Electrochemical Toxicity Sensors

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James F. Rusling University of Connecticut, Storrs, Connecticut, U.S.A.

INTRODUCTION Medical abnormalities from exposure to toxic chemicals constitute a critical public health problem in our modern world. The use of toxic chemicals in agriculture, heavy industry, and other human endeavors throughout the past century has led to widespread pollution of our environment with potential disease-causing substances.[1] New drugs also have important toxicity issues. Twenty years ago, Singer and Grunberger[2] estimated that 65,000 chemicals were commonly used in modern society, and over 200,000 new chemicals were discovered each year. Since that time, advances in automated methods of chemical synthesis including combinatorial techniques[3] have led to an explosion into the millions of new chemicals produced each year for end-use applications including drugs, agriculture, personal care, and nutrition. While only a fraction of these new compounds will turn out to be acutely toxic, those that do may have long-term possibilities of causing cancer and reproductive damage. A major mechanism of chemical toxicity involves activation of chemicals by catalytic oxidation mediated by cytochrome P450 (cyt P450) enzymes in the liver.[4–6] Lipophilic molecules bioactivated in this way often damage genetic material (i.e., DNA). They include styrene, benzo[a]pyrene, napthylamines, and many others.[7–11] Covalent DNA adducts of these activated molecules with DNA bases are important biomarkers of cancer risk in humans exposed to toxic molecules.[12–14] Conventional toxicity evaluation of new chemicals proceeds from microbiological testing to animal testing and is expensive and time consuming. While advances in speed and automation of microbiological test are on the horizon, simple, inexpensive chemical screening protocols that could be used at early stages of commercial consideration would be very useful. One such scheme could be based on enzyme bioactivation of the chemicals with detection of DNA damage from the resulting metabolites. Sensors built on this principle could be used to screen chemicals and metabolites that clearly damage DNA. These compounds could then be eliminated from further commercial consideration and testing. Such a screening process could decrease the cost of bringing new drugs and agricultural chemicals to the marketplace by eliminating toxic candidates early in their commercial development Dekkker Encyclopedia of Nanoscience and Nanotechnology DOI: 10.1081/E-ENN 120013449 Copyright D 2004 by Marcel Dekker, Inc. All rights reserved.

and lightening the burden on toxicity bioassays. Of course, new commercially viable chemicals would still need to be subjected to microbiological and animal testing before final marketing.

OVERVIEW Nanoscience can be harnessed to make sensors featuring active metabolic enzymes and double-stranded DNA in films of nanometer-scale thickness[15] for toxicity screening. Early in the quest for such toxicity sensors, we evaluated a number of film formation strategies. (For an overview, see Ref. [16]). We found that the excellent films for these applications could be made by a layer-by-layer construction method developed over the past decade.[17–20] The films are assembled by electrostatic adsorption of alternately charged layers of macromolecular ions and provide films with good stability, excellent enzyme activity, and control of thickness and architecture on the nanometer scale. Film construction relies on a series of steps in which the oppositely charged macromolecules (e.g., DNA, enzymes, and polyions) are adsorbed alternately from solutions onto an electrode. Iron heme enzymes in these films, such as cyt P450s, are easily activated for catalysis by addition of hydrogen peroxide along with substrate to a reaction solution.[15] The resulting DNA damage can be monitored by a variety of electroanalytical methods and by electrochemiluminescence, as will be seen below.

CONSTRUCTING ENZYME–DNA FILMS Preparation of enzyme–DNA films one layer at a time provides excellent control over the thickness of films designed to the specifications of the builder. Films containing two layers each of enzyme and DNA that are 20– 40 nm thick are easily made. Alternate adsorption of layers of biomolecules and polyions is a general method that has been developed over the past decade by Lvov et al.[17–20] The technique has been used to make ultrathin films of a wide variety of proteins and oppositely charged 1063

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polyions including DNA. Some polyions that have been used to make such films are shown below.

A DNA/enzyme film grown on a rough pyrolytic graphite electrode is illustrated in Fig. 1. The pyrolytic graphite has a negative charge by virtue of oxygenic functional groups on its surface, and oxidizing the surface can enhance this negative charge. The films are made as follows. First, the negatively charged electrode is immersed into a 2 mg mL
1 solution of positively charged PDDA. The polycations adsorb at roughly monolayer coverage in about 15–20 min,[18–20] effectively reversing the charge on the solid surface. The electrode is rinsed with water, then immersed in a 2 mg mL
1 solution of double-stranded (ds) DNA, and now the surface develops a negative charge. This surface is rinsed again, then immersed into a solution of enzyme (1–3 mg mL
1) in a

Fig. 1 Conceptual representation of DNA/enzyme films used for toxicity sensing. (View this art in color at www. dekker.com.)

Fig. 2 Quartz crystal microbalance frequency shifts for cycles of alternate myoglobin/ds-DNA and cytochrome P450cam /dsDNA adsorption on gold resonators coated with mixed monolayers of mercaptoproionic acid/mercaptopropanol as first layer and PDDA as second layer. DNA was from salmon testes (ST) and calf thymus (CT). Average values are shown for five replicates of [Mb/ST ds-DNA] () and four replicates of [cyt P450cam /ST ds-DNA] (.) films. (From Ref. [15] with permission. Copyright American Chemical Society.)

buffer of pH lower than the enzyme’s isoelectric point to assure a positive charge. An enzyme layer adsorbs, and the surface becomes positive. The latter two adsorption steps can be repeated as many times as desired to obtain a film of reproducible alternating layers of enzyme and DNA in a multilayer assembly. In practice, protein does not even have to be positively charged because charge patches on protein surfaces often enable good binding to oppositely charged polyions.[21] While there is extensive intermixing of neighboring layers,[17–20] this does not adversely affect the performance of most devices. In the initial stages of developing new films, it is important to monitor layer growth during or after each adsorption step with quartz crystal microbalance (QCM) weighing, surface plasmon resonance, spectroscopy, or voltammetry. Fig. 2 illustrates QCM monitoring during the construction of DNA/enzyme films on gold-quartz resonators. The frequency of the QCM resonator decreases in direct proportion to the mass on its gold coating, provided the viscoelasticity of the interface does not change.[22] Estimates of the weight of each layer and of the repeatability of the multiple adsorption steps can be obtained from QCM of dry films. Drying minimizes bias from interfacial viscoelasticity changes and absorbed

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sweep voltammetry[29] and alkylation was monitored by a.c. voltammetry.[30] In these methods, the DNA in solution was damaged and then accumulated on the electrodes by adsorption prior to analysis. Guanine and adenine bases in ss-DNA can also be oxidized on solid electrodes,[31] such as glassy carbon and pyrolytic graphite. Damage from ionizing radiation to DNA adsorbed onto carbon electrodes was detected by chronopotentiometry.[32] Guanine, with a formal potential at pH 7 of 1.3 V vs. normal hydrogen electrode (NHE) (1.06 V vs. saturated calomel electrode (SCE)), is the most easily oxidized of the four DNA bases.[33] The other bases have formal potentials up to 0.5 V more positive. Detecting nucleic acids by electrochemical oxidation depends on structure, with double-stranded (ds) DNA giving only trace oxidation peaks at best and single-stranded (ss) DNA giving irreversible oxidation peaks at about 1.0–1.2 V vs. SCE on carbon electrodes. Enhanced electrochemical signals for DNA can be obtained by catalytic electrochemical oxidation using transition metal complexes.[34] Studies by Thorp et al.[35–40] showed that Ru(bpy)23 + (bpy =2,2’-bipyridine) is an efficient electrochemical catalyst that oxidizes only guanine bases in DNA and oligonucleotides. The reaction follows the catalytic pathway below:

water. For 9-MHz quartz resonators, mass per unit area M/A (g cm
2) of the film is related to the QCM frequency shift DF (Hz) by: M=A ¼
DF=ð1:83  108 Þ

ð1Þ

where the area is that of the metal disk on the resonator in cm2. Direct scaling between DF and nominal film thickness (d) is given by:[18–20] d ðnmÞ 
ð0:016  0:002ÞDF ðHzÞ

ð2Þ

To mimic the PG electrode surface for QCM measurements of layers adsorbed on the gold-quartz resonators, we first chemisorb a mixed monolayer of mercaptopropionic acid/mercaptopropanol. This layer is represented by the first point in Fig. 2, labeled MPA. The second layer is PDDA. Quartz crystal microbalance frequency decreasing in a roughly linear fashion and at regular intervals for the multiple adsorption steps demonstrates repeatable adsorption for the two DNA/enzyme films. Relative precision of layer formation on multiple resonators within ± 15% can be achieved. Film thicknesses and component weights in Table 1 were obtained by analyzing the QCM data with Eqs. 1 and 2.

ELECTROANALYTICAL METHODS FOR DETECTING DNA DAMAGE

RuðbpyÞ2þ ¼ RuðbpyÞ3þ þ e
3 3

ð3Þ

RuðbpyÞ3þ þ DNAðguanineÞ 3

Electrochemistry of DNA

! RuðbpyÞ2þ þ DNAðguanineox Þ þ Hþ 3

Fundamental studies of the electrochemistry of DNA have shown that single-stranded (ss) DNA is more easily oxidized at guanine and adenine base sites than doublestranded (ds) DNA.[23–25] The double helical ds-DNA structure does not allow ready access of the bases to the electrode or to catalytic oxidizing agents. However, unwinding of the double helix frees the bases for closer access by oxidants, leading to faster reaction rates. Voltammetry has been applied to detecting DNA damage from strong acids,[26] strand cleavage agents,[27] and hydroxyl radicals[28] after adsorbing the DNA onto mercury electrodes. In a similar experimental approach, damage to DNA from ionizing radiation and detected by linear

DNA(guanineox) represents a guanine radical site[41,42] on DNA. Ru(bpy)33 + formed by oxidation at the electrode is cycled back to Ru(bpy)23 + by the fast chemical step in Eq. 4, which provides greatly enhanced voltammetric current over that of Ru(bpy)23 + or DNA alone. The peak current depends on the rate of the chemical step in Eq. 4. In oversimplified terms, the faster the rate of the chemical step, the larger the peak. Guanine accessibility is important in these catalytic oxidations as in direct electrochemical oxidation; that is, guanine in ssDNA reacts much more rapidly than in ds-DNA. In addition, the reaction rate at a given guanine site is

Table 1 Average characteristics of protein/DNA films from QCM results Film DNA(Mb/CT-ds-DNA)2 DNA(Mb/ST-ds-DNA)2 DNA(Cyt P450cam / ST-ds-DNA)2

ð4Þ

Total thickness (nm)

wt. DNA (Mg cm
2)

wt. Protein (Mg cm
2)

20 30 40

3.1 5.8 15

3.6 4.9 2.9

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dependent on the sequence of neighboring bases in the DNA. Electrochemical Methods For Toxicity Sensing Against the backdrop of the electrochemical research on DNA summarized above, we set out to develop methods to detect chemical DNA damage as the basis for toxicity screening. Our first attempts involved derivative square wave voltammetric (SWV) analysis of films of doublestranded (ds) DNA and ionomers Eastman AQ38S or Nafion.[43,44] Square wave voltammetric analysis was used because of its inherent high sensitivity and resolution. DNA–ionomer films were incubated with styrene oxide, which reacts with guanine bases and disrupts the DNA double helix.[7,8] Damage to DNA in these films resulted in multiple oxidation peaks that developed during the reaction. This damage was confirmed by capillary electrophoresis detection of DNA base adducts in hydrolyzed DNA reacted with styrene oxide under these same conditions. Total integrals of the derivative SWV oxidation peaks increased with time of incubation with styrene oxide. Unfortunately, direct electrochemical detection of DNA damage in films suffered from poor signal to noise ratios and data analysis that required derivative or other background corrections. Thus we explored catalytic methods of DNA oxidation (cf. Eqs. 3 and 4) to improve signal to noise in SWV detection.[45] At the same time, we began to realize that layer-by-layer growth of films had

Fig. 3 Square wave voltammetric of (PDDA/ds-DNA)2 films on oxidized PG electrodes in pH 5.5 acetate buffer containing 50 mM NaCl, with 50 mM Ru(bpy)23 + in buffer and after incubations at 37°C with saturated styrene oxide (SO) or unreactive toluene. (From Ref. [45] with permission. Copyright American Chemical Society.)

Fig. 4 The influence of incubation time with saturated styrene oxide and toluene in pH 5.5 buffer on the average catalytic peak current (less current for controls) for 5 to 15 trials per data point for (PDDA/ds-DNA)2 films. Error bars represent standard deviations. (From Ref. [45] with permission. Copyright American Chemical Society.) (View this art in color at www.dekker.com.)

clear advantages in versatility, stability, and thickness control and so was used to make films of DNA and the polycation PDDA. The best sensitivity was found with ds-DNA as the outer layer of the film and at low salt concentration. Quartz crystal microbalance of (PDDA/dsDNA)2 on gold resonators showed that the average thickness was 6 nm and that each film contained 0.23 mg of ds-DNA. [The (PDDA/ds-DNA)2 terminology represents two bilayers of PDDA/DNA.] (PDDA/ds-DNA)2 films on oxidized PG electrodes were reacted with styrene oxide, then the electrodes were washed and transferred into pH 5.5 buffer containing 50 mM Ru(bpy)23 + as the catalyst. The SWV oxidation peaks increased with incubation time with saturated styrene oxide (Fig. 3). Control (PDDA/ds-DNA)2 electrodes incubated in buffer or saturated toluene showed much smaller peaks characteristic of intact ds-DNA layers in the films. Average peak current for catalytic SWV oxidation of the ds-DNA films increased with incubation time for 30 min, then decreased slightly (Fig. 4). Each point represents a single electrode, and error bars reflect electrode-to-electrode variability. Incubation of films with toluene, for which no chemical reactions with DNA have been reported, gave catalytic oxidation peaks that showed no trends with reaction time. Catalytic SWV oxidation with Ru(bpy)23 + provided more sensitive detection of DNA damage than direct SWV oxidation. Studies of DNA and polynucleotides in solutions and films suggested that oxidation of guanine and chemically damaged adenine in partly unraveled damaged DNA were the most likely contributors to the catalytic peaks.[45]

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Fig. 5 Influence of incubation time on average SWV peak current ratios for (PDDA/ds-DNA)2 films in 20 mM Co(bpy)33 + at pH 5.5 after incubations with styrene oxide, toluene, or pure buffer. Ip,final corresponds to the peak current after each incubation; Ip.initial corresponds to peak current before incubation. (From Ref. [46] with permission. Copyright 2002 Wiley-VCH.) (View this art in color at www.dekker.com.)

Another method to monitor DNA damage in films employed a cationic electroactive probe that binds better to ds-DNA than to damaged DNA. Co(bpy)33 + was used to probe films of (PDDA/ds-DNA)2 grown layer-by-layer on PG electrodes first coated with a layer of PSS.[46] After incubation of (PDDA/ds-DNA)2 films with styrene oxide, electrodes were rinsed, placed into 20 mM Co(bpy)33 +, and the CoIII/CoII reduction peak at 0.04 V vs. SCE from the DNA-bound complex was monitored by SWV. Peak current decreased with increasing time of reaction with

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styrene oxide because of the decreasing ability of the damaged outer layer of DNA to bind Co(bpy)33 +. A single electrode can be used to measure the incubation time course. Denoting the initial SWV peak current as Ip,initial and the peak current after incubation as Ip,final, plots of the peak current ratio Ip,initial /Ip,final vs. incubation time (Fig. 5) were used to monitor DNA damage rates. This ratio helps correct for electrode-to-electrode variations. Ip,initial /Ip,final increased for ds-DNA electrodes with increasing incubation time with styrene oxide solutions. No significant variations were found for controls with or without toluene. Building films a layer at a time allowed us to incorporate catalytic metallopolyion catalysts for DNA oxidation into ‘‘reagentless’’ toxicity biosensors. Two polyions containing Ru(bpy)22 + were used that are capable of catalytically oxidizing guanines in DNA. The one shown below, denoted Ru-PVP, has 6 Ru-N bonds and reversible oxidation at 1.15 V vs. SCE.

A related polymer denoted RuCl-PVP has 5 Ru-N bonds with chloride as the sixth Ru ligand. It is reversibly oxidized at 0.75 V vs. SCE[47–49] and provided clearer

Fig. 6 Examples of data obtained with catalytic RuCl-PVP layer in films: (a) SWV of PSS/RuCl-PVP/DNA/PDDA/DNA films after incubations at 37°C and pH 5.5 with saturated styrene oxide (SO) for 5, 10, 20, and 30 min, respectively. Incubations in toluene gave no changes in peak current. (b) Influence of reaction time (37°C, pH 6.5) of PSS/RuCl-PVP/DNA/PDDA/DNA films incubated in 2 mM dimethyl sulfate (.), 2 mM methyl methanesulfonate (6), and buffer control (4) on ratio of final SWV peak current to initial peak current of PSS/RuCl-PVP. (From Ref. [49] with permission. Copyright American Chemical Society.) (View this art in color at www.dekker.com.)

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peaks on a flatter background for the catalytic reaction at this lower voltage without sacrificing very much sensitivity. Use of this catalytic polymer is illustrated in Fig. 6a, showing an increase in catalytic SWV peaks of RuCl-PVP in PSS/RuCl-PVP/DNA/PDDA/DNA films after incubation with styrene oxide. PSS/RuCl-PVP/DNA/PDDA/DNA films were also incubated with methylating agents dimethyl sulfate (DMS), a confirmed human carcinogen, and suspected carcinogen methyl methanesulfonate (MMS).[50] Films incubated with DMS or MMS gave increases in the peak current that suggested DNA damage. Fig. 6b shows changes with reaction time in the ratio of final SWV peak current of the films to the initial peak current for the PSS/RuCl-PVP layers alone. The increase in the ratio is a result of DNA unwinding caused by nucleobases that have been methylated by MMS and DMS.[51,52] Damage to DNA under these conditions was confirmed by capillary electrophoresis detection of methylated guanines and adenines in hydrolyzed DNA that had been reacted with MMS or DMS. Ru-PVP, the more powerful oxidizing agent, was used to develop a new method of DNA detection, direct electrochemiluminescence (ECL). A SWV waveform oxidized the RuII sites in the metallopolymer to RuIII. Electrochemiluminescence was measured simultaneously with catalytic SWV peaks in a simple apparatus employing an optical fiber to conduct light from the electrode to a photomultiplier detector. The principle is illustrated in Fig. 7, showing the alternating layers of ds-DNA and the ruthenium metallopolymer [Ru(bpy)2(PVP)10]2 + used in the film. Direct electrochemiluminescence (ECL) involving DNA oxidation was observed in 10-nm films of Ru-PVP assembled layer-by-layer with DNA.[53] The electrode oxidized the RuII sites in the metallopolymer to

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Fig. 8 Simultaneous SWV (downward curves) and ECL (upward curves) responses for (Ru-PVP/ds-CT DNA)2 films on PG electrodes in pH 5.5 buffer after incubations at 37°C with saturated styrene oxide. Curve labels are incubation times in min. (From Ref. [53] with permission. Copyright American Chemical Society.)

RuIII sites that reacted with guanines in the DNA. Electrochemiluminescence was measured simultaneously with catalytic voltammetric peaks at about 1.15 V vs. SCE (Fig. 8). Electrochemiluminescence was observed only when guanines were present on oligonucleotides in the films. This result along with previously proposed ECL pathways suggests that guanine radicals initially formed by catalytic oxidation of guanines by RuIII react with the metallopolymer to produce electronically exited RuII* in the film. RuII* spontaneously emits light at 610 nm and decays to ground-state RuII. Simultaneous linear growth of ECL and SWV peaks occurred after incubation with styrene oxide over 20 min. The estimated detection limit was 1 damaged DNA base in 1000, or 0.1% damage. Control incubations of metallopolymer/ds-DNA films in buffer containing unreactive toluene resulted in no significant trends in ECL or SWV peaks with time.

COMBINING BIOACTIVATION WITH DNA DETECTION Fig. 7 Conceptual cartoon of film of ds-DNA and the catalytic metallopolymer [Ru(bpy)2(PVP)10](ClO4)2 designed for simultaneous electrochemiluminescent (610 nm) and voltammetric detection of DNA. (From Ref. [53] with permission. Copyright American Chemical Society.) (View this art in color at www. dekker.com.)

A practical approach for toxicity screening combines bioactivation and DNA damage detection into a single biosensor for toxicity screening. To make these types of sensors, ultrathin films (20–40 nm thick) containing myoglobin or cytochrome P450cam and DNA were

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Fig. 9 Catalytic square wave voltammetry of PDDA/ds-DNA (/Mb/ds-DNA)2 films on rough PG electrodes before and after incubations at 37°C with 2% styrene (no styrene in controls) and 0.2 mM H2O2 in aerobic buffer (SWV amplitude: 25 mV; frequency: 15 Hz; step height: 4 mV; PDDA = polydiallyldimethylammonium ion). After incubation, electrodes were washed and placed into pH 5.5 buffer containing 50 mM Ru(bpy)23 + for the SWV analysis. (From Ref. [15] with permission. Copyright American Chemical Society.)

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grown layer-by-layer on electrodes. [Film composition is denoted by listing the components in their order of assembly, e.g., PDDA/ds-DNA(/enzyme/ds-DNA)2, where PDDA =polydiallyldimethylammonium ion.] Enzymes in the films were activated by hydrogen peroxide, generating test metabolite styrene oxide from styrene.[54] Styrene oxide formed in these nanometer-scale films reacted with double-stranded (ds) DNA in the same film, mimicking metabolism and DNA damage in the human liver. In the first demonstration of this concept, DNA damage was detected by SWV by using catalytic oxidation with dissolved Ru(bpy)23 + or by monitoring the binding of Co(bpy)33 + to the outer DNA layer in the film.[15] Later, we also detected DNA damage after enzyme-catalyzed bioactivation by using an inner RuCl-PVP layer in DNA/ enzyme films.[49] The SWV peak at 1.05 V vs. SCE for Ru(bpy)23 + increased with time of the DNA damage reaction (Fig. 9). As in the case where enzyme-free films are incubated directly with DNA damage agents, ds-DNA in the film is damaged by styrene oxide. As a result of these reactions, guanines are released from the protection of the double helix, as are adducts of adenine.[45] These species are more easily oxidized than the original ds-DNA, and the catalytic oxidation current increases. Control electrodes incubated in hydrogen peroxide without styrene showed catalytic oxidation peaks of similar heights to freshly prepared films (Fig. 9). Thus the low concentrations of hydrogen peroxide used had no measurable influence on ds-DNA. Only peaks for films that had been activated by hydrogen peroxide and styrene together increased with reaction time.

Fig. 10 The influence of reaction time with 2% styrene + 0.2 mM H2O2 on catalytic SWV peak in 50 mM Ru(bpy)23 +: (a) for PDDA/dsST-DNA(/cyt P450cam /ds-ST-DNA)2 films (.) in 50 mM Ru(bpy)23 + solution. Control (6) is for incubation with 0.2 mM H2O2. (b) PDDA/ds-DNA(/Mb/ds-DNA)2 films using CT-DNA (.) and ST-DNA () (5–15 trials per data point, CT = calf thymus, ST = salmon testes). Also shown are controls representing incubation of ST-DNA/Mb films with 2% styrene but no H2O2 (5), 2% toluene + 0.2 mM H2O2 (!), and 0.2 mM benzaldehyde + 0.2 mM H2O2 (4) (average peak current for unreacted DNA/enzyme electrodes was subtracted). (From Ref. [15] with permission. Copyright American Chemical Society.)

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Similar results were obtained using cyt P450 as the enzyme. DNA/(Cyt P450cam /ST-ds-DNA)2 films were incubated with styrene and hydrogen peroxide, and a rapid increase in SWV peak current in the first 5 min was followed by a slower increase from 5 to 30 min (Fig. 10a). Capillary electrophoresis and HPLC-MS analyses of DNA in thin films that were reacted with styrene oxide, and then hydrolyzed, confirmed formation of known styrene oxide adducts of DNA bases under similar reaction conditions. Average SWV peak currents for ds-DNA/Mb films also increased with incubation time with hydrogen peroxide and styrene at relatively larger rates for the first 5 min, then at smaller rates at longer times (Fig. 10b). Slightly larger initial rates of peak growth were found with Mb/ST ds-DNA films than with Mb/CT ds-DNA films (CT =calf thymus, ST =salmon testes). Error bars are mainly the result of film-to-film variability. No significant growth in peak current with reaction time was found for films incubated with toluene and hydrogen peroxide, styrene alone, benzaldehyde, or hydrogen peroxide alone (Fig. 10, controls). In alternative assays, Co(bpy)33 + was used as an electroactive probe. Co(bpy)33 + reduction peaks at 0.04 V decreased as DNA was damaged by enzyme-generated styrene oxide. Little change in SWV signals was found for incubations of DNA/enzyme films with unreactive organic controls or hydrogen peroxide. The catalytic SWV method was more sensitive than the Co(bpy)33 + binding assay, providing multiple measurements over a 5-min reaction time. However, the Co(bpy)33 + binding assay may be more useful in the presence of potential interferences because of its low measurement voltage where few electroactive species are reduced or oxidized.

CONCLUSION This article described how adsorption of layers or biomolecules and polyions onto electrodes can provide films of nanometer-scale thickness on electrodes for screening the toxicity of chemicals and their metabolites. Such films containing DNA and enzymes enable detection of structural damage to DNA as a basis for the toxicity screening. The most sophisticated versions of these sensors bioactivate chemicals to their metabolites, which may react with DNA. The approach mimics toxicity pathways in the human liver. DNA damage can presently be detected at levels of about 0.1% to 0.5% by methods including catalytic square wave voltammetry using soluble or polymeric catalysts, electrochemically active DNA binding probes, or electrochemiluminescence using a special metallopolymer. Catalytic polyions can be incorporated into DNA/enzyme films leading to ‘‘reagentless’’ sensors

Electrochemical Toxicity Sensors

for chemical toxicity. These sensors are suitable for detection of relative DNA damage rates in 5–10 min. We expect good future progress in applications of enzyme/DNA films to toxicity biosensors. The analytical approaches discussed above may also be adaptable to monitoring DNA oxidation, which has been suggested as a clinical marker for oxidative stress.[55] Electrode arrays could be developed to provide many tests simultaneously. Future sensor arrays could be configured to detect toxicity of metabolites generated by a range of human cyt P450s.

ACKNOWLEDGMENTS This author’s research described herein was supported by US PHS grant No. ES03154 from the National Institute of Environmental Health Sciences (NIEHS), NIH, U.S.A. The author thanks students and collaborators named in joint publications, without whom the development of the sensors described would not have been possible.

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