Biosensors and Biodetection
Series Editor John M. Walker University of Hertfordshire Hatfield, Hertfordshire, UK
For other titles published in this series, go to www.springer.com/series/7651
METHODS
IN
MOLECULAR BIOLOGY™
Biosensors and Biodetection Methods and Protocols Volume 504: Electrochemical and Mechanical Detectors, Lateral Flow and Ligands for Biosensors
Edited by
Avraham Rasooly* and Keith E. Herold† *FDA Center for Devices and Radiological Health, Silver Spring, MD, USA and National Cancer Institute, Bethesda, MD, USA † Fischell Department of Bioengineering, University of Maryland, College Park, MD, USA
Editors Avraham Rasooly FDA Center for Devices and Radiological Health Silver Spring, MD USA and National Cancer Institute Bethesda, MD USA
[email protected] Keith E. Herold Fischell Department of Bioengineering University of Maryland College Park, MD USA
[email protected] ISBN: 978-1-60327-568-2 e-ISBN: 978-1-60327-569-9 ISSN: 1064-3745 e-ISSN: 1940-6029 DOI: 10.1007/978-1-60327-569-9 Library of Congress Control Number: 2008941063 © Humana Press, a part of Springer Science+Business Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science + Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper springer.com
Preface 1. Biosensor Technologies In recent years, many types of biosensors have been developed and used in a wide variety of analytical settings, including biomedical, environmental, research, and others. A biosensor is defined by the International Union of Pure and Applied Chemistry (IUPAC) as a “device that uses specific biochemical reactions mediated by isolated enzymes, immunosystems, tissues, organelles, or whole cells to detect chemical compounds usually by electrical, thermal, or optical signals” (1). Thus, almost all biosensors are based on a two-component system: a biological recognition element (ligand) that facilitates specific binding to or biochemical reaction with a target, and a signal conversion unit (transducer). Although it is impossible to fully cover the fast-moving field of biosensing in one publication, this publication presents some of the many types of biosensors to give the reader a sense of the enormous potential for these devices. An early reference to the concept of a biosensor is from Dr. Leland C. Clark, who worked on biosensors in the early 1960s (2) developing an “enzyme electrode” for glucose concentration measurement with the enzyme glucose oxidase, a measurement that is important in the diagnosis and treatment of disorders of carbohydrate metabolism in diabetes patients. Still today, the most common biosensors used are for glucose analysis. A large number of basic biosensors, all combining a biological recognition element and a transducer, were subsequently developed. Currently, the trend is toward more complex integrated multianalyte sensors capable of more comprehensive analyses. Advances in electronics and microelectrical and mechanical systems (MEMS) have enabled the miniaturization of many biosensors and the newest generation biosensors include miniaturized multianalyte devices with high-throughput capabilities and more than 1,000 individually addressable sensor spots per square centimeter. A useful categorization of biosensors is to divide them into two groups: direct recognition sensors, in which the biological interaction is directly measured, and indirect detection sensors, which rely on secondary elements for detection. Figure 1 shows a schematic of the two groups of biosensors. In each group, there are several types of transducers including optical, electrochemical, and mechanical. For all of these technologies, the recognition ligand plays a major role. Although the most commonly used ligands are antibodies, other ligands are being developed including aptamers (protein-binding nucleic acids) and peptides. In the literature and in practice, there are numerous types of biosensors, and the choice of a suitable system for a particular application is complex, based on many factors such as the nature of the application, the label molecule (if used), the sensitivity required, the number of channels (or area), cost, technical expertise, and the speed
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A
B
Recognition Recognition Element
Recognition Element
Transducer Transducer
Ligand Target
Output Interface
None specific analytes labeled molecule
Fig. 1. General schematic of biosensors: (a) direct detection biosensors where the recognition element is label free; (b) indirect detection biosensors using a “sandwich” assay where the analyte is detected by labeled molecule. Direct detection biosensors are simpler and faster but typically yield a higher limit of detection compared with indirect detection systems
of detection needed. A primary purpose of this book is to provide more access to the technical methods involved in using a variety of biosensors to facilitate such decision making. Direct detection biosensors utilize direct measurement of the biological interaction. Such detectors typically measure physical changes (e.g., changes in optical, mechanical, or electrical properties) induced by the biological interaction, and they do not require labeling (i.e., label free) for detection. Direct biosensors can also be used in an indirect mode, typically to increase their sensitivity. Direct detection systems include optical-based systems (most common being surface plasmon resonance) and mechanical systems such as quartz crystal resonators. Indirect detection sensors rely on secondary elements (labels) for detection. Examples of such secondary elements are enzymes (e.g., alkaline phosphatase or glucose oxidase) and fluorescently tagged antibodies that enhance detection of a sandwich complex. Unlike direct detectors, which directly measure changes induced by biological interactions and are “label free,” indirect detectors require a labeled molecule to bind to the target. Most indirect sensors based on optical detection are designed to measure fluorescence. The detection system can be based on a charge coupled device (CCD), photomultiplier tube (PMT), photodiode, or spectrometer. Electrochemical transducers, which measure the oxidation or reduction of an electroactive compound on the secondary ligand, are another common type of indirect detection sensor. Several types of electrochemical biosensors are in use including amperometric devices, which measure the electric current as a function of time while the electrode potential is held constant. Ligands are recognition molecules that bind specifically with the target molecule to be detected. The most important characteristics for ligands are affinity and specificity. Various types of ligands are used in biosensors. Biosensors that use antibodies as recognition elements (immunosensors) are common because antibodies are highly specific, versatile, and bind strongly and stably to the antigen. Several limitations of
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antibodies are long-term stability, and manufacturing costs, especially for multitarget biosensor applications where many ligands are needed. Other types of ligands that show promise for high-throughput screening and chemical synthesis are aptamers and peptides. Aptamers are protein-binding nucleic acids (DNA or RNA molecules) selected from random pools on the basis of their ability to bind other molecules with high affinity. Peptides can be selected for affinity to a target molecule by display methods (phage display and yeast display). However, in general, the binding affinity of peptides is lower than the affinity of antibodies or aptamers.
2. Biosensor Applications Biosensors have several potential advantages over other methods of biodetection, especially increased assay speed and flexibility. Rapid, essentially real-time analysis can provide immediate interactive information to users. This speed of detection is an advantage in essentially all applications. Applications of biosensors include medical, environmental, public security, and food safety areas. Medical applications include clinical, pharmaceutical and device manufacturing, and research. Biosensor-based diagnostics might facilitate disease screening and improve the rates of earlier detection and attendant improved prognosis. Such technology may be extremely useful for enhancing health care delivery in the community setting and to underserved populations. Environmental applications include spill clean-up, monitoring, and regulatory instances. Public safety applications include civil and military first responders as well as unattended monitoring. Food safety applications include monitoring of food production, regulatory monitoring, and diagnosis of food poisoning. Biosensors allow multitarget analyses, automation, and reduced costs of testing. The key strengths of biosensors are the following: • Fast or real-time analysis: Fast or real-time detection provides almost immediate interactive information about the sample tested, enabling users to take corrective measures before infection or contamination can spread. • Point-of-care detection: Biosensors can be used for point-of-care or on-site testing where state-of-the-art molecular analysis is carried out without requiring a stateof-the-art laboratory. • Continuous flow analysis: Many biosensor technologies can be configured to allow continuous flow analysis. This is beneficial in food production, air quality, and water supply monitoring. • Miniaturization: Biosensors can be miniaturized so that they can be integrated into powerful lab-on-a-chip tools that are very capable while minimizing cost of use. • Control and automation: Biosensors can be integrated with on-line process monitoring schemes to provide real-time information about multiple parameters at each production step or at multiple time points during a process, enabling better control and automation of many industrial and critical monitoring facilities.
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3. Aims and Approach The primary aim of this book is to describe the basic types and the basic elements of biosensors from methods point of view. We tried to include manuscripts that represent the major technologies in the field and to include enough technical detail so that the informed reader can both understand the technology and also be able to build similar devices. The target audience for this book includes engineering, chemical, and physical science researchers, who are developing biosensing technologies. Other target groups are biologists and clinicians, who are the users and developers of applications for the technologies. In addition to supporting the research community, the book may also be useful as a teaching tool for bioengineering, biomedical engineering, and biology faculty and students. To better represent the field, most topics are covered by more than one chapter. The purpose of this “redundancy” is to try to include several alternative approaches for the topics, so as to help the reader choose an appropriate design.
4. Chapter Organization This publication is divided into two volumes: Vol. 503 is focused on Optical-Based Detectors and Vol. 504 is focused on Electrochemical and Mechanical Detectors, Lateral Flow, and Ligands for Biosensors. 4.1. Volume 503: Optical-Based Detectors
Optical detection is used in a broad array of biosensor technologies, including both direct and indirect style sensors. Volume 503 is organized in two parts. Part I focuses on direct optical detectors, while Part II concentrates on indirect optical detection. Probably, the most common approach for direct optical detection is based on evanescent wave physics, where the interaction between the evanescent wave and the bound target generates a detection signal. The most common technology in this group is surface plasmon resonance (SPR) and several chapters (see Chaps. 1–5) describe biosensors based on SPR. Other important optical direct detection methods including resonant mirror (see Chap. 6), optical ring resonator (see Chap. 7), interferometric sensors (see Chaps. 8 and 9) and grating coupler (see Chap. 10) are all included in Part I. The second part of Vol. 503 describes various indirect optical detectors. As discussed earlier, indirect detectors require a labeled molecule to bind to the target generating a signal. For optical sensors, the label molecule emits or modifies light. Most indirect optical detectors are designed to measure fluorescence. However, optical detectors can also measure optical density (densitometry), changes in color (colorimetric), and chemoluminesence, depending on the type of label used. Optical signals can be measured in various ways (described in Part II) including various CCD-based detectors, which are very versatile, inexpensive, and relatively simple to construct and use (see Chaps. 11–16 and 25). Other optical detectors discussed in Part II are photodiodes (see Chaps. 17–20), photomultipliers (see Chaps. 21–23), and spectrometers (see Chaps. 24 and 25). Photomultipliers may offer higher sensi-
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tivity, smaller footprint (the size of photodiode can be few millimeters). Spectrometers offer better interrogation of changes in light wavelengths. 4.2. Volume 504: Electrochemical and Mechanical Detectors, Lateral Flow, and Ligands
Volume 504 describes various electrochemical and mechanical detectors, lateral flow devices, and ligands for biosensors. As in Vol. 503, we describe several direct measurement sensors (in Part I), indirect methods (Parts II–III). Ligands are described in Part IV and two related technologies are described in Part V. In Part I, we describe several mechanical detectors that modify their mechanical properties as a result of biological interactions. Such mechanical direct biosensors typically sense resonance of the mechanical element, which changes when the target molecule binds to the surface. Piezoelectric biosensors (see Chaps. 1–3) employ a technology that is widely used in a variety of applications (e.g., vapor deposition of metals) and is thus readily available and relatively inexpensive. Cantilever-based systems (see Chaps. 4 and 5) can be miniaturized to micrometer dimensions with attendant benefits for system and sample size. In Part II, we describe several electrochemical detectors (see Chaps. 6–11). Electrochemical biosensors were the first biosensors developed and are the most commonly used biosensors today (e.g., glucose monitoring). Part III covers lateral flow technologies (see Chaps. 12–15). Although lateral flow devices are not “classical” biosensors, with ligands and transducers, they are included in this book because of their importance for biosensing. Lateral flow assays are simple immunodetection (or DNA hybridization) devices, which utilize competitive or sandwich assays. They are used mainly for medical diagnostics, including laboratory, home and point-of-care detection. A common format is a “dipstick” in which the test sample diffuses through a porous matrix via capillary action followed by detection by a colorimetric reagent bound to a secondary antibody. The primary antibody is bound to the matrix in a line, and the assay result is a color change at a particular location on the matrix. Lateral flow assays can be dependable and inexpensive. Part IV focuses on recognition ligands, which are key elements in any biosensor (see Chaps. 16–22). The recognition ligands bind specifically with the target molecule to be detected. Various ligands described in Part IV include antibodies, aptamers, and peptides. Antibodies are the most commonly used ligands but advances in selection methods for aptamers (SELEX) and peptides (phage and yeast display) are currently providing alternatives. Part V includes two papers on protein (see Chap. 23) and DNA preparation (see Chap. 24). These papers are relevant to the subject of biosensor technologies but did not fit elsewhere into the book organization outline.
References 1. IUPAC Compendium of Chemical Terminology 2nd Edition (1997). (1992), International Union of Pure and Applied Chemistry: Research Triangle Park, NC. 2. Clark LC Jr., Lyons C (1962) Electrode systems for continuous monitoring in cardiovascular surgery. Ann N Y Acad Sci 102:29–45.
Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xiii Contents of Volume 503. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xvii
PART I: MECHANICAL DETECTORS 1.
A Set of Piezoelectric Biosensors Using Cholinesterases . . . . . . . . . . . . . . . . . . . . Carsten Teller, Jan Halámek, Alexander Makower, and Frieder W. Scheller
3
2.
Piezoelectric Biosensors for Aptamer–Protein Interaction. . . . . . . . . . . . . . . . . . . Sara Tombelli, Alessandra Bini, Maria Minunni, and Marco Mascini Piezoelectric Quartz Crystal Resonators Applied for Immunosensing and Affinity Interaction Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Petr Skládal Biosensors Based on Cantilevers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mar Álvarez, Laura G. Carrascosa, Kiril Zinoviev, Jose A. Plaza, and Laura M. Lechuga Piezoelectric-Excited Millimeter-Sized Cantilever Biosensors . . . . . . . . . . . . . . . . Raj Mutharasan
23
3.
4.
5.
37 51
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PART II: ELECTROCHEMICAL DETECTORS 6.
7.
8.
Preparation of Screen-Printed Electrochemical Immunosensors for Estradiol, and Their Application in Biological Fluids. . . . . . . . . . . . . . . . . . . . Roy M. Pemberton and John P. Hart
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Electrochemical DNA Biosensors: Protocols for Intercalator-Based Detection of Hybridization in Solution and at the Surface . . . . . . . . . . . . . . . . . . 99 Kagan Kerman, Mun’delanji Vestergaard, and Eiichi Tamiya Electrochemical Biosensor Technology: Application to Pesticide Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Ilaria Palchetti, Serena Laschi, and Marco Mascini
9.
Electrochemical Detection of DNA Hybridization Using Micro and Nanoparticles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127 María Teresa Castañeda, Salvador Alegret, and Arben Merkoçi 10. Electrochemical Immunosensing Using Micro and Nanoparticles . . . . . . . . . . . . 145 Alfredo de la Escosura-Muñiz, Adriano Ambrosi, Salvador Alegret, and Arben Merkoçi 11. Methods for the Preparation of Electrochemical Composite Biosensors Based on Gold Nanoparticles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157 A. González-Cortés, P. Yáñez-Sedeño, and J.M. Pingarrón
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PART III: LATERAL FLOW 12. Immunochromatographic Lateral Flow Strip Tests. . . . . . . . . . . . . . . . . . . . . . . Gaiping Zhang, Junqing Guo, and Xuannian Wang 13. Liposome-Enhanced Lateral-Flow Assays for the Sandwich-Hybridization Detection of RNA . . . . . . . . . . . . . . . . . . . . . . Katie A. Edwards and Antje J. Baeumner 14. Rapid Prototyping of Lateral Flow Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alexander Volkov, Michael Mauk, Paul Corstjens, and R. Sam Niedbala
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15. Lateral Flow Colloidal Gold-Based Immunoassay for Pesticide . . . . . . . . . . . . . . . Shuo Wang, Can Zhang, and Yan Zhang
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185 217
PART IV: LIGANDS 16. Synthesis of a Virus Electrode for Measurement of Prostate Specific Membrane Antigen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Juan E. Diaz, Li-Mei C. Yang, Jorge A. Lamboy, Reginald M. Penner, and Gregory A. Weiss 17. In Vivo Bacteriophage Display for the Discovery of Novel Peptide-Based Tumor-Targeting Agents. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jessica R. Newton and Susan L. Deutscher 18. Biopanning of Phage Displayed Peptide Libraries for the Isolation of Cell-Specific Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael J. McGuire, Shunzi Li, and Kathlynn C. Brown 19. Biosensor Detection Systems: Engineering Stable, High-Affinity Bioreceptors by Yeast Surface Display. . . . . . . . . . . . . . . . . . . . . . Sarah A. Richman, David M. Kranz, and Jennifer D. Stone 20. Antibody Affinity Optimization Using Yeast Cell Surface Display. . . . . . . . . . . . Robert W. Siegel 21. Using RNA Aptamers and the Proximity Ligation Assay for the Detection of Cell Surface Antigens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Supriya S. Pai and Andrew D. Ellington 22. In Vitro Selection of Protein-Binding DNA Aptamers as Ligands for Biosensing Applications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Naveen K. Navani, Wing Ki Mok, and Yingfu Li
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323 351
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PART V: PROTEIN AND DNA PREPARATION 23. Immobilization of Biomolecules onto Silica and Silica-Based Surfaces for Use in Planar Array Biosensors. . . . . . . . . . . . . . . . . . . Lisa C. Shriver-Lake, Paul T. Charles, and Chris R. Taitt 24. Rapid DNA Amplification Using a Battery-Powered Thin-Film Resistive Thermocycler . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Keith E. Herold, Nikolay Sergeev, Andriy Matviyenko, and Avraham Rasooly Index. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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000
Contributors SALVADOR ALEGRET • Group of Sensors & Biosensors, Autonomous University of Barcelona, Barcelona, Catalonia, Spain MAR ÁLVAREZ • CIBER BBN, Instituto de Microelectrónica de Madrid, Madrid, Spain ADRIANO AMBROSI • School of Chemical Sciences, Dublin City University, Dublin, Ireland ANTJE J. BAEUMNER • Department of Biological and Environmental Engineering, Cornell University, Ithaca, NY, USA ALESSANDRA BINI • Dipartimento di Chimica, Università degli Studi di Firenze, Sesto Fiorentino, Italy KATHLYNN C. BROWN • Division of Translational Research, Department of Internal Medicine and Simmons Comprehensive Cancer Center, University of Texas Southwestern Medical Center, Dallas, TX, USA LAURA G. CARRASCOSA • CIBER BBN, Instituto de Microelectrónica de Madrid, Madrid, Spain MARÍA TERESA CASTAÑEDA • Nanobioelectronics & Biosensors Group, Institut Català de Nanotecnologia, Barcelona, Catalonia, Spain Group of Sensors & Biosensors, Autonomous University of Barcelona, Barcelona, Catalonia, Spain PAUL T. CHARLES • Center for Bio/Molecular Science & Engineering, US Naval R esearch Laboratory, Washington, DC, USA PAUL CORSTJENS • Leiden University Medical Center, Leiden, The Netherlands ALFREDO DE LA ESCOSURA • Nanobioelectronics & Biosensors Group, Institut Català de Nanotecnologia, Barcelona, Catalonia, Spain SUSAN L. DEUTSCHER • Department of Biochemistry, University of Missouri, and Harry S. Truman Veterans Memorial Hospital, Columbia, MO, USA JUAN E. DIAZ • Department of Chemistry, University of California, Irvine, Irvine, CA, USA KATIE A. EDWARDS • Department of Biological and Environmental Engineering, Cornell University, Ithaca, NY, USA ANDREW D. ELLINGTON • Department of Biochemistry, University of Texas at Austin, Austin, TX, USA ARACELI GONZÁLEZ-CORTÉS • Department of Analytical Chemistry, Faculty of Chemistry, University Complutense of Madrid, Madrid, Spain JUNQING GUO • The Provincial Key Laboratory of Animal Immunology, Henan Academy of Agricultural Sciences, Zhengzhou, People’s Republic of China JAN HALÁMEK • Department of Analytical Biochemistry, University of Potsdam, Potsdam, Germany Biophysical Engineering Group, University of Twente, Enschede, The Netherlands xiii
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JOHN P. HART • Faculty of Health and Life Sciences, University of the West of England, Bristol, UK KEITH E. HEROLD • Fischell Department of Bioengineering, University of Maryland, College Park, MD, USA KAGAN KERMAN • Department of Chemistry, University of Western Ontario, London, ON, Canada DAVID M. KRANZ • Department of Biochemistry, University of Illinois, Urbana, IL, USA JORGE A. LAMBOY • Department of Chemistry, University of California, Irvine, Irvine, CA, USA SERENA LASCHI • Dipartimento di Chimica, Università degli Studi di Firenze, Sesto Fiorentino, Italy LAURA M. LECHUGA • CIBER BBN, Instituto de Microelectrónica de Madrid, Madrid, Spain SHUNZI LI • Division of Translational Research, Department of Internal Medicine and Simmons Comprehensive Cancer Center, University of Texas Southwestern Medical Center, Dallas, TX, USA YINGFU LI • Department of Biochemistry and Biomedical Sciences and Department of Chemistry, McMaster University, Hamilton, ON, Canada ALEXANDER MAKOWER • Department of Analytical Biochemistry, University of Potsdam, Potsdam, Germany MARCO MASCINI • Dipartimento di Chimica, Università degli Studi di Firenze, Sesto Fiorentino, Italy ANDRIY MATVIYENKO • Department of Engineering, University of Maryland, College Park, MD, USA FDA Center for Devices and Radiological Health, Silver Spring, MD, USA MICHAEL MAUK • School of Engineering and Applied Science, University of Pennsylvania, Philadelphia, PA, USA MICHAEL J. MCGUIRE • Division of Translational Research, Department of Internal Medicine and Simmons Comprehensive Cancer Center, University of Texas Southwestern Medical Center, Dallas, TX, USA ARBEN MERKOÇI • Nanobioelectronics & Biosensors Group, Institut Català de Nanotecnologia, Barcelona, Catalonia, Spain MARIA MINUNNI • Dipartimento di Chimica, Università degli Studi di Firenze, Sesto Fiorentino, Italy WING KI MOK • Department of Biochemistry and Biomedical Sciences and Department of Chemistry, McMaster University, Hamilton, ON, Canada RAJ MUTHARASAN • Department of Chemical and Biological Engineering, Drexel University, Philadelphia, PA, USA NAVEEN K. NAVANI • Department of Biochemistry and Biomedical Sciences and Department of Chemistry, McMaster University, Hamilton, ON, Canada JESSICA R. NEWTON • Department of Biochemistry, University of Missouri, Columbia, MO, USA
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R. SAM NIEDBALA • Department of Chemistry, Lehigh University, Bethlehem, PA, USA SUPRIYA S. PAI • Department of Microbiology, University of Texas at Austin, Austin, TX, USA ILARIA PALCHETTI • Dipartimento di Chimica, Università degli Studi di Firenze, Sesto Fiorentino, Italy ROY M. PEMBERTON • Faculty of Health and Life Sciences, University of the West of England, Bristol, UK REGINALD M. PENNER • Department of Chemistry, University of California, Irvine, Irvine, CA, USA JOSÉ M. PINGARRÓN • Department of Analytical Chemistry, Faculty of Chemistry, University Complutense of Madrid, Madrid, Spain JOSE A. PLAZA • CIBER BBN, Instituto de Microelectrónica de Madrid, Madrid, Spain AVRAHAM RASOOLY • FDA Center for Devices and Radiological Health, Silver Spring, MD, USA National Cancer Institute, Bethesda, MD, USA SARAH A. RICHMAN • Department of Biochemistry, University of Illinois, Urbana, IL, USA FRIEDER W. SCHELLER • Department of Analytical Biochemistry, University of Potsdam, Potsdam, Germany NIKOLAY SERGEEV • FDA Center for Devices and Radiological Health, Silver Spring, MD, USA LISA C. SHRIVER-LAKE • Center for Bio/Molecular Science & Engineering, US Naval Research Laboratory, Washington, DC, USA ROBERT W. SIEGEL • Abbott, Diagnostic Division, Abbott Park, IL, USA Eli Lilly and Company, Lilly Research Laboratories, Greenfield, IN, USA PETR SKLÁDAL • Department of Biochemistry, Masaryk University, Brno, Czech Republic JENNIFER D. STONE • Department of Biochemistry, University of Illinois, Urbana, IL, USA CHRIS R. TAITT • Center for Bio/Molecular Science & Engineering, US Naval Research Laboratory, Washington, DC, USA EIICHI TAMIYA • Department of Applied Physics, Graduate School of Engineering, Osaka University, Suita, Osaka, Japan CARSTEN TELLER • Department of Analytical Biochemistry, University of Potsdam, Potsdam, Germany SARA TOMBELLI • Dipartimento di Chimica, Università degli Studi di Firenze, Sesto Fiorentino, Italy MUN’DELANJI VESTERGAARD • School of Materials Science, Japan Advanced Institute of Science and Technology (JAIST), Nomi City, Ishikawa, Japan ALEXANDER VOLKOV • Department of Chemistry, Lehigh University, Bethlehem, PA, USA SHUO WANG • Key Laboratory of Food Nutrition and Safety, Ministry of Education, Tianjin Key Laboratory of Food Nutrition and Safety, Tianjin University of Science and Technology, Tianjin, China
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XUANNIAN WANG • The Provincial Key Laboratory of Animal Immunology, Henan Academy of Agricultural Sciences, Zhengzhou, People’s Republic of China GREGORY A. WEISS • Department of Chemistry, University of California, Irvine, Irvine, CA, USA PALOMA YÁÑEZ-SEDEÑO • Department of Analytical Chemistry, Faculty of Chemistry, University Complutense of Madrid, Madrid, Spain LI-MEI C. YANG • Department of Chemistry, University of California, Irvine, Irvine, CA, USA CAN ZHANG • Key Laboratory of Food Nutrition and Safety, Ministry of Education, Tianjin Key Laboratory of Food Nutrition and Safety, Tianjin University of Science and Technology, Tianjin, China GAIPING ZHANG • The Provincial Key Laboratory of Animal Immunology, Henan Academy of Agricultural Sciences, Zhengzhou, People’s Republic of China YAN ZHANG • Key Laboratory of Food Nutrition and Safety, Ministry of Education, Tianjin Key Laboratory of Food Nutrition and Safety, Tianjin University of Science and Technology, Tianjin, China KIRIL ZINOVIEV • CIBER BBN, Instituto de Microelectrónica de Madrid, Madrid, Spain.
Contents of Volume 503 Preface Contributors Contents of Volume 504
PART I: OPTICAL-BASED DETECTORS 1.
Surface Plasmon Resonance and Surface Plasmon Field-Enhanced Fluorescence Spectroscopy for Sensitive Detection of Tumor Markers Yusuke Arima, Yuji Teramura, Hiromi Takiguchi, Keiko Kawano, Hidetoshi Kotera, and Hiroo Iwata
2.
Surface Plasmon Resonance Biosensor for Biomolecular Interaction Analysis Based on Spatial Modulation Phase Detection Xiang Ding, Fangfang Liu, and Xinglong Yu Array-Based Spectral SPR Biosensor: Analysis of Mumps Virus Infection Jong Seol Yuk and Kwon-Soo Ha Optical Biosensors Based on Photonic Crystal Surface Waves Valery N. Konopsky and Elena V. Alieva Surface Plasmon Resonance Biosensing Marek Piliarik, Hana Vaisocherová, and Jirˇí Homola Label-Free Detection with the Resonant Mirror Biosensor Mohammed Zourob, Souna Elwary, Xudong Fan, Stephan Mohr, and Nicholas J. Goddard
3. 4. 5. 6.
7.
8.
Label-Free Detection with the Liquid Core Optical Ring Resonator Sensing Platform Ian M. White, Hongying Zhu, Jonathan D. Suter, Xudong Fan, and Mohammed Zourob Reflectometric Interference Spectroscopy Guenther Proll, Goran Markovic, Lutz Steinle, and Guenter Gauglitz
9.
Phase Sensitive Interferometry for Biosensing Applications Digant P. Davé 10. Label-Free Serodiagnosis on a Grating Coupler Thomas Nagel, Eva Ehrentreich-Förster, and Frank F. Bier
PART II: INDIRECT DETECTORS 11. CCD Camera Detection of HIV Infection John R. Day 12. Simple Luminescence Detector for Capillary Electrophoresis Antonio Segura-Carretero, Jorge F. Fernández-Sánchez, and Alberto Fernández-Gutiérrez 13. Optical System Design for Biosensors Based on CCD Detection Douglas A. Christensen and James N. Herron
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Contents of Volume 503
14. A Simple Portable Electroluminescence Illumination-Based CCD Detector Yordan Kostov, Nikolay Sergeev, Sean Wilson, Keith E. Herold, and Avraham Rasooly 15. Fluoroimmunoassays Using the NRL Array Biosensor Joel P. Golden and Kim E. Sapsford 16. Biosensors Technologies: Acousto-Optic Tunable Filter-Based Hyperspectral and Polarization Imagers for Fluorescence and Spectroscopic Imaging Neelam Gupta 17. Photodiode-Based Detection System for Biosensors Yordan Kostov 18. Photodiode Array On-chip Biosensor for the Detection of E. coli O157:H7 Pathogenic Bacteria Joon Myong Song and Ho Taik Kwon 19. DNA Analysis with a Photo-Diode Array Sensor Hideki Kambara and Guohua Zhou 20. Miniaturized and Integrated Fluorescence Detectors for Microfluidic Capillary Electrophoresis Devices Toshihiro Kamei 21. Photomultiplier Tubes in Biosensors Yafeng Guan 22. Integrating Waveguide Biosensor Shuhong Li, Platte Amstutz III, Cha-Mei Tang, Jun Hang, Peixuan Zhu, Yunqi Zhang, Daniel R. Shelton, Jeffrey S. Karns 23. Detection of Fluorescence Generated in Microfluidic Channel Using In-Fiber Grooves and In-Fiber Microchannel Sensors Rudi Irawan and Swee Chuan Tjin 24. Multiplex Integrating Waveguide Sensor: Signalyte™-II Shuhong Li, Yunqi Zhang, Platte Amstutz III, and Cha-Mei Tang 25. CCD Based Fiber-Optic Spectrometer Detection Rakesh Kapoor Index
Chapter 1 A Set of Piezoelectric Biosensors Using Cholinesterases Carsten Teller, Jan Halámek, Alexander Makower, and Frieder W. Scheller Summary Piezoelectric sensors have become a versatile tool in biosensorics to study protein–protein and protein– small molecule interactions. Here we present theoretical background on piezoelectric sensors and instructions, how to modify their surface with various recognition elements for cholinesterases. These recognition elements comprise an organophosphate (paraoxon), a cocaine derivative (BZE-DADOO), and a tricyclic, aromatic compound (propidium). Additionally, a guide to the kinetic evaluation of the obtained binding curves is given in this chapter. Key words: Piezoelectric biosensors, QCM, Cholinesterase, Pesticides, Drugs, Antibody, Kinetic evaluation.
1. Introduction Sense organs have developed during evolution by adapting the organism toward the properties of their environment. They recognize chemical signals by the binding to structurally complementary receptor areas followed by the transduction of this event into electrical nerve impulses. The biosensors represent the technical counter part by coupling the recognition by a biological recognition element, e.g. enzymes, antibodies, nucleic acids, receptors and recognition systems, e.g. organelles, cells or tissues, respectively, with a chemical or physical sensor (1). Different transducer types, e.g. electrochemical, optical, piezoelectric, thermal or magnetic transducers, transfer the signal from the recognition part to the electrical domain. Biosensors have been classified according to
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_1
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the biological recognition mechanism or the mode of signal transduction or, alternatively, a combination of the two. The characteristic feature of the biosensor is (according to the definition of IUPAC) the direct (spatial) contact of the recognition element and the transducer. This integration leads to a compact functional unit, which allows reusage of the biological component and miniaturization of the sensor body. These properties allow for online measurement and are the basis of the combination of different recognition elements on one transducer array resulting in the biochip. Recently, piezoelectric sensors have received growing attention as a tool for label-free detection. Because of the progress made in the field of microelectronics and microfluidics, the piezoelectric transducers have become a competitive alternative to surface plasmon resonance (SPR) devices and grating couplers. The piezoelectric effect was discovered by the Curie Brothers in 1880 (2). They found that certain (piezoelectric) materials can develop a standing acoustic wave upon electrical stimulation. The application of a force (Greek: πιεζειν = to press) results in a charge separation and hence a polarization of the crystal. A dipole moment perpendicular to the force and proportional to its value/ amount is created. Piezoelectric materials are often composed of crystals without a center of symmetry, for example, potassium sodium tartrate (Rochelle’s salt), lithium niobate and, most frequently encountered, quartz (SiO2). Next to their chemical composition, piezoelectric sensors can be distinguished by their mode of operation – thickness shear mode (TSM) sensors and surface acoustic wave (SAW) devices being the most prominent. TSM sensors consist of a thin, circular quartz plate with electrodes (often made of gold) on both sides. Between the two electrodes, a standing acoustic wave is generated. This wave is called a shear wave, since the crystal particles are displaced perpendicular to the direction of wave propagation. The wave length of the basic resonance is double the crystal thickness l0 = 2d; hence, the corresponding resonance frequency is inversely proportional to the quartz thickness f0 μ1/d. Sauerbrey (3) was the first to use a TSM as a quartz crystal microbalance (QCM). He postulated that a mass deposition on the surface formally only changes the thickness of the resonator, while all its other mechanical properties remain constant. Therefore, the change in frequency Δf that is caused by a mass deposition Δm on the surface can be described as follows: Df Dd Dm = =f0 d rq dA
(1)
Here rq denotes the density of quartz (rq = 2.65 g/cm3). A is the piezoelectrically active area the electrode. When the resonance condition f0 = cq/2d is put into Eq. 1, it results in:
A Set of Piezoelectric Biosensors
Df = -
2 f 02 Dm. c q rq A
5
(2)
The constant cq represents the propagation speed of the acoustic wave in quartz, its value being around 3330 m/s. Equation 2 is generally called the “Sauerbrey equation.” It does not only explain the proportional relation between mass deposition and change of frequency, it also allows calculating the theoretical sensitivity of a piezoelectric sensor. For example, with a typical 10 MHz resonator with an electrode diameter of 5 mm a sensitivity of 0.86 ng/Hz can be expected. However, the “Sauerbrey equation” in general only holds true if the sensor is placed in a vacuum or in a gaseous phase. That is why the usage of piezoelectric sensors was originally restricted to the gas phase, i.e., the measurement required a reproducible drying step and comparison with the “unreacted sensor” after the interaction between the sample and the recognition layer. Thus the measuring procedure was tedious and the difference between two signals of the same order resulted in a bad reproducibility. Nevertheless, it has been successfully used in such set-ups in various publications, which have been reviewed in (4). In reality, the measured frequency difference does not only depend on the change of mass, but also on other factors such as change of viscosity, density, and temperature of the medium that is in contact with the resonator. The influence of the temperature can almost be neglected by choosing the right cutting angle for the quartz crystals. So-called AT-cut crystals (cut at an angle of 35°15′ from the z-axis) show a high frequency stability over a large temperature range (5). In 1985, Kanazawa and Gordon (6) developed and experimentally verified a model for the effects of viscosity and density changes on the measured frequency. In a flow system, such complications can usually be avoided by using dilute analyte solutions and by keeping sample buffer and running buffer the same. In the 1980s, Kösslinger et al. succeeded in performing direct measurement in aqueous media and obtained comparable analytical quality for affinity biosensors using both SPR and quartz microbalances (7). This breakthrough was the beginning of a successful development of affinity piezosensors using antibodies and, later on, also nucleic acids for molecular recognition. When compared with other transducer technologies, piezoelectric sensors have several specific features: – The production and uniformity of QCM sensors is well established. – Binding events at the sensor’s surface generate the measuring signal directly (without additional marker). – Several independent sensors can be arranged to give a sensor array.
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– The sensitive area consists of a thin gold layer. Thus the wellknown “thiol surface chemistry” can be applied (8). – The gold electrode offers the combination with electrochemical addressing of the surface immobilized biomolecules, e.g. the activity of enzymes. However, piezoelectric biosensors also have a number of disadvantages: – Only binding events of macromolecules result in a direct measuring signal (for the measurement of small molecules “competitive assay formats” are necessary). – Changes of viscosity or temperature result in a falsification of the signal. Several piezoelectric sensors for the detection of low molecular weight compounds like pesticides (9–11), toxins (12), or drugs (13) have been reported in recent years (Table 1). A piezoelectric biosensor with a reversibly coupled (via His-tag) paraoxon for the binding of tetrameric butyrylcholinesterase to the active site of the enzyme has also been described by us (14). The commercially available devices have recently been reviewed (15, 16). Here we show how to modify the piezoelectric surface to develop affinity sensors for detection of small molecular compounds using antibodies and (acetyl)cholinesterases (AChE), respectively. The chemical structures of the described surface modifications are shown in Fig. The assays are performed in a competitive mode monitoring the binding of the free antibody or the noninhibited AChE to the immobilized ligand on the sensor. Furthermore, we show the ability of the piezosensor to monitor not only the binding of enzyme to the QCM, but also the determination of the surface activity of the bound AChE by amperometric measurements.
Table 1 Overview of different bioaffinity-based piezoelectric sensors Analyte class
Analyte
Surface modification
References
Pesticides
2,4-Dichlorophenoxyacetic acid
4-Aminothiophenol + 2,4-D
(9)
Atrazine
Cysteamine + carboxylated atrazine
(10)
Paraoxon, DFP, chlorpyriphos, chlorfenvinphos
MUA + BZE-DADOO
(11)
Toxins
Cholera toxin, tetanus toxin, pertussis toxin
Octanethiol + gangliosides
(12)
Drugs
Cocaine
MUA + BZE-DADOO
(13)
A Set of Piezoelectric Biosensors O Au
S
N H
O
H N
O
O O
N O P OO O
O Au
S
N H
HO
O Au
S
N H
7
O
O N O
O OH +
N HO
O
Ni2+ +
O
(His)6 OH
N H
O P O O O
O N O
O NH2 O Au
S
N H
N N
Fig. 1. Structures of the immobilized ligands used for cholinesterase recognition; BZE-DADOO, paraoxon-aminohexanol, paraoxon-hexahistidine-Ni2+-NTA-lysine, propidium (from top), each bound to MUA on gold.
2. Materials 1. Proteins – ChE (acetylcholinesterase), from electric eel, obtained via Sigma-Aldrich. – AChE from Drosophila melanogaster, kind gift from Prof. Fournier (IPBS Toulouse, France), enzyme expression described in (17). – BChE (butyrylcholinesterase), from human serum, kind gift from Roche Diagnostics (Mannheim, Germany). – Anti-cocaine-antibody, mouse monoclonal and sheep polyclonal IgG, kind gift from Boehringer (Mannheim, Germany). – Pepsin from hog stomach, obtained from Fluka (Buchs, Switzerland). – BSA (bovine serum albumin), from Boehringer (Mannheim, Germany). 2. Chemicals – Dry acetone (Fluka, Buchs, Switzerland) and dry nitrogen were used for cleaning of sensors.
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– MUA (11-mercaptoundecanoic acid), Sigma-Aldrich (Steinheim, Germany). – MU (11-mercaptoundecanol), Sigma-Aldrich (Steinheim, Germany). – TNTU (2-(5-norbornen-2,3-dicarboximido)-1,1,3,3-tetramethyluronium-tetrafluoroborat), from Calbiochem (San Diego, USA). – NMM (N-methyl-morpholine), from Fluka (Buchs, Switzerland). – DMF (dimethylformamide), from Roth (Karlsruhe, Germany). – NTA-lysine (Na,Na-Bis(carboxymethyl)-L-lysine), from SigmaAldrich (Steinheim, Germany). – DCC (dicyclohexylcarbodiimide), from Sigma-Aldrich (Steinheim, Germany). – BZE-DADOO (benzoylecgonine-1,8-diamino-3,4-dioxaoctane), kind gift from Boehringer (Mannheim, Germany). – PAH (paraoxon-aminohexanol) and paraoxon-hexahistidine, from Biosyntan (Berlin, Germany). – Nickel sulphate (NiSO4·7 H2O), from Laborchemie (Apolda, Germany). – EDTA (ethylenediaminetetraacetic acid), from Laborchemie (Apolda, Germany). – Coomassie Brilliant Blue (Bradford’s agent), from Pierce (Rockford, USA). – Acetylthiocholine and butyrylthiocholine (iodide salt), from Fluka (Buchs, Switzerland). – Formic acid (>98% purity), from Roth (Karlsruhe, Germany). – DTNB (5,5′-dithiobis(2-nitrobenzoic acid) ), Ellman’s agent, from Fluka (Buchs, Switzerland). – DFP (diisopropylfluorophosphate), chlorpyriphos and chlorfenvinphos, all from Dr. Ehrenstorfer company (Augsburg, Germany). – Phosphate buffer (50 mM) pH 7.5 (according to Sörensen), KH2PO4 and Na2HPO4 from Sigma-Aldrich (Steinheim, Germany). – Britton-Robinson-I buffer (40 mM) pH 8.0, boric acid, acetic acid, and phosphoric acid, adjusted with NaOH, all from Sigma-Aldrich (Steinheim, Germany). – Carbonate buffer (100 mM) pH 8.5, Na2CO3, and NaHCO3, from Sigma-Aldrich (Steinheim, Germany). 3. Equipment – Piezoelectric quartz crystals (10 MHz, thickness shear mode) were provided by ICMfg (Oklahoma, USA) or Elchema (Potsdam,
A Set of Piezoelectric Biosensors
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NY, USA), respectively. The sensors exhibited a basic resonance frequency of 10 MHz and bore gold electrodes (5 mm diameter) on both sides. – Microtitre plates “Maxisorp”, from Apogent (Denmark) (96well). – Peristaltic pump and Tygon tubes (Ø 0.2 mm), from Abimed Gilson (Langenfeld, Germany). – Frequency counter UZ2400, from Grundig (Brno, Czech Republic). – Piezoelectric instrument MultiLab 3900, from Kitlicˇka Company (Brno, Czech Republic). – Ag/AgCl reference microelectrodes, from Microelectrodes, Inc. (Bedford, USA).
3. Methods The methods described below comprise 1. Characterization of the antibodies and cholinesterases used. 2. Modification of the piezoelectric sensor with various recognition elements. 3. Flow system and assay formats. 4. Kinetic evaluation. 3.1. Characterization of the Antibodies and Cholinesterases Used
For a proper quantification of the QCM results and especially for a correct kinetic evaluation, it is necessary to know the protein concentration of the antibodies and cholinesterases used. In a microtitre plate version of Bradford’s assay (18), 150 μL of the protein sample diluted in 0.9% NaCl and 150 μL of Coomassie brilliant blue are mixed in the wells of standard 96-well microtiter plates. The resulting absorption at 405 nm is compared with a standard series of bovine serum albumin in 0.9% NaCl (0–20 μg/mL). The activity of cholinesterases is usually determined by Ellman’s method (19). This assay can also be carried out in microtitre plates. Thirty microliters of the ChE dilution and 30 μL of DNTB are added to 210 μL of 40 mM Britton-Robinson-I buffer pH 8.0. The reaction is started by addition of 30 μL of the appropriate acylthiocholine substrate. A microtiter plate reader is used to monitor the development of the absorption at 595 nm. The corresponding slope is used to calculate the esteratic activity.
3.2. Modification of Piezoelectric Sensors
Before any further modification, the quartz sensors should be cleaned for 2 h in acetone and dried with nitrogen.
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3.2.1. BZE-DADOO
1. Dissolve 1.77 mg MUA (8.0 μmol) and 2.9 mg TNTU (7.9 μmol) in a mixture of 1 μL of NMM (9.6 μmol) and 100 μL DMF. 2. Incubate this mixture for 15 min at room temperature. 3. Dissolve 0.84 mg BZE-DADOO in 100 mM carbonate buffer pH 8.5 to make 200 μL of a 10 mM solution. 4. Mix 22 μL of the MUA-TNTU solution and 200 μL of the BZE-DADOO solution and incubate for 90 min at room temperature (11). 5. The MUA-BZE-DADOO conjugate thus obtained can be stored in a freezer for future use. 6. The sensing layer is created by incubation of 30 μL of the MUA-BZE-DADOO conjugate on the electrode surface for 48 h at 4°C in a wet chamber. 7. Finally, wash the sensors with distilled water. After drying, the sensors are stored in the refrigerator.
3.2.2. Paraoxon
The preparation of the paraoxon-modified sensor is similar to the BZE-DADOO sensor. 1. Weigh 0.82 mg of the paraoxon-aminohexanol conjugate (6-aminohexyl)-(ethyl)-(4-nitrophenyl)-phosphate (PAH) (2.4 μmol) and dissolve in 200 μL 100 mM carbonate buffer pH 9.0. 2. Incubate the PAH solution with the MUA-TNTU solution (prepared as described in Subheading 3.2.1). PAH should be in slight molar excess. 3. Drop 30 μL MUA-paraoxon conjugate onto the gold electrodes to form a monolayer on the sensor. 4. Incubate for 48 h at 4°C in a wet chamber. 5. Wash with distilled water and dry for storage.
3.2.3. Paraoxon–His
The paraoxon–histidine conjugate was obtained from Biosyntan (Berlin, Germany) as a solute in 20% acetonitrile/80% water/0.05% trifluoroacetic acid (14). The MUA-lysine-NTA conjugate for the crystal surface modification is synthesized as follows: 1. Weigh 1.77 mg MUA (8.0 μmol) and 2.9 mg TNTU (7.9 μmol) and dissolve both in a mixture of 1 μL NMM (9.6 μmol) and 100 μL DMF. 2. Incubate for 15 min at room temperature. 3. Dissolve 1.0 mg NTA-lysine (2.65 μmol) in 200 μL carbonate buffer pH 8.5. 4. Mix 22 μL of the MUA-TNTU solution and the 200 μL NTAlysine and incubate for 90 min at room temperature. 5. The resulting conjugate can be stored in a freezer.
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6. Drop 20 μL of the MUA-NTA-lysine conjugate on each side of the piezosensor and incubate for 24 h at 4°C in a wet chamber. 7. Wash with distilled water and dry for storage. 3.2.4. Propidium
1. Incubate the cleaned sensors in a 3:1 mix of 5 mM MUA and 5 mM MU in ethanol for 48 h. 2. Prepare a 1 mM stock solution of propidium in DMF and a 5 mM stock solution of DCC in DMF. 3. Prepare the final reaction mixture by adding 5 volume parts of the propidium stock solution, 1 volume part of the DCC stock solution, and 14 volume parts of DMF. 4. Apply 40 μL of this mixture on each sensor side to provide a sufficient excess of reagents to the surface-bound MUA. Seal the sensors in a beaker containing silicagel (to keep the reaction compartment water-free) and incubate overnight. 5. Wash the sensors after the reaction with DMF, 96% ethanol, and deionized water. Dry with nitrogen and store for future use.
3.3. Flow System and Assay Formats
The central element of the flow system is a custom-made flow-through cell made from Plexiglas that holds the modified quartz crystal sensor (see Fig. 2 A–C). The quartz plate is sealed by rubber O-rings on both sides. Only one side of the crystal is in contact with the flowing solution. The internal volume of the cell was calculated to be 8 μL. A peristaltic pump is used to draw liquid via Tygon tubes (Ø 0.2 mm) from the measuring cell. By doing this, fluid pulsations are minimized and a lower noise of the frequency baseline can be obtained. The noise of the baseline typically ranges between 20 and 50 mHz. The experiments were performed at room temperature. Phosphate buffer (50 mM) pH 7.5 (Sörensen buffer) was used for the experiments described herein. The buffer and all other solutions were thoroughly degassed before using them in the flow system (see Note 1). The flow rate was adjusted to 20 μL/min. Depending on the length of the tube between sample container and measuring cell, there usually is a delay between the change of the solution and its entering of the cell of circa 60 s (see Note 2). Before using a fresh sensor buffer should be allowed to flow for 30 min to stabilize the baseline. Additionally, a 3 min flow of 1 mg/mL BSA solution is applied to saturate any nonspecific binding sites. The regeneration agent was either 2 mg/mL pepsin in 50 mM phosphate solution adjusted to pH 2 or 500 mM formic acid (BZE-DADOO-modified sensors), respectively (see Note 3). The suggested measuring cycle consisted of the following steps (20, 21):
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A
B
cell with QCM sensor Piezoelectric device
Peristaltic pump
Samples
C
D Inlet
Outlet
Outlet
Ag/AgCl reference electrode Inlet Titanium auxillary electrode
Rubber Q-ring QCM sensor Plexiglass cell
Fig. 2. Measuring set-up; (A) photograph of the piezoelectric device and flow system, the inset shows the cell holding the quartz sensor; (B) sample QCM sensor with 10 MHz base frequency (as used throughout the described experiments); (C) cross-section through the piezo-cell showing the two rubber O-rings holding the quartz plate, only one side of the sensor is in contact with the fluid; (D) cross-section of the cell used for combined piezoelectric and amperometric measurements, the lid also hold a titanium wire electrode and the Ag/AgCl reference electrode.
●
Initial phase: 3–5 min buffer flow to achieve a stable baseline recording.
●
Binding phase: injection of sample diluted in running buffer.
●
Washing phase: 5–10 min flow of buffer.
Regeneration phase: 3–10 min flow of regeneration agent. The preliminary experiments with antibodies were monitored by piezoelectric quartz crystals connected to a gate oscillating circuit, based on the integrated oscillator driver 74LS320. The output frequency was measured using the frequency counter UZ2400 (Grundig, Brno, Czech Republic) controlled by a computer. In the experiments with cholinesterase we used the integrated MultiLab 3900 piezoelectric instrument obtained from Kitlicˇka Company (Brno, Czech Republic). This device is able to measure the difference of the measuring quartz crystal to an internal 10 MHz reference crystal. Thus the theoretical resolution of this device is very low (1 mHz). The piezoelectric devices are connected to a computer via a serial adapter cable for device control. The software LabTools 1.1 (P. Skládal, Brno, Czech Republic) was used for measurements, data acquisition, storage, and evaluation. The obtained frequency difference signal is recorded at a rate of one point per second. These data are stored in a binary format ●
A Set of Piezoelectric Biosensors
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and converted to ASCII tables of floating point values for further evaluation (e.g. MS Excel or Originlab’s Origin). Other frequency counters were reviewed by O’Sullivan and Guilbault (16). Since the gold electrode on the quartz can also be used as a working electrode, it is possible to measure the activity of the surface-bound enzyme amperometrically. Therefore, a custommade flow-through cell with integrated reference and auxiliary electrodes was constructed to hold the quartz crystal sensor (see Fig. 2D). The same thio-substrate as for the Ellman assay (see Subheading 3.1) was used for activity measurements. The esterolytic reaction generates thiocholine that can be oxidized at the working electrode at a sufficiently positive potential (+200−300 mV vs. Ag/ AgCl). A three-electrode setup with a platinum auxiliary electrode was used for amperometric measurements. 3.3.1. BZE-DADOO-Sensor for Cocaine Detection
One measuring cycle consisted of the following steps: – 1 min flow of the phosphate buffer to achieve a stable initial frequency. – 3−10 min flow of the buffer solution containing antibody (in selected experiments a mixture of antibody and cocaine was used); recording of the binding curve. – 5 min flow of the buffer to determine the resulting final change of frequency. – 4 min flow of 500 mM formic acid to dissociate the ligand–antibody complex and regenerate the sensing surface of the crystal. The determination of free cocaine was performed competitively. In a plastic microtube, the selected amount of antibody (14 μg/mL) was mixed and stirred shortly with various cocaine solutions 0.1–1,000 nM in phosphate buffer). The concentration of antibody was chosen to achieve an appropriate response of the piezosensor, i.e., here it represented a signal of approximately 35 Hz. Here the experimenter usually has to find a compromise between a large binding signal (broad signal range for the competitive assay) and an antibody concentration as low as possible (low limit of detection). The mixture was allowed to incubate for a selected time interval (for a high-affinity antibody 30 min should be enough). Then it was introduced to the flow system and the binding curve was recorded. With this system we could reach a lower limit of detection of 0.1 nM for cocaine (21).
3.3.2. BZE-DADOO-Sensor for Organophosphate Detection
One measuring cycle consisted of the following steps: – 3 min flow of the phosphate buffer to achieve a stable initial frequency. – 5 min flow of the buffer solution containing cholinesterase (preincubated with inhibitor if applicable); recording of the binding curve.
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– 5 min flow of buffer to determine the resulting change of frequency. – 5 min flow of the regeneration solution. – 5 min flow of the phosphate buffer to achieve a stable initial frequency. For regeneration, 500 mM formic acid solution was used to dissociate the BZE-DADOO–cholinesterase complex. The sensor was used for more than 40 regeneration steps. Figure 3 shows the binding records for various concentrations of butyrylcholinesterase. The binding of the enzyme to the immobilized BZE-DADOO could be detected at protein concentration down to 5 μg/mL. For the experiments with DFP, a certain BChE concentration was chosen (25 μg/mL) and incubated with different concentrations of DFP for 30 min. The results are shown in Fig. 4. The organophosphate could be detected at concentrations down to 0.1 nM. In this case an additional injection step was introduced to deposit the paraoxon-histidine on the sensor surface, which otherwise would not exhibit a recognition element for the used butyrylcholinesterase (see Note 3). For the competitive measurements 0.1 mg/mL BChE (270 nM) is mixed with various concentrations of DFP (ranging from 0.1 to 10 μM) and incubated for 30 min at room temperature (results shown in (14)). – 3 min flow of the phosphate buffer to achieve a stable initial frequency.
3.3.3. Sensor Modified with NTA-Lysine
Δf [Hz]
5 µg/mL
0
10 µg/mL 20 µg/mL
−10
50 µg/mL
−20
100 µg/mL 200 µg/mL
−30 −40 −50 0
2
4
6
8
10
t [min]
Fig. 3. Binding curves for BChE binding to a BZE-DADOO-modified piezosensor; the dashed line indicates the zero baseline. The limit of detection is 5 μg/mL (=1.4 × 10−8 mol/L) of butyrylcholinesterase.
A Set of Piezoelectric Biosensors
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0 10−5
Δf [Hz]
−2
10−6
−4 10−7 −6 10−8 −8
10−9
−10
10−10 0
0
2
4
6
8
t [min] Fig. 4. Concentration dependence for DFP; 25 μg/mL BChE were pre-incubated with various, molar concentrations of DFP (given next to the corresponding binding trace). The limit of detection is 10−10 mol/L.
– 10 min flow of 1 mM nickel sulphate to form the Ni2+–NTA chelate complex. – 10 min flow a buffer to remove excess nickel ions. – 10 min flow of the paraoxon–histidine conjugate to form the recognition layer. – 10 min flow of running buffer. – 10 min flow of BChE solution (pre-incubated with inhibitor if applicable). – 10 min flow of 0.5 M EDTA to regenerate the surface. 3.3.4. Sensor Modified with Propidium
One measuring cycle consisted of the following steps: – 3 min flow of the phosphate buffer to achieve a stable initial frequency. – 5 min flow of the buffer solution containing cholinesterase; recording of the binding curve. – 5 min flow of buffer to monitor the dissociation from the surface. – 5 min flow of the regeneration solution. In this case we had to use 2 mg/mL pepsin in phosphate solution at pH 2 to achieve a reproducible regeneration. Figure 5 shows a comparison of formic acid and pepsin as regeneration agents. In both cases 50 μg/mL Drosphila AChE was allowed to interact with the propidium-modified QCM. In case of pepsin, a
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Regeneration
Buffer
Buffer
0 −20
Δf [Hz]
−40 −60 −80 −100 Pepsin pH 2
−120
Formic acid
−140 0
5
10
15
20
25
30
t [min] Fig. 5. Influence of different regeneration solutions on the binding of Drosophila AChE, comparison of pepsin pH 2 (injected for 3 min, solid line) and formic acid (injected for 10 min, dashed line).
short injection (3 min) is sufficient to remove the AChE from the surface and to reach the baseline after switching back to running buffer. The regeneration with formic acid is less efficient. Although a longer injection time (10 min) is applied, the curve does not reach the initial baseline after switching back to running buffer. 3.3.5. Propidium-Sensor for Combined Measurements
The propidium sensor setup (see Subheading 3.3.4) can also be utilized to monitor the activity of the surface-bound acetylcholinesterase. The original measuring cycle is extended by the injection of substrate solution and an additional washing step: – 3 min flow of the carrier buffer to stabilize the baseline signal. – 3 min flow of the AChE solution; recording of the binding curve. – 5 min flow of the carrier buffer to wash unbound AChE and determine the resulting change of frequency. – 5 min of 500 μM acetylthiocholine iodide in buffer; start of the amperometric measurement and recording of the current. – 5 min flow of the carrier buffer to remove substrate and product solution. – 8 min flow of 2 mg/mL pepsin. The sensor can be used for multiple substrate injections after the AChE loading and for repeated AChE immobilization after sensor regeneration. AChE from the electric eel could be specifically bound in concentration down to 5 μg/mL (=21 nM) (22).
A Set of Piezoelectric Biosensors
3.4. Kinetic Evaluation
17
Piezoelectric sensors can be used to study affinity interactions and to derive their kinetic constants. Like surface plasmon resonance (SPR) devices or grating couplers, they are able to monitor the time course of the binding reaction. For this purpose, it does not matter which one of the binding partners is immobilized on the sensor chip as long as the mobile affinity partner is heavy enough to provide a significant mass change upon binding to the immobilized compound. Since most of the equations used are derived from their respective SPR counterparts, we will use a similar nomenclature. The immobilized affinity molecule is called “the ligand (L)”. The mobile binding partner (in the flow system) is called “the analyte (A)”. Unless stated differently, the capital letter “K” will be used to describe equilibrium constants. A small “k” will denote the respective rate constants. For a simple one-to-one binding reaction such as L+A
kon ¾¾¾ ® LA. ¬¾¾ ¾ koff
(3)
one can derive two equations from the law of mass action: – A bimolecular (second order) association equilibrium K Ass =
[LA ] = kon . [L ][A ] koff
(4)
– Or a first-order dissociation equilibrium K Diss =
[L ]× [A ] = 1 . [LA ] K Ass
(5)
The corresponding rate equation for the formation of the ligand–analyte complex is as follows: d [LA ] = kon [L ][A ] - koff [LA ]. dt
(6)
The amount of free ligand [L] can be replaced by the difference of the initial concentration of ligand [L]0 and the concentration of occupied ligand [LA]. d [LA ] = kon [L ]0 - [LA ] [A ] - koff [LA ]. dt
(
)
(7)
This rate equation has then to be translated to represent the measured frequency changes. According to the Sauerbrey equation (see Eq.2), the measured frequency difference is proportional to the mass, i.e. the amount, of analyte bound by the ligand [LA]. Δfmax is used to denote the maximum frequency shift upon complete
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saturation of all ligand molecules on the sensor surface. The concentration of the free analyte is quasi-constant, as we worked in a flow system, i.e. with a continuous supply of analyte. dDf = kon (Df max - Df )[A ]0 - koff Df dt = kon Df max [A ]0 - kon Df [A ]0 - koff Df = kon Df max [A ]0 - ékon [A ]0 + koff ù Df ë û
(8)
The complete equation to describe the binding time curves is obtained by integration of the rate equation. Df (t ) =
kon Df max [A ]0 é - (kon [A ]0 + koff )t ù × 1-e úû kon [A ]0 + koff êë
(9)
As most experiments include a certain baseline recording before the actual binding experiment, it is useful to add an offset t0 in the integrated equation. This offset time represents the time point at which the sample reaches the measuring chamber/cell and the binding events can occur. Df (t ) =
kon Df max [A ]0 æ (kon [A ]0 +koff )×(t 0 -t )ö × ç1 - e ÷ø kon [A ]0 + koff è
(10)
A simplified two-step fitting procedure has found application in many publications (21, 23). For the sake of completeness, this approach shall also be discussed here. It requires the following substitutions and results in a pseudo-first order rate equation. kobs = kon [A ]0 + koff Df Eq =
kon [A ]0 Df max kobs
k t -t Df (t ) = Df Eq éë1 - e obs ( 0 ) ùû
(11)
This method has two major disadvantages. First, it requires two fitting steps, i.e. the nonlinear regression of each individual curve with the pseudo-first order exponential equation and a linear fit of the obtained kobs values vs. the analyte concentration. Second, the two parameters kobs and ΔfEq in the exponential equation (Eq. 11) are not independent of each other, which is a major prerequisite for nonlinear regression analysis. The equation for the association phase (Eq. 10) can also be used to describe the time course of the dissociation phase. For this purpose, the concentration of the analyte [A]0 is set to zero. As an additional constraint, the parameter ΔfA is introduced. It can represent the equilibrium frequency ΔfEq or any
A Set of Piezoelectric Biosensors
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other frequency difference that has been reached during the association phase before the injection of analyte-free buffer solution. k t -t Df (t ) = Df A e off ( 0 )
(12)
In this case, t0 represents the time at which the buffer flow enters the measuring chamber and replaces the analyte solution. This equation is particularly useful for global fitting approaches. It allows deriving the dissociation rate constant koff from the dissociation phase and its subsequent use as a fixed parameter in the nonlinear regression of the association phase. As an example, Table 2 shows the rate and equilibrium constants for AChE from electric eel binding to various modified surfaces (20, 24). If you use very long injection times, the binding curve will eventually approach its equilibrium (as required by Eq. 3). As time t tends towards infinity Eq. 11 changes to the well-known Langmuir isotherm. In other words – a monomolecular layer of analyte is formed at the sensor surface. lim Df (t ) = Df max t ®¥
[A ]0 [A ]0 + K D
(13)
= Df Eq
As Eq. 13 shows, this feature can be used to derive the dissociation equilibrium constant KD directly from the frequency difference at the end of measurement ΔfEq. The main drawback of this method is that long-term experiments and large quantities of analyte are required. Figure 6 shows a simulated binding curve in a piezoelectric measurement. A normal experiment is stopped by switching back to buffer flow and thus starting the dissociation phase. It is also obvious from Fig. 6 that at least a three times longer injection of analyte is required to reach a Δf near the equilibrium.
Table 2 Kinetic and equilibrium constants for electric eel AChE obtained by fitting of the binding curves with Eqs. 1.10 and 1.12 (24) Ligand
kon
+/−
koff
+/−
KAss
Paroxon
6,327
1,027
1.16 × 10−3
5.8 × 10−4
5.45 × 106
BZE-DADOO 4,311
694
2.46 × 10−3
6.3 × 10−4
1.75 × 106
Propidium
1,939
3.52 × 10−4
8.1 × 10−5
3.69 × 107
12,987
The rate constants are stated together with their standard deviation. kon in L/(mol·s), koff in 1/s, KAss in L/mol
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Fig. 6. Simulation of binding experiment, parameters used: c = 2.5 × 10−7 mol/L, t0 = 180 s, Δfmax = −150 Hz, kon = 6,000 L/(mol s) and koff = 0.002 L/s.
4. Notes 1. When working with a flow system, bubbles in measuring cell should be avoided if possible. Therefore, all running buf fers should be degassed thoroughly before usage. If the cell is made of a clear material, e.g. Plexiglas, bubbles can be easily recognized. If you have to work with an opaque cell, frequency jumps when stopping the flow upon buffer/solution change can be good indicator of bubbles inside the cell. 2. The mentioned 20 μL/min should be considered a guide value. In several cases, e.g. larger or smaller cells, longer tubing from sample reservoir to the cell, it can be better to use a faster or slower flow rate. In any case, a constant value should be chosen for all experiments to obtain consistent results. 3. The choice of a proper regeneration agent can be crucial for successful experiments. A low pH, e.g. 0.5–1 mol/L formic acid, can be sufficient to break the affinity interaction of antibodies. In case of acetylcholinesterases interacting with propidium-modified sensors, we found the combination of low pH and enzymatic digestion of the surface-bound protein by pepsin to be more efficient. The paraoxon-hexahistidine-Ni2+NTA-lysine sensor provided a different opportunity for regeneration. After each measurement the Ni2+ ions were chelated by EDTA and a new recognition layer was applied. The usage of special “regeneration cocktails” as used for SPR devices was not tested by us, but could still provide similar results.
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Acknowledgment The authors thank the European Union (Marie-Curie-fellowship HPMD-CT-2001-00062) and the “International Max Planck Research School on Biomimetic Systems” (PhD scholarship) for financial support. References 1. Thevenot, D. R., Toth, K., Durst, R. A., and Wilson, G. S. (2001) Electrochemical biosensors: recommended definitions and classification. Biosensors & Bioelectronics 16, 121–131 2. Curie, J., and Curie, P. (1880) Développement, par pression, de l’électricité polaire dans les cristaux hémièdres à faces inclinées. Comptes rendus de l’Académie des sciences 91, 294–295 3. Sauerbrey, G. (1959) Verwendung Von Schwingquarzen Zur Wägung Dünner Schichten Und Zur Mikrowägung. Zeitschrift Für Physik 155, 206–222 4. Bunde, R. L., Jarvi, E. J., and Rosentreter, J. J. (1998) Piezoelectric quartz crystal biosensors. Talanta 46, 1223–1236 5. Janshoff, A., Galla, H. J., and Steinem, C. (2000) Piezoelectric mass-sensing devices as biosensors – an alternative to optical biosensors? Angewandte Chemie-International Edition 39, 4004–4032 6. Kanazawa, K. K., and Gordon, J. G. (1985) Frequency of a Quartz Microbalance in Contact with Liquid. Analytical Chemistry 57, 1770–1771 7. Kößlinger, C., Uttenthaler, E., Abel, T., Hauck, S., and Drost, S. (1998) Comparison of the determination of affinity constants with surface plasmon resonance and quartz crystal microbalance, in Eurosensors XII (White, N. M., Ed.), Southampton, UK, pp. 845–848 8. Mrksich, M., and Whitesides, G. M. (1996) Using self-assembled monolayers to understand the interactions of man-made surfaces with proteins and cells. Annual Review of Biophysics and Biomolecular Structure 25, 55–78 9. Halamek, J., Hepel, M., and Skladal, P. (2001) Investigation of highly sensitive piezoelectric immunosensors for 2,4-dichlorophenoxyacetic acid. Biosensors & Bioelectronics 16, 253–260 10. Pribyl, J., Hepel, M., Halámek, J., and Skladal, P. (2003) Development of piezoelectric immunosensors for competitive and direct determination of atrazine. Sensors and Actuators B-Chemical 91, 333–341
11. Halamek, J., Pribyl, J., Makower, A., Skladal, P., and Scheller, F. W. (2005) Sensitive detection of organophosphates in river water by means of a piezoelectric biosensor. Analytical and Bioanalytical Chemistry 382, 1904–1911 12. Janshoff, A., Steinem, C., Sieber, M., elBaya, A., Schmidt, M. A., and Galla, H. J. (1997) Quartz crystal microbalance investigation of the interaction of bacterial toxins with ganglioside containing solid supported membranes. European Biophysics Journal with Biophysics Letters 26, 261–270 13. Halamek, J., Makower, A., Skladal, P., and Scheller, F. W. (2002) Highly sensitive detection of cocaine using a piezoelectric immunosensor. Biosensors & Bioelectronics 17, 1045–1050 14. Makower, A., Halámek, J., Skladal, P., Kernchen, F., and Scheller, F. W. (2003) New principle of direct real-time monitoring of the interaction of cholinesterase and its inhibitors by piezolectric biosensor. Biosensors & Bioelectronics 18, 1329–1337 15. Halámek, J., Hepel, M., and Skladal, P. (2001) Investigation of highly sensitive piezoelectric immunosensors for 2,4-dichlorophenoxyacetic acid. Biosensors and Bioelectronics 16, 253–260 16. O’Sullivan, C. K., and Guilbault, G. G. (1999) Commercial quartz crystal microbalances – theory and applications. Biosensors & Bioelectronics 14, 663–670 17. Chaabihi, H., Fournier, D., Fedon, Y., Bossy, J. P., Ravallec, M., Devauchelle, G., and Cerutti, M. (1994) Biochemical characterization of Drosophila melanogaster acetylcholinesterase expressed by recombinant baculoviruses. Biochemical and Biophysical Research Communications 203, 734–742 18. Bradford, M. M. (1976) Rapid and sensitive method for quantitation of microgram quantities of protein utilizing principle of protein-dye binding. Analytical Biochemistry 72, 248–254 19. Ellman, G. L., Courtney, K. D., Andres, V., and Featherstone, R. M. (1961) A new and rapid colorimetric determination of acetylcholinesterase activity. Biochemical Pharmacology 7, 88–95
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20. Teller, C., Halámek, J., Makower, A., Fournier, D., Schulze, H., and Scheller, F. W. (2006) A piezoelectric sensor with propidium as a recognition element for cholinesterases. Sensors and Actuators B-Chemical 113, 214–221 21. Halamek, J., Makower, A., Knosche, K., Skladal, P., and Scheller, F. W. (2005) Piezoelectric affinity sensors for cocaine and cholinesterase inhibitors. Talanta 65, 337–342 22. Halámek, J., Teller, C., Makower, A., Fournier, D., and Scheller, F. W. (2006)
EQCN based cholinesterase biosensors. Electrochimica Acta 51, 5174–5181 23. Skladal, P. (2003) Piezoelectric quartz crystal sensors applied for bioanalytical assays and characterization of affinity interactions. Journal of the Brazilian Chemical Society 14, 491–502 24. Halamek, J., Teller, C., Makower, A., Fournier, D., and Scheller, F. W. (2006) EQCN based cholinesterase biosensors. Electrochimica Acta 51, 5174–5181
Chapter 2 Piezoelectric Biosensors for Aptamer–Protein Interaction Sara Tombelli, Alessandra Bini, Maria Minunni, and Marco Mascini Summary Aptamers can be considered as a valid alternative to antibodies or other biomimetic receptors for the development of biosensors and other analytical methods. The production of aptamers is commonly performed by the SELEX (Systematic Evolution of Ligands by Exponential Enrichment) process, which, starting from large libraries of oligonucleotides, allows the isolation of large amounts of functional nucleic acids by an iterative process of in vitro selection and subsequent amplification through polymerase chain reaction. Aptamers are suitable for applications based on molecular recognition as analytical, diagnostic, and therapeutic tools. The use of aptamers as biorecognition element in piezoelectric biosensors will be here reported with particular application to the detection of thrombin. Key words: Biosensor, DNA, Aptamers, Thrombin, Quartz crystal microbalance.
1. Introduction Aptamers are single-stranded DNA or RNA ligands, which can be selected for different targets starting from a large library of molecules containing randomly created sequences (1). The selection process is called SELEX (Systematic Evolution of Ligands by Exponential enrichment), first reported in 1990 (2, 3). The SELEX process involves iterative cycles of selection and amplification starting from a large library of oligonucleotides with different sequences (generally 1015 different structures). After the incubation with the specific target and the partitioning of the binding from the nonbinding molecules, the oligonucleotides that are selected are amplified to create a new mixture enriched in those nucleic acid molecules having a higher affinity for the target. After several cycles Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_2
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of the selection process, the pool is enriched in the high affinity sequences at the expense of the low affinity binders. Aptamers can offer advantages over antibodies that make them very promising for analytical applications (4, 5). The main advantage is the overcoming of the use of animals or cell lines for the production of the molecules. Antibodies against molecules that are not immunogenic are difficult to generate. On the contrary, aptamers are isolated by in vitro methods that are independent on animals, since an in vitro combinatorial library can be generated against any target. Moreover, the aptamer selection process can be manipulated to obtain aptamers that bind a specific region of the target and with specific binding properties in different binding conditions. After selection, aptamers are produced by chemical synthesis and purified to a very high degree by eliminating the batch-to-batch variation found when using antibodies. Another advantage over antibodies can be seen in the higher temperature stability of aptamers; in fact antibodies are large proteins sensitive to the temperature and they can undergo irreversible denaturation. On the contrary, aptamers are very stable and they can recover their native active conformation after denaturation. Besides antibodies, the selections process itself, with the amplification step, gives some advantages to aptamer with respect to other “nonnatural” receptors, such as oligopeptides, which cannot be amplified during their selection procedure. The additional attractive aspect of aptamers for researchers in the analytical field is the wide range of molecules for which they can be selected, including organic dyes (6), amino acids (7), antibiotics (8), peptides (9), proteins (10), vitamins (11), and also whole cells (12) or micro-organisms such as bacteria (13). With respect to their application, aptamers were selected in the past mainly for their use as therapeutic agents. In addition to the therapeutic field, aptamers have been then used in several analytical methodologies, such as affinity chromatography, capillary electrophoresis (14), mass spectrometry (15), or biosensors (1). These aptamers-based methods have been mainly employed in the clinical area for the development of diagnostic assays. A very attracting application is the exploitation of aptamers as biorecognition elements in biosensors and the majority of the reported aptamer-based biosensors make use of the thrombinbinding aptamer (15-mer, 5¢-GGTTGGTGTGGTTGG-3¢). This DNA aptamer was the first one selected in vitro, specific for a protein without nucleic acids-binding properties (5), and it has been studied as anticlotting therapeutic tool (16, 17). The thrombin-binding aptamer has been extensively investigated: its G-quartet structure has been established (18, 19) and the binding site has been identified (20). This aptamer has been coupled to different transduction principles to demonstrate the wide
Piezoelectric Biosensors for Aptamer-Protein Interaction
25
applicability of aptamers as bioreceptors in biosensors (21–27). These studies clearly indicate the suitability of aptamer-based systems for analytical applications. Thrombin (factor IIa) is the last enzyme protease involved in the coagulation cascade, and it converts fibrinogen to insoluble fibrin that forms the fibrin gel either in physiological conditions or in a pathological thrombus (28). Thrombin has also hormonelike properties, and it is involved in thrombosis and platelet activation. Therefore, thrombin plays a central role in a number of cardiovascular diseases (29), and it is thought to regulate many processes in inflammation and tissue repair at the vessel wall. The concentration of thrombin in blood can vary considerably: thrombin, not present in blood under normal conditions, can reach low micromolar concentrations during the coagulation process but low levels (low nM) of thrombin generated early in haemostasis are also important to the overall process (30). Out of the haemostatic process, circulating thrombin has been detected at high pM range in blood of patients suffering from diseases known to be associated with coagulation abnormalities. This chapter deals with the coupling of the thrombin aptamer with quartz crystal microbalance devices for the development of aptamer-based piezoelectric biosensors. The term “piezoelectric” derived from the Greek word piezen meaning “to press.” The first investigation on the piezoelectricity was performed in 1880 by Jacques and Pierre Curie (31), who observed that a mechanical stress applied to the surfaces of various crystals caused a corresponding electrical potential across the crystal, whose magnitude was proportional to the applied stress. The Curies also verified the converse piezoelectric effect in which application of a voltage across these crystals caused a corresponding mechanical strain. Application of an alternating electric field across the crystal substrate results in an alternating strain field. This causes a vibrational, or oscillatory, motion in the crystal, resulting in the generation of acoustic standing waves. Depending on various criteria, the oscillator vibrates at a characteristic resonant frequency. The most used devices in biosensors are generally bulk acoustic wave (BAW)-based employing AT-cut quartz crystals. An AT-cut crystal is typically cut at an angle of + 35°15¢ and has a zero frequency temperature coefficient at or near room temperature that results in minimal frequency changes due to temperature (32). AT-cut crystals oscillate in the thickness shear mode (33). The first quantitative investigation of the piezoelectric effect was performed by Sauerbrey, who derived the relationship for the change in frequency DF (in Hz) caused by the added mass Dm (in g): DF = -
2F02 A mQ rQ
× Dm
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Tombelli et al.
where F0 is the fundamental resonant frequency of unloaded quartz, mQ is the shear modulus of AT-cut quartz (2.947 × 1011 g cm−1s−2), rQ is the density of the quartz (2.648 g cm−3) and A is the surface area in cm2 (34). The Sauerbrey equation assumes a uniform distribution of mass on the entire electrode portion of an AT-cut quartz crystal. Mass sensitivity decreases monotonically with the radius, in a Gaussian manner becoming negligible at and beyond the electrode boundary (35). Another assumption of this equation is that the mass added or lost at the crystal surface does not experience any deformation during the oscillation: this is true for thin, rigid layers. For thicker, less rigid layers, a more complex theory is necessary. Piezoelectric crystals have been used as microbalances and as a microviscometer owing to their small size, high sensitivity, simplicity of construction and operation, light weight and the low power required (36). The quartz–crystal microbalance (QCM) has traditionally been used in many applications such as thin film deposition control, etching studies, aerosol mass measurements, and space system contamination studies (37). Recently, however, the interest in the application of piezoelectric devices in the field of analysis has increased, since it was realized that many opportunities for molecular sensing can be opened up once a suitable recognition layer or molecule is coated on the crystal. In particular, piezoelectric biosensors have found a wide range of applications in food (38, 39), environmental, and clinical (40, 41) analysis.
2. Materials 2.1. Aptamer and Target Protein
1. a-Thrombin from human plasma, MW 37 kDa (Sigma, Milan, Italy) (Product number: T6884). Thrombin is shipped as lyophilized powder containing 417 NIH Units of protein (3,093 NIH Units/mg). Thrombin stock solution (aliquots: 20 mL, 3.6 mM in binding buffer) was stored at −20°C and for the experiments further dilutions in binding buffer were stored in ice during measurements. 2. 15-mer DNA aptamer for thrombin (MWG Biotech, Milan, Italy) with the following sequence: 5¢ biotin TTTTTTTTTTTTTTTTTTTT GGT TGG TGT GGT TGG 3¢ (42). The aptamer (MW 5053 g/mol) is received lyophilized and then diluted in MilliQ water. The diluted aptamer can be stored at −20°C. The aptamer is biotinylated a the 5¢ end to allow its immobilization onto streptavidin-modified surfaces. The insertion of the polyT (20-mer) tail at the 5¢ end
Piezoelectric Biosensors for Aptamer-Protein Interaction
27
of the aptamer sequence is necessary for the optimal immobilization of the aptamer onto a solid support (see Note 1). 3. Immobilization buffer: 300 mM NaCl, 20 mM Na2HPO4, 0.1 mM EDTA, pH 7.4. 4. Binding buffer: 50 mM Tris−HCl, 140 mM NaCl, 1 mM MgCl2, pH 7.4. 5. Human serum albumin (Sigma, Milan, Italy) (Product number: A9511) was used as negative control. Albumin was shipped lyophilized and stored at 4°C. Further dilutions were prepared in binding buffer and used after preparation. 6. Human serum from clotted male whole blood (Sigma, Milan, Italy) (Product number: H1388) was used to study the effect of a complex matrix. Serum can be divided into aliquots and stored at −20°C. Human citrated plasma (Sigma, Milan, Italy) (Product number: P9523) was used as naturally containing complex matrix. 2.2. Other Reagents
1. Streptavidin, bis-2-methoxyethyl ether (diglyme), bromoacetic acid, bovine serum albumin, ethanolamine hydrochloride, Tween 20, mercaptoundecanol, N-(3-Dimethylaminopropyl)N¢-ethylcarbodiimide hydrochloride (EDAC) (Sigma, Milan, Italy). 2. Ammonium sulphate, NH3, H2O2, ethanol, NaOH, EDTA (Merck, Milan, Italy). 3. Epichlorohydrin, Milan, Italy).
N-hydroxysuccinimide
(NHS)
(Fluka,
4. Dextran T500 (Amersham Biosciences Europe, Milan, Italy). 5. NAP™ 10 Columns (17–0854–01, GE Healthcare, Uppsala, Sweden) as exchanging buffer system for plasma purification. 2.3. Instrumentation
1. The complete measuring system is illustrated in Fig. 1. 9.5 MHz AT cut quartz crystals (14 mm, 165 mm) with gold evaporated on both sides (42.6 mm2 area, Ø 7.4 mm) (International Crystal Manufacturing (USA) ). The quartz crystal (Fig. 1A) is housed inside the measurement cell such that only one side of the resonator is in contact with the solution in the cell well; in this way two measurement series can be performed on the same resonator, one for each side. The cell is made of methacrylate, which is resistant and inert toward the chemicals used in the experiment, rigid, allowing to fix it to a support with a pincer and transparent, so that it is possible to observe any anomalies (air bubbles) that could be present into the well. The cell consists in two blocks of methacrylate, which are hold together by two screws (Fig. 1B). The crystal is housed between these two blocks, and it is sandwiched between two o-rings. This kind of cell is used for batch measurements, and the solution of the reagents is inserted into the
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Fig. 1. (A) Quartz crystal and working cell; (B) Schematic view of the working cell. The crystal is hold between two σ-rings, and the sample is injected into the cell well. Only one side of the crystal is in contact with the solution; (C) Photograph of the whole measuring system with the cells and the quartz crystal analyser.
cell by pipetting it directly in the upper cell well. The volume of the well is around 1 mL but for the typical volumes used the measurements are between 100 and 200 mL. 2. The frequency variations are continuously recorded using a quartz crystal analyser, QCMagic from Elbatech (Marciana, LI, Italy) (Fig. 1C). A detailed diagram with all the measurements components is illustrated in Fig. 2. QCMagic is a measuring instrument for the recovery of the oscillation frequency of a working quartz crystal. The system is interfaced to the driving personal computer (PC) by means of a digital counter PCI board. One single PCI board can drive up to four oscillator units, thus allowing operating with up to four independent quartz crystal. QCMagic makes use of an internal reference crystal, used as a timebase comparator. Depending on the frequency of this internal timebase, each unit can be tuned to work with a specific working crystal, chosen between 1 and 10 MHz. When more units are used, they are cross-connected by means of one single cable carrying both power supply and signals. QCMagic is interfaced with a PC for data acquisition and storage. The system is driven by a software running under MS-Windows™. Oscillating frequency vs. time measurements can be performed following the data acquisition in real time on the computer screen, by means of strip chart plotting. Examples are given in Figs. 3 and 4. The analytical data are the differences between two stable frequency values (±0.5 Hz).
Piezoelectric Biosensors for Aptamer-Protein Interaction
29
Fig. 2. The components of the measuring system. 9475800 Stabilisation of the signal with water
9475600
Frequency (Hz)
9475400
Activation
9475200
Streptavidin
9475000 9474800
Buffer (A) Ethanolamine
9474600
Buffer (B)
9474400
Aptamer
9474200 9474000 0
500
1000
1500
2000
2500
Time
Fig. 3. Typical frequency variations (vs. time) recorded during the functionalization of the crystal and the immobilization of the aptamer. The entity of the immobilization can be evaluated by the difference between the frequency shift of the two steps Buffer (B) – Buffer (A).
3. Methods 3.1. Immobilization of the Aptamer onto Piezoelectric Crystals
1. The immobilization chemistry adopted follows the approach described in Tombelli et al. (43). In particular, before the immobilization of the aptamer, the electrode surface of the quartz crystal needs to be cleaned with a boiling solution consisting of H2O2 (33%), NH3 (33%), and milliQ water in a 1:1:5 ratio. The crystals are immersed in the solution for 10 min. They are then thoroughly washed with distilled water and used immediately afterwards.
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Fig. 4. Typical frequency variations (vs. time) recorded during the binding of thrombin (50 nM) to the immobilized aptamer. The reported analytical datum will be the difference in frequency between Buffer (B) and Buffer (A), which represents the baseline.
2. The freshly cleaned crystal is immersed in an unstirred 1 mM ethanolic solution of 11-mercaptoundecanol at room temperature, in the dark, for 48 h. The solution of 11-mercaptoundecanol is freshly prepared before use (2 mg of the thiol in 10 mL of ethanol). The crystal is then washed with ethanol and milliQ water and sonicated for 10 min in ethanol to remove the excess of thiol. The hydroxylic surface is treated with a 600 mM solution of epichlorohydrin in a 1:1 mixture of 400 mM NaOH and bis-2-methoxyethyl ether (diglyme) for 4 h. After washing with water and ethanol, the crystal is immersed for 20 h in a basic dextran solution (3 g of dextran in 10 mL of NaOH 100 mM). The surface is further functionalized with a carboxymethyl group using bromoacetic acid (1 M solution in 2 M NaOH for 16 h). All the reactions are performed at room temperature. The coated crystals can be stored at 4°C immersed in milliQ water for 15 days. For their use, the crystals are washed with water and placed in the cell. 3. For further functionalization, the surface of the crystal is activated prior to covalent coupling with 200 mL of a solution of NHS 50 mM and EDAC 200 mM in water (see Note 2). After 5 min, the activating solution is replaced by streptavidin 200 mg/mL in acetate buffer 10 mM, pH 5 for 20 min. The residual reacting sites are blocked with 200 mL of a solution
Piezoelectric Biosensors for Aptamer-Protein Interaction
31
of ethanolamine hydrochloride (pH 8.6, 1 M water solution). After washing with the immobilization buffer, the biotinylated aptamer is added (200 mL of a solution 1.0 mM of the probe in immobilization buffer). The immobilization is allowed to proceed for 20 min (Fig. 3). 4. The biotinylated aptamer (0.5 mM, (see Note 3) ) is thermally treated before its immobilization. The thermal treatment can unfold the aptamer strand making the biotin label at the 5¢ end available for the interaction with streptavidin on the chip surface. Before the immobilization, the biotinylated aptamer is heated at 90°C for 1 min to unfold the DNA strand and then cooled in ice for 10 min to block the DNA in its unfolded structure (44) (see Note 4). 5. The cell with the immobilized aptamer is stored at 4°C with 200 mL of binding buffer. 3.2. Binding Measurements
1. After the immobilization of the aptamer, the frequency is stabilized by keeping the crystal in contact with 100 mL of binding buffer. Thrombin (100 mL) is then added to the cell and left in contact with the immobilized aptamer for 30 min. The surface is then washed with buffer to remove the unbound protein (Fig. 4). 2. The analytical data, expressed as frequency shift, are the differences in the frequency of the crystal before the addition of thrombin and after the washing with buffer subsequent to the affinity interaction. A signal generated by the aptamer– protein interaction is considered significative when the difference between the frequency values corresponding to the two buffers is higher than 3 Hz. Different concentrations of the protein can be used to build a calibration plot. An example of calibration plot is shown in Fig. 5. 3. After each cycle of binding, the crystal surface can be regenerated by 1 min treatment with 2 M NaCl. With this treatment, the sensor-bound analyte is released at increased ionic strength that unfolds the three-dimensional structure of the aptamer without damaging the oligomer structure, and the baseline is reached again, allowing the multiuse of the sensor. The regeneration is considered successful when the frequency value of buffer (B) (Fig. 4) has reached again the frequency value corresponding to buffer (A) (Fig. 4). Generally, on the same crystal surface, 20 measurements can be performed without loss in sensitivity. 4. Negative controls can be tested to prove the specificity of the interaction. Human serum albumin is present in plasma and serum at high concentration (~50,000 mg/L), and it must be tested to prove the absence of nonspecific adsorption due to this high concentration in real matrices (see Note 5).
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Fig. 5. Typical calibration plot obtained with different concentrations of thrombin (0–200 nM) interacting with the immobilized aptamer. Thrombin is diluted in binding buffer; interaction time 30 min.
5. Standard solutions of thrombin can be added to serum or plasma to test the performance of the biosensor in complex matrices. Since serum is not containing coagulation factors, the addition of thrombin is not affecting the samples (see Note 6). On the contrary, the addition of thrombin to plasma, which contains all the proteins involved in the coagulation cascade including fibrinogen, leads to the formation of fibrin and to the rapid sample clotting. To avoid this phenomenon, fibrinogen has to be precipitated from plasma before the addition of thrombin in the preparation of spiked samples (see Note 7). After precipitation of fibrinogen thrombin can be added to plasma (see Note 8).
4. Notes 1. The procedure to fix the aptamer to a solid surface is of paramount importance to obtain an ordered and oriented layer able to assure, as much as possible, the flexibility of the bioreceptor without altering the affinity for the target molecule (45). For this reason, both the influence of a spacer in the aptamer binding behavior and of the immobilization protocol have to be studied. The spacer can consist in a polyT(20) added to the binding sequence of the aptamer. When examining the performances obtained with this aptamer on the
Piezoelectric Biosensors for Aptamer-Protein Interaction
33
piezoelectric biosensor for thrombin, it showed a good reproducibility (average CV% for the concentrations of thrombin, 100 and 200 nM: 13%) and a very good selectivity as demonstrated by the low signal (DF < 3 Hz) obtained with a high concentration of human serum albumin (77 mM HSA), used as negative control. The higher sensitivity obtained with this aptamer is probably due to the presence of the spacer that maintains the aptamer far from the sensor surface allowing the proper conformation for molecular recognition. When the biotinylated aptamer without the polyT tail was immobilized, very low sensitivity (DF for thrombin 200 nM < 20 Hz) was found, even if comparable surface density occurred. 2. The NHS and EDAC solution must be prepared immediately before use to avoid loss of activity. 3. By changing the concentration of aptamer for its immobilization, the surface capacity is not affected by the dilution from 1 to 0.5 mM of the aptamer. On the contrary, a further dilution to 0.1 mM dramatically reduced the biosensor sensitivity. 4. Even if the thermal treatment has no effect on the aptamer surface density (2.5 × 1013 molecules/cm2 with or without the treatment), the linearity in the thrombin range 0–200 nM (R2 = 0.977) and the reproducibility of the binding step (CV% = 21%) significantly improved. This result confirms that the thermal treatment ensures correct intramolecular folding. 5. To check the specificity of the sensor HSA at high concentration (77 mM), in a 1,400-fold excess respect to thrombin, is used. The interaction did not result in a measurable frequency decrease (DF < 3 Hz), demonstrating the high specificity of the sensor. 6. Thrombin was detected in serum diluted 1:100 spiked with thrombin in a concentration range 0–200 nM, with a blank value of −32 Hz. The recorded signals increased with the concentration of added thrombin demonstrating that the aptasensor was able to operate in this complex matrix. However, since serum is not containing the proteins involved in coagulation, further experiments have to be carried out with plasma, which is the matrix where normally thrombin is detected. 7. The selective precipitation of fibrinogen is based on the use of ammonium sulphate as precipitant: 250 mL of plasma are treated with 1,250 mL of 2 M ammonium sulphate and 1,000 mL of 0.1 M sodium chloride. The solution is mixed for 3–4 min, then centrifuged, and the supernatant is eluted in a NAP column for a rapid desalting and buffer exchange. The protein amount of the raw plasma and of the eluted solution was evaluated by spectrophotometric measurements at l = 280 nm to estimate the loss of protein content after precipitation of fibrinogen.
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Fig. 6. Typical calibration plot obtained with different concentrations of thrombin (0–200 nM) interacting with the immobilized aptamer. Thrombin is added to plasma (1:10 in binding buffer); interaction time 30 min.
8. Thrombin is added to plasma diluted 1:100 in a concentration range 0–200 nM. The response is linear in the tested concentration range (R2 = 0.990) and reproducibility was good (CV% = 13%). The matrix effect with a blank signal of −21 Hz is present but, despite the high complexity of the matrix, the increase in thrombin concentration could be detected (Fig. 6). References 1. Tombelli, S., Minunni, M., Mascini, M. (2005). Analytical applications of aptamers. Biosens. Bioelectron. 20, 2424–2434 2. Tuerk, C., Gold, L. (1990). Systematic evolution of ligands by exponential enrichment. Science 249, 505–510 3. Ellington, A.D., Szostak, J.W. (1990). In vitro selection of RNA molecules that bind specific ligands. Nature 346, 818–822 4. Luzi, E., Minunni, M., Tombelli, S., Mascini, M. (2003). New trends in affinity sensing: aptamers for ligand binding. TrAC. Trends Analyt. Chem. 22, 810–818 5. O’Sullivan, C.K. (2002). Aptasensors-the future of biosensing? Anal. Bioanal. Chem. 372, 44–48 6. Ellington, A.D., Szostak, J.W. (1992). Selection in vitro of single-stranded DNA molecules that fold into specific ligand-binding structures. Nature 355, 850–852 7. Geiger, A., Burgstaller, P., Von der Eltz, H., Roeder, A., Famulok, M. (1996). RNA aptamers that bind L-ariginine with sub-micromolar
8.
9.
10.
11.
12.
13.
dissociation constants and high enantioselectivity. Nucleic Acids Res. 24, 1029–1036 Tereshko, V., Skripkin, E., Patel, D.J. (2003). Encapsulating streptomycin within a small 40-mer RNA. Chem. Biol. 10, 175–187 Baskerville, S., Zapp, M., Ellington, A.D. (1999). Anti-Rex aptamers as mimics of the Rex-binding element. J. Virol. 73, 4962–4971 Wen, J.D., Gray, C.W., Gray, D.M. (2001). SELEX selection of high-affinity oligonucleotides for bacteriophage Ff gene 5 protein. Biochemistry 40, 9300–9310 Wilson, C., Nix, J., Szostak, J.W. (1998). Functional requirements for specific ligand recognition by a biotin-binding RNA pseudoknot. Biochemistry 37, 14410–14419 Herr, J.K., Smith, J.E., Medley, C.D., Shangguan, D., Tan, W. (2006). Aptamer-conjugated nanoparticles for selective collection and detection of cancer cells. Anal. Chem. 78, 2918–2924 Homann, M., Göringer, H.U. (1999). Combinatorial selection of high affinity RNA ligands
Piezoelectric Biosensors for Aptamer-Protein Interaction
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
to live African trypanosomes. Nucleic Acids Res. 27, 2006–2014 Kotia, R.B., Li, L., McGown, L.B. (2000). Separation of nontarget compounds by DNA aptamers. Anal. Chem. 72, 827–831 Cole, J.R., Dick, L.W., Jr., Morgan, E.J., McGown, L.B. (2007). Affinity capture and detection of immunoglobulin E in human serum using an aptamer-modified surface in matrixassisted laser desorption/ionization mass spectrometry. Anal. Chem. 79, 273–279 Hamaguchi, N., Ellington, A., Stanton, M. (2001). Aptamer beacons for the direct detection of proteins. Anal. Biochem. 294, 126– 131 Li, J.J., Fang, X., Tan, W. (2002). Molecular aptamer beacons for real-time protein recognition. Biochem. Biophys. Res. Commun. 292, 31–40 Macaya, P., Schultze, F.W., Smith, J.A., Roe, F.J. (1993). Thrombin-binding DNA aptamer forms a unimolecular quadruplex structure in solution. Proc. Natl. Acad. Sci. U.S.A 90, 3745–3749 Smirnov, I., Shafer, R.H. (2000). Effect of loop sequence and size on DNA aptamer stability. Biochemistry 39, 1462–1468 Paborsky, L.R., McCurdy, S.N., Griffin, L.C., Toole, J.J., Leung, L.L. (1993). The single-stranded DNA aptamer-binding site of human thrombin. J. Biol. Chem. 268, 20808–20811 Baldrich, E., Restrepo, A., O’Sullivan, C.K. (2004). Aptasensor development: elucidation of critical parameters for optimal aptamer performance. Anal. Chem. 76, 7053–7063 Radi, A.E., Acero Sanchez, J.L., Baldrich, E., O’Sullivan, C.K. (2006). Reagentless, reusable, ultrasensitive electrochemical molecular beacon aptasensor. J. Am. Chem. Soc. 128, 117–124 Gronewold, T.M.A., Glass, S., Quandt, E., Famulok, M. (2005). Monitoring complex formation in the blood coagulation cascade using aptamer-coated SAW sensors. Biosens. Bioelectron. 20, 2044–2052 Mir, M., Vreeke, M., Katakis, I. (2006). Different strategies to develop an electrochemical thrombin aptasensor. Electrochem. Commun. 8, 505–511 Zhang, H., Wang, Z., Li, X.F., Le, X.C. (2006). Ultrasensitive detection of proteins by amplification of affinity aptamers. Angew. Chem. Int. Ed. 45, 1576 –1580 Centi, S., Tombelli, S., Minunni, M., Mascini, M. (2007). Aptamer-based detection of plasma proteins by an electrochemical assay
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28.
29.
30.
31. 32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
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coupled to magnetic beads. Anal. Chem. 79, 1466–1473 Bini, A., Minunni, M., Tombelli, S., Centi, S., Mascini, M. (2007). Analytical performances of aptamer-based sensing for thrombin detection. Anal. Chem. 79, 3016–3019 Holland, C.A., Henry, A.T., Whinna, H.C., Church, F.C. (2000). Effect of oligodeoxynucleotide thrombin aptamer on thrombin inhibition by heparin cofactor II and antithrombin. FEBS Lett. 484, 87–91 Stubbs, M.T., Bode, W. (1993). A player of many parts: the spotlight falls on thrombin’s structure. Thrombosis Res. 69, 1–58 Shuman, M.A., Majerus, P.W. (1976). The measurement of thrombin in clotting blood by radioimmunoassay. J. Clin. Invest. 58, 1249–1258 Curie, J., Curie, P. (1880). An oscillating quartz crystal mass detector. Rendu 91, 294–297 Janshoff, A., Steinem, C. (2001). Quartz crystal microbalance for bioanalytical applications. Sensor Update 9, 313–354 Bruckenstein, S., Shay, M. (1985). Experimental aspects of the use of quartz crystal microbalance solution. Electrochim. Acta 30, 1295–1300 Sauerbrey, G. (1959). The use of quartz oscillators for weighing thin layers and for microweighing. Z. Physik. 155, 206–222 Hillier, A.C., Ward, M.D. (1992). Scanning electrochemical mass sensitivity mapping of the quartz crystal. Anal. Chem. 64, 2539–2554 Chang, S., Muramatsu, H., Nakamura, C., Miyake, J. (2000). The principle and application of piezoelectric crystal sensors. Mater. Sci. Eng. C 12, 111–123 O’Sullivan, C.K., Guilbault, G.G. (1999). Commercial quartz crystal. Microbalances. Biosens. Bioelectron. 14, 663–670 Kim, N., Park, I.S., Kim, D.K. (2004). Characteristics of a label-free piezoelectric immunosensor detecting Pseudomonas aeruginosa. Sens. Actuators B Chem. 100, 432–438 Mannelli, I., Minunni, M., Tombelli, S., Mascini, M. (2003). Bulk acoustic wave (BAW) affinity biosensor for genetically modified organisms (GMOs) detection. IEEE Sens. J. 3, 369–375 Skládal, P., dos Santos Riccardi, C., Yamanaka, H., Inácio da Costa, P. (2004). Piezoelectric biosensors for real-time monitoring of hybridization and detection of hepatitis C virus. J. Virol. Methods 117, 145–151 Dell’Atti, D., Tombelli, S., Minunni, M., Mascini, M. (2006). Detection of clinically relevant point mutations by a novel piezoelectric
Chapter 3 Piezoelectric Quartz Crystal Resonators Applied for Immunosensing and Affinity Interaction Studies Petr Skládal Summary Piezoelectric quartz crystals serve as resonator-based transducers for direct and real-time monitoring of immunoaffinity interactions. The measuring system is briefly characterized; several examples for immobilization of antibodies are recommended. The piezoelectric immunoassays employing direct, competitive, and displacement-based formats are demonstrated on examples. The method for kinetic characterization of immunoaffinity interactions is presented. Key words: Immunosensor, Biosensor, Flow through immunoassay, Biointeraction analysis.
1. Introduction The direct methods for real-time monitoring of immunochemical interactions employ much simplified assay formats compared with traditional approaches based on labels as enzymes, fluorophores, and radioactivity. The continuous evaluation of the progress of the interaction provides much more detailed information compared with traditional techniques, which measure the amount of the bound label only at the end of the binding process. In addition, the direct immunosensors can be used repeatedly for many assays thus reducing the running costs. Several physical transducers are capable of measuring surface mass changes resulting from formation of immunocomplexes at the sensitive area. Although mostly advanced optical systems are utilized, the piezoelectric (PZ) and acoustic devices (1) represent similar but significantly less expensive alternative.
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_3
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Several anisotropic crystals exhibit piezoelectric effect – mechanical deformation of such crystals generates oriented dipoles and electric voltage. In the opposite, alternating voltage applied on such crystals excites vibrations. At resonance frequency equal to the natural frequency of vibration, transfer of energy from the electric field to the crystal is most efficient and the energy remains conserved in the oscillating system. The typical piezoelectric quartz crystal is shown in Fig. 1. A thin plate of quartz (AT-cut) is coated with metal electrodes on both sides; gold deposited over a thin chromium layer is used as an inert surface for bioapplications. The plate is usually inserted in a holder for simplified manipulation and connection to electronics. The piezoelectric transducers are being used as chemical sensors since the discovery of the relationship between mass of adsorbed films and the resonant frequency by Sauerbrey (2): Df = -
2 f 02 Dm A rq mq
= -2.26 ´ 106 f 02
Dm . A
(1)
The change Δf of the resonance frequency f0 of piezoelectric crystals is directly proportional to the mass change Δm, the numeric constant applies to calculations using Δf in Hz, f0 in MHz, and Δm in g/cm2. The typical working frequencies are from 5 to 20 MHz, higher values are not used as the quartz plates become too thin. The system functions as a sensitive quartz crystal microbalance (QCM); improved sensitivity of recent devices resulted in the alternative name “nanobalance.” In this way, the amount of molecules bound at the sensitive area of electrodes can be easily quantitatively measured as a decrease of the resonant frequency. Even though according to the recent studies the microbalance response is more complex (3), the resonance frequency serves as a convenient signal reflecting the surface-bound molecules.
metal electrodes on the opposite sides
quartz plate wire holders contacts
Fig. 1. Piezoelectric quartz crystal resonator (left, photo of the optically polished smooth crystal) and schematic description (right). The crystal shown (part no. 151620–10) is produced by International Crystal Manufacturing, see Note 1.
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Modification of the sensing area with biomolecules provides affinity biosensors; in the particular case, when the immobilized ligand A is either antigen (hapten) or antibody, the resulting device functions as an immunosensor specific to the corresponding binding partner B. The immobilization of bioligands can include quite different strategies ranging from simple physical adsorption to covalent binding on activating sublayers and finally oriented attachment of antibodies. The generally applicable immobilization procedures (4) are suitable also for piezosensors, and some verified approaches will be mentioned in Subheadings 1–3. In addition to simple measuring of concentrations, the system is also suitable for kinetic characterization of antibodies – Subheading 7. The piezoelectric immunosensors were successfully used for measuring concentration of many different analytes (5, 6). The working formats include direct, competitive, and displacement assays – Subheadings 4–6. Large analytes (proteins, viruses, and bacteria) can be measured directly after binding to the immobilized antibody. For small molecules as haptens, the competitive measurement is needed to achieve measurable change of frequency; the analyte is mixed with antibody to form immunocomplexes, and the remaining free binding sites of the antibody can subsequently interact with the sensing surface modified with a derivative of the analyte.
2. Materials 2.1. Piezoelectric Crystals and Instruments
1. The piezoelectric crystals with gold electrodes have the basic resonant frequency of 10 MHz. The crystals with higher frequencies (up to 20 MHz) will provide a higher sensitivity (Eq. 1). For suppliers of crystals as well as other parts, see Note 1. 2. Oscillating circuit driving the crystal (Fig. 2A): The simplest construction (7) suitable for operation in liquids is based on the gate oscillator. The integrated form of this oscillator – 74LS320 (produced by Texas Instruments, part no. SN74LS320 (8)) will provide much higher energy to the crystal resulting in improved performance under variable conditions (9). 3. A counter for measuring changes of the resonance frequency can be a common device widely used in electronics. The resolution of frequency should be at least 1 Hz within 1 s interval, the upper limit of frequency should be near 50 MHz. An important feature is the possibility to connect to a personal computer to allow on-line monitoring of the affinity interaction; the easiest way is through the standard serial (RS 232C) and USB ports, the GPIB option will require a special interface
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Skládal C
+5V
C1
C1 = 100 nF L
1 2 3
OUTPUT
TANK1 TANK2
XTAL1
GND1
XTAL2
FFQ
NC
FFD
F
NC 7 8
VCC
16
C = 17 pF
15
L = 15 μH
14 Q
U1 = 74LS320
VCC⬘
F
F⬘
GND2
F⬘
Q = 10 MHz
U1
A Peristaltic pump
Buffer valves Sample
waste Regeneration oscillating circuit
frequency counter
flow cell with PZ crystal
B
computer
detector ML3620
peristaltic pump
tubing
flow cell, PZ crystal fixed inside
sample
C Fig. 2. Measuring set-up for piezoelectric immunosensors. (A) Example of the oscillating circuit for PZ crystals (Q) in solutions based on the integrated oscillator 74LS320. The capacitor C and inductor L depend on the base frequency (f0) of the particular crystal, the resonance condition (2πf0)−2 = LC should be satisfied (8). (B) Scheme of the measuring set-up. Changes of working solutions are realized either manually, using multiposition or switching (shown in the scheme) valves. (C) Real measurement based on a compact detector ML3920 (Multilab, Czech Rep.) Integrating both oscillator and counter and a miniature peristaltic pump (model PP10.1, BVT Technologies, Czech Rep.); manual changes of solutions.
Piezoelectric Quartz Crystal Resonators
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card inside the computer. Suitable general-purpose counters include models UZ2400 (Grundig), 53131A and 53132A (Agilent Technologies) and PM6690 (Fluke). 4. Suitable software for continuous reading, display, and storage of frequency: A simple program written in BASIC will be sufficient for this purpose; communication protocol will depend on the particular frequency counter. Evaluation and printing of saved data can easily be performed in most common worksheet programs. Alternatively, a specialized software LabTools was developed in Delphi in our laboratory. It communicates with the external detector through a serial port using ASCII text-based protocol; the sampling rate can be selected (0.1 and 1 s). After initialization of measurement, the frequency values are continuously acquired, stored in a datafile and displayed. Marks indicating experimental details can be introduced and various data manipulation functions (graphical editing, digital filters, and simple math operations) are provided, too. 5. Flow-through measuring cell should allow contact of one side of the crystal with working solutions. The crystal is usually sandwiched between two soft rubber o-rings; reasonable force should be applied to prevent damage of thin and brittle crystals. Suitable liquid flow cell models include models 35363 from ICM and FC-550 from Maxtek (see Note 1). Typically, the flow direction is parallel to the sensing surface (thin layer arrangement). The volume of the cell should be kept small (below 10 μL) to allow fast exchange of solutions. 6. Alternatively, the crystal might be attached with the help of glue (silicon-based) either to the bottom or side wall of a suitable vessel equipped with either stirring bar or vertical rotator. Holders for crystals suitable for immersion in solution are also available, though this design is not preferred for immunosensing due to limited diffusion. 7. Flow delivery system will consist of a peristaltic pump providing flow rates around 20–200 μL/min. Models with many rollers (−10, e.g., Minipuls MP-3, Gilson) are preferred as the noise due to pulsation will be reduced. The indicated flow rates apply for common crystals with the diameter of the working electrode around 5 mm. In fact, on the one hand, slower flow will limit consumption of valuable bioreagents; on the other hand, faster flow rate will be more appropriate for immunoaffinity studies to avoid influence of slow diffusion on kinetics. In addition, electronically controlled switching and/or multiposition valves or even more complex autoinjectors will enable automatization of performance and provide option for unattended operation. 8. Flexible plastic tubes (internal diameter 0.1–0.5 mm), materials as silicone, polyethylene, Teflon (available from common
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laboratory suppliers) are needed to connect the components of the flow system (Fig. 2b, c). 2.2. Chemicals
Most reagents are available from the Sigma – Aldrich company (cat. numbers indicated for less common compounds). Deionized water should be used for preparation of solutions. 1. Phosphate buffer, 50 mM concentration, is prepared by adding 7.8 g of NaH2PO4. 2H2O to 1 L of water and adjusting pH to 7 using solution of NaOH. 2. p-Aminopropyltriethoxysilane (APTS, A3648) dissolved in acetone at 5% vol. 3. Cysteamine (M6500) dissolved in water (10 mg/mL), prepare freshly before use. 4. 3,3′-Dithiodipropionic acid bis(N-hydroxysuccinimide ester) (DTSP, D3669) should be dissolved in dimethylsulfoxide (10 mg/mL), prepare freshly before use. 5. Glutaraldehyde (G5882), 3% solution in the phosphate buffer, can be stored frozen at −20°C. 6. Protein A (P6031) dissolved in water (1 mg/mL) can be stored frozen at −20°C. 7. Dimethylpimelimidate (D8388) dissolved in water (5 mg/mL), prepare freshly.
2.3. Working Buffers
1. Carrier solution is phosphate buffered saline (PBS), prepare by adding 7.6 g of NaCl per 1 L of the phosphate buffer. Optionally, it might be useful to add detergents (e.g., Tween-20, P1379, 0.01%) and inert protein (e.g., 250 μg/mL bovine serum albumin, 05477) to suppress nonspecific binding of proteins and other molecules from samples. 2. Regeneration solutions are rather variable; the following options are suitable for initial experiments: 200 mM glycine-HCl pH 1.5–3; 10–100 mM hydrochloric acid; 10 mM formic acid; 10–100 mM sodium hydroxide; 10–30% methanol. For small immobilized ligands, proteases are useful to cleave the associated complementary proteins; either proteinase K (P2308, 50 μg/mL in 10 mM Tris–HCl buffer pH 7.8) or pepsin (P6887, 200 μg/ mL) at pH 2 (10 mM HCl) were found useful.
3. Methods 3.1. Immobilization of Proteins on Silanized Crystals
Silanization is a widely used approach for activation of inorganic surfaces as glass and metals including gold electrodes of the PZ resonators. The deposited silane layer serves for further covalent attachment of biomolecules:
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1. Clean the PZ resonator with acetone and allow it to dry in air. 2. Incubate the cleaned resonators in acetone containing 5% APTS for 1.5 h at room temperature. 3. Take the crystal out from the APTS solution and heat in an oven at 120°C for 1 h. 4. Wash crystals carefully with acetone to remove loosely bound APTS. 5. Activate thus obtained aminogroups from APTS by incubation in 3% glutaraldehyde for 1.5 h, and then wash carefully with phosphate buffer and water. 6. Biomolecules containing free surface aminogroups (most proteins and hapten-protein coating conjugates) can be covalently immobilized by placing drops (20 μL per one electrode) of biomolecules (10–500 μg/mL final concentrations) in phosphate buffer. Incubate overnight inside a refrigerator. 7. The washed and dried modified resonators can be stored in dry state in a refrigerator for months. 3.2. Immobilization of Proteins to Thiol Monolayers on Gold
Thiocompounds (thiols and disulfides) irreversibly adsorb on gold surfaces forming self-assembled monolayers (SAM), which are convenient for further immobilization steps. 1. Clean the crystal with acetone and allow it to dry in air. 2. Incubate cleaned crystals for 2 h in the presence of thiol: either cysteamine (10 mg/mL in water) or DTSP (10 mg/mL in dimethylsulfoxide) can be used. Wash modified crystals with water. 3. Cysteamine-modified crystals should be further activated with glutaraldehyde as described above (see Subheading 1, steps 5–7), the resulting covalent attachment of protein is depicted in Fig. 3. 4. DTSP-modified crystals can be used to immobilize directly proteins through surface aminogroups by simple incubation for 2 h at room temperature. The concentration of proteins can be from 10 to 1,000 μg/mL.
3.3. Oriented Immobilization of Antibodies
1. Incubate DTSP-modified crystals (Subheading 2, step 4) with Protein A (1 mg/mL in water) for 2 h at room temperature, 20 μL per one side of the crystal. Then wash crystals with phosphate buffer and water.
Fig. 3. Immobilization of antibody on the PZ resonator. Gold modified with cysteamine SAM was activated with glutaraldehyde, protein A was covalently attached and captured Ab.
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2. Add solution of antibody (IgG, 10–500 μg/mL dissolved in phosphate buffer), 20 μL per one side of the crystal, and incubate 2 h at room temperature. Wash carefully with phosphate buffer and water. 3. Crosslink the Protein A–antibody complex formed at the crystal by incubation in dimethylpimelimidate (5 mg/mL in water) for 2 h at room temperature (see Note 2). Wash the crystals with water, allow to dry, and store in a refrigerator. 3.4. Direct Immunoassay of Biomolecules
The initial recording of baseline frequency (step 1) and final regeneration of the sensing surface (step 6) is common to several measuring procedures (Subheadings 4–7). The real trace of such analysis is shown in Fig. 4 (10). 1. The carrier solution is allowed to pass through the flow cell containing the bioligand-modified crystal fixed inside. Wait for a stable background frequency fbckg. (see Note 3). 2. The sample is added to the carrier buffer and allowed to flow for 5–10 min; longer incubation times may improve sensitivity of detection. 3. A new stable frequency f1 is obtained in the presence of carrier buffer only. 4. The absolute amount of the bound analyte can be obtained from Eq. 1 using Δf = fbckg − f1.
1
2
3
4
5
50 Hz 10 min
sample
regeneration buffer
buffer
Fig. 4. Examples of experimental traces (frequency change vs. time plot) for five sequential measurements of serum samples. The marked part is zoomed in the inset figure where individual steps of the assay cycle are shown (2 min baseline, 10 min interaction with sample, 5 min buffer zone, 4 min regeneration, and 2 min baseline). PZ sensor with immobilized anti osteoprotegerin Ab, osteoprotegerin in serum was analyzed, for procedure see (10).
Piezoelectric Quartz Crystal Resonators
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5. Alternatively, calibration curve can be constructed using standard solutions of the analyte as the dependence of Δf on concentration (see Note 4). 6. Remove any remaining immunocomplexes from the sensing surface using regenerating solutions (Subheading 2.3 item 2), and the contact time should be 1–5 min (see Note 5). 7. The next analysis can be started using the same crystal. 3.5. Competitive Immunoassay of Haptens
1. Preincubate the sample with a fixed concentration of antibody (10–100 μg/mL) for 5–15 min. 2. Pass this mixture through the flow cell containing the crystal modified with immobilized analyte (or its derivative that is recognized by the antibody). 3. The measured change of frequency will be decreasing with the increasing concentration of analyte (Fig. 5A).
3.6. Displacement Assay
1. The piezoelectric crystal modified with immobilized analyte should be saturated with the corresponding antibody. 2. The flow of sample containing the free analyte releases antibody from surface immunocomplexes and gradual increase of frequency is observed (Fig. 5B). 3. The rate of frequency increase is directly proportional to the concentration of analyte in the sample (see Note 6).
3.7. Kinetic Characterization of Antibodies
Formation of the immunocomplex AB at the surface of the piezoelectric crystal is characterized by the kinetic association ka and dissociation kd rate constants: ½¾A + B
ka kd
½¾ AB.
A
MAb + atrazine
B Peptide LQSPQQSFS
50 μg/ml 30 20
250 Hz
atrazine (µg/L)
50 Hz
10
1 0
5 min
0
0.1
Buffer
5 min
Fig. 5. Example of competitive and displacement assays. (A) Competitive assay of atrazine, binding curves obtained for mixtures of MAb (ascites fluid, 1,000× diluted) preincubated for 15 min with variable concentrations of atrazine. (B) Displacement of antibody from the sensor in the presence of increasing concentrations of the analyte – peptide representing a surface epitope of the antigen. The antigen-modified sensor was preincubated with antibody (not shown), thus a constant amount of immunocomplexes (fa ∼ 350 Hz) was present before starting each of the measurements.
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Skládal
The rate of formation of the complex d[AB]/dt can be expressed using the measured frequency f and concentration c for the free partner B: d[AB] / dt = -df / dt = ka ( f max - f )c - kd f .
(2)
The binding curves (f vs. t dependencies) were usually transformed to obtain df/dt vs. f plots that subsequently provide kinetic constants from Eq. 2 using linear regression (11). A more elegant and precise method is integration of Eq. 2 and then introduction of substitutions feq and kobs (12). The dependence of the resonance frequency f on time t can be fitted to the kinetic equation similarly as described for the optical biosensor system (13): k cf f = a max {1 - exp[-(kac + kd )]} = f eq éë1 - exp (-kobst )ùû , kac + kd
(3)
where c is concentration of the free partner B and fmax represents the capacity of the crystal (maximum change of frequency obtained after saturation of all binding sites). In this way, the binding curves can be used directly for calculation of parameters using nonlinear regression. Plot of kobs against concentration c provides values of the rate constants. A typical affinity experiment with the piezoelectric immunosensor is schematically shown in Fig. 6.
1
2
3
4
5
f0
association Frequency
dissociation Time
buffer
fa sample
feq
buffer
Fig. 6. Characterization of kinetic properties of antibodies. First, the background signal is recorded using the PZ sensor with immobilized antibody (1) in the carrier buffer only. The association reaction follows in the presence of the sample containing antigen – formation of immunocomplexes at the sensing surface (2), resulting eventually in equilibrium (3). Next, a spontaneous dissociation of the immunocomplexes (4) is observed in the absence of the sample and eventually all immunocomplexes become dissociated (5).
Piezoelectric Quartz Crystal Resonators
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First, a stable background signal f0 is obtained in the presence of buffer. Then, sample solution containing antigen is injected (association phase). The immobilized antibodies with free binding sites interact with the free antigen (of course, the situation might be inversed). A decrease of frequency is observed as surface mass on the crystal increases (the equilibrium change feq is achieved eventually). Then, buffer is injected again and the dissociation of immunocomplexes is observed. From this phase, the dissociation constant kd can be obtained independently, and fa represents the amount of surface-bound immunocomplex at the beginning of the dissociation: (4)
f = f a exp(-kd t ). Experimental procedure:
1. Prepare samples containing at least five different concentrations of the studied antigen. 2. For each of the concentrations, record first the association curve (trace of frequency obtained during flow of the sample through the cell containing a crystal modified with the corresponding binding partner) and then the dissociation curve (trace of frequency obtained when the sample is exchanged for the carrier solution), sample curves shown at Fig. 7A. 3. For each individual curve, calculate kobs and feq using the nonlinear curve fitting of the frequency vs. time plot to Eq. 3 (see Note 6). 4. Plot the values of kobs against corresponding concentrations of antibodies and obtain the kinetic rate constants using linear
buffer
5 10
sample
B
0.0008
50
0.0006 kobs
15
(s−1)
25
0.0004
25
MAb (mg / ml)
500 Hz
15
0.0002
10
A 10 min
50
5 0.0000
0
100
200 [Ab] (nM)
300
Fig. 7. (A) Experimental curves from the piezosensor analysis of antibody binding at the sensing surface with immobilized antigen – secalins. The up arrow indicates beginning of the flow of antibodies (association phase, concentrations of MAb are shown close to the curves), down arrows mark beginning of the buffer flow only (dissociation phase). (B) Plot of the kobs constant obtained by nonlinear fitting of the association parts of the binding curves against the corresponding concentrations of the antibody (Mr of antibody was assumed to be 160 kDa).
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regression (Fig. 7B); ka is equal to the slope and kd to the intercept (see Note 7). 5. The value of kd can be obtained independently from the dissociation traces by nonlinear fitting to Eq. 4. The technique described above was used to characterize several monoclonal (14) as well as recombinant (15) antibodies. In the case of polyclonal antibodies, exact kinetic parameters are not obtained (Note 8); nevertheless, the quantitative comparison of different products is possible. Moreover, fast determination of affinities is useful during screening and development of antibodies. The kinetic parameters enable to select the most suitable antibody for the planned purpose when several monoclonals are available (Note 9).
4. Notes 1. Piezoelectric crystals, oscillators, and other instrumentation for microbalance might be obtained from the following companies: International Manufacturing Company, Oklahoma City, OK (ICM, http://www.icmfg.com); Maxtek, Cypress, CA (http://www.maxtekinc.com/); CH Instruments, Austin, TX (http://chinstruments.com/chi400.html); Elchema, Potsdam, NY (http://www.elchema.com/); Seiko/EG&G, Japan; Attana, Stockholm, Sweden (http://www.attana. com/); Q-Sense, Vastra, Sweden (http://www.q-sense.com); QCM Lab, Jarfalla, Sweden (http://www.qcmlab.com/). Comparison of other piezoelectric devices was published in the literature (16). 2. The covalent crosslinking of the immobilized antibody to Protein A stabilizes the complex so that the resulting immunosensor is more stable during regeneration steps. 3. The noise level of the piezoelectric detector itself is usually negligible and most noise comes from the PZ resonator exposed to liquid environment. During stabilization of the baseline signal, variations of the mean frequency value below 0.1 Hz per minute are acceptable. Fluctuations of frequency in the course of measurements might be up to ±2 Hz, these are caused by pulsations of the peristaltic pump, temperature drifts, and mechanical and electrical disturbances. Therefore, proper thermostating and shielding significantly improves stability of signal. Thus, to achieve the signal/noise level of 3 under common conditions (measurement realized on laboratory table), the response during the interaction should exceed 5 Hz. 4. The direct assay procedure is generally suitable only for analytes with molecular weight above 20 kDa, otherwise the
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measured changes of frequency will become too small. The differential measurement ensures that different compositions of samples will not affect the measured signal. 5. Type of the regeneration buffer and incubation interval should be optimized for the particular pair of interacting biomolecules. At the beginning, it is necessary to verify that regeneration is complete and no loss of the immobilized ligand occurs. 6. The displacement assay can potentially be useful for continuous monitoring purposes; sudden increase of frequency indicates the presence of analyte in the sample. 7. For curve fitting, several mathematical programs providing the nonlinear regression procedure may be used, e.g., Origin (Microcal) and SigmaPlot (Jandell Scientific). The interaction data are imported in a simple ASCII format file consisting of a one column of frequency values with equal time spacing (mostly 1 s is sufficient for immunointeractions), the appropriate kinetic model containing desired parameters is defined and the interval of data for fitting is marked. The software then iterates the values of parameters to achieve the minimum sum of squared differences between experimental and calculated values of signal. However, critical inspection of the final parameters is always necessary. 8. For calculation of kinetic constants, the concentration of the specific IgG should be used; otherwise the calculated value of the association rate constant ka will be underestimated. 9. The calculated values of constants should always be critically interpreted as these need not agree exactly with the corresponding values obtained from experiments with both affinity partners free in solution. The immobilization of haptens (less often also antigens) can affect the conformation. In addition, the density of ligands and mass transport effects should be considered, too. The ligand can be immobilized either as a single monolayer or inside a three-dimensional dextran matrix. The latter approach provides higher levels of signal; in contrast, dissociation can be disturbed by “rebinding” effects. References 1. Grate, J. W., Martin, S. J., and White, R. M. (1993) Acoustic wave microsensors. Anal. Chem. 65, A940–A948; A987–A996 2. Sauerbrey, G. (1959) The use of oscillators for weighing thin layers and for microweighing. Z. Phys. 155, 206–210 3. Lucklum, R. and Hauptmann, P. (2006) Acoustic microsensors-the challenge behind
microgravimetry. Anal. Bioanal. Chem. 384, 667–682 4. Hermanson, G. T., Mallia, A. K., and Smith, P. K. (1992) Immobilized affinity ligand techniques. Academic Press, San Diego 5. Suleiman, A. A. and Guilbault, G. G. (1994) Review: Recent developments in piezoelectric immunosensors. Analyst 119, 2279–2282
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6. Bunde, R. L., Jarvi, E. J., and Rosentreter, J. J. (1998) Piezoelectric quartz crystal biosensors [review]. Talanta, 46, 1223–1236 7. Nomura, T., Watanabe, M., and West, T. M. (1985). Behaviour of piezoelectric quartz crystals in solutions with application to the determination of iodide. Anal. Chim. Acta 175, 107–112 8. Texas Instruments (1981) SN74LS320 crystalcontrolled oscillators, D2418, 3–801 9. Skládal, P. and Horáček, J. (1999) Kinetic studies of affinity interactions: comparison of piezoelectric and resonant mirror-based biosensors. Anal. Lett. 32, 1519–1529 10. Skládal, P., Jílková, Z., Svoboda, I., and Kolář, V. (2006) Investigation of osteoprotegerin interactions with ligands and antibodies using piezoelectric biosensors. Biosens. Bioelectron. 20, 2027–2034 11. Skládal, P., Minunni, M., Mascini, M., Kolář, V., and Fránek, M. (1994) Characterization of monoclonal antibodies to 2,4-dichlorophenoxyacetic acid using a piezoelectric quartz crystal microbalance in solution. J. Immunol. Meth. 176, 117–125
12. O’Shannessy, D. J. (1994) Determination of kinetic rate and equilibrium binding constants for macromolecular interactions: A critique of the surface plasmon resonance literature. Curr. Opin. Biotechnol. 5, 65–71 13. Morton, T. A., Myszka, D. G., and Chaiken, I. M. (1995) Interpreting complex binding kinetics from optical biosensors: A comparison of analysis by linearization, the integrated rate equation, and numerical integration. Anal. Biochem. 227, 176–185 14. Horáček, J. and Skládal, P. (1997) Improved direct piezoelectric biosensors operating in liquid solution for the competitive label-free immunoassay of 2,4-dichlorophenoxyacetic acid. Anal. Chim. Acta 347, 43–50 15. Horáček, J. and Skládal, P. (1998) Characterization of the interactions between immobilized parathion and the corresponding recombinant scFv antibody using a piezoelectric biosensor. Food Agric. Immunol. 10, 363–374 16. Henry, C. (1996) Measuring the masses: Quartz crystal microbalance (product review). Anal. Chem. 68, 625A–628A
Chapter 4 Biosensors Based on Cantilevers Mar Álvarez, Laura G. Carrascosa, Kiril Zinoviev, Jose A. Plaza, and Laura M. Lechuga Summary Microcantilevers based-biosensors are a new label-free technique that allows the direct detection of biomolecular interactions in a label-less way and with great accuracy by translating the biointeraction into a nanomechanical motion. Low cost and reliable standard silicon technologies are widely used for the fabrication of cantilevers with well-controlled mechanical properties. Over the last years, the number of applications of these sensors has shown a fast growth in diverse fields, such as genomic or proteomic, because of the biosensor flexibility, the low sample consumption, and the non-pretreated samples required. In this chapter, we report a dedicated design and a fabrication process of highly sensitive microcantilever silicon sensors. We will describe as well an application of the device in the environmental field showing the immunodetection of an organic toxic pesticide as an example. The cantilever biofunctionalization process and the subsequent pesticide determination are detected in real time by monitoring the nanometer-scale bending of the microcantilever due to a differential surface stress generated between both surfaces of the device. Key words: Microcantilever, Nanomechanical biosensors, MEMs, Immunoassay, Pesticide detection, Surface stress, Biofunctionalization, Biorecognition.
1. Introduction The working principle of the family of biosensors based on nanomechanical transducers, and specifically on microcantilevers, involves the translation of biochemical reactions into a mechanical movement in the nanometer range. In microcantilever sensors, the biochemical receptor layer is directly in contact with one of the cantilever surface. The biomolecular recognition process between the receptor layer and its corresponding analyte induce
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_4
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quantitative changes in the mechanical properties of the cantilever beam, allowing the detection of such interactions by measuring the cantilever bending or the change in its resonance frequency (1). The potential of this new kind of sensor relies on their tiny area and their ability to detect molecular interactions without labels (reducing the time and cost of sample preparation), and its low-cost fabrication and mass production by using standard semiconductor technology. In addition, their production can be easily scaled up (see Note 1). The cantilevers can be fabricated of any shape and from substantially any material utilized in microelectronics industry, i.e. crystalline or poly-silicon, silicon nitride, silicon oxide, polymer materials (see Note 2). The rectangular shape beams are the most frequently used in biological research. In biological sensors based on the bending method, it is important to have the cantilevers flat and in plane with the base surface. Initial offset or curvature of the beams complicates adjustment of the experimental setup, especially, if working with arrays of cantilever. For this reason, the most common material for cantilevers fabrication nowadays is single crystalline silicon. A large variety of biomolecular interactions have been detected with silicon microcantilevers. Our work is focused on nanomechanical biosensors based on the cantilever bending method (see Note 3), which measures the microcantilever deflection due to differences in the surface stress between the opposite sides of the microcantilever. For that method, just one side of the cantilever is sensitized with a bioreceptor layer. This specific functionalization is carried out by covering one side of the cantilever with a thin layer of gold and then using the well-known self-assembled monolayers (SAMS) chemistry for the covalent immobilization of the bioreceptor over the gold surface. The formation of consecutive biolayers over the cantilever surface can be sensed in real time due to the cantilever deflection shift produced by each layer. The cantilever motion is detected using the laser beam deflection readout technique (commonly used in atomic force microscopy), where the displacement of a laser beam reflected from the cantilever free end is registered in a position sensitive photodetector (PSD), see Note 4. In Fig. 1, a scheme of the cantilever bending method and the laser beam deflection readout is showed. The gold layer initially deposited over the cantilever surface helps to enhance the reflected beam intensity, increasing the signal to noise ratio. The sensitivity of the method depends both on the resolution of the readout electronics and on the cantilever mechanical properties, which are carefully optimized (by simulation) before fabrication to obtain maximum cantilever deflection in response to the expected bioreaction. The cantilevers are fabricated using standard silicon technology in single crystalline silicon as its
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A
Laser beam
Two-quadrants photodetector
Laser beam
(II)
B
Photodetector signal
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Beam displacement
Base signal (I) Bending signal (II)
Time
Fig. 1. (A) Simple scheme of the cantilever bending method and the laser beam displacement readout. The reflected laser beam suffer a displacement respect its original position (reflection over the flat cantilever), due to the adsorption of molecules over only one cantilever side. (B) Photodectector signal read in real time before and after the adsorption of molecules over one cantilever surface.
characteristics match the parameters of the simulated devices with the real ones. During the last years, the application of nanomechanical biosensors has been expanded to many different fields, such as genomic, proteomic, environmental control, food quality control, clinical analysis, etc. Specifically, in the proteomic field, the immunological detection is one of the most employed, for example, for detecting pathogens, discovering protein expression patterns, or isolating proteins from cellular extracts. Using an immunological detection scheme, we present the detection of a toxic chemical compound (a pesticide) as an example of the application of cantilevers biosensors in the environmental control field. There are several techniques suitable for the cantilever response read-out as the optical beam deflection, piezoresistivity, piezoelectricity, interferometry and capacitance, among the most important. However, the majority of the biochemical applications carried out with the cantilever-based sensor are based in the optical beam deflection method, because of its high sensitivity and its simplicity. But for making practical devices, this readout implementation in array platforms is technologically challenging, as it requires an array of laser sources with the same number of elements as the cantilever array. The lasers displacement could be measured by using an array of photodetectors, adding alignment complications, or using just one photodetector and sequentially switching on and off each laser source to avoid the overlapping of the reflected beams. Additional engineering work is still needed in this direction before to deliver devices for real applications.
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2. Materials 2.1. Set-Up Components
1. The cantilevers are fabricated from Silicon-On-Insulator (SOI) 100 mm standard UNIBOND™ wafers purchased from Soitec Company. They bear layers of single crystalline silicon, silicon dioxide (also called buried oxide layer), and a silicon substrate. The back side of the wafer may contain silicon dioxide. The fabrication of the cantilevers is done at Clean Room (class 100) facilities using standard microelectronics optical photolithography, reactive ion etching, wet etching, and various deposition methods. The procedure and materials used for each process are specific for different clean rooms. The dimensions of the fabricated cantilevers are: 200 μm long, 40 μm width, and 0.334 μm thick. Alternatively, the cantilevers could be commercially purchased. In this case, the ones chosen had dimensions of 200 μm length, 40 μm width, and 0.8 μm thick (ORC8-PS-W, from Olympus). 2. Fluid cell is made of glass with a cell volume of 50–100 μL (Digital Instruments, Veeco), where the cantilever is fixed. 3. Peristaltic pump (Gilson) for maintaining a constant flow during all the biochemical detection process. 4. An injection valve (model 5020, Sigma-Aldrich) allows the injection of different solutions without changing the flow speed. 5. Optical detection system composed of a diode laser of 3 mW (l = 630–670 nm), for focusing in the free-end of the cantilever and a four quadrants photodetector (model S4349, Hamamatsu), for collecting the reflected light. 6. Two x–z positioning stages, and a rotatory one, for simplifying the laser-cantilever-photodetector alignment. 7. A digital acquisition card (DAC) connected to a computer (National instruments) for reading the photodetector signal in real time. The signal acquisition is controlled by a Labviewbased application.
2.2. Surface Functionalization Based on Sams and Pesticide Detection
1. Trichloroethylene and ethanol absolute can be purchased from Panreac (no 161749); acetone from Sigma-Aldrich (179124-2L). 2. Piranha Solution contains a mix 70:30 of sulphuric acid 96% (Merk KGaA no 100732) and hydrogen peroxide 30% (Panreac, no 121076). The solution should be prepared 5–10 min before used, adding the sulfuric acid first, and followed by the peroxide. Never store piranha solutions. Piranha stored in a closed container will likely explode. Caution: Piranha solution is very dangerous. It is extremely energetic and may result in explosion or skin burns if not handled with extreme caution.
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3. Mercaptoundecanoic acid (450561-5G), N-hydroxysuccinimide (NHS) (130672-25G), N-(3-dimethylaminopropyl)N-ethylcarbodiimide hydrochloride (EDC) (E1769-1G), and ethanolamine (236381-50G) can be purchased from SigmaAldrich. EDC quickly oxidizes under air atmosphere. A good way of handling EDC is to separate it into aliquots immediately after opening the container for the first time, then store aliquots at −20°C. 4. Phosphate buffer (PBS) can be purchased from Sigma-Aldrich (P7059-1L) or home-made using monosodic and disodic phosphated salts. We recommend buying or preparing a PBS 10× (pH 7–7.5) as stock solution. Then, prepare a working solution of PBS 1× by dilution from the stock solution. When diluted to 1×, working concentration contains 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, and 2 mM KH2PO4, respectively. 5. Hydrochloric acid 0.1 M can be performed by dilution of hydrochloric acid fumant 37% (Merck, no. 1.00317.1000).
3. Methods One of the first steps to develop cantilever-based biosensor of high sensitivity toward biomolecular interactions is to simulate the cantilever response depending on its dimensions. In general, making thinner and longer cantilevers increases the flexibility, but it also amplifies the thermomechanical noise. For that reason, a compromise between the bending amplitude and the resolution needs to be reached. Once the optimum dimensions have been defined, the cantilevers can be fabricated and characterized (2). The fabrication process must produce cantilevers with well-controlled and reproducible mechanical properties, such as the spring constant, as well as initially stress-free. The spring constant can be characterized by exerting a controlled force on the cantilever with an atomic force microscopy cantilever, with a calibrated spring constant. For detecting the cantilever displacement, a home-made set-up, on the basis of the laser deflection technique, is used, which is able to detect the cantilever movement due to Brownian excitation (i.e., the motion induced in the cantilever by the random hits of thermally excited particles of the surrounding medium). The read-out configuration (e.g., laser spot size, photodetector resolution, cantilever-photodetector distance, etc.) affects the final sensitivity of the system. The next step is the application of the microcantilever as biosensor in the environmental control field, by following an immunological detection scheme. For this purpose, we present the inhibition immunoassay performed to detect the highly toxic organochlorine insecticide compound dichlorodiphenyltrichloroethane (DDT).
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Main applications of biosensors are based on antigen–antibody specific interactions. The detection of this specific interaction can be achieved (a) directly, by means of the specific immobilization of the antibody onto the transducer surface and monitoring the interaction with the antigen, or (b) indirectly, by means of an inhibition assay, where the immobilized molecule is the antigen and monitoring the inhibition of the antibody signal when mixtures of a fixed amount of antibody and variable amounts of antigen are introduced on the biosensor. For the DDT detection, an inhibition format was preferred instead of the direct detection due to the small molecular weight of the pesticide, which made the direct detection more difficult. For that reason, a hapten (DDT5) molecule, which consists of a molecule chemically equivalent to the DDT, is synthesized and coupled to a carrier protein (e.g., BSA) to be immobilized over the surface. The cantilever surface functionalization process that will be explained describes the immobilization of a hapten (or an antigen) that displays amine groups all over its surface. The receptor should be a protein, peptidic molecule, or any other molecule displaying free amine groups, or in the case of organic compounds as pesticides, if they are previously coupled to a protein carrier that supports the amine groups for the linkage. Inhibition format gives the advantage that a more stable monolayer is obtained, as proteins display several recognition sites for the antibody and an oriented immobilization is not required. In addition, they are more resistant to denaturalization than antibodies and typically, monolayers tolerate several regeneration cycles. But the main advantage is that inhibition formats have demonstrated better detection levels than direct detection formats for small analytes. Surface functionalization of cantilevers is one of the trickiest issues for the correct performance of the biosensor to reach high detection levels. The bio-receptor (BSA-DDT5) immobilization requires a selective and specific immobilization of molecules rending a densely and ordered monolayer, which must be stable enough under several regeneration cycles and free of nonspecific adsorption. This type of monolayers can be achieved using the well-known method of self assembled monolayers (SAMs). Self assembly takes place by the direct link of a thiol functional group of an alkanethiol molecule with the Au atoms of the sensor surface, rendering in a densely, ordered and very stable monolayer. Proteins are afterwards covalently attached to the surface previously functionalized with this alkanethiol monolayer. One of the main advantages of SAMs is the possibility of varying the alkyl chain length and ending group, generating different interface layers between the gold metal surface and the immobilized element. To perform the inhibition assay, samples containing a fixed concentration of monoclonal antibody (named LIB-DDT) are mixed with DDT known solutions at different concentrations. After a short incubation step, only the antibody that remained free in the mixture may couple to the bioreceptor. The signal observed in the sensor is an inversely measurement of the pesticide concentration in the sample.
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3.1. Cantilever Simulation and Fabrication 3.1.1. Cantilever Simulation
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1. The cantilever deflection, Δz, generated by a surface stress change, Δs, (due to a biorecognition process, for example) depends on the microcantilevers’ dimensions (length L and thickness h) and its material’s properties (Young’s modulus E and Poisson coefficient n) following the Stoney’s equation: Dz =
3L2 (1 - n) Ds . Eh 2
That means that for a produced Δσ, the cantilever bending will be higher for longer and thinner cantilevers, i.e., with low stiffness. The parameter that characterizes the cantilever stiffness is the spring constant, which depends on the cantilever dimensions and Young’s modulus (see Subheading 3.2). The biomolecular interactions could be difficult to detect with commercial microcantilevers due to the low surface stress induced on such microcantilevers. To improve the cantilever deflection, this parameter, and its dimensions dependency, needs to be simulated before fabrication. The simulation was performed using a finite element analysis (FEA) technique (with ANSYS software). 2. One of the most important issues for simulation of microsystems is to know the properties of the materials. In the case of microcantilevers, young modulus, Poisson’s ratio, and density are the main parameters to be considered. If monocrystalline silicon is used, as reported in this work, the mechanical properties are well-known (see Table 1). For an anisotropic material such as silicon, the Young’s modulus, Poisson’s ratio, and shear modulus depend on which crystal direction the material is being stretched. The appropriate values for each direction have to be introduced in the model. 3. Once the material is defined, the generated cantilever deflection will be larger when L/h increases (see Stoney’s equation). For that reason, a thickness of only 0.334 μm was selected. As the limit of sensitivity is determined by the signal to noise
Table 1 Silicon mechanical properties Density
r
2.33 × 103
kg/m3
Young modulus (110)
U110
1.6895 × 1011
Pa
Young modulus (100)
U100
1.3002 × 1011
Pa
Poisson’s ratio (110) (100)
n110 100
0.2785
–
Poisson’s ratio (110) (110)
n110 110
0.0625
–
Shear modulus (110) (100)
G110 100
0.7951 × 1011
Pa
Shear modulus (110) (110)
G110 110
0.5085 × 1011
Pa
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ratio, the length and width proposed was 200 μm and 40 μm, respectively, to avoid increasing the thermomechanical noise. For these chosen dimensions, the calculated deflection produced by a surface stress change of 1 mN/m (with E = 169 GPa and n = 0.2) would be about 5 nm. This value is five times larger than the one obtained for the alternatives commercial cantilevers proposed (ORC8-PS-W Olympus: 200 μ m length, 40 μm with and 0.8 μm thick, with E = 175 GPa and n = 0.25). 4. In spite of this modeling could be more time consuming, and a 3D geometrical model was chosen for the simulation of the microcantilevers, because of its more precise results. 5. The structural materials that conform the cantilevers can have intrinsic stresses and stress gradients because of the fabrication processes. These can change the mechanical performance of the microcantilevers. If stresses and stress gradients are high, they must be previously measured by using specific test structures and their values should be included in the simulations. As monocrystalline silicon will be used in the fabrication of our devices, these stresses can be neglected. 6. The spring constant of the cantilever could be simulated by applying a known force, F, at the free end of the cantilever. The force should be centered at the end of the cantilever to avoid any tilt. The spring constant, k = 0.0077 N/m, is determined by the applied force, F, and the simulated maximum displacement, z = 0.13 μm, by using the formula k = F/z. 3.1.2. Cantilever Fabrication
1. The processing starts with a SOI 4 in. wafer, which is bearing a 334 nm of single crystalline silicon on 1 μm thick silicon dioxide buried layer over the silicon substrate (see Fig. 2A).
Fig. 2. Scheme of the microcantilever fabrication process.
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The substrate is 425 μm thick (see Note 5). The cantilevers were fabricated in separate cavities with a pitch of 250 μm between them. The cavities are the through-wafer holes drilled using a deep reactive ion etching (DRIE) technique. Each cantilever was 40 μm wide and 200 μm long, the thickness of each cantilever was 334 nm (which was fixed by the thickness of the structural layer) (see Note 6). 2. The back side of the wafer is metallized with 1 μm of aluminum deposited by RF sputtering. This layer will be used as a mask material for the DRIE definition of the cavities. 3. The pattern for the cavities is defined on the wafer back side at the aluminum layer by using standard photolithography, dry and wet etching processes. Photoresist was cured for 30 min at 120°C to stand the steps of dry reactive ion etching and wet etching (Fig. 2B). A wet etching in SiOetch solution during 30 s was applied to remove the silicon dioxide from the windows. 4. The cantilevers were defined by photolithography and dry etching. The silicon dioxide buffer layer was kept to be employed as an etch stop layer for DRIE. 5. A protection photoresist layer is spun on the components side. The wafer is ready for DRIE (see Note 7). 6. The etching of the cavities (Fig. 2C) takes about 40 min in an Alcatel 601 DRIE machine (see Note 8). 7. After etching the cavities, clear silicon dioxide membranes are obtained, and a short, about 20 s, overetching without passivation is applied in an Alcatel 601 DRIE machine to remove the polymer formed in the cavities during the DRIE process. 8. The membranes of silicon dioxide are etched in vapors of Hydrofluoric (HF 49%) acid. It is a type of dry etching, because contact of the wafer with liquid HF must be avoided. The wafer is placed with the back side down on a pot with the acid. The pot has a diameter smaller than the diameter of the wafer, i.e., if the wafer has a diameter of 4 in., the pot should have a diameter of about 3.8 in. (see Note 9) 9. The wafer is rinsed with deionised water in a special bath with gentle filling and draining (see Note 10). 10. The wafer is dried at room temperature. 11. The photoresist is removed by oxygen plasma dry etching, see Fig. 2D. 12. The metallization of cantilevers is done in two steps: first, it is coated with an adhesion layer of Cr or Ti and after a gold layer is deposited by e-beam evaporation. E-beam allows a precise control of the coating thickness and the low temperature variation during evaporation makes the cantilevers less bended.
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3.2. Cantilever Mechanical Properties Characterization
1. The cantilever spring constant, k, could be calculated from the expression k = Ebh3 / 4L3, where E is the Young’s modulus, h the thickness, b the width, and L the length, respectively. Experimentally, the spring constant was determined by exerting a controlled force on the fabricated cantilever free end by using an atomic force microscope (AFM) with a calibrated cantilever (Fig. 3A). To measure the force that the AFM-tip applies over a surface, we used a force–distance curve, which is a plot of the AFM-cantilever deflection as a function of sample position, or piezoelectric displacement, along the z-axis. The fabricated cantilever spring constant is obtained from the force–distance curves slopes (Δz / ΔV) measured when exerting force over the fabricated cantilever and over a hard surface, being ΔV the photodetector’s voltage signal and Δz the piezoelectric’s displacement, in the AFM. Figure 3A shows an example of the force–distance curves measured; the AFM-cantilever deflection is higher when pressing against the hard surface (high slope) than over the fabricated cantilever, because of both cantilevers coupled-deflection. The relation obtained between the cantilevers spring constants and the force–distance curves slopes is given by: ù kAFM é DV hard / Dz hard =ê - 1ú . k ë DV fab_cant / Dz fab-cant û 2. Because of the distance, d, from the cantilever free end to the point where the force is exerted, a correction must be included in the value of the spring constant previously obtained. æ L -dö k final = k ç è L ÷ø
3
A
B ΔV
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AFM z L-d
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L
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Distance, μm
Deflection signal (V)
1,5 1
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−2 −200
−100
−0,5 0
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Vertical displacement (nm)
200
0
75
Distance, μm
150
225
Fig. 3. (A) Static method for determining the cantilever spring constant. Force–distance curves obtained over a hard surface and over a fabricated microcantilever. The inset shows the method configuration scheme. (B) A profile of silicon cantilevers fabricated by the technology described above. The cantilever is flat, but the stresses at the anchoring area produce initial cantilever displacement. The profile has been measured by an optical confocal profilometer.
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With this method, a spring constant value of 0.0067 N/m was obtained for microcantilevers of 200-μm length, 40-μm width, and 0.334-μm thick. The difference respect to the expected value (0.0079 N/m) is attributed to the compliance of the chip region where the cantilever is clamped (due to a thinning of this region, produced during the fabrication process) (2). 3. The profile of the fabricated cantilevers was measured with a confocal microscope, to check the initial curvature generated during the fabrication process. A measured profile is shown in Fig. 3C. This profile shows that the cantilever is flat; however, a biasing due to stresses at the anchoring area can be observed (see Note 11). 3.3. Set-Up Configuration
1. All the biological interactions must be performed in a liquid medium. For that reason, the cantilever chip must be fixed into a fluid cell, by using a spring system. The fluid cell has an inlet and an outlet for liquid flow. The top part of the fluid cell is made out of glass, and the bottom is closed with a coverslip using a rubber “O”-ring. 2. The flow system is mounted by connecting the peristaltic pump to the injection valve and this one to the cell flow inlet by using Teflon tubes. The length of the tubes should be reduced as much as possible to avoid losses associated with the adsorption of protein on the tube walls and the dispersion effects of two different fluids inside the tube. The loop of the injection valve has the suitable length for injecting a solution volume of 200 μL. An extra tube is used from the cell outlet to the waste glassware (see Fig. 4A, B). 3. The flow cell is mounted over a rotatory stage that allows changing the reflection angle. This rotatory movement can compensate the reflected angle changes due to different mediums (air or liquid). 4. The diode laser and the photodetector are mounted over x–z positioning stages. In this way, it is possible to focus on a specific cantilever into an array and to adjust the position along the cantilever length where the laser will be reflected, increasing the bending sensitivity. At the same time, the photodetector position could be modified for centering the reflected laser beam. Figure 4A shows a scheme of the set-up configuration. A diagram showing the optical detection system is illustrated in Fig. 4C. 5. The distance between the diode laser, the cantilever, and the photodetector was chosen by weighing up the reflected beam spot size on the photodetector, because of the beam dispersion and the spot displacement magnification that could be obtained by increasing the cantilever–photodetector distance. 6. The photodetector signal is readout and processed with a digital acquisition card connected to a computer. A home-made
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Fig. 4. (A) A schematic diagram of the experimental set-up (top view). The distance between the laser and the cantilever was fixed to the laser focal length, f, while the distance between the cantilever and the photodetector, d (d = 12 cm), was adjusted to fit the maximum spot displacement to the photodetector size (a lens can also be used for that purpose). (B) Schematic diagram (top view) of the cantilever-fluid cell, mounted over the rotatory stage. (C) Diagram and picture (front view) of the optical detection system (laser, photodetector, rotatory stage, and beam path).
software allows the continuous photodetector signal acquisition in real time. 3.4. Real-Time Detection: Surface Functionalization and Pesticide Detection 3.4.1. Gold Cantilever Cleaning Procedure
1. Gold surfaces must be thoroughly cleaned to avoid contamination of the surface and to enhance reproducibility from one measurement to another. This procedure must be performed immediately prior to performing the measurement. First, submerge consecutively for 2 min the cantilever on 2 mL of tricloroethylene, acetone, ethanol, and deionized water, respectively. 2. Dry carefully the cantilever under nitrogen flow and submerge it for 1 min on fresh piranha solution (70% H2SO4–30% H2O2). 3. Quickly rinse generously with water and again dry carefully the cantilever under nitrogen flow. 4. Cantilevers are ready to be positioned inside the flow cell to start the measurement. Once cleaned, cantilevers cannot be stored as they are susceptible to suffer contamination. New cleaning
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steps cannot be performed as successive rinses with piranha dramatically reduce the thickness of the gold layer. 3.4.2. Immobilization of the Receptor
We have demonstrated good results of DTT detection on cantilevers, using a chemistry based on glutaraldehyde (3). This molecule displays an aldehyde group in both the extremes of the molecule, and it is possible to couple the free amine groups of the bioreceptor to those from a previously immobilized SAM with amine groups (such as cystamine) by using this molecule. This renders an easy immobilization method, showed in Fig. 5A, where the first signal displacement corresponds to the formation of the SAM, the second to the glutaraldehyde bonding, and the third to the bio-receptor immobilization (the last deflection is due to the blocking of all the unreacted groups with ethanolamine). However, later experiments performed at our laboratory using SPR biosensor with the same gold-based chemistry demonstrated that this methodology displays an important drawback, as glutaraldehyde tends to form multilayers that can be partially removed during regeneration, giving less reproducibility from measurement cycle to cycle. For that reason, we shifted to a carbodimide (EDC)-mediated coupling chemistry, as enhanced results are achieved using this immobilization method (4). We strongly recommend using the EDC chemistry, and it will be detailed here. Similar cantilever behavior to the one showed in Fig. 5A has been observed with this immobilization method.
Self-Assembly of a Carboxylic-Alkanethiol onto the Gold-Coated Cantilever Surface
1. Prepare a 50 μM solution of mercaptoundecanoic acid on ethanol. The final volume might vary from one flow system to another, however, typically ranges from 200 to 400 μL.
Fig. 5. (A) Example of a cantilever surface functionalization in real time. Each one of the layers formed over the surface produce a cantilever bending. (B) Real-time monitoring of an antibody direct detection and a competitive immunoassay. The number of antibodies free in solution able to binding the cantilever surface is reduced due to the binding with the DDT free in solution. The cantilever surface was regenerated with 100 mM HCl (100 ml) to break the hapten/antibody complex; (Reprinted from (3). Copyright (2003), with permission from Elsevier).
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Hereinafter, this volume will be referenced as “one volume or 1× V” and half or double volumes as ½ V or 2× V, respectively. 2. Pump the mercaptoundecanoic acid solution through the injection loop and inject the sample into the flow cell. Flow speed must be adjusted and calibrated on each system. It is recommended a speed that keeps the sample volume in touch with the gold surface during at least 20–30 min; flow speeds in the range of 0.2–0.3 μL/s are strongly recommended. The SAM formation produce an initial deflection, mainly due to the covalent bond between the thiol group (−SH) and the gold atoms, and the interchains forces, which produce a compressive surface stress. The detected signal is equivalent to the one showed in Fig. 5A. 3. Pump 1× V of ethanol at 0.15 μL/s and proceed identically with a new injection of 1× V of a mixture ethanol/water 1:1 and finally 1× V of water. EDC–NHS Mediated Antigen Coupling (See Note 12)
1. Stabilize the system with a continuous flow of water and wait until thermal drift is null. This water flow will be continuously pumped and only interrupted with samples injections in between by turning the injection valve from the “load position,” to charge the samples into the injection loop, to the “injection position” to introduce the samples into the flow cell. 2. Prepare separately and then mix ½ V of an aqueous solution of EDC 0.4 M and ½ V of a solution of NHS 0.1 M. These solutions must be prepared immediately prior to their use. They are unstable and present short-lived in aqueous solution. 3. Pump the EDC–NHS solution into the flow cell at 0.3 μL/s. This step produce a second cantilever deflection, equivalent to the second one observed in Fig. 5A. 4. Pump immediately after the EDC–NHS solution, a previously prepared 1× V solution of the bioreceptor in the immobilization buffer (see Note 13) at 0.15 μL/s. Concentration must be optimized for each application; however, it typically ranges from 10 to 50 μg/mL. For DDT detection, 1× V of 6 μg/mL in PBS pH 7 should be used. The bioreceptor immobilization produce a new deflection, very similar to the one showed in Fig. 5A. 5. Pump 1× V of water or diluted acid (for example HCl 0.1 M) to remove the excess of the reagent and crosslinking products. 6. A second injection of the bioreceptor is recommended to ensure a maximum yield of immobilization. 7. Pump a 1× V of an aqueous solution of ethanolamine 1 M in such a way that all remaining unreacted NHS-esters could be deactivated (see last signal deflection in Fig. 5A).
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1. Restabilize the system in a continuous flow in the same buffer to the one that will be used to perform the immunodetection assay. For DDT inhibition assay, detection is performed under phosphate buffer saline solution (PBS) pH 7. 2. Prepare series of solutions (minimum 4–5) of different concentrations of the antibody in triplicate. For DDT detection, concentrations ranging from 0.5 to 10 μg/mL were used. 3. Depict the surface stress values to a calibration curve. Choose a value as high as possible, but below the saturation limit, able to give a high surface stress signal, so that at inhibition, signals would be still clear. This will be the antibody concentration that will be fixed to perform the inhibition assay.
Inhibition Assay: Detection of the Pesticide DDT
1. Ensure yourself that the system is equilibrated in the immunodetection assay buffer (see Note 14) and that the thermal drift is null. 2. Prepare and incubate for 10–15 min a series of mixture with the chosen concentration of the antibody, in our case, 5 μg/ mL and different amounts of the antigen. Concentrations of the antigen to be measured depend on the limit of detection for each specific application and antigen type. Thus, for pesticides, they should be in the range of ppb or ppt and for other molecules in the nM–pM or ng/mL–pg/mL range. 3. Pump each of the series with a regeneration step in between. Monitoring of antigen–antibody interaction can be performed at a flow speed of 0.15 L/s, while regeneration can be performed faster, at 0.3 μL/s. Regeneration solution must be optimized for each particular application (see Note 14); however, excellent results have been achieved in our lab using diluted acid such as HCl 0.1 M or 1–5 mM HCl. In Fig. 5B, three sequential injections of antibody (5 μg/mL of antibody, 5 μg/mL of antibody with 3.5 ng/mL of antigen, and 5 μg/mL of antibody), with the corresponding surface regenerations, are represented. The antibody/receptor recognition over the cantilever is again translated into a compressive bending, while the regeneration step produces the opposite effect, recovering the initial level before the antibody bonding. A reduction in the deflection signal is observed when a combination of 5 μg/mL of antibody with 3.5 ng/mL of antigen is injected. The second injection of pure antibody produces the same change in the deflection than the first one, meaning that the reduction in the signal is effectively produced by competitive assay and not by degradation on the bioreceptor layer. 4. Depict the surface stress values for antibody inhibited signal for each of the series with different antigen amounts and the fixed antibody concentration to a calibration curve. Lower limit of detection should be in the desired range for the
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application. If not, try to improve and optimize immobilization and immunoreaction conditions to reach higher signals.
4. Notes 1. This system can be easily scaled up allowing the detection of different concentrations and/or different analytes simultaneously. For biological experiments, arrays of cantilevers are normally employed (5) as one cantilever can be used as reference. Then the expected biointeraction can be diferenciated from any change of the experimental conditions. Actually, there are several commercial platforms available, based on cantilever-array sensors. 2. Other materials, such as polymers, are a cheaper alternative to increase the cantilever deflection, as the Young’s module of polymers are lower than that of silicon (6). 3. The bending detection method has a complex relation between the measured signal and the factors producing it, because of the different forces acting in the biorecognition processes. However, it reports a new type of information and represents a new alternative for biological discrimination, such as DNA single-base mismatch polymorphisms or single-molecule detection, otherwise not possible with other well-established biosensing techniques. 4. There are several techniques used for the cantilever bending detection, such as capacitive, piezoelectric, and piezorresistive; however, the laser beam deflection technique is maybe the most extended one, because of its simplicity and higher sensitivity. When working with arrays of microcantilevers, the detection of the bending by using the laser beam deflection read-out configuration could be a great challenge, because of the number of lasers and/or photodetectors required. Several strategies have been explored for solving these issues, and there are currently several systems commercially available on the basis of this redout method. An alternative configuration has been proposed by using arrays of optical waveguide cantilevers (7). The cantilevers act as a waveguide for conducting light. The exit light is collected by a second waveguide and finally with a photodetector. In this case, the cantilever bending is related to the reduction in intensity of the collected light with respect to the input light. 5. The technology for the fabrication of single cantilevers is basically the same as the technology for fabricating arrays (8). Only some processes should be appropriately chosen to satisfy the requirements for the array geometry. In our example, the cavities under the cantilevers were formed using aniso-
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tropic reactive ion etching instead of wet etching. This is due to the required distance of 250 μm between the cavities on 450 μm thick wafer, which cannot be maintained if anisotropic wet etching with bases were applied. In Fig. 6A, cantilevers extended from a border of a chip are shown. The fabrication procedure is similar to the technology for the fabrication of array of 20 cantilevers located in separate cavities and showed in Fig. 6B. The technology of array fabrication was developed to reach a 100% yield. Two thousand five hundred cantilevers per wafer were manufactured. 6. The silicon dioxide layer on the back side of the wafer may be not specified by the manufacturer. It is always recommended to measure the thickness of this layer before starting the process. 7. Protection layer of the photoresist is essential in this technological chain. If this layer is not used, the membranes of silicon dioxide would crack due to internal stress after being released from the substrate. Cracking the membranes results in a breaking on the cantilevers and brings technological yield close to zero. 8. The BOSCH deep reactive ion etching process is based on alternating etching and passivation cycles. (SF6 is used for etch cycle and C4F8 is used for the passivation cycle). With the appropriate recipe, it is possible to obtain structures with high aspect ratio and to form vertical walls in through-wafer etching. The imperfections of the technology as small lateral etching rate and notching (9) result in a small undercut from the back side and slight walls inclination by about 5°. The windows size in the photolithographic mask used for DRIE was corrected to obtain the desired cavity size at the components side. After the photolithography step and before the deep etching, it is also important to make sure that there are no impurities and rests of oxides in the openings on the back side. Otherwise, long spikes hanging from the silicon dioxide membranes, which are difficult to remove, can be formed by DRIE.
Fig. 6. (A) Photo of a chip with three cantilevers extended from the chip border. The photo has been taken before the wafer dicing. (B) An array of 20 cantilevers located in separate cavities. The cantilevers on both photographs were fabricated at the clean room facilities of CNM-IMB, Spain, from SOI wafers using the same technology.
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9. The conventional wet processing does not work well here. If the wafer was immerged into the acid directly, then HF could penetrate under the photoresist protection layer and peel it off, which would lead to a cracking of the membranes. Caution: HF should be carefully handled and could be necessary a previous training, because fluoride ion readily penetrates the skin, causing destruction of deep tissue layers, including bone. 10. Generally speaking, starting from this step, where the cantilevers are released after etching of the membranes, wet processing of the cantilevers should be avoided. Manipulating the cantilevers in liquids may result in fracture of many of them. The wafer is placed into an empty bath, which is slowly filled with deionized water. Rinsing the cantilevers with water at this stage should be done carefully; the wafer should not be moved while it is inside water. After 30 min, the bath is drained slowly and the wafer is dried at room temperature in a clean room environment. 11. Initial bending/curvatures of the cantilevers must be low to avoid high divergence of the reflected beam when the optical beam deflection method is used. This requirement is stronger in arrays of cantilevers. Two phenomena that can induce initial deflection/curvatures of a microcantilever can be identified: (a) The most common one is the gradient stress within the structure. This phenomenon induces a cantilever deflection with a constant radius of curvature that can be predicted by Stoney’s formula. This can be solved by using materials with low residual stress gradient or by compensation of the stresses with different layer materials. However, the best solution is to use single crystalline materials as monocrystalline silicon. (b) The second phenomenon is the stresses induced by the anchoring, which can induce an initial offset to the cantilever deflection. Several mechanisms cause stresses at the anchoring area that can induce the cantilever to rotate. The origin of this effect is difficult to determine and avoid. Several solutions have been presented: change the mechanical stresses at the anchoring area by the laser bending (10), or the design of a special T-shape geometry for the cantilever (11). 12. EDC-mediated coupling: This step is crucial for the success of the immobilization procedure. EDC reacts with the carboxyl groups to form an amine-reactive O-acylisourea intermediate. If this intermediate does not encounter an amine, it will hydrolyze and regenerate the carboxyl group. In the presence of N-hydroxysulfosuccinimide (Sulfo-NHS), efficiency of EDC coupling is increased by the formation of an amine-reactive Sulfo-NHS ester intermediate, which has
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sufficient stability to permit two-step cross-linking procedures, as it shows in Fig. 7. 13. Immobilization buffer: Immobilization buffer conditions can dramatically vary from one antigen type to another depending mainly on their intrinsic isoelectric point. Good immobilization signals are typically achieved on immobilization buffers displaying an equal pH to that of the isoelectric point of the protein; however, there is not a universal rule and immobilization should be tested also for an upper and lower pH buffer. 14. Immuno-detection assay buffer and regeneration conditions (12): Procedures used for regeneration are mostly empirical. The most frequent procedure, i.e., the use of low pH with low ionic strength (e.g., 10 mM glycine, pH 1.5–2.5), probably works because most proteins become partly unfolded and positively charged at low pH. As a result binding sites no longer match and the molecules repel each other. Other procedures include the use of pH 10.5, high salt concentration, or chemicals such as urea or guanidine hydrochloride. When none of these conditions work properly, a trial-and-error search is conducted to find the suitable regeneration conditions. The type of regeneration condition that is most effective for a particular antibody–antigen interaction depends on the specific mixture of ionic, hydrophobic, and hydrogen bonds involved. Other factors to be taken into account for regeneration are: the antibody type (polyclonal antibodies respond to different epitopes by slightly different mechanisms; therefore, effective removal of all polyclonal antibody from the immobilized antigen layer may require several different regeneration conditions) and affinity constant of the antibody for the antigen (the higher is the affinity constant, the hardest should be the regeneration condition, and the lower the number of regeneration cycles that can be accomplished). The protocol must be chosen according to its ability to target several binding forces simultaneously by mixing different chemicals into a regeneration cocktail. This is quite likely to increase the chances of reversibly breaking the interaction under
Fig. 7. Scheme of the chemical linkage of the bioreceptor to carboxylic groups through EDC–NHS mediated coupling.
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more gentle conditions. The ideal regeneration buffer should effectively release the antibody without irreversibly denaturing or inactivating them. In practice, all regeneration buffers cause some loss of antibody or antigen function, limiting the number of cycles that the same functionalized cantilever can be reused. pH and ionic strength conditions are gentle on antibody and protein function, while denaturants and organics are harsh. Usually, a low-pH condition is the best option to try first. If a low pH regenerates efficiently but causes irreversible functional damage, then consider testing an ionic strength buffer. If the low-pH buffer does not elute effectively, then consider testing one of the highpH conditions. To reach a condition that allow repeated regeneration of antigen or antibody (especially when using monoclonal antibody), consider testing a variety of buffer conditions to find the one that is as gentle as possible while still being effective.
Acknowledgments The authors acknowledge to the European Union (Project Optonanogen, IST-2001-37239). Authors thank Dr. Angel Montoya (Ci2B, University of Valencia, Spain) for the inmunoreagents.
References 1. Ziegler, C. (2004) Cantilever-based biosensors. Anal. Bioanal. Chem. 379, 946–959 2. Alvarez, M., Tamayo, J., Plaza, J.A., Zinoviev, K., Dominguez, C. and Lechuga, L.M. (2006) Dimension dependence of the thermomechanical noise of microcantilevers. J. App. Phys. 99, 024910 3. Alvarez, M., Calle, A., Tamayo, J., Lechuga, L.M., Abad, A. and Montoya, A. (2003) Development of nanomechanical biosensors for detection of the pesticide ddt. Biosen. Bioelectron. 18, 649–653 4. Mauriz, E., Calle, A., Lechuga, L.M., Quintana, J., Montoya, A. and Manclus, J.J. (2006) Real-time detection of chlorpyrifos at part per trillion levels in ground, surface and drinking water samples by a portable surface plasmon resonance immunosensor. Analytica Chimica Acta 561, 40–47 5. Zhang, J., Lang, H.P., Huber, F., Bietsch, A., Grange, W., Certa, U., Mckendry, R.,
Guntherodt, H., Hegner, M. and Gerber, C. (2006) Rapid and label-free nanomechanical detection of biomarker transcripts in human RNA. Nat. Nanotech. 1, 214–220 6. Ransley, J.H.T., Watari, M., Sukumaran, D., McKendry, R.A. and Seshia, A.A. (2006) Su8 bio-chemical sensor microarrays. Microelectron. Eng. 83, 1621–1625 7. Zinoviev, K., Dominguez, C., Plaza, J.A., Cadarso, V. and Lechuga, L.M. (2006) A novel optical waveguide microcantilever sensor for the detection of nanomechanical forces. J. Lightwave Technol. V.24(5) (2006) 24, 2132–2140 8. Lechuga, L.M., Tamayo, J., Álvarez, M., Carrascosa, L.G., Yufera, A., Doldán, R., Peralías, E., Rueda, A., Plaza, J.A., Zinoviev, K. et al. (2006) A highly sensitive microsystem based on nanomechanical biosensors for genomics applications. Sens. Actuators B 118, 2–10
Biosensors Based on Cantilevers 9. Kumar, S. and Pike, W.T. (2005) Technique for eliminating notching in through-wafer etching. 16th MME Micromechanics Eur. Workshop P15, 88–91 10. Zhang, X.R. and Xu, X.F. (2005) Laser bending for high-precision curvature adjustment of microcantilevers. App. Phys. Lett. 86,021114
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11. Plaza, J.A., Zinoviev, K., Villanueva, G., Álvarez, M., Tamayo, J., Domínguez, C. and Lechuga, L.M. (2006) T-shaped microcantilever sensor with reduced deflection offset. Appl. Phys. Lett. 89, 094109 12. Andersson, K., Areskoug, D. and Hardenborg, E. (1999) Exploring buffer space for molecular interactions. J. Molec. Recogn. 12, 310–315
Chapter 5 Piezoelectric-Excited Millimeter-Sized Cantilever Biosensors Raj Mutharasan Summary In this chapter, method of fabricating a cantilever biosensors and their use in measuring the presence of a protein is described. There are many variations in construction of a cantilever sensor. A simple and an easy version is described in this chapter. The specificity of the sensor is obtained by immobilizing an antibody specific to the antigen of interest. The piezoelectric cantilever sensors are very sensitive and can easily detect a 60 kDa protein at 100 fg/mL concentration. Experimental procedure for carrying out detection of a target analyte is outlined and a sample set of results is included. Key words: Biosensor, Fabrication, Resonant frequency, Immobilization, Antibody, Mass-change.
1. Introduction Cantilever biosensors have attracted considerable interest in the past decade for label-free detection of proteins and pathogens as they are described as being very high sensitivity. Excellent reviews have appeared that summarize progress (1, 2). The cantilever sensors can be broadly divided into two subcategories: (1) bending or static cantilevers, and (2) dynamic or resonating cantilevers. In the bending mode, the binding of an antigenic target to the cantilever surface changes the cantilever’s surface stress resulting in a deflection response that is proportional to the analyte. In dynamic mode, the attachment of antigen causes a resonance frequency decrease due to increase in mass. When dynamic mode is used in liquid, significant damping occurs, and bending mode is preferred when continuous measurement under liquid immersion is needed. When measurement in liquid flow condition is required, the bending mode becomes noisy. Dynamic mode yields Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_5
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good results as long as cantilever Reynolds number (2) is large, preferably greater than 105. Since Reynolds number (Re) is proportional to the square of cantilever width, response of microcantilevers whose width are of a few micrometers are highly damped in liquids, while millimeter-wide cantilevers provide relatively less damped response and can be used in liquid flow environments (4). For mechanical robustness, cantilever sensors should be much thicker than the thickness of microcantilevers which are typically ∼1 μm. Thick cantilevers exhibit such a weak bending response that the only useful approach is to use the dynamic or resonant mode for measurement. We describe here millimeter-sized cantilever sensors that the author’s lab developed for pathogen (5–12) and toxin detection (13, 14), density and viscosity (15–17) as well as for measuring fundamental adsorption phenomena (18–20). Unlike microcantilevers, they are mechanically robust and can be used under full liquid immersion and flow (21). The sensor is illustrated schematically in Fig. 1. The piezoelectric-excited millimeter-sized cantilever (PEMC) sensor is a macro-cantilever that comprises piezoelectric layer (lead zirconate titanate; PZT) layer bonded to a nonpiezoelectric layer of a few millimeters in length and 1 mm in width (9, 21). We use the direct piezoelectric effect to excite the cantilever, and the same PZT film senses the resulting response. PZT film is bonded to a base glass
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Fig. 1. (A ) Schematic arrangement of sensor in glass tube. Adapted from reference (21). (B) Cross-sectional view of flow cell with the sensor installed with O-ring.
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cantilever forming a composite cantilever. When an electric field is applied across the thickness of the PZT film, it extends along its length causing the underlying glass to bend. If the applied field is alternated periodically, the composite cantilever vibrates. The natural frequency of the cantilever depends on the flexural modulus and mass density of the composite cantilever. At resonance, the cantilever undergoes significantly higher stresses when the exciting electric field is at resonance frequency. Hence, the PZT layer exhibits a sharp change in electrical impedance and can be followed by the phase angle measurement. PEMC sensors have been found to exhibit very high mass change sensitivity of 0.3–2 fg/Hz (9, 21). The implication of this finding is that attachment of a few million 60 kDa molecules can be measured.
2. Materials 2.1. Materials for Sensor Fabrication
1. Lead Zirconate Titanate (PZT) Sheet; Piezo Systems Inc., Cambridge, MA. Catalog Number: T105-A4E-602. 2. Gold-coated cover slip. (Sigma-Aldrich, 99.999% (Au), layer thickness 100 Å, length × width × thickness 22 mm × 22 mm × 160 μm, Catalog Number: 643254). 3. Bonding agent: Loctite 409 Super Bonder Cyanoacrylate (Ellsworth Adhesives, Germantown, WI, Part No: 18030). 4. Potting compound: Loctite 1C Hysol Epoxi-Patch Adhesive (Ellsworth Adhesives, Germantown, WI. Part No: 83200). 5. Connecting wire. 30 gauge wire. (Radio Shack, Part no: 278–503). 6. Pyrex tubing, 6.3 mm OD. (Corning Inc. Part No: 237300). 7. Diamond knife. Fischer Scientific. 8. Vacuum oven. Fischer Scientific. 9. Soldering apparatus (Radio Shack, Catalog No: 64–2184). 10. Solder (Piezo Systems Inc., Cambridge, MA. Catalog number: MSF-003-NI). 11. Polyurethane (MC Clear, Wasser Corp., Auburn, WA). 12. Access to gold coating apparatus (e.g. Denton Desk II System, Denton Vacuum, New Jersey).
2.2. Materials for Sensor Surface Preparation
1. Sample containing target analyte (bovine serum albumin; BSA) (Sigma-Aldrich.com, CAS Number: 9048-46-8). 2. Antibody to target analyte – rabbit polyclonal antibody to BSA, (Sigma-Aldrich.com, Catalog No: A4338).
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3. Centrifuge tubes (1.2 mL) (Fisher Scientific). 4. Protein G (Sigma-Aldrich.com, Part No: P4689). 5. Phosphate buffered saline solution (1× PBS; 137 mM NaCl, 10 mM Phosphate, 2.7 mM KCl, pH 7.4). 6. Piranha solution (7:3 volume ratio of concentrated H2SO4 and 30% H2O2) Caution: piranha solution reacts violently with many organic materials and should be handled with great care. 7. Spin coater or a centrifuge that will run at 500 × g (Fisher AccuSpin* Model 400, Catalog No: 13-100-557). 2.3. Materials for Conducting Detection Experiment
1. Impedance analyzer – Agilent 4294A or equivalent
2.4. Materials for Flow Apparatus
1. Reservoirs for flow apparatus (Fig. 2) – 10 mL and 60 mL syringe barrels (coleparmer.com, Catalog No: WU-07940-16 and WU-07940-30).
2. Software to run impedance analyzer remotely for recording resonance frequency values as a function of time. Details given in Note 1.
2. Luer valves V1 and V2 (coleparmer.com, Catalog No: WU-30600-06). 3. Tubing – peroxide-cured silicone tubing, 3.2 mm (coleparmer. com, Catalog No: WU-06411-67). 4. Two-port manifold (MP-2 Perfusion manifold, harvardapparatus.com, Catalog No: 640206). 5. Flow cell (custom made from Plexiglas® with a 4 mm internal diameter with an inlet and outlet fitting as shown in Fig. 1).
3. Methods 3.1. Sensor Fabrication
A schematic of an assembled sensor is given in Fig. 1. Step by step instructions of construction is given below. These fabrication steps require a fair amount of dexterity. Do not be disappointed if several attempts are needed for a working sensor. 1. Using the diamond knife cut glass into 1 × 4 mm2 pieces. If you have access to a micromachining shop, have them cut to the dimension noted. 2. Cut PZT sheet into 1 × 5 mm2 pieces 3. Place PZT piece on a clean surface, and dispense 0.2 μL of Loctite 409 Super Bonder cyanoacrylate. Place the glass piece on the droplet, centering it such that approximately 1 mm of
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Sample 1 mL
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Fig. 2. Continuous flow apparatus; Adapted from (21). The flow cell was maintained at a fixed temperature. The circulating peristaltic pump was typically run at 1 mL/min flow rate. A LabView™ program running in the PC helped with data collection and analysis.
the glass protrudes beyond the PZT. Apply gentle pressure to glass piece until bonding occurs (∼20 s). 4. Strip insulation off two 10 cm long 30 gauge wire, tin it, and then make a 30° bend. 5. Using a thin wire (22 or 24 gauge wire works well) and dispense a drop of flux on PZT about 0.5 mm from the free end. 6. Carefully solder the bent 30-gauge wire at the location of flux using a small amount of solder for bonding. Do one side at a time, ensuring that excessive heat is not applied to PZT. 7. Using a glass cutter, snip a 5 cm long pyrex tube. Position the PZT-glass composite in the center of the tube cross section ensuring the solder point is within the tube. Carefully thread the wires out through the far end. You should have about 5 cm of free wire. Next step is to fix the sensor in place using epoxy.
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8. Following manufacturers instructions, mix Loctite 1C Hysol Epoxi-Patch Adhesive on a disposable surface. Dispense on either side of the sensor the mixed epoxy and inside the tube (∼2 mm) such that the sensor is fixed in place. Place a dab of the epoxy at the far end to fix the wires to the tubing so that no stress is transmitted to the sensor when moving the wires. 9. Cure the composite for 8–12 h at room temperature. 10. Using a thin artist brush, paint the PZT and the adhesive layer with a thin coat of polyurethane. Spin down at 500 × g for 5 min in an available spin coater, or in a laboratory centrifuge. 11. Cure at room temperature for 8 h followed by oven cure at 80°C overnight. 12. Prepare the piranha solution. Dip the distal end of the sensor (glass layer) in piranha solution for 3 min. Rinse in copious amounts of DI. Dry in a dust-free oven at 80°C for 1 h. 13. If the sensor has been subjected to step 12 more than three times, it is better to recoat with 100 nm gold. This can be done at most universities at the microfabrication facilities. 14. Now the sensor is ready for antibody immobilization. 3.2. Antibody Immobilization
In this section, we present antibody immobilization that takes advantage of orienting the binding region (Fab) away from the sensor surface using immobilized Protein G. Many methods that are equally effective have been reported in the literature. The reader is referred to Hermanson (22) for a complete discussion on reaction mechanisms. 1. Prepare 10 μg/mL stock solutions of antibody and Protein G. 2. Dispense 0.5 mL of protein G stock into 1.2 mL centrifuge tube. Immerse a clean sensor into the tube for 1 h. 3. Remove sensor and rinse with clean PBS. Dispense 0.5 mL of antibody solution into a clean 1.2 mL centrifuge tube. Immerse the protein G coated sensor into the tube for 1 h. 4. Remove sensor and rinse in PBS. 5. Now the sensor is ready for detecting the target analyte.
3.3. Detection Experiments
Assemble a flow apparatus as given in Fig. 2. This is similar to the apparatus we have published (9, 21). It is best done in consultation with a machine shop or a model building shop you may have access to at your campus. You will need two reservoirs (10–60 mL depending on your application; sterile syringe of 10 or 60 mL works well), tubings, and one peristaltic pump capable of pumping at 1 mL/min with minimal pulsation. Key to the
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apparatus is a sample flow cell that will house the sensor fabricated in Subheading 3.1. The one we have had good success has been published in reference (21). An enlarged figure of the flow cell is included in Fig. 1. The apparatus arrangement and flow circuit is included in Fig. 2. 1. Fill reagent reservoirs with 50 mL of PBS, and 1 mL of target BSA at 1 pg/mL. 2. Close valve V2 and open valve V1. 3. Start the peristaltic pump at 1 mL/min with an empty sensor flow cell (SFC). 4. As the SFC fills and overflows, insert the sensor, and fix it in place. 5. Insert the exit tubing from SFC into the PBS reservoir. 6. Start the data acquisition program that will collect the resonance frequency values. 7. It will take 5–15 min to achieve stabilization of baseline frequency. 8. When this occurs, open V2 and close V1, and simultaneously take the exit tubing and insert into the sample chamber. 9. As the target BSA enters the flow cell, one should see decrease in resonance frequency. Run until new steady state is achieved. 10. After reaching steady state, if release of antigen in needed, one can add another reservoir to the apparatus that contains a suitable low pH release solution. At a suitable time, this solution can be admitted in while shutting valve V1. One will see a shift up in frequency. 11. For a sample result see Fig. 3 and Note 2.
4. Notes 1. Software: The impedance analyzer, such as the Agilent 4294A, will sweep a set frequency range (sinusoidal and other) and locate maximum phase angle within that range in a contin-uous fashion. For monitoring a detection experiment, it would be useful to capture the frequency at which maximum phase angle occurs (resonance frequency) in an automatic fashion via a personal computer. Hence, the primary function of the PC is to data log resonance frequency values with time and other sensor properties (such as impedance) in a continuous fashion. Usual width of sweep is 10–2 kHz
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Fig. 3. Detection of bovine serum albumin. Resonant frequency change as BSA binds to an antibody-functionalized PZT-anchored cantilever is illustrated. In these experiments, the resonance at 902.5 kHz was used at sample flow rate of 0.5 mL/min. (A) This shows the sequential attachment of anti-BSA at 100 μg/mL, BSA detection at 1 pg/mL, and the release of the bound BSA using a pH 2 solution. The anti-BSA attachment caused a frequency decrease of 6,954 ± 21 Hz. The PBS rinse that followed did not cause any significant change in the resonant frequency, indicating that nonspecific binding, if any, was small. Upon exposure to 1 pg/mL BSA, the sensor responded with an immediate decrease in frequency and reached a steady-state change (2,022 ± 25 Hz) in ∼20 min. The PBS rinse that followed shows a slight increase in frequency, which may be a result of the removal of nonspecifically adsorbed BSA. A pH 2 release solution caused the resonant frequency to increase by 413 ± 10 Hz above the value prior to BSA attachment. After stabilization, PBS was again pumped in to obtain final resonant frequency for comparison. In PBS, the frequency increased further by 186 ± 5 Hz. The higher value obtained may be due to some release of antibody. (B) This shows the sensor response to 100 fg/mL of BSA in three different experiments. The positive control was a silanylated cantilever exposed to flowing PBS and the negative control was an unsilanylated clean sensor in presence of 10 pg/mL BSA solution. One notes that the signal to noise (S/N) ratio was a minimum of 15. Adapted from our publication (21).
(excitation voltage of 10 to 100 mV; sine wave), with the specification of central frequency. The latter will depend on the sensor and it is wise to measure it prior to the experiment and load this value in the program. If you anticipate a large frequency change during a detection experiment, it is recommended that the data acquisition program adjust the central sweep frequency after each sweep.
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2. Sample result: A sample result of BSA detection in a flow apparatus similar to the one shown in Fig. 2 is illustrated in Fig. 3. In this early result from our lab (21), the sensor was prepared by sialynation followed by antibody immobilization through carboxylic group activation chemistry (23). In this case, the immobilization was done with antibody concentration of 100 μg/mL, and the immobilization reaction was measured in situ. The flow apparatus is convenient for conducting multistep reaction, cleansing and release experiments. As shown in Fig. 3, the release of the detected antigen is achieved in situ by changing the inlet fluid from the PBS buffer to a low pH buffer. In Fig. 3b, we compare the results of three repeat experiments with controls (positive and negative). The repeat experiments were conducted with the same sensor but each time the sensor was cleaned in Piranha solution and the entire sequence of immobilization was redone. Considering the multistep preps, the response values of 290, 304, and 423 Hz for such a low target concentration (100 fg/mL) is quite good. The negative control is for the case where the sensor was prepared with the antibody but the sample did not contain BSA. Positive control is when the sensor was not prepared with the antibody, but BSA at 10 pg/mL was present in the sample.
Acknowledgment This work was partially supported by the grants USDA 200651110-03641 and EPA R-833007. The sample experiment reported was done by David Maraldo as part of his Ph.D. work. References 1. Lavrik, N. V., Sepaniak, M. J., Datskos, P. G. Cantilever transducers as a platform for chemical and biological sensors. Review of Scientific Instruments 2004, 75 (6), 2229–2253 2. Rosi, N. L., Mirkin, C. A. Nanostructures in biodiagnostics. Chemical Reviews 2005, 105 (4), 1547–1562 3. Detzel, A. J., Campbell, G. A., Mutharasan, R. Rapid assessment of Escherichia coli by growth rate on piezoelectric-excited millimeter-sized cantilever (PEMC) sensors. Sensors and Actuators B: Chemical 2006, 117 (1), 58–64 4. Sader, J. E. Frequency Response of Cantilever Beams Immersed in viscous fluids with applications to the atomic force microscope. Journal of Applied Physics 1998, 84 (1), 64–76
5. Campbell, G. A., Mutharasan, R. Detection of pathogen Escherichia coli O157:H7 using self-excited PZT-glass microcantilevers. Biosensors and Bioelectronics 2005, 21 (3), 462–473 6. Campbell, G. A., Mutharasan, R. Escherichia coli O157: H7 detection limit of millimetersized PZT cantilever sensors is 700 cells/mL. Analytical Sciences 2005, 21 (4), 355–357 7. Campbell, G. A., Mutharasan, R. PEMC sensor’s mass change sensitivity is 20 pg/ Hz under liquid immersion. Biosensors and Bioelectronics 2006, 22 (1), 35–41 8. Campbell, G. A., Mutharasan, R. Detection of Bacillus anthracis spores and a model protein using PEMC sensors in a flow cell at
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Mutharasan 1 mL/min. Biosensors and Bioelectronics 2006, 22 (1), 78–85 Campbell, G. A., Mutharasan, R. Method of measuring Bacillus anthracis spores in the presence of copious amounts of Bacillus thuringiensis and Bacillus cereus. Analytical Chemistry 2007, 79 (3), 1145–1152 Campbell, G. A., Mutharasan, R. A method of measuring Escherichia coli O157:H7 at 1 cell/mL in 1 liter sample using antibody functionalized piezoelectric-excited millimeter-sized cantilever sensor. Environmental Science and Technology 2007, 41 (5), 1668–1674 Campbell, G. A., Uknalis, J., Tu, S.-I., Mutharasan, R. Detection of Escherichia coli O157:H7 in ground beef samples using piezoelectric excited millimeter-sized cantilever (PEMC) sensors. Biosensors and Bioelectronics 2007, 22 (7), 1296–1302 Maraldo, D., Mutharasan, R. 10-minute assay for detecting Escherichia coli O157:H7 in ground beef samples using piezoelectricallyexcited millimeter-sized cantilever (PEMC) sensors. Journal of Food Protection 2007, 70 (7), 1670–1677 Campbell, G. A., Medina, M. B., Mutharasan, R. Detection of Staphylococcus enterotoxin B at picogram levels using piezoelectric-excited millimeter-sized cantilever sensors. Sensors and Actuators B: Chemical 2007, 126 (2), 354–360 Maraldo, D., Mutharasan, R. Detection and confirmation of Staphylococcal enterotoxin B in apple juice and milk using piezoelectricexcited millimeter-sized cantilever (PEMC) sensors at 2.5 femtograms/mL. Analytical Chemistry 2007, 70 (7), 1670–1677, DOI:10.1021/ac070589l Campbell, G. A., Mutharasan, R. Sensing of liquid level at micron resolution using
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self-excited millimeter-sized PZT-cantilever. Sensors and Actuators A: Physical 2005, 122 (2), 326–334 Rijal, K., Mutharasan, R. Piezoelectric-excited millimeter-sized cantilever sensors detect density differences of a few micrograms/mL in liquid medium. Sensors and Actuators B: Chemical 2007, 121 (1), 237–244 Wilson, T. L., Campbell, G. A., Mutharasan, R. Viscosity and density values from excitation level response of piezoelectric-excited cantilever sensors. Sensors and Actuators A: Physical 2007, 138 (1), 44–51 Campbell, G. A., Mutharasan, R. Monitoring of the self-assembled monolayer of 1-hexadecanethiol on a gold surface at nanomolar concentration using a piezo-excited millimeter-sized cantilever sensor. Langmuir 2005, 21 (25), 11568–11573 Campbell, G. A., Mutharasan, R. Use of piezoelectric-excited millimeter-sized cantilever sensors to measure albumin interaction with self-assembled monolayers of alkanethiols having different functional headgroups. Analytical Chemistry 2006, 78 (7), 2328–2334 Rijal, K., Mutharasan, R. A method for measuring self-assembly of alkanethiols on gold at femtomolar concentrations. Langmuir 2007, 23 (12), 6856–6863 Maraldo, D., Rijal, K., Campbell, G., Mutharasan, R. Method for label-free detection of femtogram quantities of biologics in flowing liquid samples. Analytical Chemistry 2007, 79 (7), 2762–2770 Hermanson, G. T. Bioconjugate Technique; Elsevier, San Diego, CA, 1996 Maraldo, D., Mutharasan, R. Optimization of antibody immobilization for sensing using piezoelectrically excited-millimeter-sized cantilever (PEMC) sensors. Sensors and Actuators B: Chemical 2007, 123 (1), 474–479
Chapter 6 Preparation of Screen-Printed Electrochemical Immunosensors for Estradiol, and Their Application in Biological Fluids Roy M. Pemberton and John P. Hart Summary The method of fabrication of a prototype electrochemical immunosensor for estradiol (E2) is described. Methodologies are also given for colorimetric assays, which can be used to verify and optimize reagent performance, prior to their use in the electrochemical immunoassay: these include an E2 ELISA and a colorimetric assay performed on the immunosensor surface. The electrochemical immunosensor system uses screen-printed carbon electrodes (SPCEs) upon which antibody against E2 is immobilized. Antibodies (rabbit anti-mouse IgG, then monoclonal mouse anti-E2) are immobilized by passive adsorption onto the working electrode surface. A competitive immunoassay is then performed using an alkalinephosphatase-labeled E2 conjugate. Electrochemical measurements are performed using differential pulse voltammetry (DPV) to detect the production of 1-naphthol from 1-naphthyl phosphate. The calibration plot of DPV peak current vs. E2 concentration shows a measurable range of 25–500 pg/mL with a detection limit of 50 pg/mL. The immunosensor can be applied to the determination of E2 in spiked serum, following an extraction step with diethyl ether. Key words: Screen-printed carbon electrode, Estradiol, Immunosensor, Serum.
1. Introduction The accurate and sensitive determination of estradiol (E2) concentration in biological fluids is desirable for gynecological investigations, fertility treatment assessment, and potentially for ovulation prediction. Analysis of bovine serum for the presence of estradiol is also required in the agricultural industry to monitor food-producing animals for the presence of illegally-administered hormone. Electrochemical ELISAs for E2 have been reported, in which Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_6
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competitive immunoassays conducted in microwells are followed by electrochemical measurement of an electrochemically-active product, 3,3´,5,5´-tetramethylbenzidine (TMB), at a glassy carbon electrode by flow-injection. This approach has been applied to bovine serum (1) and also to urban wastewater (2) determinations of E2. To facilitate the possibility of rapid analysis, the integration of the ELISA-type immunoassay with an electrochemical transducer measurement step can be achieved in an electrochemical biosensor format. The resulting biosensors can be disposable because of the low cost of the materials used and represent an economical means of sample analysis, particularly when based on the use of screen-printed carbon electrodes (SPCEs). In previous studies from this group, SPCEs have been tailored with surface-adsorbed specific antibodies and applied to the determination of progesterone (3, 4) or E2 (5) in competitive immunoassays employing alkaline phosphatase-labeled steroid conjugates. Other workers have also described the development of a disposable electrochemical immunosensor for 17β-estradiol using SPCEs (6). In all of these biosensor studies, the signal for these assays was obtained from the electrochemical oxidation of 1-naphthol, generated enzymatically from a 1-naphthyl phosphate substrate solution. The following sections outline the methodology and experimental procedures used in our laboratory to fabricate and test screen-printed carbon immunosensors for E2. Also included are protocols for an ELISA method and an SPCE-based colorimetric assay, which can be used to test the immunoassay reagents prior to fabricating and electrochemical testing of the immunosensors. The results of applying the immunosensors using differential pulse voltammetric (DPV) measurement in batch mode for determination of E2 in serum extracts are given as an example, as described previously (5). We have observed in our laboratory that antibody-electrodes coated in this way with anti-progesterone antibody may be used in a continuous-flow system for progesterone (4); we have also observed that the E2 immunosensors described below remain stable in a continuous-flow thin-layer cell format for at least 2 h (Pemberton et al., in preparation). The E2 immunosensors described below are therefore suitable both for batch analysis, as detailed in the following sections, and also for future applications in continuous-flow format.
2. Materials 2.1. Preparation of Screen-Printed Carbon Electrodes
1. Carbon screen-printing ink, formulation code D14 (Gwent Electronic Materials Ltd, Mamhilad, Gwent, UK).
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2. Ag/AgCl screen-printing ink, formulation code C61003P7 (Gwent Electronic Materials Ltd, Mamhilad, Gwent, UK). 3. Screen-printable dielectric ink (Gwent Electronic Materials Ltd, Mamhilad, Gwent, UK). 4. Valox™ thermoplastic polyester sheet (0.5 mm thick). 5. Screen-printing machine, e.g., DEK 248 (DEK Printing Machines Ltd., Dorset, UK). 2.2. ELISA for Estradiol
1. Medium-protein-binding 96-well ELISA plates (Greiner Labortecknik Ltd., Glos, UK). 2. Carbonate coating buffer (CB) comprising 0.015 M Na2CO3 and 0.035 M NaHCO3; final pH = 9.6. Prepare fresh in deionised water (dH2O) (see Note 1). 3. Rabbit anti-mouse IgG (rIgG) antibody, the purified fraction of antiserum (Sigma-Aldrich product no. M7023), containing 2.3 mg specific IgG per 10.0 mg of protein per milliliter. Aliquot and store at −20°C. 4. Mouse anti-estradiol monoclonal antibody (mAb), clone 2F9 (IgG) (Biogenesis Ltd., Poole, UK). Aliquot and store at −20°C at a concentration of 200 μg/mL in the presence of 0.1% NaN3. 5. Phosphate buffered saline (PBS), 0.15 M. Dissolve 8.00 g NaCl, 0.2 g KCl, 1.15 g Na2HPO4, and 0.20 g KH2PO4 in 1,000 mL dH2O. Adjust pH to 7.2 with HCl if necessary. Dispense in convenient volumes and sterilize by autoclaving. Store at room temperature (RT). 6. 17 β-Estradiol (E2) standard solutions, prepared by dissolving E2 (Cat: E-8875, Sigma-Aldrich Co. Ltd., UK) in ethanol at 1 mg/mL, followed by dilution in PBS to give a series of standards over the concentration range 0–500 pg/mL. 7. Alkaline-phosphatase–E2 (ALP–E2) conjugate (Biogenesis Ltd.); dilute 1/20 in PBS/0.1% BSA and store at −20°C. 8. Wash buffer (PBS/T): PBS containing 0.05% v/v Tween 20. 9. 0.2 M Tris-buffer, pH 9.9, prepared from SigmaFAST™ tablets. 10. Para-nitrophenyl phosphate (pNPP) tablets (SigmaFAST™). 11. Polypropylene wash bottles (500 mL) for PBS, PBS/T wash buffer, H2O, and Tris buffer reagent washes. 12. Adjustable pipette (e.g., Nichipet EX, Nichiryo, Japan), range from 20 to 200 μL. 13. Multi(8)-channel adjustable pipette (e.g., Nichipet 7000, Nichiryo, Japan), range from 50 to 300 μL. 14. Plate reader capable of measuring absorbance at 405 nm.
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2.3. Preparation of Immunosensors
• Screen-printed carbon working electrodes as described in Subheading 3.1. • Microfuge tubes (1.5 mL, polypropylene) (Cat: 616201, Greiner Bio-One Ltd., UK). • Double-sided Office Tape (25 mm width; Niceday, UK). • Cork borer (4 mm-diameter). • Carbonate coating buffer (CB) comprising 0.015 M Na2CO3 and 0.035 M NaHCO3; final pH = 9.6. Prepare fresh. • Rabbit anti-mouse IgG (rIgG) antibody, the purified fraction of antiserum (Sigma-Aldrich product no. M7023), containing 2.3 mg specific IgG per 10.0 mg of protein per milliliter. Aliquot and store at −20°C. • Polypropylene wash bottle (500 mL) containing PBS. • Mouse anti-estradiol monoclonal antibody (mAb), clone 2F9 (IgG) (Biogenesis Ltd., Poole, UK). Aliquot and store at −20°C at a concentration of 200 μg/mL in the presence of 0.1% NaN3. • Adjustable pipette (e.g., Nichipet EX, Nichiryo, Japan), range from 20 to 200 μL. • Adjustable PS-100 micropipetter (Barky Instruments, Folkestone, UK), range from 1 to 10 μL.
2.4. Colorimetric Immunoassay for E2 on SPCEs
• Screen-printed immunosensors, with added well, prepared as described in Subheading 3.3. • 17 β-Estradiol (E2) standard solutions in PBS (see Subheading 2.2, item 6). • Alkaline-phosphatase–E2 (ALP–E2) conjugate (see Subheading 2.2, item 7). • Wash buffer (PBS/T): PBS containing 0.05% v/v Tween 20. • 0.2 M Tris-buffer, pH 9.9, prepared from SigmaFAST™ tablets. • Para-nitrophenyl phosphate (pNPP) tablets (SigmaFAST™). • Adjustable pipette (e.g., Nichipet EX, Nichiryo, Japan), range from 20 to 200 μL. • Plate reader capable of measuring absorbance at 405 nm.
2.5. Electrochemical Immunoassay for E2 on SPCEs
1. Screen-printed immunosensors, with added well, prepared as described in Subheading 3.3. 2. 17 β-Estradiol (E2) standard solutions in PBS (see Subheading 2.2, item 6). 3. Alkaline-phosphatase–E2 (ALP–E2) conjugate (see Subheading 2.2, item 7). 4. Wash buffer (PBS/T): PBS containing 0.05% v/v Tween 20.
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5. 0.2 M Tris-buffer, pH 9.9, prepared from SigmaFAST™ tablets. 6. Diethanolamine-HCl buffer (DB), comprising 0.1 M diethanolamine (D-8885, Sigma-Aldrich Co. Ltd.), adjusted to a final pH of 9.85 with 1 M HCl. 7. A 5 mM solution of 1-naphthyl phosphate (monosodium salt; CAS 81012–89–7; Acros Organics, Fisher Scientific UK) in DB containing 10 mM MgCl2. 8. Screen-printed 2 mm-wide Ag/AgCl pseudoreference/ counter electrodes printed on PVC (Gwent Electronic Materials Ltd.). 9. Adjustable pipette (e.g., Nichipet EX, Nichiryo, Japan), range from 20 to 200 μL. 10. A computer-controlled potentiostat (e.g., μAutolab Type II; Eco Chemie B.V., Netherlands, supplied by Windsor Scientific Ltd., UK). 2.6. Determination of E2 in Serum
1. Human male serum (Biosera UK). 2. Diethyl ether, stabilized with copper gauze (CAS 60–29–7; Fisher Scientific UK). 3. Microfuge tubes (1.5 mL, polypropylene) (Cat: 616201, Greiner Bio-One Ltd., UK). 4. Adjustable pipette (e.g., Nichipet EX, Nichiryo, Japan), range from 1,000 to 5,000 μL 5. Whirlimix (Jencons Scientific Ltd., UK). 6. Beckman GS15R benchtop centrifuge. 7. Nitrogen (N2) (Oxygen-free; BOC, Manchester, UK) cylinder and line for solvent evaporation.
3. Methods The selectivity of immunosensors for steroid analytes is achieved with the use of appropriately selected monoclonal antibodies. The carbon working electrode provides a suitable surface for passive adsorption of proteins, and can therefore be tailored with an appropriate antibody, so that it will act as an immunoactive surface upon which an immunoaffinity assay can be performed; an electrochemical signal can then be generated by monitoring the production of an electroactive species at the underlying electrode surface. We and other workers have found that to retain maximum monoclonal antibody activity, it is desirable to use a primary antibody (rabbit IgG), which serves both to capture (e.g., from a culture medium) and to orientate the mAb. Hence this approach
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is used in the methods outlined in the following sections. No stabilizing materials are included in these methods, and we find that the immunosensors, once prepared, should be kept moist, like coated ELISA plates. They are then stable for up to 3 days if kept at 4°C. 3.1. Preparation of Screen-Printed Carbon Electrodes
The general method for preparing screen-printed electrodes is described in detail elsewhere (7–9) and illustrated diagrammatically in Fig. 1. Briefly, a screen-printing stencil is prepared using a pretensioned polyester mesh, which is coated with a layer of photo-sensitive emulsion. Photographic positives of the desired electrode template(s) are placed over the mesh, which is then exposed to strong ultraviolet (UV) light. In exposed areas, the UV light fixes the emulsion to the mesh, whereas emulsion in the covered areas remains unfixed and can be washed away to leave a negative image of the desired electrode shape(s). To print the working electrode shown in Fig. 2, the inert Valox polyester support, cut into suitably-sized card to accommodate the print, is placed into the printing machine and aligned under the stencil. The screen-printing (D14) ink suspension is placed onto the stencil at one end of the electrode image. The squeegee, held at an angle of 60° to the stencil, is then moved forwards and backwards over the image, thus forcing ink into the stencil, leaving a print on the card. The polyester card is then removed and dried at 65°C. The carbon working electrode is laid down as a 2 mm-wide conductive track, ending in a 6 × 6 mm square (Fig. 2A). Once the working electrode print has been printed onto the
Fig. 1. Diagram to show preparation of screen-printed electrodes. (A) Polyester card support, (B) screen-stencil, (C) ink suspension, and (D) squeegee (this is a variation after Fig. 3 in ref. 7).
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Fig. 2. Screen-printed electrode design: diagram of (A) working electrode and (B) working area with dielectric layer; (C) photograph of screen-printed working electrode/ dielectric with scale in cm.
required number of cards, the screen-stencil is changed for one bearing the shape of the dielectric print, and the printing process is repeated using the dielectric ink formulation. The final working area is thus defined by over-printing, using the second screen, a square dielectric layer, leaving a central hole of diameter 3 mm (Fig. 2B, C). The resulting working electrode has a geometric area of 7.1 mm2. To perform the required electrochemical measurements described below, counter/pseudoreference electrodes are required;
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these are also screen-printed from Ag/AgCl ink, using a separate screen, onto a PVC substrate. These are in the form of 2 mmwide strips (20 mm long), which can be cut out from the card as required. Screen-printing of the described working and reference electrodes, and dielectric layer was conducted by staff at Gwent Electronic Materials Ltd; screen-printed working and reference electrodes are available from Gwent Electronic Materials Ltd. 3.2. ELISA for Estradiol
1. Coat ELISA plate wells overnight at 4°C with 100 μL of the appropriate dilution of rIgG in CB (see Note 2). Use adjustable (micro)pipette(s) to prepare antibody dilutions and to coat microwells. 2. Rinse wells with PBS using wash bottle. 3. Add 100 μL of appropriate dilution of mAb 2F9 (see Note 2) in PBS per well for 2 h at RT. 4. Rinse wells with PBS, then add E2 standard solutions/samples in 50 μL volumes for 2 h at RT. 5. Add 100 μL per well of ALP–E2 conjugate at 1/50,000 (see Note 2) in PBS. 6. Incubate, RT, for a further 30 min. 7. Wash wells five times with PBS/T, twice with dH2O and once with Tris. 8. Add 150 μL per well of pNPP (1 mg/mL) in Tris. 9. Read absorbance at 405 nm after 10–60 min. 10. Examples of calibration plots are shown in Fig. 3.
3.3. Preparation of Immunosensors
1. For each immunosensor, create a reagent well by cutting the lid and the bottom off a 5 mm-diameter polypropylene microfuge tube to leave a 10 mm-long open tube. 2. For each immunosensor, cut out a 2 × 2 cm square of double-sided tape. 3. Use the cork borer to punch a hole in the center of the tape, then, having removed the covering layer from one side, attach the tape to the dielectric layer, leaving the central working electrode area exposed (Fig. 4). 4. Remove the upper covering layer from the tape and press the open tube down onto the area surrounding the working electrode, to create a reagent well on the SPCE (Fig. 4). 5. Using an adjustable pipette to deliver the required volume into the reagent well, coat working electrode of SPCE-withadded-well (Fig. 4) with rIgG (1/100 dilution) and then mAb 2F9 (1/200 dilution) according to steps 1–3, Subheading 3.2.
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Fig. 4. SPCE with added reagent well. (Reproduced from ref. 5 with permission from Elsevier Science).
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6. Rinse with PBS using wash bottle. 7. Use immediately or store in a moist box (80% humidity) at 4°C for up to 3 days. 3.4. Colorimetric Immunoassay for E2 on SPCEs
1. Add 50 μL volumes of E2 standard solutions/samples into each sensor well for 2 h at RT. 2. Add 100 μL ALP–E2 conjugate (1/50,000 in PBS) per well and incubate for a further 30 min, RT. 3. Tip out contents and using wash bottle(s) wash wells five times with PBS/T, once with dH2O, and once with Tris. 4. Shake the SPCEs/wells dry. 5. Add 150 μL pNPP (1 mg/mL) in Tris per sensor well. 6. Allow color development for 30–120 min. 7. Aspirate 100 μL volume from each sensor well into a corresponding ELISA plate well and read the absorbance at 405 nm. 8. An example colorimetric calibration plot is shown in Fig. 5. 1. Perform steps 1–4, Subheading 3.4, on all sensors in any experiment.
SPCE color assay (abs @ 405nm)
2. Taking one sensor at a time, lower a screen-printed Ag/AgCl counter/reference electrode strip into the sensor well and connect this and the working electrode to the potentiostat.
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y = 0.1081x + 0.0686 2 R = 0.9841
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[E2] (pg/ml) Fig. 5. Calibration plot for E2 standard solutions in PBS, obtained by colorimetric assay using SPCE immunosensors. Incubation time in substrate = 120 min. Inset shows correlation between this assay and the ELISA method. (Reproduced from ref. 5 with permission from Elsevier Science).
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The arrangement of the biosensor and associated instrumentation is shown in Fig. 6A , B. While recording the time, add 150 μL of 1-naphthyl phosphate solution per sensor well and incubate at RT. 3. After 10 min, use the potentiostat to apply a differential pulse waveform (step potential = 2 mV, pulse amplitude = 20 mV, step time = 0.2 s, pulse time = 0.03 s) and scan from −0.1 to + 0.7 V. 4. Repeat steps 2 and 3 for all sensors in the experiment. 5. For each sensor, record the peak height of the resulting anodic wave (see Note 3) and plot against E2 concentration. 6. Examples of the resulting DPV voltammograms and calibration plot are shown in Fig. 7.
Fig. 6. Diagrammatic (A) and photographic (B) representation of biosensor/instrumentation required for electrochemical E2 assay. (1) card bearing working electrode and reagent well, (2) Ag/AgCl counter/reference electrode, (3) substrate solution in well, (4) gold connectors, (5) potentiostat, (6) computer, (7) block to support working electrode, (8) substrate stock solution, (9) micropipettor, (10) timer.
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Fig. 7. Electrochemical immunosensor for E2 (A) Differential pulse voltammograms obtained for standard solutions of E2 in PBS over the concentration range 0–3,200 pg/mL. (B) Corresponding calibration plot. Each point represents the mean current value from two sensors. The working range is between 25 and 500 pg/mL, with a detection limit of around 50 pg/ mL. (Reproduced from ref. 5 with permission from Elsevier Science).
3.6. Determination of E2 in Serum
1. Add a 0.5 mL aliquot of serum sample to 1 mL diethyl ether in a 1.5 mL microfuge tube. 2. Whirlimix, 10 s. 3. Centrifuge at 1,000 × g for 20 min. 4. Transfer the supernatant to a suitable round-bottomed glass tube. 5. Repeat steps 1–4 (both supernatants in one tube). 6. Evaporate contents of glass tube under N2. 7. Resuspend the remaining pellet in 0.5 mL PBS. 8. Now perform the electrochemical immunoassay as in Subheading 3.5. 9. Use a PBS calibration plot (Fig. 7b) to obtain the E2 concentration (see Note 4).
4. Notes 1. Unless stated otherwise, all solutions should be prepared using deionised water having a resistivity of better than of
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1 MΩ cm. A Purite Still Plus water purification system (Purite Ltd., Oxon, UK) is a suitable source. 2. The rationale for optimising ELISA antibody and conjugate reagents is to titrate the mAb using doubling dilutions (1/10– 1/81920) in the presence of varying dilutions of the rIgG coating antibody (1/72–1/720) at several different ALP–E2 conjugate dilutions (1/2,000–1/16,000). The resulting plots (see ref. 5 for examples) are then used to select the optimum combination of reagents, which will give between 50 and 80% saturation on a calibration plot and an absorbance of at least 0.2 units. Adjustable pipettes are used to aliquot reagents into wells and doubling dilutions are best prepared in the ELISA plates using a multi(8)-channel adjustable pipette for speed of operation and to minimize errors in fluid handling. 3. The anodic DPV wave represents the oxidation of the enzyme product 1-naphthol. This occurs at a peak potential of around +125 mV vs. Ag/AgCl. Peak currents are measured by drawing a sloping baseline and measuring the vertical distance between this line and the peak. A coefficient of variation of between 13.0 and 15.6% was obtained for repeat measurements. Since the assay is competitive, the limit of detection is obtained by determining the concentration of E2, which will reliably result in a reduction in peak current compared with the mean value obtained for a blank standard containing no E2. This detection limit is obtained by calculating the mean-minus-3-times the SD of the mean for n = 4 replicate blank measurements and reading the corresponding E2 concentration from the graph. The absolute value of this mean peak current blank response and detection limit must be determined for each new batch of biosensors, since it is important to control for batch-to-batch variations. Using the instrumentation specified above, the signal/noise ratio for any given voltammogram is very high; for example, a typical baseline DPV current of 856 nA exhibits a peak-to-trough fluctuation of 3.4 nA, making current responses for each biosensor very easy to measure. 4. The diethyl ether extraction procedure is required to overcome the problems of (a) matrix effects due to, e.g., hormone-binding proteins within the serum, which may sequester E2 thereby distorting the measurable E2 concentration and (b) interfering electrochemical signals caused by the presence of serum components. Ideally, the E2 standard solutions in PBS, as well as the serum samples, should be subjected to the extraction procedure prior to electrochemical immunosensor analysis. However, in tests using serum spiked with 200 pg/mL E2, a mean recovery of 102% was obtained when the values were read against a calibration plot constructed using nonextracted E2 standards in PBS (5).
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Acknowledgments This work was supported by a grant from DEFRA.
References 1. Draisci, R., Volpe, G., Compagnone, D., Purificato, I., delli Quadri, F. & Palleschi, G. (2000). Development of an electrochemical ELISA for the screening of 17β-estradiol and application to bovine serum. Analyst 125, 1419–1423 2. Valentini, F., Compagnone, D., Gentile, A. & Palleschi, G. (2000). An electrochemical ELISA procedure for the screening of 17β-estradiol in urban wastewaters. Analyst 127, 1333–1337 3. Pemberton, R.M., Hart, J.P., Stoddard, P. & Foulkes, J.A. (1999). A comparison of 1-naphthyl phosphate and 4-aminophenyl phosphate as enzyme substrates for use with a screen-printed amperometric immunosensor for progesterone in cows’ milk. Biosens. Bioelectron. 14, 495–503 4. Pemberton, R.M., Hart, J.P. & Mottram, T.T. (2001). An electrochemical immunosensor for milk progesterone using a continuous flow system. Biosens. Bioelectron. 16, 715–723 5. Pemberton, R.M., Mottram, T.T. & Hart, J.P. (2005). Development of a screen-
6.
7.
8.
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printed carbon electrochemical immunosensor for picomolar concentrations of estradiol in human serum extracts. J. Biochem. Biophys. Methods 63, 201–212 Volpe, G., Fares, G., Quadri, F.D., Draisci, R., Ferretti, G., Marchiafava, C., Moscone, D. & Palleschi, G. (2006). A disposable immunosensor for detection of 17 beta-estradiol in non-extracted bovine serum. Anal. Chim. Acta. 572, 11–16 Hart, J.P. & Wring, J.P. (1991). Carbonbased electrodes and their application as electrochemical sensors for selected biomolecules. Anal. Proc. 28, 4–7 Wring, S.A. & Hart, J.P. (1992). Chemically modified, screen-printed carbon electrodes. Analyst 117, 1281–1285 Wring, S.A., Hart, J.P., Bracey, L. & Birch, B.J. (1990). Development of screen-printed carbon electrodes, chemically modified with cobalt–phthalocyanine, for electrochemical sensor applications. Anal. Chim. Acta. 231, 203–212
Chapter 7 Electrochemical DNA Biosensors: Protocols for Intercalator-Based Detection of Hybridization in Solution and at the Surface Kagan Kerman, Mun’delanji Vestergaard, and Eiichi Tamiya Summary An electrochemical DNA biosensor is a device that utilizes the inherent ability of two complementary strands of nucleic acids to form a double helix. The specificity of this reaction, namely hybridization, is used in the detection of target DNA sequences with a view toward developing point-of-care devices. Since the early 1990s, great progress has been made in this field, but there are still numerous challenges to overcome. This chapter describes the components of an electrochemical DNA biosensor for researchers new to the field, paying particular attention to intercalator-based DNA biosensors. We will use a well-defined electro-active DNA intercalator Hoechst 33258, as our running example. Two of the most classic DNA sensing methods: solution-based and surface-immobilized methods are discussed, along with guiding notes that would help identify and overcome possible problems in a typical electrochemical DNA biosensor experiment. Key words: Biosensor, Electrochemical, DNA, Intercalator, Hybridization, Hoechst 33258, Screenprinted electrode (SPE), Probe-modified electrode, Self-assembled DNA monolayer.
1. Introduction A biosensor is a device that combines a biological component (a recognition layer) and a physico-chemical detector component (a transducer). The transduction unit can be electrochemical, optical, piezoelectric, magnetic, or calorimetric (1). Two groups of recognition molecules form the majority of biosensors: affinitybased and catalytic-based biosensors. Affinity-based biosensors are used to bind molecular species of interest, irreversibly and noncatalytically. Examples include antibodies, nucleic acids, and Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_7
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hormone receptors. Catalytic-based sensors such as enzymes and microbiological cells recognize and bind a molecule of interest, followed by a catalyzed chemical conversion of that molecule to a product that is then detected. Electrochemical biosensors enable fast, simple, and low-cost detection, mainly because there is no need for expensive signal transduction equipment since they give electronic signals directly. They exploit both catalytic and noncatalytic-based molecules for the recognition layer. The most popular examples of these two types of molecular recognition elements are enzymes and DNA. In this chapter, we will confine our attention to DNA-based biosensors. DNA is particularly well-suited for biosensing applications, because there is a robust and specific interaction of the base pairs between complementary strands. DNA biosensors convert the Watson–Crick base-pair recognition event into a readable analytical signal (2–4). A DNA biosensor is mainly designed in such a way that a distinction can be made between single-stranded DNA (ssDNA), double-stranded DNA (dsDNA), and rarely three-stranded DNA (triplexes), providing the basis for electrochemical detection of DNA hybridization. In this chapter, we will restrict our discussions to the most common forms (dsDNA and ssDNA). There are basically four different analytical techniques for electrochemical detection of DNA hybridization: monitoring of (a) electrochemical oxidation/ reduction signals of metal nanoparticle-tagged oligonucleotides, (b) oxidation peak current of the electro-active DNA bases such as guanine and/or adenine, (c) enzymatically amplified electrochemical signal of a substrate in the presence of the enzyme-tagged hybrid, and (d) electrochemical signal of a label, which selectively binds with dsDNA/ssDNA (2, 5). In electrochemistry, DNA labeling has been exploited widely, from indirect detection platforms that use chemical mediators such as polypyridyl complexes of Ru(II) and Os(II), through use of redox-active reporter molecules (ferrocene, gold nanoparticles, semiconductor nanostructures (quantum dots: CdS, ZnS, PbS), to DNA-mediated charge transport mechanisms that utilize redox-active electrostatic (Ru(NH3)63+) and intercalative (Co(phen)33+) probe molecules (5–12). The interaction of DNA and a ligand, which can bind to DNA to form a larger complex, plays a major role in biological processes such as gene transcription and DNA replication. Intensive research on natural and synthetic ligands has revealed intercalative or nonintercalative mechanisms (13–16). A large class of molecules intercalates in the space between two adjacent base pairs of dsDNA. These molecules mostly have a polycyclic aromatic ring with a planar structure, and are often known as nucleic acid stains. Electro-active DNA intercalators include ethidium bromide, methylene blue, and benzo[a]pyrene (17, 18). Figure 1 displays the chemical drawings of the representative intercalators
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Fig. 1. Chemical drawings of the most-commonly used intercalators in the development of electrochemical DNA biosensors; (1) Cobalt(II)-Tris-(1,10-phenanthroline), (2) Ruthenium(II)-Tris-(1,10-bipyridine), (3) Methylene blue, (4) Meldola blue, (5) Ethidium bromide.
that have been commonly applied in electrochemical DNA biosensors. The fitting of an intercalator between DNA base pairs requires a separation of over 0.3 nm, inducing some local structural changes to the DNA strand such as unwinding of the double helix and lengthening of the DNA strand (18). The small molecules, which noncovalently interact with DNA, are stabilized in binding to DNA through a series of weak interactions. They include π-stacking interactions associated with intercalation of aromatic heterocyclic groups between the base pairs, and hydrogen-bonding and van der Waals interactions of functionalities bound along the groove of the DNA helix. It would be valuable to understand quantitatively the contributions from the different modes to the stabilization of the bound complex at a certain DNA site. These structural modifications lead to the functional changes in DNA, often to the inhibition of transcription and replication processes. Therefore, the intercalators are also known as potent mutagens.
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Fig. 2. Chemical drawing of Hoechst 33258.
In this chapter, we will use a DNA intercalator, Hoechst 33258 (2′-(4-hydroxyphenyl)-5-(4-methyl-1-piperazinyl)-2,5′-bi(1Hbenzimidazole), Fig. 2) as our running example. Hoechst 33258 has an electrochemical oxidation potential at ∼0.6 V (vs. Ag/ AgCl) (19–22). Its interaction with DNA results in the formation of aggregates, easily detected using atomic force microscopy (AFM) (21, 22). Excess concentration of Hoechst 33258 induces DNA condensation. DNA can be analyzed either in solution and/or as an adsorbent on an electrode surface. We will provide protocols for both analytical situations. For DNA detection process on the surface, basically, a singlestranded oligonucleotide (probe) is immobilized onto a transducer surface to recognize its complementary (target) DNA sequence, via hybridization, to form a DNA duplex (hybrid) on the electrode surface. This event is then converted into an analytical signal by a transducer. The immobilization of DNA probes on or near the electrode surface provides a suitable platform for highly integrated systems. However, an elegant surface chemistry is required for the preparation of probe modified electrodes. For electrostatic immobilization of nucleic acids on the surface of a carbon electrode, an electrochemical pretreatment of the surface is required. For example, a carbon paste electrode (CPE) is activated by applying +1.7 V (vs. Ag/AgCl) for 1 min in 0.05 M phosphate buffer solution (pH 7.4) without stirring (see Note 1). Then, DNA is immobilized on the pretreated CPE by applying a potential of +0.5 V for 5 min in DNA containing 20 mM Tris–HCl (pH 7.0) with 200 rpm stirring (see Note 2). The positively-charged electrode surface attracts the negativelycharged phosphate backbone of DNA and loads them electrostatically onto the surface (Fig. 3 A, B). The electrode is then rinsed with a blank buffer solution (20 mM Tris–HCl) for 10 s. A similar procedure as above can be used for the immobilization of oligonucleotides on the surface of a carbon electrode. The formation of self-assembled monolayer (SAM) of the thiolated nucleic acids on gold provides a very useful platform for the development of DNA sensors (10, 11). However, researchers should pay particular attention to the pretreatment and cleaning of gold electrodes to avoid defects in the SAM. First, the gold electrodes must be sequentially polished using 6 and 1-μm
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Fig. 3. (A) The electrochemical detection of hybridization on carbon electrode surface. Probe oligonucleotides are immobilized on the carbon surface with electrostatic attraction (A1), noncomplementary oligonucleotides are incubated with the probe-modified electrode (A2), no hybridization takes place between these two strands (A3), Hoechst 33258 is incubated with the probe-modified-electrode (A4), since the redox label cannot intercalate with the single-strand probe oligonucleotides on the surface, a small linear sweep voltammetric (LSV) current response is obtained (A5). The current intensity at the peak potential (∼0.6 V) is recorded relative to the background response of the blank buffer solution (dashed grey line). (B) The electrochemical detection of hybridization on a carbon electrode surface. Probe oligonucleotides are negatively charged due to their phosphate backbone and are attracted onto the positively-charged carbon electrode surface (B1), target oligonucleotides are incubated with the probe-modified electrode (B2), hybridization takes place between the complementary strands resulting in the formation of a hybrid-modified electrode (B3), the intercalator, Hoechst 33258, is incubated with the hybrid-modified-electrode (B4), since Hoechst 33258 intercalates into the doublestranded hybrids on the surface, a high LSV current response is obtained (B5). The current intensity at the peak potential (∼0.6 V) is recorded relative to the background response of the blank buffer solution (dashed grey line).
diamond pastes and 0.05-μm alumina on a polishing pad. After thorough rinsing, the electrodes are ultrasonically cleaned in distilled water for 1 min. The electrodes are then subjected to an electrochemical pretreatment step: cyclic voltammetry is applied in a 0.5 M H2SO4 solution from 0 to 1.7 V (vs. Ag/AgCl), until a
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stable redox wave of Au oxidation (∼1.2 V) and reduction (∼0.8 V) is observed. Afterwards, the electrodes are rinsed with distilled water and immersed in the solution of the thiolated oligonucleotide for ∼12 h at 4°C (see Note 3). Under these conditions, the sulfide-containing oligonucleotides are adsorbed to the gold surface involving a stable Au-S linkage. The electrodes are then washed with distilled water or preferably, a solution containing detergents (sodium dodecyl sulfate, Tween 20, PEG derivatives, etc.) to remove any nonspecifically adsorbed oligonucleotides. As for the hybridization reactions on the surface, the electrode modified with the DNA probe is immersed in a hybridization buffer (see Note 4) containing the target oligonucleotide, which is complementary to the probe, for 1 h with moderate shaking or stirring (see Note 5), forming hybrids on the electrode surface. Afterwards, the intercalator (Hoechst 33258) is exposed to the electrode surface and incubated with the hybrids on the surface for 5 min. Following the incubation step, the electrode is washed to remove the excess intercalator, and an electrochemical measurement is taken using a suitable technique, such as linear sweep voltammetry (LSV), cyclic voltammetry (CV), and/or differential pulse voltammetry (DPV). The difference between the peak current responses obtained from the intercalator after its interaction with dsDNA and ssDNA provides the basis of this electrochemical detection system (Fig. 3A, B). If the probes are not immobilized on the surface, the preparation procedure becomes greatly simplified. However, the need for a suitable reaction chamber appears to be a drawback to the development of this system. A recent design, called a DNA stick, has been used in our laboratory for detecting DNA hybridization in solution with Hoechst 33258. Hoechst 33258 aggregates with the PCR amplified DNA in solution. These changes in the anodic current signal of Hoechst 33258 at ∼0.6 V are monitored in the presence and absence of solution-phase DNA using a bare electrode. The mass transfer to the electrode surface is reduced with formation of the DNA-Hoechst 33258 aggregates, resulting in the reduced peak current intensity. Free Hoechst 33258 molecules can be transferred to the electrode surface more rapidly, and result in a high peak current intensity. Similar to the on-surface detection principle, the difference between the peak current signals obtained from the aggregated and free forms of Hoechst 33258 provides the basis of this detection system. The aggregation of DNA–Hoechst 33258 complex leads to a decrease in the voltammetric signal in proportion to the quantity of PCRamplified DNA (19–21). Over the past decade, great progress has been made in the field of electrochemical DNA biosensors, but there are still numerous challenges to overcome. Here we describe the components of electrochemical DNA biosensors and the important
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steps in their application for the detection of hybridization, using an intercalator, for researchers new to the field.
2. Materials 2.1. Screen-Printed Electrode
Planar screen-printed electrodes (SPEs) were obtained from Biodevice Technology, Co. Japan. They are used once and discarded after each measurement. SPE consists of a carbon working electrode, a carbon counter electrode, and Ag/AgCl reference electrode (Fig. 4 A). Total length of the SPE is 11 mm, and the geometric working area is 2.64 mm2. The SPEs should be kept in their plastic packages provided by the supplier, Biodevice Technology, Co. at room temperate, and cut away from their sheet when required. The SPEs are stable, unless they are damaged or used. Instead of the SPEs, other commercially available electrodes can be used, for example a glassy carbon electrode (3.0 mm i.d.,
Fig. 4. (A) Photograph of screen-printed gold and carbon electrode with the three-electrode system including a carbonbased counter electrode and Ag/AgCl-based inner reference electrode. (B) Photograph of a typical electrochemical setup with a computer-controlled potentiostat connected to a screen-printed electrode (SPE).
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product # MF-2012, Bioanalytical System, Inc.), a gold electrode (1.6 mm i.d., product # MF-2014, Bioanalytical System, Inc.), and a carbon paste electrode (3.0 mm i.d., Teflon electrode body product # MF-2010, Bioanalytical System, Inc.). Ag/AgCl-based reference electrode can be used (Product # MF-2052, RE-5B, Bioanalytical System, Inc.). The auxiliary (counter) electrode can be made of platinum wire (6 cm, product # MW-4130, Bioanalytical System, Inc.). We would recommend VC-2 voltammetry cell with Teflon electrode holder (Product # MF-1052, Bioanalytical System, Inc.) in connection with the conventional electrochemical set-up. 2.2. Electrochemistry
Electrochemical measurements were performed using a potentiostat, for example Autolab PGSTAT 12 electrochemical analysis system (Eco Chemie, The Netherlands) in connection with General Purpose Electrochemical System (GPES) software, or CH Instruments 660 Electrochemical Workstation (Austin, TX). The electrochemical set-up consisted of a SPE system is as shown in Fig. 4b.
2.3. Spectrophotometry
UV-VIS DNA-protein analyzer (Shimadzu, Japan).
2.4. Hoechst 33258
Hoechst 33258 (2′-(4-hydroxyphenyl)-5-(4-methyl-1-piperazinyl) -2,5′-bi (1H-benzimidazole) trihydrochloride, 2-[2-(4-Hydroxyphenyl)-6-benzimidazoyl]-6-(1-methyl-4-piperazyl) benzimidazole trihydrochloride, H33258) was purchased from Fluka BioChemika (Product # 14530). A stock solution is prepared by directly dissolving it in high purity water (r = 18.3 MΩ cm, Millipore, Bedford, MA, USA) and kept at 2–8°C in the dark (see Note 6). Hoechst 33258 solution (100 μM) is prepared in 50 mM phosphate buffer solution (PBS) including 100 mM NaCl (Sigma-Aldrich, Product # 71376) (pH 7.4). PBS is prepared by mixing 0.04 mol K2HPO4 (SigmaAldrich, Product # 60353) and 0.01 mol KH2PO4 (Sigma-Aldrich, Product # 60218).
2.5. Deoxyribonucleic Acid
Deoxyribonucleic acid sodium salt from salmon testes (fsDNA) was purchased from Sigma (Product # D1626). Solution of natural fish sperm DNA prepared in MilliQ water gives UV absorbance ratios of ∼1.8–1.9 at 260 and 280 nm (A260/A280), respectively, which indicates that DNA is sufficiently free of protein. Stock fsDNA is stored at 0–4°C and should be used within three days of preparation (see Notes 7–9).
2.6. Preparation of the Probe Solution
Incubate the probe/dithiothreitol (DTT; 0.1 M DTT in water) solution (purchased from Sigma-Aldrich, Product # 43816) at room temperature for 30 min, followed by removal of the DTT using a standard ethanol precipitation procedure. Determine the concentration of DNA solution and adjust it to 25 μM (make the
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final buffer composition as 50 mM PBS at pH 7.4), and use it as the probe solution. 2.7. Preparation of Hybridization Buffer Solution 2.8. Preparation of Washing Solution
Hybridization buffer contains 20 mM Tris–HCl (Sigma-Aldrich, Product # T5941) and 500 mM NaCl (pH 7.0). Washing solution contains 20 mM Tris–HCl with 300 mM NaCl and 0.1% sodium dodecyl sulphate (Sigma-Aldrich, Product # 71725) (pH 7.0).
3. Methods 3.1. Detection in Solution
Mix fsDNA solution and H33258 at a final concentration of 1 μM in 50 mM PBS and pipet an aliquot (20 μL) of this mixture on the gold or carbon SPE surface. Measure immediately using linear sweep voltammetry (LSV) after an equilibration time of 10 s, with a sample interval of 1 mV, and a scan rate of 0.1 V/s from 0 to 1 V (Fig. 5). Alternatively, cyclic voltammetry (CV) can be applied with a scan range between 0 and 1 V at a sweep rate of 0.1 V/s. For differential pulse voltammetry (DPV) measurements, the potential is scanned from 0 to 0.90 V with a step potential of 4 mV, pulse amplitude of 50 mV, and a pulse period of 0.20 s at a scan rate of 10 mV/s. The current height is recorded at the peak potential (∼0.6 V) for analytical evaluation of the measurements. All measurements should be carried out at room temperature.
3.2. Detection at the Electrode Surface
Schematic representation of the hybridization detection using an electrochemical DNA biosensor using a SPE with a gold working electrode is shown in Fig 6A, B. 1. Probe immobilization. Pipet an aliquot (20 μL) of the probe solution onto the surface of gold SPE and spread it uniformly over the area of the three-electrode system. Incubate the SPE for about ∼12 h at room temperature (Fig. 6A1, B1) and then flush with copious amounts of water to remove any nonspecifically adsorbed DNA probes. At the end of this step, a probe-modified electrode is obtained. 2. Hybridization. Prepare the oligonucleotide (the target or noncomplementary) solutions of 20 μM in hybridization buffer. For each hybridization experiment, apply 20 μL of the sample to the probe-modified electrode surface (Fig. 6A2, B2). After 1 h of incubation at room temperature, rinse the electrode sequentially with the washing solution for 5 s and blank hybridization buffer for 5 s to remove any nonspecifically bound species. At the end of this washing step, a hybrid-modified
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Fig. 5. The electrochemical detection of hybridization in solution. (A) Probe oligonucleotides do not hybridize with the noncomplementary ones and thus, the addition of the intercalator, Hoechst 33258, does not result in the formation of an aggregate. In the presence of single-stranded oligonucleotides, Hoechst 33258 can diffuse to the electrode surface and cause the formation of a high current response; (B) Probe oligonucleotides hybridizes with the complementary target oligonucleotide. An aggregate of hybrid duplexes together with the intercalator forms and prevents the access of the intercalator to the electrode surface. A small current response indicates the aggregation of hybrid duplexes with Hoechst 33258. The current intensity at the peak potential (∼0.6 V) is recorded relative to the background response of the blank buffer solution (dashed grey line).
electrode is obtained (Fig. 6B3), but no hybridization would take place in the presence of noncomplementary oligonucleotides (Fig. 6A3). 3. Intercalator binding. Put an aliquot (20 μL) of the intercalator solution onto the SPE surface (Fig. 6A4, B4). Then, incubate for 5 min at room temperature. Wash off the excess intercalator with blank PBS. 4. Electrochemical measurement. Measure the anodic peak current of the bound intercalator using LSV (Fig. 6A5, B5) after an equilibration time of 10 s, with a sample interval of 1 mV, and a scan rate of 0.1 V/s from 0 to 1 V (see Note 10). Alternatively, CV or DPV can be applied for the measurement as described in Subheading 3.1. The current height is recorded at the peak potential (∼0.6 V) for analytical evaluation of the measurements. Fig. 6. (continued) incubated with the probe-modified electrode (B2), hybridization takes place between the complementary strands resulting in the formation of a hybrid-modified electrode (B3), Hoechst 33258, is incubated with the hybridmodified-electrode (B4), since Hoechst 33258 intercalates into the double-stranded hybrids on the surface, a high electrochemical current response is obtained (B5). The peak current intensity at the peak potential (∼0.6 V) is recorded relative to the background response of the blank buffer solution (dashed grey line).
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Fig. 6. (A) The electrochemical detection of hybridization on Au electrode surface. Probe oligonucleotides with 5′-thiol are incubated on the Au surface for 12 h. Strong covalent bond between thiols and Au surface results in the formation of the probe-modified electrode (A1), noncomplementary oligonucleotides are incubated with the probe-modified electrode (A2); however, no hybridization takes place between these two strands (A3), the redox label, Hoechst 33258, is incubated with the probe-modified-electrode (A4), since the redox label cannot intercalate with the single-strand probe oligonucleotides on the surface, a small electrochemical current response is obtained (A5). The current intensity at the peak potential (∼0.6 V) is recorded relative to the background response of the blank buffer solution (dashed grey line). (B) The electrochemical detection of hybridization on Au electrode surface. Probe oligonucleotides with 5′-thiol are incubated on the Au surface for ∼12 h for the preparation of the probe-modified electrode (B1), target oligonucleotides are
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4. Notes 1. The electrochemical pretreatment of a carbon paste electrode can be performed by applying high positive bias from +1.25 to +1.75 V for a short period of time. Time periods longer than 1 min are not usually recommended. As for glassy carbon electrodes, the application of a lower bias from 0.5 to 1 V for 1–3 min is recommended. 2. Electrostatic DNA immobilization on electrode surface may require longer accumulation time periods than 5 min. Adjustment of accumulation time periods according to the concentration of DNA sample and electrode surface is recommended. For example, the immobilization of a trace amount of DNA onto a carbon paste electrode with a large surface area would certainly require a long accumulation time, while stirring the DNA containing solution. 3. The formation of a SAM on gold electrode surface may take longer than 12 h due to the surface roughness and the concentration of the thiolated oligonucleotides. In certain cases, incubation periods of 5 days for the immobilization of 1 μM 23-mer oligonucleotides on 10 μm gold microelectrodes (i.d.) have been found as the optimum period. 4. Our recommended hybridization buffers are as follows. 30 mM sodium citrate including 300 mM NaCl (pH 7.0) and 20 mM Tris–HCl including 500 mM NaCl (pH 7.0). 5. The hybridization reaction may take longer than 1 h according to the surface area, the concentration of the oligonucleotides immobilized on the surface, and the concentration of the target strand in solution. For the detection of trace amounts of target DNA, a mass transport boost using a shaker or a stirrer is highly recommended. 6. Hoechst 33258 is a hazardous chemical and should be handled with care. During the experiments, personal preventive equipment should be worn with a breathing apparatus (a mask), safety goggles, and gloves to prevent contact with skin and eyes. If swallowed, mouth should be washed out with water. In case of skin contact, flush with copious amounts of water for at least 15 min and the contaminated clothing and shoes should be removed. 7. Sperm cells from salmon testes are a good source of non-mammalian DNA. The species of salmon used is Oncerhyncus keta. The %G-C content is ∼41.2%. The melting temperature (Tm) is 87.5°C in 0.015 sodium citrate including 0.15 M NaCl. The lyophilized form can be stored at 2–8°C. The lyophilized DNA is assigned a shelf life of 5 years. DNA should be dissolved in
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autoclaved or molecular biology grade MilliQ water (nucleasefree). The solution will need to be stirred for at least 2–4 h, at room temperature, to dissolve the DNA. The aliquots of the dissolved DNA solution should be kept at −20°C. Repeating cycles of freeze/thaw should be avoided. We recommend preparing aliquots and storing these. 8. Purity of DNA or RNA may be determined by measuring the A260/A280 ratio. Buffered solutions may provide more accurate and reliable A260/A280 ratios than water. TE buffer (10 mM Tris including 1 mM EDTA, pH 8) is highly recommended for the preparation of stock solutions. A260/A280 ratios can be affected by pH and ionic strength. A lower pH will result in lower A260/A280 ratios and reduced sensitivity to protein contamination. A260/A280 measurements are expected to be 1.7–2.0 to indicate pure DNA or RNA. (Please note, the A260/A280 ratio may not be an accurate measure of nucleic acid purity. This ratio was first used to detect nucleic acid contamination in protein preparations and as such, can be a poor indicator of nucleic acid quality. Alternatively, quality can be assessed by simply analyzing the DNA or RNA by agarose gel electrophoresis or by evaluating performance (for example, by PCR amplification) ). Gently vortex DNA samples for 5 s and sonicate for 5 s. Prepare dilution tubes by adding 980 μL TE buffer to 1.5 mL Eppendorf tubes. Remove 20 μL DNA from each sample (gently pipet up and down two or three times before removing sample to mix) and mix with TE buffer to make a total volume of 1,000 μL, giving a 1:50 dilution. To prepare blank solution, take blank 1,000 μL TE buffer. Vortex all dilutions for 5 s and sonicate for 5 s. Set spectrophotometer program according to your machine. Rinse a clean, quartz cuvette twice with TE buffer, tap on bench paper to blot, and wipe with a tissue paper. Pipet blank solution into cuvette and read absorbances at 320, (background), 280, and 260 nm. Rinse cuvette twice. Pipet the first sample into the cuvette and read absorbances at 320, 280 and 260 nm. Continue by rinsing the cuvette twice and reading additional samples at 320, 280 and 260 nm. To calculate DNA concentration of each sample: (A260–A320) × 50 (DNA extinction coefficient) × dilution factor (i.e., 1,000/20) × final sample volume = DNA yield in μg. 9. For the ethanol precipitation procedure, mix one volume of the probe/DTT solution thoroughly with 0.1 volume of 3 M sodium acetate (pH 5.6) and 3 volumes of ice-cold absolute ethanol. Keep at −80°C for 30 min and centrifuge at 14,000 × g for 15 min. Wash the DNA pellet with 1 mL of 95% ethanol, centrifuge again, dry, and resuspend in water.
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10. The blank buffer solution for the measurement of the electrochemical responses should cover all three electrodes on the screen-printed electrode. An aliquot of 20–30 μL is usually enough for covering all three electrodes, but if there is a demand for working with less volume of solutions, a careful pipetting of 10 μL may also be successful.
References 1. Eggins, B. R. (ed.) (1997) Biosensors: An introduction, Wiley, New York. 2. Kerman, K., Kobayashi, M. and Tamiya, E. (2004) Recent trends in DNA biosensor technology. Measur. Sci. Tech. 15, R1–R11. 3. Murphy, L. (2006) Biosensors and bioelectrochemistry. Curr. Opin. Chem. Biol. 10, 177–184. 4. Soper, S.A., Brown, K., Ellington, A., Frazier, B., Garcia-Manero, G., Gau, V., Gutman, S.I., Hayes, D.F., Korte, B., Landers, J.L., Larson, D., Ligler, F., Majumdar, A., Mascini, M., Nolte, D., Rosenzweig, Z., Wang, J. and Wilson, D. (2006) Point-of-care biosensor systems for cancer diagnostics/prognostics. Biosens. Bioelectron. 21, 1932–1942 5. Kerman, K., Vestergaard, M., Nagatani, N., Takamura, Y. and Tamiya, E. (2006) Electrochemical genosensor based on peptide nucleic acid-mediated PCR and asymmetric PCR techniques: Electrostatic interactions with a metal cation. Anal. Chem. 78, 2182–2189 6. Kerman, K., Meric, B., Ozkan, D., Kara, P., Erdem, A. and Ozsoz, M. (2001) Electrochemical DNA biosensor for the determination of benzo[a]pyrene-DNA adducts. Anal. Chim. Acta 450, 45–52 7. Kerman, K., Ozkan, D., Kara, P., Meric, B., Gooding, J.J. and Ozsoz, M. (2002) Voltammetric determination of DNA hybridization using methylene blue and self-assembled alkanethiol monolayer on gold electrodes. Anal. Chim. Acta 462, 39–47 8. Kerman, K., Saito, M., Morita, Y., Takamura, Y., Ozsoz, M. and Tamiya, E. (2004) Electrochemical coding of single-nucleotide polymorphisms by monobase-modified gold nanoparticles. Anal. Chem. 76, 1877–1884 9. Kerman, K., Morita, Y., Takamura, Y., Ozsoz, M. and Tamiya, E. (2004) Modification of Escherichia coli single-stranded DNA binding protein with gold nanoparticles for electrochemical detection of DNA hybridization. Anal. Chim. Acta 510, 169–174 10. Li, X., Lee, J.S. and Kraatz, H.-B. (2006) Electrochemical detection of single-nucleotide
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mismatches using an electrode microarray. Anal. Chem. 78, 6096–6101 Li, X., Song, H., Nakatani, K. and Kraatz, H.-B. (2007) Exploiting small molecule binding to DNA for the detection of single-nucleotide mismatches and their base environment. Anal. Chem. 79, 2552–2555 Kerman, K., Chikae, M., Yamamura, S. and Tamiya, E. (2007) Gold nanoparticle-based electrochemical detection of protein phosphorylation. Anal. Chim. Acta 588, 26–33 Pjura, P.E., Grzeskowiak, K. and Dickerson, R.E. (1987) Binding of Hoechst 33258 to the minor groove of B-DNA. J. Mol. Biol. 197, 257–271 Teng, M.K., Usman, N., Frederick, C.A. and Wang, A.H. (1988) The molecular structure of the complex of Hoechst 33258 and the DNA dodecamer. Nucleic Acids Res. 16, 2671–2690 de, C.T., Carrondo, M.A.A.F., Coll, M., Aymami, J., Wang, A.H.J., van der Marel, G.A., van Boom, J.H. and Rich, A. (1989) Binding of a Hoechst dye to d(CGCGATATCGCG) and its influence on the conformation of the DNA fragment. Biochemistry 28, 7849–7859 Tanious, F.A., Hamelbeg, D., Bailly, C., Czarny, A., Boykin, D.W. and Wilson, W.D. (2004) DNA sequence dependent monomerdimer binding modulation of asymmetric benzimidazole derivatives. J. Am. Chem. Soc. 126, 143–153 Rohs, R., Sklenar, H., Lavery, R. and Roder, B. (2000) Methylene blue binding to DNA with alternating GC base sequence: a modeling study. J. Am. Chem. Soc. 122, 2860–2866 an, K.X., Shibue, T. and Gross, M.L. (2000) Non-covalent complexes between DNAbinding drugs and double-stranded oligodeoxynucleotides: a study by ESI ion-trap mass spectrometry. J. Am. Chem. Soc. 122, 300–307 Kobayashi, M., Kusakawa, T., Saito, M., Kaji, S., Oomura, M., Iwabuchi, S., Morita,
Protocols for Intercalator-Based Detection of Hybridization Y., Hasan, Q. and Tamiya, E. (2004) Electrochemical DNA quantification based on aggregation induced by Hoechst 33258. Electrochem. Commun. 6, 337–343 20. Chaumpluk, P., Chikae, M., Takamura, Y. and Tamiya, E. (2006) Novel electrochemical identification and semi quantification of bovine constituents in feedstuffs. Sci. Tech. Adv. Mater. 7, 263–269
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21. Saito, M., Kobayashi, M., Iwabuchi, S., Morita, Y., Takamura, Y. and Tamiya, E. (2004) DNA condensation monitoring after interaction with Hoechst 33258 by atomic force microscopy and fluorescence spectroscopy. J. Biochem. 136, 813–823 22. Saito, M., Takamura, Y. and Tamiya, E. (2005) Nanoscale time-lapse AFM imaging in solution for DNA aggregation. Nanobiotechnology 1, 361–368
Chapter 8 Electrochemical Biosensor Technology: Application to Pesticide Detection Ilaria Palchetti, Serena Laschi, and Marco Mascini Summary In recent years, electrochemical sensors and biosensors are becoming an accepted part of analytical chemistry since they satisfy the expanding need for rapid and reliable measurements. An area in which electrochemical biosensors perhaps show the greatest diversity and potential for development involves the measurement of environmentally significant parameters. The increasing number of pollutants in the environment calls for fast and cost-effective analytical requirements. In this context, biosensors appear as suitable alternative or complementary analytical tools. The aim of this chapter is to review some basic concept concerning the electrochemical biosensors and to illustrate a protocol for the detection of environmental organic pollutants on the basis of electrochemical biosensors. In particular, a method based on the inhibition of the enzyme acetylcholinesterase (AChE) for the detection of organophosphorus and carbamate pesticides will be described in detail. Key words: Electrochemical biosensor, Inhibition, AChE, Organophosphorus, Carbamate, Pesticide.
1. Introduction 1.1. General Overview
Despite the wide variation in biosensors and biosensor-related techniques that have been introduced, the consensus definition for these devices has remained fairly constant – an analytical device composed of a biological recognition element directly interfaced to a signal transducer, which together relate the concentration of an analyte (or group of related analytes) to a measurable response. Depending on the method of signal transduction, biosensors can be classified into five basic groups: optical, mass, electrochemical, thermal, and magnetic (1).
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_8
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Electrochemical biosensors have some advantages over other analytical transducing systems, such as the possibility to operate in turbid media, comparable instrumental sensitivity, and possibility of miniaturization. As a consequence of miniaturization, small sample volume can be required. Modern electroanalytical techniques (i.e., square wave voltammetry, chronopotentiometry, chronoamperometry, differential pulse voltammetry) have very low detection limit (10−7–10−9 M). In-situ or on-line measurements are both allowed. Furthermore, the equipments required for electrochemical analysis are simple and cheap when compared with most other analytical techniques (2). Basically electrochemical biosensor can be based on amperometric and potentiometric transducers, even if some examples of conductimetric as well as impedimetric biosensor are reported in literature (3–5). 1.1.1. Amperometric Transducers
L. C. Clark, Jr., is generally considered to be the pioneer of amperometric biosensors in 1962 (6). Clark trapped an enzyme that reacted with oxygen against the surface of a platinum electrode, using a piece of dialysis membrane. He then followed activity of the enzyme, glucose oxidase, by changes in oxygen concentration. Because glucose oxidase is highly specific, it reacts only with glucose, producing hydrogen peroxide and gluconic acid. This analytical scheme became the concept for the basis of the commercial glucose analyzer. Basically amperometric biosensors rely on an electrochemically active analyte that can be oxidized or reduced at a working electrode. This electrode is poised at a specific potential with respect to a reference electrode. The current produced is linearly proportional to the concentration of the electroactive product. A typical set of equipment for amperometric analyses consists of a three electrode cell, based on a working electrode, a reference electrode and an auxiliary electrode as well as the voltage source and devices for measuring current and voltage. If measurements of the current are carried out by scanning the potential, then we obtain a voltammetric system. In this case, measurements of changes in time (t) in the current (I) flowing through the system of electrodes in relation to the potential (E) applied to the working electrode are performed. The registered changes in the current allow drawing the I(t) = f [E(t)] relationship, which is called the voltammogram. In both amperometric and voltammetric biosensors the transducer is an electrode. Typical electrode material are platinum (Pt), gold (Au), carbon, and for some applications, also mercury. Like many other technologies, electrochemical sensors and biosensors have benefited from the growing power of new materials, design, and processing tools; thus many technologies are available to fabricate miniaturized, simple to operate and low cost devices. Among these, thick-film technology is one of the most used as the equipment needed is
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less complex and costly than other; moreover, thick-film electrochemical transducers can be easily mass produced at low cost and thus used as disposable; in electrochemistry, a disposable sensor offers the advantage of not suffering from the electrode fouling that can results in loss of sensitivity and reproducibility (7). Nowadays disposable thick-film electrochemical transducers were produced mainly by the screen-printing technique. Screenprinted electrodes are planar devices, based on different layers of inks printed on a plastic or ceramic substrate (Fig. 1). Many papers (8–12) in the recent years report the use of these devices for environmental as well as for clinical or food analysis, and many of these papers are related to the use of these electrodes in the field of electrochemical sensors and biosensors. One of the most used strategies in screen-printed electrode production is the use of carbon inks for the fabrication of the working electrode surface, since this material is relatively inexpensive, shows a wide working potential range, is inert, has good electrical conductivity, and has relatively high hydrogen overpotential. Moreover carbon screen-printed electrode surface can be easily modified using biomolecules, redox compounds, or catalytic particles, thus increasing the range of compounds detectable. Limitations for amperometric and voltammetric transducer include potential interferences to the response if several electroactive compounds can generate false current values. These effects have been eliminated through the use of selective membranes, which carefully control the molecular weight or the charge of compounds that have access to the electrode.
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Fig. 1. Screen-printed sensor fabrication.
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1.1.2. Potentiometric Transducers
The basic principle behind potentiometric measurements is the development of charge related to the analyte activity a1 in the sample through the Nernst relation: E = E 0 ± (RT / nF )ln a i , where E0 is the standard potential for ai = 1 mol/L, R is the gas constant, F is the Faraday constant, T is the temperature in K, n is the total number of charges on ion i, and the signal + and − are for cations and anions, respectively. Typically, a reference electrode and one working electrode both in contact with the sample are required. The use of ion-selective membranes can make these transducers sensitive to various ions (e.g., H+, F−, I−, Cl−) in addition to gases such as CO2 and NH3, including enzyme systems that change the concentration of any of these ions or gases can result in biosensors that can measure substrates, inhibitors, or modulators of the enzyme (13). The main advantage of such devices is the wide concentration range for which ions can be detected, generally 10−6–10−1 mol/L. Their continuous measurement capability is also an interesting possibility for environmental applications. The apparatus is inexpensive, portable, and is well suited for in situ measurements. The main disadvantage is that the limit of detection in some kinds of environmental samples can be rather high (10−5 mol/L) and the selectivity can be rather poor.
1.2. Environmental Application of Electrochemical Biosensors
An area in which electrochemical biosensors perhaps show the greatest diversity and potential for development involves the measurement of environmentally significant parameters (14, 15). These biosensors are composed of biological assays that have been interfaced with various signal transducers and measure the different parameters. In a number of cases, the interface of a biological assay to a signal transducer has been shown to reduce the time and complexity involved with these assays. In this chapter, we limited the description of a procedure for the detection of environmental organic pollutants, and namely organophosphorus and carbamic pesticides, on the basis of inhibition of the enzyme activity. This inhibition is evaluated by means of an electrochemical biosensor.
1.2.1. Disposable Electrochemical Biosensors for Organophosphorus and Carbamate Pesticide
Organophosphorus and carbamate compounds have come into widespread use in agriculture, since they show low environmental persistence; nevertheless, they exert a high acute toxicity. The principal effect of these compounds is the inhibition of the enzyme acetylcholinesterase (AChE), which is essential for terminating the action of the neurotransmitter acetylcholine (ACh). Actually, the intoxication by these compounds results in
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an accumulation of endogenous ACh and continual stimulation of the nervous system. Because of their toxicological activity, some of these compounds have been used also as chemical warfare agents (CWAs) (16, 17). The most frequently used methods for the unambiguous identification of organophosphorus and carbamate compounds are based on gas chromatography (GC) in combination with mass spectrometry (GC-MS) and/or tandem mass spectrometry (GC-MS/MS), liquid chromatography (LC) coupled with MS, and nuclear magnetic resonance (NMR) spectrometry (18). An alternative to the chromatographic determination of these compounds is the use of biosensor-based techniques. The bioanalytical detection of organophosphate and carbamate pesticides using cholinesterases (ChEs), either free in solution or immobilized as a biorecognition element in biosensors, has a long tradition. The promising results obtained in this research field have allowed the use of ChE in combination with a variety of transducers, such as potentiometric (19, 20), amperometric (21–23), or optical transducers (24). If a salt of acetyl or butyrylthiocholine (ATCh and BTCh, respectively) is used as substrate for the ChE enzymes, thiocholine (TCh) is produced during the enzymatic reaction. Thiol-containing compounds are known as oxidable at the surface of solid electrodes, but the oxidation generally requires high potential values on a suitable electrode (19, 25). This can be overcome using chemically-modified carbon electrodes (26–29). In this chapter, screen-printed carbon electrodes (SPCEs) were modified by incorporating in the ink an optimised percentage of cobalt(II) phthalocyanine (CoPC). As reported in the literature, among the electrochemical mediators, CoPC was indicated as one of the most suitable for the detection of thiol-containing molecules (26), and the resulting oxidation signals occur at lower voltages, thus limiting the electrochemical interference of other oxidable compounds. Using these modified SPCEs, under optimized chronoamperometric conditions, it is possible to detect pesticides, such as Carbofuran, through the study of the AChE activity as reported in (30). In the optimized conditions, the dynamic range for Carbofuran detection was 10−10–10−7 M with a detection limit of 4.9 × 10−10 M, for an analysis time of 15 min. This is an important feature, considering that the immobilization can determine a loss of the activity of the enzyme that influence the sensitivity as well as dynamic range of the pesticide detection (31). Moreover, the proposed method was less prone to electrochemical interferences since the incubation and measurement were performed in two separate steps. In the following sections is reported the methodology:
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(a) To detect organophosphorus and carbamate pesticides using an (AChE)-based electrochemical biosensor. The inhibitory effect of the pesticide determines a decrease of the catalytic activity of AChE; as a consequence, less thiocholine (TCh) was produced from acetylthiocholine (ATCh), the enzymatic substrate. Therefore, the current value, due to the oxidation of TCh at the modified carbon screen-printed electrodes (SPCEs), is lower than that recorded in a blank solution. This current decrease is correlated with the pesticide concentration. (b) To test a Cobalt(II)-phthalocyanine (CoPC) modified screen-printed carbon electrodes (SPCEs) as transducer of an (AChE)-based biosensor. (c) To test biosensor in standard pesticide solutions.
2. Materials 2.1. Chemical and Biochemical Reagents
1. AChE from Electric Eel (EC 3.1.1.7); catalogue number C3389 Sigma (Milan, Italy). 2. ATCh chloride; reagent grade 99%; catalogue number A6625 Sigma (Milan, Italy). 3. Bovin Serum Albumin (BSA); catalogue number A2153 Sigma (Milan, Italy). 4. Nafion® (perfluorinated ion-exchange resin) 5 WT % in mixture of lower aliphatic alcohols and water; catalogue number 527084 Aldrich (Milan, Italy). 5. Carbofuran (2,3-dihydro-2,2-dimethyl-7-benzo-furanol N-methylcarbamate): Aldrich (Milan, Italy) catalogue number 426008. 6. Glycine (aminoacetic acid); catalogue number G7126 Sigma (Milan, Italy). 7. Glutaraldehyde 25% v/v in water; catalogue number G5882 Sigma (Milan, Italy). 8. Acetonitrile HPLC grade: Merck (Milan, Italy). Catalogue number 1,00030 9. Sodium dihydrogenophosphate, disodium hydrogenophosphate, KCl: Merck (Milan, Italy). Catalogue number 1.03345, 1.6580, 73911 10. NaClO 14%: Merck (Milan, Italy). Catalogue number 1.05614 11. All the electrochemical measurements are performed in phosphate buffer 0.05 M pH 8.0 with 0.1 M KCl (measuring buffer). ATCh, AChE, Carbofuran, and other dilutions are prepared in the measuring buffer.
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12. The silver-based ink (Electrodag PF 410) and the graphitebased ink (Electrodag 423 SS): Acheson (Milan, Italy). The insulating mono-component ink (Vinylfast 36–100): Argon Italiana (Milan, Italy). 13. CoPC: Sigma-Aldrich Italiana (Milan, Italy). 2.2. Electrodes
1. CoPC modified screen-printed electrodes (Ecobioservices and Researchers S.r.l, http://www.ebsr.it). 2. A scheme of the CoPC modified screen-printed electrodes is shown in Fig. 1: the inks were deposited onto the polyester substrate (thick 350 μm) in a film of controlled pattern and thickness to obtain overlapping layers. At first the silver tracks were printed, then the CoPC-graphite pad was positioned over part of the silver track, to obtain the working electrode; then the counter electrode is printed using a graphite ink. Finally, the insulating layer with openings that allow the electrical contact with the circuit at one end, and the analyte solution at the other end was deposited. Thus, the screen-printed electrochemical cell consists of graphite working electrode, graphite counter electrode, and silver pseudo-reference electrode. In addition, the silver electrical contacts were covered by a graphite layer to prevent oxidation phenomena, during storage.
2.3. Electroanalytical Instrumentation
Chronoamperometric measurements are carried out using a Palmsens, portable electrochemical sensor interface, obtained from Palmsens (Houten, BV, The Netherlands). The complete measuring system is illustrated in Fig. 2.
Fig. 2. The measuring system. The CoPC sensor was housed inside the connector.
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The analytical data are the current value recorder at 30, after the potential application (0.1 V vs. Ag/AgCl).
3. Methods 3.1. Modified SPCEs Preparation
A scheme of electrochemical cell and electrode design is reported in (Fig. 1). The sensors can be obtained from Ecobioservices and Researches S.r.l. (Florence, Italy). A typical electrode modification formulation is reported in Note 1. Further explanations regarding electrode composition are reported in Note 2. Before use, the pseudo Ag reference electrode is oxidized using NaClO 14% solution, in order to avoid the oxidation of the Ag pseudoreference by thiols during measurements. For storage conditions see Note 3.
3.2. Immobilization of AChE onto CoPC-Modified SPCEs
AChE is immobilized onto the CoPC-modified working electrode surface by cross-linking with glutaraldehyde, BSA, and Nafion®. A first enzyme solution was prepared by mixing 3.3 mL of the measuring buffer with 50 μL of AChE solution and 132 mg of BSA. Then, to 300 μL of this mixture are added 6 μL of glutaraldehyde 25% v/v and 90 μL of Nafion® 5% w/w. The final reagent concentrations are AChE 7.5 U/mL, BSA 3% w/w, glutaraldehyde 0.25% v/v, and Nafion® 0.25% w/w, respectively. Finally, 7 μL of this mixture are casted onto the working area of a CoPC-modified electrode. When the enzymatic layer is dried, electrodes are dipped in a glycine solution 0.1 M for 30 min. This is a blocking treatment, necessary to saturate the surface sites not involved in the enzymatic immobilisation. Biosensors were then stored at +4°C until use.
3.3. Blank Measurements
The enzyme-modified working electrode is covered with 10 μL of buffer; after 5 min, the solution is removed. Then, 200 μL of ATCh chloride (enzymatic substrate) solution (1 mM) are casted onto the cell; after 10 min, the potential was applied and the current response at 30 s is evaluated. Chronoamperometric measurements are performed at the applied potential of +0.1 V vs. pseudo Ag/AgCl reference electrode (see Fig. 3). All potentials are referred to the screen-printed Ag/AgCl pseudo-reference electrode; the experiments are carried out at room temperature (25°C).
3.4. Inhibition Measurements
The enzyme-modified working electrode is covered with 10 μL of buffer containing the inhibitor (sample), see Note 4; after 5 min, the solution is removed and the biosensor washed with the buffer (see Note 5). Then, 200 μL of enzymatic substrate
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Measurement of current
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[carbofuran, M] Fig. 4. Typical inhibition plot of carbofuran onto AChE-based biosensor.
solution (1 mM) are casted onto the cell; after 10 min, the potential was applied and the current response at 30 s is evaluated. Chronoamperometric measurements are performed at the applied potential of +0.1 V vs. pseudo Ag/AgCl reference electrode (see Fig. 4). All potentials are referred to the screen-printed Ag/ AgCl pseudo-reference electrode; the experiments are carried out at room temperature (25°C). Inhibition percentage (I%) can be calculated as follows: [I % = 100[(I 1 - I 2 ) / I 1 ], where I2 was the oxidation current obtained for the sample solution, and I1 the oxidation current obtained for a blank measurement (incubation without pesticide).
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By plotting the inhibition percentage (I%) vs. the pesticide concentration, a typical inhibition plot can be obtained (see Fig. 4). For concentration range see Note 6.
4. Notes 1. CoPC-modified carbon screen-printed electrodes are prepared by mixing the carbon ink with CoPC powder in an amount equivalent to 5% w/w of the total carbon in the printing ink. The mixture is then homogenized. 2. As reported in literature (28), among the electrochemical mediators, CoPC was indicated as one of the most suitable for the detection of thiol-containing molecules and the resulting oxidation signals occur at lower voltages, thus limiting the electrochemical interference of other oxidable compounds. Using CoPC-modified SPCEs, under optimized chronoamperometric conditions, it is possible to detect pesticides, such as Carbofuran, through the study of the AChE activity. 3. The sensors were preferably stored in the dark at room temperature. 4. The term inhibitor used in this paragraph is referred to the carbamic pesticide Carbofuran. A stock solution of Carbofuran 10−3 M in acetonitrile was prepared; working solutions were prepared daily by diluting it in the measuring buffer. To evaluate the pesticide inhibition effect, 10 μL of carbofuran (diluted solution) were added to 500 μL of the buffer containing AChE to get the concentration range 10−11–10−6 M. 5. An important feature of the proposed method is that the sample incubation and the electrochemical measurement were performed in two separate steps, and among them a washing procedure was also included. This guaranteed that the proposed method was less prone to electrochemical interferences, since oxidable substances, eventually present in the sample, were washed off before the electrochemical measurement. Washing step is carried out by depositing and removing 10 μL of the measuring buffer onto the working electrode, for three times. 6. The investigated pesticide concentration range is 10−11–10−6 M. For concentrations higher than 10−6 M, an I% of 100 is generally obtained. The detection limit (dl) can be calculated by substituting the blank – 3× Standard Deviation in the equation of the linear portion of the inhibition curve.
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References 1. Eggins B.R., (2002) Chemical Sensors and Biosensors. Wiley, UK 2. Invitski D., Abdel-Hamid I., Atanasov P., Wilkins E., Striker S., (2000) Application of electrochemical biosensors for detection of food pathogenic bacteria. Electroanalysis, 12(5), 317–325 3. Kellner R., Mermet J.M., Otto M., Valcarcel M., Widmer H.M., (2004) Biosensors, in Analytical Chemistry: A modern Approach to Analytical Science, Second Edition, WileyVCH Verlag GmbH & Co, pp. 1078–1109 4. Cass A.E.G. (ed.) (1990) Biosensors A practical Approach. Oxford University Press, NY 5. Canh T.M., (1993) Biosensors. Chapman & Hall, London, UK 6. Clark L.C., Lyons C., (1962) Electrode systems for continuous monitoring cardiovascular surgery. Ann. N. Y. Acad. Sci., 102, 29–45 7. Alvarez-Icaza M., Bilitewski U., (1993) Mass-production of biosensors. Anal. Chem., 65, 525A–533A 8. Palchetti I., Laschi S., Mascini M., (2005) Miniaturised stripping-based carbon modified sensor for in field analysis of heavy metals. Analytica Chimica Acta, 530, 61–67 9. Laschi S., Palchetti I., Marrazza G., Mascini M., (2006) Development of disposable low density screen-printed electrode arrays for simultaneous electrochemical measurements of the hybridisation reaction. J. Electroanal. Chem., 593, 211–218 10. Farabullini F., Lucarelli F., Palchetti I., Marrazza G., Mascini M., (2007) Disposable electrochemical genosensor for the simultaneous analysis of different bacterial food contaminants. Biosens. Bioelectron., 22, 1544–1549 11. Wang J., (1994) Decentralized electrochemical monitoring of trace metals: from disposable strips to remote electrodes. Analyst, 119, 763–766 12. Centi S., Silva E., Laschi S., Palchetti I., Mascini M., (2007) Polychlorinated biphenyls (PCBs) detection in milk samples by an electrochemical magneto-immunosensor (EMI) coupled to solid phase extraction (SPE) and disposable low density arrays. Analytica Chimica Acta, 594, 9–16 13. Mascini M., Palchetti I., (2005) Enzyme Electrodes, Ion – Selective Electrodes. in Encyclopedia of Analytical Science, 4. Elsevier, Amsterdam, pp. 520–526 14. Rodriguez-Mozaz S., Lopez de Alda M.J., Marco M.-P., Barcelo D., (2005) Biosensors
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
for environmental monitorino A global perspective. Talanta, 65, 291–297 Rogers K.R., (2006) Recent advances in biosensor techniques for environmental monitoring. Anal. Chim Acta, 568, 222–231 Sanchez-Santed F., Canada F., Flores P., Lopez-Grancha M., Cardona D., (2004) Long-term neurotoxicity of Paraoxon and clorpyrifos: behavioural and pharmacological evidence. Neurotoxicol. terathol., 26, 35–317 Noort D., Benschop H.P., Black R.M., (2002) Biomonitoring of exposure to chemical warfare agents: a review. Toxicol. Appl. Pharmacol., 184, 116–126 Hooijschur E.W.J., Hulst A.G., De Jong A.L., De Reuver L.P., Van Krimpen S.H., Van Baar B.L.M., Wils E.R.J., Kientz C.E., Brinkman U.A. Th., (2002) Identification of chemicals related to the chemical weapons convention during an interlaboratory proficiency test. TrAC, 21, 116–130 Palleschi G., Bernabei M., Cremisini C., Mascini M., (1992) Determination of organophosphorus insecticides with a choline electrochemical biosensor. Sens. Actuators B, 7, 513–517 Tran-Minh C., Pandey P.C., Kumaran S., (1990) Studies on acetylcholine sensor and its analytical application based on the inhibition of cholinesterase. Bios. Bioelectron., 5, 461–471 Cagnini A., Palchetti I., Lionti I., Mascini M., Turner A.P.F., (1995) Disposable ruthenized screen-printed biosensors for pesticides monitoring. Sens. Actuators B-Chem., 24, 85–89 Palchetti I., Cagnini A., Del Carlo M., Coppi C., Mascini M., Turner A.P.F., (1997) Determination of anticholisterase pesticides in real samples using a disposable biosensor. Anal. Chim. Acta, 337, 315–321 Hernandez S., Palchetti I., Mascini M., (2000) Determination of anticholinesterase activity for pesticides monitorino using a thiocholine sensor. Intern. J. Environ. Anal. Chem., 78, 263–278 Marty J.-L., Mionetto N., Lacorte S., Barceló D., (1995) Validation of an enzymatic biosensor with various liquid chromatographic techniques for determining organophosphorus pesticides and carbaryl in freeze-dried waters. Anal. Chim. Acta, 311, 265–271 Barceló D., Lacorte S., Marty J.-L., (1995) Validation of an enzymatic biosensor with liquid chromatography for pesticide monitoring. TrAC, 14, 334–340
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26. Hart J.P., Hartley I.C., (1994) Voltammetric and amperometric studies of thiocholine at a screenprinted carbon electrode chemically modified with cobalt phthalocyanine: Studies towards a pesticide sensor. Analyst, 119, 259–263 27. Martorell D., Céspedes F., Martínez-Fàbregas E., Alegret S., (1997) Determination of organophosphorus and carbamate pesticides using a biosensor based on a polishable, 7,7,8,8-tetracyanoquino-dimethane-modified, graphite – epoxy biocomposite. Anal. Chim. Acta, 337, 305–313 28. Silva Nunes G., Skládal P., Yamanaka Y., Barceló D., (1998) Determination of carbamate residues in crop samples by cholinesterasebased biosensors and chromatographic techniques. Anal. Chim. Acta, 362, 59–68
29. Ricci F., Arduini F., Amine A., Moscone D., Palleschi G., (2004) Characterisation of prussian blue modified screen-printed electrodes for thiol detection. J. Electroanal. Chem., 563, 229–237 30. Laschi S., Ogon´czyk D., Palchetti I., Mascini M., (2007) Evaluation of pesticide-induced acetylcholinesterase inhibition by means of disposable carbon-modified electrochemical biosensors. Enzyme Microb. Technol., 40, 485–489 31. Suprun E., Evtugyn G., Budnikov H., Ricci F., Moscone D., Palleschi G., (2005) Acetylcholinesterase sensor based on screenprinted carbon electrode modified with prussian blue. Anal. Bioanal. Chem., 382, 597–604
Chapter 9 Electrochemical Detection of DNA Hybridization Using Micro and Nanoparticles María Teresa Castañeda, Salvador Alegret, and Arben Merkoçi Summary A novel, rapid, and sensitive protocol for the electrochemical detection of DNA hybridization that take the advantage of a magnetic separation/mixing process and the use of monomaleimido-gold nanoparticles of 1.4 nm diameter as label is presented. A sandwich-type assay is formed in this protocol by the capture probe DNA immobilized on the surface of magnetic beads and the double hybridization of the target (cystic fibrosis related DNA), first with the immobilized probe, and then with signaling probe DNA labeled with monomaleimido-gold nanoparticles. When the assay is completed, the final conjugate is transferred onto genomagnetic sensor surface (graphite epoxy composite electrode with a magnet inside) used as working electrode, and then the direct determination of gold nanoparticles by differential pulse voltammetry striping technique is carried out. This protocol is quite promising for numerous applications in different fields as clinical analysis, environmental control as well as other applications. Key words: Gold nanoparticles, DNA analysis, Magnetic beads, Cystic fibrosis, Genosensor, Electrochemical detection
1. Introduction Developments in nanotechnology have driven to research of nanomaterials in the 1–100 nm range offering great potential in a variety of applications such as detection of infectious diseases (1), environmental monitoring (2), detection of pathogens (3), proteomics (4), genomics (5), drug delivery (6), catalytic (7), and others bioanalysis. (8) Materials at this scale, such as metal nanoparticles (NPs), take on novel properties and functions that differ markedly from those seen in the bulk scale (8). The NPs themselves can come in a variety of shapes of which the most Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_9
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commonly prepared are: spheres (9, 10), rods (11), cubes (12), triangles (10), and ellipsoids (13). Metal NPs represent an excellent biocompatibility with biomolecules and display unique structural, electronic, magnetic, optical, and catalytic properties, which in combination with their size have made them a very attractive material in biology (14–18). The attractive physicochemical properties of gold nanoparticles (AuNPs) are highly affected by its shape and size (19, 20). The size and properties of AuNPs are highly dependent on their preparation conditions (7, 21). Dos Santos et al. have reported the synthesis of AuNPs of different shapes and sizes (22). Currently, synthesis of novel AuNPs with unique properties and with applications in a wide variety of areas is the subject of substantial research (23, 24). Among noble-metal nanoparticles, AuNPs have been the most extensively used in electrochemical biosensor applications. This is because the biochemical activity of the labeled receptor biomolecules (i.e., proteins and DNA among others) is retained when AuNPs are coupled to them (25–27). Particularly, AuNPs have been successfully used as electroactive label in the detection of DNA sequences, based on the highly specific hybridization of complementary strands of DNA (2, 5, 28–31). Microscopic magnetic beads on the micrometer size scale have become useful platforms to immobilize biomolecules at different biological assays such those related to antibodies (4), oligonucleotides (28, 30–32), and another applications (32–34). Dynabeads® M-280 Streptavidin (Dynal Biotech, Oslo, Norway) of 2.8 μm diameter, which are uniform, superparamagnetic, polystyrene beads with a monolayer of streptavidin covalently attached to the hydrophobic bead surface, are commonly used. Using a magnetic separator, the beads allow isolation and subsequent handling of target molecules in a highly specific manner. Capture, washing steps, and detection are easily performed and optimized. Herein we present an AuNPs-based electrochemical DNA hybridization detection protocol involving the use of nanoparticles – monomaleimido-Nanogold (AuNPs) 1.4 nm diameter – as labels (see Fig. 1A, B) and microparticles – magnetic beads (MB) 2.8 μm diameter – as platform for DNA probe immobilization (see Fig. 1C, D). In this approach, a DNA biosensor (genosensor) design is based on a sandwich detection strategy in which a cystic fibrosis-related DNA strand used as target is sandwiched between two complementary DNA probes: the capture probe DNA immobilized on MB via streptavidinbiotin and the signaling probe DNA modified with thiol and labeled with AuNPs via reaction of thiol group with monomaleimido so as to ensure a 1:1 AuNP-DNA probe connection.
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Fig. 1. HR-TEM images of monomaleimido-Nanogold 1.4 nm diameter (AuNPs) at: (A) ×400,000 and (B) ×500,000; and paramagnetic beads 2.8 μm diameter (MB) at: (C) ×600 and (D) ×4,000 magnifications.
Differential pulse voltammetry is used for a direct voltammetric detection of AuNPs onto magnetic graphite–epoxy composite electrode (GECE-M).
2. Materials 2.1. Apparatus
1. Electrochemical analyzer Autolab PGSTAT 20 (Eco Chemie, The Netherlands) connected to a personal computer for differential pulse voltammetry (DPV) analyses. 2. Platinum electrode (model 52–67 1, Crison, Spain); that served as an auxiliary electrode.
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3. Double junction Ag/AgCl (Orion 900200, Spain) as reference electrode. 4. Magnetic graphite epoxy composite electrode (GECE-M) as working electrode (home made as described in Subheading 3.1.2). 5. TS-100 Thermo Shaker (Spain) for the binding of streptavidin-coated paramagnetic beads (MB) with biotinylated probe (Immobilization DNA) and hybridization events. 6. MCB 1200 biomagnetic processing platform (Sigris, CA, USA), in order to carry out the magnetic separation. 7. Power supply, 3,000 V/300 mA/300 W (Code PS3003, Ecogen, S.R.L., Spain). 8. A BlueMarine 100 (Inverness Medical Ibérica, S.A.U., Barcelona, Spain) horizontal electrophoresis unit tray is used to carry out the gel electrophoresis. 9. High resolution transmission electron micrographs are taken using a Jeol JEM-2011 electronic microscope (Jeol Ltd., Tokyo, Japan). 2.2. Reagents
1. Tris (hydroxymethyl) methylamine (Tris), sodium chloride, sodium citrate, ethylenediamine tertraacetic acid disodium salt (EDTA), lithium chloride, Tween 20, boric acid, nitric acid, 65%, bovine serum albumin (BSA), (Molecular Biology reagent, Ref. B428, glycerol (G8773-500 mL), 2-Propanol and bromophenol blue sodium salt (B8026) from Sigma-Aldrich. 2. Agarose (Molecular Biology grade, Roche). 3. Xylenecyanol FF, (95600-10G, Fluka). 4. Hydrochloric acid to 37% (PanReac, Barcelona, Spain). 5. Streptavidin-coated paramagnetic beads of diameter 2.8 μm (concentration: 10 mg/mL) – Dynabeads M-280 Streptavidin – (Dynal Biotech, Norway). 6. Monomaleimido-Nanogold, 1.4 nm diameter (Nanoprobes Inc., NY.). 7. Epoxy resin (Epotek H77A) and hardener (Epotek H77B), (Epoxy Technology, Inc., USA). 8. Graphite powder of particle size 50 μm, (BDH, UK).
2.3. Oligonucleotides
1. Biotinylated probe oligonucleotide and no modified oligonucleotides from Alpha DNA, Canada. 2. Oligonucleotide modified with thiol (–SH) group is synthesized in our laboratory on an automatic Applied Biosystems DNA synthesizer, model 392, and according described procedure (35). 3. Oligonucleotides sequences used in the assay are listed in Table 1.
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Table 1 Oligonucleotides used in this protocol Name
Probe sequencea
Capture DNA (CF-A)
5´TGC TGC TAT ATA TAT-biotin-3´
Signaling DNA (CF-B)
Thiol-5´GAG AGT CGT CGT CGT3´
Target DNA (CF-T)b
5´ATA TAT ATA GCA GCA GCA GCA GCA GCA GAC GAC GAC GAC TCT C3´
One base mismatched DNA (CF-MX1)
5´ATA TAT AAA GCA GCA GCA GCA GCA GCA GAC GAC GAC GAC TCT C3´
Three base mismatched DNA (CF-MX3)
5´ATA TAT CCC GCA GCA GCA GCA GCA GCA GAC GAC GAC GAC TCT C3´
Noncomplementary DNA (CF-NC)
5´GGT CAG GTG GGG GGT ACG CCA GG3´
a
Underlined nucleotides correspond to the mismatches Target related to cystic fibrosis gene
b
3. Methods 3.1. Electrode Construction 3.1.1. Transducer Body Construction
1. Take a connection female of 2 mm of diameter, place a metallic thread, and then solder this connection in its extreme to the center of the copper disk (6 mm o.d. and 0.5 mm thickness), with the concavity up (see Fig. 2A). Previously clean the copper disk by dipping it in HNO3 solution (1:1) in order to remove copper oxide and rinsing it well with bidistilled water in order to avoid the decrease of the electrical conductivity of the transducer. 2. Introduce this connection into a cylindrical PVC sleeve (6 mm i.d., 8 mm o.d. and 20 mm longitude) (see Fig. 2B). 3. The metallic thread allows that the connection should remain fixed well in the end of the cylindrical PVC sleeve, whereas in another end there stays a cavity of approximately 3 mm deep in which will be placed the conducting paste (graphite–epoxy composite, which preparation is described at Subheading 3.2.1) and a permanent magnet (see Fig. 2C).
3.1.2. GECE-M Preparation
1. Mix manually epoxy resin and hardener in a ratio 20:3 (w/w) using a small spatula. 2. When the resin and hardener are well-mixed, add the graphite powder in the ratio 20:80 (w/w) and mix thoroughly for 30 min to obtain a homogeneous paste of graphite–epoxy composite. 3. Place the resulting conducting paste of graphite epoxy composite into the cylindrical transducer body where a neodymium magnet (diameter 3 mm, height 1.5 mm, Halde Gac Sdad, Spain, catalog number N35D315) has been introduced,
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Fig. 2. Pictures of: transducer body construction: (*A*, *B*); GECE-M preparation (*C, D*); system of three electrodes, from left to right: auxiliary, working and reference immersed into electrochemical cell (E); electrochemical analyzer Autolab PGSTAT 20 connected to a personal computer, at which DPV electrochemical detection of AuNPs was carried out (F).
2 mm under the surface of the electrode in such a way that the small neodymium magnet stays between two layers of graphite epoxy composite (see Fig. 2C). 4. Electrical contact is completed using the copper disk connected to a copper wire into a cylindrical PVC sleeve (6 mm i.d., 8 mm o.d. and 160 mm longitude) leading to the electrochemical workstation (see Fig. 2D). 5. Cure the conducting composite in a dry heat oven cured at 40°C for 1 week. Figure 2G shows a summarized scheme (not in scale) of GECE-M preparation. 6. Once the resin is hardened, prior to use, the surface of the electrode is polished with abrasive paper and then with alumina paper (polishing strips 301044-001, Orion, Spain) and rinsed carefully with bidistilled water (see Note 1). The prepared electrode will be ready for later measurements in a three electrode
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set-up (see Fig. 2E) connected with the measuring system (see Fig. 2F) as will be described in the following sections. 3.2. Buffers and Solutions Preparation
1. TTL buffer: 100 mM Tris–HCl, pH 8.0; 0.1% Tween 20; and 1 M LiCl. 2. TT buffer: 250 mM Tris–HCl, pH 8.0; and 0.1% Tween 20. 3. TTE buffer: 250 mM Tris–HCl, pH 8.0; 0.1% Tween 20; and 20 mM Na2 EDTA, pH 8.0. 4. Hybridization solution: 750 mmol/L NaCl, 75 mmol/L sodium citrate. 5. Supporting electrolyte: HCl 0.1 M as supporting electrolyte. 6. 5× Tris-Borate-EDTA Buffer (TBE) as running buffer, composition of 10× TBE buffer, for 1 L. 108 g Tris, 55 g Boric acid, 40 mL 0.5 M EDTA (pH 8.0) and MilliQ water to 1 L. The pH is 8.3 and requires no adjustment. Dilute 1 in 20 to obtain 5× TBE buffer. 7. 1× TBE buffer: 10× TBE 100 mL and MilliQ water 900 mL 8. Dyes: Bromophenol blue and xylenecyanol FF. For a 10× concentrated solution, the composition is the following: 0.2% xylenecyanol FF, 0.2% bromophenol blue, 50% glycerol, and 10× TBE buffer MilliQ Water. For preparing 100 mL add 0. 2 g of xylenecyanol, 0.2 g of bromophenol blue, 50 g of glycerol, 10 mL TBE 10× and 40 mL of MilliQ water. Add 1 μL by each 9 mL of solution. 9. BSA at 10%: Weigh 10 g of BSA powder and place it in a 125 mL flask, then add 100 mL of hybridization solution (prepared previously as in 4) to the flask. Swirl to mix the solution (see Notes 2 and 3).
3.3. Functionalization of MonomaleimidoNanogold 1.4 nm
Monomaleimido-Nanogold 1.4 nm (AuNPs) is functionalized with signaling DNA (CF-B). This oligonucleotide modified with thiol (–SH) group is directly bound to the surface of AuNPs tags. 1. The binding (see Fig. 3A) is carried out via reaction of maleimido-thiol group as has been described previously (35). Briefly: 2. Mix aliquots of lyophilized AuNPs (6 nmol) with CF-B (6 nmol) and dissolve in 10% 2-propanol. 3. Keep the mixture overnight at room temperature and store the resulting solution in refrigerator until further use. 4. The maleimido group reacts specifically with sulfhydryl groups when the pH of the reaction mixture is between pH 6.5 and 7.5 and forms a stable thioether linkage that is not reversible (see Fig. 4). 5. The obtained DNA-functionalized AuNPs carry a negative surface charge provided by the anionic thiolated oligonucleotide.
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Fig. 3. Functionalization of monomaleimido-Nanogold 1.4 nm diameter (A). Schematic representation (not in scale) of the analytical protocol (B): (I) Immobilization of the biotinylated CF-A probe onto streptavidin-coated paramagnetic beads (MB); (II) addition of the Target CF to the first hybridization event; (III) addition of monomaleimido-nanogold (AuNPs) functionalized with signalling thiolated CF-B probe to the second hybridization event; (IV) accumulation of final conjugate on the surface of the M-GECE; and (V) magnetically trigged direct DPV electrochemical detection of AuNPs tags in the conjugate.
Fig. 4. Monomaleimido-Nanogold with thiol-oligonucleotide reaction scheme.
3.4. Agarose Gel Electrophoresis of the DNA-Functionalized AuNPs
To verify the purity of the functionalization of AuNPs with CF-B, a gel electrophoresis is carried out. The sample of CF-B/AuNPs conjugate and control dyes (bromophenol blue and xylenecyanol FF) are loaded in the wells of a 2% agarose gel and 80 V is applied along the gel, with electrophoresis time of 20 min, using 0.5× tris-borate-EDTA (TBE) buffer as a running buffer. A detailed description of the procedure is given in Subheadings 3.4.1 and 3.4.2.
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1. Weigh 1 g of agarose powder and place it in a 125 or 250 mL flask (see Note 4). 2. Add 50 mL of 1× TBE buffer to the flask. Swirl to mix the solution. 3. Place the flask in the microwave. Heat on high until the solution is completely clear and no small floating particles are visible (about 2 min). Swirl the flask frequently to mix the solution and prevent the agarose from burning (see Fig. 5A). 4. Cool the solution to 55°C before pouring the gel into the plastic casting tray (see Note 5) 5. While the mixture cools, cover the ends of the gel tray with masking tape. 6. Place the plastic comb in the slots on the side of the gel tray. The comb teeth should not touch the bottom of the tray. 7. Pour the agarose mixture into the gel tray until the comb teeth are immersed about 6 mm or 1/4″ into the agarose. Pour slowly to avoid bubbles (see Fig. 5B) (see Note 6). 8. Allow the agarose gel to cool until solidified. The gel will appear a cloudy white color and will feel cool to the touch (about 20 min).
3.4.2. Gel Electrophoresis
1. Remove the comb from the wells by pulling straight up on the comb. Gently remove the tape from both ends of the gel tray. 2. Place the gel tray in the gel box with the wells closest to the negative (black) electrode. 3. Add enough 1× TBE buffer to fill the electrophoresis chamber and submerge the gel about 1/4 of inch. 4. Pipette 20 μL of control dyes into the first well and 20 μL of MB-CFA conjugate in the next well. Remember to record on the sketch the order the samples and controls were loaded.
Fig. 5. Agarose gel preparation: The mixture of agarose and ×1 TBE buffer is heated at microwave (A) until its complete dissolution. The dissolution formed is cooled up to 55°C and then poured slowly into the gel tray (B) and left to be solidified at room temperature.
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5. Close the top of the electrophoresis chamber. Plug the leads into the electrophoresis chamber. The black lead is the negative lead and should be plugged in closest to the wells. The red lead is the positive lead and should be plugged in furthest from the wells (see Fig. 6A). 6. Plug the other end of the leads into the power source and turn it on. Run the gel at 80 V until the loading dye has travelled 1/2 of the way down the gel approximately (about 20 min) (see Fig. 6B). 7. Turn off the power supply. On plug the leads and the power supply before opening the electrophoresis chamber. 8. Observe the migration of the CF-B/AuNPs conjugate toward the ‘+’ electrode and the discrete band of the conjugate, which indicate its successful preparation. 9. Choose to photograph/photocopy/scan the gel or view it on the overhead projector (see Fig. 7). 10. The obtained conjugate as resulted from functionalization could then assemble with target DNA. 3.5. Sandwich Assay Format Procedure 3.5.1. Immobilization of Capture DNA Probe onto Paramagnetic Beads
The binding of the biotinylated capture DNA probe (CF-A) with magnetic beads (MB) is carried out using a modified procedure recommended by Bangs Laboratories (36), as follows: 1. Transfer 50 μg (5 μL) of MB into 0.5 mL Eppendorf tube (see Note 7). 2. Wash the MB once with 100 μL of TTL buffer using gentle rotation or occasional mixing by gently tapping the tubes (see Note 8). 3. Separate magnetically by placing the tube on MCB 1200 biomagnetic processing platform (magnet) for 1 min (see Fig. 8) (see Note 9).
Fig. 6. Gel electrophoresis apparatus. From left to right: Cover electrophoresis chamber with the corresponding black and red leads, chamber electrophoresis and gel tray with the plastic comb inside (A); Power supply (B).
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Fig. 7. Image of the agarose gel to 2% in which the corresponding bands of control of bromophenol blue and xylenecyanol dyes (1) and DNA/monomaleimido-Nanogold 1.4 nm conjugate (2) are observed. Conditions: 80 V, electrophoresis time 20 min, using ×0.5 Tris-borate-EDTA buffer as running buffer.
Fig. 8. MCB 1200 biomagnetic processing platform (Sigris, CA, USA) in which magnetic separations are carried out.
4. Remove the supernatant with a micropipette while the tube remains on the magnet (see Note 10). 5. Resuspend gently in 20 μL TTL buffer, removing the tube from the magnet previously (see Note 11).
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6. Add 200 pmol of biotin modified capture DNA probe (CFA), (Fig. 3B-I), then adjust the volume to 100 μL by adding deionised and autoclaved water. 7. Incubate resulting MB/CF-A conjugate during 15 min at temperature of 25°C with gentle mixing in a TS-100 Thermo Shaker (see Note 12) to immobilize CF-A. 8. When the immobilization was complete separate magnetically the resulting MB/CFA conjugate (MB with the immobilized CF-A), from the incubation solution by placing the tube on the magnet for 1 min. 9. Remove the supernatant with a micropipette while the tube remains on the magnet. 10. Wash sequentially with 100 μL of TT buffer, 100 μL of TTE buffer, and 100 μL of TT buffer using gentle rotation or occasional mixing by gently tapping the tubes. 11. Separate magnetically by placing the tube on the magnet for 1 min. 12. Remove the supernatant with a micropipette while the tube remains on the magnet. 13. Resuspend gently in 50 μL of hybridization solution and it is ready for the first hybridization. 3.5.2. First Hybridization
1. Add 38 pmol (if no stated otherwise) of target DNA (CF-T) in the solution (50 μL) of the MB/CF-A conjugate obtained in the previous step (see Fig. 3B-II). 2. Adjust the volume to 100 μL by adding deionised and autoclaved water. 3. Incubate at 42°C with gentle mixing during 15 min (see Note 13). 4. When the hybridization was complete separate magnetically the obtained MB/CF-A/CF-T conjugate by placing the tube on the magnet for 1 min. 5. Wash twice with 100 μL of TT buffer using gentle rotation or occasional mixing by gently tapping the tubes. 6. Remove the supernatant with a micropipette while the tube remains on the magnet. 7. Resuspend gently in 50 μL of hybridization solution and it is ready for the second hybridization.
3.5.3. Second Hybridization
1. Add 38 pmol (see Note 14) AuNPs functionalized with CF-B in the ratio 1:1 in the solution (50 μL) of the MB/CF-A/CF-T conjugate obtained in the previous step (see Fig. 3B-III). 2. Add the necessary volume of BSA at 10% and autoclaved water to obtain a final volume of 100 μL and a final concentration of the BSA of 5% approximately (see Note 15). 3. Incubate at 42 °C with gentle mixing during 15 min.
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4. When the hybridization was complete wash the resulting MB/CF-A/CF-T/CF-B-AuNPs conjugate three times with 100 μL of TT buffer, using gentle rotation or occasional mixing by gently tapping the tubes. 5. Separate magnetically by placing the tube on the magnet for 1 min. 6. Remove the supernatant with a micropipette while the tube remains on the magnet. 7. Resuspend in 50 μL of hybridization solution and it is ready for to do the corresponding measurement. 8. Place the solution containing the final conjugate on the surface of GECE-M during 60 s which is accumulated on it due to the inherent magnetic field of the electrode (see Fig. 3B-IV and Note 16). 9. Finally carry out the direct DPV electrochemical detection of Au-NPs tags in the conjugate after the DNA hybridization event, without the need of acidic (i.e. HBr/Br2) dissolution (28, 30), according the established conditions (see Fig. 3B-V). 3.5.4. Control Assay
An identical procedure as described above except the addition of target (Fig. 3-II) to evaluate the nonspecific adsorption onto GECE-M at sandwich assay, simultaneously, was carried out.
3.5.5. Discrimination Study
To study the discrimination between CF-MX1 (one base mismatch), CF-MX3 (three base mismatch), CF-NC (noncomplementary), and the CF-T (target DNA) (See sequences in Table 1) in order to demonstrate the selectivity of the genomagnetic sandwich assay protocol should be made following the same protocol described above.
3.5.6. Conditions of Electrochemical Detection
1. The electrochemical detection is an extensively used method to analyze specific DNA sequences by means of the hybridization event due to its simplicity, selectivity, low instrumentation costs, and high sensitivity. 2. The amount of AuNPs tag was determined by DPV voltammetry as follows: 3. Choose differential pulse voltammetry (DPV) analysis mode in the Autolab software program. 4. Establish the following parameters: Deposition potential, +1.25 V; duration, 120 s; conditioning potential, 1.25 V; step potential, 10 V; modulation amplitude, 50 mV. 5. Run a blank by triplicate immersing the three electrodes: GECEM as working electrode, the Ag/AgCl as reference electrode; and the platinum electrode as auxiliary in an electrochemical cell containing 10 mL of HCl 0.1 M as supporting electrolyte (see Fig. 2E). Save the responses.
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6. Rinse the electrodes with Milli-Q water. 7. Place the sample on the surface of GECE-M during 60 s which is accumulated on it due to the inherent magnetic field of the electrode. 8. Carry out the sample measurement immersing also the three electrodes in the electrochemical cell containing 10 mL of HCl 0.1 M as supporting electrolyte. Save the response (see Fig. 2F). 9. The electrochemical oxidation of Au-NPs to AuCl4− is performed at +1.25 V (vs. Ag/AgCl) for 120 s (see Note 17) in the nonstirred solution. Immediately after the electrochemical oxidation step, DPV is performed. During this step scan the potential from +1.25 to 0 V with step potential 10 mV, modulation amplitude 50 mV, scan rate 33.5 mV/s, no stirred solution. 10. Subtract the response saved for the blank from the sample response using the Autolab software. 11. Save the result, which is an analytical signal due to the reduction of AuCl4− at potential +0.4 V (37). Use the DPV peak
Fig. 9. Typical differential pulse voltammogram (DPV) for the oxidation signals of Au during the sandwich assay to 38 pmol of CF-T (A) and sandwich assay without CF-T used as control (B). Conditions: hybridization time, 15 min; hybridization temperature, 42°C; amount of paramagnetic beads, 50 μg; electrooxidation potential, +1.25 V; electrooxidation time, 120 s; DPV scan from +1.25 to 0 V, step potential 10 mV, modulation amplitude 50 mV, scan rate 33.5 mV/s, nonstirred solution.
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height at the potential of +0.4 V as the analytical signal in all of the measurements. Figure 9A shows the typical differential pulse voltammogram (DPV) for the oxidation signal of Au during the sandwich assay to 38 pmol of CF-T. The Au reduction signal current is proportional to the amount of AuNPs, which corresponds to the concentration of hybridized DNA target. The quantitative result is obtained from the corresponding calibrate plot (not shown). Figure 9B shows the DPV response (almost negligible) to control assay owing to the fact that the sandwich is not formed.
4. Notes 1. Before each use, the surface of the electrode was wet with doubly distilled water and then thoroughly smoothed, first with abrasive paper and then with alumina paper. 2. All stock solutions are prepared using deionised and autoclaved water. 3. Store all stock solutions in refrigerator (4°C) until its use. 4. Fifty milliliters are needed for a single gel. 5. Higher temperatures will melt the plastic tray. 6. Push any bubbles to the side farthest from the wells or to eliminate them. 7. The amount of MB used in this protocol is the result of an optimization between 25 and 150 μg for the same concentration of CF-T (38 pmol). Results not shown. 8. Carry out all the washed, using gentle rotation or occasional mixing by gently tapping the tubes, approximately during 1 min. 9. Do not remove the tube from the magnet during the separation process. 10. Avoid touching the inside wall of the tube (where the beads attract to the magnet) with the pipette tip. 11. Before each different addition into Eppendorf tube, remove the tube from the magnet. 12. The influence of the time and the temperature of hybridization on DPV response is also optimized. Result not shown. 13. All the incubations were carried out at TS-100 Thermo Shaker. 14. Should be added as minimum the same concentration as CF-T DNA. 15. With the BSA used as blocking agent and the effective washing steps, nonspecific adsorption is eliminated. 16. This protocol can be adapted for other fields such as biotechnological and environmental.
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17.
The influence of time and potential of electrochemical oxidation of Au-NPs to AuCl4− upon the DPV signal also are studied in order to establish the optimal values.
Acknowledgments This work is supported by the Spanish “Ramón Areces” foundation (project ‘Bionanosensores’) and MEC (Madrid) thorough the following projects: MAT2008-03079/NAN, and ConsoliderIngenio 2010 (CSD2006-00012). References 1. Pejcic, B., De Marco, R., and Parkinson, G. (2006) The role of biosensors in the detection of emerging infectious diseases. Analyst, 131, 1079–1090 2. Cai, H., Shang, Ch., and Hsing, I. M. (2004) Sequence-specific electrochemical recognition of multiple species using nanoparticle labels. Anal. Chim. Acta, 523, 61–68 3. Lin, F. Y. H., Sabri, M., Alirezaie, J., Li, D., and Sherman, P. M. (2005) Development of a nanoparticle-labeled microfluidic immunoassay for detection of pathogenic MICROORGANISMS. Clin. Diagn. Lab. Immunol., 12, 418–425 4. Ambrosi, A., Castañeda, M. T., Killard, A. J., Smyth, M. R., Alegret, S., and Merkoçi, A., (2007) Double-codified gold nanolabels for enhanced immunoanalysis. Anal. Chem., 79, 5232–5240 5. Zhang, J., Song, S., Zhang, L., Wang, L., Wu, H., Pan, D., and Fan, Ch. (2006) Sequence-specific detection of femtomolar DNA via a chronocoulometric DNA sensor (CDS): Effects of nanoparticle-mediated amplification and nanoscale control of DNA assembly at electrodes. J. Am. Chem. Soc., 128, 8575–8580 6. Sinha, R., Kim, G. J. and Nie, S., and Shin, D. M. (2006) Nanotechnology in cancer therapeutics: bioconjugated nanoparticles for drug delivery. Mol. Cancer Ther. 5, 1909–1917 7. Cuenya, B. R., Hyeon Baeck, S., Jaramillo, T. F., and McFarland, E. W. (2003) Sizeand support-dependent electronic and catalytic properties of Au0/Au3+ nanoparticles synthesized from block copolymer micelles. J. Am. Chem. Soc., 125, 12928–12934
8. McNeil, S. E. (2005) Nanotechnology for the biologist. J. Leukocyte Biol., 78, 585–594 9. DeBenedetti, B., Vallauri, D., Deorsola, F. A., and Martínez García, M. (2006) Synthesis of TiO2 nanospheres through microemulsion reactive precipitation. J. Electroceramics, 17, 37–40 10. Shankar, S.S., Suresh, B., and Murali, S. (2005) Synthesis of gold nanospheres and nanotriangles by the Turkevich approach. J. Nanosci. Nanotechnol., 5, 1721–1727 11. Tai, H. H., Koo, H. -J., and Chung, B. H. (2007) Shape-controlled syntheses of gold nanoprisms and nanorods influenced by specific adsorption of halide ions. J. Phys. Chem. C, 111, 1123–1130 12. Hyuk Im, S., Tack Lee, Y., Wiley, B., and Xia, Y., (2005) Large-scale synthesis of silver nanocubes: The role of HCl in promoting cube perfection and monodispersity. Angew. Chem. Int. Ed., 44, 2154–2157 13. Mendoza-Reséndez, R., Bomati-Miguel O., Morales, M. P., Bonville, P., and Serna C. J. (2004) Microstructural characterization of ellipsoidal iron metal nanoparticles. Nanotechnology, 15, S254–S258 14. Hernández-Santos, D., González-García, M. B., and Costa-García, A., (2002) Metalnanoparticles based electroanalysis. Electroanalysis, 14, 1225–1235 15. Alivisatos, P. (2004) The use of nanocrystals in biological detection. Nat. Biotechnol., 22, 47–52 16. Rosi, N. L., and Mirkin, C. A. (2005) Nanostructures in biodiagnostics. Chem. Rev., 105, 1547–1562 17. Azzazy, H. M. E., Mansour M. M. H., and Kazmierczak, S. C. (2006) Nanodiagnos-
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18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
tics: A New Frontier for Clinical Laboratory Medicine. Clin. Chem., 52, 1238–1246 Katz, E., and Willner, I. (2004) Integrated nanoparticle-biomolecule hybrid systems: Synthesis, properties and applications. I. Angew. Chem. Int. Ed., 43, 6042–6108 Liz-Marzan, L. M. (2004) Nanometals: Formation and color. Materials Today, 7, 26–31 Burda, C., Chen, X., Narayanan, R., and ElSayed, M. A. (2005) Chemistry and properties of nanocrystals of different shapes. Chem. Rev., 105, 1025–1102 Miscoria, S.A., Barrera, G. D., and Rivas, G. A. (2005) Enzymatic biosensor based on carbon paste electrodes modified with gold nanoparticles and polyphenol oxidase. Electroanalysis 17, 1578–1582 Dos Santos, D. S. Jr., Alvarez-Puebla, R. A., Oliveira, O. N. Jr., and Aroca, R. F. (2005) Controlling the size and shape of gold nanoparticles in fulvic acid colloidal solutions and their optical characterization using SERS. J. Mater. Chem., 15, 3045– 3049 Panda, B. R., and Chattopadhyay, A. (2007) Synthesis of Au Nanoparticles at “all” pH by H2O2 Reduction of HAuCl4. J. Nanosci. Nanotechnol., 7, 1911–1915 Luo, Y., and Sun X. (2007) Sunlight-driving formation and characterization of sizecontrolled gold nanoparticles. J. Nanosci. Nanotechnol., 7, 708–711 Castañeda, M. T., Alegret, S., and Merkoçi, A. (2007) Electrochemical sensing of DNA using gold nanoparticles. Electroanalysis, 19, 743–753 Merkoçi, A. (2007) Electrochemical biosensing with nanoparticles. FEBS J., 274, 310–316 Luo, X., Morrin, A., Killard, A. J., and Smyth, M. R. (2006) Application of nanoparticles in electrochemical sensors and biosensors. Electroanalysis, 18, 319–326 Pumera, M., Castañeda, M. T., Pividori, M. I., Eritja, R., Merkoçi A., and Alegret, S. (2005) Magnetically trigged direct electrochemical detection of DNA hybridization
29.
30.
31.
32.
33.
34.
35.
36. 37.
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using Au67 quantum dot as electrical tracer. Langmuir, 21, 9625–9629 Ozsoz, M., Erdem, A., Kerman, K., Ozkan, D., Tugrul, B., and Topcuoglu, N. (2003) Electrochemical genosensor based on colloidal gold nanoparticles for the detection of factor V Leiden mutation using disposable pencil graphite electrodes. Anal. Chem., 75, 2181–2187 Castañeda, M. T., Merkoçi, A., Pumera, M., and Alegret, S., (2007) Electrochemical genosensors for biomedical applications based on gold nanoparticles. Biosens. Bioelectron., 22, 1961–1967 Wang, J., Xu, D., Kawde, A. N., and Polsky R. (2001) Metal nanoparticle-based electrochemical stripping potentiometric detection of DNA hybridization. Anal. Chem., 73, 5576–5581 Palecek, E., Fojta, M., and Jelen, F. (2002) New approaches in the development of DNA sensors: hybridization and electrochemical detection of DNA and RNA at two different surfaces. Bioelectrochemistry, 56, 85–90 Gijs, M. A. M., (2004) Magnetic bead handling on-chip: new opportunities for analytical applications. Microfluid. Nanofluidics, 1, 22–40 Lim, C. T., and Zhang Y. (2007) Beadbased microfluidic immunoassays: The next generation. Biosens. Bioelectron., 22, 1197– 1204 Torre, B. G., Morales, J. C., Avino, A., Iacopino, D., Ongaro, A., Fitzmaurice, D., Murphy, D., Doyle, H., Redmond, G., and Eritja, R. (2002) Synthesis of oligonucleotides carrying anchoring groups and their use in the preparation of oligonucleotidegold conjugates. Helv. Chim. Acta, 85, 2594–2607 Bangs Laboratories Inc., TechNote 101 (1999) Pumera, M., Aldavert, M., Mills, C., Merkoçi, A., and Alegret, S. (2005) Direct voltammetric determination of gold nanoparticles using graphite–epoxy composite electrodes. Electrochim. Acta, 50, 3702–3707
Chapter 10 Electrochemical Immunosensing Using Micro and Nanoparticles Alfredo de la Escosura-Muñiz, Adriano Ambrosi, Salvador Alegret, and Arben Merkoçi Summary A model immunosensor based on a labeling method using gold nanoparticles (AuNPs) and electrochemical detection is developed. Microparamagnetic beads (MB) as primary antibody immobilization platforms and AuNPs modified with a secondary antibody as high sensible electrochemical labels have been used. The carbon electrode used as transducer incorporates a magnet that allows the collection/ immobilization on its surface of the immunological sandwich attached to the MB. Briefly, the sandwich type assay consists in the incubation of streptavidin-coupled-MB with an antihuman IgG biotin conjugate, and then, the immunological reaction with the human IgG antigen takes place. After that, a gold labeled anti-human IgG reacts with the antigen, and finally the AuNPs are electrochemically detected. This approach allows the obtaining of an immunosensor with a low antigen detection limit with special interest for several applications in protein analysis. Key words: Gold nanoparticles, Protein analysis, Magnetic beads, Human IgG, Immunosensor, Electrochemical detection, Horse radish peroxidase.
1. Introduction Gold nanoparticles have been used for analytical and biomedical purposes for many years. Rapid and simple chemical synthesis, a narrow size distribution and efficient coating by thiols or other bioligands has enabled gold nanoparticles to be used as transducers for several biorecognition-binding applications. Properties such as their electron dense core, highly resonant particle plasmons, direct visualization of single nanoclusters by scattering Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_10
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of light, catalytic size enhancement by silver deposition, electrochemical properties made them very attractive for several applications in nanobiotechnology. Bioconjugated gold nanoparticles for recognizing and detecting specific DNA sequences that function as both a nano-scaffold and a nano-quencher (efficient energy acceptor) have been reported (1). Gold nanoparticles conjugated with antibodies are widely used in the field of light and electron microscopy, for visualizing proteins in biological samples (2). The sensitivity of the detection is usually improved by the silver enhancement method. Beside these applications, an increased interest is shown for their use to quench the fluorescence (3), tune the enzyme specificity (4), visualize cellular or tissue components by electron microscopy (5), electrical contacting or “wiring” between electrodes and redox enzymes (6), tailoring the DNA loading by changing the nanoparticle size (7), and for labeling DNA strands for sensor and analytical applications. The combination of biomolecules with gold nanoparticles provides interesting tools for several biological components. Oligonucleotide functionalized gold nanoparticles have become the basis for an increasing number of diagnostic applications that compete with molecular fluorophores in certain settings (8). The use of gold nanoparticles for protein analysis is also a very interested research field with special interest for applications too. Gold nanoparticle/protein conjugates are finding increasing application as biochemical sensors, enzyme enhancers, nanoscale building blocks, and immunohistochemical probes (9, 10). Nanoparticles in general and gold nanoparticles particularly offer attractive properties to act as DNA tags (11). Their sensitivity, long life-time along with multiplexing capability have led to an extensive applications in electrochemical assays in recent years (12). Most of the reported assays have been based on chemical dissolution of gold nanoparticle tag (in a hydrobromic acid/bromine mixture) followed by accumulation and stripping analysis of the resulting Au3+ solution. Due to the toxicity of the HBr/ Br2 solution, direct solid-state detection of silver precipitate on gold nanoparticle-DNA conjugates was reported by Wang et al. (13). However, this method was based on direct detection of precipitated silver, not gold nanoparticle tag itself. Direct detection of colloidal gold nanoparticles but not in connection with the detection of DNA hybridization was reported earlier by our and Costa-Gracia’s groups (14–16). A novel nanoparticle-based detection of DNA hybridization based on magnetically induced direct electrochemical detection of 1.4 nm Au67 quantum dot tag linked to the target DNA have been reported previously by our group. The Au67 nanoparticle tag is directly detected after the DNA hybridization event, without the need of acidic (i.e. HBr/ Br2) dissolution (17, 18).
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The system developed in this work establish a general electrochemical detection methodology that can be applied to a variety of immunosystems and DNA detection systems, including labon-a-chip technology, with special interest for further applications in clinical analysis, food quality and safety as well as other industrial applications.
2. Materials 2.1. Apparatus
1. An electrochemical analyzer Autolab 20 (Eco- Chemie, The Netherlands) connected to a personal computer is used for performing all voltammetric experiments. 2. A 5 mL voltammetric cell at room temperature (25°C) is used for the electrochemical experiments, using a three electrode configuration: – A platinum electrode (model 52–67 1, Crison, Spain) as auxiliary electrode. – A doble junction Ag/AgCl (Orion 900200, Spain) as reference electrode. – Graphite composite working electrodes (M-GECE) (home made as described at Subheading 3.1). 3. A TS-100 ThermoShaker (Spain) is used for all the incubations for the binding of streptavidin coated paramagnetic beads with biotinylated primary antibody. 4. MCB1200 biomagnetic processing platform (Sigris, CA, USA) is used for the magnetic separations (see Fig. 9.8 of Chap. 9). 5. Tecan Sunrise Absorbance Microplate Reader is used for performing the spectrophotometric measurements. 6. Transmission electron micrographs (TEM) are taken using a Hitachi H-700 electronic microscope (Hitachi Ltd, Japan).
2.2. Reagents
1. Streptavidin-coated paramagneticagnetic Beads (MB), 2.8 μm sized (Dynabeads M-280, Dynal Biotech., Norway). Concentration: 10 mg/mL. 2. Biotin conjugate-goat anti-human IgG (Sigma B1140, Spain, developed in goat and gamma chain specific). 3. Human IgG from serum (Sigma-Aldrich, Spain). 4. Goat IgG from serum (Sigma-Aldrich, Spain). 5. Anti-human IgG (Sigma A8667, Spain, developed in goat). 6. Hydrogen tetrachloroaurate(III) trihydrate (HAuCl4⋅3H2O, 99.9%) (Sigma-Aldrich, Spain).
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7. Trisodium citrate (Sigma-Aldrich, Spain). 8. Bovine serum albumin (BSA) (Sigma-Aldrcih, Spain).
3. Methods 3.1. Magnetic Graphite–Epoxy Composite Electrode (GECE-M) Construction 3.1.1. Electrode Body Construction
A schematic of the electrode construction is shown in Fig. 9.2 of Chap. 9. 1. Take a connection female of 2 mm of diameter, place a metallic thread and then solder this connection in its extreme to the center of the copper disk (6 mm o.d. and 0.5 mm thickness), with the concavity up. Previously clean the copper disk by dipping it in HNO3 solution (1:1) to remove copper oxide and rinsing it well with bi-distilled water. 2. Introduce this connection into a cylindrical PVC sleeve (6 mm i.d., 8 mm o.d., and 16 mm long). 3. The metallic thread allows that the connection should remain fixed well in the end of the cylindrical PVC sleeve, whereas in another end there stays a cavity of approximately 3 mm deep in which is placed the conducting composite and the permanent magnet.
3.1.2. GECE-M Preparation
1. Mix the epoxy resin (Epotek H77A, Epoxy Technology, USA) and the hardener (Epotek H77B, USA) manually in the ratio 20:3 (w/w) using a spatula. 2. When the resin and hardener are well-mixed, add the graphite powder (particle size 50 μm, BDH, U.K.) in the ratio 1:4 (w/w) and mix for 30 min. 3. Place the resulting paste into the body of the electrode prepared as described in Subheading 3.1.1, where a neodymium magnet (diameter 3 mm, height 1.5 mm, Halde Gac Sdad, Spain, catalog number N35D315) has been introduced 2 mm under the surface of the electrode. 4. Cure the conducting composite at 40°C for 1 week. 5. Once the resin is hardened, polish the electrode surface, which is wet with doubly distilled water and then smooth thoroughly, first with abrasive paper of different grades of and then with alumina paper (polishing strips 301044-001, from Orion, Spain) (see Note 1).
3.2. Buffers and Solutions Preparation
1. The phosphate buffer solution (PBS) consists of 0.01 M phosphate buffered saline, 0.137 M NaCl, 0.003 M KCl (pH 7.4) (see Note 2).
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2. Blocking buffer solution consists of a PBS solution with added 5% (w/v) bovine serum albumin (pH 7.4). 3. The binding and washing (B&W) buffer consists of a PBS solution with added 0.05% (v/v) Tween 20 (pH 7.4). 4. The measuring medium for the electrochemical measurements consists of a 0.1 M HCl solution. 3.3. Synthesis and Characterization of Gold Nanoparticles
Gold nanoparticles (AuNPs) are synthesized by reducing tetrachloroauric acid with trisodium citrate, a method pioneered by Turkevich et al. (19): 1. Boil 200 mL of 0.01% HAuCl4 solution with vigorous stirring (see Note 3). 2. Add 5 mL of a 1% trisodium citrate solution to the boiling solution (see Note 4). 3. Left the solution stirring and cooling down until the color turns deep red, indicating the formation of AuNPs. 4. Transmission electron micrographs (TEM) can be performed to characterize the AuNPs. An example of TEM images is shown in Fig. 1. Fast Fourier Transform (FFT) of crystalline planes distances, UV–Vis spectrum and energy dispersive X-ray can be also employed to achieve a better characterization of the AuNP obtained (results not shown here).
3.4. Preparation of the Conjugated AuNP/ Anti-Human IgG 3.4.1. Gold Aggregation Test
Gold aggregation test is preliminarily carried out to judge the minimum antibody concentration to be used for conjugation: 1. AuNPs solution obtained as described at Subheading 3.3 is adjusted to pH 9 with NaOH 0.01 M. 2. Prepare in water several solutions with different concentrations of anti-human IgG (ranging from 2.5 to 12 μg/mL) to a volume of 30 μL, and add it to 200 μL of AuNP suspension solution.
Fig. 1. Transmission electron micrographs of aunps (15 nm) at (A) ×50,000, (B) ×200,000, and (C) ×500,000 magnifications.
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3. After 5 min, add 30 μL of 10% NaCl solution. NaCl causes the aggregation of AuNPs and shifts the maximum absorbance peak from 520 to 580 nm. 4. The antibody concentration which prevents gold aggregation is determined by measuring the difference between the absorbance at 520 and at 580 nm and plotting it against the concentration used (see Note 5). 3.4.2. Conjugation AuNP/ Anti-Human IgG
The conjugated AuNP/anti-human IgG, is prepared by following a published procedure (20): 1. Mix 2 mL of AuNPs suspension, adjusted to pH 9 with NaOH 0.01 M, with 100 μL of 100 μg/mL anti-human IgG and incubate at 25°C for 20 min (see Note 6). 2. Add 150 μL of 1 mg/mL BSA and incubate at 25°C for 20 min (see Note 7). 3. Centrifuge at 14,000 rpm (18,000) rcf for 20 min, and reconstitute the AuNP/anti-human IgG in 2 mL of H2O milli-Q (see Note 8).
3.5. Preparation of the Sandwich-Type Immunocomplex
The binding of the biotinylated anti-human IgG with streptavidin-coated paramagnetic beads (MB) is carried out using a slightly modified procedure recommended by Dynal Biotech (21). Figure 2 is a schematic of the whole assays steps (see Note 9): 1. Transfer 150 μg (15 μL from the stock solution) of MB into 0.5 mL Eppendorf tube, wash twice with 150 μL of B&W buffer, and resuspend in 108 μL of B&W buffer (see Note 10). 2. Add 42 μL (from stock solution 0.36 mg/mL) of biotinylated anti-human IgG, and incubate the resulting solution for 30 min at temperature 25°C. 3. Separate the formed MB/anti-human IgG from the incubation solution and wash three times with 150 μL of B&W buffer. 4. Resuspend the MB/anti-human IgG in 150 μL of blocking buffer (PBS–BSA 5%) and incubate at 25°C for 60 min (see Note 11). 5. Wash three times with B&W buffer. 6. Add 150 μL of human IgG antigen solution, at different concentrations, to the MB/anti-human IgG solution, and incubate at 25°C for 30 min, forming by this way the immunocomplex MB/anti-human IgG/Human IgG. Prepare a control solution in the same way, but using 150 μL of a nonspecific antigen (goat IgG) instead of human IgG (see Note 12). 7. Wash three times with B&W buffer. 8. Finally, add 150 μL of the previously synthesized AuNP/antihuman IgG complex to the MB/anti-human IgG/Human IgG immunocomplex, and incubate at 25°C for 30 min. In
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Fig. 2. Schematic (not in scale) of (a) AuNP conjugation with anti-human IgG; (b) analytical procedure for the sandwich type assay and the electrochemical detection.
Fig. 3 are shown examples of TEM images obtained for the sandwich assay for both cases: human IgG and goat IgG. In the latest case (the nonspecific antigen) are not observed AuNPs around the MB. 3.6. Spectrophotometric Analysis: Signal Enhancement by Using AuNPs Conjugates
The conjugation of the AuNPs can be also performed with antihuman IgG peroxidase conjugate under the same experimental conditions. Then, the MB sandwich immunocomplex can be carried out without AuNP (MB/anti-human IgG/Human IgG/ anti-human IgG-HRP) and with AuNP (MB/anti-human IgG/ Human IgG/AuNP/anti-human IgG-HRP) and finally analyzed spectrophotometrically, to evaluate the benefits in using AuNPs. The spectrophotometric analysis is based in the oxidation of o-phenylenediamine (OPD) catalyzed by the horseradish peroxidase (HRP) bound to AuNPs, which generates a product with maximum absorbance at 492 nm in acidic solution. In the case of the sandwich carried out with AuNP, an enhancement of the signal is
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Fig. 3. Transmission electron micrographs (TEM) images of the MB after the sandwich type assay, carried out with a nonspecific antigen (goat IgG) (blank assay-(A)) and with 1.0 × 10−3 μg/mL of human IgG (B).
obtained (22). This simple optical-based detection procedure can also be applied as an alternative to the electrochemical method. 3.7. Electrochemical Detection
The electrochemical detection of the gold AuNPs anchored to the MB through the immunological reaction is based in a protocol that involves an accumulation step on the electrode surface, the electrochemical oxidation of AuNPs to AuCl4− and then the performing of a differential pulse voltammogram (DPV), recording the reduction process of gold, which generates a peak current that constitutes the analytical signal (14): 1. Resuspend the MB/anti-human IgG/Human IgG/AuNP/ anti-human IgG immunocomplex in 150 μL of double-distilled water. 2. Place 50 μL of the suspension on the surface of the GECE-M, and leave it there for 5 min. The inherent magnetic field of the electrode certainly improves the accumulation process, keeping the MB well immobilized and the AuNP in tight contact with the sensing surface. 3. After 5 min, transfer the electrode, without any washing steps, to an electrochemical cell containing HCl 0.1 M and oxidize the AuNPs to AuCl4− applying a potential of +1.25 V (vs. Ag/ AgCl) for 120 s in a nonstirred solution. 4. Perform differential pulse voltammetry (DPV) by scanning from +1.25 to 0.0 V (step potential 10 mV, modulation amplitude 50 mV, scan rate 33.5 mV/s, nonstirred solution), resulting in an analytical signal due to the reduction of AuCl4− at potential +0.45 V. In Fig. 4 are shown examples of the voltammograms obtained and the linear relationship between the human IgG concentration and the peak current of the gold reduction (see Note 13).
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Fig. 4. (A) Relationship between the concentration of human IgG and the peak current of the gold reduction process. (B) Differential pulse voltammograms recorded from 1.25 to 0.0 V, for human IgG concentrations between 2.5 × 10−5 and 1 μg/mL. Electrode preconditioning: 1.25 V for 120 s; deposition potential 1.25 V for 150 s; step potential 10 mv; amplitude 50 mv; scan rate 33 mv/s (vs. Ag/AgCl reference electrode).
4. Notes 1. Before each use, the surface of the electrode is wet with doubly distilled water and then thoroughly smoothed, first with abrasive paper and then with alumina paper (polishing strips 301044-001, Orion, Spain). 2. All buffer reagents and other inorganic chemicals are supplied by Sigma, Aldrich or Fluka, unless otherwise stated. All chemicals are used as received and all aqueous solutions are prepared in doubly distillated water. 3. The beaker used as electrochemical cell as well as other glasses must be washed previously overnight in an aqua-regia solution. 4. The addition of the trisodium citrate solution must be carried out quickly. 5. The minimum antibody concentration giving the highest absorbance difference corresponds to a number of protein molecules of 10 for each gold nanoparticle used in this example. This result can be also verified by theoretical calculations (22). 6. All the incubations described in this work are performed in the TS-100 ThermoShaker, with gentle mixing at 25°C. 7. The incubation with BSA is a very important step to block any remaining active surface of AuNPs. 8. All the supernatant must be extracted carefully, but not the precipitated, which must be reconstituted with H2O milli-Q. 9. The main parameters involved in this assay have been previously optimized (data not shown).
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10. All the washing steps in this procedure are performed adding the washing solution, mixing, and placing the tube on the MCB 1200 biomagnetic processing platform (magnet) for 1 min, separating the MB from the solution by applying the magnetic field, and extracting carefully the solution (no colored solution), avoiding to extract the MB (brown solution). 11. The incubation with BSA is a very important step to block any remaining active surface of MBs that could result in nonspecific adsorptions. 12. The assay performed with goat IgG allow evaluate nonspecific adsorptions. 13. The main parameters that affect the analytical signal have been previously optimized (data not shown).
Acknowledgment A.M. thanks the Spanish “Ramón Areces” foundation (project ‘Bionanosensores’) and MEC (Madrid) for the following Projects: MAT2008-03079/NAN, and Consolider-Ingenio 2010, Proyecto CSD2006-00012. A.E. thanks to the MEC (Madrid) for the Juan de la Cierva scholarship.
References 1. Maxwell, D.J., Taylor, J.R. and Nie, S. (2002) Self-assembled nanoparticle probes for recognition and detection of biomolecules. J. Am. Chem. Soc. 124, 9606–9612 2. Horisberger, M. (1981) Colloidal gold: a cytochemical marker for light and fluorescent microscopy and scanning electron microscopy. Scan. Electron Microsc. 11, 9–31 3. Huang, T. and Murray, R.W. (2002) Quenching of [Ru(bpy)3]2+ fluorescence by binding to Au nanoparticles. Langmuir 18, 7077–7081 4. You, C.C., Agasti, S.S., De, M., Knapp, M.J. and Rotello, V.M. (2006) Modulation of the catalytic behavior of α-chymotrypsin at monolayer-protected nanoparticle surfaces. J. Am. Chem. Soc. 128, 14612–14618 5. De la Fuente, J.M., Berry, C.C., Riehle, M.O. and Curtis, A.S.G. (2006) Nanoparticle targeting at cells. Langmuir 22, 3286–3293 6. Zayats, M., Katz, E., Baron, R. and Willner, I. (2005) Reconstitution of apo-glucose dehydrogenase on pyrroloquinoline quinone-
7.
8.
9.
10.
functionalized Au nanoparticles yields an electrically contacted biocatalyst. J. Am. Chem. Soc. 127, 12400–12406 Hurst, S.J., Lytton-Jean, A.K.R. and Mirkin, C.A. (2006) Maximizing DNA loading on a range of gold nanoparticle sizes. Anal. Chem. 78, 8313–8318 Lytton-Jean, A.K.R. and Mirkin, C.A. (2005) A thermodynamic investigation into the binding properties of DNA functionalized gold nanoparticle probes and molecular fluorophore probes. J. Am. Chem. Soc. 127, 12754– 12755 Ackerson, C.J., Jadzinsky, P.D., Jensen, G.J. and Kornberg, R.D. (2006) Rigid, specific, and discrete gold nanoparticle/antibody conjugates. J. Am. Chem. Soc. 128, 2635–2640 Stoeva, S.I., Lee, J.S., Smith, J.E., Rosen, S.T. and Mirkin, C.A. (2006) Multiplexed detection of protein cancer markers with biobarcoded nanoparticle probes. J. Am. Chem. Soc. 128, 8378–8379
Electrochemical Immunosensing Using Micro and Nanoparticles 11. Merkoçi, A., Aldavert, M., Marin, S. and Alegret, S. (2005) New materials for electrochemical sensing V: Nanoparticles for DNA labeling. Trends Anal. Chem. 24, 341–349 12. Katz, E., Willner, I. and Wang, J. (2004) Electroanalytical and bioelectroanalytical systems based on metal and semiconductor nanoparticles. Electroanalysis 16, 19–44 13. Wang, J., Xu, D. and Polsky, R. (2002) Magnetically-induced solid-state electrochemical detection of DNA hybridization. J. Am. Chem. Soc. 124, 4208–4209 14. Pumera, M., Aldavert, M., Mills, C., Merkoçi, A. and Alegret, S. (2005) Direct voltammetric determination of gold nanoparticles using graphite-epoxy composite electrode. Electrochim. Acta 50, 3702–3707 15. González García, M.B. and Costa García, A. (1995) Adsorptive stripping voltammetric behaviour of colloidal gold and immunogold on carbon paste electrode. Bioelectr. Bioenerg. 38, 389–395 16. Hernández Santos, D., González García, M.B. and Costa García, A. (2002) Metal-nanoparticles based electroanalysis. Electroanalysis 14, 1225–1235
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17. Pumera, M., Castañeda, M.T., Pividori, M.I., Eritja, R., Merkoçi, A. and Alegret, S. (2005) Magnetically trigged direct electrochemical detection of DNA hybridization using Au67 quantum dot as electrical tracer. Langmuir 21, 9625–9629 18. Castañeda, M.T., Merkoçi, A., Pumera, M. and Alegret, S. (2007) Electrochemical genosensors for biomedical applications based on gold nanoparticles. Biosen. Bioelectr. 22, 1961–1967 19. Turkevich, J., Stevenson, P. and Hillier, J. (1951) A study of the nucleation and growth processes in the synthesis of colloidal gold. Discuss. Faraday Soc. 11, 55–59 20. Beesley, J. (1989) Colloidal gold. A new perspective for cytochemical marking. Royal Microscopical Society Handbook No 17.Oxford Science Publications. Oxford University Press 21. Dynal Biotech, Technote 010 for product 112.05 22. Ambrosi, A., Castañeda, M.T., Killard, A.J., Smith, M.R., Alegret, S. and Merkoçi, A. (2007) Double-codified gold nanolabels for enhanced immunoanalysis. Anal. Chem. 79, 5232–5240
Chapter 11 Methods for the Preparation of Electrochemical Composite Biosensors Based on Gold Nanoparticles A. González-Cortés, P. Yáñez-Sedeño, and J.M. Pingarrón Summary Methods for the construction of electrochemical composite biosensors using gold nanoparticles and Teflon as nonconducting-binding material are described in detail. The advantages of the incorporation of gold nanoparticles to the composite electrode matrices are highlighted, giving rise to bioelectrodes with improved analytical performance in terms of stability and sensitivity with respect to other biosensor designs. Three different biosensors have been considered: a tyrosinase biosensor in which the enzyme and gold nanoparticles are incorporated into graphite–Teflon composite electrode matrices by simple physical inclusion, a progesterone immunusensor in which the antibody is directly attached to the electrode surface and amperometric transduction is carried out at a colloidal gold–graphite–Teflon–tyrosinase composite biosensor, and a mediator-less glucose oxidase biosensor constructed by bulk incorporation of the enzyme into colloidal gold-multiwall carbon nanotubes–Teflon composite electrodes. Key words: Electrochemical biosensors, Gold nanoparticles, Enzyme biosensors, Immunosensors, Composite electrodes, Carbon nanotubes.
1. Introduction Nowadays, the construction of electrochemical biosensors based on the use of gold nanoparticles constitutes an intensive research area because of the unique advantages that this nanomaterial lends to biosensing devices. So, gold nanoparticles provide a stable surface for immobilization of biomolecules with no loss of their biological activity. Moreover, they facilitate direct electron transfer between redox proteins and electrode materials, and constitute useful interfaces for the electrocatalysis of redox processes of molecules such as H2O2 or NADH involved in many biochemical reactions (1, 2). Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_11
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Also, the incorporation of nanomaterials into composite electrodes has been demonstrated to be an extremely useful strategy to construct electrochemical biosensors with an improved analytical performance. These biosensors exhibit both the characteristics of the involved nanomaterials and the advantages derived from the use of composite electrodes, such as their regeneration ability and the great versatility that they offer as a consequence of the feasible incorporation of different substances into the bulk of the electrode matrix by simple physical inclusion. Although there are several antecedents in the literature in which the advantageous features of colloidal gold and carbon paste electrode fabrication have been combined (3), this chapter is focussed on composite electrode matrices constructed using Teflon as nonconducting-binding material. Details on the preparation of several electrochemical biosensors following this methodology as well as on their analytical performance and applicability are given below.
2. Materials 2.1. Composite Electrode Matrices
1. Graphite powder (99.997% purity; Goodfellow, Cambridge Ltd., Huntingdon UK). 2. Teflon (poly(tetrafluorethylene) powder (> 40 μm, Aldrich, Saint Louis, Missouri, USA). 3. Multi-wall carbon nanotubes (MWCNTs, 30 ± 15 nm ø), with a 95% purity (NanoLab, Brighton, MA).
2.2. Colloidal Gold
1. HAuClO4 3H2O (>49% as Au, Sigma, Saint Louis, Missouri, USA). A 1% (w/w) solution was made with Milli-Q water. 2. Sodium citrate (99.0% purity, Sigma). A 1% (w/v) solution was prepared in Milli-Q water. 3. Water is obtained from a Milli-Q purification system (Millipore, Bedford, NA, USA).
2.3. Tyrosinase Biosensor
1. Tyrosinase, Tyr (from mushroom, EC.1.14.18.1, activity of 2,590 units per mg of solid) (Sigma). The enzyme was used as received. 2. Graphite and Teflon powder are the same described in Subheading 2.1. 3. Catechol (Sigma, 99%), phenol (Sigma, 99%) 3,4-dimethylphenol (Aldrich, 98%), 4-chloro-3-methylphenol (Aldrich, 99%), 4-chlorophenol (Aldrich, ≥99%), 4-chloro-2-methylphenol (Aldrich, 99%), 3-methylphenol (Aldrich, 99%), and
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4-methylphenol (Aldrich, 99%). 1.0 × 10−2 M solutions of cathecol, phenol, 4-chlorophenol, and 4-chloro-2-methylphenol are prepared in 0.1 M phosphate buffer of pH 7.4 (prepared from NaH2PO4 2H2O and Na2HPO4, ≥ 99%, Scharlab, Barcelona, Spain). 1.0 × 10−2 M solutions of 3,4-dimethylphenol, 4-chloro-3-methylphenol, 3-methylphenol, and 4-methylphenol are prepared in methanol (Scharlab, gradient HPLC grade, 99.99%). More diluted standards were prepared by suitable dilution with the 0.1 M phosphate buffer solution. 2.4. Progesterone Immunosensor
1. Monoclonal anti-progesterone (anti-Prog) (clone 2H4 rat cell culture supernatant, Sigma). A 0.01 mg/mL solution is prepared in Tris (tris(hydroxymethyl)aminomethane) (Sigma, 99%) buffer solution of pH 7.0. 2. Progesterone labeled with alkaline phosphatase (AP-Prog) (Ridgeway Science Ltd., Alvington, Gloucestershire, UK) was prepared by reaction of 9 mg of alkaline phosphatase with 0.5 mg progesterone glucoronide in a 2 mL final volume. 3. Unlabeled progesterone (Prog) (Aldrich, 98%): A stock 1 mM solution of progesterone is prepared in methanol (Scharlab, gradient HPLC grade, 99.99%) and stored at 4°C. More diluted solutions are prepared daily by dilution with 0.1 M Tris buffer solution of pH 7.0. 4. Tyrosinase, Tyr (from mushroom, EC.1.14.18.1, activity of 2,590 units per mg of solid) (Sigma). 5. Phenyl phosphate disodium salt dihydrate (Sigma, 98%): A 0.01 M phenyl phosphate solution is prepared in 0.1 M Tris buffer of pH 7.0. 6. Phenol (Merck, 99%) 0.01 M phenol solutions are prepared daily in 0.1 M Tris (pH 7.0) buffer solution. 7. Bovine serum albumin (BSA) (97%) and magnesium chloride (99%) are all from Merck. 8. Graphite and Teflon powder are the same as described in Subheading 2.1.
2.5. Glucose Oxidase Biosensor Based on Colloidal Gold–Carbon Nanotubes
1. Glucose oxidase, GOx (Sigma, 151,000 U/g solid).
2.6. Apparatus
1. Electrochemical measurements with the tyrosinase biosensor and the progesterone immunosensor are carried out with a
2. Teflon powder is the same mentioned in Subheading 2.1. 3. Glucose (>99%, Panreac, Barcelona, Spain). Stock 0.5 M solutions are prepared in 0.05 M phosphate buffer, pH 7.4, and let to stand overnight before using to allow equilibration of anomers. More diluted standards were prepared by suitable dilution with the same phosphate buffer solution.
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PGSTAT 12 potentiostat from Autolab (EcoChemie B.V., Utrecht, The Netherlands). The electrochemical software was the general-purpose electrochemical system (GPES). 2. Amperometric measurements in stirred solutions with MWCNTs-colloidal gold composite biosensors were carried out using a PalmSens Electrochemical Sensor Interface (Palm Instruments BV, Houten, The Netherlands) controlled by a Pocket PC Software. 3. A three-electrode cell (a BAS VC-2 10-mL glass electrochemical cell) consisting of a platinum wire counter electrode (BAS MW-1032), a BAS MF-2063 Ag/AgCl/3M KCl reference electrode, and the corresponding composite electrode as the working electrode, was used. All experiments were performed at room temperature.
3. Methods 3.1. Preparation of Colloidal Gold Dispersions
1. 2.5 mL of the 1% sodium citrate solution are added to 100 mL of a boiling aqueous solution containing 1 mL 1% (w/w) HAuCl4. 2. The mixture is allowed to boil for 15 min. 3. Heating is stopped and the mixture is stirred by hand with a glass rod for 15 min. 4. Colloidal gold suspension is left to reach room temperature and stored in dark glass bottles at 4°C until use. The diameter of gold nanoparticles is 16 ± 2 nm.
3.2. Preparation of the Aucoll–Graphite– Teflon Electrodes
A scheme of the fabrication procedure of composite colloidal gold– graphite–Teflon electrode matrices is depicted in Fig. 1. 1. Graphite (150 mg) and colloidal gold (900 μL) suspension prepared as described above are thoroughly mixed using a magnetic stirrer (Selecta, Barcelona, Spain) for 2 h in a dark glass flask. 2. Water is evaporated under air current at room temperature. 3. Teflon powder (450 mg) is added and thoroughly mixed by hand with a glass rod until a homogeneous mixture is obtained. 4. The mixture is pressed into pellets by means of a Carver pellet press (Perkin Elmer, Norwalk, CT, USA) at 10,000 kg/cm for 10 min. Pellets are 1.3 cm diameter and around 0.4 cm thick. 5. From each main pellet, several (five or six) 3.0 mm diameter cylindrical portions are bored with a drill. Each portion constitutes a different composite electrode.
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Fig. 1. The procedure of fabrication of composite Aucoll–graphite–Teflon electrodes.
6. Each electrode is press-fitted into a Teflon holder. 7. The electrical contact was made through a stainless steel screw. 3.3. Tyrosinase Biosensor Based on a Composite Aucoll– Graphite–Teflon Electrode
A tyrosinase bioelectrode with improved stability and sensitivity with respect to other biosensor designs is constructed by incorporation of the enzyme and gold nanoparticles into graphite– Teflon composite electrode matrices by simple physical inclusion. Amperometry at −0.10 V vs. Ag/AgCl for different alkyl and chlorophenols allows the achievement of detection limits of 3 nM for catechol and approximately 20 nM for the rest of phenolic compounds. This high sensitivity is attributed to the presence of colloidal gold in the composite matrix, which gives rise to enhanced kinetics of both the enzyme reaction with tyrosinase and the electrochemical reduction of the corresponding o-quinones at the electrode surface. Moreover, the simple renewability of the electrode surface by polishing, which is an inherent characteristic of the rigid composite bioelectrodes, coupled with the gold nanoparticles ability to attach proteins retaining their biological activity permit the fabrication of reusable and reproducible biosensors exhibiting a lifetime of at least 39 days without apparent loss of the immobilized enzyme activity (4). The composite biosensor was applied to the estimation of the phenolic compounds content in water samples of different origin.
3.3.1. Preparation of the Tyr–Aucoll–Graphite– Teflon Biosensor
1. Steps 1 and 2 of Subheading 3.2 are carried out. 2. 34.75 mg tyrosinase and 400 μL of the 0.1 M phosphate buffer solution of pH 7.4 are incorporated to the mixture by stirring during 2 h with a magnetic stirrer (Selecta) in an ice bath. 3. The resulting homogeneous mixture is dried by passing an Ar stream. 4. Teflon (415.25 mg) is added and thoroughly hand mixed using a glass rod until a homogeneous mixture is obtained.
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The final Teflon percentage is of 70% and provides suitable mechanic and conducting characteristics to the composite material. 5. Steps 4–7 of Subheading 3.2 are accomplished. 3.3.2. Monitoring of Phenolic Compounds in Water Samples
1. Aliquots of 200 μL of undiluted samples are directly added to the electrochemical cell containing 10 mL of 0.1 M phosphate buffer solution of pH 7.4. 2. Amperometric measurements under constant stirring conditions are performed with the Tyr–Aucoll–graphite–Teflon biosensor by applying a potential of −0.10 V and allowing the steady-state current to be reached. 3. The analysis of samples was carried out by applying the standard additions method by injecting appropriate volumes of 1.0 × 10−4 M phenol stock solutions prepared in the abovementioned phosphate buffer. 4. The content of phenolic compounds is expressed as a concentration of phenol.
3.4. Progesterone Immunosensor Using a Tyrosinase–Aucoll– Graphite–Teflon Biosensor as Amperometric Transducer
The synergic combination of the advantages provided by gold nanoparticle nanostructured biosensors and composite electrode matrices into which gold nanoparticles are incorporated by simple physical inclusion allows the development of a progesterone immunusensor in which the antibody is directly attached to the electrode surface, and amperometric transduction is carried out at a colloidal gold–graphite–Teflon–tyrosinase composite biosensor (see procedure below) (5). The immunosensor functioning is schematized in Fig. 2, and it is based on a sequential competitive assay between progesterone (Prog) and the antigen-labeled with the enzyme alkaline phosphatase (AP-Prog) for the binding sites of the immobilized antibodies (anti-Prog). Antibodies are preferentially linked to gold nanoparticles at the electrode surface as a consequence of the gold nanoparticles ability to adsorb proteins retaining their biological activity. Phenyl phosphate is employed as the alkaline phosphatase-substrate, generating phenol which is catalytically oxidized by tyrosinase to the o-quinone. This quinone is electrochemically reduced at −0.10 V at the tyrosinase–colloidal gold composite electrode. As it has been commented in Subheading 3.3, the presence of gold nanoparticles in the composite electrode matrix allows a high sensitivity for the detection of phenol. The calibration graph for progesterone showed a linear range between 0 and 40 ng/mL, with a detection limit of 0.43 ng/mL. Furthermore, the immunosensor lifetime is of 14 days without need of a regeneration procedure. The practical usefulness of the immunosensor was demonstrated by determining progesterone in spiked milk samples
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Fig. 2. The progesterone immunosensor using a colloidal gold–graphite–Teflon–tyrosinase composite biosensor as amperometric transducer.
at the 5.0 and 1.5 ng/mL concentration levels. Using the very simple experimental procedure described below, mean recoveries (n = 7) of 98 ± 3% and 99 ± 3%, respectively, were obtained. 3.4.1. Preparation of the Anti-Prog–Tyr–Graphite– Teflon Immunosensor
1. A Tyr-Aucoll-graphite-Teflon electrode is constructed as described in Subheading 3.3. 2. A 5.2-μL aliquot of the 0.01 mg/mL anti-Prog solution in Tris is deposited on the surface of the Tyr–Aucoll–graphite– Teflon composite biosensor and allowed to dry at air under room temperature. 3. The bioelectrode is immersed for 10 min into a 2% (w/w) BSA solution in 0.1 M Tris buffer of pH 7.0. 4. The biosensor is washed carefully with 0.1 M Tris buffer solution of pH 7.0. 5. The resulting anti-Prog-(Tyr-Aucoll-graphite–Teflon) immunosensor is incubated in 10 mL of the analyte Prog solution for 30 min. 6. The immunosensor is subsequently incubated in 10 mL of the AP-Prog solution mentioned in the item 2 of Subheading 2.4 during 40 min.
3.4.2. Determination of Progesterone with the AntiProg–Tyr–Aucoll–Graphite– Teflon Immunosensor
1. The immunosensor is immersed into the electrochemical cell containing 10 mL of the 0.1 M Tris buffer of pH 7.0 and 1 mM MgCl2. 2. 20 μL of the 0.01 M phenyl phosphate solution are added to the electrochemical cell.
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3. Amperometric measurements under constant stirring conditions of the o-quinone reduction current were carried out at −0.10 V vs. Ag/AgCl and allowing the steady-state current to be reached. 3.4.3. Determination of Progesterone in Milk with the Anti-Prog–Tyr– Aucoll–Graphite–Teflon Immunosensor
1. The immunosensor is immersed in the spiked untreated milk sample for 30 min. 2. Subsequently, it is incubated in the AP-Prog solution during 40 min. 3. Amperometry at −0.10 V is used to monitor the affinity reaction. 4. A calibration graph obtained with spiked milk that does not contain detectable endogenous progesterone is employed for progesterone quantification.
Glucose Oxidase Biosensor Based on Colloidal Gold– Carbon Nanotubes
Hybrid nanoparticles/nanotubes materials have shown to possess interesting properties, which can be profited for the development of electrochemical biosensors. In particular, gold nanoparticles– CNTs hybrids constitute biocompatible materials with important electroanalytical features because of the coupling of the abovementioned capabilities of gold nanoparticles with the electrocatalytic ability of carbon nanotubes toward the electrooxidation of several molecules of biochemical interest. The bulk incorporation of glucose oxidase into colloidal gold–carbon nanotubes composite electrodes using Teflon as the binder allows the construction of a mediator-less glucose oxidase biosensor, GOx–Aucoll–MWCNTs–Teflon, which exhibits improved analytical performance with respect to previous biosensor designs using carbon nanotubes. The biosensor is applied for the rapid determination of glucose in beverages recommended for sport practice (6).
Preparation of Glucose Oxidase–Aucoll–MWCNTs– Teflon Biosensors
The experimental procedure for preparation of glucose oxidase– Aucoll–MWCNTs–Teflon biosensors is schematized in Fig. 3. The successive steps are the following: 1. A colloidal gold dispersion is prepared as described in Subheading 3.1. 2. Colloidal gold (360 μL) is added to 50 mg of solid MWCNTs in a glass vessel and thoroughly hand mixed for 10 min with a glass rod. 3. Water is evaporated under N2 current. 4. Glucose oxidase (1 mg) is incorporated to the composite mixture by stirring with a magnetic stirrer (Selecta) for 20 min. 5. Teflon is incorporated to the mixture by adding 50 mg Teflon powder and thoroughly hand mixing until complete homogenization.
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Fig. 3. The procedure for the construction of glucose oxidase–Aucoll–MWCNTs–Teflon biosensors.
6. Portions of the resulting mixture are packed into Teflon holders (3 mm inner diameter) and pressed tightly. 7. The electrical contact is made through a stainless steel screw. 3.4.4. Determination of Glucose in Commercial Beverages Recommended for Sport Practice
1. Aliquots of the samples are appropriately diluted with 0.05 M phosphate buffer solution of pH 7.4, and transferred to the electrochemical cell containing the bioelectrode. 2. Amperometry in stirred solutions at +0.5 V vs. Ag/AgCl is employed allowing the steady-state current to be reached. 3. Samples analysis is carried out by applying the standard additions method which involved successive addition of 25 μL of a 2.5 × 10−4 M glucose solution.
4. Notes 1. All glassware used in the preparation of the colloidal gold dispersions must be carefully cleaned in a bath of a freshly prepared 3:1 HNO3 + HCl solution, and then thoroughly rinsed with Milli-Q water, previously filtered through a 22-μm microporous Nylon membrane filter (Scharlab), and dried in air. 2. Observation of color changes produced after addition of sodium citrate to the HAuCl4 solution in the procedure used for the preparation of colloidal gold is an appropriate way to verify the adequate progress of the reaction. Initially (i.e.,
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before sodium citrate addition), the tetracholoroauric acid aqueous solution is yellow. Color changes to purple upon addition of sodium citrate solution, and then it slowly changes to cherry red. This final color indicates that a correct colloidal gold dispersion is obtained. 3. An inherent advantage of the biosensor designs based on the use of rigid composite electrodes is the possibility of surface renewability. This regeneration treatment is performed by simple polishing of the electrode surface for approximately 5 s on a 150 grit SiC paper. References 1. Mena, M.L., Yáñez-Sedeño, P., Pingarrón, J.M. (2005) A comparison of different strategies for the construction of amperometric enzyme biosensors using gold nanoparticle-modified electrodes. Anal. Biochem., 336, 20–27 2. Liu, S., Leech, D., Ju, H. (2003) Application of colloidal gold in protein immobilization, electron transfer and biosensing. Anal. Lett., 36, 1–19 3. Liu, S.Q., Ju, H. (2003) Regentless glucose biosensor based on direct electron transfer of glucose oxidase immobilized on colloidal gold modified carbon paste electrode. Biosens. Bioelectron., 19, 177–183 4. Carralero, V., Mena, M.L.,Gonzalez-Cortés, A., Yáñez-Sedeño, P., Pingarrón, J.M. (2006)
Development of a high analytical performancetyrosinase biosensor based on a composite graphite–Teflon electrode modified with gold nanoparticles. Biosens. Bioelectron., 22, 730–736 5. Carralero, V., González-Cortés, A., YáñezSedeño, P., Pingarrón, J.M. (2007) Nanostructured progesterone immunosensor using a tyrosinase-colloidal gold–graphite–Teflon biosensor as amperometric transducer. Anal. Chim. Acta, 596, 86–91 6. Manso, J., Mena, M.L., Yáñez-Sedeño, P., Pingarrón, J.M. (2007) Electrochemical biosensors based on colloidal gold-carbon nanotubes-Teflon composite electrodes. J. Electroanal. Chem., 603, 1–7
Chapter 12 Immunochromatographic Lateral Flow Strip Tests Gaiping Zhang, Junqing Guo, and Xuannian Wang Summary The immunochromatographic lateral flow strip test is a one-step test that facilitates low-cost, rapid identification of various analytes at the point of care. We have developed lateral flow strip tests for the specific qualitative or semiquantitative detection of antigens, antibodies, and haptens, such as drug residues. Here, we describe in detail the preparation of three examples of the strip tests for detection of (a) the infectious bursal disease virus; (b) Trichinella specific antibodies, and (c) Clenbuterol residues in urine samples. Key words: Immunochromatography, Lateral flow strip test, Infectious bursal disease virus, Trichinella, Clenbuterol.
1. Introduction The membrane-based immunochromatographic lateral flow strip test (ILFST) represents a well-established and appropriate technique for a variety of point-of-care and field-use applications (1, 2), which was initially developed as a convenient test for pregnancy (3). As the technique eliminates the need for trained personnel and expensive equipment, ILFST has become a popular platform for the development of rapid tests since their introduction in the late 1980s. The technique is widely used for the specific qualitative or semiquantitative detection of many classes of analytes including antigens, antibodies, haptens, and even oligonucleotides (4–10). Some of the more common ILFST currently available commercially are those for pregnancy, infections including Streptococcus, Chlamydia, human immunodeficiency virus (HIV), and Hepatitis C virus (HCV).
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ILFST is based on the principles of immunochromatography. A test strip typically consists of porous materials in four zones containing different reagents: a sample application pad, a conjugate pad containing colloidal gold conjugate, a detection membrane containing narrow absorbed bands of proteins as test and control lines, and an absorbent pad (Fig. 1). For the convenience of storage and handling, the porous materials are laminated with a semirigid material of appropriate mechanical strength. The top is partially covered with a thin plastic material so as to leave a portion of the sample pad exposed for sample application (4). When applied to the sample pad, the liquid sample migrates by capillary diffusion through the conjugate pad, rehydrates the gold conjugate, and the analyte interacts with the conjugate. The complex of gold conjugate and analyte then moves onto the membrane toward the capture target, where it becomes immobilized and concentrated, producing a distinct signal in the form of a sharp red line. A second line, the control line, may also be formed on the membrane by the trapping of excess gold conjugate, indicating the test is complete (11). Sample flow Sample pad
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A Sample pad Cover film
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B Fig. 1. The lateral flow strip structure: (A) Schematic representation of the lateral flow strip. A lateral flow strip typically consists of four zones of a sample pad, a conjugate pad, a membrane containing the test and control lines, and an absorbent pad on a backing plate. (B) Lateral (top) and over (bottom) views of a lateral flow strip. The outmost layers of the ends of the strip are plastic cover films.
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A Positive Test line (+)
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: Blotted antibody/SPA;
: Analyte
B Positive Test line (−)
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: Blotted antibody;
: Blotted or free Hapten
Fig. 2. Principles of the lateral flow strip test. (A) Sandwich format: This format is used when testing for larger analytes with multiple binding sites, such as viruses and antibodies. In the case of detecting antigens, target analytes in the sample effluent are recognized by the antibody conjugate forming analyte–antibody complexes and bound to the immobilized antibody on the test line while the excess conjugate is trapped by anti-mouse IgG antibody on the control line, formation of two red colored lines on the membrane indicates a positive result. In the case of detecting antibodies, targets in the sample effluent are bound with conjugated antigens forming analyte–antigen complexes and immobilized by SPA or anti-immunoglobulin antibodies on the test line, while the excess conjugate is trapped by antibodies that recognize the conjugated antigen on the control line, formation of two red colored lines on the membrane indicates a positive result. (B) Competitive format: This is used most often when testing for small molecules with a single antigenic determinant. In this format, free analytes in the sample solution compete with the immobilized analytes on the test line to bind colloidal gold conjugated antibodies at a defined concentration. The dense of the test line is dependent on the concentration of free analytes present in the test samples.
The two predominant formats used in ILFST are the sandwich and competitive reaction schemes (12). These can be best explained graphically as shown in Fig. 2. The test strip for human chorionic gonadotropin (hCG) is a typical sandwich assay, and those for drugs of abuse are competitive or inhibition assays. In serum assays, specific antibodies are detected as an indicator of
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Reagents
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Fig. 3. Outline of a lateral flow strip manufacturing process.
various disease states. Here, we describe three types of ILFST for the detection of pathogens, specific antibodies, and haptens using infectious bursal disease virus (IBDV) (5), Trichinella (7), and Clenbuterol (CL) (9) as examples. The general manufacturing process for production of test strips includes the preparation of colloidal gold conjugates, application of reagents onto the membrane and pads, lamination of the membrane, conjugate pad, sample pad, and absorbent pad onto a support backing, cutting the prepared master cards into strips of defined length and width, and strip packaging (Fig. 3).
2. Materials 2.1. Devices
1. The XYZ 3000 platform with BioJet Quanti 3000 and AirJet Quanti 3000 dispensers (BioDot Inc., Irvine, CA) is used for membrane blotting and conjugate dispensing. 2. OV2000 Forced Air Drying Oven (BioDot Inc.) is used for drying the blotted membrane and conjugate pad. 3. LM5000 Clamshell Laminator (BioDot Inc.) is used for assembly of the master card of test strips. 4. CM4000 Guillotine Cutter (BioDot Inc.) is used to cut the assembled master cards into strips.
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1. Nitrocellulose Membrane: The Hi-Flow Plus membrane HF180 (Millipore Corp., Bedford, MA; Cat. No. SA3J044106) is used (see Note 2). 2. Fiberglass (Millipore; Cat. No. GFCP203000) is used for making sample and conjugate pads. 3. Filter paper (Millipore; Cat. No. FP10102500) is used for absorbent pads. 4. Double-sided adhesive tapes (G&L Precision Die Cutting, Inc. San Jose, Cat. No. GL-187) are used for making adhesive layers on the backing plate. 5. Plastic films with different colors are used for strip covers. 6. Plastic boards are used for strip backing plate.
2.3. Stock Solutions
1. Hydrogen tetrachloroaurate (1% (w/v) ): 0.1 g of HAuCl4 (Sigma-Aldrich, St. Louis, MO; Cat. No. G4022) is dissolved in 10 mL of triple-distilled water and stored in a brown bottle. 2. Trisodium citrate (1% (w/v) ): 0.5 g of Na3C6H5O7·2H2O (Fluka, Buchs, Switzerland; Cat. No. 71402) is dissolved in 50 mL of triple-distilled water. 3. NaCl (1 M): 5.8 g of NaCl is dissolved in 100 mL of tripledistilled water. 4. K2CO3 (0.2 M): 2.8 g of K2CO3 (Fluka; Cat. No. 60108) is dissolved in 100 mL of triple-distilled water. 5. Sodium borate (20 mM): 0.76 g of Na2B4O7·10H2O (Fluka; Cat. No. 71997) is dissolved in 100 mL of triple-distilled water. 6. Phosphate buffered saline (PBS): 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4 are dissolved in 1,000 mL of triple-distilled water.
2.4. Reagents 2.4.1. ILFST for Detection of IBDV Antigens
1. Conjugate reagent: The anti-IBDV monoclonal antibody (mAb) is purified by a protein A column from ascitic fluids of mice carrying the specific hybridoma (5), and stored at −20°C in 1 mL aliquots. 2. Test line reagent: A second anti-IBDV mAb IgG recognizing a different epitope of the viral protein is similarly purified from ascitic fluid (5), and stored at −20°C in 1 mL aliquots. 3. Control line antibody: The goat anti-mouse IgG is isolated from sera of goats immunized with mouse IgG and stored at −20°C in 1 mL aliquots.
2.4.2. ILFST for the Detection of Anti-Trichinella Antibodies
1. Conjugate reagent: The excretory-secretory (ES) antigens are purified from the supernatants of cultured Trichinella muscle larvae (7), and stored at −70°C in 1 mL aliquots.
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2. Test line reagent: Protein A, Staphylococcus aureus (SPA, Merk, Darmstadt, Germany; Cat. No. 539202) is dissolved in PBS (pH 7.2) at 10 mg/mL, stored at −20°C in 1 mL aliquots. 3. Control line reagent: The goat anti-ES IgG is isolated from sera of goats hyperimmunized with ES antigens (7) and stored at −20°C in 1 mL aliquots. 2.4.3. ILFST for Detection of Clenbuterol Residues
1. Conjugate reagent: The anti-CL mAb IgG is purified from ascitic fluid (9), and stored at −20°C in 1 mL aliquots. 2. Test line reagent: CL (Sigma-Aldrich; Cat. No. C5423) is conjugated to bovine serum albumin (BSA, Amresco Inc., Solon, Ohio, Cat. No. 0332) and used as the test line reagent (BSA-CL) (9). Briefly, 5 mg of CL dissolved in 4 mL of 0.01 M HCl (cooled to 0–5°C) is mixed with 10 mg sodium nitrite. After stirring for 6 h at 4°C, 20 mg BSA predissolved in 2 mL of 0.1 M phosphate buffer (pH 8.6) is added and the mixture stirred for a further 6 h at 4°C. The solution is kept at room temperature (RT) overnight and then conjugates are purified by filtration on Sephadex G-50 or dialyzed against PBS. 3. Control line reagent: The goat anti-mouse IgG is isolated by protein A column as above from sera of goats immunized with mouse IgG and stored at −20°C in 1 mL aliquots.
3. Methods 3.1. ILFST for Detection of IBDV Antigens 3.1.1. Preparation of Colloidal Gold Suspension
1. Add 1 mL of 1% (w/v) HAuCl4 solution in 100 mL of tripledistilled water to a clean 500 mL Erlenmeyer flask on a stirring hot plate and bring the solution to a boil (see Note 3). 2. To the boiling solution, quickly add 1 mL of 1% (w/v) of trisodium citrate under constant stirring. 3. After the color of the solution has changed from blue to dark red (within 3 min), continue to boil the solution for a further 2 min. 4. When the solution cools to RT, add triple-distilled water up to the original volume, stopper the flask to prevent evaporation, air circulation, and entry of contaminants and store the sample away from light. 5. Scan the optical density of the red solution between 500 and 600 nm, and the absorption maximum should be at 525 nm, indicating that the gold particles have an average diameter of 40 nm (see Note 4). 6. The colloidal gold suspension can be stored at RT in the dark for several months.
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1. Adjust the pH of the colloidal gold suspension to 9.0 with 0.2 M K2CO3 (see Note 5). 2. Dialyze the anti-IBDV mAb IgG against 100 volumes of 20 mM sodium borate for 24 h at 4°C with three changes of solution over this period. 3. Add 2 mL of the IgG solution at the predetermined optimum concentration (see Note 6) into 10 mL of the colloidal gold solution (pH 9.0), rapidly mix, and then incubate for 20–40 min at RT. 4. Add 1/10 volume of 10% (w/v) BSA in 20 mM sodium borate. Mix rapidly upon addition and incubate for 10–15 min at RT (see Note 7). 5. Centrifuge for 30 min at 15,000 × g at 4°C and carefully discard the supernatant. 6. Resuspend the pellet in 20 mM sodium borate containing 1% (w/v) BSA. 7. Centrifuge again, discard supernatant, and resuspend the pellet in 20 mM sodium borate containing 1% (w/v) BSA. 8. Wash one more time and finally resuspend the conjugate in 1 mL of 20 mM sodium borate containing 1% (w/v) BSA.
3.1.3. Membrane Blotting
1. Cut nitrocellulose membranes into 2.5 × 30 cm2 tapes (see Note 8). 2. Dilute the other anti-IBDV mAb IgG and goat anti-mouse IgG to 1 mg/mL in PBS (pH 7.2) for the test and control lines, respectively (see Note 9). 3. Put membrane tapes on the XYZ 3000 platform (Fig. 4). Dispense the solutions onto the membranes at 1 μL/cm as test Syringe pumps
Syringe pumps
Biojet Quanti 3000 Membrane
XYZ3000 Platform
Fig. 4. XYZ 3000 platform with BioJet Quanti 3000.
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and control lines in parallel using the BioJet Quanti 3000. The test and control lines are located at the center of the membrane with a space of 0.5 cm. 4. Dry the bloted membranes at 42°C for 30 min in the OV2000 Forced Air Drying Oven. 5. Seal the membranes in a plastic bag in the presence of desiccants and store at 2–8°C (see Note 10). 3.1.4. Preparation of Conjugate Pads
1. Cut fiberglass into 1.5× 30 cm2 strips (see Note 11). 2. Add 1 mL of the conjugate in 2 mL of 20 mM sodium borate (pH8.0) containing 2% (w/v) BSA, 3% (w/v) sucrose, 0.6 M NaCl, 0.2% (v/v) Tween 20, and 0.1% (w/v) sodium azide. 3. Put the fiberglass strips on the XYZ 3000 platform. Dispense the conjugate onto the fiberglass at 15 μL/cm using the AirJet Quanti 3000. 4. Dry the conjugate pads at 50°C for 30 min using the OV2000 Forced Air Drying Oven. 5. Seal the strips with desiccants in a plastic bag and store at 2–8°C (see Note 12).
3.1.5. Preparation of Sample Pads
1. Cut fiberglass into 1.5 × 30 cm2 strips (see Note 11). 2. Soak the strips in PBS (pH 7.2) containing 0.1 M NaCl, 0.2% (v/v) Tween 20, and 0.1% (w/v) sodium azide (see Note 13). 3. Dry the strips at 50°C for 30 min using the OV2000 Forced Air Drying Oven. 4. Seal the sample pads with desiccant in a plastic bag and store at RT.
3.1.6. Preparation of Absorbent Pads
3.1.7. Preparation of Adhesive Cards
1. Cut filter paper in 2.5 × 30 cm2 strips. 2. Seal the absorbent pads with desiccant in a plastic bag and store at RT. 1. Put double-sided adhesive tapes on one side of the support cards (see Note 14). 2. Cut the adhesive board into 7.5 × 30 cm2 strips to make adhesive cards.
3.1.8. Assembly of Master Card
1. Using a LM5000 Clamshell Laminator or manually, the precut materials are assembled into a lateral flow strip master card (Fig. 5). 2. As illustrated in Figs. 1 and 5, the blotted membrane is stamped in the middle of the adhesive backing card. The conjugate and sample pads are affixed sequentially next to the membrane with a 1–2 mm overlap at the sample end, and the
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Fig. 5. Assembly of the lateral flow strip master card. (A) Components for the strip lamination: 1, white cover film with arrows (sample end); 2, sample pad; 3, conjugate pad; 4, blotted membrane; 5, absorbent pad; 6, blue cover film (distal end); 7, adhesive backing plate. (B) The blotted membrane, conjugate, sample and absorbent pads are put in the correspondent places on the adhesive backing plate, sequentially as described in the text. (C) A master card. The sample and distal ends are covered with the white and blue films, respectively.
absorbent pad to distal end of the membrane with 1–2 mm overlap (see Note 15). 3. Cover the sample and conjugate pads at the sample end, and the absorbent pad at the distal end with different color films, respectively. 3.1.9. Cutting and Packaging
1. Cut the assembled master card into 0.3 cm strips using a CM4000 guillotine cutter. 2. Seal the test strips with desiccants in a plastic package and store at 2–8°C (see Notes 10 and 12).
3.1.10 Detection of IBDV Antigens
1. Simply break up the bursal tissue in a whirl pack with finger massage, and dilute in PBS or water. 2. Dip the IBDV antigen test strip into the solution for 10–20 s at RT. 3. Take the strip out and place horizontally for 1–2 min to observe the result. 4. If both the test and control lines turn red, the sample is recorded as positive, indicating the presence of IBDV antigens in the sample. When the control line but not the test line is colored, it is considered as negative. No colored band appears on the membrane indicates an improper testing procedure or deterioration of the strip, and the test should be repeated using a new strip (Fig. 6A).
3.2. ILFST for Detection of Trichinella Antibodies
The protocol is similar to that described above. Differences are clarified as below.
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Fig. 6. Detection of analytes using the lateral flow strips: (A) Result of IBDV antigen detection; (B) Result of anti-ES antibody detection; (C) Result of CL residue detection.
3.2.1. Conjugation of ES Antigens with Colloidal Gold
1. Adjust the pH of the stock colloidal gold solution to 8.5 with 0.2 M K2CO3. 2. Dialyze the ES antigens against 20 mM sodium borate at 4°C. 3. Prepare the ES-conjugate as described in Subheading 3.1.2.
3.2.2. Membrane Blotting
1. Dilute the SPA and goat anti-ES IgG solutions to 1 mg/mL in PBS (pH7.2). 2. Filter the solutions using 0.22 μm filters. 3. Dispense the SPA and goat anti-ES IgG solutions as test and control line reagents, using the BioJet Quanti 3000 on 2.5 × 30 cm2 nitrocellulose membranes as described in Subheading 3.1.3.
3.2.3. Detection of Trichinella Antibodies
1. Dilute blood or serum samples of human, pigs, cats, or dogs in PBS or water at 1:100. 2. Dip the ES-antibody test strip into the solution for 10–20 s at RT. 3. Take the strip out and place horizontally for 1–2 min to observe the result.
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4. If both test and control lines appear in red, the sample is recorded as positive, indicating the presence of antibodies specific to ES antigens. Appearance of only the control line but not the test line is considered as negative. No colored line appearing on the membrane indicates an improper testing procedure or deterioration of the strip, and the test should be repeated using a new strip (Fig. 6B). 3.3. ILFST for Detection of CL Residues
The protocol is similar to that described in Subheading 3.1, and differences are clarified as below. In this case, anti-CL mAb IgG is conjugated with colloidal gold and the hapten conjugate, BSACL, is used on the test line. A positive standard sample of the hapten is required for the quantitive detection.
3.3.1. Conjugation of Anti-CL mAb IgG with Colloidal Gold
1. Adjust the pH of the stock colloidal gold to 9.0 with 0.2 M K2CO3. 2. Dialyze anti-CL mAb IgG against 20 mM sodium borate at 4°C. 3. Prepare the CL antibody conjugate as described in Subheading 3.1.2.
3.3.2. Membrane Blotting
1. Dilute the BSA-CL and goat anti-mouse IgG to 1 mg/mL in PBS (pH7.2) for application as the test and control line capture reagents, respectively. 2. Filter the solutions using 0.22 μm filters. 3. Dispense the test and control line solutions using the BioJet Quanti 3000 on 2.5 × 30 cm2 nitrocellulose membranes as described in Subheading 3.1.3.
3.3.3. Preparation of Standard Reference Solutions
1. Dissolve 10 mg CL in 10 mL PBS (pH 7.2). 2. Dilute the CL solution with PBS to the final concentration of 1 ng/mL as the positive standard. 3. PBS is used as the negative standard.
3.3.4. Detection of CL Residues in Urine Samples
1. Dip CL test strips into urine samples and standard solutions for 10–20 s at RT, respectively. 2. Take the strips out and place horizontally for 5 min to observe the result. 3. Both control and test lines develop on the strip to which applied the negative standard. The test line is invisible and the control appears on the strip to which applied the positive standard. When testing urine samples, if the test line develop equal intensity to that seen with the negative standard, the sample is recorded as negative indicating no detectable level of CL. If the intensity of the test line is weaker than that of the negative standard, the sample is considered as positive
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and indicates the presence of CL residues at a concentration of less than 1 ng/mL; An invisible test line indicates that the concentration of CL residues at 1 ng/mL or higher. No colored bands appearing on the membrane indicates an improper testing procedure or deterioration of the strip, and the test should be repeated using new strips (Fig. 6C).
4. Notes 1. Materials for the lateral flow strips including membranes, fiberglass, and filter paper are commercially available from several manufactures, such as Millipore Corp., Schleicher & Schuell (S&S, Keene, NH), Whatman, Inc. (Clifton, NJ), and Pall Biosciences (East Hills, NY) etc. 2. The membrane is the most important material in a lateral flow strip. Protein-binding capacity and capillary flow rate are the most critical parameters to be considered for the membrane selection, and these are determined by polymer composition, pore size, porosity, thickness, etc. Membrane manufacturers generally offer a wide variety of material types and pore sizes and trial investigations should be undertaken to find the optimal membrane for a specific use. In practice, nitrocellulose membranes with the pore sizes of 5–15 μm and supplied as supported and unsupported forms are the most commonly used (13). 3. Containers for colloidal gold must be cleaned thoroughly. It is best that all glassware be sliconized and autoclaved before each use. It is recommended that all solutions be filtered through a 0.22 μm filter before use. Caution: The flask will be hot. Be sure to wear thermal gloves while handling. 4. The size of gold colloidal particles is directly dependent on the amount of trisodium citrate used in its preparation process, and decreases with increasing amounts of sodium citrate (Table 1). The procedure developed by Frens (14) is most commonly used to produce 40-nm gold particles, which are considered to be optimal for lateral flow tests. High quality gold particles should be monodisperse and spherical, and should include less than 5% uneven shapes (15). Colloidal gold solutions from reputable suppliers can be used to ensure that compliance with stringent quality control procedures is met. 5. In general, a protein maximally adsorbs on the gold surface at the isoelectric point (pI) of the molecule or 0.5 pH units higher (16). Therefore, the pH value of colloidal gold solution should be adjusted slightly higher than pI value of proteins prior to conjugation.
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Table 1 Sizes of gold particles correlated with addition of trisodium citrate Volumes (mL) of 1% (w/v) trisodium citrate added in 100 mL of HAuCl4 solution 0.30
0.45
0.70
1.00
1.50
2.00
Color
Blue-grey
Violet
Purple
Red
Orange-red
Orange
Absorption peak (nm)
220
240
535
525
522
518
Diameter (nm)
147
97.5
71.5
41
24.5
15
6. The optimum ratio of protein concentration to colloidal gold solution is determined prior to conjugation by the following steps: (1) Serially twofold dilute antibody solution with 25 μL of distilled water in microplate wells; (2) Add 125 μL of colloidal gold solution to each well; (3) Incubate for 5 min at RT; and (4) Add 125 μL of 1 M NaCl solution to each sample. The color of samples gradually changes from brilliant red to blue as the concentration of protein decreases. The highest dilution of the solution with no change of color contains the optimum protein for colloidal gold labeling. 7. Following the conjugation of the specific protein with colloidal gold, the conjugate must then be stabilized with a suitable agent, such as BSA, gelatin, polyethylene glycol (PEG), or casein. The stabilizer can reduce nonspecific interactions by blocking any sites on the colloidal surface that are not occupied by the specific protein, and helps provide a more-stable suspension (11). 8. There are manufacturers, such as Millipore, S&S, providing cut membranes in defined width for lateral flow strips. 9. The blotting solutions should be filtered through 0.22 μm filters to prevent clogging of the dispensers. 10. It is important to prevent rehydration of the blotted membrane. 11. There are manufacturers, such as Millipore, S&S, providing cut fiberglass in defined width and length for lateral flow strips. 12. It is important to keep the conjugate pad dry. If the pad is exposed to high humidity, water can complex with the sugar molecules, converting them to syrup and delaying particle resolubilization (13). 13. The concentrations loaded into the pad will be determined by the bed volume of the material, the volume of sample analyzed in the strip, and the variability of the samples (13).
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14. The backing materials used in test-strip assembly are generally polyester, styrene, or polyvinyl chloride (PVC) with an adhesive layer on one side. The pressure-sensitive adhesive has proven to be a reliable means of bonding together the component materials. Selection of an appropriate type of adhesive that offers the best compromise between bond strength and the extent of adhesive migration is important to improve processing and increase the shelf-life of strips. Adhesive migration especially during long periods of shelf storage can result in blocked pores, hydrophobic patches, and material rewetting problems that may interfere with the performance of the test (17). There are commercial backing cards. 15. It is very important that the extent of the overlaps be consistent so that the flow dynamics are uniform on all of the strips manufactured.
Acknowledgments This work was supported by grants from the National Basic Research Program of China (Grant No. 2005CB523203) and the Chinese High Technology Research and Development Project (Grant No. 2001AA249031). The authors thank Dr. Norman A. Gregson at King’s College, University of London for his critical review of this manuscript. References 1. Paek, S. H., Lee, S. H., Cho, J. H., and Kim, Y. S. (2000) Development of rapid one-step immunochromatographic assay. Methods 22, 53–60 2. O’Farrell, B., and Bauer, J. (2006) Developing highly sensitive, more-reproducible lateral-flow assays. Part 1: New approaches to old problems. IVD Technol. 7(6), 41–50 3. May, K. (1991) Home tests to monitor fertility. Am. J. Obstet. Gynecol. 165, 2000–2002 4. Huang, C., and Fan, E. (1998) One-step immunochromatographic device and method of use. U.S. Pat. 5,712,172 5. Zhang, G. P., Li, Q. M., Yang, Y. Y., Guo, J. Q., Li, X. W., Deng, R. G., Xiao, Z. J., Xing, G. X., Yang, J. F., Zhao, D., Cai, S. J., and Zang, W. M. (2005) Development of a one-step strip test for the diagnosis of chicken infectious bursal disease. Avian Dis. 49, 177–181
6. Ketema, F., Zeh, C., Edelman, D. C., Saville, R., Constantine, N. T. (2001) Assessment of the performance of a rapid, lateral flow assay for the detection of antibodies to HIV. J. Acquir. Immune Defic. Syndr. 27, 63–70 7. Zhang, G. P., Guo, J. Q., Wang, X. N., Yang, J. X., Yang, Y. Y., Li, Q. M., Li, X. W., Deng, R. G., Xiao, Z. J., Yang, J. F., Xing, G. X., and Zhao, D. (2006) Development and evaluation of an immunochromatographic strip for trichinellosis detection. Vet. Parasitol. 137, 286–293 8. Mapes, J. P. (2002) Indirect label assay device for detecting small molecules and method of use thereof. U.S. Pat. 6, 376,195 9. Zhang, G. P., Wang, X. N., Yang, J. F., Yang, Y. Y., Xing, G. X., Li, Q. M., Zhao, D., Chai, S. J., and Guo, J. Q. (2006) Development of an immunochromatographic lateral flow test strip for detection of β-adrenergic agonist
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11.
12.
13.
Clenbuterol residues. J. Immunol. Methods. 312, 27–33 Oku, Y., Kamiya, K., Kamiya, H., Shibahara, Y., Ii, T., and Uesaka, Y. (2001) Development of oligonucleotide lateral-flow immunoassay for multi-parameter detection. J. Immunol. Methods 258, 73–84 Chandler, J., Gurmin, T., and Robinson, N. (2000) The place of gold in rapid tests. IVD Technol. 6(2), 37–49 Weiss, A., (1999) Concurrent engineering for lateral-flow diagnostics. IVD Technol. 5(7), 48–57 Rapid lateral flow test strips: considerations for product development. (Bedford, MA: Millipore Corp., 2002)
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14. Frens, G. Controlled nucleation for the regulation of the particle size in monodisperse gold solution. Nat. Phys. Sci. 1973; 241: 20–22 15. Carney, J., Braven, H., Seal, J., and Whitworth, E. (2006) Present and future applications of gold in rapid assays. IVD Technol. 12(2), 41–49 16. Albrecht, R. M., Simmons, S. R., and Pawley, J. B. (1993) in Immunocytochemistry: A Practical Approach (Beesley, J. E., Ed.), pp. 151–176, Oxford University Press, Oxford 17. Jones, K. D., and Hopkins, A. K. (2000) Effects of adhesive migration in lateral-flow assays. IVD Technol. 6(9), 57–63
Chapter 13 Liposome-Enhanced Lateral-Flow Assays for the Sandwich-Hybridization Detection of RNA Katie A. Edwards and Antje J. Baeumner Summary Clinical and environmental analyses frequently necessitate rapid, simple, and inexpensive point-of-care or field tests. These semiquantitative tests may be later followed up by confirmatory laboratory-based assays, but can provide an initial scenario assessment important for resource mobilization and threat confinement. Lateral-flow assays (LFAs) and dip-stick assays, which are typically antibody-based and yield a visually detectable signal, provide an assay format suiting these applications extremely well. Signal generation is commonly obtained through the use of colloidal gold or latex beads, which yield a colored band either directly proportional or inversely proportional to the concentration of the analyte of interest. Here, dyeencapsulating liposomes as an alternative are discussed. The LFA biosensors described in this chapter rely on the sandwich-hybridization of a nucleic acid sequence-based amplified (NASBA) mRNA target between a membrane immobilized capture probe and a visible dye (sulforhodamine B)-encapsulating liposome conjugated reporter probe. Although the methodology of this chapter is focused on LFAs for the detection of RNA through sandwich hybridization, the information within can be readily adapted for sandwich and competitive immunoassays. Included are an introduction and application notes toward this end. These include notes ranging from the detection of nonamplified RNA and single-stranded DNA, conjugation protocols for antibodies and other proteins to liposomes, and universal assay formats. Key words: Lateral-flow assay, Liposome, RNA, DNA.
1. Introduction Clinical and environmental analyses frequently necessitate rapid, simple, durable, and inexpensive point-of-care or field tests. Lateral-flow assays (LFAs), such as those used in pregnancy tests, often fulfill these requirements. These assays rely on the appearance of a colored band, the intensity of which corresponds to the concentration of the target analyte in the sample. The goal for Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_13
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such assays is typically a qualitative “yes” or “no” answer, though some assays can provide semiquantitative results. LFAs are commercially available for the detection of a wide variety of analytes. These targets include pathogenic organisms such as Legionella pneumophilia (1, 2), Leptospira (3), Cryptosporidium parvum, and Giardia lamblia (4); viruses such as HIV (5), influenza (6), and respiratory syncytial virus (RSV) (7, 8); food allergens (9) and drugs of abuse (10); health states such as pregnancy, ovulation, or menopause (11, 12); diseases such as prostate cancer (13, 14); or biological toxins such as botulinum toxin (15, 16), and verotoxins (17, 18). Some LFAs have proven to be equivalent or better than enzyme-linked immunosorbant assays (ELISAs), direct fluorescent antibody testing, and viral cultures (5, 6, 8, 19). LFAs used in a clinical setting can obviate the need for patients to return to the clinician’s office for results (20) and can provide guidance for the administration of chemoprophylaxis after potential occupational exposure to viruses (21). In environmental or occupational settings, LFAs provide an initial assessment for mobilization of resources and containment of potential threats. LFAs for research and more fundamental applications such as probe selection, PCR-product identification, genomic sequence search, and identification of single-nucleotide polymorphisms have been suggested (22–24). Here we discuss in detail the use and preparation of LFAs for RNA detection, and the use of liposomes as signaling species in contrast to generally used latex particles and colloidal gold. 1.1. Membrane Preparation
The immobilized recognition element in nucleic acid LFAs is typically either a DNA capture probe, which is complementary to a sequence on the target (25), an antibody against one of the common probe labels such as biotin, fluorescein, or DNP (22, 24), or streptavidin or avidin, which can be used to bind biotinylated capture probes (23, 26, 27). The proteins in the protein-mediated immobilization of DNA probes are larger and bear more functional groups for interaction with the membrane resulting easily in an irreversible immobilization. In addition, the advantage to using protein-based approaches rather than direct immobilization of probe is a more universal detection scheme and allows for proper spatial orientation of capture probes. However, the latter simplifies and reduces the cost of the membrane preparation process as a protein or antibody is not needed, allows for more multiplexed analyses, and can increase assay sensitivity (24, 28). Multianalyte detection can be attained in LFAs through spatial separation of capture zones (29, 30). Examples include differentiation of Dengue virus serotype (30), immune response to disease pathogens (HCV, HIV, and TB) (31), simultaneous detection of insecticides (32), and verotoxin type identification alongside verotoxigenic E. coli (33).
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Commonly, nitrocellulose membranes are used for LFAs because of their high binding capacity, low cost, and wide availability (34). The mechanism behind protein binding to nitrocellulose is not fully understood, but is believed to be noncovalent through hydrophobic, hydrogen bonding, and electrostatic interactions (35, 36). The physical adsorption of nucleic acid probes through drying to nitrocellulose is believed to result in attachment through hydrophobic interactions, an effect that may be enhanced through modification with a poly-T tail (37). The ideal properties of the membrane include particle and pore size consistency, hydrophilicity, high protein binding, and durability. The membrane pore size is inversely proportional to the surface area available for binding and is proportional to the wicking rate (38). A fast wicking rate allows for more rapid results and lower background, but may adversely affect assay sensitivity (39) since less time for interaction between the target and the probes is available. We have also had success using polyethersulfone membranes for nucleic acid sandwich hybridization assays (22, 23, 40). For ease of handling, durability, and avoidance of adhesive interactions, a supported membrane should be chosen. These membranes are manufactured by directly casting the porous membrane material onto a plastic (polyester or cellulose acetate) backing. Alternatively, nonsupported membranes can be laminated manually prior to use. LFAs may be made more durable and user-friendly by encasing them in an injected molded plastic housing such as that shown in Fig. 1. However, we have found this to be unnecessary for laboratory use provided that a supported membrane is used.
Fig. 1. Lateral flow sandwich-hybridization assay membranes: Nitrocellulose membranes with capture probe immobilized 1.5 cm from the base of the membrane were used for the capture of target RNA using dye-encapsulating liposomes for detection. In the presence of target RNA, a sandwich hybridization complex forms with dye-encapsulating liposomes resulting in a magenta-colored band at the capture zone that is proportional to target concentration (top). In the absence of target, no band is visible as no sandwich complex has formed (bottom). Though not necessary, these LFA membranes were housed in more user-friendly packaging using a plastic cassette made using injection molding and designed for addition of liposome/target mixture in hole #1 and addition of running buffer in hole #2. Readings can be taken in the “R” hole.
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In addition, a sample pad, conjugate release pad, and wicking material are often utilized for LFAs; however, we have found these to be unnecessary for the assays described in this chapter. The biorecognition element can be applied to the membrane either manually using a pipettor, with a thin-layer chromatography (TLC) applicator, or similar approaches. Different binding characteristics, buffers (pH, salt), application rate, and biorecognition element concentration lead to differences in thickness in the capture zone line. Once the biorecognition element is applied, the membrane material is typically immersed in a blocking agent, which is used to reduce nonspecific binding of the assay components. Common agents include proteins such as bovine serum albumin (BSA), casein, and gelatin; nonionic surfactants such as Tween-20, Triton X-100, or Brij; or synthetic polymers such as polyvinylpyrrolidone (PVP), polyvinyl alcohol (PVA), and polyethylene glycol (PEG). These components compete for protein binding sites, interfere with hydrophobic interactions, and protein binding, respectively (41, 42). In addition to serving as a blocking agent, Tween-20 has also been reported to have a renaturing effect on immobilized antigens, which can improve assay sensitivity through enhanced antigen–antibody binding (43, 44) 1.2. Detection Mechanisms
LFAs typically utilize antibody-labeled colloidal gold (5, 19, 45) as a visualization method. Colloidal gold ranges in size 2–250 nm, though 30–80 nm particles are prefered for LFAs (24). A diameter of 40 nm has been reported to be optimal, allowing for clear visualization, dense packing of these small particles at the capture zone, and reduced steric hindrance for protein binding (28). These particles have a characteristic red color, allowing for visual detection or semiquantitative measurements with portable reflectometers or scanners (46–48). Proteins are associated with gold particles ionically through the particle’s negative charge with positively charged amino acids such as lysine; through hydrophobic interactions of amino acids such as tyrosine and tryptophan; and through sharing of electrons between gold and sulfur atoms of cysteine (28). When labeled with anti-biotin or streptavidin, such species allow for recognition of biotinylated biorecognition elements, such as DNA oligonucleotides or antibodies. The sensitivity of LFAs using such particles can be increased with silver enhancement (49). Alternatively, antibody-tagged latex particles (50, 51), up-converting phosphors (26), superparamagnetic particles (52), or visible dye-encapsulating liposomes (25) have been used as a signal enhancement means. The latter species are the focus of this chapter. Liposomes are vesicles formed through the association of phospholipid molecules in an aqueous environment, yielding a structure with the hydrocarbon tails forming a lipid bilayer and hydrophilic headgroups directed at both the aqueous core and
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Fig. 2. Liposome structure: (A) Biorecognition elements can be covalently conjugated to or inserted into lipid bilayers through hydrophobic interactions (not to scale.) (B) The large internal volume of unilamellar vesicles can encapsulate hundreds of thousands of hydrophilic signaling molecules and provide for their stability. (C) Surfactant introduction can provide for instantaneous signal enhancement through release of encapsulants. Fluorophores encapsulated within liposomes at high concentrations undergo self-quenching, which is overcome upon release into the surrounding medium.
external aqueous medium. One of the common methods for liposome formation is known as the reverse-phase evaporation method (53, 54). Here, phospholipids are dissolved in organic solvent; the mixture is introduced to an aqueous medium containing high concentrations of visible dye; the solvent is removed under vacuum; the resulting liposomes are passed through defined pore-size membranes to improve homogeneity and reduce the number of lipid bilayers (lamallerity); then the unencapsulated material is removed through size-exclusion chromatography and dialysis. We commonly use sulforhodamine B (SRB) dye, which is relatively inexpensive, highly soluble in water, has a high molar extinction coefficient, and has good photostability. The advantages of dye-encapsulating liposomes as a label for nucleic acid sandwich hybridization assays include long-term stability of the encapsulated signaling molecules; the substantial signal enhancement afforded by the encapsulation of hundreds of thousands of dye molecules within the large interior liposomal volume; the instantaneous signal provided through either visual detection of intact liposomes or the release of hydrophilic dye molecules from their aqueous cores upon surfactant-induced liposome lysis; and the ease of labeling through the direct incorporation of hydrophobically modified nucleic acid probes into their lipid bilayers (Fig. 2) (25, 55–57) In terms of stability, we have found that DNA reporter probe tagged sulforhodamine B-encapsulating liposomes have retained their functionality in LFAs for at least 3 years when stored at 4°C. 1.3. Assay Formats
Two main assay formats are used in lateral-flow assays: the competitive assay and the sandwich assay format. Both formats have been used for nucleic acid lateral flow assays, though the latter is more common (56, 58). The competitive assay format utilizes an analyte
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Fig. 3. Competitive Immunoassay: (A) In the absence of analyte, analyte-conjugated liposomes are captured by an analyte-specific antibody at the competition zone. (B) In the presence of analyte, analyte-tagged liposomes compete with analyte present in the sample for the antibody immobilized at the competition zone yielding a signal that is inversely proportional to the analyte concentration.
conjugated detection species, such as a dye-encapsulating liposome, which competes with analyte present in the sample for an available biorecognition element. Biorecognition elements have included antibodies (59), RNA aptamers (60), and complementary DNA probes (58). This assay format is typically used for small molecule analytes to which only a single biorecognition element can bind. Here, the signal at the competition zone is inversely proportional to the concentration of analyte in the sample (Fig. 3). In the sandwich assay format, two biorecognition elements directed to either the same epitope present multiple times on the analyte surface or the different epitopes on the target analyte are utilized. For immunoassays, this format is useful for large analytes with multiple antigenic sites. Here, one antibody is immobilized onto a porous membrane and serves as the capture antibody, while the other is conjugated to a detectable species. The response is directly proportional to the concentration of analyte in the sample. A band consisting of a secondary antibody may also be present to serve as a control that the assay components worked and the assay was run correctly. This assay format is depicted in Fig. 4.
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Fig. 4. Sandwich Immunoassay: (A) In the absence of analyte, antibody-conjugated liposomes are captured by a speciesspecific secondary antibody at the control zone, thus proving that the assay was run correctly. (B) In the presence of analyte, the anti-analyte tagged liposomes form a sandwich complex at the capture zone. Unbound liposomes are captured by a species-specific secondary antibody at the control zone.
For RNA or DNA detection, a similar assay to the sandwich immunoassay is the sandwich-hybridization format. Mechanistically, these assays rely on the target nucleic acid forming a complex between an immobilized oligonucleotide capture probe and a labeled oligonucleotide reporter probe through Watson–Crick base pairing. The capture and detection probes are carefully selected ideally with no sequence homology to other organisms, which may be present in the sample. PCR or NASBA-based strategies allow for an additional level of specificity in terms of the use of specific primers used to amplify the sequence. In Fig. 5, an example for a nucleic acid LFA with detection and control zones is shown. Assays based on sandwich-hybridization are available in several platforms, such as sequential injection analysis (55), microtiter plate assays (61), and microfluidic devices (62). The LFA biosensor assays described in this chapter rely on the sandwichhybridization of a nucleic acid sequence based amplified (NASBA) RNA target between a membrane immobilized capture probe and SRB-encapsulating liposome conjugated reporter probe. NASBA uses the enzymes avian myeloblastosis virus reverse transcriptase (AMV-RT), RNaseH, and T7 DNA dependent RNA polymerase in the presence of deoxyribonucleoside triphosphates and appropriate primers to amplify relatively few copies of target RNA into
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Fig. 5. Sandwich-hybridization assay: (A) A biotinylated DNA oligonucleotide, which is complementary to one portion of the target sequence, is mixed with streptavidin and applied to form the capture zone 1.5 cm from the base of the membrane. An unmodified DNA oligonucleotide, which is complementary to the reporter probe sequence on the liposomes, is applied to form the control zone 2.5 cm from the base of the membrane. (B) In the presence of target, a sandwich hybridization complex forms with dye-encapsulating liposomes resulting in a magenta-colored band at the capture zone. (C) In the absence of target, only the control band is visible as no sandwich complex has formed.
billions of antisense copies isothermally at 41°C within 90 min (63). The target is first denatured briefly at 65°C and a primer sequence, which is complementary to a portion of the target RNA, is allowed to anneal to the denatured sequence. This added DNA oligonucleotide is referred to as the forward primer and contains a 25-base T7 promoter on the 5′ terminus. Using the hybridized primer as a starting point, the three-enzyme mixture is added and AMV-RT utilizes the deoxyribonucleoside triphosphates (dNTPs) and ribonucleoside triphosphates (NTPs) present in the mixture to form a cDNA sequence along the length of the target at 41°C. RNAse H serves to degrade the target RNA of the RNA–cDNA hybrid, allowing a second (reverse) primer to hybridize to the cDNA, forming a T7 promoter sequence prior to AMV-RT mediated formation of a double-stranded cDNA (63). The T7 RNA polymerase can then transcribe the dsDNA sequence to produce thousands of RNA molecules that are the complementary sequence of the original target RNA. These antisense RNA molecules can then be amplified in a similar manner, with primers annealing in the reverse order, resulting in >109 copies of singlestranded RNA molecules (64). The advantages of NASBA over other amplification techniques such as PCR or RT-PCR include its rapid and selective amplification of RNA, its isothermal conditions, and the generation of single stranded RNA (65–67).
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Although RT-PCR based methods are promising for RNA amplification, they require a DNAse step to remove DNA so that it is not coamplified along with RNA (68). Unfortunately, DNAse treatment has been reported to not be completely effective in this regard, thus additional complex steps are required (69). Nucleic acid-based identification is an important tool in the study of infectious diseases, genetic abnormalities, forensics, and biowarfare agent identification (70–72). Unlike DNA or antibody targets, the use of mRNA as a target allows for the detection of only viable organisms that may have survived eradication efforts. The question of viability is important when the effectiveness of decontamination procedures against resilient organisms is considered. It is crucial to determine whether decontamination has been successful and whether activity in affected areas can be safely resumed without the threat of surviving organisms. As such, mRNA is a more suitable indicator of viability than DNA or rRNA because of its vanishingly short half-life (65, 67). One example of where detection of viable organisms is beneficial is in drinking water processing plants. The parasitic protozoa, Cryptosporidium parvum, is a common contaminant of municipal water supply systems (73), agricultural run-off (38), and marine life of commercial interest (74), and is the causative agent of Cryptosporidiosis, an infection of the GI tract leading to massive diarrhea (75). Although physical and chemical disinfection methods may have effectively killed this resistant organism in drinking water, antibody or PCR-based detection approaches do not differentiate between viable and nonviable organisms. In addition, many antibodies against C. parvum are not specific for just this organism, but other species of Cryptosporidium, thus leading to the potential for false positives and unnecessary “boil water” requirements. LFAs based on the detection of C. parvum mRNA amplified using NASBA have shown detection limits of five oocysts in 0.5 mL buffer (22, 23, 58) with ongoing studies to determine limits of detection in treated and untreated drinking water. 1.3.1. Direct Detection of rRNA
In some cases, the target RNA is of sufficient concentration to be detected without amplification, which can reduce sample processing time, expense, and allows for assays to become more feasible for point-of-care applications. Ribosomal RNA (rRNA) comprises over 80% of cellular RNA (76) and thus can serve as a suitable target for nonamplification-based detection. 16S rRNA can provide genus specific, and sometimes species specific identification of bacteria when variable regions are targeted (77, 78). Because of the large size and complex structure of 16s rRNA, these regions are often inaccessible to specific probes due to rRNA interactions with ribosomal proteins or rRNA itself (64, 79). Hence, considerations for probe access are sometimes required. Unlabeled oligonucleotide probes that can hybridize to regions adjacent to
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the labeled probe-binding region have been found to be useful for unfolding 16s RNA and thus permitting better target accessibility (78). The use of multiple capture and reporter probes can serve a similar purpose as well as to increase the number of liposomes binding per target analyte. In our hands for the detection of Streptococcus pyogenes, unlabeled probes intended to reduce secondary structure had either no or only a negative effect on the signal generated, but multiple labeled probes served to enhance our signal (80). For this liposome-based nucleic acid LFA, the limit of detection was 135 ng total RNA, which corresponds to 7.3 × 106 CFU/mL of S. pyogenes (80). This LOD is comparable to other nonamplification based methods such as electrochemiluminescence yielding detection of 105–108 CFU/mL and oligonucleotide microarrays yielding a LOD of 500 ng RNA, corresponding to 7.5 × 106 cell equivalents (81, 82). 1.4. Universal Assays
Both immunoassays and nucleic acid-based LFAs can also be made in a universal format, obviating the need for the generation of specifically labeled membranes and detection elements. This can be done through membrane immobilization of streptavidin (23) or anti-fluorescein antibodies (22) and subsequent addition of biotinylated or fluorescein-labeled capture probes, respectively, with the remaining assay components. Protein A/G (83), streptavidin/avidin (22, 84), or generic oligonucleotide (23) labels can be conjugated to the liposomes to form generic species capable of facile recognition of the Fc′ region of antibodies, biotinylated biorecognition elements, or complementary generic oligonucleotides, respectively. Anti-fluorescein, anti-biotin, or anti-digoxigenin are also options for common universal liposome tags (85). These assays are of particular interest in research laboratories where a variety of antibodies or probes need to be screened or where different analytes need to be tested. From a commercial and manufacturing standpoint, the universal format is of interest since only one type of membrane and liposome need to be prepared, which simplifies packaging of tests for any analyte.
1.5. Assay Development
The design of a nucleic acid LFA is a multistep process. Once the target organism has been selected, several tasks must be completed in addition to those already described (i.e., liposome preparation, NASBA, and membrane preparation). These include the selection of a region of mRNA that shares little homology with other organisms; the design of primers and probes with little secondary structure and efficient target binding for amplification and detection of RNA; the design of a truncated synthetic DNA target sequence to simplify assay optimization; and the selection of appropriate growth conditions and extraction procedures for RNA isolation. The latter tasks are beyond the scope of this chapter,but an example procedure for RNA isolation is provided in Subheading 3.6.
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1.5.1. Primer and Probe Design
The design of appropriate amplification primers and detection probes is essential for any nucleic acid-based detection assay. Although details are beyond the scope of this chapter, in general, it is an iterative process performed with the help of software programs such as DNAstar (DNASTAR, Inc., Madison, WI) and VisualOMP 4 (DNA Software, Ann Anbor, MI) in order to ensure primer and probes with little secondary structure and efficient binding to the target sequence. In addition, regions of low secondary structure within the target molecule can be identified using Mfold (86). The National Center for Biotechnology Information (NCBI) Basic Local Alignment Search Tool (BLAST) (87) is typically used to ensure the specificity of the selected sequences a process that in some cases can be incorporated in the initial software programs such as VisualOMP 4. This tool allows the alignment check of an oligonucleotide sequence to an extensive database of publicly available DNA and RNA sequences (GenBank) (88). If more oligonucleotide probes are required per assay, as is the case for 16s rRNA, the probe selection process becomes more complex and time consuming. A software program developed in our group addresses the need of multiple probes per assay (80). In addition, this software program is designed for the selection of primers used in NASBA and for probes needing a universal tag such as those described further below (19, 20).
1.5.2. Synthetic DNA Target Sequence
Rapid screening of experimental conditions is possible with the design of a synthetic target DNA sequence. It typically represents a part located on the target RNA between the two amplification primers that can be recognized by the reporter and capture probes. The ability of the probes to bind to the synthetic target and their limits of detection can be assessed prior to analyses with the actual target sequence. Synthetic RNA may also be used to simulate assay performance, though it is usually more costly and sensitive to degradation than its DNA counterpart. In either case, the synthetic single stranded sequence is diluted to an appropriate concentration range (typically 1–200 nM) so that quantitative dose response curves can be generated. Although a final study of the best probes with the actual target sequence is necessary, this initial screening is a useful tool in assay optimization, assessment of assay performance, and selection of optimal detection probes.
2. Materials Materials listed here are those that have been used successfully in our laboratory, though substitutions may be made. Unless otherwise specified, reagents were molecular biology grade and were
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Table 1 Example primer, probe, and target sequences for atxA from Bacillus anthracis Function
Sequence (5′→3′)
Length
atxA primer 1
AATTCTAATACGACTCACTATAGGGAGAAGGGGGAAACGGCCAATAATCA
50
atxA primer 2
TCAAATTTGCGAAGAACTTGTA
22
atxA reporter probe
CAAGATGTCCGCGTATTTAT[Chol-TEG]
20
atxA capture probe
Biotin-CTAGAAATATCGGGAAGAGAA
21
atxA synthetic target
ATAAATACGCGGACATCTTGTCTTCTCTTCCCGATATTTCTAG
43
purchased from VWR (Bridgeport, NJ). Table 1lists synthetic DNA primers, capture and reporter probes, and synthetic DNA used for the detection of atxA mRNA from Bacillus anthracis as an example. 2.1. Liposome Preparation
1. Bath sonicator (Aquasonic Model 150D, VWR, Bridgeport, NJ). 2. Rotary evaporator (Model R-114, Buchi, New Castle, DE). 3. Round bottom flask (50 mL) (Catalog # 80068-756, VWR). 4. Mini-extruder (Catalog # 610000, Avanti Polar Lipids, Alabaster, AL), including two 1 mL syringes, teflon supports and o-rings, and extruder holder/heating block. When purchasing initially, the membranes and supports listed in #5 of this section are included with this catalog number as of the date that this chapter was prepared. 5. Extrusion membranes (0.4 μm and 1.0 μm polycarbonate membranes*, 19 mm) and filter supports (Catalog # 610007, 610010, and 610014, respectively, Avanti Polar Lipids). 6. 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), cholesterol, 1,2-dipalmitoyl-sn-glycero-3-[phospho-rac-(1-glycerol)], sodium salt (DPPG) (Catalog #: 850355, 700000, and 840455, respectively, Avanti Polar Lipids). 7. Chloroform, methanol, and isopropyl ether (HPLC grade). 8. HEPES-saline buffer (10×) is composed of 0.1 M HEPES, 2.0 M sodium chloride, and 0.1% (w/v) sodium azide, adjusted to pH 7.5 with NaOH. 9. 1× HEPES-Saline-Sucrose buffer (1× HSS) is prepared by dissolving 205.4 g sucrose (0.2 M) in 300 mL 10× HEPES-saline and 900 mL water, then bringing the final volume to 3 L with deionized water. However, we usually prepare a 2 M stock solution of sucrose instead of weighing for this solution.
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10. Sulforhodamine B (0.419 g) (SRB, Catalog # S1307, Molecular Probes, Inc., Eugene, OR) is added to 0.5 mL 0.2 M HEPES, pH 7.5 and diluted to a final volume of 5 mL with deionized water. 11. 3′-Cholesteryl-TEG modified DNA reporter probe oligonucleotides (Operon Biotechnologies, Inc., Alameda, CA) are diluted to 300 μM with a 1:4 (v/v) mixture of methanol/formamide, adding the formamide to the probe first, followed by methanol. The probes are then aliquotted into 50 μL portions prior to storage at −20°C. 12. Sephadex G-50 (Catalog # G-50-150, Sigma, St. Louis, MO) and a glass chromatography column (such as VWR catalog # KT420400-1530). The column is packed by first swelling 1 g of Sephadex in 120 mL deionized water in 3 × 50 mL centrifuge tubes, and decanting off water either after centrifugation or settling by gravity then pouring. The volume is then replaced by 1× HSS and decanting procedure repeated. Overall, three 1× HSS buffer exchanges are typically sufficient. The Sephadex mixture is then poured into the chromatography column to a height of approximately 20 cm and allowed to settle while maintaining a flow of 1× HSS through the column for at least 30 min. The top of the column should be level with no gaps or bubbles throughout the remaining height. 13. Dialysis membranes, Specta/Por 2 (Catalog # 132 678, Spectrum Laboratories, Inc., Rancho Dominguez, CA). 14. Centrifuge tubes (15 mL and 50 mL) (Catalog #: 21008216 and 21008-242, respectively, VWR). 2.2. Membrane Preparation
1. TLC applicator (Linomat IV, CAMAG Scientific, Wilmington, NC). 2. Vacuum sealer (Foodsaver, San Francisco, CA). 3. Vacuum oven, capable of 23°C and 50°C. 4. Paper cutter. 5. Fine-tip tweezers (Catalog #25729-081, VWR). 6. Kimwipes, 15″×17″ (Catalog # 21905-049, VWR). 7. Nitrocellulose membranes (HanoRapid membranes, 15 m × 15.2 cm roll, Catalog # 51515, Hanomy LLC, Cheshire, CT) cut into 7.5 × 20 cm sections. Alternatively, we have had success with polyethersulfone membranes (Predator, 3 m × 9″ roll, Catalog # PREDL3R, Pall Corporation, Pensacola, FL) cut into 8 × 20 cm portions. 8. Streptavidin (Catalog # S888, Molecular Probes, Inc., Eugene, OR) is diluted to 100 μM with 50 mM potassium
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phosphate, pH 7.8 containing 1 mM EDTA and aliquotted into 50 μL portions prior to storage at −20°C. 9. 5′-Biotinylated DNA capture probe oligonucleotides (Operon Biotechnologies, Inc., Alameda, CA) are diluted to 300 μM with 50 mM potassium phosphate, pH 7.8 containing 1 mM EDTA and aliquotted into 50 μL portions prior to storage at −20°C. 10. The blocking reagent is prepared by diluting 100 mL 0.15% (w/v) casein, 100 mL 10× TBS (0.2 M Tris, 1.5 M sodium chloride, pH 7.0 containing 0.1% (w/v) sodium azide), and 100 mL 2% (w/v) polyvinylpyrrolidone (PVP) to a final volume of 1,000 mL with deionized water. 2.3. Assay Optimization Using Synthetic Target DNA
1. 10 × 75 mm culture tubes (VWR, #47729-568, Bridgeport, NJ). 2. Synthetic DNA target oligonucleotide (Operon Biotechnologies, Inc., Alameda, CA) is designed to be complementary to the cholesteryl-TEG reporter probe and biotinylated capture probe with a short linker sequence. Target is diluted to 1,000 μM with 50 mM potassium phosphate, pH 7.8 containing 1 mM EDTA and aliquotted into 50 μL portions prior to storage at −20°C. 3. Prior to running the assay, the target is diluted to 1, 5, 10, 50, 100 nM concentrations using the same buffer. 30 μL aliquots are appropriate. 4. Liposomes tagged with cholesteryl-TEG reporter probe of known optical density or phospholipid concentration. 5. Membranes with capture probe immobilized cut into 4.5 mm × 7.5–8 cm strips. 6. Hybridization buffer components: Formamide (molecular biology grade), 20× SSC (0.3 M sodium citrate, 3.0 M sodium chloride, 0.1% (w/v) sodium azide), 10% (w/v) Ficoll Type 400 prepared in water, and 2 M sucrose containing 0.1% (w/v) sodium azide. 50 mL of each component is sufficient for initial optimizations. 7. Reflectometer (λ = 560 nm, ESECO Speedmaster, Cushing, OK) or scanner with image analysis software (optional if more than qualitative results by eye are desired).
2.4. Extraction of RNA
1. RNeasy Mini Kit (Catalog # 74104, Qiagen, Valencia, CA). (a) Kit contents: 50 RNeasy Mini spin columns, 1.5 mL and 2.0 mL collection tubes, RNase-free reagents (RNasefree water) and buffers (RW1, RLT, RPE). 2. Ethanol (absolute, not denatured). 3. β-mercaptoethanol (β-ME) (14.3 M).
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4. Buffer RLT plus β-ME: Add 10 μL of β-ME per 1 ml Buffer RLT. This mixture is stable for 1 month. 5. Microcentrifuge with 2 mL tube capacity. 6. Lysozyme. 7. TE buffer (10 mM Tris–HCl, 1 mM EDTA, pH 7.5) made using RNAse free reagents. 8. 0.65 mL centrifuge tubes, RNase free. 2.5. NASBA
1. Heating blocks set at 41°C and 65°C. 2. RNAse-free 0.65 mL microcentrifuge tubes, pipette tips (0.1–10 μL volume), water, and DMSO. 3. NASBA enzyme mix (three enzyme cocktail, AMV RT, RNase H, and T7 RNAP, Catalog # NEC-1-20, Life Sciences, Petersburg, FL) aliquotted into 57 μL portions. Each aliquot is sufficient for ten NASBA reactions. 4. 5× NN buffer aliquotted into 44 μL portions. NN buffer is composed of 2.5 mM ITP, 7.5 mM GTP, 2 mM of each ATP and UTP, 60 mM MgCl2, 350 mM KCl, 25 mM DTT, 5 mM each dNTP, and 200 mM Tris. The NTP (catalog # BIO39050) and dNTP (catalog # BIO-39029) sets were ordered from Bioline Inc. (Randolph, MA). 5. The NASBA primer stocks (Operon Biotechnologies, Alameda, CA) are diluted to 100 μM with RNAse free water. 6. Primer mixtures are prepared by mixing 10 μL of each primer (A&B), 750 μL DMSO, and 230 μL water and aliquotting into 44 μL portions. 7. RNAse free water (Catalog # MB-010-1000, Rockland Immunochemical, Gilbertsville, PA) was used as a negative control.
2.6. Running an Assay
1. All components listed in Subheading 2.3 plus amplicon generated in Subheading 3.7.
3. Methods 3.1. Liposome Preparation
A flowchart of this process is shown in Scheme 1. Note that the times listed in this flowchart, and others in this chapter, reflect overall times for each step, including set-up and incubations. For specific times for each step, please refer to the text. 1. Set rotary evaporator bath and sonicator bath to 45°C. Fill the condenser on the rotary evaporator with ice and seal condenser top rim to rotary evaporator glass housing with parafilm.
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Scheme 1. Liposome preparation.
2. Prepare dye solution as described in materials section using a 15-mL centrifuge tube, cover tube with aluminum foil, and store in the 45°C sonicator bath. 3. DPPC, DPPG, and cholesterol (40.9:20.1:51.7 μmol, respectively) are added to a 50-mL round bottom flask. 4. A solvent mixture containing 3 mL chloroform, 0.5 mL methanol, and 3 mL isopropyl ether is added and the mixture sonicated for 1 min in a bath sonicator. We use sonication level 6 with the Aquasonic Model 150D bath sonicator. 5. Cholesteryl-labeled reporter probe (corresponding to 0.013 mol% of the total lipid input) solution (50 μL) is then added to the dissolved lipids and the flask vortexed until visually homogeneous (40). For the preparation of antibodytagged liposomes, the probe solution is omitted and lipid composition slightly different (see Note 1). 6. 2 mL of the 45°C sulforhodamine B dye solution is added to the lipid mixture during the first 30 s while sonicating for a total of 4 min. Swirl the flask manually during sonication (see Note 2). 7. The mixture is then placed onto a rotary evaporator at the highest rotation speed with the bath at 45°C. The vacuum should be adjusted such that bubbling or foaming of the contents does not occur. Using the Buchi R-114 rotary evaporator, a vacuum of 500 mBar for 20 min, followed by 400 mbar for 20 min is recommended. During evaporation, return unused portion of the SRB solution to the 45°C sonicator bath. 8. The mixture is then transiently vortexed preceding and following an additional introduction of 2 mL of 45°C, 150 mM SRB. Using a VWR mini-vortexer, we have found that holding the flask at a 45° angle and vortexing at level 4–5 is appropriate; however, be sure to use a stopper on the flask.
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9. The mixture is returned to the rotary evaporator for an additional 30 min under slightly higher vacuum than in step 6 (350 mBar, in our case.) Generally, the evaporation is considered complete 15 min after no further solvent is observed from the condenser. Removal of nearly all organic solvent is required for successful formation of liposomes. 10. While the liposome mixture is on the rotary evaporator, set the extruder support block of the mini-extruder on a hot plate and adjust temperature such that the temperature of the block does not exceed 65°C. 11. Set-up the mini-extruder as outlined in the manufacturer’s instructions. Label 2–50 mL centrifuge tubes for each extrusion size and clamp them in a 45°C water bath, during the extrusion process and before application to the size-exclusion column. 12. The liposomes are then extruded at 60–65°C 19 times through 1.0 μm pore membranes, followed by 19 times through 0.4 μm pore membranes. The liposomes must be maintained in the 45°C water bath during and after extrusion. 13. The level of the 1× HSS buffer volume on the Sephadex G-50 column prepared in step 12 of Subheading 2.1 is reduced to just below the level of the Sephadex. Do not allow the Sephadex to dry out. 14. The liposomes are pipetted onto the top of the column without disturbing the top of the Sephadex. This addition should be done as carefully, but rapidly, as possible using a glass Pasteur pipet in a circular path along the inside of the glass column just above the Sephadex. Important: the tubes containing the liposomes need to be kept in 45°C at all times and the transfer to the column needs to be efficient. If the liposomes cool to room temperature, they will form clumps with external dye on the column and will not separate. 15. Once all of the liposome volume has entered the column, two ∼1 mL aliquots of 1× HSS should be pipetted onto the top of the column allowing each one to enter the column before the next addition. This is done to create a separation between the liposomes/free dye and the remaining 1× HSS used for liposome elution. 16. After carefully pipetting another 1–2 mL of 1× HSS on top of the column, allow 1× HSS to flow freely (typically ∼4 mL/ min) and collect the eluting liposomes in test tubes. The liposomes will elute from the column first, forming a dark magenta band, followed by the free dye yielding a darker and larger elution volume. If the column is run properly, some separation of liposomes and free dye can usually be seen.
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17. The liposome containing fractions are then combined based either visually or on their measured optical density at 532 nm. Optical density measurements are made at 532 nm by diluting 5 μL liposomes with 995 μL of 1× HSS in a 1.5 mL spectrometer cell, or more conveniently, 1 μL liposomes in 199 μL of 1× HSS in a clear microtiter plate (Corning #3795). Be selective about which liposomes are pooled together: only the most highly concentrated fractions should be together, the intermediates together, and if needed, the low concentration fractions. This is more important if the liposomes are to be subsequently coupled to antibodies where a high liposome concentration is necessary. 18. The combined fractions are placed into dialysis bags and dialyzed overnight against the sucrose-HEPES-saline buffer. Be sure to transfer liposomes from the test tubes to the dialysis bags over a beaker to reclaim any potential spills. Replace buffer and continue dialysis until external dialysate is no longer notably pink. 19. Clean everything thoroughly, including removing the o-rings from the Teflon holders and the needles and plungers from the syringes of the extruder. 20. Store dialyzed liposomes in 15-mL centrifuge tubes at 4°C. 3.2. Membrane Preparation
A flowchart of this process is shown in Scheme 2. This procedure is for the preparation of capture probe immobilized membranes, but can be readily adapted for antibody immobilized membranes (see Note 3). 1. Set the vacuum oven to 50–55°C. Allow to stabilize for at least an hour before use. 2. Prepare blocking reagent (0.2% (w/v) PVP, 1× TBS, 0.015% (w/v) casein) or remove previously made blocking reagent from the fridge and allow to reach ambient temperature.
Scheme 2. Membrane preparation.
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3. We have found optimum dimensions of 4.5 mm × 7.5–8.0 cm per LFA membrane. The dimensions of the membrane roll or sheet often depend on the manufacturer, but we have obtained membranes in 15 cm and 24 cm widths from Hanomy LLC and Pall Life Sciences, respectively. Cut the rolled membrane into a 20 cm × 7.5–8.0 cm section with the 7.5–8.0 cm height being parallel to the original cut. Otherwise, curling of the individual membranes may be noted when running the assays. 4. Label each membrane section with a line drawn ∼0.5 cm from its top on the laminated side using a permanent marker. 5. For some types of membranes, a prewetting step may be helpful. For nitrocellulose membranes, we have used freshly prepared 10% (v/v) methanol in 1× PBS following steps 21–25 in Subheading 3.2.3, then drying in the vacuum oven for 30 min at 40°C under 15″ Hg. 6. For each 20 cm × 7.5–8.0 cm membrane section, prepare 50 μL of a 20 μM streptavidin and 60 μM biotinylated capture probe mixture in 0.4 M NaHCO3/Na2CO3, pH 9.0 containing 5% (v/v) methanol. We have found that an unmodified capture probe may also be immobilized without streptavidin using the same conditions with both suggested membrane types. However, the efficiency of the immobilization with and without streptavidin should be compared for other membrane types. 7. A control band showing that the sequence on the liposomes is capable of hybridizing and that the liposomes themselves are visible may be added. For simplicity of membrane preparation, we typically do not include this control band. However, if desired, prepare an equivalent volume of an unmodified probe that is complementary to the sequence on the liposomes for deposition as a control zone. Be sure that this sequence shares little complementarity with the capture probe sequence used at the capture zone. Otherwise, excess probes that are released during the blocking step can hybridize to the opposite zone and reduce the amount of probe available for interaction with target. 3.2.1. Automated Application of Capture Probes Using a Linomat IV Or Similar TLC Plate Applicator
The following instructions are for operation of the CAMAG Linomat IV, but should be adaptable for similar applicators using the respective manufacturer’s directions. 8. Set the Limonat IV parameters as follows: Plate width:
200
Space:
0
Start position:
0
Rate:
4 μL/s
Band width:
190
Volume:
38
9. Fill the syringe slowly with the capture probe/streptavidin mixture to avoid air bubbles, then insert syringe fully into syringe holder.
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10. Gradually lower the plunger mechanism by pressing in the front & rear buttons until the silver lever is just above the syringe plunger. 11. Press the GAS button on the front of the Linomat, then lower the silver lever onto the syringe plunger by holding down the ↓ key until you see liquid consistently bubble from the end of your syringe. You may simultaneously hold down the + and ↓ buttons (faster) if there is a fair distance between the silver lever and syringe plunger. 12. Place one of the 20 cm × 7.5–8.0 cm membrane sections so that it is lined up with the bottom and side markers on the black surface. Use the supplied flat magnets to hold down the membrane at its top and right sides. Slide the tower to 1.5 cm from the base of the membrane using the ruled marks. 13. Press Calc, then Run. Repeat steps for remaining membrane sections. If applicable, deposit the control zone sequence 2.5 cm from the base of the membrane following the same procedure. 14. Using tape with the sticky side exposed placed onto a piece of cardboard, lightly tape the top of each membrane section to the cardboard using tweezers. 3.2.2. Manual Application of Capture Probes Using a Pipettor
If a TLC or similar applicator is not available, the capture probe/ streptavidin solution may be applied manually to the lateral flow membranes. Here, a round spot for the capture and control zones would result, vs. the line seen in Fig. 1 generated using the Linomat for probe application. This procedure is useful for optimization purposes, but is more laborious than the TLC applicator procedure due to the need to apply probe and tape individual membranes. 15. Cut the membrane sections into 4.5 mm × 7.5–8.0 cm pieces. We have found it beneficial to tape a laminated piece of paper to the paper cutter with lines every 4.5 mm for a guide. Then, align one end of one section with this paper and advance the section, making perpendicular cuts every 4.5 mm. 16. Using tape with the sticky side exposed placed onto a piece of cardboard, lightly tape the top of each membrane to the cardboard using tweezers. 17. Apply 1 μL of the capture probe/streptavidin solution to each membrane 1.5 cm from the bottom of the strip using a microvolume pipettor. The more consistent the probe application is, the more consistent the LFA results will be. The probe solution will rapidly seep into the membrane. 18. If applicable, deposit the control zone sequence 2.5 cm from the base of the membrane following the same procedure.
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19. Place the taped membranes in the vacuum oven at 50–55°C and set vacuum to 15″ Hg for 90 min. 20. Remove membranes from vacuum oven and turn heat off. 21. Pour blocking reagent into a plastic Tupperware container so that it is 200
Fig. 7. Representative lateral flow sandwich-hybridization assay membranes: Nitrocellulose membranes with capture probe and a sequence complementary to the reporter probe on the liposomes immobilized 1.5 cm and 2.5 cm, respectively, from the base of the membrane used for the detection of target RNA using dye-encapsulating liposomes. In the absence of target, only the control band is visible as no sandwich complex has formed (top). In the presence of increasing concentrations of target, a sandwich hybridization complex forms with dye-encapsulating liposomes resulting in a magenta-colored band at the capture zone that is proportional to target concentration (increases from top to bottom in the figure). Here, 1 μL of the synthetic DNA sequence listed in Table 1 was diluted to 1 –200 nM, mixed with 4 μL reporter probe conjugated liposomes diluted to 1 mM phospholipid using 30% (v/v) formamide, 9× SSC, 0.2 M sucrose containing 0.2% (w/v) Ficoll as a hybridization buffer. Assays were run as described in Subheading 3.4.
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nucleotides in length. Irrespective of the extraction kit used, we suggest assessing the RNA purity and yield prior to proceeding with amplification. We use the following procedure on the RNA eluted from silica particles used in the RNeasy Mini kit (92): 1. Blank a spectrometer using TE buffer in a 1 mL quartz spectrometer cell. 2. Dilute 10 μL of filtrate with 490 μL RNase-free TE buffer in the matched spectrometer cell. 3. Measure absorbance at l = 260 and 280 nm. 4. Calculate RNA purity, RNA concentration, and total RNA yield as follows: RNA purity: A260 / A280.
(1)
RNA concentration: [RNA] = A260 × 40 g/mL_ RNA × SPectrometer_dilution_factor
(2)
RNA yield: Total_yield = [RNA] × volume_of_extract
(3)
For example, if A260 = 0.083 and A280 = 0.038 and the sample was diluted 1:10 for measurement in the spectrometer (spectrometer dilution factor), then the concentration of RNA in the sample would be 33.2 μg/mL and 260/280 ratio = 2.184. The total yield for a 50 μL extraction volume would then be 1.66 μg. We generally do not proceed with NASBA unless the A260/A280 ratio is ≥ 1.7 to avoid negative results. 5. Dilute the extracted RNA to 2 ng/μL with RNAse free water, aliquot into 5 μL portions using 0.65 mL centrifuge tubes, and store at −80°C. 3.7. NASBA
1. Mix 44 μL primer mixture with 44 μL of 5× NN buffer and 22 μL RNAse free water. This volume is sufficient for ten reactions. 2. RNA extract and RNAse free water are used as sample and negative controls, respectively. Add 5 μL volume per 0.65 mL microcentrifuge tube. In future runs, previously determined positive RNA extract may be used as a positive control for NASBA. Alternatively, positive NASBA reactions can be diluted 1:10,000-fold, stored as 20 μL aliquots at −80°C and used as strong positive controls. 3. Add 10 μL of the primer/NN mixture made in step 1 to each tube from step 2. 4. The mixture is heated at 65°C for 5 min, then at 41°C for 5 min. 5. The three-enzyme cocktail (5 μL) is added and the reaction is incubated at 41°C for 90 min. 6. The completed NASBA reactions are stored at −20°C for short term storage or −80°C for longer storage periods.
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3.8. Running an Assay
1. Add 4 μL hybridization buffer, 1 μL NASBA product, and 2 μL reporter probe conjugated liposomes to the bottom of a 10 × 75 mm culture tube. 2. Run the assay as described in Subheading 3.4 using the optimal time, temperature, and hybridization buffers.
4. Notes 1. For the preparation of liposomes to be tagged to antibodies, the cholesteryl-tagged reporter probe is omitted and 6 mol% of N-glutaryl DPPE (Catalog # 870245, Avanti Polar Lipids) is added instead. This phospholipid introduces a carboxylic acid group to the lipid bilayer allowing for facile conjugation to primary amine groups using 1-ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (EDC) (12). 2. Be sure that the solution in the flask is being sonicated moderately vigorously and that the flask is not located in a “dead spot” of the bath. We use sonication level 6 with the Aquasonic Model 150D bath sonicator. 3. Membranes are prepared as described in Subheading 3.2, except that the streptavidin and capture probe are omitted and antibodies are used instead. For sandwich immunoassays, a matched set of affinity purified antibodies directed to different epitopes of the target molecule is desirable. Ideally, the antibodies will be available at high concentration (≥1 mg/mL). For immobilization onto the membranes, a good starting antibody condition has been 1 mg/mL in 0.4 M NaHCO3/ Na2CO3, pH 9.0, though this can be adjusted depending on the application and antibody. Both antibodies of the pair should be tested for use as a capture antibody. A species-specific secondary antibody can be immobilized as a control line. We had success using the following blocking reagent composition: 0.1% (w/v) gelatin, 0.002% (w/v) Tween-20, 0.02% (w/v) PVP, and 0.005% (w/v) casein in 1× TBS. 4. For antibody conjugation, the liposomal phospholipid concentration needs to be determined using Bartlett assays (94) (see Subheading 3.3). Both antibodies of the matched set should be conjugated to liposomes to determine which serves as the better detection antibody. Liposomes should be mixed with antibodies with the latter at 0.05 mol% of the total lipid input. Then, just before use, dilute EDC to a concentration of 100 mg/mL with 0.1 M MES buffer, pH 4.65 then add five molar equivalents (based on total N-glutaryl DPPE lipid input) to the liposome/ antibody mixture. Note that EDC is very hydroscopic – use care
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to protect the reagent from moisture. Incubate the mixture at room temperature in the dark for 15 min, then purify the conjugated liposomes through a size-exclusion chromatography column packed with Sepharose CL-4B (Sigma-Aldrich #CL4B200) using 1× HSS as an elution buffer (12). 5. Try to pipet the necessary solutions directly into the base of the 10 × 75 mm test tubes and avoid touching the LFA membrane to the side of the tube after the membrane has contacted any of the solutions. Liquids on the sides of the tubes may cause the membrane to run against the side of the tube and can result in inaccurate readings. If pipettor shaft is too large to enter and deposit volumes directly on the bottom of the tube, a larger test tube may be used (12 × 75 mm, VWR #47729-570.) 6. A good starting hybridization buffer condition is 9× SSC, 0.2% (w/v) Ficoll, 30% (v/v) formamide, and 0.2 M sucrose. The ideal hybridization conditions are a balance between preventing nonspecific binding and false positives due to low stringency and allowing the target molecule to bind with sufficient strength at a reasonable rate. For a given set of oligonucleotides, optimal hybridization can be accomplished through the adjustment of three factors: formamide concentration, salt concentration, and temperature. For a high stringency hybridization, high temperature, high concentration of formamide, or low concentration of salt are needed. Conversely, low stringency conditions include those with high salt concentrations, low formamide concentrations, and low temperature. Dextran sulfate and polyethylene glycol serve as molecular exclusion agents, which effectively reduce the volume in which the probe has to hybridize. However, these agents are of little utility for probes less than 50 nucleotides (95). Instead, we have found the use of Ficoll to have an enhancing effect in the hybridization of probes and target sequence. Several references are suggested for further understanding of nucleic acid hybridization theory (71, 86, 96). 7. A good starting running buffer for immunoassays is composed of 1% (v/v) formamide, 0.2 M sucrose, and 0.5% Ficoll Type 400 in 1× PBS. Otherwise, the assays are run as described in Subheading 3.4.
Acknowledgments The authors gratefully acknowledge the review of this chapter by Kit Meyers and Barbara Leonard.
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References 1. Helbig, J. H., Luck, P. C., Kunz, B., and Bubert, A. (2006) Evaluation of the Duopath Legionella lateral flow assay for identification of Legionella pneumophila and Legionella species culture isolates. Appl Environ Microbiol. 72, 4489–4491 2. Koide, M., Haranaga, S., Higa, F., Tateyama, M., Yamane, N., and Fujita, J. (2007) Comparative evaluation of Duopath Legionella lateral flow assay against the conventional culture method using Legionella pneumophila and Legionella anisa strains. Jpn J Infect Dis. 60, 214–216 3. http://www.kit.nl/biomedical_research/ assets/images/Lepto_lateral_flow_protocol.doc 4. Johnston, S. P., Ballard, M. M., Beach, M. J., Causer, L., and Wilkins, P. P. (2003) Evaluation of three commercial assays for detection of Giardia and Cryptosporidium organisms in fecal specimens. J Clin Microbiol. 41, 623–626 5. Ketema, F., Zeh, C., Edelman, D. C., Saville, R., and Constantine, N. T. (2001) Assessment of the performance of a rapid, lateral flow assay for the detection of antibodies to HIV. J Acquir Immune Defic Syndr. 27, 63–70 6. Cazacu, A. C., Demmler, G. J., Neuman, M. A., Forbes, B. A., Chung, S. Y., Greer, J., Alvarez, A. E., Williams, R., and Bartholoma, N. Y. (2004) Comparison of a new lateral-flow chromatographic membrane immunoassay to viral culture for rapid detection and differentiation of influenza A and B viruses in respiratory specimens. J Clin Microbiol. 42, 3661–3664 7. Mokkapati, V. K., Sam Niedbala, R., Kardos, K., Perez, R. J., Guo, M., Tanke, H. J., and Corstjens, P. L. (2007) Evaluation of UPlinkRSV: prototype rapid antigen test for detection of respiratory syncytial virus infection. Ann N Y Acad Sci. 1098, 476–485 8. Slinger, R., Milk, R., Gaboury, I., and Diaz-Mitoma, F. (2004) Evaluation of the QuickLab RSV test, a new rapid lateral-flow immunoassay for detection of respiratory syncytial virus antigen. J Clin Microbiol. 42, 3731–3733 9. van Hengel, A. J., Capelletti, C., Brohee, M., and Anklam, E. (2006) Validation of two commercial lateral flow devices for the detection of peanut proteins in cookies: interlaboratory study. J AOAC Int. 89, 462–468 10. http://www.craigmedical.com/salivascreen6_drug_tests.htm 11. http://www.brittneylimited.com/PDF%20 Files/FDA_Files/Pregnancy-strip.pdf
12. Edwards, K. A., Curtis, K. L., Sailor, J., Baeumner, A. (2008) Universal liposomes: Preparation and usage for the detection of mRNA. Anal Bioanal Chem.391, 1689–1702, Epub 2008 March 8 13. Jung, K., Zachow, J., Lein, M., Brux, B., Sinha, P., Lenk, S., Schnorr, D., and Loening, S. A. (1999) Rapid detection of elevated prostate-specific antigen levels in blood: performance of various membrane strip tests compared. Urology. 53, 155–160 14. http://www.brittneylimited.com/PDF%20 Files/PSA-Prostrate-strip.pdf 15. http://www.nhdiag.com/botulism.shtml 16. Sharma, S. K., Eblen, B. S., Bull, R. L., Burr, D. H., and Whiting, R. C. (2005) Evaluation of lateral-flow clostridium botulinum neurotoxin detection kits for food analysis. Appl Environ Microbiol. 71, 3935–3941 17. Park, C. H., Kim, H. J., Hixon, D. L., and Bubert, A. (2003) Evaluation of the duopath verotoxin test for detection of shiga toxins in cultures of human stools. J Clin Microbiol. 41, 2650–2653 18. Capps, K. L., McLaughlin, E. M., Murray, A. W. A., Aldus, C. F., Wyatt, G. M., Peck, M. W., Amerongen, A., Ariens, R. M. C., Wichers, J. H., Baylis, C. L., Wareing, D. R. A., and Bolton, F. J. (2004) Validation of three rapid screening methods for detection of verotoxinproducing Escherichia coli in foods: Interlaboratory study. J Aoac Int. 87, 68–77 19. Smits, H. L., Eapen, C. K., Sugathan, S., Kuriakose, M., Gasem, M. H., Yersin, C., Sasaki, D., Pujianto, B., Vestering, M., Abdoel, T. H., and Gussenhoven, G. C. (2001) Lateral-flow assay for rapid serodiagnosis of human leptospirosis. Clin Diagn Lab Immunol. 8, 166–169 20. Kassler, W. J., Dillon, B. A., Haley, C., Jones, W. K., and Goldman, A. (1997) On-site, rapid HIV testing with same-day results and counseling. Aids. 11, 1045–1051 21. Prevention, U. S. C. f. D. C. a. (1996) in “MMWR Morb Mortal Wkly Rep”, Vol. 45, pp. 468–80 22. Baeumner, A. J., Jones, C., Wong, C. Y., and Price, A. (2004) A generic sandwich-type biosensor with nanomolar detection limits. Anal Bioanal Chem. 378, 1587–1593 23. Baeumner, A. J., Pretz, J., and Fang, S. (2004) A universal nucleic acid sequence biosensor with nanomolar detection limits. Anal Chem. 76, 888–894 24. Seal, J., Braven, H., Wallace, P. (2006) Pointof-care nucleic acid lateral-flow tests. IVD Technol.
Liposome-Enhanced Lateral-Flow Assays for the Sandwich-Hybridization 25. Rule, G. S., Montagna, R. A., and Durst, R. A. (1996) Rapid method for visual identification of specific DNA sequences based on DNA-tagged liposomes. Clin Chem. 42, 1206–1209 26. Corstjens, P., Zuiderwijk, M., Brink, A., Li, S., Feindt, H., Niedbala, R. S., and Tanke, H. (2001) Use of up-converting phosphor reporters in lateral-flow assays to detect specific nucleic acid sequences: a rapid, sensitive DNA test to identify human papillomavirus type 16 infection. Clin Chem. 47, 1885–1893 27. Corstjens, P. L., Zuiderwijk, M., Nilsson, M., Feindt, H., Sam Niedbala, R., and Tanke, H. J. (2003) Lateral-flow and up-converting phosphor reporters to detect single-stranded nucleic acids in a sandwich-hybridization assay. Anal Biochem. 312, 191–200 28. Carney, J., Braven, H., Seal, J., Whitworth, E. (2006) Present and future applications of gold in rapid assays. IVD Technol. 12, 41–49 29. Oku, Y., Kamiya, K., Kamiya, H., Shibahara, Y., Ii, T., and Uesaka, Y. (2001) Development of oligonucleotide lateral-flow immunoassay for multi-parameter detection. J Immunol Methods. 258, 73–84 30. Zaytseva, N. V., Montagna, R. A., Lee, E. M., and Baeumner, A. J. (2004) Multi-analyte single-membrane biosensor for the serotypespecific detection of Dengue virus. Anal Bioanal Chem. 380, 46–53 31. Corstjens, P. L., Chen, Z., Zuiderwijk, M., Bau, H. H., Abrams, W. R., Malamud, D., Sam Niedbala, R., and Tanke, H. J. (2007) Rapid assay format for multiplex detection of humoral immune responses to infectious disease pathogens (HIV, HCV, and TB). Ann N Y Acad Sci. 1098, 437–445 32. Zhang, C., Zhang, Y., and Wang, S. (2006) Development of multianalyte flow-through and lateral-flow assays using gold particles and horseradish peroxidase as tracers for the rapid determination of carbaryl and endosulfan in agricultural products. J Agric Food Chem. 54, 2502–2507 33. Aldus, C. F., Van Amerongen, A., Ariens, R. M., Peck, M. W., Wichers, J. H., and Wyatt, G. M. (2003) Principles of some novel rapid dipstick methods for detection and characterization of verotoxigenic Escherichia coli. J Appl Microbiol. 95, 380–389 34. O’Farrell, B., Bauer, J. (2006) Developing highly sensitive, more-reproducible lateralflow assays Part 1: New approaches to old problems. IVD Technol. 35. Hans, H. B., Karl Pflanz, E. J., and Klewitz, T. M. (2002) Qualification of cellulose nitrate membranes for lateral-flow assays. IVD Technol.
213
36. Harlow, E., and Lane, D. (1988) Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratories, Cold Spring Harbor 37. Jones, K. (2001) Membrane immobilization of nucleic acids, Part 2: Probe Attachment Techniques. IVD Technol. September 38. Sischo, W. M., Atwill, E. R., Lanyon, L. E., and George, J. (2000) Cryptosporidia on dairy farms and the role these farms may have in contaminating surface water supplies in the northeastern United States. Prev Vet Med. 43, 253–267 39. Bangs Laboratories, I. (1999) Lateral Flow Tests. http://www.bangslabs.com/ technotes/303.pdf Accessed 40. Edwards, K. A., and Baeumner, A. J. (2006) Optimization of DNA-tagged dye-encapsulating liposomes for lateral-flow assays based on sandwich hybridization. Anal Bioanal Chem. 386, 1335–1343 41. Batteiger, B., Newhall, V. W. J., and Jones, R. B. (1982) The use of Tween 20 as a blocking agent in the immunological detection of proteins transferred to nitrocellulose membranes. J Immunol Methods. 55, 297–307 42. Jones, K. D. (1999) Troubleshooting protein binding in nitrocellulose membranes Part I: Principles. IVD Technol. 32 43. Van Dam, A. P., Van den Brink, H. G., and Smeenk, R. J. (1990) Technical problems concerning the use of immunoblots for the detection of antinuclear antibodies. J Immunol Methods. 129, 63–70. 44. Zampieri, S., Ghirardello, A., Doria, A., Tonello, M., Bendo, R., Rossini, K., and Gambari, P. F. (2000) The use of Tween 20 in immunoblotting assays for the detection of autoantibodies in connective tissue diseases. J Immunol Methods. 239, 1–11 45. Smits, H. L., Chee, H. D., Eapen, C. K., Kuriakose, M., Sugathan, S., Gasem, M. H., Yersin, C., Sakasi, D., Lai, A. F. R. F., Hartskeerl, R. A., Liesdek, B., Abdoel, T. H., Goris, M. G., and Gussenhoven, G. C. (2001) Latex based, rapid and easy assay for human leptospirosis in a single test format. Trop Med Int Health. 6, 114–118 46. Chan, C. P., Sum, K. W., Cheung, K. Y., Glatz, J. F., Sanderson, J. E., Hempel, A., Lehmann, M., Renneberg, I., and Renneberg, R. (2003) Development of a quantitative lateral-flow assay for rapid detection of fatty acid-binding protein. J Immunol Methods. 279, 91–100 47. Wang, S., Zhang, C., Wang, J., Zhang, Y. (2005) Development of colloidal gold-based flow-through and lateral-flow immunoassays for the rapid detection of the insecticide carbaryl. Anal Chim Acta. 546, 161–66
214
Edwards and Baeumner
48. Klewitz, T., Gessler, F., Beer, H., Pflanz, K., and Scheper, T. (2006) Immunochromatographic assay for determination of botulinum neurotoxin type D. Sensors and Actuators B: Chemical. 113, 582–589 49. Shyu, R. H., Shyu, H. F., Liu, H. W., and Tang, S. S. (2002) Colloidal gold-based immunochromatographic assay for detection of ricin. Toxicon. 40, 255–258 50. Greenwald, R., Esfandiari, J., Lesellier, S., Houghton, R., Pollock, J., Aagaard, C., Andersen, P., Hewinson, R. G., Chambers, M., and Lyashchenko, K. (2003) Improved serodetection of Mycobacterium bovis infection in badgers (Meles meles) using multiantigen test formats. Diagn Microbiol Infect Dis. 46, 197–203 51. Birnbaum, S., Uden, C., Magnusson, C. G., and Nilsson, S. (1992) Latex-based thin-layer immunoaffinity chromatography for quantitation of protein analytes. Anal Biochem. 206, 168–171 52. LaBorde, R. and O’Farrell, B. (2002) Paramagnetic-particle detection in lateral-flow assays. IVD Technol. 8, 36–41 53. Szoka, F., Olson, F., Heath, T., Vail, W., Mayhew, E., and Papahadjopoulos, D. (1980) Preparation of unilamellar liposomes of intermediate size (0.1–0.2 μmol) by a combination of reverse phase evaporation and extrusion through polycarbonate membranes. Biochim Biophys Acta. 601, 559–571 54. Szoka, F., Jr., and Papahadjopoulos, D. (1978) Procedure for preparation of liposomes with large internal aqueous space and high capture by reverse-phase evaporation. Proc Natl Acad Sci U S A. 75, 4194–4198 55. Edwards, K. A. and Baeumner, A. J. (2006) Sequential injection analysis system for the sandwich hybridization-based detection of nucleic acids. Anal Chem. 78, 1958–1966 56. Rule, G. S., Montagna, R. A., and Durst, R. A. (1997) Characteristics of DNA-tagged liposomes allowing their use in capillary-migration, sandwich-hybridization assays. Anal Biochem. 244, 260–269 57. Rongen, H. A. H., Bult, A., and vanBennekom, W. P. (1997) Liposomes and immunoassays. J Imm Meth. 204, 105–133 58. Esch, M. B., Baeumner, A. J., and Durst, R. A. (2001) Detection of Cryptosporidium parvum using oligonucleotide-tagged liposomes in a competitive assay format. Anal Chem. 73, 3162–3167 59. Locascio-Brown, L., Plant, A. L., Chesler, R., Kroll, M., Ruddel, M., and Durst, R. A. (1993) Liposome-based flow-injection
60.
61.
62.
63.
64.
65.
66.
67.
68.
69.
70.
71.
72.
immunoassay for determining theophylline in serum. Clin Chem. 39, 386–391 Edwards, K. A. and Baeumner, A. J. (2007) Synthesis of a liposome incorporated 1-carboxyalkylxanthine–phospholipid conjugate and its recognition by an RNA aptamer. Talanta. 71, 365–372 Edwards, K. A., and Baeumner, A. J. (2006) Optimization of DNA-tagged liposomes for use in microtiter plate analyses. Anal Bioanal Chem. 386, 1613–1623 Goral, V. N., Zaytseva, N. V., and Baeumner, A. J. (2006) Electrochemical microfluidic biosensor for the detection of nucleic acid sequences. Lab Chip. 6, 414–421 Deiman, B., van Aarle, P., and Sillekens, P. (2002) Characteristics and applications of nucleic acid sequence-based amplification (NASBA). Mol Biotechnol. 20, 163–179 Clemons, W. M., Jr., May, J. L., Wimberly, B. T., McCutcheon, J. P., Capel, M. S., and Ramakrishnan, V. (1999) Structure of a bacterial 30S ribosomal subunit at 5.5 A resolution. Nature. 400, 833–840 Cook, N. (2003) The use of NASBA for the detection of microbial pathogens in food and environmental samples. J Microbiol Methods. 53, 165–174 Birch, L., Dawson, C. E., Cornett, J. H., and Keer, J. T. (2001) A comparison of nucleic acid amplification techniques for the assessment of bacterial viability. Lett Appl Microbiol. 33, 296–301 Keer, J. T., and Birch, L. (2003) Molecular methods for the assessment of bacterial viability. J Microbiol Methods. 53, 175–183 Vandamme, A. M., Van Dooren, S., Kok, W., Goubau, P., Fransen, K., Kievits, T., Schmit, J. C., De Clercq, E., and Desmyter, J. (1995) Detection of HIV-1 RNA in plasma and serum samples using the NASBA amplification system compared to RNA-PCR. J Virol Methods. 52, 121–32 Simpkins, S. A., Chan, A. B., Hays, J., Popping, B., and Cook, N. (2000) An RNA transcription-based amplification technique (NASBA) for the detection of viable Salmonella enterica. Lett Appl Microbiol. 30, 75–79 Sakallah, S. A. (2000) Molecular diagnostics of infectious diseases: state of the technology. Biotechnol Annu Rev. 6, 141–61 Iqbal, S. S., Mayo, M. W., Bruno, J. G., Bronk, B. V., Batt, C. A., and Chambers, J. P. (2000) A review of molecular recognition technologies for detection of biological threat agents. Biosens Bioelectron. 15, 549–578 Wolfe, H. J. (1988) DNA probes in diagnostic pathology. Am J Clin Pathol. 90, 340–344
Liposome-Enhanced Lateral-Flow Assays for the Sandwich-Hybridization 73. Mac Kenzie, W. R., Hoxie, N. J., Proctor, M. E., Gradus, M. S., Blair, K. A., Peterson, D. E., Kazmierczak, J. J., Addiss, D. G., Fox, K. R., Rose, J. B., and et al. (1994) A massive outbreak in Milwaukee of cryptosporidium infection transmitted through the public water supply. N Engl J Med. 331, 161–167 74. Fayer, R., Trout, J. M., Lewis, E. J., Santin, M., Zhou, L., Lal, A. A., and Xiao, L. (2003) Contamination of Atlantic coast commercial shellfish with Cryptosporidium. Parasitol Res. 89, 141–145 75. Nime, F. A., Burek, J. D., Page, D. L., Holscher, M. A., and Yardley, J. H. (1976) Acute enterocolitis in a human being infected with the protozoan Cryptosporidium. Gastroenterology. 70, 592–598 76. Brown, T. A. (2002) Genomes, Wiley, New York 77. Somer, L., and Kashi, Y. (2003) A PCR method based on 16S rRNA sequence for simultaneous detection of the genus Listeria and the species Listeria monocytogenes in food products. J Food Prot. 66, 1658–1665 78. Fuchs, B. M., Glockner, F. O., Wulf, J., and Amann, R. (2000) Unlabeled helper oligonucleotides increase the in situ accessibility to 16S rRNA of fluorescently labeled oligonucleotide probes. Appl Environ Microbiol. 66, 3603–3607 79. Ban, N., Nissen, P., Hansen, J., Capel, M., Moore, P. B., and Steitz, T. A. (1999) Placement of protein and RNA structures into a 5 A-resolution map of the 50S ribosomal subunit. Nature. 400, 841–847 80. Nugen, S. R., Leonard, B., and Baeumner, A. J. (2006) Application of a unique server-based oligonucleotide probe selection tool toward a novel biosensor for the detection of Streptococcus pyogenes. Biosens Bioelectron. 22, 2442–2448 81. Small, J., Call, D. R., Brockman, F. J., Straub, T. M., and Chandler, D. P. (2001) Direct detection of 16S rRNA in soil extracts by using oligonucleotide microarrays. Appl Environ Microbiol. 67, 4708–4716 82. Chaney, R., Rider, J., and Pamphilon, D. (1999) Direct detection of bacteria in cellular blood products using bacterial ribosomal RNA-directed probes coupled to electrochemiluminescence. Transfus Med. 9, 177–188
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83. Chen, C. S., Baeumner, A. J., and Durst, R. A. (2005) Protein G-liposomal nanovesicles as universal reagents for immunoassays. Talanta. 67, 205–211 84. Plant, A. L., Brizgys, M. V., Locasio-Brown, L., and Durst, R. A. (1989) Generic liposome reagent for immunoassays. Anal Biochem. 176, 420–426 85. Edwards, K. A., and Baeumner, A. J. (2007) DNA-oligonucleotide encapsulating liposomes as a secondary signal amplification means. Anal Chem. 79, 1806–1815 86. Zuker, M. (2003) Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res. 31, 3406–3415 87. Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990) Basic local alignment search tool. J Mol Biol. 215, 403–410 88. Benson, D. A., Karsch-Mizrachi, I., Lipman, D. J., Ostell, J., and Wheeler, D. L. (2004) GenBank: update. Nucleic Acids Res. 32, D23–D26 89. Koppel, D. E. (1972) Analysis of macromolecular polydispersity in intensity correlation spectroscopy – method of cumulants. J Chem Phys. 57, 4814–4820 90. Frisken, B. J. (2001) Revisiting the method of cumulants for the analysis of dynamic lightscattering data. Appl Opt. 40, 4087–4091 91. Fiske, C. H., and Subbarow, Y. (1925) The colorimetric determination of phosphorus. J Biol Chem. 66, 375–400 92. (2001) RNeasy Mini Handbook, Qiagen 93. Boom, R., Sol, C. J., Salimans, M. M., Jansen, C. L., Wertheim-van Dillen, P. M., and van der Noordaa, J. (1990) Rapid and simple method for purification of nucleic acids. J Clin Microbiol. 28, 495–503 94. Bartlett, G. R. (1959) Phosphorus assay in column chromatography. J Biol Chem. 234, 466–468 95. Coyne, V., James, D., Reid, S., and Rybicki, E. (1998) Detection of nucleic acids by hybridization. http://www.mcb.uct.ac.za/hybridn. htm Accessed March 24 2007 96. Boutet, V., Delaunay, V., De Oliveira, M. C., Boquet, D., Grognet, J. M., Grassi, J., and Deverre, J. R. (2000) Real-time monitoring of the hybridization reaction: application to the quantification of oligonucleotides in biological samples. Biochem Biophys Res Commun. 268, 92–98
Chapter 14 Rapid Prototyping of Lateral Flow Assays Alexander Volkov, Michael Mauk, Paul Corstjens, and R. Sam Niedbala Summary Principles and characteristics of lateral flow strip assays are reviewed. Recent technology developments permit the use of inexpensive electronic readers for interrogating lateral flow strip test results, thus avoiding the inevitable variation and subjectivity of visual inspection to assess the capture of reporter-labeled analyte on test lines of the strip. Protocols for developing lateral flow assays are described, including two specific case studies for assaying cotinine (a small-molecule metabolite of nicotine) in a competitive format, and assaying HIV antibodies in a sandwich-type assay format. Key words: Lateral flow assay, Rapid assay development, Nitrocellulose strips, Immunochromatography
1. Introduction Lateral flow assays are familiar to almost everyone in the guise of home pregnancy tests and rapid tests performed in doctor’s offices. In the laboratory, lateral flow tests are now as ubiquitous as microtiter-based assays, and lateral flow assays are used to detect large proteins or antigens, small-molecule drugs, and more recently, nucleic acids (1). Lateral flow has been mostly employed for the qualitative detection of target analytes. In clinical applications, the use of urine and saliva is common because of the convenient and noninvasive sample collection. However, lateral flow is also routinely used to detect a number of blood-based markers. In all cases, the formatting of each test is designed to meet the requirements for analytical specificity and testing location. Lateral flow tests are very often utilized outside the laboratory, often
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_14
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under conditions of high humidity and extremes in temperature. Many lateral flow tests use reporter systems that are interpreted by eye. Commonly used reporters include gold particles or colored polystyrene/latex particles. According to the type of assay, the test indicator is based on the accumulation or depletion of reporter-labeled target at capture zones defined on the lateral flow strip. The capture zones are formed by immobilizing antibodies or antigens at specified areas of the strip. For visible tests, the increase or decrease in color signal at the capture zone(s) is interpreted by the user. Although the eye is remarkably sensitive, results are influenced by ambient light conditions, and by the subjective, user-specific nature of test interpretation. Consequently, lateral flow tests use an instrumented reader to interrogate the test strips. Such instrument readers have become quite inexpensive, and are even included in over-the-counter pregnancy test kits. The advantages of lateral flow assay technologies include the speed of the test, its ease of use, and that the equipment and materials are readily available. The disadvantages include the imprecision that is due in part to the subjective visual interpretation of test results and to the variability in quality and performance of materials from different sources and suppliers. Lateral flow test performance is sensitive to various components of the test. The main components of the lateral flow test include various porous pads and membranes through which the liquid sample, buffers, and reagents migrate due to capillary forces. In practice, the wetting and spreading of the various fluids is variable and exhibits less than complete coverage of the porous materials. Some portion of sample may not contact the test zones, thus resulting in a variable loss in signal and diminishes test reproducibility. This feature and others to be discussed are inherent limitations of lateral flow assays. Nevertheless, for qualitative and semiquantitative applications, lateral flow tests are often the best choice in terms of cost-performance. A lateral flow test may be prototyped through a series of steps. Although the materials used are often common to many types of tests, they should be selected and optimized on the basis of particular test objectives and other criteria. Figure 1. shows a typical lateral flow design. The assay is composed of a nitrocellulose membrane, a sample loading pad, a conjugate pad that contains dried buffer reagents as well as the reporter conjugate, and an adsorption pad. The sample is blotted on the porous sample loading pad and is in intimate contact with the porous conjugate pad. The conjugate pad is affixed to the nitrocellulose strip. Because of capillary forces, the liquids are drawn into and wick along the porous nitrocellulose membrane. Continuous capillary flow of fluid is sustained by the adsorption pad, often also called the sump pad. The nitrocellulose membrane is striped with test and control lines, which are zones of immobilized antibody or antigen to selectively capture species in the capillary flow.
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Fig. 1. Lateral flow tests are often constructed from a series of materials that sequentially overlap. The goal is to imbed all reagents so that a flowing sample rehydrates and moves all materials up a test strip. Analytes and reagents then interact in zones placed on the strip. The result is a rapid test that provides information easily visible to the eye.
Lateral flow tests are usually designed according to either of two well-established formats: (1) the competitive assay or (2) the sandwich-type assay. A competitive assay is generally used for detection of target analytes with a molecular weight less than 3,000 Da. In format, a capture antibody is immobilized on the lateral flow strip nitrocellulose. Methods for accomplishing this step are described in more detail later. To run the assay, sample is mixed with reagent containing a reporter conjugated to a derivative of the analyte. These two compete species compete to bind to the immobilized capture antibody as migrate through the lateral flow strip. After an excess of reagent flows past the capture zone, the signal is measured. In this format, the signal generated is inversely proportional to the amount of target analyte. The sandwich assay type format is used for larger target analytes. In this format, a capture antibody immobilized on the nitrocellulose binds to one epitope on the target, while a second reporter-labeled antibody free in solution binds to a different epitope of the target. The resulting “sandwich” complex is detected using a reporter. The signal generated in this assay design is directly proportional to the amount of analyte. Although the signal intensity indicates the concentration of analyte present in the sample, in practice lateral flow assays are inherently imprecise for reasons discussed. Therefore, they are used primarily as qualitative indicators for the presence or absence of target analytes. The cutoff in signal used to discriminate between negative samples and samples containing analyte is somewhat arbitrary and usually determined by those developing the test according to permissible or desired tolerance of false positive and false-negative test results. Test results with visible reports can be
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aided by developing a color chart to interpret signal levels. In some instances, a simple instrument reader can be employed to more objectively determine positive or negative results. Two specific examples are described to illustrate the construction of a typical lateral flow test. The first is for detecting a small molecular weight analyte, cotinine, which is a metabolite of nicotine. The second example is for detecting HIV antibodies using a test capture line formed by immobilizing recombinant antigens (from a commercial source) on the nitrocellulose membrane. Lateral flow strip structures and methods of operation should be designed and implemented in view of desired performance requirements and other constraints. These design criteria inputs reflect specific parameters along with performance objectives with respect to sensitivity, specificity, reproducibility, reliability, ease-of-use, and cost. These parameters and performance metrics should be quantified, even for qualitative tests. Cotinine is the major metabolite of nicotine and is useful for the determination of tobacco smoke exposure. In this case, the test is designed to detect exposure to second hand smoke. Active smokers generally have very high levels of cotinine in their body fluids, and subjects exposed to second-hand smoke are expected to show considerably reduced levels of cotinine. To further define the test, saliva is used as the sample. This further complicates testing, as saliva is a complex mixture of mucous-submandibular gland fluids (∼75%) and low viscosity-parotid gland fluids (∼25%). The test kit will thus need a collection device that reliably delivers the oral fluid sample to the lateral flow test sample loading pad. There are numerous companies that supply materials and components necessary to construct lateral flow tests, and unfortunately, there may be considerable variability in the quality and performance of these materials and components. The most crucial material is the nitrocellulose membrane, which supports the capture zones that are interrogated when the test is read. Subheading 2 lists critical components and potential sources for the materials. It is important to note the function of each component and where each reagent will be placed on the lateral flow strip. When considering the reagents to be used, they must be capable of being dried onto the materials (i.e., the conjugate pad or nitrocellulose membrane) and stored desiccated for long periods of time. Finally, before beginning work on a lateral flow test, some specialized equipment is needed to ensure precise delivery of reagents onto the lateral flow test materials. Specific procedures for using some of this equipment are described below in the examples. Note that the goal here is to describe equipment appropriate for the research bench. There are also companies that offer equipment capable of manufacturing large numbers of test strips in highly automated fashion.
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One of the most important considerations in specifying a lateral flow assay is the biologics (antibodies or antigens) to be used in the test. For example, the antibodies that will be targeting a particular analyte should be screened for the highest affinity and specificity. Similarly, capture antigen must also be of the highest purity. Any assay will have limited performance if these components are suboptimal. Although it is possible to directly test your candidate reagents in the lateral flow format, it may be more efficient to first screen prospective materials in another format such as microtiter plates. Using microtiter plates, a large number of candidates can be quickly assessed before working them into a lateral flow test. A large number of samples and replicates can be tested much more quickly in microtiter formats compared with lateral flow testing. There are two well-established sources of antibodies: polyclonal antibodies from immunized animals and monoclonal antibodies from cell cultures. The main advantages of polyclonal antibodies are their availability and low cost. Monoclonal antibodies are more expensive, and more labor and time-consuming to produce, but they can provide much better specificity and sensitivity for lateral flow tests than polyclonals. In recent years, a third source of antibodies has emerged – recombinant antibodies generated by the means of recombinant DNA technology. These antibodies can be generated faster than poly or monoclonal antibodies, they do not require immunization of animals, and can be raised against antigens that are toxic to animals, or which are poor immunogens or nonimmunogenic molecules. As with all assays, each reagent must be adjusted to optimize test performance. A lateral flow test that is visually read is complicated by the fact that the interpretation of test results is subjective, dependent on ambient conditions and the experience of the tester. For this reason, it is preferable to use an instrumented reader to interrogate lateral flow test results. Recently, new inexpensive readers have become available. One such device (Avagotech, Menlo Park, CA) utilizes an inexpensive CCD camera, such as those found in mobile phones along with simple electronics to capture images of the lateral flow strip and its capture lines. Other more sophisticated reader devices are also available. The costs for any reader are dependant upon the capabilities of the reader hardware and software. Newer systems are now under development for fluorescence and chemilumenescent analysis. This new generation of readers is expected to increase lateral flow test analytical capabilities considerably. Our laboratory has been using an Avagotech reader for evaluation of reagents for HIV and cotinine lateral flow assays. Figure 2. shows an example of results for a sample diluted in our prototype HIV antibody detection lateral flow test strips. The Avagotech reader allows the data to be plotted and coefficients of variation to be determined, making the screening more quantitative and less subjective.
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Fig. 2. This shows the results from a series of assays for HIV antibodies or cotinine, using the Avagotech RDx reader. Each assay is placed in the reader following test development. The absolute reflectance values were plotted for each assay standard tested.
2. Materials 2.1. Materials
1. Bovine serum albumin (BSA) – min. 96%, electrophoresis grade (Sigma). 2. Keyhole limpet hemocyanin (KLH), succinylated (Sigma). 3. Sulfo-NHS – N-hydroxysulfosuccinimide (Pierce). 4. EDC – 1-Ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (Pierce). 5. NHS-activated Sepharose™ 4 Fast Flow (GE Healthcare Biosciences). 6. Protein A, essentially salt-free, lyophilized (Sigma). 7. Colloidal gold, 40 nm (Arista). 8. Antigens: gp36 (soluble) and gp41 (soluble) ver. 1 (Arista). 9. Laminated backing cards (G&L Precision Die Cutting). 10. Nitrocellulose membranes: – UniSart CN90, CN140 or CN200 (Sartorius). – High Flow Plus HF75 or HF90 (Millipore). – Mdi CNPC-SS12 (Advanced Micro Devices). ° Conjugate pads: – Cellulose fiber 1281 (Ahlstrom). – Spunbonded polyester 6613 (Ahlstrom). ° Sample pad: glass fiber 8964 (Ahlstrom). ° Absorbent pad: noncompressed cotton 222 (Ahlstrom). ° Buffers:
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– PBS – Phosphate buffered saline. 10 mM Na2HPO4, 1.76 mM KH2PO4, 2.7 mM KCl, 140 mM NaCl, pH 7.2. – Label buffer: 0.05 M Na2HPO4, 1% BSA, and 0.1% NaN3, pH 7.4. 2.2. Equipment
Striping equipment: LINOMAT 5 semiautomatic sample application device from CAMAG (Wilmington, NC USA). Cutting equipment: Model SS-4 guillotine strip cutter from AZCO Corp (Fairfield, NJ, USA).
3. Methods 3.1. Obtaining Antibodies
The most straightforward approach to procuring antibodies for a lateral flow assay is to survey commercially-available materials and the published literature. If desired antibodies are not available, one will have to generate them. Below is a description of steps required to generate polyclonal antibodies.
3.1.1. Immunization
Although large molecules can be used for immunization of a suitable animal host directly, small molecules like cotinine require covalent coupling to good immunogens such as bovine serum albumin (BSA) or keyhole limpet hemocyanin (KLH). Covalent coupling requires the presence of suitable functional groups in the hapten molecule (1). These functional groups can be introduced by chemical modification of haptens or by using hapten derivatives already containing these groups. The cotinine project utilized a derivative, trans-4-cotininecarboxylic acid, for coupling to amino groups of succinylated KLH (see Note 1). Coupling cotinine derivative to KLH can be performed with the following protocol (Number 1): 1. Dissolve 1.668 mg of trans-4-cotininecarboxylic acid (100-fold molar excess of the hapten over KLH) in 0.5 mL of anhydrous DMF (see Note 2). 2. Add 1.974 mg of sulfo-NHS and 1.742 mg EDC (1.2-fold molar excess of each compound over the hapten) to the hapten solution and stir overnight at room temperature. 3. Dissolve 5 mg of KLH in 5 mL of PBS. 4. Cool down KLH on ice and add activated hapten drop-wise with continuous stirring. 5. Continue stirring for 72 h at 4°C. 6. Dialyze the reaction mixture at 4°C against 4× 600 mL of PBS changing buffer every 2 h (Pierce dialyzes cassette with 10,000 cut-off). 7. Store conjugate at −20°C.
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Polyclonal antibodies can be easily generated in research labs equipped to handle small animals or in specialized animal care facilities existing in many large universities. Alternatively, many companies will, for a moderate fee, perform immunization and collect small samples of antiserum for antibody testing. In recent years, we have utilized Lampire Biological Laboratories Inc. (Pipersville, PA) for generation of rabbit polyclonal antibodies using cotinine-KLH as an antigen. The immunization protocol included six antigen inoculations and two bleeds for testing antibody titers. 3.1.2. Antibody Purification
Whole serum can be used in preliminary evaluation of antibody performance, but the final lateral flow assay should be developed with purified antibody. Affinity chromatography with protein A or G can purify total antibodies from the serum, but purification on immobilized antigen is preferred, since it isolates only analytespecific antibodies (2). Using only antibodies specific to the target helps minimize background and increase assay sensitivity. Antigen immobilization on solid media can be achieved in a number of ways depending on the nature of the antigen and the type of functional groups present (3, 4). For example, antigens with amino groups can be coupled to NHS or CNBractivated beads. Epoxy-activated beads can be used for coupling to hydroxy, amino, or thiol groups. We coupled cotinine-BSA conjugate to NHS-activated Sepharose via amino groups of BSA. ELISA experiments with cotinine-BSA immobilized on microtiter plates showed that antibodies could readily interact with cotinine attached to the surface of BSA molecules. The protocol (Number 2) for purification of analyte-specific antibodies by affinity chromatography on immobilized antigen includes the following steps: 1. Wash 1 mL of NHS-Activated Sepharose with 10–15 volumes of cold 1 mM HCl (see Note 3). 2. Collect Sepharose beads by centrifugation for 5–10 min at 3,000–5,000 rpm. 3. Discard supernatant and wash Sepharose twice with 10–15 volumes of PBS (see Note 4). 4. Combine washed Sepharose and 500 μL of cotinine-BSA (0.5 mg) in a 2 mL microfuge tube and mix on rotator at 4°C overnight (see Note 5). 5. Block nonreacted NHS groups by incubating the Sepharose for 2–4 h with 0.2 mL of 1 M Tris–HCl pH 8.5. 6. Transfer the medium into a 5 mL polypropylene chromatography column. 7. Wash the column with 3 mL of 0.1 M Tris–HCl pH 8.5, followed by 3 mL of 0.1 M Na-acetate pH 4.5, 0.5 M NaCl. 8. Repeat washing cycle 3–6 times.
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9. Wash the column with 10 mL of PBS. 10. Add 5 mL of anti-cotinine serum to 5 mL of PBS (see Note 6). 11. Clarify sample by centrifugation in a microcentrifuge for 10 min at maximum speed (10,000 rpm and above) (see Note 7). 12. Load supernatant onto the column. 13. Collect the flow-through fraction and reapply it onto the column (see Note 8). 14. Wash the column with 5 mL of PBS to remove unbound proteins. 15. Prior to elution add 200 μL of 1 M Tris–HCl, pH 8–9 to each of the collection test tubes to neutralize low pH in the eluted fractions (see Note 9). 16. Elute antibodies with 10 mL of 0.1 M citric acid, pH 3.0. Collect 1 mL fractions. 17. Immediately reequilibrate the column with 10 mL of PBS and wash with 5 mL of PBS containing 0.05–0.1% sodium azide for long-term storage (see Note 10). 18. In each collected fraction, measure protein concentration by absorbance at 280 nm and antibody performance by microplate ELISA. If necessary, make serial dilutions of the fractions. 19. Pool fractions positive for functionality and dialyze against 20 volumes of PBS at 4°C, making 2–3 changes of buffer every 2–4 h (see Note 11). 20. Store purified antibody at −20°C. 3.2. Testing Antibody Performance
Antibody performance should be tested at all stages of lateral flow assay development, starting with the animal bleeds. ELISA of unfractionated antisera can help eliminate poorly performing antibodies and save time in assay development. The main goal of antibody testing is to evaluate potential assay sensitivity and specificity. Sensitivity can be evaluated crudely by testing serial dilutions of antisera to find the maximum dilution that gives detection signal of acceptable strength. These results should be used only as a guide since the outcome of this experiment depends both on antibody affinity and antibody titer in the serum.
3.2.1. Testing for Assay Sensitivity
The limit of detection is determined as the lowest concentration of cotinine on the standard curve for which the following condition is true: mean sample signal plus three standard deviations (3SD) < mean blank signal minus 3SD (the 3SD error bars should not overlap). The 3SD rule can be modified depending on the exact requirements for the assay. The 3SD rule provides 99.7% confidence, 2SD – 95%, and 4SD – 99.99%. Protocol (Number 3) outlines steps for determining the assay detection limit as follows:
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1. Prepare samples with 0, 0.2, 0.5, 1, 2, 4, 6, 8, 10, and 20 ng/mL cotinine. 2. Run cotinine ELISA with eight replicates for each sample. 3. Calculate the average absorbance value for each sample and 3SD. 4. Plot the average absorbance values and 3SD error bars on a graph (see Fig. 3.) or enter the data in a table (see Table 1.). 5. Find the lowest target concentration at which the 3SD error bar does not overlap with the blank. In our example, the blank minus 3SD was 1.51 and the closest average plus 3SD was 1.47, corresponding to a 6 ng/mL concentration. The “X” marks in Table 1 points at both values. The 6 ng/mL concentration is considered the detection limit for this assay. 6. Calculate the percent displacement using the following equation: % displacement = 100 × (absorbance value of blank−absorbance value of 10 ng/mL cotinine)/absorbance value of blank (see Note 12). 3.2.2. Testing for Assay Specificity
After the antibodies have demonstrated acceptable sensitivity, one should proceed to testing for specificity (cross-reactivity) with structurally similar compounds or compounds that are most likely to be present in the assay matrix. The assay specificity was determined by running the cotinine assay in the presence of compounds structurally similar to cotinine or likely to be present in human samples. Examples included caffeine, nicotinic acid, nicotinamide, and aspirin. Protocol (Number 4) provides testing for assay specificity with the following steps: 1. Prepare calibrators containing 0, 1, and 10 ng/mL cotinine (see Note 13). 2. Prepare samples with 10,000 ng/mL of each individual compounds: caffeine, nicotinic acid, nicotinamide, and aspirin (see Note 14). 3. Run cotinine ELISA with eight replicates for each sample.
Fig. 3. This is a typical image of a lateral flow test as seen using the Avagotech RDx reader. The assay may be measured as a fixed end point as shown or kinetically monitored during test development
Test for cotinine and a control analyte
Include control method on strip
Precision around cutoff value
Definition cutoff The cutoff is defined as the concentration where there is greater than ninety-five percent (95%) confidence that a positive result will be attained
Cotinine Cutoff target
Easy to use
3.0
4.0
5.0
6.0
7.0
8.0
2.5–20 ng/mL
Ability of trained users to correctly read instructions and execute assay with negative/positive controls
10 ng/mL
≤ 30% variation
(continued)
Report showing controls used properly = 3 sites/3 naïve operators n = 10 devices at each control
Precision study reports
Approved final
1. Approved evaluation protocol for control or dye line 2. Included in product instruction
Controls assuring adequate human TBD sample volume test-end measure = showing that the device flowed properly ready for evaluation
Approved study report from inhouse and field studies Approved clinical study with XX negatives, XX positives clinical samples
Total sampling and test result ≤ XX min
TBD
Total sampling and test result < XX min
Must be a complete system for on- Complete document reports site cotinine analysis
Rapid result
Complete system must be able to collect a oral fluid sample
2.0
Verification/validation means
Must be a complete system for on-site Cotinine analysis using oral fluids
Minimum specification
1.0
Target specification
Requirement
ID
Design input
Table 1 The following table contains a list of sample requirements used when planning to develop a rapid test for cotinine, the metabolite of nicotine
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All reagents and buffers enclosed – No leaking during testing according to product instruction no leaking during use
Visible indicator to demonstrate pouch integrity
Design to minimize the risk of Sample expressed into self-concontact between the operational tained vial person and the oral fluid
Sample interference panel
Cross-reactivity (analogous and ubiquitous compounds)
Clinical agreement
Easy to use and grip
Clean, hygienic
Nontoxic
10.0
11.0
12.0
13.0
14.0
15.0
16.0
17.0
18.0
Study reports from field studies should include evaluation of leaking occurrence with normal operation
1. Report – preliminary shipping study + accelerated stability 2. Approved final report for leak test of all vials in kit
Verification/validation means
Contains no toxic components
Acceptable survey results in field tests on collection device use and grip
Biocompatibility
Study report from operator interviews
Clinical protocol summary
≥95% agreement of cotinine rapid test and reference method Approved by customer in field testing
Nonclinical cross-reactivity report
Nonclinical interference report
Substances and levels to be tested are identified
Identify substances and levels to be tested
Sample expressed into separate vial. Verified through field study reports Sample does not overflow test cassette or test strip when transferred by pouring or pipetting
Desiccant changes color if compro- Desiccant included, color indicamised tion not required
All reagents and buffers enclosed – No leaking before test is run according to temperatures and humidity no leaking before use specifications on packaging
Minimum specification
9.0
Target specification
Requirement
Design input
ID
Table 1. (continued)
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Size and physical appearance of box:
Design and artwork
Positive control 5 – 10× equivalent Contains single use positive control solution within ±25% of target of drug drug concentration
Negative controls
Shelf life
Accuracy refers to clinical sensitivity and specificity when compared with the OTI Intercept product-line and GC-MS/MS
Precision-Intra-assay (%CV) (within-run reproducibility)
Obtain the following approvals:
Test operation temperature range
Test storage temperature range
21.0
22.0
23.0
24.0
25.0
26.0
27.0
28.0
29.0
30.0
Key to the planning process was to recognize and set goals for each aspect of the tests performance
0°C to +40°C (2 weeks storage without functional losses)
18–27°C Nonclinical report
Labeling claim
≥95% negative at 0 ng/mL, ≥ 95% positive at XXX% of cutoff
≥95% negative at 0 ng/mL, ≥ 95% positive at XXX% of cutoff FDA clearance?
510(k), on file submission
Sample collection device report
Sample collection report
Tests must be “substantially equiv- FDA 510(k) clearance with Interalent” for safety and efficacy cept® as the predicate device
≥18 months at 15–27°C
Contain single use negative control
Professional and functional looking box
Rapid sampling
20.0
Oral fluid sampling time ≤2 min
Built-in sample adequacy
19.0
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4. Calculate the average absorbance value for each sample. 5. Compare average absorbance values for 1 ng/mL cotinine and potential cross-reacting compounds. A compound is considered non cross-reactive if it generates an absorbance value higher than that of 1 ng/mL cotinine sample (see Note 14). Antibodies in all four sera tested in our laboratory demonstrated no crossreactivity for tested compounds. Table 2 shows results of this experiment. 3.3. Preparation of Label Reagents
Various types of labels can be used in lateral flow assays. One of the most common is colloidal gold. Protocol 5 describes preparation of gold–protein-A conjugate used in detection of anti-HIV antibodies and cotinine. In this method, protein A is passively absorbed on gold particles. Preparation of gold–protein-A conjugate is according to the following protocol (Number 5): 1. Prepare 0.0075 mg/mL solution of protein-A in water. 2. Add 300 μL to 40 mL of 40 nm gold particles in water (1.5–1.7 OD at 540 nm). 3. Incubate 15 min at room temperature with rotation. 4. Collect particles by centrifugation at 13,000 rpm for 40 min in Beckman rotor JA-14. 5. Resuspend particles in 40 mL of the label buffer. 6. Repeat Step 4. 7. Resuspend gold particles in 1.2 mL of the label buffer. Absorbance at 540 nm should be around 40–45 OD.
Table 2 This contains a list of physical components for lateral flow tests and their function Strip component
Function
Nitrocellulose
Contains binding and control zones for test
Backing card
Acts as solid substrate for nitrocellulose and pads
Conjugate pad
Contains immobilized conjugate
Sample pad
May contain dried buffer and is sample entrance into test
Sump pad
Acts to absorb fluids as they reach the top of the lateral flow strip
Dessicants
Maintains low humidity while storing lateral flow tests
Foil envelops
Acts as barrier for storing lateral flow tests
Sources are identified for each component; however, do not consider this list exhaustive. It may be helpful to cross reference Fig. 1 with each component
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8. Spray gold on the conjugate pad and dry at room temperature (see Note 15). 9. Store the conjugate pad at room temperature in airtight plastic bags with desiccants. 3.4. Preparation of Capture Reagents and Their Immobilization on the Membrane
The choice of capture reagents mainly depends on the type of the assay being developed. Detection of anti-HIV antibodies requires antigen present on the test line. Antigen can be the native protein, its fragment, or a synthetic peptide. Our choice was a combination of two major commercially available immunogens: HIV proteins gp41 and gp36. Antigens in this form do not require any special treatment and can be sprayed onto the membrane directly. Cotinine detection requires coupling of cotinine to a large molecule to anchor it to the membrane. We used a cotinine-BSA conjugate synthesized according to Protocol 1, where BSA was substituted for KLH. A control line is usually located a few millimeters downstream from the test line. Each assay may have its own specific requirements for the control line. In our experiments with gold–protein-A conjugate and human samples, the capture line containing anti-human antibodies for HIV or anti-rabbit for the cotinine assay were the best choice. First, it can confirm that human antibodies were present in the sample, and second, a gold-labeled control line indicates adequate sample and label flow – both are very important controls when interpreting negative samples. Protocol (Number 6) describing striping the nitrocellulose membrane with capture agents and includes the following steps: 1. Cut nitrocellulose membrane into 20 × 2.5 cm strips. 2. Mount one strip at a time on LINOMAT 5 sample application device. 3. Move the dosage turret so the test line would run along the middle of the strip (see Note 16). 4. Set parameters: speed −120 nL/s, strip length −180 mm, volume −25 μL. 5. Fill the syringe with 27–30 μL of the capture reagent (1 mg/mL). 6. Spray the capture reagent. 7. Move the dosage turret 3–4 mm from the test line (see Note 17). 8. Repeat steps 5 and 6 with the control reagent – anti-human antibodies (1 mg/mL) (see Note 18). 9. Incubate striped nitrocellulose for 18–24 h at 37°C. 10. Store at room temperature in airtight plastic bags with desiccants.
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Development of a new assay may require screening of many sources for nitrocellulose membranes, sample, conjugate and absorbent pads until components providing good performance are identified (some examples are given in Subheading 2). The first three components may require pretreatment to decrease background, prevent sample loss or improve label release. Some of the examples for pretreatments: incubation in 0.5–1% polyvinyl acetate, or 20–50 mM dibasic sodium phosphate, or 0.05–0.5% Triton X-100. Addition of sucrose and trehalose onto the conjugate pad is known to facilitate label release. 3.5. Assembling Assay Components
Striped nitrocellulose must be attached to a backing card together with sample, conjugate and absorbent pads for the assay to function efficiently. The assay components can be assembled according to the following protocol (Number 7): 1. Cut laminated backing cards into 20 × 6 cm strips. 2. Peel off protective release paper to expose the adhesive surface of the card and save the paper for step 7. 3. Carefully place nitrocellulose membrane on the adhesive surface about 2 cm from the left edge (see Note 19). 4. Place conjugation pad over the left edge of the nitrocellulose with 3–4 mm overlap. 5. Place sample pad over the conjugation pad and align it with the left edge of the backing card. 6. Place absorbent pad over the right edge of the nitrocellulose and align it with the right edge of the backing card. 7. Cover the assembled card with protective release paper from step 2 and carefully roll a rubberized roller over it to ensure good adhesion and contacts. 8. Cut the card into 3–5 mm wide strips using a guillotine strip cutter. 9. Store the strips in airtight plastic bags with desiccants.
3.6. Performing Lateral Flow Assays
Commercial lateral flow assays are assembled in plastic housings to simplify sample handling and to protect users from the contact with assay components and potentially dangerous samples. Samples are usually mixed with a diluent buffer and loaded onto the strips. After absorption of the sample, strips can be washed with a second chase buffer to decrease background and to increase sensitivity of detection. In a research environment, these tests can be run in 96-well plates with nitrocellulose strips dipped into wells containing samples. Regardless of the experimental setup, it is important to read assay results at a predetermined time interval. When nitrocellulose membrane exposed to air begins to dry, it pulls liquid from the absorbent pad, which contains excess reporter.
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Table 3 This contains a suggested list of equipment to be used when constructing lateral flow assays in a research laboratory environment Equipment
Function
Laminator
Used to attach pads and nitrocellulose to backing card
Slicer
Used to divide assay cards into test strips
Striper
Used to place reagents onto various assay substrates
Listed below are the functions for such equipment as well as potential commercial sources
Detection methods depend on the type of reporter used in the assay. Colloidal gold for example can be detected visually or with instruments capable of scanning lateral flow strips. Figure 2 shows an example of such a scan taken with RDx reader from Avagotech Inc. Interpretation of lateral flow results requires reading from both test and control lines. For an assay to be conclusive, the control line must indicate that the test developed properly. Table 3 lists assay interpretations for sandwich and competitive assay formats.
4. Notes 1. Some haptens may require a short spacer (2–8 carbon atoms) for coupling to a carrier protein to produce antibodies with good performance. This linker can be introduced either by modification of hapten with a bifunctional cross linking reagent or by synthesis of the hapten derivative. Making hapten variants with 2–3 spacers of different lengths increases the likelihood of producing antibodies with desired performance. 2. Calculations were performed based on KLH molecular weight of 100,000, as recommended by the manufacturer. 3. Several milliliters of affinity medium used repeatedly can be sufficient for the development and preliminary testing of an assay. Large-scale trials or extensive optimizations may require significant amounts of purified antibody. Affinity purification on that scale may benefit from standard chromatography equipment (columns, pumps, a UV/pH monitor and a fraction collector).
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4. Coupling buffer should not contain functional groups that can react with activated medium. Phosphate, carbonate, or MES can be used for coupling amino and carboxyl groups. 5. The duration and the temperature of this incubation depend on the coupling chemistry and the stability of the ligand. Small haptens can be incubated at room temperature for 2–4 h, while proteins may require 4°C and overnight incubation. 6. The amount of serum can be established in a series of trial purification experiments. Do not load more than 1/3 of the maximum binding capacity during routing purifications. 7. Alternatively, serum can be filtered through 0.22 or 0.45 µm syringe filter. To prevent sample loss, use filters with low protein binding membranes, such as Durapore (PVDF) from Millipore. 8. Loading with lower flow rate can be used instead of reloading the flow-through fraction. 9. Low pH in the eluted fraction should be neutralized as soon as possible to prevent protein denaturation. One should adjust the amount or pH of the neutralizing buffer if pH of the elution buffer is lowered. 10. Reuse the affinity media only when processing identical samples to avoid cross-contamination. 11. Alternatively, combined antibody fractions can be desalted using size exclusion chromatography columns such as PD-10 from GE Healthcare BioSciences. 12. Concentration of 10 ng/mL is a cut-off concentration for our assay. 13. Assay results for 0 and 10 ng/mL cotinine samples are used to calculate % displacement as a quality control step. 14. Concentrations of 1 ng/mL and 10,000 ng/mL were selected to ensure that cross-reactants do not approach the absorbance value at the detection limit. 15. Alternatively, the conjugate pad can be dipped in the conjugate solution and dried. 16. Exact location of the test line can vary and is a part of assay optimization. 17. Several test lines, each for a different analyte, can be sprayed on a single membrane. Maintain at least 2–3 mm spacing between lines. 18. Great care should be taken to prevent contamination of the test line with control reagents. Use dedicated syringes and rinse them with mild detergents before and after spraying. 19. Use gloves when handling assay membranes and strips. Hold at the edges or ends. Use gloves that are talc free. Dust from gloves or human hands will adversely affect test performance.
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References 1. P.L.A.M. Corstjens, M. Zuiderwijk, M. Nilsson, H. Feindt, R.S. Niedbala, and H. J. Tanke (2003) “Lateral-flow and up-converting phosphor reporters to detect single-stranded nucleic acids in a sandwich-hybridization assay” Anal. Biochem. 312:191–200
2. Antibody Purification Handbook. (18-1037-46) GE Healthcare Biosciences 3. Hermanson, G. T. (1996) Bioconjugate techniques. Academic, New York 4. Affinity Chromatography – Principles and Methods. (18-1022-29) GE Healthcare Biosciences
Chapter 15 Lateral Flow Colloidal Gold-Based Immunoassay for Pesticide Shuo Wang, Can Zhang, and Yan Zhang Summary In recent years, immunochromatographic lateral flow test strips are used as a popular diagnostic tool. There are two formats (noncompetitive and competitive) in gold-based immunoassay. Noncompetitive gold-based immunoassay also called sandwich assay is applied for the detection of large molecular mass. For small molecular mass such as pesticide, competitive format of lateral flow colloidal gold-based immunoassay is described in this chapter. The preparation of gold colloidal and the conjugation between antibody and gold colloidal are described. Hi-flow plus nitrocellulose membranes are separately coated with goat anti-rabbit IgG (control line) and hapten-OVA conjugate (test line). Thus, the degree of intensity of color of the test line is the reverse of the concentration of pesticide in the sample and the visual result is immediately observable. Colloidal gold-based immunoassay can also be applied for multianalysis in one test strip if the detected targets show different physico-chemical properties and their haptens show great differences in chemical structure. Key words: Lateral flow, Gold-based immunoassay, Non-competitive, Competitive, Multi-analysis.
1. Introduction Lateral flow tests are also called immunochromatographic strip (ICS) tests. They have been a popular platform for rapid tests since their introduction in the late 1980s. ICS tests are used for the specific qualitative or semiquantitative detection of many analyses including antigens, antibodies, and even the products of nucleic acid amplification tests. Urine, saliva, serum, plasma, or whole blood can be used as specimens. Test specificity can also be very high. The tests use colloidal gold, dye, or latex bead conjugates to generate signal. Early rapid tests used colored latex to form the visual signal, and some current versions continue to
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_15
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use this method. Latex is originally, and still is, the prime labeling method used in agglutination tests. This is because of its predisposition to agglutinate in the presence of binding components. For rapid tests, in which stability of the conjugate is critical for avoiding false positives, this predisposition to agglutinate can become a major problem. The use of colloidal particles as versatile and efficient templates for the immobilization of biomolecules has been recognized since early 1980s. Because of its superior stability, sensitivity, and precision and reproducibility of manufacture, colloidal gold is more suitable for use in rapid tests (1–5). Colloidal gold immunoassay has been developed and applied increasingly in various research field such as for the detection of hormone (pregnancy, fertility), virus (HIV, hepatitis B and C), and bacteria (streptococcus A and B) (6, 7). The nanocolloidal gold particles could replace the enzyme to be labeled to antibody in pesticide immunoassay since it has very large surface area and good biocompatibility and stability. Compared with enzyme immunoassay (EIA), colloidal gold-based immunoassay can be completed rapidly in one-step. When antibody labeled with colloidal gold particles is combined with corresponding antigen, the colored immuno-reactant can be visually detected. There are two formats in immunochromatographic strip tests: noncompetitive and competitive reaction (8). They can be explained graphically in Notes 1 and 2. The format of noncompetitive immunochromatographic test is also called sandwich assay, which is applicable for target analyte with more than one epitope (high molecular mass analyte). For the smaller molecular mass analyses, the competitive format assay can be used (9–11). The major steps of strip preparation and analysis of competitive format assay were shown in the Scheme 1. In this type of assays, the detector reagent is typically colloidal gold-labeled antibodies against the analyte. The capture line is normally analytes conjugated to a carrier protein immobilized on the membrane. Analytes in samples will compete with analytes immobilized on the membrane for binding to the detector antibody. The more analytes present in the sample, the more effectively it will be able to block the capture of colloidal gold-labeled antibodies. Hence, an increase in the amount of analytes in samples will result in a decrease in signal in the readout zone (test line). So far, the competitive strip tests have been mainly used for the detection of low molecular mass (hapten), especially for drugs of abuse or pesticide in agricultural products. The detection limits of this competitive assay can achieve to ppb level (nanograms of analyte per gram of sample). Here colloidal gold for conjugation is used as available detection system. The signal reagent is solubilized and binds to the antigen or antibody in the sample and moves through the membrane by capillary migration. The tests can be run individually or
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Scheme 1. The major steps of strip preparation and analysis of competitive format assay.
in limited-size batches. Results can usually be read in 5–15 min. All tests include an internal procedural control line that is used to validate the test result. The benefits of immunochromatographic tests include (a) user-friendly format, (b) very short time to get test result, (c) long-term stability over a wide range of climates, and (d) relatively inexpensive to make. These features make strip tests ideal for applications such as home testing, rapid point of care testing, and testing in the field for various environmental and agricultural analytes. The assembled strips are dried and packaged, making them stable for months when properly protected from moisture and excessive heat.
2. Materials 1. HAuCl4 (0.01%): 0.01 g HAuCl4 dissolved in 100 mL deionized water. 2. Trisodium citrate (1%): 1 g trisodium citrate dissolved in 100 mL de-ionized water. 3. pH 7.2, 10 mM phosphorate buffer (PBS) and 0.05% TweenPBS. 4. Conjugate storage buffer (pH 7.2): 2 mM sodium borate containing 1% BSA and 0.1% sodium azide (1% BSA is added in order to stabilize the gold nanoparticles).
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5. pH 9.0, 5 mM borate buffer is used as dilution buffer in optimal condition studies for conjugation between colloidal gold and antibody. 6. Species-specific anti-immunoglobulin antibodies (protein A or goat anti-rabbit IgG, coated as control line) and hapten conjugated with protein (coated as test line).
All the reagents should filter with 0.45 μm membrane Nitro-cellulose Hi-Flow plus membrane (HF 0900225), Nylon+ membrane (INYC 00010), Polyvinylidene fluoride membrane (IPVH 10100), semi-rigid polyethylene sheets, and filter paper were purchased from Millipore (Bedford, MA). Hydrogen tetrachloroaurate trihydrate (CAS Number: 16961-25-4) was purchased from Sigma (St. Louis. MO). Protein A-Sepharose 4B was purchased from Amersham Biosciences (Uppsala, Sweden). Camag Linomat 5 automatic sampler (CAMAG, Switzerland, Fig. 1) and Milli-Q purified water system were also used. The synthesis of carbaryl haptens (shown in Fig. 2) was performed as previously described by Wang et al. (12). The synthesis of endosulfan haptens (shown in Fig. 3) reported by Lee et al.
Fig. 1. The photo of Camag Linomat 5 sample application.
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Fig. 2. Chemical structures of carbaryl and the haptens used for immunogen (haptenA) and used for coating on the membrane as test line (haptenB).
Fig. 3. Chemical structure of endosulfan and the haptens used for immunogen (haptenA) and used for coating on the membrane as test line (haptenB).
was accomplished according to their procedures (13). The haptens were coupled to keyhole limpet hemocyanin (KLH) for used as immunogens or coupled to OVA for used as coating on the membrane. Antibodies were produced in rabbits as described by Wang et al. (12). Female white rabbits were immunized by intradermal
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and intramuscular injections of haptens conjugated with KLH. IgG from antiserum was purified by protein A-Sepharose 4B affinity chromatography (14).
3. Methods 3.1. Membrane Selection
For immunochromatographic strip (ICS) tests, membrane with high protein-binding capacity is used as support body. Different membranes and some binding properties are presented in Table 1. Nitrocellulose Hi-Flow plus membrane is often used widely. Nitrocellulose membranes are completely neutral, and their binding properties are independent of the pH of the immobilization solution (although pH can have an effect on both the solubility and immobilization efficiency of a particular protein). The immobilization buffer with pH 7.0–7.2 is chosen. Sometimes the surfactants and detergents such as Tween-20 and Triton-X100 in very low concentration are added in the buffer to reduce the background and nonspecific binding.
3.2. Preparation and Selection of Colloidal Gold
Gold colloids were prepared by controlled reduction of gold chloride with sodium citrate using the procedure described by Frens (15). The strength of color showing was closely related to the size and quality of the colloidal gold particles. The size of the colloidal gold particles was directly dependent on the amount of trisodium citrate used in its preparation process. The results were summarized in Table 2. It was found that if the diameter of gold particles were small (40 nm) were unstable, self-coagulation occurred. For small molecular like pesticide, the optimize size selection of diameter for colloidal gold particle was 40 nm.
Table 1 Binding propertities of different membrane polymers Membrane polymer
Mechanism of binding
Nitrocellulose
Electrostatic
Polyvinylidene fluoride (PVDF)
Hydrophobic
(Charged modified) Nylon
(Ionic) Electrostatic
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Table 2 Changes of diameter size and color of formed colloidal gold with different amount of trisodium citrate Amount of 1% trisodium citrate added in a 100 mL 0.01% gold chloride solution (mL)
Diameter size Color of colloidal of colloidal gold gold particles particles (nm)
2
Reddish
15
1.6
Salmon pink
25
1
Dark red
40
0.8
Purple-red
55
0.6
Purple
75
0.46
Purple-gray
100
0.352
Blue
150
The procedures for preparing colloidal gold with 40 nm diameter were as follows: Step 1: First, 100 mL of 0.01% HAuCl4 was boiled throughly for 5 min. Step 2: Then1 mL solution of 1% trisodium citrate was added under constant stirring. Step 3: It was observed that the color of the solution had changed in less than 2 min, then it was boiled for another 5 min. Step 4: After cooling, de-ionized water was added to the initial volume. The diameter of the particle was checked with a transmission electron microscope (Fig. 4). 3.3. Optimal Condition Study for Conjugation Between Colloidal Gold and Antibody
The pH of colloidal gold solution for labeling antibody was adjusted with 0.1 M K2CO3 or 0.1 M HCL (pH 9.0 for polyclonal antibody and pH 8.2 for monoclonal antibody). Here, polyclonal antibody was used. First, antibodies were purified with protein A sepharose-4B, then dialysis in pH 7.2 10 mM PBS. For conjugation, antibody was directly absorbed on colloidal gold particle surface, mediated mainly by London-van der Waals force and hydrophobic interaction (16). To have a strong adsorption between the gold and antibody conjugate, a preliminary titration was performed. The colloidal gold was formed in solution by virtue of a balance between electrostatic repulsion and Londonvan der Waals attraction among the particles. However, on addition of ionic substance, the attracting force became greater than the counteraction, which led to an aggregation accompanying a color change from red to blue. Coating the colloidal surfaces
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Fig. 4. Colloidal gold particle with 40 nm diameter observed by transmission electron microscope.
with protein molecules, such as SPA, antibody, and BSA, could prevent this instability. Optimal conditions of antibody concentration for the coating could be determined by comparing the adsorption between 520 and 580 nm. Anti-carbaryl antibody conjugated with colloidal gold was used as example, the procedures for conjugation between colloidal gold and anti-carbaryl antibody were as follows: Step 1: Gold colloidal suspension adjusted to pH 9.0 was pipetted into a series of tubes at 1 mL per each tube. Step 2: 100 μL antibody solution diluted in a series of concentration (0–0.12 mg/mL) was added to each colloidal gold solution. Step 3: After being incubated for 5 min, each tube received 0.1 mL of 10% NaCl and was shaken for 10 min. Step 4: Absorption of each tube at 520 and 580 (A520–A580) was determined. As shown in Fig. 5, the minimal antibody to stabilize 1 mL of gold colloidal suspension was approximately 8 μg anti-carbaryl antibody. The antibody was determined to be 120% of the minimum concentration to ensure complete reaction with the colloidal gold particles. Under gentle stirring in ice bath, anti-carbaryl antibody was added drop by drop to the gold colloidal suspension within 20 min. Step 5: After overnight incubation at 4°C, the mixture was centrifuged at 10,621 g at 4°C for 30 min, and the pellet was resuspended in conjugate storage buffer (2 mM sodium borate containing 1% BSA and 0.1% sodium azide) and diluted for use.
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Fig. 5. Optimization of conjugation between anti-carbaryl antibody and colloidal gold.
3.4. Preparation of Membrane Strip for Lateral-Flow
Nitrocellulose Hi-flow plus membrane was cut into sections (3.0 × 0.5 cm2), for the detection of small molecular (such as pesticide). The procedures of preparation of membrane strip were shown: Step 1: The membrane was coated with hapten-OVA conjugates in a volume of 1 μL containing 1 μg hapten conjugate as test line with Camag Linomat 5 automatic TLC sampler. The control line was coated with 0.5 μL of anti-rabbit IgG from goat diluted 1/100 in PBS buffer (pH 7.2). The distance between the two lines was 0.5 cm. Step 2: The coated test strips were dried at 37°C for 15 min in constant temperature desiccator. Step 3: The remaining protein-binding sites of the membrane were blocked by immersing the strips in PBS containing 1% BSA at 37°C for 30 min. Step 4: The test strips were washed with PBST and dried. The coated test strips were stored in a desiccator at 4°C.
3.5. Procedure and Optimization of Lateral-Flow GoldBased Immunoassay
Lateral-flow device used in this study was shown in Fig. 6. In lateral-flow device, the dried filter acted as an absorbent actively. The main purpose of the assay was to allow visual evaluation, thus it was only used as a qualitative assay to detect contamination at a threshold level. For this, the color intensity of the test strip must be high enough to be seen and enable observation of difference in color intensity between negative control and samples. For this purpose, optimization experiments were used to determine the optimal immobilization concentration of hapten conjugate, optimal ratio of gold–antibody conjugate mixed with antigen and incubation time. Optimal immunoreagent concentration was selected as a clear appearing in the negative control with the shortest time, and the comparison of the intensity of color among samples and control could be easily distinguished by eye.
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Fig. 6. The analytical device for lateral-flow immuno-gold assay. C: control line, T: test line.
The procedures were shown in the following: series dilutions of standard solutions in 5% MeOH (prepared in PBS-0.05% Tween buffer) were mixed with corresponded PAb colloidal gold conjugate at a certain ratio (9:1 or 8:1). After incubation for 5 min, a certain volume mixture (120–150 μL) was added to sample application site of the test strip. The liquid reagent migrated by capillary toward the test line and control line. After the liquid reagent passed through the reaction zone (test line and control line), different intensity of color on the test line were separately compared with negative control. The intensity of the color of the test line was the reverse of the concentration of detected target. As shown in Fig. 7, under optimized condition different concentrations of carbaryl were detected by strip test. 3.6. Validation of the Test Strip Immunoassay by Comparing with Instrumental Analysis
The reliability of the test strip immunoassay was determined by carrying out the test with the uncontaminated samples spiked with detected target at three levels and analyzed by strip test and instrumental analysis. The results obtained by strip test should be consistent with the results obtained by instrument. The correlation between the two methods should be good. An example of validation of gold-based lateral flow immunoassay and analysis of food samples for the detection of carbaryl was shown in the following.
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Fig. 7. Lateral-flow immunogold assay for the detection of carbaryl (C, control line; T, test line; carbaryl concentration A: control; B: 100 μg/L; C: 500 μg/L; D: 1000 μg/L).
Food samples were used for carbaryl screening and determination. Carbaryl was spiked at three levels in samples and extracted using methanol, which was earlier found to be an efficient extractant for this compound in food and environmental matrices. The samples were screening by lateral-flow gold-based immunoassay and confirming by HPLC (shown in Table 3). 3.7 Colloidal GoldBased Immunoassay for Multianalysis
Colloidal gold-based immunoassay could also be applied for multianalysis. It was known that if the detected targets show different physico-chemical properties and their haptens show great differences in chemical structure, multianalysis in one test strip is feasible. The different corresponding coating antigen for each pesticide could be fixed at different sites as respective test lines on the strip. As shown in Fig. 8, using lateral-flow gold-based assay for the detection of carbaryl and endosulfan, when the mixture of two gold–antibody conjugates was added to the reaction zone, both test lines (carbaryl line and endosulfan line) and control line had color development. When gold–antibody (specific to endosulfan) conjugate was added to the test lines, only the
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Table 3 Comparison of results obtained by lateral-flow gold-based immunoassay and HPLC
Matrix Chinese cabbage
Rice (grain)
Rice (Powder)
Barley (grain)
Barley (powder)
Results obtained by gold-based lateral- Results obtained Spiked level flow immunoassay by HPLC (mg/ (mg/kg) (n = 3)a kg)b 5
−,−,−
3.95
10
±, ±, ±
6.84
15
+, +, ±
10.16
5
−,−,−
3.57
10
±, +, ±
6.76
15
+, +, +
11.07
5
−,−,−
3.94
10
±, ±, ±
8.01
15
+, +, +
10.28
5
−,−,−
3.96
10
±, +, ±
7.78
15
+, +, +
10.58
5
−,−,−
3.89
10
+, ±, +
7.18
15
+, +, +
10.53
a
(+) positive: carbaryl concentration was more than 10 mg/kg; (±) positive/ negative: carbaryl concentration was around 10 mg/kg; (−) negative: carbaryl concentration was less than 10 mg/kg b HPLC confirmation for all samples was carried out according to the method described in the previous paper (17)
color of endosulfan test line appeared, and no color developed in carbaryl test line. It was also found that there was no color development in endosulfan test line when gold–antibody conjugate (specific to carbaryl) was added. For multianalyte colloidal gold immunoassay and single pesticide (carbaryl or endosulfan) colloidal gold immunoassay, the visual detection limits of carbaryl and endosulfan were the same for both formats. The existence of endosulfan (or carbaryl) did not affect the binding between carbaryl (or endosulfan) and anti-carbaryl (or endosulfan) antibody gold conjugate by comparing the visual results of single pesticide gold-based immunoassay and multianalysis gold-based
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Fig. 8. Multianalysis flow-through gold-based immunoassay. Up line (C): control line (goat anti-rabbit IgG coated); middle line (ET): endosulfan test line (endosulfan hapten-OVA coated); down line (CT): carbaryl test line (carbaryl hapten-OVA coated). (A) The mixture of two gold–antibody conjugates was added; (B) Anti-carbaryl antibody colloidal gold conjugate was added; (C) Anti-endosulfan antibody colloidal gold conjugate was added.
immunoassay. It should be feasible to detect multiple pesticides by combining different conjugates in one NC membrane.
4. Notes 1. Noncompetitive formats This format is used when testing for larger analyte with multiple antigenic sites, such as LH, hCG, and HIV. As shown in Scheme 2, in this case, less than an excess of sample analyte is desired, so that some of the antibody–gold conjugates will not be captured at the first line (test line, antibody coated), and will continue to flow toward the second line of immobilized antibodies (the control line). These are species-specific anti-immunoglobulin antibodies (usually protein A or goat anti-rabbit IgG), specific for the conjugate antibodies on the colloidal gold particles. 2. Competitive format for small molecular mass (pesticide) This is used most often when testing for small molecules (shown in Scheme 3) with single antigenic determinants, which cannot bind to two antibodies simultaneously. The assay is designed such that if there is no pesticide present in the test, the gold– antibody will accumulate on the nitrocellulose membrane where they are trapped by the immobilized pesticide hapten conjugate while liquid migrate towards the test strip. For samples containing pesticide, the binding sites on the specific antibody molecules will be occupied first by pesticide, leaving fewer binding sites for hapten-ovalbumin (OVA) conjugate on the membrane. Consequently, less colloidal gold-labeled antibody will remain at the hapten-OVA location on the nitrocellulose
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Scheme 2. Double antibody sandwich reaction (noncompetitive).
membrane. Thus, the degree of intensity of gold-color of the test line is the reverse of the concentration of pesticide in the sample and the visual result is immediately observable. If this format is chosen, it is important to pay close attention to the amount of antibody bound to the gold particles, in relation to the amount of free antigen in the sample.
Acknowledgments The authors are grateful for financial supports from the Ministry of Science and Technology of the People’s Republic of China (project No. 2006BAD05A06), the New Century Talent Program of Ministry of Education of the People’s Republic of China (project No. NECT-04-0243).
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Scheme 3. Competitive reaction.
Reference 1. Birnbaum, S. (1992) Latex-based thin-layer immuno-affinity chromatography for quatitation of protein analytes, Analytical Biochemistry, 206, 168–171 2. Chandler, J., Gurmin, T. and Robinson, N. (2000) The place of gold in rapid tests, IVDT, March 3. Penn, S.G., He, L. and Natan, M.J. (2003) Nanoparticles for bioanalysis, Current Opinion in Chemical Biology, 7(5), 609–615 4. Dykman, L.A. and Bogatyrev, V.A. (1997) Colloidal gold in solid-phase assays, A review. Biochemistry-Moscow, 62(4), 350–356 5. Rembaum, A. and Dreyer, W.J. (1980) Immunomicrosphere: reagents for cell labeling and separation, Science, 208, 364–368
6. Syllabus of a 2-day Seminar on Solid Phase Membrane-Based Immunoassay, Paris, September 25–26, (1997), Millipore Corporation, Bedford, MA 7. Syllabus of the Latex Course, London, October1–3, (1997) Organized by Bangs Laboratories, Inc., Fishers, Inc, USA 8. Weller, M.G. (2000) Immunochromatographic techniques: a critical review, Fresenius Journal of Analytical Chemistry, 366(6–7), 635–645 9. Verheijen, R., Stouten, P., Cazemier, G. and Haasnoot, W. (1998) Development of a one step strip test for the detection of sulfadimidine residues, Analyst, 123, 2437–2441
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10. Qian, S. and Bau, H.H. (2005) Magnetohydrodynamic stirrer for stationary and moving fluids, Sensors and Actuators B, Chemistry, 106, 859–870 11. Qian, S. and Bau, H.H. (2004) Analysis of lateral flow biodetectors: competitive format, Analytical. Biochemistry, 326, 211–224 12. Wang, S., Allan, R.D., Hill, A.S., Kennedy, I.R. (2002) Rapid enzyme immunoassays for the detection of carbaryl and methoprene in grain, Journal of Environmental Science Health, B37, 521–532 13. Lee, N.J., Skerrit, J.H., Mcadam, D.P. (1995) Hapten synthesis and development of ELISAs for detection of endosulfan in water and soil, Journal of Agricultural Food Chemistry, 43, 1730–1739
14. Akerstrom, B., Brodin, T., Reis, K., Bjorck, L. (1985) Protein G: a powerful tool for binding and detection of monoclonal and polyclonal antibodies, Journal of Immunology, 135, 2589–2593 15. Frens, G. (1973) Controlled nucleation for regulation of particle-size in monodisperse gold suspensions, Nature (London) Physical Science, 241, 20–22 16. Hermanson, G.T., Mallia, A.K. and Smith, P.K. (1992) Immobilized Affinity Ligand Techniques. Academic, San Diego 17. Wang, S., Zhang, C., Zhang, Y. (2005) Development of a flow-through enzyme-linked immunosorbent assay and a dipstick assay for the rapid detection of the insecticide carbaryl, Analytica Chimica Acta, 535, 219–225
Chapter 16 Synthesis of a Virus Electrode for Measurement of Prostate Specific Membrane Antigen Juan E. Diaz, Li-Mei C. Yang, Jorge A. Lamboy, Reginald M. Penner, and Gregory A. Weiss Summary Though relatively unexploited in biosensor applications, phage display technology can provide versatile recognition scaffolds for detection of cancer markers and other analytes. This chapter details protocols for covalent attachment of viruses directly to electrodes for reagent-free detection of analytes in real-time. Customization of binding specificity leverages selections with large phage display libraries prior to covalent attachment of the selected virus to the electrode. The methods described here utilize electrochemical impedance spectroscopy (EIS) to detect molecular recognition between M13 phage bound to a Au electrode and the following analytes: prostate specific membrane antigen (PSMA), positive and negative control antibodies (p-Ab and n-Ab, respectively). Because of a thick layer built on the Au electrode, the real impedance (Zre) increases reliably with S/N ratios upon noncovalent binding to PSMA (~14) and p-Ab (~20). Key words: Biosensor, Bacteriophage, Surface modification, Electrochemical impedance, Electrode.
1. Introduction Since its invention in 1985 (1), phage display has found widespread applications in diverse fields ranging from biochemistry to materials science (2). For example, the discovery and engineering of binding motifs has been revolutionized by phage and other molecular display technologies (3–5). The display of vast libraries on the phage surface (6–8) is now routine, and current diversities approach ~1012 unique variants (2, 9). Multiple rounds of in vitro binding to an immobilized target can tailor the specificity and affinity of the displayed protein. The technique directly links the
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_16
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DNA sequence encapsulated by the phage (the genotype) and the polypeptide displayed on the surface (the phenotype). The enormous diversities afforded by phage-displayed libraries allow selection for binding to essentially any molecular target. However, phage display applications in biosensors remain incompletely explored. For example, previous research has focused upon using phage-displayed proteins as the equivalent of large antibodies to trigger a detection event (10–15). This approach requires use of phage as added reagents. Reagent-free detection requires direct attachment of the phage to the sensor for interrogation of the analyte solution. In theory, real-time sensing can take place, if binding to the phage-coated sensor generates an electrical signal for measurement. We have recently constructed a biosensor through the covalent attachment of phage onto the surface of a gold electrode, and demonstrated direct electronic recognition of the prostate cancer marker, prostate specific membrane antigen (PSMA) (16), a positive antibody (p-Ab) known to bind M13 phage, but not a negative antibody (n-Ab) that fails to bind the phage. The virus electrode technique described above requires electrochemical impedance spectroscopy (EIS), a reagent and label-free, real-time measurement, with fast response and low cost. Previous applications of EIS include sensors based upon antibodies (17, 18), enzymes (19, 20), and DNA (21, 22). Several approaches can couple an EIS measurement with detection of analytes. First, amplification can leverage a redox probe such as [Fe(CN)6]3−/4− (23–27) or [Ru(NH3)6]2+/3+ (28, 29) to increase electron transfer resistance from the build-up of biolayers upon noncovalent binding. Second, in a similar approach, gold nanoparticles (30, 31) can enhance the capacitance of the electrodes during molecular recognition. Third, in a precipitation mode (32, 33), the enzyme horseradish peroxidase (HRP) reacts with target analytes producing an impermeable layer on the electrode, increasing resistance. Fourth, using a nano-gap device (34, 35), the capacitance change upon binding can be detected at very low frequencies in the 100 Hz range. Our research focuses on the direct EIS measurements in analyte solution without reagent addition using an arbitrarily shaped electrode. Our experiments demonstrate detection by increased resistance at high frequencies from 2 to 500 kHz with useful S/N ratios up to 20-fold.
2. Materials 2.1. Microorganisms and Enzymes
1. E. coli XL1-Blue (Stratagene, La Jolla, CA, USA). 2. E. coli CJ236 (New England Biolabs, Beverly, MA, USA).
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3. Electrocompetent E. coli SS320. 4. M13KO7 helper phage (New England Biolabs, Beverly, MA, USA). 5. T4 polynucleotide kinase (New England Biolabs, Beverly, MA, USA). 6. T4 DNA ligase (New England Biolabs, Beverly, MA, USA). 7. T7 DNA polymerase (New England Biolabs, Beverly, MA, USA). 2.2. Chemicals
All chemicals and solvents (>99% purity) were purchased from Fisher Scientific, Merck or Sigma-Aldrich and used as received, unless noted. 1. DMF and ethanol (100% from Gold Shield Chemical Company) were dried with 4 Å molecular sieves obtained from Alfa. 2. Anhydrous methylene chloride (CH2Cl2) was filtered through two columns of activated basic alumina and transferred under Ar (g) according to the method described by Grubbs (36). 3. 0.5 N and 0.05 N HCl solutions were made by diluting volumetric standard 2 N HCl with nanopure water. 4. Nanopure water (resistance ≈18 MΩ, Barnstead Inc.) was used in all experiments. 5. QIAprep Spin M13 Kit (Qiagen, Valencia, CA, USA). 6. QIAquick Gel Extraction Kit (Qiagen, Valencia, CA, USA). 7. Ultrapure glycerol (Invitrogen, Carlsbad, CA, USA). 8. Ultrapure irrigation USP water (Braun Medical Inc., Irvine, CA, USA). 9. Carbenicillin (5 mg/mL in water, filter sterilized). All antibiotic stocks were stored at −20°C after filter sterilization and aliquoting. 10. Chloramphenicol (30 mg/mL in ethanol, filter sterilized). 11. Kanamycin (40 mg/mL in water, filter sterilized). 12. Tetracycline (5 mg/mL in ethanol, filter sterilized). 13. Uridine (1 mg/mL in water, filter sterilized). 14. DTT (100 mM) (Sigma, St. Louis, MO, USA). 15. dNTPs (25 mM): Solution with 25 mM each of dATP, dCTP, dGTP, dTTP (Promega Corporation, Madison, WI, USA). 16. ATP (10 mM) (Amersham-Biosciences, Piscataway, NJ, USA). 17. 10% v/v ultrapure glycerol (100 mL ultrapure glycerol in 900 mL ultrapure irrigation USP water, filter sterilized). 18. BSA Fraction V (EM Science, Gibbstown, NJ, USA). 19. Tween-20 (Sigma, St. Louis, MO, USA).
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20.
D,L-Thioctic
acid (Acros Organics).
21. N-Hydroxysuccinimide (Acros Organics). 22. Dicyclohexylcarbodiimide (Lancaster Synthesis, Inc.). 23. 4-(1-Pyrrolidino)pyridine (Lancaster Synthesis, Inc.). 24. o-Phenylenediamine dihydrochloride (Acros Organics). 25. Diamond polishing compounds (1 μm and 0.25 μm size, Ted Pella). 2.3. Medium for Growth
1. 2YT media (10 g bacto-yeast extract, 16 g bacto-tryptone, 5 g NaCl; water added to 1 L and pH adjusted to 7.0 with NaOH; autoclaved). 2. LB (5 g bacto-yeast extract, 10 g bacto-tryptone, 10 g NaCl; water added to 1 L; autoclaved). 3. LB agar (5 g bacto-yeast extract, 10 g bacto-tryptone, 10 g NaCl, 15 g granulated agar; water added to 1 L; autoclaved). 4. 2YT/carb/cmp media (2YT, 50 μg/mL carbenicillin, 5 μg/ mL chloramphenicol). 5. 2YT/carb/kan/uridine media (2YT, 50 μg/mL carbenicillin, 25 μg/mL kanamycin, 0.25–2.0 μg/mL uridine). 6. LB/carb plates (LB agar, 50 μg/mL carbenicillin). 7. LB/tet plate (LB agar, 5 μg/mL tetracycline). 8. LB/kan plate (LB agar, 25 μg/mL kanamycin). 9. SOC media (5 g bacto-yeast extract, 20 g bacto-tryptone, 0.5 g NaCl, 0.2 g KCl; water added to 1 L and pH adjusted to 7.0 with NaOH; autoclaved; add 5.0 mL of autoclaved 2.0 M MgCl2 and 20 mL of filter sterilized 1.0 M glucose).
2.4. Buffers
1. PBS (137 mM NaCl, 3 mM KCl, 8 mM Na2HPO4, 1.5 mM KH2PO4; pH adjusted to 7.2 with HCl; autoclaved). 2. PEG/NaCl (20% PEG-8000 w/v, 2.5 M NaCl; autoclaved). 3. TAE buffer (40 mM Tris-acetate, 1 mM EDTA; pH adjusted to 8.0; autoclaved). 4. TAE/agarose gel (TAE buffer, 1% w/v agarose, 1:5,000 v/v 10% ethidium bromide). 5. TM buffer (10×) (0.1 M MgCl2, 0.5 M Tris, pH 7.5). 6. 0.2% BSA in PBS (pH 7.2). The buffer was stored at 4°C. 7. PBF, pH 7.2 (4.2 mM Na2HPO4, 1.5 mM KH2PO4, 140 mM NaF, filter sterilize). 8. PBF wash buffer (0.06% BSA and 0.07% Tween-20) were added to PBF immediately before use. Make fresh as needed. 9. 0.2% BSA in PBF (pH 7.2). Stored at 4°C.
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10. PT buffer (PBS, 0.05% Tween-20). 11. PBT buffer (PBS, 0.1% BSA, 0.05% Tween-20). Stored at 4°C. 12. PBF/TW (PBF, 0.05% Tween-20). 2.5. Target Bio-Molecules
1. Anti-M13 antibody (p-Ab) was purchased from AmershamBiosciences, Piscataway, NJ, USA). 2. Horseradish peroxidase/anti-M13 antibody (Amersham-Biosciences, Piscataway, NJ, USA).
conjugate
3. Anti-Flag® M2 (n-Ab) was purchased from Sigma. 4. Prostate specific membrane antigen (PSMA) (see Note 1). 2.6. Instruments
1. Floor-model centrifuge (Sorvall) with SS-34 and SLA-1500 rotor (or equivalent). 2. 0.2-cm gap electroporation cuvet (BTX, San Diego, CA, USA). 3. ECM-600 electroporator (BTX, San Diego, CA, USA). 4. 96-well maxisorp immunoplates (NUNC, Roskilde, Denmark). 5. Ultrasonic machine: Fisher Scientific FS20. 6. 0.22 mm filter: Fisher Scientific. 7. Electrochemical Measurement: Parr 2263 (Princeton Applied Research Inc.) with software Power CV and Power Sine. 8. pH meter: Fisher Scientific. 9. Shaker: Orbital. 10. Electrode: Circular Au electrode from Bioanalysis System. 11. Microcloth: From Buehler.
3. Methods 3.1. Phage Library Synthesis 3.1.1. Constructing X8 Naïve Library
Libraries of P8 variants are constructed using an optimized version (4) of a previously described oligonucleotide-directed mutagenesis method (37). First, a mutagenic oligonucleotide (sequence: GCTACAAATGCCTATGCATAATAATGATGAGGTGGAGGATCCGGCGGA) is used to introduce stop codons at the sites to be randomized; oligonucleotide-directed site-specific mutagenesis protocols are provided in Subheading 3.1.3, and can be adapted for small scale mutagenesis to introduce the stop codons (see Note 2). The resulting “stop template” phagemid can be used as the template for library construction because the presence of stop codons eliminates wild-type (wt) protein display. Uracil-containing ssDNA (dU-ssDNA) stop template
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(purified from an E. coli dut−/ung− host) is then annealed to a mutagenic oligonucleotide (sequence: GCTACAAATGCCTATGCANNSNNSNNSNNSNNSNNSNNSNNSGGTGGAGGATCCGGCGGA) designed to replace the stop codons with NNS (N = A/G/C/T, 25% each; S = G/C, 50% each) degenerate codons that encode all 20 natural amino acids. The mutagenic oligonucleotide is used to prime the synthesis of a complementary DNA strand, which is ligated to form a covalently closed circular, double-stranded DNA (CCC-dsDNA) heteroduplex. To complete the library construction, the CCC-dsDNA heteroduplex is introduced into an E. coli dut+/ung+ host by electroporation and the mismatch is repaired to either the wild-type or mutant sequence. In an ung+ strain, the uracil-containing template strand is preferentially inactivated and the synthetic, mutant strand is replicated, thus resulting in efficient mutagenesis (> 50%). The use of a template with stop codons at all of the sites to be randomized ensures that only fully mutagenized clones contain open reading frames that can be displayed on phage. The library members can be packaged into phage particles by coinfection of the E. coli host with a helper phage. 3.1.2. Purification of dUssDNA Template
Mutagenesis efficiency depends on template purity, and thus, the use of high purity dU-ssDNA is critical for successful library construction. We use the Qiagen QIAprep Spin M13 Kit for dUssDNA purification, and the following is a modified version of the Qiagen protocol. It yields at least 20 μg of dU-ssDNA for a medium copy number phagemid (e.g., pS1607, which contains a pBR322 backbone), and this is sufficient for the construction of one library (see Note 3).
Purifying dU-ssDNA Template
1. From a fresh LB-antibiotic plate, pick a single colony of E. coli CJ236 (or another dut−/ung− strain) harboring the appropriate phagemid into 1 mL of 2YT media supplemented with M13KO7 helper phage (1010 pfu/mL) and appropriate antibiotics to maintain the host F′ episome and the phagemid. Shake at 250 rpm and 37°C for 2 h and add kanamycin (25 μg/mL) to select for clones that have been coinfected with M13KO7, which carries a kanamycin resistance gene. Shake at 250 rpm and 37°C for 6 h and transfer the culture to 30 mL of 2YT/carb/kan/uridine media. Shake overnight at 250 rpm and 37°C. 2. Centrifuge for 10 min at 15 krpm and 4°C in a Sorvall SS-34 rotor (27,000 × g). Transfer the supernatant to a new tube containing 1/5 volume of PEG/NaCl, mix gently and incubate for 5 min at room temperature. Centrifuge 10 min at 10 krpm and 4°C in an SS-34 rotor (12,000 × g). Decant the supernatant. Centrifuge briefly at 4 krpm (2,000 × g) and aspirate the remaining supernatant.
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3. Resuspend the phage pellet in 0.5 mL of PBS and transfer to a 1.5-mL microcentrifuge tube. Centrifuge for 5 min at 13 krpm in a microcentrifuge (15,000 × g), and transfer the supernatant to a 1.5-mL microcentrifuge tube. 4. Add 0.7 mL of buffer MP (Qiagen) and mix. Incubate at room temperature for at least 2 min. 5. Apply the sample to a QIAprep spin column (Qiagen) in a 2-mL microcentrifuge tube. Centrifuge for 30 s at 8 krpm in a microcentrifuge (6,000 × g). Discard the flow-through. The phage particles remain bound to the column matrix. 6. Add 0.7 mL buffer MLB (Qiagen) to the column. Centrifuge for 30 s at 8 krpm and discard the flow-through. 7. Add another 0.7 mL buffer MLB. Incubate at room temperature for at least 1 min. Centrifuge at 8 krpm for 30 s. Discard the flow-through. The DNA is separated from the protein coat and remains adsorbed to the matrix. 8. Add 0.7 mL buffer PE (Qiagen). Centrifuge at 8 krpm for 30 s and discard the flow-through. 9. Repeat step 8. Residual proteins and salt are removed. 10. Centrifuge at 8 krpm for 30 s. Transfer the column to a fresh 1.5-mL microcentrifuge tube. 11. Add 100 μL of buffer EB (Qiagen; 10 mM Tris–Cl, pH 8.5) to the center of the column membrane (see Note 4). Incubate at room temperature for 10 min and centrifuge for 30 s at 8 krpm. Save the eluant, which contains the purified dU-ssDNA. 12. Analyze the DNA by electrophoresing 1.0 μL on a 1% TAE/ agarose gel (see Note 5). 13. Determine the DNA concentration by measuring absorbance at 260 nm (A260 = 1.0 for 33 ng/μL of ssDNA). Typical DNA concentrations range from 200 to 500 ng/μL. 3.1.3. In Vitro Synthesis of Heteroduplex CCC-dsDNA
A three-step procedure is used to incorporate the mutagenic oligonucleotide into heteroduplex CCC-dsDNA, using dU-ssDNA as a template. The oligonucleotide is first 5′-phosphorylated and then annealed to a dU-ssDNA template. The oligonucleotide is enzymatically extended and ligated to form heteroduplex CCCdsDNA, which is then purified and desalted. The protocol below produces ∼20 μg of highly pure, low conductance CCC-dsDNA. This is sufficient for the construction of a library containing more than 1010 unique members (see Note 6).
Oligonucleotide Phosphorylation with T4 Polynucleotide Kinase
1. In a 1.5-mL microcentrifuge tube, combine 0.6 μg of the mutagenic oligonucleotide, 2.0 μL of 10× TM buffer, 2.0 μL of 10 mM ATP, 1.0 μL of 100 mM DTT. Add water to a total volume of 20 μL.
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2. Add 20 units of T4 polynucleotide kinase. Incubate for 1.0 h at 37°C (see Note 7). Annealing of the Oligonucleotide to the Template
1. To the 20 μL phosphorylation reaction mix, add 20 μg of dU-ssDNA template (see Note 8), 25 μL 10× TM buffer, and water to a final volume of 250 μL. 2. Incubate at 90°C for 1–2 min, 50°C for 3 min, 20°C for 5 min, and place on ice (see Note 9).
Enzymatic Synthesis of CCC-dsDNA
1. To the annealed oligonucleotide/template mixture, add 10 μL 10 mM ATP, 10 μL 25 mM dNTPs, 15 μL 100 mM DTT, 30 Weiss units T4 DNA ligase, 30 units T7 DNA polymerase. 2. Incubate at 20°C for at least 3 h. Overnight incubation is preferred. 3. Electrophorese 1.0 μL of the reaction product alongside the dU-ssDNA template. Use a 1% TAE/agarose gel with ethidium bromide for DNA visualization. If the template DNA is still present, the concentration of the mutagenic oligonucleotide can be increased. 4. Affinity purify and desalt the DNA using the Qiagen QIAquick DNA Purification Kit. Add 1.0 mL of buffer QG (Qiagen) and mix. 5. Apply the sample to two QIAquick spin columns placed in 2-mL microcentrifuge tubes. Centrifuge at 13 krpm for 1 min in a microcentrifuge. Discard the flow-through. 6. Add 750 μL buffer PE (Qiagen) to each column. Centrifuge at 13 krpm for 1 min. Discard the flow-through and centrifuge at 13 krpm for 1 min. Place the column in a new 1.5-mL microcentrifuge tube. 7. Add 35 μL of ultrapure irrigation USP water to the center of the membrane. Incubate at room temperature for 2 min. 8. Centrifuge at 13 krpm for 1 min to elute the DNA. Combine the eluants from the two columns. The DNA can be used immediately for E. coli electroporation, or it can be frozen for later use. 9. Electrophorese 1.0 μL of the eluted reaction product alongside the dU-ssDNA template. Use a 1% TAE/agarose gel with ethidium bromide for DNA visualization.
3.1.4. E. coli Electroporation and Phage Propagation
To complete the library construction, the heteroduplex CCCdsDNA must be introduced into an E. coli host that contains an F′ episome to enable M13 bacteriophage infection and propagation. Phage-displayed library diversities are limited by methods for introducing DNA into E. coli, with the most efficient method being high-voltage electroporation.
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1. Chill the 20 μg of purified DNA and a 0.2-cm gap electroporation cuvet on ice. Thaw a 350 μL aliquot of electrocompetent E. coli SS320 on ice. Add the cells to the DNA and mix by pipetting several times (avoid introducing bubbles). 2. Transfer the mixture to the cuvette and electroporate. For electroporation, follow the manufacturer’s instructions, preferably using a BTX ECM-600 electroporation system with the following settings: 2.5 kV field strength, 125 Ω resistance, and 50 mF capacitance. Alternatively, a Bio-rad Gene Pulser can be used with the following settings: 2.5 kV field strength, 200 Ω resistance, and 25 mF capacitance. 3. Immediately, rescue the electroporated cells by adding 1 mL of ice-cold SOC media and transferring to a chilled 250 mL baffled flask containing 20 mL of ice-cold SOC media. Rinse the cuvette four more times with 1 mL ice-cold SOC media. Incubate for 20 min at 37°C with shaking at 250 rpm. 4. To determine the library diversity, plate serial dilutions on LB/carb plates to select for the library phagemid (in the case of b-lactamase encoding phagemids such as pS1607). 5. Add M13KO7 (4 × 1010 pfu/mL) and incubate for 20 min at 37°C with shaking at 250 rpm. 6. Transfer the culture to a 2 L baffled flask containing 500 mL 2YT media, supplemented with antibiotic for phagemid selection (e.g. 2YT/carb media). 7. Incubate 1 h at 37°C with shaking at 250 rpm and add kanamycin (25 μg/mL). Incubate overnight at 37°C with shaking at 250 rpm. 8. Centrifuge the culture for 10 min at 10 krpm and 4°C in a Sorvall SLA-1500 rotor (16,000 × g). Transfer the supernatant to a fresh tube and add 1/5 volume of PEG/NaCl solution to precipitate the phage. Mix gently and incubate for 5 min at room temperature. 9. Centrifuge for 10 min at 10 krpm and 4°C in a SLA-1500 rotor. Decant the supernatant. Re-spin briefly and remove the remaining supernatant with aspiration. Resuspend the phage pellet in 1/20 volume of PBS. 10. Pellet insoluble matter by centrifuging for 5 min at 15 krpm and 4°C in an SS-34 rotor (27,000 × g). Transfer the supernatant to a clean tube. 11. Estimate the phage concentration spectrophotometrically (OD268 = 1.0 for a solution of 5 × 1012 phage/mL), (see Note 10). 3.2. Selecting Phage from the X8 Library Against PSMA
Phage from the X8 library described above is cycled through rounds of binding selection with PSMA (38) coated on 96-well Maxisorp immunoplates as the capture target. Phage were propagated in
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E. coli XL1-Blue with M13KO7 helper phage for further rounds of selection. 3.2.1. Coating Wells with Target Protein
1. Coat 24 wells of a 96-well Maxisorp plate with 100 μL of a 10 μg/mL solution of the target protein (e.g. PSMA). Shake at room temperature for 1 h (see Note 11). Discard the solution. 2. To block nonspecific binding to the microtiter wells, add 300 μL of 0.2% BSA in PBS to each well. Shake at room temperature for 1 h. Discard the blocking solution.
3.2.2. Phage Library Selections
1. Add a solution of the phage library (∼1012 phage/mL) in PBT buffer to the wells. Shake at room temperature for 2 h. Wash the plate five times with PT buffer. The stringency of the binding selection can be increased for successive rounds by increasing the number of washes. 2. Elute bound phage, by adding 100 μL of 100 mM HCl to each well. Shake vigorously for 5 min at room temperature. 3. Combine the eluants, and neutralize with one-third volume of 1.0 M Tris–HCl, pH 8.0.
3.2.3. Propagating Phage for the Next round of Selection
1. Add half the eluted phage to ten volumes of log-phase XL1Blue cells (OD600 = 0.5–1.0). 2. Shake at 250 rpm, 37°C for 20 min. Remove 10 μL and determine library diversity by plating 1:10 serial dilutions on LB/ carb plates to select for the library phagemid. 3. Add M13KO7 helper phage (1010 pfu/mL) and shake at 250 rpm, 37°C for 30 min. 4. Transfer the culture to ten volumes 2YT/carb/kan media and shake overnight at 250 rpm and 37°C. 5. Isolate the phage by PEG/NaCl precipitation, as described in Subheading “Purifying dU-ssDNA Template” step 2. Resuspend phage in 1/20 volume of PBS. 6. Repeat the selection process five times, using only half of the eluted phage in each round and changing the blocking agent between each round.
3.3. Screening PSMA Binding Peptide Display Levels by Spot Assay
1. Infect 150 μL of log-phase (OD600 ∼0.6) E. coli XL1-Blue cells, grown from a fresh LB/tet plate, with 1.5 μL of Round 4 eluted phage. Incubate cells for 20 min at 37°C and 250 rpm. 2. Spread 25, 50, and 75 μL of infected cells onto three separate LB/carb plates and incubate overnight at 37°C. 3. The following morning, pick 24 isolated colonies from the plate that is less crowded and place each colony into separate wells of a 96-well block containing 1 mL of 2YT/carb, supplemented with M13KO7 helper phage (1010 pfu/mL). Shake overnight at 250 rpm and 37°C.
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4. The next morning, coat, and block 24 wells of a 96-well Maxisorp immunoplate as detailed in Subheading 3.2.1. 5. Centrifuge the 96-well block containing infected cells for 20 min at 2.5 krpm and 4°C in a Beckman Allegra 6-R centrifuge with swinging trays (1427 × g). 6. Transfer 100 μL of the supernatant to the corresponding wells on the 96-well plate. 7. Shake at room temperature for 2 h. Wash the plate five times with PT buffer. 8. Decant the phage solution and wash eight times with PT buffer. 9. Add 100 μL of horseradish peroxidase/anti-M13 antibody conjugate (diluted 5,000-fold in PBT buffer). 10. Shake for 30 min at room temperature. 11. Wash three times with PT, followed by two times with PBS buffer. 12. Develop the wells with 100 μL of o-phenylenediamine dihydrochloride (1 mg/mL, 0.02% H2O2) solution. Read spectrophotometrically at 450 nm in a microtiter plate reader. 3.4. Growth of Bacteriophage
1. From a fresh LB/tet plate, pick a single colony of E. coli XL1Blue cells harboring the appropriate phagemid into 1 mL of 2YT media supplemented with M13KO7 helper phage (1010 pfu/mL), 50 μg/mL carbenicillin (for phagemid maintenance), and 5 μg/mL tetracycline (for F′ episome maintenance). 2. Shake at 250 rpm and 37°C for 6–8 h. 3. Add M13KO7 helper phage (1010 pfu/mL) and shake for 30 min at 250 rpm and 37°C. 4. Transfer the culture to 30 mL of 2YT/carb/kan media. Shake overnight at 250 rpm and 37°C. 5. Isolate the phage as detailed in Subheading “Purifying dU-ssDNA Template” steps 2 and 3. 6. Determine the phage concentration spectrophotometrically (OD268 = 1.0 for a solution of 5 × 1012 phage/mL).
3.5. Synthesis of Thioctic NHS-Ester
Thioctic NHS-ester (NHS-TE) is not commercially available, but can be synthesized by the following previously published procedure (39). 1. Add 1.0064g (4.878 mmol) of thioctic acid and 0.7062 g (6.136 mmol) NHS to a 25 mL round bottom flask containing 5 mL of dry DCM. 2. Dissolve 1.7287g of DCC (8.378 mmol) and 0.0639g of 4-pyrrolidinopyridine (0.812 mmol) in 5 mL of DCM (see Note 12). Mix until the solution is homogeneous.
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3. Transfer the DCC solution to the thioctic acid/NHS solution, and purge with N2 gas. Stir overnight at room temperature. The resultant mixture will be an opaque, light yellow. 4. Vacuum filter the solution, and transfer to a separatory funnel. The filtered solid can be rinsed with DCM to collect as much of the yellow colored product as possible. 5. Add an equal volume of water, and extract into the organic layer. Wash with water a second time. 6. Wash two times with an equal volume of 5% acetic acid, followed by two extractions with an equal volume of water. 7. Dry the organic layer over anhydrous MgSO4, and filter into a 100 mL round bottom flask under reduced pressure. 8. Rinse the filter with 2 mL of DCM, and transfer to a 100 mL round bottom flask. 9. Remove the solvent by rotary evaporation under ambient temperature. 10. Collect the flakey, light yellow solid, and dry overnight under vacuum. The dry product should be packaged in small aliquots, and stored in a light sensitive vial at −20°C in a desiccator. 11. NHS-TE was characterized by NMR and MS: 1H-NMR (400 MHz, CDCl3) d 1.56 (m, 2 H), 1.71 (m, 2 H), 1.74 (m, 2 H), 1.93 (m, 1 H), 2.45 (m, 1 H), 2.63 (m, 2 H), 2.84 (s, 4 H), 3.12 (m, 1 H), 3.19 (m, 1 H), 3.58 (m, 1 H); expected mass: 303.06, observed: 304.42 (m/z + 1), 326.38 (major, m/z + Na+). 3.6. Modifying Electrode with NHS-TE
1. Polish a circular gold electrode (3 mm diameter, Fig. 1C) with 1 μm first and then 0.25 μm diamond compounds in order on different microcloth pads. Sonicate the electrode three times in nanopure water for 3 min (see Note 13). 2. Rinse the polished electrode with nanopure water, and dry with N2 gas. 3. Store the electrode in a 10 mL test tube enclosed by a 250 mL brown glass cylinder with a bottom layer of drierite. 4. While enclosed in a desiccator, incubate the polished electrode for at least 18 h in a NHS-TE solution (16.5 mM in dry DMF). 5. Remove the NHS-TE modified electrode from a desiccator, and rinse with 100% ethanol two times before drying with N2 gas.
3.7. Constructing the Virus Electrode
1. Suspend the NHS-TE modified electrode in a phage solution (300 μL in PBS, 16–40 nM), and mix for 1 h using an orbital shaker.
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Fig. 1. Potentiostat system and working/counter electrode assembly: (A) potentiostat Princeton Applied Research, PAR 2263 controller, and (B) computer system installed with PAR Power Suite software, (C) top view of a Au disk (diameter 3 mm) working electrode and side view, showing the Pt coil counter electrode glued to the body of the working electrode.
2. Rinse the virus-modified electrode 5 min with PBF/TW buffer and then 5 min with wash buffer. 3. Dip the virus electrode in 300 μL 0.2% BSA solution, and mix for 40 min. 4. Rinse 5 min with PBF/TW and then 5 min with wash buffer. 5. Transfer the virus/BSA modified electrode into 300 μL 0.05 N HCl, and shake for another 1 min to wash away nonspecific binding phage. 6. Rinse with PBF/TW and wash buffer. 7. Test the solution used to rinse the electrode with pH paper to ensure pH ∼7.2. 8. Dip the virus-modified electrode in wash buffer for further EIS measurements (see Note 14, construction of virus electrode shown in Fig. 2A, B). 3.8. Recognizing Target Biomolecules
1. Thaw analytes (e.g., frozen p-Ab, n-Ab or PSMA aliquots) in an ice-water bath. 2. Mix the target to an appropriate dilution with the wash buffer to provide a 500 μL solution for measurement by the virus electrode. 3. Dip the virus electrode into the analyte solution, and shake for 1 h at room temperature (Fig. 2C–E for analytes n-Ab, PSMA, and p-Ab, respectively).
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Fig. 2. Virus electrode construction: (A) M13 phage is covalently tethered to a selfassembled monolayer of NHS-TE, through formation of amide bonds between free amines on the phage and the activated carboxylate. (B) Gaps in the monolayer and unreacted NHS esters are capped with BSA. (C) A control experiment is first performed by reacting with n-Ab. (D) PSMA is added next to bind molecular recognition scaffolds on the surface of the phage (hooks). (E) Last, p-Ab can react with the virus electrode.
4. Rinse the electrode with PBF/TW and wash buffer at least three times. 5. Dip the electrode in 2 mL wash buffer 1 min for further EIS measurements in a 3 mL polypropylene container (see Note 15). 3.9. Oxidizing Virus Electrode with Cyclic Voltammetry to Test Surface Passivation
1. Rinse the home-made reference electrode, a saturated calomel electrode (SCE), with nanopure water, wipe dry with a kimwipe, and dip into wash buffer solution. 2. Flame anneal the home-made Pt flat counter electrode with a propane torch, and rinse with nanopure water. Assemble, and glue the Pt counter electrode to the virus electrode as depicted in Fig. 1C. 3. Dip the counter and working electrode in wash buffer solution immediately, and avoid drying the virus electrode. (Caution: if dry, make a new virus electrode.) 4. Immerse virus/counter electrode assembly in fresh PBF buffer (pH = 7.2). 5. Connect labeled alligator clips to the corresponding electrodes (for example: green: working electrode, white: reference electrode, red: counter electrode and gray: not connected). 6. Before opening software, turn on potentiostat power. 7. Click the shortcut to “Power Suite” (if steps 6 and 7 are reversed, the program will malfunction, and cannot perform the measurement). 8. Select “Power CV” from menu, before changing and saving the file name. 9. Select multiple-cycle waveform (repeat N times).
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10. Input set up parameters: initial potential E0 (0.0 V), vertex potential E1 (−0.8 V), E2 (1 V), scan rate (20 mV/s), number of cycles (20), filter (5 Hz) and max current magnitude (40 μA). 11. Depress “Run” button and record data (Fig. 3). 3.10. Measuring Electrochemical Impedance Spectrum
1. All electrochemical impedance spectra were acquired using two electrodes: The virus electrode and the platinum counter electrode. The four electrode leads from the potentiostat were, therefore, paired as follows: The “sense” electrode and working electrode connections were applied to the virus electrode; the reference electrode and counter electrode were applied to the counter electrode. 2. Click shortcut “Power Suite” and then click software “Power Sine”, change and save the file name. 3. Set up parameters: potential amplitude = 10 mV, data quality = 3, acquire 50 points with logarithmic point spacing for each run at over the scanning frequency range from 1 MHz to 0.1 Hz at rest potential. 4. Display four windows during data collection (i) the Nyquist Plot (Zim vs. Zre), (ii) phase angle vs. frequency, (iii) Zre vs. frequency, and (iv) Zim vs. frequency.
Fig. 3. Cyclic voltammetry of a virus electrode. Cyclic voltammograms at 20 mV/s were used to examine a clean gold electrode (solid line) in PBF and a phage-covered biosensor (dash line) following experiments to assess the response to n-Ab and p-Ab. Reactions characteristic of exposed gold – including the reduction of protons (at 0.0 V) and the oxidation and reduction of the gold surface (at 0.2 and −0.55 V, respectively) – were suppressed by the covalent virus layer. This layer could be removed, gradually, by scanning the potential of this electrode to +1.0 V vs. SCE revealing the characteristic voltammetric features of the underlying gold surface. Reprinted with permission from ref. 16.
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5. Insure the working electrode can pass through the Pt coil counter electrode easily. Do not make it fit too tight to surround the electrode. 6. Flame anneal home-made Pt coil counter electrode with a propane torch, rinse with nanopure water, and assemble and fix the Pt coil to the virus electrode body using glue as before. 7. Rinse the working/counter electrode assembly by dipping it in wash buffer for 1 min. 8. Acquire the following EIS data sets in order (i) virus/BSA, (ii) acid wash (optional), (iii) n-Ab, (iv) PSMA, and (v) p-Ab 9. To initiate each acquisition, depress the “Run” key. Acquire three sets of EIS data for each solution, i–v above (n = 3). 10. All these data were analyzed to get average, standard deviation and pool deviation dDZ é å n (X - X )2 + å m (Y - Y )2 ù / (m + n - l ), if data j j =1 ë i =1 i û came from data set X and Y, then l = 2, if three data sets then l = 3) to calculate the signal to noise ratio (S/N) and error bar (2dDZ) (Fig. 4). =
4. Notes 1. PSMA is stored at 4°C, and should not be frozen. 2. For mutagenesis to introduce the stop codons, the protocols in Subheadings “Oligonucleotide Phosphorylation with T4 Polynucleotide Kinase,” “Annealing of the Oligonucleotide to the Template,” and step 1 of “Enzymatic Synthesis of CCC-dsDNA” can be scaled down by a factor of ten. The reaction product can be used directly to transform E. coli using any standard procedure. 3. The protocol can be scaled up by inoculating a larger overnight culture and purifying the ssDNA with multiple spin columns. 4. Preheating the EB buffer to 50°C prior to use can help to recover more ssDNA during the elution step. 5. The DNA should appear as a predominant single band, but faint bands with lower electrophoretic mobility are often visible. These are likely caused by secondary structure in the ssDNA. 6. All steps can be scaled-up considerably, with the exception of the annealing step. The protocol described here works well with volumes of 250 μL or less. 7. The phosphorylated oligonucleotide should be used immediately in the subsequent steps.
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Fig. 4. Electrochemical Impedance Measurement. (A, B) Nyquist plots (Zre vs. Zim) for the frequency range from 0.1 Hz to 1.0 MHz. Shown are three data sets: PBF buffer (filled diamond) is the baseline against which the impedance in n-Ab (filled circle) and p-Ab (open circle) solutions are measured. The impedance “spectrum” of this covalent virus electrode was first measured in PBF buffer, then it was measured in PBF buffer containing n-Ab, it was rinsed, and finally it was measured in PBF buffer containing p-Ab. As shown in (B), we define the virus electrode signal, ΔZre, to be Zre measured against Zre in PBF buffer. (C) ΔZre for solutions of three molecules – p-Ab (open circle), PSMA (open square), and n-Ab (filled circle) – are plotted here as a function of frequency. Error bars were obtained by repeating each impedance measurement three times and calculating the standard deviation. ΔZre and the standard deviation of ΔZre are both large for frequencies below 1 kHz. Above this frequency, ΔZre is less than 20 Ω but the reproducibility of the measured values is much better, proportionately, than at lower frequencies. To highlight this fact, one can divide ΔZre by the standard deviation, sre, to obtain an estimate of the signal-to-noise ratio (S/N). These S/N ratios, showing the appreciable values of 10–20 in the frequency range from 2 to 500 kHz, are plotted in (D). Reprinted with permission from ref 16.
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8. The dU-ssDNA template can be heated to 90°C for 2 min, then cooled slowly to room temperature to remove any secondary structures that may interfere with the annealing of the mutagenic oligonucleotide. 9. To scale up this reaction, run multiple annealing reactions of 250 μL each. 10. Use the library immediately or add glycerol to a final concentration of 10% and store at −80°C. Some displayed proteins can be denatured by freezing, which could render them unusable for selections. In general, it is best to use libraries immediately. 11. Wells can be coated overnight at 4°C with shaking on an orbital shaker. 12. DCC can be dissolved gently with a heat gun if it does not initially go into solution. Alternatively, more DCM can be added drop-wise until the solid DCC fully dissolves after swirling. 13. Do not dip the electrode into solutions more than 1 cm above the Au disk. The junction between wire and gold electrode is quite fragile, and should not be exposed to water particularly during sonication. 14. The first measurement provides a baseline for comparison with analytes PSMA and antibodies. 15. To avoid cross-contamination, use a new container for each EIS measurement.
Acknowledgments R.M.P. acknowledges funding support from the National Science Foundation (grants CHE-0111557 and CHE-0641169) and the Petroleum Research Fund of the American Chemical Society (grant 40714-AC5). G.A.W. acknowledges funding support from the NSF (grant EF-0404057). J.E.D. and J.A.L. thank the American Chemical Society (Division of Organic Chemistry) and the NIH NIGMS (Minority Supplemental Award) respectively for graduate fellowships.
References 1. Smith, G. P. (1985) Filamentous fusion phage: novel expression vectors that display cloned antigens on the virion surface. Science. 228, 1315–1317 2. Kehoe, J. W. and Kay, B. K. (2005) Filamentous phage display in the new millennium. Chem. Rev. 105, 4056–4072
3. Diaz, J. E., Howard, B. E., Neubauer, M. S., Olszewski, A., and Weiss, G. A. (2003) Exploring biochemistry and cellular biology with protein libraries. Curr. Issues Mol. Biol. 5, 129–145 4. Sidhu, S. S., Lowman, H. B., Cunningham, B. C., and Wells, J. A. (2000) Phage display
Synthesis of a Virus Electrode for Measurement of Prostate
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
for selection of novel binding peptides. Methods Enzymol. 328, 333–363 Sidhu, S. S. (2000) Phage display in pharmaceutical biotechnology. Curr. Opin. Biotechnol. 11, 610–616 Scott, J. K. and Smith, G. P. (1990) Searching for peptide ligands with an epitope library. Science. 249, 386–390 Devlin, J. J., Panganiban, L. C., and Devlin, P. E. (1990) Random peptide libraries: a source of specific protein binding molecules. Science. 249, 404–406 Cwirla, S. E., Peters, E. A., Barret, R. W., and Dower, W. J. (1990) Peptides on phage: a vast library of peptides for identifying ligands. Proc. Natl Acad. Sci. U.S.A. 87, 6378–6382 Sidhu, S. S. and Weiss, G. A. (2004) Constructing phage display libraries by oligonucleotide-directed mutagenesis, in Phage Display: A Practical Approach (Lowman, H. B. and Clackson, T., eds.), Oxford University Press, Oxford, UK, pp. 27–41 Hengerer, A., Decker, J., Prohaska, E., Hauck, S., Kößlinger, C., and Wolf, H. (1999) Quartz crystal microbalance (QCM) as a device for the screening of phage libraries. Biosens. Bioelectron. 14, 139–144 Dultsev, F. N., Speight, R. E., Fiorini, M. T., Blackburn, J. M., Abell, C., Ostanin, V. P., and Klenerman, D. (2001) Direct and quantitative detection of bacteriophage by “hearing” surface detachment using a quartz crystal microbalance. Anal. Chem. 73, 3935–3939 Hengerer, A., Kösslinger, C., Decker, J., Hauck, S., Queitsch, I., Wolf, H., and Dübel, S. (1999) Determination of phage antibody affinities to antigen by a microbalance sensor system. BioTechniques. 26, 956–964 Åström, E. and Ohlson, S. (2006) Detection of weakly interacting anti-carbohydrate scFv phages using surface plasmon resonance. J. Mol. Recognit. 19, 282–286 Goldman, E. R., Pazirandeh, M. P., Mauro, J. M., King, K. D., Frey, J. C., Anderson, G. P. (2000) Phage-displayed peptides as biosensor reagents. J. Mol. Recognit. 13, 382–387 Kelly, K. A., Waterman, P., and Weissleder, R. (2006) In vivo imaging of molecularly targeted phage. Neoplasia. 8, 1011–1018 Yang, L. -M. C., Tam, P. Y., Murray, B. J., McIntire, T. M., Overstreet, C. M., Weiss, G. A., and Penner, R. M. (2006) Virus electrodes for universal biodetection. Anal. Chem. 78, 3265–3270 Sadik, O. A., Xu, H., Gheorghiu, E., Andreescu, D., Balut, C., Gheorghiu, M., and Bratu, D. (2002) Differential impedance spectroscopy for monitoring protein immo-
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
273
bilization and antibody-antigen reactions. Anal. Chem. 74, 3142–3150 Tang, H., Chen, J., Nie, L., Kuang, Y., Yao, S. (2007) A label-free electrochemical immunoassay for carcinoembryonic antigen (CEA) based on gold nanoparticles (AuNPs) and nonconductive polymer film. Biosens. Bioelectron. 22, 1061–1067 Barton, A. C., Collyer, S. D., Davis, F., Gornall, D. D., Law, K. A., Lawrence, E. C. D., Mills, D. W., Myler, S., Pritchard, J. A., Thompson, M., and Higson, S. P. (2004) Sonochemically fabricated microelectrode arrays for biosensors offering widespread applicability. Part I Biosens. Bioelectron. 20, 328–337 Gill, A., Farace, G., Lillie, G., and Vadgama, P. (2002) Strategic issues in reliable sensing. Bioelectrochemistry. 55, 123–125 He, P., Xu, Y., and Fang, Y. (2005) A review: Electrochemical DNA biosensors for sequence recognition. Anal. Lett. 38, 2597–2623 K’Owino, I. O., and Sadik, O. A. (2005) Impedance spectroscopy: A powerful tool for rapid biomolecular screening and cell culture monitoring. Electroanalysis. 17, 2101–2113 Cui, X. Q., Pei, R. J., Wang, X. Z., Yang, F., Ma, Y., Dong, S. J., and Yang, X. R. (2003) Layer-by-layer assembly of multilayer films composed of avidin and biotin-labeled antibody for immunosensing. Biosens. Bioelectron. 18, 59–67 Ruan, C., Yang, L., and Li, Y. (2002) Immunobiosensor chips for detection of Escherichia coli O157: H7 using electrochemical impedance spectroscopy. Anal. Chem. 74, 4814– 4820 Wang, M. Wang, L., Wang, G., Ji, X., Bai, Y., Li, T., Gong, S., and Li, J. (2004) Application of impedance spectroscopy for monitoring colloid Au-enhanced antibody immobilization and antibody-antigen reactions. Biosens. Bioelectron. 19, 575–582 Wu, Z., Tang, J., Cheng, Z., Yang, X., and Wang, E. (2000) Ion channel behavior of supported bilayer lipid membranes on a glassy carbon electrode. Anal. Chem. 72, 6030–6033 Ye, J. S., Ottova, A., Tien, H. T., and Sheu, F. S. (2003) Nanostructured platinum-lipid bilayer composite as biosensor. Bioelectrochemistry. 59, 65–72 Lie, L. H., Patole, S. N., Pike, A. R., Ryder, L. C., Connolly, B. A., Ward, A. D., Tuite, E. M., Houlton, A., and Horrocks, B. R. (2004) Immobilisation and synthesis of DNA on Si(111), nanocrystalline porous silicon and silicon nanoparticles. Faraday Discusss. 125, 235–249
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29. Su, L., Sen, D., and Yu, H. Z. (2006) Voltammetric study of the ion-exchange binding of non-electroactive metal cations to DNAmodified surfaces. Analyst 131, 317–322 30. Chen, H., Jiang, J. H., Huang, Y., Deng, T., Li, J. S., Shen, G. L., and Yu, R. Q. (2006) An electrochemical impedance immunosensor with signal amplification based on Au-colloid labeled antibody complex. Sens. Actuators B 117, 211–218 31. Wang, J. B., Profitt, J. A., Pugia, M. J., and Suni, II (2006) Au nanoparticle conjugation for impedance and capacitance signal amplification in biosensors. Anal. Chem. 78, 1769–1773 32. Alfonta, L., Singh, A. K., and Willner, I. (2001) Liposomes labeled with biotin and horseradish peroxidase: A probe for the enhanced amplification of antigen-antibody or oligonucleotide-DNA sensing processes by the precipitation of an insoluble product on electrodes. Anal. Chem. 73, 91–102 33. Patolsky, F., Zayats, M., Katz, E., and Willner, I. (1999) Precipitation of an insoluble product on enzyme monolayer electrodes for biosensor applications: Characterization by faradaic impedance spectroscopy, cyclic voltammetry, and microgravimetric quartz crystal microbalance analyses. Anal. Chem. 71, 3171–3180
34. Ionescu-Zanetti, C., Nevill, J. T., Di Carlo, D., Jeong, K. H., and Lee, L. P. (2006) Nanogap capacitors: Sensitivity to sample permittivity changes. J. Appl. Phys. 99, 024305 35. Van Gerwen, P., Laureyn, W., Laureys, W., Huyberechts, G., De Beeck, M. O., Baert, K., Suls, J., Sansen, W., Jacobs, P., Hermans, L., and Mertens, R. (1998) Nanoscaled interdigitated electrode arrays for biochemical sensors. Sens. Actuators B 49, 73–80 36. Pangborn, A. B., Giardello, M. A., Grubbs, R. H., Rosen, R. K., and Timmers, F. J. (1996) Safe and convenient procedure for solvent purification. Organometallics 15, 1518–1520 37. Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987) Rapid and efficient site-specific mutagenesis without phenotypic selection. Methods Enzymol. 154, 367–382 38. Davis, M. I., Bennett, M. J., Thomas, L. M., and Bjorkman, P. J. (2005) Crystal structure of prostate-specific membrane antigen, a tumor marker and peptidase. Proc. Natl. Acad. Sci. U.S.A. 102, 5981–5986 39. Blonder, R., Willner, I., and Bückmann, A. F. (1998) Reconstitution of apo-glucose oxidase on nitrospiropyran and FAD mixed monolayers on gold electrodes: Photostimulation of bioelectrocatalytic features of the biocatalyst. J. Am. Chem. Soc. 120, 9335–9341
Chapter 17 In Vivo Bacteriophage Display for the Discovery of Novel Peptide-Based Tumor-Targeting Agents Jessica R. Newton and Susan L. Deutscher Summary A powerful strategy for targeted drug discovery is the use of bacteriophage (phage) display technology for identification of peptide-based tumor targeting agents. Peptide pharmaceuticals may possess clinically desirable properties because of their rapid blood clearance, non-immunogenic nature, and ease of synthesis. Phage display has identified hundreds of different peptide sequences that bind a desired target in vitro. Regrettably, few of these peptides offer good targeting efficacy in vivo. One reason for this is the synthesized peptide may not retain its optimal activity outside the microenvironment of the phage. Another possible explanation is that traditionally, phage selections are performed in vitro outside the complicated milieu of a living animal. Given these shortcomings, we have developed methods to select phage peptide display libraries in living mice, to identify, a priori, phage (and corresponding synthesized peptides) with ideal tumor-targeting propensity. Key words: Affinity selection, Antigen discovery, E. coli K91 BluKan, fUSE5, f88–4, Peptide-based tumor-targeting.
1. Introduction Bacteriophage (phage) display technology has become a commonly used technique for the discovery of new disease targets and targeting agents (1). Phage display is economic, flexible, and easily executed. Each phage construct in a phage display library contains a unique modified coat protein. This genetically modified protein includes a foreign protein (peptide, single chain antibody, or other polypeptide), which is fused to a coat protein and displayed on the surface of the phage particle (or virion) (2, 3). Typically random phage peptide display libraries contain peptides Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_17
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that range from 6 to 45 amino acids long. These peptides can be displayed as linear or cysteine constrained sequences and are typically fused to either coat protein III or coat protein VIII. The phage display vector designed to display foreign peptides on coat protein III is termed fUSE5 vector. This vector system can display up to five copies of the modified coat protein III on one tip of the phage virion. The phage display vector designed to display foreign peptides on coat protein VIII is designated f88–4, and displays hundreds of modified coat protein VIII proteins as well unmodified coat protein VIIIs. A phage display library can contain up to 109 random peptides; however, each individual phage clone displays a single type of peptide sequence, making deconvolution of the encoded peptide straightforward. Affinity selection is the process of screening a phage display library; it is composed of several rounds of selection, elution, and amplification of the selected fittest subpopulations of phage library (4). Selection of the library is accomplished by passing the library over the desired target and allowing the displayed random peptides to bind (or be captured), while nonbinding clones are washed away. The captured phage (or fittest subpopulation) retain infectivity and can therefore be propagated and cloned by infecting fresh bacterial host cells. Thus, a library is easily resynthesized and rescreened until molecules with a desired binding activity are obtained. Currently, there is a myriad of reported phage display derived peptides that bind cancer-associated antigens and cancer cells (1). Most of these peptides have been generated from in vitro, ex vivo, and in situ phage library selections. To date, few of these peptides have been shown to be successful in the in vivo tumor targeting and molecular imaging of cancer. Phage selections have also been performed in live animals to obtain peptides with optimal stability and targeting properties in vivo (5, 6). These in vivo selected peptides show promise in the fulfillment of the ultimate goal of in vivo molecular imaging of cancer.
2. Materials 2.1. Preparing K91 BluKan Escherichia coli Stocks
1. Soft agar stab of K91 BluKan Escherichia coli (E. coli) (From George P. Smith, Department of Biological Sciences, University of Missouri, Columbia). 2. NZY media (1×): 1% (w/v) Casein Hydrolysate Enzymatic (N-Z-Amine A) (cat# 101290, MP Biomedicals Inc, Solon, OH), 0.5% (w/v) Yeast Extract (cat# BP1422, Fisher Scientific, Pittsburgh, PA), and 0.5% (w/v) NaCl (cat# S671, Fisher Scientific, Pittsburgh, PA) (pH 7.5 with NaOH). Autoclave and store at room temperature. Antibiotic is only added at the time of experiment.
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3. NZY agar plates: 1× NZY media plus 1.5% (w/v) agar (cat# DF0140010, Fisher Scientific, Pittsburgh, PA). Dissolve agar by autoclaving the solution. After autoclaving the NZY agar the hot solution should be gently mixed. When the NZY agar is cooled enough to hold, antibiotic is added, mixed and plates are poured. 4. Kanamycin stock solution: Dissolve kanamycin sulfate (cat# K-4000, Fisher Scientific, St. Louis, MO) to 100 mg/mL in deionized H2O, aliquot, and store at −20°C. 5. 100% glycerol (cat# G30, Fisher Scientific, Pittsburgh, PA) should be autoclaved and stored at room temperature. 6. Liquid nitrogen. 7. Autoclaved sterile toothpicks or inoculating needles (cat# 08–757–135, Fisher Scientific, Pittsburgh, PA). 8. Sterile Petri dishes (cat# 08–757–9B, Fisher Scientific, Pittsburgh, PA). 9. Sterile 14 mL snap cap tubes (cat# 14–959–11B, Fisher Scientific, Pittsburgh, PA). 10. Sterile cryo-vials (cat# 05–669–56, Fisher Scientific, Pittsburgh, PA). 2.2. Determining the Infectious Units of the Stock Phage Library Solution
1. 1× NZY Media: see Subheading 2.1. 2. NZY agar plates: see Subheading 2.1. 3. Kanamycin stock solution: see Subheading 2.1. 4. Tetracycline stock solution: Dissolve tetracycline (cat# 268054, Sigma, St. Louis, MO) to 40 mg/mL with deionized H2O and sterilize by passing through a 0.45 μm filter. Add an equal volume of sterile glycerol to the tetracycline for a final concentration of 20 mg/mL. Mix thoroughly and store at −20°C. Protect this solution from light by wrapping the bottle in aluminum foil. 5. TBS (1×): 50 mM Tris–HCl, pH 7.5 (cat# O4997, Fisher Scientific, Pittsburgh, PA), 150 mM NaCl. Filter sterilize and store at room temperature. 6. TBS/gelatin: 0.1 g gelatin (cat# G-1890, Sigma, St. Louis, MO) in 100 mL 1 × TBS. Autoclave, mix, and store at room temperature. 7. Glycerol stocks of E. coli (see Subheading 2.1). 8. Microtiter plate (cat# 14–245–73, Fisher Scientific, Pittsburgh, PA). 9. Square sterile petri plate with grid on bottom (cat# 08–757– 11A, Fisher Scientific, Pittsburgh, PA). 10. Screw cap tube (50 mL) (cat# 05–539–8, Fisher Scientific, Pittsburgh, PA).
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2.3. Preclearing the Phage Library
1. Dulbecco’s PBS (1×): 2.67 mM KCl, 1.47 mM KH2PO4, 137.93 mM NaCl, 8.06 mM Na2HPO4–7H2O (cat# 14190– 300, Invitrogen, Carlsbad, California). 2. PEG/NaCl: 100 g Polyethylene Glycol 8000 (cat# BP233, Fisher Scientific, Pittsburgh, PA), 116.9 g NaCl, and 475 mL H2O. Dissolve PEG by autoclaving the solution (with an autoclavable stir bar in the bottle). Cool the PEG/NaCl solution on a stir plate with constant stirring to prevent phase separation. Reautoclave if phase separation occurs. 3. CF-1 mouse (or mouse strain of choice) (Charles River, Wilmington, MA). 4. Random peptide phage display library such as fUSE5 or f88–4 (From George P. Smith, Department of Biological Sciences, University of Missouri, Columbia). 5. Pierce Slide-A-Lyzer (3 mL) (cat# PI-66380, Fisher Scientific, Pittsburgh, PA). 6. Eppendorf tubes (1.5 and 2 mL) (cat# 05–402–25 and 05–402–7, Fisher Scientific, Pittsburgh, PA). 7. Syringe (1 cc) (14–829–25, Fisher Scientific, Pittsburgh, PA). 8. 25-gauge needle (14–821–13D, Fisher Scientific, Pittsburgh, PA).
2.4. Amplification and Purification of a Phage Display Library
1. 1× NZY media: see Subheading 2.1. 2. NZY agar plates: see Subheading 2.1. 3. Kanamycin stock solution: see Subheading 2.1. 4. Tetracycline stock solution: see Subheading 2.2. 5. PEG/NaCl: see Subheading 2.3. 6. D-PBS: see Subheading 2.3. 7. CsCl (cat# AC18950, Fisher Scientific, Pittsburgh, PA). 8. Glycerol stocks of E. coli. 9. Precleared or fittest subpopulation phage library (see Subheadings 2.3 or 2.5). 10. Screw cap tube (50 mL) (cat# 05–539–8, Fisher Scientific, Pittsburgh, PA). 11. Flask (4 L) (cat# 07–250–092, Fisher Scientific, Pittsburgh, PA). 12. Centrifuge bottles (500 mL) (cat# 14–375–355, Fisher Scientific, Pittsburgh, PA). 13. Oak Ridge centrifuge tubes (30 mL) (cat# 05–529B, Fisher Scientific, Pittsburgh, PA). 14. Beckman Ultra-Clear ultracentrifuge tubes (cat# 344059, Beckman Coulter, Fullerton, CA). 15. Pierce Slide-A-Lyzer (3 mL) (cat# PI-66380, Fisher Scientific, Pittsburgh, PA).
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1. NZY agar plates: see Subheading 2.1. 2. D-PBS: see Subheading 2.3. 3. DMPB: 50 mL Dulbecco’s Modified Eagle Medium (DMEM) (1×), 1 Complete Mini Tablet (cat# 11 836 170 001, Roche Applied Science, Mannheim, Germany), and 0.25% (w/v) BSA (cat# B4287, Sigma, St. Louis, MO). 4. DMPB + 2.5% (w/v) CHAPS (cat# C5070, Sigma, St. Louis, MO). 5. Amplified and purified precleared random peptide phage display library (see Subheading 2.4). 6. SCID mice (Harlan, Indianapolis, IN). 7. Syringe (60 cc) (14–820–11, Fisher Scientific, Pittsburgh, PA). 8. Butterfly infusion needle 25 G (14–840–41, Fisher Scientific, Pittsburgh, PA). 9. Ice buckets (cat# 02–591–44, Fisher Scientific, Pittsburgh, PA). 10. Insulated bucket (for liquid nitrogen). 11. Cutting board. 12. Weigh paper (cat# NC9798735, Fisher Scientific, Pittsburgh, PA). 13. Tweezers. 14. Razor blades. 15. Dounce and two sizes of pestle (cat# 11–850–51, Fisher Scientific, Pittsburgh, PA). 16. Eppendorf tubes (1.5 mL) (cat# 05–402–25, Fisher Scientific, Pittsburgh, PA). 17. Sterile spreader (cat# 50212551, Fisher Scientific, Pittsburgh, PA). 18. Sterile cryo-vials (cat# 05–669–56, Fisher Scientific, Pittsburgh, PA).
3. Methods There are many commercial sources of phage display libraries; however, in this chapter we describe the use of phage libraries derived from the fd-tet vector (GenBank Accession AF317217) (7); More specifically, the two most commonly used libraries, fUSE5 phage display library in the Type 3 vector (GenBank Accession AF218364), and f88–4 phage display library in the Type 88 vector (GenBank Accession AF218363). These libraries and host E. coli cells are described on the G. P. Smith Laboratory Homepage
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(http://www.biosci.missouri.edu/SmithGP/). This laboratory has a “long-standing policy of distributing our ‘kit’ (vector and E. coli host strains) and amplified random peptide libraries free to anyone who asks, including to commercial enterprises.” 3.1. Preparing K91 BluKan E. coli Stocks
K91 BluKan E. coli is the standard strain utilized for propagating fd-tet-based phage display vectors (GenBank Accession AF317217). This strain of E. coli is genetically engineered to be resistant to the antibiotic kanamycin (http://www.biosci.missouri.edu/SmithGP/PhageDisplayWebsite/Strains.doc). 1. Insert an inoculating needle or autoclaved toothpick into a stab of E. coli. 2. Streak a NZY agar plate containing 100 μg/mL kanamycin with the toothpick or inoculating needle, which is now coated with E. coli (Fig. 1). 3. Incubate the streaked plate upside down (to avoid condensation of water on the agar) overnight at 37°C. 4. Using an autoclaved toothpick, pick a well separated single colony from the plate and place the toothpick in a sterile 14 mL snap cap tube containing 2 mL of fresh NZY with 50 μg/mL kanamycin (see Note 1). 5. Grow the culture at 37°C overnight, with vigorous shaking. 6. Using sterile cryo-vials, make 200 μL aliquots of culture containing a final concentration of 15% glycerol. 7. Cap the vial, mix, flash freeze in liquid nitrogen and store at −80°C.
Fig. 1. Streaking NZY agar plate with K91BlueKan E. coli: NZY agar plates with 100 μg/mL kanamycin are streaked with K91BlueKan E. coli cells for the isolation of individual colonies and the subsequent preparation of glycerol stocks of the K91BlueKan E. coli.
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3.2. Determining the Infectious Units and Physical Units of the Stock Phage Library Solution
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1. Place a frozen aliquot of K91 BluKan E. coli glycerol stock on ice. Allow it to defrost just enough to pipette 100 μL of stock into a sterile 50 mL tube containing 5–10 mL NZY broth. 2. Grow the culture at 37°C overnight with vigorous shaking. The 50 mL tube should be secured at a 45° angle (see Note 2). 3. Inoculate 10 mL NZY with 50 μg/mL kanamycin with 100 μL of the overnight culture. Incubate at 37°C for 3 h, with vigorous shaking. 4. Two hours into the incubation period prepare a microtiter plate for serial dilution of the stock phage library solution by adding 20 μL TBS/gelatin to each well marked for use (except for the first well which will contain stock phage library solution). Also add 20 μL TBS/gelatin to two wells for use as a negative control (see Note 3). 5. Start the dilution series with 22 μL of the phage library. Transfer 2 μL of the stock phage library from the first well into the next. Mix well (Fig. 2a). 6. Using a new pipette tip, transfer 2 μL of the first dilution to the next well. 7. Repeat this process 12–14 times. 8. Monitor the turbidity of the culture from step 3 by taking OD600 readings of 1/10 dilutions (NZY as diluent and
Fig. 2. Titering phage solution. (A) Preparation of a microtiter plate for the serial dilution of stock phage solution includes the addition of 20 μL TBS/gelatin to each well marked for use (except for the first well which will contain 22 μL stock phage library solution). The stock phage solution is serially diluted by tenfold by the transfer of 2 μL from one well to the next. (B) Quantification of the infectious units is accomplished by spotting duplicate 20 μL “dots” of each dilution on a square plate containing NZY agar with 100 μg/mL kanamycin and 40 μg/mL tetracycline.
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blank). Once the OD600 reading is 0.15–0.20 place the culture on a platform rocker for 10 min of gentle rocking. 9. Add 20 μL of K91 BluKan E. coli culture to each well of the microtiter plate containing a sample. For a negative control add 20 μL of K91 BluKan E. coli culture to two empty wells. 10. Incubate the microtiter plate at room temperature for 10 min. 11. At the end of 10 min add 200 μL NZY media containing 0.22 μg/mL tetracycline. 12. Incubate at 37°C for 40 min. 13. Spot duplicate 20 μL “dots” of each dilution on a square plate containing NZY agar with 100 μg/mL kanamycin and 40 μg/mL tetracycline (see Note 4). Using a plate with a grid pattern on the bottom helps (Fig. 2b). 14. Allow the plate of 20 μL spots to air dry. 15. Incubate plate upside down at 37°C overnight. 16. The next day count the number of colonies in each spot. 17. ( ( (X colonies/0.02 mLvolume spotted) × 0.24 mLtotal vol)/0.002 mLphage) × dilution factor = Infectious Units per ume mL (IU/mL). 18. Perform a spectral scan of a 1/50 dilution of the stock phage solution from 240 to 320 nm (Fig. 3). 19. ( (Abs269 – Abs320) × 6 × 1016)/number of nucleotide bases per virion = Virions/mL (see Notes 5 and 6).
Fig. 3. Spectral scan of phage solution. Quantification of the physical units (virions/ mL) of a phage solution is achieved by performing a spectral scan of a dilute phage solution. The protein to DNA weight ratio for filamentous phage is about 6:1. Thus the UV absorption spectrum can be utilized for the calculation of phage concentration in virions/mL (8).
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Preclearing of a phage library is an important step toward the discovery of peptides with higher affinity and specificity for the desired target. This negative selection should remove many phage clones displaying peptides with the potential to bind nontarget antigens and thus complicate the affinity selection and characterization of phage clones (5) (Fig. 4). 1. Dialyze the stock phage solution against three changes of 1 L of cold, filter sterilized PBS using a dialysis membrane with a molecular weight cutoff of 10,000 Da (see Notes 7 and 8). 2. Determine the infectious units of the dialyzed phage library as described in Subheading 3.2. 3. Tail vein inject a CF-1 normal nontumor bearing mouse (or other mouse of choice) with 1013 IU (in PBS) phage library. 4. Allow phage library to circulate within the CF-1 mouse for 15 min. This period is termed the “preclearing” of the phage library (see Note 9).
Fig. 4. In vivo phage display affinity selection. A phage display library is first depleted of unwanted phage clones by preclearing the library (aka. negative selection) in a mouse. The resulting fittest subpopulation is then amplified and utilized for the subsequent three or four rounds of in vivo phage display selection.
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5. Euthanize the mouse by cervical dislocation. 6. Collect as much blood as possible through cardiac puncture using a 1-in., 25-gauge needle (see Note 10). Transfer blood from syringe to 2 mL Eppendorf tube(s) containing 1 mL PBS (see Note 11). 7. Pellet the red blood cells by low speed centrifugation 1,000– 2,000 rpm (500 × g–1,000 × g). 8. Transfer the supernatant to a new Eppendorf tube; add 15% of the total supernatant volume (v/v) of PEG/NaCl solution to the Eppendorf tube. For example, 0.15 mL of PEG/ NaCl should be added to a tube containing 1 mL supernatant. Mix well and precipitate phage overnight at 4°C. 9. Spin at 13,000 rpm (~25,000 × g) for 15 min to pellet phage and resuspend in 500 μL PBS. 10. Amplify, purify, and dialyze the resulting phage as described in Subheading 22.3.4 (see Note 12). 3.4. Amplification and Purification of a Phage Display Library
After each round of selection (negative selection or positive selection), the phage library must be amplified and purified for use in the next round of selection. It is the amplification of the resulting phage clones that generates an environment of competition for binding of phage clones to the desired target. 1. Place a frozen aliquot of K91 BluKan E. coli glycerol stock on ice. Allow it to defrost just enough to pipette 100 μL of stock into a sterile 50 mL screw cap tube containing 5–10 mL of NZY broth with 50 μg/mL kanamycin (screw the cap on tightly). 2. Grow the culture at 37°C overnight, with vigorous shaking. The 50 mL tube should be secured at a 45° angle. 3. The next morning add 100 μL of the overnight culture of K91 BluKan E. coli to 10 mL NZY broth with 50 μg/mL kanamycin in a 50 mL screw cap tube. 4. Inoculate 10 mL NZY with 50 μg/mL kanamycin with 100 μL of the overnight culture. Incubate at 37°C for 3 h, with vigorous shaking. 5. Monitor the turbidity of the culture by taking OD600 readings of 1/10 dilutions (NZY as diluent and blank). Once the OD600 reading is 0.15–0.20, place the culture on a platform rocker for 10 min of gentle rocking. 6. Transfer 2 mL of culture into a sterile 50 mL screw cap tube and add tetracycline for a final volume of 0.22 μg/mL tetracycline. Inoculate culture with the entire precleared library (from step 9 of Subheading 3.3) or all of the CHAPS eluant (from step 21 of Subheading 3.5); shake vigorously for 35 min at 37°C.
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7. Transfer the 2 mL phage infected E. coli culture to a 4 L flask containing 1 L of NZY broth with 50 μg/mL kanamycin and 20 μg/mL tetracycline and shake overnight at 37°C (see Notes 3, 13 and 14). 8. Divide the culture evenly into three 500 mL centrifuge bottles. Remove the K91 BluKan E. coli from the culture by two sequential centrifugations in a GSA rotor (or similar) at 5,000 and 8,000 rpm (4,000 × g and 10,000 × g) for 10 min each. 9. Pour the cleared supernatants into new tared centrifuge bottles and record weight in each (~1 g = 1 mL). 10. Add 15% of the total culture volume (v/v) of PEG/NaCl solution to each centrifuge bottle. Mix very thoroughly by multiple inversions and incubate solutions at 4°C overnight to allow the phage to precipitate. 11. Centrifuge the precipitated samples at 8,000 rpm (10,000 × g) in a GSA rotor for 20 min. 12. Carefully pour off the majority of the supernatant. Use a pipette to carefully remove the remaining supernatant. 13. Resuspend the pelleted phage in 10 mL of PBS and pool the phage into two 30 mL Oak Ridge centrifuge tubes. 14. Centrifuge phage at 12,000 rpm (17,000 × g) for 10 min in a SS34 rotor to pellet insoluble unwanted matter. Pour off and pipette the remaining supernatant to fresh Oak Ridge centrifuge tubes. 15. Add 15% of the total culture volume (v/v) of PEG/NaCl solution to each centrifuge bottle. Mix very thoroughly by multiple inversions and allow phage to precipitate overnight at 4°C. 16. Centrifuge at 12,000 rpm (17,000 × g) for 15 min. 17. Remove the supernatant and dissolve the phage in 10 mL PBS. 18. Centrifuge at 10,000 rpm (12,000 × g) for 10 min to remove insoluble matter. 19. Pour the supernatant into a 15 mL conical centrifuge tube (phage can be stored in refrigerator at this point). 20. Using the phage solution and PBS as diluent make a CsCl solution with a total volume of 24 mL and a density of 1.3 g/mL. 21. Dispense phage suspension into two Beckman clear ultracentrifuge tubes and centrifuge in a SW41 swinging bucket rotor for 36–48 h at 37,000 rpm (234,500 × g at rmax, 15.31 cm). 22. Collect the pure phage band, which would be toward the middle of each tube (see Note 15). 23. Dialyze the purified phage solution against three changes of ~1 L of cold, filtered sterilized PBS using a dialysis membrane with a molecular weight cutoff of 10,000 Da.
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24. Purified and dialyzed phage solution will be stable for years at 4°C (see Note 16). 3.5. In Vivo Selection of Tumor Binding Phage
1. Tail vein inject a tumor bearing SCID (or nude) mouse with 1012 IU of precleared phage library, which has been amplified and purified (Fig. 4). 2. Allow precleared phage library to circulate within the SCID mouse for one or more hours. 3. Euthanize the tumor bearing SCID mouse by cervical dislocation. 4. Perform a thoracotomy to expose the thoracic organs and to gain access to the heart. Cut a small hole in the right ventricle of the heart. 5. Insert a 25-gauge butterfly infusion needle into the left ventricle of the heart and perfuse the animal with 60 mL of PBS (see Notes 17 and 18). 6. Organs, tumor, and/or tissues of interest are then harvested, weighed, placed in labeled cryo-vials, and quick frozen in liquid nitrogen (see Note 19). 7. Tissues can now be stored at −80°C for up to 2–3 days without significant lose of phage titer. 8. Prepare the work area for tissue processing. Layout a clean cutting board with tweezers, razor blades, weigh sheets, and pestles. Next to the cutting board place a large ice bucket with DMPB, DMPB + 2.5% CHAPS, dounces, and preweighed 1.5 mL Eppendorf tubes for each tissue. Finally, place an insulated bucket filled with liquid nitrogen nearby. 9. Place the frozen cryo-vials of tissue on ice. 10. Use tweezers to remove frozen tissue from a cryo-vial and place on a clean weigh sheet. Cut off a section of tissue with a clean razor blade. Put rest of the tissue back into the cryovial and drop the cryo-vial into the liquid nitrogen. 11. Mince the section of tissue with a razor blade and transfer the freshly minced tissue to a dounce already containing 1 mL of DMPB. 12. Using first the smaller dounce, then the larger; dounce 10–20 times (see Note 20). 13. Transfer tissue homogenate to a microfuge tube (see Note 21). 14. Pellet the tissue by spinning 6,000 rpm (6,500 × g) for 3 min. 15. Pipette off the supernatant. 16. Resuspend pelleted tissue with 500 μL DMPB. 17. Repeat steps 13–15 three times. 18. Repeat steps 13 and 14. 19. Record the weight of the washed and pelleted tissue in the tube.
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20. Resuspend the pelleted tissue with 500 μL DMPB + 2.5% CHAPS. 21. Place tubes of tissues in DMPB + 2.5% CHAPS on a rotator and rotate for 1 h at 4°C (see Note 22). 22. Pellet the tissue by spinning at 6,000 rpm (6,500 × g) for 3 min. Titer supernatant (as described in Subheading 3.2) to define the amount of phage eluted from the tissue (i.e., IU of phage per gram of tissue). 23. Pick 10–20 well separated colonies from the titer results and sequence the random peptide insert (see Notes 23 and 24). 24. Amplify, purify, and dialyze the phage from the lysed tissue homogenates in step 21 (as described in Subheading 3.4). 25. Use phage library from step 23 for the next round of selection. Repeat steps 1–23 three or four times (Fig. 4). 26. Once finished with the final round of selection use the CHAPS eluted phage to infect E. coli. Spread 200 μL of the infected E. coli on NZY agar plates. Pick all the well separated colonies for sequencing and further characterization.
4. Notes 1. The lids on snap cap tubes have two stops. When growing a culture in snap cap tubes only press the lid past the first stop, so that the air is still able to freely flow into and out of the tube. 2. This overnight culture is placed at a 45° angle to insure proper aeration, thus the lid must be securely closed. 3. It is advised that a portion of the original solution of phage display library is saved back before any manipulations are preformed. 4. The fd-tet phage genome contains a tetracycline resistance gene. Thus, always amplify E. coli cells infected with phage in the presence of both kanamycin and tetracycline antibiotics (7). 5. IU/mL is usually only 5% of the V/mL concentration. 6. The phage display library is now ready for use in an affinity selection protocol. Affinity selection has also been successfully utilized in vitro and ex vivo. The affinity selection principle is the same for all types of selection. First, allow the phage library to bind to the presented target. Wash away unbound phage, elute and amplify bound phage. Repeat the selection process three to four times. For example in vitro targets may be purified, biotinylated, and immobilized on a streptavidincoated plate for affinity selection (9).
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7. We recommend the use of a Pierce Slide-A-Lyzer to minimize loss of sample. 8. From this point on, the solution of phage library must be kept sterile. Use sterile technique when removing aliquots or portions of the library (i.e., Open vial only in a sterile hood and remove aliquots using only sterile equipment). 9. This is sufficient time to allow for binding of the phage displaying random peptides to the normal vasculature and other nontumor antigens, but not enough time for the phage to extravasate into the tissues (10). 10. Too small of a needle will result in shearing of the cells. 11. Some clotting of the blood can be prevented by coating the syringe with PBS + 2 mM EDTA prior to performing the cardiac puncture. 12. In vitro and ex vivo selections also require negative selections. Instead of preclearing a phage display library in a normal mouse, the library would be precleared of phage clones that bound to the plastic of the plate and/or tube designated to hold the samples (or target) (11). Other possible negative selections could include irrelevant cell lines (for cell type specific peptides), normal cells (for abnormal or cancerous cell specific peptides), etc. 13. The f88 phage display vector requires the addition of 1 mM IPTG to the growth media as an inducer of the recombinant gene VIII. 14. Alternatively, the phage-infected E. coli can be grown and amplified in a 3 L fernbach flask (cat# 10–092, Fisher Scientific, Pittsburgh, PA). This type of flask has a higher surface area to volume ratio, which helps to aerate the culture during the period of outgrowth. 15. This band is translucent and can be difficult to see. There will usually be a sharp, flocculent band above the phage band, and a much larger flocculent band just below the phage band. These flocculent bands are relatively opaque, and are therefore much more apparent than the translucent phage band. If you have difficulty visualizing the phage band, take ~1 cm just above the lower, larger flocculent band. The phage is removed using a sterile pipette, trying to keep the pipette tip at the meniscus. 16. It is recommended that one or two aliquots of the library are kept back for long term storage. These aliquots can act as a safety net against possible contamination of the “working solution” of phage library with wild type phage. 17. The wings of the butterfly needle are used to hold the needle in the left ventricle, while 60 mL of PBS are being gently forced into the vasculature. Blood (followed by nearly clear PBS) should drain from the hole in the right ventricle.
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19. 20.
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A successful perfusion will result with the body looking bloated and pale (especially the lungs). This perfusion reduces phage from the vasculature space, thus facilitating the selection of phage displaying random peptides, which directly bind to the tissue/tumor of interest. Nontarget organs can be harvested and phage titer determined for the purpose of monitoring the selection. The number of dounce homogenations required depends on the amount of connective tissue within the organ. Dounce until tissue gives little resistance. While collecting multiple tissues and organs, clean and sterilize dounces, pestals, and tweezers between samples. Use a new razor blade for each tissue. Sterilize pestals and tweezers by dunking in a beaker of 10% bleach water for 1 or 2 min, followed by rinsing in a beaker of autoclaved deionized water. To sterilize the dounces use a cuvette washer. Again, wash first with bleach water followed by a rinse with deionized water. The 2.5% CHAPS is used to lyse cells, thus allowing for amplification of internalized phage clones. The primers for the fUSE5 vector (5´-TGAATTTTCTGT ATGAGG-3´) and the f88–4 vector (5´-AGTAGCAGAA GCCTGAAGA-3´) are designed such that the resulting DNA sequence data is “backwards.” Fd-tet phage contain single-stranded DNA. The packaged single-stranded DNA is called the plus strand, which is anticomplementary to all the viral mRNAs. Thus the primers are designed to recognize and bind to the anticomplementary sequence of the gene of interest. Therefore, when analyzing the sequence data, the sequence must be reversed and complemented (12). This is done to monitor the selection process. You should see a large variety of sequences in the early rounds. At no point should you see only one sequence. It is possible to select for phage with growth advantages instead of the desired binding properties. Another concern is that of wild-type contaminants. Presence of wild-type contaminants can be assayed by titering for plaques. Wild-type phage make large plaques, while fd-tet-derived phage make very tiny (often invisible) plaques as a result of their replication defect.
Acknowledgments The authors would like to thank Professor George P. Smith for his generous gifts of phage display libraries and Marie Dickerson
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for her technical assistance. This work was supported by a Merit Review Award from the Veterans Administration, the Department of Defense DAMD17-03-1-0130, and NIH P50 CA103130-01. For more detailed information please refer to Neoplasia 8(2006) 772–780. References 1. Newton, J.R. and S.L. Deutscher, Phage peptide display. In: Handbook of Experimental Pharmacology: Molecular imaging II, ed. W. Semmler and M. Schwaiger. 2008, Berlin: Springer, Vol. 185/2: p 145–163 2. Smith, G.P. and J.K. Scott, Libraries of peptides and proteins displayed on filamentous phage. Methods in Enzymology, 1993. 217: p. 228–257 3. Smith, G.P., Filamentous fusion phage: novel expression vectors that display cloned antigens on the virion surface. Science, 1985. 228: p. 1315–1317 4. Scott, J.K. and G.P. Smith, Searching for peptide ligands with an epitope library. Science, 1990. 249: p. 386–390 5. Newton, J.R., et al., In vivo selection of phage for the optical imaging of PC-3 human prostate carcinoma in mice. Neoplasia, 2006. 8(9): p. 772–780 6. Arap, W. and R. Pasqualini, The human vascular mapping project. Selection and uti-
7.
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9. 10.
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lization of molecules for tumor endothelial targeting. Haemostasis, 2001. 31: p. 30–31 Zacher, A.N., III, et al., A new filamentous phage cloning vector: fd-tet. Gene, 1980. 9(1–2): p. 127–140 Day, L.A. and R.L. Wiseman, A comparison of DNA packaging in the virions of fd, Xf, and Pf1. In: The Single-Stranded DNA Phages, eds. D.T. Denhardt, D. Dressler, and D.S. Ray. 1978, Cold Spring Harbor, NY: Cold Spring Harbor Laboratory. p. 605–625 Smith, G.P. and V.A. Petrenko, Phage display. Chemical Reviews, 1997. 97: p. 391–410 Zou, J., et al., Biodistribution of filamentous phage peptide libraries in mice. Molecular Biology Reports, 2004. 37(2): p. 121–129 Phage Display a Laboratory Manual, ed. I. Barbas, et al. 2001, Cold Spring Habor, NY: Cold Spring Harbor Laboratory Press Smith, G.P. Lab Homepage, http://www.biosci.missouri.edu/smithGP/
Chapter 18 Biopanning of Phage Displayed Peptide Libraries for the Isolation of Cell-Specific Ligands Michael J. McGuire, Shunzi Li, and Kathlynn C. Brown Summary One limitation in the development of biosensors for the early detection of disease is the availability of high specificity and affinity ligands for biomarkers that are indicative of a pathogenic process. Within the past 10 years, biopanning of phage displayed peptide libraries on intact cells has proven to be a successful route to the identification of cell-specific ligands. The peptides selected from these combinatorial libraries are often able to distinguish between diseased cells and their normal counterparts as well as cells in different activation states. These ligands are small and chemical methodologies are available for regiospecific derivatization. As such, they can be incorporated into a variety of different diagnostic and therapeutic platforms. Here we describe the methods utilized in the selection of peptides from phage displayed libraries by biopanning. In addition, we provide methods for the synthesis of the selected peptides as both monomers and tetramers. Downstream uses for the peptides are illustrated. Key words: Phage display, Peptides, Cell-targeting, Biopanning, Combinatorial library, Diagnostics, Therapeutics, Quantum dots.
1. Introduction The development of biosensors for the detection of different disease states is dependent on the availability of high affinity and specificity ligands for the desired cell type and/or biomarker. In many applications, the accessibility of such ligands has been the limiting factor in the development of the technology. To date, antibodies have been the most common class of ligands utilized. However, antibodies are expensive and can be difficult to modify. Additionally, if the down-stream application is to detect particular cell types (i.e., a cancerous cell vs. its normal counterpart), the antibody must bind to its target in the context of an intact cell. Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_18
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As such, our lab and others have turned toward peptide libraries as a source of cell-specific ligands (1–16). In the same fashion that phage displayed peptide libraries can be panned on purified biomolecules, whole cells can be used as the bait for the peptide library. This approach, often referred to as biopanning, results in the isolation of peptides that display high cell-specificity; ligands can be isolated that discriminate between cell types and disease states. Furthermore, cell-specific peptides can be obtained without the knowledge of a suitable cell surface biomarker. The protocol is amenable to a variety of different cell types, including primary cells. To date, we have identified cell-specific peptides for many different cell types including cells of the immune system (2, 4), pancreatic β-cells (7), cardiac cells (3), tumor cells (5, 6), and pathogen-infected cells (8). Importantly, most peptides selected in this manner are active outside of context of the phage, retaining their cell-specificity and affinity. Furthermore, we have shown that tetramerizing the peptides on a branched scaffold can greatly enhance the peptides affinity for its target cell type (5, 6, 8, 17, 18). These peptides can be employed for the delivery of fluorescent nanoparticles, as cell capture reagents for cell enrichment, and as antibody replacements for flow cytometry. As peptides are amenable to derivatization, we anticipate that these cell-specific ligands will find utility in a variety of different biosensor platforms.
2. Materials 2.1. Cell Culture and Phage Panning (Methods Outlined in Subheadings 3.1 and 3.2)
1. Tissue culture cell line or primary cell of interest. 2. Tissue culture plates (12-well) for adherent cells. 3. Polypropylene centrifuge tubes (15 mL and 50 mL) for nonadherent cells. 4. Microcentrifuge tubes (1.5 mL). 5. Cell scrapers. 6. Phage library (see Note 1) or amplification stock for each round of panning. 7. RPMI media (or any cell-specific media) without serum. 8. Chloroquine stock (100×): Dissolve 55 mg chloroquine in 10 mL PBS for a final concentration of 10 mM. Filter sterilize the solution. 9. Protease inhibitor (25×) without EDTA (Roche). 10. Phosphate buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4.
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11. PBS+: Add 0.5 mM CaCl2 and 10 mM MgCl2 to PBS in this order with stirring. 12. PBS+ with 0.1% BSA: Add 0.1 g bovine serum albumin per 100 mL PBS+. 13. 0.1 M HCl–glycine, 0.9% NaCl pH adjusted to 2.2 with glycine. 14. 1.5 M Tris–HCl, pH 8.8. 15. 30 mM Tris–HCl, pH 8.0. 2.2. Bacterial Culture and Phage Amplification and Titering (Methods Outlined in Subheadings 3.3 and 3.4)
1. Selective media for K91 bacterial stocks. We use M9-Pro minimal medium prepared as follows: Mix 7.5 g agar + 430 mL water and autoclave solution. Cool agar solution to ∼55°C. Add 25 mL 20× M9 Salts, 5 mL 20% glucose, 50 μL 1 M CaCl2, 500 μL 1 M MgSO4, 100 μL 0.1% Thiamine, 2.5 mL 0.2 mg/mL biotin, 2.5 mL 1% uridine, 8 mL 1% leucine, 8 mL 1% phenylalanine, 8 mL 1% threonine, 8 mL 1% methionine, 8 mL 1% histidine, 8 mL 1% Tryptophan, and 8 mL 1% lysine. 20× M9 salts consist of 60 g Na2HPO4, 30 g KH2PO4, 5 g NaCl, and 10 g NH4Cl. 2. LB media. 3. 100 mm and 150 mm LB-tet plates (12 μg/mL tetracycline). 4. Culture flasks for expansion and isolation of phage clones. 5. 20% PEG-8000 (Fisher Chemical) in 0.9% NaCl (see Note 2). 6. 65°C heating block. 7. Bacterial incubator with shaker. 8. Various centrifuge tubes and bottles. 9. Low (3,000 × g) and high speed (11,000 × g) centrifuges for concentrating bacterial stocks and phage isolation. 10. Spectrophotometer to monitor bacterial cultures. 11. PBS prepared as described in Subheading 2.1.
18.2.3. Quantitative Real-Time PCR for Titering (Method Outlined in Subheading 18.3.5)
1. BioRad iCycler or similar apparatus. 2. 2× Sybr® green mastermix (see Note 3). 3. Optical PCR plates for real-time PCR. 4. Optical sealing tape for real-time PCR. 5. 8-channel pipette (5–50 μL). 6. Serial dilutions of previously characterized phage preparation to generate a standard curve. 7. Specific primers to tetracycline resistance gene (see Note 4) (a) Forward primer (tetR-F1): 5′-CGAATAAGAAGGCTGG CTCTGC-3′. (b) Reverse primer (tetR-R1): 5′-GCTGTGGGGCATTTTAC TTTAGG-3′.
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2.4. Colony PCR for Sequence Determination (Outlined in Subheading 3.6)
1. General materials for PCR mastermix preparation: 10× polymerase buffer, 25 mM MgCl2, 10 mM dNTP mix, Taq polymerase (GoTaq® DNA Polymerase 5 units/μL, Promega Corp, or Choice™ Taq, Denville Scientific). 2. Thermocycler. 3. Specific primers that flank library site (a) Forward primer (fd-tet F1): 5′-GGGCGATGGTTGTTGT CATTG-3′. (b) Reverse primer (fd-tet B1): 5′-CTCATTTTCAGGGATAG CAAGCC-3′. 4. Agarose gel apparatus. 5. 100 bp ladder standards (Promega Corp, catalog # G2101 or similar). 6. Exonuclease I (10 units/μL, New England Biolabs, or other suitable vendor). 7. Shrimp alkaline phosphatase (1 unit/μL, New England Biolabs or other suitable vendor). 8. BigDye® Terminator v3.1 (Applied Biosystems Inc). 9. Ethanol (70%). 10. Hi-Di™ Formamide (Applied Biosystems Inc). 11. Sequencing stop/precipitation reagent: Prepare by mixing 125 mL 95% ethanol, 29 mL water, and 6 mL 3 M sodium acetate, pH 5.2.
2.5. Selectivity and Specificity Determinations (Outlined in Subheading 3.7)
1. Materials outlined in Subheading 2.1 for the panning and Subheading 2.2 or Subheading 2.3 for titering. 2. Isolated phage clones and a control phage clone displaying an irrelevant peptide sequence. Alternatively, a phage clone that displays no peptide (referred to as an “empty” phage) can be employed. 3. Cell lines or primary cells of interest.
2.6. Peptide Synthesis (Outlined in Subheadings 3.8–3.11)
1. Symphony Synthesizer (Rainin Instruments, Protein Technologies, Inc. Woburn, MA) or other standard solid phase peptide synthesizer. 2. Resins for solid phase synthesis: Rink Amide AM resin (substitution level 0.71 mmol/g, Novabiochem, EMD Biosciences, San Diego, CA); Fmoc4-Lys2-Lys-β-Ala-CLEAR™ Acid Resin, Fmoc4-Lys2-Lys-Lys(Biotin-PEG)-β-Ala-CLEAR™ Acid Resin and Fmoc4-Lys2-Lys-Cys(Acm)-β-Ala-CLEAR™ Acid Resin (substitution level 0.21 mmol/g, Peptides International, Louisville, KY). 3. Fmoc amino acids required to synthesize desired peptide. Prepare 200 mM amino acid solutions by dissolving 20 mmol Fmocprotected amino acids in DMF to final volume of 100 mL.
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4. Coupling reagents: 2-(1H-Benzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU), 1-Hydroxybenzotrizole (HOBt) all available from Novabiochem. Prepare 200 mM solutions as follows: Weigh out 18.965 g HBTU,6.755 g HOBt, and 11 mL NMM, Add DMF to final volume of 250 mL. 5. If desired nonnatural amino acids can be incorporated into the peptide. We routinely incorporate Fmoc-NH–(PEG)11–COOH (C42H65NO16) (Polypure, Oslo, Norway), Fmoc-Glu(biotinylPEG)–OH (C40H55N5O10S), and Fmoc-Lys(biotin)–OH (C31H38N4O6S) (Novabiochem, EMD Biosciences, San Diego, CA) (see Notes 5 and 6). 6. Piperidine in DMF (20%): 200 mL piperidine, 800 mL DMF. 7. Cleavage cocktails (see Note 7): (a) TFA: H2O:TIS (95%:2.5%:2.5%) prepared by mixing 9.5 mL trifluoroacetic acid (TFA), 0.25 mL H2O, 0.25 mL triisopropylsilane (TIS). This cleavage cocktail is used for the cleavage of linear synthesized tetrameric peptide and maleimido activated cores. (b) TFA: EDT:H2O:TIS (94%:2.5%:2.5%:1%) prepared by mixing 9.4 mL TFA, 0.25 mL ethanedithiol (EDT), 0.25 mL H2O, 0.1 mL TIS. This cocktail is employed for peptides containing a cysteine residue. 8. Diethyl ether. 9. Dichloromethane (DCM). 10. Dimethylformamide (see Note 8). 11. 3-Maleimidopropionic acid (Sigma-Aldrich Inc, St. Louis, MO). 2.7. Removal of Group from Selectively Protected Cysteine Residues (Outlined in Subheading 3.12)
1. PBS containing 0.01 M EDTA. 2. Argon for flushing solutions. 3. TFA/Anisole mixture (99:1). 4. Silver acetate (Sigma-Aldrich or other vendor). 5. Diethyl ether. 6. Dithiothreitol (0.2 M) prepared in 1 M acetic acid. 7. Guanidine hydrochloride (8 M).
2.8. Peptide Purification and Characterization (Outlined in Subheading 3.13)
1. HPLC solvent delivery system with binary gradient capability and a UV detector. 2. Reversed-phase octadecylsilica (C18) column. In our laboratory, we use the following columns: Preparative column: Vydac RP-C18 column (250 mm length×22 mm diameter, 10 μm particle size). Analytical column: Varian RP-C18 column (250 mm length×4.6 mm diameter, 5 μm particle size).
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3. Solvent filtration apparatus equipped with a 0.45 μm Teflon filter (Such as Ultra-ware filter apparatus 300/1,000 mL from Kontes glass company and 0.45 μm Teflon filters from Millipore Corp.). 4. Syringe driven filter units, 0.22 μm porosity, 13 mm (Millipore Corp.). 5. Eluent A: H2O/0.1% TFA and eluent B: acetonitrile/0.1%TFA. 6. Access to mass spectrometer and/or peptide sequencing facility. 2.9. Inhibition of Selected Phage Clone by Cognate Peptide (Outlined in Subheading 3.14)
1. Selected phage clone and corresponding synthetic peptide.
2.10. Applications of Synthetic Peptides (Outlined in Subheadings 3.15– 3.18)
1. Synthetic peptide prepared with incorporation of biotin.
2. Reagents indicated in Subheading 2.5.
2. Streptavidin-conjugated Qdots (emission wavelengths chosen to match microscope emission filter set). 3. Microscope slides and coverslips. 4. Chamber slides (8-well) (for example, VWR, Inc., catalog # 62407–296) can be used to culture adherent cells prior to staining. 5. Prolong® Gold antifade reagent with DAPI (Invitrogen). 6. Fluorescence microscope. 7. Streptavidin-coated magnetic beads (Dynabeads M280-SA, Invitrogen, 6.7 × 108 beads/mL, 1 mg beads binds 700 pmoles free biotin). 8. Cell isolation magnet (or strong magnet). 9. Streptavidin-phycoerythrin or streptavidin-FITC. 10. Flow cytometer (Cell Lab Quanta, Beckman Coulter or other suitable instrument). 11. PBS+. 12. 0.4% Formalin: 37% formaldehyde, Sigma Chemical, diluted 1:10 in PBS, immediately prior to use. 13. Ethanol (70%). 14. Fingernail polish (any color).
3. Methods The following information is based on our protocols for selection of peptide ligands for cell recognition and delivery. Selection
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of a peptide ligand, using our protocol, should be expected to take 5–6 rounds of biopanning. During each round, cells are incubated with a mixture of phage displaying different peptides. Phage that do not bind or bind only to the surface of the cell are washed away. Phage that bind to the cell and are internalized by the cell are retained. These cell-internalized phage are amplified in bacteria, isolated, and used as the input in the next round of biopanning. In each round of selection, the diversity of the phage sample is reduced while the proportion of phage displaying a peptide that mediates cell-specific binding is increased. Once a phage displayed peptide has been selected using the biopanning protocol, we characterize the binding selectivity and cell specificity of that phage clone. Our determination of selectivity compares the binding and uptake of a cell-selected phage clone with binding and uptake of a control phage clone that was randomly selected from the library. This provides a rough estimate of the affinity of the peptide. Additionally, it assures that the cellular binding is due to the selected peptide and is not the result of nonspecific phage binding. The measurement of cell specificity involves comparison of the selectivity index of a specific phage over a variety of different cell types. During the characterization process, we also prepare chemically synthesized versions of the specific peptide, monomeric and tetrameric, and test the utility of these constructs as cell-binding reagents out of the context of phage presentation. Depending on the down-stream applications of the ligand, we will incorporate a unique cysteine for chemical modification or a biotin moiety for use with streptavidin-based reagents. 3.1. Phage Panning for Adherent Cell Lines
1. Cells are seeded onto tissue culture wells 24–48 h before panning. Only a single well is required for each panning round. Once started, each round of the panning procedure requires approximately 4–5 h to complete. Additional time is required for bacterial plating for titer determination and phage amplification. 2. 24–48 h before the phage biopanning will be conducted, trypsinize cells from a propagation flask and seed cells in 12-well plate. On the day of panning, one well should be ≈90% confluent. The proper level of confluence is generally obtained by seeding 100,000–150,000 cells in a well. 3. Begin the biopanning procedure by gently removing media from the well. Wash cells with 1 mL RPMI media (or other cell-appropriate media) without serum (tip plate to accumulate liquid on one side of the well so that media can be aspirated without disturbing attached cells. Pipette wash media gently to avoid dislodging cells. Remove wash media. 4. Gently add 1 mL/well media without serum and incubate cells for 2 h at 37°C to clear cell surface receptors (referred to as “clearing the receptors”).
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5. Approximately 15 min before the end of the clearing step, prepare the phage panning solution as follows: (a) Chloroquine (10 μL) (100× stock). (b) 40 μL protease inhibitor without EDTA (25× stock). (c) 10–100 library equivalents of the phage library. The phage library used for much of our work has a diversity of 1×108 members (5, 19). Therefore, we add 1×109 – 1×1010 phage to the input sample for round one. Thus, each library member should be present in 10–100 copies in the input mixture. For each successive round of biopanning we input ∼1.5×109 phage. (d) Bring mixture to 1 mL final volume by addition of PBS+ with 0.1% BSA. 6. Remove RPMI from the cells. Wash cells once with 1 mL PBS+ with 0.1% BSA that was prewarmed to 37°C. Remove the wash solution from the cells. 7. Save 50 μL of the input phage solution for titer determination. Add the remainder of the phage solution to the cells and incubate for 1 h at 37°C in a standard tissue culture CO2 incubator. 8. After the 1 h incubation, aspirate the supernatant. We do not save this solution containing unbound phage. Wash the cells four times at room temperature: (a) Add 1 mL PBS+ with 0.1% BSA (room temperature) (b) Incubate for 5 min. (c) Aspirate buffer and repeat. 9. Acid elute/wash 1–2 times at room temperature: (a) Add 1 mL 0.1 M HCl–glycine, pH 2.2 + 0.9% NaCl. (b) Incubate for 5 min. Time could be reduced if the cell line is fragile and lysis is problematic (see Note 9). (c) If you are interested in phage that bind to the cell surface but are not internalized, you can keep this acid wash fraction when it is removed from the cells and amplify the recovered phage as detailed below. Adjust the pH of the acid wash material by addition of 1.5 M Tris–HCl, pH 8.8 after it is removed from cells. (d) Repeat acid wash once. 10. Remove the second acid wash and add 1 mL of 30 mM Tris– HCl, pH 8.0 to the cells and incubate on ice for 30 min. This hypotonic media is used to swell the cells and enhance lysis and recovery of phage. 11. Freeze cells in plate. This is a suitable place to stop the protocol if needed.
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12. Thaw cells and scrape off plate. The freeze-thaw cycle disrupts the cells and releases any internalized phage. This sample is referred to as the output fraction. Examine the well under a microscope to ensure that the cells have been disrupted. If freeze-thaw does not disrupt the cells, 0.1% Triton X-100 or other detergent can be added to the hypotonic buffer. 13. Set up amplification of output phage as well as titration of input and output phage. Titer input, acid wash (if desired), and output. Amplify output or acid wash (if desired). 3.2. Phage Panning for Nonadherent Cells
1. The panning procedure requires approximately 4 h to complete. The cells can be removed directly from a feeder flask for the panning procedure. They do not have to be seeded into a separate flask prior to the day of the panning procedure. 2. Transfer cells from feeder flask to a 50-mL centrifuge tube and pellet cells (Speed and time required for forming a good pellet will vary with cell type). 3. Resuspend cells in 10 mL media without serum and pellet again. 4. Resuspend cells in 10 mL media without serum and incubate cells for 2 h at 37°C incubator to clear the receptors. 5. Count cells during clearing and determine volume needed for 2 million cells. 6. Approximately 15 min before the end of the clearing step, prepare the phage solution as detailed in step 5 of Subheading 3.1. 7. Pellet 2 million cells and wash one time with 10 mL PBS+ with 0.1% BSA pre-warmed to 37°C. 8. Save ∼50 μL input solution for titer determination. Pellet cells and resuspend the cell pellet gently in the remainder of phage solution and incubate for 1 h at 37°C in tissue culture incubator with 5% CO2. 9. At the end of the incubation, dilute the sample to 10 mL with PBS+ with 0.1% BSA (room temperature). 10. Wash the cells four times by centrifugation and resuspension. Resuspend cells in 10 mL PBS+ with 0.1% BSA and incubate for 5 min at room temperature. Pellet cells. 11. Acid wash cells at room temperature: (a) Resuspend cell pellet in 1 mL 0.1 M HCl–glycine, pH 2.2, 0.9% NaCl. (b) Incubate 5 min (see Note 9). (c) Pellet cells and remove supernatant.
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(d) Repeat acid wash once. Remove supernatant solution. 12. Remove the second acid wash and add 1 mL of 30 mM Tris– HCl, pH 8.0 to the cells and incubate on ice for 30 min. 13. Freeze cells and thaw. The freeze-thaw cycle disrupts the cell and releases any phage. This is referred to as the output fraction. 14. Set up amplification and titration of phage. Titer input, acid wash (if desired), and output. Amplify output and/or acid wash (if desired). 3.2. Phage Panning for Nonadherent Cells
1. Between successive rounds of phage biopanning, the output phage sample (and/or the acid wash sample, if desired) must be amplified. This procedure may be performed in parallel with the phage titering as detailed in Subheading 3.3 or may be performed independently. 2. On day one, pick a single K91 bacterial colony and inoculate 10–15 mL LB media without antibiotics for each sample that will be amplified. 3. Culture bacteria at 37°C with shaking until an OD600 nm of 0.2–0.4 is obtained. 4. Spin down bacterial cells at 3,000 × g for 10 min at 4°C. 5. Resuspend the pellet in 1/10 the original volume using LB media by pipetting up and down. 6. Add your phage sample to be amplified (the entire phage sample – 50 μL aliquot removed for titration) to resuspended K91 cells and incubate for 15 min at 37°C. 7. Dispense the complete mixture of phage-infected K91 cells onto four, 150 mm LB-tet plates. Plate 1/4 of the bacterial mix/plate. 8. Allow liquid to be absorbed into plate. 9. Invert plates and incubate at 37°C overnight. 10. On day two, harvest phage. Add 10 mL LB media to each of the four LB-tet plates. Incubate for 10 min at room tempe-rature. 11. Scrape bacteria off the plate with a glass spreader. Collect all the material from the four inoculated plates in a single 50 mL centrifuge tube. Some of the media will not be recovered from the plate. 12. Add 10 mL fresh LB media to one of the four plates. Use the glass spreader to clean the plate further and transfer the wash material to the second plate. Continue until all four plates have been washed in this manner. Combine this wash material with the original harvest in the 50 mL tube. 13. Centrifuge the harvested material to obtain a firm pellet of bacterial cells (Example: 3,000 × g for 10 min at 4°C in Beckman Coulter Allegra® 25R centrifuge.).
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14. The infectious phage particles will be in the supernatant from this centrifugation step. Transfer the supernatant to a fresh centrifuge tube and measure the volume. 15. Add ¼ of the supernatant volume of 2.5 M NaCl + 20% PEG 8000 to the supernatant. Example: if volume of supernatant is 40 mL, add 10 mL of the NaCl/PEG solution. Incubate this mixture on ice for 1 h to precipitate the phage particles. 16. Collect the phage precipitate by centrifugation at 11,000 × g for 30 min at 4°C. The phage should produce a firm pellet under these conditions. 17. Pour off and discard the supernatant making sure no standing liquid is left in the centrifuge tube. Tilt the centrifuge tube to drain off any residual PEG solution. Leave the tube inverted for 1 h at 4°C. Residual PEG solution will make it more difficult to completely resuspend the phage pellet. 18. After draining for 1 h, put the tube upright and add 1 mL PBS to the pellet. Incubate on ice for 30 min. During this incubation, tilt the tube so that the pellet is completely covered with PBS. 19. Gently resuspend the phage pellet using a 1 mL pipette. Do not vortex. Vortexing concentrated phage solution may result in shearing of the phage. Mix the samples so that there are no visible chunks or cakes in the sample. Transfer the resuspended phage to a clean 1.5 mL microcentrifuge tube. 20. Pellet insoluble debris by centrifugation at 16,000 × g for 2 min in a bench top microcentrifuge. 21. Transfer supernatant to a fresh microcentrifuge tube and incubate at 65°C for 15 min in a water bath or heating block to kill any bacteria remaining in the sample. Do not extend time of this incubation or the phage will lose infectivity. 22. Pellet insoluble debris by centrifugation at 16,000 × g for 2 min. 23. Transfer supernatant to a clean tube. Discard pellet. Mix and dispense aliquots of the purified phage to clean microcentrifuge tubes. Label tube with the cell type, panning round number, date and operator’s initials. 24. Store aliquots of the phage preparation at −80°C until use. 25. Before using on cells, set up bacterial titration to determine the yield of infective phage. The titration should be performed as detailed in Subheading 3.3 for the input phage sample except that more dilute samples are required to infect with bacteria. We typically dilute amplified phage preparations 10−2, 10−4, 10−6, 10−7, and10−8. The samples diluted 10−6, 10−7, and10−8 are used to infect K91 bacterial cells and aliquots of these infections are plated. 26. For amplification of individual phage clones (see Note 10).
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3.4. Bacterial cell Culture and Phage Titration (See Note 11)
1. We maintain K91 cells on minimal media minus proline supplemented with 0.5 μg/mL thiamine. The bacteria grow slowly on these plates, generally requiring 2 days of culture to produce suitable colonies. Each day that a titration will be performed, start a liquid culture of the bacteria from a single colony. 2. Pick a single colony and inoculate 5–10 mL LB media without antibiotics. 3. Culture bacteria at 37°C with shaking until an OD600 nm of 0.4 is obtained. If culture goes past an OD600 nm of 0.6, the culture should be diluted approximately tenfold with LB media and continue culturing until the proper optical density is obtained. 4. If bacterial cells are ready before samples that will be titered, place bacterial cells on ice until needed. If placed on ice, rewarm the cells to 37°C prior to mixing with phage samples. 5. Prepare serial dilution of input phage sample (50 μL aliquot was saved for titering): (a) Add 10 μL input phage to 990 μL LB media (=10−2 dilution). (b) Add 10 μL of 10−2 input phage dilution to 990 μL LB media (=10−4 dilution). (c) Add 100 μL of 10−4 input phage dilution to 900 μL LB media (=10−5 dilution). (d) Add 100 μL of 10−5 input phage dilution to 900 μL LB media (=10−6 dilution). 6. For titration of biopanning output samples: (a) Mix the freeze-thaw cell lysate well by flicking. (b) Add 50 μL of cell lysate to 450 μL LB media (=10−1 dilution). (c) Add 100 μL of 10−1 dilution to 900 μL LB media (=10−2 dilution). (d) For the early rounds of panning, these first two dilutions should be adequate. After round three, an additional tenfold dilution is suggested. 7. Dispense 900 μL K91 bacterial cells to sterile tubes for phage infection. For input phage samples, dispense aliquots to be infected with the 10−4, 10−5, and 10−6 dilutions, respectively. For the output phage samples, dispense aliquots to be infected with the 10−1 and 10−2 dilutions, respectively. Dispense one K91 aliquot that will serve as a noninfected control.
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8. Add 100 μL of diluted phage sample to 900 μL K 91 cells. Incubate at 37°C for 15 min (see Note 12). Label LB-tet plates for each sample. We inoculate 2 LB-tet plates for each infected K91 sample, one plate with 100 μL and second plate with 50 μL. 9. Add 100 μL of each infected K91 sample onto a separate, labeled LB-tet plate and spread evenly. 10. Add 50 μL of each infected K91 sample onto a separate, labeled LB-tet plate and spread evenly. 11. Additionally, plate 100 μL of uninfected K91 cells as a contamination control. After overnight incubation, these control plates should not have any colonies. 12. Allow plates to dry before inverting. 13. Incubate at 37°C overnight to allow colonies to grow. 14. After overnight incubation, count and record the number of colonies present on each plate. There should not be any colonies on the uninfected K91 cell plates. If colonies are present on these plates, the source of contamination needs to be eliminated and the samples need to be titered again. 15. Calculate titer of each sample. For each plate, use the formula: (# of colonies × dilution factor× 10 for dilution into K91)/mL of K91 mixture plated = colony forming units (cfu)/mL in the original sample. Calculate the independent determinations for each sample and average them to obtain sample titer. 16. The output titer plates from biopanning round 3 and subsequent rounds are saved for DNA sequence analysis and determination of phage displayed peptide sequences (see Subheading 3.6). 3.5. Real-Time, Quantitative-PCR Phage Titration (See Note 13)
1. This protocol assumes the use of the BioRad iCycler. Adjustments to the protocol may be required using other devices. We use the generalized Sybr® green detection system that does not require the generation of independent-labeled probes. 2. Turn on the lamp, camera, and thermal cycler at least 30 min before a reaction starts to stabilize per manufacturer’s suggestion. 3. Prepare standards and samples. Our calibration standard for titration consists of a series of tenfold dilutions of phage. We routinely generate this calibration standard line of infectious phage that range from 100 to 1 × 109 phage/mL. We have also run ssDNA and dsDNA preparations of phage in q-PCR. 4. Prepare the master mix for the number of reactions needed.
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Component
Volume per sample (µL)
2× IQ SYBR® Green Supermix
50
10 µM forward primer
3
10 µM reverse primer
3
H2O
34
Total
90
5. Dispense 90 μL aliquots of mastermix to clean PCR tubes (1 aliquot/sample or DNA standard). Add 10 μL of standard DNA or phage sample to 90 μL of the master mix. 6. Mix the 100 μL complete reaction mixture and dispense the reaction mixture into three wells of a 96-well plate that is optically suitable for q-PCR at 25 μL/well using an 8-channel pipette. 7. Spin down the plate to exclude bubbles at the bottom of the wells. 8. Enter the PCR Protocol and Plate Setup files from the iCycler or appropriate software. We use a 3-step protocol for the individual amplification cycles: Procedure
Temperature (°C)
Time
Hot start
95
3 min
Denature
95
30 s
Anneal
55
30 s
Extension
72
30 s
Amplification cycles
40 cycles Denature before melt curve analysis
95
1 min
Annealing before melt curve analysis
55
1 min
Melt curve analysis
0.5 up
10 s
80 cycles End
4
Hold
(e) The software for the iCycler (and other real-time thermocyclers) will automatically determine the parameters of the standard line and calculate the values of phage titers for each sample. Attention should be paid to the threshold parameter established by the software, the slope of the standard line (should be close to 3.2–3.3), and the melt curve analysis, which indicates the specificity of the amplified product. The PCR products can be evaluated by agarose gel electrophoresis if there is a question about amplification specificity.
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1. Dispense 50 μL water to 16 PCR tubes. 2. Label and pick 16-well-spaced colonies on a titer plate of interest (see Note 14). 3. Using a plastic pipette tip or toothpick, stab each colony and mix it into a different tube prepared in step 1. 4. Heat the samples at 95°C for 5 min to lyse bacteria and denature proteins. 5. Cool the samples while preparing PCR master mix containing 100 μL 10× PCR buffer without MgCl2, 60 μL 25 mM MgCl2, 20 μL 10 mM dNTP mix, 20 μL 10 μM forward primer (fd-tet F1), 20 μL 10 μM reverse primer (fd-tet B1), 730 μL water and 10 μL Taq polymerase. 6. Dispense 48 μL PCR mastermix to 16 new PCR tubes. 7. Add 2 μL of lysed bacterial colony sample to each aliquot of mastermix. 8. Perform PCR. We use the following protocol: Procedure
Temperature(°C)
Time
Hot start
95
2 min
Denature
95
30 s
Anneal
55
30 s
Extension
72
30 s
Amplification cycles
35 cycles Final extension
72
5 min
End
4
Hold
9. Evaluate PCR products on 1% agarose gel. Single products of approximately 450 bp are expected. 10. Remove dNTPs and oligonucleotide primers from the PCR product by combining 20 μL of PCR product with 2 μL exonuclease I and 2 μL shrimp alkaline phosphatase (SAP). Incubate at 37°C for 30 min followed by 15 min at 85°C. 11. After exonuclease I and SAP treatment, the PCR product can serve directly as template in a dideoxy terminator DNA sequencing reaction. We use BigDye® Terminator v3.1 in the following mixture: 5 μL treated product, 1 μL 10 μM primer (fd-tet F1), 3 μL 5× reaction buffer (supplied with BigDye® Terminator), 2 μL BigDye® Terminator Mix, and 9 μL water. BigDye® reactions are prepared in a 96-well plate. 12. Perform sequencing PCR using the following conditions:
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Procedure
Temperature(°C)
Time
Hot start
95
2 min
Denature
95°C
30 s
Anneal
55°C
30 s
Extension
72°C
30 s
Amplification cycles
30 cycles Final extension
72°C
5 min
End
4°C
Hold
13. Purify BigDye reaction by adding 80 μL sequencing stop/ precipitation reagent. 14. Collect precipitates by centrifugation at 3,000 × g for 30 min. 15. Invert plate onto a paper towel to collect liquid. 16. Centrifuge inverted plate at 100 × g for 1 min to remove liquid from plate. 17. Wash precipitates with 150 μL of 70% ethanol. Centrifuge at 3,000 × g for 10 min. 18. Invert plate onto a paper towel and centrifuge at 100 × g for 1 min to remove liquid from plate. 19. Add 26 μL Hi-Di formamide™ to each well. 20. Heat to 95°C for 5 min. 21. Cool plate and load onto ABI 3100 automated sequencer or other suitable sequencer. 22. Translate DNA sequence to peptide sequence for the segment encoding pIII protein at the point of the library insertion. 3.7. Selectivity and Specificity Determinations (See Note 15)
1. The selectivity determination is performed in a manner similar to the phage biopanning protocols for adherent and nonadherent cells detailed in Subheadings 3.1 and 3.2, with some important modifications. 2. Selectivity determinations require two matched wells of adherent cells or two aliquots of nonadherent cells. The adherent cells should be seeded onto wells 24–48 h before the experiment. For nonadherent cells, use 2 million cells/phage. 3. On the day of the experiment, the receptors are cleared by incubating cells in serum-free media for 2 h. 4. Approximately 15 min before the end of the clearing step, prepare the specific phage and control phage as separate solutions, each as follows:
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(a) 10 μL Chloroquine (100× stock) (b) 40 μL protease inhibitor without EDTA (25× stock) (c) 1 × 108 phage as input (see Note 16). (d) Bring each mixture to 1 mL final volume by addition of PBS+ with 0.1% BSA. 5. Wash cells once with 1 mL PBS+ with 0.1% BSA that was prewarmed to 37°C. Remove the wash solution from the cells. 6. Save 50 μL of the input phage solution for titer determination. Add the remainder of the phage solution to the cells and incubate for 10 min at 37°C in a standard tissue culture CO2 incubator. 7. After the 10 min incubation, aspirate the supernatant directly from adherent cells. Pellet nonadherent cells for each wash cycle. 8. Add 1 mL PBS+ with 0.1% BSA (room temperature) 9. Incubate for 5 min. 10. Aspirate buffer and repeat step 8–10 4 times to remove all weakly bound phage. 11. Acid elute/wash 1–2 times at room temperature: add 1 mL 0.1 M HCl–glycine, pH 2.2 + 0.9% NaCl. 12. Incubate for 5 min. Time could be reduced if the cell line is fragile and lysis is problematic (see Note 9). 13. Repeat acid wash once. 14. Remove the second acid wash and add 1 mL of 30 mM Tris– HCl, pH 8.0 to the cells and incubate on ice for 30 min. 15. Freeze cells in plate or centrifuge tube. This is a suitable place to stop the protocol if needed. 16. Thaw cells and scrape off plate, if needed. This sample is referred to as the output fraction. 17. Set up titration of input and output phage for the control and specific phage as detailed in Subheading 3.3. 18. The following calculations are used to compare phage binding to cells: (a) Output/input ratio (O/I) = output titer of phage/input titer of phage applied to cells. (b) Selectivity index = (O/I for specific phage clone)/(O/I for control phage). (c) Specificity is determined as the selectivity index for a specific phage clone tested on a panel of cell lines and primary cells. 3.8. Monomeric Peptide Synthesis
1. Monomeric peptide syntheses are performed on a Symphony® peptide synthesizer by Fmoc solid-phase peptide synthesis but all methods reported here can be adapted to other solid phase synthesizers.
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2. For a 0.1 mmol synthesis, place 141 mg Rink Amide AM resin (substitution level 0.71 mmol/g) in the peptide synthesis reaction vessel. Place the reaction vessel in one of the positions of the synthesizer. Add 2.5 mL DMF to swell the resin. Flush with nitrogen to form a suspension of resin for 30 min (see Note 17). Drain off DMF. 3. Repeat step 2 three times. 4. Add 2.5 mL Piperidine in DMF (20%) to deprotect the resin by removing the Fmoc moieties. Flush with nitrogen for 10 min. Drain off reagent. Repeat twice. 5. Add 2.5 mL DMF to the resin. Flush with nitrogen for 30 s. Drain off DMF. Repeat six times. 6. Add 2.5 mL Fmoc-protected amino acids in DMF (200 mM) to the deblocked peptidyl resin. Add 2.5 mL HBTU, HOBt, and NMM in DMF (200 mM) to the resin. Flush with nitrogen for 45 min. Drain off reagent. 7. Add 2.5 mL DMF to the resin. Flush with nitrogen for 30 s. Drain off DMF. Repeat six times. 8. Repeat the cycle from step 4 to step 7 for the next amino acid coupling until the completion of the peptide synthesis (see Notes 18 and 19). 9. Add 2.5 mL of 20% piperidine in DMF to the resin. Flush with nitrogen for 10 min. Drain off reagent. Repeat twice. 10. Add 2.5 mL DMF to the resin. Flush with nitrogen for 30 s. Drain off DMF. Repeat six times. 11. Add 2.5 mL DCM to the resin. Flush with nitrogen for 30 s. Drain off DCM. Repeat nine times. 12. Dry the resin for 10 min. 13. Place the dry resin in a 50 mL round bottom flask. Add 5 mL cleavage cocktail TFA: EDT:H2O:TIS (94%:2.5%:2.5%:1%) to the resin. For cysteine containing peptides use TFA: EDT:H2O:TIS (94%:2.5%:2.5%:1%) (see Notes 7 and 20). Flush flask with nitrogen, stopper, and leave to stand at room temperature with occasional shaking for 3 h. 14. Remove the resin by filtration under reduced pressure through a sintered glass funnel. Wash the resin twice with 3 mL cleavage cocktail (see Note 21). 15. Combine filtrates and transfer to an appropriate sized roundbottomed flask. Concentrate the TFA and scavenger mixture quickly to a volume of approximately 3 mL on a rotatory evaporator (see Note 22). 16. Fill a 50 mL conical tube about two-thirds full with cold diethyl ether. Add the concentrated peptide/TFA solution into cold ether using a Pasteur pipette. Place the cold ether with the peptide precipitate at −80°C freezer overnight.
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17. Centrifuge the cold ether with the peptide precipitate solution at 4°C for 10 min at 2,800–3,000 × g. Carefully decant the ether from the tube (see Note 23). 18. Add 50 mL fresh diethyl ether, seal and shake the tube to resuspend the peptide. Centrifuge for 10 min under the same conditions. Repeat three times. 19. Decant the ether from the tube. Put the tube in the hood for aproximately15 min to evaporate trace amounts of residual ether. Put the tube in the vacuum desiccator and dry the peptide under vacuum for 3 h. Weigh the peptide. 3.9. Linear Tetrameric Peptide Synthesis (See Note 24)
1. Tetrameric peptides syntheses are performed on a Symphony® synthesizer (Rainin Instruments, Protein Technologies, Inc. Woburn, MA) by Fmoc solid-phase stepwise peptide synthesis on trilysine core. These instructions are easily adaptable to other automated peptide synthesis instruments. 2. For a 0.1 mmol synthesis, place 476 mg Fmoc4-Lys2-Lys-βAla-CLEAR™ Acid Resin (substitution level 0.21 mmol/g) in the peptide synthesis reaction vessel (see Note 25). Place the reaction vessel in one of the positions of the synthesizer. Add 5 mL DMF to swell the resin. Flush with nitrogen to form a suspension of resin for 30 min (see Note 17). Drain off DMF. 3. Repeat step 2 three times. 4. Add 5 mL of 20% piperidine in DMF to remove the Fmoc protecting groups on the resin. Flush with nitrogen for 10 min. Drain off reagent. Repeat twice. 5. Add 5 mL DMF to the resin. Flush with nitrogen for 30 s. Drain off DMF. Repeat six times. 6. Add 5 mL Fmoc-protected amino acids in DMF (200 mM) to the deblocked peptidyl resin. Add 5 mL HBTU, HOBt, and NMM in DMF (200 mM) to the resin. Flush with nitrogen for 45 min. Drain off reagent. 7. Repeat steps 5 and 6 (see Note 26). 8. Add 5 mL DMF to the resin. Flush with nitrogen for 30 s. Drain off DMF. Repeat six times. 9. Repeat the cycle from step 4 to step 8 for the next amino acid coupling until the completion of the peptide synthesis. 10. Add 5 mL Piperidine in DMF (20%) to the resin. Flush with nitrogen for 10 min. Drain off reagent. Repeat twice. 11. Add 5 mL DMF to the resin. Flush with nitrogen for 30 s. Drain off DMF. Repeat six times. 12. Add 5 mL DCM to the resin. Flush with nitrogen for 30 s. Drain off DCM. Repeat nine times. 13. Continue from step 12, Subheading 3.8.
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3.10. Core Synthesis for Convergent Tetrameric Synthesis
1. Maleimido activated core syntheses are performed on a Symphony® synthesizer by Fmoc solid-phase peptide synthesis. These methods can easily be adapted for other solid phase peptide synthesizers. 2. For a 0.1 mmol synthesis, place 476 mg Fmoc4-Lys2-Lys-βAla-CLEAR™ Acid Resin, Fmoc4-Lys2-Lys-Lys(Biotin-PEG) -β-Ala-CLEAR™ Acid Resin and Fmoc4-Lys2-Lys-Cys(Acm)β-Ala-CLEAR™ Acid Resin, respectively (substitution level 0.21 mmol/g) in the peptide synthesis reaction vessel (see Notes 27 and 28). Place the reaction vessel in one of the positions of the synthesizer. Add 5 mL DMF to swell the resin. Flush with nitrogen to form a suspension of resin for 30 min. Drain off DMF. 3. Repeat step 2 three times. 4. Add 5 mL of 20% piperidine in DMF as a deprotection reagent to remove Fmoc protecting groups from the resin. Flush with nitrogen for 10 min. Drain off reagent. Repeat twice. 5. Add 5 mL DMF to the resin. Flush with nitrogen for 30 s. Drain off DMF. Repeat six times. 6. Add 2.5 mL 3-maleimidopropionic acid in DMF (200 mM) to the deblocked peptidyl resin. Add 2.5 mL HBTU, HOBt, and NMM in DMF (200 mM) to the resin. Flush with nitrogen for 24 h. Drain off reagent. 7. Continue from step 10, Subheading 3.98.
3.11. Tetrameric Peptide Synthesis by Convergent Strategy (Scheme 18.I)
1. Pipette 1.5 mL 1× PBS/0.01 M EDTA in a microcentrifuge tube. Purge with Argon for 3 min.
3.12. Removal of an Acetamidomethyl Group from a Uniquely Placed Cysteine Residue
1. Dissolve 1 μmol tetrameric peptide possessing an acetamidomethyl (Acm) protecting group in 1 mL of TFA/anisole (99:1).
2. Dissolve 8 μmol of monomeric peptide containing a unique cysteine and 1 μmol of maleimido activated core in Ar-purged 1× PBS/0.01 M EDTA. Vortex it at room temperature for 2 h (see Notes 5 and 29).
2. Add 28 mg of silver acetate. Stir the solution at 4°C for 2 h. 3. Concentrate under argon to 0.5 mL. 4. Add 5 mL cold diethyl ether. Centrifuge the cold ether with the peptide precipitate solution at 4°C for 10 min. 5. Decant the ether from the tube. 6. Add 1 mL of 0.2 M dithiothreitol prepared in 1 M acetic acid. Vortex solution at room temperature for 3 h. 7. Add 0.5 mL of 8 M guanidine hydrochloride. Filter the solution by syringe driven filter unit (0.22 μm porosity). Purify the peptide by HPLC.
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Scheme 18.1. Convergent synthesis of tetrameric cell-binding peptides. A biotin or a unique cysteine can be incorporated into the peptide for use in other applications. A PEG moiety is placed between the trilysine branch and the selected peptide to increase aqueous solubility.
3.13. Peptide Purification and Characterization
1. Filter eluents through a 0.45 μm Teflon® filter before use. 2. Dissolve 15 mg of crude peptide in 2 mL of Buffer A. Filter the sample through a 0.22 μm filter.
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3. Equilibrate the HPLC preparative column under the following initial conditions. Solvent: buffer A. Flow rate: 10 mL/ min. Detection wavelength: 220 nm. 4. Once a stable baseline is obtained, inject 2 mL of the sample and use the elution profile in a linear gradient (referred to as Method A): 0–1 min, 90%A, 10%B; 1–61 min, eluent B was increased from 10 to 40% at a flow rate of 10 mL/min. 5. Collect the target peptide peak which is generally the major peak (see Note 30). 6. Lyophilize fractions containing the peptide product. If available, the fractions can be analyzed by MALDI MS to determine which fractions to collect. 7. For analytical HPLC, dissolve 1 mg of purified peptide in 1 mL of Buffer A. Filter the sample through a 0.22 μm filter. 8. Equilibrate the HPLC analytical column under the following initial conditions. Solvent: buffer A. Flow rate: 1 mL/min. Detection wavelength: 220 nm. 9. Once a stable baseline is obtained, inject 100 μL of the sample and use the elution profile in a linear gradient (referred to as Method B): 0–1 min, 90%A, 10%B; 1–51 min, eluent B was increased from 10 to 60% at a flow rate of 1 mL/min. 10. Confirm the peptide mass by mass spectrometry. For ease we typically perform matrix assisted laser desorption ionization time of flight mass spectrometry (MALDI MS) (see Note 31). 11. Edman N-terminal sequence analysis can be performed to further confirm the sequence of the peptide. 3.14. Inhibition of Phage Uptake by Free Synthetic Peptides (See Note 32)
1. Perform phage uptake experiments in the same manner as outlined in Subheading 3.7 except that the free peptide is added to the phage solution before addition to the cells. No prior addition of the peptide to the cells is required. We typically cover a peptide concentration range from 1 nM to 10 μM (see Note 33). 2. Titer the common input sample and the individual output samples as detailed in Subheading 3.3. 3. Calculate the output phage to input phage ratio in the presence of the peptide compared with the same ratio without added peptide.
3.15. Microscopy/Qdot Delivery (See Note 34; Fig. 1)
1. In a final volume of 100 μL PBS, streptavidin-Qdots (200 nM) are mixed with 600 nM biotinylated peptide and incubated for 2 h at room temperature. This incubation is performed on the day the Qdots are to be used. Control SA-Qdots can be prepared using no peptide, an irrelevant sequence peptide or a scrambled version of the specific peptide.
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Fig. 18.1. A tetrameric lung cancer targeting peptide can selectively deliver a fluorescent nanoparticle to its target cells. The peptide TP H1299.2 was isolated by biopanning on the large cell lung cancer cell line H1299. H1299 cells were incubated with 20 nM of the tetrameric TP H1299.2 peptide conjugated to SAQDot605 (D)–(F) or 20 nM tetrameric control peptide conjugated to SAQDot605 (A)–(C). Bright field images (A), (D) and nuclear staining images (B), (E) of the corresponding fields are shown. The fluorescence of the Qdot was visualized at 200-fold magnification on a Nikon TE2000 fluorescent scope. Higher magnification (×1,000) shows cell surface binding of the tetrameric TP H1299.2 peptide-SAQDot605 conjugate as well internalized particles (G)–(I). (Reproduced from ref. 5 with permission from Elsevier Science.).
2. At the end of the SA-Qdot-peptide incubation, unoccupied streptavidin sites on the Qdots are quenched by the addition of excess biotin (25 μL of a 20 μM stock) and incubated for 15 min at room temperature. 3. The mixture is diluted to 1 mL with PBS+ with 0.1% BSA to obtain a 20 nM Qdot solution for cell uptake. 4. Cells are incubated with Qdots on chamber slides or in polypropylene tubes for 10 min to 2 h. 5. At the end of the incubation, the solution containing Qdots is removed and cells are washed briefly four times in PBS+ with 0.1% BSA. The wash solution is added and then is removed without further incubation.
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6. Cells are briefly washed with 0.1 M HCl–glycine, pH 2.2. + 0.9% NaCl. Acid wash is removed after addition without further incubation. 7. Cells are washed with PBS and excess liquid is removed from the slide. 8. Cells can be viewed without fixation or they can be fixed using 0.4% formalin solution, 70% ethanol or cold acetone solution if desired. In this case, a 5 min incubation in fixative is followed by PBS washes to remove fixative solution. 9. Incubation chamber is removed from chamber slide and Prolong® Gold antifade reagent with DAPI stain is added to the samples. If suspension grown cells were used, samples can be spotted by hand onto slide or by using a CytoSpin centrifuge, if available. 10. Add cover slip to slide and seal with fingernail polish. 11. Observe samples under microscope. 3.16. Capture of Cells with Phage-Coated Tissue Culture Wells
1. Phage coated plates can be prepared by incubating phage solution (106 phage/mL) in wells at 4°C overnight. For 12-well plates, use 1 mL phage solution per well. For 96-well plates, use 0.1 mL per well. 2. Phage solution is flicked from plate and residual binding sites are masked by incubation of the wells (filled to capacity) in 0.1% BSA in PBS (1 h to overnight). 3. Nonadherent cells are removed from their culture flask and washed by centrifugation in PBS+ with 0.1% BSA. Cells are dispensed to wells containing specific phage or control phage or no phage and incubated for 10 min to 1 h. 4. Wells are washed four times with PBS+ with 0.1% BSA to remove unbound cells. 5. The number of captured cells can be determined by direct cell counting – in the well or after release using nonenzymatic cell release solution or trypsinization. Alternatively, cell numbers can be determined by lysis and assay for a specific cellular component such as ATP. ATP content can be assayed using the commercially available reagent that requires only a single reagent addition to the wells (CellTiterGlo™, Promega). 6. The captured cells can be further characterized by downstream processes for gene or protein expression.
3.17. Capture of Cells with Peptide-Coated Magnetic Beads
1. Cells are cleared by incubation for 2 h in RPMI without serum as detailed in Subheadings 3.1 and 3.2. 2. 50 μL Streptavidin-coated Dynabeads® are washed twice by suspension in 1 mL PBS and capture with a magnet for 5 min and removal of the wash media by aspiration.
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3. The magnetic beads are suspended in 1 mL PBS and allowed to react with biotin-modified synthetic peptide (50 nM) for 30 min at room temperature. 4. The ligand-coated beads are washed twice in RPMI without serum to block any remaining streptavidin sites (RPMI contains 200 mg biotin/L). 5. Magnetic beads are washed once in PBS+ with 0.1% BSA, then resuspended in 1 mL PBS+ with 0.1% BSA, and mixed with cells. 6. Cells are incubated with the ligand-coated magnetic beads for 15 min at 37°C. 7. Nonadherent cells that take up magnetic beads are captured on the magnet (5 min at room temperature). 8. Adherent cells are released from wells using enzyme-free cell dissociation buffer (GIBCO) before capture on the magnet. Time of dissociation will vary with the cell type. Some cells are easily released after a 5 min incubation on ice in dissociation buffer while other are released more effectively at room temperature or even 37°C. 9. Cells are washed by suspension in PBS+ with 0.1% BSA by release from the magnet and recapture. 10. Captured cells are then suitable for additional analysis. 3.18. Flow Cytometry
1. Cell samples are cleared by incubation in RPMI without serum for 2 h. 2. During last 15 min of clearing, peptide solution is prepared containing: (a) 10 μL Chloroquine (100× stock). (b) 40 μL protease inhibitor without EDTA (25× stock). (c) Synthetic peptide construct with biotin – concentration required varies with cell and peptide combination. No peptide addition and control peptide solutions are prepared separately. (d) PBS+ with 0.1% BSA to volume of 1 mL/sample. 3. Incubate cells with peptide for 15 min at 37°C. 4. Wash the cells in PBS+ with 0.1% BSA four times at room temperature. 5. Cells are diluted in PBS+ with 0.1% BSA containing streptavidin–phycoerythrin (SA-PE) at 1:100 dilution in the mix. 6. SA-PE conjugate is added and cells are incubated for 15 min at room temperature. 7. Stained cells are washed once and resuspended in fresh PBS+ with 0.1% BSA.
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8. Adherent cells can be released from the wells by incubation in enzyme-free cell dissociation buffer as detailed in Subheading 3.17. 9. Cell staining is evaluated by flow cytometry. Cells can be counterstained with viability marker such as Alexafluor 488-Annexin V added to resuspension buffer and incubating for 5 min at room temperature prior to loading of sample.
4. Notes 1. The phage library used in most of our selections for cell ligands displays a 20 mer peptide at the amino-terminus of the minor coat protein pIII (19). We have selected peptides from other phage display libraries (5), including the Ph.D. 12-mer library available from New England Biolabs(MJM and KCB unpublished results). 2. 20% PEG-8000 in 0.9% NaCl solution should be prepared in advance. We routinely prepare 500–1,000 mL of this solution. PEG-8000 dissolves slowly even with constant stirring. Preparation of the solution can take several hours. After all of the PEG-8000 is in solution, filter-sterilize the solution using 0.22 μm membranes (Millipore Corp., catalog # SCGP05RE). This process is also relatively slow because of the viscous nature of the solution. 3. 2× Sybr® green mixes for q-PCR are available from a number of companies. We have found the DyNAmo™-Sybr® Green qPCR kit (Finnzymes Inc., Distributed in USA through New England Biolabs) to produce the most consistent results in amplification of phage DNA recovered from mammalian cells and tissues. Some mammalian tissues appear to have a factor that inhibits the quantitative amplification of phage DNA from crude extracts. In this case we have purified total DNA from the tissue, using genomic DNA isolation kits from Qiagen, prior to qPCR. Even using purified DNA, inhibition was observed in some tissues. The recommended Sybr® green mix uses a polymerase that appears to be less subject to inhibition by these tissues. 4. Although we amplify a region of the Tet resistance gene, other constant regions of the phage genome can be amplified. 5. Fmoc-NH–(PEG)11–COOH is incorporated to increase the water solubility of the peptide if necessary. We routinely place this PEG linker between the trilysine core and the targeting peptide in the tetrameric constructs.
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6. In contrast to Fmoc-Lys(biotin)–OH, Fmoc-Glu(biotinylPEG)–OH has excellent solubility in DMF and other solvents used in solid phase peptide synthesis. The PEG-spacer restricts hindrance between the peptide and avidin, leading to better biotin binding. 7. Cleavage cocktails should always be prepared fresh. The cleavage procedure generally takes 2–4 h to perform. However, some protecting groups are quite stable to TFA depending on the location and number in a sequence, requiring up to 12 h of treatment for complete removal. 8. Amine impurities that could possibly remove the Fmoc group include dimethylamine found in DMF. It is recommended that DMF is protected under nitrogen or freshly purified before use. 9. Although most cells we have tested have been stable during the acid wash step of the protocol, some cells are lysed by this treatment (primary cardiac myocytes and A20 B cell lymphoma cells as examples). In these cases, we have deleted this step from the protocol, reduced the incubation time in the acid, or only performed a single acid wash. If necessary to isolate only internalized phage, others have treated the cells with subtilisin or other proteases to inactivate surface bound phage (16). 10. Since the output phage sample is a mixture of individual library members, we prefer to amplify the phage mixtures as colonies on large LB-tet plates. This will allow the individual members to grow without interference from competing phage. For single phage clones, we grow infected E. coli in liquid media (LB + 12 μg/mL tetracycline). The preparation of phage from liquid cultures is the same as from plates from step 13 of Subheading 3.4. 11. Titration is used to determine the number of infectious phage particle per milliliter of solution. It is used in determining the volume of a phage stock that will be added to a panning solution as well as the actual phage number in the input and output samples from a round of panning or comparative binding. Phage infection of E. coli requires that the bacteria express pili. With fd-tet phage, the phage confer tetracycline resistance on the bacteria. Bacteria without phage will not grow on the LB-tet plates. The fd-tet phage do not cause cell lysis. In fact, only phage-infected bacteria will produce isolated colonies on LB-tet plates. 12. During this step, phages are allowed to infect bacteria. Do not incubate the samples for more than 15 min. Place samples on ice immediately after removal from the incubator. Bacteria should be plated before cells have time produce progeny phage.
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13. We use real-time quantitative PCR for three distinct purposes. The first is to determine phage copy number after the phage have lost the ability to infect bacteria. For example, phage infectivity decreases rapidly after injection into a mammalian host. Second, we have used q-PCR to determine the presence of a specific phage clone in a mixture of phage. This has allowed us to add mixtures of phage in an experiment and have one phage serve as an internal control. Third, we have used q-PCR to determine phage levels in a large number of samples simultaneously. We can determine total phage copy number using sets of primers directed at a nucleotide sequence in the backbone of the phage DNA. Additionally, we can determine the copy number of a specific phage clone using one primer directed at the nucleotide sequence encoding the displayed peptide and a second primer for a sequence from the phage backbone. Not all nucleotide sequences for the displayed peptides have been suitable for generating useful clone-specific primer sequences. 14. Titration plates with well-spaced and defined colonies can be used for determination of displayed peptide sequence. We initiate sequencing with the output of the third round of biopanning. DNA sequencing is also performed to verify the identity of an amplified phage clone. Since we use an ABI 3100 DNA sequencer that has a 16 capillary array, we routinely sequence samples in groups of 16. 15. To determine the initial success of a phage display peptide selection, we compare the binding and uptake of a specific phage with the binding and uptake of a randomly chosen, control phage using the same cells employed for the selection. The randomly chosen, control phage mimics the binding characteristics of the whole phage display library in these assays. The selectivity value is the ratio of specific phage output/input and the control phage output/input. The evaluation of specificity compares the selectivity values of an individual phage across a battery of cell lines or primary cells. 16. Determinations of selectivity and specificity are comparative binding assays based on the ratio of the number of phage taken up by a cell divided by the number of phage to which the cells were exposed. A specific phage clone is being compared with a control phage that represents a nonspecific component of phage uptake by a cell sampling its environment. Therefore, it is important to match the amount of specific and control phage used as input. For these assays, the input phage should be added at 1 × 108 phage/sample. Deviations from the input phage number will distort the ratios used for comparison and produce artificially high or low selectivity indices.
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17. We recommend adjusting the N2 flow to 10 psi. Higher flow rates flush the resin to the top of the reaction vessel resulting in incomplete coupling. 18. Sometimes the coupling reaction of an activated carboxy group and a deprotected amino group is difficult to accomplish. These difficult couplings are usually sequence-dependent and not residue-specific. In these cases, a double coupling is required (i.e., repeat step 5–6 before step 7). 19. Fmoc-NH–(PEG)11–COOH, Fmoc-Glu(biotinyl-PEG)–OH and Fmoc-Lys(biotin)–OH are coupled in the same fashion as the natural Fmoc-protected amino acids. 20. Always handle thiol-containing substances in proper ventilation. These compounds have an offensive odor that can be neutralized with bleach. After the final cleavage operation, rinse all glassware, pipettes, and tubes that came into contact with scavengers with bleach before taking them out of the hood. 21. The expended resin should not be discarded but retained, in case it should prove necessary to repeat the cleavage reaction. Many times during a poor extraction step, peptide remains adhered to the resin beads and must extracted with an alternative solvent. 22. Temperature of the water bath should be below 37°C. 23. The diethyl ether washes should be retained until the yield of product has been established. If a poor yield is obtained, the washings should be evaporated under vacuum to dryness. 24. Linear synthesis of the tetrameric peptides is only recommended for shorter peptides, on the order of ten amino acids or less in length. We strongly prefer the synthetic route outlined Subheading 3.11. 25. We strongly encourage reducing the substitution of the resin for tetrameric peptide synthesized linearly, longer molecules (>30 residues) or for peptides rich in β-structural elements to a substitution value lower than 0.25 mmol/g of resin. 26. A double coupling is usually preformed to increase the stepwise coupling yield and avoid deletion contaminants. 27. For the maleimido activated core synthesis, the substitution of the resin does not have to be lower than 0.25 mmol/g. A normal substitution between 0.5 and −0.8 mmol/g will suffice the synthesis. 28. The resin of choice depends on the down-stream application of the peptide. We frequently attach the peptides to a desired support, bead, or molecule via a unique cysteine located before the trilysine branch point. The Acm protecting group can be removed as described in Subheading 3.12 without
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loss of the peptide branches. If using streptavidin conjugates, the biotin containing resin is used instead. 29. After 2 h reaction time, the reaction solution must be purified by HPLC immediately. Otherwise, self-oxidized peptide dimer side products occur as a result of disulfide formation. 30. For the tetrameric peptide synthesized by linear strategy, the RP-HPLC trace typically shows one broad main peak. For monomeric peptides, the RP-HPLC trace usually shows one clear main peak. The occurrence of peaks with longer retention times than the main peak is suggestive of incomplete removal of protecting groups. For the tetrameric peptide synthesized by convergent strategy, the RP-HPLC trace typically shows two major peaks. The peak with the lower retention time is excess amount of monomeric peptide. The other with the higher retention time is the product tetrameric peptide. 31. For monomeric peptides, the mass is confirmed by MALDI MS using a-cyano-4-hydroxycinnamic acid as the matrix using a Voyager DE™ Pro instrument in reflector mode. MALDI Mass Spectra of tetrameric peptide is obtained in linear mode using sinapinic acid as matrix. 32. To determine whether the peptide is functional outside of the phage particle, we determine whether the peptide can block cell binding of its cognate phage. This assay is particularly useful when the cellular target of the peptide is unknown. Half maximal phage blocking can be determined as a measure of peptide affinity. 33. Do not use DMSO or DMF for preparation of peptide stock solutions. These compounds appear to produce a higher level of cell uptake of phage when added to cell-phage mixtures. Incorporation of a PEG moiety improves the peptide’s water solubility and often alleviates the need for organic co-solvents. 34. Cell-binding of the selected phage clones can be determined by fluorescent microscopy utilizing anti-phage antibodies (2, 6). However, since our goal is to rapidly translate the peptide outside of the context of the phage, we most often move directly to using the free peptide for these studies.
Acknowledgments This work was supported by the National Cancer Institute of the NIH (1RO1CA106646 and R211R21CA114157-01).
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References 1. Brown, K. C. (2000) New approaches for cellspecific targeting: identification of cell-selective peptides from combinatorial libraries. Curr. Opin. Chem. Biol. 4, 16–21 2. McGuire, M. J., Sykes, K. F., Samli, K. N., Barry, M. A., Stemke-Hale, K. A., Tagliaferri, F., Logan, M., Jansa, K., Takashima, A., Brown, K. C., and Johnston, S. A. (2004) A library selected Langerhans cell-targeting peptide enhances an immune response. DNA Cell Biol. 23, 742–752 3. McGuire, M. J., Samli, K. N., Johnston, S. A., and Brown, K. C. (2004) In vitro selection of a peptide with high selectivity for cardiomyocytes in vivo. J. Mol. Biol. 342, 171–182 4. McGuire, M. J., Samli, K. N., Chang, Y., and Brown, K. C. (2006) Novel ligands for cancer diagnosis: selection of peptide ligands for identification and isolation of B-cell lymphomas. Exp. Hem. 34, 443–452 5. Oyama, T., Rombel, I. T., Samli, K. N., Zhou, X., and Brown, K. C. (2006) Isolation of multiple cell-binding ligands from different phage displayed-peptide libraries. Biosens. Bioelectron. 21, 1867–1875 6. Oyama, T., Sykes, K. F., Samli, K. N., Minna, J. D., Johnston, S. A., and Brown, K. C. (2003) Isolation of lung tumor specific peptides from a random peptide library: generation of diagnostic and cell-targeting reagents. Cancer Lett. 202, 219–230 7. Samli, K. N., McGuire, M. J., Newgard, C. B., Johnston, S. A., and Brown, K. C. (2005) Peptide-mediated targeting of the islets of Langerhans. Diabetes 54, 2103–2108 8. De, J., Chang, Y., Samli, K. N., Schisler, J. C., Newgard, C. B., Johnston, S. A., and Brown, K. C. (2005) Isolation of a Mycoplasmaspecific binding peptide from an unbiased phage-displayed peptide library. Mol. Biosyst. 1, 149–157 9. Shadidi, M., and Sioud, M. (2003) Identification of novel carrier peptides for the specific delivery of therapeutics into cancer cells. FASEB J. 17, 256–258 10. Shadidi, M., and Sioud, M. (2004) Selection of peptides for specific delivery of oligonucleotides into cancer cells. Methods Mol. Biol. 252, 569–580
11. Hong, F. D., and Clayman, G. L. (2000) Isolation of a peptide for targeted drug delivery into human head and neck solid tumors. Cancer Res. 60, 6551–6556 12. Zhang, J., Spring, H., and Schwab, M. (2001) Neuroblastoma tumor cell-binding peptides identified through random peptide phage display. Cancer Lett. 171, 153–164 13. Barry, M. A., Dower, W. J., and Johnston, S. A. (1996) Toward cell-targeting gene therapy vectors: selection of cell-binding peptides from random peptide-presenting phage libraries. Nat. Med. 2, 299–305 14. Kim, Y., Lillo, A. M., Steiniger, S. C. J., Liu, Y., Ballatore, C., Anichini, A., Mortarini, R., Kaufmann, G. F., Zhou, B., Felding-Habermann, B., and Janda, K. D. (2006) Targeting heat shock proteins on cancer cells: selection, characterization, and cell-penetrating properties of a peptidic GRP78 ligand. Biochemistry 45, 9434–9444 15. Kolonin, M. G., Bover, L., Sun, J., Zurita, A. J., Do, K. A., Lahdenranta, J., Cardo-Vila, M., Giordano, R. J., Jaalouk, D. E., Ozawa, M. G., Moya, C. A., Souza, G. R., Staquicini, F. I., Kunyiasu, A., Scudiero, D. A., Holbeck, S. L., Sausville, E. A., Arap, W., and Pasqualini, R. (2006) Ligand-directed surface profiling of human cancer cells with combinatorial peptide libraries. Cancer Res. 66, 34–40 16. Robinson, P., Stuber, D., Deryckere, F., Tedbury, P., Lagrange, M., and Orfanoudakis, G. (2005) Identification using phage display of peptides promoting targeting and internalization into HPV-transformed cell lines. J. Mol. Recognit. 18, 175–182 17. Li, S., Liu, Y. H., McGuire, M. J., and Brown, K. C. (submitted) Facile synthesis of multimeric peptides: development of cell-specific delivery systems. 18. Zhou, X., Chang, Y., Oyama, T., McGuire, M. J., and Brown, K. C. (2004) Cell-specific delivery of a chemotherapeutic to lung cancer cells J. Am. Chem. Soc. 129, 15656–15657 19. Cwirla, S. E., Peters, E. A., Barrett, R. W., and Dower, W. J. (1990) Peptides on phage: a vast library of peptides for identifying ligands Proc. Natl. Acad. Sci. USA 87, 6378–6382
Chapter 19 Biosensor Detection Systems: Engineering Stable, High-Affinity Bioreceptors by Yeast Surface Display Sarah A. Richman, David M. Kranz, and Jennifer D. Stone Summary Over the past two decades, the field of biosensors has been developing fast, portable, and convenient detection tools for various molecules of interest, both biological and environmental. Although much attention is paid to the transduction portion of the sensor, the actual bioreceptor that binds the ligand is equally critical. Tight, specific binding by the bioreceptor is required to detect low levels of the relevant ligand, and the bioreceptor must be stable enough to survive immobilization, storage, and in ideal cases, regeneration on the biosensing device. Often, naturally-occurring bioreceptors or antibodies that are specific for a ligand either express affinities that may be too low to detect useful levels, or the proteins are too unstable to be used and reused as a biosensor. Further engineering of these receptors can improve their utility. Here, we describe in detail the use of yeast surface display techniques to carry out directed evolution of bioreceptors to increase both the stability of the molecules and their affinity for the ligands. This powerful technique has enabled the production of stabilized, single-chain antibodies, T cell receptors, and other binding molecules that exhibit affinity increases for their ligands of up to 1 million-fold and expression of stable molecules in E. coli. Key words: Yeast surface display, Directed evolution, Affinity maturation, Thermal stability.
1. Introduction The design of rapid, accurate, transportable, and inexpensive detection tools for a variety of biological and environmental molecules is an extremely desirable goal. One way that this may be achieved is using biosensors, which most often combine a bioreceptor that can unambiguously detect low levels of the target molecule, and a transducer that can sense and transmit binding events (reviewed in (1, 2) ). Current research seeks to design biosensors that can
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_19
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sensitively and specifically detect infectious agents, toxins, allergens, hormones, illicit drugs, and even environmental contaminants (reviewed in (3–5) ). A successful biosensor should achieve multiple goals: it must be able to clearly detect the molecule of interest at a level of practical consequence (e.g., infectious or toxic levels); it must be able to provide reproducible results after medium to longterm storage; and it must be able to achieve a sufficiently low cost per test through ease of production, durability, and ability to be regenerated and reused. As the critical discriminating element of a biosensor, the bioreceptor molecule must exhibit exquisite stability, sensitivity, and specificity of binding (5). Although specific receptors such as monoclonal antibodies can be generated against many target molecules, further engineering of these antibodies can make them more suitable for biosensing applications (6). Yeast surface display is a powerful technique that can be applied toward increasing the desirable properties of antibodies and other biomolecules ( (7), and reviewed in (8) ). This method includes several advantages over other display and engineering systems (9). The ability to screen bioreceptor libraries by fluorescence activated cell sorting (FACS) allows quantitative comparisons between mutants to help guide the selection. In addition, ligand-binding constants to soluble target molecules may be directly measured on the yeast cell surface; these values are typically in qualitative agreement with Biacore measurements of the interaction (10). Furthermore, yeast display incorporates adaptability for alternative sorting protocols for ligands that are difficult to produce or fluorescently label in soluble form, including magnetic bead separation (11) and cell–cell conjugates (12, 13). Yeast surface display has been successfully applied to the engineering of many bioreceptor proteins including antibodies (14–16), T-cell receptors (17–19), major histocompatibility complex proteins (20–23), NK receptors (24), growth factors (25), superantigen antagonists (26), fibronectin domains (27, 28) and cytokines (29, 30). In this chapter, we describe detailed protocols for expression, mutation, and selection of optimal bioreceptor proteins by yeast surface display. Through this process, bioreceptor mutants can be engineered with greater stability for storage and regeneration conditions, as well as higher affinities for the detection of very low ligand concentrations.
2. Materials 2.1. Yeast Strain
Saccharomyces cerevisiae yeast display strain EBY100 (a GAL1AGA1::URA3 ura3-52 trp1 leu2D1 his3Δ200 pep4::HIS2
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prb1Δ1.6R can1 GAL) is commercially available from Invitrogen (cat. no. C839-00). 2.2. Yeast Display Plasmid
Yeast display plasmid pCT302 (or a variant thereof) is commercially available from Invitrogen as pYD1 (cat. no. V835-01).
2.3. Electrocompetent E. Coli Strain for Plasmid Propagation
DH10B™ (Invitrogen cat. no. 18290-015).
2.4. Restriction Enzymes
NheI (Invitrogen cat. no.15444-011), BglII (Invitrogen cat. no. 15213-010), XhoI (Invitrogen cat. no. 15231-012), and DpnI (Invitrogen cat. no. 15242-019).
2.5. Polymerase Chain Reaction (PCR)
1. dNTPs (100 mM) (Invitrogen cat. no. 10297-018). 2. Taq DNA polymerase (Invitrogen cat. no. 18038-042). 3. High-fidelity Pfu Turbo® DNA polymerase (Stratagene cat. no. 600250). 4. Bovine serum albumin (BSA) (Sigma cat. no. A4503).
2.6. DNA Purification
1. Plasmid rescue and purification from E. coli: QIAprep spin miniprep kit (QIAGEN cat. no. 27104). 2. Plasmid rescue from yeast: Zymoprep kit I or II (Zymo research cat. no. D2001 (I) or D2004 (II) ). 3. QIAquick PCR purification kit (QIAGEN cat. no. 28104).
2.7. Yeast Media
1. YPD media for propagation of EBY100. To prepare 500 mL YPD, dissolve 5 g yeast extract (BD Bacto cat. no. 212750), 10 g peptone (BD Bacto cat. no. 211677), and 10 g dextrose (EMD cat. no. 346351) to 500 mL in ddH2O. Autoclave and store at room temperature no longer than 1 month. To make 1 sleeve of YPD plates, add 7.5 g of agar to the above recipe. Autoclave. Pour plates when cool enough to handle. 2. SD-citrate-CAA media for propagation of EBY100 transformed with pCT302. To prepare 500 mL SD-citrate-CAA, dissolve 7.4 g sodium citrate (Fisher cat. no. S279-500), 2.1 g citric acid monohydrate (Sigma cat. no. C1909), 2.5 g casamino acids (BD/Bacto cat. no. 223120), 10 g dextrose, and 3.35 g yeast nitrogen base without amino acids (Sigma cat. no. Y0626) to 500 mL in ddH2O. Filter sterilize. Store at 4°C for up to 6 months. For SD-citrate-CAA plates, dissolve 91 g sorbitol (Sigma cat. no. S1876), 7.5 g agar, 7.4 g sodium citrate, and 2.1 g citric acid monohydrate to 400 mL in ddH2O. Autoclave. In a separate container, dissolve 2.5 g casamino acids, 10 g dextrose, and 3.35 g yeast nitrogen base without amino acids to 100 mL in ddH2O; add to the autoclaved
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solution when it is cool. Mix gently. Pour plates. Store plates at 4°C for up to several months. 3. SG-citrate-CAA media for induction of yeast display construct. Follow the recipe for 500 mL SD-citrate-CAA above substituting an equal mass of D-(+)-galactose (Sigma cat. no. G-0625) for dextrose. 4. Antibiotic for SD-citrate-CAA and SG-citrate-CAA cultures. Use 50 μg/mL kanamycin. Prepare 50 mg/mL stock by dissolving 500 mg kanamycin sulfate (Sigma cat. no. K4000) in 10 mL ddH2O. Filter sterilize. Aliquot and store at −20°C. 2.8. Solutions for High-Efficiency Yeast Transformation Protocol
1. 10 × TE buffer pH 7.5: To make 250 mL of the buffer, add 25 mL of a 1 M Tris Base stock (pH 7.5) and 5 mL 0.5 M EDTA pH 8.0 stock to 200 mL ddH2O, adjust the pH to 7.5 using HCl. Fill to 250 mL with ddH2O. Filter sterilize. Store at room temperature for up to several months. 2. 1 M lithium acetate: dissolve 25.5 g lithium acetate dehydrate to 250 mL in ddH2O. Filter sterilize. Store at room temperature for up to several months. 3. 1 M DTT: Dissolve 1.545 g DTT in 10 mL. Filter sterilize. For best results, make fresh each time. 4. 1 M sorbitol: Dissolve 45.55 g sorbitol to 250 mL in ddH2O. Filter sterilize or autoclave. Store at 4°C for up to several months.
2.9. Flow Cytometry/ FACS Reagents
1. Phosphate-buffered saline (PBS) containing 0.5% BSA (PBS/0.5% BSA). Prepare a 10× PBS pH 7.4 stock: For 100 mL, dissolve 0.257 g NaH2PO4 H2O, 2.249 g Na2HPO4 7H2O, and 8.765 g NaCl to 100 mL in ddH2O (pH to 7.4 using NaOH or HCl). Filter sterilize. Store at room temperature for up to several months. Add 50 mL of 10× PBS stock to 450 mL ddH2O. Dissolve 2.5 g BSA into this solution. Filter sterilize. Store at 4°C for up to several months. 2. Monoclonal antibodies: anti-HA mouse antibody HA.11 (Covance MMS-101P), anti-c-myc chicken antibody (Molecular Probes cat. no. A-21281), Alexa488-labeled goat-anti-mouse secondary antibody (Invitrogen cat. no. A-11017), Alexa647labeled goat-anti-chicken secondary antibody (Molecular Probes cat. no. A-21449), and streptavidin:phycoerythrin (SA:PE) (BD cat. no. 554061).
2.10. Equipment
1. Thermocycler. 2. Two temperature controlled incubator shakers. 3. Electroporator (Gene pulser, BioRad cat. no. 1652076). 4. Flow cytometer and FACS apparatus.
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3. Methods This section will describe a robust method for engineering a binding protein of interest using yeast surface display technology. Similar methods for gene engineering via yeast display have been reported (9, 31, 32), and slight variations may be used for particular applications to attain optimal results. 3.1. Gene Construction of Yeast Surface Displayed Bioreceptor
To engineer a more stable, higher affinity bioreceptor for biosensing applications, the protein is expressed as a fusion protein to the mating agglutinin protein Aga2 displayed on the surface of yeast. To achieve this, the bioreceptor gene can be inserted into the pCT302 vector using the in-frame NheI restriction site, and a downstream unique BglII or XhoI restriction site (see Fig. 1a) (7, 9). This plasmid allows galactose-inducible overexpression of the Aga2-gene fusion. Since the Aga2 protein is displayed on the surface of yeast linked to the Aga1 protein by a pair of disulfide bonds, a yeast strain containing a stably-integrated Aga1 gene under control of the GAL1 promoter should be used, such as EBY100 (see Note 1). Most often, the bioreceptor of interest is expressed as a single-chain construct fused to Aga2 (see Fig. 1b). For example, the binding domain of a monoclonal antibody, excluding the constant domains, may be conveniently expressed as a single-chain Fv (scFv) comprising the VH and VL genes connected by a ∼16 amino acid long flexible linker. Usually a sequence like (G3S)4 or (G4S)3 will perform well as a flexible linker that is not very proteolytically sensitive. Similarly, a T cell receptor may be expressed as a single-chain construct containing the linked Vα and Vβ chains. The pCT302 vector includes the gene expressing Aga2, followed by the affinity tag HA (sequence: YPYDVPDYA), which can be used as an expression marker. Incorporation of a C-terminal expression tag is also useful to monitor full-length expression of the gene on the yeast surface. We use the c-myc affinity tag (sequence: EQKLISEEDL) at the C-terminus. Upon induction of protein over-expression in the yeast, antibodies against these epitopes (HA and c-myc) can be used to monitor expression of the fusion by flow cytometry. Conventional cloning techniques are used to ligate the wildtype gene of interest into the pCT302 vector using a subcloningcompetent E. coli strain, such as DH10B, DH5α, or XL1-blue. The pCT302 vector confers ampicillin resistance (100 μg/mL) to the bacterial cells. The sequence of fused gene in the vector can be confirmed using primers that flank the insert (see Fig. 1a): “Splice 4/L” (forward: GGC AGC CCC ATA AAC ACA CAG TAT), “YRS” (reverse: CGA GCT AAA AGT ACA GTG GG), or “T7” (reverse: TAA TAC GAC TCA CTA TAG).
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A lac prom
T3 prom
KpnI 7 GAL1 prom
pBR322 origin
EcoRI 828
Yeast surface display gene construct diagram
HincII 978 NheI 1231 Aga-2 – linker – scTCR (bioreceptor)
EcoRI
XhoI
NheI
BglII
Splice 4/L amp marker
Aga-2
NdeI 1783 StuI 1832 AvaI 2003 XhoI 2003 AvrI 2003 BglII 2007
YRS T7 prom CEN6_ARS4
amp prom
scTCR (bioreceptor) HA (G4S)3 linker tag
c-myc STOP tag
SacI 2320 TRP1 marker
lacZ_a reporter f1 origin
AccI 3774 XbaI 3654
B
Single-chain bioreceptor
Yeast Cell c-myc biotinylated ligand Fluorescent streptavidin
HA
AGA-2 S S
S S
5 µM Yeast cell wall
Yeast Proteins (anchors)
AGA-1
approximately 50,000 copies/cell
Fig. 1. Yeast surface display elements: (A) Plasmid map of the yeast surface display vector pCT302 with a single-chain T cell receptor (scTCR) cloned in as a fusion with the yeast mating agglutinin protein Aga-2. Map was prepared with the help of PlasMapper (33 ). The open reading frame containing Aga-2 and the scTCR is expanded to show the important elements and restriction sites. (B) Diagram of a bioreceptor displayed on the surface of yeast fused to Aga-2. A detection scheme involving biotinylated ligand bound to the bioreceptor and fluorescent streptavidin may be used to analyze bioreceptor libraries on the surface of yeast.
3.2. Engineering of Bioreceptor for Improved Stability
An overall flow chart of this step of the protocol is shown in Fig. 2. The method presumes that the protein of interest is not already optimally stable, and thus it is expressed on the surface of yeast at suboptimal levels, or it unfolds irreversibly under conditions of extreme pH or high temperature.
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Clone bioreceptor gene into pCT302
Introduce mutations by PCR (error-prone amplification or directed degeneracies)
Transform library into yeast by homologous recombination
Expand library and induce expression
Stain library using conditions for desired property (folding, stability, affinity for ligand)
Repeat these steps multiple times (2-5) to identify improved mutants
Sort for best yeast binders by FACS
Repeat these steps using your improved clone as a template for the next round of mutation and selection
Isolate and characterize clones
Identification of improved, mutant bioreceptor
Fig. 2. Bioreceptor engineering by yeast surface display.
3.2.1. Introduction of Random Mutations into Bioreceptor Using ErrorProne PCR
The method for random mutagenesis of genes using error-prone polymerase chain reaction (PCR) was adapted from previous reports (34, 35). An error rate of approximately 0.5% should be expected using this protocol. For a single yeast library of random mutants of approximately 105 clones, you should prepare enough reactions to yield 50–80 μg error-prone amplified insert (between 3 and 8 reactions). 1. Prepare a 100 μL solution in a PCR tube for each reaction. The following components should be present in the solution (final concentrations listed): (a) 220 μM dATP (b) 200 μM dCTP (c) 340 μM dGTP (d) 2.4 mM dTTP (e) 0.3 ng/μL template DNA (gene of interest in pCT302 vector) (f) 250 nM Splice 4/L (forward primer) (g) 250 nM T7 (reverse primer)
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(h) 5 ng/μL Bovine Serum Albumin (BSA) (i) 3.325 mM MgCl2 (j) 0.5 mM MnCl2 (k) 1:10 dilution of Taq Polymerase Reaction Buffer (10 μL per 100 μL reaction) (l) 1 μL Taq Polymerase 2. Place PCR tube in thermocycler and run through the following program: (a) 94°C for 3 min (b) 30 cycles of: – 94°C for 1 min – 50°C for 2 min – 72°C for 3 min (c) 72°C for 5 min (d) 4°C forever 3. Check for amplification of the gene on a 1% agarose gel. 4. To reduce insert background from remaining template, digest the PCR reaction with 1 μL DpnI for 1 h at 37°C, which will fragment the template plasmid, but not the PCR product. 5. Clean up the reaction using the PCR Clean-up Kit from Qiagen. Elute each reaction in 30 μL from the Qiagen column. 3.2.2. Preparation of pCT302 Vector
This protocol will result in a linearized, dephosphorylated vector with a sequence overlap with your insert of at least 50 bp. No ligation with the insert will be necessary, as the yeast will recombine the vector and insert through homologous recombination (see Note 1 for tips on choosing an appropriate vector). 1. Starting from 50–80 μg full pCT302 vector obtained from Qiagen mini-prep or a protocol yielding similar purity, digest with NheI in appropriate buffer overnight at 37°C, while reserving a small aliquot (1 μL) uncut for later comparison on a gel. 2. Clean up the reaction after the digestion using the PCR Clean-up Kit from Qiagen. Save another 1 μL sample for gel analysis. 3. Next, digest with XhoI in appropriate buffer for at least 1 h at 37°C. Clean up the reaction using PCR Clean-up Kit, and save another 1 μL sample. 4. Digest next with BglII in appropriate buffer for at least 1 h at 37°C. Digesting with all three enzymes decreases the possibility of remaining full-length vector with the potential to re-circularize. Clean up the reaction using PCR Clean-up Kit, and save another 1 μL sample.
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5. Finally, dephosphorylate the linearized vector using Calf Intestine Phosphatase (CIP) for 1 h at 37°C. This should reduce vector background when transforming your yeast library. Clean up the reaction using PCR Clean-up Kit, and take another 1 μL sample for your gel. 6. Run all of your samples (uncut pCT302, NheI digested pCT302, NheI + XhoI digested pCT302, NheI + XhoI + BglII digested pCT302, and fully cut, dephosphorylated pCT302) on a 1% agarose gel to make sure you have not lost vector at some step. If your vector has disappeared, go back to the step before you lost the plasmid and trouble-shoot that reaction, and try again. 7. Run an additional gel comparing several dilutions of your cut, dephosphorylated vector along with your error-prone insert to determine their absolute and relative concentrations (see Note 2). 3.2.3. Preparation of Electrocompetent Yeast
This protocol has been adapted from previously reported methods (36, 37). 1. Two days before preparation of your library, inoculate 2 or 3, 3 mL cultures of sterile YPD media with a colony of EBY100 cells from a freshly-streaked YPD plate. Grow to stationary phase at 30°C (for ∼48 h), shaking at 220 rpm. If required, these cultures may be prepared further in advance and stored at 4°C. 2. The night before your library transformation, inoculate a 2 L sterile flask containing 500 mL YPD medium with 5–9 mL of starter EBY100 culture. Grow overnight at 30°C with vigorous shaking until the cells reach an OD600 of 1.3–1.5 (see Note 3). 3. Harvest the culture in sterile centrifuge bottles that can hold 500 mL, spinning at 4,000× g, 4°C, and resuspend vigorously in 80 mL sterile ddH2O. 4. To increase electrocompetence: (a) Add 10 mL of sterile, 10× TE buffer, pH 7.5 to the yeast pellet. Swirl to mix. If multiple centrifuge bottles were used, combine the cultures into a single sterile centrifuge bottle. (b) Add 10 mL of 10× (1 M) Lithium Acetate stock solution. Swirl to mix, and shake in the sterile centrifuge bottle gently (100–150 rpm) for 45 min at 30°C. (c) Freshly prepare and then add 2.5 mL of 1 M DTT while swirling. Shake gently 15 min at 30°C. 5. Dilute yeast suspension to 500 mL with sterile water. 6. Wash and concentrate the cells three times by centrifuging at 4,000–6,000 × g, and resuspend the pellets each time as follows:
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(a) First pellet – 250 mL ice-cold water (b) Second pellet – 20–30 mL ice-cold 1 M sorbitol (c) Third pellet – 0.5 mL ice-cold 1 M sorbitol (see Note 4) 7. The final volume of resuspended, electrocompetent yeast should be 1–1.5 mL (see Note 5). 3.2.4. Transformation and Characterization of Randomized Bioreceptor Library
In this method, you will transform the overlapping error-prone amplified insert and vector into the yeast, and allow them to recombine the sequences through homologous recombination. 1. Prepare a mixture of error-prone insert and linearized vector DNA at approximately a 6:1 molar ratio, or roughly a 1:1 mass ratio. The DNA concentration in the mixture should be 10–100 ng in no more than 5 μL (see Note 2). 2. In addition to your error-prone library mixture, set up the following additional controls to transform into the yeast: (a) Linearized vector alone – use the same final concentration as in the DNA mixture. (b) Error-prone insert alone – use the same final concentration as in the DNA mixture. (c) Your unmutated gene in pCT302 – these cells will serve as a control for later experiments. (d) No DNA in the transformation. 3. In a sterile, ice-cold Eppendorf tube, mix 40 μL concentrated, electrocompetent yeast with 10–100 ng of transforming DNA (≤5 μL) (see Note 6). 4. Transfer to an ice-cold, 0.2-cm-gap disposable electroporation cuvette. 5. Pulse at 1.5 kV, 25 μF, 200 Ω. The time constant reported will vary between 4.2 and 4.9 ms. Time constants that are too low (10-fold from fresh yeast. 6. Mixing the DNA with the electrocompetent yeast and allowing them to incubate together on ice for 30–45 min may improve transformation efficiency.
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7. If the conductance is too high to electroporate yeast, make sure you have adequately washed away all free ions in the electrocompetent yeast, and make sure you have prepared the DNA in salt-free ddH2O. Do not use DNA in restriction digest or dephosphorylation reaction buffers for electroporation. 8. To determine library size, count the colonies that grow up on your dilution plates. For each plate, calculate a theoretical library size by multiplying the number of colonies on the plate by the dilution factor and the total volume of resuspended library in μL divided the number of μL on the plate. For example, an error-prone library of 1 × 107 clones where the final resuspended sorbitol volume was 30 mL would yield an expected 3333, 33, and 6 colonies on your 10 μL, 10 μL at 1:100 dilution, and 10 μL at 1:500 dilution plates, respectively. If you do not get the expected numbers of colonies on your plates after dilution, it is possible that you have not adequately mixed the sorbitol-dissolved library or the diluted culture before plating. 9. If your bioreceptor cannot be probed with biotinylated ligand at its wild-type affinity and there is not a conformation-specific antibody to the protein, then the level of c-myc staining may be used as a proxy for native expression. Although some full-length protein containing the c-myc tag may be expressed on yeast even if the protein is unfolded or partially folded (45), in some cases properly folded clones may be expressed at higher levels on the yeast surface (38–40, 42). If using this technique to isolate stable mutants, it is important to frequently isolate plasmid from the yeast and analyze the sequence to make sure you did not yield sequence artifacts during homologous recombination that give high c-myc staining for reasons other than stable, native folding. 10. Optimal induction temperatures and durations should be determined for each bioreceptor. We have found that 20°C is consistently the optimal induction temperature, and the optimal induction duration ranged from 24 to 72 h, depending on the polypeptide. 11. The resuspension volume for flow cytometry will vary on the basis of the number of cells in the sample and the instrument on which you will perform the flow. For analysis, usually a volume of 400–500 μL will give an acceptable sampling rate if 0.5–1.0 × 106 cells were included per sample. For samples that will be sorted, it may be advantageous to resuspend a sample of 1 × 108 cells to a larger volume of 5–10 mL to allow for slower sampling and more careful separation. 12. When designing your flow cytometry samples, take into consideration how many fluorescent labels will appear in a single sample. If the fluorescent probes included in a single sample have some overlap in excitation and/or emission
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spectra, such as fluorescein and phycoerythrin, it is important to include valid single-stained control samples to adequately compensate the data. In addition, take care not to use multiple primary antibodies that can be detected by the same secondary reagent, such as two different mouse antibodies in the same sample if you plan to detect even one of them with a polyclonal anti-mouse secondary reagent. 13. When planning to select for higher thermostability in the bioreceptor displayed on the surface of yeast, it is important to check the percent of viability of your cells after the incubation conditions. This may be carried out by plating dilutions of cells onto SD-citrate-CAA plates after incubation at the temperatures you will use to select mutants, compared with those kept at 4°C. If a significant fraction of cells are no longer viable after the treatment (yeast cells appear to begin to lose significant viability at about 50°C), increase the initial sample size to ensure you will still sufficiently sample your library with cells that can recover and expand. 14. One general approach for mutagenesis is to direct mutations to certain region(s) of the bioreceptor, usually those regions known or believed to contact ligand. These sitedirected mutations are encoded by primers that anneal to the template and contain randomized codons at specified positions. These primers are used in a PCR-based amplification method, “splicing by overlap extension” (SOE) (43). Issues of theoretical diversity and maximum library size arise when considering how many codons to randomize. The degenerate codon NNS, where N represents any nucleotide A/T/G/C and S represents G or C, will cover all amino acids (eliminating two of three stop codons) with a theoretical diversity of 32 (4 × 4 × 2). Therefore, a library of yeast randomized at one codon using an NNS degenerate primer need only contain 32 mutants, theoretically, to fully sample all possible mutations. Yeast transformation efficiencies are such that 107–108 independent transformants can be obtained, providing sufficient library sizes to comprehensively sample all possible codon combinations in a 5-codon library (325). Other techniques that have been used include DNA shuffling (46) and “look-through” mutagenesis (47). 15. Conditions that affect the robustness of the SOE PCR include: annealing temperature, mole ratio of pre-SOE products, and competing undesired amplification. If the SOE product is in low abundance or appears contaminated by unwanted product, try a gradient of annealing temperatures in the SOE PCR program to optimize that parameter. Also, adjusting the ratio of pre-SOE product 1 to pre-SOE product 2 in the SOE reaction mixture may yield improved
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results. DpnI digestion of the pre-SOE reactions (which will fragment template DNA) prior to the SOE reaction (subjecting to a QIAgen PCR clean-up kit between digest and SOE) may also improve the PCR efficiency. 16. Staining conditions that yield maximum increase in fluorescence of a high-affinity mutant over the template/wild-type will provide optimal sorting results. A quantitative theoretical analysis of equilibrium labeling conditions of yeast display populations indicated the ligand concentration yielding maximized fluorescence discrimination on the first sort was equal to ∼ 0.05–0.1× the KD of the template or “wild-type” interaction (44). The optimal staining volume, in any equilibrium staining procedure, depends on the numbers of bioreceptors on the yeast surface and the KD of the bioreceptor/ligand interaction (and consequently the concentration of ligand in the staining mixture). The staining volume must be large enough to ensure that the number of bioreceptors does not exceed the moles of ligand; otherwise, the depletion of ligand from solution upon binding bioreceptor would have a nonnegligible effect on free ligand concentration and thus staining. In cases where the template affinity is lower, 500 μL volume is likely sufficient. However, when the template affinity is already quite high (KD < 10 nM) and the concentration of ligand used for staining is therefore low, a larger staining volume must be used so the moles of ligand at least equalize (but ideally exceed) the number of yeast surface displayed bioreceptors (assume ∼5 × 105 bioreceptors per yeast cell) (7). If the kon and koff are known, the incubation time that allows approach to equilibrium can also be determined mathematically (31, 44). 17. Although not the focus of this paper, an engineered high-affinity bioreceptor with broadened specificity (e.g., for multiple toxin subtypes) may be a desired feature of a biosensor. See GarciaRodriguez et al. (48) for yeast display engineering of a scFv for high-affinity to two subtypes of botulinum neurotoxin.
Acknowledgments The authors thank members of our laboratory and K. Dane Wittrup and members of his lab for working out many of the conditions and methods described in this chapter. This work was supported by NIH grants AI064611 and GM55767 (to DMK), the Samuel and Ruth Engelberg/Irvington Institute postdoctoral fellowship from the Cancer Research Institute (to JDS), and predoctoral fellowship ES013571 (to SAR).
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References 1. Luppa, P. B., Sokoll, L. J. and Chan, D. W. (2001) Immunosensors – principles and applications to clinical chemistry. Clin Chim Acta 314, 1–26 2. Pejcic, B., De Marco, R. and Parkinson, G. (2006) The role of biosensors in the detection of emerging infectious diseases. Analyst 131, 1079–1090 3. Stefan, R. I., van Staden, J. F. and AboulEnein, H. Y. (2000) Immunosensors in clinical analysis. Fresenius J Anal Chem 366, 659–668 4. Warsinke, A., Benkert, A. and Scheller, F. W. (2000) Electrochemical immunoassays. Fresenius J Anal Chem 366, 622–634 5. Peruski, A. H. and Peruski, L. F., Jr. (2003) Immunological methods for detection and identification of infectious disease and biological warfare agents. Clin Diagn Lab Immunol 10, 506–513 6. Goodchild, S., Love, T., Hopkins, N. and Mayers, C. (2006) Engineering antibodies for biosensor technologies. Adv Appl Microbiol 58, 185–226 7. Boder, E. T. and Wittrup, K. D. (1997) Yeast surface display for screening combinatorial polypeptide libraries. Nat Biotechnol 15, 553–557 8. Kondo, A. and Ueda, M. (2004) Yeast cellsurface display – applications of molecular display. Appl Microbiol Biotechnol 64, 28–40 9. Boder, E. T. and Wittrup, K. D. (2000) Yeast surface display for directed evolution of protein expression, affinity, and stability. Methods Enzymol 328, 430–444 10. Feldhaus, M. J., Siegel, R. W., Opresko, L. K., Coleman, J. R., Feldhaus, J. M., Yeung, Y. A., Cochran, J. R., Heinzelman, P., Colby, D., Swers, J., Graff, C., Wiley, H. S. and Wittrup, K. D. (2003) Flow-cytometric isolation of human antibodies from a nonimmune Saccharomyces cerevisiae surface display library. Nat Biotechnol 21, 163–170 11. Yeung, Y. A. and Wittrup, K. D. (2002) Quantitative screening of yeast surface-displayed polypeptide libraries by magnetic bead capture. Biotechnol Prog 18, 212–220 12. Richman, S. A., Healan, S. J., Weber, K. S., Donermeyer, D. L., Dossett, M. L., Greenberg, P. D., Allen, P. M. and Kranz, D. M. (2006) Development of a novel strategy for engineering high-affinity proteins by yeast display. Protein Eng Des Sel 19, 255–264 13. Wang, X. X. and Shusta, E. V. (2005) The use of scFv-displaying yeast in mammalian
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23.
24.
cell surface selections. J Immunol Methods 304, 30–42 Boder, E. T., Midelfort, K. S. and Wittrup, K. D. (2000) Directed evolution of antibody fragments with monovalent femtomolar antigen-binding affinity. Proc Natl Acad Sci U S A 97, 10701–10705 Kieke, M. C., Cho, B. K., Boder, E. T., Kranz, D. M. and Wittrup, K. D. (1997) Isolation of anti-T cell receptor scFv mutants by yeast surface display. Protein Eng 10, 1303–1310 Feldhaus, M. J. and Siegel, R. W. (2004) Yeast display of antibody fragments: a discovery and characterization platform. J Immunol Methods 290, 69–80 Kieke, M. C., Shusta, E. V., Boder, E. T., Teyton, L., Wittrup, K. D. and Kranz, D. M. (1999) Selection of functional T cell receptor mutants from a yeast surface-display library. Proc Natl Acad Sci U S A 96, 5651–5656 Richman, S. A. and Kranz, D. M. (2007) Display, engineering, and applications of antigen-specific T cell receptors. Biomol Eng 24, 361–373 Weber, K. S., Donermeyer, D. L., Allen, P. M. and Kranz, D. M. (2005) Class IIrestricted T cell receptor engineered in vitro for higher affinity retains peptide specificity and function. Proc Natl Acad Sci U S A 102, 19033–19038 Jones, L. L., Brophy, S. E., Bankovich, A. J., Colf, L. A., Hanick, N. A., Garcia, K. C. and Kranz, D. M. (2006) Engineering and characterization of a stabilized alpha1/alpha2 module of the class I major histocompatibility complex product Ld. J Biol Chem 281, 25734–25744 Starwalt, S. E., Masteller, E. L., Bluestone, J. A. and Kranz, D. M. (2003) Directed evolution of a single-chain class II MHC product by yeast display. Protein Eng 16, 147–156 Esteban, O. and Zhao, H. (2004) Directed evolution of soluble single-chain human class II MHC molecules. J Mol Biol 340, 81–95 Boder, E. T., Bill, J. R., Nields, A. W., Marrack, P. C. and Kappler, J. W. (2005) Yeast surface display of a noncovalent MHC class II heterodimer complexed with antigenic peptide. Biotechnol Bioeng 92, 485–491 Dam, J., Guan, R., Natarajan, K., Dimasi, N., Chlewicki, L. K., Kranz, D. M., Schuck, P., Margulies, D. H. and Mariuzza, R. A. (2003) Variable MHC class I engagement by Ly49 natural killer cell receptors demonstrated by the crystal structure of Ly49C bound to H-2K(b). Nat Immunol 4, 1213–1222
Engineering Stable, High-Affinity Bioreceptors by Yeast Surface Display 25. Cochran, J. R., Kim, Y. S., Lippow, S. M., Rao, B. and Wittrup, K. D. (2006) Improved mutants from directed evolution are biased to orthologous substitutions. Protein Eng Des Sel 19, 245–253 26. Buonpane, R. A., Churchill, H. R. O., Moza, B., Sundberg, E. J., Peterson, M. L., Schlievert, P. M., and Kranz, D. M. (2007) Neutralization of Staphylococcal enterotoxin B by soluble, high-affinity receptor antagonists. Nat Med 13(6), 725–729. 27. Koide, A., Gilbreth, R. N., Esaki, K., Tereshko, V. and Koide, S. (2007) High-affinity single-domain binding proteins with a binarycode interface. Proc Natl Acad Sci U S A 104, 6632–6637 28. Lipovsek, D., Lippow, S. M., Hackel, B. J., Gregson, M. W., Cheng, P., Kapila, A. and Wittrup, K. D. (2007) Evolution of an interloop disulfide bond in high-affinity antibody mimics based on fibronectin type III domain and selected by yeast surface display: molecular convergence with single-domain camelid and shark antibodies. J Mol Biol 368, 1024– 1041 29. Rao, B. M., Driver, I., Lauffenburger, D. A. and Wittrup, K. D. (2004) Interleukin 2 (IL-2) variants engineered for increased IL-2 receptor alpha-subunit affinity exhibit increased potency arising from a cell surface ligand reservoir effect. Mol Pharmacol 66, 864–869 30. Rao, B. M., Girvin, A. T., Ciardelli, T., Lauffenburger, D. A. and Wittrup, K. D. (2003) Interleukin-2 mutants with enhanced alpha-receptor subunit binding affinity. Protein Eng 16, 1081–1087 31. Chao, G., Lau, W. L., Hackel, B. J., Sazinsky, S. L., Lippow, S. M. and Wittrup, K. D. (2006) Isolating and engineering human antibodies using yeast surface display. Nat Protoc 1, 755–768 32. Colby, D. W., Kellogg, B. A., Graff, C. P., Yeung, Y. A., Swers, J. S. and Wittrup, K. D. (2004) Engineering antibody affinity by yeast surface display. Methods Enzymol 388, 348–358 33. Dong, X., Stothard, P., Forsythe, I. J. and Wishart, D. S. (2004) PlasMapper: a web server for drawing and auto-annotating plasmid maps. Nucleic Acids Res 32, W660–W664 34. Daugherty, P. S., Chen, G., Iverson, B. L. and Georgiou, G. (2000) Quantitative analysis of the effect of the mutation frequency on the affinity maturation of single chain Fv antibodies. Proc Natl Acad Sci U S A 97, 2029–2034
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35. Fromant, M., Blanquet, S. and Plateau, P. (1995) Direct random mutagenesis of genesized DNA fragments using polymerase chain reaction. Anal Biochem 224, 347–353 36. Meilhoc, E., Masson, J. M. and Teissie, J. (1990) High efficiency transformation of intact yeast cells by electric field pulses. Biotechnology (N Y) 8, 223–227 37. Becker, D. M. and Lundblad, V. (1996) Introduction of DNA into yeast cells, transformation by electroporation. In: Current Protocols in Molecular Biology (Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A. and Struhl, K., eds.). John Wiley & Sons, Hoboken, NJ 38. Shusta, E. V., Holler, P. D., Kieke, M. C., Kranz, D. M. and Wittrup, K. D. (2000) Directed evolution of a stable scaffold for T-cell receptor engineering. Nat Biotechnol 18, 754–759 39. Shusta, E. V., Kieke, M. C., Parke, E., Kranz, D. M. and Wittrup, K. D. (1999) Yeast polypeptide fusion surface display levels predict thermal stability and soluble secretion efficiency. J Mol Biol 292, 949–956 40. Kowalski, J. M., Parekh, R. N., Mao, J. and Wittrup, K. D. (1998) Protein folding stability can determine the efficiency of escape from endoplasmic reticulum quality control. J Biol Chem 273, 19453–19458 41. Kowalski, J. M., Parekh, R. N. and Wittrup, K. D. (1998) Secretion efficiency in Saccharomyces cerevisiae of bovine pancreatic trypsin inhibitor mutants lacking disulfide bonds is correlated with thermodynamic stability. Biochemistry 37, 1264–1273 42. Hagihara, Y. and Kim, P. S. (2002) Toward development of a screen to identify randomly encoded, foldable sequences. Proc Natl Acad Sci U S A 99, 6619–6624 43. Warrens, A. N., Jones, M. D. and Lechler, R. I. (1997) Splicing by overlap extension by PCR using asymmetric amplification: an improved technique for the generation of hybrid proteins of immunological interest. Gene 186, 29–35 44. Boder, E. T. and Wittrup, K. D. (1998) Optimal screening of surface-displayed polypeptide libraries. Biotechnol Prog 14, 55–62 45. Park, S., Xu, Y., Stowell, X. F., Gai, F., Saven, J. G. and Boder, E. T. (2006) Limitations of yeast surface display in engineering proteins of high thermostability. Protein Eng Des Sel 19, 211–217 46. Stemmer, W. P. (1994) Rapid evolution of a protein in vitro by DNA shuffling. Nature 370, 389–391
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47. Rajpal, A., Beyaz, N., Haber, L., Cappuccilli, G., Yee, H., Bhatt, R. R., Takeuchi, T., Lerner, R. A. and Crea, R. (2005) A general method for greatly improving the affinity of antibodies by using combinatorial libraries. Proc Natl Acad Sci U S A 102, 8466–8471
48. Garcia-Rodriguez, C., Levy, R., Arndt, J. W., Forsyth, C. M., Razai, A., Lou, J., Geren, I., Stevens, R. C. and Marks, J. D. (2007) Molecular evolution of antibody cross-reactivity for two subtypes of type A botulinum neurotoxin. Nat Biotechnol 25, 107–116
Chapter 20 Antibody Affinity Optimization Using Yeast Cell Surface Display Robert W. Siegel Summary Many biosensors depend on molecular recognition reagents to achieve highly sensitive and specific detection levels of an analyte of interest. Although new and improved detection platforms continue to be developed, improvements in the affinity and specificity of the molecular recognition reagents often dictate the ultimate performance level and utility of the instrument. Accordingly, large effort is placed in discovering and characterizing the reagents to be used for a biosensor application. Antibodies, owing to their unparalleled ability to bind a diverse array of antigens with high affinity and specificity, have been widely used as molecular recognition reagents in the biosensor field. The recent advent of recombinant in vitro antibody display technologies, in general, and yeast surface display, in particular, allow specific traits of a given antibody to be discreetly augmented to enhance biosensor performance. Large variegated libraries derived from existing antibodies already employed in a particular biosensor can be created and screened for mutations that confer a desired improved phenotype leading to enhanced biosensor performance. This chapter will provide a protocol for the affinity maturation of a previously isolated monoclonal antibody, the most widely used application of in vitro directed evolution. Key words: scFv, Recombinant antibody, Directed evolution, Affinity maturation, Yeast display, Mutagenesis, FACS.
1. Introduction Antibodies, owing to their unparalleled ability to bind a diverse array of antigens with high affinity and specificity, are essential recognition reagents for numerous biosensor platforms. Most diagnostic immunoassays rely upon monoclonal or polyclonal antibodies derived from animals, and this source of recognition
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_20
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reagents has served the research community for over three decades. However, one limitation of this approach is the inability to change a specific property of an isolated antibody while leaving others intact. Instead, the only recourse is to initiate another round of immunization with no guarantee of discovering a preferred alternative. The recent advent of in vitro display technologies provides the added flexibility to engineer new or augment specific attributes of interest, such as affinity, specificity, or stability, into previously isolated and otherwise desirable recognition reagents. A number of different in vitro display systems have been developed including, filamentous phage (1, 2), ribosomal (3), bacterial (4), and yeast (5). Each have their own strengths and weaknesses and is outside the scope of this work but has been recently reviewed (6). However, a central tenet of any in vitro display platform is that the phenotype of the displayed protein must be coupled to the gene encoding the displayed protein. This coupling allows diverse libraries to be sampled and the coding sequence recovered from clones identified with desired properties. Finally, selected products can be expressed in a variety of formats from intact immunoglobulin (Ig), F(ab)′2, Fab, single-chain variable fragment (scFv), to single variable domains depending on the preferred embodiment for application. Yeast display (Fig. 1) has proven to be highly effective platform for various directed evolution applications, including affinity maturation (7–9) and changes in specificity (10, 11). Recombinant antibodies are displayed on the yeast surface as a fusion protein to a cell wall component, Aga2 (5) and library generation can be facilitated by the homologous recombination system inherent in yeast (12, 13). Coupling fluorescence activated cell sorting (FACS) with cell surface display enables monitoring of both antibody expression on the cell surface and the ability of that antibody to bind antigen (Fig. 1b). The unparalleled resolution offered by FACS during the selection process enables the visualization of discrete populations that differ in affinity and/or epitope specificity and allows the quantitative recovery of desired clones (14, 15). FACS analysis also facilitates the characterization of isolated clones directly on the yeast surface eliminating the need for purified antibody. A typical affinity maturation project using yeast cell surface display is divided into several steps. First, the variable (VH and VL) domains of the parent hybridoma clone are identified. Second, the VH and VL genes are cloned into a display vector for expression as a scFv fragment on the cell surface. Third, the binding characteristics of the WT scFv are determined. Fourth, diverse variegated populations for selection are created by mutagenesis of the parent clone plasmid DNA. Fifth, clones with improved affinities are enriched from the mutant library after application of selective pressure. Sixth, selected clones are sequenced and
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Fig. 1. Yeast surface display. (A) The scFv is displayed on yeast cell surface as a translational fusion to Aga2 protein. The VH and VL domains are joined by a flexible peptide (Gly4Ser)3 linker. Surface expression can be detected using fluorescent labeled antibodies binding to the C-terminal epitope tag. Biotinylated antigen binding can be detected using fluorescent streptavidin/ avidin. Expression of the translational fusion in transformed yeast from the pYD1 plasmid is under control of the inducible Gal promoter. (B) Dual color flow cytometric analysis. ScFv expression is shown on X-axis and antigen-binding on the Y-axis. Top plot is no antigen control. Bottom plot is plus antigen control. Uninduced cells are located in the bottom left quadrant. Only cells expressing scFv are able to bind antigen and move along the Y-axis.
binding characteristics characterized. Finally, the VH and VL genes are recovered for cloning into desired expression system. As with any technical approach, many variations exist and may be of benefit for a particular application. This review intends to give the reader a rudimentary protocol that can be augmented and expanded with experience.
2. Materials 2.1. Variable Gene Isolation 2.1.1. mRNA Isolation
1. Oligotex Direct mRNA Minikit (Qiagen cat # 72022) supplied with OL1, ODB, Oligotex beads, OW1, OW2 and OEB buffers. 2. Prepare fresh OL1-BME buffer by adding 30 μL BME (Bio-Rad cat # 161-0710) to 1 mL OL1 buffer. 3. Culture of hybridoma cells producing antibody of interest.
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2.1.2. Immunoglobulin cDNA Synthesis
1. cDNA primers supplied with Mouse Ig-Primer set (Novagen cat # 69831-3). MuIgMVH3′-1 (5′-CCCAAGCTTACGAGGGGGAAGACATTTGGGAA-3′). MuIgGVH3′-2 (5′-CC CAAGCTTCCAGGGRCCARKGGATARACIGRTGG-3′), MuIgκVL3′-1 (5′-CCCAAGCTTACTGGATGGTGGGAAGATGGA-3′), MuIgλVL3′-1 (5′-CCCAAGCTTAGCTCYTCWGWGGAIGGYGGRAA-3′). 2. SuperScript III Reverse transcriptase (Invitrogen cat # 18080044) supplied with 5× First Strand Buffer and 0.1 M DTT. 3. 10 mM dNTP mix (Invitrogen cat # 18427-013).
2.1.3. VH and VL Amplification
1. Mouse Ig-Primer set (Novagen cat # 69831-3). 2. Platinum HiFi Taq polymerase (Invitrogen cat # 11304-011) supplied with 10× HiFi buffer and 50 mM MgSO4. 3. 10 mM dNTP mix (Invitrogen cat # 18427-013). 4. PureLink PCR purification kit (Invitrogen cat # K3100-02). 5. 1% ReadyAgarose minigel with ethidium bromide in 1× TBE buffer (Bio-Rad Cat # 161-3004). 6. 1× TBE running buffer: Add 100 mL of 10× TBE buffer (Bio-Rad cat # 161-0733) to 900 mL dH2O. Store at room temperature. 7. 10× loading buffer (Invitrogen cat # 10816-015).
2.1.4. VH and VL Gene Cloning and Identification
1. TOPO-TA cloning kit (Ivitrogen cat # 455-0641) supplied with pCR2.1 TOPO vector, salt solution, and OneShot TOP10 competent E. coli aliquots. 2. Xgal solution: Prepare 40 mg/mL solution of Xgal by suspending 500 mg of Xgal (Sigma cat # B9146-500) with 12.5 mL DMF (Sigma cat # D4551-250). 3. 1,000× Ampicillin stock: Prepare 100 mg/mL solution by dissolving 1 g of ampicillin (Sigma cat # A0797) with 10 mL deionized water and sterilize by filtration. Store at −20°C. 4. LB media: Dissolve 10 g tryptone, 5 g yeast extract, and 10 g NaCl in deionized water to a volume of 1 L and sterilize by autoclaving. 5. LB-Amp media: Dissolve 10 g tryptone, 5 g yeast extract, and 10 g NaCl in deionized water to a volume of 1 L and sterilize by autoclaving. Add 1 mL of 1,000× (100 mg/mL) ampicillin stock to warm media (12 h) are used. 10. Mutant libraries can be constructed by a variety of means. The simplest format has been described in this protocol. Other approaches include the use of nucleoside analogues (21) or commercially available kits (Stratagene GeneMorphII cat # 200550) that allows the mutation frequency to be tuned without bias to the identity of the mutation. Also, mutagenic DNA can always be amplified under normal conditions to increase the amount of mutagenic product. 11. The amplified mutagenic DNA can be gel purified to remove the original template from mutagenized PCR product. Practical experience has shown contamination of WT scFv pYD1 plasmid has not been problematic due to minimal amount of intact plasmid and relatively low transformation frequency of yeast. However, gel purification should be used if PCR does not generate a discrete product band of appropriate size or one wants to eliminate all WT plasmid. 12. Extensive information about yeast lithium acetate transformation is available. Refer to reference (22) or http://www.umanitoba.ca/faculties/medicine/biochem/gietz/Trafo.html. 13. Refer to (14, 16) for extensive discussion and application of optimal kinetic selection strategies. As an alternative selection strategy, an equilibrium selection in which the concentration of antigen is reduced (generally 1/10th – 1/20th) relative to the WT KD can be utilized (see ref. (23) for detailed protocol including alternative mutagenesis strategy). However, most WT scFv derived from existing hybridomas will have
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affinities in the single to subnanomolar range, which makes this approach more difficult requiring extremely large reaction volumes (see above mentioned references). 14. The mutant scFv gene can also be directly sequenced after amplification from the yeast colony. Simply transfer a portion of the colony (or cell pellet from an aliquot of liquid culture) into 20 μL of 0.1% SDS and boil for 5 min. Pellet lysate and use 1–2 μL of supernatant as template for PCR amplification reaction as described in Subheading 3.2.1 using pYD1 for and rev primers. Remove primers and unincorporated nucleotides from PCR using Exo-SAP (USB cat # 78200) following manufacturer’s directions and use as template for sequencing reaction. However, this approach is not the most robust and one will want to eventually recover the plasmid with the mutant scFv as described in Subheading 3.6.1. 15. The entire Aga2-scFv translation coding sequence is shown below. Pertinent features are highlighted in the sequence. Representative VH and VL genes are from GenBank accession number AF003725. The third position of the penultimate lysine codon in the VL gene included in the VL rev primer is denoted with a box on the antisense DNA strand.
Aga2 ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ MetGlnLeu LeuArgCys PheSerIle PheSerVal IleAlaSer ValLeuAla GlnGluLeu ThrThrIle CysGluGln IleProSer ATGCAGTTA CTTCGCTGT TTTTCAATA TTTTCTGTT ATTGCTTCA GTTTTAGCA CAGGAACTG ACAACTATA TGCGAGCAA ATCCCCTCA TACGTCAAT GAAGCGACA AAAAGTTAT AAAAGACAA TAACGAAGT CAAAATCGT GTCCTTGAC TGTTGATAT ACGCTCGTT TAGGGGAGT Aga2 ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ProThrLeu GluSerThr ProTyrSer LeuSerThr ThrThrIle LeuAlaAsn GlyLysAla MetGlnGly ValPheGlu TyrTyrLys CCAACTTTA GAATCGACG CCGTACTCT TTGTCAACG ACTACTATT TTGGCCAAC GGGAAGGCA ATGCAAGGA GTTTTTGAA TATTACAAA GGTTGAAAT CTTAGCTGC GGCATGAGA AACAGTTGC TGATGATAA AACCGGTTG CCCTTCCGT TACGTTCCT CAAAAACTT ATAATGTTT Aga2 ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ pYD1 forward primer ~~~~~~~~~~~~~~~~~~~~~~~~ SerValThr PheValSer AsnCysGly SerHisPro SerThrThr SerLysGly SerProIle AsnThrGln TyrValPhe LysLeuLeu TCAGTAACG TTTGTCAGT AATTGCGGT TCTCACCCC TCAACAACT AGCAAAGGC AGCCCCATA AACACACAG TATGTTTTT AAGCTTCTG AGTCATTGC AAACAGTCA TTAACGCCA AGAGTGGGG AGTTGTTGA TCGTTTCCG TCGGGGTAT TTGTGTGTC ATACAAAAA TTCGAAGAC Linker ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ VH for ~~~~~~~~~~~~~~~~~~~~ VH gene NheI ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ~~~~~~ GlnAlaSer GlyGlyGly GlySerGly GlyGlyGly SerGlyGly GlyGlySer AlaSerGln ValLysLeu GlnGlnSer GlyAlaGlu CAGGCTAGT GGTGGTGGT GGTTCTGGT GGTGGTGGT TCTGGTGGT GGTGGTTCT GCTAGCCAG GTGAAGCTG CAGCAGTCA GGGGCTGAG GTCCGATCA CCACCACCA CCAAGACCA CCACCACCA AGACCACCA CCACCAAGA CGATCGGTC CACTTCGAC GTCGTCAGT CCCCGACTC VH gene ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ LeuValArg ProGlyAla SerValLys LeuSerCys LysAlaSer GlyTyrSer LeuThrSer TyrTrpMet AsnTrpVal LysGlnArg CTGGTGAGG CCTGGAGCT TCAGTGAAG CTGTCCTGC AAGGCTTCT GGCTACTCC CTCACCAGC TACTGGATG AACTGGGTG AAGCAGAGG GACCACTCC GGACCTCGA AGTCACTTC GACAGGACG TTCCGAAGA CCGATGAGG GAGTGGTCG ATGACCTAC TTGACCCAC TTCGTCTCC VH gene ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ProGlyGln GlyLeuGlu TrpIleGly MetIleHis ProSerAsp SerAspThr ArgPheAsn GlnLysPhe GluAspLys AlaThrLeu CCTGGACAA GGCCTTGAG TGGATTGGC ATGATTCAT CCTTCCGAT AGTGACACT AGGTTCAAT CAGAAGTTC GAGGACAAG GCCACATTG GGACCTGTT CCGGAACTC ACCTAACCG TACTAAGTA GGAAGGCTA TCACTGTGA TCCAAGTTA GTCTTCAAG CTCCTGTTC CGGTGTAAC
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VH gene ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ThrValAsp ThrSerSer SerThrAla TyrMetGln LeuSerSer ProThrSer GluAspSer AlaValTyr TyrCysAla ArgGlyLeu ACTGTTGAC ACATCCTCC AGCACAGCC TACATGCAA CTCAGCAGC CCGACATCT GAGGATTCT GCGGTCTAT TACTGTGCA AGAGGGCTC TGACAACTG TGTAGGAGG TCGTGTCGG ATGTACGTT GAGTCGTCG GGCTGTAGA CTCCTAAGA CGCCAGATA ATGACACGT TCTCCCGAG ScFv linker ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ VH gene VL for ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ~~~~~~~~~~~~~~~ TyrAsnGly PheTrpTyr PheAspVal TrpGlyGln GlyThrThr ValThrVal SerSerGly IleLeuGly SerGlyGly GlyGlySer TACAATGGT TTCTGGTAC TTCGATGTC TGGGGCCAA GGGACCACG GTCACCGTC TCCTCAGGA ATTCTAGGA TCCGGTGGC GGTGGCAGC ATGTTACCA AAGACCATG AAGCTACAG ACCCCGGTT CCCTGGTGC CAGTGGCAG AGGAGTCCT TAAGATCCT AGGCCACCG CCACCGTCG ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ VH rev GlySer linker ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ VL gene VL for ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ GlyGlyGly GlySerGly GlyGlyGly SerAspIle GluLeuThr GlnSerPro AlaLeuMet AlaAlaSer ProGlyGlu LysValIle GGCGGTGGT GGTTCCGGA GGCGGCGGT AGCGACATC GAGCTCACT CAGTCTCCA GCACTCATG GCTGCATCT CCAGGGGAG AAGGTCATC CCGCCACCA CCAAGGCCT CCGCCGCCA TCGCTGTAG CTCGAGTGA GTCAGAGGT CGTGAGTAC CGACGTAGA GGTCCCCTC TTCCAGTAG ~~~~~~~~~~~~~~~ VH rev VL gene ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ IleThrCys SerValSer SerSerIle SerSerSer AsnLeuHis TrpTyrGln GlnLysSer GlyThrSer ProLysPro TrpIleTyr ATCACCTGC AGTGTCAGC TCAAGTATA AGTTCCAGC AACTTGCAC TGGTACCAG CAGAAGTCA GGAACCTCC CCCAAACCC TGGATTTAT TAGTGGACG TCACAGTCG AGTTCATAT TCAAGGTCG TTGAACGTG ACCATGGTC GTCTTCAGT CCTTGGAGG GGGTTTGGG ACCTAAATA VL gene ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ GlyThrSer AsnLeuAla SerGlyVal ProValArg PheSerGly SerGlySer GlyThrSer TyrSerLeu ThrIleSer SerMetGlu GGCACATCC AACCTGGCT TCTGGAGTC CCTGTTCGC TTCAGTGGC AGTGGATCT GGGACCTCT TATTCTCTC ACAATCAGC AGCATGGAG CCGTGTAGG TTGGACCGA AGACCTCAG GGACAAGCG AAGTCACCG TCACCTAGA CCCTGGAGA ATAAGAGAG TGTTAGTCG TCGTACCTC VL gene ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ NotI ~~~~~~~ AlaGluAsp AlaAlaThr TyrTyrCys GlnGlnTrp SerSerTyr ProLeuThr PheGlyAla GlyThrLys LeuGluIle LysArgPro GCTGAAGAT GCTGCCACT TATTACTGT CAACAGTGG AGTAGTTAC CCGCTCACG TTCGGTGCT GGCACCAAG CTGGAAATC AAGCGGCCG CGACTTCTA CGACGGTGA ATAATGACA GTTGTCACC TCATCAATG GGCGAGTGC AAGCCACGA CCGTGGTTC GACCTTTAG TTCGCCGGC ~~~~~~~ VL rev NotI V5 tag 6xHis ~ ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ~~~~~~~~~~~~~~~~ LeuGluSer ArgGlyPro PheGluGly LysProIle ProAsnPro LeuLeuGly LeuAspSer ThrArgThr GlyHisHis HisHisHis CTCGAGTCT AGAGGGCCC TTCGAAGGT AAGCCTATC CCTAACCCT CTCCTCGGT CTCGATTCT ACGCGTACC GGTCATCAT CACCATCAC GAGCTCAGA TCTCCCGGG AAGCTTCCA TTCGGATAG GGATTGGGA GAGGAGCCA GAGCTAAGA TGCGCATGG CCAGTAGTA GTGGTAGTG ~~~~~~~~~~ VL rev His*** CATTGAGTT TAAACCCGC TGATCTGAT AACAACAGT GTAGATGTA ACAAAATCG AC GTAACTCAA ATTTGGGCG ACTAGACTA TTGTTGTCA CATCTACAT TGTTTTAGC TG ~~~~~~~~~~~~~~~~~~~~~
References 1. Parmley, S. F., and Smith, G. P. (1988) Antibody-selectable filamentous fd phage vectors: affinity purification of target genes. Gene 73, 305–18 2. McCafferty, J., Griffiths, A. D., Winter, G., and Chiswell, D. J. (1990) Phage antibodies: filamentous phage displaying antibody variable domains. Nature 348, 552–4 3. Hanes, J., and Plückthun, A. (1997) In vitro selection and evolution of functional proteins by using ribosome display. Proc Natl Acad Sci U S A 94, 4937–42 4. Georgiou, G., Stathopoulos, C., Daugherty, P. S., Nayak, A. R., Iverson, B. L., and Curtiss 3rd, R. (1997) Display of heterologous proteins on
the surface of microorganisms: from the screening of combinatorial libraries to live recombinant vaccines. Nat Biotechnol 15, 29–34 5. Boder, E. T., and Wittrup, K. D. (1997) Yeast surface display for screening combinatorial polypeptide libraries. Nat Biotechnol 15, 553–7 6. Levin, A. M., and Weiss, G. A. (2006) Optimizing the affinity and specificity of proteins with molecular display. Mol Biosyst 2, 49–57 7. Boder, E. T., Midelfort, K. S., and Wittrup, K. D. (2000) Directed evolution of antibody fragments with monovalent femtomolar antigen-binding affinity. Proc Natl Acad Sci U S A 97, 10701–5
Antibody Affinity Optimization Using Yeast Cell Surface Display 8. Graff, C. P., Chester, K., Begent, R., and Wittrup, K. D. (2004) Directed evolution of an anti-carcinoembryonic antigen scFv with a 4-day monovalent dissociation halftime at 37 degrees C. Protein Eng Des Sel 17, 293–304 9. Razai, A., Garcia-Rodriguez, C., Lou, J., Geren, I. N., Forsyth, C. M., Robles, Y., Tsai, R., Smith, T. J., Smith, L. A., Siegel, R. W., Feldhaus, M., and Marks, J. D. (2005) Molecular evolution of antibody affinity for sensitive detection of botulinum neurotoxin type A. J Mol Biol 351, 158–69 10. Weaver-Feldhaus, J. M., Miller, K. D., Feldhaus, M. J., and Siegel, R. W. (2005) Directed evolution for the development of conformationspecific affinity reagents using yeast display. Protein Eng Des Sel 18, 527–36 11. Garcia-Rodriguez, C., Levy, R., Arndt, J. W., Forsyth, C. M., Razai, A., Lou, J., Geren, I., Stevens, R. C., and Marks, J. D. (2007) Molecular evolution of antibody cross-reactivity for two subtypes of type A botulinum neurotoxin. Nat Biotechnol 25, 107–16 12. Swers, J. S., Kellogg, B. A., and Wittrup, K. D. (2004) Shuffled antibody libraries created by in vivo homologous recombination and yeast surface display. Nucleic Acids Res 32, e36 13. Weaver-Feldhaus, J. M., Lou, J., Coleman, J. R., Siegel, R. W., Marks, J. D., and Feldhaus, M. J. (2004) Yeast mating for combinatorial Fab library generation and surface display. FEBS Lett 564, 24–34 14. VanAntwerp, J. J., and Wittrup, K. D. (2000) Fine affinity discrimination by yeast surface display and flow cytometry. Biotechnol Prog 16, 31–7 15. Siegel, R. W., Coleman, J. R., Miller, K. D., and Feldhaus, M. J. (2004) High efficiency
16.
17.
18.
19.
20.
21.
22.
23.
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recovery and epitope-specific sorting of an scFv yeast display library. J Immunol Methods 286, 141–53 Boder, E. T., and Wittrup, K. D. (1998) Optimal screening of surface-displayed polypeptide libraries. Biotechnol Prog 14, 55–62 Miller, K. D., Weaver-Feldhaus, J., Gray, S. A., Siegel, R. W., and Feldhaus, M. J. (2005) Production, purification, and characterization of human scFv antibodies expressed in Saccharomyces cerevisiae, Pichia pastoris, and Escherichia coli. Protein Expr Purif 42, 255–67 Ewert, S., Huber, T., Honegger, A., and Pluckthun, A. (2003) Biophysical properties of human antibody variable domains. J Mol Biol 325, 531–53 Corisdeo, S., and Wang, B. (2004) Functional expression and display of an antibody Fab fragment in Escherichia coli: study of vector designs and culture conditions. Protein Expr Purif 34, 270–9 Li, J., Menzel, C., Meier, D., Zhang, C., Dubel, S., and Jostock, T. (2007) A comparative study of different vector designs for the mammalian expression of recombinant IgG antibodies. J Immunol Methods 318, 113 Zaccolo, M., Williams, D. M., Brown, D. M., and Gherardi, E. (1996) An approach to random mutagenesis of DNA using mixtures of triphosphate derivatives of nucleoside analogues. J Mol Biol 255, 589–603 Gietz, R. D., and Woods, R. A. (2006) Yeast transformation by the LiAc/SS Carrier DNA/PEG method. Methods Mol Biol 313, 107–20 Chao, G., Lau, W. L., Hackel, B. J., Sazinsky, S. L., Lippow, S. M., and Wittrup, K. D. (2006) Isolating and engineering human antibodies using yeast surface display. Nat Protoc 1, 755–68
Chapter 21 Using RNA Aptamers and the Proximity Ligation Assay for the Detection of Cell Surface Antigens Supriya S. Pai and Andrew D. Ellington Summary The detection and typing of tumor cells based on differentially or similarly expressed antigens (biomarkers) have proven to be increasingly important for the diagnosis and treatment of various cancers. Sensitive techniques for the detection of cell surface antigens are therefore crucial for the early and accurate detection of cancer. Although techniques such as ELISA and tissue staining have proven their worth, these techniques often either require substantial amounts of starting material or are prone to high background and false negatives. The proximity ligation assay (PLA) has proven to be an exquisitely sensitive technique with very low background. Two probes that bind adjacent to one another on a protein target can be ligated, yielding a unique amplicon that can be sensitively detected by real-time PCR. We have now adapted PLA to cell surface protein targets using modified RNA aptamers, and have shown that aptamerbased cell surface PLA can successfully detect and differentiate between cells that differentially express a tumor antigen, the prostate specific membrane antigen (PSMA). Key words: Proximity ligation, PLA, Aptamer, Selection, Real-time PCR, Detection.
1. Introduction Aptamers are short RNA or DNA molecules analogous to antibodies in their ability to bind tightly and specifically to protein antigens. Aptamers can be selected in vitro from large nucleic acid libraries. Because of their small size and the fact that they can be readily modified during chemical synthesis, aptamers have been adapted to a variety of diagnostic and therapeutic assays in the laboratory and clinic (1–5). In particular, aptamers may prove useful for the diagnosis and treatment of cancers. The discovery and analysis of biomarkers to characterize tumorigenesis and malignancy has allowed for the Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_21
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development of a number of diagnostic assays and for the targeting of drugs and other therapeutics (3). However, although the trend toward the identification of individual protein biomarkers for different tumors has proven successful, it is quite likely that the number of tumor types that present in the human body do not always correlate perfectly with a given biomarker or even a given set of biomarkers. Rather, there are likely more probabilistic descriptions of the markers associated with the onset and progression of different tumors. To cast a wide net for such probabilistic descriptions of tumor cells, we and others have begun to select aptamers directly against transformed cells (6–10). To best utilize anti-cell aptamers for the detection and diagnosis of tumors, we have attempted to couple these nucleic acid reagents with powerful technologies for nucleic acid amplification. The proximity ligation assay (PLA) (11–14) is a technique that relies on real-time PCR but has a much lower background than other immunoPCR methods. PLA requires the adjacent binding of two reagents, forming a sort of local “sandwich” that allows nucleic acid tags on the reagents to be ligated and thus uniquely amplified. Because the formation of unique amplicons in the absence of a target is a very low probability event, PLA can be used for the detection of zeptomole amounts of protein analytes (13–16) and for the detection of single spores (17). In the current method, we demonstrate how aptamers selected against antigens on the cell surface or against the cell surface itself can be adapted to PLA for the sensitive detection of small numbers of tumor cells (Fig. 1).
2. Materials 2.1. Tissue Culture and Cell Lysis
1. Cell lines: LNCaP (ATCC CRL-1740), PC3 (ATCC CRL1435) and DU145 (ATCC HTB-81) cells were all obtained from the ATCC, Rockville, MD. 2. Media: LNCaP cells were grown in RPMI 1640 media (ATCC, Rockville, MD), PC3 cells were grown in Ham’s F12K media (ATCC, Rockville, MD), and HeLa cells were grown in Eagle’s Minimal Essential Medium, MEME (ATCC, Rockville, MD). All the media were supplemented with 10% fetal bovine serum (FBS, Gibco, Carlsbad, CA) and 100 μg/mL solution of penicillin/streptavidin (Invitrogen, Carlsbad, CA). 3. Cell harvest: The adherent cells were released into solution with trypsin (0.025%) and 1 mM ethlenediamine tetraacetic acid (EDTA), pH 7.5 (Gibco, Carlsbad, CA).
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Fig. 1. Adaptation of anti-cell aptamers to PLA. Anti-PSMA aptamers were extended at the 3′ and the 5′ end and DNA probes were hybridized to the extended aptamers. When bound to adjacent sites on a cell surface the oligonucleotides (one of which contained a 5′phosphate) could be aligned by a splint oligonucleotide and subsequently be ligated and amplified. The new amplicon is detected by real-time PCR.
4. Buffer: Cells were washed with sterile phosphate buffered saline, PBS (2.67 mM KCl, 137.9 mM KH2PO4, 1.47 mM NaCl, 8.06 mM Na2HPO4 · 7H2O) supplemented with 0.5 mM CaCl2 (PBS+). 5. Cell counting: 0.4% (w/v) Trypan blue (Biocompare, South San Francisco, CA) and hemocytometer slides (Hausser Scientific, Horsham, PA) were used for cell counts. 2.2. Synthesis of Extended Aptamers
1. Aptamer: The anti-PSMA aptamer A9 was used for cell surface PLA. 5′GGGAGGACGAUGCGGACCGAAAAAGACCUGA CUUCUAUACUAAGUCUACGUUCCCAGACGACUCGC CCGA 3′ (18). The DNA template for transcription of this aptamer had the sequence 5′ TTCTAATACGACTCACTATAGGGAGGACGATGCG GACCGAAAAAGACCTGACTTCTATACTAAGTCTACGTTCCCAGACGACTCGC CCGA 3′. ■
Primers (IDT, Coralville, IA):
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For the 3′ extension: 20 μM stocks (3′ ext.1: TTCTAATACGACTCACTA TAGGGAGGACGATGCGG), (3′ ext.2: 5′TCGG G CGAGTCGTCTG 3′), and (3′ ext. 3: 5′ CTGGT CATGCGCGGCATT TAATTCTCGGGCGAGTCGTCTG 3′). For the 5′ extension. 20 μM stocks (5′ ext.1: 5′ TCGGGCGAGTCGTCTG 3′), (5′ ext.2: 5′ GGGGAGAATA TTGAAATATAAATGGGAGGACGATGCGGACCG 3′) and (5′ext.3: 5′ TTCTAAT ACGACTCACTATAGGGGAGAATATTGAAATATAAAT 3′). ° Polymerase Chain Reaction (PCR): Aptamer A9 was PCR amplified using 10× PCR buffer (100 mM Tris–HCl, 500 mM KCl, 15 mM MgCl2, 0.1% Gelatine, pH 8.3), 4 mM dNTP stock, 20 μM primer stocks, and Taq DNA polymerase (20 U, Invitrogen, Carlsbad, CA). ° DNA visualization and purification. NuSieve 3:1 agarose (4% gel) supplemented with 0.5 μg/mL of ethidium bromide was used to analyze PCR products. 6× orange dye (recipe = 0.02 g orange G, 6 mL 50% glycerol, 1.2 mL 0.5 M EDTA pH 8.0, and 2.78 mL sterile water) was used for loading DNA samples. DNA was sized with 100 bp ladders (Invitrogen, Carlsbad, CA) and quantitated using 100-bp and 200-bp quantitation standards (100 ng/μL, GenSura laboratories, Del Mar, CA). ° Transcription: Transcriptions of the extended aptamers were performed using 10×Transcription buffer stock (400 mM Tris, pH 8.0; 120 mM MgCl2; 50 mM DTT; 10 mM spermidine chloride; 40% (w/v) polyethylene glycol 8000; 0.02% Triton X-100), 100 mM stocks of ATP, GTP, 2′ fluoro-CTP (2′F-CTP; TriLink Biotech, CA), 2′fluoro-UTP (2′F-UTP; TriLink Biotech, CA) and Y639F T7 RNA polymerase [1 U/ μL, (19)]. ° Aptamer purification: DNase I (1 U/μL, Epicenter Biotechnologies, Madison, WI), 8% polyacrylamide (7 M Urea), TEMED (Sigma, St. Louis, MO), and 10% ammonium persulfate (APS, Biocompare, CA). ° RNA precipitation: 3 M NaOAc stock, 100% ethanol, and 1 μg/μL glycogen. 2.3. Proximity Ligation Reactions
1. Aptamers: Cell-specific aptamers (in this instance, the antiPSMA aptamer A9) with 3′ and 5′ extensions: 3′ extended aptamer A9 5′GGGAGGACGAUGCGGACCGAAAAAGACCUGACUUCUAUACUAAGUCUACGUUCCCAGACGACU CGCCCGAGAAUUAAAUGCCCGCCAUGACCAG 3′
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5′ extended aptamer A9 5′GGGAGAAUAUUGAAAUAUAAAUGGGAGGACGAUGCGGACCGAAAAAGACCUGACUUCUAUACUAAGUCUACGUUCCCAGACGACUCGCCCGA 3′. 2. PLA probes: PLA probes to hybridize to the 3′- and 5′-extended aptamers were as follows: PLA probe hybridized to the 3′ end of the extended aptamer A9 5′CTGGTCATGGCGGGCATTTAATTCGTGACTTCGTG GAACTATCTAGCGTGTACGTGAGTGGGCATGT 3′ (IDT, Coralville, IA). PLA probe hybridized to the 5′ end of the extended aptamer A9 5′pGTCATCATTCGAATCGTACTGCAATCGGGTATTAT TTATATTTCAATATTCTCCC 3′ (IDT, Coralville, IA).
2.4. Real-Time PCR
■
Splint: The connecting nucleotide or the DNA splint used was of the following sequence 5′AAGAATGATGA CCCTCTTGCTAAAA 3′(IDT, Coralville, IA).
■
Buffer: PBS supplemented with 0.5 mM CaCl2 (PBS+) was used for cell dilution.
■
Ligation: The ligation reaction was carried out using 0.4 U of T4 DNA ligase, 1 U/μL (Invitrogen, Carlsbad, CA), and 80 μM ATP.
Real-time PCR was carried out using: 1. PCR Buffer: 10×AmpliTaq PCR buffer (Applied Biosystems, Foster City, CA) and 50 mM MgCl2 stock (Applied Biosystems, Foster City, CA). 2. Primers: 20 μM primer stocks (5′-GTGACTTCGTGGAAC TATCTAGCG and 5′-AATACCCGATTGCAGTA CGATTC; IDT, Coralville, IA). 3. Real-time PCR probe: 1 μM TaqMan probe stock (5′-FAMTGTACGTGAGTGGGCATGTAGCAAGAGG-BHQ, where FAM is 6-carboxyfluorescein and BHQ the Black Hole Quencher-1; IDT, Coralville, IA). 4. Deoxynucleotides: 4 mM stock of dNTPs. 5. 50× ROX reference dye (Invitrogen, Carlsbad, CA). 6. Enzyme: 1.5 U AmpliTaq Gold polymerase (Applied Biosystems, Foster City, CA). 7. Positive control: 10 pM full-length template stock (5′ AAGATTATGCTGA GTGATATCCTCCGTCATCATTCGAATCGTACTGCAATCGGGTATT 3′). 8. Real-time PCR: 7300HT Real-time PCR system (Applied Biosystems, Foster City, CA).
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3. Methods As a proof-of-principle we utilize aptamers selected against prostate-specific membrane antigen [PSMA; (18, 20)]. The aptamers contain 2′ fluoropyrimidines, which make them less sensitive to cellular nucleases and thus allow their use even in nuclease-laden environments such as serum. To adapt the aptamers to PLA they were extended with sequences that could form a ligation junction and yet not interfere with their binding function. An aptamer pair was created, one extended at its 5′ end and one extended at its 3′ end. The extended aptamers were annealed to PLA probes to generate aptamer-probe ligands; this system was modular and should allow probe sets to be changed as necessary. Since PSMA is known to be a dimer, it was hoped that the two different aptamer-probe ligands would bind adjacent to one another on the cell surface and that this would in turn lead to ligation and real-time PCR amplification. By subtracting the real-time PCR signal in the presence of cells from the signal in samples that have no cells, a metric for identifying small numbers of tumor cells in a sample was developed. Cells were incubated with aptamer-probes for an hour to facilitate aptamer binding, while ligation reactions were carried out for a much shorter time (5 min) to suppress the creation of background amplicons. Various optimizations were carried out to improve the cellspecific signal. Aptamer-probe concentrations ranging from 1 nM to 0.1 pM were attempted in the presence of a minute concentration of splint (400 pM; Fig. 2). The same probe dilution series was also performed in the presence of even smaller splint concentrations of 40 pM and 4 pM (Fig. 3). Overall, it is difficult to
Fig. 2. The A9 aptamer-probe can sensitively detect LNCaP cells versus PC3 or Du145 cells. PLA was carried out using A9 aptamer-probe concentrations of 1 nM, 100 pM, 10 pM, 1 pM and 0.1 pM with either 1,000 LNCaP, PC3, DU145 cells, or no cells. Splint concentration was kept constant at 400 pM. Delta C(T) values for all assays were calculated by subtracting the C(T) value of samples with cells from samples without cells.
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Fig. 3. Optimization of PLA conditions by changing splint concentration. (A) PLA was performed using 1,000 LNCaP or PC3 cells or no cells at a splint concentration of 40 pM. The A9 aptamer-probe concentration was varied from 1 nM to 0.1 pM. (B) PLA was carried out using 1,000 LNCaP or PC3 cells or no cells at a splint concentration of 4 pM. The A9 aptamer-probe concentration was varied from 1 nM to 0.1 pM.
predict the amounts of cells, aptamer-probes, and splints that will yield robust real-time PCR signals. This is largely because the differential formation of amplicons is dependent on several different factors: background ligation (dependent both on aptamer-probe and splint concentration); cell-specific ligation (dependent also on cell concentration); and cell-specific suppression of ligation. This latter phenomena is not generally acknowledged in PLA experiments with proteins, but is extremely important with cells. If probes bind sparsely on a cell surface, they may not be adjacent to one another for ligation and in fact may contact one another at a probability or rate that is less than what will occur in solution. Nonetheless, on the basis of multiple assays and observations, we have generally found that cell-based PLA with aptamer-probe sets work well at aptamer-probe concentrations ranging from 1 nM to 1 pM, and splint concentrations from 400 pM to 4 pM. The method eventually proved to be sensitive enough to detect a very small numbers (10–1,000) of cells even when they are present in a background of a large amount of non-cognate cells (21). 3.1. Preparation of cells for PLA
1. Prostate cancer cell lines LNCaP, PC3, and DU145 were cultured and grown in T-25 flasks (VWR, West Chester, PA) in RPMI 1640 media, Ham’s F12K media, and Eagle’s Minimal Essential Medium, MEME, respectively. All cell lines were grown in a tissue culture incubator supplied with 5% CO2 and maintained at 37°C. 2. Upon reaching a confluence of about 80%, the cells were passaged using 1 mL of Trypsin/EDTA per flask. Prior to trypsinization, media was withdrawn from harvested cell culture flasks, and the confluent cells were gently washed with 4 mL of tissue culture grade PBS to remove residual traces of media. Following the PBS wash, 1 mL of the Trypsin/EDTA
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solution was dispensed into the flasks and spread uniformly over the adherent cells. Trypsinization was enhanced by incubating the flasks at 37°C for 5–10 min (see Note 1). 3. Most cells were released into solution following 10 min of trypsinization. Cells that did not detach following trypsinization were solubilized by gently tapping the bottoms of the tissue culture flasks. An additional 5 mL of media was dispensed directly into the trypsin-cell mixture, and cell clumps were broken up by pipetting up and down a total of 11 times. 4. The cell suspension was placed in a 25 mL conical flask and centrifuged at 1,500rpm or 238 × g at 4°C for 5 min in a swinging bucket rotor. The supernatant was withdrawn from the flask, and the cell pellet was washed with 5 mL of PBS +. The cells were recentrifuged at 1,500 rpm at 4°C for 5 min in a swinging bucket rotor. The PBS+ supernatant was withdrawn, and the cell pellet was resuspended in 1 mL of PBS+. 5. To count the total number of cells present in the PBS+ solution, a 10 μL aliquot of the cell solution was mixed with 10 μL of Trypan blue dye to make a dye-cell mix with a total volume of 20 μL. The dye-cell mix was placed on each side of the hemocytometer slide grid (10 μL each side) and covered with a cover-slip for cell counting. The slide was placed under a phase-contrast microscope and live cells (unstained) present in the five main squares per grid were counted for both the grids of the hemocytometer. The total number of cells counted was divided by the total number of squares counted (ten total for both grids) and then multiplied by the cell dilution factor, which in this case was 2. Having accounted for the dilution factor, the cell count was then multiplied by 104 to calculate the total cell count in 1 mL. The final cell count is thus calculated by the following formula: Total cells counted (across both grids) ´ 2 (dilution factor) ´ total volume (1 mL) ´ 104 10
6. Since PLA can be carried out with a small number of cells, the total cell number was generally appropriately diluted by first concentrating the cells by centrifugation and then resuspending them in a concentration of PBS + that yielded the desired number of cells in 1 μL for addition to PLA reactions (see Table 1). Cells were stored on ice until used in PLA. 3.2. Synthesis of Extended Aptamers
1. A modified RNA aptamer, A9 [see (18)], that had been previously selected against the prostate-specific membrane antigen protein (PSMA) was used for the adaptation of aptamers to cell-mediated PLA. Aptamers have also been selected against whole cells and these can also be employed in PLA [see (10) for aptamer selection protocols].
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Table 1 Incubation mixes Final concentration/ Apt-pb 10 nM Stock concentration/ 100 nM Apt-pb
1 nM
0.1 nM
0.01 nM
0.001 nM
10 nM
1 nM
0.1 nM
0.01 nM
3′ Apt-pb
2 μL
2 μL
2 μL
2 μL
2 μL
5′ Apt-pb
2 μL
2 μL
2 μL
2 μL
2 μL
Cells or PBS+
1 μL
1 μL
1 μL
1 μL
1 μL
ddH2O
15 μL
15 μL
15 μL
15 μL
15 μL
Total volume
20 μL
20 μL
20 μL
20 μL
20 μL
2. To synthesize the A9 RNA aptamer with 3′ and 5′ extensions, the DNA template for the A9 aptamer was amplified in a twostep PCR using the following conditions: 95°C for 1 min, 45°C for 30 s and 60°C for 30 s; cycle ten times. (a) For adding on the 3′ extension to the A9 DNA template, 10 ng of the A9 template (5′ ACCGAAAAAGACCTGACTTCTATACTAAGTCTACGTTCCCAGACGACTCGC C CGA 3′) was amplified using 10 μL of PCR buffer, 4 μL of dNTPs, 0.5 μL of primers 3′ ext.1 (5′TTCTAATACGACTCACTATAGGGAGGACGATGCGG) and 3′ ext.2 (5′ TCGGGCGAGTCGTCTG 3′) and 1 μL of Taq DNA polymerase in a final volume of 100 μL. (b) For adding on the 5′ extension to the A9 DNA template, 10 ng of the A9 template (5′ ACCGAAAAAGACCTGACTTCTATACTAAGTCTACGTTCCCAGACGACTCGCC CGA 3′) was amplified using 10 μL of PCR buffer, 4 μL of dNTPs, 0.5 μL of primers 5′ ext.1 (5′ TCGGGCGAGTCGTCTG 3′) and 5′ ext.2 (5′ GGGGAGAATATTGA AATATAAATGGGAGGACGATGCGGACCG 3′) and 1 μL of Taq DNA polymerase in a final volume of 100 μL. 3. The sequences of the extended A9 aptamer were as listed in Subheading 2.3, step 1 4. About 5 μL of the PCR samples each were mixed with 1 μL of 6× orange dye and subjected to gel electrophoresis alongside a 100-bp marker (Invitrogen, Carlsbad, CA) to confirm the appropriate size of the product (′100 bp).
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5. Following this, 2 μL of the 3′ extended template was further amplified using 10 μL of PCR buffer, 4 μL of dNTPs, 0.5 μL of primers 3′ ext.1 (5′ TTCTAATAC GACTCACTATAG GGAGGACGATGCGG) and 3′ ext.3 (5′ CTGGTCATGCG CGGCATTTAATTCTCGGGCGAGTCGTCTG 3′) and 1 μL of Taq DNA polymerase in a final volume of 100 μL. For the 5′ extended template, 2 μL of the PCR product was further amplified using 10 μL of PCR buffer, 4 μL of dNTPs, 0.5 μL of primers 5′ ext.1 (5′ TCGGGCGAGTCGTCTG 3′) and 5′ ext.3 (5′ TTCTAATACGACTCACTATAGGGG AGAATATTGAAATATAAAT 3′) and 1 μL of Taq DNA polymerase in a final volume of 100 μL. 6. The PCR products (for the 3′ and 5′ extensions) were analyzed as mentioned in step 4. On confirmation that the PCRs were successful (product size ∼ 120 bp), the same PCRs were repeated in a 400 μL volume (each). The amplified DNA (400 μL) was mixed with 40 μL of 3 M NaOAc, 1,000 μL of 100% ethanol and 1 μL of glycogen, vortexed, and incubated at −80°C for 30 min. Following incubation, the samples were centrifuged at ~10,000 × g, 4°C for 45 min to obtain DNA pellets. The supernatant was withdrawn, and the pellets were gently washed with 400 μL of ethanol followed by centrifugation at ~10,000 × g 4°C for 30 min. The supernatant was withdrawn once again, and the DNA pellets were air-dried for 15 min to remove residual ethanol. 7. Once the pellets were air-dried, the 3′ and 5′ extended A9 DNA templates were resuspended in 20 μL of ddH2O. After resuspension, 1 μL of the DNA samples were mixed with 1 μL of 6× Orange dye and quantitated against 1 μL of 100-bp and 200-bp double-stranded DNA quantitation standards. 8. The 3′ and 5′ extended A9 aptamer templates were transcribed into RNA aptamers in a 20 μL reaction using 2 μL of transcription buffer, 1.5 μL of ATP, GTP, 2′F CTP, 2′F UTP and 1 μL Y639F T7 RNA polymerase at 42°C for 6 h. 9. The transcripts were treated with 1 μL of DNase I and incubated at 37°C for 30 min. Following DNase I treatment, the RNA was mixed with 20 μL of 2× denaturing dye, heat denatured at 65°C for 3 min and separated on a 8% denaturating gel made with 25 mL of 8% acrylamide (7 M Urea), 80 μL of APS and 25 μL of TEMED. The bands corresponding to the 3′ and 5′ extended A9 aptamers were excised and incubated overnight with 800 μL of ddH2O to elute the RNA from the gel. The RNA eluant was divided into two 400 μL aliquots and precipitated as mentioned in step 6 (see Subheading 3.2). The purified RNA was quantitated using a Nanodrop (Nanodrop, Wilmington, DE) and extinction coefficients of 893700 L/ mol.cm and 898600 L/mol cm for the 3′ extended and the 5′ extended A9 aptamers, respectively.
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3.3. Preparation of the Aptamer-Probe Conjugates for PLA
1. The transcribed and extended 3′ and 5′ aptamers were mixed in equimolar amounts (1 μM) with the 3′ and the 5′ PLA probes (1 μM), respectively, in a final volume of 100 μL of PBS + and incubated at 65°C for 5 min. The mix was cooled for 15 min at RT to enhance annealing of the probe to the aptamer and thereby generate aptamer-probe constructs.
3.4. Preparation of the Ligation Mix for PLA
1. The 1 μM stocks of the 3′and the 5′ aptamer-probes were diluted with PBS + in a tenfold dilution series to yield stocks of 100 nM, 10 nM, 1 nM, 0.1 nM, and 0.01 nM. 2. Every PLA reaction was made with either the target cells (LNCaP), nontarget cells (PC3 or DU145), or no cells (only PBS+). Accordingly, 1 μL of LNCaP, 1 μL of PC3/DU145 cells, or 1 μL of PBS + were mixed with 2 μL of the 3′and the 5′ aptamer-probe stocks in a final reaction volume of 20 μL. PLA incubation mix concentrations are displayed in Table 1 3. The reactions were incubated at RT for 1 h (see Note 2).
3.5. Proximity Ligation
1. The reaction samples incubated with aptamer-probes were ligated using 4 μL of 1 mM ATP, \1 μL of 20 nM splint, and 0.4 μL of T4 DNA ligase (raising the incubation volume to 30 μL). Table 2 depicts a single ligation reaction. 2. Ligation was allowed to proceed at 25°C for 10 min. The ligated samples were heated at 95°C for 5 min to lyse the cells and inactivate the ligase. The samples were then cooled on ice.
3.6. Real-Time PCR
1. A real-time PCR master-mix was made consisting of 1.25 μL of the forward and reverse primers, 5 μL Amplitaq buffer, 4 μL MgCl2, 2.5 μL of dNTPs, 3.75 μL of TaqMan probe, 1 μL of the ROX reference dye, 26 μL of ddH2O, and 0.3 μL of Amplitaq gold polymerase. 2. Every reaction well in the real-time PCR assay plate received 45 μL of the master-mix along with 5 μL of the PLA reaction. Every PLA reaction was assayed in triplicate (three separate additions of 5 μL).
Table 2 Ligation mix Incubation mix (from Table 1)
20 mL
ATP (1 mM stock)
4 μL
Splint (20 nM stock)
1 μL
ddH2O
4.6 μL
T4 DNA ligase (1 U/μL)
0.4 μL
Total volume
30 μL
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3. In addition to the PLA samples, 5 μL of a full length template stock was mixed with 45 μL of master-mix and served as the positive control on the real-time PCR assay plate. 4. Negative controls containing only the master-mix and no templates were also loaded onto the assay plate. 5. The set-up of a typical master-mix for a PLA reaction is described Table 3 6. Once the real-time PCR assay plate was prepared, the samples were mixed carefully with a multi-channel pipette, the wells were sealed with a Microamp full plate cover seal (ABI, Foster City, CA), and the plate was centrifuged at 1,500rpm (~238 × g) at 4°C for 1 min. 7. The prepared plate was placed in the real-time PCR system and PLA samples were assayed using the “relative quantification” plate program. The PCR cycle conditions were: 95°C for 10 min followed by 50 cycles of 95°C for 1 min and an extension step of 60°C for 1 min. 1. Once the real-time run was completed, the saved real-time assay file was opened as a “relative quantification study.” Data were displayed using the analysis button to obtain the cycle threshold C(T) values of the samples. Results from a real-time PCR run are typically represented in the form of C(T) values, which is the point at which the fluorescent intensity of the
3.7. PLA analysis
Table 3 Real-time PCR mastermix Stock concentration
Final concentration
1× (mL)
3× (mL) (for triplicates)
Forward primer
20 μM
500 nM
1.25
3.75
Reverse primer
20 μM
500 nM
1.25
3.75
dNTPs
4 mM
200 μM
2.5
7.5
MgCl2
25 mM
2 mM
4
12
10× buffer
10×
1×
5
15
TaqMan probe
1 μM
70 nM
3.75
11.25
ROX reference dye
50×
1×
1
3
Amplitaq gold enzyme
5 U/μL
1.5 U
0.3
0.9
H2O
25.95
77.85
Final volume
45
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samples crosses the intensity of the background and is represented in the form of a value termed the cycle threshold. The lower the C(T) value, the higher the number of amplicons formed or lesser the time the sample needs to amplify and cross the threshold over background. 2. To better analyze outlying data points, such as those caused by inefficiently amplified templates, the baseline was manually adjusted. The C(T) values for the reactions with cells were compared with C(T) values of samples without cells. 3. Target specific signals were represented as Delta C(T) by subtracting the C(T) values of the cell-containing reactions from the C(T) values of corresponding reactions with no cells. The triplicate Delta C(T) values were averaged and used to calculate the standard deviation. 4. Bar graphs were plotted with aptamer-probe concentrations on the X-axis and Delta C(T) values on the Y-axis. 5. The higher the Delta C(T) value, the stronger the target specific signal. In this case, reactions derived from PSMA-positive LNCaP cells showed larger Delta C(T) values than reactions derived from noncognate PC3 or DU145 cells (Fig. 3).
4. Notes 1. LNCaP cells adhere less efficiently to the walls of the flask and are prone to detach if washed vigorously with PBS. To avoid this, the PBS is placed in one corner of the flask and the flask is tilted back and forth to spread the liquid evenly on the cells without detaching them. 2. To avoid cell lysis, do not pipette the incubation mixes but mix them by tapping the Eppendorf tube with a finger followed by a quick spin in the micro-centrifuge. In addition, do not vortex once cells have been added.
References 1. Chu, T., Ebright, J., and Ellington, A. D. (2007) Using aptamers to identify and enter cells. Curr Opin Mol Ther. 9, 137–144 2. Jayasena, S. D. (1999) Aptamers: an emerging class of molecules that rival antibodies in diagnostics. Clin Chem. 45, 1628–1650 3. Lee, J. F., Stovall, G. M., and Ellington, A. D. (2006) Aptamer therapeutics advance. Curr Opin Chem Biol. 10, 282–289
4. Nimjee, S. M., Rusconi, C. P., and Sullenger, B. A. (2005) Aptamers: an emerging class of therapeutics. Annu Rev Med. 56, 555–583 5. Yan, A. C., Bell, K. M., Breeden, M. M., and Ellington, A. D. (2005) Aptamers: prospects in therapeutics and biomedicine. Front Biosci. 10, 1802–1827 6. Chu, T. C., Marks, J. W. III, Lavery, L. A., Faulkner, S., Rosenblum, M. G., Ellington,
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8.
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10.
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12.
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Pai and Ellington A. D., and Levy, M. (2006) Aptamer:toxin conjugates that specifically target prostate tumor cells. Cancer Res. 66, 5989–5992 Chu, T. C., Twu, K. Y., Ellington, A. D., and Levy, M. (2006) Aptamer mediated siRNA delivery. Nucleic Acids Res. 34, e73 Daniels, D. A., Chen, H., Hicke, B. J., Swiderek, K. M., and Gold, L. (2003) A tenascin-c aptamer identified by tumor cell selex: Systematic evolution of ligands by exponential enrichment. Proc Natl Acad Sci U S A. 100, 15416–15421 Hicke, B. J., Marion, C., Chang, Y. F., Gould, T., Lynott, C. K., Parma, D., Schmidt, P. G., and Warren, S. (2001) Tenascin-c aptamers are generated using tumor cells and purified protein. J Biol Chem. 276, 48644–48654 Shangguan, D., Li, Y., Tang, Z., Cao, Z. C., Chen, H. W., Mallikaratchy, P., Sefah, K., Yang, C. J., 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 Fredriksson, S., Gullberg, M., Jarvius, J., Olsson, C., Pietras, K., Gustafsdottir, S. M., Ostman, A., and Landegren, U. (2002) Protein detection using proximity-dependent DNA ligation assays. Nat Biotechnol. 20, 473–477 Gullberg, M., Fredriksson, S., Taussig, M., Jarvius, J., Gustafsdottir, S., Landegren, U. (2003) A sense of closeness: protein detection by proximity ligation. Curr Opin Biotechnol. 14, 82–86 Gullberg, M., Gustafsdottir, S. M., Schallmeiner, E., Jarvius, J., Bjarnegard, M., Betsholtz, C., Landegren, U., and Fredriksson, S. (2004) Cytokine detection by antibodybased proximity ligation. Proc Natl Acad Sci U S A. 101, 8420–8424
14. Gustafsdottir, S. M., Schallmeiner, E., Fredriksson, S., Gullberg, M., Soderberg, O., Jarvius, M., Jarvius, J., Howell, M., and Landegren, U. (2005) Proximity ligation assays for sensitive and specific protein analyses. Anal Biochem. 345, 2–9 15. Schallmeiner, E., Oksanen, E., Ericsson, O., Spangberg, L., Eriksson, S., Stenman, U. H., Pettersson, K., and Landegren, U. (2007) Sensitive protein detection via triple-binder proximity ligation assays. Nat Methods. 4, 135–137 16. Zhu, L., Koistinen, H., Wu, P., Narvanen, A., Schallmeiner, E., Fredriksson, S., Landegren, U., and Stenman, U. H. (2006) A sensitive proximity ligation assay for active PSA. Biol Chem. 387, 769–772 17. Pai, S., Ellington, A. D., and Levy, M. (2005) Proximity ligation assays with peptide conjugate ‘burrs’ for the sensitive detection of spores. Nucleic Acids Res. 33, e162 18. Lupold, S. E., Hicke, B. J., Lin, Y., and Coffey, D. S. (2002) Identification and characterization of nuclease-stabilized RNA molecules that bind human prostate cancer cells via the prostate-specific membrane antigen. Cancer Res. 62, 4029–4033 19. Padilla, R., and Sousa, R. (1999) Efficient synthesis of nucleic acids heavily modified with non-canonical ribose 2′-groups using a mutant T7 RNA polymerase (RNAP). Nucleic Acids Res. 27, 1561–1563 20. Chu, T. C., Shieh, F., Lavery, L. A., Levy, M., Richards-Kortum, R., Korgel, B. A., and Ellington, A. D. (2006) Labeling tumor cells with fluorescent nanocrystal-aptamer bioconjugates. Biosens Bioelectron. 21, 1859– 1866
Chapter 22 In Vitro Selection of Protein-Binding DNA Aptamers as Ligands for Biosensing Applications Naveen K. Navani, Wing Ki Mok, and Yingfu Li Summary Aptamers are single-stranded functional nucleic acids that possess cognate ligand recognition capability. These functional nucleic acids have been used for biosensing of a variety of ligands. Aptamers are isolated by “in vitro selection” or SELEX from random-sequence nucleic acid pools. For example, DNA aptamers that recognize a protein can be generated by applying a DNA library to an affinity column containing the protein target and retrieving the bound sequences after wash. These sequences are amplified and used for a new round of binding and amplification. The identity of enriched sequences are subsequently revealed by cloning and sequencing. The binding of individual aptamers to the protein can be confirmed by techniques such as gel mobility shift. This chapter will provide a detailed protocol for isolating proteinbinding DNA aptamers. Keywords: DNA; Aptamers, SELEX, Biosensing, EMSA.
1. Introduction DNA, RNA, and proteins are central to all biological reactions. It is now well-established that these macromolecules, along with small metabolites, regulate anabolism and catabolism. The ability to regulate metabolism is fundamental to living systems. A significant deviation from the norm can lead to diseased conditions. Thus, it is important to develop molecular recognition tools that can accurately sense important biological signals or entities in order for timely therapeutic interventions to be made. The development of molecular recognition elements (MREs) or biosensors that faithfully report the identities of these markers, their
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_22
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concentrations, and interactions in vitro or in vivo will pave the way for therapeutics. Furthermore, creative applications of these MREs can contribute to the basic understanding of the disease progression in specialized cases. Nucleic acids, due to their ability to form Watson–Crick base pairing, have long been exploited as MREs to develop detection tools for DNA and RNA and have evolved from classical Southern blotting technique (1) to sophisticated microarray platforms (2). The use of nucleic acids as sensing material has expanded from nucleic acid detection to the recognition of nonnucleic acid targets that include small molecules, proteins, bacteria, viruses, parasites, and mammalian cells (3). This has been realized due to the development of a process known as in vitro selection or SELEX (Systematic Evolution of Ligands by EXponential enrichment) (4, 5). The end result of the SELEX process is the identification of highly selective functional nucleic acid sequences known as aptamers. Aptamers refer to singlestranded DNA, RNA, and even modified nucleic acid molecules that have the ability to form defined tertiary structures and engage a specific target for binding. With aptamer sequences on hand, one can design various platforms for biosensing application (6). It is important to link the molecular recognition capability of aptamers to the signal transduction capability in a simple, convenient, and universal manner (7). Such a design would allow the aptamer target interaction to be readily detected with various fluorophores has been widely investigated largely because florescence technique is easy to use and there are many different strategies to convert an aptamer into a fluorescent reporter (8, 9). The key step in creating an aptamer-based biosensor is the isolation of aptamers for a given target. Since proteins are common targets for biosensor engineering, we will describe a detailed method for isolating DNA aptamers for a protein target from a random-sequence DNA library by SELEX in this chapter. In addition, we will describe general methods for the characterization of protein-binding aptamers by gel mobility shift.
2. Materials 2.1. Design, Synthesis, and Purification of DNA Libraries and Primers
1. DNA libraries: A typical DNA library is composed of sequences containing a random-sequence region of 50–60 nucleotides (nt). The library is designated as L1, with the nucleotide sequence of 5′-AGACCACAACGGTTTCCC(N60)-TAGCATAACCCCTTG-3′, is used in some of our aptamer selection projects.
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2. DNA primers: For the amphification of library L1, the following two primers can be used: P1, 5′–AGACCACAACGGTTTCCC-3′, and P2, 5′-CAAGGGGTTATGCTrA-3′. Note that P2 ends with an adenine ribonucleotide (rA). By using the ribo-terminated primer in the PCR step and digesting the double-stranded DNA product with NaOH which cleaves the lone RNA linkage embedded in the nonaptameric strand and makes this strand shorter than the aptameric strand, and subsequently conducting a PAGE-based purification step, we can retrieve the single-stranded DNA that corresponds to the aptamer strand during each selection cycle. All DNA oligonucleotides can be ordered from commercial sources; we order ours from Keck Facility at Yale University at 0.2–1.0 micromole scale synthesis. 3. 10% denaturing (8 M urea) PAGE: Mix 1681.7 g of urea, 400 mL of 10× TBE and 1 L of 40% 29:1 acrylamide/ bis-acrylamide premix (BioShop, Canada); add ddH2O to 4 L; dissolve with mild heat and stirring. Store at 4°C in amber colored bottles. Acrylamide is a known neurotoxin when unpolymerized and thus, care should be taken to avoid contact with the body. 4. Ammonium persulfate (APS, BioShop, Canada): Prepare 10% solution in water, store at 4°C. 5. N,N,N,N′-Tetrametheyl-ethylenediamine (TEMED, BioShop, Canada). This chemical is harmful if inhaled; it should be handled in fumehood. 6. Tris-borate EDTA (TBE) buffer (89 mM Tris, 89 mM boric acid, 2 mM EDTA, pH 7.5): Add 432 g of Tris base (Bioshop, BioUltraPure grade), 220 g of boric acid (Bioshop, Biotechnology grade) and 80 mL of 0.5 M EDTA, pH 8.0, to a 5 L plastic beaker; add ddH2O to a final volume of 4 L. Autoclave and store at 4°C. This solution is a 10× stock of TBE. 7. 0.5 M EDTA: Add 186.1 g EDTA (Bioshop, Biotechnology grade) to 800 mL ddH2O. Adjust the pH to 8.0 using NaOH pellets. Make the final volume to 1 L using ddH2O. Autoclave and store at 4°C. 8. Elution buffer: Mix 8 mL of 5 M NaCl, 2 mL of 1 M Tris−Cl, pH 7.5, 0.4 mL of 0.5 M EDTA, pH 8.0; add ddH2O to 200 mL. Autoclave and store at 4°C. 9. 2× Gel loading buffer: Mix 8 g of sucrose (Bioshop, UltraPure grade), 10 mg of bromophenol blue (Bioshop, ACS grade), 10 mg of xylene cyanol FF (Sigma, molecular biology grade), 400 μL of 10% sodium dodecyl sulfate (Bioshop, electrophoresis grade), and 4 mL of 10× TBE with enough ddH2O to bring the volume to 40 mL; dissolve with mild heat and stirring. To the mixture, add 44 g of urea (Bioshop, Canada,
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molecular biology grade); dissolve with mild heat and stirring. Store at 4°C. 10. Vertical Nucleic acid electrophoresis system: Our vertical PAGE system includes a power supply, glass plates, spacers, combs, clamps, and heating plates. We use adjustable slab electrophoresis unit Model ASG-400, from CBS Scientific (California) with power supply EC3000-90, from E-C Apparatus Corporation. 11. Hand-held UV shadowing lamp (VWR Scientific), razor blades, cling-film, kimwipes, ethanol and MilliQ grade distilled water. 2.2. Selection and Counter Selection
1. Partitioning matrix: The choice of matrix for the immobilization of ligand is dependent on the tag on the ligand (see Note 1). In our case, for the selection of aptamers against proteins, we find that His-tagged recombinant proteins are the best choice. Thus, we used Ni-NTA beads (Qiagen, Canada) as our matrix. 2. Selection buffer: PBS (phosphate buffered saline) diluted to 1× from a 10× stock (Dulbecco’s 10× PBS) and supplemented with 5 mM of MgCl2, 10 mM of KCl, and 0.01% (v/v) of Tween-20. 3. Binding buffer: Selection buffer supplemented with 2 μg/ mL of yeast tRNA (Invitrogen) and 125 μg/mL of BSA (Bovine serum albumin from Sigma-Aldrich). 4. Elution buffer: Selection buffer supplemented with 500 mM of imidazole (BioShop). 5. 1.5 mL low binding Eppendorf tubes (Sarstedt). 6. Rocker-shaker (Nutator-Clay Adams brand), refrigerated bench-top centrifuge (Beckman Coulter). All solutions are made in autoclaved MilliQ water, filtered through 0.22-μm syringe-filters (Millipore) and stored at 4°C.
2.3. Polymerase Chain Reaction
1. Thermocycler: Any themocycler can be used for Polymerase Chain Reaction (PCR). We regularly use the Smart-cycler™ real-time PCR machine (Cepheid), which allows us to monitor the progress of PCR amplification and decide when to stop the reaction. 2. Real time PCR tubes (available from Cepheid). 3. Low retention filter-barrier micropipette tips (MβPMolecular Bio products). 4. Biotools DNA polymerase (Biotool B&M Labs, Spain) and 10× Biotools buffer (supplied with enzyme) –1× buffer contains 75 mM of Tris−HCl, pH 9.0, 2 mM of MgCl2, 50 mM of KCl, and 20 mM of (NH4)2SO4.
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5. Syber green 50× stock: diluted from a 10,000× stock of Syber green (Molecular Probes) in dimethyl sulphoxide (DMSO, Sigma-Aldrich). 6. 10× dNTP mix (2 mM of each dNTPs): Fermentas Life Sciences; special dNTP-dGTP mix: 2 mM of dATP, dTTP, dCTP, and 0.2 mM of dGTP. 7. 0.25 N NaOH: Dissolve 2.5 g of NaOH in 250 mL of autoclaved MilliQ water. 8. 3 M sodium acetate, pH 5.2: Dissolve 40.8 g of sodium acetate-tri hydrate in 80 mL of autoclaved milliQ water, adjust to pH 5.2 with glacial acetic acid and make up the total volume to 100 mL. 9. Regular stock of [α-32P]-dGTP (GE Healthcare). Store in a secure radioisotope storage cabinet. 2.4. Testing DNA Population for Ligand Binding by Electro Mobility Gel-Shift Assay
1. 8% Native PAGE mix: To 100 mL of 40% acrylamide/ bis-acrylamide (ratio 19:1, Sigma-Aldrich), add 250 mL of 1× TBE (diluted from 10× TBE stock), 2.5 mL of 1 M MgCl2, 7.5 mL of 100% glycerol. Make up the total volume to 500 mL with Milli-Q water. Store in an amber-colored bottle. 2. 4× Gel-shift buffer: Dissolve 2 mg of yeast tRNA, 3 mg of BSA in 4× selection buffer supplemented with 10% (v/v) glycerol and in a total volume of 10 mL. This gel-shift binding buffer is aliquoted and stored in −20°C. 3. Native gel running buffer: Using the 10× TBE stock, prepare 0.5× TBE and add 5 mM of MgCl2 and 1.5% (v/v) of glycerol to the solution. 4. A radio-labeled pool of DNA. 5. Vertical gel electrophoresis apparatus, cold room or cold cabinet, Whatman paper sheets, gel dryer, phosphorimager screens (GE-Amersham), Phosphorimager (Typhoon or Storm – Molecular Dynamics), 10% (w/v) ammonium persulfate, TEMED and other accessories for normal PAGE experiment as mentioned above.
2.5. Cloning and Sequencing Selected DNA Populations
1. InsTAclone™ PCR Cloning Kit (Fermentas Life Sciences #K 1214): The kit contains a TA cloning vector (pTZ57R/T), 5× ligation buffer, T4 DNA ligase 5 U/μL, controls and Transform Aid bacterial transformation system with C medium and T-solution A and B. 2. Chemically competent E. coli host cells such as DH5α or TOP10 (Invitrogen, Canada). 3. Sterile 1.5 mL tubes, inoculating needles, spreaders, disposable petri dishes.
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4. Luria Bertani (LB) Medium (BioShop, Canada): Weigh 25 g of LB powder in to a 2 L capacity beaker. Add 800 mL of MilliQ water to the beaker and stir it until the medium dissolves completely. Make up the volume to 1,000 mL. Divide the media into four aliquots of 250 mL and store it in 500 mL capacity glass flasks. To three of the flasks, add 3.75 g of agar powder (BioShop, Canada). This LB agar medium is then autoclaved. When the LB agar medium cools to about 37–40°C add ampicillin (Sigma) to the final concentration of 100 μg/mL and pour about 15–20 mL of media in individual petri dishes. Allow the plates to cool to room temperature and store them in cool cabinet until they are to be used.
3. Methods A crucial aspect of in vitro selection experiments is the synthesis and purification of random DNA libraries and PCR primers. Great care should be taken to prevent the contamination of pipettes and working surfaces with the libraries. All the solutions and buffer stocks should be stored in the cold room. In vitro selection involves several steps, as outlined in Fig. 1. The aim of the process is to enrich for a set of oligonucleotides, which may contain potential aptamer sequences that can bind a target of interest, from a hugely diverse pool of random oligonucleotides by iterative rounds of selection and amplification. The process of partitioning is continued until a significant set of sequences show binding with the ligand of interest. The final step of selection involves cloning and sequencing of the population. The monoclonal sequences are then characterized individually to determine their affinities for their cognate ligands. Truncation approaches are undertaken to obtain the sequence of a minimal functional aptamer. The final aptamer is then used as a molecular recognition element for biosensing applications upon integrating them with suitable signal transducing platforms. 3.1. Purification of Primers and DNA Libraries
1. These instructions are specific for the use of an adjustable slab electrophoresis unit Model ASG-400, from CBS Scientific, California, in conjunction with plates with a size of 16 cm × 27 cm and 0.7 mm thick spacers. Before casting the PAGE gel, the glass plates should be thoroughly washed, wiped with 95% ethanol and let dry. Set the spacers on the notched plate and cover it with the second nonnotched plate. Clamp the sides of the plates and lay the gel horizontally on a small platform. Rinse a four well preparative comb with
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Fig. 1. The SELEX process: The circle depicts the selection cycles. The enriched population of sequences is tested for binding after 7–8 rounds of selection and following every alternate round then after. Once the binding of DNA to their target saturates, the population of sequences is cloned and sequenced. Finally, functional testing with monoclonal sequences is performed.
water and let dry. Prepare the PAGE mix by pouring 60 mL of 10% PAGE into a clean beaker and adding 600 μL of 10% APS. Caution: always wear gloves, protective goggles, and lab coat when handling these chemicals. Mix the PAGE and APS thoroughly and add 60 μL of TEMED. Working quickly, pour the gel mix into the plates while gently tapping on the top of glass plate to avoid the generation of air bubbles. Once poured, set the comb gently in between the plates and allow the gel to polymerize. 2. Meanwhile, dissolve the DNA oligonucleotides (such as L1, P1 or P2) in water and incubate them on a 37°C heating block for 20 min. Vortex the tubes briefly and spin down the liquid to bottom by a quick spin in a centrifuge. 3. Check the residual PAGE in the beaker for polymerization (usually the gel is polymerized in 30–45 min). Gently take the comb out of the gel, wash the wells immediately with water, and set the plates with polymerized gel into the electrophoresis apparatus with the backing of an aluminum plate to allow for the uniform heating of the gel. Add 1× TBE in upper and lower chamber of the apparatus, fix the electrodes, plug into
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power supply, and start the run at a constant 600 V. Pre-run the gel till the plates begin to warm up to approximately 45°C (this usually takes around 15–20 min). 4. Take the required amount of DNA (∼75 μL/well) and add 75 μL of 2× gel-loading buffer. Denature the DNA sample at 90°C for 2–3 min. Meanwhile, stop the prerunning gel and wash the urea off the wells thoroughly (with 1× TBE using a syringe). Load approximately 150 μL of DNA sample into each well and resume running the gel. 5. Run the gel till the tracking dyes, which contains bromophenol blue (lower band) and xylene cyanol (upper band), indicate that the DNA has migrated to three-fourth of the gel length. 6. Turn off the power supply and remove the plates from the electrophoresis tank. Disassemble the plates by gently prying off the top plate with a spatula. Cover both sides of the gel evenly and smoothly with SaranWrap film. Mark the sample lanes with a permanent marker. 7. Place the wrapped gel onto a fluorescent TLC plate (20 cm × 20 cm silica gel 60 F254; EM Science). Visualize the gel using a handheld UV lamp. The DNA oligonucleotide band will appear as the dark-brown shadow in the gel (xylene cyanol migrates around a 55-nt DNA fragment in a 10% denaturing PAGE gel electrophoresis). 8. Cut out the band with a sterile razor blade. Chop the gel into small pieces with razor and put them in 1.5-mL microfuge tube. Add 500 μL of elution buffer to the tube and place the tube in a thermomixer (Eppendorf) set at 37°C with shaking (1,200 rpm) overnight. 9. The next day, spin down the microfuge tube containing the gel pieces in a bench-top mini-centrifuge for 2–3 min. Gently pipette off 400 μL of the supernatant and transfer it to another 1.5-mL microfuge tube while taking care to avoid gel pieces. Add 40 μL of 3 M sodium acetate, pH 7.0 and 1 mL of cold 100% ethanol to the tube. Incubate the mixture in the −20°C freezer for at least 2 h. 10. Recover the DNA by centrifuging the tube at 14,000 rpm= 21,000×g for 30 min at 4°C. Gently pipette off the supernatant without disturbing the DNA pellet with the pipette tip. Carefully add 100 μL of 70% cold ethanol and thoroughly rinse the inside surface of the tube by gently inverting tube several times. Centrifuge the tube at 14,000 rpm (21,000 × g) for 10 min at 4°C and remove the supernatant. The pellet is washed again with 70% ethanol. Dry the pellet using a speedvac concentrator at room temperature. Dissolve the DNA oligonucleotide in 100 μL of ddH2O. Determine the DNA
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concentration based on the absorbance at 260 nm. Store the DNA oligonucleotide at −20°C till further use. 3.2. In Vitro Selection
1. Take approximately 2 nmol of the DNA library and incubate with 1 mL of selection buffer. Denature the sequences by heating the DNA at 92°C for 5 min, and the tubes are immediately chilled on ice for 10 min. The tubes are kept at room temperature for 20 min then after. This allows the DNA sequences to gradually fold. 2. Meanwhile prepare the Ni-NTA beads (the partition matrix). We have a stock of protein bound beads, which serves as our selection matrix, and we prepare the unbound beads, our counter selection matrix, before each experiment by transferring 20 μL of the beads into 500 μL of selection buffer. We then wash them three times with selection buffer by repeated centrifugation at a low speed at room temperature in a mini centrifuge and gently mixing the beads with fresh buffer each time. 3. Finally, add 100 μL of binding buffer to the beads and mix the annealed DNA library with the beads for counter-selection step. Incubate the tube on a gentle rocker-shaker for 15 min. 4. Meanwhile prepare the selection beads by adding 100 μL of binding buffer to 5 μL of the protein-bound Ni NTA beads 5. Centrifuge the counter selection tube at maximum speed and transfer the supernatant to the tube containing the protein bound beads. Incubate the beads for another 15 min on rocker-shaker. 6. Spin selection beads at maximum speed for 30 s and remove the unbound DNA. Wash the pellet with the beads with 1 mL of selection buffer and spin down the pellet after the wash. Repeat this step three times. 7. Provide the last wash after transferring the beads to a new tube (this will eliminate sequences that may be bound to the tubes). Add 200 μL of elution buffer and elute the protein along with the bound DNA library by incubating the tube on the rocker-shaker for 10 min. Pellet the beads by centrifugation, take out the supernatant, and transfer it to another tube. 8. Precipitate the DNA from the supernatant with 500 μL of 100% ethanol and spin at 21,000×g at 4°C for 30 min. Wash the pellet once with 70% ethanol and repeat the centrifugation. 9. Dry the DNA pellet, resuspend it in 20 μL water and set up PCR in a 100 μL volume using PCR primers specific for the
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library. Thirty cycles of PCR is carried out under the following conditions: 94°C for 1 min, 52°C for 45 s, 72°C for 30 s, and an additional 10 min at 72°C for final extension (see Notes 2 and 3). 10. Precipitate the PCR products with 250 μL of 100% ethanol, centrifuge the solution at 21,000 × g 4°C for 30 min and dry the pellet using a speedvac. Add 90 μL of 0.25 N NaOH to the pellet and heat the sample in heating block at 92°C for 15 min. After heating, the add 10 μL of 3 M sodium acetate to the tube and precipitate DNA with 250 μL of cold 100% ethanol (see Note 4). 11. Centrifuge the contents at 21,000 × g at 4°C for 30 min, remove the supernatant and dry the DNA pellet in a speedvac. 12. Prepare a 10% Urea−PAGE gel and prerun it at 40 mA for 15−20 min. Add 20 μL of the 1× gel loading buffer to each sample and heat DNA with dye for 2 min at 92°C. 13. Following the prerun, flush the urea out of the wells with a syringe and needle. Load the samples in the wells and continue running the gel until the xylene cyanol dye front (top dye front) migrate past two-third of the gel. 14. Wrap the gel using plastic wrap, as described previously, and detect the DNA by UV shadowing. The top band, which corresponds to the aptamer strand, is then excised with a sterile razor blade and placed into a 1.5 mL centrifuge tube. 15. Crush the band with a sterile pipette tip and add 700 μL of elution buffer to the gel. The tube is kept in a shaker overnight. 16. Steps 1−15 are repeated for about ten rounds (see Note 5). After ten rounds of selection, check the enrichment of the sequences for their ability to bind to their target ligand by conducting a gel shift assay or EMSA. The population should be radioactively labeled using a dNTP−dGTP mix containing [α-32P]-dGTP for PCR. The single-stranded DNA recovered following PCR is therefore radioactive and should be handled with all precautions. 3.3. Electromobility Gel Shift Assay
1. Cast an 8% native PAGE gel following the aforementioned protocol for preparing 10% denaturing PAGE gels. Allow the gel to polymerize for at least an hour prior to prerunning the gel in a cold room (4°C). 2. After an hour, set the plate in the gel-apparatus, gently take the comb out and flush the wells with water using a needle and syringe. Place the gel apparatus and power supply in the cold room.
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3. To the gel running apparatus, add native gel running buffer to the upper and lower chambers and start running the gel at 200 V. Prerun the gel until the effective milliamps of the gel drops to around 10 mA (it usually takes 1–2 h). 4. Meanwhile prepare the following binding mixture: 5× gel shift buffer – 4 μL Radioactive DNA – 1 μL (∼20 ng) Selection buffer – 14 μL Protein (10 μM stock) – 1 μL Prepare another tube for the DNA-only control by replacing the protein with an equal volume of selection buffer. The above protocol produces 0.5 μM of the protein of interest. However, as shown in the Fig. 2, the protein concentration can be varied, thus allowing the affinity of the selection pool for the proteins to be measured. 5. Before loading the samples into the native gel, add 6× DNA loading dye to one of the empty wells on the outside lanes. This allows for the progress of DNA migration to be tracked. 6. Run the gel at a constant current of 10 mA for another 3–4 h.
Fig. 2. Representative native gel-shift from a selected aptamer pool for protein binding. The DNA was radiolabeled by the incorporation of [α-32P]-dGTP in the PCR. The concentration of protein increases from right to left (i.e., Lane 1, no protein, Lane 2, 0.12 μM, Lane 3, 0.5 μM, Lanes 4, 5, and 6 are 1, 1.5, and 2 μM, respectively). The binding reactions were resolved on 8% native gel. The arrow depicts the DNA−protein complex, which runs slower than the DNA alone. Note the increase in intensity of the shifted band with increasing protein concentration.
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7. Stop the gel and carefully remove the plates. Mark the wells on the plate and, with the help of a scalpel, carefully open the plates. Since DNA samples on the gel are radioactive, it is important to work behind the beta-safety shields during all the operations. 8. Take a Whatman sheet and gently put it over the gel. Flip the plate and take the gel on the Whatman sheet. Cut the Whatman sheet just 2 cm more than gel size. 9. Place a piece of plastic wrap over the gel and keep it in the gel dryer for 1.5 h at 80°C. 10. After the gel is dried, place the gel in the phosphorimager cassette and expose the gel to the screen overnight. 11. Scan the gel next day using the Storm Scanner and evaluate the results. The results from a typical successful gel-shift assay are shown in Fig. 2. 12. The test for enrichment should be carried out after every alternate round of selection after seeing a signal on the native gel. 13. The progress of the enrichment should be monitored carefully and a decision is made to clone the population after no further enrichment can be seen over a couple of rounds of selection. 3.4. Cloning and Sequencing the Selected Pool
1. Prepare for the final PCR using the protocol provided in the materials section with following cycling conditions: 92°C for 1 min, 52°C for 45 s, 72°C for 30 s. After these cycles, final extension was carried out at 72°C for 10 min in order for the ends of the PCR products to be completely synthesized. 2. It is advisable (but not necessary) to purify the PCR product on an agarose gel (see Note 6). Gel purification can prevent the cloning of the primer dimers into the TOPO vectors, thus increasing the chances of cloning in the desirable sequences emanating from the library rather than random sequences and products that arose due to false priming. Gel purification can be done using commercially available kits from Qiagen, Promega or Invitrogen. The PCR product is ligated to the prepared vector following the protocol provided by the InsTAclone™ PCR Cloning Kit (MBI Fermentas). 3. Set up the ligation as follows: Vector pTZ57R/T – 3 μL (approximately 0.2 pmol ends) Purified selected population – 4 μL (0.6 pmol ends) 5× ligation buffer – 6 μL MilliQ nuclease free water – up to 29 μL T4 DNA ligase – 1 μL (5 units/μL) Incubate at room temperature (22°C) for 3–4 h or overnight at 16°C.
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4. Inoculate appropriate bacterial host strain, such as E. coli DH5α or JM109, in C-medium and let the cells grow in a 37°C shaker overnight. 5. Inoculate 2 mL of C medium with 50 μL of overnight culture and let it grow at 37°C for 30 min. 6. Meanwhile, thaw T solutions “A” and “B” on ice. Mix 500 μL of each and keep the mixture on ice. Pellet cells at 8,000 rpm (7,100 × g) for 5 min at 4°C. Decant the supernatant and add 400 μL of the T solution mixture to the pellet. Resuspend the pellet in T solution by repeatedly pipetting the solution and incubate the cells on ice for 5 min. 7. Pellet the cells again at 7,100 × g for 5 min at 4°C and decant the supernatant. Quickly add 200 μL of the T solution mix and incubate on ice for 5 more minutes. 8. Add 3 μL of the ligation reaction mixture to a pre-chilled tube and 1 μL of the vector control to another, which allows us to evaluate the transformation efficiency of the competent cells. Approximately 100 μL of the T solution with cells is transferred to each tube, which is then incubated on ice for 5 min. To transform the plasmid/vector into the cells, heat shock the cells at 42°C for 45 s. Two minutes after heat shocking, incubate the cells on ice and add 900 μL of the C medium to both tubes. The tubes are then shaken at 37°C for 30 min (see Note 6). 9. Spread 200 μL cells transformed with the ligation reaction mixture to five LB-ampicillin agar plates and 100 μL of cells from the control on one LB-ampicillin agar plate. Incubate them at 37°C for ∼12 h (see Note 7). 10. Following overnight incubation, look for colonies on both sets of plates. The control plate should have much more colonies than ligation plates. 11. Prepare the colonies from the ligation plate sequencing. Using sterile tooth picks, inoculate colonies in tubes containing 3 mL of LB broth with 50 μg/mL ampicillin. Grow these colonies in the shaker over-night at 37°C. Next morning, pellet the cells at the 13,000 rpm/2 min/4°C and process them for sequencing. We send the E. coli pellets for direct sequencing using M13 reverse or M13 forward primers (see Note 8). 12. Sequencing results are analyzed and vector sequences are deleted from the total sequence so as to retain only the data that represent the selected random sequences. We edit the sequences using Chromas software, which is available from (http://www.technelysium.com.au/chromas.html) and use BioEdit sequence alignment editor (http://www.mbio.ncsu. edu/BioEdit/bioedit.html) to align sequences and derive consensus sequences. The sequences are aligned based on their conservation and classified into different categories.
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Fig. 3. Binding curve of a DNA aptamer that is selected to bind a protein enzyme known as Bcs1. Bcs1 is a CDP-ribitol synthase (a bifunctional enzyme that catalyzes two consecutive chemical reactions leading to the formation of CDP-ribitol: the reduction of ribulose 5-phosphate using NADPH, and cytidylyltransfer using CTP) from Haemophilus influenzae. TarIJ is the corresponding CDP-ribitol synthase from Staphylococcus aureus. Radioactively labeled DNA aptamer (∼0.1 nM) was incubated with 0, 25, 50, 75, 100, and 200 nM of each enzyme in the selection buffer and the mixtures were analyzed using 8% nondenaturing polyacrylamide gel. The fraction of radioactivity in the shifted DNA band is calculated and plotted against the protein concentration.
13. The individual sequences can be tested for their affinity for the target protein by various methods like filter binding assay, gel-shift, and surface plasmon resonance. The individual sequences from each class can be synthetically prepared from DNA synthesis facilities and tested for their ligand-binding capability. A representative binding curve from which the binding affinity can be estimated is shown in Fig. 3. 14. As a second step to characterize the aptamers, the primer domains from the library can be deleted from main aptamer and truncated aptamers are tested for ligand binding capability. We use M-fold program (available from http://www. bioinfo.rpi.edu/applications/mfold/) to predict the secondary structures of each aptamer (10). The shortened and optimized aptamer sequences can be decorated with fluorophores for custom biosensing applications.
4. Notes 1. A number of methods for partitioning selected sequences from the background, which are chosen empirically by researchers, are available. For the selection of aptamers against proteins,
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the most convenient method is the affinity column using proteins that are engineered to contain tags such His-tag, Glutathione S-transferase (GST), and Intein-Chitin Binding Domain (CBD) (11). If the protein does not contain such a tag, separation of bound and unbound DNA molecules can be carried out by using nitrocellulose filters (Millipore HWAP), which will retain the protein−DNA library complex while letting the unbound DNA pass through. Other methods for separating ligands without tags include capillary electrophoresis (12), gel electrophoresis (13), sedimentation centrifugation (14), and flow cytometry (15). The in vitro selection technique has been automated recently (16) thus minimizing human interventions and making the aptamer selections against highly potent toxins and biowarfare agents a reality. 2. The selection population can be made radioactive by either 5′ end-labeling the purified DNA using [γ-32P] ATP with enzyme T4 polynucleotide kinase or by incorporating [α-32P]-dGTP during the PCR amplification. Unless specifically needed, we use [α-32P]-dGTP in PCR for labeling the population. 3. During PCR amplification of selected DNA, it is important not to over-amplify the sequences. If this goes unchecked then it may attribute to background binding and would require more selection cycles to overcome the background noise and amplify the signal. We find the real-time PCR monitoring of PCR progress to be useful for this purpose. We stop the PCR cycles 2–3 cycles after the monitoring curve reaches a plateau as shown in Fig. 4, thus avoiding the over-amplification and spurious PCR products. 4. The single-stranded DNA from the PCR can be generated in a couple of ways. This can be either achieved by labeling the reverse primer at 5′ end by biotin and separating the sense strand after PCR on streptavidin-coated magnetic beads. We find that resolving the DNA strands on PAGE using the approach given in Subheading 3.2 gives a much cleaner product and does not contribute to background signal. 5. After initial few rounds of counter-selection, it is trivial to counter-select in each round. After observing a signal on native gel-shift, the selection can even be switched onto a gel-based selection, thereby obviating the need for a bead-based selection. The gel-shifted band, which has the sequences bound to the protein, is cut out, and after elution, DNA sequences can be PCR amplified for the next round. The use of Taq polymerase for PCR can introduce a few mutations to the selected DNA species. This happens due to the low fidelity, error-prone properties of the DNA
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Fig. 4. Real time PCR progress. The bottom curve (unfilled circles) represents PCR carried out with the control sample which lacks the template; the upper curve (filled circles) represents PCR conducted with the selected population in this example. The PCR signal saturates around 21st cycle. At this point, the reaction is stopped.
polymerase. However, due to the selection pressure imposed on the population, mutations that can be tolerated are either neutral mutations (which have no effect on ligand binding) or beneficial mutations (which ameliorate the ligand binding ability of the aptamer). In fact, some researchers carrying out in vitro selection experiments intentionally to opt for mutagenic PCR during the selection rounds with an aim to increase the diversity of the sequences and achieve functional nucleic acids with better affinity (17). 6. It is advisable to have three transformation tubes for every cloning experiment. One is test sample “L” tube, second is positive control “C” tube, and third is the “no DNA” control. Control “C” tube will tell the researcher about the transformation efficiency of the competent cells. The “no DNA” control should not have any colonies on the LB-ampicillin plate. It indicates that there was no contamination of the host E. coli cells and the ampicillin drug is acting as expected. The degradation of ampicillin in the plates is indicated if the “no DNA” control plate shows a lot of colonies. 7. Care should be exercised not to leave the LB-ampicillin plates in the 37°C incubator for too long. Incubation of plates for extended times results in the formation of satellite colonies around recombinants due to degradation of ampicillin by secreted beta-lactamase. Satellite colonies make it hard to identify the authentic recombinants.
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8. Depending on the resources in hand, it is advisable to sequence as many colonies as possible. For every selection we usually sequence more than 50 colonies and categorize them. An early indication of a successful selection is given by the conservation of sequence domains, which help in manual categorization of the sequences.
Acknowledgments The aptamer research in the Li lab is funded by Genome Canada through Ontario Genomics Institute, the Canadian Institutes of Health Research, and Natural Sciences and Engineering Research Council of Canada. YL holds a Canada Research Chair.
References 1. Southern, E. M. (1975). “Detection of specific sequences among DNA fragments separated by gel electrophoresis.” J Mol Biol 98, 503–517 2. Schena, M. and D. Shalon, et al. (1995). “Quantitative monitoring of gene expression patterns with a complementary DNA microarray.” Science 270, 467–470 3. Bunka, D. H. and P. G. Stockley (2006). “Aptamers come of age − at last.” Nat Rev Microbiol 4, 588–596 4. Tuerk, C. and L. Gold (1990). “Systematic evolution of ligands by exponential enrichment: RNA ligands to bacteriophage T4 DNA polymerase.” Science 249, 505–510 5. Ellington, A. D. and J. W. Szostak (1990). “In vitro selection of RNA molecules that bind specific ligands.” Nature 346, 818–822 6. Nutiu, R. and Y. Li (2005). “Aptamers with fluorescence-signaling properties.” Methods 37, 16–25 7. Nutiu, R. and Y. Li (2003). “Structureswitching signaling aptamers.” J Am Chem Soc 125, 4771–4778 8. Nutiu, R. and Y. Li (2004). “Structureswitching signaling aptamers: transducing molecular recognition into fluorescence signaling.” Chemistry 10, 1868–1876 9. Navani, N. K. and Y. Li (2006). “Nucleic acid aptamers and enzymes as sensors.” Curr Opin Chem Biol 10, 272–281
10. Zuker, M. (2003). “Mfold web server for nucleic acid folding and hybridization prediction.” Nucleic Acids Res 31, 3406–3415 11. Dobbelstein, M. and T. Shenk (1995). “In vitro selection of RNA ligands for the ribosomal L22 protein associated with Epstein-Barr virus-expressed RNA by using randomized and cDNA-derived RNA libraries.” J Virol 69, 8027–8034 12. Berezovski, M. and A. Drabovich, et al. (2005). “Nonequilibrium capillary electrophoresis of equilibrium mixtures: a universal tool for development of aptamers.” J Am Chem Soc 127, 3165–3171 13. Smith, D. and G. P. Kirschenheuter, et al. (1995). “In vitro selection of RNA-based irreversible inhibitors of human neutrophil elastase.” Chem Biol 2, 741–750 14. Zhang, F. and D. Anderson (1998). “In vitro selection of bacteriophage phi29 prohead RNA aptamers for prohead binding.” J Biol Chem 273, 2947–2953 15. Blank, M. and T. Weinschenk, et al. (2001). “Systematic evolution of a DNA aptamer binding to rat brain tumor microvessels. selective targeting of endothelial regulatory protein pigpen.” J Biol Chem 276, 16464–16468 16. Cox, J. C. and A. D. Ellington (2001). “Automated selection of anti-protein aptamers.” Bioorg Med Chem 9, 2525–2531 17. Cadwell, R. C. and G. F. Joyce (1994). “Mutagenic PCR.” PCR Method Appl 3, S130–S140
Chapter 23 Immobilization of Biomolecules onto Silica and Silica-Based Surfaces for Use in Planar Array Biosensors Lisa C. Shriver-Lake, Paul T. Charles, and Chris R. Taitt Summary Several methods are described in which a biological recognition molecule – a critical element in any biosensor – is immobilized onto a silica or silica-based sensing substrate. Although several variations are described, the methods for covalent immobilization share a common theme and are generally composed of three steps: modification of the surface to add specific functional groups (using appropriate silanes or an amine or carboxyl-containing hydrogel), covalent attachment of a crosslinker through one of its reactive moieties, and finally, covalent linking of the biomolecule (recognition element) to the remaining reactive moiety of the crosslinker. One final method is presented in which the surface is modified with a highly hydrophobic silane and a glycolipid recognition element immobilized, essentially irreversibly, by hydrophobic interactions. All of the methods described have been successfully used to immobilize biological recognition molecules onto sensing surfaces, with full functionality in biosensor-binding assays. Key words: Silanization, Crosslinking, Biomolecules, Immunoassay, Array, Detection, Biosensor.
1. Introduction A key component of any biosensor is the biological molecules used to recognize and quantify the target of interest. These recognition species can be enzymes, antibodies, artificial and natural receptors, peptides, carbohydrates, or nucleic acids. The recognition molecules are chosen for their specificity and affinity for the analyte, as well as their ability to convert a recognition event into a measurable change in physical properties of the system. As they are immobilized onto the sensing surface in most sensor systems,
Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_23
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the means used to attach these biomolecules to the surface is a critical factor in sensor development and optimization. Retention of antigenicity, binding affinity, and/or enzymatic activity by the recognition molecule not only requires functional integrity and correct orientation of the immobilized species but may also be influenced by steric hindrance, molecular freedom of movement, and diffusional barriers. This chapter describes several methods to attach different biomolecules to silica or silica-based surfaces, although other surfaces such as indium tin oxide, tin oxide, tantalium pentoxide, and platinum may also be suitable for similar treatment (1–6). Subheadings 3.2.1 and 3.2.2 describe direct covalent linking of a biomolecule to the surface. The biomolecule may be the recognition species itself (e.g., antibody, peptide, enzymes), or may be an intermediary species, such as avidin or its derivatives, for subsequent noncovalent linking to a biotin-labeled molecule. Use of an intermediary species such as avidin enables preparation of a generic surface, with the end user able to change the (biotin-labeled) recognition species at will. An additional immobilization method (Subheading 3.2.3) utilizes hydrophobic interactions between a long chain (C18) silane layer on the sensor surface and the hydrophobic moiety of glycolipids. A separate section has been included describing preparation of hydrogel surfaces for biomolecule immobilization (Subheading 3.2.5); these surfaces are capable of binding higher quantities of immobilized biomolecules, providing higher avidity “capture” surfaces in detection assays. In addition, Subheading 3.2.4 describes general methods for creating patterned arrays of biomolecules on treated surfaces. These arrays can be used to screen samples for the presence of many targets of interest using any one of a number of multianalyte sensors described in the literature (7–19). The NRL Array Biosensor described in a later chapter is one such system that takes full advantage of the two-dimensional nature of the patterned substrate to enable simultaneous testing of multiple samples for multiple targets.
2. Materials 2.1. Cleaning Procedures (for Subheading 3.1)
1. Coplin jars: Eight slots to process batches of 16 slides. 2. Microscope slides were purchased from A.J. Daigger, Vernon Hills, IL. 3. HCl: Methanol solution: Mix hydrochloric acid (HCl) and methanol 50/50 v:v. For Coplin jars with eight slots, 100 mL is sufficient to cover the microscope slides. Personnel should
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wear safety goggles or full face shield, a lab coat, and acidresistant gloves. 4. Sulfuric acid: As this is a strongly acidic material, personnel should wear safety goggles or full face shield, a lab coat, and acid-resistant gloves. 5. KOH solution: Dissolve 10 g potassium hydroxide in 100 mL methanol. This is a very basic solution. Personnel should wear safety goggles or full face shield, a lab coat, and base-resistant gloves. 2.2. Mercapto-Modified Surfaces (for Subheading 3.2.1)
1. Glove bag was obtained from the Instruments for Research and Industry (I2R, Cheltenham, PA). The glove bag should be located in a chemical hood to reduce exposure to silane odors and toluene fumes. 2. High purity toluene and methanol were purchased from Aldrich Chemical Co., St. Louis, MO. 3. MTS/toluene solution: This solution should be prepared in a glove bag under nitrogen just prior to use to preserve the reactive thiols. Mix 1 mL mercaptopropyltriethoxysilane (MTS, Fluka Chemical Corp., St Louis, MO) with 49 mL toluene to make a 2% MTS solution (see Notes 1 and 2). 4. GMBS/EtOH solution: Dissolve 12.5 mg N-[g-maleimidobutyryloxy]succinimide ester (GMBS, Fluka Chemical Corp., St. Louis, MO) in 250 μL dimethyl sulfoxide (DMSO). Absolute ethanol (43 mL, EtOH, Warner-Graham, Cockeysville, MD) is added to this solution. This solution is prepared immediately before use, as the reactive groups degrade (see Note 3). GMBS is moisture sensitive and must be stored at 4°C in a desiccator. 5. Phosphate buffered saline, pH 7.4 (PBS, Sigma Chemical Co., St. Louis, MO, P-3813): Prepared with 18 mW water from Millipore Milli-Q system per manufacturer’s directions. 6. NeutrAvidin (Pierce Chemical Co., Rockland, IL): A stock solution of 10 mg/mL NeutraAvidin in PBS is prepared by adding 1 mL PBS to 10 mg NeutrAvidin vial. Just prior to use, 120 mL of the 10 mg/mL stock is added to 40 mL PBS. 7. Biomolecule solution (thiol–amine linking): 10–50 mg/mL (antibodies, enzymes) or 20–250 mg/mL (peptides) in PBS (see Note 4).
2.3. Amine-Modified Surfaces (for Subheading 3.2.2)
1. Amino silane solution: Mix 45 mL methanol and 5 mL deionized water. Add one drop of acetic acid. To this, add 1 mL of 3-aminopropyl triethoxysilane (Gelest, Inc., Tullytown, PA). Prepare immediately before use. 2. BS3 solution: Dissolve 1.43 mg bis(sulfosuccinimidyl) suberate (BS3, Pierce, Rockland, IL) in 50 mL of 10 mM sodium
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phosphate, pH 6.0; the final concentration of this solution is 50 mM (see Note 5). Prepare immediately before use. 3. GMBS/EtOH: Dissolve 12.5 mg N-succinimidyl-4-maleimidobutyrate (GMBS, Pierce, Rockland, IL) in 0.25 mL anhydrous dimethyl sulfoxide (DMSO); this solution can be kept for several hours. Add GMBS/DMSO solution to 43 mL absolute ethanol and mix vigorously. Once diluted in ethanol, use the GMBS solution immediately (see Note 3). GMBS is moisture-sensitive and must be stored in a dessicator at 4°C. Store opened stock bottle of EtOH in glove bag. 4. Biomolecule solution (amine–amine linking, Subheading “Amine–Amine Homobifunctional Crosslinkers”): 5–25 mg/ mL in PBS for proteins. For peptides and carbohydrates where achieving maximal surface density is important, concentrations may be as high as 1 mg/mL (in PBS). 5. Biomolecule solution (amine–thiol linking, Subheading “Amine–Thiol Crosslinking Heterobifunctional Crosslinkers”): 5–25 mg/mL (proteins) or 10–50 mg/mL (peptides and thiol-derivatized sugars) in 10 mM sodium phosphate/10 mM NaCl, pH 7.5. Tris(2-carboxyethyl)phosphine hydrochloride (TCEP, Pierce) is added to the protein/peptide/sugar solution at a 0.8 molar equivalent (11) (see Note 6). 2.4. Hydrophobic Silane-Modified Surfaces (for Subheading 3.2.3)
1. OTS solution: Add 0.4 mL octadecyltrichlorosilane (OTS, Fluka, Ronkonkoma, NY) to 100 mL anhydrous toluene. Use immediately (see Note 7).
2.5. Patterning with PDMS (for Subheading 3.2.4)
1. Plexiglas molds for PDMS: Plexiglas plates (»1/2 in. thick) are milled with a numerically controlled milling machine (Techno Isel) to form the molds for the patterning and assay PDMS flow chambers (Fig. 1). Specific measurements can be found in Rowe et al. (12) Fins approximately 1 mm wide and tall are milled in the Plexiglas to form the channels in the PDMS through which the solutions will flow.
2. Glycolipid: 1 mg/mL ganglioside GM1, GT1b, or other glycolipid in 10 mM bicarbonate, pH 8.0.
2. PDMS patterning gasket: NuSil MED-4011 (Nusil Silicone Technology, Carpinterai, CA). Mix Part A and Part B at a 10:1 w/w ratio in a large plastic disposable beaker. The mixture is degassed in a vacuum oven at room temperature (RT) until all the bubbles are removed (»30 min). The PDMS is poured into the Plexiglas mold (see Note 1). This mold is placed in the vacuum for further degassing until all the bubbles are at the surface, because bubbles near the fins can cause problems in the final PDMS flow chamber. After degassing, the mold is left at room temperature to cure, or it may be
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Fig. 1. Plexiglas and PDMS pieces for patterning. (a) Plexiglas mold for PDMS patterning gasket. The number of fins shown is 6 but up to 15 patterning channels have been designed in the same region. (b) Shown clockwise from the upper left are the PDMS patterning gasket, bottom piece of Plexiglas patterning chuck, screws, microscope slide with number etched, and the top piece of Plexiglas patterning chuck.
placed in a 60°C oven for 30 min. Some caution is advised with the oven method, as the Plexiglas mold can warp at higher temperatures. After curing, the PDMS is removed from the mold, trimmed, and washed with soap to remove any residue. For many applications, it is also useful to incubate the PDMS in 1% BSA prior to use to prevent nonspecific adsorption of molecules to the walls of the channel. 3. Patterning chucks: A chuck made of two Plexiglas plates (Fig. 1) to apply pressure evenly on the PDMS gasket was designed such that one plate has a recess to fit the slide and six tapped holes around the perimeter. The other plate is the same size and has holes that line up with the tapped holes. This plate also has access holes for the syringe needles, used to pierce the PDMS at both ends of the flow channels, for flowing the reagents. 4. Phosphate-buffered saline, pH 7.4 (PBS): Sigma Chemical Co., St Louis, MO, P-3813. Dissolve 1 packet in 1 L 18 mW water. 5. PBSTB: Phosphate buffered saline pH 7.4/ 0.05% Tween-20/ 1 mg/mL BSA. Dissolve 1 g bovine serum albumin (# A3912, Sigma Chemical Co, St Louis, MO) and 500 mL Tween 20 (Sigma Chemical Co.) in 1 L PBS to make the correct final concentrations. 6. Patterning buffer: 10 mM phosphate buffer, 10 mM sodium chloride, and 0.05% Tween 20, pH 7.4. 7. Biotin-labeled biomolecule for use with NeutrAvidin-coated slides: In sandwich immunoassays, 10–20 mg/mL biotinylated antibody in patterning buffer is routinely used for patterning. In competitive immunoassays, the biotinylated analyte concentration varies from 0.2 to 5 mg/mL.
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8. Unlabeled biomolecule for direct immobilization: 5–50 mg/mL (antibodies and other proteins) or 10–50 mg/mL (peptides and thiol-derivativized sugars) in patterning buffer. 9. Tuberculin syringes (1 mL) with 25 gauge needles. 2.6. Hydrogels (for Subheading 3.2.5)
1. MTPTS solution: Add 8.0 mL of 3-(trimethoxysilyl) propylmethacrylate (MTPTS) to a mixture of methanol (186 mL), water (5.4 mL), and acetic acid (0.6 mL). 2. DCDM solution: Add 4.0 mL of dichlorodimethylsilane (DCDM) to 196.0 mL of hexane. 3. Carboxy-hydrogel solution: Dissolve 60 mg galactose monomer, 6-acryloyl-b-O-methyl galactopyranoside (20), in 0.5 mL 18 mW Milli-Q water at a final concentration of 12% (w/v). Add this monomer solution to 30 mg of 2-acrylamidoglycolic acid monohydrate at a final concentration of 50% (w/w) of the galactose monomer concentration. Dissolve N,N-methylene bis-acrylamide (Bis) cross-linker at 13% (w/w) of the monomer concentration in 100 mL of 18 mW H2O. Heat briefly with heat gun to dissolve. Add carboxy - amine-hydrogel monomer solution to Bis crosslinker. Mix gently. Add 0.55 mg sodium persulfate and 2 mL of TEMED. Purge solution briefly with nitrogen. Use immediately. 4. Amine-hydrogel solution: Dissolve 110 mg of the galactose monomer, 6-acryloyl-b-O-methyl galactopyranoside in 0.5 mL of 18 mW H2O at a concentration of 22% (w/v). Add the monomer solution to 5.5 mg of N-(3-aminopropyl)methacrylamide at a 5% (w/w) of the sugar monomer concentration. Add 0.55 mg of N,N methylene bis-acrylamide cross-linker for a final 0.5% (w/w) of the monomer concentration. Add sodium persulfate (5.5 mL) from a 10 mg/mL solution and TEMED (2.0 mL) to the hydrogel solution. Each monomer solution is purged with a stream on nitrogen. Use immediately. 5. EDC/NHS solution: Weigh out 11 mg 1-ethyl-3-(3dimethylaminopropyl)-carbodiimide-HCl (EDC) and 28 mg N-hydroxysuccinimide (NHS) in separate vials. Keep dry. Immediately before use, pipette 0.75 mL 50 mM MES buffer, pH 6.0 into each tube and mix well. Combine the two aliquots into a single EDC/NHS solution and use without delay. Final concentrations of EDC and NHS are 40 mM and 80 mM, respectively. 6. BS3 solution: Dissolve 2.1 mg BS3 (Pierce, Rockland, IL) in 1.5 mL 10 mM sodium phosphate, pH 6.0; the final concentration is 2.5 mM. Prepare immediately before use. 7. PDMS patterning gasket (see Subheading 2.5 for preparation). 8. Phosphate-buffered saline, pH 7.4 (PBS). 9. Antibody solution: 100–300 mg/mL antibody in PBS.
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10. Blocking solution: Add 2 g bovine serum albumin (Sigma, # A3912) and 2 g casein (BDH Laboratory Supplies) to 80 mL PBS. Add 10 M NaOH, with stirring, until the pH is approximately 12 (see Note 8). Once solution is clarified, bring the pH back to neutral by adding (with stirring) first 6 M HCl, then 1 M HCl, dropwise to the solution. Once the pH is approximately 7.4, add PBS to bring the volume to 100 mL (see Note 9). 11. 1 mL tuberculin syringes and 1 mL tuberculin syringe barrels (without plunger), fitted with 1.5-in., 25-gauge needles. 12. Multichannel peristaltic pump, set to flow at a rate of 0.8 mL/ min. Inlet tubing is fitted with 25-gauge syringe needles, which will be used to connect the pump to the PDMS patterning gasket.
3. Methods For all the procedures described in this section, microscope slides are used as the silica-based support and are labeled with a diamond scribe away from the biomolecule immobilization region. Other silica-based surfaces have also been used. Two slides are placed backto-back into a Coplin jar with the etched numbers to the outside. 3.1. Cleaning
No matter which silanization method is employed, and cleaning of the solid support to generate reactive hydroxyl groups is critical for effective immobilization of biomaterials. There are several types of Si–OH groups that can form on silica surfaces. The geminal and isolated silanols are reactive, whereas the vicinal silanol and the siloxane groups are not (Fig. 2). If the surface is not properly cleaned of oils, dirt, detergents, etc., the reactive hydroxyl groups will not be formed and the silane will not be deposited in a uniform manner. Many comments against silanization methods concern the nonuniformity of the silane monolayer. The major contributors to such nonuniformity are inadequate surface cleaning and decomposure or
H
H
H O
O Si
Geminal silanol Fig. 2. Types of reactive surface hydroxyls.
O Si
Isolated silanol
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HCl/MeOH rinse
KOH/MeOH
H2SO4
rinse, dry
rinse
Boil rinse, dry
To silanization Fig. 3. Flow chart for cleaning methods.
polymerization of the silane. Two methods for cleaning silica-based supports are described; a flow chart for each cleaning method is shown in Fig. 3 (21, 22). 3.1.1. HCl/Methanol
1. Mark all slides using a diamond-tipped pencil. 2. Load slides into upright Coplin jar. Two slides can be placed back to back for each slot in the Coplin jar. 3. Mix 50 mL concentrated HCl with 50 mL MeOH in a large flask (250 mL). Pour HCl into premeasured MeOH in flask to avoid spattering of concentrated acid. Swirl flask until both components are thoroughly mixed and no schlieren lines are observed. 4. Pour HCl/MeOH mix over slides and incubate 30 min without mixing. 5. Remove HCl/MeOH mix from the Coplin jar and pour into a labeled glass bottle for waste disposal. Do not pour down the drain. 6. Rinse slides exhaustively with deionized water (no schlieren lines observed). 7. Dry slides individually under stream of nitrogen and place back into clean, dry Coplin jar. 8. Pour concentrated sulfuric acid over slides until slides are covered. Incubate for 30 min without mixing. CAUTION: Highly acidic. Use appropriate personal protective equipment. 9. Remove concentrated sulfuric acid from Coplin jar and pour into a labeled glass bottle for waste disposal. Do not pour down drain. 10. Rinse slides exhaustively with deionized water until no schlieren lines are observed. Rinse each slide individually with deionized water.
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11. Add hot 18 mW water (100°C) to the slides. Incubate for 15 min. 12. Dry each slide individually under a stream of nitrogen and place back into a clean dry Coplin jar. Each slide should look pristine. There should be no spots or cloudiness to the slide. If spots or cloudiness are present repeat from step 8. Slides should be cleaned no more than 3 days before use. 3.1.2. KOH/Methanol
1. Dissolve 10 g KOH in 100 mL methanol (MeOH) in a 250 mL flask. 2. While KOH is dissolving, etch appropriate number in upper right corner of each slide. 3. Place slides into upright Coplin jar. Two slides can be placed back to back for each slot in the Coplin jar. For metal coated slides the front is where the metal coating is located. 4. Pour KOH/MeOH over slides until slides are completely immersed. Incubate for 30 min without mixing. 5. Remove KOH/MeOH mix from Coplin jar and discard into an appropriate waste container. 6. Wash slides exhaustively with deionized water. Rinse each slide individually until no schlieren lines are visible. 7. Dry each slide individually under a stream of nitrogen and place back into clean dry Coplin jar. Each slide should look pristine. There should be no spots or cloudiness on the slide. If spots or cloudiness are present, repeat from step 3. Slides should be cleaned no more than 3 days before use.
3.2. Covalent Immobilization Procedures
Biomolecules such as antibodies, enzymes, and cells can be attached to silane-treated surfaces using the free thiols and/or amines on the surface and on the biomolecules. After treatment of clean silica surfaces with a thiol or amine-terminated silane, a homo or hetero-bifunctional crosslinker that possesses appropriate reactive groups can be used to attach a biomolecule to the surface. Figure 4 shows several illustrations of the general schemes for immobilization, described below.
3.2.1. Attachment of Biomolecules to ThiolModified Surfaces
In this section, only the heterobifunctional crosslinker that contains a maleimide for reacting to the thiols from the mercaptosilane and N-hydroxysuccinimidyl ester (NHS) to react with primary amines on the biomolecule will be described. A generalized flow chart for this procedure is shown in Fig. 5.
Thiol Silanization
In this step, the surface hydroxyl groups on the slide react with the ethoxy groups on the mercaptopropyltriethoxy silane resulting in a thiol-derivatized slide (22, 23)
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A
maleimide−linker−NHS (GMBS)
H2N−biomolecule linker−NHS
SH
S
S
H2N−biomolecule NHS−linker−NHS (BS3)
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Fig. 4. Immobilization methods. (A) Attachment of biomolecules onto thiol-derivatized surfaces (see Subheading 3.2.1 for experimental details). (B) Attachment of biomolecules onto amine-derivatized surfaces (Subheading 3.2.2) using amine–amine linking (upper pathway, Subheading “Amine–Amine (Homobifunctional Crosslinkers)”) and amine–thiol linking (lower pathway, Subheading “Amine–Thiol Crosslinking (Heterobifunctional Crosslinkers)”). (C) Noncovalent attachment of gangliosides/globosides onto hydrophobic surfaces (Subheading 3.2.3).
Clean Slide
MTS / toluene 60 min Rinse, dry G M B S / ethanol 30 min Rinse Incubate with biomolecule-NH2
Fig. 5. Protocol for attaching amine-containing biomolecules to thiol-coated surfaces.
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1. Place 16 cleaned, dried slides back-to-back in a clean, dry Coplin jar, with etched labels facing outward. 2. Place Coplin jar in glove bag. 3. For each batch of 16 slides, prepare 50 mL MTS/toluene solution immediately prior to use in the glove bag. Pour the silane/toluene mix over the slides and incubate for 1 h under nitrogen in the glove bag. 4. Remove the Coplin jar from the glove bag and place elsewhere in the chemical fume hood. 5. With forceps, remove each slide from MTS/toluene solution and rinse three times in toluene by swishing the slides 3–5 times sequentially in three 150 mL beakers filled with toluene. Caution needs to be taken when working with toluene. Proper protective gear needs to worn, and the work needs to be carried out in a chemical fume hood (see Note 10). 6. Dry each slide completely under a stream of nitrogen and place slides back-to-back in a clean, dry Coplin jar. The slides should still appear pristine. If not, the slides should not continue on to the next step as either the cleaning was insufficient or the silane has degraded. Since the thiols will rapidly oxidize within 24 h, the crosslinking step must be performed immediately after silanization. Heterobifunctional Crosslinking
During this step, the surface thiols react with the maleimide group of the crosslinker. The following step uses the linker’s NHS reactive group to bind to primary amines on the biomolecule. Crosslinking and all subsequent steps may be performed on the bench top, outside of the chemical hood. 1. For each batch of 16 slides, prepare 43 mL of GMBS/ethanol immediately prior to use. 2. Pour GMBS solution over slides and cover jar. Incubate for 30 min at room temperature. Time is critical at this step as the reactive groups on the GMBS degrade over time in the ethanol and air atmosphere. 3. Rinse the slides three times in deionized water by swishing the slides 3–5 times sequentially in three 150 mL beakers filled with water. 4. If immobilizing a single biomolecule (e.g., avidin, NeutrAvidin) over the entire slide surface, place slides back-to-back in a Coplin jar and add the 40 mL patterning buffer containing the biomolecule of interest. Incubate the slides overnight at 4°C. If using metal-coated slides, the NeutrAvidin or other biomolecule should be prepared in 10 mM phosphate buffer pH 7.4 and immobilized on the same side of the slide as the metal coating. Rinse the slides three times with PBS. They are stored in PBS (or in 10 mM phosphate pH 7.4 if metal-coated) at 4°C until patterning.
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5. If immobilizing multiple biomolecules in a patterned array, dry each slide under a stream of nitrogen and proceed immediately with patterning (Subheading 3.2.4). 3.2.2. Attachment of Biomolecules to Amine-Modified Surfaces
Biomolecules such as peptides, modified carbohydrates, Fab fragments, and other proteins can be attached to amine-modified surfaces using free thiols and amines (11, 24, 25). The cleaned surface is first treated with an amino silane, followed by a homo or heterobifunctional crosslinker possessing one or two aminespecific reactive groups and/or a thiol-specific reactive group (amine–thiol linking). After the N-hydroxysuccinimidyl ester (NHS) terminus of the crosslinker has been reacted with the surface-immobilized amines, the remaining reactive group of the crosslinker (maleimide or additional NHS) is reacted with thiol or amine moieties on the biomolecule. A generalized flow chart showing procedures for attaching the thiol and amine-containing biomolecules to amine-modified surfaces is shown in Fig. 6.
Amine Silanization
During this process, the hydroxyl groups on the surface of the slide react with the ethoxy groups of aminopropyl triethoxy silane resulting in an amine-derivatized slide (24). 1. Place 16 cleaned, dried slides back-to-back in a Coplin jar, with etched labels facing outward. 2. For each batch of 16 slides, prepare 50 mL silane/methanol/ HOAc mix immediately prior to use. Pour the silane/methanol/ HOAc mix over the slides and incubate for 1 h. 3. With forceps, remove each slide from silane/methanol/HOAc solution and rinse three times in methanol by swishing the slides 3–5 times sequentially in three 150-mL beakers filled with methanol. 4. Wash three times in deionized water (as above for methanol wash). 5. Dry each slide completely under a stream of nitrogen. Clean slide Amino silane/MeOH/HOAc 1 hour Rinse, dry Bake (100-1208C), 6-8 min
BS3, buffer pH6 30 min Rinse Biomolecule-NH2
Amine linking
Thiol linking
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Fig. 6. Protocol for attaching amine-containing biomolecules (left pathway) and thiolcontaining biomolecules (right pathway) to amine-coated surfaces.
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6. Place slides in a drying oven at 100–120°C for 6–8 min. Allow slides to cool to room temperature. 7. Place slides back-to-back in a clean, dry Coplin jar. Slides treated with amino silane may be kept under nitrogen for several weeks. These slides may be used for amine–amine crosslinking (Subheading Amine–Amine (Homobifunctional Crosslinkers) ) or amine–thiol crosslinking (Subheading Amine–Thiol Crosslinking (Heterobifunctional Crosslinkers) ) of biomolecules to the surface. Amine–Amine (Homobifunctional Crosslinkers)
During this process, the surface amine groups first react with one NHS reactive group of the crosslinker; this initial step is performed at pH 6 to minimize reaction of both NHS groups with surface amines (24). The second NHS group is later reacted with amines on the biomolecule to be attached at pH 7.5. Crosslinking and all subsequent steps may be performed outside of the chemical hood, on the bench top. 1. For each batch of 16 slides, prepare 100 mL of BS3 solution immediately prior to use. 2. Pour BS3 solution over slides and cover jar. Incubate for 30 min at room temperature. 3. Rinse three times in deionized water by swishing the slides 3–5 times sequentially in three 150-mL beakers filled with water. 4. If immobilizing a single biomolecule (e.g., avidin, NeutrAvidin) over the entire slide surface, place slides back-to-back in a Coplin jar containing 33 mL solution of the biomolecule of interest. 5. If immobilizing multiple biomolecules in a patterned array, dry each slide under a stream of nitrogen and proceed immediately with patterning (Subheading 3.2.4).
Amine–Thiol Crosslinking (Heterobifunctional Crosslinkers)
During this process, the surface amines first react with the NHS reactive group of the crosslinker (11, 25). A subsequent step uses the pendant maleimide to attach the biomolecule to the surface via its thiols. Crosslinking and all subsequent steps may be performed outside of the chemical hood, on the bench top. 1. For each batch of 16 slides, prepare 43 mL of GMBS solution immediately prior to use. 2. Pour GMBS solution over slides and cover jar. Incubate for 30 min at room temperature. 3. Rinse three times in deionized water by swishing the slides 3–5 times sequentially in three 150-mL beakers filled with water. 4. If immobilizing a single biomolecule (e.g., avidin, NeutrAvidin) over the entire slide surface, place slides back-to-back in a Coplin jar containing 33 mL solution of the biomolecule
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of interest. Incubate overnight at 4°C (proteins, peptides) or room temperature (thiol-derivatized sugars). Rinse three times with PBS and store at 4°C in PBS. 5. If immobilizing multiple biomolecules in a patterned array, dry each slide under a stream of nitrogen and proceed immediately with patterning (Subheading 3.2.4). 3.2.3. Attachment of Lipophilic Biomolecules to Hydrophobic SilaneTreated Surfaces
Gangliosides, globosides, and other lipid-linked biomolecules can be immobilized onto silica or silica-based substrates using hydrophobic interactions. A long-chain hydrophobic silane is used to coat the slide surface, in essence, creating a monolayertype structure into which the lipid moieties can intercalate. After incubation of the glyolipid with the silanized surface, the slides are cured, essentially locking the intercalated lipid chains in place. Ganglioside and globoside-patterned surfaces produced in this manner have been used to detect toxins (26, unpublished). A flowchart illustrating the procedure is shown in Fig. 7.
Hydrophobic Silanization
During this process, the hydroxyl groups on the surface of the slide react with the trichloro groups of the OTS, resulting in formation of a monolayer on the slide surface. 1. Place 16 cleaned, dried slides back-to-back in a Coplin jar, with etched labels facing outward. 2. For each batch of 16 slides, prepare 100 mL OTS solution and pour the silane/toluene mix over the slides. Incubate for 1 h under nitrogen in a glove bag. 3. Remove the Coplin jar from the glovebag and place elsewhere in chemical fume hood. 4. With forceps, remove each slide from the OTS solution and rinse three times in toluene by swishing the slides 3–5 times sequentially in three 150-mL beakers filled with toluene.
Clean Slide
OTS / toluene 60 min Rinse, dry Patterning with glycolipid overnight Rinse all channels, dry Bake (1208C), 20 min
Fig. 7. Procedure for immobilizing lipid-linked biomolecules to glass slides treated with hydrophobic silane.
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5. Dry each slide completely under a stream of nitrogen. 6. Place slides back-to-back in a clean, dry Coplin jar. Slides treated with OTS can be stored in a sealed jar (in air) at room temperature for up to a week without measurable changes in surface characteristics. Glycolipid Immobilization
During this process, glycolipid solution is incubated with the OTS-treated slide and the lipid moiety allowed to intercalate into the hydrophobic layer. 1. Rinse each slide three times in water. Place into patterning templates and pattern with glycolipid as described in Subheading 3.2.4. Let incubate overnight in the presence of glycolipid solution. 2. Rinse each channel with 1 mL deionized water under flow. 3. Remove slides from PDMS patterning template. 4. Rinse each slide three times by swishing 3–5 times sequentially in three 150 mL beakers filled with water. 5. Dry under a stream of nitrogen. 6. Bake each slide for 20 min in a drying oven at 120°C (see Note 11). Allow to cool to room temperature. Store dry at 4°C for up to one week.
3.2.4. Creation of PDMS Patterned Arrays
For multianalyte sensing, different biotinylated recognition molecules can be immobilized in each of the patterning channels. Figure 8 shows a fully assembled unit for patterning a single slide. For many of our studies, a positive control using an irrelevant biomolecule (usually anti-chicken IgY) is patterned in the outermost patterning channels and a negative control (buffer) is injected into one of the middle channels. Place slide on bottom piece of patterning chuck with etched labeled side up. 1. Place PDMS patterning gasket over the distal end (away from etched label) of the slide with channels being formed by slide and PDMS. 2. Place top piece of patterning chuck over PDMS. Finger tighten four screws (each corner) evenly. The screws should be finger tight; over-tightening will result in slide breakage; too loose and the channels leak into each other. 3. Syringes with needles and without plunger are placed on one end of each channel as an effluent reservoir. 4. Using a 1 mL syringe, inject 60–100 mL solution containing the biotinylated biomolecule (NeutrAvidin-coated surfaces) or unlabeled biomolecule (activated surface) in the patterning buffer into the appropriate channel. 5. Incubate the biomolecule with the slide overnight at 4°C.
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Fig. 8 Fully assembled unit for patterning. Solutions containing capture antibodies are in the closed syringes on the left side. Syringes on right side are inserted to allow flow through the chambers and are used as effluent reservoirs. Reprinted from ref. 10, with permission from Elsevier
6. Rinsed each channel with 1 mL PBSTB followed by air using the syringe format in step 5. 7. Remove slide from the patterning PDMS and chuck and insert into a 50 mL centrifuge tube containing 10 mg/mL bovine serum albumin in phosphate buffer, pH 7.4, for 20 min to block the slide. 8. Rinse slide with 18 mW water and dry with nitrogen. The dried patterned slide is stored at 4°C until use. 3.2.5. Preparation and Patterning of Hydrogel Thin Films
Recently, hydrogels have been used as an alternative substrate for the immobilization of biomolecules, because of their potential advantages over standard two-dimensional surfaces: increased capacity for immobilized species, low background in fluorescencebased assays, and a network conducive for affinity experiments. The following sections describe methods to prepare slides for casting of hydrogels, preparation of two types of hydrogels (amine and carboxylate-terminated), and direct covalent immobilization of biomolecules within the hydrogel matrix. Hydrogels prepared in this manner are amenable to use in biosensor assays. Figure 9
Immobilization of Biomolecules onto Silica and Silica-Based Surfaces Clean slides
Clean slides
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DCDM / hexane 1 hour Rinse, dry
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COOH hydrogel
NH2 hydrogel
BS3, phosphate pH 6 30 min Rinse Biomolecule, PBS 1 hour Rinse Repeat 5 times
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Rinse, block
Fig. 9. Procedure for immobilizing biomolecules inside COOH-hydrogels (lower left pathway) or amine-hydrogels (lower right pathway). Procedures for preparation of the top and bottom slides for casting the hydrogel slabs are shown in the upper right and left pathways, respectively.
shows a flow chart with the entire process of coating the top and bottom slides, preparing the hydrogel slab between the top and bottom slides, and patterning each type of gel. Treatment of Slides with Hydrophobic and Methacrylate Silanes
For each hydrogel prepared, two glass slides are modified using different silanes to impart differential chemical and physical properties. The slide onto which the hydrogel is cast is treated with MTPTS. The second slide, which forms the top layer of the hydrogel casting assembly, is functionalized with a silane with nonreactive pendant moieties (DCDM silane). This latter slide will eventually be removed from the assembly, leaving a hydrogel covalently attached to the MTPTS-treated slide. 1. Divide 32 acid cleaned glass slides into two groups for respective silane treatment. 2. Inscribe 16 slides with number 1–16 in top right corner of slide. 3. Inscribe the remaining 16 slides A–P in top right corner of slide. 4. Treat slides 1–16 with a MTPTS solution and incubated for1 h at 60°C. 5. Rinse MTPTS-treated slides three times with methanol and dry with a stream of nitrogen.
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6. Bake MTPTS-treated slides for 4 min at 120°C (scribed number side up). 7. Treat slides A–P with a DCDM solution for 1 h at room temperature. 8. Rinse DCDM-treated slides with hexane then dried with a stream of nitrogen. 9. Bake DCDM-treated slides for 4 min at 120°C (scribed number side up). 10. Store desiccated until further use. Preparation of Hydrogel Slabs
Hydrogel formation is accomplished via a free radical polymerization process using the initiator sodium persulfate and catalyst TEMED. The hydrogel slab is formed by creating a sandwichtype assembly composed of the MTPTS-treated slide on the bottom, the hydrogel matrix in the middle, and the DCDM-treated slide on the top. The hydrophobic nature of the DCDM-treated slide (prepared above) helps reduce the stress and shear forces upon removal from the hydrogel film, minimizing tearing or delamination of the hydrogel from the MTPTS-treated surface. Attachment of the poly(acrylate)-acrylamide film to the MTPTS-treated surface is accomplished though free radical polymerization of the polymer network and the pendant vinyl group of the MTPTSsurface. Teflon tape, which serves as the spacer between the two slides, provides control of the hydrogel polymer thickness. 1. Place the methacrylate-treated slide onto a horizontal surface and form “spacers” at each end of the slide using Teflon tape or commercial spacers. 2. Pipette 110 mL of the carboxy or amine-hydrogel solution onto the methacrylate-treated slide. Place the dichlorodimethyl (DCDM) silane-treated slide on top of the liquid, forming a sandwich-type assembly. 3. Place the assembly into an inert atmosphere (nitrogen). Polymerize overnight at room temperature. 4. Remove the DCDM-treated slide (top slide) from the MTPTStreated slide. This will reveal a thin film of highly crosslinked sugar polyacrylate hydrogel containing a carboxyl or amine-terminated moiety covalently attached to the MTPTS-treated surface. 5. Immerse the slides briefly (1.0 min) in Milli-Q water and airdry at room temperature. Store hydrogel slides semihydrated at 4°C until further use.
Immobilization of Antibodies Within Hydrogel Slabs
Biomolecules are covalently immobilized onto amine and carboxylic acid-modified hydrogel films in a method similar to that described for two-dimensional surfaces above (Subheadings 3.2.1 and 3.2.2). The key difference in the method used for
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hydrogels is the requirement for multiple cycles of cross-linking and attachment of biomolecules. For amine-terminated hydrogels, crosslinking occurs via the homobifunctional crosslinker BS3, whereas for carboxylate gels, EDC/NHS coupling chemistry is used. However, the procedure is identical for both types of hydrogels – i.e., 30 min crosslinker, wash, 60 min antibody, wash, repeat. Only the crosslinker solutions are changed: BS3 for amine hydrogels, EDC/NHS for carboxylate gels (step 5, below). 1. Place hydrogel-coated slide on bottom piece of patterning chuck with number side up. 2. Place PDMS patterning gasket over the distal end of the slide, ensuring that all channels of the patterning gasket are within the boundaries of the hydrogel slab. 3. Place top piece of patterning chuck over PDMS. Fingertighten four screws (each corner) evenly. The screws should be finger tight; over-tightening will result in slide breakage or damage to the hydrogel, whereas undertightening will result in leakage. 4. Place a syringe barrel (without plunger, fitted with a 25-gauge needle) at one end of every channel for use as a reservoir. These reservoirs will remain in place throughout steps 5–15. 5. Using a 1 mL syringe, inject 60–100 mL freshly prepared BS3 solution (amine-terminated gels) or EDC/NHS solution (carboxylate gels) into each channel. 6. Incubate for 30 min. 7. Hook each slide up to the peristaltic pump. One end of each channel will have a syringe barrel reservoir (see step 4, above), while the other end is hooked to the pump’s inlet tubing via a syringe needle. 8. Start the pump (0.8 mL/min flow rate), pulling the BS3 or EDC/NHS solution from each channel. Evacuate all channels. 9. Pipette 1 mL PBS into the syringe-barrel reservoir of each channel and continue pump flow until the entire volume has flushed through the channel and each channel is filled with air. 10. Unhook the slide from the pump. 11. Using a 1 mL syringe, inject 60–100 mL Antibody Solution into each channel. 12. Incubate for 60 min. 13. Using the same syringe, remove Antibody Solution from each channel and store on ice for subsequent rounds of patterning.
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14. Hook the slide up to the peristaltic pump, and rinse each channel with PBS, as in steps 7–10, above. This completes 1 round of patterning. 15. Repeat steps 5–14, for a total of six rounds of patterning. The final (sixth) incubation with Antibody Solution can be performed overnight at 4°C. 16. After final antibody incubation and rinsing of each channel with PBS, disassemble the slide from the patterning assembly. 17. Place the patterned slides into a Coplin jar filled with Blocking Solution. 18. Incubate with Blocking Solution for 1 h at room temperature, or overnight at 4°C. 19. Replace Blocking Solution with PBS and incubate for 30 min, with agitation. Repeat twice more for a total of three postblocking washes. 20. Allow slides to air dry. Store slides at 4°C in a moist chamber until use.
4. Notes 1. Silanization steps are performed in a N2-filled glove bag in a ventilated hood. This is to prevent air-oxidation of the thiolsilane, as well as to protect laboratory personnel from inhaling silane fumes. Triethoxy silanes are used in preference to trimethoxy silanes, due to decreased toxicity. 2. The MTS silane solution should be prepared just before use. It is very important that the silane be kept as dry as possible and that its exposure to air/oxygen is minimized to avoid decomposition/polymerization of the silane and oxidation of the thiols. Aliquoting of the silane stock with storage in dark glass vials under nitrogen is highly recommended. 3. Other heterobifunctional crosslinkers with NHS and maleimide reactive groups (e.g., BMPS, MBS, SMCC) can be used in place of GMBS. The final concentration of these other linkers in the ethanolic mix should be 1 mM. 4. The biomolecule solution should not contain any aminecontaining materials such as Tris and BSA. If present in initial protein solution, they should be removed by dialysis or another appropriate method. 5. Other homo-bifunctional NHS-based crosslinkers (e.g., DSS and EGS) may be used in place of BS3, provided the final diluted molar concentration is 50 mM.
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6. Addition of 0.8 molar equivalents of TCEP to the solution of peptide, protein, carbohydrate, etc., is crucial for linking of oxidized thiols to amine-modified surfaces. TCEP will reduce disulfide bonds in proteins, peptides, and derivatized sugars without introducing new thiols, as occurs with thiolcontaining reducing agents such as dithiothreitol (DTT) and 2-mercaptoethanol. In general, care must be exercised with this reagent, as its redox potential is significantly higher than that of DTT, and only half the concentration of TCEP is required when compared with DTT for the same application. 7. As with all halogenated silanes, trichlorosilane is highly reactive and freshly opened stocks should be used for optimal results. 8. An alternate method for dissolving casein in the Blocking Solution is slow heating to near-boiling, followed by cooling to room temperature. 9. This solution can be stored at 4°C for extended time if azide is added (final concentration 0.03 wt%). 10. Used (waste) silane and toluene are poured into labeled brown glass bottles. Glassware contaminated by silane and/ or toluene are rinsed several times with acetone; rinse acetone is discarded in same waste container as silane and toluene. 11. High temperature curing of the slides after patterning is essential for sensitive detection of ganglioside-binding toxins (26). Omission of this step has resulted in great diminished binding of cholera toxin to immobilized GM1. Curing of the OTS-treated slides before patterning will prevent immobilization of glycolipids onto the surface.
Acknowledgments The development of these methods was in part funded by the Office of Naval Research. The views expressed here are those of the authors and do not represent those of the U.S. Navy, the U.S. Department of Defense or the U.S. Government. References 1. Bhatia, S. K., Cooney, M. J., Shriver-Lake, L. C., Fare, T. L., and Ligler, F. S. (1991) Immobilization of acetylcholinesterase on solid-surfaces – chemistry and activity studies. Sens. Actuator. B-Chem. 3, 311–317 2. Duveneck, G. L., Neuschafer, D., and Ehrat, M. (1995) Process for detecting evanescently excited luminescence., Vol. International Patent Go1N 21/77, 21/64
3. Duveneck, G. L., Pawlak, M., Neuschafer, D., Bar, E., Budach, W., Pieles, U., and Ehrat, M. (1997) Novel bioaffinity sensors for trace analysis based on luminescence excitation by planar waveguides. Sens. Actuator. B-Chem. 38/39, 88–95 4. Liron, Z., Tender, L. M., Golden, J. P., and Ligler, F. S. (2002) Voltage-induced inhibition of antigen-antibody binding at conduct-
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5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
Shriver-Lake, Charles, and Taitt ing optical waveguides. Biosens. Bioelectron. 17, 489–494 Pawlak, M., Grell, E., Schick, E., Anselmettin, D., and Ehrat, M. (1998) Functional immobilization of biomembrane fragments on planar waveguides for the investigation of site-directed ligand binding by surface confined fluorescence. Faraday Discus. 111, 273–288 Sapsford, K. E., Rowe Taitt, C. A., and Ligler, F. S. (2002) Planar waveguides for fluorescence biosensors. In: Optical biosensors: Present and future (Ligler, F. S. & Rowe Taitt, C. A., eds.). Elsevier, The Netherlands, pp. 95–122 Bernard, A., Michel, B., and Delamarche, E. (2001) Micromosaic immunoassays. Anal. Chem. 73, 8–12 Delamarche, E., Bernard, A., Michel, B., and Biebuyek, H. (1997) Patterned delivery of immunoglobulins to surfaces using microfluidic networks. Science 276, 779–781 Gauglitz, G. (2005) Direct optical sensors: principles and selected applications. Anal. Bioanal. Chem. 381, 141–155 Golden, J., Shriver-Lake, L., Sapsford, K., and Ligler, F. (2005) A “Do-it-yourself” array biosensor. Methods 37, 65–72 Ngundi, M. M., Taitt, C. R., McMurry, S. A., Kahne, D., and Ligler, F. S. (2006) Detection of bacterial toxins with monosaccharide arrays. Biosens. Bioelectron. 21, 1195–1201 Rowe, C. A., Scruggs, S. B., Feldstein, M. J., Golden, J. P., and Ligler, F. S. (1999) An array immunosensor for simultaneous detection of clinical analytes. Anal. Chem. 71, 433–439 Sapsford, K. E., Rasooly, A., Taitt, C. R., and Ligler, F. S. (2004) Rapid detection of Campylobacter and Shigella species in food samples using an array biosensor. Anal. Chem. 76, 433–440 Sapsford, K. E., Taitt, C. R., Loo, N., and Ligler, F. S. (2005) Biosensor detection of botulinum toxoid A and staphylococcal enterotoxin B in food. Appl. Environ. Microbiol. 71, 5590–5592 Taitt, C. R., Anderson, G. P., Lingerfelt, B. M., Feldstein, M. J., and Ligler, F. S. (2002) Nine-analyte detection using an array-based biosensor. Anal. Chem. 74, 6114–6120 Tschmelak, J., Kumpf, M., Proll, G., and Gauglitz, G. (2004) Biosensor for seven sulphonamides in drinking, ground, and surface
17.
18.
19.
20.
21.
22.
23.
24.
25.
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water with difficult matrices. Anal. Lett. 37, 1701–1718 Tschmelak, J., Proll, G., and Gauglitz, G. (2004) Verification of performance with the automated direct optical TIRF immunosensor (River Analyser) in single and multi-analyte assays with real water samples. Biosens. Bioelectron. 20, 743–752 Moreno-Bondi, M. C., Taitt, C. R., ShriverLake, L. C., and Ligler, F. S. (2006) Multiplexed measurement of serum antibodies using an array biosensor. Biosens. Bioelectron. 21, 1880–1886 Ngundi, M. M., Shriver-Lake, L. C., Moore, M. H., Ligler, F. S., and Taitt, C. R. (2006) Multiplexed detection of mycotoxins in foods with a regenerable array. J. Food Prot. 69, 3047–3051 Martin, B. D., Lindhardt, R. J., and Dordick, J. S. (1998) Highly swelling hydrogels from ordered galactose-based polyacrylates. Biomaterials 19, 69–76 Cras, J. J., Rowe-Taitt, C. A., Nivens, D. A., and Ligler, F. S. (1999) Comparison of chemical cleaning methods of glass in preparation for silanization. Biosens. Bioelectron. 14, 683–688 Shriver-Lake, L. C. (1998) Silane-modified surfaces for biomaterial immobilization. In: Immobilized biomolecules in analysis: A practical approach (Cass, T. & Ligler, F. S., eds.). Oxford University Press, Oxford Golden, J. P., Taitt, C. R., Shriver-Lake, L. C., Shubin, Y. S., and Ligler, F. S. (2005) A portable automated multianalyte biosensor. Talanta 65, 1078–1085 Charles, P. T., Goldman, E. R., Rangasammy, J. G., Schauer, C. L., Chen, M.-S., and Taitt, C. R. (2004) Fabrication and characterization of 3D hydrogel microarrays to measure antigenicity and antibody functionality for biosensor applications. Biosens. Bioelectron. 20, 753–764 Ngundi, M. M., Kulagina, N. V., Anderson, G. P., and Taitt, C. R. (2006) Nonantibodybased recognition: alternative molecules for detection of pathogens. Exp. Rev. Proteomics 3, 511–524 Rowe-Taitt, C. A., Cras, J. J., Patterson, C. H., Golden, J. P., and Ligler, F. S. (2000) A ganglioside-based assay for cholera toxin using an array biosensor. Anal. Biochem. 281, 123–133
Chapter 24 Rapid DNA Amplification Using a Battery-Powered Thin-Film Resistive Thermocycler Keith E. Herold, Nikolay Sergeev, Andriy Matviyenko, and Avraham Rasooly Summary A prototype handheld, compact, rapid thermocycler was developed for multiplex analysis of nucleic acids in an inexpensive, portable configuration. Instead of the commonly used Peltier heating/cooling element, electric thin-film resistive heater and a miniature fan enable rapid heating and cooling of glass capillaries leading to a simple, low-cost Thin-Film Resistive Thermocycler (TFRT). Computer-based pulse width modulation control yields heating rates of 6–7 K/s and cooling rates of 5 K/s. The four capillaries are closely coupled to the heater, resulting in low power consumption. The energy required by a nominal PCR cycle (20 s at each temperature) was found to be 57 ± 2 J yielding an average power of approximately 1.0 W (not including the computer and the control system). Thus the device can be powered by a standard 9 V alkaline battery (or other 9 V power supply). The prototype TFRT was demonstrated (in a benchtop configuration) for detection of three important food pathogens (E. coli ETEC, Shigella dysenteriae, and Salmonella enterica). PCR amplicons were analyzed by gel electrophoresis. The 35 cycle PCR protocol using a single channel was completed in less then 18 min. Simple and efficient heating/cooling, low cost, rapid amplification, and low power consumption make the device suitable for portable DNA amplification applications including clinical point of care diagnostics and field use. Key words: PCR, Thermocycler, Peltier, Thin-Film Resistive Heater, Data acquisition system (DAS), Proportional-derivative-integral (PID), DNA, Food pathogens, Food testing, Point of Care (POC).
1. Introduction Polymerase chain reaction (PCR) is a powerful method for DNA amplification (1), which is routinely used for detection and analysis of DNA. Many commercial thermocyclers employ a Peltier chip (2–4), which is a solid-state technology that can be used for heating or cooling. Peltier systems are widespread and provide a Avraham Rasooly and Keith E. Herold (eds.), Methods in Molecular Biology: Biosensors and Biodetection, Vol. 504 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI:10.1007/978-1-60327-569-9_24
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very effective design in conjunction with standard commercially available PCR tubes. Peltier technology provides symmetric control of both heating and cooling in a single component and thus represents a preferred design. However, Peltier chips are made of semiconductor material that tends to be fairly massive. Thus, Peltier systems are not a likely candidate for high-speed cycling. Furthermore, standard PCR tubes have a fairly long thermal time constant (on the order of 10 s) and thus any attempt to achieve rapid PCR must start with a redesign of the reaction vessel to minimize that time constant. Air heating and cooling (5) is an alternative that has gained attention as a technique for real-time PCR, where the reaction is performed in a glass capillary (5–7). The capillary configuration provides a large surface area for heat transfer and a small cross sectional area, both of which minimize the thermal time constant. Such systems allow rapid cycling down to about one cycle per minute where the time constant of the reaction vessel (on the order of 4 s) again limits the minimum cycling rate. Some recent designs have employed infrared heating (8). This configuration requires lenses and filters, and also requires positioning of the reaction mixture at the appropriate focal distance of the optical system. Continuous flow PCR (CFPCR) microfluidic devices, which consist of a single channel that is continuously looped through different temperature zones to carry out DNA denaturing, annealing, and extension, have also been reported (9–11). The PCRs are pumped through the channel with the cycle number typically determined by the number of loops through each isothermal zone required for the PCR. Microfabrication enabled the development of an electrokinetically driven PCR in a circular continuous flow format through three isothermal zones (12) employing a 300–400 V/cm electric field for DNA circulation between the zones. Each of these designs is more complex than the one that came before, and thus they do not address our design requirements that include lowcost, low-power, and rapid cycling. Power use and integration of PCR amplification with amplicon analysis are critical elements in the design of portable thermocyclers. Several approaches for amplicon detection and analysis were developed including fluorogenic, capillary electrophoretic (CE), and electrochemical detection. Fluorogenic analysis of PCR amplicons (13–18) eliminates the need of postPCR analysis such as electrophoretic separation. The integration of fluorogenic detection during PCR amplification enables realtime monitoring (17–20). PCR amplification and capillary electrophoretic (CE) analysis were coupled in a compact instrument for microbial detection (21). A microfabricated capillary electrophoresis chip was integrated with electrochemical detection of PCR amplicons (22). A similarly integrated system consisting of a microfabricated silicon-based PCR (microPCR) chamber and
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a microfabricated electrophoretic glass chip has been developed (23). On-chip amplification using TaqMan PCR in nanoliter volumes on a highly integrated silicon microchamber array was reported (24). Rapid and robust detection of infectious disease agents, including food borne pathogens, is critical for public health security. An important group of microbial pathogens includes pathogenic E. coli, Shigella dysenteriae, and Salmonella enterica, which are some of the most common causes of food borne infectious disease worldwide, associated with a variety of diseases, including diarrhea, urinary tract infections, bacteremia, meningitis, and wound infections. These pathogens are commensal bacterial organisms ubiquitous in mammalian intestines as well as in the environment. Rapid differentiation of nonpathogenic, commensal strains of E. coli from pathogenic variants (pathovars) is a major clinical and public health concern. PCR-based tests for detecting microorganisms are increasingly being implemented in clinical laboratories (25). Such tests offer high sensitivity, specificity, and enable better characterization of the microorganisms. PCR amplification followed by microarray analysis of the amplicons was shown to be a powerful method for microbial analysis (26–35). A recent example of DNA amplification and hybridization performed in an integrated plastic device (36) required approximately 1.5–2 h to complete one assay. Recent portable PCR instruments enable microbial detection to be performed outside the laboratory and completed in a matter of minutes (25, 37). This technology may enable the development of Point of Care (POC) diagnostics, reducing the need for (38) the traditional central laboratory approach for medical diagnostics. POC allows speedy identification of infectious disease resulting in better treatment and reduction of indiscriminate use of antibiotics, a major factor in the emergence of antibiotic-resistant pathogenic microorganisms. The need for portable devices for detection of microbial pathogens led to the development of a new class of mobile, small, battery-powered instruments to perform real-time PCR in the field (37, 39). This was made possible by replacing or redesigning energy consuming components, such as the heating block and the Peltier elements, with new miniature energy efficient components such as thin-film resistive heaters, fans, and integrated LED, and silicon photodiode detectors for real time analysis of the amplicons. One of the barriers to the spread of rapid amplification technology for POC clinical use is the cost of the instrumentation, which limits the use of the technology. To make biosensor technology more accessible, recently a manuscript was published (38) describing an inexpensive array biosensor. Here we describe a simple and inexpensive battery-powered thin-film
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resistive thermocycler (TFRT), which has potential to be the base of a portable thermocycler for rapid detection of microbial pathogens in the field or in the clinic.
2. Materials 1. Film heater, Minco HK913H. 2. Cooling fan, 25 mm diameter, Radio Shack 273–240. 3. Solid state relay, Omega SSRDC100VDC12. 4. Data acquisition and control board, INET 100. 5. Power supply, 12 VDC, HY1803D Sinometer. 6. Borosilicate glass capillary, 15 mm × 1 mm OD × 0.75 mm ID. 7. Type T thermocouple wire, 30 AWG (0.25 mm diameter), Omega TT-T-30. 8. Bacterial strains: Strains used in this study are E. coli ETEC E1881C, Shigella dysenteriae, and Salmonella enterica serovar Typhimurium. These strains were obtained from the FDA Center for Food Safety and Applied Nutrition (CFSAN) bacterial collection of Ms. Christine Keys and Dr. Farukh Khambaty. 9. Ultraspec 3000 spectrophotometer (Pharmacia, Peapack NJ). 10. BLAST (Basic Local Alignment Search Tool), National Institutes of Health http://www.ncbi.nlm.nih.gov/BLAST/ 11. Oligo Design software, http://www.enme.umd.edu/ bioengineering/ 12. BSA (bovine serum albumin), powder, A2153, Sigma Aldrich, St. Louis, MO. 13. Ethidium bromide, 10 mg/mL, E1510, Sigma Aldrich, St. Louis, MO. 14. PCR grade mineral oil, DNase free, M8662, Sigma Aldrich, St. Louis, MO. 15. Gen AMP PCR System 2400 thermocycler. 16. AgaroseBP160–100, Fisher BioReagents, http://www.fishersci.com 17. TBE buffer, BP1396–86, Fisher BioReagents, http://www. fishersci.com 18. EDAS 290 digital camera/stand, Kodak, Rochester NY. 19. dNTP mix, PR-U1511, Promega Corp., Madison, WI. 20. Taq DNA polymerase, Sigma Aldrich St. Louis MO.
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21. Oligonucleotide primers, custom sequence, Operon, Huntsville, AL.
3. Methods 3.1. Thin-Film Resistive Heating Thermocycler
The primary physical characteristic that limits the cycling speed is the thermal capacitance of the heated region. To achieve highspeed thermocycling, we used a low mass capillary cartridge coupled to a thin-film resistive heater. Two CPU cooling fans were used for fast and efficient cooling of the PCR capillaries. The result is the rapid low power thin-film resistive thermocycler (TFRT) benchtop prototype shown in Fig. 1. Referring to Fig. 1, the basic elements of the TFRT prototype are: a thin-film resistive heater (A) and two miniature fans (B1 and B2) used for fast and efficient cooling of the PCR capillaries (D). The capillaries used for the PCRs are assembled on a thin (0.2 mm) microscope
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cover slip with face area 15 × 15 mm placed on a layer of aluminum foil (for more even heating) and arranged side-by-side on the flat surface placed directly on the heating surface. The capillary cassette assembly includes a fine gage Type T thermocouple (C) used for temperature control. The capillary cassette was placed directly on the heating surface of the thin-film resistive heater to minimize thermal capacitance. A Plexiglas frame was used to hold the assembly and allow access of the cooling air flow from the fans to the capillary cassette. The prototype TFRT instrument consists of a thin-film resistive heater, two fans, thermocouple, controller, software, and a computer. The heating element has an effective face area of 15 × 15 mm and a resistance of 15 Ω (Fig. 1A). The PCR capillaries are coupled directly to the heater. For cooling, two fans are positioned on opposite sides of the test section. The fans and the heating element are powered by 9 VDC (see Note 1). The system temperature is sensed by a thermocouple (see Notes 2 and 3) mounted inside a capillary and mounted on the heater in a symmetrical fashion to the capillaries that hold the PCR mixtures. For prototype development, thermocouples were mounted in each of the capillaries to map the temperature distribution on the surface of the heating element. 3.2. Measurement and Control Hardware
Figure 1A is a schematic of the temperature measurement and control circuit. A type T thermocouple (see Note 2), inserted in a glass capillary, measures the temperature of a capillary with similar characteristics to capillaries holding the reaction mixture. The thermocouple is connected to an A/D channel of the data acquisition system (DAS) in a differential configuration (see Note 3). The DAS system is also used to communicate output commands (D/A channels) from the computer to the thermocycler. The DAS system employed in this study has eight channels available that can be used as either inputs or outputs. The fans and the heater are connected to the output channels of the board via solid state relays. The computer interface employs an off-the-shelf data acquisition system and two switching relays (Fig. 1E), one for the heater and one for the fans. The system was powered by a 9 V power supply or batteries.
24.3.3. Measurement and Control Software
The control system was written in VisualBasic to read the temperature of the PCR zone and adjust the heater and fans to follow a user-specified PCR temperature profile. The control system logic includes PID (proportional-derivative-integral) functionality (see Note 4). In addition, the logic provides features to anticipate over or under-shoot following the temperature transitions to maximize control fidelity. The control program, implemented on a dedicated PC computer, drives the heater and fans via the
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DAS. The heater control system is based on PWM (pulse width modulation) running at 0.5 Hz, which represents a compromise between the desire for a high frequency to provide better control and a low frequency to allow the computer to complete overhead tasks without disrupting control (see Note 5). The temperature and the setpoint for a typical PCR run consisting of 40 cycles are shown in Fig. 2A. This plot shows the repeatability of the control system from cycle to cycle. An expanded view of a typical single cycle is shown in Fig. 2B where the heating and cooling rates can be determined. Rapid amplification was a key design goal for the TFRT system. For the case shown in Fig. 2, the time required to heat the test section from 54 to 72°C is 2.45 s corresponding to a heating rate of 7.3 K/s. For heating from 72 to 94°C the time is 3.60 s, corre-
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sponding to a heating rate of 6.1 K/s. For cooling from 94 to 54°C it takes 7.46 s corresponding to cooling rate of 5.4 K/s. The heating and cooling rates are functions of the device design and the control system settings. The largest slope of the temperature traces represents the maximum capability of the system in this configuration (the maximum slopes of both heating curves are similar). As the temperature approaches the setpoint, the control system reduces the power input to avoid overshoot (heating) or undershoot (cooling). The best performance of the system is 2 s to heat from 72 to 94°C, 2 s to heat from 54 to 72°C and cooling from 94 to 54°C in 5.5 s in a single capillary mode indicating a maximum heating rate of 9 K/s and a maximum cooling rate of 7.3 K/s. A five capillary TFRT cassette was used for amplification of four targets simultaneously (one capillary holds the control thermocouple). Temperature uniformity, which is an essential element of PCR reproducibility, was measured in a simulated PCR run. For this test, the capillaries were filled with mineral oil to simulate the PCR mixture, and each capillary carried a thermocouple similar to the thermocouple used for temperature control. Figure 3A shows the simulated PCR run consisting of ten cycles. Data analysis suggests that, in general, the temperature among the capillaries is uniform within an error band: for the 72 and 54°C levels the temperature varied by less than ±1 K around the set point and for DNA melting, the temperature is 92°C ± 2 K. Insulating the cassette was found to reduce the variability among capillaries (not shown) but PCR was found to be repeatable at the indicated variability so the insulation was not included in this study. In PCR, the most critical temperatures are the annealing and polymerization, where significant deviation especially for the annealing temperature will substantially reduce the PCR yield. The melting temperature, in general, is more tolerant to a broader range of temperatures.
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Low energy consumption is a critical design feature for operating a portable PCR instrument in the field. The energy consumption for the TFRT benchtop prototype (not including the computer and the control system) was measured at voltage inputs of 8, 9, and 10 V, and the energy consumed for heating, cooling, and for the relays was determined by integrating the power over time as the control system varied the input power. The relays were found to dissipate only 16 mW at 0.5 A. This is negligible compared with the nominal 5 W heater power. The fans dissipate approximately 1 W each but they are on for only a brief time (approximately 4 s each cycle) so that the fan energy use is also secondary to the primary energy needs of the heating element. The pulse width control system turns the heater on at full power for a fraction of each control period with the size of the fraction modulated to maintain temperature control. Thus, to determine the energy use over a PCR cycle, it is sufficient to determine the heater on-time. At a fixed power supply voltage (V) and a fixed heater resistance (R) (the heater resistance does not measurably change over the temperature range of interest), the energy use is just the on-time multiplied by the power determined as V 2/R. The energy required by a nominal PCR cycle (20 s at each temperature) was found to be 57 ± 2 J for all three power supply voltages. The control system adjusts the power input to only that needed to heat the system, which is independent of voltage. If the power is averaged over the cycle and the fan power is included, it yields an average power of 1.08 W. The majority of TFRT tests were run at a power supply voltage of 9 V with the plan to utilize a 9 V battery in a portable version. Tests were also run using a battery and the system worked well as shown in Fig. 4. A standard 9 V alkaline battery is rated
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at approximately 4.5 W-h. On the basis of an average power of 1.08 W, this implies a battery life of 4.2 h which is consistent with what we observed. It is interesting to note that a typical lithium ion laptop battery has a rating on the order of 65 W-h, so long term battery operation of this PCR device is practical using existing battery technology, and it appears feasible to power the PCR device and the computer controller from the same battery. 3.6. Design of PCR Primers
Table 1 lists the primers used to amplify genes from three bacterial pathogens. Genes selected for the detection of bacteria were uid (E. coli), ipaH (Shigella), and pfkB-thrS intergenic region (Salmonella). The NCBI database was used to find and retrieve the gene DNA sequence from a complete genome of the target organism. That sequence was then used as a search sequence against the entire database (using BLAST) to gain some understanding of its uniqueness. The sequences with high similarity were then aligned using CLUSTAL W (40–42) to identify unique conserved regions. Primers were then designed, based on those regions using Oligo Design 1.621 software (43), and are listed in Table 1. Protocol (described for the uid gene in E. coli): 1. Search Pubmed for nucleotide sequences with the search terms “uid AND coli”. Numerous whole genomes are returned. Chose Escherichia coli HS (gi:157065147).
Table 1 Primers for PCR amplification of UID (E. coli ETEC E1881C), ipaH (Shigella dysenteriae), and Salmonella specific region (Salmonella enterica serovar Typhimurium) Gene
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2. Once the Genbank sequence is displayed, search (using the broswer search feature) for “uid”. The uid region, including uidA and uidR, spans 2,793 bp and is chosen as the target. 3. Using the target sequence, run BLAST against the entire nucleotide database. The result is 17 closely identical matches with greater than 98% identity. Of these, 11 are E. coli complete genomes and 6 are Shigella complete genomes. 4. The unique E. coli sequences were then copied to a file for multiple alignment using Clustal. It was determined that the uid region was strongly conserved among all the sequences with only single point differences distributed among long stretches of identical sequence. These differences could be SNP’s or sequencing errors. In any case, they are not expected to be significant in choosing the primers. 5. The primers were chosen using Oligo Design, which has a search algorithm with multiple inputs. The primary inputs are the target melting temperature for the primers. In this case, a melting temperature of 54.5°C was selected resulting in the primer set in Table 1. The algorithm steps through the entire sequence and determines, at each location, if there is a sequence length that meets the input criteria (including melting temperature, length, hairpin potential, presence of repeats, 3′ end annealing strength, self annealing and primer dimer potential). It then steps through in the opposite direction. Candidate primer pairs that satisfy the amplicon length input specification are then displayed. 3.7. PCR Amplification of Individual Toxin Genes
The standard PCR mixture (50 μL) contained five units of Taq DNA polymerase, 1× PCR buffer supplemented with 3.5 mM MgCl2, 200 nM of forward and reverse primers, 400 mM each of dNTP (dATP, dGTP, dCTP, and dTTP), 1 mg/mL BSA (see Note 6), 0.2 μg/mL ethidium bromide, and 100 ng of DNA template (see Note 7). Typically, 3–5 μL of PCR mixture was loaded in each capillary. To prevent evaporation during thermal cycling, both ends of the capillaries were sealed with 1–2 μL of PCR-grade mineral oil. The control for the PCR amplification was performed using a Gene AMP PCR System 2400 thermocycler with the following cycle conditions: initial activation at 94°C for 5 min; 35–40 cycles at 94°C for 20 s, 54°C for 30 s, 72°C for 40 s; and the final extension at 72°C for 1 min. It is noted that the ramping times are not included in these specifications; when ramping times are included the total cycle time is 3.5 min resulting in a total time of 122 min for the control reaction (see Note 8). The nominal TFRT PCR runs were performed with the following conditions: 92°C – 40 s, 40 cycles of (92°C − 5 s, 54°C − 15 s, 71°C − 15 s), 71°C − 1 min. These specifications do include the ramping time such that the total time is 25 min. The cycle times were adjusted for longer and shorter total times, as described below.
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The presence of amplified PCR products was detected in two ways. Ethidium bromide included in the reaction mixture allowed rapid determination of PCR products in the capillaries by visualization of the capillaries under UV illumination (see Note 9). After that determination, the reaction was passed through a 1.5% agarose gel by electrophoresis in 1× TBE buffer. Gels were stained with ethidium bromide and photographed in UV light with a Kodak digital camera EDAS 290. Microbial DNA was amplified using the TFRT prototype PCR machine described above. Runs were made using a single capillary (Fig. 5) and with a cassette of five capillaries (Fig. 6). Single capillary amplification of the ipaH gene from Shigella dysenteriae was performed using the primers shown in Table 1. The gel results are shown in Fig. 5a along with the direct visualization in the capillaries in Fig. 5b. Lane 1 is a negative control with no target DNA. As a positive control, a reaction was performed in a conventional Peltier based thermocycler (Fig. 5, lane 5). The control reaction was run in a conventional thin-wall 200 μL PCR tube and was transferred to a capillary for visualization as shown in Fig. 5b. The remaining lanes (lanes 2–4) in Fig. 5 are PCRs run in the TFRT prototype in a single capillary mode with varying cycle time. The cycle times for lanes 2–4 were 37 min (20 s at 94°C, 20 s at 54°C, and 20 s at 72°C), 20 min (10, 12, 10 s), and 17.5 min (5, 12, 10 s), respectively.
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Fig. 6. DNA amplification of three foodborne pathogens using TFRT. DNA amplified using Peltier-based thermocycler lanes 1–3 and DNA amplified using the prototype thin-film resistive thermocycler (TFRT) lanes 4–6. M – 123 bp DNA ladder (see Note 9), 1 – uid (E. coli ETEC E1881C), 2 – ipaH (Shigella dysenteriae), 3 – Salmonella specific region (Salmonella enterica serovar Typhimurium), 4, 5, 6 same on the TFRT thermocycler (total time of amplification (40 cycles) was 122 and 25 min for the Peltier and TFRT thermocyclers, respectively).
To enable multiple simultaneous PCRs, a TFRT cassette of five capillaries (four PCRs plus the temperature sensor) was designed and tested. Amplification using a Peltier-based thermocycler is compared with the TFRT results in Fig. 6. For both devices the thermal protocol was similar; initial denaturation at 92°C − 40 s followed by 40 cycles of 92°C − 5 s, 54°C − 15 s, 71°C − 15 s and final elongation of 71°C − 1 min. Although both thermocyclers used the same parameters, the ramping rate of the commercial instrument is slower resulting in slower amplification (see Note 8) Three PCRs were run simultaneously for detection of three microbial pathogen DNA sequences as shown in Table 1. The amplicon size range in these experiments is 349–720 bp. As shown in Fig. 6, all target sequences were amplified in the TFRT (lanes 4–6) with comparable signals to that obtained from the commercial cycler (lanes 1–3). However, the TFRT is significantly faster (25 min) than the amplification using the commercial instrument (80 min). These results demonstrate the potential use of the TFRT system for rapid identification of multiple microbial pathogens simultaneously. Protocol: 1. Flush the capillaries with isopropyl alcohol, followed by DI water. After flushing, dry capillaries at 50°C for 30 min prior to use.
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2. Load capillaries with PCR solution by dipping one end in the previously mixed solution. Capillary forces draw the solution into the capillary. The total reaction volume in the capillary was approximately 5 μL (11.3 mm of capillary length). 3. Load 1–2 mm of mineral oil into each end of the capillary to seal the reaction against evaporation. The oil can be drawn into the capillary in a manner similar to loading the solution (i.e., by capillary forces). 4. Lay the capillaries on the TFRT heater surface. For multiple capillaries, load the capillaries in a cassette frame prior to placing on the heater surface. 5. Run the thermocycler software to execute the PCR thermal cycles. The software allows the user to specify time and temperature for each portion of the cycle. 6. After the reaction is complete, detect the amplicon in one of two ways: (1) direct visualization of the fluorescence from the capillary, or (2) extract the PCR products from the capillary and run gel electrophoresis.
4. Notes 1. During testing, the system was powered by a 9 VDC power supply. The system was also tested using a standard 9 V battery. Although the battery was found to have sufficient energy to power an entire PCR run, a practical device would need a larger battery to provide longer operational life. 2. Thermocouples were fabricated from commercial thermocouple wire by stripping the insulation approximately 1 cm from the end. The two wires were then twisted together, doubled over, and soldered. After fabrication, the thermocouple was tested for electrical continuity and for temperature measurement fidelity. Thermocouples made from the same spool of wire were found to provide closely identical signals giving a relative temperature measurement error of less than ±0.05 K. Thermocouples were made from 0.25 mm diameter wire to allow insertion of the thermocouple in the capillaries. Because of the fine wire, all operations described in this note require considerable care to avoid breaking the wire. 3. Thermocouples were attached to the data acquisition board in a differential configuration with one lead grounded to the board. Because of the exposed nature of the thermocouple junction, care must be exercised to avoid electrical grounding of the junction, which can result in a ground loop that can
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cause significant temperature measurement errors (due to the low signal level of the thermocouple). 4. The control algorithm used is a modified proportional-integralderivative (PID) algorithm. Because of the frequent setpoint changes associated with the three temperature levels in the PCR protocol, the algorithm is designed to anticipate these. In particular, the algorithm allows the user to achieve rapid heating while avoiding overshoot after temperature increases by setting an offset temperature difference. Prior to closing to within that difference of the setpoint, the heater is turned on full power. After passing the offset level, the proportional controller takes over. Once the temperature either passes the setpoint or plateaus, the integral action is turned on. 5. The control signal calculated by the PID algorithm (see Note 3) is translated into a heating profile by way of pulse width modulation. The heater receives a pulse train (frequency of 0.5 Hz) with variable on time. During the on segment of each 0.5 s pulse, the heater is on at full power. The integrated power over the full pulse is controlled to achieve the desired temperature. The thermal capacitance of the capillaries and the heater itself is sufficient to damp out any temperature fluctuations such that the temperature measured inside the capillaries is steady (does not show any 0.5 Hz ripple). 6. It was found (results not shown) that the addition of bovine serum albumen (BSA, 1 mg/mL) to the reaction significantly improved PCR yield. This is thought to be because BSA adsorbs to the glass preferentially and blocks adsorption of DNA and the polymerase on the capillary wall. 7. DNA from freshly grown cells was extracted using a phenol–chloroform extraction. The presence, concentration and purity of genomic DNA in the prepared samples were detected by measuring the absorbance at 260 and 280 nm using a spectrophotometer. 8. It is noted that, unlike the conventional thermocycler, the control system for the TFRT prototype includes the ramp time in the time at each temperature. Thus, although the system is set to a given temperature for a fixed time (e.g., 10 s for the rapid cycling case), the prototype takes as much as 7 s to make temperature transitions so the total time at the setpoint temperature is less than the value given in the protocol. 9. The PCR recipe included intercalating dye, ethidium bromide (0.2 μg/mL), to enable in-capillary detection of the amplification. Direct detection of PCR amplicons was achieved by placing the capillaries on a UV transilluminator
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(Fig. 5b), which enabled rapid analysis and demonstrated the potential of the platform for real-time detection. 10. On both Figs. 5 and 6, the image of the first two bars of the ladder are obscured such that the first easily visible bar is 369 bp. Thus, the gel results in Fig. 5 (ipaH amplicon is 515 bp) line up close to the second bar of the ladder which is 492 bp. Figure 6 has a similar interpretation. The ladder bars were obscured by the gel loading buffer dye.
Acknowledgments This work was supported in part by the FDA Office of Science and by USDA grant # 2003-35201-13784.
References 1. Saiki, R.K., D.H. Gelfand, S. Stoffel, S.J. Scharf, R. Higuchi, G.T. Horn, K.B. Mullis, and H.A. Erlich. Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science. 1988, 239(4839), 487–491 2. Wittbrodt, J. and W. Erhardt. An inexpensive and versatile computer-controlled PCR machine using a Peltier element as a thermoelectric heat pump. Trends Genet. 1989, 5(7), 202–203 3. Wilding, P., M.A. Shoffner, and L.J. Kricka. PCR in a silicon microstructure. Clin Chem. 1994, 40(9), 1815–1818 4. Khandurina, J., T.E. McKnight, S.C. Jacobson, L.C. Waters, R.S. Foote, and J.M. Ramsey. Integrated system for rapid PCR-based DNA analysis in microfluidic devices. Anal Chem. 2000, 72(13), 2995–3000 5. Chapin, K. and T.L. Lauderdale. Evaluation of a rapid air thermal cycler for detection of Mycobacterium tuberculosis. J Clin Microbiol. 1997, 35(8), 2157–2159 6. Friedman, N.A. and D.R. Meldrum. Capillary tube resistive thermal cycling. Anal Chem. 1998, 70(14), 2997–3002 7. Lagally, E.T., C.A. Emrich, and R.A. Mathies. Fully integrated PCR-capillary electrophoresis microsystem for DNA analysis. Lab Chip. 2001, 1(2), 102–107 8. Oda, R.P., M.A. Strausbauch, A.F. Huhmer, N. Borson, S.R. Jurrens, J. Craighead, P.J. Wettstein, B. Eckloff, B. Kline, and J.P. Landers.
9.
10.
11.
12.
13.
14.
Infrared-mediated thermocycling for ultrafast polymerase chain reaction amplification of DNA. Anal Chem. 1998, 70(20), 4361–4368 Kopp, M.U., A.J. Mello, and A. Manz. Chemical amplification: continuous-flow PCR on a chip. Science. 1998, 280(5366), 1046–1048 Park, N., S. Kim, and J.H. Hahn. Cylindrical compact thermal-cycling device for continuous-flow polymerase chain reaction. Anal Chem. 2003, 75(21), 6029–6033 Obeid, P.J., T.K. Christopoulos, H.J. Crabtree, and C.J. Backhouse. Microfabricated device for DNA and RNA amplification by continuousflow polymerase chain reaction and reverse transcription-polymerase chain reaction with cycle number selection. Anal Chem. 2003, 75(2), 288–295 Chen, J., M. Wabuyele, H. Chen, D. Patterson, M. Hupert, H. Shadpour, D. Nikitopoulos, and S.A. Soper. Electrokinetically synchronized polymerase chain reaction microchip fabricated in polycarbonate. Anal Chem. 2005, 77(2), 658–666 Eggerding, F.A. A one-step coupled amplification and oligonucleotide ligation procedure for multiplex genetic typing. PCR Methods Appl. 1995, 4(6), 337–345 Tyagi, S. and F.R. Kramer. Molecular beacons: probes that fluoresce upon hybridization. Nat Biotechnol. 1996, 14(3), 303–308
Rapid DNA Amplification Using a Battery-Powered Thin-Film 15. Tyagi, S., S.A. Marras, and F.R. Kramer. Wavelength-shifting molecular beacons. Nat Biotechnol. 2000, 18(11), 1191–1196 16. Nazarenko, I.A., S.K. Bhatnagar, and R.J. Hohman. A closed tube format for amplification and detection of DNA based on energy transfer. Nucleic Acids Res. 1997, 25(12), 2516–2521 17. Ririe, K.M., R.P. Rasmussen, and C.T. Wittwer. Product differentiation by analysis of DNA melting curves during the polymerase chain reaction. Anal Biochem. 1997, 245(2), 154–160 18. Wittwer, C.T., K.M. Ririe, R.V. Andrew, D.A. David, R.A. Gundry, and U.J. Balis. The LightCycler: a microvolume multisample fluorimeter with rapid temperature control. Biotechniques. 1997, 22(1), 176–181 19. Northrup, M.A., B. Benett, D. Hadley, P. Landre, S. Lehew, J. Richards, and P. Stratton. A miniature analytical instrument for nucleic acids based on micromachined silicon reaction chambers. Anal Chem. 1998, 70(5), 918–922 20. Wittwer, C.T., M.G. Herrmann, C.N. Gundry, and K.S. Elenitoba-Johnson. Real-time multiplex PCR assays. Methods. 2001, 25(4), 430–442 21. Lagally, E.T., J.R. Scherer, R.G. Blazej, N.M. Toriello, B.A. Diep, M. Ramchandani, G.F. Sensabaugh, L.W. Riley, and R.A. Mathies. Integrated portable genetic analysis microsystem for pathogen/infectious disease detection. Anal Chem. 2004, 76(11), 3162–3170 22. Ertl, P., C.A. Emrich, P. Singhal, and R.A. Mathies. Capillary electrophoresis chips with a sheath-flow supported electrochemical detection system. Anal Chem. 2004, 76(13), 3749–3755 23. Rodriguez, I., M. Lesaicherre, Y. Tie, Q. Zou, C. Yu, J. Singh, L.T. Meng, S. Uppili, S.F. Li, P. Gopalakrishnakone, and Z.E. Selvanayagam. Practical integration of polymerase chain reaction amplification and electrophoretic analysis in microfluidic devices for genetic analysis. Electrophoresis. 2003, 24(1–2), 172–178 24. Matsubara, Y., K. Kerman, M. Kobayashi, S. Yamamura, Y. Morita, Y. Takamura, and E. Tamiya. On-chip nanoliter-volume multiplex TaqMan polymerase chain reaction from a single copy based on counting fluorescence released microchambers. Anal Chem. 2004, 76(21), 6434–6439 25. Tang, Y.W., G.W. Procop, and D.H. Persing. Molecular diagnostics of infectious diseases. Clin Chem. 1997, 43(11), 2021–2038
457
26. Sergeev, N., M. Distler, S. Courtney, S.F. Al-Khaldi, D. Volokhov, V. Chizhikov, and A. Rasooly. Multipathogen oligonucleotide microarray for environmental and biodefense applications. Biosens Bioelectron. 2004, 20(4), 684–698 27. Al-Khaldi, S.F., K.M. Myers, A. Rasooly, and V. Chizhikov. Genotyping of Clostridium perfringens toxins using multiple oligonucleotide microarray hybridization. Mol Cell Probes. 2004, 18(6), 359–367 28. Volokhov, D., A. Pomerantsev, V. Kivovich, A. Rasooly, and V. Chizhikov. Identification of Bacillus anthracis by multiprobe microarray hybridization. Diagn Microbiol Infect Dis. 2004, 49(3), 163–171 29. Johnson, J., K. Jinneman, G. Stelma, B.G. Smith, D. Lye, J. Messer, J. Ulaszek, L. Evsen, S. Gendel, R.W. Bennett, B. Swaminathan, J. Pruckler, A. Steigerwalt, S. Kathariou, S. Yildirim, D. Volokhov, A. Rasooly, V. Chizhikov, M. Wiedmann, E. Fortes, R.E. Duvall, and A.D. Hitchins. Natural atypical Listeria innocua strains with Listeria monocytogenes pathogenicity island 1 genes. Appl Environ Microbiol. 2004, 70(7), 4256–4266 30. Sergeev, N., D. Volokhov, V. Chizhikov, and A. Rasooly. Simultaneous analysis of multiple staphylococcal enterotoxin genes by an oligonucleotide microarray assay. J Clin Microbiol. 2004, 42(5), 2134–2143 31. Volokhov, D., V. Chizhikov, K. Chumakov, and A. Rasooly. Microarray analysis of erythromycin resistance determinants. J Appl Microbiol. 2003, 95(4), 787–798 32. Volokhov, D., V. Chizhikov, K. Chumakov, and A. Rasooly. Microarray-based identification of thermophilic Campylobacter jejuni, C. coli, C. lari, and C. upsaliensis. J Clin Microbiol. 2003, 41(9), 4071–4080 33. Volokhov, D., A. Rasooly, K. Chumakov, and V. Chizhikov. Identification of Listeria species by microarray-based assay. J Clin Microbiol. 2002, 40(12), 4720–4728 34. Al-Khaldi, S.F., S.A. Martin, A. Rasooly, and J.D. Evans. DNA microarray technology used for studying foodborne pathogens and microbial habitats: minireview. J AOAC Int. 2002, 85(4), 906–910 35. Chizhikov, V., A. Rasooly, K. Chumakov, and D.D. Levy. Microarray analysis of microbial virulence factors. Appl Environ Microbiol. 2001, 67(7), 3258–3263 36. Liu, Y., C.B. Rauch, R.L. Stevens, R. Lenigk, J. Yang, D.B. Rhine, and P. Grodzinski. DNA amplification and hybridization assays in integrated plastic monolithic devices. Anal Chem. 2002, 74(13), 3063–3070
458
Herold et al.
37. Higgins, J.A., S. Nasarabadi, J.S. Karns, D.R. Shelton, M. Cooper, A. Gbakima, and R.P. Koopman. A handheld real time thermal cycler for bacterial pathogen detection. Biosens Bioelectron. 2003, 18(9), 1115–1123 38. Golden, J., L. Shriver-Lake, K. Sapsford, and F. Ligler. A “do-it-yourself” array biosensor. Methods. 2005, 37(1), 65–72 39. Belgrader, P., S. Young, B. Yuan, M. Primeau, L.A. Christel, F. Pourahmadi, and M.A. Northrup. A battery-powered notebook thermal cycler for rapid multiplex real-time PCR analysis. Anal Chem. 2001, 73(2), 286–289 40. Chenna, R., H. Sugawara, T. Koike, R. Lopez, T.J. Gibson, D.G. Higgins, and J.D. Thompson. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res. 2003, 31(13), 3497–3500
41. Thompson, J.D., D.G. Higgins, and T.J. Gibson. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 1994, 22(22), 4673–4680 42. Thompson, J.D., T.J. Gibson, F. Plewniak, F. Jeanmougin, and D.G. Higgins. The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 1997, 25(24), 4876– 4882 43. Herold, K.E. and A. Rasooly. Oligo design: a computer program for development of probes for oligonucleotide microarrays. Biotechniques. 2003, 35(6), 1216–1221
INDEX A
B
Absorbent pad ....................................... 170, 172, 173, 176, 177, 222, 232 Acetyl cholinesterase ......................................................... 6 AChE... ............................................... 6, 7, 15, 16, 19, 115, 118–120, 122–124 Adherent cell .........................................292, 296, 297–299, 306, 307, 315, 316, 386, 392 Adhesive card ................................................................ 176 Affinity maturation .......................................323, 337, 343, 344, 351, 352 Affinity selection ............................................275, 276, 283 AFM 60, 102 Ag/AgCl electrode .................................... 9, 12, 89, 94, 95, 105, 106, 122, 123, 130, 139, 147, 153, 160 Agarose ..................................................111, 130, 134–137, 258, 261, 262, 294, 304, 305, 330, 331, 340, 354–356, 358, 362–364, 369, 388, 410, 444, 452 Agglutination ................................................................ 238 Alkaline-phosphatase ................. 86–88, 159, 162, 294, 305 Alkanethiols .........................................................56, 63, 64 Aluminum layer ............................................................... 59 Amine groups .....................................................56, 63, 431 Amine-Amine ................................................422, 428, 431 Amine-modified surfaces ...............................421–422, 430 Amine-Thiol...........................................422, 428, 430–432 Amperometric ................................. 6, 12, 13, 16, 116–117, 157, 160, 162–165 Amplification ........................................................... 23, 24, 192–195, 209, 237, 256, 276, 278, 284–286, 292, 293, 297, 299–301 Anisotropic crystals ......................................................... 38 Antibody.................................................. 3, 6, 7, 13, 43–48, 56, 63, 65, 69, 70, 73, 75, 78, 80 purification ...................................................... 224–225 testing ...............................................186, 224, 225–230 Antigen discovery .......................................................... 275 Aptamer ............................. 23–34, 190, 385–397, 399–415 Assay sensitivity ......................................186–188, 224–226 Assay specificity ..................................................... 226–230 ATP................................................ 199, 257, 261, 262, 314, 329, 358, 368, 388, 389, 394, 395, 403, 451 AuNPs.. .................................................128, 129, 132–136, 138, 139, 141, 145, 149–152 Autofluorescence ........................................................... 342
Bacteriophage ................................. 255, 262, 265, 275–289 Bandpass filter ............................................................... 359 BAW................................................................................ 25 Beads............................................ 127–130, 134, 136–138, 140, 145, 147, 150, 185, 224, 237, 296, 314–315, 319, 324, 353, 402, 407, 413 Bending ...................................51–53, 55, 57, 61, 63, 65, 66 Binding peptide ..............................................264–265, 311 Biopanning ............................................................ 291–320 Bioreceptors ........................ 25, 52, 56, 63–65, 69, 323–347 Biotin....................................26, 31, 33, 128, 130, 131, 134, 136, 138, 145, 147, 150, 186, 188, 192, 194, 196, 198, 203, 293–297, 310, 311–313, 315, 317, 319, 320, 328, 333, 335, 341–343, 345, 353, 358–360, 365, 367, 373, 375, 379, 380, 413, 420, 423, 433 Blocking .................................................. 63, 122, 149, 150, 181, 188, 198, 202, 203, 205, 264, 425, 438 Blood.... .......................................... 25, 27, 178, 217, 237, 275, 284, 288 Bonding .......................................... 63, 65, 75, 77, 101, 187 Botulinum neurotoxin Brij................................................................................. 188 BSA blocking .................................................150, 181, 188 Bulk acoustic wave........................................................... 25 Buried layer ..................................................................... 58 Bursal disease......................................................... 169, 172 Bursal disease virus .........................................169, 172–178 BZE-DADOO ............................ 3, 6–8, 10, 11, 13–14, 19
C Cancer......................186, 255, 256, 276, 291, 313, 385, 391 Cantilever .......................................................51–70, 73–81 Capillary .......................................... 24, 170, 207, 218, 238, 246, 442, 444–446, 448, 451, 452, 454, 455 Capillary electrophoresis.......................................... 24, 442 Carbamate ......................................................115, 118–120 Carbaryl..................................................240, 241, 246–249 Carbenicillin ...................................................257, 258, 265 Carbon electrode .....................................85, 86–87, 90–92, 102, 103, 105, 119, 120, 145 Carboxylic-alkanethiol .............................................. 63–64 Casein... .......................................... 188, 198, 202, 225, 276 CCC-dsDNA ........................................................ 260–262 CCD...................................................................... 131, 327
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BIOSENSORS AND BIODETECTION 460 Index cDNA... ..................................................192, 354, 361, 362 Cell culture ..................... 159, 221, 292–293, 302–303, 391 Cell-targeting ................................................................ 291 ChEs.............................................................................. 119 Chloramphenicol ................................................... 257, 258 Cholinesterases .................................................... 3–20, 119 Chorionic gonadotropin ................................................ 171 Chromium ....................................................................... 38 Cleaning ........................................ 7, 62–63, 102, 420–421, 425–427, 429 Clenbuterol.....................................................169, 172, 174 Cloning.......................................... 327, 353–356, 362–365, 399, 403–404, 410–412 Coagulation ........................................................25, 32, 242 Colorimetric ...................................................85, 86, 88, 94 Combinatorial ......................................................... 24, 291 Competition zone.......................................................... 190 Competitive ......................................4, 6, 13, 14, 37, 39, 45, 63, 65, 85, 86, 162, 171, 185, 189, 190, 217, 219, 233, 237–239, 251, 423 Competitive immunoassay.......45, 63, 85, 86, 185, 190, 423 Composite electrode(s) ...........127, 129, 130, 148, 157, 158, 160–162, 164, 166 Conjugate pad..................................................170, 172, 173, 176, 177, 181, 218, 220, 222, 230–232 Conjugated ....................................................10, 11, 15, 43, 85–88, 92–94, 97, 127, 134–139, 145–147, 149–151, 170–179, 181, 185 Continuous flow immobilization Control line ...........................................170, 171, 173–179, 218, 231, 233, 237, 239, 240, 245–247 Cooling................................... 149, 243, 441, 442, 444–449 Cooling fan............................................................ 444, 445 Cotinine .........................................................217, 220–231 Counter electrode ............................................89, 105, 106, 121, 160, 267–270 Counter selection................................................... 402, 407 Coupling reagents.......................................................... 295 Crosslinking ............................. 64, 419, 422, 428–432, 437 Curvatures ................................................................. 52, 61 Cystic fibrosis .................................................127, 128, 131 Cytometry .....................................................292, 326, 327, 333–337, 343–345
DNA..............................................................23, 24, 26, 31, 99–112, 127–142, 146, 147, 185, 186, 188, 189–198, 206–208, 221, 256, 257, 259 analysis...................................... 127, 303, 363, 377, 441 sequencing ................................ 305, 363, 377, 403–404 dNTPs.. ..................................192, 257, 262, 305, 325, 339, 340, 389, 393–396, 403 Double sided adhesive tape ................................... 173, 176 Double-sided adhesive........................................... 173, 176
D
FACS.... ........................................... 49, 326, 329, 334–336, 342–343, 351, 352, 357–359, 360, 365–368, 372–376 Fiberglass dispensing ............................................. 172, 176 Fibrinogen ................................................................. 25, 32 Flow cell ............................................ 40, 41, 44, 45, 61, 62, 64, 74, 76, 77, 79, 359 Flow through ................................. 11, 13, 37, 41, 225, 234, 249, 261–262, 262, 361, 434 Fluidics .............................................................................. 4 Fluorescein .............................................186, 194, 335, 389
Data acquisition.......................12, 28, 79, 80, 441, 444, 446 DDT.... ...........................................................55, 56, 63–65 Dessicants ...................................................................... 230 Dextran...................................................................... 27, 30 Dichlorodiphenyltrichloroethane .................................... 55 Dielectric ink ............................................................. 87, 91 Dielectric layer........................................................... 91–93 Directed evolution ..................................323, 338, 351, 352 Displacement assay .................................................... 39, 45
E E coli.... ................................. 186, 256, 257, 260, 262–265, 275–282, 284, 285, 287, 317, 323, 325, 327, 333, 337, 354, 364, 377, 378, 403, 411, 414, 441, 443, 444, 450, 451, 453 EDC..... ...................55, 63, 64, 69, 210, 222–224, 435, 437 EDC-NHS................................................................ 64, 69 Electrical impedance ....................................................... 75 Electrochemical ................................... 3, 6, 85–97, 99–112, 115–124, 127–142, 145–154, 157–166, 255, 256, 259, 269–270, 271, 442 Electrocompetent ..........................................257, 263, 325, 331–332, 344 Electrocompetent yeast...................................331–332, 344 Electrode ........................................... 4–6, 9, 10, 12, 13, 26, 29, 38, 39, 41–43, 85–95, 99, 102–109, 110 Electrophoresis ........................................................24, 130, 134–137, 222, 304, 362–364, 369, 393, 401–406, 441, 442, 452, 454 Electroporation............................... 259, 260, 262–263, 332 ELISA.... ...................85–87, 90, 92–94, 186, 224–226, 385 Emission filter ............................................................... 296 Endosulfan .............................................240, 241, 247–249 Epoxy.... .........................74, 77, 78, 127, 129–132, 148, 224 Error-Prone PCR ........................... 329–330, 338, 358, 368 Estradiol .................................................................... 85–97 Etching ...........................................................26, 54, 59, 67 Excretory-secretory (ES) ........................173, 174, 178, 179 Extended aptamers .................................387–390, 392–394 Extrusion membranes .................................................... 196
F
BIOSENSORS AND BIODETECTION 461 Index Fluorescence .................................. 146, 221, 296, 313, 324, 334, 336, 341–342, 352, 359, 366, 368, 373–376, 378, 434, 454 Fluorescence activated cell sorting (FACS) .............................. 324, 326, 329, 334–336, 342, 351–352, 357–360, 365–368, 372–374 Fluorophore ............................................. 37, 146, 189, 335, 341, 400, 412 Foil envelops .................................................................. 230 Folding .........................................33n4, 329, 333–337, 373 Food............................................. 26, 53, 85, 147, 186, 197, 246, 247, 441, 443, 444, 453, 1187 Frequency variation ................................................... 28–30 Functionalization ............................... 29, 30, 51, 52, 54–56, 62, 63, 133–134, 136
G Gel-shift assay ................................................403, 408–410 Gelatin............................................ 181, 188, 277, 281, 388 Glucose .......................................... 116, 157, 159, 164–165, 258, 293, 334 Glucose oxidase .............................. 116, 157, 159, 164, 165 Glycolipid ....................................... 419–420, 422, 432, 433 Glycolipid immobilization............................................. 433 Gold dispersion ..................................................... 160, 164 Gold electrode ....................................... 6, 9, 10, 13, 39, 42, 102, 106, 256, 266, 269 Gold nanoparticles (AuNPs) .........................100, 127–129, 132–134, 136, 138–139, 141, 145, 146, 149, 150–152, 153 157–166, 239, 256 Graphite ........................................................117, 121, 127, 129–132, 147, 148, 157–164 Graphite electrode ..........121, 127, 129, 130, 147, 148, 157, 160–163 Graphite-based ink ........................................................ 117 Graphite–epoxy composite electrodes .......................... 127, 129, 130, 148 Graphite–Teflon electrodes ................................... 160–161 Grating ................................................... 4, 17, 40, 404, 449 Grating coupler ........................................................... 4, 17 Growth of Bacteriophage .............................................. 265
H Hapten .....................................39, 43, 45, 56, 63, 169, 171, 172, 179, 223, 237, 238, 240–242, 245, 247 Helper phage ..........................................257, 260, 264, 265 His-tagged..................................................................... 402 Hoechst 33258 ......................... 99, 102–104, 106, 108–109 Horse radish peroxidase (HRP)................................. 145, 151, 256, 259, 265 Human immunodeficiency virus (HIV) ..........169, 186, 217, 220–222, 230–231, 238 Hybridization ............99–112, 127–142, 146, 185–211, 443 Hybridoma ............................. 173, 352, 353, 360, 362, 379
Hydrogel......................................... 419, 420, 424, 434–437 Hydrophobic ................................... 69, 128, 182, 187–189, 242–243, 419–420, 422, 428, 432–433, 435–436 Hydrophobic Silane-Modified Surfaces ........................ 422 Hydrophobic silanization ...................................... 432–433
I Illumination........................................................... 452, 455 Immobilization ........... 16, 26–27, 29, 31, 37, 39, 42–43, 52, 56, 63, 64, 66, 73, 78, 102, 107, 119, 122, 128, 130, 134, 136, 138, 145, 157, 186, 194, 203, 224, 231, 238, 242, 245, 323, 402, 419–439 Immunoassay ........................................... 37, 44, 45, 51, 55, 63, 85, 86, 88, 94, 96, 185, 190, 191, 194, 237–251, 351, 419, 423 Immunochromatographic lateral flow strip test (ILFST) .............................................. 169–182 Impedance .................................. 75–77, 255, 256, 269, 271 Impedance analyzer ................................................... 76, 77 In vitro selection .............................................. 23, 399–415 Infectious bursal disease virus (IBDV) ...............................169, 172–175, 177–178 Infectious units ...............................................277, 281–283 Inhibition plot ....................................................... 123–124 Injecting system ............................................................... 61 Injection valve.......................................................54, 61, 64 Ink....... ................................... 86, 87, 90–92, 117, 119, 121 Insulating ink................................................................. 117 Integrated circuit ............................................................. 39 Intercalator .............................................................. 99–112 Interferometry ................................................................. 53 Isoelectric point ....................................................... 69, 180
K Kanamycin............................................. 257, 258, 260, 263, 277, 278, 280–282, 284, 285, 326, 333, 334, 337, 342, 359, 371, 376 Kinetic..... .............................3, 9, 17, 19, 37, 39, 41, 45–48, 161, 226, 358, 360, 367–368, 372–375, 377, 378, Kinetics analysis ............................. 358, 360, 367, 377–378
L Label reagents................................................................ 230 Label-free .......................................................4, 51, 73, 256 Labeling ........................................ 100, 145, 146, 181, 189, 229, 238, 243, 341, 342, 357–360, 367, 374–375 LabVIEW ................................................................. 54, 77 Laminating ............................................ 170, 172, 177, 187, 203–205, 222, 232, 233, 436 Laminator.......................................................172, 176, 233 Laser..... .......................... 52–55, 61–62, 312, 359, 381–382 Lateral Flow ........... 169–182, 185–211, 217–234, 237–251 Lateral flow strip test ......................................169–182, 217 Latex....................................... 185, 186, 188, 218, 237, 238
BIOSENSORS AND BIODETECTION 462 Index Lead Zirconate Titanate (PZT) .......................... 74–78, 80 LED..... ......................................................................... 443 Lens...... ................................................................... 62, 442 Library .......................................................23, 24, 259–264, 275–279, 281, 283–286, 291–292, 294, 297–298, 306, 329, 331–338, 340–342, 352, 358–360, 368, 369, 371–376, 378, 399–401, 407–408, 410, 412 Light.............................................. 10, 26, 54, 90, 146, 174, 205, 208, 218, 266, 277, 336, 341, 361–362, 366, 374, 452 Light scatter .......................................................... 205, 336 Lipid bilayers ......................................................... 188–189 Liposome ............................................................... 185–211 Luminescence ................................................................ 194 Lysozyme ...................................................................... 199
N
M
O
M13....... ....................................... 255–257, 259–260, 262, 263–265, 268, 355, 363, 411 Magnet .................................3, 99, 115, 119, 127–132, 134, 136–140, 145–148, 150, 152, 160–161, 164, 188, 204, 296, 314–315, 324 Magnetic beads.............................. 127–130, 134, 136, 140, 145, 147, 150, 296, 314–315, 324 Mask 59, 135, 314 Master Card ...................................................172, 176–177 Media............................................... 5, 116, 224, 258, 260, 263–265, 276–278, 282, 292–293, 297–300, 302, 306, 314, 325, 326, 331, 333–334, 336, 354, 356–360, 362, 365, 369, 371, 372, 376–377, 386, 391–392, 404 Medium ......................................5, 55, 61, 87, 89, 149, 189, 224, 258, 260, 279, 293, 324, 331, 359, 386, 391, 403–404, 411 Membrane .....................................................116, 169–173, 175–179, 185–188, 190–192, 194, 197, 202–205, 207, 208, 218, 220, 231, 232, 238, 240–242, 245, 250, 255–272, 283, 285, 385, 390, 392 Membrane blotting........................................172, 175–176, 178–179 Mercapto–Modified Surfaces ........................................ 421 Metal nanoparticles ........................................100, 127, 128 Metallization ................................................................... 59 Microfluidic ........................................................4, 191, 442 mRNA ................................................... 185, 193, 194, 196, 353, 360–361 MUA solution ................................................................. 10 Mutagenic library .................................. 358, 368, 369, 371, 372, 374–376, 378 Mutants ................................................. 260, 324, 329, 333, 336–338, 341–344, 352, 359, 372–373, 376–378, 380–381 Mutations ...................................... 324, 329, 333, 337, 351, 360, 372, 377–378,
Objective ............................................................... 218, 220 Optical beam deflection................................................... 53 Optical path length ......................................................... 62 Organophosphorus .........................................115, 118–120 Oriented immobilization ..................................... 43–44, 56 Oscillating circuit .................................................12, 39, 40 Oxide layer ................................................................ 54, 58 Oxidizing Virus Electrode..................................... 268–269
N-hydroxysuccinimide .........................................27, 30, 55, 64, 69, 222–224, 265–266, 268, 357, 424, 427–431, 435, 437 Nanoparticles.................................................100, 127–142, 145–154, 157–166, 239, 256, 292, 313 Nitrocellulose................................. 173, 175, 178–179, 187, 197, 203, 207–208, 217–220, 222, 230–233, 237, 242, 245 Nonadherent Cell ..................................292, 299, 300–301, 306, 307, 314, 315 NTA-lysine...................................................7–8, 10–11, 14 Nucleic acid sequence-based amplified (NASBA) .................... 185, 191–195, 199, 209–210 Nylon..........................................................165n1, 240, 242
P Paramagnetic beads .......................................129–130, 134, 136, 140, 145, 147, 150 Paraoxon ...................................................3, 6–8, 10, 14–15 Partitioning matrix ........................................................ 402 Pathogen...................................... 53, 73, 74, 127, 172, 186, 291, 292, 441, 443, 444, 450, 453 PCR........................104, 186, 191–193, 293, 294, 303–305, 325, 329–331, 338–340, 354–355, 358, 361–364, 368–370, 379, 385–391, 393–396 PDMS. ...................................................422–425, 433, 437 PEG.... ...........................104, 181, 188, 258, 260, 264, 278, 284, 285, 293–295, 301, 310, 311, 357, 358, 365 Peltier......................................................441–443, 452–453 Peptide purification ................................295–296, 311–312 Peptide synthesis ............................................294, 307–310 Peristaltic pump ..............................9, 11, 12, 40, 41, 54, 61, 77–79, 425, 437–438 Pesticide......................................3, 6, 51, 53, 54, 56, 62, 65, 115–124, 237–250 Phage Titration ..................................................... 302–304 Phase difference............................................................... 19 Phenolic Compounds ............................................ 161–162 Phospholipid .......................... 188–189, 198, 205, 206, 208 Photobleaching.............................................................. 341 Photodetector .................................................52–55, 60–62 Photodiode .................................................................... 443
BIOSENSORS AND BIODETECTION 463 Index Photodiode detector ...................................................... 443 Photolithography..................................................54, 59, 67 Photoresist ......................................................58, 59, 67, 68 PID........................................................................ 441, 446 Piezoelectric ........................................ 3–20, 23–34, 37–49, 60, 73–81, 99 Piranha solution .............................................54, 62, 76, 78 PLA.........................385–387, 389, 390–392, 395, 396–397 Planar screen-printed electrodes (SPEs) ....................................................... 105–108 Plasma 26, 27, 31–33 Plasmid .................................................. 325, 327, 328, 330, 331, 333, 337, 339, 353–355, 358, 359, 362–365, 369, 371–372, 377, 411 Platinum electrode.................... 13, 116, 129, 139, 147, 269 Polyester ............................................. 87, 90, 121, 187, 222 Polyethylene glycol (PEG) ............................104, 188, 211, 258, 260, 263–264, 264, 278, 284, 285, 293–295, 295, 301, 310–311, 311, 357, 358, 365, 370 Polyvinyl chloride (PVC).................... 89, 92, 131, 132, 148 Polyvinylidene fluoride (PVDF) ............................ 240, 242 Position sensitive photodetector (PSD) ........................... 52 Potentiometric ................................................116, 118, 119 Potentiostat....................89, 94, 95, 105, 106, 160, 267–269 Preclearing the phage library ..........................278, 283–284 Printing ink ..........................................................86, 87, 90 Probe design .................................................................. 195 Progesterone ..................................... 86, 157, 159, 162–165 Propagating phage ......................................................... 234 Propidium............................................... 3, 7, 11, 15, 16, 19 Propidium tricyclic ............................................................ 3 Proportional-derivative-integral (PID) ..........441, 446–447 Prostate specific membrane antigen (PSMA) .......255–272, 385, 387, 388, 390, 392, 397 Protease ...............................25, 42, 292, 298, 307, 315, 317 Protein A ...... 42–44, 48, 173, 174, 194, 222, 224, 230, 231, 240, 242, 243, 249 Proximity ligation .................................................. 385–397 Proximity ligation assay (PLA).............................. 385–397
R Reactive surface ......................................425, 427, 429–431 Reference electrode.............................. 9, 12, 13, 89, 91, 92, 94, 95, 105, 106, 116, 118, 121–123, 130, 132, 139, 147, 153, 160, 268, 269 Resonance........................................ 4, 9, 17, 38–40, 46, 52, 73, 75, 76, 119, 412 Resonance frequency .............4, 9, 38–40, 46, 52, 73, 75, 76 Reynolds number............................................................. 72 RNA..... ............................ 23, 185–211, 385–397, 399–401
S Saccharomyces cerevisiae ............................................324, 356 Salmonella ........................................ 441, 443, 444, 450, 453
Sample loop ..................................................................... 64 Sample pad .................................... 170, 172, 173, 176, 177, 188, 218, 220, 222, 230, 232 Sandwich ........................................... 27, 41, 127, 128, 136, 139–141, 145, 150–152, 171, 185–211, 217, 219, 233, 237, 238, 250, 386, 423, 436 Sandwich assay ......................................136–141, 151, 171, 189, 190, 219, 237, 238 Screen-printed .................... 85–97, 105–106, 117, 119–123 Screen-printed carbon electrodes (SPCEs)...................... 85–88, 90–94, 119, 120, 122 Screen-printed electrode............................90, 91, 105–106, 112, 117, 120, 121 Screen-printing ink ........................................86, 87, 90–92 Screen-stencil ............................................................ 90, 91 Screening ......................................... 48, 195, 221, 232, 247, 264–265, 276, 373 Screenprinted electrode (SPE) .................99, 105–108, 117 Secondary antibody ................ 145, 190–191, 194, 210, 326 Selecting phage...................................................... 263–264 Selective pressure .................... 352, 359, 360, 372–373, 376 Self-assembled monolayer (SAM) .................................. 43, 63, 64, 102, 110n3 Self-Assembly...................................................... 56, 63–64 Serum... ...................................................................7, 9, 27, 31, 32n1, 33n6, 42, 44, 75, 80, 85–89, 96, 120, 130, 147–149, 159, 171, 174, 178, 188, 222–225, 234, 237, 292, 293, 297, 299, 306, 314, 315, 325, 330, 386, 390, 402, 423, 425, 434, 444 Signal amplification ....................................................... 379 Signal digitalizing ...................................................... 54, 61 Silanization ........................................ 42–13, 419, 425–427, 429, 430, 432–433 Silanized crystals ....................................................... 42–43 Silicon......................................... 41, 51, 52, 54, 57–60, 442 Silicon oxide .............................................................. 52, 58 Silicon-On-Insulator (SOI) .................................54, 58, 67 Silver-based ink ..................................................... 117, 121 Single-chain Fv (scFv) ...................................247, 327, 333, 351–353, 355–360, 363–367, 369–382 single-stranded DNA (ssDNA)...............................23, 100, 104, 185, 259–262, 264, 265, 303, 357, 358, 365, 401, 408 Slicer..... ......................................................................... 233 SNPs.............................................................................. 451 Solid state relay...................................................... 444, 446 Spectrometer ..................................................202, 209, 296 Spectroscopy .......................................................... 255, 256 Spin coater................................................................. 76, 78 Sputter.. ........................................................................... 59 Stability .......................... 5, 24, 69, 157, 161, 189, 228, 238, 239, 276, 323, 324, 328, 329, 337, 338, 352, 378 Stabilization of baseline ......................................... 48n3, 79 Staining ......................................... 296, 313, 316, 333–336, 338, 341–343, 362–364, 369, 385
BIOSENSORS AND BIODETECTION 464 Index Steroid.. ..................................................................... 86, 89 Streptavidin ............................... 26–31, 128, 130, 134, 145, 147, 150, 186, 188, 192, 194, 197, 203, 204, 296, 297, 313–315, 326, 328, 341, 353, 358, 375, 386 Striper.. .......................................................................... 233 Styrene Subcloning...................................... 327, 355, 356, 364–365 Sulforhodamine B (SRB) ................... 15, 85, 189, 197, 200 Sump pad .............................................................. 218, 230 Surface functionalization .................... 30, 51, 54–56, 62–66 Surface passivation ................................................ 268–269 Surface plasmon resonance (SPR) ........................................ 4, 5, 17, 20, 63, 412 Syringe pump ................................................................ 175 Systematic Evolution of Ligands by Exponential Enrichment ( SELEX) .................23, 399, 400, 405
T T4 DNA ligase .............................. 257, 262, 356, 364, 389, 395, 403, 410 T4 polynucleotide kinase ................................257, 261–262 T7 DNA polymerase ............................................. 257, 262 Temperature control ...............................326, 446, 448, 449 Test line ................................................. 170, 171, 173, 174, 177, 179, 180, 217, 231, 234, 237, 238, 240, 241, 245–248 Tetracycline .....................257, 258, 265, 277, 278, 281–285 Tetrameric Peptide ......................... 295, 297, 309, 310, 313 Thermal stability ............................................323, 337–338 Thermocouple wire ....................................................... 444 Thermocycler ........................................................ 441–456 Thin-Film Resistive Heater ...................441, 443, 445–447 Thin-layer chromatography (TLC) ......................... 188, 197, 203, 204, 245, 406 Thioctic NHS-ester (NHS-TE)........................... 265–268, Thiol monolayers............................................................. 43 Thiol silanization................................................... 427–429 Thiol-Modified Surfaces ....................................... 247–429 Thrombin .......................................................23–26, 30–32 Titering ......................................... 224, 225, 281, 286, 287, 293, 294, 297–300, 302–305, 307, 312, 365–366
Toxin.................................................... 6, 74, 186, 323–324, 401, 432, 451–454 Trichinella ........................................169, 172–174, 177–179 Triton X-100 ..........................................188, 232, 299, 388 Trypsin ........................................... 297, 314, 386, 391–392 Tumor... .................................. 275–289, 292, 385, 386, 390 Tween... ................................................................. 239, 246 Tween-20................................27, 42, 87, 88, 104, 130, 133, 149, 176, 188, 242, 257–259, 402, 423 Tyrosinase ...............................................157–159, 161–163
U Uridine ...................................................257, 258, 260, 293
V Validation ...............................................227, 228, 246–247 Valve.... ......................................... 40, 41, 54, 61, 64, 76, 79 Vector.............................................................276, 279–280, 327–332, 340, 352, 354–357, 362–365, 369–371, 377, 403, 410 Virus electrode....................................................... 255–272 Voltammetric ........................................... 86, 103, 104, 116, 117, 129, 147, 169
W Wafer.... ................................................................54, 58, 59 Wafer dicing .................................................................... 67 Waveguide Working electrodes .................................. 12, 41, 85, 88–95, 105, 107, 116–118, 121, 122, 127, 130, 139, 147, 160, 267–270
Y Yeast display ..........................................324–327, 339, 343, 351, 352, 356, 364–365, 369 Yeast surface display ...............................323–347, 351, 353 Yeast transformation ...............................326, 356–359, 365
Z Zirconate titanate (PZT).................................... 74–78, 80
