Enzyme & Microbial Biosensors: Techniques and Protocols (Methods in Biotechnology 6)

Enzyme and Microbial

Biosensors

METHODS

IN

BIOTECHNOLOGY’”

John M. Walker, SERIES EDITOR 7. Affinity Biosensors:Techniques and Protocols, editedby Kim R. Rogers and Ashok Mulchandani, 1998

6. Enzymeand Microbial Biosensors:Techniques and Protocols, editedby Ashok Mulchandam and Kim R Rogers, 1998

5. Biopesticides:Use and Delivery, edltedby Franklin R. Hall and Juhus J, Menn, 1998 4. Natural ProductsIsolation, editedby Richard J. f Cannell, 1998 3. RecombinantProteinsfrom Plants:Production and Isolation of Clinically Useful Compounds, edltedby Charles Cunmngham and Andrew J. R. Porter; 1998 2. Protocolsin Bioremediation,editedby David Sheehan, 1997 1. Immobilizationof Enzymesand Cells,editedby Gordon F Blckerstafi 1997

Enzyme and Microbial Biosensors Techniques and Protocols Edited by

Ashok Mulchandani University of California, Riverside, CA

and

Kim R. Rogers US-EPA, Las Vegas, NV

Humana Press

Totowa, New Jersey

8 1998 Humana Press Inc. 999 Rtvervmw Drive, Suite 208 Totowa, New Jersey 07512 All rights reserved No part of this book may be reproduced, stored m a retneval system, or transmuted m any form or by any means, electromc, mechamcal, photocopymg, mtcrotilmmg, recording, or otherwise wlthout wntten pemusston from the Publtsher Methods in BlotechnoIogrM ISa trademark of The Humana Press Inc All authored papers, comments, opmrons, conclustons, or recommendations are those of the author(s), and do not necessarily reflect the views of the publisher Thus pubhcatron IS printed on acrd-free paper. a ANSI 239.48-1984 (Amertcan Standards Instttute) Permanence of Paper for Pnnted Library Matertals. Cover illustration. Fig 3 m Chapter 6, “Enzyme Btosensors Based on the Hydrogen Peroxtde Electrode,” by John Woodward Cover design by Patncra F Cleary For addtuonal coptes, pncmg for bulk purchases, and/or mformatlon about other Humana tttles, contact Humana at the above address or at any of the following numbers. Tel. 973-256-1699, Fax 973-256-8341, E-mad humana@humanapr corn, or vrsrt our Webstte http.//humanapress.com Photocopy Authorization Policy: Authonzatton to photocopy Items for Internal or persona1 use, or the internal or persona1 use of specrfic cltents, IS granted by Humana Press Inc , provtded that the base fee of US $8 00 per copy, plus US $00.25 per page, IS patd directly to the Copynght Clearance Center at 222 Rosewood Drove, Danvers, MA 01923 For those orgamzattons that have been granted a photocopy license from the CCC, a separate system of payment has been arranged and ISacceptable to Humana Press Inc. The fee code for users of the Transactional Reporting Servtce IS: [O-89603-410-0/98 $8.00 + $00.251 Pnnted in the Umted States of America

10 9 8 7 6 5 4 3 2 1

Preface In 1962 Clark and Lyons pioneered the concept of a biosensor. They proposed immobilizing enzymesat electrochemical detectors to form “enzyme electrodes” in order to expand the analyte range of ther base sensor. Smce then, the field of blosensors has greatly expanded. Some of the reasons for the expansion include both advances in signal transduction technologies and the incorporation of different biological sensing elements (Table 1). As a consequence, there are now a bewildering array of permutations of the biological sensing element and signal transducers that can be used to construct a biosensor. The purpose of the two volumes of Protocols and Techniques in Biosensors is to provide a basic reference tool and starting point for use by graduate students, postdoctoral and senior researchers, and technicians m academics, industry, and government research establishments, to enable rapid entry into the field of biosensors. There are a variety of approaches that researchers employ to select a combination of bioaffinity elements and signal transducers. One commonly used approach is to identtfy the compound or compounds of interest; identify the biological molecule that yields an appropriate recognitionlselectivtty and dynamic concentration range for the assay; and choose an assay format and signal transduction technology that will meet the analytical requirements for the proposed application, This volume, Enzyme and Microbial Biosensors: Techniques and Protocols, describes a variety of transduction technologies that have been interfaced to enzymes and microorganisms. The volume, although not an exhaustive treatise, provides a detailed “step-by-step” description for a variety of enzyme- and microbial-based biosensor techniques that will allow the novice or experienced investigator to expand mto new areas of research most appropriate for then analytical needs. Enzyme and Microbial Biosensors: Techniques and Protocols is divided into two sections covering enzyme and microbial biosensors. Chapter 1 provides an overview of the principles relevant to the design and operational features of enzyme-based biosensors. The subsequent chapters in the first section provide detailed protocols for enzyme biosensors based on electrochemical, thermal, and optical techniques. Included in the second section are techniques, such as oxygen gas electrode and optical techniques, in which the microorV

Preface

vi Table 1 Components a Biosensor

That May Be Used to Construct

Biological recognition elements Organisms Tissues Cells Organelles Nucleic acids Enzymes Enzyme components Receptors Antibodies

Transducers Potentiometric Amperometric Conductimetric Impedimetric Optical Calorimetric Acoustic

ganism is interfaced to the signal transducer. Each chapter also includes notes that provide mformation not usually reported m journal articles that can be particularly useful for those not familiar with construction and operation of a specific device or technique. We are fortunate to have assembled contributions from world-class authorities in this field and we sincerely thank them. In their enthusiasm for the field of biosensor research, they have produced articles that we believe will be of unusual help to the increasing number of researchers in this field. We are indebted to Prof. John Walker, the Series Editor for Methods in Molecular Biology TM, for his careful attention m reviewing the manuscripts mcluded in this volume. Last but not least, we warmly acknowledge the gracious support of our families. Ashok Mulchandani Kim R, Rogers

Contents Preface.. ............ . ..... . . ........ ... ............ ........ ...... .... ............... .. ...... .... V ....... ix Companion Volume Contents ....................................................... xi List of Contributors .......... ....... ................. ................... ... ............................ ....... . ... .... ............... 1 PART I. ENZYME BIOSENSORS .......................... Principles of Enzyme Biosensors . . . .. . .. . .. . .. .. . . . .. . ...-... .. .. . . .. . .. . .. . .. . .. . .. . . .. . . . .. . . . . . . . 3 Ashok Mulchandani Enzyme Biosensors Based on pH Electrode Canh Tran-Minh ....... .. ......................... ... ...... .. ... .... .. .,. ., .,..... . 15 Enzyme Biosensors Based on Gas Electrodes Marco Mascini and Gianna Marrazza .. . . .. . .. . . . . .. . . . .. . . . . . . . . . ,. , .. 23 Enzyme Biosensors Based on ISFETs Roland Ulber and Thomas Scheper .. . . . .. . . . . . . . . . .. . .. . . .. . .. . .. . . . . .. . . . .. . 35 Enzyme Biosensors Based on Oxygen Detection F. W. Scheller, D. Pfeiffer, F. Lisdat, C. Bauer, and N. Gajovic . . . 51 Enzyme Biosensors Based on the Hydrogen Peroxide Electrode . . ,..., . ., . . .. . . ,. . . . 67 John Woodward . . . . . . .. . .. . .. . ..,...,.............,......... Enzyme Biosensors Based on Mediator-Modified Carbon Paste Electrode Prem C. Pandey . . . . . . . , . . ., .. . . . . . .. . . . . . . .. . .. . .. . . . ,.. .. . .. . . . . . . . . ..,.. . . . . 81 8 Enzyme Biosensors Based on Electron Transfer Between Electrode and Immobilized Peroxidases Lo Gorton, Elisabeth Cs&egi, Tautgirdas Ruzgas, lrina Gazaryan, and Gy&gy Marko- Varga . .. . .. . .. .. . . . . .. , .. . , . ., . . . , . . 93 9 Enzyme Biosensors Based on Redox Polymers Latha Shankar, Michael G. Garguilo, and Adrian C. Michael .. . . . 121 10 Enzyme Biosensors Based on Metallized Carbon Electrodes Joseph Wang . .. .. . . .. .. . . .. .. . . . .. . . .. . . . . .., . . . . . . . . . . .. . . . . . . . .. . . 133 11 Enzyme Biosensors Based on Conducting Polymers Wolfgang Schuhmann . .. . .. . ....*.. . .. . .. . . . . . . .. . .. . ,.. . . .. . .. . . . .. . . .. .. . .. . . .. . 143 12 Enzyme Sensors Based on Conductimetric Measurement Norman F. Sheppard, Jr. and Anthony Guiseppi-Elie . . . .. .. . . . . .. . . .. 157

Vii

..,

VIII

Contents

13

Enzyme Biosensors Based on Thermal Transducer/Thermistor Kumaran Ramanafhan, Masoud Khayyami, and Bengt Danielsson . .. . . .. . . . . . .. . . . ,.. . . . . . .. .. . . . . . .. . . .. .. . 175 14 Enzyme Biosensors Based on Fluorometric Detection Ashufosh Sharma . . . .. . . . . .. . . . . . .. . .. . . . .. .. . .. .. . . . .. . . . ~....,..,,,.,, . . . . .. 187

PART II MICROBIALBIOSENSORS ..

15

....... ...... .. .. .. . .. . . . .. . .. ... ... 797

Microbial Biosensors Based on Oxygen Electrodes Klaus Riedei . . . .. . .. . .. . .. . . .. . . .. . . . . . . .. . .. .. . .. . . . . . .. . .. . .. . .. .. . .. . .. . . .. , . 199 16 Microbial Biosensors Based on Respiratory InhIbitIon Yoshiko Arikawa, Kazunori Ikebukuro, and lsao Karube . . .. . . .. . 225 17 Mlcroblal BiosensorsBased on Potentiometw Detection Aleksandr L. Simonian, Evgenia I. Rainina, and James R. Wild . .. . . . . . . .. . . . . . .. .. . . . . . . . . . . . . .. ,...,.-.,... .. . . . 237 18 Microbial Biosensors Based on Optical Detection Udayakumar Matrubutham and Gary S. Sayler .. . . . . . .. . . . .. . . . . . . . . .249 Index .. . . . . . . . . . . . . .. . .. .. .., . . .. . .. . . .. . . . .. .. . . . . . . .. . .. .,..-. .. . . . .. .. . . 257

Contents

for the companion Affinity Biosensors

volume:

Preface Companion Volume Contents List of Contributors PART I. AFFINITY BIOSENSORS

Principles of Affinity-Based Biosensors Kim R. Rogers lmmunobiosensors Based on Thermsitors Kumaran Ramanathan, Masoud Khayyami, and Bengt Danielsson Affinity Biosensing Based on Surface Plasmon Resonance Detection Bo Liedberg and Knut Johansen lmmunosensors Based on Piezoelectric Crystal Devrce Marco Mascini, Maria Minunni, George G. Guibault, and Robert Carter lmmunobiosensors Based on Evanescent Wave Excitation Randy M. Wadkins and Frances S. Ligler A Galactose-Specific Affinity Hollow Fiber Sensor Based on Fluorescense Resonance Energy Transfer Ralph Ballerstadt and Jerome S. Schultz Fiberoptic lmmunosensors with Continuous Analyte Response J. Rex Astles, W. Greg Miller, C. Michael Hanbury, and F. Philip Anderson lmmunobiosensors Based on Grating Couplers Ursula Bilitewski, Frank Bier, and Albrecht Brandenberg Receptor Biosensors Based on Optical Detection Kim R. Rogers and Mohyee E. Etdefrawi PART II. BIOSENSOR-RELATED TECHNIQUES

10

lmmunobiosensors Based on Ion-Selective Electrodes Hanna Radecka and Yoshio Umezawa 11 Biosensors Based on DNA Intercalation Using Light Polarization John J. Horvath

ix

X

12

Companion

Volume Contents

ISFET Affinity Sensor Geert A. J. Besselink and P/et Bergveld 13 Liposome-Based lmmunomigration Assays Matthew A. Roberts and Richard A. Durst 14 Isolated Receptor Biosensors Based on &layer Lipid Membranes Masao Sugawara, Ayumi Hirano, and Yoshio Umezawa 15 Eukaryotic Cell Biosensor: The Cytosensor Microphysiometer Amira T. Eldefarwi, Cheng J. Cao, Vania I. Cortes, Robert J. Mioduszewski, Darrel E. Menking, and James J. Valdes Index

Contributors 8Research Center for Advanced Science and Technology, University of Tokyo, Japan C. BAUER Ivzstztuteof Biochemzstzyand Molecular Physiology, University of Pottsdam, Berlin, Germany ELISABETH CSOREGI Department of Analytical Chemistry, Chemical Center, Lund University, Lund, Sweden BENGT DANIELSSON Department of Pure and Applied Biochemistry, University of Lund, Sweden N. GAJOVIC Institute of Bzochemzstryand Molecular Physiology, University of Potsdam, Berlin, Germany MICHAEL G. GARGUILO Department of Chemistry Universzty of Pittsburgh, PA IRINA GAZARAYAN Department of Chemical Enzymology, Moscow State University, Moscow, Russia Lo GORTON Department of Analytical Chemistry, Chemical Center, Lund University, Lund, Sweden ANTHONY GUISEPPI-ELIE 9 Department of Biomedical Engineering, The Johns Hopkins University, Baltimore, MD KAZUNORI IKEBUKURO Research Center for Advanced Science and Technology, University of Tokyo, Japan ISAO KARUBE 9Research Center for Advanced Science and Technology, University of Tokyo, Japan MASOUD KUAYYAMI Department of Pure and Applied Biochemistry, University of Lund, Sweden F. LISDAT Institute of Biochemistry and Molecular Physiology, University of Potsdam, Berlin, Germany I. LUNDSTROM Laboratory of Applied Physics, Linkoping University, Linkoping, Sweden GYORGY MARKO-VARGA Department of Bioanalytical Chemistry, Astra Draco, Lund, Sweden GIANNA MARRAZZA 9Dipartimento di Sanita Pubblica Epidemiologza e Chimica Analitica Ambzentala, Universita di Firenze, Italy MARCO MASCINI Dipartimento di SanztaPubblica Epidemiologia e Chimica Analitica Ambientala, Universzta di Firenze, Italy YOSHIKO ARIKAWA l

l

l

l

l

l

l

l

l

l

l

l

l

Xi

Contributors

xii

Center for Environmental Biotechnology, The University of Tennessee, Knoxville, TN ADRIAN C. MICHAEL Department of Chemistry, Umversity of Pittsburgh, PA ASHOK MULCHANDANI Marlan and Rosemay Bourns College of Engineering, University of Call~ornia, RiversIde, CA PREM C. PANDEY Department of Chemistry, Banaras Hindu Unzversity, Varanasl, India D. PFEIFFER BST BioSensor Technologie, GmbH Buchholzer, Berlin, Germany HANNA RADECKA Department of Chemistry, School of Science, The University of Tokyo, Japan EVGENIAI. RAININA Biochemistry and Biophysics Department, Texas A & M University, College Station, TX KUMARAN RAMANATHAN Department of Pure and Applied Biochemistry, University of Lund, Sweden KLAUS RIEDEL 9 Dr. Bruno Lange GmbH, Dusseldorj Germany TAUTGIRDASRUZGAS9Institute of Biochemistry, Vilnius, Lithuanra GARRY S. SAYLER Centerfor Envaronmental Biotechnology, The University of Tennessee, Knoxville, TN F. SCHELLER Instrtute of Biochemistry and Molecular Physiology Unzversity of Potsdam, Berkn, Germany THOMAS SCHEPER Institute of Technzcal Chemistry, University of Hannover, Germany WOLFGANGSCHUHMANN Lehrstuhl fur Analytische Chemie, RuhrUniversitat Bochum, Freuing- Weihenstephan, Germany LATHA SHANKAR9Department of Chemistry, University of Pittsburgh, PA ASHUTOSHSHARMA University of North London, UK NORMAN F. SHEPPARD,JR. Department of Biomedical Engmeering, The Johns Hopkins University, Baltimore, MD ALEXANDR L. SIMONIAN Biochemistry and Biophysics Department, Texas A & M Utuversity College Station, TX CANH TRAN-MINH Centre SPIN/Biotechnology, Ecole Natlonale Superleur des Mutes, St. Etienne, France ROLAND ULBER Institute of Technical Chemistry, University of Hannover, Germany JOSPEHWANG 9Department of Chemistry and Biochemistry, New Mexico State University, Las Cruces, NM JAMES R. WILD 9 Biochemistry and Biophysics Department, Texas A & M University College Station, TX JOHN R. WOODWARD GLI International, Inc., Milwaukee, WI UDAYAKUMAR MATRUBUTHAM l

l

l

l

l

l

l

l

l

l

l

l

l

l

l

l

l

l

I ENZYMEBIOSENSORS

1 Principles of Enzyme Biosensors Ashok Mulchandani 1.

Introduction

An enzyme biosensor is an analytical device that combines an enzyme with a transducerto produce a signal proportional to target analyte concentration.This signal can result from a changein proton concentranon,releaseor uptake of gases,such asammonra or oxygen, light emission,absorpnonor reflectance,heat emissron,and so forth, brought about by the reaction catalyzedby the enzyme. The transducer converts this signal mto a measurableresponse,such as current, potential, temperature change,or absorption of light through electrochemical,thermal, or optical means. This signal can be further amplified, processed,or storedfor later analysis. Because of their specificity and catalytic (amplification) properties, enzymes have found widespread use as sensing elements in biosensors. Since the development of the first enzyme-based sensor by Clark and Lyons (I), who mnnobihzed glucose oxidase on an oxygen-sensing electrode to measure glucose, there has been an impressive proliferation of applications involvmg a wide variety of substrates. A variety of enzymes belonging to classes of oxido-reductases, hydrolases, and lyases have been integrated with different transducers to construct biosensors for applications in health care, veterinary medicine, the food industry, environmental monitoring, and defense (2). This chapter provides an overview of the various transducers and enzymes, including the techniques applied for immobilization of these enzymes on transducers, for constructton of enzyme biosensors for wide ranging applications.

2. Transducers Table 1 gives a lrst of different transducers that have been used m enzyme-biosensor construction. The operating principles of commonly used transducers as applied to biosensors arc described below. From

Methods m Blofechnology, Vol 6 Enzyme and MIcrobra/ Biosensors Techmques Edited by A. Mulchandam and K R Rogers 0 Humana Press Inc , Totowa,

3

and Protocols NJ

Mulchanciani

4 Table 1 Transducer

Technologies

Transducertechnology/examples Electrochemical Amperometric Potentiometric Conductimetric Optical Calorimetric Luminescence Fluorescence Calorimetric Thermistor

output Apphed current Voltage Impedance Color Light intensity Light intensity Temperature

2.7. Electrochemical Transducers Three types of electrochemical transducers are used for construction of enzyme electrodes: potentiometric, amperometric, and conductometric. 2.1.1. Potentiometric Conventional potentiometric enzyme biosensors consist of an ion-selective electrode (ISE)-pH (Chapter 2), ammonium, fluoride, and so forth-or a gas-sensing electrodeO and NH3 (Chapter 3)-costed with an tmmobihzed enzyme layer. The enzymatic reaction with the analyte generates a change in potential resulting from ion accumulation or depletion. Potentiometric transducers measure the difference in potential that is generated across an ionselective membrane separating two solutions at virtually zero current flow. Examples of applications of potentiometric enzyme probes include the detection of urea, glucose, and creatinine (3,4). Table 2 gives a list of enzymes that have been used in construction of potentiometric biosensors. Because of a logarithmic relationship between the potential generated and analyte concentration, a wide detection range is possible. The requirement of a very stable reference electrode may be a limitation of these transducers (5). An improvement n-r the ISE-based biosensor technology is the advent of solid-state or silicon-based ion-selective field effect transistors (FETs). The enzyme-based field effect transistor (ENFET), which consists of an enzyme layer coated on an ton-selective FET (ISFET), offers several advantages over the ion selective electrode. It has a small compact size and requires only a finite quantity of enzyme. The immobilized enzyme layer thickness can be more easily controlled and no retaining membranes are required. An example of ENFET design can be the combination of a pH-sensitive ISFET with an

Principles of Enzyme Biosensors Table 2 Enzymes

Used with Potentiometric

Substance detected

5 BiosensorsB

Enzyme

Urea

Urease

Glucose L-Ammo acids r.-Tyrosine L-Glutamine L-Glutamic acid L-Asparagme n-amino acids Penicillin Amygdalin Nitrate

Glucose oxidase L-Amino acid oxidase L-Tyrosine decarboxylase Glutammase Glutamate dehydrogenase Asparagmase o-amino acid oxidase Penicillinase P-Glucosidase Nitrate reductase/ nitrite reductase Nitrite reductase

Nitrite

Type of ion-selective sensor Cation, pH, gas (NW, gas (Wd PH, ICation, NH4+, Igas VW Cation Cation Cation Cation PH CNNH4+ Gas (NW

aInforrnation adapted from ref. 3

immobilized hydrolase enzyme layer. The enzymatic conversion of the analyte results in a pH shift that is detected by the pH-sensitive ISFET (6) (Chapter 4). In another type of ISFET, the pF-FET, 4-fluoroaniline and oxrdases (e.g., glucose oxidase) are combined to form a new type of biosensor. The H202 produced by oxidases during the enzymatic conversion reacts with 4-fluoroaniline to generate F ions, which are detected by the pF-FET (7,s). The use of ISFETs is attractive commercially as miniaturization can be achieved. However, a disadvantage of such a system is that it requires a costly pH/mV meter since the existing instrumentation cannot be used (9). Another potentiometric enzyme-biosensor concept reported (10) is based on measurement of capacitance changes. The device, an Electrolyte Insulator Semiconductor (EIS) chip, consisted of a three-layer structure: a pH-sensitive (Si/Si02/Si3N,/Ta205) and pF-sensitive (Si/Si02/Si3N4/LaF3) layer structure that selectively detected H’ and F ions, respectively, followed by a coating of penicillinase, urease,or glucose ox&se. The fluoride-ion-based biosensorsoffered the advantage of independence with regard to buffer capacity of the solution. 2.1.2. Amperometric Amperometric enzyme biosensors form the majority of commercial biosensor devices available on the market today. In contrast to potentiometric sensors, where the potential generated across a membrane IS used to convey mformation on the analyte concentration, amperometric brosensors operate at a fixed

Mulchandani

6 Table 3 Selected

List of Enzymes

Enzyme

Useful with O2 Electrodesa

EC number

Source

Glycollate oxidase

1131

Spinach Rat liver

Lactate oxidase Glucose oxidase Xanthme oxidase

11.32 113.4 123.2

Pedlococcus sp. Aspergillus niger

Oxalate oxidase

123.4

L-Amino acid oxidase

1432

Nitro-ethane oxidase

1 7.3.1

Urate oxidase Sulfite oxidase Malate oxidase Cholesterol oxidase Ascorbate oxidase

1 7.3.3 1.8.3.1 1.1.3.2 1.1.3.6 1.10.3.3

Bovine milk

Diamond rattle snake

Hog liver Beef liver Nocardia Squash

Typical substrates Glycollate L-, o-Lactate L-Lactate S-D-Glucose Hypoxanthme Xanthme Benzaldehyde Oxalate L-Methionine L-Phenylalanme Nttroethane Aliphatic Nttrocompounds Urate Sulfite Malate Cholesterol L-Ascorbate

aInforrnation adapted from ref. II with respect to a reference electrode and the current generated by the oxidation or reduction of species at the surface of the working electrode is measured. Amperometric biosensors are based on redox enzymes; thus, their appeal is caused by the availability of a large number of oxido-reductase enzymes that can act on fatty acids, sugars,amino acids, aldehydes, and phenols (Table 3). These enzymes use molecular oxygen as an electron acceptor and produce hydrogen peroxide m the reaction with then substrates.Either the oxygen consumption (Chapter 5) or the hydrogen peroxide (Chapter 6) production can be followed as a measure of the substrate (analyte) concentration. Oxygenor hydrogen-peroxide based biosensors, however, are strongly influenced by fluctuations in dissolved oxygen concentration, which results from changes m pH, temperature, ionic strength, or partial pressure.Additionally, biosensors based on hydrogen peroxide measurement may suffer from interferences resulting from nonspecific electrochemical oxidation of compounds, such as ascorbate, uric acid, glutathione, and cysteine, at the 0.6-0.7 V (vs Ag/AgCI reference) potential required to detect H202, This has led to the development of chemically modified electrodes in which the electron acceptor dioxygen is replaced potential

Principles of Enzyme Biosensors

7

by low-mol-wt mediators, such as tetrathiafulvalene (TTF), tetracyanoquinodimethane (TCNQ), hexacyanoferrate (III), methylene blue, ferrocene (bls ($-cyclopentadienyl) iron), quinones, and N-methylphenazin-5-ium (NMP+) (Chapter 7) (12-14). When compared to the enzyme substrate reaction, the enzyme mediator reaction is generally not specific. Although effective in lowering the operating potential, mediated electrodes still suffer from some ascorbic acid and uric acid interferences. Additionally, since oxygen is a better electron acceptor than these mediators, oxygen has to be eliminated from buffers and samples. Mediators can be incorporated mto the electrode by: 1. Adsorption using a mediator-organic solvent solution that is allowed to evaporate (15); 2. Entrapment in or behind a polymer film at the electrode surface (16,17); 3. Covalent binding to monomers or polymers that can be deposited on the electrode surface (Chapters 9 and 11) (18-21); and 4. Mixing into a paste of graphite and mineral 011(Chapter 7) (14,21,22).

Hydrogen peroxide formed during the oxidase-catalyzed reactions can also be measured using peroxidase-modified electrodes (Chapters 8 and 9). In these electrodes a reduction current, resulting from either the direct or mediated electron transfer, 1s measured at low applied potential, thereby alleviating the interference problems encountered during electrochemical oxidation of H202, Measurement of H202, at low oxidation potential has been demonstrated using carbon with dispersed rhodium, ruthenium, or iridium particles (Chapter lo), which alleviates interference from ascorbic acid and uric acid, two common interferents present in biological samples. 2.7.3. Conductimetric Many enzyme-catalyzed reactions involve a change in Ionic species (Chapter 12). Associated with this change is a net change in the conductivity of the reaction solution. Since the measurement of solution conductance is nonspecific, the widespread analytical use of eonductimetric transducer has therefore been restricted. However, where the specificity does not play a significant role, conductance measurements are capable of extreme sensitivity. 2.2. Thermal Thermal enzyme sensors are based on the principle that the heat evolved in an enzymatic reaction can be utilized to calorimetrically determine the amount of substrate reacted (23,24). Thus, thermometric indicators or transducers require only a single reaction step producing sufficient or measurable heat. Molar enthalpies of enzyme-catalyzed reactions range from 4 to 100 kJ/mol. In thermal enzyme sensors, the enzyme 1sattached directly to the temperature

8

Mulchandani

transducer, a thermistor, either by crosslinking or by entrapping the enzyme m a membrane enclosmg the thermistor (25). Alternatively, the enzyme is placed in a temperature-controlled column and the heat of reaction is measured by recording the increase in temperature between the inlet and outlet as the sample flows through the column (Chapter 13). In such systems,temperature changes as low as 1O-“OCcan be measured. The mam disadvantage of such systems are nonspecific thermal effects and a baseline drift resulting from heating of the unit. The problem of nonspecific thermal effects is overcome by subtracting the temperature change caused by the flow of fhud through a column containing the enzyme immobilization matrix without the enzyme from the temperature change observed for the channel with the enzyme column. The problem of drift is alleviated by installing the complete unit m a thermostatically controlled alummum block. These additional requirements increase the overall cost of the system. 2.3. Optical In biosensors based on optical methods, the change in optical properties, such as UV/vis absorption, bio- and chemiluminescence, reflectance and fluorescence brought by the interaction of the biocatalyst with the target analyte, is monitored optically. The oxidation and reduction of the NAD(P)H during enzymatic reactions catalyzed by dehydrogenase can be monitored by measuring the NAD(P)H fluorescence (excitation at 360 nm and measurement at 450 nm) and the changes in the fluorescence intensity then related to the substrate concentration. In another approach (Chapter 14), the cofactor NAD(P)H involved in the enzymatic reaction was monitored by observing the interaction of a fluorescent dye with the cofactor. The pH-dependent fluorescence property of fluorescent indicator dyes, such as fluorescem isothiocyanate (FITC), can be used to measure the variation in pH during the enzymatic reaction to determine the analyte concentration (26,27). Gautier et al. (28) have investigated the use of luminescent enzyme systems lmked to optical transducers for the determination of sorbitol, ethanol, and oxaloacetate at the nanomolar level. A bacterial luminescence fiber-optic sensor that detected NADH was combined with various NAD(P)H-dependent enzymes--sorbitol dehydrogenase, alcohol dehydrogenase, and malate dehydrogenase-for on-line determination of sorbitol, alcohol, and malate, respectively. The bacterial luminescence enzyme and the desired dehydrogenase were coimmobilized on preactlvated polyamide membranes. This membrane was attached to the end of a fiber-optic bundle and placed in a flow through cell. The NADH formed by the reaction of the analyte with NAD in presence of the dehydrogenase enzyme was detected using the bacterial luminescence fiber-

Principles of Enzyme Biosensors

9

optic sensor. Chemiluminescence reaction of luminol With HzOz formed durmg oxidase catalyzed reactions, in presence of excess horseradish peroxidase has also been used to monitor various analytes (29). Measurement of absorbance change resulting from a chromophoric product formed in the reaction catalyzed by an enzyme rmmobilized at the common end of a bifurcated fiber-optic bundle can be used to determine the analyte concentration (30). Fiber-optic-based biosensors offer advantages of compactness, flexibility, resistance to electrical noise, and a small probe size. Calibration of optical sensors based on fluorescent or chrompohoric dyes is highly stable, especially rf measurement is made at two different wavelengths. However, these sensors suffer from the instability of the optically active dyes, 3. Enzyme Immobilization When using enzymes as biological elements in biosensors, two Important considerations have to be taken into account: operational stability and long-term use. Since both these factors are to some degree a function of the immobilization strategy used, choice of immobilization technique IS critical. Enzymes are immobilized on transducer or support matrices by physical and chemical methods (Table 4). 3.1. Physical Methods Physical methods of enzyme rmmobtlizatron, such as entrapment (31) and adsorption (32), have the benefit of applicability to many enzymes and may provide relatively small perturbation of the enzyme native structure and function. Enzymes can be entrapped in polyacrylamide, calcium alginate, agar, agarose, or chitosan polymer network or gel. Chief disadvantages of this technique are irregular pore size of the gel, lack of mechanical strength. and diffusional limitations encountered by substrates and products. In many biosensor applications, enzyme immobilized in polymer gel is placed between two precast membranes that provide the necessarybarrier against fouling and interference (33). This further increases the diffusional resistance, and sensors with such an arrangement have low sensitivity and poor lower detection limits. More recently, however, two anion exchange polymers have been used for enzyme immobilization. These polymers, Nafion and Eastman AQ, combine enzyme retention with separation of anionic-interfering species,such as ascorbic and uric acids, Since these latter polymers can be used to cast membranes directly on the surface, a layer thinner than with conventional precast membranes can be obtained (12). Direct physical adsorption of enzymeson a surface is an alternative techmque that is easily implemented. However, immobiliza-

tion using adsorptionalone generally leads to poor long-term stability of the sensor because of enzyme leakage from the surface. An additional barrier, such

10

Mulchandani

Table 4 Methods for Enzyme Method Entrapment and encapsulatron

Covalent binding

Crosslmkmg

Adsorptton

Immobilization Advantages

Disadvantages

Gentle, no dtrect chemical modtticatton; specrficrty and analyte interaction retamed Low dtffusronal resistance; strong bmdmg force between enzyme and matrix; resistant to adverse condmons of pH, tome strength Used m conJunctron with entrapment to reduce loss of enzyme

Hugh drffusion barrter; only good for small analyses, contmuous loss of enzyme

Gentle treatment of enzyme, no modrticatton of enzyme; matnx can be regenerated

Matrix not regenerable, may mvolve harsh/toxtc chenucals

Covalent links formed between protein molecules rather than matrix and protein, may mvolve harsh/toxic chemmals Very weak bonds, susceptrble to changes m pH, temperature, romc strength

as a membrane or film, is therefore required to prevent the loss of the adsorbed enzyme when the biosensor is placed in an aqueous environment. 3.2. Chemical Methods Chemical methods of enzyme immobilization include covalent binding and crosslinking using multifunctional reagents, such as glutaraldehyde and cyanuric chloride (11,34). The enzyme may be covalently attached or crosslinked to the electrode surface itself or to commercially available hydrophtltc membranes, such as Immunodyne (Pall BioSupport, Port Washington, NY) and Tmmobilon (Mlllipore, Bedford, MA), which can then be placed on the sensing area of the transducer. The chief advantage of the latter approach is that enzyme membranes can be replaced frequently and easily, thus extending the life and versatility of the biosensor (35-3 7). Another technique of physically retaining enzymes on the biosensor surface is through the use of thin electrochemically prepared polymer films (Chapter 12). These films are obtained by the electrochemical polymerization of aromatic organic compounds, such as pyrrole, thiophene, phenylenediamine, phenol, and so forth (38,39). The immobilization of enzyme in such membranes is achieved by either depositing a polymer film on the electrode surface on which the enzyme has been previously immobilized by adsorption or

Principles of Enzyme Biosensors

11

crosslinking (39), or entrapping the enzyme as a counterton from a solution of the enzyme and monomer (38). The thickness of the electrochemically deposited polymer film is usually in the nanometer range and therefore has obvious advantages in terms of increased sensitivity, In addition to immobilizmg the enzyme, these films also act aspermselective membranes to improve selectivity and provide a barrier against electrode fouling. For example, electropolymerized poly (1,2-diaminobenzene) prevents interferences from cysteine, ascorbate, and uric acid and fouling caused by protein in hydrogen-peroxidebased amperometric electrodes (33). The electrochemical polymer film can also be used to incorporate the redox mediator (16,18,40). 4. Potential Applications The main application areas of enzyme biosensors are m human and animal health care, food and fermentation industries, environmental monitoring, agriculture and defense. The use of enzyme biosensors in medical and clinical applications is expected to increase. A rapid test for such substances as amylase, glucose, lactate, paracetamol (acetaminophen), salicylate, creatine kinase, aspartate aminotransterase, and urea will be invaluable in emergencies. Measurements of urea, glucose, lactate, and creatine would be useful for continuous or high frequency use m intensive care units. Although many of these tests are presently performed in central clinical laboratories, there is now a demand for low-cost methods for monitoring these compounds in physician’s offices, outpatient departments, and in homes for diagnosis and monitoring of a patient’s condition as well as for therapy. This will require that these methods be easy to use and require minimal sample preparation. This is the area in which enzyme biosensors are expected to play a major role. The second area in which enzyme-biosensors are expected to play a major role is in environmental monitoring. There is an increasing demand for measuring several classesof substances,including toxic chemicals, such as polychlorinated biphenyls, polyaromatic hydrocarbons, phenols, dioxins, organic peroxides, and so forth, pesticides, and heavy metals in water, soil, and air. The portability and real-time output will be a key to the success.Enzyme biosensors will provide rapid, easy to use, and cost-effective on-site testing. Although the prospects are good, the development of these types of biosensors must be endorsed by regulatory bodies before there will be widespread application, These endorsement will require numerous successful field demonstrations under a wide variety of conditions encountered in the field. There are several potential applications of enzyme biosensors in the food and fermentation industry. Examples of such applications include analysis of composition (amino acids, sugars, antibiotics) and freshness measured through specific freshness indicators (such as ATP degradation products or trimethy-

12

Mulchandani

lamine accumulation as an indicator of fish freshness) of food, as well as monitoring of substrate, nutrients, and product levels in fermentation processes for potential control apphcations. The development of enzyme electrodes for in situ monitoring in fermenters, however, is hampered by the need of steam sterilization of the fermentors. 5. Future Directions The development of a highly satisfactory enzyme brosensor is hampered because enzymes are not always stable. The instability problem can be circumvented by using enzymes stable at high temperatures, naturally available from thermophilic microorganisms. Furthermore, as the structure of enzymesbecome better defined, it is possible to “tailor” enzymesthat can function under stressed environments for a long period of time. Future research m these areas IS necessary. Similarly, development of reagentless biosensors involving enzyme reactions requiring dlfmstble cofactors, such as ATP and NAD(P)H, IS presently limited because of lack of methods for efficient cofactor regeneration. An efficient regeneration at a fast rate is feasible using designer enzymes in which the cofactor is anchored and can swing between the oxrdatron and reduction sites. References 1. Clark, L. C. and Lyons, C. (1962) Electrode systerm for continuous monitoring of cardiovascular surgery Ann. NY Acad Sci 102,2945. 2. Guilbault, G G (1984) Analytical Uses of Immobilized Enzymes Marcel Dekker, New York. 3. Kaun, S. S. and Guilbault, G. G. (1987) Ion-selective electrodes and biosensors based on ISEs, m Biosensors. Fundamentals andApplicattons (Turner, A. P. F., Karube, I , and Wilson, G. S., eds.), Oxford University Press, New York, pp. 135-152. 4. Kauffmann, J.-M. and Guilbault, G. G. (1991) Potentiometric enzyme electrodes, in Biosensor Principles and Apphcatrons (Coulet, L. J and Blum, P R , eds.), Marcel Dekker, New York, pp. 63-82. 5. Luong, J. H. T , Groom, C. A., and Male, K. B. (1991) The potential role of biosensors in the food and drink mdustrtes. Biosens. Bioelectron 6,547-554. 6. Brand, U., Brandes, L., Koch, V , Kullik, T., Remhardt, B., Ruther, F , Scheper, T., Schugerl, K., Wang, S., Wu, X., Ferrett, R., Prasad, S., and Wtlhelm, D (199 1) Monitoring and control of biotechnological production processes by BioFET-FIA-sensors Appl Microbrol Biotechnol 36, 167-l 72. 7. Ho, M H. (1988) Potenttomctric biosensor based on immobihzed enzyme membrane and fluoride detector Sens Act B 15,44545 1 8. Hmtsche, R., Dransfeld, I., Scheller, F., Pham, M. T., Hoffmann, W., Hueller, J., and Moritz, W (1990) Integrated differential enzyme sensors usmg hydrogen and fluoride ion sensitive multigate FETs. Biosens Bioelect 5,321-334 9. Guilbault, G G. and Luong, J. H T. (1989) Biosensors: current status and future possibilities. Selective Electrode Rev. 11, 3-16.

Principles of Enzyme Biosensors

13

10. Beyer, M., Menzel, C., Quact, R., Scheper, T., Schiigerl, K., Treichel, W., Voit, H., Ullrich, M., and Ferret& R. (1994) Development and application of a new enzyme sensor type based on the EIS-capacitance structure for bioprocess control. Bzosensor Blosensor Bioelect 9, 17-2 1. 11. Carr, P. W. and Bowers, L. D. (1980) Immobilized enzymes, in AnaZytzcal Chemistry’ Fundamentals and Appkcations. Wiley, New York. 12. Gorton, L., Csoregl, E., Dominguez, E., Emneus, J., Jonsson-Pettersson, G , Marko-Varga, G., and Persson, B. (1991) Selective detection in flow analysis based on the combination of immobilized enzymes and chemically modified electrodes. Anal Chum Acta 250,203-248. 13. Cardow, M. F. and Turner, A. P. F. (1989) The realization of electron transfer from biological molecules to electrodes, in Baosensors: Fundamentals and Applecations (Turner, A. P. F., Karube, I., and Wilson, G., eds.), Oxford University Press, New York, pp 257-275. 14. Almelda, N. F. and Mulchandani, A. (1993) A mediated enzyme eletrode using tetrathiafulvalene and L-glutamate oxidase for the determination of L-glutamlc acid. Anal. Chum Acta 282,353-361. 15. Cass, A. E. G., Davis, G , Francis, G. D , Hill, H. A. O., Aston, W. J , Higgins, I. J , Plotkm, E. V., Scott, L D. L., and Turner, A. P. F. (1984) Ferrocene-mediated enzyme electrode for amperometric determination of glucose. Anal Chem. 56,667-67 1. 16. Kajiya, Y., Sugai, H., Iwakura, C., and Yoneyama, H. (1991) Glucose sensltlvity of polypyrole films containing immobilized glucose oxidase and hydroquinonesulfonate ions Anal. Chem 63,49-54. 17. Schuhmann, W. (1993) Non-leaking amperometric biosensors based on high-molecular ferrocene derivatives. Blosens. Bioelect 8, 191-196. 18. Foulds, N. C. and Lowe, C. R. (1988) Immobihzation of glucose oxldase m ferrocene modified pyrrole polymers. Anal Chem. 60,2473-2476. 19. Heller, A. (1992) Electrical connection of enzyme redox centers to electrodes. J. Phys. Chem. 96,3579-3587.

20. Mulchandani, A., Wang, C.-L., and Weetall, H. H. (1955) Amperometrlc detection of peroxides with poly(anilinomethylferrocene) modified enzyme electrodes, Anal. Chem 67,94-100.

21. Wang, J., Wu, L. H., Lu, Z., Li, R., and Sanchez, J. (1990) Mixed ferroceneglucose oxidase-carbon-paste electrode for amperometric determination of glucose. Anal. Chum. Acta 228,251-257. 22. Pandey, P. C., Glazier, S., and Weetall, H. H. (1993) An amperometric flowinjection analysis biosensor for glucose based on graphite paste modified with tetracyanoquinodimethane. Anal. Blochem. 214,233-237 23. Mattiasson, B., Danielsson, B., Mandenius, C. F., and Winquist, F. (1981) Enzyme thermistors for process control. Ann. NY Acad. Scz. 369,295-305. 24. Mandenius, C. F., Danielsson, B., and Mattlasson, B. (1984) Evaluation of a dialysis probe for contmuous sampling in fermentors and in complex media. Anal. Chcm. Acta 163, 135-141.

14

Mulchandani

25. Guilbault, G. G., Danielsson, B., Mandenius, C. F., and Mosbach, K. (1983) Enzyme electrode and thermistor probes for the determination of alcohols with alcohol oxidase. Anal. Chem. 54, 1582-1585. 26. Scheper, T., Brandes, W., Grau, C , Hundeck, H. G., Reinhardt, B., Ruther, F., Plota, F., Schelp, C., Schugerl, K., Schneider, K. H., Rehr, B., and Sahm, H (199 1) Applications of biosensor systems for bioprocess monitoring. Anal Chim Acta 249,25-34. 27. Rogers, K. R., Cao, C. J., Valdes, J. J., Eldefiawi,

A. T., and Aldefrawi, M. E. (1991) Acetylcholmesterase fiber-optic biosensor for detection of anticholmesterases. Fund Appl Tox~ol. 16,810-820. 28. Gautier, S. M., Blum, L. J., and Coulet, P. R. (1990) Fibre-optic biosensor based on luminescence and immobilized enzymes. Blolumm. Chemllumtn. 5,57-63. 29 Cattaneo, M. V. and Luong, J. H. T. (1993) Monitoring glutamme in animal cell cultures using a chemdummescence fiber optic biosensor. Bzotechnol. Bzoeng 41,659-665. 30. Arnold, M A (1991) Fluorophore- and chromophore-based biosensors, m Bzo-

31 32. 33. 34. 35. 36.

sensor Principles and Applications (Blum, L. J and Coulet, P R , eds.), Marcel Dekker, New York, pp. 195-212. Kierstan, M. P. J and Coughlan, M. P. (1985) Immobilization of cells and enzymes by gel entrapment, m Immobilized Cells and Enzyme (Woodward, J., ed.), IRL, Oxford, UK, pp. 39-48. Messing, R. A. (1976) Adsorption and inorganic bridge formations. Methods Enzymol. 44,148-168. Stoecker, P. W. and Yacynych, A. M (1990) Chemically modified electrodes as biosensors. Selective Electrode Rev. 12, 137-160. Weetall, H H. (1993) Preparation of immobilized proteins covalently coupled through silane coupling agents to inorgamc supports. Appl. Blochem. Biotechnol. 41,157-188. Assolant-Vinet, C H. and Coulet, P R. (1986) New immobilized enzyme membranes for tailor-made biosensors. Anal. Lett. 19, 875-885. Mulchandam, A., Male, K. B., and Luong, J. H. T (1989) Development and application of a biosensor for hypoxanthine m fish extract. Anal. Chum Acta 221,

2 15-222 37 Villarata, R L , Palleschi, G , Lubarano, G. J., Suleiman, A A., and Guilbault, G G

(1991) Amperometric aspartate electrode. Anal Chim. Acta 245,63-69. 38. Foulds, N. C. and Lowe, C. R. (1986) Enzyme entrapment in electrically conductmg polymer. J. Chem. Sot Faraday Trans 182, 1259-1264. 39. Bartlett, P. N., Tebbutt, P., and Tyrrell, C H. (1992) Electrochemical immobihzation of enzymes. 3. Immobihzation of glucose oxidase m thin films of electrochemically polymerized phenols. Anal Chem 64, 138-142 40. Reynolds, E. R., Geise, R. J., and Yacynych, A. M. (1992) Electropolymerized films for the construction of ultramicrobiosensors and electron-mediated amperometric biosensors. ACS Symp. Ser. No. 497 American Chemical Society, Washmgton, DC, pp. 186-200

Enzyme Biosensors

Based on pH Electrode

Can h Tran-Min h 1. Introduction Electrochemical transducers are often used to monitor the enzymatic reaction by potentiometric or amperometric measurements of the product or cosubstrate concentration. When the enzymatic reaction results in a change in pH, a pH electrode can be used as a potentiometric transducer. A schematic representation of an enzyme biosensor based on pH electrode is given in Fig. 1. The sensitive surface of the pH electrode is in contact with an enzymatic layer and is immersed in a solution containing the substrate. The substrate migrates toward the interior of the layer and 1s converted into reaction products, with proton H+ production or consumption, when it reacts with the immobilized enzyme. The different steps during the operation of an enzyme sensor based on pH electrode are: 1. Transport of the substratefrom the bulk of the solution toward the enzymatic layer, 2. Diffusion of the substratewithin this layer,accompanredby the enzymatictransformation of the substrateinto reactionproductwith H+ production(or consumption), 3. Migration of H” toward the pH electrode, 4. Detection of Ht by the pH electrode. Because difmston is a slow process, the enzymatic membrane must be as thin as possible to achieve rapid equilibrium of concentrations. The solution must also be well stirred to ensure a constant supply of substrate. The performances of enzyme pH electrodes will largely depend on the mnnobilization of the enzyme onto the electrode to ensure maximal contact and response. Enzymes can be immobrlized by physical entrapment (I) or by From

Methods m Biotechnology, Vol 6 Enzyme and Wcrobral Blosensors Techmques and Protocols Edlted by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa, NJ

15

16

Tran-Minh

->

P+H

Fig. 1. Schematic representation of the drffusron of the substrate S and the products P and H+ in the enzymatrc layer on a pH electrode. crosslmkmg (2), which ensures a greater long-term enzymatic stability. Enzyme pH electrodes can be constructed by dipping the electrode in a solution containing the enzyme with a crosslinking agent (immersion method) or immobrhzing the enzyme directly (3) on the ttp of the pH electrode (direct method). Prefunctionalized or reinforced membranes with a fabric base are also used to ensure the mechanical properties of enzymatic membranes (4) and to facilitate their manipulation. Membranes obtained in this way can be clipped directly onto the transducer ($6). Brosensors with a very rapid response time are obtained with aerosol vaporization, which gives active films that are thin and homogeneous (7).

2. Materials 2.1. Immersion

Method

1 pH glass electrode (mrcroelectrode can also be used). 2. 0 0 1 M sodium phosphate-buffer solution, pH 7 0 3. Enzyme solutron: 1 mg urease (EC.3.5.1.5) from Jack beans (= 800 U/mg protein) m 1 mL 0.0 1 M sodium phosphate buffer-solutron. 4. 30% bovine serum albunnn (BSA) m water 5. 25% Glutaraldehyde aqueous solutron as crosslinkmg agent

2.2. Direct Binding

Method

1. pH glass electrode with flat trp of 2 5 cm* in area. 2. 0.01 A4 sodium phosphate-buffer solution, pH 7.0

17

p H Electrode

3. Enzyme mixture: 0.2 mg butyrylcholinesterase (EC.3.1.1.8) from horse serum (= 500 U/mg protein) m 0.1 rnL phosphate-buffer solution mixed with 0 1 mL BSA (30% aqueous solution). 4 Glutaraldehyde solution: 2.5% in water, as crosslmking agent.

2.3. Reinforced

Membranes

1. 2. 3. 4. 5.

pH glass electrode equipped with a clip device. A nylon net (5 x 4 cm) of 140+m mesh size Two Teflon sheets (5 x 4 cm), 5 cm in thickness. 0 01 Msodium phosphate-buffer solution, pH 7.0. Enzyme mixture: 2 mg butyrylcholinesterase (EC.3.1.1.8) from horse serum (= 500 U/mg protein) in 1 mL phosphate-buffer solution mixed with 1 mL BSA (30% aqueous solution). 6 Glutaraldehyde solution: 2.5% m water.

2.4. Aerosol

Vaporization

1. pH glass electrode (type XG from Tacussel, Lyons, France). 2. 0.01 M sodium phosphate-buffer solution, pH 7.0. 3. Enzyme solution: 4 mg pemcillmase (EC.3.5.2.6) from Bucrllus cereus (= 400 U/mg protein) in 1 mL of phosphate-buffer solution 0.01 M, pH 7.0. 4. Glutaraldehyde solution: 2.5% in water. 5. Air-brush (type Atrium from Lefianc Bourgeois, France) using nitrogen under a pressure of 1.5 bar

2.5. Use of Enzyme pH Biosensors 2.5.7. Batch System 1. 2 3 4. 5. 6

pH meter (type pHM-64 Radiometer, Copenhagen, Denmark). Recorder (type Sefram-Servotrace, France). Enzyme biosensor using a combined pH electrode. Measuring cell with magnetic stirrer. 0.01 M sodmm phosphate-buffer solution, pH 7.0. Substrate standard solutions: 0.1 mMto 100 &in 0.01 Mphosphate-buffer solution.

2.52. Flow-Injection

Analysis (F/A)

1. All materials from the previous batch system. 2. FIA equipment, includmg pumps and injection valve.

3. Methods 3.1. Immersion

Method

1. Rinse the pH glass electrode tip with acetone and with distilled water, then dry it with filter paper. 2. Dip the whole bulb in a mixture of 1 mL enzyme solution, 1 mL albumin solution, and 0.07 mL glutaraldehyde solution for 5 s, so that the bulb of the electrode is completely covered.

Tran-Minh

18

pH glass electrode --Ii?)

0.5 Fig. 2. Immersion

method used to immobihze

enzyme on a pH microelectrode.

3. Rotate the electrode for 15 min to give a homogeneous enzymatic layer that coagulates after this period of time (Fig. 2). 4. Rinse the electrode with a phosphate-buffer solution to elute any excess crosslinking agent.

3.2. Direct Binding

Method

1 Rinse the flat tip of the pH glass electrode with acetone and with distilled water, then dry it with filter paper. 2. Turn the electrode upside down so that the flat part of the electrode is horizontal (Fig. 3). 3. Spread 10 uL of the enzyme mixture on the flat part of the electrode, then 10 I.~L of glutaraldehyde solution. 4 Perform crosslmkmg for 15 mm at ambient temperature. 5. Rinse the electrode with a phosphate-buffer solution to elute any excess crosslinkmg agent.

3.3. Reinforced 1. 2. 3 4. 5. 6.

Membranes

Lay the nylon net on one of the Teflon sheets Mix 0.2 mL of butyrylcholinesterase solutron wrtb 0.2 mL of glutaraldehyde solution. Rapidly spread this mixture on the whole surface of the nylon net Cover this surface with the other Teflon sheet and press firmly. Remove the enzyme nylon net after 30 min crosslinkmg. Rinse the enzyme nylon net with a phosphate-buffer solution to elute the crosslinking agent. 7. Cut the net m small circles with appropriate membrane dimensions. 8. Clip one membrane on the ttp of the pH electrode (Fig. 4).

19

p H Electrode

pH electrode Fig. 3. Immobilization method.

of enzymes on a pH electrode by the direct binding

PH electrode

pH sentive entj

:

m ~sheath

Enz y/math membrane

Clip

Fig. 4 Biosensor with a reinforced enzymatic membrane clipped onto a transducer

3.4. Aerosol

Vaporization

1. Rinse the tip of the pH glass electrode with acetone and with distilled water. 2. Dip it alternately three to four times in 0.1 MHCl and 0.1 A4 NaOH for 0.5 h each time. 3. Wash it with water and then dry it with filter paper. 4. Immerse it in the enzyme solution for about 20 min to ensure the adsorption of the enzyme on the glass 5. Dry the pH electrode at 4°C for 20 min and set it to rotate horizontally. 6. Vaporize the glutaraldehyde solution for 3 s onto the sensitive end of the pH electrode from a large enough distance to prevent formation of droplets (Fig. 5) 7. After crosslinkmg, rmse the enzyme electrode with a phosphate-buffer solution.

20

Tran-Minh Electrode

Nitrogen gas cylinder

Ll

Glutaraldehyde solution

Fig. 5. Deposition of thin enzymatic membranes m the construction of glass enzyme electrodes. Enzyme biosensor

Substrate ‘-

Recorder

Fig. 6. Enzyme biosensor based on pH electrode m a batch system.

3.5. Use of Enzyme pH Biosensors 3.5.1. Batch System 1. Connect the pH meter to the recorder and then to the enzyme sensor (Fig. 6) 2 Immerse the enzyme pH electrode m the various substrate standard solutions successively. 3. Record the potential changes corresponding to each substrate concentration 4. Plot the calibration curve.

3.5.2. Flow injection Analysis (F/A) 1. 2. 3. 4

Connect the pH meter to the recorder and then to the enzyme electrode. Connect the different parts of the FIA system as shown on Fig. 7. Pump the phosphate buffer as a carrier through the detection cell (flow rate: 1 mL/min) Inject 0.1 mL of standard substrate solutions at various concentrations mto the carrier stream successively so that it is transported to the detection cell. 5. Record the peak heights corresponding to each substrate concentration. 6 Plot the calibration curve

4. Notes 4.1. Immersion

Method

1. Immersion method can be used to immobilize enzymes or proteins onto electrodes of all shapes. 2. The enzyme layer thickness obtained by this method depends on the viscosity of the enzyme solution.

21

pH Electrode

Fig. 7. Enzyme biosensor based on pH electrode m a flow-mnJection system. 3. The correct binding time is to be adjusted: after too short a time, the coating is fragile and easily torn; if too long, it 1s less active and adherence 1spoor 4. The bindmg time depends on albumm and glutaraldehyde concentration. 5. This method is enzyme consuming because the enzyme solution with glutaraldehyde cannot be reused since it coagulates at the same time as the enzymattc layer.

4.2. Direct Binding Method 6 This techmque can also be applied to a spherical tip pH electrode 7. In this case, take care to spread the enzyme and the glutaraldehyde solution regularly on the whole surface of the sensitive end 8. This method is rather manual since particular care is taken to have an even layer

4.3. Reinforced

Membranes

9. Other fabrics than nylon net could be used provtded that the mesh size 1s not too large to have a continuous enzyme layer. 10 Reinforced enzymatic membranes are recommended m stirred soluttons. 11. This technique is well adapted to produce ready-made enzymatic membranes

4.4. Aerosol 12. 13. 14. 15.

Vaporization

Films obtained m this way are extremely The resulting biosensors have very short This procedure can be used to unmobilize It is also suitable for mass production of

thm (l-2 pm) response times (5-10 s). enzymes on a vanety of transducers. enzyme blosensors.

4.5. Use of Enzyme pH Biosensors 4.5.1. Batch System 16. The reference electrode may be combined with the working electrode. 17. The batch system 1sused to measure the response time of the enzyme biosensor. It can also monitor the biosensor stability and the progression of its response curve toward a steady state

22

TramMinh

18. If the sample volume 1stoo small, the consumptton of the substrate by the enzyme biosensor may result m a drift in its response. 19 The method is limited to manual use.

4.5.2. Flow-Injection Analysis 20. 2 1. 22 23

The method is readily adapted for small sample volumes. Automation of the flow-inJection system is easy High reproducibility of measurements is observed. Sample dilution can be optimized from the detection cell volume and interferences can be reduced. 24. High sample throughputs can be obtamed with fast response biosensors made from aerosol vaporization technique

References 1. Guilbault, G. G and Shu, F R. (1972) Enzyme electrodes based on the use of a carbon dioxide sensor Urea and L-tyrosine electrodes Anal. Chem 44, 2161-2165. 2. Tran-Minh, C. and Broun, G (1975) Constructton and study of electrodes using cross-lmked enzymes. Anal Chem 47, 1359-1364. 3 Beaux, J and Tran-Minh, C (1979) Inhibition de l’acetylcholmesterase immobll&e par les ions fluorure. C R Acad Sci. Pans, Ser. C 288, 545-548. 4. Mascmi, M., Iannillo, M., and Palleschi, G. (1983) Enzyme electrodes with improved mechanical and analytical characteristics obtained by binding enzymes to nylon nets. Anal Chim Acta 146, 135-148. 5. Tran-Minh, C and El Yamam, H. (1988) Enzyme sensors for determination of toxic chemicals m environmental samples, m Electrochemzcal Detectzon Techniques zn the Applzed Bzosciences, vol. 2 (Junter, G. A., ed.), Ellis Horwood, New York, pp. 131-141 6. Kumaran, S. and Tran-Minh, C. (1992) Insecticide determmation with enzyme electrodes usmg different enzyme nnmobilization techniques. Electroanalyszs 4,949-954 7. Kumaran, S., Meter, H., Danna, A. M., and Tran-Minh, C. (1991) Immobilizatton of thin membranes for the construction of glass enzyme electrodes. Anal Chem. 63, 1914-1918

Enzyme Biosensors

Based on Gas Electrodes

Marco Mascini and Gianna Marrazza 1. Introduction Enzyme biosensors based on gas electrodes are reviewed in Table 1. Gas probes exploited for assembly of biosensors have been mainly CO2 and NH, electrodes, and the range of concentration of most metabolites is 1OP5-1P2 A4. Enzymes are usually immobilized on the gas membrane in order to obtain the desired selectivity for specific metabolites The gas probes (often called potentiometric gas sensors) are constructed using an ion-selective electrode (generally a pH glass electrode) and a reference covered by a gas-permeable membrane (Fig. 1). A very thin film of a suitable electrolyte is present between the gas-permeable membrane and the surface of the ion-selective electrode. When the gaseous form diffuses from the sample solution through the gas-permeable membrane, it hydrolyzes in the thin film of internal solution, varying the concentration of some ions (H+ generally) detected by the ion-selective electrode. The variation of the potential 1s directly related to the concentration of gas existing in the sample, We can consider, for example, the NH, electrode, which is often used (as from Table 1) in the construction of several biosensors. In this case the internal solution is 0.1 MNH,Cl, and we will use a pH electrode as internal electrode. The potential of the internal cell (E,,tJ, referring to Fig. 1, will be E cell= K + 59 log [H+]

(1)

where K is a constant and the [Hf] IS calculated by the equihbrmm of NH,+ t) NH3 + H+ [H+] = K [NH,+]

[NH31 From

Methods III Botechnology, Vol. 6 Enzyme and MIcrobra/ Blosensors Techniques Edlted by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

23

and Protocols NJ

(2)

2

Uricase

Uric acid

L-Glutamate Hrstidine

Creatinme D-Ghtconate

L-Lysine Methromne r.-Phenylalamne L-Tyrosme Urea

Enzyme

Based on Gas Electrodes

Asparaginase Aspartase AMP deaminase Phospho-diesterase and AMP deaminase Creatinine deaminase Gluconate kinase and 6-phospho-n-gluconate dehydrogenase Glutamate decarboxylase Histidine decarboxylase Histidine amomalyase Lysme decarboxylase Methionine lyase Phenylalamne decarboxylase Tyrosine decarboxylase Urease

L-Asparagme Aspartate 5’-AMP Cyclic AMP

Substrate

Table 1 Enzyme Biosensors

PC02 PC02

PNH,

PC02 PC02

PNH,

PC02

PNH~

PC02 PC02

PC02

1 x ltY-22.5

x 1C3

3x10-4-1x10-2 3x1~5-1x10-* 1x10-4-2x1tY3 1x10-~-1x1~* 2.5 x l&j- 1.5 x l&2 1x1~-5x10-~ 1x10”‘-1x1&’ lxlti-1x1&2

7xlV-7x10-3

13 14 15

10 12

11

7 8 9 10

4 5 6

1 x 10-s- 1 x l&2 7x10-5-1x10-2 1.2 x ltY-24 x 1Cr3

PM3

PNH, PNH,

1

Reference 2 3

range (A4)

8x 1~s-8x10-3 7x10‘4--2x1~* 8 x 10-s-- 1.5 x l&2

Concentration

PNH, PNH,

Gas probes

Gas Electrodes

25

selective electrode

internal electrolyte

Leas-permeable membrane Fig 1. Scheme of a gas probe

When the electrode is dipped m a solution where a defined partial pressure of NH3 is present, the equilibrium in the film behind the surface of the pH electrode will shift accordingly, and the potential will be: E cell = K’ - 59lOg ([NHdsampd[N%‘l

(4)

where K’ is a constant. The concentration of the gaseous form, [NH,], will be the same both in the sample solution and in the internal thin film in contact with the pH electrode. Considering [N&+1 constant becauseof the high concentratton of internal electrolyte: Ecell = K” - 5910g [NH31

(5)

where K” is a constant. Figure 2 reports the calibration curve of several gas sensor probes as a functron of the concentratron of the species in solution. Generally when these gas electrodes are used, the pH of the sample needs to be adjusted to generate the gaseousform for which the sensoris assembled.For example,to measureNH,+ ions m the samplewe should shift the pH in alkaline medium above 10 to transform all ammonia to its gaseous form as expected from the diagram reported in Fig. 3. This is a crucial point. When we assemble biosensors, couplmg gas electrodes and immobilized enzymes,we cannot use alkaline solutions that are too strong because the activity of the enzyme will drop drastically or the activity could be permanently lost. Therefore in this coupling we should find a com-

26

Mascini and Marrazza

-200

-7

-6

-5 dissolved

-4

-3

gas concentration

-2

-1

0

log (M]

Fig 2 Calibration curve of somegasprobes. promise for the pH of the sample between the acttvtty of the enzyme and the features of the gas electrode. Figure 4 reveals that, m the case of urease with an ammonia electrode, the response time of urea changes with the pH of the sample solution. In this case a value of 8.5 was suggestedas the optimum working pH. However, from Fig. 3 we can evaluate that only 10% of the ammonia is in its gaseousform at this pH value. When we use carbon dioxide electrode the features are similar. The electrolyte used in this case is 0.1 MNaHCO, and the equilibrium is described by: CO2+ HZ0 f) HC03- + H+ (6) and therefore considermg [HC03-] constant: E = K” + 5910g[C02] (7) Compromtses in pH values are realtzed when the COZ electrode is coupled with nnmobilized enzymes. Also, m this case, the more acidic the pH of the sample is, the more the equilibrium will be shifted toward the gaseous form, C02. When immobilized enzymes are coupled on the surface of gas membranes, low pH values should be avoided, and also a compromise between activity of enzymes and features of the COZ probe should be established. To assemble an enzyme biosensor, we should immobiltze an enzyme on the surface of the gas electrodes. We can use an enzyme that cleaves a specific metabolite forming a gaseous form, NH3 or CO*. Urease is a useful example

27

Gas Electrodes 0 I -2

0 3 -4

-6 j I

I

I

I

I

I

I

0

2

4

6

8

10

12

14

PH measured species

diffusing species (G)

Ml3 or NH4

NH3

equilibria

in electrolyte

sensing electrode

H+

M-I3 + H20 f rG&++oHxNH3 + M*+ f M(NH3)xn+

M= Ag+, Cdz’,

CO2,

WO3,

Cu2+

CO2

CO2+ H20 5 H+ + HCO3-

H+

SO2

SO2+ Hz0 = H+ + HS03-

H+

HC03‘. CO32-

SOz,H2SO3Or

s032-

Fig. 3. Logarithmic

diagram for equilibria involving gas species. urea urease

) Nb+ + HC03 + OH-

(8)

When we immobilize urease on ammonia gas probe we obtain a sensor that is sensitive to urea concentration: E = K” - S log[urea]

(9)

where S is the slope of the assembled sensor (theoretically 59 mV, but we will obtain lower values, m the range of 45-55 mV). We describe below hoti to assemble a biosensor based on gas probes and how to use it (15,16) 2. Materials 2.1. Assembly

of the Gas Electrode

1. Internal solution for ammonia gas sensor: 0.1 MNH&I solution in deiomzed water. 2. Internal solution for CO2 gas sensor: 0.1 A4 NaHC03 and 0.1 M NaCl in delonlzed water.

Mascini and Marrazza

28

pH

values

8.5 and 8.0 and 7.5

9.0 9.5

-10 * urea 0

a

5 min

Fig. 4. Response time of a urea sensor as function of pH of the sample. Urea concentration in the sample increased from lo4 to 10e3 M.

2.2. Immobilization of the Enzyme on the Gas Permeable Membrane 2.2.1. Physical Entrapment 1 The enzymes are commercially available from Sigma Chemical Company (St. Louis, MO). 2. Gas-permeable membranes are made of materials such as polytetrafluoroethylene (PTFE) with 0.2-0.5urn pores, silicon rubber from Whatman (Fairfield, NJ) or Milhpore (Bedford, MA). 3. Dialysis membrane: 20-25 urn thickness, 20 kDa. Molecular wreght cutoff (MWCO) is commercially available from Spectrum Medical Inc. (Houston, TX).

2.2.2. Chemical Bonding 1. 0.1 M Phosphate buffer, pH 7 0. 2. 2.5% Glutaraldehyde in phosphate buffer. Add 1 mL of 25% glutaraldehyde (Sigma) to 9 mL 0.1 Mphosphate buffer, pH 7.0. The solution is stored at 4°C for 2 wk.

Gas Electrodes

29

3. 25% Bovine serum albumin (BSA) in 0.1 M phosphate buffer, pH 7.0. Dissolve 25 mg BSA in 10 mL phosphate buffer. The solution is prepared daily. 4. 0.1 A4 Glycine solution in 0.1 M phosphate buffer, pH 7.0 5. 0.1% Kathon in phosphate buffer, pH 7.0. Mix 100 pL Kathon CG UN N” 1760 (Rohm and Haas, UK) in 99 mL 0.1 Mphosphate buffer, pH 7.0. 6. 0.02% Sodium azide in phosphate buffer, pH 7.0. Dissolve 20 mg sodium azide in 100 mL 0.1 Mphosphate buffer, pH 7.0.

2.2.3. Use of Preactivated Membrane 1. Preactivated membrane. Immunodyne membranes from Pall Co (NY, USA); Immobilon AV from Millipore Co.; Ultrabind from Gelman Sciences, (MI, USA). 2 Enzymatic solution. Dissolve 10 mg of enzyme in 10 mL 0.1 Mphosphate buffer, pH 7.0 and use promptly. 3. 0.1 % Triton X-l 00 m phosphate buffer pH 7.0. Mix 100 pL of Triton X- 100 and 100 mL of 0.1 Mphosphate buffer, pH 7.0.

3. Methods 3.1. Assembly

of the Gas Electrode

1 Use the gas electrode equipped with an internal pH electrode with a flat surface, carrying on its body a silver/silver chloride electrode acting as reference 2. Fix the electrode into an external tube, at the base of which the gas-permeable membrane is located. 3. Pour the internal electrolyte into the tube. The electrolyte should contam 0.1 M chloride ions to fix the potential of the reference electrode and to fix at 0.1 A4 the concentration of the ion that is in equilibrium with the gaseous species 4. Use 0. I MNH&I as electrolyte for an ammonia gas sensor, and 0.1 MNaHCOs with 0.1 M NaCl as electrolyte for the carbon dioxide sensor. 5. Tighten the body of the electrode by screwing it; this operation will push the flat surface of the pH electrode against the gas-permeable membrane (see Note 1).

3.2. Immobilization of the Enzyme on the Gas-Permeable Membrane The enzyme can be immobilized often used are considered here.

in several ways. Three procedures

most

3.2.7. Physical En trapmen t 1. Fix the electrode upside down and add 2-3 mg of the lyophihzed enzyme on the gas-permeable membrane of the gas sensor 2. Place a thin dialysis membrane over the powdered enzyme. 3. Fix tightly the dialysis membrane with a suitable O-ring. 4. Stretch the dialysis membrane over the gas-permeable membrane to avoid air bubbles. 5. Put the electrode m the correct position.

Mascini and Marrazza

30

6 Dip it in the electrolyte solution to obtain a cahbration curve and to measure the sample concentration

The enzyme sensor obtained in this way generally works for a limited amount of time, i.e., 2 or 3 d, therefore, it should be used immediately

after

assembling it. 3.2.2. Chemical Bonding 1 Fix the electrode upside down with the gas membrane facing upward. 2. Put l-2 mg of enzyme on the gas membrane, 5 pL of 25% BSA in phosphate buffer, and 2-3 yL of 2.5% glutaraldehyde in phosphate buffer. 3. Mix the slurry nnmediately for 15-20 s. 4. Let it dry for 20 mm. 5. Invert the electrode when the film is formed. 6 Dip the electrode mto 0.1 M glycme solution m 0.1 A4 phosphate buffer, pH 7 0, and leave it m for 30 mm 7 Use the electrode for a calibration curve. 8 Kept it m dry state m a refrigerator, using either 0.1% Kathon or 0.02% sodium azide m buffer solutions as a preservative.

The enzyme immobilized using this procedure is very stable and can be used for several weeks, sometimes for several months. 3.2.3. Use of Reactivated

Membrane

1. Use specially preactivated membranes commercially available for munobihzmg proteins (see Note 4). 2. Dissolve the enzyme m electrolyte solutions, avoiding the use of NH2 groups or NH4 ions. 3 Use an enzyme concentratton of about 10 mg/mL m phosphate buffer. 4. Cut the membrane in the form of the gas-membrane electrode and wet it using the enzyme solution for about 30 min at room temperature. 5. Wash the membrane in phosphate buffer with some detergent, for example, 0 1% Tnton X- 100, for a few minutes to wash away the enzyme that is not covalently bonded. 6. Fix the membrane against the gas-permeable membrane, using an 0-rmg, parafilm, or some glue; generally the membrane can be fixed m the same way as the gas-permeable membrane. 7 Press it to the gas-permeable membrane

3.3. Measurement Procedure 3.3.1. Calibration Curve 1 Prepare four to five solutions at known concentration of metabohte in the range 1c5-1 0-l M; for instance, by sequential dilution five urea solutions at concentration l&l, 1t2, 10-3, l@, and 1c5 M, all m buffer suitable for enzyme reaction and for the gas sensor sensmvity (see Note 5).

Gas Electrodes

37

2. Choose the pH 8.5 in Tris buffer in the case of urea 3. Dip the enzyme electrode into 20-30 mL of each solution starting with the most dtluted one and then proceed to the more concentrated. 4. Stir the solution gently using a magnetic stirrer to measure the potential 5. Record the value obtained 60-90 s after the immersion. 6. Plot the calibration curve on semilogarithmic paper with potential values on the Y-axis and log of concentration on the X-axis, as shown m Fig. 5.

3.3.2. Measurement

of Sample Concentration

1. If the pH and the buffer used to prepare the sample are sinular to those employed for the construction of the calibration curve, dip the electrode in the sample; otherwise prepare a certain amount of buffer (20 mL) without metabolite and dip the electrode mto It. 2. When the potentials reach a stable value, add a certain amount of sample (i.e., 1 or 2 mL). 3. Measure the potential after 60-90 s by gently stirring with a magnetic device as in the preparation of the calibration curve. 4. Calculate the concentration of metabolite by interpolation of the value m mV from the calrbratton curve

3.3.3. Continuous Flow Measurement The continuous flow measurement permits evaluation of more samples and standards per unit time (see Note 7). 1. Use a suitable flow cell for the assembled electrode. The void volume should be as low as possible, generally in the range of N-100 I.IL. The flow rate should be in the range of 1 n&/mm. 2. Assemble the connections between the pump and the flow cell m order to mmrmize the void volume (m the range of 100-200 pL). 3. Use a flow rate at least three times higher than the total void volume, i.e , 1 mL/mm flow rate for a maximum void volume (tubes and flow cell) of 300 pL 4. Prepare a certain number of standard solutions m the range indicated in Subheading 3.3.1.( lc5 - 10-l M) m a suitable buffer and use thus buffer as the blank solutron. 5. Start the measurements from the most dilute standard solutron. 6. Let each solution flow for at least 2 min, then change solutions. 7. Record the potential values obtained by a flowing standard solution. 8. Plot these values for a calibration curve, mV vs log concentration. 9. Prepare the sample with the same buffer as the standard solutions. 10. Let it flow mto the cell using the peristaltic pump and record the potential value. 11. Flush the cell after each measurement using the buffer solution. 12. Evaluate the concentration of the sample by interpolation of the calibration curve.

4. Notes 1. The assembly of a gas electrode is a crucial point (see Subheading 3.1.). The layer between the gas-permeable membrane and the sensor should be as thin as

time

-4 log (cone

Fig. 5. Calibration curve for enzyme-based gas sensor.

E 0-W

(W

30 mV

I

E

10 mln -

-3 l(M)

-2

+

Gas Electrodes

2.

3.

4.

5

6

7.

33

possible. Therefore the sensing element of the ion-selective electrode is pushed against the membrane. It is important that the sensing element IS on the tip of the electrode body. The electrolyte should preferably be 0.1 M as noted, but if a more sensitive blosensor is required, a more dilute internal solution, i.e , 0.01 M, can be used. This will extend the linear part toward lower concentrations even if the response time for equilibration will increase. Therefore, a compromise must often be reached. Immobilization of enzyme by physical entrapment is the simplest way to immobilize an enzyme Always try it before other procedures. It IS a very fast procedure and avoids the risk of denaturation of the enzyme because no chemicals are added. Chemical bonding is a classic procedure, but several enzymes are deleteriously affected by glutaraldehyde treatment. Therefore, the amount of glutaraldehyde used can be varied. Often one-tenth of the recommended concentration can be used and still give good results. Some enzymes can be problematic. As a rule of thumb, always put l-2 U enzyme per cm2. Use the preactivated membrane whenever possible. Generally the membranes commercially available are thick; thin membranes are preferable because they give faster response times To obtain a good response, use an enzyme quantity for immobilization as reported in Subheading 3.2.3. In our laboratory, often the procedures of Subheadings 3.2.2. and 3.2.3. have been coupled to enhance the activity of the immobilized enzyme, i.e., to use both glutaraldehyde and BSA over the preactivated membranes. A calibration curve generated for biosensors with a slope ranging from 40 mV/ decade to 53-55 mV/decade (which is the best result) is generally accepted (see Subheading 3.3.1.). The range is from 1@-10-2 Mas from Table 1. Measurement of sample concentration should be arranged so that the composition of the buffer and the ionic strength of the sample are as close as possible to those of the standard solutions used to construct the calibration curve Continuous flow measurement in a flow cell can increase the response time because the washing of the probe is not as efficient as in Subheading 3.3.1. Therefore, always follow the procedures m Subheading 3.3.1. in order to define the response time and the recovery time of the probe, then mimmize the void volume of the flow cell and the pathway of the flowing solution The hydrodynamic conditions are of paramount importance for a fast response

References 1. Wawro, R. and Rechnitz, G. A (1976) Immobilized enzyme electrode for L-asparagine. J Membr. Scl 1, 143-148. 2. Fatibello-Filho, O., Suleiman, A., and Guilbault, G. G. (1989) Enzyme electrode for the determination of aspartate. Biosensors 4, 3 13-32 1. 3. Papastathopoulos, D S. and Rechmtz, G. A. (1976) Highly selective enzyme electrode for S-adenosine monophosphate. Anal Chem 48,862%864.

34

Mascini and Marrazza

4 Meyerhoff, M. and Rechnitz, G A (1976) An activated enzyme electrode for creatinine. Anal Chim. Acta 85,277-285. 5 Jensen, M A. and Rechmtz, G. A. (1979) Enzyme “sequence” electrode for o-gluconate. J. Membr Sci 5, 117-127 6 Arnold, M. A. and Rechnitz, G A (1980) Comparison of bacterial, mttochondrtal, tissue and enzyme biocatalysts for glutamine selective membrane electrodes Anal Chem 52,1170-l 174 7 Kovach, P M. and Meyerhoff, M. (1982) Development and application of a histidine-selective btomembrane electrode Anal. Chem 54,2 17-220. 8 Walters, R. R., Johonson, P A., and Buck, R. P. (1980) Htstidine ammonia-lyase enzyme electrode for determinatton of L-htstidine Anal. Chem 52, 1684-l 690 9. White, W. C. and Gutlbault, G. G. (1978) Lysine specific enzyme electrode for determination of lysme m grams and food stuffs. Anal Chem. 50, 1481-1485. 10. Fung, K. W , Kuan, S. S., Sung, H. Y., and Gutlbault, G. G. (1979) Methtonme selective enzyme electrode Anal. Chem. 51,2319-2324. 11 Guilbault, G. G. and Shu, F. R. (1972) Enzyme electrodes based on the use of a carbon dioxide sensor Anal Chem 44,2 161-2 166 12. Mascini, M. and Guilbault, G. G. (1977) An urease coupled ammonia electrode for urea determination in blood serum. Anal Chem 49,795-798 13. Tran-Mmh, C and Brown, G. (1975) Construction and study of electrodes using cross-linked enzymes. Anal Chem 47, 1359-l 364. 14. Kawashima, T. and Rechnitz, G. A. (1979) Potentiometrrc enzyme electrode for uric acid. Anal Chzm Acta 83,9-17 15. Mascml, M. and Palleschi, G. (1989) Design and apphcatlons of continuous momtoring probes. Se1 El Rev. 11, 191-264. 16 Mascmt, M (1995) Potentiometry. enzyme electrodes, m Encyclopedza of Analytzcal Sczence (Townsheand, A , ed.), Academic, London, UK, pp. 4 112-4 118.

4 Enzyme Biosensors

Based on ISFETs

Roland Ulber and Thomas Scheper 1. Introduction The ion-sensitive field-effect transistor (ISFET) can be regarded as a successful combination of two well developed techniques: sohd-state integrated circuits and ion-sensitive electrodes. Bergeveld (1) introduced the ISFET as a new device that combines the chemical-sensitive properties of glass-membrane electrodes with the impedance-converting characteristics of the metaloxidesemiconductor field-effect-transistor (MOSFET). In the last 25 yr, a large number of different ISFET types were developed, which differ in terms of the detectable ion and the ion-sensitive surface. Table 1 gives some examples. 1.1. Operation Principle of ISFETs As shown from Fig. 1, an ISFET consists of an ion-sensitive membrane and an FET-structure in which the metal electrode (gate) is removed and its function 1s taken over by the solution under investigation. Transistors for ISFETs are manufactured according to standard planar silicon technology, and frequently Si3N4/Si02 /Si structures are used. Each laboratory that fabricates ISFET devices uses different processing sequences and procedures, and thus a description of the complete process is both impractical and beyond the scope of this text. (For a detailed description of semiconductor processing see ref. 15.) The operation principle of an ISFET is the formation of a potential difference at the membrane/solution interface influencing the width of the sourcedrain channel of the FET. Since the value of the interfacial potential

From

Methods m Botechnology, Vol 6 Enzyme and Microbral Wosensors. Technrques Edited by A Mulchandani and K R. Rogers 0 Humana Press Inc , Totowa,

35

and Protocols NJ

Ulber and Scheper Table 1 Different Types of ISFET Detectable ions

Ionsensitive surface

K+ Na” Ag+ Ca*+ NH4+ cu*+ Ni2+ Pb*+ Cd*+ NO> ClBrFH+

Polymeric membraneiionophore Polymeric membrane/ionophore Si02 membrane PVCYronophore Polymeric membrane/ionophore As2Se3/Cu membrane As2Se3/Ni membrane Polymeric membrane/ionophore As2S3/Ag2SlCdS membrane Magnesium phosphate glasses Polymeric membranelionophore AgBr LaFs S&NJSiOJSr

References 3 3 4 5 6 7 7 8 9 10

11 12 13 14

difference depends on the concentration of corresponding ions rn solutron, the change of the FET characteristics (e.g., drain-source-current In) will be a mea-

sure of the analyte concentratton. The slope can be described by the Nernst equation. E = E" + (2.303 x RT)/(z, x F) log a,

(1)

The interested reader should consult Vlasov (16), Bergveld and Sibbald (17), or Blackburn (18) for more detailed descriptions of the physics of ISFET devices. 1.2. Enzymatically

Modified

ISFET

Many biosensors based on ISFETs were created after the first report about an enzymatic modified FET (EnFET or BioFET) for the determination of penrcillin (2). In comparison with other types of biosensors, the ISFET shows certain well-known advantages, such as miniaturization, high sensitivity, low cost and multianalyte detection potential. An ISFET can be transformed mto a biosensor by immobilizing a thm biologically active layer on top of the ionselective membrane (Fig. 2). Most reported EnFETs use pH-sensitive FETs. A few EnFETs, which use fluoride-sensitive ISFETs, have been reported. From the very beginning of studies using ISFETs, it was observed that insulators in metal-oxide semiconductors, such as Si02 and Si3N4, were pH-sensitive; therefore, the pH

ISFETs

37

&$j

St,N,

m

n+-Si

13

p-Si

m

1 semi-conductor

4 source 6 contact

Al

&@

encapsulation

SI,N, I OH

&gg

w

2 Isolator 5 drain 7 reference

3 pH-sensitive

area

electrode

Fig. 1. Diagram of apH-ISFET. ISFET is a widely investigated sensor. Different membrane materials have been studied and used in pH-FETs (I%22). Some types of pH-FETs are now commercially available and interesting EnFET applications have been described (see Table 2). The working principle of an EnFET is as follows: during the enzymatic reaction in the enzyme membrane, protons are generated or consumed. The pH changes as a result of the enzymatic reaction are measured via the pH-FET and can be correlated to the analyte concentration. 2. Materials The following electronic components and chemicals are needed for the described experiments. As solvents one needs distilled and ultrapure water.

38

Ulber and Scheper

Table 2 Substrate-Enzyme Systems Used on pH-FETa Substrate

Enzyme system

Ref.

Urea Cephalosporin C Penicillin G R,S-3-Hydroxy carbon acid esters R&Amino acid esters Glutamine

Urease Cephalosporinase Penicillm G amidase Lipase/esterase

2,23 22 2,2# 25

Glucose Ethanol Acetylcholine ATP Inosine Lactate

a-Chymotrypsinlesterase Glucose oxidasel concanavalin A Glutamine synthetase Alcohol dehydrogenasesl Aldehydedehydrogenase Acetylcholine esterase H+ ATPase Xanthme oxidase Lactate oxldase

25 26 27 28,29 30 31 32 33

aDrfferent mnnobrlrzation methods and the influence of vanous process parameters on the sensor signal are described m the practical part (methods) of thus chapter

enzyme membrane

pH-FET

Fig 2. Biosensor based on a pH-sensitive

FET.

ISFETs

39

2.1. Experimental Setup 2.1.1. Constant-Charge Circuit 1. Electronic components: TL074 CP 4 TL071 CP 4 TL072 CP 4 OP27 1 REFOl DT 1 BF256 B 4 lk0 4 1okQ 5 20 kR 18 47 kR 4 51 k0 4 1000 4 91 kR 4 50 kQ spindle 4 10 kQ spindle 8 10 yF, 35 V, tantal 2 1 uF, 35 V, tantal 1 100 nF 50 plug, 16 pol 1 plug, 32 pol 1 2. Reference electrode, e.g., Ag/AgCl electrode; Mettler-Toledo (Stembach, Germany) Type 373-M3 3. pH-Sensitive field-effect transistor, e.g., S&N4 type; abc GmbH (Puchheim, Germany) or Ta205 type; SenDx Medical (Carlsbad, CA).

2.1.2. Measuring of the pH-Sensitivity

of a pH-FET

1. 5 mL 10 mM potassium hydrogen phosphate (K2HP04) pH 2.0. 2. 5 mL 1 A4 sodium hydroxide (NaOH) in distilled water. 3. 0.146 g sodium chloride NaCl.

2.2. Immobil~zatlon 2.2.1. lmmobikation 1. 2. 3. 4.

in distilled

water;

of Enzymes on pti-FETs on Activated /SFETs

5 mL 10% nitric acid (HNOs). 1 mL 3-(triethoxysily) propylamine (y-APTES) [HzN(CH,),Si(OC,H&j. 1 mL 6% glutaraldehyde (HCO(CH,),CHO) in distilled water. 5 mg glucose oxidase (E.C. 1.1.3.4), from Aspergilhs nzger type VII (Sigma, St. Louts, MO). 5. 500 l.tL 10 mMpotassium hydrogen phosphate (K2HP04) in distilled water; pH 8.0.

Ulber and Scheper

40

2.2.2. Immobilization by Cocrosslinking with Glutaraldehyde 1. 2. 3. 4.

10 pL 10 mA4potassmm hydrogen phosphate (K2HP04) in distilled water; pH 6.0. 1 mg glucose oxrdase (E.C. 1.1.3.4), from A niger type VII (Sigma). 2 pL human serum albumin (HSA). 1.5 pL 25% glutaraldehyde (HCO(CH&ZHO) m water.

2.3. Sensor Evaluation 2.3.1. Calibration Curve of a Glucose Oxidase-Modified 1. 2. 3. 4

pH- FET

5 mL 20 mM potassium hydrogen phosphate (K2HP04) in disttlled water; pH 6 0. 0 165 g potassium ferricyamde (K,[Fe(CN),]). 0 146 g sodium chloride (NaCl). 1 mL 100 g/L glucose solution in dtstilled water

2.3.2. Influence of the Cosubstrate Concentration on the Sensor Signal 1 2 3 4

3 x 5 mL 20 mMpotassmm hydrogen phosphate (K&PO,) m distilled water; pH 6 0. 0 083, 0.041, or 0 021 g potassium ferricyamde (K,[Fe(CN),]). 3 x 0.146 g sodmm chloride (NaCl) 1 mL 100 g/L glucose solution in distilled water

2.3.3. Influence of the Buffer Capacity on the Sensor Signal 1. 1, 10, and 100 mM potassium hydrogen phosphate (K,HP04) pH 6.0, each time 5 mL. 2 3 x 0.165 g potassium ferrrcyanide (K,[Fe(CN),]) 3. 3 x 0.146 g sodmm chloride (NaCl) 4 1 mL 100 g/L glucose solution m dlstrlled water

2.4. EnFET for Enantioselective

in distilled water,

Analysis

1. 5 mL 20 mM potassium hydrogen phosphate (K2HP04), m distilled water, pH80 2. 0.146 g Sodium chloride (NaCl). 3. 450 U esterase from porcine liver (E.C.3.1.1.1) (Sigma). 4. 120 U a-chymotrypsin Type II (E.C.3 4.21.1) from bovine pancreas (Sigma) 5. 50 mg S-phenylalanine methyl ester hydrochloride (C,H&H,CH(NH,)CO,cH, . HCl). 6. 50 mg R-phenylalanine methyl ester hydrochloride (C,H&H,CH(NI$)CO,CH, . HCl)

3. Methods 3.7, Experimental Setup 3.1.1. Constant Charge Circuit Figure 3 depicts the wiring diagram of a constant charge circuit used for any kind of ISFETs. This circuit regulates the drain-source current (ID) that

n

1

B

I

C

I

0

I

I

F I

B

Fig. 3. Constant-charge circuit.

E

I

n *sJ

-1Y

I

I I

K

I

1 1

1

42

Ulber and Scheper

normally changes while changing the potential of the ion-sensitive gate during the enzymatic reaction. In changing the voltage between reference electrode and source (II,), the current between drain and source remains constant. This compensation voltage is the measuring signal. The circuit enables one to regulates the working point of the ISFET. Typical values are Uris = 2 V and In = 100 p.A. A reference electrode such as a calomel- or Ag/AgCl- based electrode is necessary for the experimental use of ISFET, since the signals must be correlated to a constant potential. Because these electrodes depend on the concentratron of the ions used in the electrode (hke Cl-), one must take care that there is no change m this parameter durmg the measurement.A calomel type of electrode 1simpossible to use becauseof the presenceof mercury by environmental analysis with ISFETs. For microelectrodes, only a Ag/AgCl electrode can be a useful candidate. 3.1.2. Measuring the pH-Sensitivity

of a pH-FET

1. Dissolve the sodium chloride m the phosphate buffer and adJust the pH value to 2.0 2. Integrate a pH-FET and the reference electrode m the buffer. 3. Switch on the constant charge circuit. Attention: Never switch on the circuit while the ISFET is out of any solution This will damage the ISFET (see Note 1) 4. While stirring (magnetic stirrer), increase the pH m steps of one pH decade up to pH 12.0 with 1 M sodium hydroxide solution. 5. Evaluate pH vs compensation voltage; typical slope (depending on the gate material) range from 45 mV/pH-decade up to 56 mV/pH decade (Fig. 4.).

3.2. Immobilization of Enzymes on pH-FETs In most cases,the EnFET is fabricated by immobihzing the enzyme in a matrix of crosslinked albumin, polyacrylamide, or triacetyl cellulose overlaid on the sensitive surface of the ISFETs. When using pH-FETs as transducers, at least a dual-gate FET is employed so that one of the FETs can act as a reference for the EnFET when its gate is coated with an enzyme-free membrane. If the difference between the two drain currents is monitored, the signal is insensitive to changes in pH of the solution, temperature, or electrical noise. 3.2.1. Immobilization on Activated ISFETs Depending on the hydroxide groups on the surface one is able to modify the sensitive area with, e.g., 3-(triethoxysily)propylamine (y-APTES) for immobilization on activated surfaces of pH-sensitive field-effect transistors 1 Clean the pH-FET for 30 mm m a solution of 10% HNOs at 80°C. 2. Silanize the pH-FET for 1 h in a solution of 10% 3-(triethoxysily)propylamine (y-APTES) in ultrapure water, pH 3.5. 3. Rinse with distilled water and dry for 2 h at 80°C. 4. Activate the silamzed pH-FET with a solution of 6% glutaraldehyde m 10 rnM potassium phosphate buffer (pH 9 0) for 3 h at 6’C

ISFETs

43 1700 , --+-

0

1600-

pH-FET (Si,N, gate)

/

\

\, 5

1600-

\o

bE9 z

\, 1400-

10

.z? tn & 1300-

etepness

66,3mVA3 \,

E 8 *200-

\

‘0

llOO-I

\

10

I

I

1

I

I

I

2

4

6

6

10

12

PH

Fig. 4. Slope of voltage vs pH of a pH-FET with a Si3N4gate. The pH-FET is now prepared for the immobilization of any enzymes on the activated surface (see Note 2). 5. Incubate the activated pH-FET with a solution of glucose oxidase (5 mg) in 500 l.tL 10 m.M potassium hydrogen phosphate and 0.5 A4 sodium chloride buffer (pH 6.0) overnight at 6°C. 6. Rinse with the sensor with distilled water and store at 6°C m 10 mM phosphate buffer (pH 6.0).

3.2.2. Immobilization by Cocrosslinking with Glutaraldehyde This method is faster than the immobilization on activated surfaces. It can be used for enzymeswith high activities, such as glucose oxidase, glucose dehydrogenase,urease, esterase,or a-chymotrypsin. 1. Dissolve the glucoseoxidase(1 mg) in abuffer (10 pL) containing 10nMpotassium hydrogen phosphate (pH 6.0). 2. Add 2.5 pL HSA (enzymatic inactive protein as spacer). 3. Add I .5 pL of 25% (w/v) glutaraldehyde solutton. 4. Apply the mixture immediately to the pH-sensitive gate surface, within 30-90 s a membrane is formed in which the enzyme is immobilized (see Note 3).

5. Washthe EnFET with distilled water; store in 10 mA4phosphatebuffer solutton (pH 6.0) (see Subheading 1.) at 6°C.

Ulber and Scheper

44

For more details of enzymatic immobilization see, e.g., Hartmeier (34), Anzai et al. (39, or Gtl et al. (36). 3.3. Sensor Evaluation 3.3.1. Calibration of an Enzymatically Modified pH-FET (Glucose Analysis) EnFETs provides a rapid and rehable method for the determination of glucose in medical and biotechnological applications (e.g., in blood samples or in a btoreactor). 1 Use a glucose oxidase modified pH-FET (see Subheadings 3.2.1. or 3.2.2.). 2 Reactron system (37). glucose + FAD GOD FADH* + 2 [Fe(CN)$-

d

D-glucono-6-lactone FAD + 2 [Fe(CN),#+

+ FADHl

(2)

2 H+

(3) 3. Prepare the measuring buffer by drssolvmg sodmm chloride (0.146 g) and potasslum ferricyanide (0 165 g) in 5 mL of 20 mA4 potassium hydrogen phosphate buffer. Adjust the pH value to 6.0 4. While stimng, increase the glucose concentration from 0 g/L up to 5/L, m steps of 0.25 g/L,, using a glucose solution of 100 g/L, m drstrlled water (pH 6.0) (see Note 4). 5 Evaluate sensor signal vs glucose concentration (Fig. 5).

Investigations of the influence of various parameters on the EnFET signal showed that pH, buffer capacity, and cosubstrate concentration affect the sensor signals. These following influences are general features of an EnFET. 3.3.2. Influence of the Cosubstrate Concentration on the Sensor Signal 1. Repeat the experrment described m Subheading 3.3.1. with different concentrations of potassmm ferrrcyamde (50, 25, and 12.5 n&I). 2 Evaluate sensor signal vs glucose concentration by different potassium ferrrcyanide concentrations (Fig. 6)

3.3.3. Influence of the Buffer Capacity The influence of the buffer capacity is also important. An increase in buffer concentration reduces the magnitude of the sensor signal, simultaneously enhancing the measuring range of the biosensor. This is explained by the increased number of buffer molecules in the enzyme-membrane buffering part of the protons generated by the enzyme reaction. 1. Repeat the experiment described in Subheading 3.3.1. with different potassium hydrogen phosphate concentratrons of 1, 10,50, and 100 mM 2. Evaluate sensor signal vs urea concentration by different buffer concentrations (Fig. 7).

ISFETs

45

n ----

60-

./=----

W---.

-B--

GOO-FET

s El 3 40.6 In “0 VJ20c oI

I

I

,

1

1

0

1

2

3

4

5

glucose

[g

I-‘]

Fig. 5. Calibration curve of a glucose oxidase-modified

pH-FET.

i 7060-

5;‘ 50L 3 40& ‘jj 30k g 20: IOo-10

I

I 0

I 1

1 2

I 3

f 4

I 5

glucose [g lml]

Fig. 6. Influence of K,[Fe(CN), oxldase-modified pH-FET.

concentration on the sensor signal of a glucose

46

Ulber and Scheper [

110 IOOgo60 Y 5

7060-

7 &

50-

'5 &

40-

2

30-

ii

20-

-a--

100 mM 1-l K,HPO,

IO-

--a--

20 mM I”

--A---

1 mM I-’ K,HPO,

o-10

'

I w

I

1 13

‘33

I 1,6

K,HPO,

I

I

283

glucose [g P]

Fig. 7. Influence of the buffer capacity on the sensorsignal of a glucose oxidasemodified pH-FET.

3.4. Special Applications

of EnFETs

3.4.1. EnFET for Enantioselective

Analysis

In the pharmaceutical industry tt is very important to produce enantiomerically pure substances, since both enantiomers of a choral bond can have different therapeutic effects. The conventional methods (e.g., using chiral columns m gas chromatography, liquid chromatography, or various nuclear magnetic resonance spectroscopic methods) used for the determination of enantiomeric excesses are time-consuming and expensive. Enzyme-field effect transistors can be used for the fast analysis of the enantiomeric excess, e.g., of amino acid esters. A combined sensor with immobilized achymotrypsin and porcine liver esterase can be used for the analysis of R,S-3-phenylalanine methyl ester. This enzyme combination was chosen since the esterase hydrolyzes the substrate nonspecrfically into the respective acid and alcohol, whereas the a-chymotrypsin converts only the Senantiomer. 1. Immobilize esterase(E.C.3.1.l. 1)on the surfaceof the pH-FET by cocrosslinking method (see Subheading 3.2.2.). 2. Immobilize a-chymottypsin (E.C.3.4.21.1)on the surface of a secondpH-FET by cocrosslinking 3. Reaction system:

47

ISFETs

40

v

n

n

0

oyo

0

o/O .p

/

.p O/

/

0

-l

II

II

II

II

II

0

20

40

60

80

S-phenylalanine

methylester

Fig. 8 Enanttoselectrvtty

II

100

[%J

of EnFETs.

pomne her esterase

R,S-phenylalamne

methylester

S-phenylalamne

methylester

->

R,S phenylalanine

+ methyl alcohol

(4)

a-chymotrypsm

->

S-phenylalanine

+ methyl alcohol

(5)

4. Measuring buffer: 20 mM L, potassium hydrogen phosphate 0.5 M L, potassium chloride, pH 8.0. 5. Calibrate the EnFET system with different ratios of R- and S-phenylalanine methyl ester from 0% S-ester up to 100% S-ester in steps of 10% S-ester. The total ester concentration should be about 1.5 g/L 6. Evaluate sensor signal vs S-ester concentrations (Fig. 8).

4. Notes 1. One can recognize damaged pH-FETs by high negative signals (higher than -3 V) or high positive signals (higher than +4 V) and by unstable signals (fluctuation higher than 3 mV/s). 2. For the immobilization of any other enzymes do the same as described for glucose oxidase. Dissolve the enzyme in an appropriate pH phosphate buffer depending on the pH optimum of the enzyme used. 3. It is very important to work as fast as possible, because the enzyme will unmobtlize very fast After 90 s the solution will be solid and no longer usable for the immobrlization. The munobrlization time depends also on the room temperature.

48

Ulber and Scheper

Best results will be obtained between 20 and 25’C. Examine the sensor under a microscope to make sure that the enzyme membrane is umform and has no defects. 4. The typical response time of a glucose oxtdase-EnFET IS about 3 mm. The response time depends on the thickness of the enzyme membrane. If the sensor signal is low (smaller than 30 mV at a glucose concentration of 5 g/L), repeat the immobilization step or try the nnmobilization on activated surfaces

References 1. Bergveld, P. (1970) Development of an ion-sensitive solid-state device for neurophysiologtcal measurements. IEEE Trans Eng BME 17,70. 2. Caras, S. and Janata, J. (1980) Field effect transistor sensitive to pemcdlm. Anaf Chem 52,1935-1937. 3. Tsukuda, K., Sebata, M., Miyahara, Y , and Miyaga, H. (1989) Long-hfe multiple-ISFETs with polymeric gates Sens Actuators 18,329 4 Perrot, H., Jaffrezic-Renault, N., Clechet, P., Wlodarski, W. B., deRooiJ, R F , and van den Vlekkert, H. H (1990) A generalized theory of an Ag+ sensitive electrolyte-msulator-semiconductor field-effect transistor with silica surface modified by chemical grafting. Sens Actuators Bl, 380 5. van den Vlekkert, H. H. and de ROOIJ, N. F. (1990) Multi sensing system basedon glass-encapsulated pH-ISFETs and a pseudo-REFET. Sens Actuators Bl, 395 6. Oesch, U., Caras, S., and Janata, J (1981) Field effect transistors sensitive to sodium and ammonmm ion Anal Chem 53, 1983 7. Vlasov, Y. G and Tarantov, Y A (1989) Development of ISFET using glassy solid electrolytes Chem Sen. Techn. 3, 173-178. 8 Battilottt, M., Mercuri, R., Mazzamurro, I , Gianmm, M , and Giongo, M (1990) Lead ion-sensitive membrane for ISFETs. Sens Actuators Bl, 438 9. Salardenne, J., Morcos, J , Ait Alial, M., and Portier, J. (1990) New ISFET sensitive membrane Sens. Actuators Bl, 385. 10. Nomura, T. and Nakagawa, G (1987) Alkah-free magnesium phoshate glasses as nitrate-ion selective materials for solid-state electrochemical sensors. BUZZ Chem Sot. Jpn. 57, 1491. 11 Wakida, S , Yamane, M , and Higashi, K. (1990) Urushi-matrix sodium, calcium, potassmm and chlortd-selektive field-effect transistors. Sens. Actuators Bl, 412.

12. Vlasov, Y., Hacklemen, D E , and Buck, R. P. (1979) Fabrication of a silver, chloride and bromide-responsive ion selective field effect potentiometric sensors Anal. Chem 51, 1570 13. Morttz, W , Meterhofer, I., and Mtller, L. (1988) Fluoride-sensitive membrane for ISFETs. Sens Actuators 15,2 11. 14 Bergveld, P. (1972) Development of an ion-sensittve solid-state device for neurophysrcal measurements. IEEE Trans. Blamed Eng. 19,342. 15 Colclaser, R. A (1980) Mxroelectronlcs Processing and Devzce Design, Wiley, New York.

ISFETs

49

16. Vlasov, Y. (199 1) Temperature coefficient of pH-sensitive ion-selective tieldeffect transistors. Mikrochzm. Acta 2, 363. 17. Bergveld, P and Sibbald, A. (1988) Analytical and Bzomedzcal Applications of ISFETs, Elsevier, Amsterdam. 18. Blackburn, G. F. (1987) BzosensorsFundamentals and Applications, Oxford University Press, New York. 19. Abe, H., Esashi, M., and Matsuo, M. (1979) A tantalum-on-sapphire microelectrode array. IEEE Trans. Electr. Dev. ED-26, 1939. 20. Esashi, M. and Matsuo, T. (1978) Integrated micro multi ion sensor using field effect of semiconductor. IEEE Trans. Biomed. Eng. BME-25, 184. 21, Vlaslov, Y, G. and Bratov, A. V. (1987) A pH-sensitive ion-selective field-effect transistor based on zirconium dtoxide film. Zh Przkl Khim 60, 755 22. Brand, U., Reinhardt, R., Ruther, F., Scheper, T , and Schugerl, K. (1990) Biofield-effect transistors as detectors m flow-injection analysis Anal Chzm Acta 238,201. 23. Anzai, J., Furuya, K., Chen, C., Osa, T., and Matsuo, T (1987) Penicillin sensors based on an ion-sensitive field-effect transistors coated with stearic acid Langmuir-Blodgett membrane. Chem. Pharm. Bull. 35(2), 693. 24 Brand, U , Scheper, T., and Schtigerl, K. (1989) Penicillin G sensor based on penicillin amidase coupled to a field effect transistors Anal. Chzm. Acta 226,87 25. Kullick, T., Ulber, R., Meyer, H. H., Scheper, T , and Schugerl, K. (1994) Biosensors based on enantioselectrve analysis. Anal Chzm Acta. 293,271 26. Kbneke, R., Menzel, C., Ulber, R., Saleemeiddin, M., and Scheper, T. (1996) Reversible coupling of glucoenzymes on fluoride-sensitive FET-biosensors based on lectin-glycoprotein binding. Biosens. Bioelectron. 12, 1229-1236. 27. Iida, T. and Kawabe, T. Eur. Pat. Appl. EP 257919 A2. 28. Kullick, T. (1994) Fortschrittberichte VD1, Reihe 8 Nr. 421, VDI-Verlag. 29. Kullick, T., Beyer, M., Henning, J., Lerch, T., Quack, R., Zeitz, I., Hitzmann, I., Scheper, T., and Schugerl, K. (1994) Apphcation of enzyme field-effect tranststor sensor arrays as detectors in a flow-injection system forsimultaneous momtoring of medium components. Part I. Preparation and calibration. Anal. Chim Acta 296,263-269. 30. Miyahara, Y., Matsu, F , Morhzumi, T., and Karube, I. (1983) Analytzcal Chemistzy Symposia Serzes, vol. 17, Elsevier, Amsterdam, pp. 501. 31. Gotoh, M., Tamiya, E , Karaube, I., and Kagawa, Y. (1987) A microsensor adenosine-5’-triphosphate pH-sensitive field effect transistors. Anal Chzm Acta 187, 287.

32. Tamiya, E., Seki, A., Karnbe, I., Gotoh, M., and Shimizu, I. (1988) Inosine sensor based on an amorphous silicon ISFET Anal Lett 21, 1785-1800 33. Dransfeld, I., Hintsche, R., Moritz, W., Pham, M. T , Hoffmann, W , and Hueller, J. (1990) Biosensors for glucose and lactate usmg fluoride ion sensitive field effect transistors. Anal Lett 23(2), 437. 34. Hartmeier, W. (1986) Immobzlisierte Biokatalysatoren, Springer Verlag, Berlm.

50

Ulber and Scheper

35 Anzai, J , Lee, S., and Osa, T (1989) Enzyme sensors based on an ISFET coated with Langmuir-Blodgett membranes. Use of polyethylenimme as a spacer for immobilizing a-chymotrypsin. Chem. Pharm. Bull 37(12), 3320 36. Gil, M. H., Ptedade, A. P., Alegret, S., Alonso, J., Martinez-Fabregas, E , and Orellana, A. (1992) Covalent binding of urease on ammonmm-selective potentiometrm membranes. Biosens Bioelectr. 7,645. 37. Shulga, A.A , Koudelka-Hep, M., and Rooij, N. F. (1994) Glucose-sensitive enzyme field effect transrstor using potassium ferricyanide as an oxidizing substrate. Anal Chem. 66 (2), 205-2 10.

5 Enzyme Biosensors

Based on Oxygen Detection

F. W. Scheller, D. Pfeiffer, F. Lisdat, C. Bauer, and N. Gajovic I. Introduction The electrochemical indication of oxygen consumption by enzyme-catalyzed analyte conversion goes back to the early 196Os, when L. Clark created the principle of enzyme electrodes. Since this time more than 30 different oxygen-consuming enzymes have been combined with the membrane-covered oxygen electrode. The experimental requirements of this combmation are quite simple. A normal Clark-type oxygen electrode loaded with an enzyme layer and an electronic amplifier allows for analytical applications (Fig. 1). However, the widespread application of enzyme electrodes based on oxygen measurement as compared with the electrochemical indication of hydrogen peroxide and redox mediators have been limited because of two major problems that influence the sensor performance at oxygen measurement. 1. Fluctuating oxygen concentrations in the measuring solution will contribute to the measuring signal. The flux of oxygen from the bulk phase to the electrode surface IS determined by all concentration gradients in front of the sensor. Therefore fluctuating oxygen concentrations m the bulk phase, e.g., dtfferent concentrations in real samples, like venous blood, will lead to erroneous results. To eliminate these disturbances, careful air saturation of all solutions at a defined temperature is necessary. The same effect will be achieved by mtroducing an aerator into a flow system or the conversion of the liquid in an aerosol. 2. The low equilibration concentration of oxygen restricts the measuring range. Most oxygenase-catalyzed reactions follow two substrate kinetics. Therefore, the oxygen concentratron will influence the reaction rate except when its concentration is considerably higher than its K, value or the concentration decrease during From

Methods m Bjotechnology, Vol 6 Enzyme and MIcrobra/ Bjosensors Technrques Edlted by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

51

and Protocols NJ

52

Scheller et al.

O-ring dialysis

membrane

enzyme membrane PET membrane membrane

holder

O2 - electrode

Fig. 1. Schematicview of a Clark-type electrode-basedenzyme sensorincluding membraneholder. the analyte conversion is negligible. The relatively low solubility of oxygen in aqueoussolutions, e.g., in air-saturated solutions at 210 pA4,allows only the determinationof micromolar concentrationsof the respectiveanalyte.Basedon a high permeability of oxygen in the carrier material usedfor enzymeimmobilization, the linear range of oxidase electrodeshas been extendedto the millimolar range.The cathodic oxygen reduction at the electrodethat is usedto generatethe measuring signal is a principal drawback. Even in the absenceof any substrate the oxygen concentrationdecreasesfrom the bulk-phasevalue to zero at the electrode surface.In the presenceof substratethe enzyme competesfor the oxygen with the indicator electrode. This processis responsiblefor the reduced linear concentrationcharacteristicsascomparedwith the H,O,-indication in which the oxygen is partly regenerated.

This chapter gives protocols for three different types of enzyme reactions that can be coupled to oxygen electrodesmay be distinguished. I. 1. Production of Hydrogen Peroxide by Oxygen-Reduction in Oxidase Catalyzed Reactions Electrochemical indication of oxidase-catalyzed reactions is possible by measuring either the consumption of oxygen at cathodic polarization or the

53

Oxygen Detection

formation of peroxide at positive potentials. Caused by the high over-potential of peroxide oxidation, reducing substances like ascorbic acid or paracetamol are cooxidized, thus generating an erroneous current signal. On the other hand the hydrophobic gas-permeable membrane applied for oxygen measurement prevents these interfering substances from reaching the indicator electrode. However, the determination of metabolites using oxygen detection IS not smted for biologic soluttons like whole blood, serum and urine because of then variable oxygen concentrations. This is especially important for blood glucose determinations, in which deoxyhemoglobin binds a part of the oxygen available in the sample. This results in falsely high glucose readings. Furthermore, possible microbial contamination may cause a reduced oxygen level of samples. A way to overcome such problems in clinical chemistry is the use of air saturation of the diluted whole blood and urine before reaching the measuring chamber and the electrode surface (1). 1.2. Oxygen Consumption by Oxygenases

(Without Peroxide Formation)

For oxygenase-catalyzed reactions or cytochrome-dependent oxidases, e.g., of the respiratory chain, measurement of oxygen consumption by using the Clark-type electrode is a well-established method both in research and in routine applications. 1.2.7. Sensitive Catecholamine

Detection Using Aerosol Technique

Determination of substancesbelow the micromolar concentration level, e.g., hormones or drugs, is accessible by substrate-recycling sensors (2). The catecholamines, epinephrine, norepinephrine, and dopamine, are of great interest because of their role as neurotransmitters and their dtagnostic value for cancer diseases.By combining an oxygen-consuming and a catecholamine (CA)-converting enzyme, such as, lactase, with an appropriate reducing enzyme (e.g., PQQ-dependent glucose dehydrogenase [GDH]), an ultrasensitive detection system can be obtained (3). catecholamine+ O2-> CA-quinone + glucose

lactase

glucose dehydrogenase

->

CA-quinone + H20

(1)

catecholamine+ gluconolactone (2) The high recycling efficiency of this system results in amplificatron factors up to 5000. To ensure this high amplification and a linear relationship between the sensor signal and the CA concentration, oxygen hmitatron has to be avoided. Because of the low concentration to be detected it is not suitable to dilute the samples drastically. Therefore the recycling sensor has been combined with an aerosol technique within a flowthrough system. The sample

54

Scheller et al.

solution is mixed with a larger volume of air m a mixing channel more than 1Ofold. Thus, the sample passesthe sensor as an aerosol. 1.2.2. Sensitive Measurement of Phenol or Alkaline Phosphatase by Measuring Oxygen in a Flow System Enzymatic oxygen consumption can be measured directly in a flow system. Tyrosinase consumes oxygen for the oxidation of phenol to qumone, which is reduced to the intermediate catechol in a coupled GDH-catalyzed reaction (Eq. 2). Alkaline phosphatase is measured after mcubation with the substrate phenylphosphate with the phenol-containing reaction mixture then analyzed as above (4). phenylphosphate

alkalme phosphatase

->

phenol + phosphate

(3)

The key factors that influence the signal and the baselme m such a flow system are analyte concentration, buffer composition, and its oxygen concentration, the flow rate, temperature and pressure. The use of an injector leads to a higher sample throughput and an Increased sensitivity of the system. 1.3. Oxidation of NADH and NADPH by Autoxidizable Mediators, NADH Oxidase, or Salicylate Hydroxylase Under Consumption

of Oxygen

To use the simple apparatus of oxygen detection also for nicotmamide

adenme dinucleotide phosphate (NAD[P]+) dependent dehydrogenases, enzyme-catalyzed oxidation of the reduced pyridme nucleotides has been coupled. Using this approach applicability of the oxygen electrode can be extended to the large field of dehydrogenase-catalyzed reactions including more than 200 enzymes of this type. Based on the Clark electrode, a novel enzyme sensor for L-malate has been developed. It comprises two enzymes: malic enzyme (L-malate dehydrogenase, oxaloacetate decarboxylatmg; MDH(dec)) and salicylate hydroxylase (SHL). The L-malate biosensor is used in a thermostatted and stirred cell. If the added sample volume is small compared with the cell content (< 1%), the temperature and the oxygen concentration of the mixture will not change noticeably, providing a stable baseline. Because L-malate occurs in high concentrations (>1c3 M) in all relevant specimens,it canbe easily determmed even if the sample is highly diluted. The reactions of the enzyme sensor are as follows: mahc enzyme

L-malate + NADP+ -> NADPH + salicylate + O2

pyruvate + CO2 + NADPH

sahcylate hydroxylase

-->

(4)

catechol + Hz0 + NADP+ (5)

The latter reaction is momtored with an oxygen electrode.

55

Oxygen Detection 2. Materials 2.1. Glucose in Prediluted

Whole Blood, Urine, and Serum

1. Basic buffer solution (for dilution and rmsing): 0.07 A4 phosphate buffer (Soerensen), pH 7.0. 2. Glucose stock solution: 100 n&f glucose in saturated benzoic acid solution. 3. Glucose calibration solutions: 3,6, 12, or 24 mA4glucose (diluted 1 to 50, e.g., 1 mL buffer + 20 yL calibration solution). 4. Measuring sample (diluted 1 to 50). 5. Glucose oxidase membrane: BST Bio Sensor Technologie GmbH, Berlin, Germany (5). Store the membrane at +4”C. 6. Polyethylene membrane, thickness = 15 urn (Metra Radebeul GmbH, Dresden, Germany). 7. Pt-Ag/AgCl oxygen electrode SM6 with platinum diameter 0.5 mm (Elbau GmbH, Berlin, Germany), KC1 electrolyte. 8. Pump I: minicassette pump MS-4 Reglo 8-100 sa (Ismatec, Glattbrugg, Zurich, Switzerland). 9. Pump II: homemade air pump. 10. Detecting device. any potentiostat available for nA currents.

2.2. Bienzymatic

Recycling

Sensors

2.2. I. Catecholamine Detection Using Aerator 2.2.1.1. BIENZYME ELECTRODE 1. Lactase from Coriolus hirsutus, E.C. 1.10.3.2. (Bach Institute of Biochemistry, Moscow, Russia; 320 U/mg, 140 mg/mL); glucose dehydrogenase from Acznetobacter cakoacetzcus (GDH, Boehringer Mannheim, Mannheim, Germany, 350 U/mg); pyrroloquinoline quinone (PQQ, Fluka, Neu-Ulm, Germany)* 0.5 mMin 0.1 MMES, pH 6.0; polyvinyl alcohol PVA 05/20 (Serva, Heidelberg, Germany). 2. Polyacrylate plates od 4 mm (to define the area of membranes to be prepared), W lamp 3. Polyethylene foil (Metra Radebeul, Dresden, Germany), cellulose dialysis membrane molecular weight cutoff (MWCO) 5 cm length) fixed as a loop m front of the flowthrough cell. 4. Homemade flowthrough cell consisting of a polyacrylamide block with a channel for aerosol flowthrough (id 88%; Fluka, Switzerland) stock solution 50 mM, dissolved in buffer 1. At 4’C stable at least 7 d 3 L-Malate (98%; Sigma) stock solution 50 mM, dissolved m buffer 1. At 4°C stable at least 3 d. 4. Hot gelatin solution 10% (w/v), in HzO: preswollen at room temperature for 20 mm, incubated in a water bath at 42°C for 60 mm 5. Enzymes: mahc enzyme (E C 1 1 1 40; suspension, 40 U/mL; Fluka), salicylate hydroxylase (lyophilisate, 3 U/mg; Sigma) 6. 1 MKCl solution. 7. Real samples: apple juice (100% fruit content), white wine, red wine. 8. Membranes: dtalysts membrane (regenerated cellulose membrane, approx 30 pm; Orwo GmbH, Wolfen, Germany), polyethylene membrane (approx 20 pm, Metra Radebeul) 9 Thermostatted homemade stirred cell (total volume, 5 cm3), with magnetic mtxer 10. Oxygen electrode SM6 (Elbau) 11 Amperometric potentiostat (EP 30, Biometra; Gottmgen, Germany) with chart recorder (Kipp & Zonen) or data-samplmg computer (Intel 386 DX 40) with homemade sampling software

3. Methods 3.1. Glucose Determination in Prediluted Whole Blood, Urine, and Serum 3.1.1. Preparation of the Glucose Electrode 1 Sandwich the glucose oxulase membrane between a polyethylene membrane and a cellulose dialysis membrane Stretch the sandwich over the membrane holder, with the polyethylene facing the electrode side, and fix it with an O-ring (Fig. 1) (Or use a commercialized measuring cell including membrane holder, BST). Fill the electrode cavity inside the membrane holder with KC1 electrolyte and mount tt onto the electrode

3. I .2. Measuring Procedure 1. Install the assembled glucose electrode in the flow system shown schematically m Fig. 2 It comprises a fluid (buffer and analyte) flow lme with a peristaltic pump and a gas (an) flow line, equipped with an air pump. The fluid stream (approx 1.5 mL/min) is pumped through a nozzle and combined with the air stream, which 1smore than 10 times the volume stream of the fluid. The resulting “spray“ of minute buffer droplets in air provides a fast saturation of the buffer/ analyte solution with oxygen, The air/fluid flow stream then passes a mixing channel and the flowthrough cell with the mounted glucose electrode. Connect a

Oxygen Detection

2 3 4. 5 6. 7. 8. 9.

59

potentiostat, poised to -600 mV, to the electrode to momtor the oxygen reduction current, Use a plotter/recorder or computer for data sampling (see Note 1) Equilibrate the measuring system with buffer (and an) for about 5 mm. Dilute the glucose stock solution to obtain 3,6, 12, and 24 mMglucose solutions. Dilute the 3,6, 12, and 24 mMglucose solutions and the sample 1 to 50 with the buffer solution. Check the concentration dependence with the diluted glucose solutions to control the linearity of the electrode. Use the 6 mM glucose solution, diluted 1 to 50, for a one-point calibration. Measure the sample, diluted 1 to 50, in the same way as the calibration solution. Calculate the glucose content of the sample (see Note 2). Store the enzyme electrode (mounted into the cell) at room temperature covered by a buffer segment.

3.2. Bienzymatic Recycling Sensors 3.2.1. Catecholamine Detection Using Aerator 3.2.1 .l. BIENZYME ELECTRODE PREPARATION 1, Prepare the PVA-solution by mcubatmg 0.2 g PVA with 1 mL water overmght. Heat the swollen polymer in a boilmg water bath for 5 mm and then cool down to room temperature. 2 Prepare the enzyme membrane by mixing 2.6 mg of the GDH with 30 PL of PQQ, allowing the holoenzyme to reconstitute for 5 mm. Add 20 uL lactase and 50 uL of the PVA solution and mix. Place 2 pL portions of this mixture on a polyacrylate plate and expose to UV irradiation (254 nm) for 30 min. Remove the membranes from the plates and store refrigerated until use (see Note 3). 3. Sandwich the enzyme membrane between a polyethylene membrane and a cellulose dialysis membrane. Stretch the sandwich over the membrane holder, with the polyethylene facmg the electrode side, and fix it with an O-ring (Fig. 1) Pipet KC1 electrolyte into the electrode cavity inside the membrane holder and mount it onto the electrode (see Note 4). 3.2.1.2. CATECHOLAMINE MEASURING PROCEDURE 1. Place the bienzyme electrodes mto the flowthrough cell and pump buffer 1 through the system for at least 30 min to equilibrate the membranes (approx 1 mL/mm). Connect the electrodes to the potentiostat (U = -0 6 V vs Ag/AgCl) 2. Introduce air into the system by turning on pump 2 (Fig. 2). (The ratio of an to buffer flow should exceed 10: 1) 3. Change buffer 1 to buffer 2; this serves as a test for the activity of the bienzyme electrode. It should result m a current decrease of 5-10 nA (with a basic current of around 80 nA). 4. ARer stabilizing of the base line introduce 3-5 mL of the catecholamine-containing solunon into the system by swltchmg from buffer 2 to sample. (Dilute unknown samples with buffer 2 to adjust pH and glucose concentration to at least 10 m Record the resultmg current decrease,1e., oxygen consumpnon, with the chart recorder (seeNotes 6-10).

Scheller et al.

60

3.2.2. Bienzyme Electrode for the Measurement of Phenol and Alkaline Phosphatase 3 2 2 1 MEMBRANE ELECTRODE PREPARATION 1. Prepare a solution of 0.2 g PVA m 1 mL water by incubating overnight at room temperature; then heat by placing in a boiling water bath for 5 mm and cool to room temperature again, 2. Prepare the bienzyme membrane by dissolving 2 0 mg GDH m 30 uL PQQ buffer After 5 mm add 1 0 mg of tyrosinase and then mix with 30 l.tL of 20% PVA solution. Spread 2 pL-portions of this mixture over an area of approx 4 mm2 on a polyacrylate plate and irradiate with UV (254 nm) for 30 mm. Remove the polymerized and dry membranes and store at 4°C. 3. On the day of use, sandwich the enzyme membrane between a polyethylene membrane and a cellulose dialysis membrane. Stretch the sandwich over the membrane holder, with the polyethylene facing the electrode side, and fix it with an O-ring (Fig. 1). Pipet KC1 electrolyte mto the electrode cavity inside the membrane holder and mount it onto the electrode and insert the latter into the flow system. 3.2.2.2.

FLOW SYSTEM

The flow system for mjection of phenol or the measurement of premcubated alkaline phosphatase(4) consistsof two parts (Fig. 3). In the first part (Fig. 3, left) two different buffer streams are mixed in a 1:1 ratio (100 uL/mm each). This changes the buffer conditions from substrate incubation (buffer 2) to detection conditions (buffer 1 + 2). In the secondpart of the system(Fig. 3, right) the buffer stream is thermostatted and the measurement of phenol with the bienzyme electrode is performed. The injector (100~uL sample loop) is placed in front of the mixer-pump 1. The pump rate of the secondpump is adjusted such that 90% of the volume streampassesthe flow cell and 10% goes directly to waste. This direct exit lme has stabilizing functions m the flow system.It comprises a 1.6~mmid elastic tubing, which scavenges an bubbles and damps the pump pulsation. The outlet (0.25~mm id tubing) serves as a flow restrictor (see Notes 11 and 12). 1. Start the system with buffer 1 and 2 (without phenylphosphate). Allow the current to stabilize for 30-60 mm (400 mV vs Ag/AgCl) It is good practice to check the oxygen sensitivity of the bienzyme electrode before measurement by a 30 s stop of the pump The optimal signals approach 80 nA/30 s. The noise of the baseline can be as low as 20 pA (see Note 13). 2. Test the system with injections of phenol standards (1.100 dilutions in buffer 2 from reservoir). 3. Add 100 pJ4 of substrate phenylphosphate to buffer 2 4. Dilute phenol standards or alkaline phosphatase standards to be measured m buffer 2 from reservoir and inject directly or after incubation. The lower detection limit for phenol is approx 20 IN and for alkaline phosphatase 3 fA4 after 1 h incubation (see Note 14).

61

Oxygen Detection

3.3. Ma/ate Sensor 3.3.1. Enzyme Membrane Preparation 1. Precipitate 0.125 mL malic enzyme (suspension in 2.9 M (NH&S04, 40 U/mL) by centrifugation at 3000g. Dissolve the precipitate in 30 pL buffer 1 and add 1 mg of salicylate hydroxylase (3 U/mg solid). Add 50 PL of the molten gelatin solution (at 42”C), mix thoroughly, and cast out the solution onto a Plexiglas support with a ptpet, to form a 1-cm2 area. Dry the membrane at room temperature for 6 h. Stored dry at 4°C the membrane is stable for at least 1 yr.

3.3.2. Preparation of the Enzyme Sensor 1 Sandwich a 2 x 2-mm piece of enzyme membrane between a dialyses and a polyethylene membrane and fix it with an O-ring on top of the oxygen electrode, which has previously been dipped in 1 MKCl solution (Fig. 1). The polyethylene membrane faces the electrode. Install the biosensor into the stirred cell (poised at 25°C) and connect it to the potentiostat. 2. Fill the cell with 2.5 mL of buffer 1 and equilibrate the enzyme sensor for at least 1 h. The base current should be constant (drift 103 This results in a very sensitive probing of glucose as is suitable to be used in a flow-inJection analysis system m which leaching of the mediator is faster compared with in a batch-type reactor. The paste electrode results m a diffusion-limited condition at the electrode solution interface that provides a wide linearity of the glucose sensor

4.2. Flow-Injection Analysis Glucose Biosensor Involving Mediator- and Peroxidase-Modified Graphite Paste 6. The amperometric response of the glucose biosensor involving mediator- and peroxidase-modified paste electrodes is based on the occurrence of the followmg coupled catalytic reactions: Glucose + O2 +

glucomc acid + H202

H202 + peroxidase (I) + HZ0 + peroxidase (II) Med (redj + peroxidase (II) + peroxidase (I) + Med (DXj

(4) (5) (6)

(7) where peroxidase (I) and peroxidase (II) are the reduced and oxidized forms of peroxidase. 7 While probing peroxidase-catalyzed reactions the TCNQ is held near its cathodic potential; thus TCNQ-’ is maintained for longer times during the electrochemical measurements and may be relatively more soluble in aqueous solutions Although TCNQ-mediated glucose biosensors provided more reproducible responses even when used in an FIA system (6) because of the high loading molar ratio of TCNQ/ peroxidase, i.e., > 103, TTF-mediated enzyme sensors provided better responses In this case TTFO is maintained for longer times while probing peroxidase-catalyzed reactions since TTFO is relatively less soluble m aqueous solutions under similar conditions of TCNQ-mediated enzyme sensor. On the other hand, when TTF is held at or near its anodic peak potential, e g., for the regeneration of gluMed

cox) + e- +

Med

(red)

90

Pandey

case oxidase, TTFf is maintained for longer times and is more soluble compared with TCNQ-’ in phosphate buffer. The relatively greater solubllity of TTF+ restricted the application of TTF m graphite paste to mediated electrochemtcal oxtdatton of the analytes. 8. When mediators are held for longer periods of time at or near their anodtc peak potential, there is a possibility of mediated electrochemical oxidation of other species present in the samples, e.g , ascorbic acid, that may severely affect the rehability of the enzyme-based biosensor for practical application depending on the concentratton of these interferences This problem can be overcome either by protecting the enzyme sensor with a suitable membrane restnctmg the transport of the interfermg analytes or by holding the mediator at or near its cathodic peak potential, thereby regenerating the reduced form of the analytes as m the case of peroxidase-catalyzed reaction. In this case peroxidase is regenerated by holding the mediator at or near its cathodic peak potential, which leads to relatively less sensrttvity of the mediator to oxidize the interfering analytes. Although, by holding the mediator a little away from the Em Oeither in the cathode or anodtc dnectron, the rano of the redox couple may have the finite value, however, tlus results m mamtarmng relanvely hrgh concentration of anions toward the cathodic side and cations toward the anodic side within the mobilized phase at the electrode/solutton interface. 9 Since most of the oxidase-catlyzed reactions under aerobic condmon generate peroxide, the peroxidase- and mediator-modified sensor could be used for onlme probing of several oxidase-catalyzed reactions. 10. Two ways can be adopted for immobihzing both oxidase and peroxidase together m a mediator-modified carbon paste: either both oxidase and peroxidase are incorporated into the graphite paste, or oxidase is nnmobrhzed m a column to give a bed for oxtdase-catalyzed reactions that produce peroxide under aerobic conditions, and peroxidase is incorporated within the carbon paste. Immobilization of both oxrdase and peroxrdase together with mediator into the carbon paste has two serious drawbacks to obtain the desired reaction step: occurrence of relatively high anaerobic condition within the paste leading to the restricted formation of peroxide within the paste, and mediated electrochemical oxldatlon of oxtdase. Accordingly, the second method, rmmobthzatton of oxrdase and peroxtdase separately, 1s desirable. This also helps m high loading of both oxidase and peroxrdase mto the immobihzed phase. Figure 6 shows the electrochemtcal regeneration of peroxrdase through TCNQ-mediated electrochemical reaction There is an increase in cathodic current on the addition of peroxide, showing thereby that TCNQ acts as an efficrent mediator for the regeneration of peroxrdase wrthm the carbon paste.

4.3. Mediator-, Dehydrogenase-, Graphite Paste

and NAD+-Modified

11. The amperometric response of a dehydrogenase-based enzymatic reaction is based on the mediated electrochemical oxldatron of NADH. The major problems associated with the development of these btosensors are:

Mediator-Modified

91

Carbon Paste Electrode

-200

0

500

mV vs ScE Fig 6. Cyclic voltammograms of TCNQ- and POD-modified graphite paste electrode in 0.1 Mphosphate buffer, pH 7.0, at the scan rate of 5 mV/s m; (A) absence and (B) presence of 5 mM H202. a. Operational stability of NADH within the carbon paste; b. Slower diffusion of NADH through and within the carbon paste; c. Effective coupling of mediator, dehydrogenase, NAD+, and NADH wrthm the carbon paste; and d. Unfavorable equilibrium toward the formation NADH of most of the dehydrogenase-catalyzed reversible reactions. One way to reduce problems (a-c) is to incorporate enzyme and NAD+ within the polyethylenimine matrix and to incorporate the modified polymer network into the carbon paste. This results m relatively hrgh stability of NAD+ within the carbon paste and efficient coupling between mediator, dehydrogenase, and NADH. 12. ADH-catalyzed reaction provides better amperometrrc response as compared with LDH-catalyzed reaction since in the former case the eqmhbrium reaction favors the formation of NADH. However, the effective couplmg between NADH and mediator leads to a raprd electrochemical reaction even to probe LDH-catalyzed reactions under similar conditions.

References 1. Hill, H. A. 0. and Sanghera, G. S. (1990) Mediated amperometric enzyme electrode, in Biosensors, A Practzcal Approach (Cass, A. E. G., ed.), Oxford University Press, New York, pp. 19-46. 2. Barlett, P. N. (1990) Conducting organic salt electrodes, in Biosensors, A Practzcal Approach (Cass, A. E. G., ed.), Oxford Universrty Press, New York, pp. 47-95.

92

Pandey

3 Heller, A. (1990) Electric wiring of redox enzyme Act Chem Res 23, 128-134. 4 Pandey, P. C , Tran-Minh, C., and Lantreibecq, F (I 99 1) Electrochemrcal studres on tetrathiafulvalenc-tetracyanoquinodimethane modified acetylcholine/choline sensor Appl Bzochem Bzotech. 31, 145-158 5. Pandey, P C , Kayastha, A. M., and Pandey, V. (1992) Amperometrrc enzyme sensor for glucose based on graphite paste-modified with tetracyanoquinodimethane. Appl Blochem Bzotech 33, 139-143 6 Pandey, P. C., Pandey, V , and Mehta, S. (1993) A glucose sensor based on graphite paste modified with tetracyanoquinodimethane. Indzan J, Chem 32A, 667-672 7 Pandey, P. C and Weetall, H. H. (1994) Application of photochemtcal reaction m electrochemrcal detection of DNA intercalation. Anal Chem 66, 12361241. 8 Albery, W J., Bartlett, P. N., and Cass, A E G. (1987) Amperometrrc enzyme electrodes Phil Trans R Sot Lond (Biol.) 316, 107-l 18 9. Pandey, P C. and Weetall, H H. (1995) Peroxrdase and tetracyanoqumodimethane modified graphite paste electrode for the measurement of glucose/lactate/glutamate using enzyme packed bed reactor. Anal Bzochem 224, 423428.

Enzyme Biosensors Based on Electron Transfer Between Electrode and Immobilized Peroxidases Lo Gorton, Elisabeth CsiSregi, Tautgirdas and Gyargy Marko-Varga

Ruzgas, lrina Gazaryan,

1. Introduction

1.1. Catalyfic Cycle and Physicochemical Properties of Peroxidases Peroxidases are widely spread in nature and are classified as oxidoreductases E.C.l. 11.1.X, where X is determined by a biologic reducer. Hemecontaining peroxidases are divided into two superfamilies, viz., plant and mammalian (see Table 1). The latter includes myeloperoxidase, lactoperoxidase, thyroid peroxidase, and prostaglandin H synthetase. The superfamily of plant peroxidases consists of yeast cytochrome c peroxidase (CCP), plant ascorbate peroxidases, fungal peroxrdases, and classic plant peroxrdases (1). Plant enzymes in general are more stable than others, and among them horseradish peroxrdase (HRP) is the most commonly used in practical analytical applications. The superfamily of plant peroxidases has been extensively studied for the last decade, and many of the enzymes have been crystallized (2-9). According to the crystal structures all plant enzymes are represented by a single polypeptide (about 300 amino acid residues) folded m 10 a-helices forming a twodomain structure with a noncovalently entrapped heme. The latter forms the active site of the enzymes and the amino acid sequences coordinating it are highly conserved through all peroxidases. Plant and fungal enzymes also contain two calcium ions and four disulfide bridges per enzyme molecule (2-7). The catalytic cycle (reactions la-c) of heme-containing peroxidases includes interaction of the native state of the enzyme with hydrogen peroxide, From

Methods m Brotechnology, Vol 6 Enzyme and MIcrobra/ Blosensors Techmques Edlted by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

93

and Protocols NJ

94

Got-ton et al

Table 1 Some Physicochemical

Properties

Orlgm of peroxldase Peroxldasea from horseradish root Fungal peroxldase from Arthromyces ramosus Chloroperoxldase from mold Caldarlomyces fumago Peroxldasefrom soybean Lactoperoxldasea from bovine milk Peroxldasefrom tobaccoleaves Cytochromec peroxldase from Saccharomyces cerewslae

of Heme-Containing

Peroxidases

Total mol wt &Da)

Isoelectric point

Carbohydrate content (%)

Ref.

42 41

3 9,8 8 34

18 5

132

19

131

41

>o

8, 9 6 35 5.2

8 >o 0

133,134 131,135 136 137

42 37 77 36 34 1

131

W~xture of lsoenzymes

resulting m the formation of an oxidized form of the enzyme usually denoted Compound I and the release of one water molecule. This initial reaction is then followed by two sequential one-electron reduction steps (reactions lb and lc) caused by reaction with an electron donor, AH2, in which an intermediate denoted Compound II is formed in the first of these two one-electron transfer steps and then finally the native state 1srecreated as a result of the second oneelectron step (10). Only iodide and hydrosulfite are known to behave as twoelectron donors (10) reducing Compound I in one step back into the native state. Native peroxidase (Fe3’) + H20, + Compound-I + Hz0 CompoundI + AH2 + Compound-II + AH*

(1’4

Compound II + AH2 + Native peroxidase (Fe3’) + AH* + Hz0

UC)

where AH* denotes the oxidized form of the electron donor AH*. Compound I contains two oxidizing equivalents: oxyferryl-heme and a free radical. In the case of CCP the free radical is localized on Trp- 19 1, whereas all fungal and plant peroxidases form a porphyrin n-cation radical (II). Oneelectron reduction of Compound I leads to Compound II with oxyferryl-heme. The reactivity of Compound II toward electron donors is usually much lower than that of Compound I (10). The only exception is the recently characterized tobacco anionic peroxidase (12). At both reduction steps electron and proton transfer occur simultaneously (13). If participation of distal residues (Arg-38 and His-42 for HRP) m the heterolytic cleavage of hydrogen peroxide is doubtless (2) for all peroxidases studied so far, the molecular determinants

Electron Transfer Between Peroxidases

95

of substrate specificity are stall unknown. By means of site-specific mutagenesis it has been demonstrated that Phe-41 in HRP controls access of the aromatic substrates to oxyferryl-heme (l&J@. Peroxidases oxidize a wide range of electron donors, such as amines and phenols. One-electron and proton transfer results in formation of substrate radicals (17), which could interact with each other formmg polymeric products or attack the enzyme itself causing its inactivation. Rapid inactivation of peroxidases in the reaction course is a well-known phenomenon and this is one of the main limitations of sensitivity m peroxidase-based assays. When reaction la proceeds on an electrode surface Compound I is reduced into ferriperoxidase by a heterogeneous electron transfer (ET) directly from the electrode material as well as by means of electron mediators. Both approaches result in a reduction current related to the concentration of hydrogen peroxide in the contacting solution. 7.2. Principles of Electrodes 1.2.1. Direct ET of Peroxidases The most simple electrode for the detection ofperoxide consists of a layer of peroxidase molecules adsorbed onto the electrode surface. If the electrode is placed into a sample and poised at a potential more negative than 0.6 V vs saturated calomel electrode (SE) then a proportionality between the registered reduction current and the peroxide concentration IS observed (mechanism presented in Fig. 1). Initially, this phenomena was discovered for HRP adsorbed on carbon black (18) and later it was demonstrated on carbon and graphite (19-3U), carbon fibers (311, gold (J&32), gold modified with viologens (33,34), and platinum (35-37) electrodes. Both iron (III) and iron (II) electrochemical transformation m the active center of HRP as well as mediatorless reduction of Compounds I and II, were found to be kinetically slow on the majority of the electrode materials (34,35,38). In general higher responses or currents for peroxide are observed at peroxidase-modified carbonaceous electrodes than on other electrode materials. It seemsas though a higher efficiency of the bioelectrocatalytic reduction of H,O, is found at electrodes rich in oxygen-containing surface functionalities, e.g., on electrochemrcally preoxidized (19), heat-treated graphite (23), and activated carbon (24). The same high responses are observed on edge-oriented pyrolytic graphite (319-41), also containing a high density of C-0 functionalities on the surface in contrast to basal-plane graphite having a low surface concentratron of C-O structures (#2). C-O functionalities, e.g., phenolic and quinoidal groups, are suspected to facilitate (mediate) ET between the carbon electrode material and peroxidases (19,23,24), In addition to HRP- and CCP-modified electrodes, bioelectrocatalytic reduction of H202 has been obtamed at graph-

96

Got-ton et al. CornRound

Electrode EappM.6 vs SCE

L

Ferrtperoxidase (Fe3-+)

Fig 1 Mechanism of direct bioelectrocatalytlc reduction of hydrogen peroxldase at peroxidase-modified electrodes. Pf is a cation radical localized on the porphyrin ring or the polypeptlde chain.

tte electrodes modified with lactoperoxidase (19,23,29), peroxidase from Arthromyces ramosus (ARP), also known as peroxidase from Coprznus cznereus (43,44), chloroperoxidase from Caldariomyces fumago (43, and soybean and tobacco peroxidases (46). Recently reversible two-electron electrochemical interconversion between resting CCP and Compound I was demonstrated on freshly polished pyrolytic graphite (47). 1.2.2. Mediated ET of Peroxidases To overcome slow heterogeneous ET of peroxidases, mediators, small redox-active molecules with inherent high heterogeneous ET rates, are frequently used to construct peroxidase electrodes (48). The reaction sequence of a mediated peroxidase-modified electrode is depicted in Fig. 2. It should be pointed out that both one-electron and two-electron mediators can be used. The one-electron donors used as mediators for bioelectrocatalytic reduction of peroxides with peroxidase-modified electrodes are ferrocenes (49-57), ‘hexacyanoferrate (II) (58-61), osmium complexed with pyridme, Os(bpy)z(py)C1 (62-64) and nickelocene (65). In these casesthe protons necessary for the peroxidase reaction (reaction lb-c) are consumed from the solution stmilarly as it happens for direct ET (Fig. 1). Exploitation of mediators of two-electron-proton donor type do not require additional protons for the peroxidase reaction cycle. The mediators of this type used for the construction of peroxtdase-modified electrodes are phenylenediamine (66,67), hydroqumone (40), catechol (68), 3-hydroxytyramine (dopamine) (691, methylene

Electron Transfer Between Peroxicfases ComDound-I (Fe4;-=O, P+)

97 Mmr(

Electrode

L

Ferriperoxidase (Fe3+)

Fig. 2. Mechanismof mediatedbioelectrocatalyticreduction of hydrogen peroxide at peroxidase-modified electrodes.M,, and M,, are the oxidized and reduced forms of mediator, respectively blue, or green and other dyes (70,71), orgamc metals (N-methylacridinium or N-methylphenazinium complexed with tetracyano-p-qumodimethane) (72,73), and tetrathiafulvalene (749. 1.3. Application of Peroxidase-Modified Electrodes Commonly HRP is used for peroxidase electrode construction, however, other peroxidases were also successfully used for electrode development (see Note 1). Peroxidase-modified electrodes have been introduced to monitor H,Oz, exploiting the advantage of operation of these electrodes at low potentials. The detection of H202 can be monitored at around 0 V vs SCE, where noise and electrochemical reactions with possibly interfering substrates are minimized. Later it has appeared that peroxidase electrodes can be additionally useful for the detection of different analytes, e.g., organic peroxides, aromatic amines, and phenolic compounds. These analytes are monitored by using electrodes modified with peroxidase alone. Additional analytes, such as glucose, L-lactate, amino acids, L-glutamate, and acetylcholme are detected by electrodes using a combination of peroxidase and additional enzyme(s). Currently, peroxidase electrodes allow H202 determination with detection limits (LDL) as low as 10 mJ4(40,50,62,64,73,75). The sensitivity of the best responding amperometric electrodes is close to 1 A/M/cm2 (23,56,62,64,76), which is about the maximal sensitivity limited by the masstransport of H202 at the electrodes (Table 2). It should be mentioned that a high reactivity of oxidized peroxidase (Compounds I and II) with a broad range of reductants, such

98

Got-ton et al.

Table 2 Configurations/Characteristics Electrode design

of Peroxidase-Modified

Condtttons of signal registration

SW~cd

Surface-modified peroxidase electrodes ARP, GR, CD1 couplmg Ss, 0 07V vs Ag/AgCl 11 HRP, heated GR, ads FI, 0 V vs Ag/AgCl 081 HRP, GR, CD1 couplmg Ss, 0 V vs Ag/AgCl 0 52 HRP, glassy carbon, ads FI, -0 3V vs AglAgCl, 0 47 mediator hydroqumone HRP, gold sols, ads Ss, 0 V vs Ag/AgCl 03 HRP, graphite, ads. FI, 0 V vs SCE 0 28 Lipophllrzed HRP, Ss, -0 05 V vs AglAgCl 0 23 TTF-TCNQ m paste, ads CCP, pyrolytic GR, ads CV,=031VvsSCE 0 15 HRP, tm(IV) oxide, Ss, 0 15 V vs AgiAgCl, 0 07 medtator ferrocene glutaraldehyde couplmg HRP, pyrolyttc GR, ads Ss, -0 01 V vs SCE 0 06 HRP, glassy carbon, ads Chronoamperometry, mediator: ferrocene Polymer-based peroxtdase electrodes HRP m Os(bpy)2+‘2+ Ss, 0 V vs SCE 0 75-l polymer, crosslmked HRP electropolymerlzed Ss, 0 15 V vs Ag/AgCl 0.01&O 14 m polypyrrole HRP electropolymertzed Ss, -0 1 V vs Ag/AgCl 0 073 m o-phenylenedtamme HRP m potentiostattcally Ss, -0 0 1 V vs Ag/AgCl 16 1O-6 deposited polypyrrole Bulk modified composite peroxtdase electrodes HRP, carbon paste Ss, 0 V vs AglAgCl, 0 85 medtator ferrocene HRP, graphite-silicone FI, -0.05 V vs AglAgCI 0 15 oil paste HRP, graphite/ paraffin FI, -0.05 V vs AglAgCl 0 07 oil paste HRP, graphite-epoxy Ss, -0.025 V vs AglAgCl 0 008 composite HRP, graphite-stlmone Ss, -0.025 V vs AglAgCl 0 004 grease paste HRP, graphite-epoxy FI, potentiometnc detection composite

Electrodes LDL (Ma 0 02 a0 2

LR (Pm

Ref 43 23 138

001

0 S-500 - - 30 0 03-l

0.5 001

- - 20 0.5-3 0 01-6

32 22 73

001

0 01-S

139 50

0 04

0 05-s

39 140

001

0 l-200

62,64,76

001

0 01-10

75

1

l-130

141

50-1750

39

0 04-10

56

- - 30

30,125

0 04

40

0 03

0.4

82,128

- - 20

142

0 5-20

142

0.75-50

143

(contmued)

Electron Transfer Between Peroxidases

99

Table 2 (continued) Peroxidase rich tissue-modified carbon paste electrodes Asparagus tissue Ss, 0 V vs SCE, mediator: ferrocene Tobacco callus tissue Ss, 0 V vs AglAgCl, mediator: ferrocene Horseradlsh root FI, -0.2 V vs Ag/AgCI, o-phenylenediamine Kohlrabi skin Ss, -0 2 V vs Ag/AgCl, mediator:ferrocene

0.066

04

0.4-60

129

0.023

0.8

5-l 10

144

03

0.3-120

67

8.4

8 4-170

130

aads= adsorptivemodification; S= sensitivity; LDL = the lower detectionlimit, LR = the linearrange; ss= steady-state,FI = flow-inJectIon,GR = graphite,CD1= carbodnmlde,CV = cychc voltammetry

as ascorbic acid, urea, phenols, and aromatic amines, opens the possibihty for their interference through the enzymatic reaction. In many of the papers it is stated that peroxidase electrodes are insensitive to different interfering substrates; however, this is probably more related to the construction of the electrodes restricting reactions of interfering reductants with the peroxidase. For any practical applications the peroxidase electrode of any configuration has to be tested for a whole variety of possible substrates in their relevant concentration range in real sample solutions (77). Peroxidase in general can react rapidly with alkyl peroxides (78); however, the enzyme seems to require a free hydroperoxy group (-OOH). Secondary and tertiary peroxides as well as endoperoxides have been tested but were unable to serve as peroxidase substrates (79). The organic peroxides that have been registered with peroxidase electrodes are presented in Table 3. The sensitivity trend (see Table 3, organized by placing the highest sensitivity first) was found to be similar for several electrode configurations (80), giving the highest sensitivity for hydrogen peroxide (see Table 2). However, a different sensitivity trend (cum01 hydroperoxide > 2-butanone peroxide > H202) was found for ARPmodified carbon fiber electrodes (81). Probably, the microenvironment of the enzyme, e.g., the immobilization procedure, can influence the activity of the peroxidase for different peroxides since it was demonstrated that modification of peroxidase with hydrophobic ferrocenes highly increased the activity for linoleic peroxide compared with the rate exhibited by native HRP (53). Similarly, a changed selectivity pattern was observed for ARP- and HRP-based electrodes when immobilized in organic media (carbon paste) (82). Organic peroxides are more soluble in organic solvents than in water. This reason caused the development of peroxidase electrodes (organic-phase peroxidase electrodes) for detection of peroxides in organic solutions or organic/ water mixtures. Strong encouragement for these developments was granted by

100 Table 3 Detection

Got-ton et al. of Organic

Peroxides

Peroxide 2-Butanone peroxide tert-Butyl peroxlbenzoate Cumol hydroperoxlde tert-Butyl peroxlacetate Trlphenylmethyl hydroperoxide Lmoleic peroxide tert-Amy1 perbenzoate tert-Butyl hydroperoxrde

Lauroyl peroxide

in Aqueous S(A/Mlcm2) 0.05a 0 00854 0.0070 0.007a 0.006a 0.006c 0.0027a 0.00150”

Solutions LDL (PM 3.1a, 6b

LR @JW 3. I-200”

3 *3@,20b 3oa

30-400a 5-1ooc

10a

O0

“Horseradtshperoxtdase-modified carbonpasteelectrodewith solublemediator(1 mM ophenylenedtamme) at anappliedpotentialof-O.2 V vs AgIAgCl, phosphate buffer, pH 7.4 (87) bCarbonfiber electrodemodified with peroxtdasefrom Arthromyces ramosus, working potential-0.05 V vs Ag/AgCl, mediator-free,phosphatebuffer solutton,pH 6 0 (82) CFerrocene-conJugated horseradish peroxldase-modified solid graphiteelectrode,working potential 0 V vs Ag/AgCl, phosphate buffer, pH 7.0 (53)

the fact that enzymes can function in nonaqueous media (83,84). It was demonstrated that the activity of HRP is a few orders of magnitude lower in orgamc solvents, e.g., ethyl acetate, compared with aqueous media (85). As a consequence, the response of peroxidase electrodes for H202 is decreased in partially organic media (80,82,86). Despite the decrease in sensitivity for H,02, extended linearity, enhanced stability, and sensitivity of peroxidase electrodes were found for the detection of organic hydroperoxides in organic solvent/ aqueous buffer mixtures (87,88). It has been shown that addition of 5-30% of aqueous buffer into the organic solvent is sufficient to obtain functioning peroxidase-modified electrodes (52,86,89,90). Detection of phenolic compounds is basedon the fact that phenols after their oxidation to phenoxy radicals (reaction la-c), can be re-reduced electrochemically (91). In this case the phenollc compound acts as a soluble mediator for the reduction of HzOz at the peroxidase modified electrode (Fig. 2). The electrodes responded to phenolic compounds such as, 2-amino-4-chlorophenol, phenol, catechol, resorcmol, p-cresol, 4-chlorophenol, 2,4-dichlorophenol,

4-chloro-3-methylphenol,and vanillin. The electrodecharacteristicsobtained for the phenolics with the most sensitive detection are presented m Table 4. It should be mentioned that peroxidase-modified electrodes have never been tested for mixtures of aromatic amines or phenohc compounds, which might appear in real samples. Rather complicated reaction kmetics have been demonstrated for the catalytic performance of peroxidase in solution containing more

Electron Transfer Between Peroxidases Table 4 The Characteristics of HRP-Modified Electrodes for the Detection of Aromatic Amines and Phenolic Aromatic amine and phenolic compound o-Dianisidine p-Ammophenol N-Acetyl-p-aminophenol Benzidme 2-Amino-4-chlorophenol 4-Chloro-3-methylphenol

p-Cresol Aniline

S(A/M/cm2) 14x9 6.1b 3.96b 0.970 0.085C 0.014c 0.0085C

701

Compounds LDL (WI 0.035a

0.02a 0.5c 2c 4c O.Old

LR W4 - - 0.5a

0.54 2-20” 4-35c 0.05-0.6d

aGlassy carbon electrode contammg HRP crosslinked with glutaraldehyde, 1 mMpotassmm hexacyanoferrate in phosphate buffer, pH 7.5 (58). bGraphlte electrode modified with tetracyanoquinodimethane and covalently nnmoblhzed HRP, 30 @4 H202 m phosphate buffer, pH 7.0 (238). “Graphite electrode with adsorbed HRP on the surface, applied potential -0 05 V vs SCE, 10 @4 H,O, in phosphate buffer, pH 7.0, FI (245). dCarbon paste electrode containing 5% of HRP, applied potential 0 1 V vs Ag/AgCl; 0 1 mM Hz02 m phosphate buffer, pH 8.0, FI (246).

than one reducmg substrate, e.g., several phenolic compounds (92-94). An alternative way would be to couple the electrode with a simple separation step using column liquid chromatography (95). A common feature of many oxidases is that they produce Hz02 on oxidation of substrates. Coupling of peroxldase-modified electrodes with different 0x1dases has been proposed to en-cumvent the problems associated with the high operating potenttal of direct electrochemical monitoring of H202. Kulys et al. (60) introduced the concept of oxidase/peroxidase bienzyme electrodes with a film containing HRP and glucose oxidase. In this electrode configuration, 0x1dation of glucose was catalyzed by glucose oxidase. Hydrogen peroxide produced in the first reaction oxidized HRP, which was re-reduced by using hexacyanoferrate (II) as a soluble mediator. The oxidized mediator in turn was reduced electrochemically m an amperometric assay.Later, different electrode configurations, e.g., mixing the oxidase and peroxidase in carbon paste (96) and wiring of peroxidase in combination with an oxidase (63,97,98) were used for the preparation of bienzyme and multienzyme electrodes. Peroxide detection in bienzyme or multienzyme electrodes was established using mediated (99) as well as direct (30) ET of peroxidases. The following oxidases have been coupled with amperometric peroxldase electrodes for detection of appropriate substrates: glucose oxidase (29,30,63,100,101), alcohol oxidase

102

Gorton et al.

(30,63,98,102-l OS),lactate oxidase (63,106-l 09), choline oxidase (63), n-amino acid oxidase (30,44,63), L-amino acid oxidase (30,110),xanthine oxidase (ill), uricase (111), cholesterol oxidase (99,112), putrescine oxidase (113), polyamine oxulase (114), bilirubin oxidase (115), glutamate oxidase (116), and pyranose oxidase (II 7). The idea of bienzyme/multienzyme sensorhas been realized with celland tissue-based electrodes, too, e.g., spinach tissue rich with glycolate oxidase and peroxidase (118). Sensors mvolvmg three or more consecutive enzymatic reactions have also been developed for detection of substratesdifficult to measure with single or bienzyme electrodes,e.g., acetylcholine (64). In conclusion, it can be stated that in general three prmcipally different constructions of peroxidase-modified electrodes can be distinguished: 1 Solid electrodes surface-modified with either adsorbed or covalently bound peroxidase, resulting in approximately one monolayer of peroxidase molecules on top of the electrode, 2. Solid electrodes surface-modified through coverage with a polymer, either redoxor electron-conductmg, into which peroxidase molecules are physically or chemttally attached (see Note 4), and 3. Bulk-modified compostte electrodes, where peroxtdase molecules are “homogeneously” distributed mto the electrode composed of a mixture of conductmg and msulatmg materials (see Notes 2 and 3). Recipes regarding preparatron of solid and composite electrodes developed in our group will be described m Subheading 3.

2. Materials 2.1. Solid Graphite Electrodes Modified with Adsorbed HRP for Detection of H202 or Phenols 1. Solid rod of spectrographic graphite (Ringsdorff Werke GmbH [Bonn-Bad Godesberg, Germany] type RWOOl, diameter 3.05 mm). 2 Fine emery paper for wet polishing (Tufbak Durite, Allar Co., Inc., Sterling Heights, MI, P 1200). 3. Horseradish peroxidase (E.C. 1.11.1.7, Sigma [St Louis, MO] type VI, obtained as a lyophilized powder, 250 U/mg sohd).

2.2, Solid Graphite Electrode Modified HRP for Detection of H202

with Covalent/y

Bound

1. Items l-3 as m Subheading 2.1. 2. Aqueous soluble carbodiimide (N-3-(dimethylammopropyl)-ll”-ethylcarbodiimide hydrochloride, Sigma).

2.3. Carbon Fiber Modified of Organic Hydroperoxides

with Adsorbed

1. Carbon fibers (Grafil HM-S/6K). 2 5-yL Glass capillary microcapsule (Drummond

HRP for Detection

Science Co., Broomell,

PA).

Electron Transfer Between Peroxidases 3 4. 5. 6.

103

Low viscosity, rapid glue (Hylo Gel, Swedish Techno Chemie, Vadstena, Sweden). Silver wire, 0.25~mm od (Alfa, Karlsruhe, Germany). Conducting carbon cement (Leit-C, Neubauer Chemikahen, Stuttgart, Germany). o-Xylene (Sigma).

2.4. Carbon Paste Electrodes Modified with HRP for Detection of H202 1. 2. 3. 4. 5. 6.

Graphite powder (Fluka, Buchs, Switzerland). Paraffin oil (Fluka, Buchs, Switzerland). Plastic syringe (ONCE/ASIK Denmark, working surface 0.053 cmm2). Silver wire, 0.25~mm od (Alfa, Karlsruhe, Germany). Horseradish peroxidase (E.C. 1.11.1.7, type VI, Sigma). Lactitol, a neutral polyol (Cortecs, Deeside, UK).

2.5. Mediatorless Coupled Oxidase-Peroxidase Carbon Paste Electrode for Detection of GIutamate 1. Items l-5 as in Subheading 2.4. 2 Polyethylemmme (50% aqueous solution, Sigma). 3. Glutamate oxidase (EC 1.4.3.11) from Streptomyces sp., 8 U/mg, Yamasa Shoyu Co. (Tokyo, Japan).

3. Methods 3.7. Preparation of So/id Graphite Electrodes Modified with Adsorbed HRP for Detection of H202 (steps 1-9) or Phenolic Compounds (steps l-10) 1 Polish the end of the solid graphite rod using wet emery paper to obtain a flat surface. 2. Wash thoroughly the flat circular end of the polished rod with deionized water. 3. Gently dry the surface with a fine filter paper followed by further drying in air. 4. Expose the graphite rod to 700°C for 1.5 min by placing it into a muffle furnace. 5. Store the electrode at ambient temperature in a desiccator until modification with enzyme. 6. Dissolve lyophilized powder of HRP in 0.1 Mphosphate buffer at pH 7.0 to get a final enzyme concentration of 20 mg/mL 7. Place 5 pL of HRP solutton on the electrode surface allowing the enzyme to adsorb for 5 min at room temperature. 8. Rinse the electrode thoroughly with deionized water. 9. Insert the peroxidase-modified electrode into the electrochemical cell containing buffer solution, apply 0 V vs SCE, and wait for the background current to reach a stable level. After that the electrode is ready for H,O, measurements, 10. Add H202 into buffer (final concentration of 50 ClM) for the determination of phenolic compound in a concentration range up to about 50 pM.

Got-ton et al.

104

3.2. Preparation with Covalent/y

of Solid Graphite Electrodes Modified Bound HRP for Detection of H202

1 Initially follow steps l-5 m Subheading 3.1. 2 Dissolve lyophilized powder of HRP in 0.1 Mphosphate buffer at pH 4.5 to get a final enzyme concentration of 20 mg/mL. 3 Dissolve carbodiimide into 0.1 M phosphate buffer at pH 5.0 to obtain a final concentration of 20 mg/mL 4. Immerse graphite electrode into carbodiimide solution, allowing the reaction to proceed for 0.5 h at room temperature, and wash with 0.1 Mphosphate buffer, pH 5.0. 5. Place 20 pL of HRP solution on carbodiimrde activated electrode surface, allowmg the mrmobilization reaction to proceed 0.5 h at room temperature. 6 Follow steps 8-9 m Subheading 3.1.

3.3. Preparation of Carbon Fiber Electrodes Modified with Adsorbed HRP for Detection of Organic Hydroperoxides 1 Insert a bundle of 20-30 carbon fibers mto a glass capillary microcapsule leaving about 4-5 mm of the fibers outside. 2 Apply a small droplet of glue at the end of glass capillary and let it rise taking care to not contaminate the fibers outside the capillary 3. Dissolve conductmg carbon cement m xylene to reduce its viscosity 4. Insert a silver wire from the other side of the captllary and apply a droplet of conducting cement to ensure contact of silver with the carbon fibers. 5. Cut the fibers at the electrode end, leaving 1.5 mm outside the tip. 6. Place the electrode mto 0.1 A4 phosphate buffer solution, pH 6.0, contammg 20 mg/mL HRP and allow adsorption of enzyme onto the carbon fibers for 10 min at room temperature. 7. Follow steps &!I m Subheading 3.1.

3.4. Preparation of Carbon Paste Electrodes with HRP for Detection of H202

Modified

1. Mix 100 mg graphite powder with 40 uL paraffin oil in mortar with pestle for 0 5 h 2 String silver wire around syringe plunder 3. Put paste from step 1 into syringe and push plunger leaving 3-4 mm at the tip of syringe free of paste. 4. Dissolve 2 mg HRP and 10 mg lactitol in 400 pL of 0.1 M phosphate buffer, pH 8.0, add 100 mg of graphite powder. 5. MIX all content from step 4 using a magnetic stirrer for 2 h at 4°C. 6. Dry the carbon powder from step 5 using a water vacuum pump for 4.5 h. 7. Place enzyme-lactitol-graphite powder into an agate mortar, add 40 pL paraffin oil, and mortar to form a homogeneous paste. 8. Fill remaining free space in the syringe tip with enzyme-modified paste (from step 7)

Electron Transfer Between Peroxiciases

105

9. Gently rub syringe tip on tine filter paper to obtain a flat surface of carbon paste electrode. 10. Insert electrode into electrochemical cell containing buffer solution, apply 0 V vs SCE, and wait for stable background current. After that, the electrode 1sready for the measurements.

3.5. Preparation of Mediator-less Coupled Oxidase-Peroxidase Carbon Paste Electrode for Detection of Glutamate 1 Heat graphite powder at 700°C for 15 s m a muffle furnace and cool it to amblent temperature m a desiccator. 2. Dissolve 1.O mg HRP, 1.O mg glutamate oxidase, and 200 pL 0.32% polyethylenimine in 200 pL of 0.1 Mphosphate buffer, pH 7.0. 3. Add 100 mg of pretreated graphite powder from step 1 and mix content for 2 h at 4’C using magnetic stirrer. 4. Dry composition for 3 h using a water vacuum pump. 5. Place modified powder into an agate mortar, add 40 pL paraffin oil, and mortar to form a homogenous paste 6. Follow steps g-10 from Subheading 3.4.

3.6. Sensor Evaluation In this section below the characterization of the electrochemical responses of some of the electrodes outlined under Subheadings 3.1.-3.5. are described. The figures (Figs. 3-7) present the electrochemical behavior of HRP-modified graphite electrodes (solid as well as carbon paste) for HzOz, cum01 hydroperoxide, and phenol, and for L-glutamate using a bienzyme electrode. Figure 3 shows cyclic voltammograms for a naked (Fig. 3A) as well as a spectrographic graphite electrode (Fig. 3B) modified with adsorbed HRP obtained in a stationary solution of 0.1 A4 phosphate buffer at pH 7.0 in the absence and presence of 4 rnM HzOz (22). The recordings were initiated at +0.6 V vs SCE and proceeded in the negative direction until 0 V where the scan direction was reversed. The scan rate was 0.05 V/s. For the naked graphite electrode the voltammograms appear virtually the same irrespective of the presence of H202, revealing the electroinactivity of H202 on naked graphite within the investigated potential range. For the HRP-modified electrode it 1svery clear that in the presence of Hz02 the reduction current drastically increases, reflecting the catalytic effect of adsorbed HRP on the electroreduction of H202, The mechanism of the catalytic reaction is outlined in Fig. 1. Figure 4 shows flow-injection peaks obtained for solid electrodes made of high-modulus type carbon fibers modified with adsorbed ARP for monitoring of cumol hydroperoxide (structure is given in the figure). The flow rate was

10 yllmin, the injection volume was 60 nL, andthe systemwas operatedat an applied potential of -0.05 V vs Ag/AgCl. The observed peak tailing was a

106

Gorton et al.

E/mV

vs

SCE

Fig. 3. Cychc voltammograms of a naked (A) and an HRP-modified (B) graphite electrodein 0.1 Mphosphate buffer, pH 6.0 (al andbJ and in the samebuffer but also contammg4 mMHzOz (a2and b2). The sweeprate was 50 mV/s. The plus (+) mdicates the start potential. consequence of the design of the flowthrough cell (81) yielding a high dispersion factor equal to 35, determined as the ratio of the current signals obtained for steady state and flow-injection measurements. Figure 5 presents calibration curves for H202 obtained wtth a carbon paste electrode modified with HRP and additionally stabilized with lactitol (82), revealing the influence of the composition of the carrier flow in the flow-injection system. The injection volume was 60 nL, and peaks were recorded with a flow rate of 10 pL/min at an applied potential of -0.05 V vs Ag/AgCl. Figure 6 shows calibration curves for (1) catechol and (2)p-cresol obtained for a carbon paste electrode modified with adsorbed HRP in the presence of 900 ).u’I~H202 m the carrrer flow (145). The results were obtained when the HRP-modified carbon paste electrode was inserted m an amperometric flow through cell connected to a single lme flow injection system. The sample was injected with a 50 pL injection loop and the carrier flow rate was 8 mL/mm.

107

Electron Transfer Between Pefcxidases

a 0 5 nA

-

I

0.1 nA

I l‘min

1 mM

500 PM

100 pM

60 p mol

3Opmol

6pmd

Fig. 4. Recorded flow-injection peaks with ARP-modified graphite fibers for detection of cum01 hydroperoxrde (structure also given m the figure)

The mechanism of the response to phenolic compounds is illustated m Fig. 2. The H202 in the carrier flow will keep the adsorbed enzyme in its oxidized state. A small background current will be registered as a result of the direct ET between the carbon particles in the paste and the oxidized HRP. When the injected sample is transported to the electrode surface the phenolic compounds will efficiently compete with the electrode as a source of electrons to the oxidized forms of HRP. As a result the phenols will be oxidized and form cation radicals, which in turn will be electrochemically re-reduced. This will be noticed as a drastically increased reduction current. The difference between this current and the background current will be a direct measure of the concentration of the phenolic compounds in the sample. Figure 7 presents calibrations curves for HzOz(-•-) and L-glutamate (-0-) for a mediatorless carbon paste electrode modified with HRP, L-glutamate oxidase, and polyethylenimine in log-log formats (116). The reaction sequence for the sensor is also shown for clarity. The sensitivity for H202 is higher than that for L-glutamate as expected for coupled enzyme systems.

Got-ton et al.

108

m

3 : w $0 2

2.5

:-

.

lfIO-

Ha0

l

- 10% 10% CH,CN CH,OH

. d

I

2:

1.5

:.

a

u

:: a

8

‘! %

m 1 n

0.5 -

I;*:

0 :

0

l

A

A

A

0

l A

,.........I..“‘.“,1

0 0

0.5

1

1.5

U-&O,1 / mM Fig. 5. Flow-injection calibration plots obtained for HzOz using mtcrocarbon paste electrodes based on HRP-modified and lactitol-stabrhzed graphite powder. Applied potential, -50 mV vs Ag/AgCl (0.01 M NaCl), mjection volume, 60 nL, flow rate, 10 uL min. The composition of the carrier is shown in the figure, where HZ0 indicates an aqueous solutton of 0.01 A4 phosphate buffer at pH 7.0. 4. Notes 1. Use of other peroxidases instead of HRP is limited for varrous reasons. CCP exhtbiting high electron exchange rates with edge-oriented graphite electrodes and mediators is not commercrally available. Additionally, because of insufficient stability already at room temperature, CCP has rarely been used for the preparation of practical sensors (5455,119). Other peroxtdases, such as, lactoperoxtdase and chloroperoxtdase, have not yet been shown to be more useful than HRP (19,23,&j. Exceptions could be the highly thermostable soybean peroxidase or peroxtdase from Arthromyces ramosus; however, electrodes modified with these peroxrdases are still scarcely studied (43,44,46,81,82,108,lI6,120) Non-heme peroxtdases having a totally different catalytic reaction cycle are scarcely reported for preparation of enzyme-modified electrodes though some data are available, e.g., regarding selenium-dependent glutathione peroxrdase (121-123). 2 Carbon paste electrodes give more freedom regarding enzyme modtficatron and addition of stabilizers and mediators as a result of the advantage of bulk moditication (96,124,125). Carbon paste electrodes as such are characterized by a lower background current than then solid electrode counterparts (126). However, pro-

Electron Transfer Between Peroxidases

109

0

Fig. 6. Flow-injection peak current dependence on concentration of (1) catechol and (2) p-cresol in the presence of 900 p.M Hz02 m phosphate buffer solutton obtained with HRP-modified carbon paste electrode at an applied potential of -50 mV vs AglAgCl. tem modificatton of the paste usually results in an increased background current, probably as a result of an increased wettability of the electrode surface (127). Almost all composite electrodes exhibit lower sensitivities when compared with surface-modified electrodes. The procedure of enzyme mcorporation into carbon paste is very important m the case when a mediatorless design is used. To avoid msulation of the enzyme from the carbon particles by the pasting 011,the peroxtdase has to be adsorbed on the carbon before the oil is added (30). Studies of different additives and graphite powders revealed that the sensitivity of medtatorless electrodes can be increased 10-17 times by some additives and proper choice of the source of graphite. The best additive so far was found to be lactitol (128). 3. Carbon paste electrodes modified with peroxtdase rich tissue have also been described (Table 2) pointing out a cheap source of the enzyme. It seems as though tissue-modified electrodes are intentionally based on mediated ET of peroxtdases even though less efficient mediatorless currents can be observed (129). The lower detection limits are restricted to higher values for these kmd of electrodes than those obtained for other types of peroxidase electrodes based on purified enzyme preparations. Reducing substrates such as, catechol, dopamine, ascorbic acid, and so forth, in concentrations higher than 10 pMhave been demonstrated to be mterfering for these electrodes (129,130). 4 The stability of peroxidase electrodes depends on the desrgn of the electrode. HRP has been found to be very stable in solution. However, peroxtdase-modtfled

Got-ton et al.

L-glutamate

a-ketoglutarate

5

4 2 2

3

B 2

1

1

0

log

2 conc./pM

3

4

Fig 7. Calibrattons curves (log-log plots) obtained for a glutamate oxidase and HRP-modified carbon paste electrode stabilized with polyethylenimine. Signs represent (-a-) HzOz and (-0-) L-glutamate peak currents. The carrier in the flow-injection systems consisted of 0.1 M phosphate buffer at pH 7 0 (flow rate; 0.7 mL/mm), the injection volume was 50 pL, and the system was operated at -50 mV vs Ag/AgCl electrodes, especially when based on adsorption of the enzyme, are usually characterized by a decreased operational stability compared with the enzyme m solution. Usually a better stability is obtained when the enzyme is covalently mnnobiltzed or entrapped into a polymer matrix.

Acknowledgment The authors thank the followmg organizations for financial support: the European

Community

EC No. EV5V-CT93-0354,

the Swedish

Board

of

Industrial and Technical Development (NUTEK), the Swedish Natural Science Research Council (NFR), and the Swedish Research Council for the Engineering Sciences (TFR). The Swedish Institute and the Swedish Royal Academy of Science are acknowledged by Tautgirdas Ruzgas and Irma Gazaryan for financial

support.

Electron Transfer Between Peroxidases

111

References 1. Welinder, K. G. (1992) Superfamily of plant, fungal and bacterial peroxtdases. Curr. Opin. Strut

Btol 2,388-393.

2. Finzel, B. C., Poulos, T. L., and Kraut, J. (1984) Crystal structure of yeast cytochrome c peroxidase refined at 1.7 A resolution. J. Btol. Chem 259, 13,027-13,036 3. Patterson, W. R. and Poulos, T. L. (1995) Crystal structure of recombinant pea cytosolic ascorbate peroxidase. Biochemistry 34,433 l-434 1. 4. Poulos, T. L., Edwards, S. L., Wariisht, H., and Go!d, M. H. (1993) Crystallographic refinement of lignin peroxidase at 2 A J Biol Chem. 268, 4429-4440.

5. Sundaramoorthy, M., Klshi, K , Gold, M. H., and Poulos, T. L (1994) The crystal structure of manganese peroxidase from Phanerochaete chrysosporium at 2.06 A resolution. J Brol. Chem. 269,32,759-32,767. 6. Petersen, J. F. W., Kadziola, A., and Larsen, S. (1994) Three-dimensional structure of a recombinant peroxrdase from Coprinus cmereus at 2.6 A resolution. FEBS Letts. 339,29 l-296 7. Kunishima, N., Fukuyama, K , Matsubara, H., Hatanaka, H., Shibano, Y., and Amachi, T. (1994) Crystal structure of the fungal peroxidase from Arthromyces ramosus at 1.9 A resolution. structural comparisons with the lignin and cytochrome c peroxidases. J MOE Biol 235,33 l-344. 8. Schuler, D. J., Ban, N., van Huystee, R. B., McPherson, A., and Poulos, T. L. (1996) The crystal structure of peanut peroxidase Structure 4,3 11-321 9 Henriksen, A., Svensson, L. A, Smith, A T., Burke, J. E., Thorneley, R N F , Welmder, K G., and Garhede, M (1993) Crystallographtc studies of peroxidases from horseradish and barley, in Plant Peroxtdases* Bcochemcstry and Physiology (Welinder, K. G., Rasmussen, S. K., Penel, C., and Greppin, H., eds.), Geneva University, Geneva, pp 5-8. 10. Dunford, H. B. (1991) Horseradish peroxidase: structure and kinetic properties, in Peroxtdases in Chemtstty and Btology (Everse, J., Everse, K. E., and Grisham, M. B., eds.), CRC, Boca Raton, FL, pp. l-23. 11. Erman, J. E., Vitello, L B., Mauro, J. M., and Kraut, J. (1989) Detection of an oxyferryl porphyrmp-cation-radical intermediate in the reaction between hydrogen peroxide and a mutant yeast cytochrome c peroxidase. evidence for tryptophan- 19 1 involvement in the radical site of Compound I. Bzochemistry 28,7992-7995 12. Gazaryan, I. G. and Lagrimim, L. M. (1996) Anionic tobacco peroxidase overexpressed in transgenic plants. I. Purificatton and unusual kinetic properties Phytochemistry 41,1029-l 034. 13. Dunford, H. B and Adeniran, A. J. (1986) Hammett correlation for reactions of horseradrsh peroxtdase Compound II with phenols. Arch Btochem Btophys. 251,536-542.

14. Newmyer, S. L. and Ortiz de Montellano, P. R. (1995) Horseradish peroxidase His420Ala, His420Val and Phe410Ala mutants. Histidme catalysis and control of substrate access to the heme iron, J. Bzol. Chem. 270, 19,430-19,438.

112

Got-ton et al.

15. Gazaryan, I. G., Doseeva, V. V., Galkin, A. G., and Tishkov, V. I. (1994) Effect of single-point mutations Phe41His and Phe143Glu on folding and catalytic properties of recombinant horseradish peroxidase expressed in E colt. FEBS Letts 354,248-250. 16. Smith, A. T., Sanders, S. A., Thorneley, R. N F., Burke, J F., and Bray, R. C (1992) Characterization of a haem active-site mutant of horseradish peroxidase, Phe4 lVa1, with altered reactivity towards hydrogen peroxide and reducing substrates. Eur J Btol. Chem. 207, 507-519. 17. Harvey, P. J. and Candelas, L. P. (1995) Radical cation cofactors m hgnm peroxidase catalysis. Biochem. Sot Transacttons 23,262-267 18 Yaropolov, A. I., Tarasevich, M. R., and Varfolomeev, S. D. (1978) Electrochemical properties of peroxidase. Bioelectrochem Bioenerg. 5, 18-24. 19 Razumas, V., Jasaitis, J., and Kulys, J. (1984) Electrocatalysls on enzyme-modlfied carbon materials. Bloelectrochem. Bzoenerg. 12,297-322. 20. Kulys, J. and Samalms, A. (1984) Dependence of the efficiency of bioelectrocatalytic processes on the electrode surface state. Bloelectrochem. Btoenerg 13,163-169 2 1, Bogdanovskaya, V. A , Tarasevich, M. R., Hmtsche, R., and Scheller, F (1988) Electrochemical transformations of proteins adsorbed at carbon electrodes. Bzoelectrochem. Bzoenerg. 19,581-584. 22. Jonsson, G. and Gorton, L (1989) An electrochemical sensor for hydrogen peroxide based on peroxide adsorbed on a spectrographic graphite electrode Electroanalyszs 1,465468. 23. Csoregl, E., Jonsson-Pettersson, G., and Gorton, L. (1993) Mediatorless electrocatalytic reduction of hydrogen peroxide at graphite electrodes chemically modlfied with peroxidases. J. Biotechnol 30,3 15-337. 24. Ho, W. O., Athey, D., McNeil, C. J., Hager, H J., Evans, G P , and Mullen, W. H (1993) Mediatorless horseradish peroxidase enzyme electrodes based on activated carbon. potential application to specific binding assay.J Electroanal. Chem 351,185-197 25. Bogdanovskaya, V. A., Khorozova, E., Vorob’ev, V. G., Tarasevlch, M. R , and Shterev, G I. (1990) Enzymatic and electrochemical reactions involving peroxldase. Elektrokhlmlya 26,573-579 26. Bogdanovskaya, V. A. (1993) Bloelectrocatalysls. problems and prospects Elektrokhzmlya 29,44 l-447 27. Bogdanovskaya, V. A., Fridman, V. A., Tarasevich, M. R., and Scheller, F (1994) Bloelectrocatalysis by lmmobllized peroxldase* the reaction mechanism and the possibility of electrochemical detection of both inhibitors and activators of enzyme. Anal Lett. 27,2823-2847. 28. Yaropolov, A, I., Cherednikova, T V., Em&us, J , Marko-Varga, G , and Gorton, L (1995) Investigation of the electrochemical properties of horseradish peroxldase adsorbed on solid graphite in the presence of monoclonal antibodies against the enzyme Bloelectrochem Bioenerg 38,40 1404 29. Gorton, L , Bremle, G , Csdregi, E., Jonsson-Pettersson, G., and Persson, B. (199 1) Amperometnc glucose sensors based on immobilized glucose-oxidlzmg enzymes and chemically modified electrodes Anal Chum Acta 249,43-54

Nectron Transfer Between Peroxidases

113

30. Gorton, L., Jonsson-Pettersson, G., Csoregi, E., Johansson, K., Dominguez, E., and Marko-Varga, G. (1992) Amperometric biosensor based on an apparent direct electron transfer between electrodes and immobilized peroxtdases. Analyst 117,1235-1241. 3 1. Csdregi, E., Gorton, L., and Marko-Varga, G. (1993) Carbon tibres as electrode materials for the construction of peroxidase-modrfied amperometric biosensors. Anal. Chim. Acta 273, S-70. 32. Zhao, J., Henkens, R. W., Stonehuemer, J , O’Daly, J P., and Crumbliss, A. L. (1992) Direct electron transfer at horseradish peroxidase-colloidal gold moditied electrodes. J. Electroanal Chem. 327, 109-l 19 33. Razumas, V., Gudavicius, A., and Kulys, J. (1983) Redox conversion of peroxidase on surface-modified gold electrode. J Electroanal Chem. 151,3 1 l-3 15 34. Razumas, V., Gudavicms, A., and Kulys, J. (1986) Kinetics of peroxidase redox conversion on vrologen-modified gold electrodes. J Electroanal. Chem. 198,8 l-87

35. Durliat, H., Courterx, A., and Comtat, M. (1989) Reactions of horseradish peroxidase on a platinum cathode. Bloelectrochem. Bioenerg. 22, 197-209 36. Adeyoju, O., Jwuoha, E. I., and Smyth, M. R. (1994) Amperometric determmation of butanone peroxide and hydroxylamine via direct electron transfer at a horseradish peroxtdase-modified platmum electrode. Anal Proc 31, 177-l 79 37. Comtat, M. and Durhat, H. (1994) Some examples of the use of thm layer spectroelectrochemistry in the study of electron transfer between metals and enzymes. Blosens. Bloelectron. 9,663-668. 38. Cosgrove, M., Moody, G. J., and Thomas, J. D. R. (1988) Chemtcally nnmobilrsed enzyme electrodes for hydrogen peroxide determination. Analyst 113,18 1 l-l 815. 39. Wollenberger, U., Bogdanovskaya, V., Bobrm, S , Scheller, F., and Tarasevich, M. (1990) Enzyme electrodes using bioelectrocatalytic reduction of hydrogen peroxide. Anal. Lett. 23, 1795-1808. 40. Dominguez-Sanchez, P , Tut&t-Blanco, P., Femandez-Alvarez, J M., Smyth, M R., and O’Kennedy, R. (1990) FlOW-inJeCtlOn analysis of hydrogen peroxide using a horseradish peroxidase-modified electrode detection system. Electroanalysts 2,303-308. 41. Armstrong, F. A., Bond, A. M., Buchi, F. N., Hamnett, A., Hill, H. A. O., Lannon, A. M., Lettington, 0. C., and Zoski, C. G. (1993) Electrocatalytrc reduction of hydrogen peroxide at a stationary pyrolytic graphite electrode surface in the presence of cytochrome c peroxidase: a description based on a microelectrode array model for adsorbed enzyme molecules. Analyst 118,973-978. 42. McCreery, R. L (199 1) Carbon electrodes: structural effects on electron transfer kinetrcs, in Electroanalytical Chemrstry (Bard, A. J., ed.), Marcel Dekker, New York, pp. 22 1-374. 43. Kulys, J. and S&mid, R. D. (1990) Mediatorless peroxtdase electrode and preparation of bienzyme sensors. Blelectrochem Bloenerg. 24305-3 11. 44 Johansson, E., Marko-Varga, G., and Gorton, L. (1993) Study of a reagent- and mediatorless biosensor for d-ammo acids based on co-immobihzed d-amino acid oxidase and peroxidase in carbon paste electrodes. J, Biomater Appl. 8, 146-173.

Got-ton et al.

114

45. Ruzgas, T., Gorton, L., Emneus, J., Csoregi, E , and Marko-Varga, G. (1995) Direct bioelectrocatalytic reduction of hydrogen peroxide at chloroperoxidase modified graphite electrode. Anal. Proc. 32,207,208. 46 Lindgren, A. (1995) Development of a peroxidase based biosensor for deterrmnation of phenolic compounds. Master Thesis, Lund University, Lund, Sweden. 47. Mondal, M. S., Fuller, H. A,, and Armstrong, F. A. (1996) Direct measurement of the reduction potential of catalytically active cytochrome c peroxidase compound I: voltammetric detection of a reversible, cooperative two-electron transfer reaction. J Am Chem Sot 118,263,264. 48. Bartlett, P. N., Tebbutt, P., and Whitaker, R G. (1991) Kinetic aspects of the use of modified electrodes and mediators in bioelectrochemistry. Prog. React Kinet. 16,55-155.

49 Epton, R., Hobson, M. E., and Marr, G. (1978) Oxidation of ferrocene and some substituted ferrocenes m the presence of horseradish peroxidase. J Organomet. Chem. 149,23 l-244. 50. Tatsuma, T , Okawa, Y., and Watanabe, T. (1989) Enzyme monolayer- and bilayer-modified tin oxide electrodes for the determination of hydrogen peroxide and glucose Anal Chem 61,2352-2355 51. Smit, M H. and Cass, A. E. G. (1990) Cyanide detection using a substrateregenerating, peroxidase-based biosensor. Anal Chem 62,2429-2436. 52. Schubert, F., Saini, S , Turner, A. P. F , and Scheller, F (1992) Organic phase enzyme electrodes for the determination of hydrogen peroxide and phenol. Sens Actuators B7,408-411. 53. Tsai, W.-C. and Cass, A. E. G. (1995) Ferrocene-modified horseradish peroxidase enzyme electrodes a kinetic study on reactions with peroxide and linoleic hydroperoxide. Analyst 120,2249-2254. 54. Cooper, J. M., Alvarez-Icaza, M., McNeil, C. J , and Bartlett, P. N. (1989) A kmetic study of an amperometric enzyme electrode based on immobihzed cytochrome c peroxidase J. Electroanal. Chem. 272,57-70 55. Cooper, J. M , Bannister, J. V., and McNeil, C. J (1991) A kinetic study of the catalysed oxidation of 1,3-dimethylferrocene ethylamine by cytochrome c peroxrdase. J Electroanal Chem 312, 155-163. 56. Dominguez-Sanchez, P., Miranda-Ordieres, A. J., Costa-Garcia, A., and TunonBlanco, P. (1991) Peroxidase-ferrocene modified carbon paste electrode as an amperometric sensor for the hydrogen peroxide assay.Electroanalysis 3,281-285. 57. Wang, J., Reviejo, A. J., and Angnes, L. (1993) Graphrte-Teflon enzyme electrode. Electroanalyszs 5,575-579. 58. Kulys, J. and Vidzmnaite, R (1983) The development of high sensitive enzyme electrodes for the determination of aromatic ammes Anal Lett 16, 197-207 59 Scott, D. L., Paddock, R. M , and Bowden, E. F (1992) The electrocatalytic enzyme function of adsorbed cytochrome c peroxidase on pyrolytic graphite. J Electroanal. Chem. 341,307-321 60. Kulys, J., Pesliaktene, M., and Samalms, A. (1981) The development of bienzyme glucose electrodes. Bzoelectrochem. Bzoenerg 8, 8 l-88

Electron Transfer Between Peroxidases

115

61. Wang, J. and Varughese, K. (1990) Polishable and robust biological electrode surfaces. Anal. Chem 62,3 18-320 62. Vreeke, M., Maidan, R., and Heller, A. (1992) Hydrogen peroxide and l3-mcotmamide adenine dmucleotide sensing amperometric electrodes based on electrical connection of horseradish peroxidase redox centers to electrodes through a three-dimensional electron relaying polymer network. Anal. Chem. 64,3084-3090. 63. Ohara, T. J., Vreeke, M. S., Battaglini, F., and Heller, A. (1993) Bienzyme sensors based on electrically wired peroxidase. Electroanalysrs $825-83 1. 64. Garguilo, M. G., Huynh, N., Proctor, A., and Michael, A C (1993) Amperometric sensors for peroxide, choline, and acetylcholine based on electron transfer between horseradish peroxidase and a redox polymer. Anal Chem 65, 523-528. 65. Liu, H., Qian, J., Liu, Y., Yu, T., and Deng, J. (1995) Nickelocene-mediating sensor for hydrogen peroxide based on bioelectrocatalytic reduction of hydrogen peroxide. Anal. Proc 32,475-477. 66 AdeyoJu, O., Iwuoha, E I , and Smyth, M. R. (1995) Kinetic characterization of the effects of organic solvents on the performance of a peroxidase-modified electrode in detecting peroxides, Thiourea and Ethylenethiourea. Electroanalyszs 7, 924-929. 67. Wang, J. and Lm, M. S. (1989) Horseradish-root-modified carbon paste bioelectrode. Electroanalysis 1,4348. 68. Pantano, P., Morton, T. H., and Kuhr, W. G. (1991) Enzyme-modified carbonfiber microelectrodes with millisecond response times J Am Chem Sot 113, 1832-1833. 69. Yang, X , Guilbault, G. G., and Suleiman, A. A. (1997) New dissolved mediator for immobilized horseradish peroxidase electrode. Talanta, m press 70. Lei, C. and Deng, J. (1996) Hydrogen peroxide sensor based on coimmobilized methylene green and horseradish peroxidase in the same montmorillonite-modified bovine serum albumin-glutaraldehyde matrix on a glassy carbon electrode surface. Anal. Chem. 68,33&I--3349. 71. Liu, Y., Liu, H., Qian, J., Deng, J., and Yu, T. (1995) Regenerated silk fibroin membrane as immobilization matrix for peroxidase and fabrication of a sensor for hydrogen peroxide utilizing methylene blue as electron shuttle. Anal Chum Acta. 316,65-72. 72. Kulys, J., Samalius, A., and Svirnnckas, G.-J. (1980) Electron exchange between the enzyme active center and organic metal. FEBS Lett. 114, 7-10. 73. Korell, U. and Spichiger, U. E. (1994) Novel membraneless amperometric peroxide biosensor based on a tetrathiafulvalene-p-tetracyanoquinodimethane electrode. Anal. Chem. 66,5 1O-5 15. 74. Bifnlco, L., Cammaroto, C., Newman, J. D., and Turner, A. P. F. (1994) TTFmodified biosensors for hydrogen peroxide. Anal. Lett. 27, 1443-1452. 75. Tatsuma, T., Gondaira, M., and Watanabe, T. (1992) Peroxidase-incorporated polypyrrole membrane electrodes. Anal. Chem. 64, 1183-l 187.

116

Got-ton et al.

76 Horrocks, B. R., Schmrdtke, D., Heller, A., and Bard, A. J. (1993) Scanning electrochemical microscopy. Enzyme ultramtcroelectrodes for the measurement of hydrogen peroxide at surfaces. Anal. Chem 65,3605-3614. 77 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 m liquid chromatography coupled to oxidase immobihzed enzyme reactors Anal Chem 67, 1326-133 1. 78 Anm, H and Yonetani, T. (1992) Mechanism of action of peroxidases, m Degradatzon of Enwonmental Pollutants by Mcroorganlsms and Thew Metaloenzymes (Siegel, H. and Siegel, A., eds.), Marcel Dekker, New York, pp. 219-241. 79. Marklund, S. (1971) Hydroxymethylhydroperoxide as inhibitor and peroxtdase substrate of horseradish peroxidase. Eur. J. Biochem. 21,348-354 80 Mulchandam, A., Wang, C -L., and Weetall, H H. (1995) Amperometric detection of peroxides with poly(amhnomethylferrocene)-modtfied enzyme electrodes Anal Chem 67,94-100. 81. Csoregi, E., Gorton, L., Marko-Varga, G., Tudos, A. J., and Kok, W. T. (1994) Peroxidase-modified carbon fiber mtcroelectrodes m flow-through detection of hydrogen peroxide and organic peroxides. Anal Chem. 66,3604-36 IO. 82. Popescu, I. C , Csoregi, E., and Gorton, L. (1996) Peroxtdase-modified carbon paste microelectrode as amperometric &detector for peroxides m partial aqueous media. Electroanalyszs 8, 10 14-l 0 19 83. Klibanov, A. M. (1989) Enzymatic catalysis in anhydrous organic solvents. Trends Blochem Scl 14, 141-144. 84. Wang, J. (1993) Orgamc-phase btosensors-new tools for flow analysis: a short review. Talanta 40, 1905-1909 85. Yang, L. and Murray, R. W. (1994) Spectrophotometric and electrochemical kinetic studies of poly(ethylene glycol)-modified horseradish peroxidase reactions in organic solvents and aqueous buffer. Anal. Chem 66, 2710-2718. 86 Wang, C.-L. and Mulchandam, A. (1995) Ferrocene-comugated polyamlmemodified enzyme electrodes for detection of peroxides in organic media. Anal Chem 67,1109-l 114. 87. Wang, J., Freiha, B., Naser, N., Gonzalez-Romero, E., Wollenberger, U., Ozsoz, M., and Evans, 0 (199 1) Amperometrtc biosensmg of organic peroxides with peroxidase-modified electrodes. Anal Chim Acta 254,8 l-88. 88. Wang, J., Wu, L -H , and Angnes, L. (1991) Organic-phase enzymatic assays with ultramtcroelectrodes. Anal Chem. 63,2993,2994. 89. Schubert, F., Saini, S , Turner, R S , and A P F (1991) Mediated amperometric enzyme electrode incorporating peroxidase for the determination of hydrogen peroxide in organic solvents. Anal. Chim. Acta 245, 133-138. 90. Lm, Y., Liu, H., Qian, J., Deng, J., and Yu, T. (1995) Feature of an amperometnc ferrocyanide-mediated H,O, sensor for organic-phase assay based on regenerated silk fibroin as nnmobihzation matrtx for peroxidase Electrochzm Acta 41,77-82

Electron Transfer Between Peroxidases

117

9 1. Ruzgas, T., Gorton, L., Emnkus, J., and Marko-Varga, G. (1995) Kinetic models of horseradish peroxldase action on a graphite electrode J Electroanal. Chem 391,41-49.

92. Gazaryan, I. G., Loginov, D. B., Lialulm, A. L , and Shekhovtsova, T. N. (1994) Determination of phenols using various peroxidases. Anal Lett 27, 2917-2930. 93. Courteix, A. and Bergel, A (1995) Horseradish peroxidase catalyzed hydroxylation of phenol: il. kinetic model. Enzyme Mlcrob Technol 17, 1094-l 100 94. Courteix, A. and Bergel, A. (1995) Horseradtsh peroxldase-catalyzed hydroxyiation of phenol: I. thermodynamic analysis. Enzyme Mlcrob Technol. 17,1087-l 093. 95. Marko-Varga, G., Em&us, J., Gorton, L , and Ruzgas, T. (1995) Development of enzyme-based amperometnc sensors for the determination of phenohc compounds. Trends Anal. Chem 14,319-328 96. Gorton, L. (1995) Carbon paste electrodes modified with enzymes, tissues, and cells. Electroanalysts 7, 23-45. 97. Maidan, R. and Heller, A. (1992) Elimmatlon of electrooxldizable interferantproduced currents in amperometric biosensors. Anal Chem 64,2889-2896 98. Vijayakumar, A. R , Csoregl, E., Heller, A., and Gorton, L. (1996) Development of an alcohol blosensor based on various coupled oxldase-peroxldase systems Anal. Chum. Acta 327,223-234.

99 Boguslavsky, L., Kalash, H., Xu, Z., Beckles, D., Geng, L., Skotheim, T , Laurmavicms, V., and Lee, H. S. (1995) Thin film blenzyme amperometric biosensors based on polymeric redox mediators with electrostatic bipolar protecting layer. Anal. Chum Acta 311, 15-2 1. 100. Marcmkeviciene, J. and Kulys, J. (1993) Bienzyme strip-type glucose sensor. Biosens. Bloelectron.

8,209--212.

101. Scheller, F. W., Schubert, F., Neumann, B., Pfelffer, D., Hmtsche, R., Dransfeld, I., Wollenberger, U., Renneberg, R., Warsinke, A., Johansson, G., Skoog, M., Yang, X., Bogdanovskaya, V., Buckmann, A., and Zaitsev, S. Y. (1991) Second generation blosensors. Blosens. Bloelectron. 6,245-253. 102. Buttler, T., Johansson, K., Gorton, L., and Marko-Varga, G. (1993) On-line fermentation process monitoring of carbohydrates and ethanol using tangential flow filtration and column liquid chromatography. Anal Chem. 65,2628-2636. 103. Johansson, K., Jonsson-Petterson, G., Gorton, L., Marko-Varga, G., and Csdregi, E. (1993) A reagentless amperometric biosensor for alcohol detection in column liquid chromatography based co-immobilized peroxldase and alcohol oxidase in carbon paste J Biotechnol. 31,301-3 16. 104. Marko-Varga, G., Johansson, K., and Gorton, L. (1994) Enzyme-based blosensor as a selective detection unit m column liquid chromatography. J. Chromatogr. A660,153-167.

105. Gibson, T. D., Hulbert, J. N., Parker, S. M., Woodward, J. R., and Higgins, I. J. (1992) Extended shelf life of enzyme-based biosensors using a novel stabllization system. Bzosens Bioelectron. 7,701-708.

118

Got-ton et al.

106. Spohn, U., Narasaiah, D., Gorton, L., and Pfeiffer, D. (1996) A btenzyme carbon paste electrode for the amperometric detection of l-lactate at low potentials. Anal. Chem. Acta 319,79-90

107 Spohn, U , Narasaiah, D., and Gorton, L. (1996) The influence of the carbon paste composition on the performance of an amperometrtc bienzyme sensor for L-lactate. Electroanalyszs 8, 507-5 14 108. Spohn, U., Narasaiah, D., and Gorton, L. (1997) Reagentless l-lactate sensors based on carbon paste electrodes modified with different lactate oxidases and peroxidases. J Prakt. Chem. 339,607-614. 109 Narasaiah, D., Spohn, U., and Gorton, L. (1996) Simultaneous determmation of D- and L-lactate by enzyme modified carbon paste electrodes. Anal Lett 29, 181-201. 110. Kacanikhc, V., Johansson, K., Marko-Varga, G., Gorton, L., Jonsson-Pettersson, G., and Csoregi, E (1994) Amperometric biosensors for detection of L- and Damino acids based on co-immobilized peroxidase and l- and d-amino acid oxidases m carbon paste electrodes. Electroanalysis 6,381-390. 111. Kulys, J., Laurmavicms, V., Pesliakiene, M , and Gureviciene, V (1983) The determmation of glucose, hypoxanthine and uric acid with use of bi-enzyme amperometrtc electrodes. Anal. Chum Acta 148, 13-18. 112. Crumbliss, A. L , Stonehuerner, J G , Henkens, R W , Zhoe, J , and O’Daly, J P. (1993) A carrageenan hydrogel stabilized colloidal gold multi-enzyme biosensor electrode utihzmg immobihzed horseradish peroxidase and cholesterol oxidase/cholesterol esterase to detect cholesterol m serum and whole blood. Biosens. Bloelectron. 8,33 l-337. 113. Yang, X. and Rechmtz, G. A. (1995) Duel enzyme amperometric biosensor for putrescine with interference suppression. Electroanalysis 7, 105-108. 114. Lm, M. S., Hare, M., and Rechnitz, G. A. (1992) Multienzyme contaming tissue-based and ferrocene-mediated bioelectrode for the determmation of polyamines Electroanalysu 4, 521-525. 115. Wang, J. and Ozsoz, M. (1990) A pohshable amperometric biosensor for bilirubin Electroanalysis 2,647-650 116 Ghobadi, S., Csoregi, E., Gorton, L., and Marko-Varga, G (1996) Carbon paste electrodes based on co-immobilized peroxidase and l-glutamate oxidase for determinatton of l-glutamate. Current Separatzons 14,94-102. 117. Liden, H., Buttler, T., Jeppsson, H., Volt, J., Marko-Varga, G., and Gorton, L. (1995) Two amperometric biosensors as liquid chmmatogmpmc detectors for on-line monitoring of carbohydrate consumption and ethanol production in bioprocesses.Proceedings of Transducer’s 9YEurosensors N, June 2529, Stockholm, Sweden, pp. 474477 118 Oungpipat, W. and Alexander, P W (1994) An amperometric bi-enzyme sensor for glycolic acid determination based on spinach tissue and ferrocene-medianon. Anal Chum Acta 295,37-46 119 Frew, J. E., Harmer, M A., Hill, H. A. O., and Labor, S. I. (1986) A method for estimation of hydrogen peroxtde based on mediated electron transfer reactions of peroxtdases at electrodes. J Electroanal Chem 201, I-10.

Electron Transfer Between Peroxidases

119

120. Vreeke, M. S., Yong, K. T., and Heller, A. (1995) A thermostable btosensor of hydrogen peroxtde. Anal. Chem 67,4247-4249. 121. Hua, C., Walsh, S., Smyth, R., Svancara, I., and Vytras, K. (1992) Voltammetric behavior of dihydronicotmamide adenine dmucleottde phosphate at enzymemodified electrodes Electroanalysis 4, 107-l 10 122. Hua, C , Smyth, M. R., and O’Fagain, C. (1991) Determination of glutathtone at enzyme-modified and unmodified glassy carbon electrodes. Analyst 116,929-93 1. 123. Wrmg, S A. and Hart, J. P. (1992) Chemically modified, screen-printed carbon electrodes. Analyst 117, 1281-1286. 124. Wang, J., Ciszewski, A , and Naser, N. (1992) Strtppmg measurements of hydrogen peroxide based on biocatalytic accumulatton at mediatorless peroxidaselcarbon paste electrodes. Electroanalysis 4,777-782 125. Gorton, L., Csoregi, E., Dominguez, E., Emneus, J., Jonsson-Pettersson, G., Marko-Varga, G., and Persson, B. (1991) Selective detection m flow analysts based on the combination of immobilized enzymes and chemically modified electrodes. Anal. Chum Acta 250,203-248. 126. Kalcher, K., Kauffmann, J.-M., Wang, J., Svancara, I., Vytras, K., Neuhold, C., and Yang, Z (1995) Sensors based on carbon paste in electrochemical analysts: a review with particular emphasis on the period 1990- 1993. Electroanalysrs 7,5--22. 127. Kulys, J., Gorton, L., Dominguez, E., Emneus, J., and Jarskog, H. (1994) Electrochemical characterization of carbon pastes modified with proteins and polycations. J. Electroanal. Chem. 372,49-55. 128. Popescu, I. C., Zetterberg, G., and Gorton, L. (1995) Influence of graphite powder, additives and enzyme immobilization procedures on a mediatorless hrpmodified carbon paste electrode for amperometric flow-injection detection of H202, Biosens Bioelectron. 10,443-46 1 129 Oungpipat, W., Alexander, P. W., and Southwell-Keely, P. (1995) A Reagentless amperometric biosensor for hydrogen peroxide determination based on asparagus tissue and ferrocene mediation. Anal Chim. Acta 309,35-45 130. Chen, L., Lm, M S., Hara, M., and Rechnitz, G A. (1991) Kohlrabi-based amperometric biosensor for hydrogen peroxide measurement. Anal Lett 24, 1-14. 13 1. Everse, J., Everse, K. E., Grisham, M. B. (eds.) (199 1) Peroxzduses in Chemutv and Biology, ~01s. 1 and 2, CRC, Boca Raton, FL. 132. Shinmen, Y., Asami, S., Amachi, T., Shimtzu, S., and Yamada, H. (1986) Crystallization and characterization of an extracellular fungal peroxidase. Agrzc Bzol. Chem. 50,247-249. 133. Sessa, D. J. and Anderson, R. L. (1981) Soybean peroxtdases: purification and some properties. J Agrzc. Food Chem 29,960--965. 134. Gillikin, J. W and Graham, J. S. (1991) Purification and development analysis of the maJor anionic peroxidase from the seed coat of glycme max. Plant Physlol 96,2 14-220. 135 Polis, D and Shmukler, H. W. (1953) Crystalline lactoperoxidase J Bzol Chem 201,475-500.

Got-ton et al.

120

136. Lagrimini, L. M., Burkhart, W , Moyer, M., and Rothstem, S (1987) Molecular cloning of complementary DNA encoding the lignin-forming peroxidase from tobacco molecular analysis and tissue-specific expression. Proc Natl. Acad Sci USA 84,7542-7546. 137 Takio, K , Titani, K , Ericsson, L. H., and Yonetani, T (1980) Primary structure of cytochrome c peroxidase The complete ammo acid sequence Arch Bzochem Bzophys 615,6 15-629 138. Kulys, J., Bilitewski, U , and Schmid, R. D. (1991) The kmetics of stmultaneous conversion of hydrogen peroxide and aromatic compounds at peroxidase electrodes Bloelectrochem Bzoenerg 26,277-286. Armstrong, F A and Lannon, A M (1987) Fast mterfacial electron transfer 139 between cytochrome c peroxtdase and graphite electrodes promoted by aminoglycostdes: novel electroenzymtc catalysis of H,O, reduction J Am Chem Sot 109,7211,7212 140 Vidal, J. C., Yague, M. A., and Castillo, J. R. (1994) A chronoamperometric sensor for hydrogen peroxide based on electron transfer between immobilized horseradish peroxidase on a glassy carbon electrode and a diffusmg ferrocene mediator Sens Actuators B 21, 135-141 141 Deng, Q and Dong, S. (1994) Mediatorless hydrogen peroxide electrode based on horseradish peroxidase entrapped m poly(o-phenylenediamme) J Electroanal. Chem. 377, 191-195. 142 Wollenberger, U., Wang, J., Ozsoz, M., Gonzalez-Romero, E., and Scheller, F (199 1) Bulk modified enzyme electrodes for reagentless detection of peroxides Bloelectrochem. Bloenerg 26,287-296. 143. Zulfikar, Hibbert, D B , and Alexander, P. W (1995) A tubular graphite-epoxy electrode incorporatmg horseradish peroxidase as a potentiometric sensor for hydrogen peroxide. Electroanalyszs 7,722,725 144. Navaratne, A. and Rechnitz, G A. (1992) Improved plant tissue-based biosensor using m vitro cultured tobacco callus tissue. Anal Chum Acta 257,59-66 145. Ruzgas, T., Emntus, J., Gorton, L , and Marko-Varga, G. (1995) The development of a peroxtdase biosensor for monitoring phenol and related aromatic compounds. Anal Chum. Acta 311,245-253. 146. Dommguez-Sanchez, P., O’Sullivan, C. K , Miranda-Ordieres, A J , TunonBlanco, P., and Smyth, M R (1994) Flow InJection amperometric detection of aniline with a peroxidase modified carbon paste electrode. Anal. Chim. Acta 291,349-356

Enzyme Biosensors

Based on Redox Polymers

Latha Shankar, Michael G. Garguilo, and Adrian C. Michael 1. Introduction Enzymes can be immobilized near electrode surfaces by trapping them m a crosslinked polymer. Gregg and Heller (I) introduced an mteresting extension of this technique by incorporating the polymer into the sensortransduction mechanism. The strategy was to attach redox complexes to the polymer backbone and use the resultant redox polymer to mediate electron transfer between the immobilized enzymes and the substrate electrode. The principles behind the operation of these enzyme electrodes are essentrally identical to those behind sensors that use soluble mediators. The main difference IS that the redox functtonallttes pendant to the crosslinked polymer cannot participate in molecular diffusion, Rather, the electron transport process occurs by a series of self-exchange reactions between neighboring redox centers, a process referred to as electron hopping (2). The polymeric mediator offers several important features. Perhaps the most important is the opportunity to construct very small self-contained sensors. Tightly binding the mediator and the enzymes to the electrode eliminates the need for a dialysis membrane to trap the enzymes and eliminates the need to add soluble mediator to the sample. These features have been invaluable to our work on microsensors for measurements m the brain tissue of living animals (3), in which rt is vital to make the sensors as small as possible and it is not feasible to add reagents to the sample. Several factors impinge on the choice of redox complex: it must be possible to attach the complex to a crosslinkable polymer; the complex must From- Methods m Biotechnology, Vol. 6 Enzyme and Wcrobral Blosensors Technrques Edlted by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

121

and Protocols NJ

122

Shankar, Garguilo, and Michael redox polymer

cross linker

OP

(W)2cI

SchemeI electrode

SchemeII. readily participate in electron self-exchange as well as crossexchange with enzymes and electrodes; the complex must be chemically stable in two oxidation states; and it must possess an appropriate half-wave potential. An osmium-centered polypyridyl complex meets these criteria. Scheme I reports the structure of the crosslinkable redox polymer, the preparatton and use of which IS described in detail in this chapter (in Scheme 1, bpy is 2,2’bipyridine). A poly(viny1 pyridine) backbone is derivatrzed with the osmium-centered redox complex and ethylamme side chains that, m conJunction with the diepoxide also in Scheme I, provides the crosslinking. The redox polymer and the diepoxide are water soluble and, therefore, compatible with aqueous enzyme solutions. Heller’s group has used this, and related, polymeric mediators to construct sensors that mcorporate a variety of oxidase enzymes (4). Our own work has focused on microsensors for the measurement of choline in the extracellular fluid of the brain tissue in living animals. This demands an extremely small device, so the substrate electrode used is the end of a single carbon fiber with a 1O-pm diameter. Although a variety of oxidase enzymes participate in electron transfer with the polymeric mediator, choline oxidase (ChOx), which exhibits high cosubstrate specificity, appears to be an exception. The choline sensors, therefore, operate according to the dual-enzyme mechanism depicted m Scheme II (5). Immobilized choline oxidase generates hydrogen peroxide,

723

Redox Polymers

which is reduced by horseradish peroxidase (HRP). The polymer then medrates the electroreduction of HRP. Because the polymer mediates the reduction of HRP, rather than the OXIdation of an oxidase, the sensor is operated at potentials that are negative with respect to the half-wave potential of the mediator, approx 0.3 V vs saturated calomel electrode (SCE). Operating the sensor at negative potentials prevents the detection of easily oxidizable compounds present in brain fluid, such as ascorbate (vitamin C). Although the low potential prevents oxidation of ascorbate at the electrode, ascorbate can still interfere with the transduction mechanism by reducing the mediator, the HRP, or the peroxide. Two steps are taken to prevent this interference. First, ascorbate OXIdase is coimmobilized in the crosslinked polymer. Because ascorbate oxidase produces water rather than peroxide, it consumes ascorbate without contributing to the amperometric signal. Second, a layer of Nafion is cast over the crosslinked polymer layer. Nafion is permselective against the ascorbate anion. The resulting sensors can be used to quantitate micromolar concentrations

of choline

in the presence of 1OO-fold higher concentra-

tions of ascorbate. We have recently described in vivo experiments carried out with these microsensors (3). 2. Materials 2.1. Synthesis of the Crosslinkable

Redox Polymer

The following solvents are needed: acetonitrile, concentrated HCl, diethyl ether, N,N’-dimethyl formamide (DMF), ethanol, ethylene glycol, methanol, and ultrapure water. 2.7.7. Synthesis of [Os(bpy)&I$I, Where bpy = 2,2’-Bipyridine (6)

- 2H,O,

1. 1.9 g potassium osmium (IV) hexachloride K2[OsC16] (Alfa, Johnson Mathey, Ward Hill, MA). 2. 1.3 g 2,2’-bipyridine

(Sigma, St. Louis, MO).

2.7.2. Synthesis of Os(bpy)&I,

(6)

1. 1.O g [Os(bpy),C12]C1.2H,0, prepared according to Subheading 3.1.1. 2 2.0 g sodium drthionite, Na2S203, in 200 mL ultrapure water (Aldrich, Milwaukee, WI).

2.7.3. Derivitization of Po/y(viny/ Pyridine) (7) 1. 0.494 g Os(bpy)& prepared according to Subheading 3.1.2. 2. 0 430 g poly(4-vinyl pyridine) (Polysciences, Warrington, PA). 3 1.5 g 2-bromoethylamine hydrobromide.

124

Shankar, Garguilo, and Michael

4. 2 g ammonium hexafluorophosphate (NH,PF,) (Aldrich) 5. 5.2 g anion exchange beads (Bio-Rad [Richmond, CA] AG 1X4, chloride form).

2.2. Fabrication of Substrate Electrodes 2.2. I. Glassy Carbon Electrodes 1. Glassy carbon rod, 3-mm diameter by l-cm length (e g., type V-10 from Atomergic Chemicals, Farmmgdale, NY). 2. Brass rod, 3-mm m diameter by 7-cm length 3. A cylmdrtcal block of Teflon, l-cm diameter by 7-cm length 4 Napped pohshmg cloth and deagglomerated alumma mlcropohsh with particle sizes of 5.0, 1.O, and 0 3 pm (Buehler, Lake Bluff, IL).

2.2.2. Carbon Fiber Microelectrodes I lo-pm-diameter carbon fibers (P-55, Union Carbide, New York, NY) 2 Spurr low-viscosity epoxy: mix thoroughly vmyl cyclohexene dioxide (1 g), DER resin 736 (0 6 g), and nonenyl succmic anhydride (2.6 g). Just before use, add diammomethanol (150 uL) and mix thoroughly again Cure overnight at 70°C. This epoxy is purchased as a kit from Polysciences 3. Mercury 4 Pyrex tubmg, 2-mm od 5 Fused silica tubmg (0 25-mm id) (Supelco, Bellefonte, PA) 6. Micropipet puller (e.g., Narishige [Tokyo, Japan] Model PB-7).

2.3. Electrode Modification 2.3. I. Deposition of Redox- Polymer and Enzymes 1 2 3. 4 5. 6. 7.

Nanopure water (Barnstead Nanopure detomzation system, Dubuque, IA) Crosslinkable redox polymer, prepared accordmg to Subheading 3.1. Choline oxidase, from Alcaltgenes (Sigma). Horseradish peroxidase type II (Sigma) Ascorbate oxidase, from Cucurbitas (Sigma) Choline chloride (Aldrich). Phosphate-buffered saline (PBS): 150 mM NaCI, 100 mM Na2HP0,, adjust pH to 7.4 with NaH,PO, (J. T. Baker, Phillipsburg, NJ). 8. 4-(2-hydroxyethyl)-l-piperazine-ethanesulfonic) (HEPES) acid and the sodium salt (Sigma). 9. HEPES buffer, pH 8.0. titrate a 10 mM solution of the acid with the sodium salt. 10. Poly(ethylene glycol400 diglycidyl ether) (Polysciences).

2.3.2. Deposition of Nafion 1 Nafion perfluorinated ion-exchange powder, 5 wt% solution m mixture of lower aliphatic alcohols and 10% water. (Aldrich). 2. 2-Propanol as solvent.

Redox Polymers

125

3. Methods 3.1. Synthesis of the Crosslinkable Redox Polymer 3.1.7. Synthesis of [Os(bpylzCI&I~2H~0 (6) 1 Mix K,[OsCl,] (1 9 g) and 2-2’-brpyridine (1.3 g) m DMF (40 mL) in a 100~mL round-bottom flask fitted with a reflux condenser. 2. Reflux for 1 h with stnring. The solution darkens in about 15 mm. 3. Allow to cool to room temperature (-2 h). Solid KC1 forms in the dark solution 4. Filter through a sintered glass funnel. The dark red filtrate IS used for the following steps. Discard the solid KCl. 5 Transfer the filtrate to a 1-L beaker and add ethanol (20 mL) 6. Slowly add diethyl ether (500 mL) The product forms a dark red precrprtate. 7 Collect the red solid, [Os(bpy)2C12]Cl.2H20, and dry in air

3.1.2. Synthesis of Os(bpy)&I,

(6)

1. Dissolve [Os(bpy),Cl,]Cl*2H,O (1 g) m 20 mL of DMF and 10 mL methanol m a 500-mL beaker. 2. While stirring, add the dithionite solution dropwtse over a period of 45 min The color of the solution changes from dark red to dark purple. 3. Cool the purple solutron in an ice bath for 0 5 h and scratch the sides of the beaker with a glass rod to obtain a dark purple precipitate. 4. Collect the solid Os(bpy),C& and store desmcated under nitrogen 5. Characterize a sample of the product dissolved in water by UV-VIS spectrophotometry. Peaks with molar absorptivities near 10,000 will be noted with h,,, values of 383,466, and 558 nm. A broad charge transfer band will be noted with h,,, near 840 nm 6. Characterize a sample of the product dissolved in acetonitrile containing 1 M tetraethyl ammonium perchlorate by cyclic voltammetry. A redox couple with a half-wave potentral near -100 mV vs SCE will be observed. Evidence of a second redox couple with a half-wave potential near +800 mV vs SCE indicates formation of the trzs-bpy complex of osmium, which is not a problem unless it is a major component of the sample.

3.7.3. Derivatization

of Po/y(viny/ Pyridine) (7)

1. Attach a reflux condenser to a lOO-mL botlmg flask; purge with N2 Add 0.494 g Os(bpy)&&, 0.430 g poly(Cviny1 pyridme), and 18 mL ethylene glycol to the flask Reflux under N2 for 2.5 h 2. Allow the solution to cool to room temperature. 3. Add 30 mL DMF and 1 5 g bromoethylamine hydrobromtde and stir overnight at 45°C 4. Pour the dark solution into rapidly stirred acetone. Collect the dark solid that precipitates, which is the chloride form of the crosslinkable redox polymer 5. Dissolve the solid in -25 mL water and filter. To the filtrate, add NH4PF6 dissolved in -25 mL water Allow to stand for 0.5 h.

Shankar, Garguilo, and Michael

126

6 Collect the solid, which is the hexaflurophosphate form of the polymer, and store m a desiccated chamber. 7. Dissolve the solid m 20 mL acetonitrile and 50 mL water. Stir over anion exchange beads for 3 h. Filter and evaporate the filtrate to 10 mL. 8. Freeze-dry the aqueous filtrate to obtain dry redox polymer as a dark brown solid. 9. Store the product m an evacuated container and keep it away from hght

3.2. Fabrication of Substrate Electrodes 3.2.7. Glassy Carbon Electrodes Although glassy carbon electrodes are commercially available from a vanety of sources, we use electrodes constructed in-house according to the followmg procedure. These electrodes are economical and quite easy to make, especially if the assistance of a machinist is available. 1 Drill a hole through the center of the Teflon block with a diameter that closely matches the glassy carbon rod Press tit the glassy carbon rod mto the hole, it must be a very tight fit so that a water-tight seal forms between the Teflon and the carbon. Make the end of the glassy carbon rod flush with the end of the Teflon block. Press the brassrod into the oppositeend of the Teflon block until it makes electrical contact to the glassy carbon. 2 Sand the face of the glassy carbon disk and surrounding Teflon msulator with progressively finer sand paper. Use 600 grit m the final step Sanding will be necessary after application of the crosslinked redox polymer After sanding, check for the presence of remaining redox polymer by recordmg a cyclic voltammogram. Sand again until no redox polymer remains. 3 Polish the electrodes to a mirrorlike finish with a suspension of alumina micropolish on napped finishing cloth. Start with the 5.0~pm particle size, followed with the 1.0~pm, and finally with the 0 3-pm particle size Between polishing steps, vigorously rinse the electrode surface under a Jet of ultrapure water from a squirt-bottle. Although 0.05~pm mlcropollsh IS avallable, its use IS not recommended; it is difficult to wash the 0.05~pm particles from the electrode. 4. Somcate the electrodes to remove final traces of the alumma micropolish, rinse in ultrapure water, and dry m air.

3.2.2. Carbon Fiber Microelectrodes Microelectrodes basedon carbon fibers mounted mto pulled glasscapillary tubes have become a routine substrate electrode for carrying out measurements of neurotransmitters and related substances,such as choline, in the bram of anesthetized animals. Preparation of these microelectrodes is described in this section. 1 Work in an area that is free of drafts with a white, brightly lit surface. The carbon fibers are delivered m a bundle containing, typically, approx 1000 fibers. Cut a sec-

tion of thebundleabout 15-cmlong Isolateasinglefiber from thebundle.Note: The fibers stick together, so a pair of fibers IS easily mistaken for a single fiber.

Redox Polymers

127

2. Using forceps, immerse a glass tube, about lo-cm long, in a graduated cylinder filled with acetone. Hold the tube with one end slightly above the surface of the acetone, allowing capillary action to fill it to the top. Hold a carbon fiber about 1 cm from the end and guide it into the immersed tube. Raise the tube so that the acetone drains away; the fiber will stick to the inside surface of the tube. Gently let go of the fiber without pulling it away from the glass (keep fingers clean and dry so the fibers do not stick to your skin). Reimmerse the tube in the acetone and help the fiber float through the tube until it is protruding from both ends. 3, Mount the tube with the carbon fiber inside m a capillary puller. Preadjust the heating and pulling controls so that the glass collapses around the fiber with a 5-10 mm taper. Inspect the pulled tubes under a microscope: the glass tip must be snug around the fiber, and the fiber must extend far enough inside the tube to allow electrical connection. 4. Although the glass will appear to be sealed around the fiber, the seal is not adequate and it is necessary to back fill the capillary with epoxy. The epoxy is injected into the barrel of the electrode and fills the tip by capillary action. Air m the tip must be able to escape. Tugging gently on the fiber creates a small gap between the fiber and the glass that allows air to escape but does not allow the epoxy to flow out of the trp. 5. Fill a disposable syringe with the premixed Spurr epoxy. Attach a needle with a section of fused silica tubing glued inside; the fused silrca tubing must be long enough to reach inside the pulled glass tubes. Deposit a small droplet of Spurr epoxy inside the tapered part of the glass of the glass tube. After a short time, the Spurr epoxy will fill the electrode tip by capillary action. Cure the epoxy. 6. Fill the barrel of the microelectrode with mercury and insert a nichrome wire for electrical hook-up. Seal the open end of the glass tube by tightly wrapping with a strip of Paratilm to hold the hook-up wire in place and prevent mercury spills. Trim the fiber to the desired length, between 100 and 400 pm.

The procedure

described here for the preparation

of choline

sensors was

devised through a systematic investigation into how the sensor response is influenced by the absolute and relative amounts of redox polymer, crosslinking agent, and enzymes deposited on the electrode. The discussion here is limited only to the procedure currently in use in this laboratory, 3.3.1. Deposition of Redox- Polymer and Enzymes 1. For each electrode to be modified, prepare a mixture by combining, in the order specified, the following items (see Note 1): a. 20 pL of a 1 mg/mL solution of the redox polymer dissolved in water,

b. 4 pL of a 3 mg/rnL solution of the diepoxide crosslinking agent in water, c. 10 pL of a 4 mg/mL solution of ascorbate oxidase in HEPES buffer, d. 10 uL of a 3 mg/mL solution of HRP in HEPES buffer, and e. 10 pL of a 2 mg/mL solution of choline oxidase in HEPES buffer.

128

Shankar, Garguilo, and Michael

2a. To modify a 3-mm-diameter glassy carbon electrode, transfer a 5-uL ahquot of the mrxture to the electrode surface with a plastic prpet; the transfer must be completed within 5 min of preparing the mixture. Use the prpet to spread the solutton over the entire electrode surface. The hydrophobrc nature of the Teflon insulation will help contam the solution to the vicinity of the electrode. 2b. To modify a carbon fiber microelectrode, a 2-pL ahquot of the mixture IS used The aliquot is measured mto a mrcroprpet tip and then the plunger of the prpet 1s depressed, causmg the aliquot to form a droplet at the end of the prpet. Under a microscope, the droplet IS held m contact with the carbon fiber electrode for several minutes. During this time, the water m the mixture evaporates and the components of the mixture deposit onto the electrode. The droplet must be held m place with the mrcroprpet long enough for the solution to become too vrscous to flow, otherwrse surface tension forces will cause the droplet to travel up the glass body of the electrode (if this happens, the droplet can usually be dragged back to the carbon fiber with the prpet). See Note 2. 3 Stand the modified electrode m au for 48 h. During this time the cross lmkmg reaction takes place. After 48 h, rinse the sensor m ultrapure water for 15 mm, and dry m an for 2 h (the times mentioned here are important).

3.3.2. Deposition of Nafion If the sensors are to be used in the presence of ascorbate, which is found in of about 400 p&f, it is necessary to coat

most brologic tissues at concentration the sensor with a Nafion overlayer.

1. All Nafion solutions should be somcated for approx 5 mm pnor to use because the Nafion tends to accumulate on the walls of the container. Dilute the as-received 5% Nafion solutron with 2-propanol. Make 0.5, 1, and 3% Nafron solutions in 2-propanol. 2a. To apply Nation to the modified glassy carbon electrodes, spread 2 pL of the 0.5% solutron over the enzyme-containing layer with a pipet. 2b. Nation IS applied to carbon fiber microsensors by dip-coatmg. Dip the microsensor m the 0.5% Nafion solution for 10 s, then hold it m an for 20 s. Repeat ten times. Wart for 10 min and repeat the ten dips m 1% Nafion solution Wait 10 min. Next, dip in the 3% Nafion solution for 20 s and hold in air for 40 s. Repeat three times. 3 Stand the sensor in air for 34 h and then store in PBS until use Hereafter, keep the sensors wet.

3.4. Sensor Evaluation In this section we briefly characterize the electrochemical responses of the glassy carbon electrodes modified with the redox polymer according to the procedure described m Subheading 3.3. The electrochemical behavior of the modified carbon fiber electrodes is essentially identical, except that the currents are approximately three orders of magnitude smaller as a consequence of their smaller surface area.

Redox Polymers

729

4 3 3” Ee! 2

1

0 -1

600

400 200 E (mV vs. SCE)

0

Fig. 1. Cyclic voltammograms (10 mV/s) recorded with a glassycarbon disk electrode (3-mm diameter) modified with a crosslinkedpolymer layer containing HRP andcholine oxidase.Voltammogramswere obtained in phosphate-bufferedsalinewith and without 5 mM hydrogen peroxide. Figure 1 reports cyclic voltammograms obtained with a modified glassy carbon electrode in a stationary solution of PBS in the presence and in the absence of 5 rnM Hz02. The recording of the voltammograms was initiated at +600 mV vs SCE and proceeded initially in the negative potential direction. The direction of the potential scan was reversed at 0 mV vs SCE. The potential sweep rate was 10 mV/s. In the absence of peroxide, the unperturbed voltammogram of the redox polymer is observed. The voltammogram, which has the symmetric appearance expected when the redox couple is confined to a thin layer, shows that the half-wave potential of the mediator is near +300 mV vs SCE. In 5 Mperoxide, the voltammogram takes on the appearance expected for a catalytic process mediated by the redox polymer. Although data are not shown here, there is no evidence of catalysis if HRP is not immobilized in the film, confirmmg that the nonenzymatic oxidation of the osmium complex by peroxide is very slow. Figure 1 demonstrates that the sensor will respond to peroxide whenever the applied potential is held at values negative with respect to the half-wave potential of the mediator.

730

Shankar, Garguilo, and Michael

Cyclic voltammetry is a useful way to examine the modified electrodes in stationary solution, but at 10 mV/s the procedure is too slow for routme work. More often, these sensors are operated in an amperometric fashion, i.e., with the potential at a constant value. A convenient approach for characterizing the amperometric behavior of these sensors is to mount them m a flow system equipped with a chromatography-style injection valve. As with any electrochemical detector, It 1simportant to have pulse-free flow through the system, which 1seasily established with a gravity feed from an elevated buffer reservoir. The injection valve allows samples of different composltion to be sequentially introduced to the sensor and then removed in a convenient manner. Since the concentration of sample at the sensor changes very rapidly with this technique, the temporal response of the sensor can also be examined. The top panel of Figure 2 shows the full time course of the response of a choline sensor to the inJection of a sample bolus containing choline (50 CLM) into the flow system. During this experiment, a constant potential of-100 mV vs SCE was applied to the sensor. The sample valve was actuated at the 10-s time point, and after a delay of about 4 s as the sample travels to the sensor, the amperometric signal changes in about 5 s to a new steady-state value. Using the difference between the average current measured before the injection and 20 s after the injection as a quantitative measure of the response, these sensors typically exhibit a detection limit near 1 w choline and respond in a linear fashion up to millimolar concentrations. In addition to calibrating the sensor, it is important to evaluate the extent to which the sensor suffers interference by ascorbate. The lower panel of Figure 2 clearly shows why this is necessary.This panel shows the quantitative response of two different sensors, one coated with Nation and one not coated with Nafion, to the injection of sample solutions containing a fixed concentration of choline (20 r-LM>and varying concentrations of ascorbate (O-400 @4). Whereas the response of the Nafion-coated sensor to choline is unaffected by the ascorbate, this 1snot true of the uncoated sensor. Our recent experience shows that cholme sensors with this type of ascorbate interference cannot be used for in vivo experiments because of the high ascorbate levels present in brain extracellular fluid. 4. Notes 1 Dissolve the redox polymer in water at least 24 h in advance,but do not use solutions for more than 1 wk. In this recipe, the quantitiesof enzymelisted refer to the as-receivedsolid, which we use without purification. The order of mixing of the componentsis important sinceit influences the performance of the final sensor.Choline oxidasemust be the last componentaddedbecause,on addition

131

Redox Polymers

-6 0

100

200

300

400

ascorbate, mM

Fig. 2. (A) The time course of the response of a choline sensor mounted in a flow system to the injection of a sample solution containing 50 plM choline The sample valve was opened at the 10-s time pomt and held open for 30 s. (B) A comparison of the quantitative response to 20 $4 choline obtained with a Nafion-coated choline sensor to that obtained with a sensor lacking a Nafion coatmg. of this enzyme, a precipitate will slowly form in the solutton. The exact composition of the precipitate is unclear. If the precipitation is allowed to proceed for more than about 5 mm, the sensitivity of the sensor to enzyme substrate will be decreased. Also, the deposited film will not be uniform, making it difficult to coat the sensor with Nafion (see Subheading 3.3.2.) 2. Examme the sensor under the microscope to make sure that the coating of the enzyme solution is uniform and that no cracks appear after the apphcatton of the enzyme droplet.

References 1. Gregg, B. A. and Heller, A. (1991) Redox polymer films containing enzymes: 2. Glucose oxidase containing enzyme electrodes. J Phys. Chem 95,5974-5980.

132

Shankar, Garguilo, and Michael

2 Dalton, E. F , Surridge, N. A., Jermgan, J. C., Wilbourn, K. O., Facci, J. S., and Murray, R W (1990) Charge transport m electroactive polymers consistmg of fixed molecular redox sites. Chem Phys 141, 143-157. 3 Garguilo, M. G. and Michael, A. C. (1995) Optimization of amperometrrc microsensors for momtormg cholme in the extracellular fluid of brain trssue Anal Chim Acta 307,291-299. 4. O’Hara, T. J , Rajagopalan, R., and Heller, A (1994) “Wired” enzyme electrodes for amperometrtc determmation of glucose or lactate m the presence of interfering substances Anal Chem 66,245 l-2457. 5. Garguilo, M. G. and Michael, A. C. (1994) Quantitation of choline in the extracellular fluid of brain tissue with amperometric microsensors Anal Chem. 66, 262 l-2629. 6 Lay, P. R., Sargeson, A. M., and Taube, H. (1986) crs-bis(2,2’-Bipyridme-N,N’)

complexes of ruthenmm(III)/(II) and osmmm(III)/(II), m Inorganzc Syntheszs, vol. 24 (Shreeve, H. J. M., ed.), Wiley, New York, pp. 291-299. 7. Gregg, B A and Heller, A. (1991) Redox polymer films contammg enzymes 1. A redox-conductmg epoxy cement synthesis, characterizatton, and electrocatalytic oxidatron of hydroqumone J Phys Chem 95,5970-5975

Enzyme Biosensors Based on Metalked Carbon Electrodes Joseph Wang 1. Introduction 7.7. Selectivity of Amperomefric Enzyme Electrodes Amperometric biosensors based on enzyme electrodes satisfy many of the requirements for clinical assays,environmental monitoring or process control. Unfortunately, the inherent specrficity of the biocatalytic (recognition) reaction is compromised by the partial selectivity of the electrode transducer. Coexrstmg electroactive compounds,present m complex samples,such as,whole blood or fermentation broths,may result in overlapping current signals.For example,the clinical monitoring of glucose or lactateis commonly compromised by anodic contributions from easily oxidizable constituentsof biologic flu&, suchas,urrc or ascorbic acids. The degreeof such interference is strongly influenced by the operating potential. Several routes have been proposed to minimize the interference of oxidizable components, and hence to enhance the sensor selectivity. An early and still useful strategy is to use discriminative or permselective coatings that effectively exclude potential interferences from the transducer surface (1). Different polymers, mixed or multilayers, with transport properties based on charge, size, or polarity, have been used against coexisting oxidizable constituents (2-5”. Effective, yet incomplete, rejection has been reported in most cases.Alternatively, it is possible to use artificial electron acceptors, such as, ferkocene derivatives (6) or ferrrcyanide (7), that shuttle electrons from the redox center of the enzyme to the transducer surface. Redox polymers, “electrically wired” to the enzyme, represent another approach for shuttling electrons between the enzyme and the transducer (s). Yet, despite the lowering of the operating potential, mediated electrodes still suffer from some ascorbic or uric From. Methods m Bfotechnology, Vol. 6 Enzyme and Microbial Bfos8nSOfS Technrques Edlted by A Mulchandani and K R Rogers 0 Humana Press Inc , Totowa,

733

and Protocols NJ

134

Wang

acid interferences; in addition, the toxicity of some mediators limits their use in in viva applications.Another, less-usedroute is the biocatalytic destruction of potential interferences (9,10), that requires the coimmobilization of additional enzyme(s). The goal of this chapter is to review a relatively new and effective strategy for preventing common interferences m amperometric biosensmg based on the use of metallized-carbon transducers. Such focus on the transducer ehmmates major interferences in the first place, and hence circumvents the need for rejecting or destroying them. The remarkable selectivity of these metaldispersed carbon transducers is attributed to their strong, preferential, electrocatalytic action toward the detection of the enzymatically produced peroxide or reduced nicotinamide adenine dinucleotide (NADH) species. Such activity allows tunmg of the operating potential to the optimal potential range, (+O.l to -0.2 V) at which unwanted reactions do not occur. These new firstgeneration biosensors also offer high sensitivity, fast response, and reduced sensor complexity (as expected from the absenceof interferant-eliminating layers). The development, performance characteristics, and biosensing applications of these devices are reported in the following sections. The references are not meant to be exhaustive; they show examples of particular applications, of which there may be a few others (see Note 1). 1.2. Which Metal Center? Early work on metal-dispersed carbon biosensors focused on the utility of platinized carbon for enhancing the sensitivity of oxidase- or dehydrogenasebased devices (11-13). Palladium-dispersed or sputtered particles displayed similar improvements (1415). The dramatically improved sensitivity toward the liberated peroxide or NADH species is attributed to the tine dispersion of the metal microparticles m the carbon matrix. Such dispersion results in highly catalytic surfaces, as long as the particle size is comparable to that of the electrical double layer (1617). Unfortunately, the dispersed platmum or palladium particles are also highly electrocatalytic toward interfering substances.Accordingly, our group and that of Turner have been searching for other noble metal catalytic centersthat offer strong preferential electrocatalyticaction toward the liberated peroxide or NADH species,but not toward coexistmgoxidizable constituents, In the systematic development of these interference-free transducers, we have asked ourselves several fundamental questions: 1 Which metal centers(or combinationof metals)offer preferential electrocatalytic detection of the peroxide or NADH products? 2 Why do thesemetalsoffer suchan attractive behavior‘? 3. Is there a trend m the overvoltage lowering? 4. What is the mechanism by which these metal centers promote the redox reactions of these species? 5. What is the influence of the preparation conditions on the electrocatalytic action?

Metallized Carbon Electrodes

Fig. 1. Scanning electron micrograph of a glucose microsensor prepared by codepositing rhodium and glucoseoxidaseonto a carbonfiber electrode(Reproduced with permission from ref. 21).

Such systematic investigation has led to the identification of dispersed rhodium (16,17), ruthenium (18,19) and iridium (19) particles as selective promoters of the peroxide (and to a lesserextent NADH) reactions (seeNote 2). In the following sectionswe will discussproceduresfor preparing different types of metallized-carbon biosensors(seeNote 3). Thesewill include metallized-carbon enzyme microelectrodes fabricated by one-step codeposition of metal microparticles and the biocatalyst (13,20-23), which allows a controllable localization of the enzyme onto small electrode surfaces (Fig. l), metallized biocomposites that couples the advantagesof carbon paste biosensors (versatility, bulk modification, renewability, and low background [24]) with the effective electrocatalytic action of three-dimensionally dispersed metal particles (25), and mass-producible thick-film metal-dispersed carbon strips (15,26,27) (see Note 4).

136

Wang

2. Materials The following materials lized-carbon biosensors.

are needed for the preparation

2.7. Mefaiiized-Carbon

Paste Enzyme Electrodes

1, 2. 3. 4. 5. 6

of the various metal-

Mineral oil (Nu~ol, from Aldrich, Milwaukee, WI) Metal-on-graphite particles (from Aldrich or Alfa [Ward Hill, MA]). Relevant oxidase enzyme (from Sigma, St. Louis, MO, or Fluky, Buchs, Switzerland) Polyethylimine (PEI) enzyme stabilizer (from Sigma). Glass or Teflon 2-mm id 7-cm long tube (electrode body). Copper wire (1 -mm diameter, g-cm long) for electrical contact.

2.2. Metaliized-Carbon

Screen-Prlnted Biosensors

1. Commercial carbon ink (from Gwent [Gwent, UK], Ercon [Wareham, MA], or Acheson Inc [Woodbury, CT]). 2. Metal powder (from Aldrich or Alfa). 3. Oxidase enzyme (from Sigma or Alfa). 4. Silver/silver-chloride mk (for printing the reference electrode from Ercon or Acheson) 5. Insulator ink (from same sources). 6. Alumina ceramics or plastic (PVC) substrates. 7. Nafion (Aldrich) or Eastman AQ (Eastman) polymer solutions 8. Semiautomatic screen printer (from MPM Inc., [Franklin, MA] or DEK Inc [Dorset, UK]).

2.3. Metallized-Carbon 1. 2 3. 4 5. 6. 7. 8. 9.

Enzyme Microekctrode

Carbon fiber (10 pm diameter, Union Carbide, Chicago, IL). Glass capillary (l-mm id) Copper wire (0.5~mm diameter). Metal standard solution (Rh, Ru, Pt, Pd, AAS grade from Aldrich) Relevant oxidase enzyme (Sigma). Distilled water. 0.1 M Potassium hydroxide solution. Mercury. Electrochemical cell (Model VC-2, BAS Inc., W. Lafayette, IN).

3. Methods

3.1. Preparation of Mefallized-Carbon

Paste Enzyme Electrodes

1. Hand mix 60 mg mineral oil and 40 mg of the metal-on-graphite particles. 2. Hand mix 5 mg of the enzyme with 2 mg of PEI stabilizer. 3 Add the enzyme-stabilizer mixture to the metallized-carbon paste and mix thoroughly for 30 min. 4. Pack a portion of the resulting paste into the electrode cavity of the glass or Teflon sleeve. 5. Establish electrical contact to the inner surface with the copper wire. 6. Smooth the outer carbon paste surface on a weighing paper Caution: Metal-on-carbon particles may be harmful rf inhaled or swallowed. The exact paste composition depends on the desired viscosity.

137

Metallized Carbon Electrodes 3.2. Preparation

of Metallized-Carbon

Screen-Printed

Biosensors

1. Add the metal powder to the commercial carbon ink (at 5% w/w) and mix thoroughly. 2. Screen print the resulting ink on the substrate through a patterned stencil 3. Dry and cure the resulting strip according to the manufacturer’s recommendations (usually 30 min at 15OT). 4. Print and cure similarly the silver/silver chloride reference electrode 5. Print and cure the insulatmg layer. 6. Allow to cool. 7. Immobilize the enzyme on the exposed working electrode by casting 50 pL of 1% Nation or Eastman AQ polymer solution containing 1000 U/n& enzyme.

3.3. Preparation

of Metallized-Carbon

Enzyme Microelectrode

1. Insert a single carbon fiber into a l-mm id glass capillary, leavmg 5 mm outside. 2. Seal the fiber carefully with epoxy resin, and provide electrical contact (to the inner fiber) with a copper wire and mercury droplet. 3. Prepare 2 mL of solution containing 100 mg/L of the metal (in distilled water), and adjust the pH to 5.0 (with potassium hydroxide). 4. Add 1000 U of the oxidase enzyme to the metal solution. 5. Immerse the carbon fiber electrode, along with the reference and counter electrodes, into the solution. 6. Codeposit the enzyme and metal potentiostatically for 10 mm at a potential of-O.7 V with stirring. 7. Disconnect the applied potential and rinse the enzyme microelectrode for 15 s with distilled water. 8. Store the electrode m phosphate buffer (pH 7 4) until use

Caution: Handle the mercury carefully. 3.4. Operation of Metallized Carbon Biosensors The various metal-dispersed carbon enzyme electrodes described above commonly operate in the amperometrtc or chronoamperometric modes (see Note 4). The experiments are carried out in a 5-10 mL batch cell or by injectmg into the flow through electrochemical detector. Microfabricated strips may serve as self-contained cells onto which sample droplets are placed (see Note 2). Chronoamperometric measurements are commonly performed by immersing the biosensor in the quiescent sample solution, and stepping the potential from open circuit to the desired value (corresponding to the peroxide or NADH reactions). Amperometric measurements are carried out by applying the desired potential while stirring or flowing the solutron, and allowing the background current to decay before measurements of the sample and standards. The selection of the operating potential for both operating modes should be facilitated by recording first the hydrodynamic voltammograms, i.e., current-potential plots for both the target substrate, as well as for the potential interferences (see Note 1). Figure 2 displays a typical response of the palladium-dispersed strip

138

Wang

0.2

0.4

Potential

0.6

0.0

01)

t b 2t *-b

2 min

a

0.5v

Time Fig. 2. (Top) Hydrodynamic voltammograms for glucose at the metalhzed and nonmetalhzed strips. (Bottom) Amperometnc response of the palladium-dispersed carbon strip glucose sensors to 5 x 1C3 A4 glucose (a), 1 x l@Murlc acid (b), 1 x lOA Macetaminophen (c), and 1 x l@Mascorbic acid (d), using applied potentials of 0.3 and 0.5 V.

139

Metallized Carbon Electrodes a b c d $. JI 4 $

Time Fig. 3. Amperometnc response of the rhodium-dispersed glucose oxidase carbon paste biosensor to additions of 2.5 x 1CY5A4 acetaminophen (a), uric acid (b), ascorbic acid (c), and 1 x 1t3 M glucose (d). Operating potential, -0.1 V. (Reproduced with permission from ref. 16).

electrode toward the glucose substrate and common interferences at two dlfferent operating potentials, together with hydrodynamic voltammograms for glucose for the metallized and nonmetallized surfaces (see Note 3). 4. Notes 1. The metallized-carbon enzyme electrodes, described above, are aimed pnmarily at enhancing the selectivity by tuning the operating potential to the optimal region at which interferences from coexisting electroactive species are eliminated (Fig. 3). The electrocatalytic action also results in a greatly enhanced sensitivity. 2. Since these bioprobes still rely on the use of oxygen (as a cofactor), attention should be given to the oxygen-dependence issue. An appropriate membrane coverage may be useful to alleviate the oxygen limitation, extend the linear range, and reject large surface-active species from the surface. Such coverage may, however, result in longer response times. 3. As with other types of enzyme electrodes, attention should also be given to the issue of long-term stability, associated with the gradual loss of the biocatalytic activity. The stability of the electrocatalytic action should also be considered. 4. The specific type, configuration, and size of the metallized carbon biosensor should depend upon the specific application, e.g., sensor strips for single-use applications or flow-detector biocomposite for high-speed analysis.

140

Wang

References 1 Wang, J. (1991) Modified electrodes for electrochemical sensors. Electroanalyszs 3,255-259. 2. Wang, J. and Wu, H. (1993) Permselective lipid-poly(o-phenylenediamme) coatmg for amperometric btosensing of glucose. Anal. Chzm Acta 283,683-688 3. Palmtsano, F., Guerriert, A., Quinto, M., and Zambonm, P. G (1995) Electrosynthesized btlayer polymeric membrane for effective ehmmation of interference m amperometric biosensors. Anal Chem. 67, 1005-l 009. 4 Sasso, S., Pierce, R , Walla, R., and Yacynych, A. M (1990) Electropolymerized 1,2-dtaminobenzene as a means to prevent interferences and foulmg and to stabilize tmmobtltzed enzyme m electrochemtcal btosensors Anal Chem 62, 1111-1117. 5 Bartlett, P N and Cooper, J M (1993) A review of the mnnobthzatron of enzymes m electropolymertzed films J Electroanal Chem 362, l-12 6 Cass, A., Davis, G , Francis, G , Hill, H. A., Aston, W., Higgins, J , Plotkin, E., Scott, L , and Turner, A. P (1984) Ferrocene-mediated enzyme electrode for amperometric determination of glucose. Anal. Chem 56, 667-67 1. 7 Lewis, B. (1992) Laboratory evaluatton of the glucocard blood test meter Clzn Chem 28,2093-2096 8. Taylor, C , Kenaust, G., Katakis, I., and Heller, A (1995) “Wtrmg” of glucose oxidase wtthm a hydrogel made with polyvinyl tmidazole complexed with [(OS-4,4’-dimethoxy-2,2’-bipyridme)Cl]. J. Electroanal Chem 396,51 l-5 15. 9. Wang, J., Naser, N., and Wollenberger, U. (1993) use of tyrosmase for enzymatic ehmmation of acetaminophen interference in amperometrrc sensing. Anal. Chzm Acta 281, 19-24. 10 Maiden, R. and Heller, A. (1992) Elimination of electrooxidizable mterferantproduced currents in amperometric btosensors. Anal. Chem 64,2889-2896. 11. Ikariyama, Y., Yamauchi, S., Yukiashi, T., and Ushioda, H. (1989) Electrochemical fabrication of amperometrm microenzyme sensor, J Electrochem Sot 136,702-706.

12 McNeil, C., Spoors, J., Cooper, J., Alberti, K., and Mullen, W (1990) Amperometrtc biosensor for rapid measurement of 3-hydroxybutyrate m undiluted whole blood and plasma. Anal. Chzm Actu 273,99-105 13. Wang, J., Li, R., and Lin, M. S. (1989) Flow cell based on glucose oxtdase-modtfied carbon fiber ultramtcroelectrodes. Electroanalyszs 1, 15 l-l 54. 14. Jonsson, G. and Gorton, L. (1987) An amperometric glucose electrode based on adsorbed glucose oxidase on palladium/gold modified graphtte. Anal Lett. 20,839-855. 15. Wang, J. and Chen, Q. (1994) Screen-printed glucose strip based on palladmmdispersed carbon ink. Analyst 119, 1849-l 85 1. 16 Wang, J , Lm, J., and Lu, F (1994) Htghly selective membrane-free, mediatorfree glucose biosensor. Anal. Chem 66,3600-3603 17. White, S., Turner, A. P., Schmid, R , Bilitewskt, U., and Bradley, J (1994) Investtgation of platmzed and rhodimzed carbon electrodes for use m glucose sensors. Electroanalyszs 6,625-632.

Metallized-Carbon Electrodes

141

18. Wang, J., Fang, L , Lopez, D., and Tobias, H. (1993) Highly selective and sensitive amperometrtc brosensing of glucose at ruthenium-dispersed carbon paste enzyme electrodes. A&. Lett 26, 18 19-l 830. 19. Wang, J., Romero, E., and Reviejo, A. (1993) Improved alcohol biosensor based on ruthenium-dispersed carbon paste enzyme electrodes. J Electroanal Chem 353,113-120. 20. Wang, J. and Angnes, L. (1992) Mimaturized glucose sensors based on electrochemical codeposition of rhodium and glucose oxidase onto carbon-fiber electrodes Anal. Chem 64,45W59. 2 1 Sakslund, H., Wang, J., and Hammertch, 0. (1994) A critical evaluation of a glucose biosensor made by codeposition of palladium and glucose oxidase on glassy carbon, J Electroanal Chem. 374,71-79 22. Tamiya, E., Sugiura, Y., Takeuchi, T., Suzuki, M., Karube, I., and Akiama, A. (1993) Ultra micro glutamate sensor using platinized carbon-fiber electrode. Sens Actuat. BlO, 179-184 23. Sakslund, H.,Wang, J , Lu, F., and Hammerich, 0. (1995) Development and evaluation of glucose microsensors based on electrochemical codeposition of ruthenium and glucose oxidase onto carbon fiber microelectrodes. J Electroanal Chem. 397,149-155. 24. Gorton, L. (1995) Sensors based on carbon paste m electrochemical analysis. Electroanalyszs 7,5--45. 25. Wang, J , Naser, N., Angnes, L., Wu, H., and Chen, L (1992) Metal-dispersed carbon paste electrodes. Anal Chem 64, 1285-1288. 26. Newman, J., White, S., Tothill, I., and Turner, A. P. (1995) Catalytic materials, membranes and fabrication technologies suitable for the construction of amperometric biosensors. Anal Chem 67,4594-4599. 27. Wang, J., Chen, Q., Pedrero, M., and Pingarron, J. (1995) Screen-printed amperometric biosensors for glucose and alcohols based on ruthenium-dispersed carbon inks. Anal Chim. Acta, 300, 111-I 16.

11 Enzyme Biosensors Wolfgang

Based on Conducting

Polymers

Schuhmann

1. Introduction A major concern in the development of amperometric biosensors is the tight and reproducible immobilization of enzymes with high activity on the electrode surface. Additionally, aiming on the development of miniaturized amperometric enzyme electrodes, it should be possible to predefine the site for the immobilization

of the enzyme without using manual deposition

techniques,

In this respect, conducting polymers like polypyrrole, polythiophene, polyaniline, and polymdole show distinct advantages over nonconducting polymers because they can be electrochemically grown exclusively on the surface of an electrode. Several reviews summarizing this research area have been published m the last few years (1-5). The formation of conducting polymers is usually attained by means of an electrochemically induced polymerization of the monomer in 02-free solution. The first step is the oxidation of monomers at an appropriate electrode potential, leading to radical cations in the vicinity of the electrode, which can react in a second step with another monomeric or oligomeric radical cation, or a neutral monomer under formation of a dimeric radical cation or a dimeric diradical dication. The diradical dication may lose two protons, leading to the dimer (or to higher ohgomers). Since the thus obtained dimers and ohgomers have lower oxidation potentials than the monomer itself, the polymer formation process leads predommantly to a chain propagation, and after a critical chain length for precipitation is attained, the deposition of the polycationic polymer occurs on the surface of the anode. The intermediate radical cations may undergo reactions with nucleophilic species in the solution, preventing chain propagation. Thus, exclusion of oxygen and the proper choice of the From

Methods m Biotechnology, Vol 6 Enzyme and Mwob/a/ Blosensors Technrques Edlted by A Mulchandam and K R Rogers 0 Humana Press Inc , Totowa,

143

and Protocols NJ

144

Schuhmann

polymerization solution is indispensable for reproducible deposition of conducting polymer films. Local changes of pH as a result of the liberation of protons during the polymerrzatron reaction have to be taken mto account. The resulting polymer has a net positive charge that is neutralized by incorporation of anions from the electrolyte. For the reproducible formation of conductmg polymer films the electropolymerization potential determines-together with the temperature, the monomer concentration, the properties of the solvent, and the electrolytethe chain length of the polymer and thus the properties and morphology of the obtained polymer film The thickness of the polymer film can be estimated by measuring the charge transferred during the electrochemical film formation. The electrodeposition can be performed by controllmg the potential, which 1s the optimal method for producing homogeneous films. Thus, control of the deposition potential is indispensable for the reproducible formation of conductmg polymer films, and potentiostatic deposition regimens are highly recommended. Galvanostatic deposition procedures have been described that facilitate the calculation of the charge and hence the estimation of the film thickness. However, depletion of monomers in the diffusion zone around the electrode surface may lead to an uncontrolled high potential and a deterioratron of the film conductivity as a result of overoxidation processes. One major advantage of electrochemtcal production of conductmg polymers over deposition techniques necessary for nonconductmg polymers is the exact localization of the polymerization reaction together with the control of the properties of the conducting polymer film during the electrochemical deposition process. However, sometimes the polymer film is growing on only a few active spots of the electrode surface leading to variations of the film thickness. Electrochemical platimzation of the electrode surface by means of reductive deposition of small platinum crystallites from H2PtC1,-solution increases significantly the number of active spots on the surface and hence leads to a simultaneous covering of the whole electrode surface by the growing polymer film. The homogeneity of the polymer film can be estimated from the formation of a diffusion barrier for redox species with increasing film thickness. Obviously, formation of functionahzed modified electrode surfaces by means of electrochemical polymerization procedures can be applied for the irnmobilization of biologic recognition elements on electrode surfaces. In principle, there are two different approaches to immobilize enzymesusing conducting polymers. The first and most widely used is to entrap the enzyme within the growing rarmtied polymer network during its electrochemical formation (6-l 0). The second 1s to use a two-step procedure consisting of the formation of a mnctionalized conducting-polymer film followed by the covalent binding of the biocatalysts to the functionalities at the polymer surface (II). Because of the, m general, large size

Conducting Polymers electrode with alatinum cl

U

V

enzyme

Fig. 1. Schematicrepresentationof the enzymedistributionin polypyrrole-based amperometricbiosensors.(Left) Enzymeentrappedduringthe electrochemicalpolymer formationprocess(one-stepprocedure).(Right) Enzymecovalentlybound at a functionalizedconductingpolymer(sequentialtwo-stepprocedure). of enzymesas comparedwith the pore size of the polymer, the enzymesare in this caseexclusively bound to the outer polymer surfacewhereasthey should be evenly distributed within the film after the first approach(Fig. 1). In the caseof entrapmentof enzymeswithin the growing polymer film, the slow diffusion of the high-mol-wt enzyme leadsto fast depletion of its concentration within the reaction zone in front of the electrode. Hence,to increasethe immobilized enzyme activity it is advantageousto use a potentiostatic pulse profile that enablesthe enzyme and the monomer to diffuse from the bulk of the deposition solution toward the electrode surface according to the existing concentration gradient and to reestablish high monomer and enzyme concentrations next to the electrodesurfacebefore the subsequentpotentiostatic pulse. As has been pointed out before, nucleophilic attack on the intermediate radical cations may interfere with the polymerization process.The nucleophilicity of side-chainsat the enzyme itself may lead to decreasedchain propagation probability and consequently to significant changesof film porosity and film morphology as compared with polymer films obtained in the absence of the enzyme. Thus, entrapment of enzymeswithin a growing conducting polymer film has to be optimized with respectto the individual properties of the enzyme in question and its maximum concentration in the deposition solution,

146

Schuhmann

A second possibility for the immobilization of enzymes at the conductingpolymer-modified electrode surface makes use of fimctionalized conducting polymer films as partners for the formation of covalent bonds. Either an already functionalized monomer can be polymerized or copolymerized with the unsubstituted parent monomer under direct formation of the functionallzed surface or an unsubstituted polymer film can be derivatized in a heterogeneous reaction on the electrode surface subsequent to the formation of the conducting-polymer film. These functionalized electrode surfaces are subsequently used for covalent binding of suttable enzymes in a second step. This sequential procedure has obvious advantages over a one-step entrapment process because of the inherent possibility to choose the optimum reaction conditions for each single step of the procedure. The conducting polymer film can be grown from organic solvent, functlonahzation may be performed under excluston of water and oxygen, and finally it is possible to switch to an environment that 1sbest suited for the covalent binding of the enzyme in question. Especially, the formation of the polymer under conditions that may be deleterious for biologic compounds expands the number of potentially usable conducting polymers significantly. In addition, labile enzymes, which may be deactivated during entrapment because of uncontrolled low pH values near the electrode surface, may be used in conducting-polymer-based biosensors. One possibility for the derivatization of already grown polypyrrole films m a heterogeneous reaction mvolves the nitration of polypyrrole using in sztu generated acetyl nitrate and the subsequent electrochemical reduction of the formed pyrrole-bound nitro groups under formation of poly(3-aminopyrrole) (12-14). Glucose oxidase has been immobilized via amide bonds generated from activated carboxylic side chains of the enzyme and the polymer-linked amino functions. As expected from the location of the enzyme at the outer surface of the polymer film, the response characteristics of these electrodes are dependent on the thickness of the polymer layer. The surface area and thus the amount of immobilized enzyme activity increaseswith increasing polymer film thickness, whereas the diffusion distance for H202 gets larger, leading to a lower probability for an HzOrmolecule to reach the electrode surface. Another possibility for obtammg functionalized conductmg polymer films for covalent attachment of enzymes is based on the polymerization of N-substituted pyrrole derivatives and subsequent covalent binding of the enzyme (1516). Similar modified electrodes have been obtained using amino-functionalized azulene derivatives (17), amino-functionahzed dithienylpyrrole derivatives (18), and functionalized thiophene or bithiophene derivatives (19). The sequential twostep process allows tailoring of the properties of the underlying conducting polymer film before binding the enzyme. Since the porosity and morphology of the polymer films can be varied by adjusting proper deposition conditions, the

147

Conducting Polymers

inherent size-exclusionproperties of the polymer film can be used to prevent interfering compounds from being oxidized at the metal-electrode surface (20,21). In the following sections the entrapment of glucose oxidase within a growing polypyrrole film, as well as the covalent binding of glucose oxidase at an amino-mnctionalized polypyrrole film, will be described in detail. 2. Materials 2.1. Electrode Pretreatment 1. P&electrode molten m soft glass exposing a hrghly polished platinum 1-mm diameter to the electrolyte. 2 HNOs concentrated. 3. H2S04 concentrated. 4. A1203 polishing paste with grain sizes of 6,3, 1, and 0.3 urn. 5. Polishing cloth.

drsk of

2.2. Platinization 1. 2 nuI4 H,PtCl, in HZ0 (4 mg/mL). 2. 02-free water.

2.3. Electrochemical

Deposition

of Po/ypyrrole

1. 2. 3. 4.

100 mMKC1. Pyrrole: best available purity, at least 99%. AlzOs-column (neutral; length, 5 cm; width, 0.4 cm). Electrochemical cell consisting of a three-necked flask with glass valve for connection to a high-vacuum/argon line. 5. Platinum counter electrode (coiled l-mm-diameter platinum wire molten in soft glass with a length of 2 cm) 6. Saturated calomel electrode as reference electrode with a saturated KC1 internal electrolyte separated from the deposrtron solution by means of a VYCOR diaphragm.

2.4. Entrapment 1. 2. 3. 4.

of Glucose Oxidase in Polypyrrole

Films

100 mA4KCl. Pyrrole: best available purity; at least 99%. AlzOs-column (neutral; length, 5 cm; width, 0.4 cm). Electrochemical cell consistmg of a three-necked flask with glass valve for connection to a high-vacuum/Argon line. 5. Platinum counter electrode (coiled 1-mm-diameter platinum wire molten in soft glass with a length of 2 cm). 6. Saturated calomel electrode as reference electrode with a saturated KC1 internal electrolyte separated from the deposition solution by means of a VYCOR diaphragm. 7. Glucose oxidase from Aspergillus niger (Sigma [St. LOUIS, MO], Type X, 125 U/mg).

Schuhmann

148 2.5. Functionalization of Polypyrrole and Covalent Immobilization of Glucose Oxidase 1 2. 3 4.

CH,CN (distilled and stored over molecular sieve 4 A under argon m the dark) Tetrabutylammoniump-toluenesulfonate (TBATos). l-Cyclohexyl-3-(2-morpholinoethyl)carbodiimide-metho-~-toluenesulfonate (CCD). Acetic anhydride.

5*

cu(No3)2

6 Glucose stock solution (anomerrzation was allowed to take place before use)

3. Methods 3.1. Hectrode

Pretreatment

1 Immerse the electrode m concentrated HNOs for 10 min in an ultrasonic bath. 2. Rinse the electrode with water (5 mm m the ultrasonic bath)

3. Pohsh the electrode surface on a pohshmg cloth using alumina paste with 6-, 3-, and 1-pm, and finally 0.3~pm grain size. 4. Rinse the electrode with water m an ultrasonic bath (5 mm) 5 Remove any lipid by washing the electrode in 10 M NaOH. 6 Rinse m water m the ultrasonic bath. 7 Rinse with concentrated H2S04 for 10 min m the ultrasonic bath 8 Rinse with water 9. Apply potential cycles with a scan rate of 100 mV/s m 0.1 MH$O, avoidmg Clm the electrolyte (Hg/HgS04 reference electrode; +650 mV vs normal hydrogen electrode [NHE]) a Scan-610 to +lOOO mV b. Scan-810 to 1600 mV. c. Subsequent scans -610 to +lOOO mV until the cychc voltammogram represents the bare platinum surface (Fig. 2; see Note 2) 10. Hold the electrode potential for 1 min at -210 mV vs Hg/HgS04. 11, Rinse with water

3.2. Platinization 1, Insert the pretreated electrode m a flask (see Note 3), which allows the positioning of the workmg electrode, a reference electrode, and a Pt-coil as a counter electrode in a volume of about 1 nL. 2. Connect the high vacuum/argon line to the glass valve of the flask. 3. Evacuate the flask three hmes to a residual pressure of 1v3 mbar and fill it with argon. 4. Transfer 4 mL of an 02-free 2 mA4 HzPtC16-solution m HZ0 (4 mg/mL) under exclusion of O2 in the electrochemical cell. 5. Apply three potential cycles between +500 and -400 mV vs saturated calomel electrode (SCE) with a scan rate of 10 mV/s (Fig. 3) 6. Remove the platimzation solution from the flask by means of a Pasteur pipet and wash the electrodes and the flask three times with O,-free water (see Note 4). 7. Dry the electrode using the vacuum line (final pressure is 1(Y3 mbar).

149

Conducting Polymers

-850

-550

-250

60

350 650 950 Potential [mV vs SCE]

Fig. 2. Cychc voltammogram of a Pt-electrode m 0.5 A4H,S04 between O2 and Hz evolutron (scan rate of 100 mV/s, Hg/HgS04 reference electrode; 1. Scan -610 to +l 000 mV, 2. Scan. -8 10 to 1600 mV; subsequent scans -610 to +I 000 mV. Cycling the electrode has to be continued until the cyclic voltammogram shows the separated waves for the Pt-hydride formation (see arrows).

3.3. Electrochemical

Deposition

of Polypyrrole

1. Purify pyrrole by passing 0.5 mL pyrrole (as received) through a neutral A&O,column (5 +0.4 cm) to remove any colored components (see Note 5). 2. Dissolve 56 pL pyrrole in 1.944 mL of 100 mA4KCl (final concentration of pyrrole is 100 m.M). 3. Apply a potentiostatic pulse profile (see Note 6) to the working electrode with 875 mV for 1 s (depositton phase) and 0 mV for 5 s (resting phase).

3.4. Entrapment

of Glucose Oxidase in Polypyrrole

Films

1. Purify pyrrole by passing 0.5 mL pyrrole (as received) through a neutral A1203column (5 cm +0.4 cm) to remove any colored components (see Note 5). 2. Dissolve 56 pL pyrrole m 1.944 mL of 100 mA4 KC1 (final concentratron of pyrrole is 100 mA4). 3. Add 1 mg/mL glucose oxtdase. 4. Apply a potentiostatic pulse profile (see Notes 6 and 7) to the working electrode with 875 mV for 1 s (deposition phase) and 0 mV for 5 s (resting phase). Figure 4

Schuhmann

150

-4 00 -400

-300

-200

-100

0

100

200

300

400

Potential

600 b+Jl

Fig 3 Cyclic voltammogram obtained during the platinization surface (scan rate of 10 mV/s, +500 and 400 mV vs SCE).

of the cleaned Pt

shows the deposition current over time (value taken at the end of each deposition phase) for deposition of polypyrrole in the presence of increasing concentrations of glucose oxidase.

3.5. Functionaliza tion of Polypyrrole of Glucose Oxidase

and Covalent

lmmobiliza tion

1. Purify pyrrole by passing 0.5 mL pyrrole (as received) through a neutral A1203column (5 +0.4 cm) to remove any colored components (see Note 5). 2. Dissolve 56 pL pyrrole m 1.944 mL of 100 mM CH-JN (final concentration of pyrrole is 100 mM) containing 100 mA4 TBATos (see Note 8). 3. Apply a potentiostatic pulse profile (see Note 6) to the working electrode with 875 mV for 1 s (deposition phase) and 0 mV for 5 s (resting phase) 4. Nitrate (see Note 9) the formed polypyrrole film by immersing the electrode into a solution of 700 mg Cu(NO& 3H20 m 20 mL acetic anhydride for 5 mm at 20°C under argon atmosphere.

151

Conducting Polymers 1000 ii E e! 2 "

800

800

400

200

0

i-l-

0

60

120

180

240

300

Time [s]

Fig. 4. The course of the deposition current with time (values taken at the end of each deposition phase) IS shown for deposition of polypyrrole m the presence of increasing concentrations of glucose oxidase (GOD). Concentrations of GOD were 0.5 (m), 1.0 (O), and 5.0 mg/mL (A). 5. Rinse the electrode three times wish O,-free CH,CN. 6. Reduce the formed nitro groups electrochemtcally by cycling three times the potential between +500 and -2500 mV vs SCE with a scan rate of 10 mV/s in a CHsCN solution containmg 100 mA4 TBATos (see Note 10). 7. Activate the carboxyhc side chains of 10 mg/mL glucose oxidase using a 100 mA4 solution of CCD in 100 rnM acetate buffer (pH 4.5) containing 100 mM glucose. 8. Immerse the modified electrode surface in the activated enzyme solutton for 3 h. 9. Rinse the electrode surface extensively with 100 mMKCl to remove any adsorbed enzyme.

3.6. Chsrscferizefion

of the Obtained Glucose Sensors

1. Immerse the prepared polypyrrole/enzyme electrode in a three-electrode electrochemical cell containing 100 rnM phosphate buffer (pH 7.4) as electrolyte together with an SCE reference electrode and a Pt-wtre counter electrode.

Schuhmann

152

80

60

0

5

IO Glucose

16 Concentration

20 [mMJ

Fig. 5. Callbratton graphs for polypyrrole-entrapped glucose oxidase sensors (mcreasmg enzyme concentratton durmg the electrochemical deposition procedure). Concentrations of GOD were 0.5 (e) and 2.5 mg/mL (A). 2. Apply a constant potential of 600 mV vs SCE between the reference electrode and the working electrode by means of a potentiostat 3, Allow the background current to decay until a constant and low value is obtained (in general Cl nA, typically about l-3 h) 4. Add ahquots of an anomenzed glucose stock solution (typtcal concentration steps are 1 m/14) and record the current through the electrochemtcal cell using a strip recorder Figure 5 shows the cahbratton graphs for different polypyrrole-entrapped glucose oxidase sensors, demonstrating the effect of increasing enzyme concentration durmg the electrochemical deposition procedure on the formatton of the polymer film.

4. Notes 4.7. Electrode Pretreatment 1 Electrode pretreatment IS very important for the reproducible deposition of conducting polymers on the respective surfaces. To obtain a conducting polymer

Conducting Polymers

153 personal computer with AD/DA card for application of the pulse profile and data aquisition

h-vacuum/ line

volume 1 ml

Fig. 6. Electrochemicalcell for the deposition of conducting polymers under inert gas atmosphere. film with reproducible film thicknessand morphology over the whole electrode area, it is important to standardizemethods of electrode polishing, removal of any lipid residues,and platinization. 2. Cycling theelectrodein 0.5MH2S04 betweenO2andHZevolutionshouldbe continued until the cyclic voltammogramshowsthe separatedwaves for the P&hydride formation (a typical exampleis shownin Fig. 2). However,the ideal surfacefor the electrochemicalpolymerizationis neitherthe Pt-hydridenor the Pt-oxide.Thus,it is necessaryto conditionthe electrodeat-2 10mV vs Hg/HgS04beforeuse.

4.2. Pla tin&a tion 3. Figure 6 showsthe cell usedfor electrochemicaldeposition of conducting polymers under inert gasatmosphere.A freshly cleanedand pretreatedelectrodesurface has in general only very few spotswith enhancedsurfaceactivity at which the electrochemicalpolymerizationprocessusually startsasthe nucleationenergy is lowered. The proposedmethod of platinization leadsto a uniform deposition of platinum clusterswith an averagesize of about 10 nm. Becauseany of these platinum cluster may serveasnucleation site and henceasa starting point for the

154

Schuhmann

conducting polymer deposition, the polymerization reaction takes place uniformly on the whole electrode surface leading to a uniform film thickness. 4. Since the obtained platinized electrode surface is highly active, it should not be transferred through air. A transfer chamber with continuous Argon flush to transfer solutions and the electrode under controlled condtttons from one flask to the other should be used.

4.3. Electrochemical

Deposition

of Polypyrrole

5. The quality of pyrrole is of high importance for the reproducible formation of polypyrrole films. Sometimes, even distillation and chromatography over Al,O, does not lead to satisfying results. Possibly, even minor impurities of nucleophilic compounds may efficiently interfere with the chain propagation reaction during the electrochemical polymerization. It saves a lot of time to buy the purest compound available. In general, it is sufficient to purify the necessary amount of pyrrole before use with a small A&O3 column. It is important that no colored compounds are eluted from the column. 6 Polypyrrole can be deposited at constant potential and at constant current. However, m both cases the concentration of the monomer at the electrode surface is dependent on the diffusion from the bulk of the solution to the reaction zone adjacent to the electrode surface. Thus, the probability of chain propagation, which is mamly determined by the concentration of radical canons m the ieaction zone, is decreased. To circumvent these problems, a potentiostatic pulse regimen permitting the monomer concentration to be reestablished at the electrode surface during a resting phase between subsequent short pulses with applied deposition potential should be used. Because of the increase of the electrode surface area with growing film thickness the current at the very end of a single deposition pulse is increasing concomitantly

4.4. Entrapment

of Glucose Oxidase in Polypyrrole

Films

7 The enzyme is entrapped within the growing polymer network. Since the mass transport by means of diffusion is even slower for the large enzyme molecules, the potenttostatic pulse deposition mode increases significantly the number of entrapped enzyme units Additionally, the enzyme itself bears nucleophihc side chains that trap electrochemically generated radical cations and hence decrease the probabihty for chain propagation Thus, the enzyme concentration, which should be high for the entrapment of a high enzyme activity, has a significant influence on the film formation itself and on the morphology of the deposited polymer. However, using the pulse deposition mode allows entrapment of a suff~cient enzyme activity using small enzyme concentrations in the deposition solution

4.5. Functionalization of Glucose Oxidase

of Po/ypyrro/e

and Covalent

Immobilization

8. The two-step procedure for the unmobilization of enzymes at conducting polymer films allows the deposition of the conducting polymer from organic sol-

Conducting Polymers

155

vents, giving a higher degree of freedom to tailor-make the film properties Polypyrrole from CH,CN using Tos-as counteranionis known to posseshigh film densityand a smooth polymer surface (CH,CN should only be used under a hood). 9. Nitratton of the depositedpolypyrrole film occurs in a heterogeneouspolymeranalog reaction with acetylnitrate, a compound that is highly explosive. By generating acetylnitrate in sztu from acetic acid and Cu(NO& we did not encounter any problems. 10. The electrochemical reduction of the formed nitro groups to the amino functions (a six-electron process) needs potentials of at least -1000 mV vs SCE

Acknowledgment The author is grateful to Harms-Ludwig Schmidt, Lehrstuhl fur Allgemeine Chemie und Biochemie, TU Munchen for the continuous support of the work, and to Johanna Strohmeier and Heidi Wohlschlager for their excellent technical help during the past years. References 1. Bartlett, P. N and Cooper, J. M (1993) A review of the immobilization of enzymes in electropolymerized films. J Electroanal Chem 362, 1-12. 2. Deshpande, M. V. and Amalnerkar, D. P. (1993) Biosensors prepared from electrochemically-synthesized conducting polymers. Prog. Polym SCL l&623--649. 3. Bartlett, P. N. and Birkin, P R. (1993) The application of conducting polymers m biosensors. Synth. Met. 61, 15-21. 4. TroJanowicz, M. and Krawczyk, T. K. V. (1995) Electrochemical biosensors based on enzymes immobihzed in electropolymerized films (review). Mzkrochrm Acta 121,167-181.

5. Schuhmann, W. (1995) Conducting polymer based amperometric enzyme electrodes (review). Mkrochim Acta 121, l-29. 6 Umana, M. and Waller, J. (1986) Protein-modified electrodes. The glucose oxidaselpolypyrrole system. Anal Chem. 58,2979-2983. 7. Foulds, N. C. and Lowe, C R (1986) Enzyme entrapment m electrically conductmg polymers. Immobihsation of glucose oxidase m polypyrrole and its apphcation in amperometrtc glucose sensors. J. Chem Sot. Faraday Trans. 82, 1259-1264. 8 Fortter, G., Brassard, E., and Belanger, D. (1988) Fast and easy preparation of an amperometric glucose biosensor. Biotechnol Tech. 2, 177-l 82. 9. Fortter, G , Brassard, E., and Belanger, D. (1990) Optimization of a polypyrrole glucose oxidase biosensor. Biosens Bioelectron 5,473-490. 10. Bartlett, P. N. and Whitaker, R. G. (1987) Glucose oxidase immobilized m polyN-methylpyrrole, in electrochemical immobilisation of enzymes, part II J Electroanal Chem. 224,37-G. 11. Schuhmann, W. (1994) Conducting polymers and their application in amperometric biosensors, in ACS Symposium Series. Diagnostic Biosensor Polymers, vol. 556

(Usmam,A M and Akmal, N., eds.), American Chemrcal Society,Washington, DC, pp. 110-123.

156

Schuhmann

12 Schuhmann, W., Lammert, R., Uhe, B., and Schmidt, H -L (1990) Polypyrrole, a new possibihty for covalent binding of oxtdoreductases to electrode surfaces as a base for stable biosensors. Sens Actuators Bl, 537-541. 13 Schuhmann, W (1991) Functtonahzed polypyrrole. A new material for the construction of biosensors. Synth. Met. 414,2!9-32. 14. Schuhmann, W. and Schmidt, H.-L. (1992) Amperometric biosensors for substrates of oxidases and dehydrogenases, in Advances in Biosensors vol. II (Turner, A. P F , ed.), JAI, London, UK, 79-l 30 15 Schalkhammer, T., Mann-Buxbaum, E., Ptttner, F , and Urban, G. (1991) Electrochemtcal glucose sensors on permselecttve nonconducting substituted pyrrole polymers. Sens Actuators B4,273-28 1. 16. Schalkhammer, T., Mann-Buxbaum, E., Urban, G., and Pittner, F. (1990) Electrochemtcal biosensors on thin-film metals and conductmg polymers. J Chromatogr. 510,355-366.

17. Schuhmann, W , Huber, J , Mirlach, A., and Daub, J. (1993) Polyazulenes 5 Covalent binding of glucose oxidase to functionalized polyazulenes-the 1st application of polyazulenes in amperometric biosensors. Adv Mater. 5, 124-126 18 Rockel, H., Huber, J., Gleiter, R., and Schuhmann, W. (1994) Synthesis of functionahzed poly(dithienylpyrrole) derivatives and their application m amperometric biosensors. Adv. Mater 6,568-57 1. 19. Hiller, M., Kranz, C., Huber, J., Bauerle, P., and Schuhmann, W. (1996) Amperometric biosensors by immobthzatton of redoxenzymes at polythiophenemodified electrode surfaces. Adv. Mater. 8,2 19-222. 20. Wang, J., Chen, S.-P., and Lm, M. S. (1989) Use of different electropolymerizatton conditions for controlling the size-exclusion selectivity of polyamlme, polypyrrole, and polyphenol films. J. Electroanal Chem 273,23 l-242. 2 1. Cosmer, S , Deronzier, A., and Roland, J. F (1991) Controlled permeabihty of functionalized polypyrrole films by use of different electrolyte amon sizes in the electropolymerization step. J Electroanal Chem 310,7 l-87.

12 Enzyme Sensors Based on Conductimetric Measurement Norman F. Sheppard, Jr. and Anthony Guiseppi-Elie 1. Introduction 1.1. Principles Enzyme sensors based on conductimetrtc measurement exploit the fact that the changes in substrate and product concentrations resulting from the catalytic action of some enzymes may be accompanied by a net change in solution electrical conductivity. As discussedby Lawrence (I), this conductivity change may result from a number of mechanisms. These Include: 1. The generation of ionic groups; 2. The separattonof unlike charges; 3. Proton generation and buffering; 4. Changes in the size of charge-carrying groups; and 5 Changes in the degree of association of ionic species.

For any given enzyme and corresponding substrate, one or more of the above mechanisms may produce a measurable conductivtty change. The mechanism that, by far leads to the greatest conductivrty change is the generation of ionic species. Many amidohydrolase enzymes convert a neutral substrate to charged products. One example is the tetramertc enzyme L-asparaginase (E.C.3.5.1.1), which catalyzes the hydrolysis of the amino actd L-asparagine (Asn; mol wt =132, pK, = 2.02, p&,=8.80) to L-aspartic acid (pK,= 2.19, pKt, = 9.67, pK, = 4.25) and ammonia, as follows: HOOC-CH(NH,~H,-CONH,

HOOC-CH(NH,+CH,~OO

From

+ Hz0 +

+ NH4+

Methods m Brofechnolcgy, Vol 6 Enzyme and Mjcrobral Bosensors Techniques E&ted by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

157

(1)

and Profoco/s NJ

158

Sheppard and Guiseppi- Elie

Similarly, the enzyme L-glutaminase (E.C.3.5.1.2) catalyzes the hydrolysis of L-glutamine (Gln; mol wt = 146, pK, = 2.17, pKb = 9.13) to glutamrc acid (pK, = 2.19, pK,.,= 9.67, pK, = 4.25) and ammonia, as follows: HOOC-CH(NH,)-CH&H+ONHZ HOOC--CH(NH,)-XH,-cH,O~

+ HZ0 + + NH;

(2)

A third example of the enzyme-catalyzed generation of ions is the hydrolysis of urea by the enzyme urease (E.C.3.5.1.5). The overall reaction results in the conversion of urea (mol wt = 60) to ammonium, bicarbonate and hydroxide ions. NH2CONH2+ 3H20 + HCO: + OH- + 2 NH; (3) In each of the cases above, the neutral substrate molecule is transformed by the enzyme transmutation mto charged products that increase the solution conductivity. The change in conductivity is largest, on a per substrate-molecule hydrolyzed basis, for the enzyme urease. Urease will be used to demonstrate the fabrication and testing of a conductimetric enzyme sensor in the protocol below. The second important mechanism by which a conductivity change may be generated as a result of enzyme action is the separation of unlike charges. Ammo acid oxidases (AAO) (E.C.1.4.3.2) and ammo acid decarboxylases (E.C.4.1 .l.) cleave zwitterionic ammo acids mto separate anions and cations. The AA0 enzymes catalyze the conversion of a D- or L-amino acid to the corresponding 2-keto acid, RCHNH;

COO- + O2 + Hz0 + RCOCOO- + NH; + H202

(4)

The ammo acid decarboxylases catalyze the following reaction: RCHNH; COO- + Hz0 + RCH,NH; HCO, (5) Esterases,such as acetylcholinesterase (E.C.3.1.1.7), result in the formation of carboxylic acids. Under neutral conditions, where the pH 1smuch greater than the pK, of the acid formed, the product dissociates to form a carboxylate ion and a proton, as follows: CH3COOCH2CH2N+(CH&+ HZ0 -+ HOCH2CH2N+(CH& + CH$OO- + H+ (6) If the solution in which this reaction is carried out is buffered, the magnitude of the resulting

conductance

change will depend on whether the buffer, in its

ionized form, is posrtively or negatively charged. The conductance of the former will be increased upon binding

a proton, whereas that of the latter will

decrease. For example, rf the above reaction is carried out in Tris (Trrs(hydroxymethyl)aminomethane; pK, = 8.06) buffer, it will be described by the eqmhbrium,

Conductimetric Measurement (CH20H)$NH,

159 (7)

+ H+ cs (CH20H),CNHf

the net reaction of which results in the formation of two additional ions: CH3COOCH2CH2N+(CH3)3 + Hz0 + (CH20H)$NH2 (CH&HOCH2CH2N+ + CH$OO- + (CH20H)$NH;

-+ (8)

If however, the reaction were carried out in tricine (N-tris(hydroxymethy1) methylglycine; pK, = 8.15): (CH,0H)3CNHCH2CO@

+ H+ f)

(CH20H)$NHCH2COOH

(9)

leading to a net reaction: CH3COOCH2CH2N+(CH3)3 + Hz0 + (CH20H)3CNHCH2C00(CH& HOCH$H,N+ + CH$OO-+ (CH20H)$NHCH2COOH

+ (10)

where neutralization of the ionized form of the buffer by the proton results in no net change in the number of ions present. In this case, there is a change in the size of the charge-carrying groups (mechanism 4), which may result in a net conductancechange owing to the differing mobilities of reactantsand products. The catalytic action of the enzyme hexokinase (E.C.2.7.1.1), which catalyzes the conversion of ATP to ADP, results m a conductivity change related to a number of the abovementioned factors (1,3-5). ATP4- + Glucose + ADP3- + Glc-6-P2- + Hf

(11)

Factors that influence conductivity in this reaction include the generation of an ionic molecule, glucose-6-phosphate,and the generation of a proton. Cleavage of a phosphate from ATP results in a change in the charge of the molecule, as well as its size, which affects mobility. Finally, magnesium ion, a divalent cation, chelates ATP and ADP. As a consequence of these factors, the conductance change accompanying the hexokinase-catalyzed conversion is a function of both the magnesium concentration and the buffer identity. In summary, there are a number of mechanisms by which an enzymatic reaction can lead to a conductivity change. The magnitude of the conductivity change, and to some extent the suitability of a particular enzyme for the construction of a conductimetric biosensor, depends on the reaction mechanism, the background conductivity, and the nature of the buffer used.

7.2. Conductivity Measurement Electrical conductivity measurement is one of the oldest methods of the analytical chemist. Conductimetric biosensors are based on the measurement of enzymatically catalyzed changes in conductivity, but the devices differ in important ways from the analytical chemist’s conductivity cell. The following discussion is intended to review the basics of conductivity measurement and

Sheppard and Guiseppi- Elie

160

illustrate these differences. More detailed discussions of the theory and practice of conductivity measurement can be found in Bard and Faulkner (2) and Kell and Davey (3). The transport of ions in solution, driven by an electric field, produces an electric current. The current density, J, expressed as current per unit area, IS directed along the electric field vector, E. For sufficiently small field strengths, the current density is duectly proportional to the magnitude of the electric field. As indicated in the expression below, the proportronahty constant is the electrical conductivity, (T,also referred to as the specific conductance, K. J=oE

(12)

The electrical conductivity is an intrinsic property of the electrolyte. The current carried by an electrolyttc solution depends on the concentratrons of ions present, c, then valences, z, and electrophorettc mobrlmes, p. The conductivity can be expressed as, (13)

where the sum is taken over the types of ions present and F is Faraday’s constant. The changes in conductivity that provide the transduction mechanism used in conductimetric biosensors result from changes in ion concentrations and mobilities accompanymg enzymatic catalysis. The electrical conductivtty of a solution 1smeasured by establishing a known electric field in the bulk of the solution and measuring the current as a result of migration of ions in the field. The electric field is typically established by applying a voltage between a pair of metal electrodes immersed m the solution. The resulting migration of ions to the electrodes may lead to electrode polarization, which reduces the electric field m the bulk of the solution and leads to errors in the measurement. To mmtmrze electrode polarization, an alternating voltage IS used to rapidly reverse the direction of the field before charging can occur. Electrode polarization is also minimized through the use of platinum black electrodes, formed by the electrodeposition of platinum on platmum or gold electrodes. This platimzation process increases the surface area of the electrode, and reduces electrode polarization by increasing the capacitance of the electrode. To ensure that the measured current reflects only the ionic conducttvtty of the solution, the amplitude of the voltage applied between the electrodes should be sufficiently small, on the order of 100 mV or less. The aim IS to prevent Faradaic reactions from occurring, so that the oxidatton or reduction of electroactive speciesat the electrodes does not contribute to the measured current. Figure 1 illustrates the important differences between a typical conductivity cell, used to measure solution conductivity, and a conductrmetrrc biosensor,

V

SigI.lalS

i

V

Fig. 1. Compawon

conductimetric biosensor

m\

conductivity cell

I

of conductivq

/lb

device circuit

cs = GK

G=l/R

equations

IYI =+ciG

cel1 to conductlmetnc hosensor.

equivalent

Sheppard and Guiseppi- Elie

162

designed to measure enzyme-catalyzed changes in conductivity. The electrodes of a conductivity cell are usually constructed of platimzed platinum to mimmize electrode polarizatton. Under these condttions, the applied voltage will produce an alternating current that is in phase with the voltage. To the mstrumentation supplying the voltage and measuring the current, the pair of electrodes and intervening electrolyte are electrically equivalent to a resistor. The ratio of the measured current to the applied voltage is known as the conductance, G. The conductance has units of Siemens, and IS the inverse of the electrical resistance. The electrical conductivity of the electrolyte is the product of the conductance and a geometrical factor known as the cell constant, K, which has units of inverse length, and is specific to a given set of electrodes. o=GK

(14)

For closely spaced, plane-parallel electrodes, the cell constant IS equal to the spacing between the electrodes divided by then area. The cell constants for other types of electrodes, such as coplanar interdigitated electrode arrays, are also functions of the electrode dimensions, but in general can only be calculated using sophisticated analyses or numerical methods (4). The cell constant of such an electrode pair may be determined experimentally using one or more standard solutrons of known conductivity. Conductrmetric biosensors are constructed by mnnobillzmg an enzyme m close proximity to a conductimetrlc transducer. A transducer conststs of a pair of electrodes, commonly in a coplanar arrangement, such as an interdigitated electrode array. However, unlike conduct&y cells, the electrodes are rarely platinized. Even when driven with an alternating voltage, electrode polarization results, and the current will lag behind the voltage. The conductimetric biosensor thus appears to the measuring circuitry as electrically equivalent to a resistor m parallel with a capacitor. (The sensor can also be represented by a series combinatron of a resistor and a capacitor, or more complicated equrvalent circuits in which the elements represent physical features of the system, such as, the bulk conductance, double-layer capacitance, and Faradaic resistance [2/.) The current can be described by its magnitude and its phase with respect to the applied voltage, and can be resolved mto a component in phase with the applied voltage, and a component 90” out of phase. The ratio of the amplitude of the in-phase component to the amplitude of the applied voltage 1s the conductance, whereas the ratio of the out-of-phase component to the applied voltage is the susceptance. The susceptance IS equal to the product of the capacitance, C, and the angular frequency, u). The behavior of this equivalent circuit is commonly described using a complex quantity known as the admittance, Y, the real part of which is the conductance, and the imaginary part being the susceptance.

Conductimetric Measurement

Top

View

163

Cross-section

Fig. 2. Schematicillustration of a conductimetrlcurea sensor,using a planar mterdigitated electrode array as the conductimetrictransducer. Y=G+joC

(15) Depending on the type of measurement instrumentation used (see Note 5), the investigator may be presented with the magnitude (and possibly phase) of the current, or the admittance of the sensor computed from the voltage and current. With but one exception (5), investigators to date have not sought a quantitative determination of conductivity, but have used the admittance or the current as a signal that reflects the catalytic activity of the enzyme. To first order, these measures provide kinetic information equivalent to that which would be obtained from the conductivity, since the electrode geometry and therefore the cell constant (K in Eq. 14) remains fixed. 1.3. Enzyme-Based Conductimetric Sensors The catalytic reaction mechanisms described above in Subheading 1.1. can be exploited to construct conductimetric enzyme sensors. In their simplest form, these devices consist of a conductimetric transducer onto which the enzyme of interest is immobilized. A typical sensor design is illustrated schematically in Fig. 2. The conductimetric transducer is a set of planar mterdtgitated electrodes, patterned on an insulating substrate using microelectronic or printed-circuit fabrication technology. The electrodes are typically prepared from a noble metal, such as gold or platinum. The electrodes are coated with a sensing layer containing one or more immobilized enzymes. The enzyme may be immobilized to the surface of the device using a number of methods. Reticulation, the crosslinkmg of an enzyme or enzyme-protein solution using glutaraldehyde, has been most commonly used (6-13). Other approaches include direct covalent attachment to the silica surface of microsensors (5), and entrapment of enzyme within electropolymerized films (14-17). Conductimetric enzyme biosensors have been constructed for a number of analytes. Sensors for urea, based on the urease-catalyzed hydrolysis of urea, have been studied by a number of investigators, because of the relatively large

164

Sheppard and Guiseppi- Elie

conductance change resulting from the conversion of a neutral molecule into ionic products (5-11,18,19). These sensors have been applied to the clinical analysis of urine samples (7), and to the on-line monitoring of the urea concentration m spent dialysate during hemodialysis (11). Sensors for glucose based on crosslinked glucose oxidase sensing layers (19,20) have been constructed, as have been devices in which the enzyme was immobilized in a conducting polymer (14). Sensors for other analytes, such as, penicillm (15), ammo acids (8), L-asparagine (7), creatinine (7), and acetylcholine- and butylcholine chlorides (21), have been reported. Conductimetric enzyme sensors may be operated in a kinetic mode or at steady state. When first exposed to a sample contaming the analyte of interest, the catalytic action of the immobilized enzyme leads to the formation of reaction products. These products accumulate within the sensing layer, and through one or more of the mechanisms described m Subheading l.l., produce a change m the conductance of the sensor. The accumulation of products at the surface of the device is accompanied by diffusion of products away from the sensor surface. Eventually, a steady state is reached where the rate of generation of products balances the diffusive flux away from the sensor surface. In kinetic operation, the initial rate of change of the measured current (or conductance) is used to provide a measureof the reaction rate, and therefore the substrate concentration. This approach is most common when the sensor is used in a system, such as a flow-injection analysis system, for consecutive assays of multiple samples. Alternatively, the steady-state conductance of a conducttmetric biosensor also provides a measure of the analyte concentration, and is used where continuous monitoring is desired Shul’ga et al. (19) presented a comparison of the kinetic and steady-state responses of conductimetric sensors for urea and glucose, and found that the kinetic response was less sensitive to differences in the buffer capacity of the sample. 1.4. Major Procedures The major procedures involved in the construction of a conductimetric enzyme sensor include preparation of the conductimetric transducer; immobilization of the enzyme to form the sensing layer; and testing of the sensor. The procedures for the design and fabrication of conductimetric transducers based on microfabricated (6,7,9,10,12,13) and screen-printed electrodes (11) used in recent publications are beyond the scope of the chapter. The interested reader is referred to the text by Lambrechts and Sansen (22) and references therein. The conductimetric transducers used for the protocol are commercially available, microfabricated, interdigitated electrode arrays (Fig. 3). To prepare the probe, the silica surface of the interdigitated electrode array is treated with an aminosilane to improve adhesion of the immobilized enzyme sensing layer.

Conductimetric

Measurement

165

Fig. 3. Commercially available conductimetric transducers (photo courtesy of ABTECH Scientific).

Proceduresfor the deposition of platinum black on the electrodesare provided. A number of methods have been used for immobilizing enzymes to conductimetric sensors; this protocol makes use of reticulation, a procedure that is widely used for the construction of enzyme-basedbiosensors(23). The testing of the sensorrequires instrumentation to measurethe current resulting from an applied AC voltage, or equivalently, the electrical admittance of the sensor. The use of an impedanceanalyzer or a potentiostat/lock-in amplifier combination for thesemeasurementswill be described. 2. Materials 2.1. Electrodes/Electrode

Preparation

1. Planar interdigitated electrode arrays (IME- 1550FD-Pt, ABTECH Scientific, Yardley, PA) (seeNote 1). 2. 3Aminopropyl trimethoxy silane. 3a. Platinizing solution, commercially prepared (YSI #3 140, Yellow Springs Instrument Co., Yellow Springs,OH), or

166

Sheppard and Guiseppi- Elie

3b 3 g of Platinum (IV) chloride (hydrogen hexachloroplatinate (IV) hydrate; FW 409.82), 1.25 mg lead acetate (to saturation) in 100 mL of 0 01 MHCl at 25°C

2.2. Enzyme Immobilization 1. 2. 3. 4.

Enzyme: Urease E C.3.5.1.5 (Sigma [St. Louis, MO] Type IX, 80,000 U/mg). Bovine serum albumin (BSA) (Sigma, Fraction V) Glutaraldehyde, 25% aqueous solution (Sigma). 10 mA4 Tris-HCl, pH 7.5, buffer (buffer A).

2.3. Sensor Evaluation 1 0.5 MUrea solution m 10 mMTns-HCl,

3. Methods 3.1. Electrode

pH 7.5 (stock solution).

Preparation

1. Clean the conductimetric transducer by sequential washing in acetone, isopropanol, and water, as follows: a. Somcate the conductimetric transducer in acetone for 60 s. In this and subsequent steps, suspend the device in the solvent such that the solvent does not contact the plastic of the probe. b. Sonicate in 2-propanol for 60 s. c Sonicate in deionized water for 60 s 2 Silanize the surface with aminosilane (see Note 2) a. Prepare a 2% sllane solution by addmg 400 pL of 3-ammopropyl tnmethoxysilane to 20 mL of a solution of 95% ethanolR% water. b Immerse the conductimetric transducer into the silane solution for 10 mm, at room temperature c Rinse thoroughly with ethanol d Place the transducer in an oven for 10 mm at 110°C to cure the silane e. Repeat the solvent cleaning procedure above (step la-c). 3. Deposit platinum black on the mterdigitated electrode array (optional, see Note 3). a. Electrically connect the mterdlgltated electrode arrays on the transducer by twisting the ends of the four insulated leads together. Connect this as the working electrode of a three-electrode electrochemical cell with a platinum mesh counter electrode and Ag/AgCl reference electrode. b. Cycle the interdigitated array electrodes from -1 to -2 V with respect to the Ag/AgCl reference electrode for 3 min m phosphate-buffered saline. The purpose of this cathodic cleaning is to remove any aminosilane bound to the electrodes. c. Rinse the transducer thoroughly in deionized water, and reassemble the electrochemlcal cell in the platmlzmg solution. d. Use a galvanostat to pass a reducing current of 6 mA (equivalent to a current density of 40 mA/cm2) for 20 s. If only a potentlostat is available, platmlzatlon will occur at a potential of approx -0.1 V vs Ag/AgCi reference. Startmg

Conciuctimetric Measurement

167 5mM

2mM

1 mM

30

60 90 Time(seconds)

120

Fig. 4. Conductance, measured at 1 kHz, of a conducttmetric urea sensor after immersion in urea solutions of 1,2, and 5 mm urea, prepared in 10 mA4 Tris buffer. from this value, manually adjust the potential to reach the desired current for the specified time. e. Inspect the mterdtgttated electrode arrays under a microscope for uniform grayblack coating on electrodes.

3.2. Enzyme Immobilization 1. 2. 3. 4.

Prepare 1 mL of 150 mg/mL solutton of urease m buffer A (Solution A). Prepare 1 mL of 150 mg/mL solution of BSA in buffer A (Solution B). Prepare 1 mL of a 5% (v/v) solution of glutaraldehyde m buffer A (Solution C). Combine 100 pL of solution A, 100 uL of B, and 50 uL of C, and stir until well mrxed, trymg not to introduce air bubbles mto the mixture 5. Supporting the electrode horizontally, paint a thin (~1 mm) film of the enzyme/ BSA/glutaraldehyde mixture over the active area of the interdigitated electrode array using a nylon brush or cotton swab. The mixture should be fluid enough such that it spreads into a uniformly thin film (see Note 4). After the film has gelled, immerse and store the electrode in buffer A.

3.3. Sensor Evaluation 1. Fill a magnetically stirred, thermostatted vessel with 50 mL of buffer A. Imtiate moderate stirring, and maintain the temperature at the desired value (e.g., 37°C) Immerse the sensor in the buffer solution and secure with a clamp. 2. Program the impedance analyzer to measure conductance at a frequency of 10 kHz, with a lOO-mV excitation amplitude. (see Note 5 for a discussion of other instrumentation systems for use with the sensor.) 3 Connect the leads from the electrodes to the impedance analyzer. 4. After mtttatmg the data recording system (strip chart, computer), add an ahquot of concentrated urea stock solution to bring the urea concentratron m the vessel to the desired concentration. The conductance (or current) should rise as ptctured

168

Sheppard and Guiseppi-Elie

in Fig. 4, approaching a steady state at a time ranging from 1 to 10 min or more, depending on the thickness of the enzyme layer. 5. Additional aliquots of urea may be added to produce a stepwtse increase in the steady state response traceable to increases in the urea concentration. 6. After completion of the tests, the sensor should be removed from the vessel, rinsed thoroughly with buffer A, and stored in Tris buffer at 4°C

4. Notes 4.7. Electrode Preparation 1. Electrode fabrtcatton. The commercially available platinum mterdtgltated electrode arrays specified for use in this procedure provtde a convenient probe for constructing conductimetric biosensors. Readers interested m custom fabrication of mterdigitated electrode arrays for use in conductimetric blosensors may wish to refer to discussions of the design constderations of these devices (410). As a lower cost alternative to microfabricated conductimetric transducers, Mikkelsen and Rechmtz (8) describe the construction of a simpler device formed by embedding platinum wire m a silicone rubber matrix. 2. Srlamzation refers to the use of a stlane couplmg agent to covalently attach a primary amine fnncttonahty to the silica substrate of the planar interdlgttated electrode array. The reactton of glutaraldehyde with these primary amines during the reticulation (rmmobilization) step leads to covalent attachment of the crosslmked enzyme gel to the sensor substrate, improving the adhesion of the sensing layer and the lifetime of the sensor The chemistry of the coupling of 3aminopropyl trlmethoxy sllane (APTS) to the silica surface of the conductimetrlc transducer is illustrated schemattcally m Fig. 5 The methoxysllane linkages of the APTS are first hydrolyzed to silanols, which react with silanols on the silica surface of the conductlmetrtc transducer substrate, as well as with those of other ammostlanes. These reactions lead to the formatton of a crosslmked network of srlanes containing primary amines, covalently attached to the transducer surface Interested readers may wish to consult Plueddemann (24) or Allara (25) for further details on the chemistry of silane coupling agents. 3. Platinization. The purpose of platmrzing the electrodes is to reduce the mterfactal impedance between the electrodes and the crosslmked enzyme gel. A reduced interfacial impedance will increase the sensitivity of the measurement, improve the reproducibthty, and facilitate quantitative analysis of device operation. However, because some care is required to prevent shorting of the electrodes while platmrzing the mrcrofabricated interdtgttated electrode arrays, this step should be considered optional. The current and time specified m step 3.1.3.d in Subheading 3.1. are those recommended for the ABTECH 1550-FD-Pt electrode array. The use of other electrodes will require calculation of the current and time, using the guidelines that follow. The reduction of electrode impedance achieved when platmizing 1s directly related to the total amount of platinum electrodeposited. Recalling Faraday’s law, this is proportional to the charge passed, which is equal to the

Conciuctimetric Measurement

169

product of the current and the electrodeposition time. A total charge density of 0 5 A-s/cm2 IS considered a light deposit of platinum black, whereas a value of 50 A-s/cm2 is considered heavy (26). A value of 10 A-s/cm2 is recommended by Kell and Davey (3) to provide a balance between reduced impedance and mechamcal stability of the deposit. The charge density specified in this protocol for the IME 1550-FD-Pt device is approx 0.8 A-s/cm2. This conservative value was chosen to prevent shorting of the interdigitated electrode array by the deposited platinum. The nature of the platinum black deposit depends on the rate at which it is deposited, which is proportional to the current density. A current density of 10 mA/cm2 is typically used for platinization (3), although somewhat higher current densittes may be needed for electrodes with lateral dimensions ~1 mm (26). Satisfactory platinization of the IME 1550-FD-Pt device has been achieved using a current density of 40 mA/cm2.

4.2. Immobilization 4. Reticulation. The most challenging procedure in this protocol is the preparation of the sensing layer by cocrosslinking the enzyme and albumin on the surface of the conductimetric transducer The experience of the authors, and that of other investigators (3, is that there can be considerable variability m the rate at which the crosslinkmg proceeds to form a gel, most hkely caused by variations m the reactivity of the glutaraldehyde. As such, the recommended amount of glutaraldehyde solution (solution C) in the formulation should be considered as a starting point. The recipe should be adjusted by using greater or lesser amounts of solution C to lead to gelation after approx 5 mm. This provides a sufficient time for mixing and coating of the electrode array. Crosslinking that is too rapid leads to thick and nonuniform films of enzyme. The sensing layer can be removed should the quality of the immobilized enzyme layer appear poor (e.g., too great a thickness, bubbles) or for the construction of a new sensor. The tirst step is to mechanically remove the bulk of the reticulated protein gel layer using a soft bristle brush or a moistened paper towel Next, the transducer should be sonicated for 2 min in detergent. Note that both of these procedures may remove the platinization from the platinized digits of the transducer, and so may necessitate replatimzation. Furthermore, the platmum particles may slough off and become trapped within the interdigitated space. If the particle is lodged so as to cause a short, the transducer will need to be discarded The final step in reconditioning a conductimetric transducer is the removal of traces of covalently bound organics (if ammo silane is used) or adsorbed polymer or protein, Immerse the transducer in 50 mL of 4: 1 H2S04:H202 for 5 mm, then rinse thoroughly m deionized water. The preparation of the sensor can be mimediately restarted at the silanization step (step 2, Subheading 3.1.).

4.3. Instrumentation 5. Impedance analyzers. Two types of instrumentation systems have been used to interface conductrmetric biosensors (the choice of which appears to depend on

Sheppard and Guiseppi-Elie

170 NH3

CH30

) -Fi-

-3 CH30H OCH3

P

Fig. 5. Schematic illustration of the reaction leading to covalent coupling of 3aminopropyl trimethoxy silane to oxidized silicon surface. whether the investigator is an engineer or a chemist!) The first of these are impedance analyzers, commonly used to measure the circuit properties of electronic components. The second type of mstrumentation system is a potentiostat equipped for electrochemical-impedance spectroscopy. The former is more convenient to use, although the latter may be more readily available to the reader These mstrumentation systems are illustrated schematically m Fig. 5, and briefly described below Impedance analyzers (Fig. 6A) measure the electrical impedance of devices by applying a smusoidal voltage to the terminals of the device/sensor and measuring the amplitude and relative phase of the resultmg current. Automated analyzers resolve the in-phase and out-of-phase components of the current and, given the users selection of an equivalent circuit representation, compute the specified parameters (e.g., conductance and susceptance). A second type of instrumentation system used to interface conductimetric biosensors is an electrochemical-impedance spectroscopy system. These are constructed around a potentiostat having the capability of adding an externally applied signal to the working electrode potential and of providing an output of the working electrode current. As sketched m Fig. 6B, a signal generator is used to supply a small amplitude sinusoidal voltage, typically at an audio frequency, to this external input. The external current output of the potentiostat, which takes the form of a smusoidal voltage, is connected to the sample input of a lock-m amplifier (the output of the signal generator is connected to the reference input) Typically, the lock-m amplifier is set up to measure the m-phase (0’) and out-ofphase (90”) components of the signal, from which an equivalent resistance and capacitance can be calculated. An equivalent approach, taken by Mikkelsen and Rechnitz (8), was to directly measure the magnitude of the current by adjusting the phase angle of the lock-in amplifier so as to obtain the maximum output,

171

Conductimetric Measurement Impedance

Analyzer

sensor

Signal Generator Wavetek 187

Potentiostat EG&G PAR 362 sensor

I

Lock-in Amplifier EG&G PAR 5101

I

C

sensor

Fig. 6. Instrumentation systemsused for interrogation of enzyme-basedconductimetric sensors.(A) impedanceanalyzer; (B,C) electrochemical-impedancespectroscopy systems. thereby matching the phase angle of the current. Alternatively, some electrochemical-impedancespectroscopysystems(Fig. 6C) usespecial-purposeinstrumentation, suchasthe SolartronFrequencyResponseAnalyzer, which combines the signal generatorand lock-in detectioncircuitry in a single chassis.

Acknowledgments The authors wish to thank Matthew Lesho and Timothy Hendricks for their assistance. This work was supported in part by a National Science Foundation Presidential Young Investigator Award to NFS (ECS-9058419). A. G. E. thanks ABTECH Scientific, Inc. for support.

172

Sheppard and Guiseppi-Elie

References 1 Lawrence, A. J. (1971) Conductimetrrc enzyme assays. Eur. J Btochem. 18,221. 2. Bard, A. J. and Faulkner, L. R. (1980) Electrochemtcal Methods* Fundamentals and Applicattons, Wiley, New York. 3 Kell, D. B. and Davey, C. L. (1990) Conductimetric and impedimetric devices, m Bzosensors* A Practzcal Approach (Cass, A. E. G., ed ), JRL Press at Oxford Umversity Press, London, pp. 271. 4. Sheppard N. F , Jr., Tucker, R. C., and Wu, C. (1993) Electrical conductivity measurements using microfabricated interdigited electrodes Anal Chem 65, 1199. 5 Sheppard, N. F , Jr , Mears, D. J , and Guiseppi-Elie, A (1996) Model of an imrnobihzed enzyme conductimetric urea biosensor. Btosens Bioelec 11, 18-24 6. Watson, L. D., Maynard, P., Cullen, D. C., Sethi, R. S , Brettle, J., and Lowe, C. R (1988) A microelectromc conductimetric biosensor Biosensors 3, 10 1. 7. Cullen, D. C., Sethi, R. S , and Lowe, C. R. (1990) Multianalyte miniature conductance biosensor Anal Chim. Acta 231,33 8 Mikkelsen, S R. and Rechnitz, G. A. (1989) Conductometric transducers for enzyme-based biosensors. Anal. Chem. 61, 1737. 9. Pethig, R. (1991) Dielectric-based biosensor Biochem. Sot Trans 19,21 10. Jacobs, P., Suls, J., and Sansen, W. (1994) Performance of a planar differentialconductivity sensor for urea. Sens Act B 20, 193 11. Jacobs, P., Sansen, W., and Hombrouckx, R. (1994) Continuous monitoring of the spent dialyzate urea level using a disposable biosensor Clinical evaluation, ASAIO J 40, M393. 12. Dzydevich, S. V., Shul’ga, A. A., Soldatkin, A. P., Hendji, A. M., JaffrezicRenault, N. and Martelet, C (1994) Conductometric biosensors based on cholinesterases for sensitive detection of pesticides. Electroanalysts 6,752 13. Hendji, A. M. N., Jaffrezic-Renault, N., Martelet, C., Shul’ga, A. A., Dzydevich, S. V , Soldatkin, A P., and El’skaya, A. V. (1994) Enzyme biosensor based on a micromachmed interdigttated conductometric transducer: application to the detection of urea, glucose, acetyl- and butyrylcholme chlorides, Sens Act B B21, 123.

14 Hoa, D. T., Kumar, T. N S., Punekar, N. S., Srinivasa, R S., Lal, R., and Contractor, A Q. (1992) Biosensor based on conducting polymers. Anal Chem 64,2645 15 Nishizawa, M., Matsue, T., and Uchida, I. (1992) Pemcillin sensor based on a microarray electrode coated with pH-responsive polypyrrole Anal Chem 64,2642. 16 Guiseppi-Elie, A., Tour, J M., Allara, D. L., and Sheppard, N F., Jr. (1996) Bioactive polypyrrole thm films with conductimetric response to analyte, m Electrtcal, Optical and Magnettc Properties of Organtc Solrd-State Matertals (Dalton, L., Jen, A. K. Y , Wnek, G. E , Rubner, M. F , Lee, C Y -C., and Chiang, L. Y., eds.), Materials Research Society, Pittsburgh, PA, pp. 439. 17. Guiseppi-Elie, A., Wallace, G. G., and Matsue, T. (1996) Chemical and biological sensors based on electrically conducting polymers, in Handbook of Conductive Polymers, 2nd ed. (Skotheim, T., Elsenbaumer, R., and Reynolds, J. R., eds.), Marcel Dekker, New York, pp. 129-143.

Conductimetric Measurement

173

18. Lawton, B. A , Lu, H., Pethig, R., and Wei, Y. (1989) Physlco-chemical studies of the actvlty of urease and the development of a conductimetric urea sensor. J. Mol Liquids 42,83. 19. Shul’ga, A. A., Soldatikm, A. P., El’skaya, A V., Dzyadevich, S. V , Patskovsky, S. V., and St&ha, V I (1994) Thin-film conductometric biosensors for glucose and urea determmation. Bzosens. Bioelec. 9,2 17. 20. Soldatkin, A. P., El’skaya, A. V., Shul’ga, A. A., Jdanova, A S., Dzyadevich, S. V., Jaffrezic-Renault, N., Martelet, C., and Clechet, P. (1994) Glucose-sensitive conductometric biosensor with additional Naiion membrane: reduction of influence of buffer capacity on the sensor response and extension of its dynamic range. Anal. Chum. Acta 288, 197. 21. Nyamsi-Hendjii, A M , Jaffrezic-Renault, N , Martelet, C., Shul’ga, A A., Dzydevich, S. V., Soldatkin, A P , and El’skaya, A. V. (1994) Enzyme biosensor based on a micromachined interdigitated conductometric transducer application to the detection of urea, glucose, acetyl- and butylcholme chlorides Sens Act B 21, 123 22 Lambrechts, M. and Sansen, W. (1992) Biosensorsa Mwroelectrochemlcal Devices, Instnute of Physics Publishing, New York. 23. Wilson, G. S., and Thevenot, D. R. (1990) Unmediated amperometrtc enzyme electrodes, in Blosensors. A Practical Approach (Cass, A E. G., ed.), IRL, New York, pp. 287-3 19. 24. Plueddemann, E. P. (199 1) &lane Coupling Agents, 2nd ed., Plenum, New York. 25. Allara, D. L. (1995) Crmcal issues in applications of self-assembled monolayers. Blosens. Bioelec lo,77 1. 26. Geddes, L. A. and Baker, L E. (1989) Principles of Applied Blomedlcal Instrumentatlon, 3rd ed , Wiley-Interscience, New York

13 Enzyme Biosensors Based on Thermal Transducer/Thermistor Kumaran

Ramanathan,

Masoud Khayyami, and Bengt Danielsson

1. Introduction 1.1. Calorimetric Biosensor Calorimetry is the measurement of heat accompanying a chemical or biochemical reaction (1,2). The heat changes could be endothermic (absorption of heat) or exothermic (evolution of heat). In the early 1950s the technique of microcalorimetry was devised to measure the heat accompanymg a biochemical reaction (3). After the initial elucidation of the concept, scanning microcalorimeters were designed to follow the heat changes accompanymg folding and unfolding of protein molecules. The conventional microcalorimeters used sensitive thermocouples as the measurement device whereas the modern calorimeters, designed in the early 1970s; used thermistors, with a temperature sensitivity of 1W5-106 “C. The evolution of calorimetric sensors(45) included heat conduction, isoperibol, and more recently, isothermal calorimetry. The unique advantage of isothermal calorimetry involving thermistors is the fact that the heat evolved or absorbed during a biochemical reaction is a direct measure of the reaction, In the recent past, the concept has been extended to the design of miniaturized thermistor-based calorimeters, 50 mm in length and 15 mm in diameter or smaller. The thermistor-based calorimeters popularly known as “enzyme thermistors” (ET) were designed to cover a broader range of applications (6), which included determination of metabolites, monitoring of enzyme reactions, bioprocess monitoring, characterization of immobilized biocatalysts, bioseparations, reactions in nonaqueous medium, environmental control, and more recently investigation of ionic interactions. From

Methods m E/otechnology, Vol 6. Enzyme and Mfcrobial Biosensors Techniques EdWd by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

775

and Protocols NJ

Ramanathan, Khayyami, and Danielsson

176

Table 1 Linear Concentration Ranges of Substances with Calorimetric Sensors Using Immobilized Analyte

Enzyme(s) used

Ascorbic acid ATP (or ADP) Cellobiose Cholesterol Creatinine Ethanol Glucose Glucose

Ascorbate oxtdase Pyruvate kmase + hexokinase @Glucosidase + glucose oxtdaselcatalase Cholesterol oxidase/catalase Creatinine lmmohydrolase Alcohol oxidase Hexokmase Glucose oxidase/catalase

L-Lactate L-Lactate L-Lactate (or pyruvate) Oxalate Penicillin G Pyruvate Sucrose Urea

Lactate-2-monooxygenase Lactate oxidaselcatalase Lactate oxidaselcatalase + lactate dehydrogenase Oxalate oxidase P-Lactamase Lactate dehydrogenase Invertase Urease

Measured Enzymes Linear range W4 0.0 l-O.6 10 nA4-a 0.05-5 0.01-3 0.0 l-10 0 0005-l 0.01-25 0 0002-l O.OOl-75c 0.005-2 0.0002-1 10 lllwa 0 005-0.5 0.002-200 0.01-10 0.05-100 0.005-200

Enthalpy change (-kJ/mol)

53 + 100

28 (759 80 + 100

ca 225

67 (115b) 47 (159 61

aWlth substrate recycling % Trls buffer CWlth benzoqumone as electron acceptor

The metabolites determined include alcohols, glucose, and lactate, using alcohol oxidase, glucose oxidase, and lactate oxidase, respectively. The detection limit for these compounds is in the submicromolar range m pure solutions (Table 1). The normal procedure IS to coimmobilize the oxrdases with catalase to increase total heat production, reduce oxygen consumptton, and eliminate hydrogen peroxide. Cellobiose has been measured by using P-glucosidase m combination with glucose oxidase and catalase Determination of cholesterol and cholesterol esters has been accomplished using cholesterol oxidase and cholesterol

esterase, and triglycerides

by using Iipase. Typical

sample matrices

are blood, serum, and fermentation broth. Oxalate in urine has also been measured using oxalate oxidase (7). Substrate recycling offers a route for highly sensitive measurements. An example is lactate (or pyruvate) determination with a coimmobtlized lactate dehydrogenase/lactate oxidase (LDHILOD) column that repeatedly oxidizes lactate to pyruvate and reduces pyruvate to lactate.

Thermal Transducerflhermistor

177

Each cycle produces considerable amount of heat. A similar approach has been employed for determination of nicotinamide adenine dinucleotide/methemoglobin reductase screening LDH (NAD+/NADH) by coenzyme recycling using plus glucose-6-phosphate dehydrogenase and ATP/ADP by coupling pyruvate kmase with hexokinase as the recycling enzymes (6). By operating at excessive glucose supply, a hexokinase column was used for indirect assay of ATP within micromolar sensitivity. For btoprocess monitoring the ET has been employed for the assay of penicillin m fermentation broth using p-lactamase or penicillin acylase for the reaction, Also alcohol generated in alcohol fermentation using yeast has been monitored using alcohol oxidase for monitoring ethanol. Other metabolites that have been monitored during fermentations include lactate, glycerol, acetaldehyde, sucrose, and glutamme. The characterization of nnrnobil~zed mvertase has been carried out and the technique was successfully coupled to the catalytic activity determination m immobilized cells. Similarly the results of this technique were useful in selection of Trigonopsis variabilis strains for high cephalosporin-transforming activity. Also, the cephalosporin-transforming activity of o-amino acid oxidase isolated from yeast was identified in a similar manner. The thermometric signal was proportional to the number of cells as well as the amount of n-amino acid oxidase immobilized in the ET microcolumn. The ET has also been coupled to a thermometric enzyme-linked immunosorbent assay(ELISA) procedure (TELISA) for the determination of hormones, antibodies, and other biomolecules generated during the fermentation process(seeChapter 2). Predominantly alkaline phosphatasehas been used as the enzyme label for such an assay. Genetically engineered enzymeconjugates, e.g.,human proinsultilkalme phosphatase conjugate, were used for the determination of insulin or proinsulin. Monitoring of specific proteins eluting out of chromatographtc columns has been demonstratedusing the ET as a direct on-line monitor for purification of proteins or enzymes.As an example,LDH was recovered from a solutton by affinity bindmg to N6-(6-aminohexyl)-AMP-sepharose gel andthe signal from the ET was usedto regulate the addition of the AMP-Sepharose suspensionto the LDH solution. Significant interest has also been generated in monitoring enzyme activity in nonaqueous solvents. For example, the reactions of immobilized lipoprotein lipase with tributyrin in a buffer-detergent system and cyclohexane have been compared using an ET. Also it was demonstrated that horse radish peroxidase produced a considerably higher temperature signal m toluene than in water. Furthermore, addition of dtethyl ether m small amounts was found to enhance this effect. In an analogous approach the reaction of chymotrypsin m 10% dimethyl formamide (DMF) for hydrolysis (exothermic) and synthesis (endothermic) of peptide bonds has been monitored using the ET.

178

Ramanathan, Khayyami, and Danielsson

Furthermore, the ET has also been successfully applied to the monitoring of heavy metal (Hg 2+,Cu2+, Co2+, Zn2+, and Ag+) toxicity in the environment by measuring the inhibition of urease activity down to ppb levels of the metal ions. The restoration of the activity was also tested on chelation of the metal ions using strong chelating agents. More recently it has been demonstrated that the thermistor approach can be used to monitor specific interactions of fluoride ions with silica-packed columns in the flow-injection mode. 1.2. Construction of the ET The overall detection system consists of two components, i.e., the ET calorimeter and the flow-injection system (FIA). The earlier versions of the ET were made of Plexiglas and later it was modified by using metal blocks for temperature control Instead of water baths m the former models (Fig. 1). Essentially the ET consistsof a plastic column, which can hold varying amounts of the enzyme depending on the volume of the column (usually 100 IU per column). 2.10. Glutaraldehyde Glutaraldehyde tends to polymerize on storage and should be distilled before use for coupling or stored in small portions in the freezer. Purification of glutaraldehyde can also be carried out by using the following steps: 1 2. 3 4. 5.

Mix 700 uL of 25% glutaraldehyde with 1 mL cooled activated charcoal. Shake the mixture vigorously several times durmg 10 mm. Centrifuge at 1OOOgfor 5 mm and collect the supematant in a 1-mL Eppendorftube. Repeat at least four times. Take 50 nL glutaraldehyde from step 4 in a quartz cuvet and dilute with 950 pL of 0.1 A4 phosphate buffer, pH 7.0.

6. Measurethe absorbanceat 280 nm (a) and 235 nm (b) using phosphatebuffer as reference. The ratio b/a should be ~0.23.

2. Il. Buffers The nature and type of the buffer could vary depending on the type of analyte being detected and the enzyme reaction(s). A common buffer is 0.1 A4 phosphate buffer, pH 7.0-7.5. If a proteolytic reaction is connected with the enzymatic reaction, the sensitivity of the measurements can be increased by using buffers with high protonation

enthalpy (such as Tris buffer). The major limlt-

Ramanathan, Khayyami, and Danielsson

182

ing factor for column life is usually mechanical obstruction. It is therefore a good practice to filter the solutions used, as well as the samples,through a l-5-~ filter and to prevent microbial growth in the solutions by some bacterlostatlc agent, for example sodium azide (NaN3) (see Note 3). 3. Methods 3.7. Immobilization

of the Enzyme

1 To 1 mL of sedimented y-amino-propyltriethoxysilanized and glutaraldehyde activated CPG with a mesh size of 4Q-80 and mean pore diameter 550 A, add 4 mg or 1000 U of a suitable enzyme 2. Usually unmobtlization is allowed to proceed overnight at 4°C with gentle mixing 3. Wash the preparation extensively on a glass filter with 0 1 A4 phosphate buffer, pH 7 0. This preparation is stable for several months For increased stabrhty the Schtff s bonds between the enzyme and the carrier can be reduced with sodium borohydride

3.2. Reversible Immobilization In some situations reversible immobilization may be conventent, for instance if the enzyme is very unstable or in inhibition studies where there is an irreversible inhibition of the enzyme.The reversible immobilization canbe accomplishedusing antibodies to bind the enzyme or, if the enzyme is a glycoprotein which is most often the case, by using a lectin (concanavalm A). Spent enzyme can stmply be washed out with glycme solution at pH 2.2 and fresh enzyme added in situ. 3.3. Packing the Column The upper end of the plastic column is connected to a pump and filled with buffer, and the mnnobthzed enzyme preparation suspended in buffer 1sadded to the column in small volumes with a Pasteur pipet. It is practical to make an extension of the column with a piece of tubmg of the same diameter as the column. During packmg excess buffer is sucked out with the pump and the column gently tapped to make the carrier material settle properly and avoid cavities m the column packing (see Note 4) 2. After filling the column with immobilized biocatalyst and attaching the filter, rt is inserted mto the apparatus and the buffer 1scrrculated through the system to equilibrate the column

3.4. Enzyme Activity Measurement To characterize the column the activity of the enzyme before and after the nnmobilization can be measured. For example, if the enzyme is LDH the following procedure can be used: 1 The enzyme obtamed from commercial sources may be a suspension of the enzyme m ammonmm sulfate. If so, it has to be purified by dialysis or gel tiltration against 0 1 Mphosphate buffer, pH 6 8.

Thermal Transducer/Thermistor

183

2. For determmatlon of LDH activity a stock solution containing 0.2 mMNADH and 1 Mpyruvate in 0.1 Mphosphate buffer, pH 6.8, is prepared and stored in the dark because NADH is light sensitive and unstable at pH 7.0 if stored for long. 3. Take 3000 pL of the solution prepared in step 2 in a quartz cuvet and monitor the absorbance at 340 run 4. Add 30 pL of the LDH solution and record the change m absorbance at 340 nm for 10 min. The decrease in absorbance is caused by the consumption of NADH during the formation of lactate from pyruvate and provides a measure of the activity of LDH in 30 yL of LDH in 3.03 mL. Similarly, the protein content of the enzyme solution could be determined by using the classical Folin-Lowry technique. 5. Perform a similar analysis of the immobilized LDH by taking a known weight of the preparation and incubatmg the NADH/pyruvate solution in a stirred cuvet or circulating the reaction solution with a pump through a flow through cuvet in a closed system. The activity could be expressed per milligrams of the CPG/silica Alternatively, the remaining activity or protein in the solution left after immobllization can be determined and subtracted from the initial value. Obviously, any of these techniques used for comparing bound enzyme with enzyme in solution cannot give an exact result, but are useful in comparing different immobilization preparations and procedures.

3.5. Injection

of the Sample

Samples are normally injected with a chromatographic tion valve with a 25-500~PL sample loop.

type of sample mjec-

1. Place the injection knob on the injection valve in the load position. Aspirate enough sample with the syringe attached to the loop to completely fill the mjection loop with some margin (see Note 5). 2. Turn the knob to the injection position. This will permit the circulation buffer to flow through the loop and carry the analyte m the flow stream to the reactor. 3. When the recorder pen 1sreturning back to the baseline the next mJectlon can be made. In some arrangements an automatic sample injection valve is used that 1s operated with a timer or a computer. To further increase the flexibility of the system a selector valve can be placed before the injection valve to select different sample or standard solutions.

3.6. Recording the Signal The output of the instrument in the form of a temperature peak can be recorded at a suitable sensitivity on the amplifier and recorder depending on the magnitude of the signal. An appropriate set of standard solutions with known concentrations is injected to construct a calibration curve. Alternatively, a chromatography integrator or a computer can be used to calculate the sample concentration after calibration. The linear range depends on the sample volume and can be as wide as 1 pM-1 M if no limitations in reactant concentrations occur. Oxidase reactions in which

Ramanathan, Khayyami, and Danielsson

184

oxygen is used as an electron acceptor are limited by the oxygen concentration and the operating concentration range is only linear up to 1mA4at large sample volumes (see Note 6). 3.7. Sensor Evaluation 1. Linear range/peak height: The thermograms can be evaluated by peak hetght or peak area determination. This could be performed manually or by use of an mtegrator or computer This enables close to an end-point determination. The linear range obtained is an mdication of the actual activity of the enzyme column. 2. Sensitivity* Within the linear range the sensitivity is rather independent of the activity of the enzyme column, but is controlled by the injected sample volume The highest sensitivity is obtained at steady state, i.e., at continuous sample introduction Smaller sample volume increases the linear range The sensitivity can be increased by adding sequentially operating enzymes or by usmg coenzyme or substrate recycling. The most sensitive, useful temperature range is hmrted by fluctuation of the signal because of friction and turbulence withm the column. Typically 4-2 x lti K is contributed by such nonspecific effects 3. Stability: a. Using FIA technique with repeated injections stable recordings and deternnnations can be made for several weeks with daily calibrations. With a specific enzyme good operational stabihty is normally achieved with little change m the performance for more than thousands of samples or several months of continuous experimentation. With contmuous monitormg the stability 1s acceptable for a day’s run for concentrations >5 mM. b The life of the column is largely determined by mechanical obstruction that gradually increases back-pressure of the column. The obstruction is partly a result of attrition from column material caused by pump pulsation It is therefore important that pumps with minimal pulsation be employed. Alternatively, pulse dampers can be included m the flow system to improve the

operational lifetime of the column. It 1sfurthermore recommendedto have an in-line

filter in the flow line or to use only filtered buffer and sample

solutions. 4. Notes 1 The recording should produce a symmetric peak with minimum broadening of the peak to facilitate the direct conversion of the peak height as a measure of the analyte concentration. 2. Alternatively other materials such as agarose (Sepharose), superporous agarose, nylon tubing, nylon powder, polyvinylalcohol, polyacrylamide (Eupergit C), and cellulosic materials could be used for the nnmobtlization of the enzyme or whole cells Support materials carrying OH groups can be activated with CNBr or tresyl chloride. Some materials are available m an epoxide form (such as Eupergit or Biosynth) to which the enzyme can be directly added. Cells can also be entrapped m small algmate or gelatin beads.

185

Thermal Transducer/Thermistor .

A

ov

di

/

0

1

3

2 Glucose

4 4

(MM)

Fig 2. (A) The thermometric signal in the form of symmetrrc peaks for varying concentrations of glucose between 0.1 and 4 mA4using 0.1 A4 phosphate buffer as the circulation medium, at pH 7.0 and flow rate of 0.74 mL/min. (B) The peak heights from A plotted as a function of glucose concentration injected using a sample loop size of 40 pL to obtain a linear cahbratron graph with correlation coefficient R = 0.99 3. All buffers should be at least partially degassed in an ultrasonication bath under vacuum (water aspirator) to minimize disturbances caused by air bubbles trapped m the flow system. 4. In an alternatrve procedure the rmmobihzed preparation could be suspended m buffer and pumped into the column with the inlet at the base of the column. 5. It is important that the analyte solution does not contain any air bubbles because this would affect the thermal signal and the sample amount. 6. The nature of the peak should be comparable to the ones illustrated in Fig. 2.

References 1. Spink, C. and Wadso, I. (1976) Calorimetry as an analytical tool in biochemistry. Meth. Biochem. Anal 23, 1-159 2. Grime, J. K. (1985) Analytical Solution Calorimetry Wiley, New York. 3. Privalov, P. L. (1974) Thermal investigations on biopolymer solutions and scanning microcalorimetry. FEBS Lett. rlO(Suppl.) 5 140-5 153. 4. Danielsson, B. and Mosbach, K. (1986) Theory and application of calorimetric sensors, in Bzosensors: Fundamentals and Applicahons (Turner, A. P. F., Karube, I., and Wrlson, G. S., eds.), Oxford Universrty Press, Oxford, UK, pp. 575-595

5

186

Ramanathan, Khayyami, and Danielsson

5 Damelsson, B. and Mosbach, K. (1988) Enzyme therrrnstors Meth Enzymol. 137, 181-197 6. Danielsson, B. and Mattiasson, B. (1996) Thermistor-based biosensors, m Handbook of Chemical and Biological Sensors (Taylor, R. F. and Schultz, J. S., eds.), Instttute of Physics Publishing Ltd., Philadelphia, pp. 1-17. 7 Danielsson, B. and Winqutst, F. (1990) Thermometric sensors, m Btosensor. A Practzcal Approach (Cass, A. E G , ed.), Oxford University Press, Oxford, UK, pp. 191-208

Enzyme Biosensors Ashutosh

Based on Fluorometric

Detection

Sharma

1. Introduction 1.1. Fluorescence-Based Enzyme Methods Measurement of specific analytes employing the combination of fluorimetry and the enzymatic reaction offers a number of advantages including high specificity, ease of measurement, and low cost. The use of fluorescence-based enzyme techniques can provide the basis for a variety of broanalytical and blosensor assays.Fluorescence-based enzyme biosensors typicahy employ dehydrogenases and the oxidases to target specific analytes by measuring the changes in the fluorescence emission behavior of a product or an indicator fluorophore. Depending on the type of enzyme employed, several options are available to achieve the desired transduction. These may include measuring (1) the direct fluorescence of an enzyme product or cofactor, or (2) changes in the fluorescence of an indicator fluorophore resulting from an interaction with a product of the enzymatic reaction. The method described in this chapter uses the interaction of an enzyme cofactor with an indicator fluorophore as a measure of substrate concentration. This format also involves a unique photoreaction that regenerates the cofactor. 1.2. Indicator-Based Enzyme Methods Enzyme reactions are coupled to fluorescence detection through either the disappearance of a substrate or the appearance of products. The substrates or products typically linked to fluorescence measurement include H+, Hz02, 02, or enzyme cofactors. The interaction of these products with an indicator fluorophore can allow the enzymatic reaction to be monitored in addition to providing a means of measuring substrate concentrations. From’ Methods m Botechnology, Vol 6 Enzyme and Ucrob/al Elosensors Technrques Edited by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

187

and Protocols NJ

Sharma

188

Modulation of fluorescence as a result of the changes m the pH within a bioreactor has been used to develop a variety of blosensors, including those for glucose (I), penicillin (2), ammonia, and urea (3). Kiefer et al. (4) described a lactose sensor based on a lipid bilayer containing the protein lactose permease. Lactose permease 1sa cotransporter for lactose and H+. When lactose is present on one side of the membrane, cotransport produces an increase in the concentration of H+ on the opposite side of the membrane. Changes m pH are typotally measured using a fluorescent dye indicator. Enzyme-catalyzed reactions that produce H202 have also been monitored by luminescence techniques. One example is the chemlluminescence reaction of luminol with H,O, in the presence of excesshorseradish peroxldase (HRP) ($5): HRP

lummol + 2H24 + OH- ->

ammophthalate + 3H,O +N2 + hght (1)

Other chemilummescent reactions have also been used to measure a number of analytes (6-9). In these reactions, the emitted light intensity during the steady state of the reaction IS directly related to the quantity of H202 present. Although these techmques are highly sensitive m detecting the presence of hydrogen peroxide, they suffer from the disadvantage of susceptiblhtyto mnterferenceby hydroquinone, bilirubm, uric acid, and asparticacid. Furthermore, thesesensorshave a limited lifetime becauseof the stoichiometric consumption of lummol. The quenching of indicator fluorescence by an enzymatic substrate or product has also been reported. Most of these schemesare based upon the ability of oxygen to produce quenching of molecular fluorescence. The early work of Kautsky (10) was used by Lubbers and Opltz (11) to develop a glucose biosensor based on oxygen quenching of an indicator fluorophore. Numerous oxygen-sensmg schemes are now available (12-19) that use indicator fluorescence quenching. However, the oxygen dependence of oxidase-catalyzed reactions makes them prone to errors caused by changes m oxygen concentration, nonlinear response, short sensor life, and so forth. In addition, oxygen-based sensor configurations are further complicated by the redox chemistry of H202, which 1soften involved in these reactions. Because of the broad range of NADPH-lmked enzymes, one of the most extensively exploited fluorescence-based enzyme assaysrelies on changes in the fluorescence of this cofactor. This assaymethod is based on the difference observed in the blue fluorescence of NADH at 460 nm (excitation at 340 nm) between the oxidized and reduced cofactor. Using this format, biosensor schemes for analytes such as lactate (20,21), and ammonia (22), have been developed. An example of NAD(P)H-linked assays include the dehydrogenase reactions in which a substrate (S) 1sconverted to the product (P) with the reduction of mcotinamlde adenine dinucieotide (phosphate), NAD(P)+, to NAD(P)H.

Flurometric Detection

189 s-dehydrogenase

S+NAD(P)+->P+NAD(P)H+H+

(2)

The progress of the enzyme assaymay be monitored optically by observing either changes in fluorescence of the cofactor NAD(P)H or changes in a fluorescence indicator resulting from its interaction with NAD(P)H. Monitormg the NADH fluorescence at 460 nm, the enzymatic conversion of substrate to product, or vice versa, may be followed directly without perturbation of the enzymatic process. The NADH detection-based enzyme biosensors are inherently sensitive; however, to be effective, the cofactor must be incorporated into the reagent phase and must also be part of a reversible reaction scheme to offer an effective sensor lifetime. Problems of sample autofluorescence as a result of the UV excitation wavelengths employed m this form of sensor are often also present and seriously degrade the effectiveness of the sensor measurement. In another approach, which will be described in some detail m this chapter, the enzyme-catalyzed reaction can be monitored by observing the interaction of a fluorescent indicator with the cofactor NAD(P)H (23). A specific case of this approach is described by a photoreaction between NAD(P)H and a fluorophore, resulting in a change in intensity of the indicator thionme (24). This method shows certain advantages because excitation and emission wavelengths of the indicator are shifted to longer wavelengths than those used with NAD(P)H. As a consequence, the background fluorescence typically found in biologic matrices is reduced. A typical reaction scheme is grven below: enzyme+

S + NAD+ + F enzyme+ hv .-> NADH + F’ ->

hv

P +NADH NAD+ + F

+ F’

(3) (4)

In an example of this reaction sequence,NAD+ is initially reduced to NADH, then oxidized back to NAD+, when ethanol is converted to acetaldehyde (Fig, 1). The concentration of NADH generated is directly dependent on the concentration of ethanol and the equilibrium constant of the enzymatic reactron. Subsequently, NADH being a quencher of thionine fluorescence allows the indirect measurement of ethanol (substrate) concentrations through the change in thionine fluorescence (25-27). The photoactivation of thionine (F) to the excited state (F*) allows the catalytic cycling of NAD+. The overall reaction process for the substrate-induced quenching (SIQ) of thionine by ethanol is empirically given by the substrate-induced quenching relation (25): (IO)/(I) = 1 + KfsrQ[ethanol10

(5)

190

Sharma ethanol

NAD+ +-

3I ADH

acetaldehyde

NADH

-

Fig. 1. Schematic of substrate-induced quenching by ethanol.

where I, and I are the fluorescence intensities in the absence and presence of the quencher, [ethanol&, is the initial ethanol concentration, and KzsrQis the substrate induced quenching constant at a time t. 2. Materials The following materials apply to SIQ probe construction. 1 2. 3. 4 5. 6 7.

Thionine (AR grade) (see Note 1). Ethanol (spectroscopic grade). P-NAD+ (Sodium salt, from Yeast, grade VII). Alcohol dehydrogenase (Bakers Yeast) E.C. 1.1.1.1 (Sigma, St.Loms, MO) Sodium bicarbonate (NaHCOs). Sodium carbonate (Na,COs). Polytetrafluoroethylene (PTFE) membrane 0.75 pm pore size (BOSS Leicester, UK).

3. Methods 3.1. Preparation of Reagents 3.1.1. Buffer and Indicator Solutions 1. Dissolve NaHCOs (8.401 g) m distilled water to a volume of 0.95 L. 2 Adjust pH with 1 MNaOH to pH 9.0 * 0.1. 3. Add distilled water to a final volume of 1 L (carbonate buffer, 0.1 M final concentration) 4. For the indicator solution, dissolve 2.8 mg thtonme m distilled water to a final solution volume of 100 mL (100 pA4 final concentration) and store the indicator solution in the dark (see Note 5).

3.1.2. Preparation of Enzyme and Cofactor Solutions 1. Prepare the enzyme/NAD+ solutions in lo-mL batches. 2. Weigh 54.8 mg NAD+ and dissolve it m a small amount of carbonate buffer. 3. Add carbonate buffer to make a final volume of 10 mL (8 mM final NAD+ concentration) 4. Add 466 U of alcohol dehydrogenase (ADH) to the NAD+ solutions described in steps l-3 and store the stock solutions at or below 4°C before the assay.

191

Flurometric Detection fluorescence excitation

fluorescence emission optical window I

\

reservoir chamber

b

I

/

P

/

PTFE O-ring d

sample in d

sample out

Fig 2. A reservoir-system SIQ biosensor (not to scale).

3.1.3. Preparation of PTFE Membrane 1 Cut a l-cm* piece of 0.75~mm thick PTFE membrane (see Note 3). 2 Wash it first with toluene, then with methanol, and finally with water. 3 Air dry before use.

3.2. Substrate-Induced

Quenching

(S/Q) Biosensor

Protocol

1. Prepare the SIQ assay mixture by adding 5 mL thronine stock solution to 5 mL enzyme/NAD+ assay mixture and store the mixture in an amber vial m the dark before the assay. 2. Assemble the bioprobe chamber (Fig. 2) using the following components: a glass plate 1.5 x 1.5 cm2, that was placed in a prefabricated (Plexiglas) receptacle with a cap (l-cm*) that served as a window, and a precut l-mm-thick O-ring (1 x 1 cm*) that was made of PTFE and fixed onto the glass plate using an optical epoxy. 3. Transfer 100 pL of the SIQ mixture mto the cavity of the O-ring. 4. Cover the O-ring by the PTFE membrane 5. Close the receptacle with the cap. 6. Snap-fit the bioprobe into the prefabricated flow cell made of stainless steel

3.3. S/Q Biosensor

Assay Protocol

The ethanol SIQ assay is carried out in a series of experiments performed with fresh SIQ mixture in the reservoir chamber of the biosensor for each experiment. Ethanol is added to the distilled water in the sample bottle to yield ethanol concentrations ranging from 0 to 10 rnit4. The ethanol solution is pumped through the sample chamber and the fluorescence in the reservoir chamber

is monitored.

When the substrate (ethanol)

is pumped

through

the

Sharma

192

sample chamber, the fluorescence intensity (for the SIQ assay) decreases exponentially with rime from an initial value to a minimum value. This value is dependent on the inmal concentration of substrate.When the fluorescence intensity attams a minimum, then the assayis said to have reached the assayend point. 1 Securethe flow cell containing the bioprobe tightly using the flow cell mount of

2. 3 4. 5. 6 7. 8

the front-face excitation fluorimeter. The sensor response is monitored using front-face excitation at an emrssron wavelength of 620 nm by using an excrtatton radiation of 560 nm for preset time periods Purge the complete sample flow circuit by pumping through 400 mL of dtstrlled water and allow the cncurt to pump dry. Add 100 mL of distrlled water to the sample bottle of the btosensor sample flow crrcun Start the peristaltic pump at a flow rate of 0 1 L/mm. Inmate the data-logging program once the flow m the sample crrcurt stabilizes and all an bubbles are evacuated Run the fluonmeter for 100 s to develop a “baselme” correspondmg to initial relative intensity (Is). Extract &35 uL ethanol from the sample vial using a gas-tight syringe. Inject the ethanol sample solution into the sample bottle and swirl to ensure adequate mixing.

3.4. Biosensor Evaluation In the reservoir cavity the analyte takes part m the enzymatic reaction, resulting

in the production

of NADH.

Thionine

present in the reservoir cavity

solution is excited by light entering via an optical window in the side of the reservoir chamber. The excitation light is in the visible band (560 nm, green), hence the optical window is made of glass, not quartz or other expensive UVtransmitting material. The NADH present in the reservoir cavity as a result of the enzymatic reaction produces fluorescence quenching of thionine. This results m a reduction in the intensity of the fluorescence emission light leaving the reagent cavity via the optical wmdow. The fluorescence emission light is detected and processed to determine the relative fluorescence intensity from the SIQ assay mixture (see Note 6). The fluorescence intensity of the SIQ mrxture decreases with time until the assayreaches a minimum corresponding to final relative intensity (4. The actual quenching value for the assay is calculated by dividing the initial intensity I0 by I. Figure 3 shows the effect of 9.7 rnM (0.035% v/v) ethanol on the fluorescence of the thionine in the assay mixture at the assay end point. The profile of the thionine spectrum remains the same after fluorescence quenching. Peak fluorescence

was at 6 18 run.

The fluorescence response obtained with the reservoir SIQ biosensor for a 9.7 mm ethanol concentration dissolved in water in a closed flow loop (flow rate of 0.1 L/mm) is shown in Fig. 4. After the addition of ethanol to the SIQ

193

Flurometric Detection

60-

9.7 mM ethanol

60-

wavelength (nm)

Fig. 3. Effect of various ethanol concentrations on the emission spectra of the thiomne component of the SIQ assay at assay equilibrium Assay mixture is 23.3 U/mL ADH, 4 mMNAD+, 50 l.tMthionine in 0.05 Mcarbonate buffer, pH 9.0, at 2OT Fluorescence data were collected at an emission wavelength of 580-690 nm for a front-face excitation wavelength of 560 nm

the fluorescence decreases to a mmtmum that is dependent on the initial ethanol concentration. The time to reach a minimum fluorescence value for the ethanol SIQ assayunder the prescribed conditions for higher concentrations of ethanol is a3000 s (50 min). The ethanol SIQ biosensor was tested for the range O-9.7 mA4 (O-0.035% v/v) ethanol. The quenching value was obtained by dividing the initial fluorescence mtensity m the absence of ethanol to that measured at assay equilibrium for a given amount of ethanol. Figure 5 shows the SIQ plot for the quenching of thionine by ethanol at assay minima. For the SIQ assay component concentrations used, the biosensor exhibited a sensitive response to ethanol with a maximum SIQ constant of 225.7 A4-‘. The magnitude of quenching was favorable with a half-quenching concentration of 8 mA4 ethanol. The behavior of the sensor is similar to that of the bioassay performed in a cell, except that the diffusion of ethanol through the PTFE membrane is slow, therefore affecting the equilibrium time. The assayhas a long timescale to mmimum reading of ~50 min. Clearly, the slow response is a result of the PTFE membrane. The biosensor response and sensitivity would be improved by a thinner and more permeable membrane (see Notes 4 and 7). The reservoir SIQ brosensor has been demonstrated to be capable of measuring ethanol concentrations in the range O-l 0 mM. However, as is typical of reservoir-type biosensors, the sensor response was slow. Both sensitivity and response time were influenced to a high degree by the properties of the semiperassay mixture,

194

Sharma

0

500

1000

1500

2000

2500

3000

3500

Fig. 4. Plot of relative intensity versus time for the reservoir-form of the ethanol SIQ assay Values to right of data traces are concentrations of ethanol m mA4. Assay mixture is 50 $4 thiomne, 4 mM NAD+, and 23.3 U/mL ADH, in 0 05 M carbonate buffer, pH 9.0, at 20°C. Fluorescence data were collected at an emission wavelength of 6 18 nm for a front-face excitation wavelength of 560 nm. 0 222018-

== -0 -O

16141210 0

1 2

I 4

'

, 6

I

, 8

, 10

ethanol cont. in mM Fig. 5. SIQ plot for the quenching of thiomne by ethanol at assay equilibrium. Assay mixture was comprised of 50 flthiomne, 4 mMNAD+, and 23 3 UlmL ADH m 0.05 M carbonate buffer, pH 9 0, at 20°C Fluorescence data were collected at an emisston wavelength of 6 18 nm for a front-face excitation wavelength of 560 nm.

meable membrane. It was necessary to replenish the SIQ reagent in the reagent chamber. This sensor showed no slgmficant cross-sensitivity to methanol, propanol, nonanol, ethandiol, 1,3-butanediol, or glycerol. 4. Notes 1. Check the reagents for then purity by performing thin-layer chromatography (TLC) for the indicator and recording respective absorption and fluorescent spectra. Make sure that they are m agreement with the literature.

F/urometric Detection

195

2. Enzyme and NAD+ solutions should be prepared daily. 3. Do not stretch the membrane. 4. Oxidation of thionine results in the recovery of fluorescence, which is achieved via a number of different reaction routes discussed in ref. 8. In the solution oxygen provides such a possibility. However, various competing processes result in nonlinearity in the recovery response, and also lead to fluctuations in the signal. 5. Some amount of thionine appears to become immobilized onto the proteins in use. This should lead to some changes in the indicator behavior. We have not experienced serious problems because of this aspect. However, it necessitates that higher concentrations of thionine should be used to obtain reasonably strong fluorescence signals. 6. Various possibihties exist to construct sensors using this method. These include a. Immobilized thionine, cofactor, and the enzyme; b. Immobilized thionine but solution-phase cofactor and enzyme; c. All elements m solution phase; and d. Covalent bonding of the indicator to the enzyme or cofactor itself. Such formats coupled with the order of the reagent layer should provide different performance. 7. Our recent experience suggests excellent performance with the liquid phase reagent entrapped inside a membrane barrier to construct fiber-optic sensors.

References 1. Trettnak, W., Leiner, M. J. P., and Wolfbeis, 0. S. (1988) Fibre-optic glucose sensor with a pH optrode as the transducer. Bzosensors 4, 15-26. 2. Kulp, T J , Cammins, I., Angel, S. M., Munkholm, C., and Walt, D. R (1987) Polymer nnmobihsed enzyme optrodes for the detection of penicillin. Anal Chem. 59,284!%2853. 3 Xie, X., Suleiman, A. A , and Guilbault,

G. G (1991) Determination of urea by a fiber-optic fluorescence biosensor. Talanta 38, 1197-1200. 4. Kiefer, H , Klee, B., John, E., Stierhof, Y., and Jahnig, F. (1991) Biosensors based on membrane transport proteins. Biosens. Bloelec. 6,233-237. 5. Bostick, D. T. and Hercules, D. M. (1975) Quantitative determination of blood glucose using enzyme induced chemiluminescence of luminol. Anal. Chem. 47, 447-452 6. Zhou, X. and Arnold, M. A. (1995) Internal enzyme fiber-optic biosensors for

hydrogen peroxide and glucose. Anal. Chim. Acta 304, 147-156. 7. Abdel-Latif, M. S. and Guilbault, G. G. (1988) Fiber-optic sensor for the determination of glucose usmg micellar enhanced chemiluminescence of peroxyoxalate reaction. Anal Chem 60,2671-2674. 8. Genovesi, L., Pedersen, H., and Sigel, G. H. (1988) The development of a generic hydrogen peroxide sensor with application to the detection of glucose. SPIE-Chem. Biochem Environ. Appl Fibers 990,22-27. 9. Jianzhong, L., ZhuJun, Z., and Ling, L. (1994) A simplified enzyme-based fiber optic sensor for hydrogen peroxide and oxidase substrates. Talanta 41, 1999-2002.

196

Sharma

10. Kautsky, H. (1939) Quenching of lummescence by oxygen. Trans Faraday Sot 35,216-219 11. Lubbers, D. W. and Opitz, N. (1983) Optical fluorescence sensors for continuous measurement of chemical concentrations in biological systems Sew. Actuators 4, 64 l-654. 12. Wolfbeis, 0. S., Offenbacher, H., Kroneis, H., and Marsoner, H (1984) A fast responding fluorescence sensor for oxygen. Mcrochzm Acta [Wzen] l2, 153-158. 13, Peterson, J. I., Fitzgerald, R. V., and Buckhold, D. K. (1984) Fiber-optic probe for m-vivo measurement of oxygen partial pressure. Anal Chem 56,62-67. 14. Lippitsch, M E., Pusterhofer, J., Lemer, M. J P , and Wolfbeis, 0. S. (1988) Fibre-optic oxygen sensor with the fluorescence decay time as the mformation carrier Anal Chim Acta 205, l-6. 15. Goswami, K., Klamer, S. M., and Tokar, J M (1988) Fiber optic chemical sensor for the measurement of partial pressure of oxygen, SPIE-Chem Brochem Environ Appl. Fzbers 990,ll l-l 15 16 Carraway, E. R., Demas, J. N., and DeGraff, B. A. (1991) Photophysics and oxygen quenching of transition-metal complexes on fumed silica. Langmuzr 7, 2991-1998 17. Sharma, A. and Wolfbeis, 0. S (1988) Fiber-optic oxygen sensor based on fluorescence quenching and energy transfer. Appl Spectrosc. 42, 1009-1011 18. Khmant, I., Belser, P., and Wolfbeis, 0. S. (1994) Novel metal-organic ruthenium(I1) Ditmm complexes for use as longwave excitable luminescent oxygen probes. Talantu B 985-99 1. 19 Sharma, A (1994) Excimer fluorescence quenching based oxygen sensor Proc SPIE 2131,598. 20. Wangsa, J. and Arnold, M. A. (1988) Fiber-optic biosensors based on the fluorometric detection of reduced mcotmamide adenine dmucleotide. Anal Chem 60, 1080-1082. 2 1. Scheper, T. and Btickmann, A F (1990) A fiber optic biosensor based on fluorometric detection using confined macromolecular mcotinamide adenine dmucleotide derivatives. Blosens. Bloelec 5, 125-l 35. 22 Kar, S. and Arnold, M. A. (1992) Fiber-optic ammonia sensor for measuring synaptic glutamate and extracellular ammonia. Anal. Chem. 64,2438-2443. 23. Sharma, A (199 1) Method and optical probe for the determmation of NADH m a sample, UK patent application no 9 12 15749. 24. Sharma, A. (1992) Photolytic oxidation of reduced mcotinamide adenine dinucleotide. Spectrochlm. Acta 48A, 893-897 25 Sharma, A. and Quantrill, N S M (1994) Measurement of ethanol using fluorescence quenching. Spectrochim Acta A 50, 1163. 26. Sharma, A. and Quantrill, N. S. M. (1994) On the substrate induced quenching of a fluorophore and its applications m enzyme assay. Proc SPIE 2131,563. 27 Sharma, A. and Quantrill, N. S. M. (1996) Ketoglutarate assay based on fluorescence quenching by NADH. Btotechnol. Prog. 12,4 13.

MICROBIAL BIOSENSORS

15 -Microbial Biosensors Based on Oxygen Electrodes Klaus Riedel 1. Introduction Microbial sensors are based on microorganisms in intimate contact with a transducer, which converts the biochemical signal into a quantifiable electrical response signal. The aim of this combinatton is the sensitive determination of a large spectrum of substances in various fields, especially m brotechnology and pollution control. The use of microbial cells in place of isolated enzymes offers several advantages over enzyme electrodes, such as, elimination of the tedious enzyme extraction and purification steps, avotdante of the need for a cofactor, and increased stability. The microbial sensors show an increased stability because of the enzyme environment is optimized by evolution and well suited for recovery. These sensors are essentially living and may be fed and kept alive for a long period. Furthermore, the whole cell may perform multistep transformations that could be difficult, if not impossible, to achieve with single enzymes. However, microbial sensors suffer from the multireceptor behavior of intact cells, resulting in a rather poor selectivity. This ability to recognize a group of substanceshas been exploited for the determination of complex variables, such as the sum of biodegradable compounds in waste water (BOD) (I,21 and mutagenicity of compounds (3). Moreover, the enormous wealth of microorganisms with a wide spectrum of metabolic types is an inexhaustible reserve for many uses of biosensors. A particular advantage is the ability to measure the respiratory activity of microorganisms and its alteration as a result of the presence of a tested substance. This allows a relatively simple transduction of the substrate response of microorganisms by an oxygen electrode. From

Methods m Biotechnology, Vol 6 Enzyme and Mwobral Biosensors Technrques EdIted by A Mulchandani and K R Rogers 0 Humana Press Inc , Totowa,

199

and Protocols NJ

200

Riedel

Fig 1. Design of microbial sensor (15) 1, isolator; 2, electrolyte, 3, anode; 4, cathode, 5, Teflon membrane; 6, munobilized microorganisms, and 7, dialysis membrane.

The microbial sensors herem reported measure the change m respiratory activity of microorganisms that are monitored directly by an electrochemical device. This sensor type is called a respiratory electrode. 7.1. Construction of a Microbial Sensor The design of the cell biosensor shown in Fig. 1 is m principle identical to an enzyme sensor. The main parts of such a biosensor are the microorganisms

as the recognition system and an oxygen electrode as the physical transducer. The parts are separated by a gas-permeable membrane. The cells are immobilized using an outer semipermeable membrane covering the sensor. 1.2. Immobilization of Microbes The basis of a biosensor is the intimate contact between the biocatalyst and the transducer element. The most convenient and successful way to construct the sensor is by immobilization of the biocatalyst. In general, the immobilization of the microorganisms for analytical purposes should result m the following effects: 1 Increased working stability of the organisms and the btosensor, 2 Reusability of the organisms because of their increased storage stability, and 3, Long, predictable, half-life of the actrvity of the mnnoblhzed organisms because they become an integrated constituent of the analytical device. Moreover, the oxygen permeabrhty through the unmobrl~zed mtcroorgamsm membrane must be good

Methods of immobilization include physical methods (adsorptton, entrapment in supports) and chemical methods (covalent binding to supports). However, chemical methods, to our knowledge, have not been successful for

Oxygen Electrodes

201

immobilization of microbial cells because of loss of biologic actwtty. The immobilization with the broadest range of applications is the entrapment of microorganisms in polymers forming gel membranes to entrap the microbial cells such as, agar, gelatin, collagen, polyacrylamide, and polyvinylalcohol (4). Physical methods (adsorption, entrapment in supports) for immobihzation of microorganisms dominate. The simplest and most widely used approach is the physical adsorption through centrifugation or filtration of a microbial suspension onto a membrane or sheet of acetylcellulose (5), filter paper (56) or nylon (7). Microorganisms adsorbed onto filter paper have high sensitivity compared with entrapped microbes (8). In the case of adsorption, the matrix only gives mechanical support to the microbes; the pores of the support are filled up with living organisms. The porous support gives a defined thickness to the microbial layer. The second immobihzation method with broad application is the entrapment of microorganisms in biologic (agar and collagen (91) or chemical (polyacrylamide [9/ or polyvinylalcohol (2j) macromolecules. A very promismg approach is the use of so-called prepolymers of ENT (poly(ethyleneglyco1) and ENTP type (poly(propyleneglyco1) or modified polyvmylalcohols for entrapment of microbial cells (10). These polymeric gels hinder leakage of the orgamsms but allow access to substrates and oxygen, In most cases the gel membranes increase diffusion resistance, thus slowing down the response of the sensor. It is possible to prepare a thin layer of immobilized cells with a response time Cl min (2). 1.3. Function of Microbial Sensors Microbial sensors show principal differences from enzyme sensors.The difference between an enzyme biosensor and a microbial sensor IS that in the case of the microbial sensor the substrate must be transported through the cell membrane, which forms a diffusion barrier. Solutes can pass only via specific translocation systems, either with an active transport system or by facilitated diffusion; passive transport by diffusion is of minor importance. Active transport, which allows accumulation of substratesagainst a concentration gradient, requires highly specific carrier proteins and consumes metabolic energy. Therefore the coupling to the cell energy-transducing systems,especially to the respiratory chain, is an important aspect of active transport. The function of a microbial respiratory sensor is described as follows: 1. Oxygen diffuses from the an-saturatedsolution through the dialysis membrane, the membranecontaining the microorganisms,and the Teflon membraneand is then reduced at the cathode A small proportion of the oxygen ISconsumedby the microorganisms.The steadystatecurrentrepresentsthe oxygendiffusion throughthe composedmembraneand reflects the endogenous respiration of the microorganisms

Riedel

202

addhon of

‘JS

time Cminl Fig. 2. Typical responsecurve of microbial sensor(principles of measurement)(4). (1) endpoint measurement(respiration rate R), and (2) kinetic measurement(acceleration of respiration rate A). 2. If assimilable substrateis addedto the measuringsolution, the substratepermeatesthrough the dialysis membrane,is taken up by the microbial cells,and subsequently degraded.Theseprocessesare causedby an increaseof respiration rate resulting n-ra decreasein the dissolved oxygen concentration and the current decreasesuntil a new steadystateis reached In principle, there are two possibilities of measurement (Fig. 2): (1) endpoint measurement (steady state mode), m which the differences m current (AI) reflects the respiration rate of the substrates (RS), and (2) the kmetrc measurement (first derivative of the current-time curve corresponding to the acceleration of respiration [A]). The first method has been most frequently used in microbial sensors. Here a relatively high concentratron of biomass and thick membranes are required. Furthermore, because of the diffusional resistance imposed by the microbial cell membranes, the response times for microbial sensors are longer than for enzyme electrodes. Response times of comparable magnitude to those of enzyme sensors are reached only wrth kmetrcally controlled microbial sensors. For these sensors very low microbe loadings and suitable immobilization of microorganisms, as well as thin membranes have to beused (Fig. 3) (11). The sensitivity of this type of sensoris primarily determined by the cell activity, andnot by drfGsrona1limitation. This meansthat the substratetransport into the cell and substrateasslrmlationshould be the rate-limiting process(12). The response of a microbial sensor is related to the physiologtc state of the microorganisms. The physiology of microorganisms used in the mtcrobtal sensor is optimally characterized by conditions of extreme nutrtent limitation. Their

Oxygen Electrodes

203

1

2

3

4

cell Loadmg Im g dry welghtIcm21

Fig. 3 Influence of cell loading (Bacillus subtzlu) on the change of current after addition of 0.15 mmol/L glucose (II)

metabolism is in a standby stateso as to guarantee the survival of the cell. These physiologic

limitations

determine both the stability and sensitivity

of biosensors.

1.4. Additional Considerations and Applications Many kinds of microbial sensors have been described in the literature. Among the respiratory indicators used for microbial sensors, oxygen is the dominant one. These sensorshave a high potential for analytical application in three fields: the determination of substrates (Table l), the measurement of complex parameters (like BOD), and the investigation of physiologic state or bioactivitity of microorganisms. One disadvantage of microbial sensors for substrate determination is their low selectivity. Biochemical knowledge makes it possible to alter the selectivity and sensitivity of microbial sensors. Approaches for enhancing the selectivity and sensitivrty are: 1. The induction of desired transport and/or metabolic systems, 2. The inhibition or suppression of undesired transport mechanisms or metabolic pathways,

3. The coupling of enzymeswith immobilized microbial cells to form hybrid sensors for elimination of interfering substances or the formation of specific products, 4. The transfer of desired plasnud-controlled pathways into suitable microbial strains; and 5. The exclusion of undesired substrates by dialysis membranes

Glucose Fructose Sucrose Maltose Lactose Organic acids Acetic acrd Ammo acids L-glutamrc acid L-tryptophan Peptrdes Aspartame Angiotensm Gonadotropin releasing hormone Steroids Cholesterol Androstendrone Testosterone

0 l(k) 0 l(k) 0 l(k) 0.1(k)

0 5-0.75 (k) 0 5-0.75 (k) 0 5-0.75 (k)

0 01-O 15 4 x 10-4-7 x lcr’ O-07-0.6 (2-15) x 1O-3 (2-15) x 1O-3

0.0150 13 0 015-O 13 0.0150.13

B. subtzlis Pseudomonas Juorescens

B subtzlzs B. subtzlis B. subtilis

Nocardza erythropolzs N. erythropolzs N. erythropolzs

3-5(s)

8(s)

0.08-l .2

Trzchosporon

brasszcae

Bacillus subtilis Saccharomyces cerevzsiae Brevibacterzum lactofermentum B. lactofermentum B lactofermentum B subtzlzs E. coli + GOD

Response time, min 10(s) 10(s) 0 l(k) HO(s) 1O(s) 10(s) 1O(s) 0.1(k) 3(s)

Measurmg range, mM 0.0125-0.125 0.025-o I25 0.05-O 6 0.01-l .o 0.1-1.0 0.1-I 0 0 l-l.0 0.050.5 0.15-l 4

Pseudomonasjluorescens

Electrodes

Carbohydrates Glucose

Based on Oxygen

Mtcroorganisms

Sensors

Analyte

Table 1 Microbial

30 30 30

28 29 29

18 27

26

21 22 11 23 24 24 24 17,20 2.5

Ref

Trimethylamine Other compunds Phosphate Iron (II, III)

Uric acid Creatmme

Urea

NO,

NH,

Antibiotics Nystatin N-compounds NH,+

(332

Vitamms/cofactors L-ascor-acid Cobolamin (B,,) Alcohols Methanol Ethanol agglomerans

(526) 1O-3 S-70 0.06-2.2

Chlorella vulgaris Thlobaclllus ferrooxldans

16)

0.025-0.5 0.4-8.8

48 49 OS-5

Us)

45 46

44

8 41 42 43

39 40

38

33 34 35 36 37

47

(continued)

31

32

3.5

3(s)

7(s)

0.1(k) 4(s) 10(s) 3(s)

0.01-0.15 0.005-2.5 0.01-0.6 0.15-5 2-200

8(s) 7(s)

0.002-0.08 0.45-l 0

Nitrosomonaseuropaea Nitrirfying bacteria from activated sludge B. subtilis Nitrifying bacteria Nitrobacter spec. Nitrifymg bacteria from activated sludge Nitnfying bacteria from activated sludge + urease Altenarla tennis Nztrobacter spec. + Nltrosomonasspec. + creatrnase Pseudomonasaminovorans

60(s)

2(s) 2(s) 2-3(s)

10(s) 10(s)

2.5-3(s) 120(s)

O-54 U/cmp3

0.06-o 7 0.05-0 5 co.4 2 x l(r5--2 x lo”’ 0.2-5

0.004-0 7 (5-25) l@s

Saccharomycescerevislae

Unidenttfied bacterium Trzchosporon brasswae Acetobacter xylinum Candida wnli Pseudomonas

Enterobacter E coli

0.02-O. 15 0 02-0.8 0.002-0 04 0.004-0.04 0 02-0.08 0 4CLO.20 0 2-l .o 0 2-3 0 2-3 0 7-l .35 0.0002-0.02 0 004-O 04

Trlchosporon cutaneum Rhodococcus spec Truzhosporon belgeliz (cutaneum) Rhodococcus spec Rhodococcus spec Pseudomonas putida Alcallgenes eutrophus

Alcaligenes eutrophus Pseudomonas putida Pseudomonas putida Pseudomonas Jtuoresence Pseudomonas putzda

‘Measuring principle steady state or endpomt (s), kmetx (k)

Benzoate 3-chlor-benzoate 2,4-dlchlorphenoxyacetlc acid Biphenyl PCB benzene naphthahne Caprolactam

Chlorophenols

0 013-6 6

Methylomonas flagellata

Measurmg range, mM

Methane Aromatics Phenol

Electrodes

Microorganisms

Based on Oxygen

Analyte

Table 1 (continued) Microbial Sensors

0.25(k) 0.25(k) 2-10(s) 26) 1.5-2.0(s)

0 25(k) 0.25(k) 0.25(k) 0 25(k) 0.25(k) 0.25(k) 0.25(k)

1-2(s)

Response time, mm

56,57 56,57 58 59 60

52 53 54 52 55 56,57

51

50

Ref

Oxygen Electrodes

207

However, microbial sensors suffer from the “multireceptor” behavior of intact cells, resulting in a rather poor selectivity. This abrhty to recognize a group of substances is exploited for the determination of complex variables, such as the sum of biodegradable compounds in waste water. Biochemical oxygen demand (BOD), as a indicator of the amount of biodegradable organic compounds, is a widely used parameter for the control of waste water. The main disadvantage of this parameter is the necessary time of 5 d. A more rapid estimation of BOD is possible by using a microbial sensor containing whole cells immoblhzed on an oxygen electrode. The prerequisite for the use of microorganisms for BOD sensors is a wide spectrum of substrates. BOD sensors have been developed using the various microorganisms (13). The apphcatron of the btosensor technique offers an elegant possrbllity for the study of respiration by mrcroorgamsms. For such investigations it IS necessary that the signal reflects the changes of respiration rate without limitation by substrate diffusion. For this reason, very low microbe loading, resultmg in response times of about 15-30 s, ISused (II). Thus kinetically controlled respiratron electrode allows the efficient measurement of changes in respiration rate after substrate addition. This signal, which is designed as acceleration of respiration, is connected with the substrate uptake, as indicated by Its behavior with glucose, a-methylglucose, and glucose-6-phosphate, as well as the inhrbmon of glucose signal by chloromercuribenzoate (CMB), dinitrophenol (DNP), and sodium fluoride (NaF) (12). The change of respiration rate designated as acceleration of respiration seems to be a suitable indicator for the characterizatron of microbes. This parameter is an overall value encompassing the uptake of substrates as well as their oxidative degradation. It seems to be related to the limiting step of this metabolic chain. We have suggested that the acceleration of respiration is mainly connected with the substrate uptake, for example glucose (12). The rate of this process parallels to some extent the physiological state of the microorganisms. Microbial cultures can have different properties and potentials for product formation during a growth cycle. To express these different properties the term physiologic state was proposed by Malek (14). This term comprises the sum of the brochemical, morphological and especially physiologrc features and actlvities characterized by the most important indicators for the multiplication or activities of the organisms (14). The respiration of mrcroorganisms has been used to characterize the physrologic state. The application of the blosensor technique offered an elegant posiibility for the study of respiration by mrcroorganisms in relation to various substrates. For characterizing the physiological state of microorganisms it is necessary that the signal reflects the changes of respiration rate without limrtatlon by substrate diffusion.

Riedel

208 CORRE A 3020t! E ' 1.22 08! I.$; = Q3-5 g o-2-

WI

zE 0.8.- t

4

,,0’

‘O-0

\

8

12

16 tCh1

20

24

10.8 i

20

Fig. 4. Change of respiration rate (R) and acceleration of respiration rate (A) of glucose (0.15 mmol/L) durmg the growth cycle of B subtzh. For estimatton of R and A, microbtal samples have been taken and used for mtcrobtal sensors. Usmg the second Faraday law and regarding the biomass concentratron the resptration parameters A and R have been calculated as [nA/mm . mg DW) and [nA/min2 . mg DW) (615).

This method was demonstrated by the alteration of glucose sensltlvlty of B. during fermentation (Fig. 4) (6). Furthermore, the physiological state of the methylotrophic yeast Candida boidinzi was examined during growth in a

subtilis

209

Oxygen Electrodes 100 80

200

-B

./

ii

a

\ .

./

100 -

2

aE s

‘0

./

r 2

l-

I

/ L

[ _ 1 _ &./y 4

8

/I 12

---.

I

-

, 16

20

2&

28

1

tthl

Fig. 5. Course of acceleration of respiration A caused by glucose (0.4 mmol/L) methanol (4 mmol/L) of CandIda boldmr at growth cycle (6). pH-stated glucose medium. Afterward, the exhaustion lized to methanol by the yeast (Fig. 5).

and

of glucose is metabo-

2. Materials

2.1. Microorganisms 1. Microbial culture media: a. Bacrllus subtilis: 2.0% glucose, 0.5% casamino acids, 0.5% NH&I and 20 mL salt solution (0 4% FeCl, . 3Hz0, 0.13% CaC&, 0.2% ZnSO, . 7Hz0, 2.46% MgS04. 7Hz0, 3,7% KCl/L 0.134 mol phosphate buffer (15). b. Issatchenkia orlentalis 0.3% malt extract, 0.3% peptone, 0.3% yeast extract and 1.O% glucose (16). 2. Sources of microoganisms are national and international strain collections, such as ATCC-American Type Culture Collection, Rockville, MD. CSB-Centraalbureau voor Schimmelcultures, Baarm, The Netherlands. CNCM-Collection Nationale de Cultures de Microorganismes, Institut Pasteur, Paris Cedex 15, France

Riedel

210

DSM-Deutsche Sammlung von Mikroorganismen und Zellkulturen GmBH, Braunschweig,Germany. A further possibility is the isolation of microorganisms with specific capabilities from desired sources, such as soil or water. 2.2. Protective

Membranes

The outer membranes serve to keep the immobrhzed microorganisms on the oxygen electrode and prevent loss from the sensor. This is necessary, because it is not possible to stably link the microorganisms on the immobilization matrix using bifunctional reagents. Bifimctional reagents typically cause the destruction of microbial cells. In general the outer membrane must be < 1O-l 5 pm thick and mechanically as well as chemically stable. Especially suitable are so-called nuclear pore trace membranes based on polycarbonate or polyphtheretalate from Nucleopore Corp. or Oxyphen GmbH. The preferred membranes have a thickness of 10 pm and a pore size of 0. l-l .O pm. In addition, dialysis membranes are used, such as cellulose acetate. Commercially available sources of membranes include the following: l l l l l

Amicon, Danvers, MA. Mlllipore Ltd Elkton, MD Nucleopore Corp., Pleasanton, CA. Oxyphen GmbH Dresden, Dresden, Germany. Pall, Glen Cove, NY.

2.3. Electrochemical Devices of Microbial Sensors (Oxygen Electrode) Commercially available conventional dissolved-oxygen electrodes according to Clark are suitable transducers for microbial respiration sensors This electrode type consists of a precious metal cathode (gold or platinum) and a silver anode, both immersed m a solution of saturated potassium chloride. A gas-permeable membrane is used to separate the electrode from the unmobilized microorganisms. A polarization voltage of -600 mV selectively reduces oxygen via the followmg reactions: cathode: O2 + 2Hz0 + 4e- + 4OH-

(1)

anode:4Ag + 4Cl- + 4AgCl+ 4e-

(2)

The reductron current of oxygen is proportional to the dissolved oxygen concentration. Suppliers of oxygen sensors include the following:

Oxygen Electrodes

-+37--l

211

1 -ii Blosensor

Buffer

Standard Sample

Fig 6 Schematic diagram of measurement biosensor system l l l l l

ABB Kent, Stonehouse, UK Oxford Electrodes, Roschester, UK. Radiometer, Copenhagen, Denmark. Yellow Springs Instruments Ltd., Aldershot, UK. Denki Kagaku Keiki (DKK), 7842 Japan.

2.4. Measuring System/Instrumentation The following equipment is necessary: 1 A thermostatted measuring chamber with magnetic stirrers, 2. A potentiostat, and 3. A recorder.

It is advantageous in many casesto use a personal computer to control the measuring system as well as registration and calculation of measuring values. Such a measurmg system is shown in Fig. 6. Commercially available systems can also be used. Such commercial systems are most specific for determined parameters, for example glucose or BOD. But it is possible to configure these systems to accommodate

desired microbial

sensors. Commercially available biosensor systemsinclude the followmg: l l

AUCOTEAM GmbH, Berlin, Germany. Dr. Bruno Lange GmbH Berlin, Dusseldorf, Germany.

Riedel

212 l l l

Granfield Biotechnology Ltd., Newport Pagnell, Bucks, MK16 9QS, UK. Pmfgerltewerk Medingen GmbH, Dresden, Germany. Yellow Springs Instruments Ltd., Yellow Springs, OH. Potentiostats

l l l l l l l

and other instruments are available from the followmg sources:

Aber Instruments, Cefn Llan, Aberystwath, UK Bioanalytical Systems Inc., West Layfayette, IN Hewlett-Packard Ltd , Bracknell, Reading, UK. Knauer, Berlin, Germany. Oxford Electrodes, Hoo, Rochester, Kent, UK. Radiometer-Tacussel, Vrlleurbanne, France Sensrtor AB, Lingkoping, Sweden.

2.5. Reagents 1. Immobrlization reagents. Polyvinylalcohol solutton (PVA): 0.5 g PVA (Sigma, St LOUIS, MO) dissolved m 10 mL distilled water (5%). 2. Buffer: 0.1 M phosphate buffer pH 6.8; 22.823 g K2HP04 3 HZ0 and 13 61 g KH,P04 m aqua dest. 3 Calibration solution. BOD: Soluttons contammg either glycerol (404 mg/L, 85% glycerol liquid) with a BODS value of 275 mg/L BODS or glucose (458 mg/L) with a BOD, value of 275 mg/L BOD are employed as the standard soluttons for calibration of the BOD sensor 4. Inhrbrtors. 0.1 mmol/L chloromercuribenzoate (Sigma), 50 mmol/L NaF (Sigma) 5. Antrmrcrobral reagents: 0.1 g Kathon CG (Rohm and Haas Company, Philadelphia, PA) in 100 mL distilled water (0 1%) or 0 1 Euxyl K 400 (Synopharm GmbH, Hamburg, Germany) m 100 mL aqua dest. a. Compositton of Kathon CG: 5-chloro-2-methyl-4-rsothtazolm-3-one, 2methyl-4-isothiazolm-3-one, and Mg salts. b. Composttron of Euxyl K 400: 5-chloro-2-methyl-4-isothiazolm-3-one, 2methyl-4-isothrazolin-3-one, benzylalcohol, and Mg salts. 6. Substrates (all from Sigma)* glucose, glucose-6-phosphate, a-methylglucose, maltose, and glutamic acid.

3. Methods 3.1. Preparation

of Microbial

Sensor (BOD Biosensor)

1. Cultivate the yeast Issatchenkza orlentalis under aerobic conditions (rotatmg shaker) at 30°C for 24 h until the stationary phase. See Note 1 for further mrcrobra1 species for BOD brosensors. 2. Centrifuge the culture medium and resuspend the sediment with 0.1 Mphosphate buffer, pH 6 8, to get a biomass concentratron of 0.2-1.0 g wet wt/mL (corresponding to approx 4 times the dry weight: 0.05-0.25 mg dry wt/mL). See Note 2 for other procedures to prepare microbial sensors

213

Oxygen Electrodes

Oxygen electrode

Teflon membrane

a

Dialyse membrane with immobilized microorganisms

0

“OVling

Fig. 7. Construction of microbial sensor. 3. Add an aliquot of polyvinylalcohol to the microbial suspensionof a PVA solution (5%) and mix the suspensionwith a vortexer. SeeNote 3. 4. A volume of 20 pL of this suspensionis droppedonto a capillary membrane(see Note 4) to get an areawith a diameterof approx 6 mm. The areaof immobilized microorganismsmust be greaterthan the areaof the cathode. 5. Store the membrane containing the entrapped cells at room temperature for 10 min and then for 24 h at 4°C. This membranecan be stored at 4°C for a long time (approx 3 mo). 6. Placethe immobilized microorganismsmembraneon the Teflon membraneof an oxygenelectrodeandfix it in placeusingeithera boredcapor ‘W-ring (seeFig. 7).

3.2. Sensor Optimization 3.2.1. Biomass Loading 1. Preparethe sensoras describedin Subheading 3.1., step 2, using various biomassconcentrations. 2. Calculation of biomassloading (L, in mg DW/cm2) L=(MWx

v)(4IrxI-?)

(3)

where is the MWmoist weight of biomassper milliliter, D W is the dry weight of biomass, V is the volume, r is the radius of immobilized area,and 4 is the calculation factor of moist to dry weight (MW/4 = DW).

Riedel

214

3.2.2. Improvement 3.2.2.1

of Selection

INFLUENCE OF SELECTIVITY BY INDUCTION OF DESIRED METABOLIC ACTIVITY OF MICROORGANISMS

The mduction of desired activity can be achieved by two different methods: l

l

Cultivatton of the mrcroorgamsms with the correspondmg substrate, for example, maltose (17), glutamate (18j, or phenol and benzoate (19). Incubation of the sensor with the correspondmg substrate The prerequlstte for such influence of sensitivity 1s a knowledge of the genotype of the microorganisms (17).

3.2 2.2. INFLUENCE OF SELECTIVITY BY CULTIVATION 1. The bacterium Badus subtilis is cultivated in a semisynthetic medium with the correspondmg substrate (0.5% maltose) under aerobic condrtrons (rotating shaker) at 30°C for 18-24 h until the stationary phase. 2. Imrnobrhzation and preparation of a biosensor as described in Subheading 3.1., steps 2-6. 3.2 2.3. INFLUENCE OF SELECTIVITY BY INCUBATION OF THE SENSOR 1. Cultrvation, immobilization ofB subth and sensor preparation as described m Subheading 3.2.2.2., but without maltose. 2. The sensor is incubated with tested substrate maltose (1 0 mA4) for 3 h See Note 5 3.2.2 4. INFLUENCE OF SELECTIVITY BY INHIBITION OF UNDESIRED ACTIVITIES

The principle of mcreased selectivity by inhrbition of undesired activities is explained with the example of glutamic acid determination (18). Normally the determination of glutamic acid in the presence of glucose is not possible, because the signal of glucose is greater than the signal of glutamrc acid. The glucose activity can be reduced by blocking the glucose uptake system with chloromercuribenzoate (CMB). CMB is a thtol reagent and inhibits the glucose carrier irreversibly. A further inhibition of the glucose signal is achieved by reversible inhibition of glycolysls usmg NaF 1. B subtzlzs is grown on a rotation shaker at 30°C for 18 h m a semisynthetic medium containing as C-source glutamrc acid (0.5%). 2 Immobilization and prepare sensor as described in Subheading 3.1. 3. Incubate the biosensor with 0.1 mA4 CMB for 1 h (See Note 6). 4. Measure glutamic acid samples with 50 mA4 NaF containing buffer.

3.2.3. Hybrid Sensor By combining microorganisms with enzymes it is possible to improve selectivity as well as to determine polymers, such as starch, proteins and lipids,

215

Oxygen Electrodes

which the microorganisms cannot uptake. In the last case the microorganisms are combined with hydrolases, for example B. subtilis with glucoamylase (20). The procedure is as follows: 1. Cultivate and immobilize of B subtilis as described in Subheading 3.2.2.2. 2. Enzyme (glucoamylase E C.3.2.1.3) is crosslinked in a mixture with albummglutaraldehyde on silk (1 g/cm2 mounted with dialysis membrane m front of a microbial layer.

3.3. Sensor Evaluation 3.3.1. Measurement Modes Response 1 Place the microbial sensor in a thermostatically controlled and stirred measuring chamber. In general this chamber contains 2 mL of 0.01-O. 1 Mphosphate buffer. The buffer IS oxygen-saturated by stirring. The working conditions are related to the microorganisms used. In general the temperature is 30°C and the pH is 6.8. 2. The current output of the microbial sensor is measured with apO,-meter or a biosensor measurement system and can be calculated with a computer (See Notes 7-l 1).

3.3.2. Calibration 1. Add aliquots of standard analyte solution into the measuring chamber to generate a series of various concentration steps. Appropriate concentrations range from* 0.0 l-2 mM For calibration of a BOD sensor glucose or glycerol with determmed BOD, are suitable (see Note 12). 2. Determine the linear range of the calibration by plotting the signal (kinetic measurement: dIldt or endpoint measurement AI) vs concentration of substrate 3. Calibration equations for linearity range are as follows. c=a+kxS

(4)

a = (B x cs)l(S - B)

(5)

k = (cs)/(S - B)

(6)

where c is the substrate concentration, a is the offset, k 1sthe slope, S is the signal (current [nA or nA/min], B IS the blank signal, and cs is the Standard concentration 4. Calculate of the limit of detection and the limit of determination as follows: The limit of detection and the limit of determination are calculated by the blankvalue procedure from variation of the blank probe and empirical factors. Limit of detection is given by: yL=B+4,65sB

and limit of determination

(7)

is given by: yD=B+

14, 1 sa

where sn is the variation of the blank probe.

(8)

Riedel

276 3.3.3. Sample Measurement

Mode of sample can be manual with a pipet or automatic with a dosing pump, When the current is stable, mject 100 pL of the sample solution into the measuring chamber. The signal is compared with the calibration data and the substrate concentration is calculated. Generally, a recovery time of 5-l 0 min 1srequired between measurements. The recovery time is related to the substrate concentration and the time of influence of the sample. High substrate concentrations and long measurement times cause long recovery times. 3.4. Physiologic

Effect

3 4.1. Determination

of Substrate Spectrum (Substrate Screening)

1. Inject of testsubstratesolutions(100 &) in a concentrationrangeof 0.1-2.0 mM into the measuring systemwith microbial sensor. 2 A concentrationseriesallows the application of Michaelis-Menten kinetics It IS possibleto calculatethe apparentl$.,,and V,,, (seeNote 13). 3.4.2. Investigation

of inhibitor Effect

In principle, there are two possibrlities for the investigation of inhibitor effects: 1 In the first place a desired substrateis injected, for example glucose, and after reaching a steadystatecurrent the inhibitor is added.The decreaseof current is evidence of an inhibitor effect. 2. The inhibitor is addedto the measuringchamber.After reaching a constantcurrent the substrateis injected.The substratesignal obtainedafter inhibitor is comparedwith the substratesignal without inhibitor. The difference is evidence of an inhibitor effect. 3.4.3. Investigation

of Substrate Uptake

The principles of investigation of substrate uptake are described with the example of glucose. Usually, for the study of glucose transport the glucose analog a-methylglucose is used. For the localization of this effect, the signals of glucose, a-methylglucose (o-MG), and glucose-6-phosphate (G-6-P) have been compared using a sensor containing B. subtills (12). The glucose analog a-MG is taken up by the phosphoenolpyruvate sugar transport system similar to glucose but without being metabolized. a-MG was therefore used to study the glucose uptake. Phosphoenolpyruvate, which is a product of glucose degradation, is used to phosphorylate glucose during its transport and G-6-P is accumulated in the cell. On the other hand, G-6-P is taken up by a transport system that is independent of glucose. a-MG Initiates a decrease in current (Fig. 8). In other words, a-MG causes an increase of

Oxygen Electrodes

217

c0, t 3 Fig. 8. Current-time curves of a B. subtzlis sensor for glucose, a-methylglucose MG), and glucose-6-phosphate (g-G-P) (0.15 mM) (12).

(a-

respiration. This indicate that the change of respiration ts raised only by the uptake of U-RIG. Compared with glucose, the acceleration of respiration 1s smaller than for a-MG because the microbial cells cannot regenerate the phosphoenolpyruvate pool. If endogeneous reserves are exhausted the respiration rate again reaches the initial value of respiration. The procedure is as follows: 1. Inject 0.15 n-&f glucose into the measurement chamber and the register the response curve. 2. After rinsing the measuring chamber, inject 0.15 mM glucose-6-phosphate. 3. a-methylglucose m a manner that is analogous to glucose and glucose-6-phosphate is investigated. After a-methylglucose injection it is necessary to activate the sensor with glucose (see Note 14).

3.4.4. Characterization

of the Physiological State of Microorganisms

1. Take samples of 100 uL up to 1000 uL of culture suspension, depending on biomass concentratton, at different fermentation times. Sediment the culture suspension by centrifugation in a special centrifuge tube (Fig. 9) on a very thm paper layer (e.g., Joseph paper or cigaret paper). The diameter of this paper must be greater than the diameter of the cathode. 2. Sandwich the paper layer with the centrifuged microorganisms between the gaspermeable membrane of a Clark electrode and a dialysis membrane. 3. After preparing the electrode, the respiration 1s permanently slowed down as a result of the consumption of endogenous reserve substrates. After about 15-45 min a steady-state of oxygen concentration inside the microbial membrane is reached corresponding to the endogenous respiration (indicated by the steady state current. 4 If an assimilable substrate for the characterization of microbes is added into the measuring solution the oxygen concentration (and the current) decreases until a new steady state is reached. The difference between the steady state current corresponding to endogenous respiration and after uptake of substrate reflects the

Riedel

218

Frg. 9. Specral centrrfugatron tube (15). 1, tube; 2, thin paper layer; 3, screw. respiratron rate, RS in nA for the substrate: RS=I=IE--IS

(9)

where IE is the current without substrate (endogenous respiration), and Is 1s the current with substrate. Using the second Faraday law and regarding the biomass concentratton (mg DWImL), the respiration rate can be calculated m terms of oxygen consumption per time m nmol OJ(min mg DW). RS = (I . 60)/(zo . F DE’)

(10)

where z. is the electron number per mol of oxygen, and F 1sthe Faraday constant (96,5 16 coulombs). The acceleration of respiration, A (equal to the change of respiration rate with time), after addition of substrate is measured as the initial rate of current change per mm (r, m nmol0, [mm2 x mg DW]). A = (dlldt) = (dR)l(dt)

(11)

5. It is promising, the substrate to dose in a concentratton range of 0.1-2 0 and the parameter of Michaelis-Menten kmettcs to calculate. 4. Notes 1. It is possible to use following microbial species for BOD biosensors. Hansenula anomala, Trichosporon cutaneum, Pseudomonas putida, Rhodococcus erythropolis, Bacdlus hchenlformu, Bacdlus polymyxa, and B subtilis, as well as combinations of two or more microbial strains, such as B. subtilisJB. lichenlformls, Enterobacter speaeslcdrobacter species, and I. onentaldR. erythropolu.

2. Other procedures to prepare mmrobtal sensors are: (1) filtration of culture medmm through a porous cellulose acetate membrane (millipore type HA with

Oxygen Electrodes

3.

4. 5

6

7.

8 9.

10.

11.

219

pore stze 0.45 pm and a thickness of 150 pm) by suction from a water pump (not suitable to prepare kmetically controlled sensors, because the diffusion resistance is high); (2) direct transfer of microorganisms from an agar plate to the spacer dialysis membrane by an inoculation loop (a screening method and does not allow introduction of a defined quantity of cells into the sensor). The PVA-molecules are not linked with bifunctional reagents, such as glutaraldehyde. This reagent destroys the cell membrane of microorganisms. A further advantage of immobilizatron with PVA is the formation of a very thin layer A capillary membrane with a pore size of 0.6 pm, a diameter of 25 mm, and a thickness of 10 ym is suitable (Type Oxyphen). An incubation time of 3 h is usually achieved by bacterial sensors. Biosensors contaming yeast require an mcubation of 12 h. In each case incubation with the given substrate caused a specific increase of sensitivity for this substance only Prerequisite is the adequate genotypical potency of the mtcroogamsms. The achieved level of activity is constant (17). It is important that the inhtbitors are used m relatrvely small concentrations, thereby achievmg a partial inhibition. High inhibitor concentrations can abolish the metabolism. When a new biosensor 1s prepared with a microbe-immobilized membrane the sensor is not immediately active. The activation required a time of approx 2-24 h under working condmons. This activation time is related to the activity of the cells and the biomass loading on the immobilized membrane. First, the current is near 0 and increases with time. In this period the current changes continuously. When the output current becomes stable, measurement is started. Buffers with organic compounds, such as citric acid, acetate or Tris, are not suitable because the organic compounds are substrates for sensor microorgamsms. Moreover, this buffer can introduce contamination. The stability of a microbtal sensor is related to tts working conditions. In general, numerous measurements implied high stabihty; a low number of substrate determinations and long pauses tmplyed low stability. Normally, microbial sensors should be stored in buffer between use (overnight) at room temperature or at 4°C. If the activity of the btosensor decreases, the microorganisms can typically be activated by incubation with nutrient medium for some hours until the activity is recovered by growth of new cells A large growth of biomass must be hmdered, because a high loading causes a high diffusion resistance. This results in a low signal. Prevention of contamination is important for the working of the measurement system containing microbial sensor. Contamination influences the measurement results by degradation of substrate and by oxygen consumptton. Especially, m a dosage system, microbial films can build up on the walls of the tubes It is possible to conserve the measuring system with antibacterial reagents, which Inhibits the growth of contaminating microorganisms under anaerobic conditions. The microorganisms of the sensor are stable under aerobic conditions. One example of an antimicrobial reagent is Kathon CG or Euxyl K 400. These reagents are used in

220

Riedel

concentrations of 0 1%. The measuring system ts washed with these reagents, then the measuring chamber is emptied and filled with buffer. 12. The commonly used GGA standard (150 mg glucose and 150 mg glutamtc acid, equal to 220 mg/L BOD,) for BOD+ahbratton IS not suitable for a microbral sensor, because (1) this standard IS instable because of microbial contammation, and (2) the glutamic acid reaction of microorganisms 1sdecreased in the presence of glucose because of glucose repression. 13. The kmetx coefficients apparent KIM and Vmarare only for the tested biosensor itself 14. After a-methylglucose injection tt is necessary to activate the sensor with glucose

References I. Karube, I , Matsunaga, T , Mitsuda, S , and Suzukt, S (1977) Mtcrobtai electrode BOD sensor. BzotechnoL Bzoeng. 19, 1535-1547. 2. Rtedel, K., Renneberg, R , Kuehn, M., and Scheller, F (1988) A fast esttmation of BOD with mtcrobtal sensors. AppI Mxrobloi. Bzotechnol 28,3 16-3 18 3. Karube, I. and Suzuki, S. (198 1) Preliminary screening of mutagens wtth a microbial sensor. Anal Chem. 53, 10261026. 4 Riedel, K , Renneberg, R , Wollenberger, U , Kaiser, G , and Scheller, F (1989) Microbral sensors’ fundamentals and application for process control. J Chem. Tech Brotechnol. 44, U-106 5 Matsunaga, T,, Karube, I., and Suzuki, S. (1980) A specific microbial sensor for formic acid. Eur. J Appl Mwobzol Brotechnol 10,235-243 6. Riedel, K , Liebs, P., Renneberg, R., and Scheller, F (1988) Characterizatton of the physiological state of microorgamsms using the respiration electrode. Anal. Lett 21,1305-1322 7. Kulys, J. and Kadziausktene, K. (1980) Yeast BOD sensor Bzotechn Bzoeng 22, 22 l-226 8 Riedel, K., Huth, J , Kuhn, M., and Ltebs, P. (1990) Amperometrrc determmation of ammonmm ions with a microbial sensor J Chem. Tech. Bzotechnol 41, 109-l 16. 9. Matsunaga, T., Karube, I , and Suzuki, S. (1978) Rapid determmatton of mcotmic acid by unmobilized Lactobaclllus arabznosus Anal Chum Acta 99,233-239 10 Fukui, S and Tanaka, A. (1984) Application of biocatalysts mnnobiltzed by prepolymer methods. Adv Blochem Eng Biotechn 29,2-33. 11. Rtedel, K., Renneberg, R., and Liebs, P. (1988) Biochemtcal basis of kmetically controlled microbial sensors. Bioelectrochem Bzoenerg. 19, 137-l 44. 12. Riedel, K (1991) Biochemical fundamentals and improvement of selectivuy of mtcrobial sensors-a mimreview Bloelectrochem Bzoenerg 25, 19-30. 13 Riedel, K. (1998) Application of biosensors to environmental samples, in Commercial Biosensors. Appbcatlon to Cluwal, Bzoprocess and Envzronmental Samples (Ramsay, G., ed ), Wiley, New York, pp. 267-294. 14. Malek, J. (1966) Introduction, in Theoretical and Methodologzcal Basis of Contmuous Culture of Mlcroorganums (Malek, J. and Fencle, Z., eds.), Publishing House of the Czechoslovak Academy of Sciences, Prague, pp. 9-30.

Oxygen Electrodes

221

15. Riedel, K., Liebs, P., and Renneberg, R. (1985) An electrochemtcal method for the determination of cell respiration. J. Basic Mzcrob 25, 5 l-56. 16. Rtedel, K , Uthemann, R., Yang, X., and Renneberg, R. (1998) Deter mmation of BOD in waste water with a commercial combination sensor containing Rhodococcus erythropoks and Issatchenkia orientalzs. Biosens. Bioelectron, submitted. 17. Riedel, K., Renneberg, R., and Scheller, F. (1990) Adaptable microbial sensors. Anal. Lett 23, 757-770.

18. Riedel, K. and Scheller, F. (1987) Inhibitor-treated microbral sensor for the selective determination of glutamic acid. Analyst 112,34 1,342. 19 Riedel, K., Hensel, J., and Ebert, K. (1991) Biosensoren zur Bestunmung von Phenol und Benzoat auf der Basis von Rhodococcus-Zellen und Enzymextrakten (m German), Zbl Bakt 146,425-434. 20. Renneberg, R., Riedel, K., Liebs, P., and Scheller, F. (1984) Microbial and hybrid sensors for determmation of alpha-amylase activity. Anal. Lett. 17,349-358. 2 1. Karube, I , Mitsuda, S., and Suzuki, S. (1979) Glucose sensor using immobilized whole cells of Pseudomonasjuorescens. Eur J Appl. Mzcrobiol Bzotechn 7,343-350. 22. Vais, H., Onancea, F., Faghi, A. M., Delcea, C., and Margmeanu, D. G. (1985) Amperometric electrode for glucose with immobilized bacteria (Pseudomonas jluorescens).

Rev. Roumazne Biochzm 22,57-62.

23. Mascini, M. and Memoli, A. (1986) Comparrson of microbial sensors based on amperometric and potentiometric electrodes. Anal. Chzm Acta 182, 113-l 22 24. Hikuma, M., Obana, H., and Yasuda, T. (1980) Amperometric determmation of total assimilable sugars in fermentation broths with use of mnnobihzed whole cells Enzyme Mzcrobiol Techn 2,234-238. 25 Svorc, J., Mieartms, S., and Barhkova, A. (1990) Hybrid biosensor for the determination of lactose. Anal. Chem. 32, 1626-l 63 1. 26. Hikuma, M., Kubo, T , Yasuda, T., Karube, I., and Suzuki, S. (1979) Amperometric determination of acetic acid with munobihzed Trzchosporon brasszcae. Anal

Chzm. Acta. 109,33-38.

27. Vmcke, B. J., Devleeschouwer, M. J., and Patriarche, G J (1985) Bacterial electrode for the analytical use of the L-tryptophane oxidative metaboltsm of Pseudomonas fluorescens. J Pharm. Belg. 40,357-365. 28. Renneberg, R., Riedel, K , and Scheller, F. (1985) Microbial sensor for aspartame. Appl Microbial. Bzotechnol. 21, 180,181. 29. Riedel, K., Renneberg, R., Kleine, R., Kruger, M., and Scheller, F. (1988) Microbial sensor for peptides Appl Microbial. Biotechnol. 28,272-275. 30. Wollenberger, U., Scheller, F., and Atrat, P. (1980) Microbial membrane electrode for steroid assay. Anal. Lett 13, 120 1-12 10. 3 1. Vincke, B. J., Devleeschouwer, H. J., and Patnarche, G. J. (1985) Determmatron of Lascorbic acid with bacterial tissue and enzyme electrodes. Anal Lett 18,1593-l 606 32. Karube, I., Wang, Y , Tamiya, E., and Kawarai, M. (1987) Microbial electrode sensor for vitamin B,,. Anal. Chzm. Acta 199,93-97. 33. Karube, I., Suzuki, S., Okada, T., and Hikuma, M (1980) Microbial sensors for volatile compounds. Biochimie 62, 567-574.

222

Riedei

34 Hikuma, M , Kubo, T , Yasuda, T., Karube, I., and Suzuki, S. (1979) Microbial electrode sensor for alcohols Biotechn Bioeng 21, 1845-1853. 35. Divies, C. (1975) Ethanol oxidation by an Acetobacter xylznum microbial electrode. Ann. Microbial (Paris) 126A, 175-I 86 36. Mascini, M., Memoli, A., and Olana, F (1989) Microbial sensor for alcohol Enzyme Mzcrobiol

Technol 11,297-301.

37 Suzuki, S. and Karube, I. (I 987) An amperometric sensor for carbon droxide based on nnmobihzed bacteria utihzmg carbon dioxide Anal Chzm. Acta 199,85-91. 38 Karube, I., Matsunaga, T., and Suzuki, S. (1979) Mtcrobtassay of nystatm using yeast electrode Anal. Chim Acta 109,39-44. 39. Hikuma, M., Kubo, T , Yasuda, T., Karube, I., and Suzuki, S. (1980) Ammonia electrode with unmobihzed mtnfying bacteria. Anal. Chem 52, 1020-1024. 40 Okada, T., Karube, I , and Suzuki, S. (1982) Ammonium ion sensor based on munobihzed mtrifymg bacteria and a cation exchange membrane. Anal Chum Acta 135, 159-165.

41 Karube, I., Okada, T., and Suzuki, S. (1981) Amperometric determmatton of ammonia gas with unmobihzed nitrifying bacteria. Anal Chem. 53, 1852-l 855. 42 Karube, I., Okada, T , Suzuki, S., Suzuki, H , Hrkuma, M , and Yasuda, T (1982) Amperometric determmation of sodium nitrite by a mtcrobtal sensor. Eur J Appl Mtcrobiol

Bzotechnol

15, 127-132.

43. Okada, T., Karube, I., and Suzuki, S. (1983) NO, sensor which uses uumobthzed nitrate oxidizing bacteria Bzotechn Bzoeng 25, I64 1-165 1 44 Okada, T., Karube, I., and Suzuki, S. (1982) Hybrid urea sensor using mtrtfymg bacteria Eur J, Appl Mzcrobtol Biotechnol. 14, 149-154 45 Wollenberger, U (1989) unpublished data 46. Kubo, I., Osawa, M., Karube, I., Matsuoka, M , and Suzukt, S. (1983) Hybrid biosensor for clinical analysis. Proceedings of the Internattonal Meettng on Chemical Sensors, Fukuoka, Japan, pp. 660-665. 47 Gamatt, S., Luong, J. H. T., and Mulchandam, A. (1991) A mtcrobtal btosensor for trimethylamine usmg Pseudomonas aminovorans cells. Biosens. Btoelectron 6, 125-131. 48. Matsunaga, T., Suzuki, S., and Tomoda, R. (1984) Photomicrobial sensors for selective determination of phosphate. Enzyme Microbial Techn. 6,355-357. 49. Mandl, M. and Macholan, L (1990) Membrane biosensor for the determination of iron (II, III) based on immobilized cells of Thiobacillus ferrooxidans. Folta Mtcrobtol

35,363-367.

50. Okada, T., Karube, I., and Suzuki, S. (1981) Microbial sensor system which uses Methylomonas sp. for the determmation of methane. Eur J. Appl. Mtcrobiol. Biotechn

12, 102-106.

51. Neujahr, H. Y. and Kjellen,

K. G. (1979) Bioprobe electrode for phenol. Bzo-

technoi Btoeng. 21,671-678.

52. Riedel, K., Hensel, J., and Ebert, K. (1991) Biosensoren zur Bestimmung von Phenol und Benzoat auf der Basis von Rhodo-coccus-Zellen und Enzymextrakten (in German). Zbl Bakt 14fi425-434.

Oxygen Electrodes

223

53. Riedel, K., Beyersdorf-Radeck, B., Neumann, B., and Scheller, F. (1995) Microbial sensors for determination of aromatics and their chloroderivates. Part III: Determination of chlorinated phenols using a biosensors containing Z’richossporon beige& (cutaneum). Appl. Mtcrobiol. Biotechnol. 43,7-g. 54. Riedel, K., Hensel, J., Rothe, S., Neumann, B., and Scheller, F. (1993) Microbial sensors for determination of aromatics and their chloroderivates. Part II: Determination of chlorinated phenols using a Rhodococcus containmg biosensors. Appl Microbial. Biotechnol. 38,556-559. 55. Riedel, K., Naumov, A. V., Boronin, L. A , Golovleva, L. A., Stein, J , and Scheller, F. (1991) Microbial sensors for determination of aromatics and their chloroderivates. Part I: Determination of 3-chlorobenzoate using a Pseudomonas containing biosensors. Appl. Microbial Bzotechnol 35,557-562. 56. Beyersdorf-Radek, B., Riedel, K., Neumann, B., Scheller, F., Schmid, R. D. (199 1) Development of microbial sensors for determination of xenobiotics GBF Monographs 17,55-60. 57. Beyersdorf-Radek, B., S&mid, R. D., Riedel, K., Neumann, B., and Scheller, F. (1991) Microbial sensors for the determination of aromatics and their chloroderivates. Proc Symp Environm Biotechnol Oostende 66th Event of the European Federation of Btotechnology (Verachtert, H. and Verstraete, W F., eds.), 65-68. 58. Tan, H.-M., Cheong, S.-P., and Tan, T.-C. (1994) An amperometric benzene sensor using whole cell Pseudomonasputida ML2. Biosens Bioelectron 9, l-8. 59. Kbmg, A., Zaborosch, C , Muscat, A., Vorlop, K. D., and Spener, F. (1996) Microbial sensors for naphthaline using Spingomonas sp. Bl or Pseudomonas tluorescens WW4. Appl. Mtcrobiol. Btotechnol 45,844-850 60. Riedel, K., Naumov, A. V., Grishenkov, V. G., Boronin, A. M., Stein, J , Scheller, F., and Mtiller, H.-G. (1989) Plasmid-containmg microbial sensors for caprolactam. Appl. Microbtol. Btotechnol. 31,502-5&I.

Microbial Biosensors Based on Respiratory Inhibition Yoshiko Arikawa, Kazunori Ikebukuro,

and lsao Karube

1. Introduction Biosensors generally provide a rapid and convenient alternative to conventional methods for momtormg chemical substancesin fields as diverse as medicine, environmental monitoring, and food processing. A biosensor is a device that combines a biologic-sensing element with an electronic signal-transducing element. As biologic-sensing elements, enzymes, antibodies, receptors, organelles, and microorganisms, as well as animal and plant cells, can be used. Among these biologic-sensing elements, microorganisms have several advantages, as listed below (I): 1. Wide range of suitablepH andtemperature. 2 Long lifetime. 3. Low cost(becauseno isolation of the active enzymeis needed) Purified biomolecules, such as enzymes, are used as the sensing element in most biosensors. Although enzymes show high specificity for their substrates, they are generally expensive and have poor stability. Therefore mlcroorganisms have been employed as the sensing element of biosensors to solve these problems. They are suitable for on-line environmental monitoring that requires long-term stability. Microbial biosensors have been developed for assaying biochemical oxygen demand (BOD), a value related to the total content of organic material in waste water (2). Microbial sensors, of course, have some disadvantages, as listed below: 1. Long responsetime. 2. Long time to return to baseline. 3. Lower selectivity (becausea microorganism containsmany enzymes)

From* Methods m Bloiechnology, Vol 6. Enzyme and MIcrobra/ Blosensors. Technrques Edlted by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

225

and Protocols NJ

Arikawa, Ikebukuro, and Karube

226 Immobilized

microorganism

membrane

‘02

/ Organic

compound

Fig. 1 Principle of microbtal sensorusing oxygen electrode.(A) without organic compounds,(B) with organic compounds Therefore it is necessary to optimize measurement and storage conditions to shorten the measurement time of microbial sensors. The low selectrvity of microbial sensors can be complemented by use of membranes, such as a gaspermeable membrane (3). See Riedel et al. (4) for comprehensive reviews. More recently developed microbial sensors are reviewed by Karube et al. (5’. 1.1. Principle

of Microbial

Sensors

The principle of microbial sensors is shown in Fig. 1. A typical microbial sensor consists of an oxygen electrode to which a membrane containing munobilized aerobic microorganisms is attached. The respiratory and metabolic functions of the microorganisms are used to detect particular substances. Immobihzation of the microorganisms is a key step m the fabrication of microbial sensors.Immobilizatton should not harm the microorganism, and rt should, ideally, improve their stability for use in continuous monitormg. When analytes are degraded by nmnobilized microorganisms, respiratory activity increases and dissolved oxygen is consumed. The oxygen concentration is monitored using the oxygen electrode. A large decrease m current coming from the oxygen electrode indicates high metabolic activity of the immobilized microorganisms, which in turn indicates a high concentration of analyte. Some sensors use mtcroorganisms whose respiration is inhibited by the target analyte, as in the case of sensors for cyanide or pesticides.

Respiratory Inhibition 7.2. Immobilization

227 of Microorganisms

Immobilization methods must not harm the immobilized microorganisms and yet provide a sensor with a good stability under the prevailing conditions of temperature, ionic strength, pH, redox potential, and chemical composition. Major methods of immobilization are listed as follows: 1. 2. 3 4

Physical adsorption method. Entrapment method. Crosslinking method. Covalent binding method.

Methods of microorgamsm lmmobllization are usually modified versions of methods used for immobilizing enzymes. Immobilization of microorganisms is also carried out in reactors used in the food industry (e.g., beer and wine production) and in waste water treatment reactors. The most widely used methods of immobilization are physical adsorption or entrapment methods. Since chemical methods often damage the cell membrane and decrease biological

activity,

those methods have seldom been successfully

applied. See Nomura and Karube (6) for comprehensive reviews. 1.3. Oxygen Electrodes One of the most suitable transducers for microbial sensors is an electrode, especially the Clark-type oxygen electrode. The oxygen electrode measures the change in dissolved oxygen concentration resulting from the respiration of the immobilized microbes. Clark-type electrodes are either polarographic or galvanic in nature. Polarographic electrodes consist of a platinum cathode and a silver anode, both of which are immersed in an electrolyte solution of saturated potassium chloride. A suitable polarization voltage between the anode and the cathode 1s -0.6 or -0.7 V, and oxygen is selectively reduced at the cathode at this voltage. The reactions occurring in the polarographic system are as follows. Cathode: O2 + 2Hz0 + 4e- -> Anode: 4Ag + 4Cl-->

4OH4AgCl+

(1) 4e-

(2)

These chemical reactions result in a current that IS proportional to the concentration of dissolved oxygen. Galvanic electrodes consist of a lead anode and a silver or platinum cathode. The combination of the anode and cathode makes a potential difference so that the reaction shown below occurs spontaneously. Cathode: O2 + 2Hz0 + 4e--> Anode: Pb + 2OH-->

4OH-

(3)

PbO + Hz0 + 2e-

(4)

Arikawa, Ikebukuro, and Karube

228

Although galvanic electrodes are simple and economical, they have a slower response time and poorer stability compared with polarographic electrodes.

1.4. Microbial Sensors for Cyanide Determination Yeast can be used as a microorganism for such sensors, and thus cyanide can be measured by respiratory inhibition using an oxygen electrode. Cyanide is extremely toxic because it inhibits the respiration of life forms (7). The respiratory chain in the inner mttochondrral membrane m yeast contains the enzyme complexes through which electrons pass from NADH to OZ. Compounds such as potassium cyanide and sodium cyanide bind to heme a in cytochrome a and cytochrome a3in the cytochrome oxidase complex. The resultant inactivation of heme a causes a decrease in the respiration. The cyanideinduced decrease m respiratory activity causes a decrease in oxygen consumption. The cyanide concentration can be measured from the change m the oxygen consumption of the immobilized microorganisms with an oxygen electrode. Cyanide is mamly discharged from industries,such aselectroplating plants, coke plants, chemical industries, and petroleum refineries (8), and there have been several accidentsof cyanide contamination of rivers. Cyanide may enter the environment from many sources, m the form of water or air pollution. Therefore, a cyanide-detection systemshould be urgently established for river water analysis. Cyanide is commonly determined using spectrometric methods (!W2). A spectrofluorimetric method was also reported (13) but these methods require laborious and trme-consuming procedures. Flow-Injection analysis (FIA) was applied to these methods for rapid analysis (14,15), but these methods require many chemical reagents and expensive instruments. A cyanide electrode has also been reported (161, but it suffers from interference problems. Therefore, conventional analysis methods are not suitable for monitoring cyanide in the envnonment. A mtcrobral sensor for cyanide detection is one of the prospective methods for monitoring cyanide (Fig. 2) and the fabrication of such a microbial biosensor is described in this chapter. A batch-type microbial sensor is mainly described, but a flow system for rt and its reactor type are also described. 2. Materials

2.1. Solutions and Media 1. 2 3. 4. 5. 6. 7.

0.01 MTris-HCI buffer (pH 7.0). 0.01 MTris-HCI buffer (pH 7.0) containing 150mg/L glucose 0.9% Sodium chloride solution. 2% Sodium algmate solution. 1% Calcium chloride solution. N medium consistingof 8 g/L nutrient broth. YM medium consistingof 10g/L glucose,5 g/L peptone,3 g/L yeastextract,and 3 g/L malt extractat pH 5.8.

229

Respiratory Inhibition immobilized yeast membrane

i

0,

~

_-

oxygen electrode

0 0 OQ 0

electric current

\

I

Pt cathode

CN- addition

immobilized yeast’membrane oxygen electrode

electric current -*

I\

Pb anode Pt cathode

Fig. 2. Principleof microbialsensorfor cyanidedetermination.

2.2. Strains Several microorganisms can be examined to test which is the best one for cyanide detection. 1. 2. 3. 4. 5. 6.

Micrococcus luteus IF0 3342. Escherichia coli IF0 14249. Pseudomonasaeruginosa IF0 1095. SaccharomycescerevisiaeIF0 0337. Trichosporon cutaneumIF0 1046. Bacillus subtilis IAM 1069.

kf. uteusIF0 3342, E. coli IF0 14249,P. aeruginosa IF0 1095,S. cerevisiae IF0 0337, and T. cutaneum IF0 1046 were obtained from the Institute for Fermentation (IFO), Osaka, Japan. B. subtih’s IAM 1069 was obtained from IAM Culture Collection, Institute of Molecular and Cellular Biosciences, University of Tokyo, Japan. In addition, incubators are necessary for cultur-

230

Arikawa, Ikebukuro, and Karube

ing these microorganisms and a centrifuge IS required for their collection. S. cerevisiae should exhibit greater respiratory activity than the other microbes and have a higher sensitivity to cyanide (I 7).

2.3. Components of the Microbial Sensor For fabrication of a microbial sensor the following materrals are necessary: 1. A porous cellulose-nitrate membrane on which to nnmobllize mrcroorgamsms

(e.g., pore size,0.45 pm; Tokyo Rosh1Ltd., Tokyo, Japan) 2. A water pump for gentle suction to immobilize the microorgamsms on the cellulose-nitrate membrane. 3. A 200-mesh nylon membrane to fix the immobilized microbe onto the oxygen

electrode. 4 A Clark-type oxygen electrode (e.g., Type U-l, ABLE Corp., Tokyo, Japan)

For fabrication of a reactor-type microbial sensor, the followmg additional materrals are necessary: 5 Porous glass beads on which to immobilize microorganisms (pore size, 120 pm; sphere diameter, 0.4-l .O mm; Schott Nippon KK, Tokyo, Japan) 6. Sterile 20-mL syringe. 7 Sterilized cotton

2.4. Measurement System The measurement is made in a batch-reaction cell. A schematic diagram of the measurement system is shown m Fig. 3. The components are as follows: 1 Circulating WaterJacket (volume, 30 mL) to control the temperature of the sample solutron. 2 Magnetic stirrer and stir bar to stir the buffer solution during the measurement 3 10-m Resistor to convert electric current to voltage.

4 Electronicrecorder (e.g.,Type EPR-15lA, TOA ElectronicsLtd., Tokyo, Japan). For fabrication of a flow-type mrcrobial sensor system, the followmg additional materials are necessary. 5. Flow cell for Clark-type oxygen electrode (e.g., Type U-l, ABLE Corp., Tokyo, Japan). 6. Peristaltic pump to dehver the solutions into the sensor system (Mimpuls 3, Gilson, France) 7 Silicon tubes (id, 2 mm; od, 4 mm).

3. Methods 3.1. Growing the Microorganisms 1. Inoculate each bacterial stram into the appropriate media. Use N medmm for the growth of B subtih IAM 1069, M. luteus IF0 3342, E colt IF0 14249, and

231

Respiratory Inhibition oxygen electrode I

immobilized thermostated

Fig. 3. Schematicdiagram of the microbial cyanide biosensor. P. aeruginosaIF0 109.5.Culture thesestrainsin 100mL of N medium aerobically

at 38°C for longerthan 18h. Rotationrate is approx 120rpm. Use YM medium for the growth of S. cerevisiaeIF0 0337 and T. cutaneumIF0 1046. Culture these strainsin 100mL of YM mediumaerobicallyat 28°C for longerthan 18h. Rotation rate is approx 120 rpm. The microbesto be immobilized should be undergoing their final phaseof exponentialor their first stageof stationarygrowth. 2. Harvestthesecells by centrifugationat 3000gfor 10min. Discardthe supernatant. 3. Resuspendthe resulting cell pellets in 10mL of 0.01 MTris-HCl buffer (pH 7.0) and centrifuge again at 3000g for 10 min. Discard the supernatant.Repeatthis proceduretwice. Use the resulting pellet to immobilize the cells. The weight of wet cells may be from 10 to 100mg.

3.2. Immobilization of the Microorganisms and Fabrication of the Microbial Electrode 1. Resuspendthe cell pellets in 2 mL of 0.01 M Tris-HCl buffer (pH 7.0). 2. Load the cell suspensiononto a porous cellulose-nitratemembranewhile applying gentle suction from a water pump. The cells are trappedon the membrane. 3. Sandwichthe microorganismstrappedon the membranewith anothercellulosenitrate membrane. 4. Placethe sandwichedmembranesincorporatingthe immobilized microorganisms on the Teflon membranecover of a Clark-type oxygen electrode.

232

Arikawa, Ikebukuro, and Karube

5. Cover it with a protective layer of 200-mesh nylon membrane. The resultmg electrode is used as a microbial sensor.

3.3. Immobilization of the Microorganisms onto the Beads for a Fabrication of the Reactor-Type Microbial Sensor There are two methods used for immobilization of microorganisms beads, the physical adsorption method and entrapment method.

onto the

3.3.7. Physical Adsorption Glass, charcoal, collagen, and chitm-chitosan mg microbes. An example of the protocol is as follows:

beads are used for immoblliz-

1 Add 20 mL of porous glass beads (pore size, 120 pm; sphere diameter, 0.4-l .Omm; Schott Nippon KK, Tokyo, Japan) to 100 mL of an overnight culture of S cerevzszae and keep standing at 4°C for 24 h. 2. Wash the unmobihzed-yeast beads thoroughly with stenlized water 3 Pack the unmoblllzed-yeast beads into a sterihzed 20-mL syringe, both ends of which are stuffed with sterllrzed cotton This syringe packed with immoblhzedyeast beads serves as an immobilized-yeast reactor (see Note 1).

3.3.2. Entrapment Several materials, such as, agar, algmic acid, K-carrageenan, gelatin, collagen, polyacrylamide, and polyvinylalcohol can be used as entrapment materials. The general method of Immobilization, using algimc acid gel as an example, is described as follows: 1. Suspend microorganisms (2-2 5 g wet wt) in 10 mL of 0.9% sodium chloride solution. 2. Mix the cells-salme suspension m 12.5 mL of 2% sodium algmate solution. 3. Place the final mixture in a syringe fitted with a needle and add dropwise 300 mL of stirred 1% calcmm chloride solution to yield calcnnn alginate beads (see Note 2).

3.4. Measurement

System

A schematic diagram of the batch-type measurement system is shown m Fig. 3. 1. Fill a thermostatically controlled, water-Jacketed vessel with 30 mL of 0.01 A4 Tris-HCl buffer (pH 7.0) contammg 150 mg/L glucose. Control the temperature of the buffer solution at 30°C. Addition of glucose IS indispensable to get a stable response to cyanide. It seems that pH of the solution does not greatly affect the response to cyanide but it is preferable to use a buffer solution of pH 7.0 or 8.0 because toxic hydrogen cyanide gas 1s generated at low pH (see Note 3).

Respiratory inhibition

233

2. Connect the microbial sensor to the recorder. A IO-k0 resistor should be connected m parallel with the microbial electrode. 3 Place the microbial sensor into the buffer solution m a water jacket. The solution should be constantly stirred during the measurement to ensure that tt is saturated with oxygen. Measure the output of the microbial electrode with an electromc recorder 4. Add the cyanide solution to the buffer solution when the output current becomes stable. Record the current change. The response of the microbial sensor to cyanide is determined by calculatmg the difference between the current before and after the addition of cyanide. Less than 2 mm should be required for the current from the mtcrobtal sensor to stabthze after addition of cyanide to the buffer solution 5, Make a calibration graph by measuring different concentrattons of cyamde When S cerevzszae is used as an munobihzed mtcroorganism, and the measurement 1s carried out in 0.01 A4 Tris-HCl buffer (pH 7.0) containing 150 mg/L glucose at 3O”C, a linear response should be observed m the range of 0.3 @fto 15 pA4 cyanide.

3.5. Flaw-Type

Measurement

Instead of the batch-type

System

system, a flow-type

microbial

sensor can be fabrr-

cated. A flow-type microbial sensor is sometimes more suitable for continuous monitoring

than the batch-type

one. There are two types of flow-type microbial

sensors,One is the membrane-type flow system and the other is the reactor type. A membrane-type microbial sensor (flow system)can be fabricated as follows: 1. Place a bottle of 0.0 1 A4 Tris-HCl buffer (pH 7 0) contaming 150 mg/L glucose m a thermostatic water bath Stir the buffer solutton constantly during the measurement to ensure that it 1s saturated with oxygen 2. Connect the bottle containing the buffer solution to a flow cell containing the microbial electrode with slhcon tubes (id, 2 mm; od, 4 mm). The buffer solutton is pumped to the mtcrobtal electrode by peristaltic pump (Mmtpuls 3, Gtlson, France) and the outlet of the flow cell fed to waste. 3 The measurement procedure IS the same as described m Subheading 3.3.

In the case of a reactor-type microbial sensor, a reactor that contams immobilized microbe beads is used instead of a microbial electrode that has an immobilized microbe membrane. Better stability is expected with a reactortype microbial sensor than with a membrane-type microbial sensor. 4. Notes 1. The bindmg forces prevalent in physical adsorption are hydrogen bonds, van der Waal’s forces, and electrostatic forces. Physical adsorption methods are simple and they do not harm the microorganisms as much. The dtsadvantage of this method 1sthe leakage of the immobtltzed microorganisms See Subheading 3.3.1.

234

Arikawa, Ikebukuro, and Karube

2. Alginic acid gels do not harm the munobilized microbes and have good permeability to the analyte. For this reason these gels are used in a wide range of experiments However, alginic acid gels suffer from instability, especially in the presence of phosphate ion. Polyacrylamide is often used as an alternative material See Subheading 3.3.2. 3. As little as 0.15 g of potassium cyanide can be lethal. Hydrogen cyanide is more toxic than potassium cyamde Hydrogen cyanide might be formed below pH 5 0; therefore, keep the pH of the solution above 7.0 during the experiments. This sensor system is very sensitive, and thus can measure a potassium cyanide solution with a concentration = 1 55x + 2.1, R* = 0.975.

3.4.4. OF Detection in a Sample Sample preparation: Take sample, add CHES to obtain a final concentration of 1 mA4, 100 mA4NaCl and 50 mA4 MgCl,; adjust pH to 9.0 (see Note 2).

Po ten tiome tric Detection

pH sensitjve ISPET

243

C

Sealing

Biocatalyst

Fig. 3. Schematicdiagramof (A) a flowthrough biosensor,(B) a flowthrough measurementcell and (C) minireactor. (A) (1) buffer tank, (2) 4-way valve; (3) peristaltic pump; (4) thermo-controlledblock; (5) minireactor;(6) measurementcell; (7) pH-sensitive ISFET probe; (8) pH meter; (9) recorder.Parts4-6 are constructedof Plexiglas. 1. Fill the measurementcell with a sampleand register changesin pH for 10 min. 2. Calculateparaoxonconcentration in thesampleusingthecalibrationcurve(seeNote 6).

3.5. Flowthrough System Biosensor 3.5.1. Construction of Flowthrough System 1. The schematicdiagram of the flowthrough system is presentedin Fig. 3A and consistsof a buffer tank, four-way valve, a peristaltic pump, a temperature-controlled block with measurementcell, a minireactor 5, and a pH-sensitive ISFET probe. Transducer signals are measuredby a pH meter, and registered by a recorderor computer.The flowthrough measurementcell (Fig. 3B) hasa volume of 100 ltL and the pH probe is hermetically fixed into the cell by a plug and silicon O-ring. All connectingtubes (Tygon) should be as short as possible (the internal volume of the systemis 2.5 mL).

244 3.5.2. Measurements

Simonian, Rainina, and Wild with the Flowthrough System

1 Calibrate the pH-sensmve ISFET probe with standard buffer solutions before assembling with the measurement block. Wash thoroughly with distilled water and dry. 2. Fill the minireactor with immobilized biocatalyst; it must be fully packed with biocatalyst. Remove air bubbles. 3. Insert the packed mmireactor mto the temperature-controlled block, install the pH-sensitive ISFET probe into the measurement cell, thermostabihze system (30”(Z), and switch on the peristaltrc pump (flow rate 1.0 mL/mm). Remove any air bubbles from the flow system. 4. Check the flowthrough system stability. Turn the valve 90’ and close the flow loop. Record the pH signal for 20 min The pH level should be 9 00 _+0.05 durmg this time. Turn the valve 90” back and open the flow loop

3.5.3. Calibration of the Flowthrough Biosensor 1, Fill the flowthrough system with working buffer contammg 1 rnM paraoxon (5 mL), turn the valve 90” and close the flow loop. Record pH variation for 20 min. Calculate ApH,, = pHlnltlal - pHafter 20,,,,,, 2 Substnute the working buffer containing paraoxon with working buffer alone Turn the valve 90” to open the flow loop. Wash the system thoroughly until the pH level reaches 9.00; the system is ready for a new measurement (see Note 5). Figure 4 presents a typical experimental curve obtained as a result of procedures described m step 1 3. Repeat the procedure described in step 1, with different paraoxon concentrations in the range l-1000 pA4 and plot a calibration curve. After each measurement repeat the procedure described m step 2. 4 Use DeltaGraph software to plot the calibration curve as ApH2s vs paraoxon concentration (concentration axis will be logarithmrc). Figure 5 shows the dependence of ApHZo on paraoxon concentration for this flowthrough system. The inset graph demonstrates a linear relationship for paraoxon m the range l-100 pM. The equation ~sj(x) = 0.007x + 0.32, R2 = 0 964.

3.5.4. OP Detection in a Sample Sample preparation:

Prepare sample as indicated

in Subheading 3.4.4.

1. Fill flowthrough system with sample (5 rnL). Turn the valve 90” and close the flow loop. Record pH variation over 20 mm. 2. Calculate paraoxon concentration m the sample according to the calibration curve (see Note 6).

3.6. Repefitiwe Use of Biocatalyst 1 The same biocatalyst may be reused for 20 or more consecutive analyses if the system is thoroughly washed after each measurement and stored m appropriate conditions when not use.

Po ten tiorne tric Detection

245

paraoxon 101

I

6

5

io

is

io

is

30 35

time, min. Fig. 4. A typical experimental time response of flowthrough brosensor. Experimental conditions: working buffer, initial pH 9 0, 500 mg biocatalyst wt in minireactor, 1 rnMparaoxon, temperature 30°C flow rate 1 .OmL/min. The arrow indicates solution replacement.

-r

1

“’

““‘I

10

“I

100

dO0

[paraoxon], PM Fig. 5. The dependence of ApH on paraoxon concentratron. Experimental conditions: working buffer, initial pH 9.0,500 mg biocatalyst wt in minireactor, 30°C flow rate 1.0 mL/min. The mset graph demonstrates a linear relattonshtp for paraoxon m the range l-100 mM. The equation is&v) = 0.007x + 0.32, R2 = 0.964.

Simonian, Rainina, and Wild

246

3.7. Storage of Biocatalyst Properly prepared biocatalyst is stable for at least 3 mo. To maintain OPH activity of the immoblhzed cells, take the following precautions: 1 Store the biocatalyst in storage buffer at 6-8”C, and change the buffer weekly 2 To store the packed minireactor, flush it with fresh storage buffer and hermetltally close with caps (Fig. 3C) Replace buffer regularly, at least weekly.

4. Notes 1 For convenience, prepare 10 mMCHES, pH 9 0, for use as a stock solution. Immediately before analysis, dilute the stock buffer, adjust pH If necessary, and use for one working day. The system should be closed to eliminate contact between air and

buffer, becauseof the high CO2solublbty in aqueoussolution. 2. The sensltlvlty of potentlometrlc blosensors depends primarily on the buffer capacity of the solution used for analysis-the lower the buffer capacity the higher the sensitivity (16,18). However, such diluted buffers are very unstable. This limitation can be mimmlzed by mtroducmg additional neutral ions into the buffer, such as NaCl and MgCl*, at concentrations that provide a stable (1 mA4 CHES) buffer, without influencing the biosensor sensitivity. 3 This step must be done very thoroughly to achieve uniform distribution of cells in the very viscose polymer solution Keep mixture at room temperature at least for 30 min for removal of air bubbles 4. Adhere to the instructions for the pH electrodes and ISFET probes used for the analyses and storage. Electrode-sensitive components might bind analyte, which would lead to a loss of a stable signal. To eliminate the adsorbed analyte from the surface, clean electrodes as described by the manufacturers. 5 The accuracy of measurements is of fundamental importance To achieve a stable system, paraoxon or other analytes must not be adsorbed on the surface of any part of the blosensor or the results will be compromised. Thus, the mamfold parts of both the flow and batch systems in contact with OPs should be thoroughly washed after 35-40 measurements with a 2% (w/v) NaOH solution followed by an extensive dlstllled water rinse Tygon tubing should be used because of its low adhesion characteristics. 6 When performing analysis with the real samples for either batch or flow blosensors, assure that all of the components present m the sample do not affect the physical transducer. For this reason, repeat the procedures described m Subheading 3.4.2., step 2 and Subheading 3.5.2., steps 3 and 4 without biocatalyst.

Acknowledgment This research was developed with the financial support of a University Research Initiative from the US Army Research Office, NATO Grant LG 941411, Research Enhancement Program from the Texas A&M University System, and by the NRC-CAST programmatic support of E. I. Rainina.

Poten tiome tric Detectlon

247

References 1. Bousse, L., Kirk, G., and Sigal, G. (1990) Biosensors for detection of enzymes immobilized in microvolume reaction chambers. Sens Actuators Bl, 555-560. 2. Moo-Young, M., ed. (1988) Bioreactor Immobilized Enzymes and Cells. Fundamentals and Appkcattons. Elsevier Applied Science, London, UK. 3. Karube, I. and SangMok Chang, M. E (199 1) Microbial biosensors, in Bzosensor PrincipZes and Applications (Blum, L. J. and Coulet, P. R., eds.), Marcel Dekker, New York, pp. 267-301. 4. Tran-Mmh, C. (1993) Biosensors. Chapman & Hall, London, UK. 5. Rechnitz, G. A., Kobos, R. K., Riechel, S. J., and Gebauer, C. R. (1977) A bioselective membrane electrode prepared with hvmg bactertal cells. Anal Chzm Acta 94, 357-365. 6. Hukuma, H., Obana, H., Yasuda, T , Karube, I., and Suzuki, S. (1980) A potenttometrrc microbial sensor based on unmobihzed Escherzchza colz for glutamic actd, Anal. Chim. Acta 116,61-67. 7. Jensen, M. A. and Rechnitz, G. A. (1978) Bacterial membrane electrode for h-cysteine. Anal. Chim. Acta 101, 125-130. 8 Rechnitz, G A , Riechel, T L , Kobos, R K., and Meyerhoff, M E (1978) Glutamate selective membrane electrode that uses living bacterial cells. Scrence 199,440,44 1 9. Di Paolantonio, C. L., Arnold, M. A., and Rechnitz, G. A (1981) Serine-selective membrane probe based on immobilized anaerobic bacterta and a potentiometric ammonia gas sensor. Anal Chim Acta 128,121-127. 10. Matsumoto, K., Seijo, H., Watanabe, T., Karube, I., Satoh, H., and Suzukt, S. (1979) Immobihzed whole cell-based flow-type sensor for cephalosporms. Anal. Chum. Acta 105,429-432. 11. Dave, K., Miller, C., and Wild, J. (1993). Characterization of organophosphorus hydrolases and the genetic manipulation of the phosphortriesterase from Pseudomonas dimtnuta. Chem. Biol Interac 87,55--68 12. Lozinsky V. I., Faleev, M. F , Zubov, A. L., Ruvinov, S. B., Antonova, T. V., Vainerman, E. S., Belikov, V. M., and Rogogin, S. V. (1989) Use of PVA-cryogel entrapped Citrobacter intermedtus cells for continuous production of 3-fluoro+ tyrosine. Biotechnol. Lett. 11(l), 43-48 13. Rainma, E. I., Varfolomeyev, S. D., Dave, K., and Wild, J. R. (1994) Cryounmobrhzation of organophosphate neurotoxin degrading bacteria in poly(viny1) alcohol. Proceedings of the 1993 ERDEC Scienttfk Conference on Chemical Defense Research, US Army, Edgewood, MD, pp. 895-900. 14. Rainina, E. I., Badalian, I. E., Ignatov, 0. V., Fedorov, A. B., Simonian, A. L., and Varfolomeyev, S. D. (1996) Cell biosensor for detection of phenol in aqueous solutions. Appl. Bzochem. Bzotechnol. 56, 117-127. 15. Simonian, A. L., Rainina, E. I., Lozinsky, V. I., Badalian, I. E., Khachatrian, G. E., Tatikian, S. Sh., Makhlis, T. A., and Varfolomeyev, S. D. (1992) A biosensor for L-prolme determination by use of immobilized mtcrobial cells. Appl Bzochem. Btotechnol. 36, 199-210.

248

Simonian, Rainha, and Wild

16 Rainina, E. I., Efremenko, E N., Varfolomeyev, S D., Simonian, A L., and Wild, J. R. (1996) The development of a new biosensor based on recombinant E colt for direct detection of organophosphorous neurotoxms. Blosens Bioelec@on. 11,991-1000 17 Lozmsky, V. I. and Zubov, A. L. (1995) Russian Patent No. 2036095 18. Kumaran, S. and Tran-Minh, C. (1992) Determination of organophosphorous and carbamate msecticides by flow-inJection analysts. Anal Blochem 200, 187-194.

Microbial Biosensors Based on Optical Detection Udayakumar Matrubutham and Gary S. Sayler 1. Introduction In general, a biosensor is an analytical tool with a biologic reactive component interfaced or integrated to a physicochemical transducer. A microbial biosensor based on optical detection employs whole-cell microorganisms to catalyze, sense, and transmit optical signals. These signals may correspond to light emission, reflection, fluorescence, or absorption. The microorganisms can be immobilized either on the transducer or in a separate compartment with the light signal transmitted through an optical medium to the transducer. Although cell-free, noncatalytic biosensors have been proposed, development of whole cell optical biosensors based on bioluminescence and chemiluminescence have been reviewed adequately (1,2). Few research groups have constructed and demonstrated the application of bioluminescent reporter bacteria in biosensing environmental processes (3). Bacterial bioluminescence is a fascinating natural phenomenon that has enabled the development of gene reporters in molecular studies.Primarily, the bioluminescence genes in the marine bacteria VibrioBshceri are used in the construction of ZUXgene reporters; however, genes from other bacteria have also been used in a few cases(4). In bioluminescent reporters, a promoterless Zuxgene cassette is fused to an inducible gene promoter that when induced encodes enzymes for bioluminescence. Typically, the gene cassettemay carry 1uxCDABE or IuxAB genes. The former encodes the enzymes luciferase and fatty acid reductase, synthetase, and transferase, whereas the latter encodes only luciferase. When using ZuxAB in the gene fusion, an external source of aldehyde substrate must be supplemented to the luminescence reaction, because it is an important com-

From

Methods m Blotschnology, Vol 6 Enzyme and Ucrobfaal Bfosensors Technques Edited by A Mulchandanl and K R Rogers 0 Humana Press Inc , Totowa,

249

and Protocols NJ

Matrubutham and Sayler

250

A

zz4x

1

1 PROMOTER1 C

Lipids

1 D

1A

1 B

1E

1

f

Synthetase/Reductase

: t Myristoyl-ACP

AMP+ Transferase I-120

t t Acetyl-CoA

B

Naphthalene -+

0 Salicylate

Salicylate

Salicylate ++

TCA cycle

NahR Regulatory protein

Promoter I

Fig. 1.(A) Bioluminescence reactionindicatingtheregenerationof fatty acicl-aldehyde coupleand the emissionof light; (B) Schematicrepresentationof an nab-Zux fusionwith the mechanismanddirectionof geneexpressionindicatedby arrows. ponent otherwise regeneratedin vivo. Figure 1 provides an overview of the bioluminescence reaction with a ZuxCDABE cassetteand indicates the cyclic regeneration of the fatty acid-aldehyde couple with the emission of bluegreen light (490 nm). The figure also depicts the strategic insertion of the Zux genesin the creation of a Zuxreporter, viz., nab-ZUXfor a naphthalenebiodegradation system (5). The Zuxgene reporters have been shown to be useful in understanding plant pathogenesis,rhizosphere bacterial colonization, mutagenesisand dynamics of geneinduction and expression(S-9). In recentyears,the bioreporter n&z-ZUX

Optical Detection

251

of the naphthalene biodegradation system has been demonstrated as a model system in investigating environmental processes, such as bioavailability and biodegradation, and in depicting the advantages of Zuxbioreporters and microbial optical biosensors in real-time process monitoring (1UJ1). The reporter gene nah-Zux had been constructed by a transpositional fusion of the 1uxCDABE gene cassettewith the nahG promoter of an NAH7-archtypal naphthalene degradation pathway in Pseudomonasfluorescens HK44 (5). The bacteria emits light when induced with naphthalene or its metabolic intermediate, salicylate. A few other lux fusions have also been created for studying different environmental systems.For instance, the tod-Zux and mer-lux fusions enable the detection of toluene and mercury, respectively (12,13). Bioluminescence is detectable with any light-detection device. Review of recent studies on the development and use of optical biosensor systemsmay be resourceful for a novice in this area (14). The fundamental features of any light device should include its ability to capture and multiply low quantities of photons. In general, the photomultiplier tube in a scintillation counter should be adequate for most laboratory techniques. However, specialized instruments may be required, depending on the application of the optical biosensor. The design of the optical component of a specialized instrument must accommodate the intensity and location of the light source. For example, the sensitivity of the light detector and the transmittance capacity of the conducting medium are crucial in the detection and transmission of low-intensity light over a long distance. Nevertheless, the essential components in designing an optical biosensor include a photon-detection unit (e.g., photomultiplier), an optional light-transmission device (e.g., liquid light guide or fiber-optic cables) and a converter unit to transform the light signals into electrical readings. The’reporter cells may be immobilized on the sealed end of the liquid light guide as for on-line biosensor or positioned in a container in series to the light guide. Figure 2 provides a schematic presentation of such a light-detection system. The biosensor tip represented in that figure had been designed for real-time light monitoring with P. jluorescens HK44 (9). Alternate photon-detection devices like the avalanche photodiode and CCD camera could also be used in these systems instead of the photomultiplier, if cost is not a prohibitive factor. The liquid light guide and fiber-optic bundle perform similar functions, using internal reflection to guide energy along the entire path to the photomultiplier. The liquid guide, however, has higher transmittance than fused silica bundles of equal size and is also very flexible. The guide is primarily a plastic tube tilled with a transparent, nontoxic fluid. The tube has sealed ends with polished windows to provide efficient interlocking capability with stainless steel adapters. As shown in Fig. 2B, the biosensor tip made with the sealed end

Matrubutham and Sayler

252

A I.........~

I

I

0 0 I

Photomultiplier

Digital readout unit

B

“...’ .‘;iLight tight box I Liquid light guide

Coaxial cable

Stainlesssteel ferrule

I Polishedwindow end

less steel mesh

Liquid guide

I Adapter coupler

Fig. 2. (A) Essential components and assemblageof the optical detection system; (B) Schematicof a biosensortip designedwith a liquid light guide for on-line analysis.

of the liquid light guide could be placed inside a stainlesssteel ferrule and held strongly in any position (9). Finally, dependingon the biosensorapplication, the microbial sensorcan be constructed in a suitable bacterial host and the light-detection system developed for either continuous or discontinuous monitoring of light signals. Assay development and ingenious biosensor techniques are delineated in this chapter. Assuming that the performer possessesthe appropriate bioluminescent reporter bacteria, the protocols for bacterial immobilization, optical detection, the design of a light-detection system, and the ultimate setup of the biosensor are provided. 2. Materials 2.1. Biological

Component

1. Bacterial culture medium: Any suitable broth medium to grow the bioreporter bacteria; for example,P. fEuore.scens HK44 is usually grown in YEPG medium.

253

Optical Detection

YEPG consists of the following in g/L distilled water. Yeast extract, 0.2, polypeptone, 2.0, glucose,1.O,andNH,NO,, 0.2. 2. Alginate solution: Sterile, 3% Na-alginate (low viscosity, Sigma, St.Louis, MO) dissolved in 0.85% NaCl (seeNote 3). 3. Strontium chloride: Sterile, 0.1 A4in distilled water. 4. Others:Sterile glycerol, sterile emptyglassbeaker (250 mL) andstir bar, cheesecloth, sterile IO-mL disposablesyringes,26-gageneedles,rotary shaker. 2.2. Optic& Component 2.2.1. Light-Detection Equipment 1. Light detectorandamplifier:Photomultiplier(Oriel,Stratford,CT, USA,model77340). 2. Light transmitter: Liquid light pipe (Oriel, model 77554) or glass fiber-optic bundle. 3. Output readout: Digital detectionunit (Oriel, model 7070). 2.2.2. Light-Detection

System for Growing Cell Assay

The following is a description for a very simple but efficient light detection system. As shown in Fig. 2A, connect the photomultiplier to the digital detection unit with a coaxial cable. The detection unit must be in series to an electrical power input. Interface one end of the liquid light guide to the photomultiplier and position the other end inside a light-tight box, through a light tight aperture. The sealed end of the liquid light guide inside the box should be in series with any holder or clamp device for holding the sample container. The box can be made of any material and must provide an access port for placing and removing the sample in the holder. 2.2.3. Light-Detection

System for On-Line Analysis

Figure 2B presents a schematic of a biosensor tip for on-line monitoring of light signals. The liquid light guide can be coupled with any on-line system with proper adapters. Depending on the type of system,a stainless steel adapter can be procured to fit the free end of the liquid guide (the other end is interfaced with a photomultiplier). As shown in the figure, snugly fit the sealed end of the liquid guide with a stainless steel ferrule. The ferrule can be held in place with a set-screw. Cut a piece of stainless steel wire mesh (104 urn; Spectrum, Houston, TX) to fit over the polished window end of the liquid guide. This mesh can be held in place with a wire string or an O-ring and fitted inside a shallow groove machined on the ferrule. The adapter coupler can be used in connecting the end of the liquid guide to a three-way T-connector, such that the tip of the light guide will be on-line to the inflow and outflow of any continuous system. The adapter coupler car he obtained in any size depending on the type of connectors.

Matrubutham and Sayler

254 3. Methods 3.1. Growing

Cell Assay

1. Initiate a fresh bacterial culture from an overnight starter culture and allow it to reach mid-log phase. 2. Transfer 2 mL of the culture to a 25-mL mineralization vial with a tightlid (see Notes 5 and 6). 3. Add 2 mL of test solution to achieve the desired final concentration of the test compound. 4. Shake the vials in a rotary shaker at desired rpm (see Note 7). 5. Monitor bioluminescence over a period of time with the light-detection system for a batch analysis (see Fig. 2A). See Note 10. 6. Record optical density of cultures at the end of the assay to normalize light data to the actual cell numbers per sample. 7. Check spurious light production with suitable controls (see Notes 9 and 11). 8. Always perform the assay at ambient conditions (see Notes 1 and 2).

3.2. On-Line Biosensor 3.2. I. On-Line Biosensor Probe Tip Preparation Refer to Fig. 2B, ref. 9. 1. Prepare an exponential phase culture, wash cells in saline (0.85% NaCl) followed by centrifirgation, and resuspend in saline (see Note 4). 2. Mix 12.5 mL of the cell suspension with 25 mL of low-viscosity alginate solution (3.5%) and 7.5 mL of sterile glycerol. 3. Aliquot the mixture into 1.5-mL Eppendorf tubes and store at -7O’C. 4. Inject the mixture into the space between the wire mesh and the liquid light pipe (shown in Fig. 2B) inside the ferrule with the help of a 1-mL syringe fitted with 26-gage needle without trapping any air bubbles. 5, Immediately immerse the tip into a stirred, sterile 0.1 A4 &Cl2 solution in a beaker and keep immersed for 1 h at room temperature to congeal the alginate-SrC12 matrix. 6. The probe tip is now ready and can be installed on-line for monitoring (see Note 8).

3.2.2. Preparation of Beads The biosensor tip may contain beads of immobilized cells instead of the matrix. The beads can also be used in biosensor design, such as porous metallic sensor module, and installed in any environment of interest. The protocol to prepare the alginat&rCl, beads is as follows and can be scaled up as needed: 1. Prepare exponential phase culture, wash, and resuspend in saline as in the previous protocol. 2. Mix 1 part cell suspension with two parts alginate solution and keep the mixture on ice. 3. Start stirring 0.1 M SrC12 solution in a sterile glass beaker on a stir plate at room temperature.

Optical Detection 4. 5. 6. 7. 8. 9.

255

Transfer the alginate-cell mixture mto a lo-mL disposable syringe. Push the nnxture slowly through a 26-gage needle to form drops mto the stirring &Cl,. The drops will solidify into algmate beads. Allow the beads to stir in SrC12 for at least 45 min. Remove the beads by decanting the solution through sterile cheesecloth. Transfer the beads into a sterile tightly capped container (disposable centrifuge tube) and store at 4’C until use.

4. Notes 1 Perform biolummescence assays in pH regimens ranging between 6.0 and 7 0. 2. Keep the temperature ambient (23-28%). 3. Before autoclavmg, dissolve the alginate in saline by stnrmg overmght Otherwise, algmate clumps will form and alter the viscosity. 4. Glycerol-alginate stocks of the bacteria may be substituted with freshly prepared mixtures since storage can affect the biosensor performance. 5 For growing cell assays, it is preferable to keep the volume at a minimum and have a 1: 1 ratio of cell suspension to test solution. 6 It is also preferable to work with a low density of cells because the lux reaction demands a hrgh level of oxygen. 7 Always return the samples back to the rotary shaker smce growing cells need oxygen and so does the lux reaction. 8. Avoid using phosphate buffers because they disrupt the integrity of alginateSrC12 matrix or beads 9. Avoid the uncontrolled use of materials dissolved in organic solvents m the broluminescence reaction, because solvents tend to perturb bacterial membrane lipids and increase the amount of light emissron (see ref. 15). 10 Monitor biolummescence in short time intervals, because subtle variations m the light synthesis can be missed easily. 11. Validate the induced light production with molecular analysis for lux transcripts (mRNA analysis), elimmatmg the possibility of falsely induced hght and confirming the bioreporter strain construction.

Acknowledgment Critical note comments by Staci Kebrmeyer, Bruce Applegate, and Janeen Thonnard are highly appreciated. The work was supported by the DOE grant DE-FG05-94ER6 1870.

References 1. Bronstein, I., Fortin, J., Stanley, P. E., Stewart, G. S. A. B., and Krrcka, L. J. (1994) Chemiluminescent and biolummescent reporter gene assays. Anal Biochem. 219, 169-181. 2. Schugerl, K., Hitzman, B., Jurgens, H., Kulick, T., Ulber, R., and Weigal, B (1996) Challenges in Integrating biosensors and FIA for on-lme monitoring and control. Trends Biotech. 14,2 1-3 1.

Matrubutham and Sayler

256

3. van der Lehe, D., Corbisier, P., Baeyens, W., Wuertz, S., Diels, L., and Mergeay, M. (1993) The use of biosensors for environmental monitormg. Presented in EEROVLAB workshop on Environmental Technology, Nov 2 l-25, Grobbendonk, Belgium. 4 Stewart, G. S. A. B. and Wtlhams, P. (1992) lux genes and the applications of bacterial bioluminescence. J. Gen. Mlcrobiol. 138, 1289-1300. 5. King, J. M. H , DtGrazia, P M., Applegate, B., Burlage, R., Sansevermo, J , Dunbar, P., Larimer, F., and Sayler, G. S. (1990) Rapid, sensitive biolummescence reporter technology for naphthalene exposure and biodegradation Science 249,778-781. 6 Shaw, J. J. and Kado, C. I. (1986) Development of a Vibno biolummescence gene-set to monitor phytopathogemc bacteria during the ongoing disease process in non-disruptive manner. &o/Technology 4,560-564. 7. Shaw, J. J., Settles, L. G., and Kado, C I. (1988) Transposon Tn443 1 mutagenesis of Xanthomonas campestrls pv. Campestrls. characterization of a non-pathogenic mutant and cloning of a locus for pathogemcity. Mol. Plant-Mzcrobe Interact. 1,3!%45

8 Shaw, J. J , Dane, F., Geiger, D., and Kloepper, J. W. (1992) Use of biolummescence for detection of genetically engineered microorganisms released mto the environment. Appl. Environ. Mlcroblol. 58,267-273. 9. Brennerova, M. V and Crowley, D. E. (1994) Direct detection of rhlzosphere colonizing Pseudomonas sp. using an Escherichia ~011rRNA promoter in a Tn7-lux system. FEMS Mlcroblol Ecol 14,3 19-330 10. Heitzer, A, Malachowsky, K., Thonnard, J. E., Bienkowskt, P. R , White, D C , and Sayler, G. S. (1994) Optical biosensor for environmental on-lme monitoring of naphthalene and salicylate bioavatlability with an immobilized bioluminescent catabolic reporter bacterium. Appl Environ Mzcrobzol. 60, 1487-l 494. 11. Hettzer, A., Webb, 0. F., Thonnard, J. E., and Sayler, G. S. (1992) Specific and quantitative assessment of naphthalene and salicylate bioavailability by using a bioluminescent catabolic reporter bacterium. Appl Enwon Microblol. 58, 2046-2052.

12 Applegate, B., Lackey, L., McPherson, J., and Menn, F. (1993) A biolummescent reporter for the co-oxidation of tnchloroethylene (TCE) by the toluene dioxygenase m Pseudomonas putida F 1. Abstracts of the 93rd General Meeting of the American Society for Microbiology, Q- 108, May 16-20, Atlanta, GA 13. Selifonova, 0 , Burlage, R., and Barkay, T (1993) Bioluminescent sensors for detection of Hg(I1) in the environment. Appl Envrron. Microbrol. 59,3083-3090. 14. Burlage, R. and Kuo, C. T. (1994) Living biosensors for the management and manipulation of microbial consortia. Annu. Rev Microbrol, 48,291-309 15. Heitzer, A., Applegate, B., Pinkart, H., Webb, 0. F., Phelps, T. J., Sayler, G. S , and White, D. C (1997) Catabolic bioluminescent reporter bacteria: phystological aspects of environmental applications. J, Mxroblol Methods, in review.

Index A Activated ISFETs enzyme immobilization, 39,40, 43 Adsorption enzyme immobilization, 9 microbial immobilization, 20 1 respiratory inhibition microbial biosensors, 232, 234 Aerator catecholamme detection, 55,59,60 flow system, 55,56,60 Aerosol technique sensitive catecholamine detection, 53, 54 Aerosol vaporization pH electrode enzyme biosensors, 17, 19, 21 Alcohol measurement hydrogen peroxide electrode biosensors, 75 Alginic acid gels, 234 Alkaline phosphatase flow system scheme, 57f Alkaline phosphatase measurement bienzyme electrodes, 56,57,63,64 Alkyl peroxides peroxidase, 99 Ammonia gas probes, 27 Amperometric enzyme biosensors, 5-7

Amperometric enzyme electrodes selectivity, 133, 134 Arginine detection Streptococcus

faecium,

238

Aromatic amine detection HRP-modified electrodes, 101t Autoxidizable mediators NADH oxidation, 54

B Batch mlection analysis, 78 Batch-system biosensors potentiometric detection microbial biosensors, 241243 Bienzyme electrodes alkaline phosphatase measurement bienzyme electrodes, 56, 57,

63,64 enzyme recycling biosensors, 55-

57,59,60,62-64 phenol measurement, 56,57,63,64 Biologic oxygen demand (BOD),

207,225,238 biosensors, 2 18 Bioluminescence detection, 25 1 Bioluminescent reporters, 249 c Caliorimetry defined, 175

257

258 Calorimeters thermistor-based, 175 Calorimetric biosensors, 175-l 78 Carbon dioxide electrodes, 26 Carbon fiber electrodes organic hydroperoxrde detection, 102, 103, 104 Carbon fiber microelectrodes fabrication redox polymers enzyme biosensors, 124,126,127 Carbon paste electrodes, 108-l 09 glutamate detection, 103-l 05 hydrogen peroxide detection, 103-105 Catalytic cycle peroxidases, 93-95 Catecholamme detection aerator, 55, 59-62 Cell-based biosensor organophosphorus neurotoxins, 239 Cephalosporin detection whole-cell-based flow biosensor,238 Chemical binding enzyme immobilizatton gas electrodes enzyme biosensors, 28-29, 30 Chloroperoxidase, 108 Clark-type electrode based enzyme sensor schematic, 52f Conductimetric biosensors construction, 162 enzyme-based, 157-l 7 1 vs conductivity cell, 160-162 Conductimetric transducers, 7 Conducting polymers electrochemical production advantages, 144

Index enzyme biosensors, 143-l 55 enzyme immobrlization, 145, 146 formation, 143, 144 Conductivity measurement, 159163 Constant-charge circuit, 4 I ISFET enzyme biosensors, 39, 40,42 Covalent bonding enzyme mrmobilization, 10, 11 conducting polymer formatron, 146 polypyrrole deposmon, 147, 149, 154 Crosslmkable redox polymer synthesis, 123-125 Crosslmkmg enzyme immobilization, 10, 11 pH electrode enzyme biosensors, 16 Cryoimmobihzation potentiometric detection microbial biosensors, 24 1 Cyanide determination microbial sensors, 228 D

Direct binding method pH electrode enzyme biosensors, 16-l&21 E

E. co/i glutamic acid detection, 238 E. coli growth potentiometric detection microbial biosensors, 240 Eastman AQ, 9 Electrical conductivity measurement, 159-l 63

index Electrochemical transducers, 4-7 Electrochemical-impedence spectroscopy system conductimetric measurement enzyme biosensors, 170, 171 Electrolyte Insulator Semiconductor (EIS) chip, 5 Enantioselective analysis, 40,46,47 Environmental monitoring enzyme biosensors, 11 Enzymatically modified ISFETs, 36,37 Enzyme biosensors advantages, 199 amperometric, 5-7 applications, 1l-l 2 conductimetric measurement, 157-171 conducting polymers, 143-155 defined, 3 enzyme immobilization, 9-l 1 fluorescence quenching, 188 fluorometric detection, 187-l 95 future, 12 gas electrodes, 22-33 hydrogen peroxide electrode, 67-78 indicator-based, 187 ISFETs, 35-48 mediator-modified carbon paste electrode, 8 l-9 1 metallized carbon electrodes, 133-139 oxygen detection, 5 l-64 peroxidase electron transfer, 93-110 pH electrode, 15-22 potentiometric, 4-5 principles, 3-l 2 redox polymers, 12 l-l 3 1 thermal transducer/thermistor, 175-185

259 transducers, 3-9 Enzyme immobilization adsorption, 9 chemical methods, 10, 11 conducting polymer formation, 145, 146 controlled pore glass (CPG), 83, 85,86 covalent binding, 10, 11 crosslinking, 10, 11, 16 entrapment, 9 enzyme biosensors, 9-l 1 gas permeable membrane, 28-30,33 pH electrode enzyme biosensors, 15,16 pH-FET, 39,40,43-45,47 physical methods, 9, 10 thermal transducer/thermistor enzyme biosensors, 182 vs cell immobilization, 237 Enzyme Instability, 12 Enzyme membrane functions hydrogen peroxide electrode enzyme biosensors, 69, 70 Enzyme sensors vs microbial sensors, 20 1 Enzyme thermistors, 175 construction, 178 invertase characterization, 177 penicillin assay, 177 TELISA, 177 Enzyme-based conductimetric biosensors, 163, 164 urine analysis, 164 Enzyme-based tield effect transistor (ENFET), 4, 5 applications, 46,47 enantioselective analysis, 40,46, 47

260 F

Fermentation industry enzyme biosensors, 11, 12 Ferrocene mediated-enzyme biosensors, 82 Fiberoptic-based biosensors advantages, 9 Flow injection analysis (FIA) glucose biosensors, 85-87, SSf, 89,90 pH electrode enzyme biosensors, 17, 20, 22 potentiometric detection microbial biosensors, 243,244 respiratory inhibition microbial biosensors, 233,234 thermal transducer/thermistor enzyme biosensors, 178 Flow system aerator, 55, 56, 60 alkaline phosphatase scheme, 57f Fluometric detection enzyme biosensors, 187-l 95 substrate-induced quenching (SIQ) biosensor protocol, 191,192 Food mdustry enzyme biosensors, 11, 12 G Gas electrode assembly, 27-29,32,33 Gas electrodes enzyme biosensors, 22-33 enzyme immobihzation, 28-30 Glass pH electrodes, 237 Glassy carbon electrodes, 124, 126 Glucose measurement flow injection analysis, 85-87, 88f

Index mediator-modified carbon paste electrode enzyme biosensors, 83-90 oxygen detection enzyme biosensors, 55,58, 59,61,62 pH-FET, 44,45 Glucose microsensor scanning electron micrograph, 135f Glutamate detection carbon paste electrodes, 103-l 05 Glutamic acid detection E. co/i, 238 Glutamine measurement hydrogen peroxtde electrode biosensors, 75 Glutaraldehyde enzyme immobihzation, 40, 43, 44, 181 Graphite paste mediator-modified carbon paste electrode enzyme biosensors, 82, 83 H

Home care enzyme biosensors, 1I HRP-modified electrodes aromatic amine detection, 101t phenol detection, IO 1t Hybrid sensors oxygen electrode, 2 14, 2 15 Hydrogen peroxidases bioelectrocatalytic reduction, 96f Hydrogen peroxide bioelectrocatalytic reduction, 97f detection carbon paste electrodes, 103105 solid graphite electrodes,102, 103

Index electrode enzyme biosensors, 6778 commercial systems, 73 construction, 67-69, 73, 74 enzyme membrane functions, 69,70 production in oxygen catalyzed reactions, 52, 53 voltammogram, 68f I

Immobilon, 10 Immunodyne, 10 Impedance analyzers conductimetric measurement enzyme blosensors, 169, 170 Inhibitor effect oxygen electrode microbial sensors, 2 16 Intensive care unrts enzyme biosensors, 11 Invertase characterization ET, 177 Ion-selective electrode @SE)-based biosensor technology, 4 Ion-sensitive field-effect transistors (ISFETs) enzyme biosensors, 35-48 operation, 35-37 sensor evaluation, 40,44-46 types, 36t L

Lactoperoxidase, 108 Lactose biosensors, 75 Light-addressable potentlometric sensors (LAPS), 237 Liquid light guide, 25 1 Lux gene reporters, 250, 25 1

261 M

Malate enzyme biosensors, 58, 61, 64 Mediated-enzyme biosensors disadvantages, 8 1, 82 Mediator-modified carbon paste electrode enzyme biosensors, 81-91 Metal-oxide semiconductor fieldeffect-transistor (MOSFET), 35-48 Metallized carbon biosensors operation, 137-l 39 Metallized carbon electrodes enzyme blosensors, 133-l 39 enzyme microelectrode, 136, 137 enzyme paste electrodes, 136, 137 metal center selection, 134, 135 screen-printed biosensors, 136, 137 Microbial biosensor, advantages, 225 BOD cyanide organophosphorus, 228 cyanide determination, 228 disadvantages, 203,225, 226, 238 electrochemical devices, 2 10, 2 11 function, 201-203 optical detection, 249-255 oxygen electrodes, 199-220, 204t-206t, 226% 227,228 potentiometric detection, 237246

principle, 226 respiratory inhibition, 225-234 vs enzyme sensors, 201

262

Index

Microbial immobilizanon, 200,20 1,227 adsorption, 20 1 chemical methods, 200, 201 entrapment, 20 1 physical methods, 200, 201 Mlcrocalorimeters, 175 N NAD(P)H fluorescence measurement, 8 NADH detection-based enzyme biosensors, 189 NADH oxrdatron autoxidizable mediators, 54 NADH oxidase, 54 sahcylate hydroxylase, 52 Nafion, 9 Nah-lux reporters, 250, 25 1 NH3 electrode, 23, 24t, 25 Non-heme peroxidases, 108 0 Optical biosensors, enzyme, 249-255 microbial, 249-255 Optrcal transducers, 8, 9 Orgamc hydroperoxide detection carbon fiber electrodes, 102-l 04 Orgamc peroxides detection aqueous solutions, 1OOt Organophosphorus neurotoxins cell-based btosensor, 239 Oxidase/peroxrdase brenzyme electrodes, 101, 102 Oxrdo-reductase enzymes, 6 Oxygen catalyzed reactions hydrogen peroxide production, 52,53

Oxygen detection enzyme biosensors, 5 1-64

alkaline phosphatase measurement, 54 disadvantages, 51, 52 microbial sensors, 203 phenol measurement, 54 Oxygen electrode couplmg enzyme reactions, 52-54 Oxygen electrode microbial sensors, 199-220,204t-206t measuring instrumentation, 211,2 I2 microbial sensor optimization, 213-215 mrcrobral sensor preparation, 212,213 mrcroorgamsms, 209, 2 10 physrologrc effect, 2 16-2 18 protectwe membranes, 210 Oxygen electrodes microbial sensors, 227, 228 Oxygenases, 53, 54 P Penicillin assay ET, 177 Peroxidase electrodes stabihty, 109, 110 Peroxidase electron transfer enzyme biosensors, 93-l 10 Peroxidase-modrfied electrodes application, 97-102 configurations, 98t, 99t Peroxidases alkyl peroxides, 99 catalytic cycle, 93-95 direct ET, 95, 96 mediated ET, 96, 97 physiochemrcal properties, 93-95 pH electrode enzymebrosensors,15-22 enzyme immobrlization, 15, 16 operation, 15

index pH-FET diagram, 37f enzyme immobilization, 39,40, 43-45 pH-sensitivity measurement, 39, 42-44 substrate enzyme systems, 38t Phenol detection, 100, 101 bienzyme electrodes, 56, 57, 63, 64 HRP-modified electrodes, 101t solid graphite electrodes, 102, 103 Platinization conductimetric measurement enzyme biosensors, 168, 169 covalent polymers enzyme biosensors, 147-149, 153 154 Polypyrrole deposition covalent polymers enzyme biosensors, 147, 149, 154 Potentiometric detection enzyme biosensors, 4,5 enzymes, 5t Potentiometric detection microbial biosensors, 237-246 Preactivated membranes gas electrodes enzyme biosensors, 29, 30,33

R Redox polymers enzyme biosensors, 121-131 crosslinkable redox polymer synthesis, 123-125 electrode modification, 124, 127, 128 sensor evaluation, 128-l 31

263 Reinforced membranes pH electrode enzyme brosensors, 17, 18,21 Respiratory inhibition mrcrobral biosensors, 225-234 Reticulation conductimetric measurement enzyme biosensors, 169 s S-phenylalanine methylester, 47f Salicylate hydroxylase NADH oxidation, 52 Sensor evaluation ISFETs, 40,44-46 redox polymers enzyme biosensors, 128-13 1 thermal transducer/thermistor enzyme biosensors, 184 Silanization conductimetric measurement enzyme blosensors, 132 Solid graphite electrodes hydrogen peroxide detection, 102,103 phenol detection, 102, 103 Streptococcus faecium arginine detection, 238 Substrate uptake oxygen electrode microbial sensors, 2 16,2 17 Substrate-induced quenching (SIQ) biosensor protocol fluometric detection, 191, 192 schematic, 190f Substrate-recycling sensors, 53 Sucrose measurement hydrogen peroxide electrode biosensors, 74

264

Index

T TELISA enzyme thermosistors, 177 Thermal transducer/thermistor enzyme biosensors, 175-I 85 enzyme activity measurement, 182, 183 enzyme immobilization, 182 flow-injection system, 178 reverse immobilization, 182 Thermal transducers, 7-9 Thermistor thermal transducer/thermistor enzyme biosensors, 179 Thermistor-based calorimeters, 175 Transducers, 3-9 amperometric, 5-7 conductimetric, 7 electrochemical, 4-7 optical, 8, 9

potentiometric, 4, 5 thermal, 7, 8 types, 4t

U Universal Sensors, 72, 72t Urea sensors response time, 28f Urine analysis enzyme-based conductimetric biosensors, 164

W Whole-cell-based flow biosensor cephalosporin detection, 238 Y Yeast biosensor, 228 YSI membrane biosensors, 7 1t YSI 2300 Stat Plus analyzer, 74