Biosensors: a practical approach

Biosensors

Practical Approach Series For full details of the Practical Approach titles currently available, please go to www.oup.com/pas. The following titles may be of particular interest: Kinetic Analysis of Macromolecules (No 267) Edited by Kenneth Johnson

This new volume has been specifically designed to aid in the understanding of 'modern enzymology'. Of particular benefit to students and researchers are the hints and tips in each chapter for solving some of the most difficult problems encountered in modern enzymology. September 2003 0-19-852493-5 (Hbk); 0-19-852494-3 (Pbk) Phage Display (No 266) Edited by Tim Clackson and Henry B. Lowman

This book allows researchers to design and undertake all aspects of a phage display project, from designing an experimental strategy and constructing a library to performing selections and analyzing the results. January 2004 0-19-963874-8 (Hbk); 0-19-963873-X (Pbk) The Internet for Molecular Biologists (No 269) Edited by Clare Sansom and Robert Horton

This book is designed to help molecular biologists who are more at home at a laboratory bench than in front of a computer keyboard, to use the internet more effectively. Sansom and Horton provide a broad introduction to using Internet based computing resources including core databases, online resources plus tools and techniques for exploiting and authoring internet-distributed information. January 2004 0-19-963887-X (Hbk); 0-19-963888-8 (Pbk)

No. 268

Biosensors Second edition

A Practical Approach Edited by

Jonathan M. Cooper Department of Electronics, University of Glasgow, Glasgow G12 8LT, UK

Anthony E.G. Cass Department of Biological Sciences, Imperial College London, London SW7 2AZ, UK

OXPORD UNIVERSITY PRESS

OXFORD UNIVERSITY PRESS

Great Clarendon Street, Oxford OX2 6DP It furthers the University's objective of excellence in research, scholarship, and education by publishing worldwide in Oxford New York Auckland Bangkok Buenos Aires Cape Town Chennai Dar es Salaam Delhi Hong Kong Istanbul Karachi Kolkata Kuala Lumpur Madrid Melbourne Mexico City Mumbai Nairobi Sao Paulo Shanghai Taipei Tokyo Toronto Oxford is a registered trade mark of Oxford University Press in the UK and in certain other countries Published in the United States by Oxford University Press Inc., New York © Oxford University Press 1990, 2003 The moral rights of the author have been asserted Database right Oxford University Press (maker) First edition published 1990 Second edition published 2004 All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without the prior permission in writing of Oxford University Press, or as expressly permitted by law, or under terms agreed with the appropriate reprographics rights organization. Enquiries concerning reproduction outside the scope of the above should be sent to the Rights Department, Oxford University Press, at the address above You must not circulate this book in any other binding or cover and you must impose this same condition on any acquirer A catalogue record for this title is available from the British Library Library of Congress Cataloging in Publication Data (Data available) ISBN 0 19 963846 2 (Hbk) 0 19 963845 4 (Pbk) 10 9 8 7 6 5 4 3 2 1 Typeset by Newgen Imaging Systems (P) Ltd., Chennai, India Printed in Great Britain on acid-free paper by The Bath Press, Avon

Preface

Over the past 20 years, the field of biosensor research have had a significant impact in both laboratory research and the commercial sector. Over that period, biosensors have revolutionized the care and management of diabetes and have had important impacts in several other areas of clinical diagnostics. Europe, North America, and Asia-Pacific have all seen the rise of small- and mediumsized companies seeking technical and application niches in the manufacture or use of biosensors. The current activity in both gene and protein "biochips" can be seen as the latest set of tools that allow users who are not analytical science practitioners to make technically complex and reliable measurements with the minimum of intervention. Similarly, the concern about the dissemination of chemical or biological weapons and the need for their rapid and reliable detection will need to be met by devices that have many characteristics in common with biosensors. When the first edition of this book was published some 13 years ago, the field was dominated by electrochemical devices and the focus was primarily on clinical diagnostics. This dominance was partly a consequence of electrochemical (primarily amperometric) biosensors having been the first examples to be intensively investigated, but also because they formed the basis of the first mass produced and marketed devices (for blood glucose measurement). Whilst electrochemical methods are still major influences in biosensor research, optical methods have come increasingly to the fore and this trend is reflected in the current volume. Other significant developments have included the use of protein engineering methods to design and produce proteins that are better suited for the role as device component rather than constituents of a living cell, and the application of production methods such as screen printing that can manufacture very large number of sensors with a high degree of reproducibility. This latter is a prerequisite as biosensors move from laboratory demonstration to commercial product. The application focus of biosensors has also broadened with time and whilst clinical diagnostics probably remains the single biggest area, roles are also being found in environmental (including food) monitoring, personal security (including warfare), drug discovery, and basic biological research. v

PREFACE

Despite these changes, the field remains one of interdisciplinary challenge and exemplifies the current worldwide trend in engaging physical scientists and engineers in activities at the forefront of the biological sciences. As the latter move toward a more quantitative approach, the need for sophisticated tools for measurement and analysis of biological systems become ever more pressing and many of the ideas that drove and continue to drive biosensor research are directly relevant to this endeavor. The editors of this revised edition exemplify the discipline "hopping" that is characteristic of research in biosensors. One of us (JMC) trained as a biochemist and now holds a chair in an Electronics Department whilst the other (AEGC) trained as a chemist and holds a chair in a Biological Sciences Department. The contributors, who have generously donated their time and experience in writing the chapters that follow, demonstrate not only the progress in biosensors research over the past few years but also the experimental approaches that have enabled this progress. Finally, we would particularly like to acknowledge the contributions to biosensor research made by Professor Pierre Coulet. One of the pioneers of this field, Pierre has recently retired from his chair at the Universite Claude Bernard and we wish him a healthy and prosperous retirement, working on his farm in the Ardeche. JMC AEGC

vi

Contents

Protocol list xi Abbreviations xiii Contributors xv 1

2

Redox hydrogel-based electrochemical biosensors 1 Adam Heller 1 Electron conducting redox polymers in biosensors 1 1.1 Relationship to mediator-based sensors 3 1.2 The electrocatalytic activity of redox hydrogels and the "wiring" of enzymes 2 1.3 Dependence of the diffusivity of electrons on crosslinking and the advantage of composite electrodes 2 1.4 The redox polymers and their electrochemistry 3 1.5 Crosslinkers and crosslinking 4 2

Enzyme electrodes 4 2.1 Electron transfer between enzyme and polymer redox centers 4 2.2 Microscopic homogeneity and salt effects in the redox polymer-enzyme system 6 2.3 Optimal compositions 6 2.4 Special matrices 7

3

Specific sensor examples 7 3.1 Amperometric and potentiometric biosensors of substrates of "wired" redox enzymes 7 3.2 Sensor measuring the turnover rate of hydrolytic and other non-redox enzymes 8 3.3 In vivo glucose sensors 10 3.4 Affinity sensors 30

Hybridization at oligonucleotide sensitive electrodes 19 Daren J. Caruana \ Introduction 19 2

Function of oligonucleotide sensitive electrodes 20

3 Hybridization efficiency and sensitivity 21

vii

CONTENTS

3

4

Probe oligonucleotide structure and dynamics 22 4.1 Surface concentration 22 4.2 Probe length and orientation 23 4.3 Attachment of probe 25

5

Hybridization conditions 30 5.1 Temperature 31 5.2 Ionic strength 32 5.3 Base mismatch 33 5.4 Mass transport 34 5.5 Nonspecific adsorption 34 5.6 Other factors 36

6

Hybridization kinetics

7.

Summary 38

36

Screen-printing methods for biosensor production 41 Xian-En Zhang \ Introduction 41 2

Screen-printing technology 42 2.1 Materials and methods 42 2.2 Apparatus 47 2.3 Printing patterns 48 2.4 Printing process 49

3 Applications 51 3.1 Clinical diagnosis 51 3.2 Food analysis bioprocess control 52 3.3 Environmental monitoring 52 3.4 Other approaches 54 4 4

Conclusion 55

Kinetic modeling for biosensors 59 Philip Bartlett and Chee-Seng Toh \ Introduction 59 1.1 The purpose and practice of modeling 59 1.2 Enzyme kinetics 60 1.3 Basic electrochemistry 63 2

Modeling 69 2.1 The flux diagram for the membrane|enzyme|electrode 70 2.2 Simplifying assumptions 70 2.3 The flux equations 71 2.4 Solution of flux equations 73 2.5 Deriving a complete kinetic model 78 2.6 Experimental verification of approximate analytical kinetic models 81 2.7 Numerical simulation methods 82

3 Applications 89 4

viii

Kinetic modeling in other types of biosensors 89 4.1 Potentiometric enzyme electrodes 90 4.2 Optical and photometric biosensors 90 4.3 Immunosensors 91

CONTENTS

5 Conclusions 92 List of symbols 92 5

Bio-, chemi-, and electrochemiluminescence for fiber-optic biosensors 97 Loïc J. Blum and Pierre R. Coulet \ Introduction 97 2 Design of the biosensor 97 2.1 Optical waveguide 97 2.2 Setup 98 2.3 Light-emitting reactions 300 2.4 Preparation of the sensing layer 303 3 Examples of determinations with the luminescence sensors 105 3.1 ATP determination 305 3.2 NADH determination 305 3.3 Extension to other analytes using dehydrogenases as auxiliary enzymes 105 3.4 H2O2 determination 306 3.5 Extension to other analytes involving H2O2 detection 307 4 Concluding remarks 108

6

Determination of metal ions by fluorescence aniostropy: A practical biosensing approach 109 Richard Thompson, Badri Maliwal, Hui Hui Zeng, and Michele Loetz Cramer 1 Introduction and rationale 109 1.1 Why fluorescence anisotropy to determine metal ions? 309 2 Theory of anisotropy-based determination of metal ions 111 2.1 "Reagent" approaches 333 2.2 "Reagentless" approach 332 3 Fluorescent aryl sulfonamides for zinc(II) determination 114 4 Removal of zinc from carbonic anhydrase (CA) 115 5 Avoidance of metal ion contamination 117 6 Determination of Zn using a reagent approach 119 7 Determination of Cu and other ions by using a reagentless approach 123 8 Calibration of anisotropy 124

7

Fluorescence-based fiber-optic biosensors 131 David R. Walt, Caroline L Schauer, Shannon E. Stitzel, Michael S. Fleming, and Jason R. Epstein

\ Introduction 131

1.1 Fiber polishing 333

2 Single-analyte detection using an enzymatic sensing layer 134 2.1 Enzymatic sensing layer 334 2.2 PAN gel immobilization 335

ix

CONTENTS

3 Multi-analyte arrays 136 3.1 Immobilization via polymer photodeposition 136 3.2 Microwell array platform preparation 138 3.3 Live-cell array fabrication 146 4 8

Conclusions 151

Functional analysis of ion channels: Planar patch clamp and impedance spectroscopy of tethered lipid membranes 153 Michael Mayer, Samuel Terrettaz, Laurent Giovangrandi, Thierry Stora, and Horst Vogel \ Introduction 153 2

Planar patch clamp 154 2.1 Concept of patch clamp on a chip 354 2.2 Formation of planar bilayers on a chip 357 2.3 Chip-based planar bilayers: single-channel measurements of alamethicin pores 165

3 Impedance spectroscopy of tethered lipid membranes 168 3.1 Basics of impedance spectroscopy 168 3.2 Measuring technique and electrochemical cell 370 3.3 Hybrid lipid layer 370 3.4 Tethered lipid bilayers 174 3.5 Lipid bilayer tethered via surface-attached proteins 377 3.6 Highly insulating tethered lipid bilayers for single-channel experiments 179 9

Protein engineering for biosensors 185 Gianfranco Gilardi \ Introduction 185 2

Rational protein engineering 187 2.1 Modeling and calculations on protein structures 188 2.2 Site-directed mutagenesis 199

3 Directed evolution 222 3.1 Random mutagenesis: error prone PCR 224 3.2 Recombination: DNA shuffling 226 3.3 Functional screening of the library 230 4

Functional characterization of the mutants 234

5

Other aspects of protein engineering 234

6

Concluding remarks 238

Index

x

241

Protocol list

Probe oligonucleotide structure and dynamics Determination of solution concentration of single stranded oligonucleotide and assessment of purity of oligonucleotide 26 Attachment of oligonucleotides 28 CNBr activated coupling to an edge plan graphite electrode 29 Reactive electrophoretic deposition 30 Hybridization conditions Determination of melting temperature 32 Screen-printing technology Preparation of a screen-printed glucose electrode 50 Design of the biosensor Enzyme immobilization on collagen membranes 102 Enzyme immobilization on membranes Immunodyne ABC type from Pall-Gelman 103 Enzyme immobilization on Ultra Bind type membranes 103 Chemical modification of HRP with His-tag 104 Enzyme loading of IDA sepharose beads 104 Preparation of the sensing layer with a photocross-linkable polymer (PVA-SbQ) 104 Fluorescent aryl sulfonamides for zinc(ll) determination Synthesis of ABD-N 115 Removal of zinc from carbonic anhydrase (CA) Preparation of apoprotein 116 Avoidance of metal ion contamination Preparation of solutions free of contaminating metal ions 118 Determination of Zn using a reagent approach Determination of zinc concentration with ABD-N: a reagent approach 322 Calibration of anisotropy Determination of Cu and others by using a reagentless approach 324 Measurement and calibration of anisotropy 128 Single-analyte detection using an enzymatic sensing layer Silanization/functionalization of a fiber tip 335 PAN-enzyme sensors 135 xi

PROTOCOL LIST

Multi-analyte arrays Preparation of acryloylfluorescein (7) 137 Fabrication of an enzyme/pH array 137 Chemical etching of a germania-doped imaging fiber bundle 339 Internal encoding 340 External encoding 340 Enzyme immobilization by physical absorption 343 Enzyme immobilization with glutaraldehyde (15, 18) 342 Amine amplification with PEI (12, 13) 143 Periodate oxidation (19, 20) 143 Preparation of ssDNA probe labeled microspheres (13) 145 Preparation of molecular beacon-modified microspheres (23) 146 Fabrication of a fiber-optic based living array of cells 348 Cell viability assay with BCECF-AM 150 Cell viability test via pH sensitive nanospheres 150 Planar patch clamp Fabrication of silicon chips 357 Dry cleaning and activation of silicon chips 358 Wet cleaning and activation of silicon chips 359 Surface modification of the silicon chips 360 Preparation of giant unilamellar vesicles 363 Formation of PLBs on a silicon chip 162 Fabrication of Sylgard® pads 165 Impedance spectroscopy of tethered lipid membranes Preparation of a Ca2+-sensitive hybrid lipid layer electrode 372 Preparation of an OmpF-containing lipid bilayer tethered on a gold electrode 375 Formation of a proteolipid bilayer tethered by SLIC to a gold electrode 178 Rational protein engineering Protein sequence alignment 388 Creation of three-dimensional models 393 Molecular graphics: display and basic calculations on protein structures 394 Calculation of a map of electrostatic potentials on a protein surface 397 Hot start PCR 207 Agarose gel electrophoresis of DNA 209 Purification of PCR products from agarose gels 233 Megaprimer PCR 213 Restriction digest of DNA 215 Ligation of a DNA insert in a vector 236 Transformation of a plasmid in a bacterial host 217 Isolation of a plasmid from a bacterial host 238 Directed evolution QuikChange® PCR mutagenesis 223 Error prone PCR 225 DNA shuffling 227 High throughput screening for NAD(P)H-dependent activity 233 Other aspects of protein engineering 234 Fluoresence labelling of the S37C mutant of the maltose binding protein 235

xii

Abbreviations

BCA Bicinchoninic acid BEE Benzoin ethyl ether BLM Black lipid membrane bp base pair BSA Bovine serum albumin CVFF Consistent valence forcefield ECL Electrochemiluminescence ESFF Extensible systematic forcefield FIA flow low injection analysis HF Hydrofluoric acid HRP Horseradish peroxidase IPTG Isopropyl-1-thio-BD-galactopyranose ITCHY Iteractive Truncation for the Creation of Hybrid Enzymes LB Luria-Bertani broth NADH Nicotinamide adenine dinucleotide ND Neutral density PAA poly (acrylamide)-co-poly-(N-vinylimidazole) PAN Poly (acrylamide-co-N-acryloxysuccinimide) PBS Phosphate-buffered saline PCR Polymerase chain reaction pdb Protein data bank PEI Polyethyleneimine PLB Planar lipid bilayer PMT Photomultiplier tube PNA Peptide nucleic acids PVI poly(N-vinylimidazole) PVP poly(4-vinylpyridine) RMSD Root mean square deviation SBB Sodium borate buffer SCE saturated calomel electrode SEEP Staggered Extension Protocol SHIPREC Sequence Homology Independent Protein Recombination xiii

ABBREVIATIONS

SNP SPR SSPE TB UV

xiv

single nucleotide polymorphism surface plasmon resonance Saline sodium phosphate EDTA Terrific broth Ultraviolet

Contributors

Phil Bartlett

Michael S. Fleming

School of Chemistry University of Southampton Southampton SO17 1BJ United Kingdom

Department of Chemistry Tufts University Medford, MA 02155 USA

Loïc J. Blum Ldboratoire de Génie Enzymatique et Biomoléculaire Université Claude Bernard Lyon 1 69622 Villeurbanne CEDEX

Daren J. Caruana The Department of Chemistry University College London London WC1H OAJ United Kingdom

Pierre R. Coulet Ldboratoire de Génie Enzymatique et Biomoléculaire Université Claude Bernard Lyon 1 69622 Villeurbanne CEDEX

Michele L. Cramer Department of Biochemistry and Molecular Biology University of Maryland School of Medicine 655 West Baltimore Street Baltimore, Maryland 21201-1559

Gianfranco Gilardi Department of Biological Sciences Imperial College London, Technology and Medicine London SW7 2AY United Kingdom

Laurent Giovangrandi Ecole polytechnique fédérate de Lausanne (EPFL) Ecublens, CH-1015 Lausanne

Adam Heller The University of Texas at Austin Department of Chemical Engineering 1 University Station C0400 Austin, TX 78712-1062

Badri Maliwal Department of Biochemistry and Molecular Biology University of Maryland School of Medicine 655 West Baltimore Street Baltimore, Maryland 21201-1559

Jason R. Epstein Department of Chemistry Tufts University Medford, MA 02155 USA

Michael Mayer Ecole polytechnique fédérale de Lausanne (EPFL) Ecublens, CH-1015 Lausanne

xv

CONTRIBUTORS

Caroline L. Schauer

Chee-Seng Toh

Department of Chemistry Tufts University Medford, MA 02155 USA

Department of Chemistry National University of Singapore Singapore 117543

Shannon E. Stitzel

Ecole polytechnique fédérale de Lausanne (EPFL) Ecublens, CH-1015 Lausanne

Department of Chemistry Tufts University Medford, MA 02155 USA

Thierry Stora Ecole polytechnique fédérale de Lausanne (EPFL) Ecublens, CH-1015 Lausanne

Samuel Terrattaz Ecole polytechnique fédérale de Lausanne (EPFL) EcuUens, CH-1015 Lausanne

Richard B. Thompson Department of Biochemistry and Molecular Biology University Of Maryland School of Medicine 655 West Baltimore Street Baltimore, Maryland 21201-1559

xvi

Horst Vogel

David R. Walt Department of Chemistry Tufts University Medford, MA 02155 USA

Hui Hui Zeng Department of Biochemistry and Molecular Biology University Of Maryland School of Medicine 655 West Baltimore Street Baltimore, Maryland 21201-1559

Xian-En Zhang Wuhan Institute of Virology Chinese Academy of Sciences Wuhan 430071, Hubei, China

Chapter 1 Redox hydrogel-based electrochemical biosensors Adam Heller University of Texas at Austin, USA.

1 Electron conducting redox polymers in biosensors 1.1 Relationship to mediator-based sensors Redox hydrogel-based sensors are a subgroup of mediator-based sensors. Unlike their older cousins, the diffusional mediator-based sensors, their electron transport mediating redox centers are polymer-bound. When the resulting redox polymers are crosslinked on electrodes, they become insoluble, but swell in water to form redox hydrogels (1–4). These hydrogels can be as soft as Jell-O, or as tough as leather, depending on the extent of their crosslinking (1, 5). Upon hydration and swelling, the mobility of their segments is increased. This increased segmental mobility translates to an increase in the frequency of electron-transferring collisions between the tethered redox centers, and increases the electronic conductivity of the redox hydrogels. The diffusivity of electrons in the redox hydrogels is typically in the 10~ 6 -10~ 10 cm 2 s" a range (6-9). In addition to conducting electrons, the redox hydrogels, like other hydrogels, also conduct ions (10-18). Because they are permeable to water-soluble species, water-soluble chemicals like nitrite and biochemicals like ascorbate and dopamine can be electrooxidized or electroreduced in their three-dimensional matrices (19-21). When redox enzymes are co-immobilized in the redox hydrogels, their reaction centers can also be electrooxidized or electroreduced. The enzyme-containing redox hydrogels catalyze, therefore, the electrooxidation and the electroreduction of the substrates of the enzymes (22-54). Enzymes with FAD, FMN, PQQ., heme, and copper-containing redox centers have been co-immobilized and their centers are electroactive. Examples of the electrooxidized biochemicals include glucose, fructose, cellobiose, lactate, cholesterol, glycerol-3-phosphate, pyruvate, phenols, primary and secondary alcohols, histamine and other amines, D-amino acids, and glutamate. The electroreduced chemicals include O2 and H2O2. 1

ADAM HELLER

1.2 The electrocatalytic activity of redox hydrogels and the "wiring" of enzymes The electrocatalytic activity of the films formed by crosslinking depends on electron-transferring collisions between redox centers of the co-immobilized enzymes and those of the redox polymer (7-18). When these collisions are frequent enough to assure the efficient collection of the electrons from the enzyme reaction centers by the redox polymer, or the efficient delivery of electrons by the redox polymer to the reaction centers of the enzyme, the enzyme is said to be electrically "wired" and the redox polymer is said to "wire" the enzyme. The crosslinked redox polymers are poor electron conductors when dry: hydration is of essence for the electron transport to be fast and for the current density to be high. When the electron diffusivities reach, after hydration, 10~ 6 -10~ 9 cm 2 s" a and the co-immobilized redox enzyme turns over at a rate of >100 s ~ \ the limiting current density of substrate-electroreduction/oxidation on a semi-infinite planar enzyme electrode is typically of 10~ 3 -10~ 4 Acm~ 2 . Because their electron-transfer mediating centers are not leached, the "wired" enzyme electrodes can be used in microelectrodes (55-59), in experiments in vivo (60-68), and in flow systems (69-79).

1.3 Dependence of the diffusivity of electrons on crosslinking and the advantage of composite electrodes Nanocomposite enzyme electrodes can be made without covalent crosslinking by sequentially adsorbing countercharge films of a redox polymer, usually a polycation and enzymes which is often a polyanion at neutral pH (80-85). In the absence of covalent crosslinking, these films disintegrate in use and their components are slowly leached. Crosslinking by forming covalent bonds prevents the disintegration. While leaching of the redox polymer is prevented by crosslinking, excessive crosslinking reduces the mobility of the segments of the immobilized polymer and thereby the diffusivity of electrons and the current density (5). Excessive crosslinking-related losses are observed not only when the crosslinker forms covalent bonds, but also when ionic bridges are formed. Such bridges are formed in polycationic redox polymer films by large, little-hydrated anions like C1O4~, PO 4 3 ~, and citrate (5, 9,15). Small, well-hydrated anions like Cl~ are less effective crosslinkers. Even though increasing the concentration of NaCl does not reduce the diffusivity of electrons, such an increase can be damaging when it causes the decomposition of electrostatic complexes formed of a polyanionic enzyme and a polycationic redox polymer (86-88). The formation of an electrostatic complex prevents phase-separation of the enzyme and the redox polymer. In the case of polymers formed by the polymerization of weak bases, lowering of the pH increases the fraction of the bases that are protonated and thereby the charge on the polymers. This increases their hydration and thereby the mobility of their segments and the diffusivity of electrons. Plots of the dependence of the electron diffusivity on pH reveal a transition between the less electron-conducting 2

REDOX HYDROGEL-BASED ELECTROCHEMICAL BIOSENSORS

non-protonated and the more electron-conducting protonated forms of the redox polymers (6, 7). The density of positive charge on the crosslinked redox polymer is similarly increased upon increasing the fraction of the transition metal complexed bases (7-9). Even when the electron diffusivity is diminished by excessive crosslinking, high current densities of substrate electrooxidation or electroreduction can be maintained in hydrophilic graphite-based composite electrodes (89). Current densities of >2 mAcm~ 2 of glucose electrooxidation are reached when the composite is made with carbon graphite particles or carbon fibers, when the particles or fibers are of 10 |j,m diameter and are hydrophilic (89). When the particles are hydrophobic or smaller, the current density diminishes because the mass transport in the channels that are narrow or are not wetted by water is poor. This also raises the hydration, the segmental mobility, and the diffusivity of electrons. Nevertheless, when more than 20% of the heterocyclic nitrogens of poly(4-vinylpyridine) (PVP) or poly(N-vinylimidazole) (PVI) are complexed, for example, with [Os(bpy)2Cl]+'2 + , the density of ions and counterions is so high that the polymer becomes rigid and its electronic conductivity decreases.

1.4 The redox polymers and their electrochemistry The redox polymers that were most extensively used in the "wiring" of enzymes were made by attaching redox functions to PVP (1, 8, 9,17,19, 22-25, 90), PVI (32,

91-93), and poly(acrylamide)-co-poly(N-vinylimidazole) (PAA) (34). The redox functions were usually coordinatively pyridine or imidazole-attached complexes of Os 2+/3 + , such as [Os(bpy)2Cl] + /2+ (where bpy = 2,2'-bipyridine). Typically between 20% and 7% of the rings of the polymer were coordinated with the redox polymer, most often about 10%. An example of a redox polymer formed of a PVP is shown in Figure 1. The redox potentials of the polymers depend predictably on their chemistry. The following trends are consistent with the predictions of Lever (94) and are experimentally observed in the redox polymers. The more exoergic the binding of the ligands in the complex, the more positive its redox potential. Because the coordinative binding of osmium to vinyl pyridine is stronger than to N-vinylimidazole, the redox potentials of PVP-based polymers are positive by about 0.1 V relative to the related PVI-based complexes. Substitution of chloride ions in the inner coordination sphere of a complex by a nitrogencontaining heterocyclic ligand such as a pyridine or an alkylimidazole shifts the potential in the oxidizing direction. The redox potentials are more positive for Ru 2+ ' 3 + complexes than they are for Os2+'3+ complexes. The latter are positive relative to the Co2 + '3+ complexes when the ligands are similar. For a particular ligand, such as pyridine, 2,2'-bipyridine or 2,2',6',2"-terpyridine, an electron withdrawing substituent shifts the potential in the positive direction and an electron adding substituent shifts the potential in the negative direction. Thus, the potentials are shifted increasingly in the negative direction in the series of dicarboxylate, unsubstituted, methyl, methoxy and amine 3

ADAM HELLER

Figure 1 An enzyme wiring redox polymer derived of PVP. About one-sixth of the rings of the polymer are complexed with [Os(bpy)CI] +/2+ and about one-sixth are quaternized with bromoethylamine. Electrons are transferred through collisions between the tethered but mobile osmium-based redox centers. Quaternization makes the polymer water soluble and provides reactive primary amines for condensation with aldehydes produced by periodate oxidation of glycoenzymes.

4,4'-disubstituted 2,2'-bipyridines. When the same redox couple is incorporated in different polymers, the redox potential shifts in the positive direction as the polymer is made more polycationic or less polyanionic. Attaching of the redox function to a flexible and long spacer arm increases the volume element in which the couple can collide with a neighboring function and increases the diffusivity of electrons.

1.5 Crosslinkers and crosslinking A particularly convenient way of crosslinking polymers of nitrogen-containing heterocyclics is by reacting these with di- or tri-functional epoxides (1). Polyethylene glycol diglycidyl ether is a convenient crosslinker because of its solubility in water. Its epoxides react at ambient temperature with nitrogens of the heterocyclic rings. The polymers are also crosslinked by oxidizing their Os3+ redox centers to a higher oxidation state osmium compound, for example, by electrooxidation above >0.8 V versus SCE (SCE is the potential of the saturated calomel electrode).

2 Enzyme electrodes 2.1 Electron transfer between enzyme and polymer redox centers The exchange of electrons between a redox polymer and an enzyme is governed by the classical rules of electron transfer. In the multi-step electron transfer 4

REDOX HYDROGEL-BASED ELECTROCHEMICAL BIOSENSORS

process between an electrode and a substrate-reduced or substrate-oxidized enzyme reaction center, the transport of electrons is rapid only if all steps are thermodynamically downhill. Because the electrons cascade to or from the electrode, the redox potential of the polymers must be reducing with respect to the potential of the electrode and oxidizing with respect to the potential of the enzyme when a substrate is electrooxidized. The opposite is true when a substrate is electroreduced (25). The faster the self-exchange between the redox centers of a polymer, the faster the rate of electron transfer to or from an enzyme's redox center. The rate decreases exponentially with the distance between the fixed reaction center of the enzyme and the mobile redox couple tethered to the polymer. At equal distance, the exchange rates are higher when the reorganization energies are smaller. When the octahedral (or other) structure of a complex of a transition metal ion is preserved upon its oxidation and reduction, the reorganization energy is smaller, and electrons are more rapidly transferred when the distance is fixed and transferred to a more remote center when at an equally high rate. This is the case if the coordination is strong and the interatomic distances and angles do not change excessively when the complex is oxidized or reduced. Complexing of the metal ion thus provides for a higher self-exchange rate. However, because complexing also increases the size of the redox couple it can increase the distance of closest approach to an enzyme's deeply buried reaction center. When the reaction centers of the enzyme are deeply buried in its protein or glycoprotein, the redox couple approaches the center mainly through the channel in which the substrate of the enzyme diffuses. Because the rate of electron transfer drops by about a factor of 2.7 for each added angstrom, polymers with less bulky complexes are usually required for effective "wiring" of redox enzymes. By far the most studied enzyme has been glucose oxidase, the structure of which is known. The oxidation of the FADH2 centers of this enzyme is effectively mediated by both positively and negatively charged redox couples. Fe(CN)63~ '4~ is an effective and widely used redox mediator, as is the ferrocene/ferrocinium couple. The osmium complex based redox couples, which are larger, also mediate very effectively the exchange of electrons between the FADH2 centers of glucose oxidase and electrodes. When they are polymer-bound, their effectiveness depends, however, on their size: The bulkier the complexes, the less they approach the reaction centers of the enzyme and the slower the transfer of electrons from these to the redox polymer. Slow electron exchange implies, in the case of FADH2 enzymes, competition by oxygen: The electrons flow not only through the redox polymer to the electrode, but also to O2, the natural co-substrate of the enzyme, reducing it to hydrogen peroxide. Thus, the competition of oxygen for electrons of the redox polymers is a measure of the rate of electron transfer between the enzyme and the redox polymer. It is distinctly slower for very large complexes, even though these have high self-exchange rates. Because the closest distance of approach between a redox center of a polymer and the reaction center of the enzyme is through the narrow channel in which 5

ADAM HELLER

the substrate enters and the product leaves the enzyme, the rate of electron transfer depends on the flexibility of the polymer's backbone. High transfer rates and, therefore, high current densities are observed when the polymers are flexible. The electron-conducting, but rigid polypyrroles and polythiophenes yield high current density enzyme electrodes only when they were modified with pendant flexible hydrophilic or ionic redox functions (95-96).

2.2 Microscopic homogeneity and salt effects in the redox polymer-enzyme system Since the gain in entropy is small when two macromolecules are mixed, polymers are usually not miscible. They become miscible only when a bond is spontaneously formed, in an exoergic reaction, between the two macromolecules. The proteins and glycoproteins of enzymes are usually charged at neutral pH. Their charge is defined by their isoelectric points and on the pH of the solution. Phase separation of the redox polymer and the enzyme is avoided when the charge of the redox polymer is opposite to the charge of the enzyme (86-88). Glucose oxidase and lactate oxidase, which are polyanions at neutral pH, are effectively "wired" by redox polycations, including polymers made by attaching osmium complexes to PVP or PVL When the backbone is a polyanion, as it is in the case of polyacrylic acid, it is more difficult to avoid the separation of the phases and the polyanionic enzymes are less effectively connected to the electrodes by the polyanionic redox polymer. When microscopic homogeneity and effective "wiring" are provided by electrostatic interactions between the redox polymer and the enzyme, the quality of the "wiring" depends on the ionic strength of the solution used. Large salt effects are observed, because at high salt concentrations the charges of the interacting macromolecules are effectively screened by ions. In a concentrated NaCl solution, the dominant anions near the redox polycation are not carboxylate functions of the enzyme protein, but Cl~ anions. Similarly, the dominant cations in the proximity of the polyanionic enzyme are not the cations of the redox polymer, but Na + cations. When the charges of the enzyme and the polymer are balanced and the salt concentration is low, an insoluble enzyme-polymer adduct precipitates. This adduct re-dissolves at high salt concentration where the enzyme is not effectively "wired." (86-88) There is a clear optimum of NaCl concentration for the wiring of enzymes, which happens to coincide for glucose oxidase and its "wires" with the ~0.1 M physiological salt concentration at neutral pH. The loss of effective "wiring" and therefore in current density at high ionic strength can be overcome by crosslinking the enzyme and the polymer when the ionic strength is low. Such crosslinking prevents phase separation by preventing the dissociation of the enzyme and its "wiring" redox polymer.

2.3 Optimal compositions In well-crosslinked electrostatic adducts of enzymes and redox polymers, where phase separation is unlikely, the current densities reach an optimum when 6

REDOX HYDROGEL-BASED ELECTROCHEMICAL BIOSENSORS

enzyme is incorporated in the redox polymer, then decline when the fraction of the enzyme becomes excessive. The optimal enzyme concentration is that concentration where the rate of supply of electrons or holes (electron vacancies) by the substrate to the enzyme equals the rate of their removal from the enzyme to the electrode. When the polymer is in excess, there is not enough enzyme to supply electrons or holes to the wire. When the enzyme is in excess, the electroncurrent carrying capability of matrix is insufficient. For ~100 kDa enzymes turning over at about 1000 times per second the optimal enzyme weight fraction is about 0.3. For slower enzymes it is higher. Comparison of enzymes with different well-bound co-factors suggests that, in general, those with heme and PQQ functions transfer electrons more readily to redox polymers than enzymes with FAD and FMN co-factors. The reason is that the heme and PQQ. functions are usually closer to their periphery. The feasibility of preventing the escape of the weakly bound NADH and NADPH co-factors from the protein through forming specific chemical bonds between the NADH or the NADPH and either the redox polymer or the enzyme has been extensively studied. At this time it does not appear that such systems with immobilized NADH or NADPH can yield high (~10~ 3 Acm 2 ) current densities.

2.4 Special matrices The redox polymers and enzymes can be co-immobilized by the sol-gel method in hydrated silica matrices, in which the enzymes are stabilized (97, 98). These matrices are, however, insulating and hamper, in the absence of carbon or metal additives, the direct electrical communication between the enzymes and the electrodes (99-103). Electrodes can be conveniently made with carbon pastes (104-106), formed of graphite and mineral oil, where stabilization by poly(ethyleneimine) has been reported (105). The hydrated wired enzymes have been used in organic solvents and CO2 (107-109).

3 Specific sensor examples 3.1 Amperometric and potentiometric biosensors of substrates of "wired" redox enzymes When the reaction center of an enzyme is connected through its wire to an electrode, its turnover is observed as an electrical current. Examples of enzymes that were electrically wired include glucose oxidase, PQQ-glucose dehydrogenase, fructose dehydrogenase, cellobiose oxidase, lactate oxidase, cholesterol oxidase, glycerol-3-phosphate oxidase, pyruvate oxidase, tyrosinase, alcohol oxidase, amine oxidase, D-amino acid oxidase, glutamate oxidase, horseradish peroxidase, soybean peroxidase, and laccase. The resulting electrodes measured amperometrically the concentrations of the substrates of their enzyme.

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3.2 Sensor measuring the turnover rate of hydrolytic and other non-redox enzymes When multiple enzymes were incorporated in the redox hydrogel so as to form in situ a precursor the substrate of the electrically connected enzyme, the concentration of the precursor was amperometrically monitored. For example, by co-incorporating cholesterol oxidase and cholesterol esterase in the redox hydrogels, the concentration of the cholesterol esters was monitored. In this case, hydrolysis of the cholesterol esterase, catalyzed by the esterase, yielded cholesterol, which was electrooxidized by the wired cholesterol oxidase. In a second example, co-incorporation of acetylcholine esterase, choline oxidase, and horseradish peroxidase allowed the monitoring of acetylcholine esterase inhibitors. In this case, the acetylcholine esterase hydrolyzes acetyl choline to choline; the choline reacted with oxygen to yield betaine aldehyde and hydrogen peroxide, in the reaction catalyzed by choline oxidase; and the hydrogen peroxide was electroreduced on the electrically connected reaction centers of horseradish peroxidase. The reaction centers of choline oxidase were not wired. If they were connected, the two parallel connections of the choline oxidase and that of the horseradish peroxidase would have produced opposite currents and the measurements would not have been meaningful (41, 42, 110, 111). The amperometric enzyme electrodes formed by connecting the redox centers of enzymes to electrodes to redox hydrogels have, like all other enzyme electrodes, characteristic apparent Michaelis constants and maximal current densities. In the linear part of the current vs concentration diagram, the entire influx of the electroreduced or electrooxidized substrate is reacted. When the influx of substrate exceeds the rate at which it is electroreduced or electrooxidized, the current density no longer increases linearly with its concentration. In this case, the sensitivity is reduced and eventually the electrode "saturates," and the current no longer increases when the concentration of the substrate is increased. In order to extend the linear range of response to higher concentrations, one can overcoat the sensors with a membrane that limits the influx of the substrate. Because the enzyme electrodes have high current densities and it is attractive to miniaturize the electrodes, their overcoating with a membrane causes a particular problem. Pressure sealing of a membrane with an elastomeric O-ring, is obviously irrelevant, because O-rings and pressure seals are not available for very small electrodes. Forming the membrane in situ by casting from a solvent also poses a special problem because it is the residual solvent fraction that determines the pore size and distribution defining the relative rates of nucleation and diffusion, which depend on difficult to control parameters such as temperature gradients, vapor pressure gradients, and the presence or absence of nucleation centers. The most reproducible micromembranes are formed by sequential chemisorption reactions of polymers that react with each other. For example, a membrane of a glucose sensor can be formed by successive chemisorption of a polycation/ polyanion/polycation... sequence (68). The thickness and the mass transport

8

REDOX HYDROGEL-BASED ELECTROCHEMICAL BIOSENSORS

characteristics of these layer-by-layer formed membranes are relatively easy to control, allowing precise control of the permeabilities of the substrate, the products, and of all interferants. The layer-by-layer assembled micromembranes allow simultaneous optimization of all biosensor characteristics, including their linear range, their apparent engineering life (the life seen by the user), their drift (the variation of the sensitivity with time), and the sensitivity to interferants (68). Because the typical current densities are of 10~ 4 -10~ 3 AcrrT2 and because currents of 10"9 A can be conveniently measured using inexpensive potentiostats, the typical electrode dimensions are of 10~ 4 -10~ 6 cm2. The diameter of the smallest "wired" enzyme electrodes made thus far was of a few micrometers (55-59). These electrodes had microelectrode characteristics, the diffusion of the electrons to and from the electrode, as well as the substrates, and products being radial (24-26). Biosensors made with polycationic redox polymers have a problem not encountered in sensors made with diffusional mediators. The charge of the cations of the polymers is balanced by the charge of mobile anions in the solution, and the density of these anions in the redox hydrogel greatly exceeds their concentration in the solution. The permeability of the anions in the redox polymers is the product of their diffusivity and their concentration in the film. The high concentrations of anions and the high diffusivity makes the permeability to anions high. This high permeability exacerbates interference by anionic interferants, particularly by urate and ascorbate. While urate interference can be reduced, even eliminated, by poising the electrodes at potentials negative of 100 mV vs Ag/AgCl, interference by ascorbate is severe and can only be eliminated by a membrane in which the solubility of ascorbate is much lower than the solubility of glucose. This is the case when a membrane is formed of multiple layers of polyanions and polycations, the charges being internally balanced. In the absence of an excess of cationic sites, the anions are not particularly soluble in the membrane (68). When the potential at which the electrode is poised exceeds 100 mV (vs Ag/ AgCl), the electrooxidation of urate is particularly damaging. Because the concentration of urate in the redox polymer can be high, urate can electropolymerize and precipitate in the redox polymer film. The polymer formed is much less soluble in water than urate itself. Its presence is seen not only in the reduced current density but also in the reduction of the thickness of the electroactive layer. Transition metal ions, such as Zn2+ and Fe2+ can adversely affect the electrodes by coordinatively crosslinking pyridines and imidazoles of the polymers wiring the enzymes. The crosslinking reduces the diffusivity of electrons, and the thickness of the electroactive film shrinks. This detrimental effect can be avoided by incorporating transition metal ion coordinating centers in the membrane. When the membrane is made of sequential layers of polyacrylic acid and a polycation, then incorporation of polyfvinyl pyridines) in the polycationic layer stops the influx of Fe2 + and Zn2 + and stabilizes the electrode (68).

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ADAM HELLER

3.3 In vivo glucose sensors The tailoring of the characteristics of sensors made by connecting reaction centers of glucose oxidase with redox polymers to electrodes through their overcoating with precisely engineered micromembranes opens the way to implantable biosensors with well-defined characteristics. Specifically, sensors of about 5 x 10 ~ 4 cm2 area have been engineered to simultaneously provide a sensitivity of about 3 nAmM"1; a linear range of 1-30 mM; a drift of < 5% per day; and operational stability sufficient to eliminate the need to recalibrate the sensor for 3 days. When these sensors are overcoated with thin glucose permeable film of poly(ethyleneoxide) and then are heparinized, they accurately measure the subcutaneous and intravenous glycemia in animals. The implanted sensors have no leachable components and are insensitive to changes in partial pressure of oxygen. Because the typical resistance of the skin is about 100 kQ and the currents are of ~10 ~ 8 A, counter-reference electrodes larger than 1 cm2 can be external, that is, placed on the skin. For currents of < 50 nA, the potential drop is about < 1 mV, which is insufficient to perturb the measurement (68).

3.4 Affinity sensors By attaching affinity reagents to redox enzymes, it is possible to construct amperometric electrodes tracking affinity reactions. The current increase results in the electrode becoming electrocatalytic when electrical contact is established between the electrode and the reaction centers of the enzyme with which one of the members of the affinity couple is labeled (112-120). Although the affinity reactions can be readily observed with any redox enzyme, it is convenient to use peroxidases, because their heme centers are easy to wire and their turnover rate is high. Their substrate, H2O2, can be generated internally in the affinity sensing layer (117). Hydrogen peroxide is generated by reacting choline with dissolved oxygen to form betaine aldehyde and hydrogen peroxide, a reaction catalyzed by co-immobilized choline oxidase, the reaction centers of which are not wired. The affinity reactions can be carried out on rotating electrodes, so as to prevent the transport of the affinity reagents to the surface of the electrodes from controlling the time required for the detection. The rate of the affinity reaction depends on the transport of the affinity reagents in the redox hydrogels which can be slow for reagents larger than 100 kDa, the detection requiring 10-20 min. In hybridization sensors it is convenient to use soybean peroxidase as the labeling enzyme. Unlike horseradish peroxidase, soybean peroxidase is stable enough to be used for temperatures up to 75 °C (35). Thus, when the enzyme is bound to a DNA segment and if the segment hybridizes to a segment of DNA or RNA immobilized in the redox polymer is targeted, good specificity can be achieved by running the hybridization reactions near the melting temperature of the hybrid. Because the melting temperature of hybrids with defects is lower, one can readily distinguish between perfect and imperfect hybrids. Using the DNA modified redox polymers, one can detect in minutes the presence of about 10

REDOX HYDROGEL-BASED ELECTROCHEMICAL BIOSENSORS 40,000 copies of the enzyme labeled DNAon the electrode. Furthermore, a single mismatch in an 18 base pair sequence can be readily observed (119). Beyond the target hybridization of a redox hydrogel bound DNA sequence with an enzyme labeled sequence, it is also possible to observe the hybridization of a long (~100 base) DNA or RNA sequence with a sequence bound to the redox hydrogel. In this case the long sequence is hybridized to the sequence on the electrode; the presence of the long sequence is then confirmed by hybridizing another part of it with a second complementary sequence of the now immobilized long chain, this one labeled. This scheme is applicable equally to the amperometric detection of DNA and RNA. Because the abundance of RNA organisms is greater, by about 10,000, than the abundance of DNA, the method appears at this time to be suitable for the detection of organisms.

References 1. Gregg, B. A. and Heller, A. (1991) Redox polymer films containing enzymes. 1. A redox-conducting epoxy cement: synthesis, characterization, and electrocatalytic oxidation of hydroquinone, J. Phys. Chem., 95(15), 5970-5. 2. Hillman, A. R., Loveday, D. C., Glidle, A., Vos, J. G., Clarke, A. P., Kelly, D., and Bruckenstein, S. (1993) Electrochemical quartz crystal microbalance studies of redox polymer films, Macromol Symp., 80, 323-35. 3. Calvo, E. J. and Etchenique, R. (1999) Viscoelastic changes in Os-containing poly(allylamine) based redox hydrogels for amperometric enzyme electrodes. ]. Phys. Chem. B, 103, 8944-50. 4. Etchenique, R. A. and Calvo, E. J. (1999) Gravimetric measurement in redox polymer electrodes with the EQCM beyond the Sauerbrey limit, Electrochem, Commun., 1,167-70. 5. Binyamin, G. and Heller, A. (1999) Stabilization of wired glucose oxidase anodes rotating at 1000rpm at 37 °C, J. Electrochem. Soc., 146, 2965-7. 6. Aoki, A. and Heller, A. (1993) Electron diffusion coefficients in hydrogels formed of crosslinked redox polymers, J. Phys. Chem., 97, 11014-19. 7. Aoki, A., Rajagopalan, R., and Heller, A. (1995) Effect of quaternization on electron diffusion coefficients for redox hydrogels based on poly(4-vinylpyridine), J. Phys. Chem., 99, 5102-10. 8. Rajagopalan, R., Aoki, A., and Heller, A. (1996) Effect of quaternization of the glucose oxidase "wiring" redox polymer on the maximum current densities of glucose electrodes,/. Phys. Chem., 100, 3719-29. 9. Rajagopalan, R. and Heller, A. (1997) Electrical "wiring" of glucose oxidase in electron conducting hydrogels, Ch. 7. In Molecular electronics (ed. J. Jortner and M. Ratner), pp. 241-54, Blackwell Science Publishers. 10. Pickup, P. G., Kutner, W., Leidner, C. R., and Murray, R. W. (1984) Redox conduction in single and bilayer films of redox polymer, J. Am. Chem. Soc., 106, 1991-8. 11. Oyama, N. and Hatozaki, O. (1988) Redox active membranes and electron transfer, Kagaku (Kyoto), 43, 280-1. 12. Forster, R. J., Vos, J. G., and Lyons, M. E. G. (1991) Controlling processes in the rate of charge transport through osmium 2,2'-bipyridine authors poly(4-vinylpiperidine) complex [Os(bipy)2(PVP)nCl]Cl redox polymer-modified electrodes, J. Chem. Soc., Faraday Trans., 87, 3761-7. 13. Mathias, M. F. and Haas, O. (1992) An alternating current impedance model including migration and redox-site interactions at polymer-modified electrodes,/. Phys. Chem., 96, 3174-82.

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ADAM HELLER 14. Surridge, N. A., Zvanu, M. E., Keene, F. R., Sosnoff, C. S., Silver, M., and Murray, R. W. (1992) Effects of mixed-valent composition and bathing environment on solid-state electron self-exchanges in osmium bipyridine redox polymer films, /. Phys. Chem., 96, 962-70. 15. Mathias, M. F. and Haas, O. (1993) Effect of counterion type on charge transport at redox polymer-modified electrodes, /. Phys. Chem., 97, 921-5. 16. Kelly, D. M. and Vos, J. G. (1994) Osmium and ruthenium poly(pyridyl) redox polymers as electrode coatings, Ekctroact. Polym. Ekctrochem., 2, 173-232. 17. Sharp, M. and Larsson, H. (1995) Studies of the temperature dependence of charge propagation rates in quaternized poly(4-vinylpyridine) polymers containing electrostatically bound and coordinated redox sites, Electroanal. Chem., 386, 189-95. 18. Sun, L, Jiao, K., and Weber, S. G. (1998) Charge transport through osmium-containing redox polymers in nitrophenyl-based solvents: effect of solvent size, /. Phys. Chem. B, 102, 1945-50. 19. Doherty, A. P. and Vos, J. G. (1992) Electrocatalytic reduction of nitrite at an [Os(bipy)2(PVP)10Cl]Cl-modified electrode, /. Chem. Soc., Faraday Trans., 88, 2903-7. 20. Doherty, A. P., Stanley, M. A., and Vos, J. G. (1995) Electrocatalytic oxidation of ascorbic acid at [osmium(2,2'-bipyridyl)2-(poly-4-vinylpyridine)10Cl]Cl modified electrodes: implications for the development of biosensors based on osmium-containing redox relays, Analyst (Cambridge, UK), 120, 2371-6. 21. Ju, H., Ni, J., Gong, Y., Chen, H., and Leech, D. (1999) Electrocatalytical oxidation and determination of dopamine at redox polymer/Nafion modified electrodes, Anal. Lett, 32, 2951-64. 22. Gregg, B. A. and Heller, A. (1990) Cross-linked redox gels containing glucose oxidase for amperometric biosensor applications, Anal. Chem., 62, 258. 23. Heller, A. (1990) Electrical wiring of redox enzymes, Ace. Chem. Res., 23, 128. 24. Gregg. B. A. and Heller, A. (1991) Redox polymer films containing enzymes. 2. Glucose oxidase containing enzyme electrodes,/. Phys. Chem., 95(15), 5976-80. 25. Heller, A. (1992) Electrical connection of enzyme redox centers to electrodes, /. Phys. Chem., 96, 3579-87. 26. Katakis, I. and Heller, A. (1992) L-a-glycerophosphate and L-lactate electrodes based on the electrochemical "wiring" of oxidases, Anal. Chem., 64, 1008-13. 27. Elmgren, M., Lindquist, S.-E., and Henriksson, G. (1992) Cellobiose oxidase crosslinked in a redox polymer matrix at an electrode Surface—a new biosensor,/. Electroanal. Chem., 341, 257-73. 28. Wang, D. L. and Heller, A. (1993) Miniaturized flexible amperometric lactate probe, Anal. Chem., 65, 1069-73. 29. Ye, L., Hammerle, M., Olsthoorn, A. J. J., Schuhmann, W., Schmidt, H.-L, Duine, J. A., and Heller, A. (1993) High current density "wired" quinoprotein glucose dehydrogenase, Anal. Chem., 65, 238-41. 30. Vreeke, M., Maidan, R., and Heller, A. (1992) Hydrogen peroxide and (3-nicotinamide adenine dinucleotide sensing amperometric electrodes based on electrical connection of horseradish peroxidase redox centers to electrodes through a three-dimensional electron relaying polymer network, Anal. Chem., 64 3084-90. 31. Elmgren, M., Lindquist, S.-E., and Sharp, M. (1993) Charge propagation through a redox polymer film containing enzymes—effects of enzyme loading, pH and supporting electrolyte,/. Electroanal. Chem., 362, 227-35. 32. Ohara, T. J., Rajagopalan, R., and Heller, A. (1994) "Wired" enzyme electrodes for amperometric determination of glucose or lactate in the presence of interfering substances, Anal. Chem., 66, 2451-7. 33. Prayda, M., Jungar, C. M., Iwuoha, E. L, Smyth, M. R., Vytras, K., and Ivaska, A. (1995) Evaluation of amperometric glucose biosensors based on co-immobilization of glucose

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34. 35. 36. 37. 38.

39. 40. 41.

42. 43. 44. 45. 46. 47. 48. 49. 50. 51.

oxidase with an osmium redox polymer in electrochemically generated polyphenol films, Anal. Chim. Ada, 304, 127-38. De Lumley-Woodyear, T., Rocca, P., Lindsay, J., Dror, Y., Freeman, A., and Heller, A. (1995) Polyacrylamide based redox polymer for connecting redox centers of enzymes to electrodes, Anal. Chem., 67, 1332-8. Vreeke, M. S., Yong, K. T., and Heller, A. (1995) A Thermostable biosensor of hydrogen peroxide, Anal. Chem., 67, 4247-9. Robinson, G., Leech, D., and Smyth, M. R. (1995) Electrically "wired" tyrosinase enzyme inhibition electrode for the detection of respiratory poisons, Electroanalysis, 7, 952-7. Vijayakumar, A. R., Csoregi, E., Heller, A., and Gorton, L. (1996) Alcohol biosensors based on coupled oxidase-peroxidase systems, Analytica Cliimica Acta, 327, 223-34. Ruzgas, T., Csoregi, E., Katakis, I., Kenausis, G., and Gorton, L. (1996) Preliminary investigations of an amperometric oligosaccharide dehydrogenase-based electrode for the detection of glucose and some other low molecular weight saccharides, J. Mol. Recognit., 9, 480-4. Hedenmo, M., Narvaez, A., Dominguez, E., and Katakis, I. (1996) Reagentless amperometric glucose dehydrogenase biosensor based on electrocatalytic oxidation of NADH by osmium phenanthrolinedione mediator, Analyst (Cambridge, UK), 121,1891-5. Larsson, T., Elmgren, M., Lindquist, S.-E., Tessema, M., Gorton, L, and Henriksson, G. (1996) Electron transfer between cellobiose dehydrogenase and graphite electrodes, Anal. Chim. Acta, 331, 207-15. Katakis, I. and Heller, A. (1997) Electron transfer via redox hydrogels between electrodes and enzymes, In Frontiers in biosensorics, I, Fundamental aspects (ed. F. W. Scheller, F. Schubert, and I. Fedowitz), pp. 229-41, Birkhauser Verlag, Basel, Switzerland. Kenausis, G., Chen, Q., and Heller, A. (1997) Electrochemical glucose and lactate sensors based on "wired" thermostable soybean peroxidase operating continuously and stably at 37 °C, Anal. Chem., 69, 1054-60. Tessema, M., Csoregi, E., Ruzgas, T., Kenausis, G., Solomon, T., and Gorton, L. (1997) Oligosaccharide dehydrogenase-modified graphite electrodes for the amperometric determination of sugars in a flow injection System, Anal. Chem., 69, 4039-44. Paredes, P. A., Parellada, J., Fernandez, V. M., Katakis, L, and Dominguez, E. (1997) Amperometric mediated carbon paste biosensor based on D-fructose dehydrogenase for the determination of fructose in food analysis, Biosens. Bioekctron., 12, 1233-43 Daigle, F. and Leech, D. (1997) Reagentless tyrosinase enzyme electrodes: effects of enzyme loading, electrolyte pH, ionic strength, and temperature, Anal. Chem., 69, 4108-12. Trudeau, F., Daigle, F., and Leech, D. (1997) Reagentless mediated laccase electrode for the detection of enzyme modulators, Anal. Chem., 69, 882-6. Stigter, E. C. A., De Jong, G. A. H., Jongejan, J. A., Duine, H. A., Van DerLugt, J. P., and Somers, W. A. C. (1997) Electron transfer and stability of a quinohemoprotein alcohol dehydrogenase electrode,/. Chem. Technol. Biotechnol, 68, 110-16. Daigle, F., Trudeau, F., Robinson, G., Smyth, M. R., and Leech, D. (1998) Mediated reagentless enzyme inhibition electrodes, Biosens. Bioelectron., 13, 417-25. Somers, W. A. C., Stigter, E. C. A., Van Hartingsveldt, W., and Van Der Lugt, J. P. (1999) Enantioselective oxidation of secondary alcohols at a quinohemoprotein alcohol dehydrogenase electrode, Appl. Biochem. Biotechnol, 75, 151-61. Alkasrawi, M., Popescu, I. C., Mattiasson, B., Csoregi, E., and Laurinavicius, V. (1999) A redox hydrogel integrated PQ_Q;glucose dehydrogenase based glucose electrode, Anal. Common., 36, 395-8. Belay, A., Collins, A., Ruzgas, T., Kissinger, P. T., Gorton, L, and Csoregi, E. (1999) Redox hydrogel based bioenzyme electrode for L-glutamate monitoring, J. Pharm. Biomed. Anal, 19, 93-105.

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ADAM HELLER 52. Gajovic, N., Binyamin, G., Warsinke, A., Scheller, F. W., and Heller, A. (2002) Operation of a miniature redox hydrogel-based pyruvate sensor in undiluted deoxygenated calf serum, Anal. Chem., 72, 2963-68. 53. Niculescu, M., Frebort, I., Pec, P., Galuszka, P., Mattiasson, B., and Csoregi, E. (2000) Amine oxidase based amperometric biosensors for histamine detection, Electroanalysis, 12, 369-75. 54. Niculescu, M., Nistor, C, Frebort, I., Pec, P., Mattiasson, B., and Csoregi, E., (2000) Redox hydrogel-based amperometric bienzyme electrodes for fish freshness monitoring, Anal. Chem., 72, 1591-7. 55. Pishko, M. V., Michael, A. C., and Heller, A. (1991) Amperometric glucose microelectrodes prepared through immobilization of glucose oxidase in redox hydrogels, Anal. Chem., 63, 2268-72. 56. Rohde, E., Dempsey, E., Smyth, M. R., Vos, J. G., and Emmons, H. (1993) Development of a flow-through electrochemical detector for glucose based on a glucose oxidasemodifled microelectrode incorporating redox and conducting polymer materials, Anal. Chim. Ada, 278, 5-16. 57. Horrocks, B. R., Schmidtke, D., Heller, A. and Bard, A. J. (1993) SECM 24. Enzyme microelectrodes for the measurement of hydrogen peroxide at surfaces, Anal. Chem., 65, 3605-14. 58. Sakai, H., Baba, R., Hashimoto, K, Fujishima, A., and Heller, A. (1995) Local detection of photoelectrochemically produced H2O2 with a "Wired" horseradish peroxidase microsensor, J. Phys. Chem., 99, 11896-900. 59. Niwa, O., Kurita, R., Liu, Z., Horiuchi, T., and Torimitsu, K. (2000) Subnanoliter volume wall-jet cells combined with interdigitated microarray electrode and enzyme modified planar microelectrode, Anal. Chem., 72, 949-55. 60. Csoregi, E., Quinn, C. P., Schmidtke, D.W., Lindquist, S.-E., Pishko, M.V.,Ye, L, Katakis, I., Hubbell, J. A., and Heller, A. (1994) Design, characterization, and one-point in vivo calibration of a subcutaneously implanted glucose electrode, Anal. Chem., 66, 3131-8. 61. Csoregi, E., Schmidtke, D. W., and Heller, A. (1995) Design and optimization of a selective subcutaneously implantable glucose electrode based on "wired" glucose oxidase, Anal. Chem., 67, 1240-44. 62. Quinn, C. P., Pishko, M. V., Schmidtke, D. W., Ishikawa, M., Wagner, J. G., Raskin, P., Hubbell, J. A., and Heller, A. (1995) Kinetics of glucose delivery to subcutaneous tissue in rats measured with 0.3 mm amperometric microsensors, Am. ]. Physio!., 269 (Endocrinol. Metab. 32), E155-E161. 63. Schmidtke, D., Pishko, M. V., Quinn, C. P., and Heller, A. (1996) Statistics for critical clinical decision making based on readings of pairs of implanted sensors, Anal. Chem., 68, 2845-49. 64. Schmidtke, D. W. and Heller, A. (1998) Accuracy of the one-point in vivo calibration of "wired" glucose oxidase electrodes implanted in jugular veins of rats in periods of rapid rise and decline of the glucose concentration, Anal. Chem., 70, 2149-55. 65. Wagner, J. G., Schmidtke, D. W., Quinn, C. P., Fleming, T. F., Bernacky, B., and Heller, A. (1998) Continuous amperometric monitoring of glucose in a brittle diabetic chimpanzee with a miniature subcutaneous electrode, Proc. NatlAcad. Set, 95, 6379-82. 66. Schmidtke, D. W., Freeland, A. C., Heller, A., and Bonnecaze, R. T. (1998) Measurement and modeling of the transient difference between blood and subcutaneous glucose concentrations in the rat after injection of insulin, Proc. NatlAcad. Set, 95, 294-9. 67. Heller, A. (1999) Implanted electrochemical glucose sensors for the management of diabetes, Ann. Rev. Biomed. Eng., 1, 153-75. 68. Chen, T., Friedman, Keith A., Lei, I., and Heller. A. (2000) In situ assembled mass transport controlling micromembranes and their application in implanted amperometric glucose sensors, Anal. Chem., 72, 3757-63.

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REDOX HYDROGEL-BASED ELECTROCHEMICAL BIOSENSORS 69. Csoregi, E., Laurell, T., Katakis, I., Heller, A., and Gorton, L. (1995) On-line glucose monitoring using microdialysis sampling and amperometric detection based on "wired" glucose oxidase in carbon paste, Microchimica Acta, 121, 31-40. 70. Huang, T., Yang, L., Gitzen, J., Kissinger, P. T., Vreeke, M., and Heller, A. (1995) Detection of basal acetylene line in rat brain microdialysate, J. ChromatographyB, Biomed. Appl., 670, 323-27. 71. Yang, L., Janle, E., Huang, T., Gitzen, J., Kissinger, P. T., Vreeke, M., and Heller, A. (1995) Application of "Wired" peroxidase electrodes for peroxide determination in liquid chromatography coupled to oxidase immobilized enzyme reactors, Anal. Chem., 67, 1326-31. 72. Kato, T., Liu, J. K., Yamamoto, K., Osborne, P. G., and Niwa, O. (1996) Detection of basal acetylcholine release in the microdialysis of rat frontal cortex by high performance liquid chromatography using a horseradish peroxidase-osmium redox polymer electrode with pre-enzyme reactor, J. Chromatogr., B: Biomed. Appl., 682, 162-6. 73. Yang, L. and Kissinger, P. T. (1996) Determination of oxidase enzyme substrates using cross-flow thin-layer amperometry, Electroanalysis, 8, 116-21. 74. Tessema, M., Larsson, T., Buttler, T., Csoregi, E., Ruzgas, T., Nordling, M., Lindquist, S.-E., Pettersson, G., and Gorton, L. (1997) Simultaneous amperometric determination of some mono-, di-, and oligosaccharides in flow injection and liquid chromatography using two working enzyme electrodes with different selectivity, Anal. Chim. Acta, 349, 179-88. 75. Tessema, M., Csoregi, E., Ruzgas, T., Kenausis, G., Heller, A., Solomon, T., and Gorton, L. (1997) Oligosaccharide dehydrogenase modified graphite electrodes for the amperometric determination of sugars in flow injection system, Anal. Chem., 69, 4039-44. 76. Niwa, O., Horiuchi, T., Kurita, R., and Torimitsu, K. (1998) Online electrochemical sensor for selective continuous measurement of acetylcholine in cultured brain tissue, Anal. Chem., 70, 1126-32. 77. Osborne, P. G., Niwa, O., and Yamamoto, K. (1998) Plastic film carbon electrodes: enzymic modification for online, continuous, and simultaneous measurement of lactate and glucose using microdialysis sampling, Anal. Chem., 70, 1701-6. 78. Larsson, N., Ruzgas, T., Gorton, L, Kokaia, M., Kissinger, P., and Csoregi, E. (1998) Design and development of an amperometric biosensor for acetylcholine determination in brain microdialyzates, Electrochim. Acta, 43, 3541-54. 79. Yao, T. and Ogawa, H. (2000) Highly sensitive and selective detection of pyruvate by amperometric flow-injection analysis based on enzymatic substrate recycling and sensitive detection of hydrogen peroxide, J. How Injection Anal., 17, 37-42. 80. Pishko, M. V., Katakis, L, Lindquist, S.-E., Heller, A., and Degani, Y. (1990) Electrical communication between graphite electrodes and glucose oxidase/redox polymer complexes, Mo!. Cryst. Liq. Cryst., 190, 221. 81. Pishko, M. V., Katakis, L, Lindquist, S.-E., Ye, L., Gregg, B. A., and Heller, A. (1990) Direct electrical communication between graphite electrodes and surface adsorbed glucose oxidase/redox polymer complexes, Angew. Chem. Intl. Ed., 29, 1, 82. 82. Sirkar, K., Revzin, A., and Pishko, M. V. (2000) Glucose and lactate biosensors based on redox polymer/oxidoreductase nanocomposite thin films, Anal. Chem., 72, 2930-36. 83. Narvaez, A., Suarez, G., Popescu, I. C., Katakis, L, and Dominguez, E. (2000) Reagentless biosensors based on self-deposited redox polyelectrolyte-oxidoreductases architectures, Biosens. Bioelectron., 15, 43-52. 84. Cheng, L., Liu, J., and Don, S. (2000) Layer-by-layer assembly of multilayer films consisting of silicotungstate and a cationic redox polymer on 4-aminobenzoic acid modified glassy carbon electrode and their electrocatalytic effects, Anal. Chim. Acta, 417, 133-42.

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ADAM HELLER 85. Li, W., Wang, Z., Sun, C, Xian, M., and Zhao, M. (2000) Fabrication of multilayer films containing horseradish peroxidase and polycation-bearing Os complex by means of electrostatic layer-by-layer adsorption and its application as a hydrogen peroxide sensors, Anal. Chim. Ada, 418, 225-32. 86. Katakis, I., Ye, L, and Heller, A. (1994) Electrostatic control of the electron transfer enabling binding of recombinant glucose oxidase and redox polyelectrolytes, J. Am. Chem. Soc., 116, 3617-18. 87. Katakis, I., Ye, L., Kenausis, G., and Heller, A. (1994) Design of redox polyelectrolyte "wires" for glucose electrodes, Polym. Mater. Sci. Eng., 71, 592-3. 88. Katakis, L, Vreeke, M., Ye, L., Aoki, A., and Heller, A. (1996) Electron conducting adducts of water soluble redox polyelectrolytes and enzymes. In Advances in molecular & cell biology, Volume 15B, (ed. E. Edward Bittar, B. Danielsson, and L. Billow), pp. 391-409, JAI Press Inc. 89. Gary Binyamin, Jason Cole, and Adam Heller (2000) Mechanical and electrochemical characteristics of composites of wired glucose oxidase and hydrophilic graphite. J. Electrochem. Soc., 147, 2780-3. 90. Kenausis, G., Taylor, C., and Heller, A. (1996) Wiring of glucose oxidase and lactate oxidase within a hydrogel made with poly(vinyl pyridine) complexed with [Os(4,4'dimethoxy-2,2'-bipyridine)2 Cl] +/2 + , J. Chem. Soc., Faraday Transactions, 92, 4131-6. 91. Taylor, C., Kenausis, G., Katakis, L, and Heller, A. (1995) "Wiring" of glucose oxidase within a hydrogel made with polyvinyl imidazole complexed with [(Os-4,4'-dimethoxy 2,2'-bipyridine)Cl]+/2 + , J. Electroanal. Chem., 396, 511-15. 92. Ohara, T. J., Rajagopalan, R., and Heller, A. (1993) Glucose electrodes based on crosslinked [Os(bpy)2Cl]+ '2+ complexed poly (1-Vinyl-imidazole) films, Anal. Chem., 65, 3512-17. 93. Kenausis, G., Taylor, C., Rajagopalan, R., and Heller, A. (1996) "Wiring" of lactate oxidase within a low redox potential electron conducting hydrogel, J. Mol. Recog., 9, 626-31. 94. Lever, A. B. P. (1990) Electrochemical parametrization of metal complex redox potentials, using the ruthenium(III)/ruthenium(II) couple to generate a ligand electrochemical series, Inorg. Chem., 29, 1271-85. 95. Schuhmann, W., Kranz, C., Huber, J., and Wohlschlaeger, H. (1993) Conducting polymer-based amperometric enzyme electrodes. Towards the development of miniaturized reagentless biosensors, Synth. Met, 61, 31-5. 96. Gajovic, N., Habermuller, K., Warsinke, A., Schuhmann, W., and Scheller, R W. (1999) A pyruvate oxidase electrode based on an electrochemically deposited redox polymer, Electroanalysis, 11, 1377-83. 97. Heller, J. and Heller, A. (1998) Loss of activity or gain in stability of oxidases upon their immobilization in hydrated silica: Significance of the electrostatic interactions of surface arginine residues at the entrances of the reaction channels, J. Am. Chem. Soc., 120, 4586-90. 98. Chen, Q., Kenausis, G. L, and Heller, A. (1998) Stability of oxidases immobilized in silica gels, J. Am. Chem. Soc., 120, 4582-85. 99. Park, T.-M., Iwuoha, E. L, Smyth, M. R., and MacCraith, B. D. (1996) Sol-gel-based amperometric glucose biosensor incorporating an osmium redox polymer as mediator, Anal. Commun., 33, 271-3. 100. Park, T.-M., Iwuoha, E. L, and Smyth, M. R. (1997) Development of a sol-gel enzyme inhibition-based amperometric biosensor for cyanide, Electroanalysis, 9, 1120-3. 101. Park, T.-M., Iwuoha, E. L, Smyth, M. R., Freaney, R., and McShane, A. J. (1997) Sol-gel based amperometric biosensor incorporating an osmium redox polymer as mediator for detection of L-lactate, Talanta, 44, 973-8.

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REDOX HYDROGEL-BASED ELECTROCHEMICAL BIOSENSORS 102. Kane, S. A., Iwuoha, E., and Smyth, M. R. (1998) Development of a sol-gel based amperometric biosensor for the determination of phenolics, Analyst (Cambridge, UK), 123, 2001-6. 103. Park, T.-M. (1999) Amperometric determination of hydrogen peroxide by utilizing a sol-gel-derived biosensor incorporating an osmium redox polymer as mediator, Anal. Lett., 32, 287-98. 104. Pravda, M., Adeyoju, O., Iwuoha, E. I., Vos, J. G., Smyth, M. R., and Vytras, K. (1995) Amperometric glucose biosensors based on an osmium (2+/S + ) redox polymermediated electron transfer at carbon paste electrodes, Electroanalysis, 7, 619-25. 105. Jezkova, J., Iwuoha, E. I., Smyth, M. R., and Vytras, K. (1997) Stabilization of an osmium bis-bipyridyl polymer-modified carbon paste amperometric glucose biosensor using polyethyleneimine, Ekctroanalysis, 9, 978-84. 106. Parellada, J., Narvaez, A., Dominguez, E., and Katakis, I. (1997) A new type of hydrophilic carbon paste electrodes for biosensor manufacturing binder paste electrodes, Biosens. Bioelectron., 12, 267-75. 107. Iwuoha, E., Smyth, M. R., andVos.J. G. (1994) Amperometric glucose sensor containing nondiffusional osmium redox centers: analysis of organic-phase response, Electroanalysis, 6, 982-9. 108. Iwuoha, E. I. and Smyth, M. R. (1996) Organic phase enzyme electrodes: kinetics and analytical applications, Biosens. Bioelectron., 12, 53-75. 109. Dressman, S. F., Garguilo, M. G., Sullenberger, E. V., and Michael, A. C. (1993) Characterization of the electroenzymic reduction of hydrogen peroxide in carbon dioxide by voltammetry with a chemically modified microelectrode, J. Am. Chem. Soc., 115, 7541-2. 110. Heller, A., Maidan, R., and Wang, D. L. (1993) Amperometric biosensors based on 3-dimensional hydrogel-forming epoxy networks, Sensors and Actuators, 13, 180-3. 111. Ohara, T. J., Vreeke, M. S., Battaglini, F., and Heller, A. (1993) Bienzyme sensors based on "electrically wired" peroxidase, Electroanalysis, W. Simon Memorial Issue, 5, 825-31. 112. Vreeke, M., Rocca, P., and Heller, A. (1995) Direct electrical detection of dissolved biotinylated horseradish peroxidase, biotin, and avidin, Anal. Chem., 67, 303-6. 113. Lu, B., Iwuoha, E. I., Smyth, M. R., and O'Kennedy, R. (1997) Development of an "electrically wired" amperometric immunosensor for the determination of biotin based on a non-diffusional redox osmium polymer film containing an antibody to the enzyme label horseradish peroxidase, Anal Chim. Acta, 345, 59-66. 114. Lu, B., Iwuoha, E. I., Smyth, M. R., and O'Kennedy, R. (1997) Development of an amperometric immunosensor for horseradish peroxidase (HRP) involving a nondiffusional osmium redox polymer co-immobilized with anti-HRP antibody, Anal. Commun., 34, 21-4. 115. de Lumley-Woodyear, T., Campbell, C. N., and Heller, A. (1996) Direct enzymeamplified electrical recognition of a 30-base model oligonucleotide, J. Am. Chem. Soc., 118, 5504-5. 116. de Lumley-Woodyear, T., Campbell, C. N., Friedman, E., Friedman A., Georgiou, G., and Heller, A. (1999) Rapid amperometric verification of PCR amplification of DNA, Anal. Chem., 71, 535-8. 117. Campbell, C. N., de Lumley-Woodyear, T., and Heller, A. (1999) Towards Immunoassay in whole blood: separationless sandwich-type electrochemical immunoassay based on in-situ generation of the substrate of the labeling enzyme, Fresenius]. Anal Chem., 364, 165-9. 118. Danilowicz, C. and Manrique, J. M. (1999) A new self-assembled modified electrode for competitive immunoassay, Ekctrochem. Commun., 1, 22-5.

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ADAM HELLER 119. Caruana, D. J. and Heller, A. (1999) Enzyme-amplified amperometric detection of hybridization and of a single base pair mutation in an 18-base oligonucleotide on a 7|xm diameter microelectrode, J. Am. Chem. Soc., 121, 769-74. 120. de-Lumley-Woodyear, T., Caruana, D. J., Campbell, C. N., and Heller, A. (1999) Reactive electrophoretic activation of a microelectrode for enzyme-amplified recognition and for melting-temperature determination of a simple oligonucleotide, Anal. Chem., 71, 394-8.

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Chapter 2 Hybridization at oligonucleotide sensitive electrodes Daren J. Caruana Department of Chemistry, University College, London, 20 Gordon St, London WC1H OAJ, UK.

1 Introduction DNA hybridization on solid surfaces was developed on membranes as a routine biotechnological technique in the 1970s (1). With the invention and refinement of new surface analytical techniques, oligonucleotide detection by surface confined hybridization is the fundamental principle behind the function of most oligonucleotide sensitive devices. These devices are capable of high throughput DNA analysis for applications such as sequencing, genetic diagnosis, drug phenotyping, computing, and gene expression analysis. Regardless of application, a common aspect of biosensor devices is the conversion of a physiochemical process confined at a surface into an interpretable signal. In the case of oligonucleotide sensitive electrodes, the transducer element of the device differentiates between single and double stranded oligonucleotides at a solid interface. The challenge is to define the optimum conditions for maximum hybridization efficiency, to obtain a high degree of sensitivity. The aim here is to discuss the important aspects of oligonucleotide attachment to conducting surfaces and the process of hybridization to the complementary strand for the fabrication of oligonucleotide sensitive electrochemical sensors. Conceptually, hybridization at a surface may appear to be the same as the equivalent process in bulk solution; however, a unique set of conditions needs to be defined. The emphasis of this chapter will be to provide practical information to aid in the determination of optimum hybridization conditions. Different applications or electrode models will require different set of conditions for optimum hybridization and detection sensitivity. Therefore, it is unconstructive to provide a set of experimental conditions, which will not always apply for every situation.

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DAREN J. CARUANA These variables will be divided into two groups: (a) Characterization of the surface confined probe; defining the environment of the probe, attachment method and surface structure. (b) Hybridization conditions defining the conditions under which association of a solution-phase target oligonucleotide with a surface tethered probe oligonucleotide. Many of these experimental variables are interrelated and all must be carefully considered to determine optimum conditions for hybridization. Discussion will be confined to oligonucleotides of 14-50 nucleotides in length.

2 Function of oligonucleotide sensitive electrodes Random collision between two complementary strands is the fundamental requirement for hybridization to take place on a solid surface. The probability or frequency for collision will dictate the kinetics of this process. Once collision has occurred, the annealing must then take place to produce a hybrid. If the resulting hybrid is completely complementary, the hybrid will be stable. Hybridization will still take place if the sequences are not completely complementary, but the hybrid will be less stable than the completely complementary hybrid. Differentiation between interactions resulting in complementary hybrids and other interactions leading to less stable non-complementary hybrids may be carried out by carefully defining the experimental conditions. This is the key to the function of oligonucleotide sensitive electrodes that rely on hybridization, and is the subject of this chapter. Although the discussion here will be confined to DNA, which is readily available by custom synthesis, there are a number of other nucleic acid analogs that may also be used. RNA may be used but is experimentally difficult to manipulate due to its susceptibility to degradation from RNAses that are more abundant than DNAses. Uncharged analogs such as peptide nucleic acids (PNA), and other non-phosphorous containing analogs are potentially suitable for use as probes in oligonucleotide sensitive electrodes (2). The structure of the nucleotide duplex is an intertwined structure of two antiparallel molecules. The three-dimensional shape is maintained by an array of individually weak interactions which are strong collectively, and which stabilize the molecules in a rigid familiar double helical structure. The interactions are mainly hydrogen bonds, hydrophobic, electrostatic, van der Waals, and dipole interactions. Hybridization involves the formation of these interactions to produce a close packed helix. The formation of these forces holding the two strands together, most notably the hydrogen bonding between the individual bases and ionic interactions (Figure 1) can be translated into a range of physiochemical experimental parameters. These parameters in some way contribute to the formation or disruption of the individual weak interactions and determine which hybrids remain on the surface and which are removed. 20

OLIGONUCLEOTIDE SENSITIVE ELECTRODES

Figure 1 Showing the Watson-Crick hydrogen bonding between (a) A-T, (b) G-C base pairs, and (c) the phosphate deoxyribose backbone (3).

3 Hybridization efficiency and sensitivity In bulk solution, the process of hybridization of two complementary oligonucleotide strands is a specific interaction between two molecules and occurs homogeneously in solution. In the case of surface hybridization, the process may be looked upon as a specific adsorption process. Essentially the same interactions are taking place but may be characterized differently. The function of these oligonucleotide devices may be characterized by the hybridization efficiency and hybridization rate, which are not normally issues when describing hybridization process in the bulk solution. Hybridization efficiency is a measure of the proportion of probe oligonucleotides that successfully hybridize to the complementary strand under optimum hybridization conditions at the surface. The hybridization rate describes the flux of target strand to the surface to facilitate hybridization. The equilibrium constant for the hybridization process for two complementary single stranded oligonucleotides: Probe + Target -> Hybrid is given by:

where K is the equilibrium constant. Experimental hybridization data may also be expressed as a ratio, v, of hybridized to single stranded probe oligonucleotide:

21

DAREN J. CARUANA The quantitative determination of the proportion of hybridized oligonucleotides on a surface is not trivial, and requires the use of sensitive quantitative analytical techniques such as radiolabeling or fluorescence. If indirect methods of detection are used, then only relative amounts of hybridization may be obtained. Ultimately the extent of hybridization may be determined by the stringency of the hybridization conditions and surface concentration of the tethered oligonucleotide.

4 Probe oligonucleotide structure and dynamics Although the structure of double stranded DNA is well studied, single stranded DNA is much less defined especially at a solid surface. It is tempting to picture a solid surface decorated with probe single stranded oligonucleotides standing up straight pointing toward the solution ready to hybridize with the complementary target single stranded oligonucleotide to form the duplex. In reality the probe structure and surface density are difficult to characterize due to the minute amount present on the surface. The probe architecture will substantially affect the hybridization efficiency and, therefore, the sensitivity of the oligonucleotide sensitive electrode. The probe structure at the surface will be dependent on a number of parameters including the surface charge, electrical potential, hydrophobicity, probe attachment method, and also solution physicochemical or chemical conditions. These variables, specifically for probe structure, will be discussed in relation to how these properties affect hybridization efficiency. The discussion will enable the experimentalist to identify a unique set of conditions for a unique surface and application. The probe concentration, length, spacer group, and surface orientation will be discussed in some detail.

4.1 Surface concentration Steric hindrance to hybridization as a consequence of high surface packing of oligonucleotides on the surface has a direct impact on the hybridization efficiency (4). There are methods of oligonucleotide surface modifications that lead to a highly dense layer of single stranded oligonucleotide which may lead to hybridization efficiencies of less than 5%. Therefore, strict control of surface concentration is enormously important to conserve a reasonable hybridization efficiency and detection limit. There is a relationship between probe length in base pairs, and surface distribution based only on nearest neighbor interactions contributing to steric interactions for monolayer coverage. As a rough "rule of thumb" a relationship may be formulated between the number of molecules that may be attached to the surface, assuming the probe oligonucleotide exists as a stretched linear molecule and the oligonucleotide is attached at only one point. The equations given in Table 1 provide an estimate of the theoretical optimum surface concentration that will minimize steric interactions between probe oligonucleotides. Another consideration is the difference in length of the target 22

OLIGONUCLEOTIDE SENSITIVE ELECTRODES Table 1 Relating length of probe oligonucleotide in base pairs with maximum close packed (^/3) surface coverage to minimize nearest neighbor interactions. Roughness is r= (actual surface area)/ (geometric area) Attachment point

Equation

Variables

Terminus

b is length in bases and r is roughness factor

Terminus via spacer

b is length in bases, r is roughness factor, and s is spacer length in A

Midpoint

b is total length in bases and r is roughness factor

Midpoint via spacer

b is length in bases, r is roughness factor, and s is spacer length in A

strands that can influence near neighbor hybridization. When the target strand is smaller than the probe, the hybridization rate increases roughly proportionately to the difference in length. When the target strand is longer, the hybridization rate is reduced. Accurate quantitative measurement of surface concentration of single stranded oligonucleotides may be carried out by surface sensitive techniques in combination with radioisotope (32P) or fluorescence labeling methods (5).

4.2 Probe length and orientation The availability of the probe strand to the solution has a direct affect on both the rate and efficiency of hybridization. The lengths of the probe and the spacer group determine the secondary structure that the molecule can exhibit, and the orientation of the probe molecule is largely dependent on the surface properties of the support, as shown in Figure 2. Attachment of the oligonucleotide through a spacer group reduces nearest neighbor steric effects. The length and properties (charge, hydrophobicity, flexibility) of the spacer are important and influence the structure of the probe layer. The longest aliphatic spacers available commercially are already attached to oligonucleotides at 12C in length. Longer aliphatic chains become less soluble and may behave as surfactants. Glycols and amino-based spacers are more water soluble and are available with different lengths and terminal reactive groups. As yet there has been no comprehensive study on the surface properties and spacer properties on the hybridization efficiency; however, it is clear that attachment via a spacer improves the hybridization efficiency, although an optimum length has not been determined. The simplest spacer is to use a poly(dT) which extends the oligonucleotide and places the important sequence further from the surface. However, this does not overcome problems of secondary structure of probe molecules. 23

DAREN J. CARUANA

Figure 2 Schematic of single stranded oligonucleotides attached to the surface (a) stretched out to the solution, (b) with the charged phosphate backbone interacting with the surface, (c) with the bases interaction with the surface, and (d) via spacer groups.

Figure 3 Schematic representation of a probe oligonucleotide with a hairpin structure.

Single stranded oligonucleotide molecules probably behave as a polyelectrolyte with either no fixed or random secondary structure. For single stranded oligonucleotides with lengths less than 8-10 bases, the molecules will exhibit rather little secondary structure. An oligonucleotide longer than 11 nucleotides may fold back on itself and form a hairpin, if the sequence supports it, as shown in Figure 3. The occurrence of such hairpin structures dramatically reduces the availability of probe to the solution and reduces hybridization efficiency. Hairpin

24

OLIGONUCLEOTIDE SENSITIVE ELECTRODES Table 2 Surface sensitive techniques suitable for the determination of structure of probe oligonucleotides Description/requirement

Neutron reflection

Requires Synchrotron Radiation Provides high resolution surface density and thickness in the axis and involved data analysis perpendicular to the surface plane

Scanning probe microscopy

Atomic Force or scanning tunneling microscope

X-ray photoelectron Scanning electron microscope with XPS capability spectroscopy Fluorescence

High sensitivity photon counting mode equipment

References

Comments

Technique

9

X, Y, and Z spatial surface structure

6, 10, 11

Thickness of surface layers and elemental density

4, 12

Inter- and intramolecular interactions. An indirect method requiring chemical labeling

13

structures are easily destabilized by heating (the melting temperature may be calculated using the formula in Section 5.1) or with the addition of denaturing agents such as formaldehyde or glyoxal (ethanedial). Hairpin structures are frequently associated with RNA molecules. The physicochemical nature of the surface (i.e. hydrophobicity, charge, electrical potential) will affect the structure of the probe in various ways. Electrostatic forces between the surface and the negatively charged backbone of oligonucleotides do influence the structure of the molecule. Intuitively, negatively charged surfaces repel the oligonucleotides away from the surface, whilst positively charged surfaces cause the molecule to lie down with the phosphate groups in contact with the surface. Interestingly, studies using positively charged surfaces decorated with oligonucleotides of 24 bases appear to form non-helical structures not found free in solution (6). The influence of an electrochemical potential applied to the surface will have a similar effect to chemical surface charges. The effect of electrochemical potential on a surface decorated with a dense covering of short double stranded oligonucleotide has been observed (7), however, the effect on single stranded oligonucleotides has not been investigated in detail. Hydrophobic surfaces tend to interact with the bases making the oligonucleotide unavailable for hybridization, as shown in Figure 2 (8). The lack of information on the fine structure of submonolayer single stranded oligonucleotide is mainly due to technical difficulty with observing small changes in structure in situ. Such studies require refinement of sophisticated surface sensitive techniques and heavy investment of personnel and money. A list of techniques that have been applied to these studies with the information that they provide is given in Table 2.

4.3 Attachment of probe The immobilization of probe onto a solid support has been used as an analytical tool in molecular biology after it was first described by Gillespie and 25

DAREN J. CARUANA Speigelman (14). Nitrocellulose, nylon, or polystyrene supports were used to bind fragments of DNA and RNA with lengths greater than 500 nucleotides. The attachment to these membranes was purely physisorption and behaved well at low temperatures. There are currently several commercially available membranes designed to covalently bind nucleic acids either by UV radiation or by chemical treatment of activated membranes. For electrochemical-based detection of hybridization, the methods for attachment have been adapted for attachment onto solid conducting surfaces. Regardless of how DNA or hybridization is detected, the method chosen for the attachment of single stranded oligonucleotides onto the appropriate surface depends on the nature of the surface. Several approaches have been developed for surface attachment and may be broadly divided into two: The first involves the base-by-base synthesis of oligonucleotides on the surface using highly efficient chemical reactions (15,16). The second involves direct attachment of ready synthesized single stranded sequences onto the surface. Both approaches have been extensively developed; however, the former involves a larger degree of investment in specialist equipment and is an exceedingly attractive method for DNA array fabrication due to the light addressable nature of the synthesis. Certainly, the attachment of the ready synthesized oligonucleotides on the surface is the most convenient, and will be discussed on a practical level here. The choice of method is frequently dependent on the nature of surface, whether a hard or soft (polymeric coated solid) conducting surface is used. For surface modification, the concentration of probe in solution and the purity of the DNA may be measured as described in Protocol 1. There are many sources of custom sequence oligonucleotides for different lengths, linker groups and functional groups at 5', 3' and internal positions. The product purity may be specified; either ethanol or phenol precipitated which is minimum purity or HPLC, PAGE and ion exchange purified for higher purity. For oligonucleotide sensitive electrode preparation precipitated purity is probably sufficient.

Protocol 1

Determination of solution concentration of single stranded oligonucleotide and assessment of purity of oligonucleotide Equipment and materials UV spectrophotometer Quartz cuvette of path length 1 cm Buffer containing: 0.1M Trizma® base buffer (Sigma Aldrich chemical company), 10 mM EDTA (ethylenediaminetetraacetic acid), 2 mM NaCl, adjusted to pH 7.4 26

All pipette tips and glassware should be sterilized or thoroughly cleaned to remove nuclease contamination.

OLIGONUCLEOTIDE SENSITIVE ELECTRODES Protocol 1 continued

Method 1 Add 1 ml of buffer to a cuvette and zero (blank) the spectrometer. 2 Place the sample in a similar cuvette and take absorbance reading at 260 and 280 nm, and 230 and 325 nm. 3 The concentration is given by the following equation:

where N is the number of bases indicated by the subscript.

4 If the ratios A26o/A28o and A230lA325 are between 1.8 and 1.9, and 1.9 and 2.0, respectively, the sample is highly purified. Lower ratios may contain protein or phenol (phenol is used to precipitate the DNA) contamination. This procedure works for both DNA and RNA either single or double stranded.

4.3.1 Attachment to gold by chemisorption Attachment via a thiol onto gold is a very popular method due to the availability of thiol terminated single stranded oligonucleotides (5). There are no complex chemical steps or chemistry involved and is relatively easy to control in terms of oligonucleotide surface concentration. Protocol 2 describes the method used for the deposition of oligonucleotides onto clean gold surfaces. The surface concentration of oligonucleotides may be controlled by using a mixture of different thiol bearing molecules in the deposition mixture. The gold pre-treatment is extremely important. It is recommended that thorough cleaning of the surface should be carried out by repetitive voltammetric cycling in a weak acid (H2SO4 0.05M) solution. This is a controlled electrochemical technique, which ensures that the surface is prepared reproducibly. Alternative methods of producing a clean surface would be to sputter gold on the surface of a flat substrate (if using glass as the substrate, the surface needs to be coated with Cr to aid adhesion). The surface of sputtered gold using a lowpressure gold sputterer in an argon atmosphere is reported to be predominantly Au [111] crystal face exposed to the liquid (although this does not mean that the surface is atomically flat). The gold electrodes produced in this way are convenient because the surface is relatively clean as long as care is taken to reduce exposure to polluting environments.

4.3.2 Non-covalent or physisorption This method was developed for the hybridization of DNA on nitrocellulose membranes for techniques such as Southern blots. The adsorption of oligonucleotides normally longer than 500 bases works well for high area surfaces where surface concentration and hybridization efficiency are not of concern. This method is difficult to control and relies on the adsorption of the unmodified 27

DAREN J. CARUANA oligonucleotide on a clean surface. There are methods developed for electrochemical hybridization detection at Indium doped tin oxide surfaces with oligonucleotides in the order of 400-1500 bases, which are strongly adsorbed (6,17).

Protocol 2

Attachment of oligonucleotides Materials Oligonucleotide modified with a thiol terminated 3' or 5' purchased from Genosys or other custom synthesis source. Sterile pH 7.2 HEPES (0.01M) buffer

Cysteamine as the diluent thiol All pipette tips and glassware should be sterilized or thoroughly cleaned to remove nuclease contamination.

Method 1 Make the appropriate mixture of thiol modified oligonucleotide and diluent thiol cystamine in HEPES buffer. 2 Place the clean electrode surface into an aqueous solution containing the thiol modified oligonucleotide and leave at room temperature for 24 h. 3 Remove and wash the electrode thoroughly with deionized water. 4 The electrode should be modified immediately prior to use.

4.3.3 Chemical attachment A variety of chemical attachment methods are available. All effectively produce covalently attached oligonucleotide on a range of conducting surfaces. The first step is to introduce reactive chemical groups on the chosen surface. This is not always straightforward and frequently determines the electrode material. There are several methods, which are specific for different surfaces or pre-treatment of surfaces (18). The surface of conducting silicon after silanization or other chemical treatments to introduce reactive anchor groups are very numerous. Characterization of the efficiency of each chemical step and the number density of sites introduced by each method is difficult and requires sensitive surface analysis. Electrochemical methods of surface modification of edge plane graphite with oxygen containing reactive groups may be used for oligonucleotide attachment through CNBr coupling (see Figure 4(a) and Protocol 3).

4.3.4 Deposition onto a polymer coated electrode Using a polymeric support on a conducting surface is a convenient way of increasing the hybridization signal by deposition of more probe oligonucleotides due to the higher active area. Optimization of this method is difficult, as hybridization within the polymer matrix will depend on the microenvironment of the polymer such as density, charge, hydrophobicity, thickness, etc. 28

OLIGONUCLEOTIDE SENSITIVE ELECTRODES

Figure 4 Reaction schemes for the (a) CNBr activated coupling and the (b) reactive electrophoretic deposition on electrodes baring reactive amino groups, respectively.

Protocol 3

CNBr activated coupling to an edge plan graphite electrode Materials Cyanogen bromide, sodium hydroxide Amino terminated single stranded oligonucleotide

Sterile 0.1M sodium bicarbonate buffer pH8.5 Ice and magnetic stirrer.

Method 1 To 25 ml of deionized water in a fume hood add 5 g of CNBr and adjust the pH to 11.0 with 5% NaOH and place the oxidized edge plane graphite electrode in the solution and stir for 20 min. 2 Remove the electrode and place in 20 ml ice-cold 0.1M sodium bicarbonate buffer pH 8.5 and add amino terminated single stranded oligonucleotide (ca. 300-500 |j,g) and leave stirring for 48 h at 4 °C. 3 Wash electrode in potassium phosphate buffer and use immediately. The DNA solution can be used only once. For control of surface concentration, use an amine (such as CH3CH2NH2) added to the DNA solution to dilute the surface concentration.

Attachment of oligonucleotides to the polymer support may be done in a variety of ways, before, during, or after deposition of the polymer matrix. Electropolymerization of pyrrole monomer units modified with oligonucleotide at an electrode surface is an example of codeposition (19). Protocol 4 outlines an 29

DAREN J. CARUANA electrophoretic method of deposition of oligonucleotide with a 5' chemically reactive group, which forms a covalent bond to an amine on the polymer backbone (20, 21). These techniques are very attractive where spatial segregation of the oligonucleotides is important. The drawback is that characterization is not easy and the determination of the optimum hybridization efficiency is difficult.

Protocol 4

Reactive electrophoretic deposition Equipment and reagents Bench top centrifuge CH instruments potentiostat with chronoamperometric capabilities Electrode of diameter between 10 and 25 |im pre-coated with polymeric matrix with reactive amino groups

l-[3-(dimethylamino)propyl]-3-ethylcarbodiimide hydrochloride (EDC) (Aldrich. Cat. No. 16,146-2) Single stranded oligonucleotide (15-35 bp) from Genosys Microamplicon tube (Amicon) with a 3000 Da cutoff membrane.

Method 1 450-550 |j,g of probe oligonucleotide is dissolved in 50 |j,l of pH 7.0 methylimidazole (0.02 M) buffer. 2 To the oligonucleotide solution was added 50 |j,l of EDC (0.2M) and was stored at 4 °C for 16 h. 3 450 |jl of deionized water was added to the mixture and placed into a microamplicon tube to reduce the volume to ca. 50 |jl by spinning at 50,000 rpm (approx. 15 min). This procedure was repeated twice to remove excess buffer and EDC. The final concentration of EDC and methylimidazole will be ca. 2 x 10~ 4 M and 2 x 10 ~ 5 M, respectively. 4 The 50 |jl of oligonucleotide was then transferred to an electrophoretic cell with 100 |jl of deionized water. 5 The microelectrode was placed in the cell containing the oligonucleotide approx. 1 mm from the counter electrode. A constant potential of 0.9 V for 5 min was applied to the working electrode.

Hybridization conditions In practical terms, the hybridization is dependent on the thermodynamic stability of the duplex formed and the collision frequency; both are interrelated and difficult to define independently. However, both are controlled by the physicochemical conditions under which hybridization takes place (22). The kinetics of this reaction in solution is well studied and, for the simple case it is described 30

OLIGONUCLEOTIDE SENSITIVE ELECTRODES

by a second order rate equation. Hybridization kinetics at a solid surface is experimentally more difficult to monitor in real time, and is comparatively less well understood compared to the equivalent process in solution (23). Nevertheless, the physical and chemical parameters that dictate the stability of the surviving hybrid need to be considered for optimizing hybridization efficiency. There is no single protocol that is appropriate for all applications of oligonucleotide sensitive electrodes. There are many variables to be considered when designing the hybridization conditions that will either affect the stability of the hybrids, the rate at which the hybrids form, or both. The discussion here will focus on the effect of temperature, number of base pair mismatches, ionic strength, formamide concentration, nonspecific adsorption, and mass transport on the hybridization stability and hybridization efficiency.

5.1 Temperature By far the most important variable is temperature. Hybridization temperature is always discussed relative to the melting temperature, Tm, for a given oligonucleotide at which an equilibrium is reached where 50% of single stranded oligonucleotides are hybridized. The melting temperature, Tm, of an oligonucleotide is dependent on the length in base pairs base composition and salt concentration, which are related according to the following relationship (24):

where (%G + C) is the content of G and C in the oligonucleotide and I is the length in base pairs. There are many different versions of melting temperature equations and different equations apply at slightly different conditions. There is no equation which will estimate the melting temperature for all lengths of oligonucleotides. The equation given above holds for 14 to ca. 72 nucleotides for sequences in solution at pH between 5 and 9. As the length in base pairs increases, the melting temperature becomes more independent of the length. Theoretical equations also give Tm values based on nearest neighbor and thermodynamic changes in entropy and enthalpy when hybridization occurs (25, 26). There have been few studies to assess the effect of surface confinement of the hybridization process. Krull et al. (13) have shown that when a single base pair mismatch in the middle of a model 20 nucleotide changed the melting temperature 6-10 °C rather than 3.8-6.1 °C change in solution. The melting temperature for oligonucleotides in solution may be determined experimentally by measuring the absorbance at 260 nm using a spectrophotometer whilst increasing the temperature linearly, as outlined in Protocol 5.

31

DAREN J. CARUANA

Protocol 5

Determination of melting temperature Equipment and reagents UV spectrophotometer with a programmable temperature controlled micro-cuvette holder Quartz cuvette of path length 1 cm, small volume (200 pi) Buffer containing: 0.1M Trizma® base buffer (Sigma-Aldrich chemical company), 1 mM EDTA (Ethylenediaminetetraacetic acid), 2 mM NaCl, adjusted to pH 7.4

Two complementary oligonucleotide samples All pipette tips and glassware should be sterilized or thoroughly cleaned to remove nuclease contamination.

Method 1 Zero the spectrophotometer with buffer at 260 nm. 2 Place 200 |j,l single stranded oligonucleotide dissolved in buffer in the cuvette to obtain an absorbance between 0.05 and 0.1. 3 Mix equimolar amounts of the two complementary single stranded oligonucleotides in buffer and place the solution in the cuvette and thermostat at ca. 30 °C below the calculated melting temperature for 30 min. 4 Increase the temperature of the cuvette at 1 °C min"1 and monitor the absorbance at 260 nm. 5 At the point of melting, the absorbance will increase over a range of 15 °C. The Tm is the temperature when the absorbance is halfway between starting and the finishing values.

Ionic strength Ionic strength affects the stability of hybrids, rate of hybridization, and interactions between probe molecules attached to the surface. High ionic strength improves the stability of hybrids and increases the rate of hybridization when hybridization is not limited by mass transport. Sodium chloride is commonly used in hybridisation buffers, but the effects are more pronounced with divalent cations such as Mg2 + . Quantitative effects of salt concentration on the rate of hybridization and hybrid stability have not been determined in detail for oligonucleotides at surfaces, but probably reflect the trend observed in solution. 1M sodium chloride is frequently used, as higher ionic strength will tend to stabilize mismatched hybrids and slow down the hybridization rate (for filter hybridization above 0.1M NaCl, the hybridization rate is not markedly affected) (27). The high salt concentration is important to reduce the electrostatic interactions between single and double stranded molecules attached to the surface. When using peptide nucleic acids, the ionic strength is less important. 32

OLIGONUCLEOTIDE SENSITIVE ELECTRODES

In the presence of quaternary ammonium salts such as tetraethylammonium or tetramethyl ammonium salts, the hybridization is independent of base composition, only length is important (28). This is very useful when an array of probes with different sequences (%G & C) is spatially localized on the surface. In the presence of TEA+ or TMA + , the Tm is dependent only on the number of base pair mismatches and length of probe (28). Typical concentration ranges of TMAC1 used in hybridization buffers are between 2M and 3M which can lower the melting temperature by 10-15 °C for a 20 base pair oligonucleotide.

5.3 Base mismatch The reassociation requires that all the bases meet up and form hydrogen bonds when the sequences are complementary. Hybridization will also occur when base sequences are not completely complementary leading to base pair mismatch. The degree of association strongly depends on the number and nature of the mismatch. The stability is dependent on the following factors: (a) A mismatch occurring on the center section of the molecule is less stable than molecules with mismatches occurring at the periphery. (b) The number of mismatches as a percentage of the length. If multiple mismatches are dispersed, the stability of the hybrid will be less than if they were clustered. (c) The identity of the mismatched nucleotide can yield different stability due to greater steric disruption to the duplex. The order of stability for a single position from most stable is T-A, A-T> G-T, G-A, > A-A, T-T, C-T, C-A (28). Very frequently differentiating between single base mismatch and complementary duplexes is required for many oligonucleotide sensitive electrodes. As a rule, the shorter the probe, the easier it is to differentiate between them by melting temperature differences. In an analytical sense, mismatched hybrids may be considered as nonspecific interactions, and their occurrence needs to be reduced. As discussed above, there are several degrees of mismatching and there are no quantitative rules to characterize their stability in terms of melting temperature. As a general guide, with every 1% mismatch bases in a DNA duplex, the Tm is reduced by 1±0.5 °C. For an estimate of the hybridization temperature for the differentiation of complementary and mismatched sequences, the following equation maybe used:

where M is the minimum number of mismatches and L is the total number of base pairs. A target strand of 20 base pairs with one mismatch will have a melting temperature ca. 5 °C less than the Tm of the fully complementary duplex.

33

DAREN J. CARUANA

5.4 Mass transport The probability for a target strand to collide with the complementary strand tethered to a surface is increased by physically increasing the flux of the solution containing the target to the surface. The rate of mass transport may be improved by engineering of the hybridization cell, shaking, stirring, or affecting migration of the target. Reducing the volume and/or increasing the probe-modified surface to volume ratio will improve the diffusional mass transport to the surface. The hybridization process involves not only planar diffusion toward the surface, but two-dimensional diffusion across the surface of the probe modified interface (8). The architecture of the oligonucleotide decorated surface is important. If the surface is porous or covered by a polymeric matrix, then diffusion will be heavily impeded thus reducing the collision frequency. Optimum surface area without impeding the diffusion will require nanoscale surface design to increase the solution availability of probe. Migration in an applied electric field has been studied in the context of improving the mass transport to the surface. However, there have been reports of other effects associated with the electric field at the surface where hybridization takes place. The physical basis of these are not clear but may be a direct or indirect effect of the electric field on the hybridization.

5.5 Nonspecific adsorption From a detection point of view, lowering nonspecific adsorption with the surface is important to maintain a measurable signal-to-noise ratio. The surface of the oligonucleotide sensitive layer maybe viewed as having two adsorption sites: the probe, a specific binding site, and the area around the probe which is nonspecific. The aim would be to restrict the binding to the probe to form a duplex. The importance of nonspecific adsorption is dependent on the method of detection. When detection relies on labeled target (fluorescent or enzyme) brought to close proximity to the electrode, then nonspecific adsorption is important and will lower sensitivity. Other complications may arise regarding the label used for hybridization indication, such as the nonspecific adsorption of the label itself to sites on the surface. If the detection is specific to duplex formation, then nonspecific adsorption is less important. The extent of adsorption of oligonucleotide molecules is dependent on the nature of the surface surrounding the probe oligonucleotides, the pH, ionic strength, and temperature (29). There are strategies that have been developed for filter hybridization to reduce nonspecific adsorption, which are effective for high molecular weight immobilized probes. These strategies include buffers (such as Denhardt's buffer) containing high ionic strength buffer salts, surfactants, and large molecular weight molecules such as Ficoll (a synthetic non-ionic sucrose polymer) polyvinylpyrrolidone, BSA, and also nonhomogeneous heat denatured sheared DNA (Salmon sperm DNA or yeast tRNA). For low molecular weight probe molecules on the 34

OLIGONUCLEOTIDE SENSITIVE ELECTRODES

surface, such strategies are not so effective and would mask much of the surface and reduce hybridization efficiency. Strategies that are effective in reducing nonspecific adsorption: (a) Introducing a wash step after hybridization has taken place to remove nonspecific adsorption. This approach is restricted to post-hybridization detection, not real time. (b) Maintaining the surface, predominantly, negative charged by surface modification, or addition of surfactant such as SDS. (c) Using polymeric coated surfaces with low surface Gibbs free energies of adsorption such as poly(acrylamide). Figure 5 shows the response from an acrylamide-based redox polymer decorated with 18 nucleotide probe. Addition of enzyme (Soyabean Peroxidase, SBP) labeled noncomplementary target at 550 s gives a low signal indicating small amount of nonspecific adsorption. When enzyme labeled complementary stand is added to the hybridization solution at 1450 s, a large amperometric response results from the in situ hybridization (30).

Figure 5 Current-time plot of the catalytic current of a microelectrode coated with the probe-bearing redox polymer, (a) 10 jil of 40 nM SBP-labeled target with four mismatched bases were introduced at 550 s, followed by (b) 10 jil of 40 nM SBP-labeled perfectly matching target at 1450 s. Stirred 1 ml pH 7 HEPES buffer, 1M NaCI with 1.0 mM H202; -0.06 V (Ag/AgCI) thermostated at 45 °C stirred. (Reproduced with permission from ref. 21.) 35

DAREN J. CARUANA

5.6 Other factors Other factors that may affect the stability of the hybrids are pH and formamide concentration. Within the pH range 5-9, there is no effect in the melting temperature. However, pH effect on surface hybridization has not been studied extensively, it is likely to affect the ionizable groups on the electrode surface rather than a direct effect on the hybridization. This is of importance when using polymer-modified surfaces with an increased surface area. Formamide decreases the stability of oligonucleotide hybrids by lowering the Tm. This is a useful property when temperature is an issue for detection or if attachment of probe to the electrode surface is non-covalent. In solution 30% formamide reduces the melting temperature by ca. 20 °C. When formamide is present, the term — 0.72 (%formamide) is added to the equation on page 33.

6 Hybridization kinetics The kinetics of the hybridization of a single strand target in solution is dependent on the mass transport to the surface bearing the probe oligonucleotides. If the sequences are completely complementary, the hybridization may be described by a Langmuir-type isotherm for all sites being equal. The diffusion limited first-order Langmuir model:

where t is the time, r and T0 are the coverage and the total coverage at infinite time, and kLD is the rate constant. This equation appears to fit the hybridization transient well when mass transport is diffusion limited. There are two cases that may be limiting; either the rate of mass transport or the rate of surface confined hybridization. If the concentration of the target is low relative to the probe concentration, the limiting factor would be the mass transport; if however the target concentration is high, then hybridization rate would be limiting. The rate of hybridization is dependent on the hybridization conditions chosen and the application for the oligonucleotide sensitive electrode. Manipulation of hybridization rate is important when concentration of target is required. The dynamic range can be achieved when the hybridization rate is mass transport limited. This is a simplified picture and the rate is ultimately dictated by the stringency of the conditions chosen for hybridization: (a) Temperature. A maximum rate of hybridization occurs at 20-25 °C below the Tm in solution. As the temperature approaches Tm, the hybrids tend to dissociate leading to a low overall rate of hybridization. The temperature dependence on the rate of surface-confined hybridization has not been studied in detail but it is likely that the trends would be similar. (b) Concentration of target. As the single stranded target concentration increases, the hybridization rate will increase. However, high concentration will lead to nonspecific adsorption/binding which will depress the signal-to-noise ratio. 36

Table 3 Summary of effect of experimental variables on the hybridization and stringency of conditions for sequence discrimination Variable

Hybrid stability

Hybridization rate

Attached probe

High stringency

Low stringency

Temperature

10 °C Below 7m hybrid is stable

Maximum rate is ca. 25 °C below 7m Closer to the 7m the rate is slower

No effect

Temperature close to the 7m above the 7m of the single base pair mismatch

Temperature approx. 20 °C lower than 7m

Ionic strength

High ionic strength high stability

Rate increases x 4 up to 0.4M. No significant change up to 1M

High ionic strength leads to screening of anionic charges on adjacent probes

1M NaCI

Above 0.4M NaCI

Formamide

Reduce 7m by 0.72 (%Formamide)

No effect

Unknown

Lowering of 7m destabilizes complementary and mismatched hybrids equally

Not required

DAREN J. CARUANA (c) Target. If the target is double stranded in the test solution, melting before hybridization occurs at the surface of the electrode. This will reduce the rate of hybridization due to a competing hybridization process in the solution. (d) Formamide. From filter studies, formamide does not appear to affect the hybridization rate but shifts the optimum temperature by the same amount as the melting temperature. Formamide at 30-50% in the hybridization conditions reduces the stability of hybrid 30-42 °C depending on the length of the molecule. (e) Ionic strength. At high ionic strength (1M NaCl) hybridization rate is fast due to stabilization of duplex once hybridization has taken place. At low ionic strength, < 0.1M Na+ , a twofold increase in concentration increases the rate by 5-10-fold. (f) Dextran sulphate. 10% dextran sulfate gives a 10-fold increase in reassociation rate in solution. The effect is believed to be associated with volume exclusion raising the effective concentration. For short sequences hybridization accelerators are probably not effective.

7 Summary In summary the process of hybridization is a complex interaction dependent on an array of physicochemical and chemical variables, see Table 3. The topics covered in this chapter ensure that the variables are identified and optimized to maximize hybridization. The detection of the hybridization process at a surface using electrochemical-based methods has attracted a great deal of interest to date (31). Regardless of which technique is used for detection of hybridization, the design of the probe modified surface and the choice of conditions are important to consider. The surface design influences the detection sensitivity (hybridization efficiency) and hybridization conditions will dictate the function and sequence selectivity. The current emphasis in oligonucleotide hybridization research is to develop robust detection methodologies that support analysis of individual micron-sized pixels in an array of different sequences of oligonucleotides and to avoid labeling target strands with an indicator molecule.

References 1. Southern, E. M. (1975). J. Mo!. Bio!., 98, 503. 2. Eckstein, E. (1991). Oligonucleotides and analogues, practical approach series. Oxford University Press. 3. Stryer, L. (1995). Biochemistry, 4th edn, p. 789. W.H. Freeman & Co., New York. 4. Shchepinov, M. S., Case-Green, S. C., and Southern, E. M. (1997). Nucleic Acid Res., 25, 1155.

5. Herne, T. M. and Tarlov, M. J. (1997). J. Am. Chem. Soc., 119, 8916. 6. Lemeshko, S. V., Powdrill, Y., Belosludtsev, Y. Y., and Hogan, M. (2001). Nucleic Acids Res., 29, 3051.

7. Kelly, S. O. et al. (1998). Langmuir, 14, 6781. 38

OLIGONUCLEOTIDE SENSITIVE ELECTRODES 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31.

Chan, V. et al. (1998). J. Coll. Int. Sci., 203, 197. Levicky, R., Herne, T. M., Tarlov, M. J., and Satija, S. K. (1998). J. Am. Chem. Soc., 120, 9787. Sam, M. et al. (2001). Langmuir, 17, 5727. Huang, E. et al. (2001). Langmmr, 17, 1215. Hartwich, G., Caruana, D. J., de Lumley-Woodyear, T., Wu, Y., Champbell, C. N., and Heller, A. (1999). J. Am. Chem. Soc., 121, 10803. Watterson, J. H. et al. (2000). Langmuir, 16, 4984. Gillespie, D. and Speigelman, S. (1965). J. AM. Biol, 12, 829. Pease, A. C. et al. (1994). Proc. Natl. Acad. Sci. USA, 91, 5022. Maskos, U. and Southern, E. M. (1993). Nucleic Acid Res., 21, 2267. Armistead, P. M. and Thorp, H. H. (2000). Anal. Chem., 72, 3764. Halliwell, C. M. and Cass, A. E. G. (2001). Anal. Chem., 73, 2476. Korri-Youssoufi, H. et al. (1997). J. Am. Chem. Soc., 119, 7388. de Lumley-Woodyear, T., Caruana, D. J., Champbell, C. N., and Heller, A. (1999). Anal. Chem., 71, 394. Caruana, D. J. and Heller, A. (1999). J. Am. Chem. Soc., 121, 769. Anderson, M. L. M. (1999). Nucleic acid hybridization, p. 9. BIOS Scientific Publishers Ltd, Oxford, UK. Bej, A. K. (1996). In Nucleic acid analysis, principles and bioapplications, (ed. C.A. Dangler) p. 1, Wiley-Liss, NY. Bolton, E. T. and McCarthy, B. J. (1962). Proc. Natl. Acad. Sd. USA, 48, 1390. Breslauer et al. (1986). Proc. Natl. Acad. Sci., 83, 3746. Sugimoto et al. (1996). Nucleic Acids Res., 24, 4501. Anderson, M. L. M. and Young, B. D. (1985). In Nucleic acid hybridisation, (ed. B. D. Hames and S. J. Higgins) p. 73, A practical approach series. IRL Press. Oxford. Anderson, M. L. M. (1999). Nucleic acid hybridisation, p. 78. BIOS Scientific Publishers Ltd, Oxford UK. Gani, S. A., Mukherjee, D. C., and Chattoraj, D. K. (1999). Langmuir, 15, 7130. de Lumley-Woodyear, T., Campbell, C. N., and Heller, A. (1996). J. Am. Chem. Soc., 118, 5504. Kuhr, W. (2000). Nature Biotech., 18, 1024.

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Chapter 3 Screen-printing methods for biosensor production Xian-En Zhang Wuhan Institute of Virology, Wuhan, China.

1 Introduction Conventional enzyme electrodes use bulky electrochemical devices as transducers, such as the oxygen electrode and hydrogen electrode. Applications of the sensors are attractive but their market development is slow and limited due to the fact that these sensors, either electrodes or enzyme membranes, are made expensively and need frequent calibration and regular maintenance. In addition, the interferences from variation in oxygen tension or unspecific electro-active substances are also problems. Therefore, cheaper, more reliable and "friendly" biosensors are required to promote exploitation of the market. This had not been realized until the invention of the mediated enzyme electrode and introduction of screen-printing technology. The mediated enzyme electrode was first demonstrated in 1984 (1). The authors employed the ferrocene/ferricinium ion couple to replace molecular oxygen to shuttle the electron between an enzyme redox center and electrode. Its minimal sensitivity to oxygen concentrations, to change in pH and to nonspecific electroactive substances meant that it was possible to incorporate it in a device with a wide range of applications, especially in vitro or in vivo monitoring in whole blood and fermentation process, where oxygen tension fluctuate. This was the start of the second generation of biosensors, concluded by Professor Scheller in the First Congress on Biosensors (1990). The screen-printing technology is conventionally used in both the electronic and printing industries. It once became the core part of a patent for preparation of the thick film electrochemical sensor in 1981 (2). The technology has advantages of design flexibility, process automation, good reproducibility, a wide choice of materials, and reduced expense. It had been thus sought as an alternative method for mass production of biosensors at low cost. In 1987, a few groups reported their disposable enzyme sensors separately (3-5). All these sensors were based on the mediated and screen-printed

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enzyme electrode. An early example was a pen-sized glucose meter that used ferrocence-mediated enzymatic electrochemistry and screen-printed technology to give a digital glucose readout only 30 s after the application of whole blood. The sensor was for "one-shot" measurement and was disposable. The authors claimed that "this pen marks the beginning of a new generation of devices involving direct electrochemistry which allows it to be fast, accurate, precise, and silent." At the time, two types of the ExacTech (later known as Medisense ExacTech) personal glucose monitors, "pen-type" and "credit card-type" soon became commercially available. "Within two-years of launching into this S billion world market, the product had achieved a 20% market share in the United States and were making substantial inroads into the market worldwide," claimed Higgins in the First World Congress on Biosensors (6), and the annual sales reached half a billion US dollars in the middle of 1990s as estimated by Turner of Cranfield University in the 4th World Congress on Biosensors (1996). As a result the personal blood glucose monitor for people suffering from diabetes is known as the best biosensor examples. This early stage progress was followed by a wide variety of investigations referring to the thick film biosensors, giving a strong push on the second generation biosensors. In 1994, 10 years after the publication of the first mediated enzyme sensor, the inventors shared the Mullard Prize awarded by the Royal Society for their most original contribution to the field. The progress of the field has been documented from various aspects (7-13). The aim of this chapter is to further outline the techniques and applications of screen-printed biosensors.

2 Screen-printing technology The screen-printing process may be a hand operation, a semi-automatic or automatic machine process. Due to the widespread establishment of screen-printing processes in industry, fabrication devices and various materials are commercially available.

2.1 Materials and methods 2.1.1 Substrate matrix The substrate matrix is the supporting material that provides surface for printing of the functional and constructional parts of the sensor. Since the electronic signal generated by the biosensors is small, the substrate must be inert. The cost of the substrate is also a consideration because the sensor is a "one-shot" or disposable measuring device. Some materials have been exploited for this purpose. PVC, which features dielectric properties, chemical inertness, low cost, and workability, is the most commonly used substrate (14-18). Ceramic (aluminum oxide, A12O3) is also frequently used for its excellent properties such as strength, hardness, heat resistance, and corrosion resistance (19-21). Other possible materials are polycarbonate (22, 23), nitrocellulose (24), and glass fiber (25). 42

SCREEN-PRINTING METHODS FOR BIOSENSOR PRODUCTION

2.1.2 Inks The inks or pastes are deposited sequentially through masks or screen on the substrate to form the functional and structural parts of the sensors. A wide range of inks with different physical and chemical properties (e.g. viscosity, conductivity, thermal resistance, and water resistance) can be found to meet diversified requirements in biosensor fabrication (13). They can be classified as two main categories, conductive inks and dielectric inks. The conductive inks form conductive tracks and the electrodes of the sensors. They are made of conductive materials, binding agent, solvent, and additives. The conductive material maybe gold, platinum, silver, or carbon (graphite) powder, which is dispersed in a binding agent. The carbon paste (CP) consists of a mixture of graphite powder and an organic binder. The electrodes made with CP show relatively low background current, a wide operating potential window, convenient modification, renewability, and low cost (26). Because of these fascinating properties, carbon paste electrodes (CPEs) are in extensive use in electroanalysis. Electrochemical performance of a series of CPEs was investigated for screen-printed sensors (27). The binding agent may be epoxy resin, alkyd resin, acrylic resin, polyurethane resin, or phenolic resin, which are dissolved in relevant organic solvents (e.g. mineral oil, ester, alcohol, ketone, etc.). Choice of a solvent is based on its solubility for the binding agent and volatility. Additives may be needed to improve the performance of the ink (such as viscosity, adhesion, curing time, etc.). For instance, a surfactant helps to disperse the carrier powders in the ink so as to prevent them from agglutination and sedimentation. Another example is the drier, which can shorten the curing process by stimulating oxidation of unsaturated bonds of oil molecules or inactivating the natural antioxidants (e.g. proteins, phospholipids, vitamins, which may coexist in the diluting agents). The drier is mainly composed of metal ions and organic acid. Cobalt, manganese, and zinc are the most important ones in this respect. The dielectric ink, usually made with polymer or ceramic, forms the insulation or encapsulating layer of the sensor. It has to be compatible with the substrate, that is, PVC ink is for PVC substrate and is not favorable to ABS substrate. The commercial inks are normally provided as concentrated packages, which need to be diluted before application. Diluents which usually come with the inks increase fluidity and decrease viscosity of the inks. Immobilization of inks after printing is called the curing process, which may be carried out at ambient temperature, by thermal curing or by ultraviolet radiation (UV). Curing at ambient temperature is regarded as an all-purpose method because it is a mild process, although it takes hours or days, longer than other methods. Thermal curing is employed when the substrate is ceramic, it makes stronger binding between inks and substrate and it reduces the curing time. However, the thermal curing is not suitable for the substrates that are not thermally resistant, such as PVC and other plastic materials. Ultraviolet inducible immobilization is a quick curing process and usually takes only a few seconds. It is usually composed of one or more UV curable

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binding agents (e.g. polyurethane resin and epoxy resin or alkyd resin), padding material (e.g. graphite powder), photosensitive agent, and adjuvant. The UV curable binding agent molecules adsorb the energy from radiation to cause so-called "thermal polymerization," the cross-linking reaction between its unsaturated carbon bonds, or "oxidation polymerization," the polymerization reaction between peroxides or hydroxides and the binding agent (28, 29). Photosensitive agents induce the immobilization of the binding agent, which takes a key role during the UV curing process. Benzophenone and benzoin dimethyl ether are the common photosensitive agents. Addition of a sensitizing agent (e.g. nitrile bases) may assist further in the curing. Valuable studies on the method have been carried out by some investigators during last few years (30-33). They showed that UV curing is a highly efficient method and that it needs no expensive facility and consumes less energy. Most importantly, its high printing speed allows the real online production of biosensors. However, some problems for UV curing remain to be solved. For example, quick immobilization of the ink brings about a high inside stress of the coating, which may lower the adhesion force to the substrate. Other drawbacks include costliness and short storage time of the ink prior to printing. Conducting and dielectric inks are well prepared by the commercial suppliers. They meet most of the requirements of the screen-printed sensors fabrication. Direct use of the commercial ink is recommended. However, in some cases, formulations have to be custom-made for specific purposes, for example, to modify the working electrodes, following indications in the literatures. Table 1 lists the basic information of the inks for biosensor fabrication, either commercial products or custom-made formulas.

2.1.3 Sensing element The sensing element is the analyte-specific part of biosensors. From a catalytic point of view, any enzyme could act as a sensing element. However, some enzymes are very expensive to obtain and are not affordable for disposable biosensors. And yet, most enzymes show poor thermal stability. Their shelf lives are too short to be used as sensing elements because the "one-shot" biosensor has no need to be calibrated by the customer, loss of enzyme activity during storage causes significant determination error. Additional stabilization treatment may need even for glucose oxidase that is widely used as a model enzyme for biosensors. Up to date, oxidases and dehydrogenase are the most frequently used sensing elements. Electrochemical properties of many of these enzymes have been well characterized on the electrodes. As mentioned above, a variety of mediators, such as ferrocenes, benzoquinones, and active dyes, may be added to reduce the working potential. As an option, some biological elements, other than enzymes, may be used as sensor elements, for example, antibodies (38-42) and microbial cells (43). The sensing element is immobilized onto the surface of the electrodes through covalent attachment, cross-linking or adsorption. Automated immobilization can be achieved either by ink jet techniques or screen-printing 44

Table 1 Inks that are typically used for screen-printed biosensors Composition or commercial product

Application

Immobilization

References or supplier

70% graphite powder and 30% paraffin, or 60% graphite powder and 40% light mineral oil (w/w). 1-4% (w/w) mediator, such as ferrocene (for oxidases) or Meldola's Blue (for dehydrogenases) may be added to reduce working potentials

Working electrode

Air dry or thermal curable if enzyme and mediator is not incorporated

(34)

70% graphite powder and 30% paraffin, or 60% graphite powder and 40% light mineral oil (w/w).

Padding for conducting track

Air dry or thermal curable

Carbon/graphite ink, SS, or Electrodag

Working electrode or conducting pad

71 °C, 2-5 mm

Acheson

Ag paste

Silver pastes, Electrodag series

Working electrode and conducting track

71-95 °C, 2-5 mm

Acheson

Ag-Pd paste

No. 7474 or QM 22

Reference electrode and conducting track

850 °C, 30 mm on AI203 substrate DuPont, Bad Homburg, Germany

Reference electrode

71-107 °C, 2-5 mm

Acheson

Reference electrode

Air dry

(35)

Ink

Conductive Carbon paste

Ag/AgCI paste Silver/Silver chloride pastes, SS, and PE series 0.2 g AgCI with 1 g Ag ink

Table 1 (Continued) Composition or commercial product

Application

Immobilization

References or supplier

PVC ink, SS seires

Insulating layer

Air dry

Zhongyi inks

Insulating layer, compatible with Au

80 °C, 20-25 min and UV exposure 1-3 s. Compatible with 96% alumina ceramic substrate and Au conductorz

DuPont

3-4% (w/v) hydroxyethyl cellulose added with Triton 100 (0.01%) and polyethylene glyco (3%)

Out dialysis membrane

Air dry

(35)

CA

2% (w/v) cellulose acetate

Out dialysis membrane

Air dry

Gafquat

4% cationic form of Gafquat. Gafquat 755N (Internations Specialty Products Ltd.)

Stabilize enzyme membrane

Air dry

Ink

Dielectric PVC paste

Ceramic paste Fodel 6050 dielectric paste

Encapsulating HEC

(36)

SCREEN-PRINTING METHODS FOR BIOSENSOR PRODUCTION

carbon-containing inks. The latter is thought to be a particularly favorable application for the production of biosensors (13) and is used for the glucose sensors produced by Medisense Inc. (Cambrdge, MA). To print the sensing layer, the sensing element is integrated into the ink to form the sensing element ink (30, 44-46). The ink has to be carefully prepared to ensure that the sensing element is dispersed in the ink properly. Otherwise, a determination error would occur. One drawback of the printing method is that the large amount of enzyme ink must be applied at a time, which may not be economical on the laboratory scale. The ink jet may be a more flexible method, it can be performed easily either in the mass production or laboratory scale. Both methods are applied in the later stage of fabrication so that the sensing element is not exposed to the high temperatures that may be needed during the initial thermal curing processes. In some cases, coatings of semipermeable membranes (e.g. Nafion or cellulose acetate) (47, 48) can be printed to the surface of the sensor so as to prevent the enzyme from leaking and to reduce the influence of possible interference from the sample solution.

2.2 Apparatus The screen-printing machine may be horizontal, vertical, or desk mounted. The printing surface may be flat, rotary, or cylindrical and the printing process may be direct, indirect, or electrostatic deposition. But the flat and direct printing machines are most convenient for biosensor fabrication. With the growing interests in screen-printing, the microprocessor-controlled automation process has largely replaced the hand-made process. An up-to-date automated screen-printer is equipped with advanced microcomputer controls with multi major functional program presettings. As shown in Figure 1, the main parts of the machine normally include a printing table with various printing sizes, pneumatic frame clippers making prepress preparation or printing frameshift much easier and faster, a squeegee that is driven by pneumatic element, a frequency regulator that controls the printing speed, and a micro adjuster that leads to high precision. Figure 1 is a diagram of the core part of the screen-printing machine.

Figure 1 Screen-printing process. Negative pressure is generated by an air compressor.

47

XIAN-EN Z H A N G

Figure 2 Diagram of screen-printing process for preparation of biosensors.

In InkJet printing, the liquid dispenser that delivers the sensor element to each sensor strip operates at controlled volumes. It is usually integrated into a platform to combine motion control with line and dot dispensing. This allows one program to control dispense speed, line concentration, drop size, and position. Biojet QuantiSOOO™ (BioDot Inc.) is an example used in the author's laboratory (Figure 2). The device is based on the combination of a solenoid valve to form drops and a syringe pump to meter reagent from a positive volume displacement. The system is capable of drop-on-demand dispensing precisely defined drop sizes (as low as 4.16 nl/drop) that are determined by the stepping resolution of the syringe pump (driven by a high resolution stepper motor) and synchronizing the stepper increments with the opening and closing of the solenoid valve. This state-of-the-art device provides non-contact dispensing of individual drops, which are quantitatively defined in either line or dot formats. Figure 3 shows the ink jet nozzle with a group of screen-printed electrodes on the substrate board.

2.3 Printing patterns The sensor may be a three-electrode (working electrode, counter electrode, and reference electrode) or a two-electrode configuration (without counter electrode). Conventional cyclic voltammetric experiments incorporate a three-electrode system avoiding current flow through the reference electrode. Instead, for small electrodes, the current flowing in the cell will not be large and therefore will not perturb the reference potential significantly, it may be convenient to use only a two-electrode configuration. If a multiple measurement is considered, the sensor can be designed as a multi-working electrode pattern. A group of printing masks is prepared using photolithographic technique, each mask provides a pattern for printing the relevant layer. The mask is made with nylon, polyester, or stainless steel meshes (100-300 meshes per inch with the fiber thickness 30-60 |j,m) that are stretched over a metal (e.g. duralumin) or wooden frame through an assembling machine. The mesh is coated with a photosensitive gel, 48

SCREEN-PRINTING METHODS FOR BIOSENSOR PRODUCTION

Figure 3 Ink jet nozzle with the screen-printed electrodes. The nozzle is jetting the enzymemediator solution onto each working electrode. The nozzle is fixed in position while the platform that carries the screen-printed electrode board moves from right to the left, driven by a high resolution stepper motor, ensuring the finely controlled volume size of enzymemediator solution to be deposited on the correct position of each electrode surface.

on which a master plate is then conglutinated tightly. The plate is made from a template by a photographic method. During exposure, the coating at the transparent area of the plate is solidified, leaving the covered area glutinous. The latter is to be removed by a solvent. After washing and drying, the mask is checked carefully. Additional modification steps may be required to ensure the quality of the mask, for example, in order to increase the solidity of the mask, repeated exposure or a hardening agent is applied occasionally.

2.4 Printing process Inks are deposited sequentially on a clean substrate by forcing them through a group of masks with the squeegee. Each layer is deposited following the corresponding mask. Drying or curing is performed between two printing stages. The ink that contains the sensing element generally forms the final layer, however it may be covered with a layer of semipermeable membrane when necessary. Figure 3 shows the diagram of a printing process, taking a two-electrode configuration glucose sensor as an example. Details are demonstrated in Protocol 1 that is routinely operated in our laboratory. The optimum screen-printing process may be different as it depends on the materials to be deposited, the intended application and the type of machine employed. Evaluation of the screen-printed 49

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electrochemical enzyme sensors can be performed with cyclic voltammetry and chronoamperometry methods.

Protocol

Preparation of a screen-printed glucose electrode A model S-600M screen printer (Ever Bright Printing Machine Ltd., Hong Kong) is used to print the electrodes throughout. All drying processes are undertaken at room temperature. 1 Clean the PVC substrate (150 mm x 150 mm x 0.5 mm) with tap water, anhydrous ethanol, and distilled water sequentially. Each sheet of the substrate allows a group of 45 sensors to be printed. 2 Fix the cleaned PVC substrate on the printing table of the printing machine by negative air pressure. 3 Place the first mask, which is designed for printing conductive strips, over the substrate on the table against registration stops, ensuring accurate pattern fitting. 4 Pour the silver ink onto the screen. Spread the ink over the surface of the screen with the squeegee so that the ink is pushed through the mask. 5 Let the ink solvent evaporate at ambient temperature. Print a carbon pad on to the silver strips by the same printing procedure with the second mask. Allow the ink to dry. The carbon pad protects the silver strip from oxidation through being exposed to air. 6 Print the reference electrode, with the third mask, by applying a silver ink mixed with finely ground silver chloride in the ratio of 2 g AgCl per gram of ink, onto one end of a graphite pad. The ink is again left to dry at room temperature. 7 Print the insulating layer, with the fourth mask, by applying a PVC ink, leaving terminals and active surfaces exposed. The active surface is a circular area that forms a planar, two-electrode electrochemical cell. One semicircle is the carbon electrode and the other is the silver/silver chloride reference electrode. 8 Prepare a saturated 1,1'ferrocenedimethanol solution in distilled water. Dissolve Aspergillus niger glucose oxidase into the ferrocene solution, making the final concentration 10 units GOD/|j,l. 9 Drop 1-2 |j,l of the enzyme-ferrocene solution onto each surface of the carbon electrodes by using the BioDot liquid dispenser. 10 The electrodes are then allowed to dry in a desiccator at room temperature. 11 Print an outer membrane (HEC) on the surface of the electrode cell to cover the enzyme and ferrocene layer. The HEC solution is prepared by mixing 3.5% (w/v) hydroxyethyl cellulose, 0.02% Triton XI00, and 3% polyethylene glycol in distilled water. 12 After the water had evaporated under ambient conditions, the sensors can be stored over silica gel at 4 °C.

50

SCREEN-PRINTING METHODS FOR BIOSENSOR PRODUCTION

3 Applications As they are inexpensive, reliable and rapid, screen-printed biosensors have been recognized so far as the most important technique pushing biosensors into practical use. During last 10 years, hundreds of screen-printed biosensors have been reported, with applications in a wide variety of analytical problems in medicine, pharmaceuticals, the environment, food, bioprocess, security, and defense. Some of them are made as pocket devices for home care of the patients and for field tests. Many of them are believed to soon become commercially available.

3.1 Clinical diagnosis Clinical importance of the screen-printed biosensor has been well demonstrated by the pocket blood glucose meter that can be found in the market in many styles. Most of the meters are designed as a one-step process. Only one drop of finger blood is needed for the detection. The result can be obtained in less than 1 min. With these devices millions of diabetics can test their blood glucose level at home, therefore, careful control of the disease is thus feasible so as to greatly improve their life quality. Subsequent studies were carried out for improving the sensor performance further. A detection limit to 0.02 mM was achieved by the enzyme electrodes prepared from platinized Vulcan XC-72 carbon particles (49), and an increased linear range up to 45 mM was achieved in the author's laboratory (to be published). A highly selective glucose sensor using copper (Il)-hexacyanoferrate and GOD with a carbon ink has been described (50). The dispersed metal-hexacyanoferrate catalyst offers a marked decrease in the overvoltage for the oxidation of the enzymatically liberated hydrogen peroxide, and gave a stable response at physiological pH. Such efficient catalytic activity allows tuning of the detection potential to a region (around — 0.1 V) where interfering reactions are negligible. Such an operation eliminates the need for an anti-interference membrane barrier, and along with the one-step dispersion of the enzyme and electrocatalyst, greatly simplifies the sensor fabrication. One problem encountered is the effect of temperature on the enzyme electrode. Since the enzyme electrodes provided to the customers were pre-calibrated by the manufactures at a certain temperature, errors will occur if the detection was performed at a different temperature, because the response signals varies with variation in temperature. Automatic temperature compensation was built based on careful investigation of the temperature coefficient of the enzyme electrode (51), making the device more reliable. Other important metabolites that have been measured with the screen-printed sensors are lactate (16, 34, 52), uric acid (53, 54), urea and creatinine (55), cholesterol (56), progesterone (57). In particular, sensors for urea and creatinine may fulfill a clinical demand in kidney dialysis, the treatment for renal disorder patients. During the dialysis process, concentration of blood urea and creatinine are used as control parameters. A bedside measurement sensor will provide the 51

XIAN-EN ZHANG

useful information to the doctor in a timely fashion and reduce the treatment costs significantly.

3.2 Food analysis bioprocess control In addition to the importance of glucose and lactate as major analytes in clinical diagnosis, they are also parameters in many foods, beverages, and bioprocess control. A screen-printed lactate sensor was constructed and applied with a flow injection system based on dialysis, to off-line and online monitoring of Geotridmm candidum cultivation producing lactate oxidase (58). By online injection analysis of undiluted fermentation broth over a period of 12 h prior to the inoculation, no significant loss of sensor response was observed which demonstrates the good operational stability of the system even with complex real samples. The stability of the sensors was due to use of UV polymerizable carbon ink and optimized working condition. Recently, a novel flow injection analytical system with a screen-printed enzyme sensor incorporating Os-complex mediator has been developed for continuous determination of glucose, giving a through put of about 40 samples per hour. The glucose biosensors retained their constant response after more than 100 injections and storage over a month. The design is suitable for automatic and rapid determination of glucose (59). Multipurpose biosensor is particularly useful in the analysis of foods and beverages. Its feasibility increased since the introduction of screen-printing technology, with which combination of multifarious sensing probes can be easily designed. The first example was a design for simultaneous determination of glucose and ethanol (60), followed by the sensors for determinations of biogenic amines for the fish freshness estimation (61, 62), maltose and glucose for dry beer quality control (35), starch and glucose for starch hydrolysis process monitoring (63), sucrose and glucose for analysis of soft drinks (48), and glucose and glucoamylase activity for control of glucoamylase production (64). The results showed great interests in the food and fermentation industries. Table 2 gives details of some applications.

3.3 Environmental monitoring With the growing concern of human beings in relation to their living environment, detection of toxic pollutants becomes more important than ever for environmental evaluation and protection. For instance, the effects of chemical pesticides on human health has drawn global attention due to their inappropriate use worldwide. Some pesticides are almost non-degradable, they can accumulate in the human body through the food chain, causing chronic poisoning. Some are very toxic, causing acute poisoning to animals and humans. Organophosphate pesticides, known as the strong nerve inhibitors, have dominated the pesticides market for many years and have caused numerous cases of acute poisoning due to careless intake of polluted vegetables, fruits, tea, or water. They are now prohibited in many countries. Field or local detection of 52

SCREEN-PRINTING METHODS FOR BIOSENSOR PRODUCTION Table 2 Some screen-printed biosensors and their performances Analyte

Sensor element

Electrode configuration

Performance

Lactose in milk

p-galactosidase, glucose oxidase

Pt working electrode Ag/Pd reference electrode

2-25 mmol/l SD: (65) 8.81% (n = 16) Stability, longer than 3 months

Lactic acid

Lactate dehydrogenase, NAD(+)

Meldola blue modified carbon working electrode and an Ag/ AgCI combined reference/counter electrode

1-20 mmol/l CV = 8.7% (n = 6)

(34)

Alcohol in beverages

Alcohol dehydrogenase, NAD(+)

Meldola's blue modified carbon working electrode and an Ag/ AgCI combined reference/counter electrode

35 mmol/l 90% residual activity after 49 days, stabilized by trehalose

(15)

NADH and lactate

NAD(+ ) and lactate dehydrogenase

NADH, 3-60 nmol/l Lactate, 0-20 mmol/l

(16)

Uric acid and hypoxanthine fish freshness

Xanthine oxidase

1-50 nmol/l CV: 2%

(62)

Maltose and glucose

Carbon working Amyloglucosidase/ glucose oxidase (A/G) electrodes Ag/AgCI reference electrode

Starch and glucose

GOD Glucoamylase

Carbon working electrode Ag/AgCI electrode

Lysine in fermentation

Lysine oxidase entrapment into polyurethane hydrogel

Pt working electrode Ag/AgCI pseudo reference electrode, Carbon counter electrode

Gentamicin in milk

Anti-gentamicin antibody labeled with GOD

Flow electrochemical cell

Carbon working electrode

Reference

Maltose, -20 mmol/l (35) Glucose, -40 mmol/l CV: 3.5-5.29% Starch, 0.4% (w/v) Glucose, -20 mmol/l

(48)

(66)

0-10 ng/kg CV: = 13.2%

(67)

these pesticides is of particular importance. Sensors based on the screen-printed technology have been constructed to detect some of the important pesticides, such as 2,4-dichlorophenoxyacetic acid (68-70), organophosphate pesticides (71-79), carbamates (78, 80), polychlorinated biphenyls (81, 82). The sensors for organophosphate pesticides are based on the inhibition, through binding reactions, of cholinesterase or acetylcholinesterase that play an important role in nerve conduction. The principle was introduced to construct biosensors for more than 20 years but none of these sensors were reliable due to reasons of cost and poor reproducibility. The screen-printed biosensors may be the answer 53

XIAN-EN ZHANG

to the problem. Other applications using screen-printed biosensors include determination of nitrogen compounds which are one of the major factors causing eutrophication. Local determination of these compounds in water provides useful information for the control of eutrophication. A research team in England reported a group of screen-printed sensors used for the determination of ammonium ion (83, 84) and sulfite (85). The proposed biosensors were evaluated on samples of unspiked and spiked river water; the recovery and precision data indicated that the devices could be expected to give reliable data in these waters. Another example is quick estimation of biological oxygen demand (BOD). BOD indicates the level of organic pollution in the water. Conventional methods need 5 days to operate. With the BOD sensor, a combination of living microbial cells and oxygen electrode, the result can be obtained in just 20 min (86, 87). A disposable BOD sensor, based on mediated oxidation of organic material by the cells, was proposed (88). This protocol, however, needed further study to answer the question on how a chemical mediator interacts with the microbial cells.

3.4 Other approaches DNA sensor technology is one of the notable progresses in the biosensor field. A disposable electrochemical sensor for the detection of short DNA sequences is described. Synthetic single stranded oligonucleotides have been immobilized onto graphite screen-printed electrodes through binding of avidin-biotinylated oligonucleotide and adsorption at a controlled potential. The probes were hybridized with different concentrations of complementary sequences. The formed hybrids on the electrode surface were evaluated by differential pulse voltammetry and chronopotentiometric striping analysis using daunomycin hydrochloride as an indicator of the hybridization reaction. The DNA sensor was able to detect 1 ng/ml of target sequence responding to point mutation human diseases (89). Another DNA sensor was made of an ion-exchange film-coated screen-printed electrode adapted to the bottom of a polystyrene microwell. An alkaline phosphatase label was used to hydrolyze the monoester phosphate salt of [(4-hydroxyphenyl)aminocarbonyl]-cobaltocenium. This anionic substrate is transformed into a cationic electroactive product, which is then accumulated by ion exchange at the electrode surface to give an amplified electrochemical response. Detection limits of 10 amol/ml was thus achieved for amplified cytomegalovirus DNA. The sensor was also used to assay human chorionic gonadotropin hormone (hCG). The method was 10-40-fold more sensitive than the conventional absorption spectrophotometry using p-nitrophenyl phosphate as the substrate (90). Based on this work, it is likely that we can exploit the screen-printed oligonucleoitide array for mass DNA sample analysis. Biomimetic or molecular imprinting is being increasingly recognized as a versatile technique for the preparation of synthetic polymers containing tailor-made recognition sites. This is achieved by co-polymerizing functional and cross-linking monomers in the presence of an analyte, which acts as a molecular template. After elution of the template, complementary binding 54

SCREEN-PRINTING METHODS FOR BIOSENSOR PRODUCTION

sites are revealed that allow specific rebinding of the analyte (91, 92). The method was proposed for biosensor purposes in 1990 (93) and has shown its utility in recent years (94, 95). However, such sensors need to be mass-produced cost effectively. Attempts were made by a few investigators. A biomimetic sensor based on the combination of a screen-printed peroxidase electrode and an iron porphyrin complex iron(III)-meso-tetrakis-(pentafluorophenyl)-beta-tetrasulfonatoporphyrin chloride was proposed for aliphatic hydrocarbons, using octane as the model analyte. The biomimetic compound was immobilized in a polyelectrolyte matrix. Catalytic reaction was described in the homogeneous system and on the sensor surface (96). The binding principle was used to build an imprinted polymer-based sensor for herbicides (2,4-dichlorophenoxy acetic acid). The method involves a competitive binding step of the herbicides with a unrelated electrochemically active probe, 2,4-dichlorophenol that showed very high non-specific binding to the imprinted sites. The imprinted polymer particles were directly coated onto the screen-printed carbon working electrode. Following incubation of the modified electrode in a solution containing the analyte and the probe, the bound fraction of the probe was quantified by differential-pulse voltammetry. This system provides a cheap, disposable sensor for rapid determination of environmentally relevant and other analytes (91).

4 Conclusion The screen-printing technology has shown its great utility to biosensor researchers. During the last 10 years, it has become a mature, widespread sensor technology, drawing biosensors out of "the tower of ivory," for use by the people. With the technology, biosensor design can be more flexible: disposable or reusable, single or multiple purpose, planar or spherical sensing surface, mediated or direct electrode process. Therefore, further application of the technology in the biosensors field is very promising. It is, however, pointed out that, at present, most of the screen-printed biosensors are disposable; successful developments of the sensors largely rely on the cost and stability of the sensing elements. This is an intrinsic problem of biological nature. Many genetic expression vectors have been developed for high-level expression of the enzymes or other functional proteins, and protein engineering provides a powerful tool for improving the enzyme toward the desired properties, which may be a potential resolution to the problems. (See Chapter 9, this volume.)

Acknowledgment The author thanks the Comet Bioelectronic Tech Company (Wuhan, China) for its consultancy in the preparation of screen-printed biosensors, and Zhi-Ping Zhang, Wei Li, and Li-Qun Chen for valuable discussions.

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58

Chapter 4 Kinetic modeling for biosensors Philip N. Bartlett and Ghee-Seng Toh Department of Chemistry, University of Southampton, Southampton, S017 1BJ, UK.

1 Introduction 1.1 The purpose and practice of modeling The purpose of a kinetic model of a biosensor is to identify the key experimental factors (such as the rates of reactions, rates of mass transport, loading of the bio-recognition component, etc.), which determine the response, or output, of the sensor and to then provide a link between these key experimental factors and the concentration of the analyte and the sensor response. Thus, modeling provides a mathematical description of the physical processes occurring within the system. By applying kinetic modeling to enzyme electrodes and biosensors, we can derive a number of advantages. It is clear that several different kinetic processes are involved in the overall functioning of any enzyme electrode or biosensor. In an enzyme electrode these include the reactions between the enzyme and its substrate, between the enzyme and the mediator, and between mediator and electrode, as well as the various mass transport processes which bring reactants to the electrode and take products away. On its own, any single measurement cannot give any insight into the relative significance of the different kinetic steps. If, however, a set of data from the sensor, obtained under a sufficiently wide range of different conditions, is analyzed and compared to a kinetic model for the sensor, this will yield more intimate knowledge of the system. In turn, this will provide the experimenter with the tools with which to plan future experiments or with which to rationally improve sensor performance for a specific application. In contrast, in the absence of a suitable kinetic model for a particular sensor, it will be necessary to carry out a large amount of trial and error experimental work in order to achieve a comparable understanding of the system. This is very time consuming and it will have to be repeated, effectively re-inventing the wheel, each time a new but related system is studied. Developing a kinetic model is not only an efficient experimental approach, it also gives insight into the mechanisms and processes involved in the operation of 59

PHILIP N. BARTLETT AND CHEE-SENG TOH

the sensor. This is especially true when we approach the analysis using approximate analytical methods (see below). This is a highly satisfying experience for the individual, and it can produce a rich reward in terms of the new knowledge that can be obtained from such a pursuit. In this chapter, we will take the reader through a basic understanding of enzyme kinetics, electrochemistry, and the basic mathematical tools necessary for an understanding of kinetic analysis found in the literature on enzyme electrodes. Toward the end of the chapter, the reader will find a table (Table 2) summarizing much of the published work in this field from 1992 to the present. This will be useful for those who wish to take the subject further and to apply these approaches to their own data for enzyme electrodes. In what follows we concentrate heavily on amperometric enzyme electrodes. This is because there is a much greater amount of literature on the modeling of this type of biosensor than others. However, the same general principles will apply in developing models for other biosensor types including potentiometric and conductimetric biosensors as well as optical and gravimetric sensors. In all cases it is necessary to consider the interplay of mass transport to, or from, the sensor surface and its interaction with the kinetics of the biorecognition process and the transduction mechanism. At the end of the chapter we briefly review the modeling of other biosensor types.

1.2 Enzyme kinetics 1.2.1 Equilibrium and the steady state At equilibrium, the concentrations of reactants, intermediates, and products are constant; however, this does not mean that there is no reaction going on. Equilibrium is a dynamic state. At equilibrium, the rates of the forward and reverse reactions (i.e. the products of the appropriate rate constant and reactant concentrations for each step) must be in balance. In general, enzyme electrodes do not operate at equilibrium, rather they operate in the steady state where there is a constant supply of fresh reactant to the electrode. In the steady state, the rates of change in the concentrations of the reaction's intermediates are negligible so that the rates of reaction are unchanging with time. Under these conditions we can equate the fluxes (i.e. the number of moles of reaction occurring per unit area in unit time) for the different reaction steps in order to derive useful kinetic information about the system. For biosensors these reactions generally occur at or near the sensor surface rather than uniformly throughout the solution; consequently, the concentrations of reactants and products at the sensor surface will be different from those in the bulk. As a result, there will be concentration gradients set up and net fluxes of material to and from the electrode surface.

1.2.2 Michaelis-Menten kinetics The simplest description of steady-state enzyme kinetics is based on the work of Michaelis and Menten (1). It assumes that the substrate first forms a complex 60

KINETIC MODELING FOR BIOSENSORS

with the enzyme in a reversible step and that equilibrium is maintained between the enzyme, E, and substrate, S, and the enzyme-substrate complex, ES. The irreversible breakdown of this enzyme-substrate complex then yields the product, P.

The second assumption in this model is that the concentration of the enzymesubstrate complex is constant, so that it is a steady-state process. Clearly this is not true immediately after the reactant and enzyme are first mixed together when the concentration of intermediate is building up (called the pre-steady state phase) and will only be true as long as the concentration of substrate is not significantly depleted by the course of the reaction. The following, more generalized, mechanism in which the forward and backward rate constants for the formation of the ES complex are explicitly included, was proposed by Briggs and Haldane (2):

If eE is the total concentration of enzyme and eES is the concentration of the enzyme-substrate complex, then the concentration of uncomplexed enzyme is (eE — eEs). Assuming that the concentration of substrate is much greater than the concentration of enzyme (which is generally the case), then the concentration of uncomplexed substrate can be taken as equal to the initial concentration of substrate, s. Then we can write

At steady state, deES/dt = 0 (strictly the approximation we make is that deES/dt is small compared to the reaction flux k^s). Then,

The velocity of reaction, v, is given by

so that which can be rewritten in the form of the Michaelis-Menten equation

where kcate^ is the maximum reaction velocity and KM — (k_i+ kcat)/ki is the Michaelis constant. 61

PHILIP N. BARTLETT AND CHEE-SENG TOH

Figure 1 Normalized plot of steady-state velocity against substrate concentration for an enzyme reaction obeying Michaelis-Menten kinetics. The dotted lines show the point at which the concentration of substrate equals KM.

This model reduces to the simple form of Michaelis-Menten kinetics, with

1.2.3 Analysis of enzyme kinetic data Analyses of enzyme kinetics are often based on the Michaelis-Menten equation (7). Figure 1 shows a typical plot of reaction velocity, v, against substrate concentration, s. From Figure 1 we can see that at low concentrations the reaction velocity increases linearly with substrate concentration. This is because when s
KINETIC MODELING FOR BIOSENSORS Table 1 Linear plots derived from the Michaelis-Menten equation Equation

Plot

Slope

Intercept

Lineweaver-Burk

1/v against 1/s

KM//fcate^

lAcatej:

1 | kM ncat e^ Kcat e^- s Hanes s KM , s

s/v against s

l//fcate^

KlvlAcate^

Eadie-Hofstee

v against v/s

KM

kcatej;

1 v

V

K ca t 6^

n cat 6^

v = /we E -^

used, can lead to significant errors in the estimates of KM and kcate£. The Hanes and Eadie-Hofstee plots represent attempts to overcome this problem (see a standard text on enzyme kinetics for further details (3, 4)). These methods for producing linear plots from the Michaelis-Menten equation all date from the era when computers were not so widely available. Nowadays the easiest, and statistically more correct, method to analyze enzyme kinetic data using the MichaelisMenten model is to use a nonlinear least squares fitting routine (with weighting factors if appropriate) to directly fit the experimental data to Equation (7).

1.2.4 The significance of KM for biosensor applications For cases where k c at
1.3 Basic electrochemistry 1.3.1 Mass transport In an enzyme electrode, the enzyme catalyzed reaction occurs in a localized region at, or close to, the electrode surface. Consequently there is an interplay and interaction between the enzyme kinetics and mass transport of material to and from the electrode surface. In general, mass transport can occur by three 63

PHILIP N. BARTLETT AND CHEE-SENG TOH

processes: migration, convection, and diffusion. Migration is the movement of ions in an electric field. It does not occur for neutral molecules (such as glucose) and is not significant in solution if there is an excess of an inert electrolyte present (such as 0.15 moldm~ 3 NaCl). Convection is the bulk movement of solution caused by stirring and can be important for biosensors. Unfortunately simply using a magnetic stirrer or stirring rod to agitate the solution does not generate calculable, reproducible mass transport conditions. However, methods do exist to produce well defined and calculable mass transport (such as the rotating disc, wall-jet, or channel flow geometries). Diffusion is the movement of species down a concentration gradient and is always present when there are variations in concentration from one region to another.

1.3.2 Pick's first law Pick's first law describes the relationship between the flux due to diffusion, J (mol cm" 2 s~ a ), and the concentration gradient. In one dimension: where D (cm2 s a ) is the diffusion coefficient. Consider a situation in which the potential at an electrode is stepped to a value well above the equilibrium potential for substance S, such that it is irreversibly oxidized at the electrode in an unstirred solution. The concentration of S will instantaneously go to zero at the electrode surface when the potential is changed. The resulting time dependent concentration profiles for S are shown in Figure 2. The concentration gradient is steepest at the electrode/solution

Figure 2 Normalized concentration profile for the irreversible reaction of S at an electrode in an unstirred solution under non-steady-state conditions. The profiles are calculated for 2 -1 D=1*10-5 cm D=1*10 cm ss

64

KINETIC MODELING FOR BIOSENSORS

interface and the flux of S toward the electrode is greatest near the electrode. This high flux of S near the electrode is not matched by a similarly high flux of S further away from the electrode, see Figure 2. Therefore, the concentration profile for S will continue to change with time such that the concentration gradient at the electrode surface declines. For this semi-infinite one-dimensional case, a steady-state is never achieved. However, if we carry out the same experiment under conditions of forced convection (e.g. if we use a rotating disc electrode), where there is mass transfer of S toward electrode by both convection and diffusion, a steady state will be set up. In fact it turns out that close to the electrode surface diffusion is the dominant form of mass transport but away from the electrode convection dominates. This is because in any stirred system there is a stagnant boundary layer at the electrode surface whose thickness depends on the particular conditions. When using a magnetic stirrer, the thickness of the boundary layer is poorly defined and is not necessarily fixed. In contrast, for the rotating disc electrode the thickness of the boundary layer is well defined, calculable, and easily varied experimentally. This makes the rotating disc an ideal experimental system to study electrode reactions. For the rotating disc electrode, the diffusion layer thickness, XD, is given by:

where v (cm2 s a ) is the kinematic viscosity and W (Hz) is the rotation speed. At the rotating disc, in the steady state, the concentration profile can be described by a linear concentration profile within a stagnant layer where diffusion dominates and a constant value outside this region where convection maintains the concentration at its bulk value. (In fact the concentration profile is not strictly linear within the diffusion layer because diffusion and convection are not as sharply spatially separated as our simple description implies. Nevertheless, this simple description gives an accurate result for the flux of material at the electrode surface.) For a potential step at a rotating disc electrode, as soon as the potential is changed, the concentration of S goes to zero at the electrode surface. The concentration gradient of S at the electrode surface then decreases as the concentration polarization of S spreads out across the diffusion layer. For the first part of the experiment the response is the same as that for the stationary electrode shown in Figure 2. However, the difference comes when the concentration profile spreads out to reach the edge of the diffusion layer because now convection becomes important bringing fresh material up to the outside of the diffusion layer. This means that the system reaches a steady state in which the flux of material reacting at the electrode is balanced by the fluxes of material crossing the stagnant layer by diffusion and being brought to the edge of the stagnant layer by convection. A typical steady-state concentration profile for S at a rotating disc electrode is shown in Figure 3. The rate-limiting step under these conditions is diffusion across the stagnant layer and the steady-state flux, Jss, 65

PHILIP N. BARTLETT AND CHEE-SENG TOH

Figure 3 Normalized steady-state concentration profile for the irreversible reaction of S at an electrode at a rotating disc electrode. Calculated for D=lx 10~5 cm2 s"1, Hz, and v=0.01 crr^s"1.

is given by where sbuik is the bulk concentration. The current is directly related to the flux. If the reaction of each molecule of S at the electrode involves n electrons, the current density i (Acm" 2 ) is given by: where n is negative for oxidation (removal of electrons) and positive for reduction (addition of electrons) so that, by convention, oxidation currents are positive and reduction currents negative. Since current is directly proportional to the concentration gradient, the current changes will follow closely to the changes in the concentration gradient with time for our potential step experiment. For the rotating disc the current will start out large and will then fall to a constant value in the steady state whereas for the electrode in the unstirred solution the current will fall to zero. The current transients are shown in Figure 4. In reality instrumental limitations and double-layer charging will limit the curents at short times whereas random convection caused by thermal gradients, density gradients, and other disturbances will intervene at long times making the current at the stationary electrode erractic. In order to model enzyme electrode behavior, it is essential to understand the relationship between current, flux, and concentration gradient, as well to understand the conditions under which a steady state is achieved.

1.3.3 Pick's second law Pick's first law describes the relationship between flux and concentration gradient and, as we will see later, is essential in the kinetic modeling of the 66

KINETIC MODELING FOR BIOSENSORS

Figure 4 Current transients for the irreversible reaction of S at an electrode in (a) unstirred solution, (b) at a rotating disc electrode. Note that the effects of double-layer charging are not included in the diagram. The transients are calculated for sbu|k = l mmol dm" 3 ; D=l x 10~5 cm2 s"1; n = l; w = 9 Hz; and v=0.01 cm2 s"1.

steady-state amperometric current for an enzyme electrode. In the last section we gave a qualitative description of the evolution of these concentration gradients, and hence flux and current, with time following a potential step. The mathematical relationship which describes the rate of change of the concentration with time, and hence the basis for the quantitative description of this behavior, is Pick's second law. In one-dimension, for a species S, Pick's second law can be written as:

This second-order partial differential equation in essence states that the change in concentration within a volume element of solution as a result of diffusion is given by the difference in flux between the amount of S entering the volume element and the amount leaving, Figure 5. The solution of Equation (13) requires knowledge of the boundary conditions. For the simple case of oxidation of S at a stationary electrode in an unstirred solution following a potential step to a potential at which reaction of S is mass transport limited, the boundary conditions are

67

PHILIP N. BARTLETT AND CHEE-SENG TOH

Figure 5 Normalized concentration profile for the irreversible reaction of S at an electrode in an unstirred solution illustrating Pick's second law. At any point on the curve ds/dt<0 because fluxout >flux in . Hence the profile evolves with time as shown in Figure 2.

Solution of Equation (13) with these boundary conditions yields the Cottrell Equation (5) From Equation (17) we can see that the current density is proportional to t a'2 and, as anticipated from our earlier discussion, falls to zero at long times.

1.3.4 Coupled chemical reaction So far we have only considered the situation where there is mass transport by either diffusion or, in the case of the rotating disc electrode, by diffusion and convection. For the enzyme electrode, mass transport is coupled with chemical reaction. For example, consider the case where an electroinactive substrate, S, is converted by an enzyme into an electroactive product, P, which is detected at the electrode surface. An example here might be the hydrolysis of p-aminophenylphosphate to p-aminophenol catalyzed by alkaline phosphatase (6). We can represent the reactions as:

Now we must incorporate the reaction term into the expression for the mass transport. When we include the homogeneous kinetic step, Equation (13) becomes

where s is the concentration of substrate S. The first term on the right-hand side describes the contribution from diffusion and the second term describes the chemical reaction step. As we shall see later, enzyme electrode kinetics can be described by an equation of this form where the enzyme reaction rate replaces 68

KINETIC MODELING FOR BIOSENSORS

the chemical reaction term in Equation (20). In order to solve Equation (20) to obtain the time-dependent concentration profile for the substrate S, we must specify the boundary conditions. We can write two similar differential equations for the reaction products P and P'

In the steady state ds(x,t)jdt—0 and we can simplify Equation (20)

Integration of this equation with appropriate boundary conditions will yield the time-independent solution describing the steady state. In general, in order to solve coupled diffusion-reaction problems of the type described above, we need to integrate the differential equations which describe the reactions involved. However, in the special case where diffusion and reaction occur in distinct and physically separate regions, for example, where diffusion occurs through a membrane and the enzyme catalyzed reaction occurs in a thin layer behind the membrane, we can treat the steady-state problem by equating fluxes for the different processes involved (diffusion, enzymatic reaction, electrochemical reaction), and apply the steady-state assumption to all the intermediate species at each point in space. An illustration of these two approaches to the analysis of steady-state problems in amperometric enzyme sensors is given in Section 2.

2 Modeling Deriving a kinetic model is a process of describing the different kinetic steps involved in the chemical system in the form of mathematical equations. Therefore, the obvious place to begin is with an understanding of the different processes involved in the system under study. A useful starting point, particularly for someone who is new in the field or exploring a new system, is to look for treatments of similar systems in the literature. Later in this chapter we provide a review of the literature to assist in this. In this section we consider two different types of amperometric enzyme electrodes. First we will consider an electrode in which the enzyme is entrapped, together with a redox mediator, behind an inert membrane, which acts as a diffusion barrier (we will refer to this as a membrane] enzyme|electrode, where the vertical lines indicate interfaces between different phases). This type of enzyme electrode has been extensively modeled (7-11). Second we consider an electrode configuration in which the enzyme is entrapped within a membrane at the electrode surface together with some redox mediator (an enzyme membrane|electrode). 69

PHILIP N. BARTLETT AND CHEE-SENG TOH

2.1 The flux diagram for the membrane|enzyme|electrode It is essential to start by including all the possible kinetic steps in the model before going on to make any simplifying assumptions. A detailed flux diagram including the different mass transport and reaction steps is a good starting point. The flux diagram can then be simplified as various assumptions are included. For the membrane| enzyme|electrode, in which the enzyme and mediator are trapped behind an inert membrane at the electrode surface, the flux diagram must include the following processes. In the thin enzyme layer behind the membrane: (a) electron transfer between the electrode and the mediator (b) the redox reaction between the mediator and enzyme (c) the reaction between the substrate and enzyme. In addition there are the following mass transport processes: (a) diffusion of the substrate within the membrane (b) diffusion of the product within the membrane (c) partition of the substrate between the solution and the membrane (d) partition of the product between the solution and the membrane (e) transport of the substrate in the external solution (f) transport of the product in the external solution. In setting up this model, even at this early stage, we have already made some assumptions so that the flux diagram is not unnecessarily complicated. For example, we have chosen to model a system in which enzyme and mediator are trapped within a thin film behind the membrane, as a result we do not need to consider transport of either species through the membrane or in the external solution. The resulting flux diagram is shown in Figure 6.

2.2 Simplifying assumptions In many cases we can simplify the kinetic model considerably by making appropriate assumptions. However, it is important to consider carefully whether the assumptions you make are valid for the particular system or set of experiments. For now let us make the following assumptions to simplify the model we are considering: (a) We will assume that the enzyme layer is sufficiently thin that we can neglect concentration polarization for all of the different species within this layer. This is the essential assumption that allows us to separate the mass transport and reaction parts of the problem and thus to use the method of equating the steady-state fluxes below. (b) We will assume that the enzyme reaction is irreversible, so that the product does not affect the forward enzyme reaction. As a consequence, we can omit all the terms involving the product of the enzyme reaction. 70

KINETIC MODELING FOR BIOSENSORS

Figure 6 Flux diagram showing the different kinetic processes for an amperometric membrane|enzyme|electrode. Rate constants are represented as k, while partition coefficients are represented as K. The subscripts S and P denote the substrate and product of the enzyme reaction, respectively. E is the enzyme and M is the mediator. The subscripts red and ox indicate the reduced and oxidized forms of the enzyme or mediator, and subscript D indicates a diffusion process.

(c) We will assume that the electrochemical regeneration of the mediator at the electrode is rapid and is not rate-limiting. Hence, we can set the concentration of Mox within the enzyme layer equal to the total concentration of the mediator. As a result the enzyme-mediator reaction becomes pseudo-first order, where the pseudo first-order rate constant, k'E, is the product of the mediator concentration and the second-order rate constant for the reaction between the enzyme and mediator, k'E = kE[M]. This assumption will not hold true in all cases, for example, if the electron transfer kinetics for the mediator are slow or if the electrode potential is not chosen so that there is a sufficient overpotential to drive the reaction. (d) Finally, for simplicity, we will assume that the partition coefficient for the substrate into the membrane is unity, Ks — 1. Hence the substrate concentration within the membrane at the membrane|solution interface, smemiOUt, will be equal to the concentration of substrate in the exterior solution at the membrane surface, s0, and the concentration of substrate in the membrane at the membrane|enzyme layer interface, smem in, will be equal to the concentration of substrate within the enzyme layer, Siayer. The resulting simplified flux diagram is shown in Figure 7.

2.3 The flux equations The flux equations corresponding to the processes represented in the simplified flux diagram can now be written down. There are two important points to note when writing flux equations, the relative signs of the fluxes and the units. 71

PHILIP N. BARTLETT AND CHEE-SENG TOH

Figure 7 Simplified flux diagram for a membrane|enzyme|electrode. The symbols and terms used are the same as those in Figure 6.

In the case of homogeneous reactions we are familiar with thinking about the rate or reaction in terms of the change of concentration per unit time (mol dm ~ 3 s ~ a ). However, for reactions at electrode surfaces we are interested in the flux because this is directly related to the current. Flux is defined as the change in the number of moles per unit area per unit time (molcm" 2 s" 1 ). When writing rate equations, the theoretical basis for the derivation of flux equations is the principle of conservation of mass. By using fluxes, we do not need to consider the volumes of the system, which would be necessary if we used reaction rates. In order to convert reaction rates into fluxes, we need to include an appropriate distance. For example, the first-order enzyme catalytic rate constant for the conversion of reactant into product, kcat, is defined in the enzyme rate equation as In order to calculate the corresponding flux we need to include the film thickness, I, so that In the steady state, the fluxes for the various processes in our simplified flux diagram (Figure 7) must all be equal and can be written in terms of the current density, i, as follows

72

KINETIC MODELING FOR BIOSENSORS

where we have taken the modulus of the current to avoid the complication of specifying the sign of the current for oxidation or reduction, and k1; k_ 1; kcat, and eES have the same meaning as in the conventional enzyme kinetics described in Section 1.2. An additional parameter, the enzyme layer thickness, I, is included in the enzymatic and mediator rate constant terms, so that all the reactions are expressed in terms of fluxes. k'E is the pseudo first-order rate constant for the re-oxidation of ered by mediator in the film and is equal to kE[M], where [M] is the concentration of mediator species.

2.4 Solution of flux equations By expressing Siayer in terms of smem,out; CES. ^red. £ox in terms of eE; and ka and k_ a in terms of kcat and KM, we can solve the simultaneous equations to derive the following analytical solution:

where KM = (kCat + k-i)/ki has the same meaning as the Michaelis constant in conventional enzyme kinetics. If we want to include the diffusion process within the Nernst diffusion layer in the external solution at the outside of the membrane, we can include one more flux equation where k DiS is the mass transfer rate constant within the Nernst diffusion layer. Substituting Equation (32) into (31), we obtain

2.4.2 The advantages of using reciprocal expressions It is often advantageous to express the current in a reciprocal form as in Equations (31) and (33). Expression of the analytical solution in a reciprocal form has several advantages. First, it is often easy to identify the different limiting cases within the reciprocal equation. A limiting case is a simplified form of the expression which applies when one or other of the processes is much slower than all the others and is therefore the rate limiting step in the overall process. For example, we can see that the second term of Equation (33) describes the limiting case when all the enzyme present is in the form of the enzyme-substrate complex, so that the flux is limited by the maximum enzyme catalytic rate (kcateE). Looking at the third term we can see that it describes the case when all the enzyme is rapidly reduced by substrate such that there is only reduced enzyme present and the flux is limited by the rate of reaction between the enzyme and mediator (kge^. In contrast, when the first term is important, the 73

PHILIP N. BARTLETT AND CHEE-SENG TOH

flux appears to be limited by both enzyme kinetics and substrate diffusion. If the first term in Equation (33) is the dominant term, we can discard the second and third terms (this corresponds to the situation in which both the breakdown of the enzyme-substrate complex, ES, and reaction of the enzyme with the mediator are fast). Rearrangement of the first term in Equation (33) then gives:

which can be rearranged to give

Each of the three terms in Equation (35) is readily understood. The first term describes the enzyme kinetics at low substrate concentrations (i.e. sbuik < KM). The second term describes the diffusion limited flux of S through the membrane and the final term the diffusion limited flux of S across the Nernst diffusion layer. Using the same method, Equation (33) can be rearranged into the following form

Looking at Equation (36) we can see how mass transport, which determines the ratio Sbuik/Siayer, can have an influence on both the enzymatic and the mediator reaction rates. When mass transport of substrate across the Nernst diffusion layer and through the membrane is much faster than the enzyme kinetics, the substrate concentration within the film is maintained at the bulk value, Sbuik —siayer- When mass transport of substrate into the enzyme layer is much slower than the enzyme reaction, the concentration of substrate in the enzyme layer, Siayer, tends to zero. Thus, the second advantage of using reciprocal expressions is that we can better understand how changes in the experimental variables can bring about a change in the rate-limiting step from one process to another. Conversely, if we understand the limiting kinetic processes involved in the system, we can derive the reciprocal expression by inspection. The third advantage of using reciprocal expressions is that we can readily obtain meaningful data from the gradient and intercepts of a double reciprocal plot of 1/i against 1/Sbuik- However, since double reciprocal plots always over emphasize the data at low concentration at the expense of the high concentration data, a better alternative is to use nonlinear least squares fitting by using suitable curve fitting software.

2.4.3 Solving the coupled diffusion/reaction problem The solution for the membrane|enzyme]electrode problem by equating fluxes, as described above, starts by assuming that mass transport and the enzyme 74

KINETIC MODELING FOR BIOSENSORS

catalyzed reactions occur in spatially distinct regions. In fact, reaction and diffusion will often occur together within the same region of space. Under these circumstances we cannot proceed by equating the fluxes to obtain the steadystate solution but rather we must obtain solutions for the coupled diffusion reaction equations which describe the system. This kind of treatment is essential for electrodes in which the enzyme and mediator are immobilized together within the same film. Common examples in the literature include cross-linked redox hydrogels containing immobilized enzymes (12-16) and enzyme immobilized throughout films on electrode surfaces such as in cross-linked films (17,18) or electrochemically polymerized films (19-26). In order to illustrate the approach to solving this type of problem, we will present a much simplified example; more detailed treatments can be found in the literature. To simplify the problem we will begin by assuming that the mediator-enzyme reaction is very rapid, such that the enzyme is always in the oxidized state. We make the same assumption as we made above (Section 2.2). We also assume that the partition coefficient of substrate into the film is unity and that the effects of mass transport in the bulk solution are insignificant so that the concentration at the outside of the film is equal to the bulk concentration. The differential equation describing diffusion and enzyme catalyzed reaction within the enzyme membrane is obtained from Equation (20) by replacing ks(x, t) by the enzyme kinetic term,feCateEs/(^M+ s)- This gives:

In the steady state, dsjdt — 0, hence

Equation (38) is a nonlinear second-order differential equation which cannot be solved analytically. To overcome this problem, we can find approximate analytical solutions by making different limiting assumptions. The two obvious assumptions which we can make in order to simplify Equation (38) are either that the substrate concentration is much greater than KM (i.e. s > KM) or that the substrate concentration is much smaller than KM (i.e. s
75

PHILIP N. BARTLETT AND CHEE-SENG TOH Integration gives

where Aa is an integration constant. Applying the boundary condition that, since there is no reaction of the substrate at the electrode surface,

gives Aa = 0 and

For an enzyme membrane | electrode in which the mediator is trapped within the layer at the electrode surface, the current density, i, is directly related to the flux of substrate entering the membrane layer,

where 1 is the thickness of the enzyme membrane. Hence, we can write an expression for the steady-state current density at the enzyme membrane | electrode under conditions of high substrate concentration

This is the current density when the saturated enzyme kinetics are rate limiting and is identical to the corresponding expression obtained for the membrane] enzyme|electrode in Section 2.4.2. We can obtain the corresponding substrate concentration profile if we integrate Equation (42) and apply the boundary condition at the external solution enzyme membrane interface. Since we have assumed that the substrate concentration at the outside of the enzyme membrane is equal to the bulk concentration and there are no partition effects, we have:

Thus, we obtain:

In this case we find that the concentration profile is parabolic. Note that Equation (46) appears to imply that if 1 is large enough the concentration of S could be negative! This arises because we have assumed that s^KM. Thus, our approximate treatment is only valid if s > KM all the way through the membrane. 76

KINETIC MODELING FOR BIOSENSORS

We now turn to the other approximation. When the substrate concentration is much smaller than KM, the enzyme kinetic term reduces to (kcatCz/KivOs and Equation (38) becomes

This is a linear second-order differential equation for which we can write a general solution of the form: where A2 and B2 are integration constants and

is the kinetic length, that is, it is the distance that the reactant S can diffuse within the enzyme layer before its concentration is reduced to 1/e (36.79%) of its original value by reaction with the enzyme in the unsaturated enzyme kinetic regime. Differentiation of Equation (48) gives

Applying the same boundary condition as before, so that at x = 0, ds/dx = 0, we find that A2 — 0. Then applying the boundary condition for the concentration of substrate at the outside of the enzyme layer, that is, at x — I, s — sbuik, we find that

Thus, we obtain the following expressions for substrate concentration profile:

and substrate concentration gradient:

Again, the steady-state current density is given by

so that for this case

Equation (55) itself has two limiting forms. When J/Xk
PHILIP N. BARTLETT AND CHEE-SENG TOH

Figure 8 Normalized concentration profiles for the substrate calculated from Equations (46) and (52) for s>K M and s
Equation (55) becomes

which is the current density when the unsaturated enzyme reaction is rate limiting and is identical to the corresponding expression obtained for the membrane] enzyme|electrode in Section 2.4.2. When l/X k >l, tanh(l/Xk)^l and Equation (55) becomes

This is a new expression and corresponds to the situation where all of the substrate is consumed in a reaction layer at the outside of the enzyme membrane. A corresponding expression does not arise in the treatment of the membrane] enzyme|electrode because there we explicitly assume that there is no concentration variation for the substrate across the enzyme membrane layer. By plotting the concentration profiles described by Equations (46) and (52), we can see how the substrate concentration, s, changes with distance, x, as the ratio (kcate2/J
2.5 Deriving a complete kinetic model The preceding sections (2.1-2.4) give some idea of how to approach kinetic modeling through the use of reasonable assumptions and limiting cases. We have seen how steady-state analyses can be used to derive approximate analytical solutions. One can then obtain useful kinetic data from the system by analyzing 78

KINETIC MODELING FOR BIOSENSORS

the data either by constructing suitable straight line diagnostic plots or by curve fitting. However, we can go further. We can use this approximate analytical approach to generate solutions for all the possible limiting cases and then combine these into a case diagram which shows how these limiting cases are interrelated. A case diagram is really a map that shows how the behavior of the system changes as the different experimental parameters are varied and it can be used as a map to optimize the performance of a device for a specific application.

2.5.1 Case diagrams The final step in developing approximate analytical solutions is to present the whole model in the form of the case diagram. In this way, one can visually grasp and understand how the physical processes affect the performance of the biosensor. Thus, the case diagram is a highly condensed description of the behavior of the system. It must be remembered, when looking at the case diagram, that although the approximate analytical solutions are accurate for the different limiting cases, the solutions are much less accurate at the boundaries between the different cases. To illustrate this latter point, we will apply this approach to simple Michaelis-Menten kinetics for a reaction in homogeneous solution. This is a very simple situation and one for which an accurate analytical solution is available against which we can compare the expressions for the two limiting cases. Figure 9 shows a plot of the full Michaelis-Menten equation in the form of a one-dimensional case diagram. The full Michaelis-Menten equation is

Figure 9 Illustration of the one-dimensional case diagram for Michaelis-Menten enzyme kinetics. The solid line shows the full analytical solution, the two broken lines correspond to the two limiting cases.

79

PHILIP N. BARTLETT AND CHEE-SENG TOH

In case I, when s
and the reaction rate is first order with respect to the substrate. In case II, when s>K]v[> Equation (58) becomes

where the reaction rate is zero order with respect to the substrate concentration. At the boundary between the two cases, where s — KM, we find that although these two approximate solutions are equivalent (i.e. if we substitute s = KM in Equation (59) we obtain Equation (60)), the two approximate expressions do not agree with the full equation. On the other hand, away from the boundary the approximate expressions are very good approximations to the full equation. As a more complex example consider the enzyme membrane | electrode problem considered briefly in Section 2.4.3. Starting from the approximate analytical solutions we can derive a case diagram which describes the different solutions, this is shown in Figure 10. For this system there are four limiting cases. Three of these correspond to the simple limiting cases derived in Section 2.4.3 and given by Equations (44), (56), and (57). For thin films, when 1 < Xk, the reaction occurs uniformly throughout the enzyme layer. If s < KM (case I), the reaction is first order in S, if s > KM (case III), the enzyme is saturated and the reaction is zero order in S. For thicker films, when 1 > Xk, the enzyme-substrate reaction occurs in a thin layer at the outside surface of the enzyme layer and the concentration of substrate falls substantially as we go into the film. For low substrate concentrations, when s < KM (case II), the

Figure 10 Case diagram for the enzyme membrane|electrode problem showing the four cases with the corresponding concentration profiles. 80

KINETIC MODELING FOR BIOSENSORS

reaction is first order in S and so the concentration of S falls exponentially as we go into the enzyme layer. When s > KM, we find a new, fourth, case not discussed in Section 2.4.3. In this case (case IV), the reaction kinetics will be zero order in S at the outside of the film but as the concentration of S falls, it will change to first order at some point within the film. The corresponding concentration profiles are shown in Figure W. This type of model has been used by Bartlett et al. (27-29) to describe the oxidation of nicotinamide adenine dinucleotide (NADH) at polyfaniline) electrodes. Other examples of case diagrams and their applications can be found in the literature (14, 30, 31).

2.6 Experimental verification of approximate analytical kinetic models Deriving kinetic models without subsequent verification against real experimental data is a theoretical exercise of little value for real applications. Quite often, it is the comparison of the model to experimental data that helps to refine the model and to produce an improved understanding of the factors which determine the performance of the real biosensor. Before rushing ahead with experimental measurements, the researcher needs to think about the design of the experiments. Although one can derive a considerable amount of information from a single experiment if one already has a valid theoretical model, a single experiment can never be sufficient to "prove" or "disprove" the validity of a particular model. In order to do this, a systematic series of experiments must be carried out. This series of experiments should be designed to systematically test the dependence of the biosensor response on each experimental variable over as wide a range as feasible. That is, if factors such as enzyme loading, rotation speed, and film thickness are variables in the kinetic model, it will be necessary to carry out series of experiments to investigate the dependence of the biosensor output on each of these variables in turn. This immediately implies that this type of study can only be carried out on biosensors which can be reproducibly fabricated so that the results from one sensor can be compared quantitatively with those from another sensor. The approach toward experimental measurements which will allow us to establish a meaningful kinetic model is as follows. (a) Set up an initial model for the biosensor and define those experimental variables which are likely to affect the performance of the electrode (e.g. enzyme loading, substrate concentration, membrane thickness). (b) Choose a starting point—it is best, if possible, to start with conditions close to those for similar biosensors already described in the literature. If there is no appropriate equivalent, one has to be guided by the initial model and the results of preliminary experiments. (c) Compare the values from the first exploratory experiments with those derived from the initial model. Try to find out which step is rate limiting. 81

PHILIP N. BARTLETT AND CHEE-SENG TOH

If the initial model does not fit, it will need to be modified to include new factors or assumptions which were not considered in the initial model. Further exploratory experiments may then be required to explore the revised model. (d) Once one is happy that the model appears to be the correct one, carry out a detailed series of experiments systematically varying each experimental parameter. Try to vary each parameter over as wide a range as practicable so that all the different cases are explored. (e) Analyze the data using the model to produce a set of quantitative results for the model parameters (e.g. rate constants, mass transport coefficients, etc.). (f) Critically compare the quantitative results obtained from the model both from one experiment to another and with existing values available in the literature. Only when all the experimental data fits the model, when the quantitative values obtained from each series of experiments are consistent, and when the quantitative values for rate constants, diffusion coefficients etc. are reasonable in view of the published literature should one be satisfied that the model is reasonable. (g) Even at this stage one should only accept the validity of the model with caution. If there are other possible experimental ways to test the model these should be investigated if at all possible.

2.7 Numerical simulation methods So far we have concentrated on the use of approximate analytical solutions. A second, complementary, approach is to use numerical simulation techniques to model the processes involved in the system. Numerical techniques can provide accurate treatments for the behavior close to the boundaries between limiting cases. The disadvantage is that these numerical approaches do not provide as much insight into the behavior of the biosensor. Thus, the combined use of the two approaches is often more powerful than either on its own particularly for complex situations, for example, see ref. (14). Below, we will briefly describe the general principles behind the use of numerical simulation. A full discussion of the different techniques is beyond the scope of the present chapter. For further information, the reader should consult some of the more specialized texts (32).

2.7.1 Explicit numerical methods The first use of digital simulation in electrochemistry was based on the explicit finite difference method, pioneered by Feldberg (33). In many ways this is the simplest method to understand and to implement but it also suffers a number of limitations, particularly when one wishes to simulate diffusion and coupled kinetics. The first step in setting up a numerical simulation is to transform the problem into a discrete form. The explicit finite difference method treats the problem as an array of discrete "boxes" where the boxes are sufficiently small such that the concentration of species within each box is considered to be constant, see Figure 11. 82

KINETIC MODELING FOR BIOSENSORS

Figure 11 Schematic diagram showing the relationship between the change in concentration (s t+it -s t ), over a discrete time, St, and the net flux, (J\ + i — J\-i), across a discrete distance, x.

As an illustrative example of the method, we will consider a potential step at an electrode in a solution of the reduced form of an electroactive species, S, where the potential is stepped from a potential at which there is no reaction to one at which the oxidation of S is mass transport limited, this is the same situation considered in Sections 1.3.2 and 2. The potential step brings about a sharp change in the concentration near the electrode surface. If we divide the space in front of the electrode into a number of boxes then the concentration differences between the boxes create the driving force for diffusion of species into and out of the boxes. We can estimate the change in concentration over time within each box by calculating the net flux of species moving into or out of the box.

where c5sf is the change in concentration in the ith box over a time interval §t, A is the area of the electrode, V—ASx is the volume of the box, andjjinet the net flux into the box. Hence, for the box at a distance i$x from the electrode, where all the boxes are assumed to be of the same width, 8x, the change in concentration over time §t will be

83

PHILIP N. BARTLETT AND CHEE-SENG TOH

where Jj+1 andjj_! are the fluxes across the boundaries with the two adjacent boxes. Then using the discrete form of Pick's first law, we have

and

Thus, from Equation (62)

Rearranging gives

Equation (66) gives an explicit expression of the concentration in the ith box from the electrode surface at time step (t + §t) in terms of the concentrations in the ith box and the two boxes on either side of it at time t. In practice it is useful to recast Equation (66) in a dimensionless form by defining a dimensionless diffusion coefficient, DM such that

and introducing a dimensionless concentration, , defined as the ratio of the concentration to the bulk value (<j> — s/sbuik). Equation (66) then becomes

Thus by starting from the known concentration profile at time zero we can use Equation (68) to calculate the evolution of the concentration profile with time. This is the basis of the explicit finite difference method. Using the dimensionless formulation in Equation (68) has the advantage that we do not need to carry out a new simulation every time we change the bulk concentration or the substrate diffusion coefficient; rather, we can carry out the dimensionless simulation once and then calculate the specific results for particular concentrations and diffusion coefficients by appropriate scaling of the result. So far we have only considered the simulation of the diffusion part of the problem. In the explicit finite difference method, reaction kinetics can be included by adding a second calculation at each time step in which we calculate the change in concentration within each box as the result of the homogeneous kinetics. Full details can be found in the literature (32). A limitation of the explicit method is that the simulation is only stable for restricted values of the dimensionless diffusion coefficient, DM. For a 84

KINETIC MODELING FOR BIOSENSORS

one-dimensional simulation, the condition is that DM be less that 0.5. From Equation (67) we can see that this imposes a restriction on the relative sizes of the time step, 8t, and box size, 8x, that we can use. In turn this can be a severe limitation when we include reaction kinetics in our simulations. For this reason a number of more sophisticated simulation methods are often applied and we briefly introduce these below.

2.7.3 The Crank-Nicholson method Fully explicit methods, although simple to understand and to program, have the disadvantage that they are not unconditionally stable. The Crank-Nicholson method (34) uses a semi-implicit approach which calculates concentrations at time (t + §t/2) by linear interpolation between the known concentrations at time t and the unknown concentrations at time (t + §t). A comparison of this method to the fully explicit method described above is shown in Figure 12. As an illustration, consider three points describing the concentrations of a species at a particular point in space but at different times. Given an approximate value for the rate of change of the concentration of S in the ith box at time t, dSj(t)/dt, we can estimate the value of the concentration in the same box at time (t + St)

As we saw in the previous section (Equation (66)), dsf(t)/dt can be approximated from the discretized form of the second derivative (over space) in the diffusion equation. It is obvious from Figure 12 that this method suffers from poor accuracy when the concentration of S is changing rapidly, that is, when dsf(t)/dt is large. In

Figure 12 Schematic diagram showing the estimation of s,,t+itfrom s/,t using the explicit method, and the estimation of s/j+st from s/j+st/2 using the Crank-Nicholson (CN) method. 85

PHILIP N. BARTLETT AND CHEE-SENG TOH

the Crank-Nicholson method, the gradient at (t + 8t/2), that is, dsf(t + 8t/2)/dt, is used to estimate st(t + §t) from st(t) as follows:

As shown in Figure 12, this gives a more accurate estimate of st(t + §t) as compared to the fully explicit method. In the Crank-Nicholson method, the value of Sj(t + §t/2) is approximated by linear interpolation as follows:

so that Sj(t + c5t/2) is expressed in terms of st(t) and st(t + 8t) and is not calculated, but simply used as a point of reference to obtain a better estimation of st(t + 8t). As in the explicit method, c5t(dsf(t+c5t/2)/dt) is used to approximate for the second derivative (over space) in the diffusion equation, expressed in terms of both the known concentration, st(t) and unknown concentration, st(t + 8t), using the above expression for s4(t + <5t/2). Using this method, we obtain a set of n simultaneous equations and from the known boundary values (usually at i — 0 and i — n +1), the n unknown concentrations at (t + 8t) can be solved. In theory this method is stable for all values of DM.

2.7.4 Other simulation methods In addition to the two approaches described here, there are many other more sophisticated approaches that can be used in the digital simulation of electrochemical systems and that can help to improve the accuracy and efficiency of the simulation. In general, these improvements are achieved at the cost of additional mathematical complexity in setting up the simulation. For example, Joslin and Fletcher (35) introduced the idea of using unequal box sizes by using a transformation function for the distance variable. This improves the accuracy of the simulation, especially for systems with sharp changing concentration profiles, such as that near the electrode surface during a potential step experiment, by making the boxes smaller in the region where the concentration profile changes rapidly. Another method, known as the hopscotch method (36), provides a simple alternative to the Crank-Nicholson method. It explicitly calculates the unknown concentrations at time (t + 8t), from known concentrations at time t. From these new concentration values, it then implicitly derives the concentration values in every other box (e.g. for all the even values of i) for the same time step, (t + 8t). In the next time step, concentrations for the even i values are calculated explicitly and those for the odd i values implicitly. Thus, the calculation continues, alternately treating any given box explicitly and then implicitly. This approach is fairly easy to implement and stable for all values of DM but it does also have disadvantages (32). Another approach is to use polynomial curve fitting, as in the orthogonal collocation method (37, 38). This approach is different to the finite difference methods described above. It uses a trial polynomial function to describe the concentration profiles and exactly fits the trial polynomial to the concentration 86

Table 2 Summary of the literature on the kinetic modeling of enzyme electrodes Authors

Model

Expt. type

Real system

Method

Ref.

Albery, Kalia, and Magner, 1992

lla

SS

/

AS

(11)

Bartlett, Tebbutt, and Tyrrell, 1992

la

ss

/

AS

(20)

Bacha, Bergel, and Comtat, 1993

la

SS/T

X

DS

(52)

Calvo, Danilowicz, and Diaz, 1993

la

SS

/

AS

(53)

Marchesiello and Genies, 1993

la

SS

/

AS

(54)

Randriamahazaka and Nigretto, 1993

la

CV

X

DS

(55)

Schulmeister and Pfeiffer, 1993

lla (multi-layer)

SS

X

DS

(10)

Tatsuma and Watanabe, 1993

la

SS

X

DS

(56)

Lyons, Lyons, Fitzgerald, and Bartlett, 1994

la

T

/

AS

(23)

Battaglini and Calvo, 1994

la

SS

/

AS/DS

(57)

Martens and Hall, 1994

la

SS

X

DS

(58)

Sorochinskii and Kurganov, 1994

lip

SS

X

AS

(59)

Rhodes, Shults, and Updike, 1994

lla (multi-layer)

SS/T

DS

(60)

Albery, Driscoll, and Kalia, 1995

lla

SS

Bacha, Bergel and Comtat, 1995

la/Ha

SS/T

Gros and Bergel, 1995

la

SS

/ / / /

Bartlett and Pratt, 1995

la

SS

X

Martens, Hindle, and Hall, 1995

lla

SS

(8)

la

CV

DS

(17)

Chen and Tan, 1995

lla (multi-layer)

SS

AS

(61)

Lyons, Greer, Fitzgerald, Bannon, and Bartlett, 1996

la/lp

SS

/ / / /

AS

Kong, Liu, and Deng, 1995

AS

(30)

Cambiaso, Delfino, Grattarola, Verreschi, Ashworth, Maines, and Vadgama, 1996

lla (multi-layer)

SS/T

X

DS

(62)

Gooding and Hall, 1996

la (permeable electrode)

SS

/

DS

(63)

AS

(7)

DS

(9)

DS

(22)

AS/DS

(14)

Table 2 (Continued) Authors

Model

Expt. type

Real system

Method

Ref.

Jobst, Moser, and Urban, 1996

lla (multi-layer)

SS

X

DS

(64)

Desprez and Labbe, 1996

la

ss

X

AS

(65)

Sheppard, Mears, and Guiseppi-Elie, 1996

Ic

T

DS

(18)

Chen and Tan, 1996

lla (multi-layer)

T

/ /

DS

(66)

Krishnan, Atanasov, and Wilkins, 1996

la

SS

X

AS/DS(67)

Sorochinskii and Kurganov, 1997

lip

AS

(68)

la/H

ss ss

X

Somasundrum, Tongta, Tanticharoen, and Kirtikara, 1997

X

AS

(21)

Zhu and Wu, 1997

la

AS

(69)

la

ss ss

X

Gooding, Hall, and Hibbert, 1998

/

AS for thin film/DS for thick film

(70)

ss ss ss ss

/ /

DS

(71)

AS

(72)

X

AS

(73)

/

AS

(74)

Neykov and Georgiev, 1998

la

Gajovic, Warsinke, Huang, Schulmeister, and Scheller, 1999

la

Coche-Guerente, Desprez, Diard, and Labbe, 1999

la

Coche-Guerente, Desprez, Labbe, and Therias, 1999

la

(Votes Electrode types la/lp/lc: amperometric/potentiometric/conductimetric enzyme-membrane [electrode, lla/llp: amperometric/potentiometric membrane|enzyme|electrode, H: Homogeneous case, amperometric only. Experiment types T: Transient response, SS: Steady state, CV: Cyclic voltammetry. Real system: Indicates experimental results from an actual enzyme electrode were compared with the simulation. Simulation methods DS: Digital simulation, AS: Approximate analytical solution.

KINETIC MODELING FOR BIOSENSORS

at a number of carefully selected points. These points are selected according to polynomial theory so that they minimize the errors between the polynomial and the actual concentration profile. Although this method is technically more demanding as it requires sophisticated integration tools, it is however, very efficient in terms of computing time.

3 Applications In Table 2 we have compiled the literature on digital simulation and approximate analytical models as applied to enzyme electrodes from 1992 to the present. For a survey of earlier literature, one may refer to the paper by Bartlett and Pratt (39). Table 2 also includes information on the electrode configuration (membrane | enzyme|electrodes, enzyme membrane|electrodes, or homogeneous kinetics) and experiment type (steady state, transient, or cyclic voltammetry). By a membrane|enzyme|electrode, we mean the situation in which the enzyme is entrapped behind a membrane at the electrode surface. By a enzyme membrane | electrodes we mean the situation in which the enzyme is entrapped within the membrane at the electrode surface. Homogeneous kinetics refers to the situation where both the enzyme and mediator are free to diffuse in the bulk solution. Different information can be obtained from each of these configurations and different modeling approaches are employed. For example, from electrochemical measurements on homogeneous solutions, we can obtain information on the enzyme-substrate kinetics or the enzyme-mediator kinetics, depending upon which is the slower step. In the membrane|enzyme]electrode configuration, mass transport and enzyme catalyzed reaction can often be considered to occur in different regions of space provided the enzyme layer is thin enough. As a result, the problem is less complicated as compared to the enzyme membrane |electrode situation. Of these three types of electrode configurations, the enzyme membrane | electrode is currently the most popular.

4 Kinetic modeling in other types of biosensors The discussion above has concentrated heavily on amperometric enzyme electrodes. This is because these are at present the most widely modeled type of biosensors. Nevertheless, the kinetic modeling methods discussed above can be applied generally to other types of biosensors and in this final section, we will consider some of these. The main difference between amperometric enzyme electrodes and other types of biosensors is in the type of transducer employed and the signal measured. In addition to amperometric measurement, biosensors utilizing potentiometric, conductimetric, capacitative, optical, photometric, thermal, and mass transduction have all been described in the literature. Figure 13 shows the types of kinetic process involved in the response of a generalized biosensor.

89

PHILIP N. BARTLETT AND CHEE-SENG TOH

Figure 13 Schematic diagram of a generalized biosensor showing the various kinetic steps involved in its response toward a substrate.

4.1 Potentiometric enzyme electrodes In a potentiometric enzyme electrode, the transducer measures the interfacial potential established at the electrode surface by reactants or products of the enzyme catalyzed reaction. In many ways the situation is similar to that of an amperometric enzyme electrode. The substrate is converted into products at, or close to, the enzyme electrode surface, hence we still need to consider the interplay of mass transport and enzyme kinetics. The crucial difference is that there is no net electrochemical reaction so the boundary conditions at the electrode surface are different (now ds/dx is zero at the electrode surface) and the electrode potential is related to the surface concentrations through the Nernst equation. Thus, the output potential of the electrode is related to the logarithm of the steady-state concentration of the species being detected by the electrode. Once again both approximate analytical approaches and numerical methods can be used to model this type of biosensor.

4.2 Optical and photometric biosensors Optical and photometric transducer-types have been widely employed in biosensors because of their high sensitivity and potential simplicity, especially when configured to use fiber optics. Optical measurements can be based on changes in refractive index, fluorescence, chemiluminescence, or absorbance. Optical biosensors frequently make use of a dye indicator or a dye indicator-substrate complex. For example, using the integral form of Beer-Lambert law for the absorbance of dye indicator molecules, the transducer output is linearly related to the concentrations of the indicator and indicator-substrate complex

90

KINETIC MODELING FOR BIOSENSORS

integrated over the path length, I, of the sensor

where gln and glns are the molar extinction coefficients of the free and complexed indicator dye, respectively, and Aft) is the absorbance at time t. The success of these techniques depends on the ability of the dye indicator to bind specifically and reversibly to the analyte molecule: If we assume that the reaction between the substrate and the dye is fast, the reaction between the dye indicator and the analyte can be described by an equilibrium constant:

Then expressing the concentrations of indicator and indicator-substrate complex in terms of substrate concentration, and substituting into equation (72), a time-dependent expression relating substrate concentration to absorbance can be derived for a flow-cell detection system (40). Note that analyses of optical biosensor responses in the literature seldom take into consideration the mass transport of substrate in the membrane or the bulk solution (41-43). Apparent Michaelis constants can be obtained by plotting the initial rate of change of absorbance against substrate concentration (43).

4.3 Immunosensors A popular use of optical transduction is in the area of immunosensors using fluorescently labeled antibodies. When the antibody binds to the antigen, the environment of the fluorescent label changes, this change in the physicochemical environment of the label causes a corresponding change in the fluorescence emission. Both steady-state and time-resolved methods have been used for analytical detection (44), as well as to determine the kinetics of the antibody-antigen binding reaction (45). If the labeled antibody is immobilized at the sensor surface, or within a membrane, modeling the response will require a consideration of mass transport of the antigen as well as the binding kinetics. Another optical technique which can be used to study the binding interactions of biomolecules is the surface plasmon resonance (SPR) technique. In this case changes in the thickness, or refractive index, of a thin sensing film deposited upon a thin metal film in which surface plasmons are excited, causes a shift in the angular position of the reflected intensity minimum. The SPR technique is very sensitive to small changes in the optical properties at the metal surface. Thus, the technique can be used to determine the amount of analyte bound to biorecognition elements immobilized on the sensor surface (46-48). Other types of transducers used in immunosensors include capacitance (49, 50) and impedance measurements (51). In all of these immunosensors, mass transport of the analyte in the membrane or the bulk solution is generally ignored because the biorecognition element (in 91

this case an antibody) is often present as a monolayer with no overlayer. In addition, the binding reaction is often much slower than mass transport (e.g. it takes 35 min for equilibrium to be reached in the formation of the human serum albumin antigen-antibody complex (44)).

5 Conclusions In this chapter we have given an introduction to the modeling of biosensors, concentrating particularly on amperometric biosensors. The key part played by the interplay of mass transport and nonlinear kinetics has been illustrated. This interplay leads to a richness and complexity in the behavior of these systems which, with the aid of modeling, can be understood and thus controlled.

List of symbols Symbol

Meaning

Units

A

Electrode area

cm2

A(t)

Absorbance at time t

—

A±;A2

Integration constants

—

B2

Integration constant

—

D

Diffusion coefficient

cm 2 s

DM

Dimensionless diffusion coefficient

—

Ds, Dp, or Dp/

Diffusion coefficient of S, P, or P', respectively

cm 2 s

E

Enzyme

—

Eox

Enzyme in the oxidized state

—

Ered

Enzyme in the reduced state

—

ES

Enzyme-substrate complex

—

e

Total concentration of enzyme

mol dm ~ 3

eEs

Concentration of enzyme-substrate complex

mol dm ~ 3

eox

Concentration of oxidized enzyme

mol dm ~ 3

e

Concentration of reduced enzyme

mol dm ~ 3

In

Indicator dye

—

InS

Indicator dye/substrate complex

—

/

Current density

Acm~2

J

Flux

mol cm" 2 s" 1

J/,in

Flux into the fth box (for digital simulation)

mol cm" 2 s" 1

J/.net

Net flux into or out of the fth box (for digital simulation)

mol cm" 2 s" 1

-//.out

Flux out of the fth box (for digital simulation)

mol cm" 2 s" 1

JSS

Steady-state flux

mol cm" 2 s" 1

Ks

Partition coefficient for S intomembrane

—

Kp

Partition coefficient for P into membrane

—

Keq

Equilibrium constant for the reaction

—

E

92

red

1

1

Symbol

Meaning

Units

KM

Michaelis constant

mol dm" 3

K|vl,app

Apparent Michaelis constant obtained from kinetic analyses of biosensor

mol dm" 3

k

First order rate constant for conversion of S to P

s- 1

k±

Rate constant of forward step in an enzyme reaction

mol" 1 drr^s"1

k-i.

Rate constant of backward step in an enzyme reaction

mor1 drrr's"1

'feat

Rate constant for breakdown of ES to yield E and P

s- 1

Mass transfer rate constant of product and substrate in membrane, respectively

cms" 1

kt

Second-order rate constant for the reaction between the enzyme and mediator

mor1 drrr's"1

k'E

Pseudo first-order rate constant for the reaction between s- 1 the enzyme and mediator

k'

Heterogeneous rate constant for the oxidation of P to P'

cms" 1

k'of, k'D:S

Mass transfer rate constant of product and substrate in solution, respectively

cms" 1

L

film thickness

cm

M

Redox mediator

—

n k'

IS mem,s mem,P>• n

N

Number of electrons per molecule oxidized or reduced

—

P

Product of enzyme reaction

—

P'

Product of electrochemical reaction

—

P

Concentration of product

mol dm" 3

P'

Concentration of electrochemical product

mol dm" 3

Po

Product concentration at the outside surface of membrane

mol dm" 3

Bulk concentration of product

mol dm" 3

Product concentration within the enzyme layer Product concentration at the membrane enzyme layer interface

mol dm" 3 mol dm" 3

Product concentration at the external solution membrane interface

mol dm" 3

s

Substrate

—

s

Concentration of S

mol dm" 3

s,

Concentration in the fth box

mol dm" 3

So

Concentration of S at the outside surface of membrane

mol dm" 3

s

Bulk concentration of substrate

mol dm" 3

Slayer

Substrate concentration within the enzyme layer

mol dm" 3

Substrate concentration at the membrane enzyme layer interface

mol dm" 3

Substrate concentration at the external solution membrane interface

mol dm" 3

t

Time

s

V

Volume

cm3

Pbulk Player Pmemjn

Pmem,out

bulk

Smemjn

Smem,out

PHILIP N. BARTLETT AND CHEE-SENG TOH Symbol

Meaning

Units

^max

Maximum rate of enzyme reaction at saturating substrate concentration

mol dm 3 s

V

Rate of enzyme reaction

mol dm" 3 s" 1

1

W

Rotation speed of rotating disk electrode

Hz

XD

Diffusion layer thickness

cm

X*

Kinetic length

cm

X

Distance from planar electrode

cm

£

Molar extinction coefficient

mol" 1 dm3 cm" 1

V

Kinematic viscosity

crr^s- 1



Dimensionless concentration

—

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25.

94

Michaelis, L. and Menten, M. L. (1913). Biochem. Z., 49, 333. Briggs, G. E. and Haldane, J. B. S. (1925). Biochem. ]., 19, 338. Cornish-Bowden, A. (ed.) (1976). Principles of enzyme kinetics. Butterworths, London. Dixon, M. and Webb, E. C. (ed.) (1979). Enzymes, 3rd edn. Longman, London. Bard, A. J. and Faulkner, L. R. (ed.) (2001). Electrochemical methods. Fundamentals and applications, 2nd edn. Wiley, New York. Thompson, R. Q., Porter, M., Stuver, C., Halsall, H. B., Heineman, W. R., Buckley, E., and Smyth, M. R. (1993). Anal. Chim. Ada, 271, 223. Albery, W. J., Driscoll, B. J., and Kalia, Y. N. (1995). J. Electroanal. Chem., 399, 13. Martens, N., Kindle, A., and Hall, E. A. H. (1995). Biosens. Bioelectron., 10, 393. Bacha, S., Bergel, A., and Comtat, M. (1995). Anal. Chem., 67, 1669. Schulmeister, T. and Pfeiffer, D. (1993). Biosens. Bioelectron., 8, 75. Albery, W. J., Kalia, Y. N., and Magner, E. (1992). ]. Electroanal. Chem., 325, 83. Rajagopalan, R., Aoki, A., and Heller, A. (1996). J. Fhys. Chem., 100, 3719. Surridge, N. A., Diebold, E. R., Chang, J., and Neudeck, G. W. (1994). ACS Symposium Series, 556, 47. Bartlett, P. N. and Pratt, K. F. E. (1995). J. Electroanal. Chem., 397, 61. Niculescu, M., Nistor, C., Frebort, L, Pec, P., Mattiasson, B., and Csoregi, E. (2000). Anal. Chem., 72, 1591. Linke, B., Kerner, W., Kiwit, M., Pishko, M., and Heller, A. (1994). Biosens. Bioelectron., 9, 151. Kong, J. L., Liu, H. Y., and Deng, J. Q. (1995). Anal. Lett., 28, 1339. Sheppard, N. F., Mears, D. J., and GuiseppiElie, A. (1996). Biosens. Bioelectron., 11, 967. Bartlett, P. N., All, Z., and Eastwickfleld, V. (1992). J. Chem. Soc., Faraday Trans., 88, 2677. Bartlett, P. N., Tebbutt, P., and Tyrrell, C. H. (1992). Anal Chem., 64, 138. Somasundrum, M., Tongta, A., Tanticharoen, M., and Kirtikara, K. (1997). J. Electroanal. Chem., 440, 259. Gros, P. and Bergel, A. (1995). J. Electroanal. Chem., 386, 65. Lyons, M. E. G., Lyons, C. H., Fitzgerald, C., and Bartlett, P. N. (1994). J. Electroanal. Chem., 365, 29. Besombes, J. L, Cosnier, S., and Labbe, P. (1996). Talanta, 43, 1615. Nakaminami, T., Ito, S., Kuwabata, S., andYoneyama, H. (1999). Anal. Chem., 71, 1928.

KINETIC MODELING FOR BIOSENSORS 26. Wu, Q., Storrier, G. D., Pariente, F., Wang, Y., Shapleigh, J. P., and Abruna, H. D. (1997). Anal. Chem., 69, 4856. 27. Bartlett, P. N. and Wallace, E. K. N. (2000). J. Electroanal. Chem., 486, 23. 28. Bartlett, P. N. and Simon, E. (2000). Phys. Chem. Chem. Phys., 2, 2599. 29. Bartlett, P. N., Birkin, P. R., and Wallace, E. N. K. (1997). J. Chem. Soc., Faraday Trans., 93, 1951. 30. Lyons, M. E. G., Greer, J. C., Fitzgerald, C. A., Bannon, T., and Bartlett, P. N. (1996). Analyst, 121, 715. 31. Albery, W. J., Bartlett, P. N., Driscoll, B. J., and Lennox, R. B. (1992). J. Electroanal. Chem., 323, 77. 32. Britz, D. (ed.) (1988). Digital simulation in electrochemistry, 2nd edn. Springer-Verlag, New York. 33. Feldberg, S. W. (1969). In Ekclroanalylical chemistry (ed. A. J. Bard), Vol. 3, p. 199. Marcel Dekker Inc., New York. 34. Crank, J. and Nicolson, P. (1947). Proc. Cambridge Phil. Soc., 43, 50. 35. Joslin, T. and Fletcher, D. (1974). J. Electroanal. Chem., 49, 171. 36. Gourlay, A. R. (1970). ]. Inst. Maths. Appl, 6, 375. 37. Whiting, L. F. and Carr, P. W. (1977). J. Electroanal. Chem., 81, 1. 38. Villadsen, J. V. and Stewart, W. E. (1967). Chem. Eng. Set, 22, 1483. 39. Bartlett, P. N. and Pratt, K. F. E. (1993). Biosens. Bioekclron., 8, 451. 40. Hisamoto, H., Tsubuku, M., Enomoto, T., Watanabe, K., Kawaguchi, H., Koike, Y., and Suzuki, K. (1996). Anal. Chem., 68, 3871. 41. Dremel, B. A. A., Schmid, R. D., and Wolfbeis, O. S. (1991). Anal. Chim. Acta, 248, 351. 42. Kar, S. and Arnold, M. A. (1992). Anal. Chem., 64, 2438. 43. Yerian, T. D., Christian, G. D., and Ruzicka, J. (1988). Anal. Chim. Acta, 204, 7. 44. Bright, F. V., Betts, T. A., and Litwiler, K. S. (1990). Anal. Chem., 62, 1065. 45. Ingersoll, C. M. and Bright, F. V. (1997). Anal. Chem., 69, 403A. 46. Jung, L. S., Shumaker-Parry, J. S., Campbell, C. T., Yee, S. S., and Gelb, M. H. (2000). J. Am. Chem. Soc., 122, 4177. 47. Nikitin, P. L, Beloglazov, A. A., Kochergin, V. E., Valeiko, M. V., and Ksensevich, T. I. (1999). Sens. Actual., B, 34, 43. 48. Kabashin, A. V., Kochergin, V. E., and Nikitin, P. I. (1999). Sens. Actual. B, 54, 51. 49. Bataillard, P., Gardies, F., Jaffrezic-Renault, N., Martelet, C., Colin, B., and Mandrand, B. (1988). Anal. Chem., 60, 2374. 50. Gebbert, A., Alvarez-Icaza, M., Stocklein, W., and Schmid, R. D. (1992). Anal. Chem., 64, 997. 51. Souteyrand, E., Martin, J. R., and Martelet, C. (1994). Sens. Actual. B, 20, 63. 52. Bacha, S., Bergel, A., and Comtat, M. (1993). J. Electroanal. Chem., 359, 21. 53. Calvo, E. J., Danilowicz, C., and Diaz, L. (1993). J. Chem. Soc., Faraday Trans., 89, 377. 54. Marchesiello, M. and Genies, E. (1993). J. Ekclroanal. Chem., 358, 35. 55. Randriamahazaka, H. and Nigretto, J. M. (1993). Electroanalysis, 5, 221. 56. Tatsuma, T. and Watanabe, T. (1993). Anal. Chem., 65, 3129. 57. Battaglini, F. and Calvo, E. J. (1994). J. Chem. Soc., Faraday Trans., 90, 987. 58. Martens, N. and Hall, E. A. H. (1994). Anal. Chem., 66, 2763. 59. Sorochinskii, V. V. and Kurganov, B. L. (1994). Biosens. Bioekctron., 9, 481. 60. Rhodes, R. K., Shults, M. C., and Updike, S. J. (1994). Anal. Chem., 66, 1520. 61. Chen, Y. and Tan, T. C. (1995). AicheJ., 41, 1025. 62. Cambiaso, A., Delflno, L, Grattarola, M., Verreschi, G., Ashworth, D., Maines, A., and Vadgama, P. (1996). Sens. Actual. B, 33, 203. 63. Gooding, J. J. and Hall, E. A. H. (1996). ]. Eleclroanal. Chem., 417, 25.

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64. 65. 66. 67. 68. 69. 70. 71. 72.

96

Chapter 5 Bio-, chemi-, and electrochemiluminescence for fiber-optic biosensors LoTc J. Blum and Pierre R. Coulet Laboratoire de Genie Enzymatique, Universite Claude Bernard, Lyon I, 69622 Villeurbanne Cedex, France.

1 Introduction For many years, our laboratory has been involved in the development of biosensors first based on electrochemical transduction (1). One of the key components in the design of a biosensor is the sensing layer associated with the transducer and involving a biological element which confers molecular recognition properties to the analytical system. Among the several types of transducing modes which have been proposed (2), optical systems, namely those including fiber-optics, appear particularly promising (3). In the present chapter, we will focus on optical biosensors involving bio-, chemi-, and electrochemiluminescence (ECL). In luminescence reactions, light is emitted when specific molecules involved in the reaction return to the ground state after having been excited in the course of the biochemical or chemical reaction. The emission of light is triggered by the molecular recognition of the analyte by the biological element, generally an enzyme. This light is transmitted through a waveguide, here a fiber-optic, to a light detector, here the photomultiplier tube (PMT) of a luminometer. The principle of luminescent sensors is illustrated in Figure 1.

2 Design of the biosensor 2.1 Optical waveguide Two types of fiber-optics were used as waveguides.

2.1.1 Glass fiber-optic bundle The fiber-optic system was a 1-m long bundle of glass fibers with a diameter of 8 mm. A screw cap maintains the enzymatic disk in close contact with the end 97

LOIC J. BLUM AND P I E R R E R. COULET

Figure 1 Principle of a luminescent sensor. LER: Light-emitting reaction, PMT: photomultiplier tube.

Figure 2 Sensing tip of the fiber-optic biosensor.

of the bundle (Figure 2). The emitted light is conducted from the enzymatic tip through the fiber-optic to the (PMT) of a luminometer. This connecting end was designed to fit exactly into the light-tight chamber of the luminometer (4).

2.1.2 Liquid core fiber-optic A liquid core fiber-optic from L. O. T. Oriel (Courtaboeuf, France) can be used, core diameter 5 mm, overall diameter 7 mm, connected to the PMT of a luminometer (5).

2.2 Setup Either type of fiber-optic can be associated with an enzymic sensing layer to constitute the biosensor. The fiber-optic end bearing this layer can be either immersed in a batch reaction vessel or connected to a flow cell for Flow Injection Analysis measurements (FLA). The other end is connected to the luminometer.

2.2.1 Measurements in a batch system The biosensor is immersed in a 3.5-ml stirred and thermostated reaction medium. Samples were introduced with a syringe through a septum in the reaction vessel protected from ambient light by a black PVC jacket. Temperatures of 23 °C, 25 °C, or 30 °C were selected depending of the type of assays used (Figure 3).

2.2.2

Measurements in a flow system

The flow system consists of one-channel peristaltic pumps (model P-l, Pharmacia) an injection valve (model 5020, Rheodyne) on which a 10 or 30 (il sample loop was fitted, depending on the study, and a specially designed flow cell with a 125 or 250 jol inner volume containing a magnetic bar, 4 or 7 mm in diameter (Figure 4(a) and (b)). 98

Figure 3. Jacketed reaction cell for batch measurements, (a) Fiber-optic bundle, (b) 0-ring (c) thermostated jacket, (d) screw cap, (e) reaction medium, (f) light-tight PVC jacket, (g) septum, (h) syringe needle guide, (i) enzyme membrane, and (j) stirring bar.

Figure 4 Flow system setup. (A) Detail of the flow cell, (a) liquid core fiber-optic, (b) plexiglass window, (c) enzyme membrane, (d) light-tight flow cell, (e) stirring bar. (B) Overall setup. PP1, PP2: peristaltic pumps, FO: fiber-optic; FC: flow cell, W: waste, IV: injection valve.

LOIC J. BLUM AND P I E R R E R. COULET

Figure 5 Flow system for ECL measurements. GCE: glassy carbon electrode, SL: sensing layer, MB: magnetic bar, Pt: platinum electrode, FC: flow cell, FO: fiber-optic.

For electrogenerated chemiluminescence, the cell was modified enabling a glassy carbon (BAS, USA) 3 mm in diameter to be adapted (6). The glassy carbon electrode was polarized against a pseudo reference platinum electrode connected to a PRG-5 polarograph (Tacussel-Radiometer Analytical) (Figure 5).

2.3 Light-emitting reactions Bioluminescence is a special form of chemiluminescence occurring in numerous kinds of living organisms and depending on specific protein systems, generally enzymatic, which can be isolated and used in the design of fiber-optic biosensors (7). Bioluminescence reactions involving mainly ATP and NAD(P)H are extremely sensitive leading to very powerful analytical devices.

2.3.1 Bioluminescence (a) ATP can be detected with luciferase (EC 1.13.12.7) from the firefly Plwtinus pyralis in the presence of the substrate luciferin, Mg2+, and molecular oxygen.

(b) NAD(P)H can be determined using the bioluminescent marine bacteria bi-enzyme system from Vibrio fischeri or Vibrio harveyi. The first enzyme oxidoreductase (EC 1.6.8.1) catalyzes the oxidation of NAD(P)H into NAD(P)+ with the concomitant reduction of FMN into FMNH2 which serves as a substrate for the bacterial luciferase (EC 1.14.14.3) with oxygen and a long chain aldehyde (R-CHO) as cosubstrates. 100

LUMINESCENCE AND FIBER-OPTIC BIOSENSORS

2.3.2 Chemiluminescence The most common Chemiluminescence reaction uses luminol (5-amino-2,3-dihydro-l,4-phthalazinedione) in alkaline conditions and in the presence of horseradish peroxidase (HRP, EC 1.11.1.7). Some related hydrazides can also be used instead of luminol.

2.3.3 Electrochemiluminescence Hydrogen peroxide can be detected without the need of peroxidase. The luminol ECL generated by a glassy carbon electrode polarized at + 425 mV vs a Platinum pseudo reference electrode can be used.

2.3.4 Extension of the analytical properties Suitable auxiliary enzymes such as dehydrogenases can be coupled to the light emitting enzyme reactions allowing their extension to numerous compounds for possible detection through the fiber-optic biosensor.

As dehydrogenases can be used in combination with the light-emitting bi-enzyme system of marine bacteria, so also oxidases leading to the production of H2O2 can be associated with the different types of chemiluminescent detection, allowing the oxidase substrates to be easily quantified.

Examples of determinations with dehydrogenases and oxidases will be given later in this chapter.

2.4 Preparation of the sensing layer Several types of polymeric supports have been used for preparing the sensing layer, either from natural origin like collagen, or synthetic-like derivatized polyamide or polyvinylalcohol.

2.4.1 Immobilization techniques Note that in the different protocols that will be described, several activation steps are chemical reactions that must be conducted carefully in the fume hood. 101

LOi'C J. BLUM AND PIERRE R. COULET

Enzyme Immobilization on collagen membranes Originally developed in our laboratory more than two decades ago (8, 9) a mild method of activation of collagen films produced locally at that time, based on the acyl-azide formation from available carboxyl groups is still currently used for preparing highly stable enzyme layers. Briefly, the carboxyl groups were first esterified in acidic methanol, then immersed in a hydrazine aqueous solution leading to the formation of hydrazides and finally treated with nitrous acid for the subsequent formation of acyl-azides ready for coupling through available amino groups of the enzyme.

Protocol 1

Enzyme immobilization on collagen membranes 1 Cut out from commercial collagen films,a disks of the required diameter matching the diameter of the fiber-optic end. Note: The following steps must be carried out in a fume hood.

2 Immerse the disks in 60 ml methanol containing 0.2M HC1 for 3-5 days at room temperature (20 °C) in a flask closed with parafilm. Note that a longer time treatment may damage the disks. 3 Treat the disks overnight with 1% aqueous hydrazine solution at room temperature. 4 Soak the disks for 20 s in 0.5M NaNO2/0.3M HC1 at 4 °C. 5 Mop up rapidly the excess of reaction medium with filter paper and incubate the disks at 4 °C for 3 h in the enzyme solution (generally at a concentration between 0.5 and 10 mg/ml depending on the specific activity) in Phosphate Buffered Saline (PBS) (10 mM phosphate buffer, 0.14M NaCl, 1 mM KC1, pH 7.4). 6 Wash the disks extensively for 30 min in 0.1M phosphate buffer, pH 7 containing 1M KC1. And then for 10 min in PBS. 7 Store the enzymatic disks in PBS at 4°C (other storage conditions are possible according to the type of enzymes, either at — 20 °C in 20% glycerol-PBS or in a dried state). a

Possible supplier: ICN for Cellagen™ membranes, USA.

Enzyme immobilization on preactivated poly amide membranes The use of commercially available preactivated nylon membranes prepared by Pall Co. for immunochemical applications had been originally introduced by our group in the design of biosensors (10). These membranes exhibit a high density of covalent binding sites able to react with -OH, -COOH, or -NH2 groups available at the enzyme surface. New preparations from this supplier or from other sources are now available and can be utilized using a similar binding protocol. 102

LUMINESCENCE AND FIBER-OPTIC BIOSENSORS

Protocol 2

Enzyme immobilization on membranes Immunodyne ABC type from Pall-Gelman 1

Cut out from Immunodyne ABC type membranes disks of the required diameter matching the diameter of the fiber-optic. 2 Apply dropwise on each side of the disk 10 (il of a suitable enzyme solution (typically at a concentration of 10 mg/ml, which can be adjusted depending on the availability and activity) in 0.1M phosphate buffer, pH 6. 3 After 5 min at room temperature, wash the disks successively for 10 min in 0.1 M phosphate buffer, pH 6, then for 20 min in 0.1 M phosphate buffer, pH 7 containing 1 M KC1. 4 Store the enzymatic disks in PBS at 4°C (other storage conditions are possible according to the type of enzymes, either at — 20 °C in 20% glycerol-PBS or in a dried state).

Protocol 3

Enzyme immobilization on Ultra Bind type membranes 1 2

3 4 5

Cut out from Ultra Bind type membranes, disks of the required diameter matching the diameter of the fiber-optic. Immerse the disks for 10 s in an enzyme solution (at a concentration of 10 mg/ml, which can be adjusted depending on the application) in 0.1 M phosphate buffer, pH6. Withdraw the impregnated disks and let them dry for 15 min at room temperature. Wash the disks as indicated above for the ABC type membrane Store the enzymatic disks in PBS at 4°C (other storage conditions are possible according to the type of enzymes, either at — 20 °C in 20% glycerol-PBS or in a dried state).

Immobilization of "His-tag" enzymes Many enzymes expressed in microorganisms can be genetically engineered by adding Histidine residues (His-tag) to their C- or N-terminal residues, in order to facilitate their purification. Glycoproteins can be also modified chemically and histidine residues grafted onto the carbohydrate part of the molecule after periodate oxidation. As an example, HRP and glucose oxidase (Gox) have been modified following this procedure. Related work must be mentioned here dealing either with preparation of enzyme-IgG conjugates after periodate oxidation (11) or enzyme immobilization on metal-chelate carriers (12,13). Advantage can be taken from this type of modification to immobilize the derivatized enzyme 103

LOIC J. BLUM AND PIERRE R. COULET

directly on metal-chelate supports which can be embedded in a reticulated sensing layer adaptable to the fiber-optic biosensor. As an example, peroxidase was chemically modified with histidine, then immobilized through site directed adsorption on iminodiacetate (IDA) sepharose beads which were entrapped in polyfvinylalcohol) bearing styrylpyridinium groups (PVASbQ) photocross-linked, then forming the sensing layer. This entrapment method is derived from a previous one implying the direct coating of the sensitive tip of biosensors with PVA-SbQ through a controllable step-procedure (14).

Protocol 4

Chemical modification of HRP with His-tag Note: It is recommended that the reactions are performed in a fume hood. 1 2 3 4 5

Dissolve 5 mg of HRP in 200 jol of distilled water and 80 jol of a 0.1 M NaIO4 solution. Mix and stir for 20 min. Add 40 |ol of 0.2 M L-histidine in 0.1 M carbonate buffer, pH 9 to the activated peroxidase solution and make up with the same buffer to 800 |ol Incubate at room temperature under stirring or shaking for 2 h. Add 20 (il of a freshly prepared 0.1 M NaBH4 solution in water, mix and keep at room temperature for 15 min. Separate HRP-His from low molecular weight species by desalting on a Sephadex G-25 M column.

Protocol Protocol 5

Enzyme loading of IDA sepharose beads 1

Equilibrate 500 [d of IDA-sepharose beads with 30 mM sodium diethyl barbiturateHC1 buffer, 30 mM KC1, 1.5 mM MgCl2, pH 8.5 (veronal buffer). 2 Filter and pack a 0.5-ml plastic column with the beads and perfuse with a 100 mM NiCl2 solution in veronal buffer. 3 Wash with veronal buffer and introduce the HRP-His preparation (Protocol 4) into the column. 4 Wash the column with veronal buffer and store it at 4°C. The enzyme bound sepharose beads in the column are ready for subsequent use.

Protocol 6

Preparation of the sensing layer with a photocross-linkable polymer (PVA-SbQ) 1

104

Prepare in distilled water a small volume (0.5-1 ml) of a diluted solution (1:10 v/v) of PVA-SbQ_from Toyo Gosei Kogyo (polymerization degree 2300; SbQ_ content 1.06%).

LUMINESCENCE AND FIBER-OPTIC BIOSENSORS

Protocol 6 continued 2 Mix 1.8 mg of wet HRP-IDA sepharose beads (Protocol 5) with 250 |ol of the PVA-SbQ. solution. 3 Spread 20-50 jol of this mixture depending on the disk diameter, on a plexiglass transparent disk, the diameter of which matches the end of the fiber-optic. It is recommended to prepare several disks at the same time. 4 Expose the disks under a tungsten light for 90 min followed by ultraviolet (UV) light at 254 nm for 30 min to ensure polymerization. 5 Store the disks in 30 mM veronal buffer.

3 Examples of determinations with the

luminescence sensors

To illustrate the various possibilities of the luminescence sensors, a selection of results is given. For ATP, NADH, and H2O2 determination, the enzymatic systems described above have been used with different types of sensing layers. Extension to other analytes has been explored using auxiliary enzymes. The various types of enzymic membranes and luminescent detections either in a batch mode or with flow systems have been applied.

3.1 ATP determination With the immobilized firefly luciferase on preactivated polyamide membranes, ATP could be assayed over a wide dynamic range, from 2 x 10 ~ al M to 1 x 10 ~6M with the batch system. The measurements were performed in 0.05 M Tris-acetate buffer at pH 7.75, containing 30 mM MgCl2, 2 mM dithiothreitol (DTT), and 0.2 mM luciferin. The time necessary to reach a steady-state response depended on the ATP concentration and was about 1 min for the lowest concentration.

3.2 NADH determination Using the bacterial oxidoreductase-luciferase system bound to preactivated polyamide membranes, the linear dynamic range was between l x l O ~ 9 M and 3 x l O ~ 6 M with a detection limit of 3xlO~ 1 0 M. The optimum conditions selected were 0.1 M phosphate buffer, pH 7 and a final concentration of FMN and decanal as long chain aldehyde of 20 |omol and 0.005%, respectively. Bovine serum albumin (BSA) (0.2%) and 2 mM DTT were added in the reaction medium Using the batch system, the time necessary for the light emission to reach the plateau was dependent on the NADH concentration and was between 1 and 3 min with a stable signal for several minutes.

3.3 Extension to other analytes using dehydrogenases as auxiliary enzymes By co-immobilizing suitable dehydrogenases with the bacterial oxidoreductaseluciferase system, the biosensor specificity could be extended to other analytes, 105

LOIC J. BLUM AND PIERRE R. COULET

that is, the substrates of these dehydrogenases. This has been performed with sorbitol dehydrogenase (EC 1.1.1.14) from sheep liver and alcohol dehydrogenase (EC 1.1.1.1) from yeast, for the assay of sorbitol and ethanol, respectively (15). Briefly, the three enzymes were co-immobilized on the same preactivated polyamide disk. When dealing with multi-enzymatic systems, It must be stressed that careful attention must be paid to compatibility of both co-substrates concentrations and optimum pH values of the different enzymes. There is no general rule and conditions vary with each system. For sorbitol determination, the reaction medium was optimized as follows: 0.1 M phosphate buffer, pH 7.3 containing 0.2% BSA and 2 x l O ~ 4 M DTT; 2 x 10 ~5M FMN; 0.005% decanal in 0.1% Triton X-100; 1 mM NAD + . The linear range of assays performed in a batch system was 2 x 1 0 ~8M to 2 x 10 ~5M at 23 °C with a response time at the steady state of 6-7 min. The coefficient of variation for 10 replicate measurements was 6% at 4.4 x 10 ~7M. In another approach (16), a compartmentalization of the sensing layers was obtained with the three-enzyme sequence organized in two different stacked disks, one bearing the sorbitol dehydrogenase and the other, the bacterial bioluminescent bienzyme system. Covalent immobilization of the enzymes have been performed on Immunodyne ABC type membranes from Pall Co. With a well-balanced ratio of the different enzymes in the two layers, the sensitivity of the resulting biosensor was noticeably improved. This was attributable to the effect of hyperconcentration of the common intermediate, NADH, in the microcompartment existing between the two enzymatic disks. This biosensor has been used for the determination of D-sorbitol in foodstuffs and pharmaceutical samples. Both the reproducibility and operational stability were satisfactory and the results were in good agreement with those given by a colorimetric reference method. Furthermore, they were obtained in a shorter time. For ethanol determination, the reaction medium was similar to the medium used for D-sorbitol determination with the co-immobilized three-enzyme system, except for NAD+ concentration here equal to 5.5 mM. The linear range of assay was 4 x 10 ~ 7 M to 7 x 10 ~5M. The bioactive disks kept at -20 °C in 0.1 M phosphate buffer, pH 7 containing 1% BSA, 20% glycerol, and 0.2 mM DTT kept their full initial activity after at least 3 months of storage.

3.4 H202 determination The fiber-optic sensor with peroxidase bound to a preactivated polyamide membrane as a sensing layer has been used for hydrogen peroxide determination based on the chemiluminescence of luminol. The linear dynamic range was 2 x 10 ~8M to 2 x 10 ~5M. Luminol concentration was chosen equal to 0.05 mM and a pH value of 8.5 was selected.

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LUMINESCENCE AND FIBER-OPTIC BIOSENSORS

3.5 Extension to other analytes involving H202 detection 3.5.1 Without auxiliary enzymes A chlorophenol sensor has been developed based on the ability of some halophenols to enhance the peroxidase catalyzed luminol chemiluminescence. The HRP was immobilized on a collagen membrane and among the chlorophenols tested, the lowest detection limit was obtained with 4-chloro-3-methylphenol and was equal to 0.01 (omol (i.e. 1.4 (ig/1 in the sample), determined in a flow system (17).

3.5.2 Using oxidases as auxiliary enzymes One of the interests of this approach is the possibility of co-immobilizing auxiliary enzymes leading to the formation of H2O2 as a product, together with the suitable luminescent system. This enables various compounds to be assayed at a very low level. For instance, glucose can be assayed using Gox co-immobilized with peroxidase on a preactivated polyamide membrane.

Results obtained with the flow system showed that the calibration plot was linear from 0.25 nmol to 0.25 (jmol. The biosensor was tested for the analysis of glucose in soft drinks and results were in good agreement with those obtained with a standard spectrophotometric method (18). An ECL-based fiber-optic biosensor for glucose and lactate has been also developed (19). Gox and lactate oxidase were immobilized either on polyamide or collagen membranes. With the polyamide membranes as the enzymatic support, the detection limits for glucose and lactate were 150 and 60 pmol, respectively, whereas with collagen membranes the corresponding values were lowered to 60 and 30 pmol, respectively. For both analytes, measurements could be performed over four decades of concentration. A good agreement was obtained between the biosensor method and a reference method for assays in natural media like sera for glucose or whey for lactate. Recently a chemiluminescent choline biosensor using histidine-modified peroxidase immobilized on metal-chelate substituted beads embedded in a PVA-SbQ sensing layer involving immobilized choline oxidase has been described and used in a flow system (20). This analytical system exhibits very good performances, notably an unusually long operational stability over 200 measurements with a coefficient of variation of 3.5%. We presume that the pre-immobilization of the HRP on IDA-sepharose greatly improves the sensing layer stability.

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4 Concluding remarks We have been convinced for many years that the performance of biosensors strongly depends on the characteristics of the sensing layer and we have spent time in designing and improving these layers. They were adapted first to electrochemical devices and then to optical biosensors. Shifting from micrometric to self-assembled nanometric bioactive layers for nanodevices, we expect new developments in the future but key characteristics such as sensitivity and stability will remain a challenging goal in every case.

References 1. Coulet, P. R. (1992). In Advances in biosensors (ed. A. P. F. Turner), p. 151, JAI Press Ltd., London. 2. Blum, L J. and Coulet, P. R. (ed.) (1991). Biosensor principles and applications. Marcel Dekker, New York. 3. Wolfbeis, O. S. (1991). Fiber optic chemical sensors and biosensors, Vol. I & II. CRC Press, Boca Raton, FL 4. Blum, L. J., Gautier, S. M., and Coulet, P. R. (1988). Anal. Lett., 21, 717. 5. Marquette, C. M., Ravaud, S., and Blum, L. J. (2000). Anal. Lett., 33, 1779. 6. Marquette, C. M., Degiuli, A., and Blum, L. J. (2000). Appl. Biochem. Biotech., 89, 107. 7. Blum, L. J. (1997). Bio- and chemi-luminescent sensors. World Scientific, Singapore. 8. Coulet, P., Julliard, J., and Gautheron, D. (1973). Brev. Invent. Fr., 2.235.133. 9. Coulet, P. R., Julliard, J. H., and Gautheron, D. C. (1974). Biotechnol. Bioeng., 16, 1055. 10. Assolant-Vinet, C. H. and Coulet, P. R. (1986). Anal. Lett., 19, 875. 11. Foulds, N. C., Frew, J. E., and Green, M. J. (1990). In Biosensors. A practical approach (ed. A. E. G. Cass), p. 97, IRL Press, Oxford. 12. Coulet, P. R., Carlsson, J., and Porath, J. (1981). Biotechnol. Bioeng., 13, 663. 13. Chaga, G. (1994). Biotechnol. Appl. Biochem., 20, 43. 14. Leca, B., Morelis, R. M., and Coulet, P. R. (1995). Mikrochim. Acta, 121, 147. 15. Gautier, S. M., Blum, L. J., and Coulet, P. R. (1990). J. Biolumin. Chemilumin., 5, 57. 16. Michel, P. E., Gautier, S. M., and Blum, L. J. (1997). Enzyme Microb. Technol, 21, 108. 17. Degiuli, A. and Blum, L. J. (2000). J. Med. Biochem., 4, 32. 18. Blum, L. J. (1993). Enzyme Microb. Technol. 15, 407. 19. Marquette, C. A. and Blum, L. J. (1999). Anal. Chim. Acta, 381, 1. 20. Tsafack, V. C., Marquette, C. M., Pizzolato, F., and Blum, L. J. (2000). Biosens. Bioelectron., 15, 125.

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Chapter 6 Determination of metal ions by fluorescence anisotropy: A practical biosensing approach Richard Thompson, Badri Maliwal, Hui Hui Zeng, and Michele Loetz Cramer Department of Biochemistry and Molecular Biology, University of Maryland School of Medicine, Baltimore, Maryland 21201.

1 Introduction and rationale Fluorescence anisotropy is a well-known experimental technique which has hitherto been little used for chemical analysis or sensing, the major exception to this being fluorescence polarization immunoassay (1, 2). Yet it has important advantages among photoluminescence techniques (fluorescence, phosphorescence) that argue for its use in this regard. Note that "polarization" and "anisotropy" measure the same phenomenon, the polarization of fluorescence emission; the two measurements are normalized differently, but are algebraically interconvertible with ease.

1.1 Why fluorescence anisotropy to determine metal ions? With the proliferation of techniques for determining metal ion concentrations free in solution, it is reasonable to discuss why a new technique, particularly a new fluorescence technique, should be useful. Several mass and optical spectroscopic and electrochemical techniques offer determination of metal ions with high sensitivity, precision, and accuracy, but most of these techniques are usable only on discrete samples and do not provide information in real time. For Cu(II), in particular, ion selective electrodes exist with high sensitivity and selectivity which permit a continuous readout (3), but the response time of these devices is fairly slow at low levels. For biological applications, in particular, the value of fluorescence-based techniques has been their adaptability to fluorescence microscopy, to permit imaging of metal ion levels and fluxes in the frame of reference of the living cell. This approach has been most fruitful in the study of 109

R I C H A R D THOMPSON ET AL.

calcium (4-6), but has been useful for other analytes as well (7). Key to these advances was the development of "wavelength-ratiometric" probes, where the concentration of the analyte (and the contrast of the image) is based on the ratio of fluorescence intensities at two different wavelengths. The virtues of this approach are that it is largely insensitive to variations in excitation intensity, probe concentration changes due to washout or bleaching, specimen thickness, and some inner filter effects. However, this approach has drawbacks as well. For imaging, one would prefer to change excitation rather than emission wavelengths, which for many useful probes largely precludes the use of laser excitation due to the lack of rapidly tunable laser sources, and makes it very difficult to construct a confocal laser scanning microscope. Also, optical devices such as acousto-optical tunable filters capable of reasonably fast scanning (milliseconds or less) of lamp excitation or emission have optical tradeoffs such as small numerical apertures which make their use suboptimal. Also, with wavelength ratiometric approaches, one must excite or emit at particular wavelengths where absorbance or emission is submaximal, and one cannot use light from the entire wavelength band being ratioed. In part to address these issues, fluorescence lifetime-based sensing (8-10) and imaging (11,12) were introduced, which offer many of the same advantages, but without the need to change wavelengths. Although several groups have demonstrated such microscopes and a version has just been introduced commercially, the need for either high frequency modulation of excitation or very short light pulses has made such instruments rather expensive and complex. Like wavelength-ratiometric measurements, but unlike lifetime measurements, anisotropy is a steady-state measurement which does not require short pulses or high frequency electronics. As a steady-state measurement, it can be performed with simple, even handheld fluorometers. Indeed, using the classic T-format optical configuration of Weber (13), anisotropy is explicitly ratiometric, and even when the more common L-format is used, the ratiometric advantages are nearly obtained (Thompson and Gryczynski, in preparation). Fluorescence anisotropy microscopy was demonstrated some time ago (14, 15). Unlike wavelength-ratiometric approaches, laser sources are not only usable but desirable, due to their typically high polarization and monochromaticity. Consequently confocal optics would appear to be easier to implement, although this has not yet been done. We have demonstrated metal ion sensing by anisotropy using two photon excitation (16). Moreover, the polarization measurement may use the whole bandpass, and optical devices are available which can rapidly switch between which polarization orientation of light they will transmit, or which transmit both simultaneously (Wollaston prism). From these purely optical considerations, it would appear that anisotropy-based indicator systems offer significant advantages for microscopy.

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METAL IONS AND FLUORESCENCE ANISOTROPY

2 Theory of anisotropy-based determination of metal ions 2.1 "Reagent" approaches The principle of determination of metal ions by measuring a change in fluorescence anisotropy is straightforward. We shall consider first the case of a macromolecule, which binds the metal ion, and a separate, fluorescent "reagent," whose binding to the macromolecule depends on the binding of the metal ion. In this case the macromolecule is apocarbonic anhydrase II, which exhibits relatively high affinities in its active site for Zn(II) (4 pM) and Cu(II) (0.1 pM); moderate affinity for Cd(II) (10 nM), Ni(II)(10 nM), and Co(II) (100 nM); and negligible affinity for other metal ions, notably Ca(II) and Mg(II). When Zn(II) is bound in the active site, certain aryl sulfonamides have been found to bind to the metal as a fourth ligand, displacing Zn-bound water (17) and inhibiting the enzyme; such inhibitors are important in treating glaucoma, and hundreds are now known (18). Cd, Ni, and Co promote sulfonamide binding only weakly when they are present in the active site, and Cu seems not to at all (19-21). Several aryl sulfonamide inhibitors have been described that are fluorescent (22-26). We found that if the aryl sulfonamide does not bind appreciably in the absence of the metal (27), the fluorescence changes accompanying the binding of the aryl sulfonamide reflected the fractional occupancy of the binding site by the metal, and thus the free metal ion concentration (28). Since the fluorescent aryl sulfonamide is a small molecule (<1000 Da) with a short rotational correlation time (<100 ps), if its lifetime and quantum yield are commensurate, we would expect it to exhibit a substantial increase in fluorescence anisotropy upon binding to the carbonic anhydrase, with its much longer rotational correlation time (15 ns (23)). (see Figure 1). Quantitatively, the anisotropy of

Figure 1 Anisotropy-based metal ion determination using a reagent. In the absence of Zn(ll), apo-carbonic anhydrase does not bind the fluorescent aryl sulfonamide, and it exhibits low anisotropy because its rotational correlation time is much shorter than its lifetime. In the presence of zinc the sulfonamide binds rigidly to the protein, exhibiting a high anisotropy because its lifetime is shorter than the rotational correlation time of the protein.

Ill

R I C H A R D THOMPSON ET AL.

the bound and free forms can be predicted (in the absence of segmental motion) by the Perrin equation:

where r is the observed anisotropy, r0 is the limiting anisotropy at the excitation wavelength in use, T is the lifetime, and § is the rotational correlation time. When the active site is partially saturated with Zn(II), it is also partially saturated with the fluorescent inhibitor, and what one observes is a mixture of anisotropies from the two forms: where robs is the observed anisotropy, r is the anisotropy, / is the fraction, and Qis the quantum yield of the free and bound states. For best results the lifetimes of the free and bound states should be between the rotational correlation times of the fluorescent inhibitor in its free and bound states; fortunately, this is the usual case. From a practical standpoint, the analysis is straightforward. Fixed amounts of the apoprotein and fluorescent aryl sulfonamide are added to a sample containing free zinc, and the anisotropy is measured. As we pointed out, the choice of concentrations of apoprotein and sulfonamide has an impact on the precision and detection limit of the determination. In particular, the detection limit is set not by the apoprotein's affinity for zinc, but rather by the ability to distinguish small proportions of the bound sulfonamide reagent in the presence of a large excess of free reagent. In the case of ABDN described in Protocol 4, the affinity constant of the sulfonamide for the holoprotein is about 0.8 |oM, and consequently the ABDN concentration must be in this range if most of the holoprotein is to have ABDN bound to it; otherwise, a significant fraction of holoprotein (and thus zinc) would remain "invisible." The emission shift and quantum yield increase of the bound form of ABDN make it possible to distinguish a rather small proportion, yielding a detection limit of less than 100 nM. From the standpoint of the metal, binding is in the stoichiometric regime (29): the free metal ion concentration is well above the binding constant to the apoprotein, and consequently any free metal ion is scavenged by the protein. Thus, the fractional saturation simply represents the total amount of free (and exchangeable) metal ion found in the sample. The concern here is that 1 nmol free zinc (well above the KD) in a test tube will only saturate 0.1% of 1 |oM apoprotein present there. In most cases the amounts of zinc likely to be present may be anticipated, and appropriate concentrations selected.

2.2 "Reagentless" approach A somewhat different approach from the "reagent" approach discussed above is the "reagentless" approach, which as might be expected does not require a separate diffusible reagent. Rather, a suitable fluorescent label is covalently attached to the protein, and exhibits an anisotropy commensurate with its

112

METAL IONS AND FLUORESCENCE ANISOTROPY

Figure 2 "Reagentless" anisotropybased metal ion determination. In the absence of metal, the lifetime of the fluorescent label is longer than its rotational correlation time, and anisotropy is low. Metal binding to the active site partly quenches the fluorophore by energy transfer or other mechanism, reducing its lifetime with respect to its rotational correlation time, and increasing the anisotropy.

Figure 3 Simulation of metal ion-dependence of fluorescence anisotropy in a reagentless format. Without metal ions the fluorescent label's lifetime equals the rotational correlation time of 15 ns, with a relative quantum yield of 1.0, and a result ing anisotropy of 0.2. Binding of metal ions to the binding site results in 25% ( +), 50% (o), 75% (A), or 90%(«) quenching, with a commensurate drop in lifetime. As metal ion concentration increases (metal ion concentration expressed in terms of the metal ion's KD), anisotropy increases. (Copyright American Chemical Society, reproduced with permission.)

lifetime and rotational correlation time (often that of the protein) Figure 2. Binding of the metal ion induces a change in the fluorescence lifetime (or more rarely, in the rotational correlation time), which changes the fluorescence anisotropy. 113

R I C H A R D THOMPSON ET AL.

The magnitude of these effects can be predicted using the Perrin equation (Equation 1); in favorable cases they can be quite large (30) Figure 3. The principal design issues lie in the selection of the fluorescent label, and its placement so as to assure an optimum response (30). The main advantage of the reagentless approach is the freedom from having to use a high concentration of the fluorescent sulfonamide, and maintain it in close proximity to the apoprotein; by contrast, the reagentless approach brings the label with the apoprotein stoichiometrically. Also, quenching is a more versatile phenomenon, in that more different kinds of metal ions can be determined in this way by their propensity to quench the fluorescence of a particular label.

3 Fluorescent aryl sulfonamides for zinc(ll) determination At present probably the best fluorescent aryl sulfonamide for determining zinc by fluorescence anisotropy is ABD-N. ABD-N is readily synthesized from two commercially available compounds, and the workup after the reaction is agreeably simple. ABD-N is synthesized from ABD-F (31) and ethanolamine. The reaction is a typical condensation of a sulfonyl halide with an amine to form a sulfonamide; in this case the amine is much cheaper than the sulfonyl chloride, so the high pH needed to keep the amine deprotonated (especially in the presence of the HF produced in the reaction) is easily provided by using a substantial excess of the amine. The reaction requires a fairly polar solvent to co-dissolve the sulfonyl fluoride and the ethanolamine; DMF has been satisfactory, and we have not tried others. The reaction mixture should turn a deep yellow-orange within a few minutes, and excitation with a longwave UV light elicits bright greenish fluorescence. The progress of the reaction may be followed by thin layer chromatography on silica gel (Rf of product = 0.78 in 20% methanol in methylene chloride), but in practice stirring for a few hours is always adequate. We have found it convenient to workup the reaction by dilution with water followed by freezing and lyophilization (freeze-drying), since the ethanolamine, DMF, and HF are volatile under these conditions. If the mixture melts during lyophilization either the vacuum is inadequate or not enough water was used to dilute the mixture. Lyophilization is complete when the outside of the flask is warm, with no ice on it; care should be taken to protect the vacuum pump oil from the corrosive HF and ethanolamine by protecting it with a cold trap containing dry ice-ethanol or liquid nitrogen. ABD-N is quite water soluble, so the classical method of sulfonamide purification by extraction of the product from a neutral mixture, followed by raising the pH to make the water soluble sulfonamide anionic and extracting into the aqueous phase is thus ineffective. We have not observed unreacted ABD-F or the hydrolysis product sulfonic acid on thin layer chromatography, so in general the product should be of adequate purity. Purity may be verified by checking the absorption and emission spectra (26) in the presence and absence of holocarbonic anhydrase II. The compound should be 114

METAL IONS AND FLUORESCENCE ANISOTROPY

stored frozen in the dark and dessicated; concentrated (>1 mM) solutions in ethanol are reasonably stable, but dilute solutions seem not to be.

Protocol 1

Synthesis of ABD-N Equipment and reagents ABD-F (Molecular Probes, Portland, OR; Cat no. F-6053) Ethanolamine (Aldrich, cat no. 39,813-6) 5 ml Wheaton V-bottom vial (Aldrich Zll.511-8) Dimethylformamide Triangular magnetic stirring bar (Pierce cat no. 16000) 100 ml pear-shaped flask

100 ml pear-shaped a flask Silica-gel TLC plates, no fluorescent indicator (Aldrich Z18.531-0) TLC development chamber (Aldrich Z24,390-6) TLC eluent: 20% methanol in methylene chloride Lyophilizer Dry ice-ethanol bath, or liquid nitrogen bath.

Method 1 Dissolve 5 mg ABD-F in 1 ml DMF in V-bottom vial with stirring. 2 Add 200 |ol ethanolamine; stir at room temperature. 3 Monitor progress of the reaction by spotting aliquots of the reaction mixture on silica gel TLC sheets and chromatographing; reaction is usually complete in 1 h. 4 Add reaction mixture to pear-shaped flask and dilute with 50 ml water; freeze in a shell on the inside of the flask using the dry ice-ethanol or liquid nitrogen baths as available. 5 Put pear-shaped flask on lyophilizer overnight, or until complete.

4 Removal of zinc from carbonic anhydrase (CA) Carbonic anhydrase binds zinc extremely tightly (Ku = 4 pM (32)) and because of the slow dissociation rate constant zinc dissociates very slowly indeed (Ti/2 = 8200 min (33)). Consequently, even putting CA together with an excess of a strong zinc chelator like phenanthroline at a low pH (e.g. pH 5.5) does not thoroughly remove the zinc for several days. Happily, Hunt found that dipicolinic acid (pyridine-2,6-dicarboxylic acid; DPA) would catalyze the release of the zinc very effectively, even though DPA is not an especially strong chelator (34). In practical terms, removal of zinc need only take hours, or overnight with very little effort. In either case the holoprotein is dissolved in DPA-containing buffer and then separated from it by dialysis, either actively in a centrifuging device, or passively. Ordinarily, the zinc is removed by the DPA treatment in less than an hour, so in the passive method the dialysis can begin immediately, since

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it takes some hours to reduce the DPA to an ineffective level. The DPA is the separated from the protein by dialysis or gel filtration, in part because it can impart weak fluorescence and its absorbance interferes with determination of the protein concentration by spectrophotometry. Once the DPA is removed, special precautions are necessary to prevent the apoprotein from binding to contaminating zinc in solutions and on glassware; the precautions are listed in Protocol 3. The apoprotein is conveniently stored in HEPES sodium sulfate buffer; the presence of high concentrations of chloride or phosphate may interfere with sulfonamide binding (35). HEPES is chosen as the buffer becaus of its low affinity for zinc and availability in highly purified form; also satisfactory are Bicine (SigmaUltra grade, Sigma cat. No. B-8660) and MOPS (SigmaUltra grade, Sigma cat. No. M-5162). Note that the protein is stable for long periods at neutral pH in the apo-form, and indefinitely in the holo form (36), but that it is less stable at pHs below 6. Note that the apo and holo forms may be lyophilized. Verification that the zinc has been removed can be done by adding a small aliquot of the apoprotein to a solution of dansylamide (DNSA) in aqueous buffer, and exciting with an ultraviolet source. DNSA in the presence of the apoprotein emits weak greenish emission, whereas if any holoprotein is present strong bluish fluorescence is observed (23, 27). Small proportions of holoprotei can be removed (if necessary) by passage over an arylsulfonamide affinity chromatography column: the apoprotein passes through unretarded, whereas holoprotein binds tightly to the resin. Readers are urged to consult Protocol 3 to avoid re-binding zinc to the newly made apoprotein.

Protocol 2

Preparation of apoprotein Directions are given for both active and passive dialysis methods. The active method is faster (about 2 h, mainly determined by the centrifugation), but more labor-intensive; the passive method is less trouble and easier to scale up.

Equipment and reagents Bovine carbonic anhydrase II (catalog no. C2522, Sigma Chemical; CA II from other sources or CA I may be substituted) 100 ml 10 mM Tris 50 mM 2,6-dipicolinic acid (Aldrich P6380-8), pH 7.2 15 ml volume Amicon Centriprep concentrator (Model YM-10, 10,000 MWCO, cat. No.4321) Sephadex G-25 minicolumn (PD-10; cat. No. 17-0851-01, Amersham Pharmacia Biotech AB) DNSA; (Aldrich cat. No. 21,889-8)

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• Spectrapor dialysis tubing, 10,000 molecular weight cutoff (passive method) 2 1 50 mM HEPES (Ultrapure Bioreagent, J.T. Baker cat. No. 4018-04), 100 mM sodium sulfate, pH 7.3, Chelexed (passive method) Aminobenzenesulfonamide-agarose affinity resin (Sigma cat No. A-0796) 0.5 x 5 cm chromatographic column Spectrophotometer Spectrofluorometer (ThermoSpectronics AB-2, or equivalent).

METAL IONS AND FLUORESCENCE ANISOTROPY

Method (active) 1 2 3 4

5

6

10 mg CA is dissolved in 15 ml Tris-DPA buffer, added to the Centriprep tube, and centrifuged at 3000 rpm for 30 min, or until complete. The filtrate (inner tube) is discarded, and fresh Tris-DPA buffer is added up to the line; the Centriprep is recentrifuged as above. The PD-10 column is washed with approximately 10 ml of HEPES sodium sulfate buffer. Approximately 1 ml of the retentate is applied to the PD-10 column and eluted with HEPES sodium sulfate buffer; the apoprotein concentration in the excluded volume is determined by spectrophotometry (g2go = 49,000 M ~ a cm"1) Success of zinc removal is assessed by adding 0.5 |oM apoprotein to a 10 |oM solution of DNSA and measuring the emission spectrum (excitation at 330 nm, emission scanned from 400 to 650 nm) in comparison to that of the DNSA alone. Significant emission at 450 nm indicates the presence of holoprotein. Small proportions of holoprotein may be removed by passage of the apoprotein preparation over the 0.5 x 5 cm column of aminobenzensulfonamide affinity resin eluted with HEPES sodium sulfate.

Method (passive) 1

10 mg (or more) CA II is dissolved in a minimum volume (approx. 1 ml) Tris-DPA buffer, incubated for 1 h at room temperature, and introduced into the dialysis tubing. It is dialyzed overnight at 4 °C or for 4 h at room temperature vs 11 of HEPES sodium sulfate buffer in a plastic beaker. The dialysis is repeated at least one more time. 2 Completion of the zinc removal and elimination of any remaining holoprotein are carried out as in steps 5 and 6 above.

5 Avoidance of metal ion contamination A corollary of the high affinity of apocarbonic anhydrase for certain metal ions is the propensity for contamination with metal ions. CA can be saturated (in a sufficiently large volume) with copper or zinc ions at picomolar levels. As a result it is necessary to consider other sources of contamination. For instance, A.C.S. reagent grade buffer salts are typically specified to contain "heavy metals" at the level of a few parts per million. Buffers at typical millimolar levels made using these salts will therefore have metal ion concentrations in the nanomolar regime, which is unsatisfactory. Often the specification for the buffer salt is given as "X ppm heavy metals (as Pb)," which means that the lead levels have been determined as an indicator of purity, but not necessarily other metal ions of interest. Other grades of material from various suppliers are of adequate purity; for example, "Ultrapure," "Suprapur," and "electronic" grades are typically satisfactory, and often provide detailed analyses to assure this. This is important 117

R I C H A R D THOMPSON ET AL.

if the buffer constituent cannot be freed of metal ion contamination as described below, for instance, if it is a metal ion itself, such as calcium or magnesium. Typically, it is necessary to remove metal ion contamination by chromatography over a chelating resin such as Bio-Rad Chelex-100 (Bio-Rad Laboratories, Hercules, CA 94547). This resin contains iminodiacetate residues immobilized on polymer beads, which tightly bind most metal ions; details of its use are given below. Similarly, the water in which the buffer is made must be comparably clean; fortunately, the 18 Mfi water produced by many laboratory water systems is usually satisfactory. Other materials are potential sources of contamination as well. Most glassware tends to leach metal ions into solution, which can be minimized by rinsing in acid such as 1M HC1; however, as a general rule solutions once freed of contaminating metal ions are never permitted to contact glass again. More convenient is the use of disposable plasticware, which generally has little or no contamination; caution must be exercised, however, as some mold release agents contain zinc. We have found that SAMCO "Pasteur" pipets, Eppendorf Microfuge tubes, and Bio-Rad metal-free pipet tips (catalog BR-41) are all satisfactory. Of course, no such procedures are necessary until the solution to be handled has been stripped of metal ions. Removing metal ion contaminants from solution by passage over a Chelex column is straightforward. The resin is poured into a 2 x 30 cm (or other suitable size) chromatographic column in distilled water, and a few column volumes of water are passed through it, and it is allowed to run "dry" to minimize dilution of the buffer. The buffer is then passed through at a rapid rate; we typically find that a modest pressure head of 30 cm provides an adequate flow. After passing the buffer through, the column is again allowed to run dry, then rinsed with a few column volumes of water. This is important with buffers or fluids that will support bacterial growth, such as citrate buffers or artificial cerebrospinal fluid. At some point the column resin can become saturated with metal ions, and then the resin must be discarded or regenerated.

Protocol 3

Preparation of solutions free of contaminating metal ions Equipment and reagents • Bio-Rad Chelex-100 biotechnology grade chelating resin, Cat No. 143-2832 • Calcium chloride tetrahydrate, Suprapur grade, EM Science cat. No. 2384-2 EM Industries, Gibbstown, NJ 08027

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• Magnesium sulfate heptahydrate, Puratronic grade, cat No. 10801, Alfa Aesar, Ward Hill, MA 01835 • 2 x 30 cm chromatographic column

METAL IONS AND FLUORESCENCE ANISOTROPY

Protocol 3 continued Metal-free pipet tips (Bio-Rad BR-41: catalog 223-9041) Eppendorf 1.5 ml natural polypropylene microfuge tubes (cat. No. 2236 411-1) SAMCO transfer pipets (cat. No. 694 Q-PET; Samco Scientific, 1050 Arroyo Ave., San Fernando, CA 91340-1822) 1 1 polycarbonate bottles (Nalgene cat. No. 2205-0032)

1 1 Teflon bottles (Nalgene cat. No. 1600-0032) 1 1 polypropylene bottles (Nalgene 2006-0032) 2 1 polycarbonate bottles (Nalgene 2205-0210).

Method (removal of metal ion contaminants from solutions and buffers) 1

Pour the Chelex resin (dry or wet) into the 2 x 30 cm (or other convenient size) chromatographic column. Rinse with 5-10 column volumes of double-distilled (or better) water (30-50 cm pressure head is adequate). 2 Let the water run out of the column until the resin is "dry." 3 Add buffer to be stripped of metal ions to the column, and let it run through the column into a plastic bottle. 4 Rinse remaining buffer out of column with distilled water as above.

Method (preparation of buffers that contain metal ions such as Ca or Mg) 1

Prepare buffer except for Ca and Mg salts, and remove contamination by Chelex treatment as above. 2 Add Suprapur grade CaCl2 and/or Puratronic grade MgSO4.

6 Determination of Zn using a reagent approach Free zinc concentration down to the nanomolar range may be readily determined by fluorescence anisotropy using apocarbonic anhydrase and ABDN as a reagent. To an aqueous sample containing free zinc ion near neutral pH are added apocarbonic anhydrase and ABDN (usually in ethanol or DMF). Suitable concentrations of apocarbonic anhydrase and ABDN are in the range of a few micromolar each, for the reasons given in Section 2.1. Care should be taken to not add more than a few percent of the organic solvent containing the ABDN to the reaction mixture to avoid precipitating the protein. Samples should be incubated for at least 20 min, and preferably 1 h (or 1 h at 35 °C) for samples suspected of having nanomolar concentrations of zinc. The reason for this is the slow rate constant for association of zinc with the (wild-type) apoprotein. The slowness of the reaction can be circumvented by the use of fast CA variants invented for the purpose by Professor Carol Fierke and her colleagues at the University of Michigan. This is illustrated in Figure 4 (37). Anisotropy is read with excitatio at 430 mn, emission at 560 mil or shorter; the anisotropy is wavelength 119

R I C H A R D THOMPSON ET AL.

Figure 4 Fluorescence intensity of apo-CA (human wild-type, •; or E117A variant, (o) plus ABD-N in the presence of 1 nM free Zn at room temperature, expressed as a function of time. (Copyright Society of Photooptical Instrumentation Engineers, reproduced with permission.)

Figures Schematic of determination of Zn( 11) by apo-CA and ABD-N; inset shows the emission spectrum of apo-CA plus ABD-N in the presence and absence of Zn(ll).

dependent owing to the shift upon binding Figure 5. The free zinc concentration is determined by reference to a calibration curve depicted in Figure 6. The calibration should be performed on the same fluorometer with the same wavelengths and slit widths, as the anisotropy is wavelength dependent.

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METAL IONS AND FLUORESCENCE ANISOTROPY

Figure 6 Fluorescence anisotropy of apo-CA plus ABD-N as a function of free Zn(ll) concentration, measured at emission wavelengths of 560 (o) and 680 (y) nm. (Copyright Elsevier Science B.V., reproduced with permission.)

In the same way that it is difficult to prepare stable solutions with low concentrations of protons by adding small amounts of bases or acids, it is similarly difficult to prepare solutions with low concentrations of free metal ions. In biological systems as well as natural waters, a large fraction of metal ions are bound to various molecules having a wide range of affinities for the metal ion in question, as well as a wide range of association and dissociation kinetics. It is thus necessary for calibration purposes to prepare solutions that buffer the metal ion concentrations (6). These buffers contain ligands which, in a manner analogous to pH buffers, bind the free metal ion such that it is 95% or more bound, leaving a small proportion free in solution. The amount of free metal is adjusted by changing the total metal ion concentration, care being taken that the concentration of ligand is substantially greater than the total metal ion concentration. For simple buffers a pH buffer is included, chosen not only for pH buffering in the given range, but low affinity for the metal of interest. For zinc buffers MOPS and HEPES are often used as pH buffers, and depending on the zinc concentration range, one might use Bicine or nitrilotriacetic acid as the ligand. Simple buffers may be formulated as described (38), but buffering metal ions in media where other chelators and metal ions may be present such as artificial sea waters and cerebrospinal fluids requires a more sophisticated program such as MINEQL (Environmental Research Software, Hallowell, ME). Sample recipes for a particular range of zinc concentrations is given in Table 1; note that the recipe must be recalculated for differing pH, temperature, or ionic strength. 121

R I C H A R D THOMPSON ET AL. Table 1 Zinc ion buffer recipe. 15 mM EGTA, 10 mM MOPS pH 7.0, 25°C

[Zn(ll)], free, pM

[Zn(ll)], total, |xM

1250

3691 2104 1131

625 312 156

78.1 39.1 19.5 9.77 4.88 2.44 1.22 0.61

588 300 151

76.1 38.1 19.1 9.55 4.78 2.39

ml 0.0499M Zna to add to 500 ml 36.98 21.08 11.34 5.89 3.00 1.52 0.762 0.382 0.191 0.0957 0.0479 0.239

Note lt is convenient to use a volumetric standard solution of zinc (e.g. Aldrich cat. No. 31,962-7) of known (assayed at 0.0499 M) concentration to prepare these buffers. a

Protocol 4

Determination of zinc concentration with ABD-N: a reagent approach Equipment and reagents ABD-N (synthesized in Protocol 1) Apo-carbonic anhydrase (stripped of zinc in Protocol 2) 50 mM HEPES, pH 7.3, Chelexed (see Protocol 3) or other suitable buffer

Sample containing free zinc Free Zn calibration standards (formulated as in Table 1).

Method 1 Add apo-CA to the sample and aliquots of the calibration standards to approximately 3 uM. 2 Take emission spectrum (excitation 430 nm, emission 500-700 nm); if significant background intensity is present, measure the fluorescence intensities at the four polarizations at excitation 430 nm, emission 560 nm, and subtract these values from those measured in the presence of ABD-N to correct for the background. 3 Add ABD-N to approximately 5 |oM. 4 Wait 1 h (or less at warm temperatures, higher Zn concentrations, or fast apo-CA variant, as discussed above). 5 Measure anisotropy with excitation 430 nm, emission 560 nm.

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Figure 7 Fluorescence anisotropy of apo-N67C-ABD-T as a function of the concentrations of Cu(ll) (O), Zn(ll) (+), Cd(ll) (o), Ni(ll) (A), and Co(ll) (•). (Copyright American Chemical Society, reproduced with permission.)

7 Determination of Cu and other ions by using a reagentless approach In some situations the fact that the fluorescent sulfonamide is a separate molecule from the protein which binds the metal can lead to concern. For instance, if a specimen has some mechanical filtration property within its matrix, the concentrations of ABD-N and apoCA may change as they diffuse through the specimen at differing rates. Certainly a receptor or macromolecule which preferentially bound one of the two would alter the relative concentrations and thus the response to zinc. Clearly it would be more desirable to have the fluorescent moiety attached to the protein. The fluorescent moiety could be chosen to exhibit a lifetime change upon binding of the metal, and thus an anisotropy change as depicted in Figure 2. We designed a fluorescent aryl sulfonamide, ABD-T (7-(5maleimidyl)-pentylaminobenz-2-oxa-l, 3-diazole-4-sulfonamide), capable of being coupled to apocarbonic anhydrase, which exhibits substantial changes in its fluorescence lifetime and therefore anisotropy when metals bind to the protein Figure 7. The sulfonamide moiety does not appear to bind to the metal itself. The protein has certain amino acid(s) on its surface replaced with cysteinyl residues; the position of such cysteinyl residues may be chosen to optimize the response knowing the physics of quenching by Forster transfer (39). The desirable parameters may be chosen using simulations like that in Figure 3. Variants such as 123

R I C H A R D THOMPSON ET AL.

N67C, L198C, and S131C (30, 40) have proven most useful in this regard, but others are certainly feasible. Conjugation of the protein variant with the thiol-reactive fluorophore ABD-T is done using standard procedures. Reduction of the cysteinyl thiol is assured by use of the reducing agent TCEP, rather than a classical thiol like p-mercaptoethanol or dithioerythritol (DTE), because it does not react with the reagent. The reagent at present is custom synthesized, but is likely to become commercially available shortly. Other covalent labels (ABD-F (30) and dansylaziridine (41)) worked, but with poorer response than ABD-T.

8 Calibration of anisotropy Several approaches have been described for calibration of anisotropy measurements. Naturally, this entails measurement of a sample having known anisotropy in one's instrument. Typical fluorescence anisotropy instruments used in research have accuracy and precision in the range of ±0.002 or so (42), under ideal conditions where excitation noise and detector noise are not excessive; the latter condition is typically met with a bright sample requiring only moderate detector gain. Kawski has described a much more precise instrument (100-fold better), but these are not commercially available.

Protocol 5 Protocol 5

Determination of Cu and others by using a reagentless approach Equipment and reagents ABD-T, custom synthesized TCEP (tris-(2-carboxyethyl)phosphine, hydrochloride) Molecular Probes catalog T-2556 50 mM HEPES buffer, pH 7.5 Metal ion buffers for the desired range

Reagents and equipment for zinc removal, in Protocol 2

N67C-CA 5 ml Reactivial with triangular stir bar Spectrofluorometer fS-mercaptoethanol.

Method 1

Dissolve 3 mg (0.1 (omol) of N67C-CA in 4 ml buffer in the Reactivial with gentle stirring, add 0.6 mg TCEP and gently stir for 1 h. 2 Add 0.5 mg ABD-T and stir for 4 h; quench by adding 50 [d p-mercaptoethanol if desired. 3 Dialyze the reaction mixture against buffer, or dipicolinate buffer for zinc removal as in Protocol 2. 124

METAL IONS AND FLUORESCENCE ANISOTROPY

Protocol 5 continued 4 Add 50 nM (or more) apo-N67C-ABD-T to sample containing free metal ion or calibration buffer; equilibration occurs in minutes (Cu, probably Ni, Co, perhaps Cd) to an hour for Zn (see Figure 4). 5 Measure anisotropy in spectrofluorometer with excitation at 430 nm, emission at 550 nm.

The most common samples for calibration are scattering solutions, because dilute scattering solutions have an anisotropy of 1.0000. However, the use of scattering solutions as high accuracy standards requires some care, as discussed below. The most widely used scattering solutions for this purpose are suspensions of glycogen, colloidal silica (DuPont Ludox), or commercial coffee creamer. Any of these are satisfactory as long as they do not fluoresce and are well dispersed in the solvent (usually water) so that the intensity of the scattering does not vary as aggregates drift through the excitation beam. Such samples are not stable and must be prepared fresh. For accurate results, the concentration of the scatterer must be so low as to be almost imperceptible. This is because if a sufficient proportion of light is scattered twice instead of once, the anisotropy is reduced. Moreover, the actual amount of scatterer required varies with excitation due to the v4 dependence of scattering. Thus while a useful solution for excitation around 400 nm might look only slightly more cloudy than tap water, one used at 800 nm should be overtly cloudy because the strength of scattering is 16-fold less. Some have proposed the use of the O-H stretching Raman band of water (or hydroxylic solvents) as a scattering sample, and inasmuch as Raman is indeed a scattering phenomenon, it can be used. This approach has the virtue that the sample (clean water) is easy to obtain, and double scattering events are not observed. However, except in UV the Raman scatter is sufficiently weak such that the detector gain must usually be high, which may increase noise and reduce precision. With bright, monochromatic laser sources this is less of a concern. Also, care must be taken to assure that no fluorescence is present in the sample (HPLC grade water or better is preferred), as clearly a few percent of fluorescence admixed with the Raman scatter will corrupt the anisotropy measurement. To maximize the strength of the Raman signal excitation in UV at approximately 350 nm, and emission at 402 nm is preferred; often fluorometer manufacturers describe conditions (wavelengths, slit widths) whereby the Raman signal-to-noise ratio may be maximized, as a performance test. Using a fluorescent sample as anisotropy standard has the virtues that (1) it more precisely mimics the actual samples to be tested, and is less prone to error from excitation scattered elsewhere in the optical train or specularly reflected and (2) the intensity (and often wavelength) may be chosen to give the best precision of which the instrument is capable. Making an anisotropy sample with a value arbitrarily close to zero is straightforward. Many simply use 125

R I C H A R D THOMPSON ET AL.

Figure 8 Absorbance (dashed line) and fluorescence emission (solid line) spectra of Erythrosin B in pH 7.2 buffer.

Figure 9 Absorbance (dashed line) and fluorescence emission (solid line) spectra of Phloxine B in pH 7.2 buffer.

fluorescein in a fluid solvent at low concentration, but consideration of the Perrin equation reveals that the anisotropy with excitation at the peak is about 0.005, which is unsatisfactory. The problem may be avoided by excitation at a wavelength where the limiting anisotropy (r0) is near 0.000; for fluorescein this wavelength is about 350 nm at low temperature (temperature dependent) (43).

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METAL IONS AND FLUORESCENCE ANISOTROPY

For intermediate values of the anisotropy, the best choices are fluorophores with short lifetimes, perhaps in viscous solvents. Thus, fluorophores like Erythrosin B or Phloxine B with lifetimes due to their halogenation of much less than 1 ns exhibit substantial anisotropy in fluids as viscous as water at room temperature. These standards are also useful for imaging and microscopy applications. Cuvettes are preferably made of synthetic fused silica (e.g. Suprasil, Spectrosil) rather than fused quartz; the latter has modest levels of fluorescing impurities, which can add background. Samples should be made fresh for the day's measurements, as fluorophores degrade rapidly in dilute solution; it is most convenient to put a small, unweighed amount in ethanol and then dilute this to give an appropriate optical density, rather than weighing material out. An emission spectrum is taken to assure that the instrument is performing satisfactorily (Erythrosin B: 500-650 nm, Figure 8; Phloxine B (525-675 nm, Figure 9): fluorescein: 490-640 nm). For these standard compounds the intensity is high enough that relatively small slit sizes (2 or 4 nm bandpass on excitation and emission) can be used, while still keeping the PMT voltage at a fairly low range. If the signal is low, open up the slits first, if this can be done without letting in stray excitation light due to the small Stokes' shifts of these fluorophores. If the signal is still too weak, increase the voltage on the photomultiplier tube (PMT); for the commonly used Hamamatsu R928 PMT, voltages much above 1100 Vresult in the noise level increasing faster than the signal level, as judged by the dark current. Next, an emission spectrum must be taken of the solvent or buffer alone under the same conditions. This is very important, for several reasons. First, it verifies the integrity of the sample. Second, it checks for impurities in the solvent. Third, it checks for the presence of Rayleigh or Raman scatter in the emission, which are major sources of artifact in anisotropy measurement. This is attributable to the fact that the completely polarized scattered light contributes disproportionately to the anisotropy. In an intensity measurement, the admixture of 5% scattered light would be barely perceptible, but add 5% scattered light to a fluorescence anisotropy measured at 0.05, and the value increases to 0.10, a 100% error. Excitation and/or emission filters may be used to avoid this stray light. Following the spectra, the anisotropy is measured at the suggested excitation and emission wavelengths. For fluorescein at the given conditions, the anisotropy is 0.000; for Phloxine B, it is 0.055, and for Erythrosin B, it is 0.243. While in principle one can precisely measure the anisotropy by measuring the intensity at each of the four possible polarizer orientations with prolonged signal averaging, in practice we find it better to signal average at each position for a few seconds and perform several (4-10) cycles through all four positions, averaging the results. This permits the identification of "flyers," or other anomalies which might jeopardize the data set. For very weak signals, we take advantage of the fact that the "G-factor" used to correct for the non-ideality of the emission optical train in transmitting the two orthogonal polarizations is solely a property of the emission optical train. Thus, it can be measured using a different excitation wavelength, or even a different fluorophore under conditions where the signal-to-noise ratio is much better and higher precision is achievable, and 127

R I C H A R D THOMPSON ET AL.

the value applied to measurements with the dimmer sample. This avoids superimposing the noise of the G-factor measurement on the overall anisotropy measurement. As stated above, the ordinary accuracy and precision of anisotropy measurements in modern instrumentation is about ±0.002, but can be much better. Precision or accuracy much poorer than this under conditions of adequate signal requires explanation.

Protocol 6

Measurement and calibration of anisotropy Equipment and reagents Eiythrosin B (Aldrich cat no. 19,826-9) solution in water, O.D. <0.2 at 525 nm Phloxine B (Aldrich cat. No. 19,827-7) solution in water, O.D. <0.2 at 548 nm Fluorescein sodium (cat. No. 16,630-8) solution in water, O.D. <0.05 at 495 nm Water from a Milli-RQ_ water purification system (Millipore, Bedford, MA), but other water purer than double distilled (such as HPLC grade) is also likely to be satisfactory Cuvettes (synthetic fused silica) from NSG

Precision Cells, 195G Central Ave.,Farmingdale, NY 11735 (Type 23C); Starna Cells, P.O. Box 1919 Atascadero, CA 9342 <www.starna.com> (e.g. cat No. 21-Q;10); or Hellma Glastechnische-Optische Werke, Postfach 1163, D-79371, Mullheim, Baden, Germany; ; other brands are certainly acceptable Spectrophotometer Spectrophotofiuorometer.

Method 1 Take emission spectra of standard(s) with excitation and emission as follows: Erythrosin B, excitation at 500 nm, emission 520-650 nm; Phloxine B, excitation 510 nm, emission 530-675 nm; and fluorescein, excitation 400 nm, emission 490-640 nm). 2 Take emission spectrum of water or buffer alone, under the same conditions. 3 Measure anisotropy of standards at the following wavelengths: Erythrosin B: excitation 525 nm, emission 550 nm; Phloxine B: excitation 550 nm, emission 570 nm; and fluorescein: excitation 350 nm, emission 510 nm. Average at least 10 replicate measurements of the intensity at each of the four polarizer orientations.

Acknowledgments The authors wish to thank the Office of Naval Research, National Institutes of Health, and National Science Foundation for their support; Krystyna Gryczynska for drawing the figures; and Ignacy Gryczynski for helpful discussions.

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References 1. Dandliker, W. B., Kelly, R. J., Dandliker, J., Farquhar, J., and Levin, J. (1973). Immunochemistry, 10, 219-27. 2. Jolley, M. E., Stroupe, S. D., Schwenzer, K. S., Wang, C. J., Lu-Steffes, M., Hill, H. D., Popelka, S. R., Helen, J. T., and Kelso, D. M. (1981). Clin. Chem., 27, 1575-9. 3. Belli, S. L and Zirino, A. (1993). Anal. Chem., 65, 2583-9. 4. Tsien, R. Y. (1989). In Annual review of neuroscience, Vol. 12, p. 27, Annual Reviews, Inc., Palo Alto, CA. 5. Grynkiewicz, G., Poenie, M., and Tsien, R. Y. (1985). J. Bio!. Chem., 260, 3440-50. 6. Nuccitelli, R. (1994). In Methods in ceU biology (ed. L. Wilson and P. Matsudaira), Vol. 40, p. 364, Academic Press, New York. 7. Haugland, R. P. (1996). Handbook of Fluorescent Probes and Research Chemicals, Sixth edition Molecular Probes, Inc., Eugene, Oregon, p. 679. 8. Lippitsch, M. E., Pusterhofer, J., Leiner, M. J. P., and Wolfbeis, O. S. (1988). Anal. Chim. Acta, 205, 1-6. 9. Szmacinski, H. and Lakowicz, J. R. (1994). In Topics in fluorescence spectroscopy Vol. 4: probe design and chemical sensing (ed. J. R. Lakowicz), Vol. 4, pp. 295-334, Plenum, New York. 10. Lakowicz, J. R., Szmacinski, H., and Thompson, R. B. (1993). In SPIE conference on ultrasensitive laboratory diagnostics (ed. G. E. Cohn), Vol. 2388, pp. 2-17, SPIE, Los Angeles, CA. 11. Szmacinski, H., Lakowicz, J. R., and Johnson, M. L. (1994). In Numerical computer methods (ed. M. L. Johnson and L. Brand), Vol. 240, pp. 723-48, Academic Press, New York. 12. vandeVen, M. and Gratton, E. (1993). In Optical microscopy: emerging methods and applications (ed. B. Herman andj. J. Lemasters), pp. 373-402, Academic Press, New York. 13. Weber, G. (1956). J. Opt. Soc. Am., 46, 962. 14. Dix, J. A. and Verkman, A. S. (1990). Biophys. ]., 57, 231-40. 15. Fushimi, K., Dix, J. A., and Verkman, A. S. (1990). Biophys. J., 57, 241-54. 16. Thompson, R. B., Maliwal, B. P., and Zeng, H. H. (2000). J. Biomed. Optics, 5, 17-22. 17. Nair, S. K, Elbaum, D., and Christiansen, D. W. (1996). J. Biol Chem., 271, 1003-7. 18. Maren, T. H. (1977). Am. J. Physio!., 232, F291-7. 19. Lindskog, S. and Thorslund, A. (1968). Euro. J. Biochem., 3, 453-60. 20. Moratal, J. M., Martinez-Ferrer, M.-J., Jimenez, H. R., Donaire, A., Castells, J., and Salgado, J. (1992). J. Inorg. Biochem., 45, 231-43. 21. Einarsson, R. and Zeppezauer, M. (1970). Acta Chem. Scand., 24, 1098-102. 22. Elbaum, D., Nair, S. K., Patchan, M. W., Thompson, R. B., and Christiansen, D. W. (1996). J. Am. Chem. Soc., 118, 8381-7. 23. Chen, R. F. and Kernohan, J. (1967). J. Bio!. Chem., 242, 5813-23. 24. Thompson, R. B., Maliwal, B. P., and Zeng, H.-H. (1999). In SPIE conference on clinical diagnostic systems and technologies (ed. G. Cohn), Vol. 3603, pp. 14-22, Society of Photooptical Instrumentation Engineers, San Jose, CA. 25. Thompson, R. B., Maliwal, B. P., and Fierke, C. A. (1998). Anal. Chem., 70, 1749-54. 26. Thompson, R. B., Jr., W. O. W., Maliwal, B. P., Fierke, C. A., and Frederickson, C. J. (2000). J. Neurosd. Methods, 96, 35-45. 27. Thompson, R. B. and Jones, E. R. (1993). Anal. Chem., 65, 730-4. 28. Thompson, R. B. and Patchan, M. W. (1995). J. Fluores., 5, 123-30. 29. Weber, G. (1975). Energetics of Ligand Binding to Proteins. In Advances in protein chemistry (eds C. B. Anflnsen, J. T. Edsall and F. M. Richards) Vol. 29, pp. 1-83. 30. Thompson, R. B., Maliwal, B. P., Feliccia, V. L., Fierke, C. A., and McCall, K. (1998). Anal. Chem., 70, 4717-23. 31. Toyo'oka, T. and Imai, K. (1984). Anal. Chem., 56, 2461-4.

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R I C H A R D THOMPSON ET AL. 32. Huang, C.-c, Lesburg, C. A., Kiefer, L L, Fierke, C. A., and Christiansen, D. W. (1996). Biochemistry, 35, 3439-46. 33. Kiefer, L. L. and Fierke, C. A. (1994). Biochemistry, 33, 15233-40. 34. Hunt, J. B., Rhee, M. J., and Storm, C. B. (1977). And. Biochem., 79, 614-7. 35. Lindskog, S., Henderson, L. E., Kannan, K. K, Liljas, A., Nyman, P. O., and Strandberg, B. (1971). In The enzymes (ed. P. D. Boyer), Vol. 5, pp. 587-665, Academic Press, New York. 36. Thompson, R. B., Zeng, H. H., Loetz, M., and Fierke, C. (2000). In In-vitro diagnostic instrumentation (ed. G. E. Cohn), Vol. 3913, pp. 120-7, SPIE, San Jose, CA. 37. Thompson, R. B., Zeng, H.-H., Loetz, M., and Fierke, C. A. (1999). In SPIE conference on advanced materials and optical systems for chemical and biological detection (ed. M. Fallahi and B. I. Swanson), Vol. 3858, pp. 161-6, Society of Photooptical Instrumentation Engineers, Boston, MA. 38. McCall, K. A. (2000). Metal Ion Specificity and Affinity in Carbonic Anhydrase Variants. In Department of biochemistry, pp. 190 Duke University, Durham, NC. 39. Thompson, R. B., Ge, Z., Patchan, M. W., and Fierke, C. A. (1996). J. Biomed. Optics, 1, 131-7. 40. Thompson, R. B., Ge, Z., Patchan, M. W., Huang, C.-c., and Fierke, C. A. (1996). Biosens. Bioelectron., 11, 557-64. 41. Thompson, R. B., Maliwal, B. P., Feliccia, V., and Fierke, C. A. (1998). In Systems and technologies for clinical diagnostics and drug discovery (ed. G. E. Cohn), Vol. 3259, pp. 40-7, SPIE, San Jose, CA. 42. Jameson, D. M., Weber, G., Spencer, R. D., and Mitchell, G. (1978). Rev. Scientific Instr., 49, 510-14. 43. Lakowicz, J. R. (1999). Principles of fluorescence spectroscopy. Kluwer Academic/Plenum Publishers, New York.

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Chapter 7 Fluorescence-based fiber-optic biosensors David R. Walt, Caroline L. Schauer, Shannon E. Stitzel, Michael S. Fleming, and Jason R. Epstein Max Tishler Laboratory for Organic Chemistry, Tufts University, Medford, MA 02155.

1 Introduction This chapter describes optical biosensors that combine analyte-sensitive indicators with optical fibers for detecting chemical and biological species. Fiber-optic biosensors are more versatile because they are flexible in terms of their platform design and signal transduction. Optical transduction mechanisms include fluorescence, absorbance, and reflectance. Using these mechanisms, sensors can be designed to exploit optically detectable differences in intensity, lifetime, spectral shape, polarization, and wavelength (1). Optical fibers come in a range of lengths and diameters, and can be adapted for either single- or multianalyte detection, and can incorporate a variety of sensing chemistries. Because light is transmitted over long distances in a fiber, a detector can be located at a distance from the sensor tip, allowing the fiber to be used for remote or hazardous applications. In addition, fibers can be miniaturized, allowing the sensor to be used for in situ, or in vivo applications. Fiber-optic based sensors also have the advantage that they are not susceptible toward electromagnetic interferences. An optical fiber is comprised of a plastic, glass, or fused silica core, and a clad, which is a thin layer of lower refractive index material surrounding the core (2) (Figure 1). The different refractive indices of the clad and core cause light introduced at the fiber's proximal end to be totally internally reflected toward the distal end. A fluorescent indicating layer located on the fiber's distal tip responds to the presence of analyte by changing its optical properties. The change in the indicator's properties modulates the light, which is transmitted back through the fiber to a detector. The intensity changes or spectral properties of the returning light correlate with the analyte concentration. Fiber-optic sensors are created using either a single core fiber or an imaging fiber bundle. A single core fiber comprises one fiber with a surrounding cladding. 131

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Figure 1 Excitation light is introduced into the fiber. Since the core and the cladding have different refractive indices, the light is totally internally reflected the length of the fiber. Isotropic emitted light enters the fiber and is carried back to a detector.

An imaging fiber bundle is a coherent bundle of 3,000-100,000 individually clad fibers, each 3-10 (am in diameter. Each fiber's position is maintained throughout the bundle's length as shown in Figure 2. An imaging fiber bundle is able to transmit light signals and images concurrently (3,4), while a single core fiber can carry light, but not images. Due to the advantages associated with monitoring a signal and image simultaneously, this chapter will primarily focus on the use of imaging fibers, although the sensing chemistries can also be applied to single core fibers. Fiber optical biosensors can incorporate single or multiple analyte sensing formats. To detect a single analyte, a sensing layer is immobilized on the fiber tip as described in Section 2. The fiber tip is first functionalized with a surface reactive group, such as a primary amine or reactive acrylate. The reactive group is then used to couple a specific sensing material to the surface. Typical biosensors employ enzymes as the sensing material in conjunction with a fluorescent indicator, which monitors the enzymatic reactants or products, such as pH (5) or O2 (6). Multi-analyte sensing can be accomplished using an array of sensors on an imaging fiber. Biosensor arrays are fabricated by immobilizing the biological sensing agents on the imaging fiber tip, and are described in Section 3. One method for creating a multi-analyte sensing array is to generate local sensing regions by immobilizing polymer spots on a fiber tip (Section 3.1). An alternative approach involves etching the fiber to form an array of microwells on the fiber face, in which functionalized beads or cells are deposited. 132

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Figure 2 An optical imaging fiber comprises individually cladded optical fibers. When fused into a bundle, the entire bundle can transmit a coherent image, with each individual fiber in the bundle acting as a pixel in the entire image. With this image, a gradient index mask was used to filter light to transmit the image.

Fiber-optic biosensors use similar equipment to measure the fluorescence response for both single- and multi-analyte sensing formats. As seen in Figure 3, the instrumentation consists of a 75-W xenon arc lamp, computer controlled shutter and filter wheels, dichroic mirror, optical filters, and a CCD camera. The light sources and detectors can be adjusted to fit the experimental conditions. Although many of the procedures described in this chapter involve an optical fiber substrate, the immobilization techniques are universal, and can be transferred to other analytical methods, including membrane immobilization and other generic array platforms.

1.1 Fiber polishing Prior to fabricating a fiber-optic sensor, both ends of the optical fiber are polished to create a smooth surface with minimal scratching. A smooth surface ensures that a uniform surface is available for coupling sensing elements to the fiber surface, and facilitates visualization of transmitted light signals through the fiber. Individual fibers can be polished manually or multiple fibers can be machine polished simultaneously. The fiber ends are first polished with a series of aluminum oxide lapping films (12, 9, 3,1, 0.3 (am). The lapping films are kept wet during polishing. The polished fiber is sonicated in deionized water to remove any residual lapping material and allowed to dry. After polishing, the fiber tip is functionalized to facilitate adhesion of the sensing layer to the surface. Surface functionalization is described in Protocol 1. 133

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Figure 3 Instrumentation which is used for fluorescence measurements and imaging. A modified epifluorescence microscope was converted from a vertical to a horizontal configuration.

2 Single-analyte detection using an enzymatic sensing layer Enzyme-based sensors utilize an enzyme's inherent selectivity toward a single species or class of compounds. Many enzymatic reactions consume or generate acid or oxygen in response to the presence of a specific compound. Enzyme-based biosensors couple the enzymatic reaction with a fluorescent pH (7) or O2 (6) sensitive dye to determine the amount of analyte present. For example, esterases hydrolyze esters into alcohols and carboxylic acids. Therefore, by monitoring the amount of acid produced, one can determine the amount of hydrolysis.

2.1 Enzymatic sensing layer Co-immobilizing an enzyme and indicator within a polymer layer onto an imaging fiber bundle tip creates an enzymatic sensing layer. The fiber tip's silica surface must first be silanized to modify the silica surface with an amine, acrylate, or other reactive group. By adding a reactive group to the fiber tip's surface, the enzyme-containing polymer layer can be covalently attached to the fiber. These modifications are accomplished by allowing the silica hydroxyl groups on the fiber tip to react with organosilanes (Protocol 1) and then allowing the polymer precursors to react with the reactive residues attached to the surface. A transparent polymer layer is necessary to collect visual data. 134

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Protocol 1

Silanization/functionalization of a fiber tip 1 Place a polished fiber tip in a 10% 3-aminopropyl triethoxysilane/acetone solution for 2 h to add an amine functionality to the fiber tip. Alternatively, a 10% 3-(trimethoxysilyl)propyl methacrylate/acetone solution can be used to add an acrylate functionality to the fiber tip (5). 2 Rinse the fiber tip well with acetone and air cure for over an hour. Be careful not to touch the silanized tip.

2.2 PAN gel immobilization Poly (acrylamide-co-N-acryloxysuccinimide) (PAN) has been used to immobilize both enzymes and indicators without altering their respective properties (8). PAN has an N-succinimidyl ester functionality, which reacts with primary amines to create a cross-linked polymer matrix. Enzymes that have primary amines on the amino acid side chains, and indicators with amine-reactive groups can be immobilized in the PAN polymer as seen in Protocol 2. The PAN/enzyme layer is then attached to a fiber by reacting with an amine-functionalized fiber tip. While preparing the enzyme-sensing layer, an enzyme substrate is added at a concentration of 10 times Km to protect the enzyme's active site. Excess substrate ensures that active site residues are not modified. Once the polymer gel solution is formed, it is spin-coated onto the fiber face. In this process, an aliquot of the solution is dripped onto a fiber tip, and the resulting gel thickness is dependent on the rotation speed of the spin coater, as well as the amount of polymer cross-linking.

Protocol 2

PAN-enzyme sensors 1

Prepare a solution of HEPES/Na-HEPES by combining 9.5 ml of 0.3M HEPES [HA] pH ~5 (3.6 g/50 ml) and 9.0 ml of 0.3M Na-HEPES [A-], pH ~9 (3.9 g/50 ml dH2O).

2 Prepare a solution of 0.5M triethylenetetramine (TET) by adding 1.24 ml of TET to 8.8 ml of dH2O. 3 Prepare an enzyme substrate solution by dissolving lOx Km of enzyme substrate in HEPES/Na-HEPES buffer. 4 Prepare the enzyme solution by combining 500 units lyophilized enzyme and 200 |ol enzyme substrate solution. 5 Add 100 |ol of enzyme substrate solution to 25 mg of PAN, mixing with a stir rod to break up clumps. 6

Stir for 30 s, then add 10.6 |ol of 0.5M TET.

7 Stir for another 30 s, then add 25 jol enzyme solution.

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Protocol 2 continued 8 Stir for an additional 10 s, remove 10-20 |ol aliquot of solution, and place on an amino-silanized fiber tip (Protocol 1). Spin at 2000 rpm for 10 s. The remaining mixture should gel in 1-3 min. 9 Cure for 1 h at room temperature. 10 React the fiber tip for 15 min with an amine-reactive dye (such as Nhydroxysuccinimide or isothiocyanate derivatives) in enzyme substrate solution in the dark. 11 Wash the fiber tip 2x in HEPES/Na-HEPES buffer for about 30 min each. 12 Wash the fiber tip at least 3x in testing buffer to equilibrate. Store at 4°C until ready to use. Fiber-optic enzymatic sensors are suited for combined imaging and chemical sensing. For example, a PAN polymer-based pH sensor is used to monitor the acid release from sea urchin eggs upon fertilization, while simultaneously visualizing the eggs (9). PAN polymer is also used to immobilize acetylcholine esterase anda pH sensitive dye to measure in vivo activity of a mouse fibroblast (NIH 3T3) (3) These examples illustrate the ability to measure localized chemical dynamics in real-time, making PAN/enzyme sensors suitable for in vivo monitoring.

3 Multi-analyte arrays Imaging fiber bundles have both single- and multi-analyte detection applications. For multi-analyte detection, there are two different types of sensor formats. The first approach involves photopolymerizing discrete polymer sensors in multiple positions on an imaging fiber tip. The second method uses a chemical etchant to form an array of microwells on the fiber face, in which individual microsphere sensors are distributed. This section discusses these two different sensor formats.

3.1 Immobilization via polymer photodeposition Polymer photodeposition onto a fiber occurs via free radical polymerization initiated with ultraviolet (UV) light. The distal end of a methacrylate-silanized fiber (Protocol 1) is first placed into a solution containing a mixture of monomers, cross-linkers, enzymes, indicators, substrates, and photoinitiator. The indicators contain either a polymerizable functional group or a modifiable functional group to enable incorporation into the polymer matrix. For example, fluorescein is modified with an acryloyl group to form acryloylfluorescein (Protocol 3), which is pH sensitive and can be covalently immobilized in the polymer matrix. The photodeposition of polymer spots onto the fiber tip takes advantage of the imaging fiber's discrete light pathways (7, 10) (Figures 1 and 2). A specialized photodeposition system is used to control the position of each polymer spot on the fiber tip. The deposition system consists of a mercury-xenon arc lamp, appropriate collimating lenses and filters, a pinhole mask, an electronic shutter, and an inverted microscope connected to a CCD-camera (5, 7). 136

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The deposition is accomplished by using a pinhole mask to illuminate a discrete spot on the fiber's proximal end. Only the illuminated fibers carry light to the distal tip, so the photoinduced polymerization of the sensing polymer occurs only at the illuminated regions. The polymer spots produced on the fiber tip are dependent on the size and shape of the pinhole mask. An optical biosensor for penicillin is fabricated using photopolymerized immobilization of the enzyme penicillinase along with a pH indicator (7). The same fiber also has immobilized sites containing only pH sensors without enzyme. The fluorescent intensities of the enzyme-containing sites are compared with enzyme-free sites, enabling the penicillin concentration to be determined at any sample pH (7). By measurin both pH and penicillin, penicillin is determined without the need to maintain a constant pH. Biosensors used for fermentation monitoring encounter problems where the pH cannot be maintained. This dual-analyte sensor is used to monitor the fermentation ofPenicillium chrysogenum (3).

Protocol 3

Preparation of acryloylfluorescein (7) 1

Fluoresceinamine isomer I (180 mg, 0.519 mmol) and acryloyl chloride (45 |ol, 0.55 mmol) are added to dry acetone (20 ml), and the reaction mixture is stirred for 1 h in the dark. 2 Filter the resulting precipitate and then wash with acetone, followed by dichloromethane.

Protocol 4

Fabrication of an enzyme/pH array The following procedure is optimized for penicillinase, but can be adapted for use with other enzymes. 1 A pH sensitive monomer stock solution is prepared consisting of 10 ml of hydroxyethyl methacrylate (ophthalmic grade), 200 [d of ethylene glycol dimethacrylate, and 1 ml of acryloylfluorescein (Protocol 3) in n-propanol (5 mg/ml). 2 500 [d of the above monomer stock solution is combined with 30 mg of benzoin ethyl ether (BEE). 3 The enzyme is added to the above solution with an additional amount of BEE. Enzymes catalyze reactions at different rates, therefore, the amount of enzyme added should be enough so as not to limit the sensitivity of the sensor. For example, to prepare a penicillin sensitive polymerization solution, 6.5 mg of lyophilized penicillinase is added to 1 ml of the above working pH sensitive solution with an additional 60 mg BEE.

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Protocol 4 continued

4 The enzyme/pH sensitive polymerization solution is sonicated in an ice bath with a microtip probe from Heat Systems, Inc. at an output of 3 for 60 sec in the pulse mode (0.1 s On/0.4 s Off). Sonication is a means to ensure proper mixing, and the ice bath is used to prevent heating the suspension. 5 The solution is deoxygenated for 10 min with N2 and is used immediately. 6 While the solution is deoxygenated, a pinhole is placed between the mercury light source and the fiber to selectively illuminate the desired area. The illuminated fiber area depends on the pinhole size used and the amount of light focused on the fiber. 7 With a 1.0 neutral density (ND) filter in the optical axis, the illuminated area is focused to the desired size and the light flux maximized and measured at 188 mW/cm2. 8 With a 3.0 ND filter in place, the fiber's proximal end is positioned to illuminate the designated area. This positioning is performed after the fiber's distal tip is functionalized with a methacrylate group according to Protocol 1. 9 A small volume of the penicillin/pH sensitive polymerization solution is drawn into a capillary tube and the distal end of the fiber is placed in the tube. 10 The exposure time is set for 40 s with the electronic shutter. Exposure time must be determined and optimized for each preparation. 11 After the shutter is closed, the distal end of the fiber is washed with ethanol. 12 Repeat steps 8-11 at a different location on the fiber tip to create an array of polymer matrices (spots). The sensor arrays are stored in 5.0 mM phosphate (200 mM KC1) pH 7.0 buffer at 4°C until used. For other applications, the biosensor is modified to include an oxygen-sensing site by incorporating a solution of 1 mg/ml Ru (Ph2phen)32 + in dichloromethane with a siloxane working solution (6). The siloxane working solution is made by combining 200 |ol of (80-85%) dimethyl (15-20%)(acryloxypropyl) methylsiloxane copolymer with 545 jol of dichloromethane and 12 mg BEE. 3.2 Microwell array platform preparation The second multi-analyte approach involves an array of microwells etched into the fiber face (11) (Protocol 5). The microwells are used to house individually addressable sensing elements, typically microspheres coupled with a specific sensing chemistry. Various microsphere sensor arrays are made using the microwell format, including enzyme and DNA sensors (12-15). An imaging fiber bundle with a germania doped silica cladding and a silica core is chemically etched to create an array of microwells based on the different etch rates of the core and clad materials. Prior to etching, the fiber ends must be polished (Section 1.1) well to avoid pitting in the fiber during etching. Varying the etching times and etchant solution concentrations provides wells of different depths (11). After washing the fiber, the well depth is determined by scanning force microscopy. A chemical etch is performed using a buffered hydrofluoric acid (HF) solution discussed in Protocol 5 (16).

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Protocol 5

Chemical etching of a germania-doped imaging fiber bundle 1

In a 1.5 ml plastic eppendorf tube, combine 100 [d of HF (50%), 0.2 g of ammonium fluoride, and 600 [d of deionized water. 2 Hold the imaging fiber's distal face in the HF acid solution for 80 s, which results in approximately 5 (am deep wells. 3 Rinse the etched fiber in deionized water for 30 s followed by sonication for 30 s in deionized water to remove any salts that may have built up during etching. Caution: Hydrofluoric acid (HF) is extremely caustic!

3.2.1 Microsphere array fabrication and encoding Microsensors are made from commercially available silica and polymer microspheres and are available in a range of sizes and functionalities. The microsphere size should be complementary to the microwell diameter, and the functionality can be modified to incorporate the desired sensing chemistries. Enzymes are attached to the microspheres through multiple methods (Section 3.2.2). DNA microsensors are prepared by attaching either single stranded DNA sequences or molecular beacons to surfaces (Section 3.2.3). Microsphere sensors are usually stored as aqueous suspensions. A microsphere array is fabricated by placing an aliquot of the aqueous microsensor suspension on the etched fiber face. The microspheres then distribute into the wells as the solution evaporates, forming individually addressable optical sensors. Extra sensors not occupying wells are removed with an anti-static swab. Cotton swabs should not be used since they leave debris behind, which interferes with the assay. If the microsensors are sensitive to drying, the array can be assembled wet. Teflon capillary tubing extending 3-4 mm off the etched fiber end is used to hold 5 |ol of the sensor bead solution. Keeping the fiber upright, the bead suspension is pipetted into the tubing, allowing the beads to settle into the wells for 30 min. The array is then dipped into 1.5 ml of the appropriate buffer and the capillary tubing is removed with tweezers. The fiber is equilibrated in buffer for 5 min. The fiber is then placed in another 1.5 ml of fresh buffer for another 5 min before use. Many different sensor types can be placed into an etched imaging fiber bundle by simply mixing the different sensor types prior to creating the array as discussed above. This array fabrication process leads to a random distribution of the sensor elements throughout the array. For this reason, the position of each microsphere sensor in the array must be determined after fabrication through various encoding schemes (12, 16, 17). Depending on the specific application, array encoding can occur either prior or subsequent to assembly. In one encoding scheme, which relies on each sensor type producing a unique well-defined response to a specific analyte, encoding occurs after array fabrication. 139

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Microsensors are positionally registered by exposing the array to a test analyte. The signals are compared to known responses from each sensor class. Because there is no need for additional encoding, the sensors are considered "self-encoding." Another method for positionally registering the sensor class is to use an optical bar code. Prior to protein or DNA coupling, microspheres are encoded with dyes either internally, externally, or both to distinguish the different probe types in the array (12, 16, 17). Spectral overlap of all dyes should be avoided.

Protocol 6

Internal encoding 1

200 |ol (10.7% microspheres by weight) of a stock suspension of 3.1-|om diameter microspheres famine functionalized, 87% polystyrene, 13% divinylbenzene; Bangs Laboratories, Fisher, IN) are filtered, and then washed with dry THF. 2 Combine the beads in a microcentrifuge tube with 200 [d of europium (III) thenoyltrifluoroacetonate.3H2O (Acros, New Jersey) dye (ex/em 365/615) in THF and mix for 2 h. Dye concentrations of 1 M and dilutions ranging from 1:1 to 1:1000 are used to generate multiple distinguishable encoding levels. Filter the bead suspensions, wash with methanol, and store in 0.01% Tween in ultra-pure water.

Protocol 7

External encoding 1

External encoding is performed on 10 |ol (10.7% microspheres by weight) of microsphere stock famine functionalized, 87% polystyrene, 13% divinylbenzene; Bangs Laboratories, Fisher, IN). Rinse the stock three times with BT buffer (0.1M boric acid, 0.1M NaOH, 0.13M NaHCO3, and 0.01% Tween, pH 9). 2 Prepare an initial stock solution of Cy5 dye in 25 |ol DMF (one vial of dye is enough to functionalize 1 mg of antibody, Amersham Life Sciences, Inc, Pittsburgh, PA, ex/em 620/700). This stock solution is diluted to any distinguishable level, typically in the range of 1:1 to 1:100. The desired concentration is used in step 3. 3 Suspend the beads in 100 jol of BT buffer and 5 |ol of the Cy5 dye solution. Mix the beads with the dye solution for 2 h. 4 After the external encoding step, wash the beads three times with BT buffer and three times with 0.01M PBST buffer (phosphate buffered saline, 0.01% Tween, pH 7.4). The encoded beads can be stored for up to a year in the PBST buffer at 4 °C. Alternatively, another dye can be used separately or in conjunction with the Cy5 dye to increase optical encoding of the beads. Rhodamine-based dyes are typically used, such as N-succinimidyl esters of carboxytetramethylrhodamine (TAMRA; Molecular Probes, Eugene, OR) (ex/em 535/580). Bead suspensions are made in 100 |ol of BT buffer consisting of TAMRA concentrations of 0.1, 0.4, and 3.0 mM. The beads are mixed with the dye solution for 2 h. 140

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3.2.2 Enzyme-labeled microspheres Mixtures of enzymatic sensors are used to generate an array capable of multianalyte detection. Enzymes are either physically adsorbed to microspheres or covalently attached to the surface. The protocols listed below offer several methods for preparing enzyme-labeled sensors for use in the imaging fiber array platform. Enzymes can be immobilized onto nearly any type of surface including polymer, silica, or metal surfaces (such as gold). An array is constructed by first etching an imaging fiber as described in Protocol 5, then by adding an aliquot of a suspension of enzyme coated microspheres to the fiber tip. The solution is allowed to air dry for 2 min, and then excess beads are removed with an anti-static swab. The array is rehydrated prior to use by placing the etched end into an appropriate buffer. As previously described (Protocols 2 and 4), enzyme activity can be monitored by measuring enzymatic reaction products such as O2 or pH with fluorescent indicators. If multiple microsensor types are included in the array, the beads should be encoded as described in either Protocol 6 or 7, prior to enzyme immobilization.

Physical adsorption Proteins/enzymes may be adsorbed onto a surface using a variety of methods. The preparation of a concentrated enzyme solution (1-5 mg/ml) with a pH near its isoelectric point (pi) results in a protein with no net negative or positive charge. The buffer used to prepare the enzyme solution should not contain any detergents and should have a low ionic strength. The microspheres are cleaned to remove any impurities or other components that could interfere with adsorption to their surface. A general method for adsorbing proteins/enzymes to microsphere surfaces is detailed below in Protocol 8.

Protocol 8

Enzyme immobilization by physical absorption 1 Weigh 2 mg of dry silica or polymer microspheres (3-5 (am diameter) or measure a volume of a microsphere suspension that corresponds to approximately 2 mg of microspheres. 2 Wash the microspheres with PBST by centrifuging, removing the supernatant and resuspending the microspheres in an appropriate buffer. Repeat four times. 3 Exchange the protein into an appropriate buffer using either dialysis or ultrafiltration (choice of method depends on the stability of the protein). Bring the protein concentration up to approximately 2 mg/ml. A bicinchoninic acid assay (BCA) or simple absorbance reading at 280 nm may be used to estimate protein concentrations if unknown. 4 Centrifuge the microspheres and remove the supernatant. Add 500 jol of the protein solution to the microspheres, and place the solution on a shaker to gently mix at 4 °C for 2 h.

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Protocol 8 continued 5 After mixing, clean the microspheres as above with an appropriate storage buffer (usually a higher ionic strength buffer with a low level of a non-ionic detergent to prevent microsphere aggregation). Test the microspheres for retention of enzyme activity.

Covalent immobilization Enzymes are covalently immobilized onto microspheres using a variety of reagents. Immobilization reagents react with surface side chains, typically amine-containing amino acids such as lysine or with carboxylic acid containing residues such as aspartic or glutamic acids. The relative side-chain position in the protein's three-dimensional structure may affect protein binding. In many cases, immobilizations of large proteins (i.e. greater than 200 kDa) results in recovery of low specific activity due to alterations in protein structure. Post-translational modifications to the enzyme may also affect immobilization, as many proteins are heavily glycosylated, etc. These carbohydrate structures can dramatically alter the protein's surface interactions. In such cases, enzyme immobilization is accomplished by linking the protein to a surface via its carbohydrate chains (Protocol 11). Specific examples are given in the enzyme immobilization protocols described below. The number of free amine groups on a microsphere surface can vary greatly. This variation directly affects the amount of enzyme that may be coupled per unit surface area. For this reason, the type of microsphere used for the immobilization is very important. Polymer microspheres, for example, typically have fewer surface amine groups than amine-labeled silica spheres. To immobilize an enzyme on polymer spheres, the number of amine groups may need to be amplified (Protocol 10). The number of surface amine groups is amplified by reacting the amine-modified microsphere with glutaraldehyde followed by polyethyleneimine (PEI). The resulting amine groups are then reacted again with glutaraldehyde to yield the correct functionality for protein coupling.

Protocol 9

Enzyme immobilization with glutaraldehyde (15, 18) 1

Measure 2-4 mg of 3-5 (am diameter amine-labeled silica or amine-modified polymer microspheres in an eppendorf tube. Clean the microspheres in PBST by five cycles of centrifuging, removing the supernatant, and resuspending in 50 mM phosphate buffer pH 6.9 containing 0.01% Tween-20. 2 Activate the cleaned microspheres by incubating in a 2.5% aqueous solution of glutaraldehyde in 50 mM phosphate buffer pH 6.9 containing 0.01% Tween-20. Approximately 1 ml of this solution is added for every 2-4 mg of beads. 3 Cover the tube with aluminum foil and place on a vortex shaker. Vigorously mix the suspension for two hours at 4°C.

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Protocol 9 continued 4 Remove excess glutaraldehyde by five cycles of centrifuging, supernatant removal, followed by resuspension in ultra-pure water. Repeat the wash an additional five cycles using 50 mM phosphate buffer pH 7.7 containing 0.01% Tween-20. If amine amplification is needed, see Protocol 10. 5 Prepare a 2 mg/ml solution of enzyme in 50 mM phosphate buffer pH 7.7 containing 0.01% Tween-20. Remove the supernatant from the microspheres prepared in the previous step, and then combine them with 1.0 ml of the enzyme solution. Mix and place on a vortex shaker. Allow the glutaraldehyde-activated beads and enzyme to mix for 2 h at 4 °C in the dark. 6 After 2 h, remove excess enzyme solution by washing three times with pH 7.7 phosphate buffer, followed by five times with 50 mM phosphate buffer pH 7.4 containing 0.01% Tween-20. Store at 4°C until use.

Protocol 10

Amine amplification with PEI (12, 13) 1 After step 4 from Protocol 9, microspheres are activated by incubating in a 2.5% aqueous solution of PEI in 50 mM phosphate buffer pH 6.9 containing 0.01% Tween-20. Approximately 1 ml of this solution is added for every 2-4 mg of beads. 2 Mix the suspension for 2 h at 4 °C on a vortex shaker. 3 Remove excess PEI by five cycles of centrifuging and supernatant removal, followed by resuspension in ultra-pure water. Repeat the wash an additional five cycles using 50 mM phosphate buffer pH 7.7 containing 0.01% Tween-20. 4 Proceed with a second addition of glutaraldehyde (step 2-6 from Protocol 9).

Protocol 11

Periodate oxidation (19, 20) 1

Suspend 2 mg of amine-labeled silica or polymer microspheres in 1 ml of 0.13M carbonate buffer pH 9.6 containing 0.01% Tween-20. Wash the microspheres three times by centrifuging, removing supernatant, and resuspending in carbonate buffer. 2 Dissolve 1 mg of enzyme in 130 jol of a 1.68 mg/ml solution of sodium periodate (in PBS pH 7.4). Shake at 4 °C for 20 min. Add 6 jol of ethylene glycol, mix immediately, and shake for 10 min. Add this solution to the microsphere suspension from step 1. Shake for 2.5 h at 4 °C on a vortex mixer. 3 After 2.5 h, prepare a fresh solution of sodium cyanoborohydride (2.5M in carbonate buffer). Add 26 |ol of this solution to the microsphere suspension and shake for 30 min. After 30 min, add 65 |ol of a l.OM solution of glycine in carbonate buffer pH 9.6. Shake the suspension another 30 min.

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Protocol 11 continued 4 Centrifuge the suspension at 5000 x g for 2 min and then remove the supernatant. Resuspend with 1 ml of carbonate buffer pH 9.6 containing 0.01% Tween-20. Save the supernatant to test for enzyme activity. Wash the microspheres five times with 1.0 ml of PBS (pH 7.4) containing 0.01% Tween-20. Test the microspheres for enzym activity as appropriate. Store the enzyme-labeled microspheres at 4 °C in the dark. Note: Reagent volumes and concentrations may be scaled to prepare larger amounts of microspheres. Use caution when handling sodium cyanoborohydride, as it is very toxic. Weigh in a hood. Dispose of all sodium cyanoborohydride solutions after treating with a concentrated solution of iron (II) sulfate heptahydrate.

3.2.3 DNA detection and sequence analysis The Human Genome project is a driving force for the development of new analytical methods for DNA-DNA and protein-DNA interactions. High-density oligonucleotide probe arrays, such as the DNA chip (21), have become popular tools for advanced diagnostic and genetic analyses. Oligonucleotide arrays are particularly applicable to single nucleotide polymorphism (SNP) and genetic defect detection. Oligonucleotide arrays are currently deposited on planar two-dimensional silicon substrates using micro-contact or ink jet printing. Computer controlled arraying robotics place specific probe sequences into an array format on the chip surfaces. Although arrays produced by this method have proven to be very useful in genetic analysis, their production is time consuming and expensive, and they are inflexible in design and applications. Optical fiber based DNA sensors and arrays are amenable to such in situ and in vivo applications. The following sections describe optical fiber based methods for the detection of DNA hybridization and/ or DNA sequence analysis.

3.2.4 A native and fluorescently labeled DNA detection In the optical fiber based method, a single stranded (ss) DNA probe is immobilized onto either a single core fiber or encoded microspheres (12, 13). The nonfluorescent DNA probe(s) are complementary to the target ssDNA, such that fluorescently labeled target ssDNA is detected upon hybridization with the immobilized probe. Fluorescence from the target sequences is quantified using a CCD camera or photomultiplier tube (PMT). Numerous methods exist for labeling DNA, with the most common being to introduce the labels during conventional polymerase chain reaction (PCR) amplification procedures using a fluorescently labeled primer. Alternatively, a reactive group can be incorporated into the DNA sequence, which specifically reacts with the labeling dye. Native ssDNA may also be detected by competition experiments with fluorescently labeled targets (12), which are identical in sequence to the target of interest. Methods for optical fiber based DNA sensor preparation and DNA detection are outlined in the protocols below.

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Protocol 12

Preparation of ssDNA probe labeled microspheres (13) 1

2 3 4 5

6

DNA probes are activated by combining 20 nmol of the 5'-amino-terminal oligonucleotide of interest in 180 [d of 0.1M sodium borate buffer (SEE, pH 8.3) with 40 nmol of cyanuric chloride in 40 |ol acetonitrile. The solution is mixed thoroughly and the reaction is allowed to continue for 2 h at 4°C. Unreacted cyanuric chloride is removed with three cycles of ultra-filtration with a 3500 MW cut-off microconcentrator device. The activated oligonucleotides are covalently linked to the microspheres in step 5. 5 |ol of an encoded bead suspension (Protocol 6 or/and 7) are washed three times with 0.02M phosphate buffer (PBS, pH 7.0). The washed beads are then combined with 150 |ol of 5% glutaraldehyde in PBS and shaken for 1 h. After mixing, the beads are rinsed three times in PBS. Add 150 |ol of 5% PEI (in PBS) to the beads and mix again for 1 h. The bead suspension is then washed three times with PBS buffer and three times with SBB. Add 100 |ol of the 150 |oM cyanuric chloride activated oligonucleotide probe and shake overnight at 4°C. Note: The probe solution can be saved for reuse. The oligonucleotide-functionalized beads are washed three times with SBB. To block any unreacted amine sites, a solution of 0.1M succinic anhydride in DMSO/ SBB (90/10) is added to the beads and shaken for 1 h at 4 °C. The beads are then rinsed three times with SBB and three times with saline sodium phosphate EDTA buffe (6X, SSPE), in which the DNA probe labeled microspheres are stored.

Molecular beacons Molecular beacons (22) are artificially synthesized single-stranded sequences of DNA. In solution, they fold into hairpin-like structures, which results from the hybridization of complementary DNA sequences at the 3' and 5' ends of the beacon. In addition, a fluorescent dye (e.g. fluorescein) is incorporated at the 3' end of the beacon and a quencher (e.g. 4-(4-dimethylaminophenylazo benzoic acid) (DABCYL) is incorporated at the 5' end. Folding of the beacon brings the fluorescent dye and quencher into proximity, resulting in a large decrease in fluorescent emission from the dye. The sequence in between the 3' and 5' complementary regions is called the loop region. The nucleotide sequence in this region is complementary to a DNA target sequence of interest. Complementary hybridization of ssDNA to the loop region can destabilize the hairpin structure resulting in dehybridization of the complementary 3' and 5' ends. Dehybridization causes the separation of the 3' and 5' ends of the molecular beacon, removing the quenching effect of the quencher. Fluorescence signal enhancements (20-50 x ) often result by removing this quenching. The unique design of molecular beacons (Figure 4) allows for the detection of DNA hybridization without the need for fluorescently labeled targets. The ability to detect 145

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non-fluorescentiy labeled targets greatly simplifies the detection and quantification of specific DNA sequences in biological samples.

Protocol 13

Preparation of molecular beacon-modified microspheres (23) 1 Internal and external encoding is performed on the microspheres as demonstrated in Protocol 6 and/or 7. The amine-functionalized microspheres (87% polystyrene, 13% divinylbenzene, Bangs Laboratories, Fisher, IN) are coated with biotin in step 2 after the encoding. 2 An 18 mM stock solution is made with 6-( (6-( (biotinoyl) amino) hexanoyl) amino hexanoic acid, succinimidyl ester in DMSO. This solution is then diluted 1:10 in 0.15M sodium bicarbonate buffer with 0.01% Tween-20 (pH 8.3). 3 300 \jl of the diluted stock solution from step 2 are combined with 2 mg of microspheres, and shaken for 30 min. The microspheres are then washed three times in 0.01M PBST buffer. 4 Microspheres are passivated with a filtered 0.5% solution of bovine serum albumin (BSA). 1.0 ml of BSA solution is added to the microspheres and the suspension is shaken for 3 h at 4°C. The microspheres are then washed three times with 1.0 ml of 0.01M PBST buffer. 5 A 5 mg/ml stock solution of NeutraLite avidin is prepared in 0.01M PBS buffer containing 2 mM of sodium azide. 6 The stock solution of NeutraLite avidin is diluted 1:4 in PBST buffer. 300 (ol of this dilution is combined with 2 mg of beads from step 3. The suspension is mixed on a shaker for 24 h at 4°C. The microspheres are then washed three times in 0.01M PBST. 7 A 33 (xg/ml solution of biotinylated molecular beacons in 0.01 M PBST is prepared. 2 \il of this solution is added to 5 \sl of bead stock, and mixed for 1 h. The microspheres are then washed three times with 0.01M PBST. The molecular beacon labeled microspheres can be stored for up to a year in the 0.01M PBST buffer at 4°C.

Molecular beacon-modified microspheres are used to prepare high-density arrays with application in the detection of multiple DNA targets. Such arrays are prepared as indicated in Section 3.2 and Protocol 5. 3.3 Live-cell array fabrication Living cells can efficiently "sense" their environment and thus are useful as biosensors. Cells are used in many different applications, including water purity testing, detection of metal ions, oxoanions, sugars, and toxic organic compounds, as well as in high-throughput drug screening. For example, some 146

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Figure 4 (a) Diagram of a fiber-optic microarray with molecular beacon (MB) functionalized— microspheres in the hairpin configuration randomly distributed on the fiber where the fluor and the quencher are in close proximity, (b) The same fiber with the MB hybridized to its complement, removing the quencher away from the fluor and generating a signal, (c) Diagram of the array containing three different MB probes and the effect on the imaging system when complements hybridize. The sensing regions are not drawn to scale. Tens of thousands of sensors can occupy an array at one time.

cells are responsive to heavy metals such as mercury or lead, and may be useful for detecting the presence of such metals in environmental water samples. Daunert and co-workers used recombinant bacteria to fabricate a living biosensing system for simple carbohydrates (24). The cell-based system is used as a toxicity screen of antiseptics and antioxidants (25). In these sensor systems, cellular processes are linked to reporter genes, allowing cellular activity to be monitored. While techniques currently exist for monitoring single cells or populations of cells, fiber-optic based arrays of cells allow several hundred individual cells to be monitored simultaneously and repetitively. The cells are encoded with different fluorescent probes (Section 3.3.1) allowing different cell populations within the array to be identified. Dispersing encoded living cells into complementary sized fiber-optic microwells creates cell arrays (Section 3.3.2). Since each well in the array is optically wired within the fiber bundle, cells in the array can be individually addressed and their fluorescence properties recorded over time (26). Once the cells are distributed into the etched fiber, cell viability tests are run to identify living cells (Section 3.3.3).

3.3.1 Cell encoding In order to create a randomly distributed array of living cells, the cells are first encoded with fluorescent dyes. A variety of dyes are available, which can be used to encode the cells through standard protocols (26). The dye serves two purposes: it allows cells to be located within the array, and it allows different cell types to be identified. Because each dye has a different excitation and emission spectrum, 147

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the encoding of the cells provides a unique "fingerprint" of the cell type for cell identification. Although many types of cells can be used, fiber-optic arrays are demonstrated with NIH 3T3 mouse fibroblast cells (26). Subconfluent monolayers of NIH 3T3 mouse fibroblast cells are cultured using standard protocols (27) in 100 mm disposable Petri dishes and subsequently encoded.

3.3.2 Cell array fabrication The array is fabricated from the encoded cells and the etched fiber. Once the fiber is etched (Protocol 5) and the cells are encoded, the cell array is fabricated. The different cell types are combined into one cell suspension, added to the fiber face, and then allowed to settle into the fiber wells. When cells are in solution, they are rounded, easily fall into the wells, and adhere to the well bottom. The first step of Protocol 14 is to prevent medium evaporation, which occurs very quickly due to the small well volume (~85 fl), and results in cell death.

Protocol 14

Fabrication of a fiber-optic based living array of cells 1 An etched fiber bundle is held vertically in a vial filled with DMEM+ culture medium. The vial is capped with a rubber stopper and the whole ensemble is placed in a sonication bath. A vacuum is pulled on the vial while sonicating for 15 min to remove air bubbles. 2 A portion of the cell suspension is drawn into a 1.5 mm diameter capillary tube. 3 The etched face of the fiber is inserted into the capillary tube and secured into place with a thin strip of laboratory film. Care is taken to secure the fiber in the capillary tube quickly so that the medium in the wells does not evaporate. Keep the fiber upright for a minimum of 1.5 h to allow cells to fill the wells on the tip of the fiber. 4 Remove the array from the capillary tube and immediately place in DMEM to avoid evaporation of the medium.

3.3.3 Cell viability After the cells are encoded and the cell array is fabricated, the viability of the cells is monitored. There are several methods to determine cell viability including both commercially available kits and other methods (26). As seen in Figure 5, the different cell viability tests produce different fluorescence readings. Although the resulting live/dead information is the same, the mechanisms of detection are different. One method is to use a LIVE/DEAD Viability/Cytotoxicity Kit, which is available from Molecular Probes (Eugene, OR). This standard protocol kit uses two fluorescent indicators, calcein acetoxymethyl ester (calcein-AM) and ethidium homodimer-1 (EthD-1) to monitor both intracellular esterase activity and plasma membrane integrity. Live cells absorb the cell-permeant calcein-AM, and cleave the lipophilic blocking groups from calcein-AM with nonspecific esterases. 148

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Figure 5 Cell viability methods. Each method uses different detection mechanisms to determine if the cells in an array are alive or dead.

Cleaving the blocking groups creates the fluorescent calcein, which leads to an increase in fluorescence and indicates that the cells are alive. Cell death is indicated when the fluorescence of EthD-1 increases due to binding of the indicator to nucleic acids in DNA. EthD-1 has low membrane permeability, so it only binds to DNA when the cell membrane has been compromised, an indication of cell death. Another cell viability test that also monitors cell esterase activity uses 2', 7'-bis (2-carboxyethyl)-5(and 6)-carboxyfluorescein acetoxymethyl ester (BCECF-AM) (Molecular Probes). BCECF-AM works in a similar manner to calcein-AM, with the lipophilic blocker groups being cleaved by cell esterases to create BCECF. BCECF is fluorescent and indicates that the cells are alive; however, it should be noted that BCECF is pH sensitive, while calcein is not. 149

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Protocol 15

Cell viability assay with BCECF-AM 1 Prepare a solution of 0.1 |oM BCECF-AM in DMEM or serum-free media. 2 Bring the array of cells into focus on a fluorescence microscope system and locate the cell positions via the encoding dye wavelengths. 3 Incubate the array in the indicator solution for 1 min at room temperature. 4 Remove the indicator solution and rinse the array tip with DMEM. Place the array tip in fresh DMEM solution. 5 Excite the BCECF-AM indicator with 495 nm light and monitor the fluorescence at 530 nm. An increase in the fluorescence indicates that the cells are alive, which takes approximately 10 min. A third cell viability test is based on the decrease in pH due to cellular metabolism. This indicator system is not commercially available. In Protocol 16, pH sensitive, nanometer-sized beads are fabricated and distributed into the wells with the cells. The beads are held in place near the cells with a gel coating. The beads do not passively enter the cells, but are in proximity to monitor changes in extracellular pH due to cellular activity. The fluorescent indicator immobilized on the bead decreases in fluorescence as the pH is lowered by cellular metabolism.

Protocol 16

Cell viability test via pH sensitive nanospheres 1 2 3 4 5 6

Rinse 10 jol of 100 nm diameter amino-functionalized polymer beads (1.921 x 1014 spheres/ml) with 10 mM phosphate-buffered saline (PBS) at pH 7.4. Add 300 [d of 4.28 mM fluorescein isothiocyanate dye solution to the washed beads. Allow the dye to react for 30 min. Wash the dyed nanometer beads with PBS buffer three times until all excess dye is removed. Combine 10 [d of pH sensitive beads with 50 jol of 1% w/v alginic acid sodium salt (medium viscosity). Bring the array of cells into focus on a fluorescence microscope system and locate the cell positions via the encoding dye wavelengths. Dip the array tip into the nanometer beads/alginic acid mixture and then immerse in a 0.1M calcium chloride solution to induce gelation of the alginic acid.

7 Once the alginic acid reacts, immerse the array in DMEM. Monitor the fluorescence of the nanometer beads at 495/530 nm. A decrease in fluorescence as a function of time indicates that the cells are alive. A significant decrease should be measurable within 5-10 min.

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4 Conclusions While there are currently many types of optical biosensors, optical fibers provide a versatile platform for either single- or multi-analyte sensing applications. The utility of the optical fiber based biosensors in applications ranging from monitoring biological processes to sensing using multiple analyte arrays is demonstrated in this chapter. Sensors are fabricated to collect visual and chemical data concurrently or to collect multi-analyte data using high-density arrays. Fiber-optic biosensors encounter complications associated with bleaching of the fluorescent indicator, leaching of the sensing elements and indicators, or physical changes to the sensor's composition. Normalizing the data, covalently attaching the sensing elements and indicators, and using the sensors in buffered media address these complications. Advantages to fiber-optic sensors include the ability to perform remote sensing, their lower sensor costs, their flexibility in sensor design, and their miniaturization capabilities. The described sensors have fast response times, which are ideal for real-time monitoring, and have applications in pollution monitoring, process control, and medical diagnostics. The protocols given in this chapter are the foundation for making fiber-optic sensors. By combining these building blocks with ingenuity, biochemical sensors can be made for almost any type of sensing application imaginable.

References 1. Albert, K. J., Lewis, N. S., Schauer, C. L, Sotzing, G. A., Stitzel, S. E., Vaid, T. P., and Walt, D. R. (2000). Chem. Rev., 100, 2595-626. 2. Walt, D. R. (1998). Ace. Chem. Res., 31, 267-78. 3. Bronk, K. S., Michael, K. L., Pantano, P., and Walt, D. R. (1995). Anal. Chem., 67, 2750-7. 4. Michael, K. L., Taylor, L. C., and Walt, D. R. (1999). Anal. Chem., 71, 2766-73. 5. Healey, B. G., Foran, S. E., and Walt, D. R. (1995). Science, 269, 1078-80. 6. Ferguson, J. A., Healy, B. G., Bronk, K. S., Barnard, S. M., and Walt, D. R. (1997). Anal. Chim. Ada., 340, 123-31. 7. Healey, B. G. and Walt, D. R. (1995). Anal. Chem., 67, 4471-6. 8. Pollack, A., Blumenfeld, H., Wax, M., Baughn, R. L, and Whitesides, G. M. (1980). J. Am. Chem. Soc., 102, 6324. 9. Micheal, K. L. and Walt, D. R. (1999). Anal. Biochem., 273, 168-78. 10. Barnard, S. M. and Walt, D. R. (1991). Nature, 353, 338-40. 11. Pantano, P. and Walt, D. R. (1996). Chem. Mater., 8, 2832-5. 12. Ferguson, J. A., Boles, T. C., Adams, C. P., and Walt, D. R. (1996). Nature Biotech., 14, 1681-4. 13. Ferguson, J. A., Steemers, F. J., and Walt, D. R. (2000). Anal. Chem., 72, 5618-24. 14. Walt, D. R. (2000). Science, 287, 451-2. 15. Steemers, F. J. and Walt, D. R. (1998). In Book of abstracts, Vol. 216. The American Chemical Society Washington DC. 16. Michael, K. L., Taylor, L. C., Schultz, S. L., and Walt, D. R. (1998). Anal. Chem., 70,1242-8. 17. Czarnik, A. W. (1997). Curr. Opin. Chem. Bio., 1, 60-6. 18. Walt, D. R. and Agayn, V. I. (1994). Trends in Anal. Chem., 13, 425-30.

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DAVID R. WALT ETAL 19. Szurdoki, F., Michael, K. L, Taylor, L. C., Schultz, S. L, and Walt, D. R. (1998). In Book of Abstracts, Vol. 216. ACS, Boston, MA. 20. Tijssen, P. (1985). Practice and theory of enzyme immunoassays, Vol. 15. Elsevier, Amsterdam, The Netherlands. 21. Carlson, R. and Brent, R. (1999). Nat. Biotech., 17, 536-7. 22. Tyagi, S. and Kramer, F. R. (1996). Nat. Biotech, 14, 303-8. 23. Steemers, F. J., Ferguson, J. A., and Walt, D. R. (2000). Nat. Biotech., 18, 91-4. 24. Shetty, R. S., Ramanathan, S., Badr, I. H. A., Wolford, J. L., and Daunert, S. (1999). Anal. Chem., 71, 763-8. 25. Guan, X., Ramanathan, S., Garris, J. P., Shetty, R. S., Ensor, M., Bachas, L. G., and Daunert, S. (2000). Anal. Chem., 72, 2423-327. 26. Taylor, L. C. and Walt, D. R. (2000). Anal. Biochem., 278, 132-42. 27. Freshney, R. I. (1983). Culture of animal cells: a manual of basic technique, 2nd edn. A. R. Liss, New York.

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Chapter 8 Functional analysis of ion channels: Planar patch clamp and impedance spectroscopy of tethered lipid membranes Michael Mayer, Samuel Terrettaz, Laurent Giovangrandi, Thierry Stora, and Horst Vogel EPFL, Lausanne, Switzerland.

1 Introduction Ion channels play essential roles in cellular processes such as maintenance of the membrane potential, signal transduction, and osmoregulation and are therefore directly or indirectly targeted by many clinically used drugs (1). Despite th importance of ion channels, the molecular mechanisms of their function are largely unresolved. A nearly unlimited number of mutant proteins provided by combinatorial genetics (2, 3) and huge libraries of potentially active compounds produced by combinatorial chemistry (4) offer enormous possibilities for unraveling molecular details of channel function and for finding new medicines. Electrical recordings of ion channel activity provide the most detailed insight into the function of these important proteins (1, 5, 6); however, presently available techniques are not ideally suited for efficient screening of the existing large numbers of compounds. In this chapter, two novel electrical methods for functional analysis of ion channels are presented. Both methods are suited for miniaturization and open the door to fully automated bioanalytical approaches in the micro- and nanometer scale. The first part describes automated formation of planar lipid bilayers (PLB) across (sub-)micrometer-sized holes in silicon wafers. Such bilayers are suited for functional incorporation of ion channels and recording of the ionic flux through single-channel proteins in real time (7, 8). This so-called "planar patch clamp" technique potentially offers substantial advantages compared to traditional patch clamp and PLB recording, both in terms of 153

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extremely facile handling as well as in improved sensitivity. Here, first results from channel-forming peptides will be reported. The second part of this chapter concerns ion channel activity in tethered lipid membranes on gold electrodes measured by electrical impedance spectroscopy. These tethered membranes show exceptional mechanical stability, can be (pre-) fabricated in microarrays, and are ideally suited for the investigation of ligand— receptor interactions by combined electrical and optical detection (9-11). Here we summarize recent experiments performed with natural and artificial ligand— gated ion channels.

2 Planar patch clamp In 1976, Neher and Sakmann introduced the patch clamp technique, which allows recording of the passage of ions through single-channel proteins embedded in the membrane of living cells (12-14). Due to this scientific breakthrough, which was awarded with the Nobel Prize in 1991, an ever-increasing number of diseases could be attributed to a malfunction of ion channels: cystic fibrosis, epilepsy, anxiety, cardiac arrhythmia, and hereditary hearing loss, to mention a few (1, 15, 16). Consequently these proteins are increasingly important as targets for new medicines (6). Patch clamp measurements rely on a stable, electrically insulating contact (seal) between the tip of a glass micropipette and a patch of a cell membrane (17). To resolve ion currents flowing through single-channel proteins, which are located in the membrane patch area, the seal resistance between the pipette and the cell membrane must be in the order of 109 fi ("giga-seal") (18). Seal resistances < 1 Gfi result in current noise, which is higher than the electrical events to be measured (typically 1-100 x 10 ~ 12 A). An alternative method to perform electrical recordings of single ion channels is the so-called PLB, also called black lipid membrane (BLM) technique (19-21). For this approach, ion channels are integrated into a PLB, which is spread across a small hole in a Teflon® septum separating two fluid compartments (22, 23 Ag/AgCl electrodes access both compartments electrically. Like in patch clamp an appropriate high-gain amplifier can resolve the ionic current flowing through ion channels in the membrane. The patch clamp and the PLB technique are extremely valuable tools to stud ion channels and the patch clamp especially is used extensively in fundamental research and in pharmaceutical development of therapeutic compounds. Both techniques, however, require time-consuming manual procedures and are not ideally suited for automation, that is, for the efficient screening of a large number of compounds and channel proteins (6).

2.1 Concept of patch clamp on a chip In the approach described here, planar silicon microchips are used to replace the classical "patch pipet." Instead of manually moving the tip opening of 154

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Figure 1 Principle of planar patch clamp, (a) A concentric electric field E attracts negatively charged vesicles toward the aperture in the diaphragm of the silicon chip, (b) Upon contact with the chip surface, the vesicle opens and spreads due to electrostatic attraction to the positively charged surface of the chip, (c) The result is a single PLB spanning the aperture (see close-up). Ion currents / flowing through membrane-incorporated channel proteins can hence be monitored in real time.

a micropipette to a biological membrane, the membrane is brought in contact with a fixed aperture in a silicon chip. Lipid bilayers are automatically delivered to the aperture by electrophoretic positioning of negatively charged lipid vesicles (Figure l(a)). Due to strong electrostatic interaction (Figure l(b)) the vesicles spread on the positively charged chip surface forming an aperture-spanning planar bilayer with seal resistances up to 200 Gfi (Figure l(c)) (8, 24). Using microchips (Figure 2(a, b)) instead of pipettes has several advantages: (1) The aperture geometry is controlled and diameters between 0.03 and 40 (am can be fabricated reproducibly (Figure 2(c)) (8, 23, 25-27). (2) The thin diaphragm (< 0.2 (am) minimizes the access resistance of the aperture, and therefore its contribution to the electrical current noise (23, 28, 29). (3) The planar geometry of the chips allows the combination with optical techniques, like fluorescence-based single molecule detection (8, 26, 30-33). (4) The compact chip design is mechanically more robust than a patch pipette attached to 155

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Figure 2 (a) Schematic diagram of a silicon chip for planar patch clamp measurements. The fabrication of silicon chips with these characteristics is described in Protocol 1. (b) Scanning electron microscopic image of the sample cavity at the bottom of the chip, and (c) of an aperture with a diameter of 1.2 \im in a diaphragm with a thickness of 150 nm. (d) Schematic side view of the complete recording setup. The silicon chip is mounted on a Sylgard® pad containing a channel that is filled with electrolyte.

a pipette holder. Additionally, the small area of the bilayer increases its stability (23, 28, 34) and reduces the electrical capacity compared to PLB (24, 25). (5) As a future development, recording chips with multiple apertures can be used for parallel measurements (23). 156

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2.2 Formation of planar bilayers on a chip Here we concentrate on the formation of PLBs using unilamellar giant vesicles. For efficient seal formation, the surface of the chips has to interact strongly with the vesicles and the average diameter of the vesicles has to exceed 5-10 (am. Chip production, cleaning, surface modification, and the formation of giant vesicles is explained in the following protocols. With the modified chips and the giant vesicles in hand, the planar bilayers can be formed automatically by electrophoretic vesicle positioning.

2.2.1 Chip fabrication and pretreatment To perform single-channel recordings with high resolution on silicon chips, the microstructures have to fulfill several requirements. The surface of the chips must be suited for surface modification; therefore, a top layer of SiO2 is deposited (Figure 2(a)), which allows chemical (silanes) and physical modification (physisorption of polycations). Additionally, the electrical capacitance of the chips must be as low as possible (< 35 pF) to minimize the capacitive contribution to the overall noise characteristics of the recording setup (8, 23-25, 28, 35, 36). Chips with low capacitance can be produced by adding a relatively thick insulating layer of SiO2 (0.8-1.5 (am, see Figure 2(a)) to the bottom of the chips (24). The capacitance is further reduced by a Sylgard® pad (see Figure 2(d)), which minimizes the contact area between the electrolyte (buffer) and the chip surface (24).

Protocol 1

Fabrication of silicon chips Equipment and reagents Silicon chip processing requires heavy equipment (clean rooms, plasma reactors, furnaces, mask aligner, wet bench, etc); it is assumed that a specialized laboratory will do this work (e.g. GeSiM mbH, Germany; Institut fur Mikrotechnik Mainz GmbH, Germany; MicroFAB Bremen GmbH, Germany, or Paul Scherrer Institut, Switzerland). Equipment and reagents may have to be adapted to the available facilities.

Method Note: The process described below presents only the main steps and does not include steps like the definition of the alignment marks or photoresist deposition and removal. Such procedures will be standard in any chip fabrication facility. 1 Starting material: n-type silicon wafer, thickness of 380 (am, double side polished, electrical resistance of 1 ficm. 2 Formation of a 1.5 (am thick SiO2 film by wet thermal oxidation.

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Protocol 1 continued 3 Removal of the SiO2 on the front side by etching with buffered hydrofluoric acid (BHF). 4 Deposition of a low stress Si3N4 layer by low pressure chemical vapor deposition (LPCVD), thickness of 120 nm. 5 Deposition of a low temperature oxide (LTO) layer (LPCVD SiO2), thickness 1 (am. 6 Removal of oxide (LTO) on the backside of the chip (BHF etching). 7 Photolithography of cavity patterns, backside. 8 Dry etching (reactive ion etching, RIE) of LPCVD Si3N4, backside. 9 Wet etching (BHF) of backside SiO2. 10 Anisotropic terra methyl ammonium hydroxide (TMAH) etching of the silicon down to the Si3N4-LTO diaphragm. 11 Removal of the oxide (LTO) on the front side (BHF etching). 12 Formation of an 800-nm thick SiO2 layer by wet thermal oxidation. 13 LTO deposition (LPCVD SiO2), thickness 40 nm. 14 Photolithography of the aperture patterns, front side. Aperture diameters between 0.6 and 7 (am are recommended. 15 Dry etching (RIE) of LTO-Si3N4-LTO layer, front side. 16 Protection of the chip surface by coating with acetone-soluble photoresist on both sides. 17 Sawing to yield final chip dimensions of 3 mm x 3 mm x 0.38 mm.

Due to strong contamination resulting from the final sawing procedure, we recommend to coat the wafer with an acetone-soluble photoresist (step 16 of Protocol 1). In this way, chips are also protected during transport and can be cleaned before use as described in Protocol 2 or 3. Efficient cleaning is important, since close contact between the lipid bilayer and the chip surface is required to obtain high seal resistances (> 1 Gfi) (17, 37).

Protocol 2

Dry cleaning and activation of silicon chips Equipment and reagents • Oxygen Plasma Asher/Cleaner Tegal Plasmaline 415 (Petaluma, CA, USA)

• Fluoroware® boxes, 2 (Entegris, Chaska, MN, USA).

Method 1

158

Separate the chips from the sawed wafer by cutting the supporting foil with a scalpel blade.

FUNCTIONAL ANALYSIS OF ION CHANNELS

Protocol 2 continued 2 Immerse the chips (still attached to the supporting foil) in a bath of acetone. Chips will lift off from the foil after 10-20 s. 3 Transfer the chips in a new bath of acetone. 4 Rinse the chips with a stream of acetone and transfer them into two additional, fresh baths of acetone. 5 Rinse the chips with a stream of acetone, and dry them with a stream of nitrogen. Store the chips in hermetic Fluoroware® boxes. 6 Clean and activate the chips in an oxygen plasma (pressure of 800 mTorr, power of 100 W) for 2-3 min. In case the chips are not immediately modified according to Protocol 4, they can be stored in Fluoroware® boxes and re-activated using step 6 just prior to surface modification.

Protocol 3

Wet cleaning and activation of silicon chips Reagents • NH4OH solution (28% NH3, Fluka, Switzerland)

• SC-2 solution (6:1:1 H 2 O 2 : 32% HC1)

• SC-1 solution (5:1:1 H 2 O 2 : NH4OH solution)

• Dilute HF (100:1 vol. H 2 O 2 :48% HF).

vol.

H2O : 30%

vol.

H 2 0:30%

Warning: HF is extremely dangerous, take all necessary safety precautions!

Method 1 2 3 4 5 6 7 8 9

Separate the chips from the sawed wafer and clean them in acetone following steps 1 to 5 of Protocol 2. Immerse the chips for 20 min in SC-1 solution heated to 75-80 °C. Rinse the chips with deionized water. Dip the chips for 10 s in dilute HF. Rinse the chips with deionized water. Immerse the chips for 20 min in SC-2 solution at room temperature. Rinse the chips with deionized water. Dip the chips for 10 s in dilute HF. Rinse the chips with deionized water and dry them with a nitrogen stream. In case the chips are not immediately modified according to Protocol 4, they can be stored in Fluoroware® boxes and re-activated using step 8 and 9 prior to surface modifications.

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The cleaning method described in Protocol 2 is recommended, however, if an oxygen plasma cleaner is not available, a wet cleaning protocol (Protocol 3) can be used. Stable planar membranes with giga-seals are obtained by appropriate electrostatic attraction between the chip surface and the negatively charged giant vesicles. Therefore, the chips are pretreated with positively charged poly-L-lysine (see Protocol 4).

Protocol 4

Surface modification of the silicon chips Reagents •

Poly-L-lysine (Mr = 70,000-150,000) from Sigma, USA.

Method 1

Immerse freshly plasma- or wet-cleaned chips (Protocol 2 or 3) in a solution of 0.01-0.1% poly-L-lysine for at least 5 min until use. 2 Rinse the chips thoroughly in a stream of deionized water for at least 1 min. 3 Dry the chips in a stream of nitrogen. 4 The chips can now be stored in air (in a Fluoroware® box) or used immediately for the measurements.

Figure 3 Laser scanning microscopic fluorescence images of giant unilamellar vesicles, prepared according to Protocol 5. (a) Vesicle preparation before removal of vesicles smaller than 5-10 \m\ by dialysis, (b) Vesicle preparation after 18 h dialysis over a nylon net filter. Note that small vesicles inside giant vesicles (multivesicular vesicles) cannot be removed by dialysis. However, these enclosed vesicles do not interfere with the process of vesicle-derived seal formation. The vesicles were stained with TRITC DHPE. Excitation at 543 nm, fluorescence detected using a LP560 nm filter. Scale bars: 50 \im. From ref. (8), with permission.

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FUNCTIONAL ANALYSIS OF ION CHANNELS

2.2.2

Formation of giant unilamellar vesicles

Vesicle-derived lipid bilayers spanning the chip aperture can only be obtained with giant vesicles (Figure 3), which have diameters larger than the aperture. Therefore, the preparation of vesicles with diameters >10 (am is crucial for efficient seal formation. Generally, the formation of unilamellar giant vesicles is promoted by a high percentage (> 10%) of negatively charged lipids (38, 39), by gentle lipid film hydration (40) at elevated temperature (30-70 °C) (41,42), and by hydration of the lipid film in a solution of low ionic strength (41). To adjust the osmotic properties of the vesicles, sorbitol can be added to the hydration solution in concentrations up to 0.4 M. Protocol 5 was optimized to yield vesicles with diameters >20 (am.

Protocol 5

Preparation of giant unilamellar vesicles Equipment and reagents • Rotary evaporator (Rotavapor R114, Biichi, Switzerland). • Sorbitol, asolectin, and cholesterol were obtained from Fluka, Switzerland. l-Palmitoyl-2-oleoyl-sn-glycero-3-[phosphorac-(l-glycerol)] (POPG) was purchased from Avanti Polar Lipids Inc., Alabaster, USA and N-(6-tetramethylrhodaminethio-

carbamoyl)-!, 2-dihexadecanoyl-sn-glycero3-phosphoethanolamine (TRITC DHPE) from Molecular Probes, Eugene, USA. Stock solutions in chloroform methanol (7: 3): 10 mg/ml asolectin; 7 mg/ml POPG; 10 mg/ml cholesterol; 0.1 mg/ml TRITC DHPE. Store the stock solutions at - 20 °C.

Method 1

2

3

4 5

6

Mix the lipid stock solutions (total amount of lipids: 1.25 mg) in a thoroughly cleaned 10 ml round flask to a molar ratio of 63.7% asolectin, 25% POPG, 11% cholesterol, and 0.3% TRITC DHPE. Dry the lipids at 45 °C in a rotary evaporator at 500 mbar until no more solvent is visible (this takes usually less than 30 min), then dry at room temperature at 20 mbar for at least 2 h to obtain a sufficiently solvent-free, homogeneous lipid film. Pre-hydrate the lipid film for approx. 2 min with a stream of nitrogen, which is enriched with water vapor. To obtain N2 enriched with water, bubble N2 through a heated flask (80 °C) filled with deionized water. Hydrate the lipids with 10 ml of 0.2M sorbitol in deionized water overnight at 37 °C. Harvest the turbid and slightly rose-colored vesicle cloud with a pipette (avoid excessive shear stress). The total volume of the harvested cloud varies between 1.5 and 4 ml. Store the vesicles at 4 °C.

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2.2.3

Size separation of vesicles

Even an optimized protocol for the formation of giant vesicles yields a heterogeneous size distribution of vesicles (Figure 3(a)). Vesicles < 5 (am diameter inhibited seal formation presumably due to contamination of the highly attractive chip surface (these vesicles diffuse faster and are present in large numbers). Therefore, a size separation method is required to remove vesicles with diameters < 5 (am. Efficient and gentle size separation of giant vesicles (see Figure 3(b)) is obtained by dialysis. The raw vesicle suspension (1 ml) is filled into a tube made of a nylon net filter (20 (am mesh size, Millipore, Bedford, USA) and dialyzed at 4°C for at least 16 h in 0.6 1 of the same sorbitol solution as used for lipid film hydration (0.2 M). The sorbitol solution is agitated gently (to avoid excessive convective flow through the nylon mesh) during dialysis.

2.2.4

Automated seal formation

By using negatively charged vesicles, properly directed electrical fields can provide precise electrophoretic positioning. Here, concentric electrical fields (see Figure 4) are used for positioning, because they result in a focused movement of charged objects toward the point of the highest electric field. A concentric electrical field is created around the small aperture in the chips upon application of a potential between the cis and the trans compartment (Figure 2(d)). By accessing the buffer compartments with Ag/AgCl electrodes, the main voltage drop after application of a potential occurs within and near the aperture. The resulting radially symmetrical field directs the electrophoretic movement of charged objects to the spatially fixed aperture, as illustrated in Figure 5(a). Upon delivery of a giant vesicle to the aperture, the strong electrostatic interaction with the chip surface leads to vesicle spreading (Figure 5(b)) (8). The result is a solvent-free unilamellar lipid bilayer spanning the aperture. Reconstitution of channel-forming peptides or ion channels into this planar bilayer allows single-channel recording with a current resolution superior to standard PLB recordings and comparable to patch clamp.

Protocol 6

Formation of PLBs on a silicon chip Equipment and reagent • Bilayer Lipid Membrane Amplifier data acquisition card, National InstruBLM-120, equipped with a 10 mV/pA headments Corp., Austin, USA). stage, Bio-Logic Science Instruments SA, • Buffer: 85 mM KC1, 2 mM Hepes pH 7.4 Claix, France. (adjusted with KOH). • Oscilloscope (e.g. TDS 210, Tektronix, Inc., USA) or PC (equipped with AT-MIO-16E-1

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FUNCTIONAL ANALYSIS OF ION CHANNELS

Protocol 6 continued

Method 1

2 3 4 5 6 7 8 9 10

Put the chip with the cavity side upward in a clean Fluoroware® box. Place a Sylgard® pad (see Protocol 7) on the chip such that the hole in the pad is positioned over the cavity. Fill the cavity with a droplet of buffer. Fill the liquid channel below the chip (trans compartment, Figure 2(d)) with buffer (electrolyte). Place the chip on the channel with the cavity side facing the channel. Add 10 [d of buffer to the as side of the chip. Immerse the Ag/AgCl electrodes in the as and trans compartment and connect them to the amplifier. Adjust the voltage offset and evaluate the series resistance of the chip. Apply —60 mV to the as side. Add 1-4 |al dialyzed vesicle suspension to the as side. After seal formation (see Figure 5(b)) wait for 1 min to evaluate the seal resistance and the stability of the planar bilayer.

AITSYS 5 . 5 . 3

Figure 4 Finite element simulation of the electric field E around a 2 jxm aperture in a diaphragm with a thickness of 100 nm and an applied potential of 100 mV. The gray part in the center of the aperture indicates that the electric field exceeds IT.OOOVm" 1 . Due to the walls of the cavity on the bottom side of the chip (Figure 2 ( a ) ) , the electric field on each side of the chip is slightly different. The simulation was done with ANSYS software from ANSYS Inc. (Canonsburg, PA, USA).

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11 If desired, the buffer in the as and trans compartment can be replaced by a buffer with higher ionic strength (e.g. 0.5 M KC1, 10 mM Hepes pH 7.4). 12 After a seal resistance >1 Gfi and a stable baseline is reached, reconstitute channelforming peptides or proteins of interest. 13 Depending on the current amplitude and the kinetics of the recorded current traces, the low-pass filter of the amplifier should be adjusted to 0.1-10 kHz. A good starting range for the filter cutoff frequency is 1-3 kHz (the sampling rate should be at least four times higher than the cutoff frequency) (43). More detailed information about data acquisition can be found in literature (44) or online (http:// www.axon.com/MR_Axon_Guide.html) in the "Axon Guide" (45).

Figure 5 Electrophoretic vesicle positioning in two different configurations, (a) Coulter counter-like current modulations /(t) due to small vesicles (undialyzed sample) passing through the aperture of a chip with a "non-attractive" surface (unmodified Si3N4 diaphragm), (b) When negatively charged and dialyzed giant vesicles are added to a positively charged chip surface, an abrupt step to zero current is observed due to closure of the aperture by a single bilayer. Resulting seal resistances are typically 1-200 GS1. From ref. (8), with permission.

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FUNCTIONAL ANALYSIS OF ION CHANNELS

Protocol 7

Fabrication of Sylgard

pads

Equipment and reagents • Oven at 60 °C

• 30 and 90 mm plastic Petri dishes.

• Sylgard® 184, base and curing agent (Dow Corning, Midland, USA)

Method 1 Prepare ca. 6 g of Sylgard® elastomer in a 30 mm Petri dish, with a base to curing agent ratio of 10:1. 2 Devide the elastomer into two 90 mm Petri dishes (~3 g each). Store this sample for 30 min at room temperature to allow complete coverage of the bottom and removal of air bubbles. 3 Cure the elastomer at 60 °C overnight. 4 Cut the desired patterns (ca. 7 x 7 mm) with a scalpel blade. Punch a hole (diameter ca. 1.3 mm) in the center of the pad using a clean metal tube with a sharpened tip opening. 5 Peel off the pads with tweezers when needed.

Chip-based planar bilayers: single-channel measurements of alamethicin pores Ion channels are only functionally active if they are embedded in an appropriate lipid membrane. An important parameter influencing ion channel function is the chemical composition of the lipid bilayer (e.g. the proportion of cholesterol or charged lipids) (46). PLB experiments not only make it possible to study the function and the characteristics of reconstituted single ion channels but also to assess the effect of the lipid composition on ion channel activity. Additionally PLBs are very useful to study those ion channels that are not accessible with patch pipettes, like channels located in intracellular membranes or ion channels from bacteria. Although there is extensive BLM literature dealing with the functional reconstitution of ion channels into planar bilayers (46-51) this task requires experience in handling fragile membrane proteins. The principle of single-channel recording can, however, be demonstrated with channel-forming peptides or self-integrating ion channels. Both, channel-forming peptides and self-integrating ion channels have recently gained increasing attention for biosensor development (8, 37, 52-58). Examples of channel-forming peptides are melittin (a component of the bee venom) and the antibiotic alamethicin (Figure 6(a, b)). These peptides self-integrate into bilayers and form voltage-gated ion channels by oligomerization (Figure 6(c, d)) (59-61). In the case of alamethicin, well-defined 165

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Figure 6 Alamethicin structure and channel formation, (a) Alamethicin structure according to Fox and Richards (59), showing an a-helical structure of the peptide. (b) Space-filling model of the same structure, (c) Model of voltage-gated alamethicin activation (59). Upon application of a voltage V, alamethicin peptides integrate in the lipid bilayer. (d) Aggregation of alamethicin peptides leads to the formation of pores. The pore diameter (and consequently the single-channel conductance) increases with increasing number of assembled monomers (60, 61).

Figure 7 Chip-based single-channel recording of an alamethicin pore, (a) The l-t diagram recorded at a potential of -130 mV in 0.2 M KCI and 8 mM Hepes pH 7.4 shows the typical activation pattern comprising up to seven conductance states. The dotted line represents the current baseline while the channel is closed, (b) Current histogram derived from alamethicin traces recorded under conditions given in (a). The conductance states Oi to 07 are shown. C reflects the current baseline. From ref. (8), with permission.

multiple open states of single pores are found resulting in up to nine conductance states (21, 62, 63). Figure 7 shows typical alamethicin events recorded with a vesicle-derived lipid bilayer on a chip. Upon seal formation, alamethicin was added to the cis side of the bilayer (final concentration in the cis compartment: 3 ng/ml) and after 10-50 s voltage-dependent activation with up to seven conductance states was observed (64). 166

FUNCTIONAL ANALYSIS OF ION CHANNELS

Single-channel recording provides unique information about the function and the kinetic properties of ion channel proteins. Important parameters, which are accessible by PLB or patch clamp experiments, are: (a) the single-channel conductance (related to the amplitude of the ionic current flowing through a single open channel) (b) the existence of sub-conductance levels (c) the open state lifetime (d) the closed state lifetime (e) and the opening and closing probability. The single-channel conductance is an important parameter since mutated channels might have an altered conductance, which can cause diseases, like a subtype of cystic fibrosis (14). The open and closed state lifetimes and consequently the opening probability might depend on the applied voltage as in the case of voltage-gated ion channels. On the other hand, ligand-gated ion channels have a higher opening probability in the presence of specific ligands (e.g. the neurotransmitter acetylcholine increases the opening probability of the acetylcholine receptor) (17). The third category of ion channels comprises mechanosensitive channels, whose opening probability is altered by mechanical stress applied to the cell membrane (65). In the case of alamethicin, the opening probability of ion channels increases strongly with increasing applied potential, and the single-channel conductance of the various open states are listed in Table 1. As the example of alamethicin demonstrates, single-channel recordings can yield detailed information about the function of channel proteins. Table 1 Conductance states of alamethicin pores, measured at - 170 mV in 0.5 M KCI in an asolectin/POPG/cholesterol (70%: 25%: 5%) bilayer, compared with previously reported conductance values in asolectin and l,2-diphytanoyl-sn-glycero-3-phosphocholine (DphytPC) bi layers Open state

Conductance (planar patch clamp) (pS)

Conductance (PLB) in bilayers of Asolectin (pS)a

DphytPC (pS)b

1

129

40

88

2

423

370

499

3

0

824

880

7

1430 0

1868 8

1990 0

2611 1

4

1247

5

1694 4

1160

(Votes a

According to Schlue and Hanke, recorded at +100 mV (21).

"Accoring to Sansom, recorded at -125 mV (63).

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Ion channels take part in vital functions of the cell and we only begin to understand their involvement in genetic and autoimmune diseases (1). We believe that the planar patch clamp technology presented here will open new possibilities to investigate the function of ion channel proteins. Due to the miniaturized format, the automated formation of giga-seals and the potential for parallel recordings, this chip technology should be particularly useful for high-throughput assay formats.

3 Impedance spectroscopy of tethered lipid membranes There is a fast growing interest in the realization of long-lasting, stable lipid layers on solid supports for application in molecular (bio)electronics and (bio)sensors, as well as for fundamental studies using surface-sensitive techniques (10, 66, 67). In cases where only the physical and chemical properties of a lipid membrane/water interface are of interest, supported lipid monolayers are the system of choice. Such lipid monolayers are most easily produced by self-assembly on hydrophobic supports (e.g. gold surfaces covered by a monolayer of thioalkanes or silanized oxide surfaces). After incorporating certain receptor molecules in the outer lipid monolayer, specific ligand-receptor interactions may be detected by measuring changes of the optical or the electrical properties of the molecular layer (9, 68). A classical example where ligand receptor interactions using biological components were measured on the lipid membrane surface, is the binding of cholera toxin to its ganglioside receptor (9). This concept was some time later adapted and commercialized by a surface plasmon resonance biosensor company (69). In this context, selective detection of Ca2+ by ionophores in a supported hybrid lipid layer is reported. Tethered lipid Mayers are required to preserve the function of integral membrane proteins. In addition to their biological importance, ion channel proteins are particularly interesting in the field of biosensors because they provide an intrinsic amplification of ligand binding (52, 53). The modulation of the electrical resistance of a tethered lipid bilayer by ligand binding to a particular membrane receptor protein is illustrated with the pore-forming protein OmpF. Antibody detection with tethered lipid layers containing a novel synthetic ligand-gated channel (SLIC) is described at the end of this chapter (70). The formation of molecular layers, which are attached chemically to a gold electrode either by the phospholipids or by the proteins, is of particular importance and techniques to create hybrid lipid layers and tethered lipid bilayers are described in detail in the different protocols.

3.1 Basics of impedance spectroscopy The use of a supporting inert electrode excludes a simple amperometric measurement as presented above, unless ions can be reduced or oxidized at the electrode. Experiments with redox proteins on conductive supports have been reviewed recently (71). In the absence of faradic reactions, molecular interactions can be investigated with alternating current methods. The basics of electrical 168

FUNCTIONAL ANALYSIS OF ION CHANNELS Table 2 Definitions of the impedance Z and of the admittance Y.

Resistor Capacitor RC parallel RC series

Notes "Real part of the impedance. "Absolute value of the imaginary part of the impedance. °Real part of the admittance. d

Imaginary part of the admittance.

The impedance Z is defined as the ratio of the applied voltage to the resulting current, where Vm is the amplitude of the voltage, /m the amplitude of the current, 6>m the phase delay, uj, the radial frequency and / the imaginary number ( — I)0'5.

impedance spectroscopy have been already discussed in this series (72). We will therefore concentrate on aspects related to supported lipid membranes. In a typical impedance spectral analysis, a small-amplitude sinusoidal voltage is applied between two electrodes at successive frequencies and the current response is measured. If the response is linear, the resulting current has the same frequency as the stimulus and the ratio of the amplitude Vm/Im is constant at a given frequency. The impedance Z, and the admittance Y, are usually described by complex numbers, whose real and imaginary parts represent their components for a phase delay of 0° and 90°, respectively (Table 2). The convention of plotting the absolute value of the imaginary part of the impedance is used throughout this text (the imaginary part of the impedance of a capacitor is actually negative). The impedance depends on the frequency and this spectrum can often be fitted by an electrical equivalent circuit composed of resistors and capacitors. The real and imaginary parts of the impedance and admittance of basic circuit elements are given in Table 2. A supported lipid mono- or bilayer is classically modeled by a capacitor and a resistor in parallel. The capacitance describes the dielectric properties of the electrically insulating lipid layer sandwiched between two conductive phases, namely the electrode and the electrolyte. The resistance critically depends on electrically conducting defects in the lipid layers. More complicated models consisting of a larger number of circuit elements or using non-ideal circuit elements such as the constant phase angle element have been proposed but will not be considered here (9, 53, 73). Data analysis and interpretation are central issues in impedance spectroscopy. In particular, there is an intrinsic ambiguity in data fitting in the sense that many equivalent circuits have exactly the same mathematical forms and cannot be distinguished from a single spectrum. Spectra taken under different conditions are required to establish a unique model (74).

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3.2 Measuring technique and electrochemical cell There are several ways to record an impedance spectrum as discussed in detail elsewhere (75, 76). Recent technical developments improved and facilitated such measurements considerably. Impedance modules are now offered on standard electrochemical analyzers such as the BAS 100 from Bioanalytical Instrument and software for sophisticated data analysis is available commercially. One of the major improvements of modern instrumentation, as far as the investigation of insulating molecular layers is concerned, is the extension of the measurable impedance limit to the Teraohm range as in the Solartron 1296 dielectric interface. This is of particular importance if it is desired to decrease the electrode size to the micrometer range. Impedance measurements can, however, be performed by simply using a 2-phase-channel lock-in amplifier such as SR 850 from Stanford Research Systems. Sinusoidal voltages are applied to the electrochemical cell by the internal function generator in a programmable automated frequency sweep. The currents in phase and 90° out of phase with the applied ac voltage are directly recorded on the two channels of the lock-in amplifier and a highly accurate phase-sensitive detection is performed digitally. Our set-up includes a homebuilt current-to-voltage converter with a gain of 104-107 and the lock-in amplifier measures the input voltage. An additional external function generator or a dc offset can be used. It is important to realize that the current, which is proportional to the admittance (Table 2), is actually measured and that the impedance is then calculated. An impedance spectrum will include contributions from the whole electrochemical cell, that is, the electrodes, the interfaces, and the electrolyte. It is good practice to minimize the effect of all elements except the working electrode. Therefore, a large (ca. 1 cm2 effective area) and highly conductive counter electrode should be used. If the electrolyte contains chloride ions, an AgCl-coated Ag wire is recommended. A high concentration of the electrolyte (usually KC1 > 0.1M decreases the effect of the electrical resistance of the aqueous phase and of the interfacial capacitances. Under these conditions, the impedance of a small working electrode (in our case: 3.34 mm2) coated with an electrically insulating lipid layer will dominate the spectrum. An electrochemical system with two electrodes is sufficient because only a negligible current will actually flow through the Ag/AgCl electrode.

3.3 Hybrid lipid layer 3.3.1 Description A hybrid lipid layer consists of a phospholipid monolayer deposited on a hydrophobic monolayer grafted to an underlying support as shown in Figure 8. Monolayers of remarkable electrical insulation have been produced by the chemisorption of thioalkanes of sufficient hydrocarbon chain length on gold electrodes (77). Phospholipids are then self-assembled on this hydrophobic su face. Formation and characterization of thioalkane monolayers and hybrid lipid 170

FUNCTIONAL ANALYSIS OF ION CHANNELS

Figure 8 Schematic view of a two-electrode electrochemical cell with an Ag/AgCI counter electrode and a gold working electrode coated with a chemically attached monolayer of tetradecanethiol and a second phospholipid (POPC) monolayer. An ion-selective electrode is obtained if an ionophore, such as the Ca2+ -ionophore ETH1001, is incorporated in the phospholipid monolayer.

layers have been described in detail in recent reviews (10, 78, 79). Hybrid lipid layers are typically used for investigating the binding of proteins and polypeptides to the lipid membrane surface (80). In the absence of electrical amplification (e.g. the introduction of charges in a low-dielectric-constant layer), electrical (capacitance) measurements are, however, less sensitive than optical methods for the detection of hydrated protein layers at these surfaces. This was shown for the binding of cholera toxin to its receptor ganglioside GM1, incorporated into a phospholipid monolayer by simultaneous surface plasmon resonance and electrical impedance measurements (9). The coating of the thioalkane surface with phospholipids can be performed according to different methods (10, 81). Self-assembly techniques from vesicles or from a detergent solution are distinguished by their simplicity. In the following protocol, a very simple dilution technique from ethanol produces vesicles in situ that will spontaneously cover the hydrophobic thioalkanes in the presence of water. The formation of a mixed layer of phospholipids and ionophores is described hereafter.

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Protocol 8

Preparation of a Ca2+-sensitive hybrid lipid layer electrode Equipment and reagents • Gold electrode • Solution of ca. 1 mM tetradecanethiol (Fluka, Switzerland) in ethanol • Solution of 1 mM palmitoyl-oleoyl phosphatidyl choline, POPC (Fluka) in ethanol

• Solution of 1 mM Ca2+ ionophore, ETH 1001 (Fluka) in ethanol.

Method Incubate a clean gold electrodea in the thioalkane solution in ethanol for about 1 h to form a self-assembled thioalkane monolayer. Overnight immersion results in better insulating layers and is recommended. 2 Remove the excess of thioalkane by washing with ethanol. 1

3 Mount the electrode in the electrochemical cell and cover the electrode with a 1 mM solution of POPC and ETH1001. In our electrochemical cell as little as 10 (il of solution is required. 4 Agitate the solution by using a syringe and wait for 5 min. 5 Dilute with several milliliters of water. a

We used freshly evaporated gold electrodes on glass supports which we transferred immediately from the vacuum chamber into the thiol solution. Cleaning procedures with a plasma discharge or with an oxidizing cleaner such as piranha solution (3:2 H2SO4/H2O2) or chromic acid (saturated potassium dichromate in concentrated sulfuric acid) are recommended if an aged gold electrode is used. Warning: These are dangerous cleaning solutions and care must be taken in solution handling. Methods for preparing gold surfaces are described elsewhere in detail (78).

3.3.2 Impedance spectroscopy data The impedance spectra shown in Figure 9 are typical of an electrically blocking electrode and can be modeled by the equivalent circuit shown in the inset. The capacitor C accounts for both the hybrid lipid membrane and the electrode/ electrolyte interface. Under usual conditions of high electrolyte concentration, the capacitance of the interface is negligible as compared to the membrane capacitance CM. The serial resistor Rs accounts mainly for the resistance of the electrolyte. The resistance R in parallel to the capacitor is dominated by the charge transfer resistance RCT. Its value is very large and cannot be obtained with precision under our measuring conditions because practically no Faradic currents are flowing in this electrochemical cell in the absence of redox ions. The simple model of Figure 9 describes the electrochemical cell equally well when the 172

Figure9 Real (o) and imaginary (G) part of the impedance spectrum of a 3.34 mm2 disk gold electrode covered with a hybrid lipid membrane (composed of a first tetradecanethiol and a second POPC monolayer). The electrical equivalent circuit is presented in the inset. The hybrid lipid membrane and the electrode/electrolyte interface are modeled by the same RC parallel circuit. The capacitance of the membrane, dominates the imaginary part of the impedance spectrum at all measured frequencies while the resistance of the membrane is masked by the extremely high value of the charge-transfer resistance. The resistance of the electrolyte Rs, can be obtained from the real part of the impedance at high frequencies.

Figure 10 Traces of the imaginary part Y, of the admittance of a hybrid lipid bilayer membrane containing the ionophore ETH 1001 measured at 120 Hz as a function of time. The admittance is modulated by the Ca2+ concentration: (a) 10~5 M, (b) 10~4 M, (c) 10~ 3 M,(d) 10~ 2 M, (e) lO^M MgCI2 in the surrounding buffer. The working electrode is a disk n-doped silicon electrode (ca. 1 mm2) covered by a 3.5 nm Si02 layer derivatized by (3,3'dimethylbutyl)dimethyl (dimethylamino)silane. The sensing membrane is formed by self-assembly on the hydrophobic silane monolayer by diluting an ethanol solution of POPC:ETH1001 (1:1) with buffer. From ref. (82), with permission.

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gold electrode is coated with a self-assembled thioalkane monolayer or with thioalkane and an additional phospholipid layer. Therefore, the changes of the measured capacitance can be used to quantify the layer formation. Substantial changes of the capacitance are obtained by incorporating ionophores into the supported membrane. By ion complexation of the ionophore, electrical charges are incorporated in the low-dielectric-constant environment of the sensitive layer. The number of charged complexes and thus the membrane capacitance depend on the ion composition of the electrolyte. Figure 10 shows the real-time response of a membrane prepared by self-assembly of equimolar amounts of POPC and ETH1001 (82). The measured admittance is directly pro portional to the capacitance of the molecular layer at a sufficiently low frequency. The membrane is initially in contact with 0.1 M MgQ2, which is then replaced by a solution of 0.1 M MgCl2 containing the indicated amount of Ca2+ . The capacitance changes abruptly to a new value depending on the calcium content of the electrolyte solution. After washing with the calcium-free 0.1 M MgQ2, the signal returns to its original value. Crown-ethers derivatized with alkyl chains have been used to give K + sensitivity to supported phospholipid monolayers (83, 84). Similarly, the change of capacitance of a chelator-modified gold electrode has been used to detect traces of divalent metal ions (0.5 nM Cu2 + ) (85).

3.4 Tethered lipid bilayers 3.4.1 Description Phospholipid bilayers chemically bound to the gold support are obtained by selfassembly of sulfur containing synthetic lipids. These thiolipids are phospholipids comprising at their polar head groups a hydrophilic spacer, which is terminated by a -SH group (Figure 11) (52, 86-88). The hydrated spacer decouples the lipid bilayer from the gold surface. The resulting aqueous phase between the electrode surface and the lipid bilayer has been designed to accommodate extracellular parts of transmembrane proteins. G protein coupled receptors and ion channels have been reconstituted in a functional active form into these gold supported lipid bilayers (11, 53, 87). Protocol 9 has been optimized for the incorporation of the trans-membrane protein OmpF in tethered lipid bilayers. OmpF is a pore-forming protein from the outer membrane of Escherichia coli whose conductance can be modulated by binding the bacterial toxin Colicin N (53). The proteolipid layer is formed in two steps. First, a mixed monolayer of thiolipid and a short i^hydroxy (or) carboxythioalkane is formed by coadsorption from a detergent solution. The small hydrophilic thiols serve to dilute the thiolipid molecules laterally and to prevent direct contact and thus denaturation of membrane proteins at the gold surface. Second, the proteolipid bilayer is then completed by incubating the monolayer-covered gold electrode with a mixed detergent solution of dioleoyl phosphatidyl choline (DOPC) and OmpF, and finally diluting this solution below the critical micelle concentration of the detergent.

174

FUNCTIONAL ANALYSIS OF ION CHANNELS

Figure 11 Schematic view of a two-electrode electrochemical cell with an Ag/AgCI counter electrode and a gold working electrode coated with an OmpF-containing lipid bilayer, which is tethered to the gold electrode via synthetic thiolipids. The membrane protein OmpF forms ion channels in the bilayer, which are closed by the selective binding of the toxin Colicin N. From ref. (53), with permission.

Protocol 9

Preparation of an OmpF-containing lipid bilayer tethered on a gold electrode Reagents • Gold electrode (45 nm thick gold film evaporated on a glass slide) • Buffer 1 (0.3 M NaCl, 20 mM sodium phosphate, pH 7.4) • Buffer 2 (48 mM octylglucoside (Bachem) in buffer 1)

• Solution of 1 mM sulfanylpropionic acid (Fluka) in buffer 2 • Stock solution of approximately 10 ~ 5 M OmpF in buffer 2 (purification protocol described elsewhere) (89) • Solution of DOPC (0.5 mg/ml) in buffer 2.

• Solution of thiolipids (0.5 mg/ml buffer 2) (synthesis of thiolipids described elsewhere) (87)

Method 1

Leave the gold electrode overnight in a mixture of thiolipids and sulfanylpropionic acid at a molar ratio of 2 :1 in buffer 2. 175

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Protocol 9 continued

2 Wash with buffer 2. 3 Mount the gold electrode in the electrochemical cell and wash again with buffer 2. 4 Add a solution of 0.5 mg/ml DOPC and 5 (xg/ml OmpF in buffer 2 and incubate for about 30 minutes. 5 Dilute stepwise with buffer 1, thereby forming a tethered lipid bilayer at subcritical micellar concentrations of DOPC and OmpF.

Figure 12 Real (O) and imaginary (D) part of the impedance spectrum of a 3.34 mm2 disk gold electrode covered with a tethered lipid bilayer. An extra RC parallel circuit element has to be added in series to the equivalent circuit presented in Figure 9 in order to account for a second time constant. The resistances of the gold/water interface (charge transfer) RCT, of the membrane RM, and of the electrolytic solution Rs, are spectrally well resolved. CM and CDL denote the capacitance of the membrane and the interface, respectively. The electrical resistance RM of the bilayer can be obtained from the difference between the values of Zr in the plateau region around 100 Hz and at high frequencies. From ref. (53), with permission.

3.4.2 Impedance spectroscopy data The impedance spectra of a tethered lipid bilayer are shown in Figure 12 (53). As a main difference to the spectra of Figure 9, two charging processes are recognized. The impedance of the lipid bilayer is well separated from the interfacial impedance. This is believed to be due to the aqueous phase between the electrode and the lipid bilayer. As a main consequence, the resistance of the lipid bilayer is separated from the extremely high charge transfer resistance at the gold electrode and can be measured in the frequency range between 10 Hz and 1 KHz. (A more complex circuit was used in the original literature (53) but left the resistance of the bilayer, R M > virtually unchanged.) The obtained 176

FUNCTIONAL ANALYSIS OF ION CHANNELS

Figure 13 Interaction between OmpF in lipid bilayers and Colicin N added to the surrounding buffer solution. Conductance changes of OmpF in BLMs (•) and in tethered membranes by impedance spectroscopy (G); changes of the SPR signal due to the binding of Colicin N to OmpF in tethered membranes (o). In the impedance experiments, the initial and the final conductances were 1.54 x 10 ~ 3 SI"1 cm2 and 1.35 x 10~3 SI"1 cm2, respectively. From ref. (53), with permission.

resistance depends on the protein content of the supported membrane and a low protein-to-lipid ratio is required. It should be noted that the measured layer resistances are still a few orders of magnitude lower than the electrical resistances reported for PLB experiments (see first part of this chapter). Nevertheless, the modulation of the resistance of supported lipid layers containing a channel or a pore by a biological function, such as ligand binding, has been demonstrated by different groups (52, 53, 90). The conductance decrease of a supported lipid bilayer containing OmpF by the selective binding of the toxin Colicin N is shown in Figure 13. The membrane protein OmpF forms ion channels in the bilayer, which are closed by the selective binding of the toxin Colicin N. The specificity of the effect is confirmed by parallel SPR and PLB measurements.

3.5 Lipid bilayer tethered via surface-attached proteins 3.5.1 Description In order to create a robust device, both the lipid bilayer and the functional molecule might be chemically attached to the supporting gold surface. To immobilize proteins in an oriented manner, functional groups have to be introduced into the protein, which will then be recognized by the complementary binding partner on the sensor surface. Histidine-tags are commonly used to bind proteins to surfaces derivatized with the nitrilotriacetic group (10). 177

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Figure 14 Schematic representation and amino acid sequence (one-letter-code) of a synthetic ligand-gated ion channel (SLIC) chemically bound to a gold electrode and then mixed with a supported lipid bilayer. "X" is used for the amino acid ornithine that allows the synthesis of branched polypeptides. The conductive pathway can be closed reversibly and specifically by the antibody Sp3E9.

Another possibility is the site-specific modification of proteins by biotin, which are then bound to surface-immobilized streptavidin (10, 11). Here we report on the direct immobilization of an artificial protein to a gold surface via cysteine groups. An example of a new class of synthetic ligand (antibody)-gated channel protein (SLIC) made of branched polypeptides is shown in Figure 14. The amino acid sequence is shown in one-letter-code. Conceptually, a SLIC comprises a ligand-binding and a channel-forming region. As a ligand-binding region we have chosen the peptide (NANP)3, which is recognized by the specific monoclonal antibody Sp3E9. As will be demonstrated below, antibody binding modulates the channel activity of SLIC. This opens the door for new highly sensitive immunosensing on the basis of simple electrical detection methods. The NANP sequence is the major B cell epitope of the circumsporozoite protein of Plasmodium falciparum, the parasite causing human malaria. The channel-forming part is built by four melittin peptide segments, which are chemically attached to the branched spacer. The melittin sequence was chosen because of its well-known channel-forming properties (58, 70, 91).

Protocol 10

Formation of a proteolipid bilayer tethered by SLIC to a gold electrode Reagents • Gold electrode (45 nm thick gold film evaporated on a glass slide) . Buffer 1 (0.1 M KC1, 10 mM sodium phosphate, pH 7.4) • Buffer 2 (48 mM octylglucoside (Bachem) in buffer 1)

178

. Stock solution of SLIC (10 ~ 5 M) in 0.1 M KC1, 100 mM sodium phosphate, pH 9.0. • Solution of thiolipids (0.5 mg/ml buffer 2) • Solution of phospholipids (0.5 mg POPC/ml buffer 2).

FUNCTIONAL ANALYSIS OF ION CHANNELS

Protocol 10 continued

Methods 1 Mount the gold electrode in the electrochemical cell. 2 Add a dilute aqueous solution of 10 ~ 8 M SLIC at pH 9 and let the molecules selfassemble for about 1 h.a 3 Add the thiolipid from buffer 2 and wait until a steady-state impedance value is obtained (ca. 1 hour). 4 Wash with buffer 2. 5 Add POPC from buffer 2 and wait at least 15 min. 6 Dilute stepwise with buffer 1 to remove the excess of detergent. a

Note that most membrane proteins have to be introduced from a detergent solution at a neutral pH. A prior coating of the gold electrode with a short hydrophilic spacer is often needed to avoid direct contact of the protein with the gold surface, which might induce protein denaturation.

3.5.2 Impedance spectroscopy data The impedance spectra are qualitatively similar to those presented in Figure 12 and can be fitted with the same equivalent circuit (70). Only the real part of the impedance is plotted in Figure 15. Increasing the thiolipid-to-protein ratio to approximately 2200 increases the membrane resistance, R M > sufficiently so that the charging current of the membrane dominates the intermediate frequency range. The resistances of the solution, of the membrane and of the interface (charge transfer) are then well separated in the impedance spectra. Here, the important finding is the increase of RM in the presence of nanomolar concentrations of the antibody Sp3E9 in the bulk solution. The major part of the response can be reversed by washing with buffer because the fraction of rebound antibodies Sp3E9 is very small at this low density of surface-bound antigens. Unspecific adsorption of IgG to these layers is found to be negligible.

3.6 Highly insulating tethered lipid bilayers for single-channel experiments 3.6.1 Description We recently obtained gold electrode-tethered lipid bilayers with exceptionally high electrical resistances (92). We reached values above 2 x 108 fi, which translate into a membrane resistance of 7 Mficm2. For comparison, the highest electrical resistances of unsupported lipid bilayers have been reported for PLB with values around 10 Mficm2. Due to the low density of defects of the tethered membrane, the effect of a few channels can be resolved thus opening the way to single-channel experiments on this highly stable and versatile platform. In turn a minute quantity of analyte, here antibodies, can be measured which is of great interest for bioanalytics. 179

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Figure 15 Real part of the impedance spectra of a SLIC-containing lipid bilayer at various concentrations of antibody Sp3E9 (CAb, from bottom to top): O M , 1 0 ~ 9 M , 3 x l O ~ 9 M, 10~8 M, and 3 x 10~8 M. The increase of the layer resistance as a function of the antibody concentration in the bulk solution is seen at frequencies between 10 Hz and 1 KHz. From ref. (70), with permission.

The increase in membrane resistance is mainly due to an optimized lipid composition. A new thiolipid consisting of a single phytanoic acid coupled to a hydrophilic spacer terminated by a thiol group is used to tether the first lipid monolayer to a gold electrode (92). The lipid bilayer is then completed by a additional monolayer of 1,2-diphytanoyl-srt-glycero-3-phosphocholine (DphytPC) and cholesterol formed by dilution of a detergent solution of DphytPC/cholesterol (9/1) below the critical micelle concentration of octylglucoside. In order t maintain a high membrane resistance, a very small number of channel molecules should be incorporated in the lipid bilayer. In the experiments reported here, the SLIC of Figure 14 inserts into a preformed highly insulating tethered lipid bilayer at low concentration from the aqueous phase, in analogy to protocols used in classical PLB experiments. SLIC has such a high affinity to the tethered lipid bilayer that no dc voltage is required for its bilayer insertion.

3.6.2 Impedance spectroscopy data The impedance spectra are qualitatively similar to the ones presented in Figure 12 and can be fitted with the same equivalent circuit using similar parameter values. The only difference lies in the substantial increase (more than four orders of magnitude) of the membrane resistance (92). In Figure 16(a) the incorporation of SLIC in a tethered lipid bilayer is reflected in a drop in membrane resistance measured on the real part of the impedance at 0.01 Hz. Upon addition of 0.7 |oM SLIC in the aqueous phase, the layer resistance decreases immediately and saturates in about 30 min. The slow kinetics allows interruption of the bilayer 180

FUNCTIONAL ANALYSIS OF ION CHANNELS

Figure 16 (a) Real part of the electrical impedance (measured at 0.01 Hz) of a tethered lipid bilayer after addition of 7 x 10~7 M (i) of SLIC to the aqueous phase. Single-channel recordings on PLBswere done at -30 mV in 1 M NaCI, 10 mM Tris-HCI, pH 7.4 as described in refs. (24, 58). The single-channel events shown correspond to a conductance level of 90 pS at buffer conditions used for IS. (b) Real part of the electrical impedance (measured at 0.01 Hz) of the tethered lipid bilayer containing SLIC after the addition (i) of 10~6 M of the antibody Sp3E9. From ref. (92), with permission.

insertion process by washing with buffer and thus keeping the number of SLIC molecules in the membrane as low as possible in a controlled way. Single conductance states of up to 90 pS are measured for SLIC in freestanding PLBs. The decrease of the membrane resistance after incorporation of SLIC in tethered lipid membranes shown in Figure 16(a) can be explained by the presence of only about 200 open SLIC channels with a conductance of 90 pS. Selective antibody binding to SLIC in the lipid bilayer increases the membrane resistance as a function of the concentration of the antibody Sp3E9 in the aqueous phase (Figure 16(b)). The observed response corresponds to a total closure of 95 SLIC channels with a conductance of 90 pS. With the presently achieved 181

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signal-to-noise ratio, the closure of a few individual channels could be detected. The kinetics of opening and closing of single-channels, however, cannot be measured with an ac voltage at low frequency. Further optimization such as a decrease of the time constant of the membrane using, for example, microelectrodes will be required to record channel dynamics in tethered lipid bilayers.

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Chapter 9 Protein engineering for biosensors Gianfranco Gilardi Imperial College, London, UK.

1 Introduction Protein engineering is defined here as the ensemble of the methods that allow the modification of the gene that codifies for a protein with the purpose of generating mutants or variants (multiple mutants generated by random mutagenesis) with the desired properties. Traditionally the area of biosensors has been the domain of chemists who already have explored chemical modification of proteins over the past few decades. The advantage of protein engineering over traditional chemical modification lies in its ability to introduce controlled modifications in specific positions. Molecular biology has revolutionized the way we can design proteins for biosensing: it allows a high degree of control over the amino acid sequence by precise variations at the DNA level. This enables one to design properties such as binding, catalytic turnover rates, substrate specificity, and stability. Recombinant DNA technology also allows for the creation of fusions of proteins and peptides with combined functions such as tagging for immobilization and binding or catalysis. The early 1990s saw the first applications of protein engineering specifically for application to biosensors. The increased knowledge of protein structure-function, combined with the larger number of structures available and genes cloned have allowed increasingly wider applications to biosensing. An early example is the engineered S337C mutant of the maltose binding protein where the surfaceexposed serine 337 was mutated into a unique cysteine able to covalently link thiol-specific flurophores such as LANBD (1, 2). These acted as reporter groups for maltose binding that resulted in an increase in the fluorescence emission signal (Figure 1). This approach was soon applied to other binding proteins for detection of other ligands such as phosphate ions (3) and glucose (4), and it has recently been extended to engineered antigen-antibody systems (5). Mutagenesis has now become a widely used strategy, applied to both fundamental and applied fields of research oriented toward a variety of purposes. 185

Figure 1 Rational protein engineering combined with fluorescence labeling of the maltose binding protein (MBP) mutant S337C. (a) A unique cysteine was engineered by site-directed mutagenesis in position 337, where the maltose binding protein undergoes a large conformational change upon binding its ligand. (b) The thiol-specific fluorophores IANBD and acrylodan are covalently linked to the unique cysteine 337 where they act as reporter group for ligand binding, (c) Upon maltose binding, the fluorescence emission at 519 nm of IANBD (on excitation at 480 nm) increases and a binding curve is obtained (see Protocol 17).

PROTEIN ENGINEERING FOR BIOSENSORS

On the basis of these, different branches of protein engineering can be identified. The most traditional branch is that of analytical protein engineering that uses mutagenesis for understanding fundamental issues of protein structure and function of existing proteins. Most relevant to biosensors is rational protein engineering that uses the established knowledge of well-studied proteins to create site-directed mutants displaying the desired functions, and directed evolution that uses random mutagenesis and screening or selection methods to identify variants with new properties. Another emerging branch is the de novo protein engineering that uses the established knowledge on protein folding and structure prediction to create new proteins from first principles. This chapter will cover selected aspects of protein engineering relevant to the area of biosensors. It will illustrate the fundamental concepts and provide the basic protocols to enable the non-expert to design and set up mutagenesis experiments for various purposes.

2 Rational protein engineering Rational protein engineering relies on the existing information about the threedimensional structure of the target protein and the implication of specific residues in its function. When this information is not available, a possible alternative is the directed evolution of the target protein. Directed evolution is based on random mutagenesis and it is most applicable when a suitable selection or screening method can be identified (Figure 2). This approach will (a)

(b)

Structure-based design Analysis of three-dimensional structures/models Site-directed mutagenesis

Random mutagenesis Error prone PCR/homologous gene family

DMA sequencing

Recombination DMA shuffling

Expression

Functional screening of library Selection (in vivo)/ Screen ing (in vitro)

Functional characterization

DMA sequencing

Structure determination

Functional characterization

Structure determination

Figure 2 Flow chart showing the experimental stages involved for (a) rational design and (b) directed evolution of proteins. The need for structural and functional information for the rational design and an efficient functional screening are respectively evidenced for the two approaches to guide the iterative cycles shown by the arrows. Ideally the two approaches are interconnected.

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be described in Section 3, but as shown in Figure 2, it can be used in combination with rational design in series of iterative cycles. When a three-dimensional structure of the protein is not available, rational protein engineering can still be based on a three-dimensional model of the target. The model can be validated by traditional biochemical studies on the native protein or by the lengthy process of establishing the role of different residues by alanine scanning mutagenesis. The final output is the generation of a structural and functional model that allows the identification of functionally important residues to be mutated by site-directed mutagenesis to achieve the desired property.

2.1 Modeling and calculations on protein structures Protein modelling is a discipline in its own right and only aspects relevant to the design of biosensors will be covered in this section. The reader is referred to critical reviews in this area (6-8 and references therein); this section will provide the fundamental principles and protocols necessary for the construction of a three-dimensional model of a target protein and the procedures for calculations on the three-dimensional structures. The models derived by using the procedures described in this paragraph are generally good enough for at least initial engineering studies. Protein modelling generally relies on the vast amount of structural and functional information stored in databases available on the web. Based on these, the non-expert can easily have access to programs that allows them to build threedimensional models of target proteins. One example of such program is the comparative model building service of SWISS-MODEL available at http://www. expasy.ch/. The service allows the user to build a three-dimensional model of a protein based on the amino acid sequence that is submitted by the user by email. The server uses sequence alignment and homology or comparative modelling methods (9) to produce the three-dimensional coordinates of a model of the target protein that are sent back to the user by email in the form of a protein data bank (pdb) file. The standard format of the pdb file allows the user to visualize and study the three-dimensional model with a large number of molecular graphics programs, for example, the Swiss-PdbViewer (9) or Insight II by Biosym/MSI (http://www.accelrys.com/). This latter program was used to produce the three-dimensional protein structures shown in this chapter. The quality of the three-dimensional model built by SWISS-MODEL on the basis of the published structures of similar proteins can then be checked for the deviations from ideal bond geometry, angle, and length. The quality of the three-dimensional model can be assessed using the Biotech Validation Suite for Protein Structures (10; http://biotech.ebi.ac.uk:8400). The output of this server is a series of Ramachandran plots that show the occurrence of the residues in the most geometrically favored regions. The G factor is given as an overall indicator of quality of the stereochemical properties of the model, including torsion angles and covalent geometry. When necessary, the quality of the 188

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model can be improved by energy minimization by using programs such as the module Discover 3 of Insight II. Protocols for the preparation of three-dimensional models for refinement by molecular dynamics and energy minimization are available on the Accclrys web site (http://www.accelrys.com/). Once a reliable three-dimensional structure or model of the protein of interest has been generated, a wide number of possibilities exists to study and interrogate the structure to allow informed and quantitative considerations on the position and functional roles of its residues. The three-dimensional structure can be represented in a number of convenient ways to suit the purpose of the investigation. Figures 3 and 4 show the sequence alignment and the three-dimensional models obtained in this laboratory for the three main human cytochrome P450 (CYP) enzymes. These enzymes are part of an ongoing research program in this laboratory that aims at the construction of optical and electrochemical sensors for environmental and pharmaceutical purposes (11-13). CYP form a large class of haem-thiolate monoxygenases involved in xenobiotic metabolism in human and nearly all living organisms. The three-dimensional models of Figure 4 have formed the basis for protein engineering studies, both by rational design and directed evolution.

Protocol 1

Protein sequence alignment Equipment A standard computer connected to the web with web-browser software.

Method 1

Download the amino acid sequences for the bacterial CYP102 (P450 BM3 from Bacillus megaterium), rat CYP2C5, and human CYP2C9, CYP2C19, CYP2D6, CYP2E1, and CYP3A4 from the SwissProt database (http://ca.expasy.org/sprot). These sequences are chosen on the basis that only the X-ray crystal structure of CYP102 and CYP2C5 are known and published to date. They will be used to validate the results of the three-dimensional models of the unknown human CYP.

2 Submit the sequences in fasta format to Multialign (http://prodes.toulouse.inra.fr/ multalin/multalin.html). This creates a multiple sequence alignment from a group of related sequences using progressive pair-wise alignments as described in (14). The alignment obtained is shown in Figure 3, where the key stretches of sequence involved in substrate binding and catalysis are identified by comparison with the known CYP102 and CYP2C5 structures. 3 Download the pdb files of the three-dimensional X-ray structures of CYP102 and CYP2C5 from the Brookhaven National Laboratory (http://www.rcsb.org) and open them using the SwissPdbviewer program (9) available at (http://ca.expasy.org/spdbv/). Derive the secondary structure elements for CYP102 and CYP2C5 for these structures. 189

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Protocol 1 continued 4 Predict the secondary structure elements for the human CYPs using the 3D-PSSM service available at (http://www.sbg.bio.ic.ac.uk/~3dpssm/). This predicts the secondary structure for the aligned amino acid sequences and it also provides a confidence level for the predictions. Alternatively, the PSIPRED service available at (http://bioinf.cs.ucl.ac.uk/psipred/) can be used, but no confidence score is provided as feedback. 5 Compare the information on the secondary structure available for CYP102 and CYP2C5 with that predicted for the human CYPs to validate the sequence alignments. Intervene editing where necessary.

Figure 3 Sequence alignment between class II cytochrome P450s. The sequences of cytochromes of known three-dimensional structure, the bacterial CYP102 (P450 BM3) and the rabbit CYP2C5, are aligned with the human CYP2C9, 2C19, 2E1, 2D6, and 3A4 of unknown structure to date. The validity of the alignment is supported by the position of the highly conserved structural elements such as the cysteine ligand and the ExxR motif.

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Figure 4 Carbon-a three-dimensional models for the human CYP2D6 (black) and CYP2C9 (grey), superimposed to the three-dimensional structure of rabbit CYP2C5 (light grey). The models were built as described in the text, following the structure alignment shown in Figure 3.

Protocol 2

Creation of three-dimensional models Equipment Computer connected to the web and equipped with the Insight II Biosym/MSI software with the Builder and Discover modules. Other similar molecular modeling software will be equally good, but the procedure will need to be modified according to the specific program used.

Method Step 1. Creation of the three-dimensional model 1 Submit by email the amino acid sequences of human CYP2C9, CYP2C19, and CYP2D6 to the comparative model building service SWISS-MODEL available at the expasy site at http://www.expasy.ch/spdbv/. The server will build models for the submitted sequences by sequence alignment and homology or comparative modeling using a program called PROMOD II (9). 2 The SWISS-MODEL service will email back to you a pdb file along with any relevant information or problems for the sequence submitted. The models are built by the server using the published CYP X-ray crystal structures, mostly the closely related CYP2C5 (17) and the haem domain of CYP102 (15). 3 Submit the pdb file to the WHATCHECK service available at http://biotech.ebi.ac. uk:8400/. This will check the models for any geometry and bond length problems. 4 Address the potential problems, then place the haem from the CYP2C5 structure (17) in the human CYP models by using the SwissPdb viewer program (9) available through the same website as the SWISS-MODEL service.

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Protocol 2 continued 5 Proceed to refinement of the model by energy minimization. Step 2. Energy minimization of the three-dimensional model

6 Load the pdb files into Insight II and open the Builder module. 7 Break the bonds between the haem pyrrole nitrogens and the haem iron, as well as that between the iron and the cysteine sulfur. Bonds are broken by selecting them and pushing delete. 8

Select the consistent valence forcefield (CVFF) with the forcefield/select command and clear the potentials and charges. Note that in Insight II, the residue library has to be set to "cvffa.rlb" by using the session/environmental variables command, before the forcefield CVFF is selected.

9 Replace the iron in the haem with a sodium atom using the atom/replace command of the Builder module. 10 Add the hydrogens to the protein model using the modify/hydrogens command and select a pH of 7, switch off the lone pairs option and set the capping mode to charged. 11 Delete the two hydrogen atoms attached to the carboxyl groups of the haem's propionates. 12 Change the bonds in the outer ring of carbons on the haem from Kekule structures to partial double bonds using the modify/bond command of Builder. Change the bonds of the haem's pyrrole nitrogens to partial double bonds. 13 Fix the potentials, partial charges and formal charges using the CVFF residue library by selecting the forcefield/potentials command. 14 Select the forcefield Extensible systematic forcefield (ESFF) from the forcefield/ select command and clear the potentials but keep the charges. 15 Restore the haem's iron together with the bonds to the pyrrole nitrogens and the cysteine ligand by using the modify/bond command. 16 Assign the ESFF parameters by using the forcefield/potentials command and setting the potential action to fix, the partial charge action to accept and the formal charge action to accept. 17 Change the porphyrin's pyrrole carbons to have an atom potential type of c5p using the atom/potential command. 18 Check the partial charge assignments by using the forcefield/potentials command and fixing the potential action to accept, the partial charge action to fix and the formal charge action to accept. 19 Select the Discover 3 module and start the refinement. 20 Initially fix the haem, alpha carbons and side chains to minimize the hydrogens previously added. Minimize the models for 300 iterations (called steps by the program) followed by 10 fs of molecular dynamics. Further minimize for 300 steps. Then fix the haem and the alpha carbons and minimize the side chains of models for 300 steps. 192

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21 Submit the model to the Procheck service (http://biotech.ebi.ac.uk:8400/), that will check the models for bond length and geometry problems. Residues without geometry problems are then tethered and the haem is fixed before further minimization and molecular dynamics are carried out. This is repeated for many cycles until energy minimized models are obtained. Once the three-dimensional X-ray or NMR structure or a reliable three-dimensional model for the target protein has been obtained, molecular graphics is an invaluable instrument to investigate the structure and decide on the mutants to be constructed. The following protocols provide guidelines on how to display and perform calculations on three-dimensional protein structures, taking as an example CYP102. For example, the three-dimensional structure of the haem domain of CYP102 can conveniently be represented as a ribbon displaying the secondary structure of the carbon-a backbone to allow visualization of the buried haem prosthetic group and the conformational change that occurs upon substrate binding (Figure 5(a)). The structure can be used not only to locate key residues of interest as targets for mutagenesis, but also to perform calculations to interpret or extrapolate principles for design of new mutants. One example is the Connolly algorithm (18) that allows calculatation of the solvent-accessible surfaces of proteins. This can be performed within the Insight II program by choosing the radius of a probe that is rolled over the whole surface of the protein. A radius of 1.4 A is typically chosen to define a sphere that approximates a water molecule. The algorithm provides values of areas (in A2) for the whole or (b) 25-,

Figure 5 (a) Super-imposition of carbon-abackbone of the substrate-free and palmitoleatebound structures of the haem domain of CYP102. The haem and the palmitoleate are shown in CPK, respectively. The cysteines400 (axial ligandtothe haem iron), 62, and 156 are shown in CPK respectively, (b) Example of results from the accessible surface area (ASA) calculated for the three cysteines by using the Connolly algorithm (18) using a probe of radius 1.4 A. Dark and light bars correspond to calculations performed on the substrate-free and palmitoleatebound structures, respect ively. The level of details of the information is shown by the areas for the whole residues (C62, C156, and C400) as well as for the sulfur atom only (C62-S, C156-S, and C400-S).

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re-entrant area for all or for the selected residues. For example, the surface accessibility of the three cysteine residues of the haem domain of CYP102 are shown in Figure 5(b). The following protocols describe the procedure to be followed to display a protein structure using Insight II, to overlay two structures and to calculate the surface accessibility of specific residues.

Protocol 3

Molecular graphics: display and basic calculations on protein structures Equipment This protocol illustrates how to display and perform basic calculations using the program Insight II by Biosym/MSI (http://www.accelrys.com/) installed on a suitable Silicon Graphics workstation, such as an SGI Indigo2 IRIX 6.2. Most of the other molecular graphics program will perform the same calculations, operating in a very similar manner, often using a PC or Mac environment. In all cases, the pdb files containing the three-dimensional coordinates of the atoms of the protein under investigation must be available. The pdb files are available for download from the PDB web site (http://www.rcsb.org/pdb/). The substrate-free (2hpd.pdb) (15) and palmitoleate-bound (Ifag.pdb) (16) structures of haem domain of CYP102 are chosen as examples. The results are shown in Figure 5.

Method Step 1. How to load and display a molecule 1 Open a Unix shell of an SG workstation. 2 Type: >InsightII—You are now in the Insight II environment, Viewer module. 3 Select the Molecule pulldown, >get. 4 Click on file type PDB under the PDB Directory User. Select one of the two following pdb files. 5 2hpd.pdb: substrate-free structure; call the object FREE, 6 Ifag.pdb: palmitoleate-bound structure; call the object BOUND. 7 Set the heteroatom on to display "non-protein" moieties, such as the haem and the substrate, called "haem" (residue 460) and "pam" (residue 465) in the pdb files, respectively. 8 Later, if you want to load the two structures together, use the connect object to select the object you want to move. Click connect object first, and then click on the molecule to be selected. 194

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Protocol 3 continued Step 2. How to display and color selected residues 9 Under molecule pulldown:

10 render to display atoms/residues as CPK, sticks and so on (NB: selecting CPK will slow down the program), 11 display to show or hide part of the structure. 12 colour to selectively color part of your molecule. 13 ribbon turns the backbone in a ribbon. 14 To select part of your structure set Molecule pick level to molecule and type the selected residue behind the molecule name in the command window as "molecule name: residue name or number." For example, type "BOUND:pam" to select the palmitoleate. 15 Using display you can show only the ribbon (set atoms off)', specific residues can then be selectively shown. To color different parts of the molecule choose color pulldown, type: 16 FREE: 1-455 for the substrate-free structure shown in yellow ribbon; 17 FREE: 460,400 for the haem and cysteine axial ligand shown in red and green CPK respectively (note that Insight II does not require a space between commas and numbers); 18 BOUND: 1-455 for the palmitoleate-bound structure shown in blue ribbon; 19 FREE: 460,465,400,62,156 for the haem, palmitoleate, cysteine axial ligand, and surface cysteines 62 and 156 shown in red, gray, green, magenta, and cyan CPK, respectively. Step 3. How to calculate surface accessibility 20 Open a Unix Shell under the Desktop menu. 21 Type >jot and minimize the window: you will need this window to check the value of Connolly surface calculations. 22

Select surface under the molecule menu.

23

Set create on.

24

Select surface type Connolly or van der Waals.

25 Display as Dots at quality Medium. 26 The probe radius can be changed to check the level of exposure of the residues. The surface can be quantified (in A2); values can be seen in jot (open the jot window you should have set up at point A) as molecule name_surf.out. Be careful: the data will be overwritten in the same file every time you calculate a surface, even if different residues or part are selected. Write down the values displayed as these will be reset at each single step. 27 The values found for the three cysteines 62,156, and 400 for the FREE and BOUND structures are shown in Figure 5(b).

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Protocol 3 continued Step 4. How to superimpose molecules 28 To compare conformational changes occurring upon substrate binding, it is possible to use the command superimpose under the transform pulldown, taking as a reference the haem (called "hem" in the pdb files):

29

Set superimposition mode to atom and label mode off.

30

Source spec.: type "BOUND:hem;

31

Target spec.: type "FREE:hem.

32 Execute and then click on end definition and execute to visualize the superimposition the program just calculated. The information line at the bottom should give you the output root mean square deviation (RMSD) in Angstrom. Step 5. How to measure distances 33 Set the measure pulldown to distances. 34 Set the monitor on. 35

Set monitor mode on add.

36 The atom can be specified as described (molecule name:residue number:atom type i.e. BOUND: 400: CB for the C|3 of residue 400) or they can be selected by clicking on the atom with the mouse left button. 37 The distance will be reported on the screen. Step 6. How to replace a residue 38 Select the Biopolymer module from the star pulldown at the top left corner. 39 Select residue pulldown and replace. 40 Type the number of the residue to replace (molecule name: residue number) in the box. 41 Click on residue and select the amino acid desired from the list. Another useful operation that can be performed to guide the design of mutants is the calculation of a map of electrostatic potentials on a protein surface. It is widely recognized that electrostatics play a key role in molecular interactions such as protein/protein, protein/ligand, enzyme/substrate as well as protein/surface interactions in electrochemical or other immobilized systems. Therefore, the characterization of such interactions is often an important element in the rational protein design process. Electrostatic potentials can be calculated using programs such as the DelPhi module within Insight II. This can calculate the electrostatic potentials on the points of a cubical three-dimensional grid that is defined around a given object containing a number of pre-defined point charges. This grid is divided into a solvent and a solute space. The main feature of this system is that the solvent space (water) has a much higher dielectric constant (g) than the solute (protein). This is because the peptide dipoles in the protein have much less orientational freedom than those in the solvent. DelPhi models this system using the 196

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Poisson-Boltzmann equation together with a finite difference approximation (19). Protocol 4 illustrates how to calculate the surface electrostatic potentials for the haem domain of CYP102 using DelPhi. The results are shown in Figure 6.

Protocol 4

Calculation of a map of electrostatic potentials on a protein surface Equipment Same as Protocol 3, with the addition of the module DelPhi to the Insight II software. To perform a DelPhi calculation, a number of variables need to be defined. The dielectric constants for the solute (protein) and the solvent (water) can be set to 2.00 and 78.6, respectively; the ionic strength of the solvent can be set at 0.1 OM or any other suitable value, and the solvent radius of 1.40 A with an ionic radius of 2.00 A. All radii are set to their van der Waals values as follows: H (1.10), C (1.55), N (1.40), O (1.35), S (1.81), P (1.88), Fe (1.95). Only formal charges are taken into account: Glu, Asp ( — 0.50 on each carboxyl-oxygen), Lys (+1.00 on NQ, Arg (+ 0.50 on each of NH1 and NH2), +/ - 1.00 on N- and C-terminus, Fe ( + 2.00), porphyrin nitrogens ( — 0.50), and phosphate oxygens ( — 0.67). The haem propionates taken to be fully protonated. These formal charges are to be implemented manually in a *.crg file (* refers to object name).

Method 1

Load the molecule into Insight II by going to the molecule menu and selecting get. Select pdb as file type and select the molecule from the left-hand side of the menu, select heteroatom and press execute. For the substrate-free of the haem domain of CYP102, use the pdb file Ifag.pdb.

2 Open the Builder module. 3 Open the environment menu and select environmental variables. Select the residue library, select get and press execute. Type cvffa.rlb in the interpretation box and press set and execute. The residue_library = cvffa.rlb should appear at the bottom of the Insight II window. 4 Open the modify menu and select hydrogens. Enter the molecule name, set a pH of 7 and set the capping mode to charged. Press execute: the hydrogens should appear on the molecule in white). 5 Open the DelPhi module. 6 From the DelPhi setup menu pick the boundary option and select full-columbic, press execute. 7

From the DelPhi setup menu select the grid option. Switch on the molecule region option and add the name of the molecule. The grid size boundary space option should be on with a size of 15 A.

8

Select the grid resolution point spacing option and use a grid point size of 1 A. Press execute (a white box should appear around the molecule). 197

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9 This will result in a grid resolution of 1.0 A, with a minimal distance between the molecular surface and the grid boundary of 15 A. Note that for such resolution, the number of grid points increases rapidly with object size, which in turn has its effect on the computing time needed to complete the calculation. The origin is centered on the grid, and not the object. The potential values on the boundary points (edge of the grid) are approximated according to Debye-Hiickel theory. This approximation is denoted as "full_coulombic," as selected in point 6. The need for this approximation stems from an essential feature of an Poisson-Boltzmann computation: it calculates the potential value of each grid point in respect to the surrounding ones. This is not possible for the points that lie on the edge of the grid. Hence the need to approximate their values. However, the quality of the approximation has an effect on the entire computation. The best method for doing this is known as "focussing": first a computation is carried out on a crude grid that encompasses the grid of interest. The results of this computation are then incorporated in a second, more accurate, one. The approximation according to Debye-Hiickel is slightly more crude than the one mentioned above, but was deemed to be sufficient for this application. 10 Return to the DelFhi setup menu and select the solute option. Switch on the charge (current charges) and radius options (vdw radii) and set the solute dielectric to 2. Press execute. 11 From the setup menu, select solvent and set the solvent dielectric to 78.6, the solvent radius to 1.4, the ionic strength to 0.1 or another suitable value, and the ionic radius to 2. Press execute. 12 Select iterations from the setup menu and enter 2000 in the non-linear iterations option, switch on the converge criteria and enter an RMS difference of le-05 and a max difference of 0.0001. Press execute. 13 This will apply the Poisson-Boltzmann algorithm in its nonlinear form with a 2000 iterations limit and convergence limit of 1.0 E — 5; these settings should be more than sufficient to reach convergence. The computed potential energy (kcal mol ~ a e ~ a ) is the total electrostatic energy, which represents the energy that is involved in moving the molecule in a given charged state from vacuum to the solvent environment. This form of potential energy is denoted as "total_plus_grid". 14 Go to the templates option and select charge edit. Switch on the get option and select protein formal. Press execute. 15 Go to the charge edit menu and select the list option. Press execute. A list of the rules used to assign formal charges to the protein should appear in the Unix shell window. 16 Return to Insight II and select the add option from the charge edit menu and for the haem of the CYP102 add the following charges: +2 for Fe and -0.5 for the prophyrin nitrogens (named N A, N B, N C, and N D). Press execute. 17 Go to the charge edit menu and select the list option. Press execute. The rule list will reappear in the Unix shell window, ensure the charges you have added are included. 18 In the DelFhi menu select run, enter an appropriate job name. Press execute. 198

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Protocol 4 continued 19 The output of the computation consists of three files: *.prm that contains the parameter set of the computation; *.atm that contains the atomic coordinates of the object; *.grd that contains the potential values per grid point. 20 Once the DelPhi has completed the calculations, the results are displayed using the Connolly surface of the molecules colored in the DelPhi spectrum.

21 To do this, select the grid menu and go to get. Enter the name of the grid file you just created (*.grd). Press execute. 22

In the Insight II menu, select molecule and go to the surface option. Select the Connolly surface type, enter the name of your molecule, a solid style in display surface, and a low quality surface. Press execute.

23 Once the Connolly surface has been calculated and rendered, go to the molecule menu and select colour. Switch on the attribute surface. Enter the name of your molecule and select grid as the colour method, select the DelPhi spectrum and enter the name of the grid file you created with DelPhi in the scalar grid name option. Press execute. 24 The surface of your molecule should be colored in blue for the positive charges, in red for the negative charges, and in white for the neutral regions.

An example in which electrostatic calculations have played a key role in understanding the interactions between two redox proteins is shown in Figure 6. The haem domain of CYP102 from Bacillus megaterium is able to form an electron transfer complex with flavodoxin from Desulfovibrio vulgaris through electrostatic interactions of a patch of positively charged residues located near the haem ligand cysteine 400 and the negatively charged residues located around the FMN. These are evidenced in the open complex shown in Figure 6( b). The electron transfer complex is stabilized by five H-bonds between the residues Glu 16, Phe 107, Tyr 17, Asp 129, Glu 110 of flavodoxin and HislOO, Arg 131, Ser 108, Gin 109, ArglSl of CYP102, respectively. The model of the complex has guided the engineering of a DNA gene fusion, where the flavodoxin is linked to the CYP102 P450 domain through a suitable loop. The fusion is exploited for the electronic coupling between the P450 domain and the surface of electrodes via the flavodoxin module (12). 2.2 Site-directed mutagenesis On the basis of the knowledge of the structure/function relationship of the protein under investigation, the rational design cycle may begin according to the scheme of Figure 2. Rational design relies on controlled amino acid changes in precise positions. These changes are achieved by site-directed mutagenesis. The pre-requisite for site-directed mutagenesis is the availability of the cloned gene of the target protein and its expression at high levels, typically tens to hundreds of milligrams per liter of culture, of the mutants in a host cell that does not produce the wild-type protein. Site-directed mutagenesis can be used to replace, delete, or insert one or more amino acids. In all cases, a DNA polymerase is used to insert an

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Figure 6 (a) Ribbon diagram of the complex between the haem domain of CYP102 from Bacillus megaterium and flavodoxin from Desulfovibrio vulgaris. (b) View of the open complex obtained by a ±90° rotation of the two molecules to show the interface of contact between the proteins. The Connolly surface shows the electrostatic potentials calculated using DelPhi. Positive potentials are shown in black, negative potentials in grey, and neutral in white with a contour scale of ±5 kcal/mol.

oligonucleotide primer containing one or more mismatches in reference to the wild-type DNA template. Several methods have been developed over the last 15 years to achieve these objectives; all of these have advantages and disadvantages related to their yields, costs, or length and complexity of the procedure. In general, all site-directed mutagenesis methods share the need for a mutagenic oligonucleotide that encodes for the desired mutation(s). Figure 7 shows a schematic diagram for site-directed mutagenesis. The gene encoding for the protein of interest is cloned in a suitable plasmid vector by using unique restriction sites, that are specific DNA sequences recognized by commercially available endonucleases. The dsDNA is then used to produce a ssDNA template either by propagation in a Ml 3 phage (Kunkel or phosphorothioate method) or simply by heath denaturation (polymerase chain reaction, PCR method). The ssDNA template is annealed with the mutagenic oligonucleotide that functions as a primer for the synthesis of the complementary mutant strand by a DNA polymerase. An ideal DNA polymerase should not have a 5'- to 3'-exonuclease or strand displacement activity, so that the mutagenic primer is not displaced once the mutant strand synthesis is completed; moreover, the use of a high fidelity enzyme with only one mis-incorporation per 107 bases will decrease the risk of undesired secondary mutations. The T4 DNA polymerase meets these requirements and for this reason it has been used extensively for site-directed mutagenesis. The T4 DNA ligase is then used to link the nicks at the 5'-ends of the primers and the resulting heteroduplex DNA is propagated by transformation in Escherichia coli. At this point the theoretical 50% efficiency of mutagenesis is lowered by host repair mechanisms, therefore, a selection strategy must be used to improve the efficiency of mutant strand recovery at rates as high as possible (80-90%). Various methods of selection have been developed over the years; here I will illustrate some alternatives including the use of 200

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Figure 7 Schematic diagram showing the overall steps necessary to perform a sitedirected mutagenesis.

uracil-containing DNA (Kunkel method), incorporation of dNTP analogs (phosphorothioate method), as well as two examples of the more recent and widely used PCR method (megaprimer and QuikChange™ methods). The great advantage of the PCR method is that selection is not necessary due to the rapid and exponential synthesis of the mutant strand by a heat stable DNA polymerase.

2.2.1 Kunkel method This is one of the earliest mutagenesis methods developed in 1985 by T. A. Kunkel (20, 21). The procedure consists of a combination of in vitro DNA synthesis and in vivo selection steps. Based on the incorporation of deoxyuridine, the Kunkel method relies on the in vivo degradation of a uracil-containing template strand. It can be used with any vector that can produce ssDNA, requiring the isolation and manipulation of ssDNA and a dut~ung~ Escherichia coli strain. For this reason, the wild-type ssDNA is replicated in vivo by a dut~ung~ Esdierichia coli host strain such as BW313 that lacks the dUTPase (dut~) and uracil N-deglycosidase (ung~) activity, allowing the incorporation of a fraction of dUTP instead of dTTP. This ssDNA is used as template for the in vitro mutagenic oligonucleotide-directed 201

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synthesis of the mutant DNA strand, resulting in a dsDNA. This is then transformed in a dut + ung + Escherichia coli strain, for example, JW101, that will degrade the wild-type template strand that contains uracil (U) instead of thymidine (T).

2.2.2 Phosphorothioate method The phosphorothioate method is purely in vitro, allowing a more direct control of the experimental conditions. It is based on the resistance to restriction enzyme digestion by a mutant DNA strand containing a thiol-modified dNTP analog (22, 23). The mutant strand is synthesized from the mutagenic oligonucleotide in the presence of dCTPaS using a ssDNA wild-type template in the presence of T7 DNA polymerase and ligase. The heteroduplex DNA is then nicked by the Nci I restriction endonuclease that will produce nicks only on the CCCGG repeats of the wild-type strand. The T5 DNA exonuclease is used to completely digest the nicked wild-type strand. The remaining DNA fragments are used as primers for the repolymerization by DNA polymerase I in the presence of T4 DNA ligase, creating a heteroduplex mutant DNA with a mutagenesis efficiency greater than 80%.

2.2.3 PCR methods The method based on PCR is the preferred approach available today. It provides a quick and efficient strategy to generate mutants by amplification of a dsDNA template using synthetic oligonucleotide primers that contain the desired mutation(s). The PCR methods use a heat stable DNA polymerase such as the Taq polymerase from Thermus aquaticus or the Vent from Thermococcus litoralis or the Pfu from Pyrococcus furious. The choice of the best heat stable DNA polymerase depends on the needs of the specific experiment, as different enzymes have different optimal temperature, level of fidelity, and proof-reading activity (Table 1). Most protocols for PCR site-directed mutagenesis are based on the method of Higuchi et al. (24). In all cases the enzyme carries out the synthesis of a complementary DNA strand of DNA from the 5'- to the 3'-end using a ssDNA template, starting from a dsDNA region, where a synthetic oligonucleotide primer has annealed. At least one of the primers must contain the sequence for the desired deletion(s), insertion(s), or replacement(s). Two primers are necessary, each complementary to opposite strands of DNA, initiating the synthesis of the region flanked by them. The elements necessary in the mix for a PCR reaction are an excess of the deoxynucleotides (dNTPs) that provide both the energy and the nucleosides required for the synthesis of the DNA, an excess of the oligonucleotide primers, and a very small amount of the DNA template. The reaction is carried out by first heating the sample to separate the DNA strands (denaturation), followed by cooling to allow binding of the primers on the DNA template (annealing) and setting to the temperature for optimal polymerase activity (extension). The heating/cooling process is repeated in cycles by using an automated thermocycler able to achieve the required temperatures very rapidly. The major advantage of the PCR method is that the new DNA strand containing the 202

Table 1 Properties of the DNA polymerases most often used in protein engineering Polymerases

Organism origin

Optimal temp. (°C)

Error rate ( x l O 6)

Exonuclease activity

Comments

DNA polymerase 1

Escherichia coli CM 5199, a lysogen carrying \polA transducing phage

37

9

5'->3'

Nick translation of DNA

3' —> 5'( proof-reading activity)

Second strand synthesis of cDNA Can withstand up to 95 °C

Hot Tub DNA Polymerase

Thermus ubiquitous

75

Klenow fragment of DNA polymerase

Escherichia coli carrying a plasmid in which two-thirds of the 3'-end of the Escherichia coli DNA polymerase 1 gene (Klenow fragment) is cloned

37

40

3'->5'

DNA sequencing by the Sanger dideoxy method and oligolabeling. In vitro mutagenesis. Generates blunt ends.

T4 DNA polymerase

Escherichia coli carrying a T4 DNA polymerase overex pressing plasmid

37

<1

3' — >5'

Removal of 3'overhangs In vitro mutagenesis Fill-in of 5'overhangs.

T7 DNA polymerase

Escherichia coli BL21 carrying the plasmid pAR1219 that contains the 77 gene under the control of the lacUVS promoter

37

15

—

Site-directed mutagenesis 3'-end labeling.

Taq DNA polymerase

Thermus aquaticus

75

285

—

DNA amplification by PCR DNA sequencing Can withstand up to 95 °C

Tth DNA polymerase

Thermus thermophilus

75

—

DNA sequencing, especially templates with high secondary structure Lacks an intrinsic nuclease activity.

Table 1 (Continued) Polymerases

Organism origin

Optimal temp. (°C)

Error rate ( x l O 6)

Exonuclease activity

Comments Has intrinsic reverse transcriptase activity in the presence of Mn2+ . Can withstand prolonged incubations up to 95 °C

Pfu DNA polymerase

Pyrococcus furiosus

72

1.3

5'->3' 3' —> 5'( proof-reading activity)

VentR DNA polymerase

Escherichia coli that carries the Vent DNA polymerase gene from the archaea Thermococcus litoralis

75

57

3' —>5'( proof-reading activity)

Thermal cycle sequencing.

High temperature dideoxy-sequencing.

PROTEIN ENGINEERING FOR BIOSENSORS Table 2 Exponential growth of the new dsDNA as a function of the number of PCR cycles Cycles

No. of mutated dsDNA

1

0

5

8

10

256

15

8192

20

262,144

25

8,388,608

30

268,435,456

mutagenic primers will accumulate exponentially at each cycle, whilst the wildtype DNA template will remain in trace amounts as initially introduced by the experimentalist. For this reason it is crucial to use only minimal amounts of template DNA. A quantitative example of number of mutant strands generated at each cycle is given in Table 2. Some key aspects of the PCR reaction must be carefully planned. For example, although increasing the number of cycles leads to an exponential increase in the amount of the product, typically 20-25 cycles are sufficient to generate amenable quantities of the products. Increasing the number of cycles also increases the probability that errors are introduced in a random fashion and they are subsequently amplified by the polymerase. Modern PCR thermal cycles are provided with a hot-top assembly that prevents condensation on the thin walls and lid of the PCR tubes. This will cause changes in the reaction conditions that can result in undesired error prone amplification. In the event that no hot-top assembly is available, a typical 50 jol PCR reaction can be covered with 30 [d of mineral oil to achieve the same results. Care must be taken during the following steps in not pipetting any of the oil present on the surface. The design of mutagenic oligonucleotide primers is crucial for the success of the PCR reaction. The parameters to be evaluated in the primer design process are the length of the sequence, the degree of similarity to the template and the presence of secondary structure. The length of the primers varies depending on the number of variations introduced. A primer of 20 base pair (bp) is generally sufficient for a single base mismatch. The primer must be long enough to hybridize the target DNA at a unique position. There are a number of software packages freely available on the web that allow one to automatically check the sites for hybridization of a primer to a given template (see Table 3). A 100% identity over 10-15 bases must be present on both stretches at the 5'- and 3'-ends of the DNA surrounding the mismatch. A full match between template and primer is particularly important at the 3'-end of the primer because the DNA polymerases typically will not extend from a mismatched or poorly hybridized 3'-end. On the other hand, non-complementary bases may be added to 205

Table 3 Some useful web sites for protein engineering and PCR applications Web address

Information

www.expasy.ch

Range of proteonomic tools, from translation to homology modeling

http://biotech.embl-heidelberg.de:8400/

Checking protein models for problems

www.bmm.icnet.uk/~3dpssm/

Recognition of protein folds

www.firstmarket.com

Restriction map generator

http://www.chemie.uni-marburg.de/ becker/welcome.html

Provides useful links to useful software for PCR and primer design

http://www.ebi.ac.uk/software'software.html

Archives of software for molecular biologists

http://endeavor.med.nyu.edu/download/molbio/ http://bioinformatics.weizmann.ac.il/mb/ software.html http://alces.med.umn.edu/VGC.html

Useful for oligonucleotide Tm and protein coding regions

http://bimas.dcrt.nih.gov/sw.html

Useful links to software sites and molecular biology protocols

http://www.yk.rim.or.jp/~aisoai/soft.html and http://iubio.bio.indiana.edu/soft/molbio/ Listings.html

Useful collection of www links for information and services to molecular biologists

http://www.ncbi.nlm.nih.gov/

Links to GenBank® and sequence databases

Table 4 Codon usage by Escherichia co// calculated out of a total of 1,363,279 codons. Each group shows the count, % frequency, and codified amino acid. Note that AUG and UUG are the most frequently used start codons. UAA, UAG, and UGA are stop codons. 3rd position

1st position

2nd position U

C

A

G

U

30,407 (2.23) Phe

11,523 (0.85) Ser

22,048 (1.62) Tyr

7,062 (0.52) Cys

c

U

22,581 (1.66) Phe

11,766 (0.86) Ser

16,669 (1.22) Tyr

8,846 (0.65) Cys

C

18,943 (1.39) Leu

9,793 (0.72) Ser

2,706 (0.20) Stop

1,260 (0.09) Stop

A

18,629 (1.37) Leu

12,195 (0.89) Ser

326 (0.02) Stop

20,756 (1.52) Trp

G

15,018 (1.10) Leu

9,569 (0.70) Pro

17,631 (1.29) His

28,458 (2.09) Arg

U

15,104 (1.11) Leu

7,491 (0.55) Pro

13,272 (0.97) His

29,968 (2.20) Arg

C

5,316 (0.39) Leu

11,496 (0.84) Pro

20,912 (1.53) Gin

4,860 (0.36) Arg

A

71,710 (5.26) Leu

31,614 (2.32) Pro

39,285 (2.88) Gin

7,404 (0.54) Arg

G

A

41,375 (3.03) lie 34,261 (2.51) lie 5,967 (0.44) lie 37,994 (2.79) Met

12,223 (0.90) Thr 31,889 (2.34) Thr 9,683 (0.71) Thr 19,682 (1.44) Thr

24,189 29,529 45,812 14,076

(1.77) (2.17) (3.36) (1.03)

Asn Asn Lys Lys

11,982 (0.88) Ser 21,907 (1.61) Ser 2,899 (0.21) Arg 1,694 (0.12) Arg

U C A G

G

24,910 20,800 14,850 35,979

20,808 34,770 27,468 45,862

43,817 25,996 53,780 24,312

(3.21) (1.91) (3.94) (1.78)

Asp Asp Glu Glu

33,731 (2.47) Gly 40,396 (2.96) Gly 10,902 (0.80) Gly 15,118 (1.11) Gly

U C A G

206

(1.83) (1.53) (1.09) (2.64)

Val Val Val Val

(1.53) (2.55) (2.01) (3.36)

Ala Ala Ala Ala

PROTEIN ENGINEERING FOR BIOSENSORS

the 5'-end of either or both primers. The possibility of these additional mutations offers the opportunity to add unique restriction sites to the PCR products to facilitate subsequent sub-cloning steps. In this case, extra bases must be added at the 5'-end to allow the direct digestion of the PCR product by restriction endonucleases. The length of the primer is also important because it determines the melting temperature, Tm, of the oligonucleotide. This parameter is usually determined experimentally by the supplier of the oligonucleotide, but at the stage of the design of the primers a rough estimate can be made by using the formula: Tm = 4 x (nG + nC) + 2 x (nA + nT),

where nA, nT, nG, and nC indicate the number of these basis in the primer sequence. Most primers exhibit Tm between 55 °C and 65 °C, but ideally the length of both primers should be adjusted to have a Tm as similar as possible. Another important parameter is the presence of a secondary structure in the primers. The presence of a secondary structure interferes with the hydridization of the primer with the template, and therefore should be avoided. It arises from the self-complementarity within the primer that anneals to itself (self-annealing). As this is often caused by a high content of G and C bases, the GC content should not exceed 50% of the total number of bases. A number of software packages are freely available to guide the design of primers and the calculation of Tm (Table 3). Moreover, the codon chosen for the design should be checked against the list of codons used by Escherichia coli, to ensure it is not in the list of the rarely used codons that would impair translation of the gene into the mutant protein (Table 4). Protocol 5 describes a typical PCR experiment.

Protocol 5

Hot start PCR This protocol illustrates a general PCR experiment for amplification of a 1 kb DNA fragment flanked by two oligonucleotide primers n.l and n.2.

Equipment and reagents 1

Plasmid DNA template, 50 ng/nl. This is the typical concentration of plasmid DNA template obtained from a commercial plasmid purification kit such as the QIAprep Miniprep (Qiagen Ltd, UK). As a rule of thumb, the concentration of dsDNA can be estimated from the absorbance at 260 nm using the formula: A2eo x 50 = Hg/ml.

2 The purity of the dsDNA preparation can be estimated from the ratio: A260/A280>1.8.

207

GIANFRANCO GILARDI Protocol 5 continued

3 Oligonucleotide primers at 50 pmol/|ol The concentration of the stock solution of synthetic oligonucletides is provided by the manufacturer. As a rule of thumb, the concentration of ssDNA can be estimated from the absorbance at 260 nm using the formula: A26o x 33 = (Xg/ml. 4 Mix of the four deoxynucleotides, ATP, TTP, CTP, GTP, generally called dNTP mix, at 2.5 mM each. These are commercially available as a mix already made at 20 or 25 mM (Amersham Biosciences Ltd, UK). 5 DNA polymerase at 2.5 U/|ol (U, units). Depending on the specific experiments, different DNA polymerases can be used. Taq DNA polymerase (5 U/nl, Amersham Biosciences Ltd, UK) can be used for standard mutagenesis or error prone PCR experiments. Pfu (2.5 U/nl, Stratagene) or Vent (2 U/nl, New England Biolabs Inc) can be used to achieve DNA amplification with high fidelity. 6 DNA polymerase buffer. A 10 x solution of this buffer is supplied by the manufacturer together with the specific DNA polymerase. 7 PCR block. Various types of PCR thermocyclers are commercially available. They can be programmed to perform the desired number of denaturation, annealing, extension cycles. More sophisticated blocks allow to program gradients of temperatures so that different annealing temperatures can be tested at once. In all cases the use of a heated cover is recommended to avoid condensation of the reaction mixture. When this option is not available, 30 |ol of mineral oil should be added on top of the reaction mixture.

Method For a 50 |ol reaction: 1 Add in a 0.5 ml PCR tube: 1 |ol plasmid template (50 ng) 1 |ol oligonucleotide primer n.l (50 pmol/|ol) 1 |ol oligonucleotide primer n.2 (50 pmol/|ol) 4 nl dNTP mix (2.5 mM each) 5 [d 10 x DNA polymerase buffer 37 |ol water. 2 Mix the reagents and briefly spin in a microcentrifuge to collect the mix at the bottom of the tube. 3 Incubate in the PCR block at 94 °C for 5 min. 4 Chill on ice for 2 min. 5 Add 1 nl of DNA polymerase (2.5 U).

208

PROTEIN ENGINEERING FOR BIOSENSORS

Protocol 5 continued

6

Perform 25 cycles as follows:

7 denaturation: 94 °C for 30 s, 8

annealing: 60 °C for 1 min,a

9 extension: 72 °Cb for 1 min,c 10 At the end of the cycles incubate at 72 °C for 7 min, then place on ice. 11

Load 10 (il of the PCR product on an agarose gel (See Protocol 6) and visualize the bands of the product with a UV transilluminator.

12 When the conditions are optimized,11 load all the PCR products on an agarose gel. After separation, excise the bands from the gel and purify the PCR products from the agarose (see Protocol 7). a

This should be 2 °C lower than the lowest Tm of the oligonucleotide primers.

b

Set the original extension temperature according to the DNA polymerase used (see Table 1). c

This depends on the length of the DNA strand to be synthesized. A typical extension rate is 1 min/kb of target length.

d

lf no PCR product is formed, lower the annealing temperature 2-3 °C from the previous set value and run the reaction again. If nonspecific PCR products are present in the form of many bands or smear on the agarose gel, raise the annealing temperature by 2-3 °C and run the reaction again, until a clean band of the desired product is observed.

Protocol 6

Agarose gel electrophoresis of DNA Submarine agarose gel electrophoresis is a widely used method for the separation and purification of DNA fragments of 100-10,000 bp in length. The range of the separation is determined by the percentage of agarose used in the preparation of the gel (Table 5). The DNA is visualized with ethidium bromide, a highly toxic compound; all operations must be carried out using gloves and with the precautions listed by the supplier.

Equipment and reagents 1

50 x TAB buffer stock solution: 242 g Tris, 57.1 ml concentrated acetic acid, 100 ml of 0.5M EDTA at pH 8.0, make to 11 with water. To obtain a 1 x , dilute 20 ml of the 50 x TAB stock with 980 ml of water. This corresponds to 40 mM Tris-acetate, 1 mM EDTA, pH.

2

Stock ethidium bromide solution, 10 mg/ml. 209

GIANFRANCO GILARDI Protocol 6 continued

3 1% agarose gel stock solution: 1 g agarose in 100 ml of 1 x TAB buffer. Other agarose percentage can be prepared varying the amount of agarose. Generally the solution is prepared in heat-resistant bottles, so that when the gel is solidified it can be re-dissolved warming the bottle in a microwave oven. The agarose stock solution will be usable for 1 week. 4

6 x loading buffer: 0.25% (w/v) bromophenol blue, 40% (w/v) sucrose. Make to volume with water. 5 Submarine gel tank, for example, the GNA100 by Amersham Biosciences Ltd, UK.

Method 1 Place masking tape to block the two open edges of the gel tray. 2 Mix a sufficient volume (12 ml for a 5 x 7.5 cm standard gel tray with 25 jol wells) of 1% melted agarose stock solution with 2 jol of stock ethidium bromide in a 15 ml tube. 3 Pour the gel in the tray and place the comb. 4 Allow the gel to cool and set. 5 Remove the masking tape and place the gel tray in the gel tank. 6 Fill the tank with 1 x TAB buffer until the gel is fully covered by a 3 mm layer of buffer. Gently remove the comb avoiding formation of air bubbles in the wells. 7 Prepare the DNA samples by mixing with the appropriate volume of 6 x loading buffer. For a typical 25 jol well, mix in a tube 20 jol of DNA sample with 4 jol of loading buffer in a tube. Use a suitable pipette to load the mix at the bottom of the well. Also prepare suitable molecular weight markers to be loaded for reference. 8 Run the samples at a constant voltage of 120 V for approximately 30-40 min. 9 To visualize the separation, place the gel tray on a UV transilluminator (use a protection shield). The DNA will be made visible by the intercalated ethidium bromide and appear as bright orange bands.

Table 5 Range of separation of agarose gel electrophoresis

210

% Agarose gel

Optimum resolution of linear DNA (kb)

0.5

30-1.0

0.7

12-0.8

1.0

10-0.5

1.2

7-0.4

1.5

3-0.2

2.0

1-0.1

PROTEIN ENGINEERING FOR BIOSENSORS

Protocol 7

Purification of PCR products from agarose gels Equipment and reagents Isolation of PCR products and of DNA in general from agarose gels is a straightforward procedure which is greatly facilitated by the availability of commercial kits that allow the whole procedure to be carried out in less than 20 min. Some examples of these kits are the GENECLEAN by BIO101, the QIAquick by Qiagen, and the Wizard by Promega. Low melting point agarose must be used for this procedure, as all the kits rely on melting the agarose and binding the DNA to silica in high concentrations of a chaotropic salt. This is followed by a washing step and elution of the DNA in low salt or pure water. It is thought that high concentrations of chaotropic salt disrupt the water molecules structured around the negatively charged silica, allowing the formation of a cation bridge between the silica and the DNA phosphate backbone. Removal of salt causes rehydration of the silica and release of the DNA. Commercial kits are capable of to isolating a wide range of sizes of DNA, from less than 100 bp to more than 10 kbp. However, different sizes of DNA will bind to silica under various salt concentrations, pH, and wash conditions, therefore it is recommended to look carefully at the different kits available and assess which is most suitable for the particular size of DNA you are using.

Method There are the many variations on the procedure, depending on the kit used. However, all kits proceed as follows: 1

Excise the DNA band from the agarose gel stained with ethidium bromide and transfer it to an eppendorf tube. This must be done very rapidly using a razor blade over a UV transilluminator. As the UV light is mutagenic, the speed of the process ensures that no unwanted mutations are introduced at this stage on the DNA to be used. An appropriate face shield must be used.

2 Weigh the excised band to determine the volume of chaotropic salt to be added. Generally DNA will bind to silica when the salt concentration is above 4M and the pH is kept between 6.0 and 7.4. Smaller DNA strands (< 200 bp) bind at lower pH, therefore in this case 10% acetic acid can be added to keep the pH around 6.0-6.5. For example, the GENECLEAN kit provides a 6M Nal solution and its protocol requires 3 volumes of this solution to 1 volume of agarose. 3 Melt the low-melting temperature agarose by incubating the tube at 55 °C in a water bath for 5 min. Mix the content of the tube to help melting. 4 Add the silica suspension. Generally this is a 1:1 water: silica suspension provided in the kit. For example, GENECLEAN provides the so-called GLASSMILK. Take care to resuspend the silica before pipetting 10 [d of the GLASSMILK. This should be more than sufficient, as typically 10 [d of GLASSMILK will bind 1-2 [ig of DNA.

211

GIANFRANCO GILARDI Protocol 7 continued

5 Incubate 5 min at room temperature to allow the DNA to bind to the silica. Mix every 1-2 min to ensure that the silica remains resuspended. 6 Separate the DNA bound to the silica from the bulk solution. This can be done either by spinning the tube 5 s at full speed in a microfuge, or by using small spin columns provided with the kit. In the former case the silica must be carefully recovered at the end of the tube, in the latter it is retained by the spin column. The QIAquick kit columns already contain the silica in the column itself. 7 Wash the silica with a high salt concentration solution. This varies between the kits. For example, GENECLEAN provides a concentrated solution of NaCl, Tris, and EDTA with 50% ethanol. The silica must be washed 2-3 times with 10-50 volumes of wash solution. When spin columns are provided, they can either be inserted on an eppendorf tube and spun, or they can be fixed on a vacuum manifold apparatus and the wash removed by suction. 8 Elute the DNA. Add 50 (il of pure sterile water or TEA buffer to the silica, wait 2 min, then spin the columns on clean and sterile eppendorf tubes to recover the solution containing the DNA. Usually more than 80% of the DNA is recovered in one step. This is generally sufficient, as a second elution will result only in an additional ^10% of DNA. The silica and the columns are then discarded.

PCR megaprimer method The megaprimer PCR is a widely used method for site-directed mutagenesis. Generally three oligonucleotide primers are used in two steps of amplification (Figure 8). The first step uses one mutagenic and one flanking primer. The mutagenic primer contains the mismatches necessary to introduce the desired mutation(s). The flanking primer should be designed to include a restriction site (R1) to allow future sub-cloning of the product of mutagenesis. The PCR product of the first step (PCR1) is purified from agarose gel electrophoresis. It is then used as "megaprimer," given its unusually high length, for the second step of PCR (PCR2) together with a second flanking primer designed to allow amplification of the entire gene or to the nearest restriction site (R2). This second restriction site will be used in combination with R1 to allow sub-cloning. If the mutagenic primer can be designed to contain a restriction site, only the first step of PCR will be necessary and the product of the first PCR reaction can be cloned directly in the expression vector. The insertion of a new restriction site (R3) associated with the presence of the specific sequence of the desired mutation will allow an easy screening procedure to select for the clones that contain the mutant plasmid. In all cases it is strongly recommended to confirm the presence of the desired mutation and check the absence of undesired ones by DNA sequencing of the full length of the gene. The size of the megaprimer varies depending on the requirements of the specific gene and mutation to be introduced. Generally it is much longer than a typical oligonucleotide primer, varying between ~100 and ~900 bp. The difficulty of this method lies in achieving good amplification in the second 212

PROTEIN ENGINEERING FOR BIOSENSORS

Figure 8 (a) Schematic diagram of a PCR experiment. Two oligonucleotide primers are designed to anneal on opposite strands flanking the sequence to be extended by a thermostable DNA polymerase. In order to facilitate the sub-cloning of the DNA products, two restriction sites R 1 and R2 are included, (b) Steps for a megaprimer PCR experiment. The oligonucleotide primer containing the desired mutation is used to give a PCR product, that is then used as a megaprimer to extend the full length of the target sequence.

PCR step using the megaprimer. For this reason, the experiment should be designed so that PCR1 is as short as possible, to avoid exceedingly high Tm and the risk of secondary structure causing self-annealing. On the other hand, the size of PCR1 should be such that the recovery from the agarose gel electrophoresis is not a limiting factor. A typical experiment is reported in Protocol 8 (25).

Protocol 8

Megaprimer PCR Equipment and reagents See Protocol 5.

Method Step 1. PCR1.

1

For a 100 ul reaction, add in a 0.5 ml PCR tube: 1 ul plasmid template (50 ng) 2 ul oligonucleotide primer n.l (50 pmol/ul) 2 ul oligonucleotide mutant primer n.2 (50 pmol/ul) 8 ul dNTP mix (2.5 mM each)

213

GIANFRANCO GILARDI Protocol 8 continued

0 |ol 10 x DNA polymerase buffer 75 |ol water.

2 Perform the PCR experiment as described in Protocol 5, but add 2 jol of DNA polymerase (5 units). 3 Load all the PCR product on a 1.5% agarose gel (use a wide comb to obtain large bands that are easier to excise from the gel).a 4 Perform the separation as described in Protocol 6. 5 Excise the band corresponding to the megaprimer (PCR1 product) by cutting the band with a sharp blade. Minimize the exposure of the megaprimer to the UV light to avoid undesired mutations. 6 Purify the megaprimer from the agarose as described in Protocol 7. 7 Determine the concentration of pure megaprimer as described in Protocol 5. Step 2. PCR2 8 For a 100 |ol reaction, add in a 0.5 ml PCR tube: 1 |ol plasmid template (50 ng) 2 |ol oligonucleotide primer n.3 (50 pmol/|ol) 40 |ol megaprimer (50 pmol/|ol)b 8 |ol dNTP mix (2.5 mM each) 10 |ol 10 x DNA polymerase buffer 75 |ol water. 9 Perform the PCR experiment as described in Protocol 5, but add 2 [d of DNA polymerase (5 units). 10 Cool the PCR products on ice. 11 Load 10 |ol of the PCR product on a 1% agarose gel to check that the correct product has been formed. 12 When the Taq DNA polymerase is used, add 1 |ol of the Klenow fragment (5 U/|ol) directly to the PCR reaction and incubate at 37 °C for 1 h. 13 Perform a restriction digest as described in Protocol 9, using the suitable restriction enzymes flanking the mutated amplified region.0 14 After separation by agarose gel electrophoresis (Protocol 6), purify the product of digestion from the gel (Protocol 7). 15 Perform a ligation in a vector suitable for expression (Protocol 10). 16 Transform (Protocol 11) the resulting plasmid in a strain of Escherichia coli compatible with the promoter carried by the chosen expression vector (e.g. Escherichia coli BL21 (DE3) for a pT7 plasmid carrying the T7 promoter). 17 Spread on LB plates containing the appropriate antibiotic and incubate at 37 °C overnight.

214

PROTEIN ENGINEERING FOR BIOSENSORS

Protocol 8 continued

18

Pick 5-10 colonies and grow overnight at 37 °C on liquid LB medium with the appropriate antibiotic.

19 Isolate the plasmid from the colonies (Protocol 12), perform a restriction digest (Protocol 9), and separate on agarose gel electrophoresis (Protocol 6) to verify that the plasmid contains the correct DNA insert. 20

Submit the positive clones to DNA sequencing to verify the presence of the mutation. This can be done routinely by sending the plasmid with the relevant primers to DNA sequencing services.

a

In some cases it may be better to first load only 10 (il of the PCR reaction products, to check the efficiency of the reaction and the purity of the megaprimer prior to large scale separation. In an ideal case, all the primers of the reaction have been converted in the megaprimer only. In this case one may proceed directly to the second step of PCR, without purification an agarose gel. When this convenient short cut is chosen, one must be aware of the risk of recovery of the very small amount of the wild-type used as template in the first step.

b

Ideally the concentration of the megaprimer should be determined spectrophotometrically from the absorbance at 260 nm. Then 100 pmol should be added to achieve equal concentration with primer n.3.

C

A sufficient number of bases must be present at the terminals to allow restriction endonucleases to function. When this proves to be difficult, an intermediate cloning step maybe necessary. This consists in the blunt end ligation of the PCR product into a high copy number plasmid such as the pBluescript vector. The fragment can then be easily digested using the relevant restriction endonucleases.

Protocol 9

Restriction digest of DNA Equipment and reagents • 1.5 ml microcentrifuge tubes and standard bench microcentrifuge • Heating block at 80 °C • Restriction endonucleases from various suppliers, for example, Amersham

Biosciences, New England Biolabs, Promega, Stratgene • Water bath at 37 °C • Ice bucket • Reaction buffer provided with the restriction endonucleases.

Method 1

For a 20 [d reaction, add in a 1.5 ml microcentrifuge tube: 16 nl DNA (0.1-1.0 ^g)a 2 |ol 10 x buffer for the chosen restriction endonucleaseb 1 |ol each of the two restriction endonucleases (5-10 units of each).c

215

GIANFRANCO GILARDI Protocol 9 continued 2 Make sure that the enzymes are added last and the tube is always kept on ice except for the incubation period.

3

Mix gently using a pipette.

4

Briefly spin using a microcentrifuge to collect the mixture at the bottom of the tube.

5

Incubate for 1-3 h at 37 °C.d

6

Stop the reaction by incubating 3 min at 80 °C.

7 Cool on ice, briefly spin, and load 5-10 ul on an agarose gel to check the products (Protocol 6). a

A typical DNA miniprep should contain this concentration of DNA that is easily visualized on agarose gel.

b

Consult with the requirements of the suppliers of the enzymes. Usually the buffer is supplied with the enzymes. The correct concentration and buffer compatibility between different enzymes is crucial to avoid nonspecific cleavage of DNA.

C

10% of the total volume of the reaction mixture, because the glycerol used to store the enzymes will promote nonspecific cleavage of DNA.

d

Length and temperature of incubation varies depending on the efficiency of the enzyme. Consult the recommendations of the enzyme suppliers.

Protocol 10

Ligation of a DNA insert in a vector Equipment and reagents • 0.5 ml microcentrifuge tubes and standard bench microcentrifuge

• Sterile water. • Heating block at 65 °C

• Linearized vector

• PCR product or insert • 10 x T4 ligase buffer containing 10 mM ATP . T4 ligase (3-6 U/ul)

Method 1

For a typical 30 ul experiment reaction, add in a 0.5 ml microcentrifuge tube: 2 ul of linearized vector (approx 50 ng) 20 ul of PCR product or insert (approx 250 ng) 3 ul 10 x T4 ligase buffer containing 10 mM ATP 1 ul T4 ligase (3-6 U/ul) 4 ul sterile water.

2 In parallel, set up a control reaction as follows: 2 ul of linearized vector (approx 50 ng) 3 ul 10 x T4 ligase buffer containing 10 mM ATP

216

PROTEIN ENGINEERING FOR BIOSENSORS

Protocol 10 continued

1 |il T4 ligase (3-6 U/nl) 24 (il sterile water. 3 Proceed for the two reactions as follows: 4 Mix the water, vector, and PCR product (insert) in a 0.5 ml microcentrifuge tube. 5 Heat the sample at 65 °C for 10-15 min to uncoil any supercoiled DNA. 6 Cool the sample on ice and touch spin to collect all the liquid to the bottom of the tube. 7 Add the buffer followed by the ligase enzyme. Incubate the sample overnight at 16 °C for blunt-end or 4°C for sticky-end ligations. 8 Use half the reaction mixture to transform competent cells (Protocol 11) and store the remainder at — 20 °C in case a second transformation is needed. For blue-white screening of the insert, see the note in Protocol 11.

Protocol 11

Transformation of a plasmid in a bacterial host Equipment and reagents Standard LB media are made using per liter 10 g of tryptone, 10 g of sodium chloride, and 5 g of yeast extract; the pH was then adjusted to seven using 0.1M sodium hydroxide. For agar plates 15 g of agar is added per liter of LB. Ampicillin stocks contain 100 mg/ml of ampicillin and are used to give a final concentration of 100 (xg/ml and stored at — 20 °C when not in use. IPTG stocks contained 1M IPTG and were diluted to give a working concentration of 1 mM and stored at — 20 °C when not in use.

Method Step 1. Competent cell preparation An easy procedure to produce Escherichia coli competent cells is the so-called calcium chloride method. This method yields around 2 x 107 transformed colonies per microgram of plasmid DNA. 1 Pick one Escherichia coli colony from a freshly grown (overnight at 37 °C) plate, and transfer it into 10 ml of LB broth. Incubate over night at 37 °C with vigorous shaking. 2 Pick 7 |ol of the overnight culture and inoculate 7ml of fresh LB broth, which is then incubated at 37 °C with vigorous shaking for 2 h. 3 Spin the cells at 4000 rpm for 10 min at 4 °C. 4 Remove the supernatant and keep the cell pellet on ice at all times. 5 Resuspend the cells in 3.5 ml of ice cold 50 mM CaCl2.

217

GIANFRANCO GILARDI Protocol 11 continued

6 Leave on ice for 30 min. 7 Spin the cells at 4000 rpm for 10 min at 4°C and remove the supernatant. 8 Resuspend the pellet with 700 |ol of ice cold 50 mM CaCl2 and store on ice. 9 The cells will be competent within a couple of hours, with a peak after 24 h. Step 2. Transformation procedure 10 Add 1 |ol (50 ng) of the desired plasmid to 200 [d of the competent cells. 11 Incubate on ice for 30 min. 12 Quickly heat shock the cells for 90 s at 42 °C. 13 Immediately transfer the tube to an ice bucket and incubate for 2 min. 14 Add 800 |ol of fresh LB broth and incubate for 45 min at 37 °C to allow the bacteria to recover and express their antibiotic resistance marker encoded by the plasmid. 15 Spin the cells at 4000 rpm for 10 min at 4°C and remove the supernatant. 16 Resuspend the pellet with 200 |ol of fresh LB. 17 Plate the cells onto agar plates containing the antibiotic for which the plasmid confers resistance. 18 Incubate the plates overnight at 37 °C to allow the colonies to grow.a a

When the transformants are expected to contain a plasmid that contains a ligated insert, a so-called blue-white colony screen may be helpful in identifying the colonies containing the insert. This can be achieved by using the vector pBluescript that contains the cloning site on the fS-galactosidase gene. When the insert is present, the fS-galactosidase is not expressed and the colonies will appear white. When the insert is not present, the fS-galactosidase is expressed and the colonies will yield a blue precipitate. The screen is achieved by adding 80 (ig/ml of 5-bromo-4-chloro-3-indolyl-|3-D-galactopyranoside (X-gal, prepared in dimethylformamide), plus 20 mM isopropyl-1-thio-fS-D-galactopyranoside (IPTG prepared in sterile water) to the agar plate. Alternatively, 100 (il of 10 mM IPTG and 100 (il of X-gal can be spread on the agar plate 30 min prior to plating the transformations. Do not mix the IPTG and the X-gal before pipetting them onto the agar plates because they may precipitate.

Protocol 12

Isolation of a plasmid from a bacterial host Equipment and reagents Plasmids must be isolated from bacterial cultures derived from one single colony picked from fresh agar plate, not older than one week. The Luria-Bertani (LB) broth is usually preferred to Terrific Broth (TB) because the latter would yield 2-5 times the number of cells that may present problems in the efficiency of alkaline bacterial lysis, purity of plasmid DNA (contamination from genomic DNA) and overloading of the DNA binding membranes used in DNA purification.

218

PROTEIN ENGINEERING FOR BIOSENSORS

Protocol 12 continued

LB broth, 1 1 10 g NaCl 10 g tryptone 5 g yeast extracts 1 ml of 1M Tris-Hcl at pH 7.0. TB broth, 1 1 12 g tryptone 24 g yeast extracts 8 ml of 50% glycerol 2.31 g KH2PO4 2.54 g K2HPO4. Generally the protocols for plasmid isolation are adjusted for culture volumes of 1-5 ml. One single colony is grown overnight for 12-16 h, in the presence of a suitable antibiotic (see Table 6). The antibiotic selection must be kept at all stages of growth and it is important to realize that long incubations cause depletion of the antibiotic. For example, the resistance to ampicillin is caused by the enzyme p-lactamase encoded by the plasmid-linked bla gene. As p-lactamase hydrolyzes ampicillin, the levels of this antibiotic are continuously depleted from the culture medium. This is evidenced by the presence of satellite colonies in agar plates. Escherichia coli DH5a and XLl-Blue are highly recommended for plasmid preparations. Strain HB101 and its derivatives such as the TG and JM series are known to produce large amounts of carbohydrates that are released during the plasmid isolation procedure and can inhibit subsequent enzyme activities if not completely removed. Another important factor to be considered is the copy number of plasmid per cell that is dependent on the origin of replication born by the plasmid itself (see Table 7). Copy numbers higher than 200 per cell are classified as "high," whilst lower than 100 are "low." This must be kept in mind when expecting a high yield of plasmid preparation. Traditional methods for plasmid isolation are based on the alkaline lysis of the bacteria followed by phenol extraction and ethanol precipitation of the plasmid DNA. The tedious steps of lysis, repeated extraction, and precipitation are today replaced by kits that require three simple steps: lysis, adsorption of the plasmid DNA to a membrane, washing, and elution of the plasmid DNA. The protocol reported here is based on the widely used QIAprep® kit provided by Qiagen. Although the DNA yields depend on many factors such as plasmid copy numbers, bacterial growth, size of the culture, a typical preparation by QIAprep® yields 5-20 (ig of dsDNA in 50 jol of final elution volume. This is good enough for most applications. Interestingly, Qiagen can also provide kits for high throughput plasmid preparations in 96-well plates; 96 preparations can be achieved in less than 45 min.

Method Step 1. Alkaline lysis of bacteria 219

Protocol 12 continued

1

Spin down the 1-5 ml overnight culture at maximum speed for 5 min using a conventional microcentrifuge. 2 Resuspend the pellet in 250 |ol of lysis buffer PI by vortexing, taking care that no lumps of cells are observed. The buffer contains RNase to improve the purity of the plasmid DNA (usually RNA co-purifies with the plasmid). 3 Add 250 |ol of buffer P2, mix gently by inversion, allow to lyse for no more than 5 min. This buffer contains sodium hydroxide that causes bacterial lysis due to the alkaline conditions.

4 Add 350 |ol of buffer N3 to terminate the lysis, mix gently by inversion. This buffer contains guanidine hydrochloride to fully precipitate the proteins and acetic acid to neutralize the pH. The solution should become cloudy. 5 Centrifuge for 10 min at full speed. Step 2. DNA adsorption to membranes 6 Carefully remove the supernatant and load it to the centrifuge spin columns containing the DNA binding membrane. 7 Centrifuge for 30-60 s, discard the flowthrough. 8 Wash with 500 |ol of buffer PB, centrifuge for 30-60 s, discard the flowthrough. This buffer contains guanidine hydrochloride and isopropanol and it is necessary for removing the nuclease activity and the carbohydrates from strains such as HB101, TG, and JM series (see above). Step 3. Washing and elution of the plasmid DNA 9 Wash with 750 ul of buffer PE, centrifuge for 30-60 s, discard the flowthrough. This buffer contains ethanol that ensures that the DNA remains bound to the membrane while being washed. The same considerations made on DNA binding to resins made in Protocol 7 are valid here. 10 Centrifuge again at full speed for 1 min to remove any residual buffer. This is very important to elute a clean plasmid DNA in the following step. 11 Place the spin column on a clean and sterile eppendorf tube. 12 Add 50 ul of sterile water to the top of the spin column, let stand for 1 min to promote DNA release, centrifuge for 1 min. Maximum elution efficiency is obtained at pH 7.0-8.5. 13 Collect the flowthrough containing the pure DNA plasmid and store at — 20 °C. Table 6 Preparation of most commonly used antibiotics. All solutions must be stored at -20°C Antibiotic

Stock solution

Working concentration (|xg/ml)

Dilution

Ampicillin sodium salt

50 mg/ml in water

100

1:500

Chloramphenicol

34 mg/ml in ethanol

170

1:200

Kanamycin

10 mg/ml in water

50

1:200

Streptomycin

10 mg/ml in water

50

1:200

Tetracyclin HCI

5 mg/ml in ethanol

50

1:100

PROTEIN ENGINEERING FOR BIOSENSORS Table 7 Origin of replication and copy numbers of the most commonly used plasmids and cosmids Plasmids

Origin of replication

Copy number

pUC vectors

ColEl

500-700

pBluescript vectors

ColEl

300-500

pGEM vectors

pMBl

300-400

pTZ vectors

pMBl

>1000

pBR322 and its derivatives

pMBl

15-20

pACYC and its derivatives

p!5A

10-12

pSClOl and its derivatives

pSClOl

~5

SuperCos

ColEl

10-20

pWE15

ColEl

10-20

Cosmids

QuikChange ® PCR method The QuikChange® is the most rapid and efficient method available to date to produce site-directed mutants. It can be performed in one day and it consists of a PCR reaction carried out directly on the plasmid DNA template, without the need of sub-cloning steps or specialized vectors, or unique restriction sites or multiple transformations. The procedure consists of three steps: PCR reaction, digestion and transformation. QuikChange® is provided in a kit by Stratagene® (www.stratagene.com) and the principle of the procedure is shown in Figure 9. The first step consists in a PCR reaction that uses the high fidelity PfuTurbo DNA polymerase with two 5'-phosphorylated primers containing the desired mutations, each complementary to the same region of opposite strands of the vector. The incorporation of the oligonucleotide primers by PCR extension generates a mutated plasmid containing staggered nicks. The mutated strands are selected by digestion of the template wild-type, methylated plasmid with the restriction endonuclease Dpn I. This enzyme digests the sequence 5'-Gm6ATC-3' and is specific for methylated DNA as the starting plasmid isolated from the commonly used dam+ Escherichia.coli strains. The final step consists of the transformation of the mutated nicked dsDNA into the XL-1 Blue supercompetent cells that will repair the nicks in the mutated plasmid. The QuikChange® kit comprises a control reaction to be carried out with the oligonucleotide primers and the pWhitescript™ plasmid provided. This plasmid contains a stop codon (TAA) at the position of glutamine 9 (CAA) of the p-galactosidase gene. This results in an inactive p-galactosidase that gives white colonies on LB-ampicillin agar plates containing IPTG and X-gal. The mutagenesis control reaction is designed to introduce oligonucleotide primers that restore the CAA codon, resulting in an active enzyme. Following transformation, the colonies containing the successful mutagenesis can be screened for the p-galactosidase 221

G I A N F R A N C O GILARDI

TM

Figure 9 Schematic procedure for the QuikChange

PCR mutagenesis method.

blue phenotype. The mutagenesis efficiency (£) can be calculated with the formula: E — (No. of blue colonies)/(Total No. of colonies) x 100. A typical successful experiment should result in a 80% efficiency of mutagenesis.

3 Directed evolution Directed evolution is a new and powerful approach of protein engineering particularly suitable when little information is known on the structure/function relationship of the target protein/enzyme. It relies on iterative cycles of random mutagenesis, recombination, and functional screening of a library of variants from which mutants with improved or new properties are identified (Figure 2). The possibility of generating enzymes with high chemical and thermal stability, solubility in organic solvents, activity toward new substrates, specificity and enantio- or regie-selectivity in catalysis is a very attractive target for the biosensors of the future. Essentially this approach mimics in vitro the process of natural molecular evolution that is able to generate a potentially infinite plethora of new proteins with 222

PROTEIN ENGINEERING FOR BIOSENSORS

new functions. Therefore, it becomes obvious that there is a great potential that directed evolution offers to the bioanalytical area, where the experimentalist will no longer be limited by the availability and the properties of natural proteins.

Protocol 13

QuikChange

PCR mutagenesis

Equipment and reagents See Protocol 5. The Stratagene kit contains all the reagents except the plasmid template and the oligonucleotide primers. An important step in all PCR experiments is the choice of the oligonucleotide primers. Some variations from the general procedure given in Section 2.2.3 are necessary when using the QuikChange® method. Both primers should contain the desired mutation and must anneal to complementary sequences on opposite strands of the plasmid. They should be 25-45 bases long with the desired mutation in the middle, with a minimum GC content of 40% and terminating with one or more C or G bases. They must be purified by FPLC or PAGE and their Tm should be equal to or greater than 78 °C. Two different empirical formulae can be used to calculate Tm: Tm = 81.5 + 0.41 (%GC) - 675/N - %mismatch can be used for substitutions, where N is the length in bases, %GC and %mismatch are whole numbers; Tm = 81.5 + 0.41 (%GC) - 675/N can be used for insertions or deletions, where N does not include the bases that are being inserted or deleted.

Method Step 1. PCR reactions 1 For a 50 jol reaction, add in a 0.5 ml PCR tube: 1 |ol plasmid template (125 ng) 2 |ol oligonucleotide primer n.l (125 pmol/|ol) 2 |ol oligonucleotide mutant primer (50 pmol/|ol) 4 nl dNTP mix (2.5 mM each) 5 |ol 10 x DNA polymerase buffer 35 |ol water 1 |ol Pfu DNA polymerase (2.5 U/|ol). 2 Perform the following cycling reactions. 3 1 cycle at 95 °C for 30 s. 4 12-18 cyclesa as follows: denaturation: 95 °C for 30 s annealing: 55 °C for 1 minb extension: 68 °C for 2 min/kb of plasmid length. 223

GIANFRANCO GILARDI

Protocol 13 continued

5 Place on ice for 2 min to cool the reaction to < 37 °C. Step 2. Dpn I digestion of the wild-type template 6 Add 1 [d of a 10 U/|ol stock of Dpn I directly to the reaction mix. 7 Mix gently using a pipette. 8 Briefly spin using a microcentrifuge to collect the mixture at the bottom of the tube. 9 Incubate for 1 h at 37 °C. Step 3. Transformation 10 Transform a 2 jol aliquot of the reaction mix according to the procedure described in Protocol 9. If possible use the commercial XLlO-Gold Ultracompetent Escherichia coli cells that are capable of highly efficient transformations as they are both endonuclease (endAl) and recombination (recA) deficient. This will increase the yield of transformants. If standard competent cells are used (prepared as described in Protocol 9), use the whole reaction mix or increase the concentrations of DNA by ethanol precipitation. a

Adjust the number of cycles according to the following: 12-16 cycles = point mutations, single amino acid changes; 18 cycles = multiple amino acids deletions or insertions. b This should be 2°C lower than the lowest Tm of the oligonucleotide primers. Directed evolution is based on the generation of genetic diversity by DNA recombination, also called DNA shuffling. This process mirrors the Darwinian evolution where random mutagenesis, recombination, and selection allow the addition of beneficial mutations and the removal of deleterious ones. This section will describe the three key steps of the iterative cycles of directed evolution: random mutagenesis, recombination, and functional screening. As indicated in Figure 2, once a trend in the structure/function properties of the target enzyme has been rationalized, the directed evolution approach can be set to work in tandem with the rational design one.

Random mutagenesis: error prone PCR Molecular diversity can be created in a number of ways, for example, by error prone PCR and combinatorial cassette mutagenesis. In combinatorial cassette mutagenesis, a relatively short DNA sequence is mutagenized by synthetic oligonucleotide primers (26). This method is relatively straightforward, but it is useful only when the targeted amino acids are in the same stretch of primary sequence. It relies on the synthesis of a large number of oligonucleotide primers, and therefore it is combinatorial only within the limits of the synthetic oligonucleotides, that may be labor-intensive and expensive. Error prone PCR is by far the most used approach to produce random mutants. It was developed by Leung in 1989 (27, 28) who exploited the ability of thermostable DNA polymerases lacking proof-reading activity, for example, Taq polymerase (see Table 1), to introduce random mistakes during the extension 224

PROTEIN ENGINEERING FOR BIOSENSORS

of the new DNA strand. Taq has the highest known error rate, in the range of 0.1 x 10~ 4 to 2 x 10~ 4 per nucleotide per pass of the polymerase. The rate of mis-incorporation depends on the specific reaction conditions, therefore, the standard PCR protocol needs to be modified to control the error rate. For example, the MgCl2 concentration is increased to 7 mM to stabilize non-complementary pairs, MgCl2 is added at 0.5 mM to diminish the template specificity of the polymerase, the concentrations of dCTP and dNTP is increased to 1 mM to promote mis-incorporation, and the amount of Taq DNA polymerase is increased to 5 U to promote the chain extension beyond the positions of base mismatch. Generally the procedure proposed in Protocol 14 works well for DNA templates up to 1 kb; longer target sequences can be divided into two or more fragments.

Protocol 14

Error prone PCR Equipment and reagents See Protocol 5. Stock reagents are: 10 x PCR buffer: 70 mM MgCl2 500 mM KC1 100 mM Tris-HCl pH 8.3 0.1% gelatin 10 x dNTPs: 2 mM dGTP, 2 mM dATP, 10 mM dCTP, 10 mM dTTP MnCl2 : 5 mM.

Method 1

For a 100 |ol reaction, add in a 0.5 ml PCR tube: 1 |ol DNA template (50 ng) 2 |ol oligonucleotide primer n.l (50 pmol/|ol) 2 |ol oligonucleotide primer n.2 (50 pmol/|ol) 10 nl 10 x dNTP mix 10 [d 10 x DNA polymerase buffer 63 [d water.

2 Mix well. 3 Add 10 [d of 5 mM MnCl2, mix well ensuring that there is no precipitate. 4 Add 5 U of Taq DNA polymerase to a final volume of 100 |ol. 5 The mixture is heated for 5 min at 94 °C and then subjected to thermal cycling as follows (30-80 cycles depending on the specific requirements): 94 °C for 1 min 55 °C for 1 min 225

GIANFRANCO GILARDI Protocol 14 continued

72 °C for 1 min. 6 Isolate the amplified DNA fragment of the correct size by agarose gel electrophoresis, digest and ligate with the appropriate vector.

Error prone PCR is able to produce large libraries of random mutants in a sequential fashion where deleterious mutations are accumulated in conjunction with beneficial ones. This can seriously limit the evolutionary process because of the high probability that the effects of the few beneficial mutations are lost due to the many deleterious ones. From a biological point of view, sequential random mutagenesis is equivalent to an asexual evolutionary process.

Recombination: DNA shuffling The breakthrough in directed evolution was made by Stemmer in 1994 with the introduction of DNA shuffling that allows the in vitro formation of recombinant genes from a set of parental ones (29, 30). DNA shuffling requires the availability of a pool of homologous genes that can be derived from a naturally occurring protein family or from a library of mutants produced by error prone PCR, as described in the previous paragraph. Error prone PCR can introduce a number of point mutations in a gradual fashion. It lacks the essential features of block changes typical of DNA shuffling that also enables the removal of deleterious mutations by back-crossing with parental or wild-type DNA (Figure 10). Block changes are introduced with the

Figure 10 Recombination of mutant or homologous genes by DNA shuffling.

PROTEIN ENGINEERING FOR BIOSENSORS

fragmentation of the pool of genes using DNase I. The fragments are purified by agarose gel electrophoresis and reassembled into a full length gene by repeated cycles of PCR without added primers (Protocol 15). During this step the fragments prime each other on the homologous regions, resulting in recombination when fragments derived from one parental gene prime on another one, causing a template switch. The recombinant genes obtained with this procedure can then be used for other rounds of mutation and recombination.

Protocol 15

DNA shuffling Equipment and reagents See Protocol 5 and 14.

Method Step 1. DNAse I digestion 1 Dilute the genes or PCR products with the random mis-incorporations (Protocol 14) to 2-4 ng in 10 mM Tris-HCl (pH 7.4), 1 mM MgCl2. 2 Equilibrate the mixture at 15 °C for 5 min in a thermocycler. 3 AddO.SU of DNAse I. 4

Digest for 2 h at 15 °C.a

5

Stop the digestion by heating at 90 °C for 15 min.

6

Purify fragments of ca. 50 bp by 2% agarose gel electrophoresis.

Step 2. Fragment reassembly 7 Reassembly is achieved by Taq polymerase in the absence of primers. A 100 [d PCR mixture contains: 10-30 ng/|ol of fragments 0.2 mM each dNTPs 2.2 mM MgCl2 50 mM KC1 10 mM Tris-HCl (pH 9.0) 0.1% Triton X-100 2.5 U Taq polymerase. 8

Perform the PCR cycles (30-45) as follows:

96 °C for 3 min 50-55 °C for 1 min 72 °C for 1 min. 9 Incubate the mixture at 72 °C for 5 min.

227

GIANFRANCO GILARDI Protocol 15 continued

10 Reassembly of fragments of 100-200 bp size usually yield PCR products of the correct size; smaller fragments of 10-50 bp show some heterogeneity in the size of the PCR product, but this is normally rectified by restriction digest of the PCR product.

Step 3. Amplification of the reassembled fragments 11 A typical 100 |ol PCR reaction contains: 30 mmol of each primer 0.2 mM each dNTP 2.5UTaq/P/it (1:1) 1 x Taq buffer. 12 Perform the PCR cycles as follows. 13 Incubate at 96 °C for 2 min. 14 Perform 20 cycles as: 94 °C for 30 s 55 °C for 30 s 72 °C for 1 min. 15 Isolate the amplified DNA fragment of the correct size by agarose gel electrophoresis, digest and ligate with the appropriate vector. a

Time may need to be adjusted down to 10-20 min depending on the length of the target DNA.

Key features of DNA shuffling are: the requirement of considerable quantities of DNA templates with homologous regions able to prime each other; the presence of sequences of homology separated by regions of diversity; the block changes typical of sexual recombination brought about by template-switch or cross-over. These features have been subject of studies that led to modifications and improvements of original basic protocol. This paragraph reports only some examples of these new approaches. The Staggered Extension Protocol (StEP) has dramatically decreased the amount of the DNA template required by DNA shuffling (31). This method allows the template switching during the synthesis to yield chimeric genes without the requirement of the DNase I fragmentation step (Figure 11 (a)). The DNA templates are mixed with one or more primers and subjected to cycles of denaturation and short annealing/extension steps. The growing PCR products, or fragments, randomly prime to different templates and further extensions will contain template switching. The process is repeated until full length genes are obtained and purified on agarose gel electrophoresis. Parental-like genes are eliminated by digestion with Dpn I as described for the QuikChange method (see section on Quikchange® PCR method, Protocol 13).

228

PROTEIN ENGINEERING FOR BIOSENSORS

Figure 11 Procedures for the (a) StEP, (b) the ITCHY, and (c) the SHIPREC methods.

DNA shuffling generally relies on homologous gene recombination where template switching occurs at points of high sequence identity. Therefore, the DNA shuffling methods described so far can recombine only closely related sequences, that are more than 70% identical. Although many interesting target proteins show very similar functions and high three-dimensional structural homology, they often share only a very low sequence identity. In these cases it is convenient to refer to the Iteractive Truncation for the Creation of Hybrid Enzymes (ITCHY). This method allows the generation of N- and C-terminal fragment libraries of two genes by progressive truncation of the coding sequences with exonuclease III, followed by ligation of the products to make a single crossover hybrid library. This strategy yields cross-overs throughout all the coding region, as shown in Figure 11 (b) (32). Only a small number of cross-overs will connect the two parent genes at sites where the sequences align. Another method able to generate libraries of hybrid genes from distantly related sequences is the Sequence Homology Independent Protein Recombination (SHIPREC). A key feature of SHIPREC is that it ensures cross-overs at positions sharing a similar structural environment, maximizing the probability of obtaining functional proteins (33). The SHIPREC procedure starts with the fusion of two genes to give a heterodimer (Figure ll(c)). This is then digested by DNase I in the presence of Mn 2+ that favors double-strand breaks over nicks, producing an ensemble of fragments of random lengths. Fragments of length corresponding to that of either gene parent are isolated and treated with SI nuclease or T4 polymerase to produce blunt ends. The chimeric genes are first circularized by intra-molecular blunt end ligation, followed by linearization by restriction 229

GIANFRANCO GILARDI

digestion. This creates a library of chimeric DNA sequences where the gene that was at the 5' position in the dimer will now be at the 3' position and will donate the C-terminus of the hybrid protein. The cross-over will therefore be distributed over the entire length of the gene. This approach has been applied to produce soluble hybrids of the membrane associated human cytochrome P450 1A2 and the soluble haem domain of the bacterial cytochrome P450 BM3, that share only 16% sequence identity. The SHIPREC method successfully produced cross-overs mainly at structurally related sites along the sequences. The resulting folded and partially soluble hybrid P450s had cross-overs close to one of the termini, where there is no structural similarity. Simple cross-overs in the interior of the proteins were too disruptive to produce a folded and active protein.

3.3 Functional screening of the library A key requirement to proceed in the iterative cycles leading to the best recombinant protein is the availability of a screen or selection procedure that is sensitive to the properties target in the in vitro evolution process. This step is not trivial, as the vast majority of the mutations are deleterious and only very few will lead to the improved target property. The number of possible variants (V) of a protein that can be created by introducing M substitutions simultaneously over N amino acids is given by the formula: V = 19M[N!/(N-M)!M!]. This means that even for a small protein, when only 10 positions are targeted for 3 simultaneous changes of amino acids, there are 823,080 possible variants. The ability to identify one possible active variant among this large library is close to the limit of most screen or selection methods. Traditionally the term selection is reserved for the methods of searching a library based on an enzymatic function that confers a growth or survival advantage to the host organism. An example of selection is given by Stemmer in the first application of DNA shuffling (30). In this work Stemmer used a mixture of /Mactamase genes to carry out three cycles of DNA shuffling and two cycles of back-crossing with wild-type DNA. Escherichia coli XLl-Blue containing the plasmid expressing the mutagenized /Mactamases was grown on agar plates containing increasing amount of the antibiotic cefotaxime. Only the Escherichia coli expressing the /Mactamase with improved activity toward cefotaxime could grown on plates containing increasing amounts of the antibiotic. This allowed a 32,000-fold increase of the minimum inhibitory concentration from 0.02 (xg/ml typical of the wild-type to 640 (xg/ml typical of the evolved enzyme. Generally the use of a host organism as a means for biological selection is very efficient and it allows searches of libraries containing more than 10s variants. There is, however, a high risk that, under the selective pressure, the host organism itself develops alternative metabolic mechanism unrelated to the target activity. On the other hand, a screening procedure involves the construction of an arrayed protein library and the application of a rapid and high throughput 230

PROTEIN ENGINEERING FOR BIOSENSORS

Figure 12 Screening procedures developed for (a) Cytochrome P450cam (36), (b) Cytochrome P450 BM3 (37) and (c) more generally for NAD(P)H dependent oxidoreductases (13).

assay sufficiently sensitive and specific to enable the identification of positive variants. The development of the screening assay is usually the critical step in the directed evolution of a particular enzyme. Although in some cases visual screens based on color or halo-formation are rapid and straightforward, they are nonquantitative and often insensitive to small changes in the enzymatic properties. Preference is given to quantitative assays based on 96 (or more) well plates. This format not only is quantitative, but it is also amenable to automation using robotic arms, liquid handling systems and plate readers with automated data acquisition. There is a number of examples of screening procedures described in the literature (see reviews 34, 35). In keeping with this chapter's theme on cytochrome P450, this paragraph will give three examples of screening procedures developed for two members of this class of enzymes. Cytochrome P450s are haem-thiolate monoxygenases able to catalyze the insertion of one atom of oxygen from O2 into a wide variety of substrates with concomitant production of water, using the reducing equivalents of NAD(P)H normally provided through a redox chain involving one or two electron transfer proteins. They are involved in key reactions of detoxification of xenobiotics and biosynthesis of steroids, generating much interest in the bioanalytical area. Arnold and coworkers (36) developed mutants of P450 cam from Pseudomonas putida that are able to utilize the electrons and oxygen provided by hydrogen peroxide, eliminating the need for expensive cofactors and complex redox chains. P450 cam was evolved to acquire hydroxylating activity toward the un-natural substrate naphthalene. The screening procedure shown in Figure 12(a) relied in the co-expression of the horseradish peroxidase (HRP) mutant 231

GIANFRANCO GILARDI

Asn255Asp (HRP1A6). This mutant was previously obtained by directed evolution to allow expression of HRP in an active form in Escherichia coli. After 16 h growth on agar/TB plates, the P450 cam variants and HRP1A6 were replicated using a nitrocellulose membrane and transferred onto fresh agar plates containing 6 mM naphthalene and 10-30 mM hydrogen peroxide. The naphthols generated by the P450 cam variants were converted by HRP1A6 into fluorescent dimers and polymers that were detected by fluorescence digital imaging. Different naphthols obtained by diverse P450 cam variants of altered regio-specificity of hydroxylation are polymerized by HRP1A6 in products with different fluorescence spectra. These characteristics were used to select P450 cam variants tuned to different regio-specificity in catalysis. Variants with improved properties were subjected to in vitro recombination by StEP, yielding to several mutants with ~20 fold improvement in naphthalene hydroxylation activity over the wild-type. Another interesting monoxygenase is P450 BM3 from Bacillus megaterium. This enzyme contains the haem catalytic domain fused with the FMN/FAD reductase on the same polypeptidic chain. Wild-type P450 BM3 hydroxylates fatty acids of chain length between C12 and CIS. Arnold and co-workers engineered this enzyme to hydroxylate short chain alkanes such as octane (37). The screening procedure used an octane analogue, 8-pnpane, that upon terminal hydroxylation produces an unstable hemiacetal. This decomposes into the aldehyde and p-nitrophenolate that is detected by its yellow color at 410 nm (Figure 12(b)). Two round of error prone PCR and the screening of 2000-3000 colonies produced a variant with activity 2-3 fold higher than the wild-type. Both the screening procedures described so far are specific for the substrate under investigation, and in the case of octane the procedures are also dependent on the position of the hydroxylation. A general screening procedure developed using P450 BM3, but applicable to all NAD(P)H-dependent enzymes has been developed in our laboratory (13). The screening procedure relies on the conversion of the NADP+ resulting from enzymatic activity into a fluorescent product upon treatment with alkali (Figure 12(c)): for this reason, it has been named "alkali assay." The procedure can be applied to purified enzyme as well as to whole Escherichia coli cells expressing the target NAD(P)H-dependent enzyme. The wide applicability, the use of whole cells, the simplicity of procedure, the suitability for a microtiter plate format are very important properties for high throughput screening purposes. The alkali assay offers a powerful tool for important areas of applications such as screening for NAD(P)H-linked enzymatic activity of molecules of pharmacological and biotechnological interest and screening libraries of random mutants of NAD(P)H-dependent enzymes to allow design of new biocatalysts. The overall process is shown in Figure 13 and the procedure is described in Protocol 16.

232

PROTEIN ENGINEERING FOR BIOSENSORS

Protocol 16

High throughput screening for NAD(P)H-dependent activity Equipment and reagents Standard LB media are made using per liter 10 g of tryptone, 10 g of sodium chloride, 5g of yeast extract. Ampicillin stocks contain 100 mg/ml of ampicillin and are used to a final concentration of 100 (xg/ml and stored at — 20 °C when not in use. IPTG stocks contained 0.05M IPTG and are used to a final concentration of 0.5 mM and stored at — 20 °C when not in use. NADPH stocks contained 15 mM NADPH and are used to a final concentration of 1.5 mM and stored at — 20 °C when not in use. 9M NaOH buffer, 0.3M HC1 buffer, and 0.1M KH2PO4 and the pH of the latter was adjusted to eight using 10M KOH. The buffers are stored at room temperature. Microtiter plates and a fluorescence and/or absorbance microtiter plate reader are required.

Method 1

From a freshly grown (overnight at 37 °C) pick one Escherichia coli colony and transfer it into 5 ml of LB broth supplemented with ampicillin. Incubate overnight at 37 °C with vigorous shaking (230-250 rpm).

2 Dilute 1:100 the overnight culture in fresh LB supplemented with ampicillin, and incubate at 37 °C with vigorous shaking for 3-3.5 h. 3 Transfer 200 [d aliquots of the liquid culture in the well(s) of the microtiter plate and add IPTG to induce protein expression. 4 Incubate for 2 0 h a t 3 7 ° C with vigorous shaking. 5 Spin the cells at 4000 rpm for 10 min at 4 °C. 6 Remove the supernatant and resuspend the cells in 200 |ol 0.1M KH2PO4 pH 8. Check cell density by determining the absorbance at 600 nm using a microtiter plate reader after mixing the plate for 10 s. The optimal absorbance is ~0.3. If necessary, dilute the cells further in 0.1M KH2PO4 pH 8. 7 Add the substrates, including a positive and a negative control and mix the plate prior to incubating for 1 h at room temperature. If checking activity against a natural substrate the incubation length is 1 h. If checking activity against a nonnatural substrate, the incubation length is 2 h. 8 Add 1.5 mM NADPH and mix the plate prior to incubating for 1 h at room temperature. If checking activity against a natural substrate, the incubation length is 1 h. If checking activity against a non-natural substrate, the incubation length is 4 h. Cover the plates in foil because NADPH is photo-sensitive. 9 Stop the reaction by transferring 80 jol aliquot (mix first) to a new microtiter plate to which 80 |ol 0.3M HC1 was previously added and after mixing incubate at room temperature for 15 min. This step essentially lowers the pH to 1.0-2.5. 233

GIANFRANCO GILARDI Protocol 16 continued

10 Transfer 80 |ol aliquot to a new microtiter plate to which 270 jol 9M NaOH was previously added and after mixing, cover in foil and incubate in dark for 2.5 h. This step essentially increases the pH to 14.8. 11 After mixing, read the plate in the range from 340 to 550 nm and check for active mutants.

Functional characterization of the mutants Once the presence of the desired mutation has been confirmed by DNA sequencing, the protein can be expressed and purified. The yields should reach levels of at least 10 mg of pure protein per litre of bacterial culture to allow reasonable applications in the biosensor area. In the first analysis, the functional performance of the mutant should be checked with a number of complementary physical-chemical methods. The detailed functional characterization of the mutants aims at establishing what effect, direct or indirect, the mutation has on the function of the protein. Although X-ray crystallography and NMR spectroscopy are the methods of choice for a detailed analysis of the effect of the mutation on the structure and its consequences on function, the application of these techniques is not always quick and straightforward. Information on stability and folding can be obtained from circular dichroism, fluorescence spectroscopy, and calorimetry. Depending on the presence of cofactors or prosthetic groups, UV-vis and EPR spectroscopy can also provide useful information on the active site of enzymes. Stopped flow spectroscopy can also provide a detailed description of the enzyme kinetics. As a whole these investigations should confirm that the desired function has been achieved without significant perturbation of the other properties. This allows the establishment of general principles for the design in iterative cycles as shown in Figure 2. This task is not trivial because the effect of a mutation on the overall three-dimensional scaffold of the protein is not always obvious.

Other aspects of protein engineering This chapter has so far covered the most widely used methods for engineering proteins. There are many other aspects that cannot be adequately covered in the space limits of this chapter, but I would like to briefly address some of them in this paragraph, pointing the reader toward the cited literature. Often, after the expression of mutants constructed by molecular biology methods, some level of chemical modification of the protein may be required. This is the case, for example, for the site-specific labeling of engineered unique cysteines as shown in Figure 1. Protocol 17 describes how to label the unique cysteine 337 of the S37C mutant of the maltose binding protein with the thiol-reactive fluorophore LANBD.

234

PROTEIN ENGINEERING FOR BIOSENSORS

Figure 13 Screening process for the identification of active NAD(P)H-dependent oxidoreductases in a library of random mutants.

Protocol 17

Fluoresence labelling of the S37C mutant of the maltose binding protein This method describes the labelling of the unique cysteine 337 of the S37C mutant of the maltose binding protein with the thiol-reactive fluorophore IANBD. Only minor changes on parameters such as length of incubation with the fluorophore may be needed to adapt this protocol to other mutants or other thiol-specific fluorophores. The final binding curve obtained from this experiment is shown in Figure 1.

235

GIANFRANCO GILARDI Protocol 17 continued

Equipment and reagents The mutant S337C is constructed, expressed, and purified as described in (1, 2). Dithiothreitol (DTT) is supplied by Sigma; prepare a stock solution 0.85 mg/ml. The source of the fluorophore IANBD (N-((2-(iodoacetoxy)ethyl)-N-methyl)-amino-7-nitrobenz-2-oxa-l,3-diazole) is Molecular Probes Inc. The IANBD stock solution is 0.9 mg/ml in dimethylformamide (DMF). The maltose is supplied by Sigma and the stock solution: 10 mM maltose in 10 mM potassium phosphate buffer, pH 7.0. Fluorescence emission measurements are made at room temperature on a standard fluorimeter, for example a Perkin-Elmer LS 50.

Method 1 Determine the concentration of the pure S337C mutant. This can be done spectrophotometrically using an s28o for mbp of 78,570 M^cm" 1 . 2 Work out the volumes so that 2 ml of protein are chosen to be labeled. 3 Incubate the S337C on ice for 30 min with five fold excess of dithiothreitol. This step is necessary to ensure that the cysteine is reduced and free to react with IANBD in the following step. We have seen a tendency of S337C to form dimers via intermolecular disulfide bonds. From the concentration of the protein, work out the volume of DTT to be taken from the stock solution. Typically this is around 20 |ol when a pure fraction of S337C is used directly after purification on a DEAE column as described in (2). 4 Add the appropriate volume of the IANBD stock solution to reach a 10-fold excess in relation to the protein. Usually this is around 100 |ol; larger volumes are not recommended because the IANBD is dissolved in DMF and this may affect the protein. Keep the mixture on ice for 30 min. This time is very crucial for efficient labeling of the protein and may need to be adjusted. Reproducible results are only obtained by control over the time and the temperature of this step. 5 After the reaction, promptly separate the excess free fluorophore by gel filtration on a Sephadex G25 column equilibrated with ice-cold 10 mM potassium phosphate buffer, pH 7.0. Perform this step in a cold room. 6 Collect 1 ml fractions, carefully keeping them on ice. 7 Measure the absorbance at 280 nm and 478 nm and plot an elution profile. The first peak that eluted in the void volume is the labeled S337C (called S337C-NBD), the second is the free fluorophore. 8 Calculate the molar ratio of fluorophore-to-protein (Cf/Cp) using the formula: C

f/ C p = (Anaxf

X

e280p)/(-A280 X emaxf -Amaxf X e 2 gof),

where the Cf is the molar concentration of fluorophore, Cp is the molar concentration of protein, Amaxf is the value of the absorbance maximum of the fluorophore (478 nm for IANBD, g2gop is the molar extinction coefficient of the

236

PROTEIN ENGINEERING FOR BIOSENSORS

Protocol 17 continued

protein at 280 nm, A280 is the total absorbance of the sample at 280 nm, gmaxf and fi2gof are the molar extinction coefficient of the fluorophore at the absorbance maximum and at 280 nm, respectively. In the case of IANBD bound to S337C, the relevant constants are: g280 S337C = 78,570/M cm; g280 S337C-NBD = 80,315/M cm; g478 IANBD = 26,000/M cm; g280 IANBD = 1745/M cm. 9 Select the fractions with Cf / Cp = 1 and pool them. These will be used for the fluorescence titrations with the ligand, maltose. 10 Adjust the concentration of S337C-NBD to be 1 |oM using 10 mM potassium phosphate buffer, pH 7.0. This is needed to allow the fitting to the model described by the hyperbolic binding curve described below. 11 Collect the emission spectra scanning from 500 to 600 nm upon excitation at 480 nm. An emission peak for the fluorophore will be observed at 530 nm. Store this first spectrum obtained in the absence of maltose. 12 Add aliquots of the 10 mM maltose stock solution in the same buffer by using a high-precision Hamilton syringe. Typical additions are 10 aliquots of 1 jol each, followed by 5 x 2 (0,1 and 3 x 10 |ol Collect and store the emission spectra at each step, but direct readings of emission at 530 nm are also adequate. 13 Plot the reading of the fluorescence emission at 530 nm against the concentration of maltose added in the cuvette. Fit the data to the equation: AFobs = AFmax[Mal]/(KD + [Mai]), where AF obs is the observed change in fluorescence intensity, AFmax is the maximum attainable change in fluorescence intensity, [Mai] is the molar concentration of maltose and Ku is the dissociation constant. A typical Ku for the S337C-NBD for maltose is 62 ± 0.2 |oM Figure 1. Proteins can also be created from first principle by solid state synthesis (38, 39). The four helix bundle fold is a popular target for the de novo design of model cytochrome and a range of binding proteins because of the relative simplicity of their intra-chain hydrogen bonding pattern and the well-understood hydrophobic effect that drives their folding. On the other hand, the progress in creating E-structures has been slower due to the complex long-range inter-chain patterns of hydrogen bonding that characterize this type of folding (40). A variety of programs are now available in guiding the rational de novo design of proteins, allowing to predict the folding of a particular sequence. An alternative approach to the de novo rational design of proteins is the combinatorial synthesis. Here libraries of peptides and proteins are generated either by chemical solid phase synthesis or by combinatorial synthesis of genes that are then expressed by Escherichia coli. For example, Haehnel and co-workers have developed a method for the combinatorial synthesis of four helix bundles attached to a cellulose membrane via a cleavable linker and assembled to a cyclic

237

GIANFRANCO GILARDI

peptide (41). The immobilized peptides can be easily screened for metal binding and redox activity for exploitation in electrochemical devices (42). Libraries of peptides folding into helical bundles can also be obtained with DNA methods using, for example, a binary pattern of polar and non-polar amino acids, in which the characteristic of the residue is specified but the exact side chain is varied randomly. This can be achieved by constructing a library of synthetic genes containing patterns of degenerate NAN codons (in which N denotes any base), where polar amino acids are required (lysine, histidine, glutamic acid, glutamine, aspartic acid, asparagine), and NTN where apolar amino acids are specified (methionine, leucine, isoleucine, valine, phenylalanine). This approach is successfully used by Hecht and co-workers for the creation of peptides with peroxidase activity (43). A whole range of possibilities are available for tagging proteins to facilitate their purification or immobilization or to obtain combined activities. Suitable plasmids allow to clone the gene of the target protein next to strings of typically 4-6 histidines (His-tags) or biotin-acceptor peptides, designed to be either at the N- or C-terminus of the protein. This has been used for a wide range of applications, for example, the immobilization of peroxidases (44, 45). On a larger scale, gene fusion is another method emerging in the bio-analytical area. Two examples of fusions are the green fluorescent protein and the maltose binding protein. These can provide useful systems for detecting gene expression or for immobilization on solid supports. The molecular Lego developed in our laboratory uses gene fusion to link electron transfer and catalytic modules of redox proteins and enzymes to build artificial electron transfer chain for amperometric devices (11, 12). Protein engineering has also contributed to achieve the controlled immobilization of protein on surfaces. This is a very important achievement for the rational design of sensing surfaces. Cysteine residues are very interesting targets for this purpose owing to their low occurrence in natural proteins and to the specific reactivity of the thiol of the side chain, in particular toward fluorescence and redox labels as well as gold surfaces. Hill and co-workers attached mutants of P450cam to gold surfaces via a unique, surface exposed cysteine (46). Scanning tunneling microscopy has proven to be an invaluable technique to study the coverage of proteins on atomically flat gold (47). The same approach has been used for other proteins such as azurin (48, 49).

6 Concluding remarks Proteins and enzymes are undoubtedly an attractive target for the analytical biotechnology area. Their dimensions are intrinsically in the nanoscale, they are formed by precise scaffolds folded to perform specific and often coupled functions such as binding, stereo-specific catalysis, pumping, and self-assembling. All these functions can be detected by an array of different biophysical methods that have made much progress allowing to reach the level of single molecule detection. This chapter has shown how the field of protein engineering 238

PROTEIN ENGINEERING FOR BIOSENSORS

has refined many different strategies to specifically tune or even create de novo the biological element to perform a wide range of desired functions imposed by the experimentalist. We are no longer limited by the properties or the availability of natural proteins. There is a very bright future for protein engineering in nanobiotechnology.

Acknowledgments I would like to acknowledge all the members of my group, past and present. Particular thanks to Professors Gerard Canters and Tony Cass for introducing me to the fields of protein engineering and biosensors.

References 1. Gilardi, G., Zhou, L. Q.., Hibbert, L, and Cass, A. E. G. (1994). Anal. Chem., 66, 3840-7. 2. Gilardi, G., Mei, G., Rosato, N., Agro, A. F., and Cass, A. E. G. (1997). Protein Eng., 10, 479-86. 3. Brune, M., Hunter, J. L., Corrie, J. E. T., and Webb, M. R. (1994). Biochemistry, 33, 8262-71. 4. Marvin, J. S. and Hellinga, H. W. (1998). J. Am. Chem. Soc., 120, 7-11. 5. Renard, M., Belkadi, L., Hugo, N., England, P., Altschuh, D., and Bedouelle, H. (2002). J. Molec. Bio!., 318, 429-42. 6. Schonbrun, J., Wedemeyer, W. J., and Baker, D. (2002). Curr. Opin. Struct. Bio!., 12, 348-54. 7. Risen, S. C. G. and Thornton, J. M. (2002). Curr. Opin. Struct. Bio!., 12, 374-82. 8. Hardin, C., Pogorelov, T. V., and Luthey-Schulten, Z. (2002). Curr. Opin. Struct. Bio!., 12, 176-81. 9. Guex, N. and Peitsch, M. C. (1997). Electrophoresis, 18, 2714-23. 10. Macarthur, M. W., Laskowski, R. A., and Thornton, J. M. (1994). Curr. Opin. Struct. Bio!., 4, 731-7. 11. Sadeghi, S. J., Meharenna, Y. T., Fantuzzi, A., Valetti, R, and Gilardi, G. (2000). Faraday Discuss., 116 135-53. 12. Gilardi, G., Meharenna, Y. T., Tsotsou, G. E., Sadeghi, S. J., Fairhead, M., and Giannini, S. (2002). Biosens. Bioelectron., 17, 133-45. 13. Tsotsou, G. E., Cass, A. E. G., and Gilardi, G. (2002). Biosens. Bioelectron., 17, 119-31. 14. Corpet, F. (1988). Nucleic Adds Res., 16, 10881-90. 15. Ravichandran, K. G., Boddupalli, S. S., Hasemann, C. A., Peterson, J. A., and Deisenhofer, J. (1993). Science, 261, 731-6. 16. Li, H. Y. and Poulos, T. L. (1997). Nature Struct. Bio!., 4, 140-6. 17. Williams, P. A., Cosme, J., Sridhar, V., Johnson, E. F., and McRee, D. E. (2000). Mo!ec. Cell, 5, 121-31. 18. Connolly, M. L. (1983). Science, 221, 709-13. 19. Nicholls, A. and Honig, B. (1991). J. Comput. Chem., 12, 435-45. 20. Kunkel, T. A. (1985). Proc. Not!. Acad. Sci. USA, 82, 488-92. 21. Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987). Meth. Enzymol, 154, 367-82. 22. Taylor, J. W., Ott, J., and Eckstein, F. (1985). Nucleic Acids Res., 13, 8765-85. 23. Nakamaye, K. and Eckstein, F. (1986). Nucleic Acids Res., 14, 9679. 24. Higuchi, R., Krummel, B., and Saiki, R. K. (1988). Nucleic Acids Res., 16, 7351-67. 25. Sarkar, G. and Sommer, S. S. (1990). Biotechniques, 8, 404-7. 26. Derbyshire, K. M., Salvo, J. J., and Grindley, N. D. F. (1986). Gene, 46, 145-52. 27. Leung, D. W., Chen, and Geddel, D. V. (1989). Technique, 1, 11-15.

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GIANFRANCO GILARDI 28. Cadwell, R. C. and Joyce, G. J. (1995). Mutagenic PCR. In PCR primer: A laboratory manual (ed. C. W. Dieffenbach and G. S. Dveksler, pp. 583-9, CSHL Press, New York. 29. Stemmer, W. P. C. (1994). Proc. Natl. Acad. Sd. USA, 91, 10747-51. 30. Stemmer, W. P. C. (1994). Nature, 370, 389-91. 31. Zhao, H. M., Giver, L, Shao, Z. X., Affholter, J. A., and Arnold, F. H. (1998). Nature Biotechnol, 16, 258-61. 32. Ostermeier, M., Shim, J. H., and Benkovic, S. J. (1999). Nature Biotechnol, 17, 1205-9. 33. Sieber, V., Martinez, C. A., and Arnold, F. H. (2001). Nature Biotechnol, 19, 456-60. 34. Petrounia, I. P. and Arnold, F. H. (2000). Curr. Opin. Biotechnol, 11, 325-30. 35. Cirino, P. C. and Arnold, F. H. (2002). Curr. Opin. Chem. Biol, 6, 130-5. 36. Joo, H., Lin, Z. L, and Arnold, F. H. (1999). Nature, 399, 670-3. 37. Farinas, E. T., Schwaneberg, U., Glieder, A., and Arnold, F. H. (2001). Adv. Synth. Catal, 343, 601-6. 38. Wilken, J. and Kent, S. B. H. (1998). Curr. Opin. Biotechnol, 9, 412-26. 39. Borgia, J. A. and Fields, G. B. (2000). Trend. Biotechnol, 18, 243-51. 40. DeGrado, W. F., Summa, C. M., Pavone, V., Nastri, F., and Lombardi, A. (1999). Annu. Rev. Biochem., 68, 79-819. 41. Rau, H. K, Dejonge, N., and Haehnel, W. (2000). Angewandte Chemie-International Edition, 39, 250-3. 42. Willner, L, Heleg-Shabtai, V., Katz, E., Rau, H. K., and Haehnel, W. (1999). J. Am. Chem. Soc., 121, 6455-68. 43. Moffet, D. A., Certain, L. K., Smith, A. J., Kessel, A. J., Beckwith, K. A., and Hecht, M. H. (2000). J. Am. Chem. Soc., 122, 7612-13. 44. Presnova, G., Grigorenko, V., Egorov, A., Ruzgas, T., Lindgren, A., Gorton, L, and Borchers, T. (2000). Faraday Discuss., 116 281-89. 45. Zhang, J. K. and Cass, A. E. G. (2000). Anal Chim. Acta, 408, 241-7. 46. Lo, K. K. W., Wong, L. L, and Hill, A. O. (1999). FEBS Lett., 451, 342-6. 47. Davis, J. J., Hill, H. A. O., and Bond, A. M. (2000). Coord. Chem. Rev., 200, 411-42. 48. Chi, Q. J., Zhang, J. D., Friis, E. P., Andersen, J. E. T., and Ulstrup, J. (1999). Electrochem. Commun., 1, 91-6. 49. Chi, Q.. J., Zhang, J. D., Nielsen, J. U., Friis, E. P., Chorkendorff, L, Canters, G. W., Andersen, J. E. T., and Ulstrup, J. (2000). J. Am. Chem. Soc., 122, 4047-55.

240

Index ABD-F 114, 124 ABD-N 112, 114-15, 119, 123 determination of zinc concentration 122 ABD-T 123-4 Accelrys web site 189 acetone-soluble photoresist 158 acetylcholine 167 esterase 8, 58 acousto-optical tunable filters 110 acrylic resin 43 acrylodan 186 acryloylfluorescein 136-7 A.C.S. reagent grade buffer salts 117 active dyes 44 affinity sensors 10 agarose gel electrophoresis 210, 228 AgCl-coated Ag wire 170 alamethicin 165-7 Albery, W. J. 87 alkyd resin 43 aluminium oxide, A12O3 42 amine amplification 143 ammonium ion 53 amperometric enzyme electrodes 70, 89 types of 69 analytical protein engineering 187 anionic interferants 9 anisotropy calibration of 124-8 theory of determination of metal ions based on 111-14 antibiotics, preparation of most commonly used 220 antibody-antigen binding reaction 91 apoCA 123 apocarbonic anhydrase 117, 123 apolar amino acids 238 apoprotein, preparation of 116-17 aqueous microsensor suspension 139 Arnold, F. H. 231 aryl sulfonamides 111-12 affinity chromatography column 116 ascorbate 9 Ashworth, D. 87 asolectin 167 aspartic acids 142 Atanasov, P. 88 automated immobilization 44 azurin 238

back-crossing 230 Bannon, T. 87 Bartlett, P.N. 59, 80, 87, 89 BAS 100 170 Battaglini, F. 87 BCECF-AM (2',7'-bis (2-carboxyethyl)-5(and 6)carboxyfluorescein acetoxymethyl ester) 149, 150 Beer-Lambert law 90 benzoin dimethyl ether 44 benzophenone 44 benzoquinones 44 Bergel, A. 87 Bicine 116, 121 binding agent 43 Bioanalytical Instruments 170 BioDot Inc. 48 Biojet QuantiSOOO™ 48 biological oxygen demand (BOD), quick estimation of 54 biological selection 230 bio luminescence 100 bioluminescent marine bacteria 100 biomimetic or molecular imprinting 54 bio mimetic sensor 55 Bio-Rad Chelex-100 118 Bio-Rad metal-free pipet tips 118 biorecognition process and transduction mechanism, kinetics of 60 biosensing 109 biosensors amperometric and potentiometric 7 arrays 132 design of 97 made with polycationic redox polymers 9 potentiometric and conductimetric 60 Biosym/MSI 188 Biotech Validation Suite for Protein Structures 188 biotin-acceptor peptides 238 black lipid membrane (BLM) technique 154 literature 165 Blum, L. J. 97 bovine serum albumin (BSA) 34, 105 Briggs, G. E. 61 buffers that contain metal ions such as Ca or Mg, preparation of 119

Bacha, S. 87 Bacillus megaterium 199, 232 haem domain of CYP102, and flavodoxin from Desulfovibrio vulgaris, ribbon diagram of complex between 200

CA117, 119 Ca(II) 111 calcein acetoxymethyl ester (calcein-AM) 148 Calvo, E. J. 87

241

INDEX Cambiaso, A. 87 carbamates 52 carbonic anhydrase 111 carbon paste (CP) 43 carbon paste electrodes (CPEs) 43 carbon-a three-dimensional models for the human CYP2D6 191 Caruana, D. J. 19 Cd(II) 111 cell array fabrication 148 encoding 147 viability 148-9, 150 assay with BCECF-AM 150 test via pH sensitive nanospheres 150 cellulose acetate 46 ceramic 42 channel-forming peptides 165 chelating resin 118 chemiluminescence 101 chemiluminescent choline biosensor 107 Chen, Y. 87 chimeric DNA sequences, library of functional screening 230-4 chimeric genes 229 chip fabrication and pretreatment 157 chlorophenol sensor 107 cholera toxin 168 cholesterol 51, 180 esterase 8 oxidase 8 choline oxidase 8, 10 cholinesterase 53 clad, refractive indices of 131 CnBr coupling 28 Co(II) 111 cobalt 43 Coche-Guerente, L. 88 Colicin N 174-5, 177 collagen 101 membranes, enzyme immobilization on 102 colloidal silica 125 combinatorial cassette mutagenesis 224 combinatorial chemistry 153 combinatorial genetics 153 combinatorial synthesis 237 commercial coffee creamer 125 complementary DNA 202 composite electrodes, advantage of 2-3 Comtat, M. 87 concentration change in, and net flux, schematic diagram showing relationship 83 gradient 64 conductive inks 43-4 Connolly algorithm 193 surface 199-200 contamination metal ions, preparation of solutions free of 118 convection and biosensors 64

242

cosmids see plasmids Coulet, P. R. 97 Coulter counter-like current modulations 164 coupled chemical reaction 68 coupled diffusion/reaction problem solving 74 Cramer, M. L. 109 Cranfield University 42 Crank-Nicholson (CN) method 85-6 creatinine 51 crosslinkers 4 crosslinking 1-2, 75 Cu and other ions, determination using reagentless approach 123-4 Cu(II) 109, 111 Cuvettes 127 CYP102 193-4, 197, 199 cysteine 238 337 234-5 cysteinyl residues 123 cysteinyl thiol 124 cystic fibrosis 167 cytochrome, human (CYP) P450 92, 231, 232

enzymes 189, 230 sequence alignment 190 Danilowicz, C. 87 dansylamide (DNSA) 116 dansylaziridine 124 Darwinian evolution 224 Daunert, S. 147 dehybridization 145 dehydrogenases 101 Delfino, L. 87 DelPhi 197, 200 see also Insight II Deng, J. Q, 87 Denhardt's buffer 34 deoxynucleotides (dNTPs) 202 analogs 201 Desprez, V. 88 Desulfovibrio vulgaris 199 flavodoxin from 200 see also Bacillus megaterium dextran sulphate 38 dialysis 51, 116 Diard, J. P. 88 Diaz, L. 87 2,4-dichlorophenoxyacetic acid 52 dielectric inks 43-4 diffusion and biosensors 64 limited first-order Langmuir model 36 digital simulation, use in electrochemistry 82 4-(4-(dimethyllaminopherylazo) benzoic acid) (DABCYL) 145 dioleoyl phosphatidyl choline (DOPC) 174 1,2-diphytanoyl-sn-glycero-3phosphocholine (DphytPC) 167,180

INDEX dipicolinic acid (pyridine-2,6-dicarboxylic acid; DPA) 115-16 dipole interactions 20 directed evolution 187, 222-34 protocols error prone PCR 225 DNA shuffling 227-8 high throughput screening for NAD(P)H-dependent activity 233 QuikChange® PCR mutagenesis 223-4 steps of iterative cycles of 224 disposable biosensors 44 dithioerythritol (DTE) 124 DMF 114, 119 DNA array fabrication 26 detection 144 double stranded see dsDNA hybridization on solid surfaces 19 nonhomogeneous heat denatured sheared 34 polymerase 199, 203-5, 213 probe, non-fluorescent 144 sensor technology 54 shuffling in directed evolution 227-8 DNA-DNA interactions 144 DNAses 20 drier 43 Driscoll, B. J. 87 dsDNA 22, 200, 202 exponential growth as function of number of PCR cycles 205 DuPont Ludox 125 DUTPase(dut~) 201 Eadie-Hofstee plots 63 electrocatalytic activity of redox hydrogels and "wiring" of enzymes 2 electrochemical-based detection of hybridization 26 electrochemical cell, two-electrode with Ag/AgCI counter electrode and gold working electrode, schematic view 175 electrochemical systems method, digital simulation of 86 electrochemiluminescence (ECL) 97, 101 fiber-optic biosensor based on 107 measurements, flow system for 100 electrons conducting redox polymers in biosensors 1 diffusivity of 2 transfer between enzyme and polymer redox centers 4 electrophoretic vesicle positioning in two different configurations 164 electroreduced chemicals 1 electrostatic interactions 20 electrostatic potentials 196 electroxidized biochemicals, examples of 1

enzyme electrodes 4, 8 basic electrochemistry 63-9 behavior model 66 immobilization on collagen membranes 102 on preactivated polyamide membranes 102 kinetics 60 data analysis 62 equilibrium and steady state 60 membrane|electrode problem, case diagram for 80 overcoating with membrane 8 wiring redox polymer 4 enzyme-based sensors 134 enzyme-containing redox hydrogels 1 enzyme-labeled microspheres 141 enzyme-mediator kinetics 89 enzyme/pH array, fabrication of 137-8 enzymes covalent immobilization 142 with high chemical and thermal stability 222 immobilization 103 with glutaraldehyde 142-3 by physical absorption 141 loading of IDA sepharose beads 104 sensors, disposable 41 enzyme-substrate (ES) complex, 61, 74 kinetics 89 epoxy resin 43 Eppendorf Microfuge tubes 118 EPR spectroscopy 234 Epstein, J. R. 131 error prone PCR 224, 226 in directed evolution 225 Erythrosin B 127 Escherichia 232 Escherichia coli 174, 200, 237 XLl-Blue 230 codon usage by 206-7 Escherichia.coli dam+ 221 Escherichia coli, dut~ung~ 201 ETH 1001 173-4 ethanol 119 ethanolamine 114 ethidium homodimer-1 (EthD-1) 148-9 ExacTech 42 excitation light introduced into fiber 132 external encoding in fluorescence based fiber-optic biosensors 140 Feldberg, S.W. 82 ferrocene/ferricinium ion couple 41 ferrocenes 44 enzymatic electrochemistry mediated by 42 fiber optic biosensors 132-3 advantages 151 bio-, chemi-, and electrochemiluminescence for 97 243

INDEX fiber optic biosensors (Continued) enzymatic 136 fluorescence-based 131 light-emitting reactions 100-1 preparation of sensing layer immobilization techniques 101 protocols chemical modification of HRP with His-tag 104 enzyme immobilization on collagen membranes 102 enzyme loading of IDA sepharose beads 104 preparation of sensing layer with photocross-linkable polymer (PVA-SbQ) 104 sensing tip of 98 setup 98 fiber-optic based arrays of cells 147-8 fiber-optic microarray with molecular beacon (MB) functionalised microspheres 147 fiber polishing 133 Pick first law 64-6, 84 second law 66-8 Ficoll 34 Fierke, C. 119 filter hybridization 34 studies 38 firefly 100 fish freshness estimation 52 Fitzgerald, C. A. 87 flavodoxin 199 Fleming, M. S. 131 flow-cell detection system 91 Flow Injection Analysis (FIA) 98 measurements in a batch system 98 in a flow system 98-100 fluorescein 126-7, 136, 145 fluorescence intensity of apo-CA 120 lifetime 113 sensing 110 measurements and imaging instrumentation used for 134 microscopy 109 polarization immunoassay 109 fluorescence anisotropy of apo-CA plus ABD-N as a function of free Zn(II) concentration 121 of apo-N67C-ABD-T as a function of the concentrations of Cu(II), Zn(II), Cd(II), Ni(II) and Co(II) 123 determination of metal ions by 109 microscopy 110 fluorescence based fiber optic biosensors 137, 141 protocols acryloylfluorescein, preparation of 137

244

amine amplification with PEI143 cell viability: assay with BCECF-AM 150; test via pH sensitive nanospheres 150 chemical etching of germania-doped imaging fiber bundle 139 enzyme immobilization with glutaraldehyde 142-3 by physical absorption 141 external encoding 140 fabrication of enzyme/pH array 137-8 of fiber-optic based living array of cells 148 internal encoding 140 PAN-enzyme sensors 135-6 periodate oxidation 143 preparation of molecular beacon-modified microspheres 146 silanization/functionalization of fiber tip 135 ssDNA probe labelled microspheres, preparation of 145 fluorescent aryl sulfonamide 123 for zinc(II) determination 114-15 fluorescent calcein 149 fluorescent dyes 145, 147 fluorescent sulfonamide 123 fluorophores 127 flux diagrams for membraneenzymeelectrode, simplified 70 of different kinetic processes for amperometric membrane] enzyme|electrode 70 "flyers", identification of 127 formaldehyde 25 formamide 36, 38 Forster transfer 123 Fox, R. O., Jr. 166 free amine groups on a microsphere surface 142 free radical polymerization 136 free zinc concentration 119-20 functional characterization of mutants 234

functional screening 2224 fused quartz 127 fused silica core 131 Gajovic, N. 88 gel filtration 116 gene fusion 238 generalized biosensor, schematic diagram of 90 Genies, E. 87 Georgiev, T. 88 germania-doped imaging fiber bundle, chemical etching of 139 germania doped silica cladding 138 G-factor 127, 188

INDEX giant unilamellar vesicles 160-2 formation of 157, 161 Gilardi, G. 185 Gillespie, D. 25 Giovangrandi, L 153 glass fiber 42 glass fiber-optic bundle 97 glucoamylase activity 52 production, control of 52 glucose 52, 107 blood, pocket meter 51 electrooxidation 3 monitors, personal 42 oxidase 5-6 sensors 46 glutamic acids 142 glycogen 125 glycols 23 glycoproteins 103 glyoxal (ethanedial) 25 gold electrode 168, 174 tethered lipid bilayers 179 surfaces 168, 177, 238 Gooding, J. J. 87 graphite screen-printed electrodes 54 Grattarola, M. 87 gravimetric sensors 60 green fluorescent protein 238 Greer, J. C. 87 Gros, P. 87 Gryczynski 110 Guiseppi-Elie, A. 88 Haehnel, W. 237 haem-thiolate monoxygenases 189, 231 Haldane, J. B. S. 61 Hall, E. A. H. 87 Hamamatsu R928 PMT 127 Hanes plots 63 Hanke, W. 167 heavy metals 117 Heller, A. 1 HEPES121 sodium sulfate 116 Hibbert, D. B. 88 Higgins, I. J. 42 Higuchi, R. 202 Hill, H. A. O. 238 Hindle, A. 87 4-6 histidines (His-tags) 103, 177, 238 holoprotein 116 hopscotch method 86 horseradish peroxidase (HRP) 8, 10 with His-tag, chemical modification of 104 mutant Asn255Asp (HRP1A6) 231 HPLC grade water 125 Huang, T. 88 human cytochrome see cytochrome

human diseases, point mutation 54 Human Genome project 144 human serum albumin antigen-antibody complex 92 Hunt,]. B.115 hybrid lipid layer description 170-1 hybridization base mismatch 33 of DNA on nitrocellulose membranes 27 kinetics at solid surface 31 mass transport 34 nonspecific adsorption 34-5 at oligonucleotide sensitive electrodes 19 process of 38 of redox hydrogel bound DNA sequence 11 sensors 10 sequence discrimination 37 target, double stranded 38 temperature 31 hydrated silica matrices 7 hydrogen bonds 20 electrode 41 peroxide 10 hydrophobic interactions 20 hydroxides 44 IANBD 185-6 thiol-reactive fluorophore 234-5 imaging fiber bundle 132, 136 iminodiacetate residues 118 immobilization of "His-tag" enzymes 103 of large proteins 142 of peroxidases 238 via polymer photodeposition 136 impedance modules 170 spectral analysis 169 spectroscopy 153 basics of 168 data 172 measuring technique and electrochemical cell 170 of tethered lipid membranes 168-82 ink jet printing technique 44, 48-9 inks typically used for screen-printed biosensors 45-6 Insight II 188-9, 193-4, 196 internal encoding in fluorescence based fiber-optic biosensors 140 in vitro DNA synthesis 201 in vitro evolution process 230 in vivo glucose sensors 10 in vivo selection steps 201 ion channels activity electrical recordings of 153 in tethered lipid membranes on gold electrodes 154 245

INDEX ion channels (Continued) self-integrating ionic strength 32, 38 ionophore 173 ion selective electrodes 109 Iteractive Truncation for Creation of Hybrid Enzymes (ITCHY) 229 Jobst, G. 88 Joslin, T. 86 Kalia, Y. N. 87 Kawski 124 kidney dialysis 51 kinematic viscosity 65 kinetic modeling of biosensors 59 applications 89-92 list of symbols 92-4 of enzyme electrodes, summary of literature on 87-8 immunosensors 91-2 optical and photometric biosensors 90 potentiometric enzyme electrodes 90 kinetic models case diagrams 79-81 deriving complete 78 experimental verification of approximate analytical 81 simplifying assumptions 70 Kirtikara, K. 88 KM significance for biosensor applications 63 Kong, J. L 87 Krishnan, P. 88 Krull 31 Kunkel method 200-2 Kunkel, T. A. 201 Kurganov, B. L. 87 Labbe, P. 88 labelling enzyme 10 lactate 51-2, 107 lactate oxidase 6 Langmuir model 36 Langmuir-type isotherm 36 light-emitting bi-enzyme system of marine bacteria 101 Lineweaver-Burk plot 62-3 lipid bilayer selective antibody binding to SLIC in 180 for single-channel experiments, highly insulating, tethered 179-82 tethered via surface-attached proteins description 177 impedance spectroscopy data 179 liquid core fiber-optic 98 Liu, H. Y. 87 live-cell array fabrication 146 246

LIVE/DEAD Viability/Cytotoxicity Kit 148 luminescence and fiber-optic biosensors, enzyme immobilization on membranes Immunodyne ABC type from Pall-Gelman 103 on Ultra Bind type membranes 103 luminescence sensors analytes involving H2O2 detection 107 dehydrogenases as auxiliary enzymes analytes using 105 determinations with 105-6 oxidases as auxiliary enzymes 107 luminol chemiluminescence 107 luminometer 97-8 Luria-Bertani (LB) broth 218 Lyons, C. H. 87 Lyons, M. E. G. 87 lyophilization (freeze-drying) 114 Magner, E. 87 Maines, A. 87 7-(5-maleimidyl)-pentylaminobenz-2oxa-1,3-diazole-4-sulfonamide) 123 Maliwal, B. 109 maltose 52 maltose binding protein (MBP) 238 mutant S337C 185-6, 234 manganese 43 Marchesiello, M. 87 marine bacteria 101 Martens, N. 87 mass transport 63 Mayer, M. 153 Mears, D. J. 88 mediated enzyme electrode 41 mediator-based sensors 1 Medisense Inc. 46 megaprimer 212 melittin 165, 178 membrane | enzyme | electrode configuration 89 flux diagram for 70 flux equations, solution of 73 problem solving 74 Menten, M. L. 60 (3-mercaptoethanol 124 metabolites measured with screen-printed sensors 51 metal ions 5 contaminants, removal from solutions and buffers 119 contamination, avoidance of 117-18 and fluorescence anisotropy, protocols ABD-N, synthesis of 115 anisotropy, measurement and calibration of 128 apoprotein, preparation of 116-17 contamination metal ions, preparation of solutions free of 118 reagentless approach, determination of Cu and others using a 124-5

INDEX zinc concentration determination with ABD-N 122 Mg(II) 111 Michaelis constants 8, 91 in conventional enzyme kinetics 73 Michaelis, L. 60 Michaelis-Menten enzyme kinetics 60-2, 75, 79 one-dimensional case diagram for 79 equation, linear plots derived from 63 micromembranes, reproducible 8 microspheres sensors 139 molecular beacon-modified 146 ssDNA probe labelled 145 microwells 132 array platform preparation 138 migration and biosensors 64 MINEQ.L 121 modelling kinetic 69-89 purpose and practice of 59 molecular beacons 145 molecular diversity 224 molecular evolution, natural 222 molecular probes 148-9 monochromatic laser sources 125 monoclonal antibody Sp3E9 178 monolayer of thioalkanes 168 MOPS 121 Moser, I. 88 Mullard Prize 42 multi-analyte arrays 136-50 sensing 132 multipurpose biosensor 52 mutagenesis 185 mutagenic oligonucleotide 200 primers, design of 205 NAD(P)H 100, 231 dependent activity in directed evolution, high throughput screening for 233 dependent enzymes 232 Nation 46 NAN codons 238 nanocomposite enzyme electrodes 2-3 nearest neighbour interactions 22-3 Neher, E. 154 Nernst diffusion layer 73-4 equation 90 Neykov, A. 88 Ni(II) 111 nicotinamide adenine dinucleotide (NADH) oxidation at poly(aniline) electrodes 81 see also NAD(P)H Nigretto, J. M. 87 NIH 3T3 mouse fibroblast cells 148 nitrilotriacetic acid 121, 177

nitrocellulose 26, 42 NMR spectroscopy 234 N-succinimidyl esters of carboxytetramethylrhodamine (TAMRA) 140 functionality 135 NTN238 nucleic analogs 20 nucleotide duplex, structure of 20 numerical methods, explicit 82 numerical simulation methods 82 oligonucleotide probes attachment to gold by chemisorption 27 in base pairs, length of 23 deposition onto a polymer coated electrode 28 non-covalent or physisorption 27 protocols attachment of ligonucleotides 28 CnBr activated coupling to an edge plan graphite electrode 29 determination of solution concentration of single stranded oligonucleotide and assessment of purity of oligonucleotide 26 reactive electrophoretic deposition 30 oligonucleotides 207 arrays 144 chemical attachment 28 devices 21 hybrids, stability of 36 primers 212-13, 224 strands, hybridization of two complementary 21 oligonucleotide sensitive electrodes 21 determination of melting temperature 32 function of 20 hybridization conditions 30 efficiency and sensitivity 21 OmpF 168, 175 in lipid bilayers 177 trans-membrane protein 174 "one-shot" biosensor 42, 44 optical fiber 131, 133 based DNA sensors 144 optical sensors 60, 104 optical transduction mechanisms 131 optical waveguide 97 organic solvents 43 organophosphate pesticides 52 orthogonal collocation method 86 Os-complex mediator 52 osmium 4 oxidation polymerization 44 oxygen electrode 41 P450 see cytochrome PAN see poly (acrylamide-co-Nacryloxysuccinimide) patch clamp on chip, concept of 154 247

INDEX patch pipet 154 PCR without added primers 227 amplification procedures 144 experiment, schematic diagram of 213 megaprimer method 212 methods 200-2 site-directed mutagenesis 202 thermal cycles 205 penicillin 137 Penicillium chrysogenum 137 peptide nucleic acids (PNA) 20, 32 peptides folding into helical bundles 238 periodate oxidation fluorescence-based fiber-optic biosensors 143 peroxidases 10 peroxides 44 Perrin equation 112, 114, 126 pesticides 52-3 Pfeiffer, D. 87 PjuTurbo DNA polymerase 221 pH range 36 phenanthroline 115 phenolic resin 43 Phloxine B 127 phospholipid (POPC) 171, 174 phosphorothioate method see Kunkel method Photinus pyralis 100 photocross-linkable polymer (PVA-SbQ), preparation of the sensing layer with, fibre-PLB see planar lipid bilayers photoluminescence techniques 109 photomultiplier tube (PMT) 127 of luminometer 97-8 photosensitive agents 44 planar bilayers on chip 157, 165 planar lipid bilayers (PLB) 153-4, 156, 165 formation of 157 planar patch clamp 153-68 principle of 155 protocols dry cleaning and activation of silicon chips 158 fabrication of silicon chips 157-8 fabrication of Sylgard® pads 165 giant unilamellar vesicles, preparation of 161 PLBs, formation on silicon chip 162 surface modification of silicon chips 160 wet cleaning and activation of silicon chips 159 planar silicon microchips 154 plasmids and cosmids, most commonly used 221 Plasmodium falciparum 178 PLB see planar lipid bilayers Fletcher, D. 86

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Poisson-Boltzmann algorithm 197-8 polar amino acids 238 poly(4-vinylpyridine) (PVP) 3-4 poly (acrylamide-co-Nacryloxysuccinimide) (PAN) 135 enzyme sensors 135-6 gel immobilization 135 polymer-based pH sensor 136 polyamide, synthetic-like derivatized 101 polycarbonate 42 polychlorinated biphenyls 52 poly(dT) 23 poly-ethyleneimine (PEI) 142 poly-L-lysine 160 polymer microspheres 142 photodeposition 136 polymerase chain reaction see PCR polynomial curve fitting 86 poly(N-vinylimidazole) (PVT) 3 polyurethane resin 43 poly(vinylalcohol) 101 bearing styrylpyridinium groups (PVA-SbQ) 104, 107 polyvinylpyrrolidone 34 POPC see phospholipid Pratt, K. F. E. 87, 89 primers 207 printing process 48-9 probe attachment of 25 length and orientation 22-5 oligonucleotides hybridization kinetics 36-8 structure and dynamics 22-30 progesterone 51 protein-DNA interactions 144 protein engineering for biosensors 185, 187, 189, 191, 197, 234-5 protocols calculation of a map of electrostatic potentials on a protein surface 197-9 creation of three-dimensional models 191-3 fluoresence labelling of the S37C mutant of the maltose binding protein 235-7 molecular graphics, display and basic calculations on protein structures 194-6 protein sequence alignment 189-90 proteins controlled immobilization on surfaces 238 modeling and calculations on protein structures 188 physical adsorption 141 rational design of, de novo 237 Pseudomonas putida 231 PVC 42-3

INDEX pWhitescript™ plasmid 221 Pyrococcus furious 202 quaternary ammonium salts 33 Quickchange™ PCR method 201, 221, 228 mutagenesis in directed evolution 223-4 schematic procedure for 222 Ramachandran plots 188 Raman band of water 125 scatter 125, 127 signal excitation in UV 125 random collision between two complementary strands 20 random mutagenesis, error prone PCR 224-6 Randriamahazaka, H. 87 rational design 199 flow chart showing experimental stages 187 rational protein engineering 187-222 Rayleigh scatter 127 reaction schemes for CNBr activated coupling and electrophoretic deposition on electrodes 29 "reagent" approaches 111-12 "reagentless" anisotropy-based material ion determination 113 approach 112 determination of Cu and others by using a 124-5 reassociation 33, 38 reciprocal expressions, advantages of using 73 recombinant DNA technology 185 recombination 224 DNA shuffling 226-30 redox hydrogel based electrochemical biosensors 1 containing immobilized enzymes, cross-linked 75 redox polymer-enzyme system microscopic homogeneity and salt effects in 6 optimal compositions 6 redox polymers 1 cross linked 2 and their electrochemistry 3 and enzymes 7 redox proteins 199, 238 Rhodamine-based dyes 140 Rhodes, R. K. 87 Richards, F. M. 166 RNAses 20 RNA sequence 11 rotating disc electrode 65, 68

rotation speed 65 rotational correlation time 113 Sakmann, B. 154 SAMCO "Pasteur" pipets 118 Sansom, M. S. 167 saturated calomel electrode (SCE) 4 Schauer, C. L. 131 Scheller, F. W. 88 Scheller 41 Schlue, W. R. 167 Schulmeister, T. 87-8 screen-printed biosensors, applications 51 in clinical diagnosis 51 in environmental monitoring 52-4 in food analysis bioprocess control 52 and their performances 53 screen-printed lactate sensor 52 screen-printing apparatus 46 for biosensors 41 diagram of preparation 48 protocol for screen-printed glucose electrode, preparation of 50 process 46 technology 42-50 segmental mobility 1 semipermeable membranes 46 sensing element 44-7 sensor examples, specific 7 measuring turnover rate of hydrolytic and other non-redox enzymes 8-9 made with diffusional mediators 9 "self-encoding" 140 Sequence Homology Independent Protein Recombination (SHIPREC) 229-30 sequential chemisorption reactions of polymers 8 Sheppard, N. F. 88 Shults, M. C. 87 silanization/functionalization of fiber tip 135 silanized oxide surfaces 168 silica core chemically etched 138 silicon chip for planar patch clamp measurements, schematic diagram of 156 single analyte sensing 132, 134 single stranded DNA see ssDNA single stranded oligonucleotides attached to surface, schematic of 24 complementary, equilibrium constant for hybridization process for two 21 synthetic 54 single-channel conductance 167 recording of alamethicin pore, chip-based 166-7

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INDEX SiO2 157 site directed mutagenesis 199-222 PCR method protocols hot start PCR 207-9 megaprimer PCR 213-15 purification of PCR products from agarose gels 211-12 submarine agarose gel electrophoresis of DNA 209-10 protocols isolation of plasmid from bacterial host 218-20 ligation of DNA insert in a vector 216-17 restriction digest of DNA 215-16 transformation of plasmid in bacterial host 217-18 schematic diagram for 200 site-directed mutants 187 sodium chloride 32 Solartron 1296 170 sol-gel method 7 solid state synthesis, proteins created by 237 Somasundrum, M. 88 sorbitol determination 106 Sorochinskii, V. V. 87 Southern blots 27 soybean peroxidase (SBP) 10, 35 Sp3E9 179-80 spacers, amino-based 23 Spectrosil 127 Speigelman, S. 26 SR 850 170 ssDNA (single stranded DNA) 22, 200 artificially synthesized 145 native 144 template 202 Staggered Extension Protocol (StEP) 228-9, 232 Stanford Research Systems 170 starch 52 steady-state enzyme kinetics 60 Stemmer, W. P. C. 226, 230 StEP see Staggered Extension Protocol steric hindrance to hybridization 22 steroids 231 Stitzel, S. E. 131 Stora, T. 153 Stratagene 221 substrate electrooxidation or electroreduction 2-3 matrix 42 sucrose 52 sulfite 53 sulfonamide, affinity constant of 112 sulfonyl halide 114 supported lipid monolayers 168 Suprasil 127 surface concentration 22 of single stranded oligonucleotides 23

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surface plasmon resonance (SPR) technique 91 SWISS-MODEL, three-dimensional model built by 188 Swiss-PdbViewer 188 Sylgard® pad 157 synthetic ligand-gated channel (SLIC) 168, 178, 180 T4 DNA polymerase 200 Tan, T. C. 87 Tanticharoen, M. 88 Taq polymerase 224 Tatsuma, T. 87 TCEP (tris-(2-carboxyethyl)phosphine, hydrochloride) 124 Tebbutt, P. 87 Terrettaz, S. 153 Terrific Broth (TB) 218 tethered lipid bilayers 168 description 174-6 impedence spectroscopy data 176-7 protocols Ca2+ -sensitive hybrid lipid layer electrode, preparation of 172 OmpF-containing lipid bilayer tethered on a gold electrode, preparation of 175 proteolipid bilayer tethered by SLIC to gold electrode, formation of 178-9 tetradecanethiol 171 tetraethylammonium 33 T-format optical configuration 110 Therias, S. 88 thermal curing 43 thermal polymerization 44 Thermococcus litoralis 202 Thermus aquations 202 thioalkane 171, 174 thiol-specific fluorophores 185-6 Thompson, R. 109-10 three-dimensional model of target protein, construction of 188 three-electrode system sensor 48 Toh, C. S. 59 Tongta, A. 88 toxicity screen of antiseptics and antioxidants 147 transducer 89 transition metal ions 9 Turner 42 two photon excitation 110 two-electrode configuration sensor 48 electrochemical cell, schematic view of 171 Tyrrell, C. H. 87

INDEX ultraviolet (UV) light 136, 234 curing process 44 inducible immobilization 43 Updike, S. J. 87 uracil-containing DNA 201 uracil N-deglycosidase (ung~) 201 urate 9 Urban, G. 88 urea 51 uric acid 51 Vadgama, P. 87 van der Waals interactions 20 Verreschi, G. 87 vesicles 161-2 see also giant unilamellar vesicles Vibrio fischeri 100 Vibrio Iwrveyi 100 Vogel, H. 153 Walt, D. R. 131 Warsinke, A. 88 Watanabe, T. 87 Watson-Crick hydrogen bonding 21 "wavelength-ratiometric" probes 110 Weber, G. 110 web sites for protein engineering and PCR applications, some useful 206

Wilkins, E. 88 "wired" redox enzymes, substrates of 7 Wollaston prism 110 World Congress on Biosensors First 41-2 Wu, H. H. 88 xenobiotics 231 XL-1 Blue 221 X-ray crystallography 234 structure for target protein, three-dimensional 193 Zeng, H. H. 109 Zhang, X. E. 41 Zhu, K. 88 zinc 43, 112, 116 buffers 121 from carbonic anhydrase (CA), removal of 115 chelator 115 concentration, determination with ABD-N 122 determination using reagent approach 119 ion buffer recipe 122 Zn(II) 111-12 schematic of determination 120

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