 μL (see Note 13). 2. Add FITC to each biomarker tube such that the dye-to-biomarker molar ratio matches the values in Table 1. Wrap the tube in aluminum foil or another material to prevent light exposure. 3. Incubate the biomarker with the FITC solution 1 h to overnight at room temperature in a dark location (see Note 14). 4. Use a centrifugal cutoff filter to remove unattached dye after labeling. The cutoff molecular mass for the filter should be less than that of the biomarker; however, larger cutoff values allow the dye to be removed more easily (see Note 15). Filter each biomarker about 4 times for 15 min at 12,000 rpm (8000  g), refilling the filter with sample buffer each time. 5. Serially dilute the filtered biomarker samples by factors of 10 down to the low-nanomolar range in sample buffer. Store samples covered at 4  C.

3.3

Performing μCE

1. Rinse a t-shaped microdevice three times with running buffer, then fill all channels and reservoirs with running buffer. Each reservoir should hold ~10 μL of buffer. 2. Place the filled device on the microscope stage and secure in place to prevent unwanted movement. Place each electrode into the corresponding reservoir (Fig. 3a). 3. Remove buffer from the sample reservoir, then refill with ~10 μL of fluorescent sample. 4. Focus the laser 1–2 cm down the separation channel from the intersection. Apply the injection voltages for 30–90 s (Fig. 3b).

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Fig. 3 μCE device and electrode setup for voltage application. (a) Photograph of device and Pt electrode placement. (b) Schematic for injection voltage application. (c) Schematic for separation voltage application. Voltages other than those pictured here (500 and 1500 V) may be used, but the injection voltage should be lower than the separation voltage. Arrows indicate the flow direction of the green sample

Fig. 4 Representative μCE result obtained using the described experimental conditions

5. Switch to the separation voltages (Fig. 3c) and simultaneously begin recording data on the computer. Results such as those shown in Fig. 4 should be obtained. Turn off the voltages when data collection is completed (see Note 16). 6. Rinse out the device reservoirs and channels three times with running buffer. The device can then be filled with running buffer and used again. 7. When finished, rinse out the device reservoirs and channels. Vacuum out the liquid and store the microchip in a dry location.

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Notes 1. Extreme caution should be used when handling HF etching solution. HF is highly corrosive and toxic. Proper personal protective equipment should be used and precautions taken to prevent exposure during use. As HF is an SiO2 etchant, it should be stored in plastic, not glass, containers. 2. KOH is corrosive, and appropriate personal protective equipment should be used when preparing and handling KOH solutions. KOH is hygroscopic, so pellets should be weighed immediately before use. Pellets should be dissolved in water in a plastic, not glass, container to prevent etching of the container. The dissolving process is exothermic and should be performed in a fume hood. When heating the KOH solution for etching, the KOH container should be placed in a secondary water bath container on the hot plate and should be stirred constantly to prevent bumping. 3. Poly(methyl methacrylate) (PMMA) is an alternative material to COC for μCE. However, since the properties of PMMA are different from COC, a few adjustments in procedures should be made. In Subheading 3.1.2, adjust the baking times in steps 2, 4, and 6 to 30, 28, and 32 min, respectively. Additionally, the photografting solution in Subheading 2.1, step 9, should be 5% PEGDA and 1% 2,20 -azobis(2-methylpropionamidine) dihydrochloride initiator in water. The photografting exposure time in Subheading 3.1.3, step 2, should be lowered to 10 min. 4. Alternatively, μCE devices can be fabricated using 3D printing [8, 9] or purchased commercially. 5. Some PTB biomarkers separate more easily in μCE than others. Peptides 2 and 3 have nearly identical electrophoretic mobilities, and are thus very challenging to separate. Defensins yield several peaks in μCE that may interfere with other biomarkers. Finally, the thrombin-antithrombin III complex is not available commercially; the complex must be formed synthetically from its respective components. 6. Caution should be used with high voltages. All electrodes should be treated as live electrodes until it is confirmed that power is off. To prevent accidental injuries, power supplies should be set to trip when the electrical current exceeds a set limit; typical μCE currents under these conditions are 200 rpts]

8

8

9

200

4

Full-Mutation Female [9+9+9=29 / >200 rpts]

4

200

9 9

700bp

1000bp

700bp

1000bp

Fig. 2 Representative Capillary Electropherogram Profiles of Normal, Premutation, and Full-Mutation Samples. X-axis represents fragment length in base pairs or bp and Y-axis represents Relative Fluorescence Units (RFUs). Red peaks in electropherograms are from the ROX-labeled internal size ladder. The CGG repeat size and AGG interruption pattern of FMR1 alleles are indicated on top of each electropherogram in square brackets. “+” sign indicates the presence of an AGG interruption. The stutter pattern characteristic to expanded FMR1 alleles are observed in premutation and full-mutation samples. Numbered arrows indicate the number of CGG repeats. Inset panels in premutation and full-mutation samples show the zoomed in view of the amplicon peaks. Full-mutation male and female samples had stutter peaks extending beyond 200 CGG repeats and a full-length peak around 1050 bp

4

Notes 1. Repeated freezing and thawing of the reagents may affect the amplification efficiency of TP-PCR. Reduce the number of freeze-thaw cycles by preparing smaller aliquots of primers, dNTPs, and genomic DNAs and storing them in multiple 0.6 mL microcentrifuge tubes. Genomic DNA extracted from a wide variety of sources (peripheral blood, lymphoblastoid cell lines, saliva, buccal swab, and dried blood spots) can be tested using the described TP-PCR protocol. However, genomic DNA specimens should be quantified and checked for impurities before running the TP-PCR assay. Presence of impurities in the extracted DNA and using DNA at concentrations below the recommended 100 ng may affect the electropherogram pattern and/or the Relative Fluorescence Units (RFUs) of the generated amplicon peaks. If a significant drop in RFUs or an irregular

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electropherogram pattern is observed due to impurities in DNA, perform phenol-chloroform purification of the DNA and repeat the TP-PCR experiment with purified DNA specimen and freshly prepared reagents. 2. We highly recommend including at least two positive controls (fragile X samples with a FM expansion) and a negative control (or blank with water or 1 x TE buffer added instead of genomic DNA) in each TP-PCR experiment. While the positive controls will inform if the assay conditions are optimal for detecting and sizing expanded full-mutation alleles, the negative controls will expose any underlying contaminations. 3. Please ensure that the reagents are used at the recommended final concentrations (column 2 of Table 2). Also note that the ratio of dGTP and dCTP to dATP and dTTP is 5:1 and that the TP primer is supplied at a 1000-fold dilution compared to the Fam-F and Tail primers. Pipetting errors that alter the final concentrations of the TP-PCR reagents may affect the electropherogram pattern and the ability of TP-PCR assay to size expanded FMR1 alleles. Ensure that the pipettes are serviced and calibrated to avoid inaccurate pipetting and suboptimal performance of the TP-PCR assay. 4. Rare deletions and point mutations that account for nearly 1% of the fragile X cases will not be detected by this assay that targets the analysis of FMR1 CGG repeat expansions. 5. Polymorphism within the annealing position of the locusspecific primer may result in the failure to amplify expanded allele and generate false-negative results. 6. Due to the overlapping amplicon peaks from two FMR1 alleles in heterozygous females, determination of the exact number and pattern of AGGs in samples with complex interruption patterns can be difficult. References 1. Yu TW, Berry-Kravis E (2014) Autism and fragile X syndrome. Semin Neurol 34:258–265 2. Hill MK, Archibald AD, Cohen J, Metcalfe SA (2010) A systematic review of population screening for fragile X syndrome. Genet Med 12:396–410 3. Monaghan KG, Lyon E, Spector EB, erican College of Medical G, Genomics (2013) ACMG Standards and Guidelines for fragile X testing: a revision to the disease-specific supplements to the Standards and Guidelines for Clinical Genetics Laboratories of the American College of Medical Genetics and Genomics. Genet Med 15:575–586

4. Saul RA, Tarleton JC (1993) FMR1-related disorders. In: Adam MP, Ardinger HH, Pagon RA, Wallace SE, LJH B, Stephens K, Amemiya A (eds) GeneReviews®. University of Washington, Seattle, WA 5. Usdin K, Hayward BE, Kumari D, Lokanga RA, Sciascia N, Zhao XN (2014) Repeatmediated genetic and epigenetic changes at the FMR1 locus in the Fragile X-related disorders. Front Genet 5:226 6. Willemsen R, Levenga J, Oostra BA (2011) CGG repeat in the FMR1 gene: size matters. Clin Genet 80:214–225

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7. Tassone F et al (2012) FMR1 CGG allele size and prevalence ascertained through newborn screening in the United States. Genome Med 4:100 8. Maenner MJ et al (2013) FMR1 CGG expansions: prevalence and sex ratios. Am J Med Genet B Neuropsychiatr Genet 162B:466–473 9. Seltzer MM, Baker MW, Hong J, Maenner M, Greenberg J, Mandel D (2012) Prevalence of CGG expansions of the FMR1 gene in a US population-based sample. Am J Med Genet B Neuropsychiatr Genet 159B:589–597 10. Jin P, Warren ST (2000) Understanding the molecular basis of fragile X syndrome. Hum Mol Genet 9:901–908 11. Nolin SL et al (2003) Expansion of the fragile X CGG repeat in females with premutation or intermediate alleles. Am J Hum Genet 72:454–464 12. Yrigollen CM, Durbin-Johnson B, Gane L, Nelson DL, Hagerman R, Hagerman PJ, Tassone F (2012) AGG interruptions within the maternal FMR1 gene reduce the risk of offspring with fragile X syndrome. Genet Med 14:729–736 13. Fernandez-Carvajal I, Lopez Posadas B, Pan R, Raske C, Hagerman PJ, Tassone F (2009) Expansion of an FMR1 grey-zone allele to a full mutation in two generations. J Mol Diagn 11:306–310 14. Tassone F (2015) Advanced technologies for the molecular diagnosis of fragile X syndrome. Expert Rev Mol Diagn 15:1465–1473 15. Nolin SL, Glicksman A, Ding X, Ersalesi N, Brown WT, Sherman SL, Dobkin C (2011) Fragile X analysis of 1112 prenatal samples from 1991 to 2010. Prenat Diagn 31:925–931 16. Nolin SL et al (2013) Fragile X AGG analysis provides new risk predictions for 45-69 repeat alleles. Am J Med Genet A 161A:771–778 17. Chen L et al (2010) An information-rich CGG repeat primed PCR that detects the full range of fragile X expanded alleles and minimizes the need for southern blot analysis. J Mol Diagn 12:589–600 18. Chen L et al (2011) High-resolution methylation polymerase chain reaction for fragile X analysis: evidence for novel FMR1 methylation patterns undetected in Southern blot analyses. Genet Med 13:528–538

19. Filipovic-Sadic S et al (2010) A novel FMR1 PCR method for the routine detection of low abundance expanded alleles and full mutations in fragile X syndrome. Clin Chem 56:399–408 20. Hantash FM et al (2010) Qualitative assessment of FMR1 (CGG)n triplet repeat status in normal, intermediate, premutation, full mutation, and mosaic carriers in both sexes: implications for fragile X syndrome carrier and newborn screening. Genet Med 12:162–173 21. Rajan-Babu IS, Law HY, Yoon CS, Lee CG, Chong SS (2015) Simplified strategy for rapid first-line screening of fragile X syndrome: closed-tube triplet-primed PCR and amplicon melt peak analysis. Expert Rev Mol Med 17:e7 22. Rajan-Babu IS, Teo CR, Lian M, Lee CG, Law HY, Chong SS (2015) Single-tube methylation-specific duplex-PCR assay for rapid and accurate diagnosis of Fragile X Mental Retardation 1-related disorders. Expert Rev Mol Diagn 15:431–441 23. Zhou Y, Law HY, Boehm CD, Yoon CS, Cutting GR, Ng IS, Chong SS (2004) Robust fragile X (CGG)n genotype classification using a methylation specific triple PCR assay. J Med Genet 41:e45 24. Zhou Y, Lum JM, Yeo GH, Kiing J, Tay SK, Chong SS (2006) Simplified molecular diagnosis of fragile X syndrome by fluorescent methylation-specific PCR and GeneScan analysis. Clin Chem 52:1492–1500 25. Rajan-Babu IS, Chong SS (2016) Molecular correlates and recent advancements in the diagnosis and screening of FMR1-related disorders. Genes (Basel) 7(10):87 26. Warner JP, Barron LH, Goudie D, Kelly K, Dow D, Fitzpatrick DR, Brock DJ (1996) A general method for the detection of large CAG repeat expansions by fluorescent PCR. J Med Genet 33:1022–1026 27. Lyon E, Laver T, Yu P, Jama M, Young K, Zoccoli M, Marlowe N (2010) A simple, high-throughput assay for Fragile X expanded alleles using triple repeat primed PCR and capillary electrophoresis. J Mol Diagn 12:505–511 28. Fu YH et al (1991) Variation of the CGG repeat at the fragile X site results in genetic instability: resolution of the Sherman paradox. Cell 67:1047–1058

Part VI Applications in Oncology

Chapter 15 A Capillary Electrophoresis UV Detection-Based Method for Global Genomic DNA Methylation Assessment in Human Whole Blood Angelo Zinellu, Elisabetta Sotgiu, Salvatore Sotgia, and Ciriaco Carru Abstract Quantitative analysis of DNA methylation patterns is of special importance in several developmental and pathological situations. The development of simple and robust methods to assess DNA methylation is required to facilitate its measurement and interpretation in clinical practice. We describe a highly reproducible CE-UV method for the separation and detection of cytosine and methylcytosine, after formic acid hydrolysis of DNA extracted from human whole blood. Hydrolyzed samples were dried and successively dissolved with water and then injected into the capillary without sample derivatization procedures. The use of a run buffer containing 50 mmol/L BIS-TRIS propane (BTP) phosphate buffer at pH 3.25 and 60 mmol/L sodium acetate buffer at pH 3.60 (4:1, v/v) allowed a baseline analytes separation within 12 min. Precision tests indicated an elevated reproducibility with an inter-assay CV of 1.98%. Key words Capillary electrophoresis, DNA methylation, 5-Methylcytosine

1

Introduction The term “epigenetics” refers to heritable changes in gene expression and chromatin structure under exogenous stimuli that does not involve changes to the underlying DNA sequence [1]. DNA methylation occurs on cytosines that precede a guanine nucleotide or CpG sites and it is considered one of the major forms of epigenetic modifications that play an important role in gene expression and cellular differentiation [2]. This modified cytosine is designed as the fifth base of human DNA and it constitutes approximately 1% [3] of the bases in mammalian genomes. Three active DNA methyltransferases DNMT1, DNMT3a, or DNMT3b catalyze the addition of the methyl group from the methyl donor S-adenosylmethionine (SAM) to DNA cytosine to form 5-methylcytosine [4, 5]. The inclusion of a methylated cytosine in a promoter region has a regulatory effect on gene transcription since it can prevent the binding of transcription factors to DNA [6]. Methylation at the cytosine

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_15, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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residue of the CpG site is involved in gene silencing, tissue differentiation, genomic imprinting, and X chromosome inactivation. Aberrant DNA methylation is besides related to several diseases such as cancer cardiovascular disease and neurological disorders [7–9]. Therefore, due to the impact that DNA methylation alteration may have on human health, literature search for techniques able to easily detect and measure methylation DNA pattern. Among the analytical approaches for measuring total DNA methylation, acidic hydrolysis of DNA by means of formic acid followed by quantification of the resulting free nucleobases provides significant advantages in terms of speed, expense, and ease of assay. Once produced, free nucleobases can be measured by a wide variety of methods and techniques such as immunoassays [10], thin-layer chromatography [11], gas chromatography [12], capillary electrochromatography [13], capillary zone electrophoresis (CZE) [14], and in particular reversed-phase high-performance liquid chromatography [15]. However it has been described as DNA methylation values showed a relatively narrow distribution with a particularly low biological variation (interindividual CV less than 5%) [16]. It has been previously reported as the statistical power (to detect differences between groups or to evaluate correlations between variables) decreases significantly with increasing measurement imprecision, and this effect is already evident with a CV assay of 3% [17]. This implies that high measurement precision is essential to improve statistical power of studies. In this work we describe a new, simple, and easy analytical assay by CE for the detection of 5-methylcytosine and cytosine after DNA acidic hydrolysis for the evaluation of global DNA methylation degree in whole blood. The method that we proposed ensures an inter-assay CV of about 2%, as suggested for the measurement of analytes with a narrow distribution [17].

2 2.1

Materials Equipment

1. Capillary electrophoresis system (Beckman MDQ) equipped with a Diode Array Detector. The system was fitted with a 30 kV power supply with a current limit of 250 μA. 2. e-CAP uncoated capillary tubing 100 μm I.D., 375 μm O.D. 3. Eppendorf 5810 R centrifuge. 4. Vortex mixer. 5. Thermo Scientific Heraeus Pico 17 microcentrifuges. 6. Jouan speed vac RC1010. 7. Milli-Q system. 8. ino-Lab pH-meter, 9. Falc magnetic stirrer. 10. Ultrasonic Cleaner Branson 1510.

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11. Captair Toxicap 1200. 12. New classic MF balance. 13. Thermo Forma -86C ULT Freezer. 14. EDTA glass tubes. 15. Millex-GP Syringe Driven Filter Unit 0.22 μm. 16. 32 Karat Software Version 5.0. 2.2

Reagents

1. Cytidine. 2. 5-Methylcytidine 3. 99% Formic acid 4. Sodium acetate, 5. NaOH. 6. BIS-TRIS propane. 7. Acetic acid glacial. 8. 37% Hydrochloric acid 9. 1 M Phosphoric acid 10. QIAamp DNA Blood Mini Kit. 11. 0.22 μm Nylon filters (47 mm diameter).

2.3

Solutions

All solutions are prepared using ultrapure water (prepared by purifying deionized water to attain a sensitivity of 18 M Ω cm at 25
C) and analytical grade reagents. All reagents are prepared and stored at room temperature (unless indicated otherwise). 1. 0.5 M NaOH: weigh 2 g NaOH in a 50 mL beaker and add 40 mL of deionized water. Stir the solution until the NaOH is completely dissolved. Transfer the solution in a 100 mL graduated cylinder and add water to a final volume of 100 mL. 2. 0.1 M HCl: slowly add 0.821 mL of 37% HCl to 25 mL of deionized water in a 100 mL graduated cylinder. Adjust the final volume of solution to 100 mL with deionized water. 3. 50 mM BIS-TRIS propane pH 3.25: weigh 1.4115 g of BTP in a 100 mL beaker and add 90 mL of deionized water. Stir the solution until the powder is completely dissolved. Titrate the solution with 1 M phosphoric acid until pH reaches 3.25. Transfer the solution in a 100 mL graduated cylinder and add deionized water to a final volume of 100 mL (see Note 1). 4. 1 M phosphoric acid: slowly add 3.421 mL of 85% H3PO4 to 12.5 mL deionized water in a 50 mL graduated cylinder. Adjust the final volume of solution to 50 mL with deionized water. 5. 60 mM sodium acetate pH 3.60: weigh 0.6804 g of sodium acetate in a beaker and add 90 mL of water. Stir the solution until the powder is completely dissolved. Titrate the solution with acetic acid glacial until pH reaches 3.6. Transfer the

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solution in a graduated cylinder and add deionized water to a final volume of 100 mL (see Note 2). 6. CE run buffer: mix 40 mL of 50 mM BTP pH 3.25 with 10 mL of 60 mM sodium acetate pH 3.6 (see Note 3). 7. 90% Formic acid (w/w): slowly add 100 mL of 99% formic acid to 13 mL of deionized water. 8. Standard mixture for calibration curve (top calibrator: cytidine 80 μmol/L, 5-methylcytidine 4 μmol/L): separately weigh 10 mg of cytidine and 5-methylcitidine in a 1.5 mL Eppendorf tube and add 1 mL of deionized water. Thoroughly vortex-mix the solution until complete dissolution of standard. Transfer 194.6 μL of cytidine and 10.3 μL of 5-methylcytidine in a 100 mL graduated cylinder and add deionized water to a final volume of 100 mL.

3

Methods

3.1 Preparation of Calibration Curve

1. Serially dilute the top calibrator 4 times with water to obtain 200 μL of the five calibrators: (a) 80 mM cytidine and 4 μM 5-methylcytidine. (b) 40 μM cytidine and 2 μM 5-methylcytidine. (c) 20 μM cytidine and 1 μM 5-methylcytidine. (d) 10 μM cytidine and 0.5 μM 5-methylcytidine. (e) 5 μM cytidine and 0.25 μM 5-methylcytidine. 2. Transfer 100 μL of the supernatant in a 1.5 mL vial with a screw cap and dry it in a speed-vac. 3. Add 100 μL of 90% formic acid. 4. Thoroughly vortex-mix the solution. 5. Put the vial in a block heater at 130
C for 80 min. 6. After cooling vials, dry solution in a speed-vac (see Note 4). 7. Add 100 μL of water to dried samples for CE injection (see Note 5). 8. Thoroughly vortex-mix the solution before injection on CE (see Note 6).

3.2 Preparation of Biological Sample

1. Collect blood by venipuncture into vacuum collection tubes containing EDTA and centrifuge immediately at 3000  g  5 min at 4
C. 2. Extract DNA by QIAamp DNA Blood Mini Kit in accordance to the instructions of “blood and body fluid protocol” furnished from the supplier (see Note 7). 3. For CE, transfer 3 μg of extracted DNA in a 1.5 mL vial with a screw cap and proceed as in Subheading 3.1, steps 2–8.

Blood Global Genomic DNA Methylation Assessment by CE-UV

3.3

CE Method

217

1. Perform the analysis in an uncoated fused silica capillary, 100 μm I.D. and 60 cm length (50 cm to the detection window) (see Note 8). 2. Between runs capillary is rinsed for 1 min with 0.1 mM HCl and equilibrated with run buffer for 1 min. 3. Inject 240 nL of sample by hydrodynamic injection at 13.8 kPa for 5 s. 4. Carry out the electrophoretic run in a run buffer containing 50 mM BIS-TRIS propane phosphate buffer at pH 3.25 and 60 mM sodium acetate buffer at pH 3.6 (4:1, v/v), 20 kV (at normal polarity) for 12 min. Set cartridge temperature at 20
C. Detection must be performed at 282 nm wavelength. Figure 1 shows typical electropherograms of cytosine and 5-methylcytosine standard mixture (a) and whole blood DNA extracted sample (b) after formic acid hydrolysis (see Note 9).

Fig. 1 Typical electropherogram of (a) standard cytosine (1) and 5-methylcitosine (2) and (b) whole blood DNA extracted sample after formic acid hydrolysis (3 ¼ guanine)

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Notes 1. Solution must be stored at 4
C and it may be used for 4 weeks. 2. Filter through a 0.22-μm nylon filter to remove any particulate matter prior to its use. Solution must be stored at 4
C and it may be used for 4 weeks. 3. Solution must be prepared fresh each day. 4. Dried standards should be stored at 80
C for 6 months. 5. Use of an ultrasonic bath allows to facilitate sample resuspension after dry procedure. 6. Build calibration curves by plotting corrected area vs. the concentration of nucleosides standard solutions. 7. Check extracted DNA at 260 and 260/280 nm UV absorption in order to assess, respectively, DNA concentration and purity. An optical density ratio 260/280 ranging between 1.7 and 1.9 is considered indicative of acceptable purity. If necessary DNA could be further purified by 3 M sodium acetate pH 5.5 and 100% ethanol precipitation. Briefly, mix DNA with sodium acetate, so that the final salt concentration was 0.3 M, and add 2 vol of 100% cold ethanol. Store the samples at 20
C overnight, then centrifuge for 1 h at 3000  g, at 4
C. Discard the supernatant and wash the pellet with 70% cold ethanol. After centrifugation, discard the supernatant and dry the pellet in a speed-vac. Proceed as in Subheading 3.2, step 3. 8. New capillaries are rinsed for 5 min with 0.1 mmol/L HCl, 5 min with 0.5 mmol/L NaOH, and 5 min with water. Rinse cycle is repeated for three times. 9. The percentage of methylated to total cytosine was calculated using the formula: μmol methylcytosine/(μmol methylcytosine + μmol cytosine)  100.

Acknowledgements This work was supported by CNR—DSBM UNISS Flagship InterOmics (cod. PB05). References 1. Portela A, Esteller M (2010) Epigenetic modifications and human disease. Nat Biotechnol 28:1057–1068 2. Lou S et al (2014) Whole-genome bisulfite sequencing of multiple individuals reveals complementary roles of promoter and gene body

methylation in transcriptional regulation. Genome Biol 15:408 3. Dahl C, Guldberg P (2003) DNA methylation analysis techniques. Biogerontology 4:233–250

Blood Global Genomic DNA Methylation Assessment by CE-UV 4. Gruenbaum Y, Cedar H, Razin A (1982) Substrate and sequence specificity of a eukaryotic DNA methylase. Nature 295:620–622 5. Okano M, Xie S, Li E (1998) Cloning and characterization of a family of novel mammalian DNA (cytosine-5) methyltransferases. Nat Genet 19:219–220 6. Gifford CA, Meissner A (2012) Epigenetic obstacles encountered by transcription factors: reprogramming against all odds. Curr Opin Genet Dev 22:409–415 7. Jones PA, Baylin SB (2002) The fundamental role of epigenetic events in cancer. Nat Rev Genet 3:415–428 8. Muka T et al (2016) The role of epigenetic modifications in cardiovascular disease: a systematic review. Int J Cardiol 212:174–183 9. Qazi TJ et al (2018) Epigenetics in Alzheimer’s disease: perspective of DNA methylation. Mol Neurobiol 55:1026–1044 10. Piyathilake CJ et al (2000) Immunohistochemical evaluation of global DNA methylation: comparison with in vitro radiolabeled methyl incorporation assay. Biotech Histochem 75:251–258 11. Barciszewska MZ, Barciszewska AM, Rattan SI (2007) TLC based detection of methylated cytosine: application to aging epigenetics. Biogerontology 8:673–678

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12. Romerio AS et al (2005) Measurement of DNA methylation using stable isotope dilution and gas chromatography-mass spectrometry. Anal Biochem 336:158–163 13. Zhang M, El Rassi Z (1999) Capillary electrochromatography with novel stationary phases: II. Studies of the retention behaviour of nucleosides and bases on capillaries packed with octadecylsulfonated-silica microparticles. Electrophoresis 20:31–36 14. Sotgia S et al (2008) Rapid quantification of total genomic DNA methylation degree by short-end injection capillary zone electrophoresis. J Chromatogr A 21:145–150 15. Kok RM et al (2007) Global DNA methylation measured by liquid chromatography–tandem mass spectrometry: analytical technique, reference values and determinants in healthy subjects. Clin Chem Lab Med 45:903–911 16. Zinellu A et al (2017) Evaluation of global genomic DNA methylation in human whole blood by capillary electrophoresis UV detection. J Anal Methods Chem 2017:4065892 17. Teerlink T (2005) Measurement of asymmetric dimethylarginine in plasma: methodological considerations and clinical relevance. Clin Chem Lab Med 43:1130–1138

Chapter 16 Capillary Electrophoresis Analysis of Prostate-Specific Antigen (PSA) Noemi Farina-Gomez, Diana Navarro-Calderon, Angel Puerta, Monica Gonzalez, Jose´ Carlos Diez-Masa, and Mercedes de Frutos Abstract The Capillary Electrophoresis (CE) profile of isoforms (peaks) of a glycoprotein can be useful to show alterations in its posttranslational modifications (PTMs) linked to diseases. These changes can modify the electrophoretic mobility of these isoforms in a minor extent and, therefore, very reproducible CE methods are needed to detect them. In this chapter, a method for the analysis of prostate-specific antigen (PSA) by Capillary Zone Electrophoresis (CZE) with UV detection is detailed. High reproducibility in the separation of a large number of PSA isoforms is achieved by performing capillary conditioning in acid media and by using a background electrolyte (BGE) at pH 8.0 formulated with decamethonium bromide and urea. Key words Reproducibility, Capillary conditioning, Prostate-specific antigen, PSA, Isoform, Proteoform, Glycoform, Cancer biomarker

1

Introduction Glycoproteins can present different posttranslational modifications (PTMs) which give rise to several proteoforms of a given protein. Alterations in PTMs, including glycosylation, of proteins have been found to be related to diseases and namely with cancer [1, 2]. Some of these modifications in a given glycoprotein are translated into changes in its size and/or charge. Therefore, capillary electrophoresis (CE) can be an appropriate technique to monitor these changes in the glycoprotein. In this way, the CE profile of a glycoprotein has been related, for example, to vascular pathologies [3, 4]. In the case of prostate cancer (PCa), altered glycosylation of the prostate-specific antigen (PSA) has been described [5, 6]. PCa is the most common cancer in men and the second by number of deaths in men of developed countries [7–9]. The most efficient approach to fight PCa is detection, surveillance, and early intervention, if needed [10]. However, the PCa biomarker approved by the Food and Drugs Administration (FDA) and usually employed in

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_16, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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clinics (serum PSA concentration) has limited selectivity and sensitivity, and, therefore, better PCa markers need to be found. In this sense, the CE profile of PSA can be of help to show alterations in PSA glycosylation or other PTMs due to prostatic diseases. To make use of this approach, it has to be taken into account that the changes in migration of the CE peaks of the glycoprotein due to PTMs alteration can be very minor. Therefore, the CE methods used must be highly reproducible and able to provide resolution of a quite large number of isoforms. The term “isoform” in this context refers to each CE peak, which can include one or more proteoforms [11] of the glycoprotein. Capillary zone electrophoresis (CZE) methods to analyze PSA isoforms have been described in the bibliography [12]. The CZE-UV method previously developed in our laboratory used to provide very repeatable separation of 8 PSA isoforms [13]. However, later on, we observed lack of migration time repeatability and of resolution in several instances. This fact is exemplified in Fig. 1A, where the first PSA analysis performed on each one of three capillaries from the same lot is shown. The first PSA peak was not seen and migration time was different for each capillary column. In this chapter the protocol to be followed to achieve highly reproducible CE-UV separation of a large number of PSA isoforms is detailed. In this optimized protocol capillary conditioning is performed with HCl, which, as shown in Fig. 1B, provided very good repeatability in migration time and resolution for the first PSA analysis performed in three capillaries of the same lot. Resolution of a higher number of isoforms than in the precedent method [13] is achieved by modifying the BGE pH and composition [14]. The reproducibility, high resolution, and long-term performance of the method make it appropriate for its future use to find prostate cancer markers based on comparison of the CE profiles of isoforms of PSA isolated from individuals with different prostatic pathological conditions.

2

Materials Sample

Human PSA standard purified from human seminal plasma (Certified Reference Material BCR®613).

2.2 Sample Preparation

1. Milli-Q water from purification system was used for the whole procedure.

2.1

2. 2
Phosphate buffered saline (2
PBS) (see Note 2). 3. Disposable gloves (see Note 3). 4. Centrifugal filter devices with nominal Mr 10,000 cutoff membrane and 0.5 mL volume Microcon® 10 YM-10 (Millipore). 5. Centrifuge Biofuge Stratos (Heraeus Instruments).

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Fig. 1 First analysis of PSA in (A) three capillaries from the same lot conditioned with NaOH (see Note 1), (B) three capillaries from the same lot conditioned with HCl (see Subheading 3.2). BGE: 5 mM sodium tetraborate, 10 mM sodium dihydrogen phosphate, 2 mM decamethonium bromide, pH 9.0. Separation at 25 kV and 35  C. Injection of PSA (1 mg/mL) at 35 mbar for 30 s. UV detection at 214 nm. Capillary column 50 μm ID (78.5 cm total length and 70 cm effective length). Asterisk (*) denotes baseline disturbance 2.3 Capillary Column Conditioning and Capillary Electrophoresis Separation

1. Nylon filters (25 mm diameter, 0.2 μm pore size) attached to plastic syringes are used to filter all the solutions except the samples. 2. 37% HCl is used to prepare 1 M HCl and 0.1 M HCl solutions. 3. Optimum background electrolyte (BGE): 5 mM sodium tetraborate, 10 mM sodium dihydrogen phosphate, 2 mM decamethonium bromide, 3 M urea, adjusted to pH 8.0. BGE is prepared weekly from stock solutions (see Note 4). The BGE and the stock solutions are stored at 4  C. Aqueous stock solutions are as follows: 100 mM decamethonium bromide,

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100 mM sodium dihydrogen phosphate, 62.5 mM sodium tetraborate, and 6 M urea. From them, the corresponding volumes to prepare 50 mL BGE are taken. The solution pH is adjusted at 8.0 with 1 M HCl and the volume is completed to 50 mL with water in a volumetric flask (see Note 5). 4. Uncoated fused-silica capillaries (78.5 cm total length and 70 cm effective length) of 50 μm ID are from CM Scientific Limited (Silsden, UK) (see Note 6). 5. Glass inserts deactivated (0.25 mL) from Agilent Technologies are used for PSA samples of low volume. Polypropylene vials with polyurethane caps (1 mL) from Agilent are used for water, BGE, HCl, and for the waste (see Note 7). 6. An Agilent G7100 CE system equipped with a UV–Vis diodearray detector is used. System control and data collection are carried out by 3D-CE ChemStation software (Agilent Technologies).

3

Methods

3.1 Sample Preparation

1. Allow the ampoule of lyophilized standard of PSA to equilibrate for one hour at room temperature prior to opening. 2. Open the ampoule. 3. Prepare 1 mg/mL PSA solution by adding 71 μL of water to the 71 μg of PSA standard in the ampoule. 4. Gently shake the ampoule for approximately 5 min. 5. Aliquot the PSA solution in 10 μL fractions using long tips for the pipette, for example, Gel saver II tips from Starlab. Store them in 0.65 mL low binding tubes from Sorenson BioScience at 20  C until use. 6. This standard sample does not contain additives. For other PSA samples which contain salts, they must be removed before performing CE analysis. To this aim, the solvent of the sample is exchanged to water using a 10 kDa cutoff centrifuge filter device previously passivated (see Note 8). The procedure is as follows: (a) allow the centrifuge to cool down at 4  C, (b) centrifuge the sample at 14000
g for 35 min at 4  C, (c) after that, rinse the PSA retained on the membrane of the filter device by centrifuging 3
0.3 mL of water at 14000
g for 35 min at 4  C, (d) to concentrate the sample, perform additional centrifugation without adding water at 14000
g for 40 min at 4  C, (e) when there is almost no liquid on the membrane, set the device in the recovery mode (upside down) and centrifuge it at 1000
g during 3 min at 4  C to recover the sample in the minimum volume, that usually is about 7–10 μL. As it can be

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225

Fig. 2 Effect of the presence of salts in the PSA sample on the CE profile. (a) PSA standard directly diluted on water (PSA concentration 1 mg/mL). (b) PSA standard diluted 1:1 in 2
PBS (PSA concentration after dilution 0.5 mg/mL). (c) PSA standard after removal of the PBS previously added (PSA concentration 1.3 mg/mL). Capillary conditioned with HCl. BGE: 5 mM sodium tetraborate, 10 mM sodium dihydrogen phosphate, 2 mM decamethonium bromide, 3 M urea, pH 8.0. Separation at 25 kV and 35  C. Rest of conditions as in Fig. 1

seen in Fig. 2, efficient removal of salts in this type of samples allows achieving the CE profile of PSA, which was precluded to be observed by the presence of salts. 3.2 Capillary Column Conditioning (See Note 9)

1. Conditioning of a brand new capillary. Before using a capillary for the first time carry out an initial conditioning, which includes initial preconditioning (see Subheading 3.2, step 2) and stabilization cycles (for the steps in a stabilization cycle see Subheading 3.2, step 3). Carry out the stabilization cycles until relative standard deviation (RSD) for migration time of the electroosmotic flow (EOF) marker (tEOF) for three consecutive injections is lower than 0.5%. The signal of water is used as the EOF marker (see Note 10). Fig. 3 shows an example of the 21 electropherograms obtained for water during the stabilization cycles of a brand new capillary and its migration time (tEOF) along the number of stabilization cycles. In this example RSD of tEOF for the analyses number 19, 20, and 21 is 0.40%. 2. Initial preconditioning. Flush the following liquids through the capillary at 1 bar: 1 M HCl (30 min), water (5 min), 0.1 M HCl (5 min), water (5 min), and BGE (30 min). 3. Stabilization cycles. Each stabilization cycle corresponds to one analysis and one between-analyses conditioning (see Subheading 3.2, step 4). For the analysis, water is used as sample and the CE method is the same one used for PSA analysis (see Subheading 3.3) (see Note 11).

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Fig. 3 Variation of the migration time of the EOF marker (water) along the number of stabilization cycles. (A) Electropherograms of the water peak in the 21 analyses needed to stabilize the capillary; the number on top of each peak indicates the stabilization cycle number. (B) Plot of the tEOF versus the number of stabilization cycles. Injection of water at 35 mbar for 30 s. Rest of conditions as in Fig. 2

4. Between-analyses conditioning. Flush the following liquids through the capillary at 1 bar: water (5 min), 1 M HCl (1 min), water (5 min), and BGE (5 min). 5. Daily conditioning. Flush the following liquids through the capillary at 1 bar: 1 M HCl (2 min), water (5 min), and BGE (30 min).

CE of PSA

3.3 Capillary Electrophoresis Separation

227

1. At the beginning of every day, carry out the daily conditioning (see Subheading 3.2, step 5). 2. Inject the sample at the anodic end of the capillary at 35 mbar for 30 s (see Note 12). 3. Perform separation at 25 kV and 35  C with the UV detection at 214 nm. 4. Before injecting the next sample perform the between-analyses conditioning, as indicated in Subheading 3.2, step 4. 5. Renew the BGE vials for each analysis (see Note 13). 6. At the end of the day the capillary is stored with water inside by rinsing it for 5 min at 1 bar. 7. From the electropherogram, one can calculate, for each PSA isoform, relative migration time (tm/tEOF), effective electrophoretic mobility (μeff), percentage of corrected peak area (% Acorr) and resolution with the adjacent peak (see Note 14). The separation of 10 PSA isoforms with very good precision achieved by this optimized CE method is illustrated by the migration time, percentage of corrected area and resolution values and their intra-day and inter-day RSD (%) values shown in Tables 1 and 2. 8. Clean the electrodes and pre-punchers of Agilent G7100 CE system weekly (see Note 15).

4

Notes 1. Capillary conditioning with NaOH was performed as follows: the brand new capillary was rinsed at 1 bar with 1 M NaOH (30 min), water (5 min), 0.1 M NaOH (15 min), water (5 min), and BGE (30 min); after this initial preconditioning several stabilization cycles were carried out until the RSD value (n ¼ 3) of the migration time of the EOF marker was lower than 0.5%. Each stabilization cycle consisted of one injection of water and one between-analyses conditioning. For betweenanalyses conditioning the capillary was rinsed at 1 bar with water (5 min), 0.1 M NaOH (10 min), water (5 min), and BGE (5 min). The BGE was 5 mM sodium tetraborate, 10 mM sodium dihydrogen phosphate, 2 mM decamethonium bromide at pH 9.0. 2. PBS consists of 0.01 M disodium hydrogenphosphate/sodium dihydrogenphosphate, 0.138 M sodium chloride, 2.7 mM potassium chloride, pH 7.4. Ten times concentrated PBS (10
PBS) was prepared and from it 2
PBS was obtained daily by diluting 1 part of 10
PBS with 4 parts of water. 10
PBS was made as follows: a solution A consisting of 0.1 M Na2HPO4, 1.38 M NaCl, and 27 mM KCl and a solution B

14.45

15.33

15.66

16.32

16.68

17.22

17.82

18.37

18.87

20.19

1

2

3

4

5

6

7

8

9

10

0.44

0.36

0.41

0.40

0.40

0.32

0.38

0.23

0.35

0.34

0.34

3.43

a

5.08

1.07

1.95

1.77

1.79

3.55

0.02

8.19

1.70

2.37

1.29

4.45

9.23

6.42

8.12

55.80

1.12

11.41

1.09

RSD ¼ Relative standard deviation

12.70

RSD Average %a

RSD Average %a

20.20

18.90

18.39

17.84

17.24

16.70

16.35

15.66

15.34

14.45

12.71

0.10

0.08

0.09

0.08

0.08

0.08

0.07

0.09

0.08

0.06

0.05

RSD Average %a

tm

% Acorr

tm

EOF

Isoform

Day 2

Day 1

Intra-day n ¼ 3

1.50

1.31

4.22

9.85

5.96

8.04

55.33

1.20

11.26

1.32

8.40

4.77

1.67

1.77

1.52

1.23

0.22

2.51

0.97

6.72

RSD Average %a

% Acorr

20.36

19.02

18.52

17.95

17.36

16.81

16.45

15.76

15.44

14.54

12.79

0.24

0.22

0.21

0.20

0.18

0.21

0.20

0.19

0.17

0.16

0.15

RSD Average %a

tm

Day 3

1.13

1.18

4.26

9.08

6.01

7.84

56.41

1.22

11.57

1.28

6.72

9.00

1.14

2.52

2.87

1.53

0.93

7.07

1.57

6.05

RSD Average %a

% Acorr

20.25

18.93

18.43

17.87

17.27

16.73

16.37

15.69

15.37

14.48

12.73

0.48

0.42

0.45

0.42

0.44

0.40

0.42

0.35

0.39

0.38

0.37

RSD Average %a

tm

Inter-day n ¼ 3

Table 1 Intra- and inter-day precision for migration time (tm) and percentage of corrected area (% Acorr) of PSA isoforms

1.23

1.26

4.31

9.38

6.13

8.00

55.85

1.18

11.42

1.23

17.77

7.10

2.89

4.17

3.99

2.59

0.97

6.88

1.72

9.83

RSD Average %a

% Acorr

228 Noemi Farina-Gomez et al.

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229

Table 2 Intra- and inter-day precision for resolution (Rs) of each pair of consecutive PSA isoforms Intra-day n ¼ 3

Isoforms

Inter-day n ¼ 3

Day 1

Day 2

Day 3

Rs

Rs

Rs

Rs

Average

RSD%a

Average

RSD%a

Average

RSD%a

Average

RSD%a

1–2

3.44

0.72

3.28

5.19

3.34

5.61

3.35

4.33

2–3

1.16

5.88

1.11

6.71

1.10

5.55

1.13

5.82

3–4

2.33

6.06

2.34

2.80

2.29

0.97

2.32

3.49

4–5

1.18

2.51

1.15

4.39

1.18

2.25

1.17

3.04

5–6

1.57

4.84

1.63

3.98

1.64

4.81

1.61

4.38

6–7

1.74

2.23

1.70

4.03

1.73

2.79

1.72

2.87

7–8

1.74

2.28

1.75

1.87

1.77

0.46

1.75

1.68

8–9

1.62

3.16

1.65

0.68

1.67

1.12

1.65

2.21

9–10

3.56

2.24

3.39

6.31

3.68

3.67

3.54

5.16

RSD ¼ Relative standard deviation

a

consisting of 0.1 M NaH2PO4, 1.38 M NaCl, and 27 mM KCl were prepared; solutions A and B were mixed until reaching pH 6.85. Be aware that pH is modified by the dilution process. For this reason, 10 x PBS was made at pH 6.85. When 10
PBS was diluted to PBS, the final pH was 7.4. 3. Always use gloves to protect yourself and to protect the sample from the proteases which could be on your hands. 4. Decamethonium bromide and urea solutions are less stable than the rest of BGE components, for long times. For this reason, independent stock solutions are prepared for these components. The initial pH of the BGE mixture before pH adjustment must be above 8.2; otherwise, degradation of the decamethonium bromide solution should be suspected. 5. The use of this BGE formulated with 3 M urea and at pH 8.0 provides separation of up to 10 PSA peaks with higher resolution and shorter analysis time than when using a BGE of the same pH and components without urea, as it can be seen in Fig. 4. Both BGEs at pH 8.0 allow separating a higher number of PSA peaks than the BGE at pH 9.0 without urea, as shown in Fig. 1. 6. Capillaries are purchased in reels and cut to the desired length just before their use. Detection window is made by burning the polyimide coating.

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Fig. 4 Comparison of CE analysis of PSA isoforms performed with BGE (a) without and (b) with 3 M urea. Rest of conditions as in Fig. 2

7. When BGE with urea is used, urea tends to form salt deposits on the electrodes and on the pre-punchers of the CE instrument, especially at the capillary outlet. During the analyses, this precipitate can disturb the separation or/and contaminate the solutions set in the vials. In order to diminish the precipitation of urea on the outlet electrode and on the outlet pre-puncher during the capillary column conditioning, there should always be water in the waste vial (Fig. 5). In any case, special attention should be paid to clean the electrodes and the pre-punchers when BGEs containing urea are used (see Note 15). 8. Before their use, centrifuge filter devices are passivated during 12 h at +4  C by filling them with a solution of 5% (w/v) Brij® 35 to avoid any nonspecific adsorption of PSA to the device [15]. After that, discard the detergent solution and rinse 5 times the filter device with water. 9. For this CE instrument model, apply the command “wash inlet electrode” to avoid contamination of the washing solutions and of the sample, arising from the solution used in the previous step of the method. This command should be introduced in every step of the method in which the capillary inlet and the inlet electrode are changed from one vial containing a solution to a different one. If the prior vial contains water, applying the command is not necessary. This command consists of the application of slight vacuum to the inlet vial while lowering this vial so that the liquid stored in the space between the capillary and the electrode is removed. 10. The peak corresponding to water is positive in separations using the BGE which contains urea (see Figs. 2, 3, 4b and 6) and it is negative in separations using BGEs which do not contain urea (see Figs. 1 and 4a).

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231

Fig. 5 Drawing of the outlet pre-puncher A and pictures of the outlet pre-puncher after one-day running analyses with a BGE containing urea when there was no water in the waste vial (top-view B1, and view of the lateral connector B2) and when water was added in the waste vial (top-view C1, and view of the lateral connector C2)

11. The sample and the BGE vials are renewed for each new injection of water. 12. To avoid PSA degradation, keep the sample at room temperature the shortest time possible. If a cooling system for the sample tray is not installed in the CE instrument, the PSA sample has to be taken from the refrigerator and placed in the tray about 4 min before injection to allow it to reach room temperature. Similarly, the PSA sample has to be removed from the sample tray and placed in the refrigerator just after each injection has taken place. If a cooling system is installed, do not place the BGE vials in the sample tray; instead, use the replenishment system (if there is one) to avoid salts precipitation. 13. Renewal of BGE for each PSA analysis is necessary to avoid the detrimental effect of BGE electrolysis on PSA separation. This

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Fig. 6 Effect of BGE electrolysis on PSA separation. (a) First analysis and (b) second analysis of PSA using the same BGE vials. Rest of conditions as in Fig. 2

detrimental effect produces undesired change in BGE pH and decrease in resolution of the PSA peaks that take place when the same BGE vials are used for more than one analysis, as it can be observed in Fig. 6. 14. Relative migration time for each isoform is calculated as the ratio of the migration time of the isoform (tm) to the migration time of the EOF (tEOF). Effective electrophoretic mobility (μeff) is calculated as:   lL 1 1  ð1Þ μeff ¼ V t m t EOF where l and L are the effective length and the total length of the capillary, respectively, and V is the applied voltage. Corrected area for each isoform is calculated as the ratio of the peak area to the migration time. Corrected area percentage for each isoform is calculated as 100 times the ratio of the corrected area of the isoform to the sum of corrected area for all the isoforms. Resolution (Rs) between each pair of consecutive isoforms is calculated as: Rs ¼ 1:18

ðt m2  t m1 Þ ðw 1 þ w 2 Þ

ð2Þ

where wi is the peak width at their half-height and assuming that the peaks have Gaussian shape. 15. As mentioned in Note 7, the use of salts and urea-containing BGE produces salt deposits in the electrodes and pre-punchers of the Agilent G7100 CE system. These deposits can cause arching and lead to current leakage, buffer contamination, and/or carry-over. Therefore, cleaning the electrodes and pre-punchers regularly is necessary. In our experience, cleaning

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should be carried out weekly. The cleaning consists in flushing the electrodes and pre-punchers with water to remove crystals, sonicate them in water for 7 min, flush them with isopropanol, and finally use compressed air to dry the electrodes and pre-punchers. It is important to make sure that the electrodes and the pre-punchers are completely dry before reinstallation to avoid current leakage and arching in the following analyses.

Acknowledgments Financial support from the Spanish Ministry of Economy and Competitiveness (project CTQ2013-43236-R) is acknowledged. Noemi Farina-Gomez and Monica Gonzalez acknowledge the Ph. D. JAE-pre grant and the JAE-doc grant, respectively, from CSIC, co-financed by the European Social Fund (ESF). Diana NavarroCalderon acknowledges her contract in the frame of the Youth Guarantee Implementation Plans, financed by the ESF and the Youth Employment Initiative (YEI). References 1. Varki A, Kannagi R, Toole B, Stanley P et al. (2015–2017) Glycosylation changes in cancer. In: Varki A, Cummings RD, Esko JD (eds) Essentials of glycobiology, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. https://doi.org/10.1101/gly cobiology.3e.047, https://www.ncbi.nlm.nih. gov/books/NBK453023 2. Gaunitz S, Nagy G, Pohl NLB, Novotny MV (2017) Recent advances in the analysis of complex glycoproteins. Anal Chem 89 (1):389–413. https://doi.org/10.1021/acs. analchem.6b04343 3. Puerta A, Diez-Masa JC, Martin-Alvarez PJ, ˜ on J, Martin-Ventura JL, Barbas C, Tun Egido J, de Frutos M (2011) Study of the capillary electrophoresis profile of intact a-1acid glycoprotein isoforms as biomarker of atherothrombosis. Analyst 136(4):816–822. https://doi.org/10.1039/c0an00320d 4. Puerta A, Martin-Alvarez PJ, Ongay S, DiezMasa JC, de Frutos M (2013) Immunoaffinity, capillary electrophoresis, and statistics for studying intact alpha 1-acid glycoprotein isoforms as atherothrombosis biomarker. Methods Mol Biol 919:215–230. https://doi.org/ 10.1007/978-1-62703-029-8_20 5. Vermassen T, Speeckaert MM, Laumen N, Rottey S, Delanghe JR (2012) Glycosilation of prostate specific antigen and its potential diagnostic applications. Clin Chim Acta

413:1500–1505. https://doi.org/10.1016/j. cca.2012.06.007 6. Llop E, Ferrer-Batalle´ M, Barrabe´s S, Guerrero PE, Ramı´rez M, Saldova R, Rudd PM, Aleixandre RN, Comet J, de Llorens R, Peracaula R (2016) Improvement of prostate cancer diagnosis by detecting PSA glycosylation-specific changes. Theranostics 6(8):1190–1204. https://doi.org/10.7150/thno.15226 7. Torre LA, Bray F, Siegel RL, Ferlay J, LortetTieulent J, Jemal A (2015) Global cancer statistics, 2012. CA Cancer J Clin 65(2):87–108. https://doi.org/10.3322/caac.21262 8. Torre LA, Siegel RL, Ward EM, Jemal A (2016) Global cancer incidence and mortality rates and trends-An update. Cancer Epidemiol Biomark Prev 25(1):16–27. https://doi.org/ 10.1158/1055-9965.epi-15-0578 9. Jemal A, Ward EM, Johnson CJ, Cronin KA, Ma J, Ryerson B, Mariotto A, Lake AJ, Wilson R, Sherman RL, Anderson RN, Henley SJ, Kohler BA, Penberthy L, Feuer EJ, Weir HK (2017) Annual report to the nation on the status of cancer, 1975–2014, featuring survival. J Natl Cancer Inst 109:9. https://doi. org/10.1093/jnci/djx030 10. Tomasetti C, Li L, Vogelstein B (2017) Stem cell divisions, somatic mutations, cancer etiology, and cancer prevention. Science 355 (6331):1330–1334. https://doi.org/10. 1126/science.aaf9011

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11. Smith LM, Kelleher NL, The Consortium for Top Down Proteomics (2013) Proteoform: a single term describing protein complexity. Nat Methods 10(3):186–187. https://doi.org/10. 1038/nmeth.2369 12. Donohue MJ, Satterfield MB, Dalluge JJ, Welch MJ, Girard JE, Bunk DM (2005) Capillary electrophoresis for the investigation of prostate-specific antigen heterogeneity. Anal Biochem 339(2):318–327. https://doi.org/ 10.1016/j.ab.2005.01.043 13. Garrido-Medina R, Diez-Masa JC, de Frutos M (2011) CE methods for analysis of isoforms of prostate-specific antigen compatible with online derivatization for LIF detection.

Electrophoresis 32(15):2036–2043. https:// doi.org/10.1002/elps.201000524 14. Farina-Gomez N, Puerta A, Gonzalez M, Diez-Masa JC, de Frutos M (2016) Impact of capillary conditioning and background electrolyte composition on capillary electrophoresis analysis of prostate specific antigen isoforms. J Chromatogr A 1443:254–261. https://doi. org/10.1016/j.chroma.2016.03.037 15. Puerta A, Diez-Masa JC, de Frutos M (2004) Use of immunodotting to select the desorption agent for immunochromatography. J Immunol Methods 289(1–2):225–237. https://doi. org/10.1016/j.jim.2004.04.021

Chapter 17 Prostate Protein N-Glycosylation Profiling by Means of DNA Sequencer-Assisted Fluorophore-Assisted Carbohydrate Electrophoresis Tijl Vermassen, Nico Callewaert, Sylvie Rottey, and Joris R. Delanghe Abstract DNA sequencer-assisted fluorophore-assisted carbohydrate electrophoresis allows for accurate profiling of the asparagine-linked (N-) glycosylation patterns, a posttranslational modification present on many soluble and membrane proteins. This technique has been extensively tested to identify N-glycosylation patterns associated with serum proteins. Here we describe the use of DNA sequencer-assisted fluorophore-assisted carbohydrate electrophoresis to identify the N-glycosylation patterns of prostate proteins in urine. Key words Fluorophore

1

N-Glycosylation,

Carbohydrate electrophoresis, Capillary electrophoresis, Urine,

Introduction Aberrant glycosylation of proteins is a fundamental characteristic of tumorigenesis and aggressive clinical behavior. Efforts for improvement of diagnostic biomarkers in various pathologies have therefore focused on searching for disease-specific and cancer-specific protein glycoforms [1–4]. Various techniques exist to assess the N-glycan profile such as capillary electrophoresis, liquid chromatography, mass spectrometry, and lectin microarrays [5]. Extensive research using capillary electrophoresis microfluidics systems has resulted in clinically usable assays, e.g., liver pathologies [6–11]. Recently, a great deal of research has been performed into the search for novel biomarkers for prostate pathologies, in particular for prostate cancer. Assessment of the N-glycan profile related to prostate pathology can be a tremendous asset as possible diagnostic (and prognostic) biomarker [12]. Here, we describe the use of DNA sequencer-assisted fluorophore-assisted carbohydrate electrophoresis, in adaptation of the protocol described by Laroy

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3_17, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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et al. [13], which can easily allow for determination of the urine prostate protein N-glycan patterns. In short, urinary prostate protein N-glycans were released from urine using the on-membrane deglycosylation method and labeled with 8-aminopyrene-1,3,6trisulphonic acid (APTS). Subsequently, the glycans were desialyted overnight at 37
C by the addition of an in-house sialidase. Two microliters of the desialyted stock (10 μL) together with a reference maltooligosaccharide ladder were analyzed with a multicapillary electrophoresis-based ABI3130 sequencer. The peaks were further analyzed with GeneMapper version 3.7 software by normalizing peak height intensities to the total intensity of the measured peaks. Analysis of these N-glycan profiles could be of help as possible biomarker for various prostate pathologies [14–16] or to gain further insights in the tumor biology of prostate cancer [17]. Because of the strict 1:1 stoichiometry of the APTS labeling of N-glycans, this protocol facilitates straightforward quantification of the analytes. Moreover, using high-throughput DNA sequencerassisted fluorophore-assisted carbohydrate electrophoresis, up to 136 samples can be analyzed each day which makes it an ideal technique for use in routine clinical practice.

2

Materials Throughout the procedures, ultrapure water obtained via Milli-Q Integral Water Purification System is used.

2.1 N-Glycan Release from Urinary Proteins

1. Vacuum manifold for filtration plates. 2. Eppendorf microcentrifuge tubes. 3. Paper tissues. 4. Parafilm. 5. Aluminum foil. 6. 96-well PCR plate or PCR tubes. 7. Polyvinylidene fluoride (PVDF) membrane-lined 96-well filtration plate. 8. On-membrane denaturing buffer: 8 M urea, 360 mM Tris–HCl pH 8.6, 3.2 mM ethylenediaminetetraacetic acid. Store at room temperature. 9. 100% methanol. Store at room temperature. 10. 0.1 M 1,4-dithiothreitol (DTT) solution: 0.231 g DTT powder in 15 mL on-membrane denaturing buffer. Store at 4
C (see Note 1). 11. 0.1 M iodoacetic acid solution: 0.279 g iodoacetic acid powder in 15 mL on-membrane denaturing buffer. Store at 4
C (see Note 1).

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12. 1% polyvinylpyrrolidone (PVP) 360 in water. Store at 4
C (see Note 1). 13. Peptide N-glycosidase (PNGase) F working solution: 0.1 μL in-house PNGase F stock solution in 50 μL 10 mM Tris-acetate pH 8.3. Store at 4
C (see Note 2). 2.2 Labeling and Cleanup of the N-Glycans

1. SpeedVac centrifugal evaporator: essential equipment in order to evaporate the reaction mixture to dryness (vacuum). 2. APTS labeling solution: 1:1 mixture of 20 mM APTS working solution in 1.2 M of citric acid and 1 M NaCNBH3 in dimethyl sulfoxide (see Note 3). 3. 0.45 μm MultiScreenHTS-HA filter plate 4. Sephadex G10 resin. 5. 100 μL multiscreen column loader system.

2.3 Digestion of the N-Glycans

1. Exoglycosidases digestion mix: 100 mM NH4Ac pH 5 buffer, exoglycosidase (in-house or purchased), ultrapure H2O in 1:1:8 stoichiometry. Store at 4
C (see Note 4).

2.4 Analysis of the N-Glycans

1. ABI 3130 genetic analyzer: optimized to enable both DNA and N-glycan analysis. 2. MicroAmp optical 96-well reaction plate. 3. Reference maltooligosaccharide ladders (dextran and RNase B) (see Note 5, Fig. 1). 4. POP-7 polyacrylamide linear polymer. 5. Running buffer supplied by the manufacturer.

3

Methods A schematic overview of all major steps is shown in Fig. 2.

3.1 On-Membrane NGlycan Release from Urinary Proteins

1. Add 500 μL urine and 1 mL on-membrane denaturing buffer into the Eppendorf microcentrifuge tubes and mix gently. Incubate the mixture for 1 h at 50
C for total denaturation (see Note 6). 2. Wet the PVDF membrane by adding 300 μL methanol. Throughout this protocol, liquids are removed from the wells by applying a vacuum under the plate, unless specified otherwise (see Note 7). 3. Wash the PVDF membrane three times with 300 μL water and once with 50 μL on-membrane denaturing buffer. 4. Load the denaturated samples onto the membrane and apply vacuum under the plate (see Note 8).

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Fig. 1 Reference maltooligosaccharide ladders. On top a dextran ladder is depicted. Dextran is a complex branched polysaccharide and is composed of chains of varying lengths. Dextran ladders are therefore used to identify the number of structural units in each identified N-glycan. Below, a RNase B ladder is depicted. RNase B is a high-mannose glycoprotein that can be used as a positive control and to identify mannose-rich Nglycans, which are most commonly found intracellular (Figs. 3 and 4)

5. Wash the wells twice with 50 μL on-membrane denaturing buffer (see Note 9). 6. Reduce the disulfide bounds by adding 50 μL DTT solution to each well. Incubate the membrane for 1 h at 37
C (see Note 10). 7. Wash the PVDF membrane three times with 300 μL water. 8. Carboxymethylate the free sulfide groups by adding 50 μL iodoacetic acid solution to each well. Incubate the membrane for 30 min at room temperature in the dark (see Notes 10 and 11). 9. Wash the PVDF membrane three times with 300 μL water. 10. Block the unbound parts of the membrane by adding 100 μL 1% PVP 360 to each well. Incubate the membrane for 1 h at room temperature (see Notes 9 and 10). 11. Wash the PVDF membrane three times with 300 μL water (see Note 12). 12. Release the N-glycans from the glycoproteins by adding 50 μL PNGase F working solution to each well. Incubate the membrane for 3 h at 37
C (see Notes 10 and 11).

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Fig. 2 Overview of the protocol with all major steps indicated during the N-glycosylation analysis of urinary prostate glycoproteins

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13. Transfer the samples to a 96-well PCR plate or to PCR tubes (see Note 13). 14. Evaporate the mixture to dryness (vacuum) at 50
C for approximately 1 h (see Note 14). 3.2 Labeling and Cleanup of Released N-Glycan

1. Label the released N-glycans by adding 1 μL APTS labeling solution to each well. Vortex the samples, centrifuge for 10 s at 750  g, and incubate overnight at 37
C (see Notes 3 and 15). 2. Quench the reaction by adding 5 μL water to each sample. Vortex the samples and centrifuge for 10 s at 750  g. 3. Pack the wells of a MultiScreenHTS-HA filter plate with Sephadex G10 to a height of 1.2 cm by using a 100-μL Multiscreen column loader system as follows: load the first 100 μL of resin, wet it with 100 μL of water. Repeat this loading and swelling step once more. Prepare the plate by centrifuging for 10 s at 750  g and adding 100 μL to the wells. Repeat twice (see Note 16). 4. Place a PCR plate under the MultiScreenHTS-HA filter plate and load the samples into the packed wells. Elute three times, each time adding 10 μL of water and centrifuging for 10 s at 750  g (see Note 17). 5. Evaporate the mixture to dryness (vacuum) at 50
C for approximately 1 h (see Notes 14 and 18). 6. Resolve the samples in 10 μL water. 7. Repeat steps 4–5 for optimal cleanup of the labeled N-glycans. 8. Resolve the samples in 10 μL water, vortex the samples, and centrifuge for seconds at 750  g (see Note 19).

3.3 Extensive Digestion with Exoglycosidases of Labeled N-Glycans

1. Load 5 μL labeled native N-glycans onto a 96-well PCR plate. 2. Evaporate the mixture to dryness (vacuum) at 50
C for approximately 15 min (see Note 14). 3. Digest the dried samples by adding 2 μL sialidase digestion mix to each well. Centrifuge for 10 s at 750  g and incubate overnight at 37
C (see Note 15). 4. Add 8 μL water (see Note 20). 5. Load 2 μL desialyted stock onto a 96-well PCR plate (see Note 21). 6. Evaporate the mixture to dryness (vacuum) at 50
C for approximately 15 min (see Note 14). 7. Digest the dried samples by adding 2 μL exoglycosidase digestion mix to each well. Centrifuge for 10 s at 750  g and incubate overnight at 37
C (see Notes 4 and 15). 8. Add 8 μL water.

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Table 1 Equipment-specific parameters Parameter

Value

Oven temperature

60
C

Pre-run voltage

15 kV

Pre-run time

180 s

Injection voltage

1.2 kV

Injection time

16 s

Run voltage

15 kV

Run time

1000 s

In case of insufficient N-glycan yield, the analysis can be improved by increasing the injection time. This will have limited impact on the electropherogram compared to an increase of the injection voltage

3.4 Analysis of Native and/or Deglycosylated NGlycans

1. Load 2 μL of each sample onto the MicroAmp optical 96-well reaction plate (see Note 22). 2. Evaporate the mixture to dryness (vacuum) at 50
C for approximately 15 min (see Note 14). 3. Resolve the samples in 15–20 μL water. 4. Load 15 μL of the reference maltooligosaccharide ladders in two separate wells. 5. Vortex the plate and centrifuge for 10 s at 750  g (see Note 23). 6. Load the plate into the ABI 3130 genetic analyzer equipped with a standard 36-cm capillary array filled with the POP-7 polyacrylamide linear polymer. Use the running buffer supplied by the manufacturer for DNA sequencing. 7. Run the ABI 3130 genetic analyzer according to the equipment-specific parameters (Table 1). 8. Analyze the data with the GeneMapper software using the factory-provided settings for amplified fragment length polymorphism (AFLP) analysis. An overview of the extensive structural analysis of a typical sample is depicted in Figs. 3 and 4.

4

Notes 1. The DTT and 1% PVP 360 solutions are preferable made immediately before use and stored temporarily at 4
C until use. It is however also possible to store work solutions of DTT and 1% PVP 360 for some time at 20
C. The iodoacetic acid solution should always be made immediately before use and stored temporarily at 4
C in the dark until use.

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2. The volumes used to prepare the PNGase F working solution can vary depending on the number of units PNGase F in the stock solution. It is important to apply 100 PNGase F units per sample for optimal deglycosylation of the sample. Current calculations have been made starting from a PNGase F stock solution containing 1000 PNGase F units per μL. Regular review of PNGase F activity is therefore necessary to determine the right number of PNGase F units in the stock solution. 3. APTS is acquired as a powder. All the powder has to be dissolved in 191 μL 1.2 M citric acid and the 100 mM APTS stock solution has to be stored at 20
C. The 20 mM APTS working solution is obtained by diluting 1:5. During the brief moment that the 100 mM APTS stock solution is used, it has to be kept constantly on ice. Mixing the 20 mM APTS working solution with the 1 M NaCNBH3 solution must only be done prior to use of the APTS labeling solution. It is advisable to prepare both the 20 mM APTS working solution and the 1 M NaCNBH3 solution separately and to store at 4
C wrapped in aluminum foil until labeling of the N-glycans. Be advised that in acidic conditions, NaCNBH3 releases HCN gas. It is therefore crucial to work in a hood or in a well-ventilated place when preparing and adding the APTS labeling solution. 4. Type of exoglycosidase depends on type of experiment. Exoglycosidases are stored at 4
C or at 20
C. Exoglycosidase digestion mixes are freshly made prior to addition to the mixtures. Keep both the exoglycosidases and exoglycosidase digestion mixes on ice during the experiment. In case of an extensive digestion, it is also possible to add an exoglycosidase digestion mix with more than one exoglycosidase. For instance, when performing a digestion on the desialyted stock with both the α-1,3/4/6-fucosidase and β-1,4-galactosidase, the mix has to be prepared as mentioned in Subheading 3.3, step 7 in a 1:1:1:7 stoichiometry. Be cautious that some of the purchased enzymes may have their own buffer and can only be used serial. Most commonly used exoglycosidases in N-glycan sequencing are given in Table 2. 5. Dextran is a complex branched polysaccharide made of many glucose molecules and is composed of chains of varying lengths. Dextran ladders are used to identify the number of structural units in each identified N-glycan. RNase B on the other hand is a high mannose glycoprotein that can be used as a positive control and to identify mannose-rich N-glycans (see Fig. 1). 6. Denaturation can be performed for longer than 1 h and at higher temperatures. In-house experiments have shown that 1 h at 50
C are the minimum requirements for this protocol. In addition, the on-membrane N-glycan release method is most useful for volumes with up to 10 μg of glycoproteins, otherwise

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Fig. 3 Structural analysis of mannose-rich glycoproteins. X-axis depicts elution time; Y-axis represents relative fluorescence units. On top, a dextran ladder is shown to determine the amount of structural units in each N-glycan structure. A RNase B ladder is depicted below to determine which glycoforms correspond with

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the in-solution method should be used. Always add twice the volume of on-membrane denaturing buffer to the sample volume. In case of low volume samples (e.g., 10 μL), always add 50 μL on-membrane denaturing buffer. Next, pre-analytical conditions are of the utmost importance for analysis of N-glycans. Beware for albuminuria, as albumin shows high affinity for the PVDF membrane and will therefore compete with the glycoproteins for binding to the membrane. Otherwise, storage temperature of the urine specimens is crucial to obtain a correct
N-glycan profile. Optimal storage temperature is 20 C (for up to 1 week) whereas higher storage temperatures will result in incorrect N-glycan profiles (Table 3 and Fig. 5). 7. Only treat the membranes in the wells that you want to use. Seal the others with adhesive tape for 96-well plates.

Table 2 Exoglycosidases typically used in the sequencing of N-glycans Exoglycosidase

Source

Specificity

α-Sialidase

Arthrobacter ureafaciens

α-2,6 > α-2,3 > α-2,8-bound sialic acid

α-Sialidase

Recombinant from Salmonella typhimurium

α-2,3-bound sialic acid

β-1,4-Galactosidase

Recombinant from Streptococcus pneumonia

β-1,4-bound galactose

α-1,3/4/6-Fucosidase

Bovine kidney

α-1,2-, α-1,3, α-1,4 or α-1,6-bound fucose

α-1,3/4-Fucosidase

Almond meal

α-1,3 or α-1,4-bound fucose

β-N-Acetylhexosaminidase

Jack bean

β-bound N-acetylhexosamine

α-Mannosidase

Jack bean

α-1,2, α-1,3 or α-1,6-bound mannose

α-1,2-Mannosidase

Trichoderma reesei

α-1,2-bound mannose

This list is not comprehensive

ä Fig. 3 (continued) high-mannose N-glycans. After sialidase digest, a total of 13 peaks could be seen on the Nglycosylation profile (A to M). Further digestion with Trichoderma reesei α-1,2-mannosidase indicated that peak E has at least one external α-1,2-bound mannose unit. Additional digestion with Jack Bean α-mannosidase, which cleaves all α-1,2/3/6-bound mannose, reduced the number of structural units by 4 resulting in the core structure (A). This proved that peak D and peak E consisted of 4 and 5 α-bound mannose units and can therefore be identified as Man5 and Man6. Identification of all other peaks is shown in Fig. 4. Glycan symbols are those suggested by the Consortium for Functional Glycomics (http://glycomics. scripps.edu/CFGnomenclature.pdf)

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Fig. 4 Structural analysis of complex branched glycoproteins. X-axis depicts elution time; Y-axis represents relative fluorescence units. On top, a dextran ladder is shown to determine the amount of structural units in

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8. Low volumes (less than 250 μL) can be processed directly. Larger volumes require reloading onto the membrane until the entire volume has been processed. On top, it is clearly visible when the liquids have fully migrated through the PVDF membrane. It is very important to regulate the vacuum under the plate carefully so that the liquid is passed through the

Fig. 5 Effect of urine storage temperature on the N-glycosylation profile of urinary prostate proteins. Influence of pre-analytical storage temperature (20
C, 4
C and room temperature) was assessed. X-axis depicts elution time; Y-axis represents relative fluorescence units. Overall, peaks intensity of core-fucosylated structures decreased with increasing storage temperature (highlighted in red) despite for one profile (sample B at room temperature) in which the peaks intensity increased with increasing storage temperature (highlighted in green). It is therefore of the utmost importance to store urine samples at 20
C or even at 80
C in order to obtain a correct profile during N-glycosylation analysis ä Fig. 4 (continued) each N-glycan structure. A RNase B ladder is depicted below to determine which glycoforms correspond with high-mannose N-glycans. Two mannose-rich glycoforms (Man5 and Man6, blue peaks) were identified using α-mannosidases (Fig. 3). After sialidase digest, N-glycans were digested with bovine kidney α-1,3/4/6-fucosidase to remove all fucose units which identified peaks A, C, D, F, G, I, and K as structures containing one fucose unit. Next, N-glycans were further degalactosylated by use of Streptococcus pneumoniae β-1,4-galactosidase to remove all galactose units which allowed to identify the bi- (peak A to G), tri- (peak H an I), and tetraantennary structures (peak J and K). In the last enzymatic step, the basic structure (peak B) was obtained by removing all N-acetylhexosamine units by means of Jack Bean hexosaminidase, which confirms the multiantennary N-glycans and ratifies the complex N-glycosylation character of urinary prostate proteins. Identified glycosylation structures are enlisted at the bottom of the figure. Glycan symbols are those suggested by the Consortium for Functional Glycomics (http://glycomics.scripps.edu/ CFGnomenclature.pdf)

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Table 3 Coefficient of variation for different urine storage temperatures 4
C

Room temperature

Overall core-fucosylation

11.7%

30.2%

Biantennary core-fucosylation

15.3%

34.1%

Branched biantennary core-fucosylation

22.1%

39.3%

Triantennary core-fucosylation

7.2%

14.4%

Tetraantennary core-fucosylation

2.2%

5.8%

Total amount of biantennary structures

9.7%

13.0%

Total amount of triantennary structures

14.3%

14.2%

Total amount of tetraantennary structures

27.8%

43.9%

UGM

27.4%

72.7%

N-Glycosylation

ratio

High variance is observed when urine samples are stored at different temperatures (N ¼ 4)

membrane in approximately 1 minute. It is therefore possible to start with only 400 μL urine in Subheading 3.1, step 4. This will reduce the time needed to remove the liquids with sufficient yield in released N-glycans. 9. If needed, the protocol can be paused at this time and the plate can be stored overnight at 4
C if you cannot proceed immediately. At the first pause point, the PVDF membrane is washed twice with 50 μL on-membrane denaturing buffer although the liquids will not be removed after the second wash step. At the second pause point, blocking of the membrane is done overnight instead of during 1 h. Be advised that the entire on-membrane N-glycan release takes at least 7.5 h (manual work not included). 10. The DTT solution, iodoacetic acid solution, 1% PVP 360 solution, and PNGase F working solution can be prepared, or thawed if stock available, during the previous incubation step. Always make additional working solution to correct for volume loss due to pipetting error. For instance: in case of 20 samples, calculate the needed volume for 25 samples. Always place the PVDF plate in a closed plastic box together with two wet paper tissues during the incubation steps. This will prevent the sample to evaporate. 11. For the carboxymethylation step, it is required to place the PVDF plate in the dark by wrapping it into aluminum foil. On the other hand, it is required to place a parafilm on top of the PVDF plate during the release of the N-glycans to ensure minimal evaporation of the samples. In both cases, one has to be cautious not to apply pressure on top of the wells as this can

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result in migration of the reaction mixture through the PVDF membrane and subsequent loss of N-glycan yield. 12. As the membrane is now fully bound, wash steps might take longer compared to previous wash steps. 13. Most optimal and least time consuming is to transfer the samples to a 96-well PCR plate using a multichannel pipet. In case of lower number of samples, one can also opt to transfer all samples separately to PCR tubes. 14. Duration of evaporation will depend on the number of samples. Higher number of samples will result in a longer evaporation time. Evaporation time can be reduced by preheating the SpeedVac centrifugal evaporator in advance to the desired temperature. Do not heat the samples above 50
C as this will result in a degeneration of the N-glycans. Dried N-glycans can be stored at 20
C for at least 1 year. 15. It is not necessary to remove the PNGase F before APTS labeling, because its presence does not influence the electropherogram in the size range of 3–25 glucose units. However, as labeling is performed in an excess of ATPS, cleanup steps are necessary to remove all unbound ATPS molecules which will interfere in the analysis of the N-glycans. Moreover, the PCR tubes or PCR plate are preferable placed upside down during the overnight incubation. This will prevent the samples from evaporating. 16. MultiScreenHTS-HA filter plate can easily be stored at room temperature. It is therefore more practical to pack the whole plates during the digestion step of the N-glycan release. When stored plates are used, only prewet the columns that will be used during the cleanup process by adding 100 μL water and centrifuge for 10 s at 750  g. It is also advisable to always pack two MultiScreenHTS-HA filter plates as the second plate can be used during the second phase of the cleanup process and as counterbalance during centrifugation. 17. N-Glycans will be collected in the eluate by the principle of sizeexclusion chromatography. The unbound APTS molecules will migrate deep into the resin whereas the labeled N-glycans will migrate from the resin. 18. Due to the excess of unbound ATPS molecules, the eluate will have a fluorescent yellow color. This makes it easy to see whether or not the samples have dried completely. 19. Samples are now ready in their native form. At this point, Nglycans can be analyzed directly on the ABI 3130 genetic analyzer, stored at 20
C for more than 1 year or further digested for more extensive structural analysis [16].

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Fig. 6 Difference between native and desialyted N-glycosylation profile of urinary prostate proteins. X-axis depicts elution time; Y-axis represents relative fluorescence units. On top, the typical native N-glycan profile is depicted whereas below the desialyted profile of the same sample has been shown. Due to the negative charge each sialic acid unit contains, structures will elute faster from the capillary resulting in a more complex profile. Desialylation of the N-glycans results in uncharged structures which will elute from the capillary based on size of each structure, allowing for optimal structural analysis. Based on the current profile, it is clear that the majority of the complex N-glycans are secreted in their sialyted form

20. Samples are now ready in their desialyted form. At this point, desialyted N-glycans can be analyzed directly on the ABI 3130 genetic analyzer, stored at 20
C for more than 1 year or further digested for more extensive structural analysis [16]. Note that desialylation is a necessary step toward the structural analysis of the obtained N-glycans. In their native form, N-glycans can contain up to 4 negatively charged unit of sialic acid. This interferes with the electropherogram due to the shorter elution time for N-glycans making it difficult to interpret the electropherogram, especially in case of complex tetraantennary N-glycans. As all units of sialic acid units are removed during the desialylation step, interpretation of the electropherogram is simplified (Fig. 6). 21. Be cautious that one only has 10 μL of desialyted stock. In case of multiple digestions, it may be required to repeat the desialylation step again to obtain a larger stock volume. 22. Loaded samples can vary between native forms and extensively deglycosylated N-glycans, depending on the type of analysis. Make sure samples are always arranged to permit simultaneous loading of 4 capillaries. If needed other wells have to be filled with water during Subheading 3.4, step 3. 23. The plate is now ready to be analyzed by means of the ABI 3130 genetic analyzer. Plates can be stored temporarily at 4
C depending on the availability of the ABI 3130 genetic analyzer. Place the plate upside down during storage to prevent

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evaporation of the samples. Repeat Subheading 3.4, steps 2–3 after storage to make sure sufficient sample volume is present for analysis. References 1. Dwek MV, Brooks SA (2004) Harnessing changes in cellular glycosylation in new cancer treatment strategies. Curr Cancer Drug Targets 4:425–442 2. Arnold JN, Saldova R, Hamid UM et al (2008) Evaluation of the serum N-linked glycome for the diagnosis of cancer and chronic inflammation. Proteomics 8:3284–3293 3. Drake PM, Cho W, Li B et al (2010) Sweetening the pot: adding glycosylation to the biomarker discovery equation. Clin Chem 56:223–236 4. Meany DL, Chan DW (2011) Aberrant glycosylation associated with enzymes as cancer biomarkers. Clin Proteomics 8:7 5. Vanderschaeghe D, Festjens N, Delanghe J et al (2010) Glycome profiling using modern glycomics technology: technical aspects and applications. Biol Chem 391:149–161 6. Blomme B, Francque S, Trepo E et al (2012) N-glycan based biomarker distinguishing non-alcoholic steatohepatitis from steatosis independently of fibrosis. Dig Liver Dis 44:315–322 7. Callewaert N, Van Vlierberghe H, Van Hecke A et al (2004) Noninvasive diagnosis of liver cirrhosis using DNA sequencer-based total serum protein glycomics. Nat Med 10:429–434 8. Vanderschaeghe D, Guttman A, Callewaert N (2013) High-throughput profiling of the serum N-glycome on capillary electrophoresis microfluidics systems. Methods Mol Biol 919:87–96 9. Vanderschaeghe D, Laroy W, Sablon E et al (2009) GlycoFibroTest is a highly performant liver fibrosis biomarker derived from DNA

sequencer-based serum protein glycomics. Mol Cell Proteomics 8:986–994 10. Vanderschaeghe D, Szekrenyes A, Wenz C et al (2010) High-throughput profiling of the serum N-glycome on capillary electrophoresis microfluidics systems: toward clinical implementation of GlycoHepatoTest. Anal Chem 82:7408–7415 11. Verhelst X, Vanderschaeghe D, Castera L et al (2017) A glycomics-based test predicts the development of hepatocellular carcinoma in cirrhosis. Clin Cancer Res 23:2750–2758 12. Vermassen T, Speeckaert MM, Lumen N et al (2012) Glycosylation of prostate specific antigen and its potential diagnostic applications. Clin Chim Acta 413:1500–1505 13. Laroy W, Contreras R, Callewaert N (2006) Glycome mapping on DNA sequencing equipment. Nat Protoc 1:397–405 14. Vermassen T, Van Praet C, Lumen N et al (2015) Urinary prostate protein glycosylation profiling as a diagnostic biomarker for prostate cancer. Prostate 75:314–322 15. Vermassen T, Van Praet C, Poelaert F et al (2015) Diagnostic accuracy of urinary prostate protein glycosylation profiling in prostatitis diagnosis. Biochem Med (Zagreb) 25:439–449 16. Vermassen T, Van Praet C, Vanderschaeghe D et al (2014) Capillary electrophoresis of urinary prostate glycoproteins assists in the diagnosis of prostate cancer. Electrophoresis 35:1017–1024 17. Vermassen T, D’Herde K, Jacobus D et al (2017) Release of urinary extracellular vesicles in prostate cancer is associated with altered urinary N-glycosylation profile. J Clin Pathol 70:838–846

Chapter 18 Aptamer-Based Microchip Electrophoresis Assays for Amplification Detection of Carcinoembryonic Antigen Shulin Zhao Abstract Microchip electrophoresis (MCE), regarded as a miniaturized version of capillary electrophoresis (CE), has exhibited prominent advantages in terms of low sample consumption, rapid analysis times, easy operation, efficient resolution of compounds, and increased throughput. This technology has led to more research focus on analysis particularly in hospital settings for clinical diagnostics. However, since the channels in microchip are very small, achieving the desired assay sensitivity on a microfluidic platform remains a challenge. Here, we describe aptamer-based MCE assays for amplification detection of carcinoembryonic antigen (CEA) in human serum. Key words Microchip electrophoresis, Aptamer, Laser induced fluorescence detection, Carcinoembryonic antigen

1

Introduction Carcinoembryonic antigen (CEA) as one of the most widely used tumor marker is used in the clinical diagnosis of colorectal, pancreatic, gastric, and cervical carcinomas [1–4]. Meanwhile, the CEA level in serum is also related to the stage of the tumor, the outcome of therapy, and the prognosis [5], so it can be used as a marker to directly evaluate curative effects, recrudescence, and metastasis [6]. Various immunoassays, ranging from the radioimmunoassay [7], enzyme-linked immunosorbent assay [8], timeresolved fluoroimmunoassay [9], chemiluminescence immunoassay [10], and MCE-based immunoassay [11] to impedimetric immunoassay [12], were developed for CEA detection. However, the conventional immunoassay requires a relatively long assay time, and involves troublesome liquid-handling procedures and many expensive antibody reagents. Aptamers are single-stranded oligonucleotides selected in vitro by the systematic evolution of the ligand by the exponential enrichment (SELEX) process from random-sequence nucleic acids

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libraries [13]. Compared to other recognition elements, such as antibodies, aptamers have many advantages, such as simple synthesis, good stability, high affinity, excellent selectivity, and wide applicability, making them suitable candidates for biological application [14]. In order to obtain high sensitivity for the detection of trace biomarkers, aptamer-based amplification assays have been paid more and more attention [15]. Microchip electrophoresis (MCE), regarded as a miniaturized version of capillary electrophoresis (CE), has exhibited prominent advantages in terms of low sample consumption, rapid analysis times, easy operation, efficient resolution of compounds, and increased throughput. In the past decade, many academic laboratories have demonstrated the potential of MCE as a powerful analytical tool leading to the next generation in chemical separation/detection technologies [16]. This technology has led to more research focus on analysis particularly in hospital settings for clinical diagnostics [17] including the detection of disease biomarkers [18], DNA [19], RNA [20], and proteins [21]. However, since the channel in microchip for the detection was extremely small, the sensitivity of MCE methods reported for clinical analysis was in the range of 10
6–10
9 M (limit of detection, LOD), which was hardly sufficient for quantifying trace components in biological samples. Therefore, effectively detecting the separated trace analytes by MCE remains still a challenge. In this work, we combine the advantages of aptamer-based amplification, magnetic beads (MBs), and MCE techniques to develop a novel fluorescence signal amplified strategy based on MBs assisted target-induced strand circle for improved assay sensitivity in MCE-laser induced fluorescence (LIF) detection. We used CEA as the model analyte and have demonstrated the feasibility of proposed signal amplification strategy in MCE-LIF assays. The new strategy opens new avenues on the developing high sensitive MCE-LIF method and shows a potential application in early clinical diagnostics of cancer disease.

2

Materials Prepare all solutions using Milli-Q ultrapure water (18 M Ω cm at 25  C) and analytical grade reagents.

2.1 Solutions and Equipment

1. All solutions used in MCE are filtered through a 0.22 μm nylon membrane (see Note 1). 2. The sequences of oligonucleotides are 50 -NH2-ATA CCA GCT TAT TCA ATT-30 (containing an aptamer of CEA); 30 -T CGA ATA AGT TAA CGA CTC CTC GTT A-50 (complementary DNA strand); 50 -FAM-ATT GCT GAG GAG C-30 .

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Fig. 1 Dimensions and layout of the glass/PDMS microchip used in this work: S sample reservoir, B buffer reservoir, SW sample waste reservoir, BW buffer waste reservoir

3. The sequences of fluorescein amidite (FAM) labeled DNA is 50 -FAM-ATT GCT GAG GAG C-3. 4. 50 mg/mL Carboxyl-modified magnetic beads (MBs): diameter is 75 μm. 5. 10  NEB buffer solution, pH 7.9: 100 mM Tris–HCl solution containing 500 mM NaCl, 100 mM MgCl2, and 10 mM dithiothreitol. 6. 20 mM Tris–HCl buffer solution, pH 7.4: use to dilute the oligonucleotides to give a stock solution of 20 μM (20 μM 50 -NH2-ATA CCA GCT TAT TCA ATT-30 solution and 20 μM 30 -T CGA ATA AGT TAA CGA CTC CTC GTT A-50 . The solution was mixed and heated to 95  C for 3 min, and slowly cooled down to room temperature to obtain dualstranded DNA before use). 7. 20 mM phosphate buffer solution, pH 7.4 containing 0.8% hydroxypropyl methylcellulose (HPMC) and 15 mM tetrabutylammonium phosphate (TBAP) was used as the MCE running buffer. 8. Human blood samples were centrifuged at 16,000  g for 15 min to obtain serum. These samples were stored at
20  C until analysis, diluted 10 folds with 10 mM phosphate buffer solution, pH 7.4 before analysis. 9. Microchip: The design of the glass/PDMS microchip is illustrated in Fig. 1. The glass layer with microchannels (all channels etched in glass substrates were 25 μm deep and 45 μm wide) is fabricated by using a standard photolithography and wet chemical etching technique [22]. The PDMS surface for bonding to the etched glass slide is prepared from Sylgard 184 (PDMS) silicone elastomer mixed with its curing agent at 10:1 (w/w).

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Fig. 2 The MCE-LIF detection system developed by our laboratory 2.2 Setup for MCELIF Assay

1. A microfluidic chip fluid intelligent electric drive instrument with 8-channel high-voltage circuit capable of supplying 0~5000 V DC was used for the voltage output and control of the MCE. Figure 2 illustrates the setup. 2. The fluorescence signal was collected on a chromatography workstation. The general principle of the detection system is as follows. The MCE-LIF detection system was placed in a dark box to reduce background noise. The semiconductor laser (30 mW) emits 473 nm light that is reflected by a dichroic mirror and focused on the channel (detection window) of microchip by an eyepiece to excitated fluorescence of the analyte. The emitted fluorescence is collected and focused on the objective lens by the same optical system and filtered by the cutoff filter, is finally collected by a photomultiplier tube (PMT) and converted into an electrical signal that is processed by the chromatography workstation.

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3.1 Preparation of dsDNA Immobilized MBs

1. The dsDNA was immobilized on the MBs according to the procedure described by Bi et al. [23] with a slight modification. 2. A suspension of MBs (200 μL, 50.0 mg/mL) in a 5 mL eppendorf tube was separated magnetically. 3. The MBs were washed three times with MES buffer (3  300 μL), and then suspended to a final volume of 200 μL in same buffer solution. 4. A 0.2 M NHS solution (200 μL) and a 0.8 M EDC solution (200 μL) were added to the eppendorf tube, and the mixture was incubated at room temperature for 2 h to activate the carboxylate groups on the MBs. 5. The MBs were then washed three times with 10 mM phosphate buffer solution (3  300 μL), and resuspended to a final volume of 2.0 mL. 6. An aqueous solution of the dsDNA (200 μL, 6.25  10
7 M) was then added to 500 μL activated MBs solution, and the resulting suspensions were allowed to stand for 24 h at 25  C for the immobilization of the dsDNA on the surface of the activated MBs. 7. Finally, the resulting MBs-dsDNA conjugates were separated magnetically, and washed three times with 10 mM phosphate buffer solution (3  300 μL), and then resuspended in 400 μL of 10 mM phosphate buffer solution containing 1.0% BSA. 8. The solution was incubated at 25  C for 2 h to eliminate the risk of unspecific binding, separated magnetically. Then, MBs-dsDNA conjugate was washed three times with 10 mM phosphate buffer solution (3  300 μL), and resuspended to a final volume of 1.0 mL with 10 mM phosphate buffer solution. This suspension was stored at 4  C for further use.

3.2 Amplification Reaction

1. A 10 μL volume of standard CEA solution or sample solution was mixed with 90 μL of MBs-dsDNA conjugate solution in a 0.5-mL centrifuge tube. 2. This mixture solution was then incubated for 1 h at 37  C. 3. The resulting solution was separated magnetically, and 95 μL of supernatant was transferred into a 0.5-mL centrifuge tube. 4. Subsequently, 5 μL of 50 μM FAM labeled DNA probe solution was added, and incubated for 1 h at 37  C. 5. Finally, 11 μL of 10  NEB buffer 2 and 3 μL of 10 u/μL Nb. BbvCI solutions were added, and incubated for 2 h at 37  C. The resulting solution was analyzed by MCE-LIF.

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3.3 MCE-LIF Procedure

1. Before repetitive runs are started, the microfluidic channel was rinsed sequentially with 0.1 M NaOH, water, and MCE running buffer solution for 5 min each. Prior to electrophoresis, all reservoirs were filled with the MCE running buffer solution (see Note 2). 2. Replace the electrophoretic buffer solution in reservoir S with the sample solution (the resulting solution of amplification reaction). 3. For loading the sample solution, a set of electrical potentials were applied to four reservoirs: reservoir S at 500 V, reservoir B at 250 V, reservoir BW at 350 V, and reservoir SW at grounded. The sample solution was transported from reservoir S to SW in pinched mode. After 15 s, potentials were switched to reservoir B at 2500 V, while reservoir BW was grounded for separation and detection. 4. During separation, the sample between the S reservoir and SW reservoir was drawn back by applying a pinched voltage of 1500 V to prevent the sample entering the separation channel.

3.4 Calibration of the MCE-LIF System

1. Rinse the microchannels sequentially with 0.1 M NaOH, water, and MCE running buffer for 5 min each (see Note 3). 2. Fill reservoirs B, S, SW, and BW with the MCE running buffer. 3. Apply vacuum to reservoir BW (see Note 4). 4. Replace the MCE running buffer solution in reservoir S with sample solution (the resulting solution of amplification reaction with CEA standard solution). 5. Apply a set of electrical potentials to reservoirs as following: reservoir S at 500 V, reservoir B at 250 V, reservoir BW at 350 V, and reservoir SW at grounded to inject the sample. Duration: 15 s (see Note 5). 6. Change the potentials applied as following: reservoir B at 2500 V, reservoir S at 1500 V, reservoir SW at 1500 V, and reservoir BW at ground. At the same time, start to record the MCE-LIF electropherogram (as shown in Fig. 3). Duration: 1.5 min. 7. Plot peak height from FAM labeled DNA segment (peak 1) against CEA concentrations to obtain a calibration curve and the equation from linear regression for CEA.

3.5 Quantification of CEA in Human Serum

1. Do steps 1–3 described in Subheading 3.4. 2. Replace the MCE running buffer solution in reservoir S with a sample solution (the resulting solution of amplification reaction with human serum which was diluted 10 folds with 10 mM phosphate buffer solution at pH 7.4). 3. Do steps 5–6 described in Subheading 3.4.

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Fig. 3 Electropherograms for CEA standard solutions at various concentrations. The concentrations of CEA were 0 (a), 0.13 (b), 0.30 (c), 0.40 (d), 0.60 (e), 0.80 ( f ), 2.0 (g), 4.0 (h), 6.0 (i), and 8.0 ( j) ng/mL. Peak identification: (1) FAM labeled DNA segment (CEA); (2) FAM labeled DNA. MCE conditions were as following: 2500 V separation voltage and a running buffer containing 20 mM phosphate, 0.8% HPMC, and 15 mM TBAP at pH 7.4 A

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Fig. 4 Electropherograms obtained from the serum samples of a healthy volunteer (a) and a cancer patient (b). Peak identification: (1) FAM labeled DNA segment (CEA); (2) FAM labeled DNA. MCE conditions were as following: 2500 V separation voltage and a running buffer containing 20 mM phosphate, 0.8% HPMC, and 15 mM TBAP at pH 7.4

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4. Record the MCE-LIF electropherogram (as shown in Fig. 4a, b), and determine the CEA concentration from the peak height from FAM labeled DNA segment (peak 1) using the calibration equation obtained above for CEA.

4

Notes 1. Filtering all solutions before use in MCE is very important since channels in the microchip are very small in size and they are easily blocked by minute particles in solutions. 2. Vacuum was applied to the reservoir BW in order to fill the separation channel with the electrophoretic buffer. 3. Rinse the microchannels with 1 M NaOH for 30 min for the first use of the microchip. 4. Apply a vacuum to reservoir BW to fill the channels with the MCE running buffer solution. Check to make sure there are no air bubbles in the channels. 5. Under the potential conditions applied, the sample solution (the resulting solution of amplification reaction with CEA standard solution) is transported from reservoir S to SW. That is, the sampling channel that involves a small section of the separation channel is filled with the sample solution (the resulting solution of amplification reaction with CEA standard solution).

References 1. Kimura H, Matsuzawa S, Tu CY, Kitamori T, Sawada T (1996) Ultrasensitive heterogeneous immunoassay using photothermal deflection spectroscopy. 2. Quantitation of ultratrace carcinoembryonic antigen in human sera. Anal Chem 68:3063–3067 2. Seker D, Kaya O, Adabag A, Necipoglu G, Baran I (2003) Role of preoperative plasma CA 15-3 and carcinoembryonic antigen levels in determining histopathologic conventional prognostic factors for breast cancer. World J Surg 27:519–521 3. Shen GY, Wang H, Deng T, Shen GL, Yu RQ (2005) A novel piezoelectric immunosensor for detection of carcinoembryonic antigen. Talanta 67:217–220 4. Liu Y, Jiang H (2006) Electroanalytical determination of carcinoembryonic antigen at a silica nanoparticles/titania sol-gel composite membrane-modified gold electrode. Electroanalysis 18:1007–1013

5. Pan J, Yang QW (2007) Antibodyfunctionalized magnetic nanoparticles for the detection of carcinoembryonic antigen using a flow-injection electrochemical device. Anal Bioanal Chem 388:279–286 6. Walter K, Norbert N, Jochen S, Rudolf P, Herbert H (1988) Is there any clinical relevance of serial determinations of serum carcinoembryonic antigen in small cell lung cancer patients. Cancer 62:1348–1354 7. Pergters J, Schmide-Gayk H, Peters B, Armbruster FP, Quentmeler A, Mathlas D (1989) lmmunoradiometric assay of carcinoembryonic antigen with use of avidin-biotin labeling. Clin Chem 35:573–576 8. Lin JH, Yan F, Ju HX (2004) Noncompetitive enzyme immunoassay for carcinoembryonic antigen by flow injection chemiluminescence. Clin Chim Acta 341:109–115 9. Yuan JL, Wang GL, Majima K, Matsumoto K (2001) Synthesis of a terbium fluorescent

Aptamer-Based Microchip CE for Detection of CEA chelate and its application to time-resolved fluoroimmunoassay. Anal Chem 73:1869–1876 10. Dungchai W, Siangproh W, Lin JM, Chailapakul O, Lin S, Ying XT (2007) Development of a sensitive micro-magnetic chemiluminescence enzyme immunoassay for the determination of carcinoembryonic antigen. Anal Bioanal Chem 387:1965–1971 11. Ye F, Shi M, Huang Y, Zhao S (2010) Noncompetitive immunoassay for carcinoembryonic antigen in human serum by microchip electrophoresis for cancer diagnosis. Clin Chim Acta 411:1058–1062 12. Hou L, Tang Y, Xu M, Gao Z, Tang D (2014) Tyramine-based enzymatic conjugate repeats for ultrasensitive immunoassay accompanying tyramine signal amplification with enzymatic biocatalytic precipitation. Anal Chem 86:8352–8358 13. Ellington AD, Szostak JW (1990) In vitro selection of RNA molecules that bind specific ligands. Nature 346:818–822 14. Shangguan D, Li Y, Tang Z, Cao ZC, Chen HW, Mallikaratchy P, Sefah K, Yang CJ, Tan W (2006) Aptamers evolved from live cells as effective molecular probes for cancer study. Proc Natl Acad Sci U S A 103:11838–11843 15. Xue L, Zhou X, Xing D (2012) Sensitive and homogeneous protein detection based on target-triggered aptamer hairpin switch and nicking enzyme assisted fluorescence signal amplification. Anal Chem 84:3507–3513 16. Shang F, Guihen E, Glennon JD (2012) Recent advances in miniaturization-the role of microchip electrophoresis in clinical analysis. Electrophoresis 33:105–116

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Part VII Application in Virology

Chapter 19 Highly Sensitive SDS Capillary Gel Electrophoresis with Sample Stacking Requiring Only Nanograms of Adeno-Associated Virus Capsid Proteins Chao-Xuan Zhang and Michael M. Meagher Abstract Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) has been the method of choice in the past decades for size-based protein analysis. However, in general it requires the protein concentration in mg/mL level and thus is not practical for trace level protein analysis, not to mention the lengthy laborintensive procedures. The SDS capillary gel electrophoresis (SDS CGE) method reported herein requires only nanogram-sized proteins loaded onto the autosampler. A sample stacking technique (e.g., headcolumn field-amplified sample stacking (HC FASS)) was employed, providing three orders of magnitude sensitivity enhancement compared to conventional SDS CGE. This method has been used routinely in purity analysis and characterization of adeno-associated virus (AAV) intermediates and finished gene therapeutics of AAV vectors. The sensitivity achieved is comparable to the currently most sensitive sizebased protein assay silver-stained SDS PAGE. The highly sensitive sample stacking SDS CGE can be used for other types of proteins as well. Key words SDS capillary gel electrophoresis (SDS CGE), Head-column field-amplified sample stacking, FASS, Adeno-associated virus (AAV) protein, High sensitivity

1

Introduction Adeno-associated virus currently appears as a preferred platform for gene therapy with tremendous opportunities [1–3]. AAV vectors encoding factor VIII (FVIII) or factor IX (FIX) have been developed for clinical trials in patients with hemophilia of great therapeutic potentials [4, 5]. It is thought that the capsid of non-enveloped AAV is composed of 60 copies of three capsid proteins VP1 (87 kDa), VP2 (73 kDa), and VP3 (61 kDa) in a 1:1:10 stoichiometry [6]. AAV capsid proteins protect the therapeutic transgene from environment and are critical for viral infectivity and vector potency of AAV therapeutics. Analysis of AAV capsid proteins is challenging because the protein concentration is very low and often only nanograms of proteins are available for

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analysis. Conventional sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) has been used as the method of choice for protein analysis in the past decades [7]. However, it generally requires protein samples at mg/mL protein concentration levels. Thus, conventional SDS-PAGE is not practical for quality control testing of therapeutic AAV vectors because of the sample amount required, not to mention the quantitative variations due to the labor-intensive procedures. SDS capillary gel electrophoresis (SDS CGE) has gained increased popularity for size-based protein separation. It is an attractive alternative to traditional SDS-PAGE because samples can be analyzed automatically and chromatogram-like peaks, rather than images, are ready for quantification. However, current SDS CGE suffers from low sensitivity, which is not better than SDS-PAGE. Generally SDS CGE requires micrograms of proteins at mg/mL concentrations loaded onto the autosampler [8]. Sensitivity is often a challenge in developing capillary electrophoresis (CE) methods for solving real-world problems. In order to improve the sensitivity in CE, a more sensitive and of course more expensive detector such as laser induced fluorescence (LIF) is commonly employed. Online sample pre-concentration techniques in CE provide convenient approaches for sensitivity improvement without modification of the instrument and without additional cost [9–11]. Field-amplified sample stacking (FASS) is simple, yet effective [12]. In FASS, samples prepared in low-conductivity solutions may be injected electrokinetically or hydrodynamically. Thus, FASS can be classified into head-column FASS (for electrokinetic injection) and in-column FASS (for hydrodynamic injection) [13]. For in-column FASS, large sample volume is injected hydrodynamically and stacking occurs inside the capillary after injection. The sample matrix must be removed from the capillary prior to separation. In contrast, head-column FASS (HC FASS) is injected electrokinetically and stacking occurs concurrently at the head of the capillary during injection. The operation is simple because little sample matrix is injected. Although numerous online sample pre-concentration techniques have been reported as proof-of-concept studies, only a few of them are effective in real-world applications. Two decades of applications have demonstrated that HC FASS provide much larger sensitivity enhancements than other sample stacking techniques [14–24]. The working mechanisms of HC FASS have been studied theoretically and experimentally [25–27]. In a recent study, we have demonstrated that HC FASS can provide three orders of magnitude sensitivity enhancement in SDS CGE of AAV capsid proteins [22]. It requires only low nanograms of AAV proteins loaded onto the autosampler. In HC FASS of AAV proteins, a short water plug is introduced at the head of the capillary prior to the sample injection. Sample prepared with low conductivity is electrokinetically injected

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into the capillary filled with high conductivity running buffer and a low conductivity pre-injection water plug. During injection, the water zone will experience a much higher electric field than the running buffer zone. The negatively charged protein-SDS complexes move quickly from the sample vial into the water zone at the head of the capillary and slow down at the boundary to the running buffer. During injection/stacking, the EOF inside the capillary pushes the water zone toward the capillary inlet, avoiding sample solvent entering the capillary. Thus, there is no sample matrix injected and no limit on sample injection volume (see Note 1). The sample stacking SDS CGE method reported herein delivers high sensitivity, requiring only 10–50 ng of proteins loaded in the autosampler. The actual amount injected is much less than what is loaded in the autosampler (see Note 2). This is not a proof of principle study. This method has been routinely used for purity analysis of AAV intermediates and cGMP manufactured AAV vector products for Phase I/II clinical trials.

2 2.1

Materials Reagents

1. SDS-MW assay kit containing uncoated fused silica capillaries, SDS-MW gel buffer, acidic wash solution (0.1 N HCl), basic wash solution (0.1 N NaOH). 2. 99% Sodium dodecyl sulfate (SDS). 3. 99% 2-mercaptoethanol (ME). 4. Amicon Ultra-0.5 Centrifugal Filter Unit with 30 K Nominal Molecular Weight Limit (NMWL) ultrafiltration membranes.

2.2

AAV Samples

1. AAV intermediates. 2. Therapeutic AAV vectors. Both intermediates and vectors were provided by the Process Development group at Department of Therapeutics Production & Quality, St. Jude Children’s Research Hospital. The total protein concentration was measured by the bicinchoninic acid assay (BCA).

2.3

CE System

2.4 Solution Preparation

A Beckman Coulter PA800 Plus Pharmaceutical Analysis CE system equipped with a UV absorbance detector, set at 214 nm and Beckman 32 Karat software were used. 1. SDS 150 mg/mL solution: Weigh 1.52 g SDS and add about 8 mL deionized water. Vortex until a clear solution is formed. Add more deionized water to give a final volume of 10 mL. Store at room temperature (see Note 3).

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2. SDS 0.5 mg/mL solution: Weigh 0.051 g SDS and add 100 mL of deionized water. Vortex mix and store at room temperature (see Note 3). 3. Matrix exchange solution: Add 0.5 mL of ME to 9.5 mL of SDS 0.5 mg/mL solution. Prepare fresh on the day of use.

3

Methods

3.1 Sample Preparation

1. Add AAV sample of approximately 100–500 ng of total proteins, 6 μL of 150 mg/mL SDS, and 3 μL of 2-Mercaptoethanol (ME) into a centrifuge tube and vortex. 2. Incubate the mixture at 50  C for 3 min. 3. Transfer the incubate mixture and 450 μL matrix exchange solution to an Amicon Ultra-0.5 micro-centrifuge tube. 4. Centrifuge at 14,000  g for 10 min at 20  C. Discard the filtrate. 5. Add 450 μL matrix exchange solution to the Amicon tube. 6. Centrifuge at 14,000  g for 10 min at 20  C. Discard the filtrate. 7. Place the Amicon Filter upside down into a clean Amicon collection tube. 8. Centrifuge at 200  g for 0.5 min. Discard the filter. Incubate the mixture at 50  C for 3 min. 9. Add 970 μL deionized water into the collection tube containing the sample and vortex. Transfer 100 μL of the sample (about 10–50 ng of total proteins) onto the CE autosampler for SDS CGE analysis within 1 day (see Note 4).

3.2 SDS CGE with Sample Stacking

1. Autosampler temperature is set at 10  C. Capillary cartridge temperature is set at 25  C. 2. At the beginning of each batch of analysis, an uncoated capillary (50 μm I.D., 30.2 cm total length, 20 cm from injection to detector) is rinsed sequentially with: 0.1 N NaOH (10 min at 20 psig), 0.1 N HCl (5 min at 20 psi), D.I. water (2 min at 20 psig), SDS-MW Gel Buffer (10 min at 70 psig). 3. Prior to each sample injection, the capillary is rinsed sequentially with: 0.1 N NaOH (3 min at 70 psig), 0.1 N HCl (1 min at 70 psig), D.I. water (1.5 min at 70 psig), SDS-MW Gel Buffer (10 min at 70 psig) (see Note 5). 4. A short D.I. water plug is injected by applying a pressure of 20 psig for 0.4 min. 5. Samples were injected by applying a negative voltage of for 60 s (see Note 6).

5 kV

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267

Fig. 1 Typical electropherograms of three replicate runs of an AAV sample. Approximately 50 ng of total AAV proteins were loaded on the autosampler. The peak migrated before the VP3 may be for an unidentified component of AAV capsid

6. For separation, a negative voltage of 15 kV was applied for 30 min with a pressure of 20 psig on both ends of the capillary. 7. The method is programed to increment to fresh solution/ water vials every 6 injections. 8. At the end of each analytical batch, the capillary was rinsed sequentially with: 0.1 N NaOH (10 min at 70 psig), 0.1 N HCl (5 min at 50 psig), D.I. water (2 min at 50 psig), SDS-MW Gel Buffer (10 min at 70 psig) (see Note 7). 3.3

CE Data Analysis

Typical electropherograms of three replicate runs of an AAV sample is presented in Fig. 1. Three peaks corresponding to AAV capsid proteins VP3, VP2, and VP1 are obtained. Other unknown peaks could be observed for impurities or unknown components of AAV capsids. 1. Use Beckman 32 Karat software for data acquisition and analysis. 2. Peak integration is performed with appropriate parameters. 3. Purity is calculated by summarizing the peak areas of VP1, VP2, and VP3 and dividing by the total peak area (see Note 7).

4

Notes 1. Theoretically, stacking efficiency (e.g., sensitivity) in HC FASS is largely determined by the conductivity ratio of buffer zone to water zone and water zone length fraction. Note that both

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parameters changes with time during injection as the water zone is being pumped out of the capillary by EOF. A longer water zone will allow more analytes injected before the transient water zone disappears due to diffusion and EOF. However, too large a water zone will decrease the amount of analytes injected due to the reduced electric field strength in the water zone. Thus, it is important to select an optimum water zone length. In this work, a water zone of around 10 mm generated the highest peaks. Different CE systems and different capillary sizes require different parameters to achieve an optimum water zone length. 2. Sample stacking SDS CGE requires only tens of nanograms of proteins loaded in the autosampler. Actual injected amount is estimated to be at pictogram levels. The sensitivity achieved with the sample stacking SDS CEG is comparable to silverstained SDS PAGE, which is commonly regarded as the most sensitive SDS PAGE assay. 3. If SDS precipitates from the SDS 150 mg/mL solution, discard it and prepare a fresh solution. 4. Treated samples should be analyzed immediately or stored at 10  C for no more than one day. 5. Fresh buffer and deionized water should be used for every six injections or less. 6. Injection time may be increased for even higher sensitivity. However, peaks may become slightly broader when injection time is over 180 s. 7. The sample stacking SDS CGE method can be used for analysis of other proteins as well.

Acknowledgements Dr. Timothy Lockey, Dr. Bryan Piras, Dr. Jason Drury, Dr. Susan Sleep, and Dr. Clifford Froelich are greatly acknowledged for providing AAV samples and analytical support. This work was supported by American Lebanese Syrian Associated Charities (ALSAC). References 1. Rodriguez-Merchan EC (2018) What’s new in gene therapy of hemophilia. Curr Gene Ther 18:107–114 2. Lykken EA, Shyng C, Edwards RJ, Rozenberg A, Gray SJ (2018) Recent progress and considerations for AAV gene therapies

targeting the central nervous system. J Neurodev Disord 10:16 3. Bass-Stringer S, Bernardo BC, May CN, Thomas CJ, Weeks KL, McMullen JR (2018) Adeno-associated virus gene therapy: translational progress and future prospects in the

Highly Sensitive SDS CGE of AAV Capsid Proteins treatment of heart failure. Heart Lung Circ 27 (11):1285–1300 4. Allay JA, Sleep S, Long S, Tillman DM, Clark R, Carney G, Fagone P, McIntosh JH, Nienhuis AW, Davidoff AM, Nathwani AC, Gray JT (2011) Good manufacturing practice production of self-complementary serotype 8 adeno-associated viral vector for a hemophilia B clinical trial. Hum Gene Ther 22:595–604 5. Piras BA, Drury JE, Morton CL, Spence Y, Lockey TD, Nathwani AC, Davidoff AM, Meagher MM (2016) Distribution of AAV8 particles in cell lysates and culture media changes with time and is dependent on the recombinant vector. Mol Ther Methods Clin Dev 3:16015 6. Halder S, Van Vliet K, Smith JK, Duong TT, McKenna R, Wilson JM, Agbandje-McKenna M. (2015) Structure of neurotropic adenoassociated virus AAVrh.8. J Struct Biol 192:21–36 7. Eckard AD, Dupont DR, Young JK (2018) Development of two analytical methods based on reverse phase chromatographic and SDS-PAGE gel for assessment of deglycosylation yield in N-glycan mapping. Biomed Res Int 2018:3909674 8. Karageorgos I, Gallagher ES, Galvin C, Gallagher DT, Hudgens JW (2017) Biophysical characterization and structure of the Fab fragment from the NIST reference antibody, RM 8671. Biologicals 50:27–34 9. Voeten RLC, Ventouri IK, Haselberg R, Somsen GW (2018) Capillary electrophoresis: trends and recent advances. Anal Chem 90:1464–1481 10. Liu Y, Wang W, Jia M, Liu R, Liu Q, Xiao H, Li J, Xue Y, Wang Y, Yan C (2018) Recent advances in microscale separation. Electrophoresis 39:8–33 11. Breadmore MC, Wuethrich A, Li F, Phung SC, Kalsoom U, Cabot JM, Tehranirokh M, Shallan AI, Abdul Keyon AS, See HH, Dawod M, Quirino JP (2017) Recent advances in enhancing the sensitivity of electrophoresis and electrochromatography in capillaries and microchips (2014–2016). Electrophoresis 38:33–59 12. Chien RL, Burgi DS (1992) On-column sample concentration using field amplification in CZE. Anal Chem 64:489A–496A 13. Zhang CX, Thormann W (1996) Headcolumn field-amplified sample stacking in binary system capillary electrophoresis: a robust approach providing over 1000-fold sensitivity enhancement. Anal Chem 68:2523–2532

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14. Zhang CX, Aebi Y, Thormann W (1996) Microassay of amiodarone and desethylamiodarone in serum by capillary electrophoresis with head-column field-amplified sample stacking. Clin Chem 42:1805–1811 15. Zhang CX, Thormann W (1998) Headcolumn field-amplified sample stacking in binary system capillary electrophoresis. 2. Optimization with a preinjection plug and application to micellar electrokinetic chromatography. Anal Chem 70:540–548 16. Song JZ, Chen J, Tian SJ, Sun ZPJ (1999) Assay for the determination of low dosage form of formoterol dry syrup by capillary electrophoresis with head-column field-amplified sample stacking. J Pharm Biomed Anal 21:569–576 17. Wey AB, Zhang CX, Thormann W (1999) Head-column field-amplified sample stacking in binary system capillary electrophoresis. Preparation of extracts for determination of opioids in microliter amounts of body fluids. J Chromatogr A 853:95–106 18. Chou YW, Huang WS, Chen CC, Lin SJ, Wu HL, Chen SH (2005) Trace analysis of zotepine and its active metabolite in plasma by capillary electrophoresis with solid phase extraction and head-column field-amplified sample stacking. J Chromatogr A 1087:189–196 19. Yang Y, Boysen RI, Hearn MT (2006) Optimization of field-amplified sample injection for analysis of peptides by capillary electrophoresis-mass spectrometry. Anal Chem 78:4752–4758 20. Ito E, Nakajima K, Waki H, Miseki K, Shimada T, Sato TA, Kakehi K, Suzuki M, Taniguchi N, Suzuki A (2013) Structural characterization of pyridylaminated oligosaccharides derived from neutral glycosphingolipids by high-sensitivity capillary electrophoresismass spectrometry. Anal Chem 85:7859–7865 21. Theurillat R, Sandbaumhu¨ter FA, BettschartWolfensberger R, Thormann W (2016) Microassay for ketamine and metabolites in plasma and serum based on enantioselective capillary electrophoresis with highly sulfated γ-cyclodextrin and electrokinetic analyte injection. Electrophoresis 37:1129–1138 22. Zhang CX, Meagher MM (2017) Sample stacking provides three orders of magnitude sensitivity enhancement in SDS capillary gel electrophoresis of adeno-associated virus capsid proteins. Anal Chem 89:3285–3292 23. Ciura K, Pawelec A, Buszewska-Forajta M, Markuszewski MJ, Nowakowska J, Prahl A, Wielgomas B, Dziomba S (2017) Evaluation of sample injection precision in respect to

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sensitivity in capillary electrophoresis using various injection modes. J Sep Sci 40:1167–1175 24. Wang Z, Guo H, Chen M, Zhang G, Chang R, Chen A (2018) Separation and determination of corynoxine and corynoxine B using chiral ionic liquid and hydroxypropyl-β-cyclodextrin as additives by field-amplified sample stacking in capillary electrophoresis. Electrophoresis 39 (17):2195–2201. https://doi.org/10.1002/ elps.201800129. Epub ahead of print 25. Sˇesta´k J, Thormann W (2017) Insights into head-column field-amplified sample stacking: Part I. Detailed study of electrokinetic

injection of a weak base across a short water plug. J Chromatogr A 1502:51–61 26. Sˇesta´k J, Thormann W (2017) Insights into head-column field-amplified sample stacking: Part II. Study of the behavior of the electrophoretic system after electrokinetic injection of cationic compounds across a short water plug. J Chromatogr A 1512:124–132 27. Sˇesta´k J, Theurillat R, Sandbaumhu¨ter FA, Thormann W (2018) Fundamental aspects of field-amplified electrokinetic injection of cations for enantioselective capillary electrophoresis with sulfated cyclodextrins as selectors. J Chromatogr A 1558:85–95

INDEX A Acylcarnitines ...................................................... 140–143, 146, 147, 149–152, 158, 160, 161 Adeno-associated virus (AAV) protein................ 263–268 Affinity chromatography affinity cartridge ........................................................ 59 affinity cartridge packing .......................................... 59 α1-Acid glycoprotein (AGP) .......................25–38, 41–55 Amino acids Asn297....................................................................... 78 branched-chain ........................................... 15–23, 153 5-methylcytosine ................................... 213, 214, 217 8-Aminopyrene-1,3,6-trisulfonic acid (APTS) labeling........................87, 89, 236–238, 242, 248 Analyte concentrator....................................................... 63 Antibody antibody immobilization ................................. 58, 195 Aptamer ..........................................................58, 251–258 Auto-antibody detection .............................................. 122

B Background electrolyte (BGE)................................17–21, 23, 58–61, 63, 64, 67–69, 71, 73, 97, 98, 101–104, 106, 107, 129, 130, 132, 144, 145, 147, 149, 150, 159–161, 167, 169, 222–226, 229–232 Binding analysis ............................................................. 112 Bioanalyzer ...................................................................... 36 Biomarker cancer ....................................................................... 221 glycan ................................................................ 77, 236 Branched-chain amino acids ............................ 15–23, 153

C Cancer biomarker.......................................................... 221 Capillary activation ............................................. 17, 61, 97, 101 conditioning .................. 67, 149, 150, 222, 225, 226 cutter........................................... 17, 69, 98, 103, 143 fused silica capillary ............................................17, 33, 37, 46, 54, 61–65, 69, 98, 103, 110, 113, 115, 116, 119, 129, 141, 143, 150, 157, 159, 168, 217, 224, 265 window making ....................................................... 157

Capillary electrophoresis basic principles......................................................... 5, 7 calibration .................................................................. 19 capillary zone electrophoresis (CZE)............ 214, 222 clinical applications ................................................. 4, 7 history .......................................................................... 4 μCE ............................................................................ 20 SDS capillary gel electrophoresis (SDS CGE) ......................................... 36, 263–268 Capillary electrophoresis systems ABI 3130xl genetic analyzer ................................. 203, 237, 241, 248, 249 Agilent 7100.............................................21, 107, 143 Beckman P/ACE MDQ ............................44, 46, 129 Beckman P/A800 Plus ........................................... 265 FIDAlyzer ................................................................ 111 laboratory built ....................................................... 195 Microlyne................................................................. 189 Sheathless CESI 8000............................................. 168 Capillary modification..................................................... 42 Carbohydrate electrophoresis.............................. 235–250 Carcinoembryonic antigen (CEA) ...................... 251–258 CE-ESI-MS ................................................................... 149 Cell culture ........................................................... 95, 96, 98 incubation..........................................................95–107 CE-MS coupling .................................................. 139–161 Chemokines.........................................185, 188, 191–196 Chip gel electrophoresis ...........................................26, 36 Clinical chemistry..................................................v, 7, 186 Clinical proteomics ............................................................v Clinical samples ................................................... 7–10, 18, 95–98, 100–103, 107 Contactless conductivity detector (CE-C4D)............................................. 15, 95–107 Cystic fibrosis (CF) ....................................................... 142 Cytokines ............................. 25, 185, 186, 188, 191–196

D Data analysis with GeneMapper software ..........203, 205, 236, 241 Deglycosylation ....................................... 25–38, 236, 242 Diagnosis point-of-care diagnostics ........................................ 175 Diode-array detector................................... 129, 214, 224

Terry M. Phillips (ed.), Clinical Applications of Capillary Electrophoresis: Methods and Protocols, Methods in Molecular Biology, vol. 1972, https://doi.org/10.1007/978-1-4939-9213-3, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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CLINICAL APPLICATIONS

272 Index

OF

CAPILLARY ELECTROPHORESIS: METHODS

Dissociation constant ...................................112, 119–121 DNA analysis ................................................................... 79 DNA sequencer-assisted fluorophore-assisted carbohydrate electrophoresis (DSA-FACE) ............................................ 235–250 Donepezil ............................................................. 127, 132 Dried blood spots collection ........................................................ 148, 149 extraction ........................................................ 148, 149

E Electrokinetic injection .......................6, 43, 72, 161, 264 Electropherogram ............................................. 19–21, 23, 35, 47, 49–51, 53, 79, 88, 89, 98, 100, 101, 104, 106, 107, 131, 132, 150, 152, 153, 159, 170, 192, 194, 201, 203, 205–209, 217, 225, 226, 241, 248, 249, 256–258, 267 Electrophoretic injection ..........................................41–55 Electrospray ionization ............................... 139, 144, 146 Elution buffer......................................187, 192, 193, 196 Enantiomers ........................................127, 128, 131, 132

F Field-amplified sample stacking (FASS).............. 264, 267 Flow-induced dispersion analysis (FIDA)........... 109–122 Fluconazole (FLU) .................... 127, 128, 130, 132–135 Fluorescein 5-isothiocyanate (FITC)......... 176, 180, 183 Fluorescence detection .............................................29, 43 Fragile X-associated primary ovarian insufficiency (FXPOI)............................................................. 199 Fragile X-associated tremor/ataxia syndrome (FXTAS) ............................................................. 199 Fragile X mental retardation 1 (FMR1) ............. 199–208 Fragile X syndrome (FXS) ............................................ 199 Fused silica.................................................. 17, 33, 37, 46, 54, 61–65, 69, 98, 103, 110, 113, 115, 116, 119, 129, 141, 143, 150, 157, 159, 168, 217, 224, 265

G Gel electrophoresis................................................ 3–5, 26, 36, 200, 201, 263–268 Glucose ......................................................... 7, 83, 87, 89, 90, 95–107, 141, 242, 248 Glycoforms ..................................... 41–55, 235, 243, 246 Glycomics ............................................................. 244, 246 Glycoprotein................................................25–38, 41–55, 77, 79, 221, 222, 238, 242, 243, 245 Guthrie cards .......................................142, 144, 148, 188

H Head-column field-amplified sample stacking (HC FASS) ............................................... 264, 267 Highly-sulphated α,γ-cyclodextrins (CDs).................. 128

AND

PROTOCOLS

High resolution .............................. 72, 79, 144, 160, 222 History of capillary electrophoresis.................................. 4 Hjerten, S. ......................................................................... 5 Hot embossing............................................ 176, 179, 183 Human plasma ..................................15–23, 44, 118, 166 Hydrodynamic injection ..................................... 6, 19, 43, 51, 70, 144, 150, 160, 161, 217, 264

I Immunoaffinity capillary electrophoresis .............. 43, 57–73, 185–196 extraction ..............................................43, 57–73, 186 Immunoaffinity capillary electrophoresis antibody immobilization .......................................... 58 elution buffer........................................................... 196 extraction cartridge .............................................62, 64 extraction disk ............................... 190–192, 195, 196 Immunoglobulin G (IgG) ........................................77–89 Immunoglobulin purification...................................83, 84 Immunosubtraction ................................ 62–65, 192, 196 Inborn errors of metabolism (IEM) ................... 139–161 Inflammation ............................26, 77, 78, 185, 189, 194 Injection electrokinetic injection..................6, 43, 72, 161, 264 electrophoretic injection .....................................48, 49 head-column field-amplified sample stacking........ 267 hydrodynamic injection .......................................6, 19, 43, 51, 70, 144, 150, 160, 161, 217, 264 multiplexed injection ........... 141, 147, 150, 152, 155 sample stacking............................16, 21, 57, 263–268 short end injection ......................................... 127–136 Internal standard (IS) ............................................. 17, 51, 82, 97, 101, 146, 149, 150, 153, 159, 168 Isoform separation ...................................... 110, 222, 226

L Lab-on-a-chip......................... 25–39, 175–198, 251–259 Lactate .....................................................................95–107 Laser induced fluorescence (LIF).................................v, 6, 25–38, 43, 82, 119, 141, 186, 264 Ligand binding assays .......................................... 109, 110

M Magnetic beads (MBs)............... 59–61, 67, 70, 252, 255 Mass spectrometer ABI Voyager DE PRO MS ....................................... 31 Agilent 6550 quadrupole time-of-flight MS ......... 143 Agilent 6230 time-of-flight MS ............................. 143 Mass spectrometry CE-MS .............................. 57–73, 139–161, 165–171 IA-SPE-CE-MS .........................59, 60, 65–68, 70, 73 MSI-CE-MS .................................................. 142, 144, 148–150, 152–155, 157, 159–161

CLINICAL APPLICATIONS

OF

CAPILLARY ELECTROPHORESIS: METHODS

Metabolic profiling.......................................149, 165–171 Metabolites ........................................................79, 95, 96, 141–150, 152, 153, 155–161, 165–170 Metabolomics ..................... v, 7, 145, 148, 158, 165–167 Methylamidation .......................................................79, 89 Microcartridge.......................... 58, 59, 62–64, 69, 71–73 Microchip capillary electrophoresis ................................. 175, 252 conditioning .............................................................. 34 electrophoresis.........................25, 175–184, 251–258 gel electrophoresis................................. 26, 33, 34, 36 microdevice fabrication.................................. 175–184 microlyne borofloat glass chip................................ 189 power supply ............................................................. 27 SU-8 microchips ................................................. 25–38 Microdialysis .....................................................95, 96, 107 Microfabrication............................................................ 176 Microfluidics........................................................ 175, 179, 189, 235, 253, 256 Molecular identification of virus strain ............... 263–270 Molecular interactions ......................................... 117–119 Moving boundary electrophoresis ................................... 4

N Newborn infants ......................................... 185, 186, 188 Newborn screening .............................................. 139–161 N-glycan analysis ........................................ 81, 82, 84–86, 88, 236–238, 240, 241, 244, 249 N-glycosylation ............................ 79, 239, 244, 246, 249 Nucleic acids..................................................... v, 3, 5, 251

P Pathogen detection .............................................. 263–268 PCR T-PCR............................................................. 199–208 Photolithography ................................................... 26, 253 Plasma human ................................. 15–23, 44, 118, 166, 222 Poly(ethylene glycol) .................................................... 176 Polymer matrix ................................................................ 36 Poly(methyl methacrylate) (PMMA) device fabrication .................................................... 182 Preconcentration ...............................................57, 60, 63, 73, 142, 144, 150, 157, 160 Preparation of antibody fragments ..........................58, 67 Pressure-assisted analysis.................................... 16, 18, 19 Prostate specific antigen (PSA) ........................... 221–233 Proteins protein-antibody interaction .................................... 78 protein profiling ............................................. 235–250

AND

PROTOCOLS Index 273

protein-protein interaction ..................................... 110 proteoform ...............................................59, 221, 222 Purification ................................................. 26, 31, 43, 79, 82–84, 86, 87, 90, 209, 222, 236

R Repeat expansion ........................................ 200, 201, 209

S Samples biomass-limited .............................................. 165–171 preparation .........................................................44, 47, 50, 79, 87, 88, 113, 119, 148, 149, 171, 216, 222, 224, 225, 266 Sample stacking .................................16, 21, 57, 263–268 SDS capillary gel electrophoresis (SDS CGE).............. 36, 263–268 Serum.............................................................4, 5, 7–9, 30, 42–44, 46–48, 50–52, 54, 57–73, 78, 83, 109, 110, 113, 121, 168, 185, 222, 251, 253, 256–258 Sheathless-CE................................................................ 168 Sheathless interface .............................................. 157, 166 Sheath liquid............................................................ 60, 61, 64, 67, 69–71, 141, 143–145, 149–151, 157, 159–161, 166 Solid-phase extraction (SPE)........................... 57–73, 149 Statistical analysis .......................................................... 145 SU-8 microchips ............................................................. 25 Sweat collection............................................................. 144

T Taylor dispersion analysis (TDA) ................................. 110 Time-of-flight mass spectrometer (TOF MS) ............. 157 Tiselius, A. ..................................................................... 4, 5 Transthyretin (TTR) ............................................ 7, 57–73 Treatment of biological samples .................................... 97 Triplet-repeat primed PCR (TP-PCR) ............... 199–208

U Urine...........................................7, 9, 236, 244, 247, 249 UV-vis detector ......................................... 6, 96, 110, 224

V Virus identification............................................... 263–268

Z Zone electrophoresis......................................................... 4