Cytochrome P450 biosensors—a review

Biosensors and Bioelectronics 20 (2005) 2408–2423

Review

Cytochrome P450 biosensors—a review Nikitas Bistolasa , Ulla Wollenbergera , Christiane Jungb , Frieder W. Schellera,∗ b

a Department of Analytical Biochemistry, University of Potsdam, Karl-Liebknecht-Street 24-25, 14476 Golm, Germany Max-Delbr¨uck-Center for Molecular Medicine, Protein Dynamics Laboratory, Robert-R¨ossle-Strasse 10, 13125 Berlin, Germany

Received 25 August 2004; received in revised form 10 November 2004; accepted 10 November 2004 Available online 5 January 2005

Abstract Cytochrome P450 (CYP) is a large family of enzymes containing heme as the active site. Since their discovery and the elucidation of their structure, they have attracted the interest of scientist for many years, particularly due to their catalytic abilities. Since the late 1970s attempts have concentrated on the construction and development of electrochemical sensors. Although sensors based on mediated electron transfer have also been constructed, the direct electron transfer approach has attracted most of the interest. This has enabled the investigation of the electrochemical properties of the various isoforms of CYP. Furthermore, CYP utilized to construct biosensors for the determination of substrates important in environmental monitoring, pharmaceutical industry and clinical practice. © 2004 Elsevier B.V. All rights reserved. Keywords: Cytochrome P450; Bioelectrocatalysis; Electrochemistry; Modified electrodes

Contents 1.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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2.

The reaction cycle of CYP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3.

Protein electrochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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4.

Direct electron transfer of CYP in electrochemical sensors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Bare electrodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Clay modified electrodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Phospholipid modified electrodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Electrodes modified with multilayer films . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. CYP bionsensing at high temperatures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6. Spectroelectrochemistry and surface plasmon resonance of CYP. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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5.

Summary and conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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6.

Future perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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∗

Corresponding author. Tel.: +49 331 977 5121; fax: +49 331 977 5150. E-mail address: [email protected] (F.W. Scheller).

0956-5663/$ – see front matter © 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.bios.2004.11.023

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Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. Introduction Cytochromes P450 CYP form a large family of heme enzymes that catalyze a diversity of chemical reactions such as epoxidation, hydroxylation and heteroatom oxidation. The enzymes are involved in the metabolism of many drugs and xenobiotics and are responsible for bioactivation. Many of these compounds are even inducers for CYP expression in different organs. (Poulos, 1995; Ortiz de Montellano, 2004; Lewis, 1996). The catalytic abilities of the CYP family have attracted the interest of enzyme engineers already in the 1970s (Brunner and Loesgen, 1977). Derived from their physiological function, intact hepatocytes or microsomes were selected as biocatalytic components in extracorporal detoxification reactions. Later on, the CYP mediated specific hydroxylation of a broad spectrum of substrates, including highly inert alkanes or steroids in a preparative scale, were the target. However, these studies had to face complications both due to the limited stability of the labile multi-enzyme system and the need of the regeneration of the cofactor NADPH or NADH. Considerable improvement has been achieved by the development of appropriate methods of immobilization of CYP. To simplify the technology, the multi-component system has been restricted to the substrate converting part, i.e., the terminal oxidase. In this way the need of the electron transferring proteins containing the redox active flavine or FeS cluster should be avoided (Table 1). Regeneration of the reduced cofactor can be realized by enzymatic reduction of NADP+ , e.g., by using glucose-6phosphate dehydrogenase. This represents the classical concept of preparative application of dehydrogenases. Alternatively, the electrons can be supplied directly to the terminal oxidase by reduced mediators, e.g., of the viologene type. These substances are well suited to reduce the prosthetic group of CYP due to their negative redox potential. However,

Table 1 The isoforms of CYP enzyme that are used for sensor construction, the type of electrodes together with the different electrode modifications are displayed P450 enzymes studied

Electrodes

Electrode modifications

CYP 1A1 CYP 1A2 CYP 2B4 CYP 2C19 CYP 2D6 CYP 2E1 CYP 3A4

Au Platinum Tin oxide Glassy carbon Pyrolytic graphite Edge-plane graphite Carbon cloth

Bare Clay Phospholipids Multilayers Screen printed Antimony doped Thiol

CYP 17A CYP 102 CYP 101 CYP 176A1 CYP 11A1 CYP 4A1 CYP 4A1

dioxygen – co-substrate for the substrate conversion – reacts with these potent reductants thus consuming in a parasitic reaction the reduced mediator to form hydrogen peroxide. Some progress has been achieved by using Co2+ sepulchrate as mediator, which reacts only slowly with the ambient oxygen (Estabrook et al., 1996). Attempts have also been made to attach CYP-enzymes to electrodes by introducing electroactive bridges covalently coupled to the protein (Lo et al., 1999; Shumyantseva et al., 2000). Such redox relays have been introduced at specifically selected sites generated by protein engineering or randomly, e.g., CYP2B4 with covalently attached riboflavin (Shumyantseva et al., 2000). A list of CYP bionsensors based on mediated electron transfer is presented in Table 2. The ultimate approach is the direct (mediatorless) electron supply from a redox electrode to the redox active group of the CYP (Table 2). This concept was established by us (Scheller et al., 1977) in parallel with the development of the ‘promoted’ direct electron exchange of modified electrode for c-type cytochromes (Eddowes and Hill, 1977; Yeh and Kuwana, 1977). In addition to the potential application of CYP for preparative purposes, its ability to metabolize a broad spectrum of endogenous substances, e.g., fatty acids, steroid hormones, prostaglandins like mediators and foreign compounds, e.g., drugs and environmental toxins has made this enzyme family interesting as recognition element for biosensing. The highly specific conversion of endogenous substances like steroid hormones makes substrate determination in complex media feasible. The CYP catalysis leads usually to detoxification with following excretion of drugs but may also form reactive products and activate procancerogens. Adverse effects in multi-drug treatments have been seen in many patients originating from overlapping substrate specificities or inhitibory effects of the different isozymes or polymorphic enzymes. Therefore, for drug or xenobiotics risk assessment the measurement of the substrates specificity and concentration in conjunction with distribution of CYP isozyme and polymorphic enzymes based on biosensors would be of high clinical relevance. The reaction stoichiometry offers two traditional principles to couple the substrate conversion to a transducer to give a biosensor: (i) measurement of the oxygen consumption by a Clark-type electrode; (ii) consumption of the reduced co-substrate by measuring the optical absorbance of NAD(P)H. However, both reaction partners (O2 and NAD(P)H) are not only consumed in the CYP catalyzed substrate

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Table 2 Survey of CYP biosensors using electrochemical and optical biosensors Technique

Electrode/modification

E◦ (formal potential)

Substrates tested – catalysis

Comments

1. Bare electrodes Kazlauskaite et al. (1996)

Recombinant CYP101

CV

EPG

−526 mV (ls), −390 mV (hs)

Binding of camphor seen. Catalysis not described here

Lo et al. (1999)

CYP101—WT, mutants

CV

EPG, Au

−428 to −449 mV

No catalysis

Fantuzzi et al. (2004)

CYP2E1

CV, chronoamperometry

GC, Au

−334 mV

p-Nitrophenol

Arg 72,112, 364 and Lys344 that interact with Pdx interact also with bare EPG. Surface cys replaced with Ala. Electrochem. due to O2 reduction to a certain extent. 1 electron transfer, catalysis with different modifications (see below)

2. Electrode modifier: phospholipids Iwuoha et al. (1998) CYP101

CV, amperometry

DAB-BSAglutaraldehyde GCE

−260 mV (ls)

Catalysis with camphor, adamantanoneand fenchone (direct ET)

Zhang et al. (1997)

CYP101

CV, SWV

DMPC—PG, DDAB - PG

−250 mV (DDAB), −357 mV (DMPC)

Catalysis seen with O2 and TCA

Fantuzzi et al. (2004)

CYP2E1

CV, chronoamperometry

Bare, thiol, DDAB – GC, Au

−334 mV

Catalysis with p-nitrophenol

Oku et al. (2004)

CYP119

CV

DDAB-plastic formed carbon electrode

−250mV vs. SSE (20 ◦ C), −50 mV vs. SSE (80 ◦ C)

No catalysis

Aguey-Zinsou et al., 2003

CYP176A1

CV, potentiometry

DDAB-EPG electrode

−360 mv

No catalysis

Fleming et al., 2003

CYP102

CV

DDAB-EPG electrode

−250 mV

Catalysis with O2 and H2 O2

CV (direct electrochem.), electrolysis (reductase mediated) CV, QCM, product analysis

Multilayers with PSS Carbon cloth

−310 mV

Catalysis with styrene

Multilayers with PEI, PDDA, PSS—Au

−250 mV

Catalysis with styrene

3. Multilayers on electrodes Estavillo et al. (2003) CYP1A2

Lvov et al. (1998)

CYP101

Calibration curve done with amperometry. Km = 1,41–3,9 mM. Also use of Co(Sep)3+  = 7,2 mol cm−2 (DMPC),  = 4,9 mol cm−2 (DDAB), ks = 25(DMPC) s−1 , ks = 26 (DDAB) s−1 CYP2E1 electrochemistry and catalysis with different electrodes and modifications Electrochemistry also observed at 80 ◦ C., E (at 20 ◦ C) = 90mV, E (at 80 ◦ C) = 30mV Fe2+ /Fe3+ redox potential unaffected by subsrate binding. pH dependence −59 mV/pH unit ks: 221 s−1 no shift of E◦ with substrate. Slope of pH change (3–8): −33 mV/pH unit, (8–10): −126 mv/pH unit Electrolysis done at −600 mV at 4 ◦ C. Catalase inhibited catalysis. Cationic PEI and PDDA and anionic PSS films were used. Au-MPS-(PEI/PSS-CYP101) - CYP101 pH5,2

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Sensors based on direct electron transfer Reference CYP species

CYP101

QCM, CV, electrolysis, product analysis

Multilayers with PEI - PG

−250 mV

Catalysis with styrene

Rusling et al. (2000)

CYP101

CV, SWV

−270 mV

Catalysis with O2 and H2 O2

Lei et al. (2000)

CYP101

CV

PEI multilayers – rough PG electrode Clay—GC

−368 mV

No catalysis

Zu et al. (1999)

CYP101

CV, electrolysis, product analysis

Au-MPS—PDDA, DDAB, multilayers carbon cloth

−250 mV

Catalysis with styrene and cis-β-methylstyrene

Shumyantseva et al. (2004)

CYP2B4

CV, chronoamperometry

Clay/detergent-GC

−292 to −305 mV

Catalysis with aminopyrine, benzphetamine

Joseph et al. (2003)

CYP3A4

QCM, CV, SWV, amperometry electrolysis, product analysis

Au – MPS – PDDA multilayers

342 mV CV), 335 mV (SWV)

Catalysis with verapamil and medazolam

Nicolini et al. (2001)

CYP11A1

CV

Langmuir Blodgett films (mono- and multilayers), ITO glass plate

−295 to −318 mV

No catalysis

4. Various modifiers Iwuoha et al. (2004)

CYP2D6

CV, amperometry

Polyaniline doped GCE

−120 mV

Catalysis with fluoxetine

Bistolas et al. (2004)

CYP101

Spectroelectrochemistry

−380 mV

No catalysis

Davis et al. (2000)

CYP101 (K334C mutant)

CV, SWV

Dithionite and aldrithiol Au Polycrystalline Au electrode

None mentioned

No catalysis

P450-biosensors based on mediated electron transfer 1. Bare electrodes Estabrook et al. (1996) CYP17A, CYP4A1, CYP3A4, CYP1A2, CYP102

Fusion protein—electrolysis (O2 monitorin)

Strip of mesh of Pt gauze attached to Pt wire

None

Catalysis with progesterone and pregnenolone

Optimization of multilayers conditions. Influence of pH studied.  = 0,1 nmol cm−2 , Turnover rate = 6,3 h−1  = 0,15 nmol cm−2  = 3,54 pmol cm−2 , ks = 5–152 s−1 . CYP101 immobilised and in solution, Product turnover rates higher in multilayers that in solution Product analysis,  = 40.5 pmol cm−2 , kcat = 1.54 min−1 Amperometry done at −500 mV vs. Ag/AgCl. Response time = 15–25 s, Km = 271–1082 ␮M. ks = 0,45 s−1 , E0  = −470 mV vs. Ag/AgCl, Binding of cholesterol seen. Binding also characterized with X-Ray, QCM,CD, Ellipsometry, Brewster angle microscopy Km = 3,7 ␮mol/L., E◦ shifted anodically in the presence of substrate. Native state of enzyme during electrolysis. STM, QCM, K334C mutant greater affinity for Au, and enhanced electrochemistry compared to the wild type. Wild type more mobile than mutant (from QCM study).

N. Bistolas et al. / Biosensors and Bioelectronics 20 (2005) 2408–2423

Munge et al. (2003)

Set potential −650 mV and the decrease in oxygen content was monitored. Use of Co(Sep)3+ . 2411

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Table 2 (Continued ) Sensors based on direct electron transfer Reference CYP species

Technique

Electrode/modification

E◦ (formal potential)

Gilardi et al. (2002)

CYP102, CYP2E1

Staircase CV

GC disc

−500 mV

PfCYP1A2, RfCYP2B4, RfCYPscc

CV, amperometryspectrophotometry

Screen printed -thick film Rh-graphite electrode

−555 mV (cholesterol free), −357 mV (cholesterol bound)

Aminopyrine, aniline. 7-ethoxyresorufin, 7-pentosyresorufin

Shumyantseva et al. (1999)

CYP2B4

Electrolysis

Screen printed -thick film graphite electrode

Reduction at −507 mV

No catalysis but direct ET was observed

Shumyantseva et al. (2000)

CYP1A2, CYP2B4

CV, amperometry spectrophotometry

Screen printed -thick film Rh-graphite electrode

−547 mV (RfCYP2B4), −557 mV (RfCYP2B4)

Catalysis with aminopyrine, aniline, 7-ethoxyresorufin, 7-pentosyresorufin at −500 mV

3. Various modifiers Iwuoha et al. (1998)

CYP101

CV, amperometry

DAB-BSAglutaraldehyde GCE

−260 mV (ls)

Catalysis with camphor, adamantanoneand fenchone (direct ET)

Reipa et al. (1997)

CYP101

CV, AC voltammetry, spectroele-ctrochemistry

Antimony-doped - Tin oxide

−437 mV (Pdx-normal −427 mV)

Catalysis seen with camphor.

Reipa et al. (2002)

CYP101 (Y96F mutant)

Spectroelectrochemistry

Nano-crystalline Sb-dobed tin oxide electrode

WT: −414 mV to −550 mV, Y96F: −448 mV to −492 mV

Catalysis with camphor and styrene

Mayhew et al. (2000)

CYP101 (Y96F mutant)

Spectroelectrochemistry

Antimony-doped tin oxide electrode

None

Catalysis with styrene.

2. Screen printed electrodes Shumyantseva et al. (2001)

Substrates tested – catalysis

Comments Neomycin promoter, Fusion protein: Use of heme (from CYP102 and CYP2E1)-flavodoxin (from Desulfovibrio vulgaris)

Calibration curve done with amperometry. Km = 1,41–3,9 mM. Use of Co(Sep)3+ Use Pdx as mediator. No product in absence of Pdx. Turnove rate 0,5 s−1 NADH turnover rates: WT: 852 nmol−1 s−1 (cam bound), 56 nmol−1 s−1 (styrene bound), Y96F: 29 nmol−1 s−1 (cam bound), 130 nmol−1 s−1 (styrene bound), Phenosafranine used as mediator Use of Pdx as mediator in electrolysis. Product quantification using spectrophotometry: Turnover of product 8 min−1 (WT), 70 min−1 (Y96GF) using NADH, PdR and Pdx

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Use of 11A1 P452B4, CYP1A2 and CYPscc with covalently attached riboflavin (Rf) to improve catalysis, Catalytic substrate reduction measured at −500 mV Truncated flavo-CYP2B4, complex with CO formed under reduction conditions. Use of riboflavin. P452B4 with covalently attached riboflavin for catalysis, increased catalytic rates of substrates.

Faulkner et al. (1995)

CYP4A1

Thiol—Au

None

Optical biosensors based on P450 and ISFET-biosensor Ivanov et al. (1999) CYP2B4

SPR, Spectrophotometry

EDC/NHS Dextran coated cuvette

kon = 0.5–4 × 106 M−1 s−1 , No catalyis koff = 0.5 s−1

Makings and Zlokarnik (2000)

CYP3A4, CYP2C19

Fluorescence

Optical molecular sensor

None

Catalysis with various substrates

Ivanov et al. (2000)

CYP2B4

Optical

EDC/NHS Dextran coated cuvette

None

No catalysis

Ivanov et al. (2001)

CYP2B4, CYP11A1, CYP101

Optical

EDC/NHS Dextran coated cuvette

None

No catalysis

Ivanov et al. (2001)

CYP2B4

Opical biosensor spectropho-tometry

DLPE, DSPE phospholipids aminosilane cuvette

Hara et al. (2002)

CYP1A1

Voltage output, fluorescence

IFSET

Potentials are referred to SCE if not otherwise stated.

Catalysis with lauric acid

No catalysis

None

Catalysis with dichlorophenols

Fusion protein, No direct ET between CYP and Au electrode, Co(Sep)3+ , Electrolysis at −450 mV vs. NHE Interprotein ET occurs between CYP2B4, cytb5 and NADH-CYP reductase through complex formation and random collision. Use of an optical probe which attaches to the substrate and give rise to fluorescence Formation of ternary complex between CYP2B4, NADPH-cyt CYP reductase and cytb5 Formation of binary-ternary complex between CYP2B4, CYP11A, CYP101 and their respective redox partners. kon and koff CYP2B4, cytb5 and NADH-CYP reductase incorporated readily into the phospholipid layer. Fusion enzyme

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Mediated electrochemistry, Electrolysis

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Fig. 1. Catalytic cycle of CYP. The first step is the binding of RH substrate followed by one electron reduction. The binding of substrate involves a shift of the spin state of heme from low to high spin. After activation of the heme with O2 and a second one-electron reduction, an oxygen atom is transferred to the substrate leading to the formation of ROH product.

conversion but also in a parasitic “uncoupling” hydrogen peroxide release via NAD(P)H without product formation. Therefore, the indication of the hydroxylated product gives the only real measure of the enzyme activity. Only for a few substances, e.g., aniline (Renneberg et al., 1978) this can be performed by direct electrochemical quantification. However, coupling of direct cathodic reduction of the terminal oxidase with substrate turnover can overcome the NAD(P)H dependent uncoupling. The generation of “catalytic currents” is therefore the direct indicator of CYP dependent electrocatalysis. The different approaches of this concept will be presented in this review.

2. The reaction cycle of CYP Since the first three-dimensional structure of the bacterial CYP101 was elucidated by Poulos (Poulos et al., 1985) several other structures have been resolved including microsomal ones, CYP2B4 (Scott et al., 2003; Werck-Reichhart and Feyereisen, 2000). The active center is the iron-protoporphyrin IX with an axial thiolate of a cysteine residue as fifth iron ligand. In the absence of a substrate at the beginning of the cycle (Fig. 1) CYP is in the hexa-coordinated low-spin ferric form with water being the sixth ligand. The overall reaction of substrate hydroxylation of the CYP monooxygenase function is the insertion of one atom of the oxygen molecule into an substrate RH, the second atom of oxygen being reduced to water while consuming two reducing equivalents under formation of ROH (Eq. (I)). RH + O2 + 2e− + 2H+ → ROH + H2 O

(I)

The electrons are delivered by flavoproteins or ferredoxinlike proteins and NAD(P)H in a complex electron transfer chain. The most generally accepted mechanism for substrate hydoxylation by CYP includes the following steps although

several details remain still unsolved (Auclair et al., 2002). Substrate binding to the hexa-coordinated low-spin ferric enzyme excludes water from the active site, which is causing a change to the 5-coordinate high-spin state. The decrease of polarity is accompanied with a positive shift of the redox potential by about 130 mV that makes the first electron transfer step thermodynamically favourable. The transfer of one electron from a redox partner reduces the ferric iron to the ferrous enzyme. This can now bind molecular oxygen forming a ferrous-dioxygen (FeII -O2 ) complex. The second electron is transferred along with a proton gaining an ironhydroperoxo (FeIII –OOH) intermediate. The O O bound is cleaved to release a water molecule and a highly active ironoxo ferryl intermediate. This intermediate abstracts one hydrogen atom from the substrate to yield a one-electron reduced ferryl species (FeIV OH) and a substrate radical or reacts in a concerted reaction with the substrate C H bond without intermediate radical formation. Then, it follows immediately the enzyme-product complex formation and release of the product ROH to regenerate the initial low-spin state. The iron-oxo intermediate may however also induce the formation of protein radicals (Sch¨unemann et al., 2002).

3. Protein electrochemistry The transfer of electrons between and within proteins is an essential feature for many physiological processes, including biological energy transfer, metabolism and enzymatic catalysis. The mechanism of electron transfer usually involves protein–protein interactions as it is the case of CYP, where electrons are transferred, i.e., from putidaredoxin reductase to putidaredoxin and then to CYP101. In enzymes an appropriate conformational arrangement is important for the binding of the substrate to the active site and the transfer of charge to the enzyme. The use of electrochemistry allows investigating the electrochemical properties of redox enzymes and their mechanism by observing the direct electron transfer in real time. Much of the knowledge of the mechanism of electron transfer in proteins is based on the Marcus theory (Marcus and Sutin, 1985). The factors that govern a highly specific and directional protein-mediated electron transfer are the magnitude of the electronic interaction between the donor and the acceptor centres, and the contribution of the Franck–Condon (FC) factor involving the differences between the redox potentials of electron donator and acceptor. The rate constant of electron transfer (kET ) can therefore be expressed using the following equation: kET =

2π |HDA |2 FC h ¯

(1)

The magnitude of the electronic interaction as described by the electronic coupling matrix element (HDA ) depends on the distance (r) and the nature of the donor and acceptor redox

N. Bistolas et al. / Biosensors and Bioelectronics 20 (2005) 2408–2423

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centres and can be described by

2

2  



|HDA |2 =
H0DA
exp [−2β(d − 3)] =
H0DA
exp −β d (2) ◦

where HDA is the value of the matrix element at a predefined ˚ and β is the coefficient that describes the dedistance (3 A) cay of electronic coupling with d. The Franck–Condon (FC) factor depends on the thermodynamic driving force for the reaction (G◦ ) and the reorganisation energy (λ):   2 (G0 + λ) 1 exp − (3) FC = √ 4λkB T 4πλkB T where kB is the Boltzmann’s constant. From Eq. (3) can be seen that ln KET increases as the driving force G◦ increases until a maximum is reached at −G◦ = λ. Beyond this point by increasing the G◦ further causes a decrease in the ln KET providing λ remains constant. The driving force is derived from the difference in midpoint reduction potentials of the electron donor and acceptor centres, which is modulated by the protein environment surrounding the redox active sites. The reorganization energy is associated with the rearrangement of the atomic nuclei of the reactants into the configuration that they occupy in the products. Although the above description of the Marcus theory refers for the homogenous electron transfer, it can be also applied to the heterogeneous electron transfer between the protein and the electrode. In this case a redox enzyme is adsorbed on the surface of an electrode. The electronic interaction (coupling) can be varied by using different selfassembled monolayers thus changing the distance and the orientation of the protein from the electrode. In addition, increasing the applied potential increases the driving force and therefore the number of protein molecules that allows electron exchange. Several ways have been used to optimize electron transfer between the redox protein and the electrode particularly using chemically modified electrodes in combination with electrochemical techniques like cyclic voltammetry (CV) and square wave voltammetry (SWV) in addition to amperometry. This is particularly important in the case of heme proteins like CYP in which the electrochemically active heme centre is buried in the protein structure and it is surrounded by an amino acid chain in order to gain a hydrophobic environment for catalysis.

4. Direct electron transfer of CYP in electrochemical sensors On unmodified electrodes enzymes tend to denature and to passivate the electrode. However, CYPs naturally are involved in electron transport pathways of protein redox partners, which require specific docking sites. Therefore, electri-

Fig. 2. Schematic representation of CYP sensor. The enzyme could either be adsorbed or immobilized on a variety of metal and non-metal electrodes (see text). Addition of substrate leads to the formation of product, in this case hydroxylation, in the presence of O2 .

cal contact to CYP-enzymes should be possible at suitable surface modifications of electrodes. The electrochemistry of CYP has been investigated using a variety of metal electrodes (Table 1) such as Au, Pt and Tin oxide, as well as non-metal electrodes such as glassy carbon (GC), pyrolytic graphite (PG), edge-plane graphite (EPG), and carbon cloth (CC). Although direct electron transfer has been observed on bare electrodes, modifying the electrode with an appropriate medium like a polymer or a polyelectrolyte, in order to attain native structure and appropriate orientation so increasing electron transfer between the enzyme and the electrode has been very popular in recent years (Fig. 2). The bioelectrocatalysis by proteins and enzymes such as cytochrome c, CYP, glutathione peroxidase and cellobiose dehydrogenase (a heme-flavo enzyme) at modified electrodes has been recently reviewed (Scheller et al., 2002). In the first biosensor based on the direct electron transfer between the electrode and CYP (Scheller et al., 1977), solubilized CYP from rabbit liver showed a polarographic reduction step at a mercury electrode of −580 mV versus SCE and was partially reduced in constrast to the microsomal CYP which was not detectable. Catalytic currents were obtained following demethylation and hydroxylation of 1.8 mM benzphetamine, 2 mM p-nitroanisole and 2 mM aminopyrine. Furthermore, aniline (Renneberg et al., 1978) and steroids such as deoxycorticosterone (Scheller et al., 1979) could also be electrocatalytically determined using CYP from liver microsomes at a mercury electrode with an apparent Km (K m ) for aniline of 18 mM. In the last decade attempts have also been made to construct CYP biosensors based on the direct electrode transfer using bare electrodes (Kazlauskaite et al., 1996; Lo et al., 1999). 4.1. Bare electrodes Hill and co-workers have used bare EPG to characterize the unpromoted electrochemistry of recombinant CYP101 (Kazlauskaite et al., 1996). In this work, cyclic voltammetric (CV) measurements were carried out in the presence and in the absence of the substrate camphor at 6 ◦ C. Strictly anaerobic conditions were used to prevent formation of the binding of oxygen to Fe2+ and the possibility of second electron transfer. The results indicated reversible oxidation and reduction.

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The interaction of the CYP101 with the bare EPG has been proposed by the authors to be possible via the positively charged Arg-72, Arg-112, Arg-364 and Lys-344 residues on the surface of CYP101. The formal potential (E◦ ) was −526 mV versus SCE with the camphor-free CYP101 and an E◦ of −390 mV versus SCE for the camphor-bound form. These values are in agreement with the redox potential of CYP101 in solution for the substrate-free form with an E◦ of −547 versus SCE and substrate-bound form, with an E◦ of −394 versus SCE (Sligar and Gunsalus, 1976). The binding of camphor to the active site of the enzyme shifts the spin state of the heme prosthetic group from low to high. The authors also claim that a catalytic response upon camphor addition was observed, although details are not given. In order to investigate the role played by amino acids on the immobilisation of CYP101 on the electrode surface, Lo and co-workers have carried out site-directed mutagenesis to modify the surface of the enzyme (Lo et al., 1999). In particular, the wild type (WT) CYP101 contains eight cysteine residues and cysteines 58, 85, 136, 148, and 334 are at or near the protein surface, with Cys-334 being the most exposed one (Poulos et al., 1985). In this study these cysteine residues were replaced by chemically inert alanines. Electrochemistry of WT CYP101 and the mutants could be observed with bare EPG electrodes with formal potentials ranging from −428 to −449 mV versus SCE at high scan rate (10 V/s). However, a quasi-reversible or irreversible electrochemistry was obtained with bare Au electrodes. The electrochemistry of cysteine free CYP101 was in both cases indistinguishable with that of the WT and the single cysteine mutant enzyme, which indicate that electron transfer, is not affected by these residues. Furthermore, electrochemistry of CYP2E1 could be seen with bare GC electrodes with a midpoint potential of −334 mV versus SCE, indicating that CYP2E1 was adsorbed on the electrode surface (Fantuzzi et al., 2004). The heterogeneous electron transfer rate (ks ) was found to be 5 s−1 , which is rather low. 4.2. Clay modified electrodes At bare electrodes the rather low electron transfer achieved between the protein and the electrode limits their use for the construction of efficient CYP biosensors. Modification of electrodes with compounds that facilitate electron transfer, prevent denaturation of protein and cause appropriate orientation of the protein have thus been widely used. Fast heterogeneous electron transfer has been observed when GC electrodes are modified with sodium montmorillonite (SMC) (Lei et al., 2000). Sodium montmorillonite is a member of the general mineral group of clays, which among others also includes smectite, laponite, kaolinite, talc, goethite and orche. Clay minerals are layer type aluminosilicates, ubiquitous in geologic deposits, terrestrial weathering environments, and marine sediments. Clay particles are also very small, colloidal in size, so their behaviour is controlled by surface

forces. Their colloidal and rheological properties have reviewed previously by Luckham (Luckham and Rossi, 1999). Direct evidence of clay-mediated charge transfer has previously been shown by Teng (Teng et al., 1997) for montmorillonite K10 (iron-containing clay). Although SMC contains no iron atoms in its structure (the general formula is Na0.67 (Si)8 (Al3.3 Mg0.67 )-O20 (OH)4 ), charge transfer may be mediated by either Si or Mg. In addition to SMC, other clays such as kaolinite, talc, goethite and orche have also been used to modify electrodes and investigate the electrochemistry of heme proteins. In particular voltammetric studies were carried out for c-type cytochromes, such as cytochrome c, c3 and c553 , where their electrochemical response were investigated using kaolinite, talc, goethite and orche modified PG electrodes (Sallez et al., 2000). Investigation of the electrochemistry of CYP101 (Lei et al., 2000) was carried out in our group using SMC-Pt modified electrodes. The approach was also used for CYP106A2 (unpublished). In particular GC electrodes were modified with SMC that was mixed with colloidal Pt nanoparticles. CYP101 was then adsorbed on the electrode and its redox activity was examined. Direct electron transfer between the electrode and the heme group of the substrate-free enzyme was observed, with an E◦ of −383 mV versus SCE. Similar results were also obtained when the GC electrode was modified with SMC and Pt colloid mixed with CYP101. The E◦ lies in this case at −399 mV SCE. These values deviate strongly from the redox potential of the substrate-free enzyme in solution, which lies at −547 mV versus SCE. Other groups have also obtained large deviations on the formal potential of the substrate free enzyme by using polyion-protein multilayers ranging from −238 to −357 mV versus SCE (Zhang et al., 1997; Lvov et al., 1998; Munge et al., 2003). The reason for that may be that the interaction of CYP either with the clay or with surface of the electrode cause slight alteration to heme environment as to make it less stable, thus shifting the E◦ more positive. A positive shift of the redox potential may be indicative for low to high spin-state conversion that has been ascribed to strong interaction of CYP with surfaces (Niki, 2002). The positive shifts of the redox potential are generally observed when water is excluded from the heme pocket as in the case of camphor binding (Poulos et al., 1986; Jung et al., 2003) and therefore we suggested that the adsorption process leads to a dehydration of the CYP structure. Although the spectrum of CYP101 is not affected by the clay, CO difference spectrum has shown a small increase in P420, the denatured form of CYP, by 4%. The heterogeneous electron transfer rate constants reached values as high as 152 s−1 for CYP101 and 300 s−1 for CYP101 mixed with clay comparing to rates between 27 and 84 s−1 reported for the transfer of the first electron from putidaredoxin to CYP101. This similarity suggests that the negatively charged clay (Ege et al., 1985) obviously mimics the electrostatics of the natural redox partner putidaredoxin and may hold the CYP in a productive orientation. In this orientation the active site of the adsorbed

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CYP101 is still accessible for small iron ligands like CO and dioxygen (Lei et al., 2000). Moreover, liver microsomal phenobarbital induced CYP2B4 has been incorporated in montmorillonite on glassy carbon electrodes (Shumyantseva et al., 2004). In contrast to CYP101, CYP2B4 has a flavoenzyme as redox partner and does not need an iron-sulphur helping protein for delivery of electrons. Using cyclic voltammetry at low scan rates a reduction peak is observed at around −430 mV versus (Ag/AgCl). The electron transfer reaction is obviously very slow. This process is enhanced in the presence of a non-ionic detergent such as Tween 80. CYP2B4 is a membrane bound enzyme and detergent is needed to monomerize CYP2B4 (Kiselyova et al., 1999) as was confirmed also by AFM-studies. From the cyclic voltammograms the amount of electroactive protein of 40.5 pmol cm−2 was calculated. Cyclic voltammetry demonstrates also a reversible one-electron surface redox reaction with a formal potential of about −302 versus SCE and a heterogeneous electron transfer rate constant of 80 s−1 . As in many of the published cases, the formal potential of CYP at the surface determined by the heterogeneous redox reaction is more positive than the redox potential in solution. Therefore, an influence of the detergent on the heme environment as above can also be discussed. The studies of direct heterogeneous electron transfer have been carried out in most cases by cyclic and square wave voltammetry. In these studies the first of the two electrons required for the catalytic reaction has been transferred despite the authors do not see the shift of the reduction potential upon substrate addition as has been reported by (Kazlauskaite et al., 1996) and is known for the reaction in solution. In all cases catalytic oxygen reduction is observed but only rarely catalytic substrate conversion could be achieved. CYP101 predominantly catalyzes the regio- and stereospecific hydroxylation of (1R)-camphor to exclusively 5-exohydroxycamphor. Other compounds than camphor, such as compounds of environmental and industrial interest have also been identified as substrates for CYP101. During catalysis electrons are transferred from NADH to CYP101 through putidaredoxin reductase (PdR) and putidaredoxin (Pdx). The negatively charged group of Asp38 in Pdx forms a salt bridge with Arg112 in the positively charged patch (Arg112 and Arg109, Arg79) in CYP101 to shuttle electrons between PdR and CYP101 (Roitberg et al., 1998). Thus, a ferredoxin in DET contact to an electrode may deliver the reducing equivalents to CYP. On indium–tin oxide electrodes CYP101 conducts camphor hydroxylation mediated by putidaredoxin (Reipa et al., 1997) and dehalogenation of haloalkanes with spinach ferredoxin in the presence of polylysine as promoter proceed (Witz et al., 2000). Acceleration of styrene epoxidation and dehalogenation of hexachloroethane, carbon tetrachloride and other polyhalomethanes was successful with mutated CYP101 (Reipa et al., 2002; Mayhew et al., 2000; Walsh et al., 2000). We succeeded in developing biosensors based on mediator-free CYP2B4 catalysis by immobilizing monomer-

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Fig. 3. Relationship of catalytic current, obtained with CYP2B4 bionsensor after addition of aminopyrine, with increasing concentrations of aminopyrine.

ized CYP2B4 in montmorillonite (Shumyantseva et al., 2004). When substrates were added to air saturated buffer solution, there was an increase of the reduction current. A typical concentration depedence measured in chronamperometry is shown for aminopyrine and benzphetamine (Fig. 3). The reaction was inhibited by metyrapone. This indicates that CYP2B4 possess catalytic activity in the presence of substrate. A further evidence was delivered by product analysis. After 1 h of controlled potential electrolysis at −500 mV versus SCE formaldehyde was measured. The apparent catalytic rate related to the amount of electroactive protein is kcat = 1.54 min−1 which is comparable to the value kcat = 3.5 min−1 of the microsomal system (Shumyantseva et al., 2001). 4.3. Phospholipid modified electrodes The majority of CYP enzymes are located in a hydrophobic environment in the endoplasmic reticulum of cells, although cytosolic enzymes also exist, such as CYP101 (Lewis, 2001). In order to mimic the physiological environment of CYP enzymes, a number of groups have used phospholipids, such as didodecyldimethylammonium bromide (DDAB), dimeristoyl-l-␣-phosphatidylcholine (DMPC), dilauroylphosphatidylethanolamine (DLPE) and distearoylphosphatidylethanolamine (DSPE), for the construction of biosensors. Phospholipid layers form stable vesicular dispersions that bear structural relationship with the phospholipid components of biologically important membranes. By this way a membranous environment is created that facilitates electron transfer between the enzymes redox centre and the electrode. Using this approach a CYP101 biosensor was created for monitoring drug conversion (Iwuoha et al., 1998). The biosensor comprised a GC electrode modified with CYP101 contained in DDAB vesicle dispersion. Glutaraldehyde in the presence of bovine serum albumin was used as a cross-linking agent. CV measurements of the CYP electrode in air-free buffer indicated direct electron exchange between the heme group of CYP101 and the electrode. The E◦ was found to be −260 ± 10 mV versus SCE at a scan rate of 500 mV s−1 and a peak separation of 36 mV. When 3 mM of ethanolic

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solution of camphor was added a catalytic response was observed in an aerobic measuring buffer. The peak cathodic potential and current were −375 mV versus SCE and 44 ␮A in the absence of camphor and −350 mV versus SCE and 50 ␮A in presence of camphor, respectively. Under anaerobic conditions addition of 3 mM ethanolic solution of camphor produced a catalytic response with a peak cathodic potential of −430 mV versus SCE and a peak cathodic current of 14 ␮A. This indicated a typical fast reversible electrochemistry of heme Fe3+/2+ redox species coupled to subsequent process, such as hydroxylation of camphor or H2 O2 production by uncoupled turn-over. The authors therefore claim that even in degassed buffer the oxygen in the aerobic ethanolic camphor solution was sufficient for the reaction to take place. In a similar study Zhang and co-workers in addition to DDAB, used DMPC to incorporate CYP101 and then modify PG disc electrodes (Zhang et al., 1997). CV responses with DMPC showed reversible electrochemistry with an E◦ of −357 ± 4 mV versus SCE, whereas E◦ with DDAB was −238 ± 10 mV versus SCE. These values are more positive than the redox potential of CYP101 in solution, that is −547 mV versus SCE, as a result of protein–lipid interactions and/or possible lipid-dependent electrical double layer effects on electrode potential (Bard and Faulkner, 1980). CV measurements in the presence of CO showed a positive shift of the E◦ from −357 to −297 mV for DMPC and from −238 mV versus SCE to −291 mV versus SCE for DDAB, whereas for the protein in solution the redox potential changes from −547 to −394 mV versus SCE (Zhang et al., 1997; Jefferson and Griffin, 1972). These results indicate that electron transfer involves the heme Fe3+/2+ couple of the enzyme. The heterogeneous electron transfer rate (ks ) was calculated in this case to be 25–26 s−1 . The authors have obtained catalytic response with 50 mM trichloroacetic acid in apparently oxygen-free measuring buffer. 4.4. Electrodes modified with multilayer films In order to improve the direct electron transfer between Au electrodes and heme proteins like CYP, Rusling and co-workers have investigated the direct electrochemistry of CYP101 using layer-by-layer films of CYP101 with polyions (Lvov et al., 1998; Munge et al., 2003). In these studies Au and pyrolytic graphite (PG) electrodes were modified by sequential adsorption of poly(styrenesulfonate) (PSS) and/or branched poly(ethyleneimine) (PEI) and CYP101 thus creating CYP101-multilayer films. Quartz crystal microbalance (QCM) investigation has revealed regular and reproducible layer formation. The thickness and the amount of CYP101 for bilayers on Au were 15 nm and 0.1 nmol cm−2 . The association of CYP101 with the cationic PEI and the anionic PSS was possible due to the distribution of positively and negatively charged amino acids on the surface of the protein. Direct electron transfer between the heme group of CYP101 and the Au electrode was observed in both cases, although voltammetric peaks were larger with PSS than with PEI. The

E◦ was −250 mV versus SCE at neutral pH. The positive shift in comparison to the redox potential of CYP101 in solution (−547 mV versus SCE) may be due to the surfactant head group charge and its association with the protein’s surface. Furthermore, oxidation of styrene was successfully catalysed by polyion films containing cytochrome CYP101. GC–MS product analysis (styrene oxide) showed a turnover number of 9.3 h−1 , which is larger than the turnover number when CYP101 was in solution, that is 0.35 h−1 . Catalysis of styrene as well as benzaldehyde oxidation has also been observed by the same laboratory using proteinpolyion modified carbon cloth (CC) electrodes (Zu et al., 1999). The formal potential was the same as that reported with CYP101-multilayer modified Au electrode (Lvov et al., 1998), but the turnover number for the catalysis of styrene was slightly lower 7.2 h−1 . However, using CYP101 in solution the turnover was 7.0 h−1 , which approached that of immobilised CYP101. It was also significantly larger than the turnover number observed with Au electrode, i.e., 0.35 h−1 (Zhang et al., 1998). All turnover numbers were calculated after product analysis was carried out with GC–MS. A reason for the larger turnover number with CC electrodes could be the 20-fold larger active surface area of the CC electrode as compared to the Au electrode. Addition of 3000 units of catalase destroyed the H2 O2 and thus decreased the turnover number significantly to 0.2 h−1 . This reflects the fact that catalysis of styrene or benzaldehyde with CYP101 is carried out through H2 O2 . Conversion of styrene-to-styrene oxide has also been studied with a biosensor based on CYP1A2 (Estavillo et al., 2003). Cytochrome CYP1A2 is the main enzyme that metabolizes caffeine but it is also relatively active in converting styrene, although not as active as CYP2E1 and CYP2B6. In this study, however, PSS-CYP1A2 multilayers were grown on CC electrodes until films denoted PSS/(cyt CYP/PSS)2 were obtained. The E◦ obtained was −310 mV versus SCE, which is similar to values obtained in the other works with multilayers described above. Oxidation of 10 mM styrene was monitored using a PSS-CYP1A2 modified CC electrode poised at −600 mV versus SCE. Results obtained indicated a turnover rate of 39 h−1 , which is significantly higher than 9.3 and 7 h−1 obtained with CYP101 (Lvov et al., 1998; Zu et al., 1999). Assays carried out with CYP101 and CYP1A2 in conventional solution reactions with electron donors and reductase showed turnover rates that lies at 10 h−1 for CYP101 and 17 h−1 for CYP1A2. In addition to CYP101 and CYP1A2, CYP2E1 has also been studied with phospholipid modified GC and Au electrodes. Reversible electrochemistry was observed both with DDAB and poly(diallyldimethylammonium chloride) (PDDA) modified GC electrodes with an E◦ of −329 mV versus SCE. The electron transfer rate was 2 s−1 for DDAB and 1 s−1 for PDDA, which is lower than the similar study done by Zhang and co-workers. The authors have even examined the direct electrochemistry of CYP2E1 using Au electrodes modified with thiol and/or PDDA. Using a

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mercaptopropionic acid (MPA)-PDDA modified Au electrode an E◦ of −530 mV versus SCE and a ks of 2 s−1 were obtained, which with a cystamine-maleimide modified Au electrode the E◦ and ks were −421 mV versus SCE and 10 s−1 . Although the electron transfer rate is faster when a cystamine-maleimide modified Au electrode than the other Au modified electrodes was used, it is still not as fast as the electron transfer rate obtained with GC modified electrodes. Besides catalyzing styrene and benzaldehyde, CYP enzymes play an important role in the metabolism of endogenous compounds as well as in pharmacokinetics and toxicokinetics. That’s why Joseph have developed a biosensor with human CYP3A4 as a novel drug-screening tool (Joseph et al., 2003). It was constructed by assembling enzyme films on Au electrodes by alternate adsorption of a layer of CYP3A4 on top of a layer of PDDA. The biosensor was applied to the detection of verapamil, midazolam, quinidine and progesterone. QCM monitoring showed that the protein concentration on the surface of the biosensor was 27 pmol cm−2 . Electrochemical investigation of the enzyme-bound film revealed well-defined anodic and cathodic redox peaks with an E◦ of −146 mV versus SCE, which indicates reversible oxidation and reduction of the heme group. Furthermore, the enzyme was catalytically active as indicated by the concentration dependent catalytic responses obtained using verapamil, midazolam, quinidine and progesterone. Product analysis after the electrolysis also confirmed the catalytic activity of the enzyme. A linearity of calibration up to 2.85 mM with verapamil was obtained. Kapp m values calculated from amperometric data for midazolam, progesterone and quinidine were 0.547, 0.271 and 1.082 mM, respectively. The response time to reach 95% of the steady state was approximately 15–25 s. Catalytic response has also been obtained with fluoxetine using CYP2D6 on a polyaniline-doped GC electrode. The enzyme exhibited reversible electrochemsitry with a E◦ of −120 mV versus SCE, which upon increasing concentrations of fluoxetine a cathodic shift was observed up to 350 ␮M of substrate. Linearity was however observed only until 1.0 ␮M. At higher concentration saturation was attained. The limit of detection was 1nM and the Kapp m value 3.7 ␮M (Iwuoha et al., 2004). 4.5. CYP bionsensing at high temperatures An interesting aspect of CYP electrochemistry is the electron transfer ability of a CYP from thermophilic organisms at high temperatures. CYP119 is found in Sulfolobus tokodaii strain 7 and its crystal structure was recently determined (Oku et al., 2004). CYP119 has an unusually high denaturation point of around 90 ◦ C and tolerance for extreme pH values and pressures for extended periods in solution (Yano et al., 2000; Koo et al., 2000, 2002; Puchkaev et al., 2002). Using DDAB modified plastic formed carbon (PFC) electrodes Oku and co-workers adsorbed CYP119 and have shown the electrochemistry at 20 ◦ C and at 80 ◦ C (Oku et al., 2004). In

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the first case a quasi-reversible reduction and oxidation was observed with an E◦ of −250 mV versus Ag/AgCl and a peak seperation (E) of 90 mV at a scan rate of 0.2 V s−1 . The E◦ at 80 ◦ C was shifted to −50 mV versus Ag/AgCl and the E lied at 30 mV. This result clearly shows that CYP119 is redox active even at higher temperatures, whereas the ordinary CYPs have a relatively low thermostability. Dehalogenation of hepatotoxic and carcinogenic solvents like chloroform and methyl chloride have been electrocatalytically induced using this thermophilic enzyme (Blair et al., 2004). Blair and co-workers have immobilized CYP119 in a methyltriethoxysilane sol–gel film on graphite and also adsorbed in dimethyldidodecylammonium poly(p-styrene sulfonate) (DDABPSS) modified graphite electrode. The enzyme retains about 93% of its activity when adsorbed in DDABPSS at 30 ◦ C with only a moderate loss at temperatures between 80 and 90 ◦ C. E◦ was −220 mV versus SCE at 30 ◦ C and −285 mV versus SCE at 80 ◦ C. Catalytic currents were observed with 10 mM CH2 Cl2 , CHCl3 and CCl4 with electron turnover ranging from 4.5 s−1 (CH2 Cl2 ) to 52.1 s−1 (CCl4 ). Mutagenesis studies have shown that catalysis with CYP119 was improved when the side chain of Thr214, which although not highly conserved in other CYP systems is next to the highly conserved and catalytically important Thr213, was replaced by Val (Koo et al., 2002). In particular, kcat for lauric acid hydroxylation, increased from 0.36 min−1 (wild type) to 2.08 min−1 (T214V mutant) and even to 8.80 min−1 when the additional D77R mutation was introduced. Aspartic acid (D) is also near the active site of CYP119 (Koo et al., 2000). Measurements were carried out in the presence of Pdx, Pdx reductase and NADH. Styrene epoxidation was also improved in the same study with Vmax increasing from 0.05 min−1 (wild type) to 0.20 min−1 (T214V). 4.6. Spectroelectrochemistry and surface plasmon resonance of CYP Although direct electron transfer between the electrode and CYP has been established, one important but still not solved question is whether cytochromes CYP remains native when interacting with electrodes. Spectroelectrochemistry allows getting an insight into the structural changes accompanied with the electrochemical redox cycling. Spectroelectrochemistry of wild type high-spin CYP101 has been carried out by using mediator (Reipa et al., 1997; Mayhew et al., 2000; Reipa et al., 2002) and mediator-free (Bistolas et al., 2004) electrode system. The spectroelectrochemistry of CYP101 in the presence of mediators such as Pdx and phenosafranine was studied using antimony-doped tin oxide electrode. Although electron transfer was observed between the electrode, via the mediator to CYP101, no spectra were taken in the presence of CO, which is the best indicator for the presence of the protein’s native state. We therefore investigated CYP101 with spectroelectrochemistry on a mediatorfree environment with the purpose to indicate that the protein

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retains its native state in the electrochemical cell during electrolysis. Reversible oxidation and reduction was observed using both 4,4 -dithiodipyridin and sodium dithionite modified Au capillary electrodes (Bistolas et al., 2004). An E◦ of −380 mV versus SCE was determined, which is similar to the CV studies of CYP101 (Lei et al., 2000) and the redox potential of high-spin CYP101 −394 mV versus SCE. The spectra of CYP101 in the presence of either carbon monoxide or metyrapone, both being inhibitors of CYP101 catalysis, displayed the spectroscopic pattern that is characteristic of the binding of these two ligands to the heme center of the enzyme. This is the first spectroscopic proof that the CYP101 remains in the native state during electrolysis. In order to elucidate the mechanisms whereby interprotein electron transfer takes place in CYP systems and how the system is operated, Ivanov and Archakov have constructed an optical biosensor to study binary and ternary complex formation of CYP2B4, CYP101 and CYP11A1 with their respective redox partners (Ivanov et al., 1999, 2000, 2001a, 2001b, 2001c). The method was based on the design of an optical evanescent sensor, a resonant mirror (Ivanov et al., 2001a). Proteins were immobilized on the carboxymethylated dextran-coated cuvette. The results indicated that on one hand, the complex formation of CYP2B4 with its redox partners was demonstrated to occur due to hydrophobic interactions of protein fragments with the phospholipid layer. On the other hand electrostatic interactions were important for the complex formation of CYP101 with its redox partners. Complex formation of CYP11A1 with its redox partners were problematic to be seen due to the two orders higher lifetime of the binary complexes than the hydroxylation cycle. This has ruled out the effective functioning of adrenodoxin as electron carrier in the complex. The associations rate constants for the CYP2B4, NADPH-CYP reductase and cytochrome b5 complexes formed were found to be close to the diffussion limit −0.5 to 4 × 106 M−1 s−1 —while their dissociation constants did not exceed 0.5 s−1 (Ivanov et al., 1999). For CYP101, putidaredoxin reductase and putidaredoxin complex the association rate constant was 0.34 × 106 M−1 s−1 , while the dissociation constant was 0.047 s−1 . It was also shown that the interprotein electron transfer occurs not only during complex formation but also upon random collision. This was due to hydrophobic and charged amino acids on the surface of the CYP, which play a dominant role in the formation of productive electron transfer complexes.

ing of the electrode surface for appropriately orienting the terminal oxidase towards the electrode have been successfully adopted from cytochrome c. In this respect, the electrostatic attraction on molecular level has been used. However, due to the well-known low conformational stability pronounced structural changes are plausible in the process of embedding in the matrix at the electrode. These deviations from the behaviour in solution are obviously reflected by the anodic shift of the structure-free protein and it is reflected by the smaller anodic peak in CVs. The electron transfer at electrodes modified with monolayers of phospholipids or mercaptolalkanol/acids should resemble in analogy to cytochrome c—the other sphere type. The mode of electron transfer via conducting polymers or multilayers like clay nanoparticles or polyelectrolytes seems widely unclear. For the fast heterogenous electron transfer with glucose oxidase – an intrinsic redox enzyme with the prosthetic group buried deep within the protein fabric – the coupling of redox relays to the protein has been established (Degani and Heller, 1987). This concept has been successfully transferred to the CYP electrochemistry by Archakov’s group by binding riboflavin to the protein surface (Shumyantseva et al., 2000). On the other hand, the concept of ‘redox wire’ (Willner and Katz, 2000) transferring the electrons via immobilized mediators from the electrode to the prosthetic group has not yet been realized for the electrochemical substrate hydroxylation by CYP. Interpretations of the reaction mechanism are complicated by the fact that substrate conversion requires oxygen whilst the reduction of the CYP heme is mashed by the cathodic oxygen reduction. Most probably, both reduction of the prosthetic group and of oxygen proceed in parallel at the electrode. To unravel the complexity, the influence of catalase and of CYP inhibitors is a useful diagnostic criterion. (i) Substrate conversion by the by-product H2 O2 is suppressed by the addition of catalase. (ii) Substrate hydroxylation via the two-e-reduction of the iron–oxygen complex is effected by the presence of inhibitors, e.g., metyrapone. These criteria have not yet been strictly applied in the majority of investigations.

6. Future perspectives 5. Summary and conclusions Regarding protein electrochemistry the number of papers on CYP ranks third after cytochrome c and glucose oxidase. The binding sites for electron transferring proteins at the molecule surface which are essential for the fast electron transfer and subsequent oxygen activation characterize the CYP family to the intrinsic redox proteins according to Ikeda’s (Ikeda, 1992) classification. Therefore, the engineer-

In most cases biosensors based on CYP can catalyze a variety of substrates, as has been described above. In recent years profit has been taken from the use of protein engineering. This implies the introduction of specific attachment regions and electron transfer regions to overcome the kinetic limitations. Furthermore, engineering of the protein surface could provide means of immobilizing the protein directly on the electrode, thus shortening the distance between the redox center and the electrode and achieving faster electron transfer.

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Recent work indicates the potential of engineering sites for surface binding and redox active dyes, de-novo designed redox systems and genetic chimeras (Gilardi et al., 2001, 2002; Wong and Schwaneberg, 2003). Nature offers almost unique biocatalysis of the CYP family exhibiting both (i) almost absolute specificity for recognizing a given substrate also in mixtures of very similar compounds, e.g., steroid hormones, and (ii) conversion of highly different compounds following the same reaction type, e.g., demethylation of various drugs. Using genetic engineering both recognition types may be optimized for the analytical system under investigation. In this respect, arrays carrying different CYP isoforms will be developed for characterizing the metabolism of drug converting organs, e.g., the liver of different species. Full interaction of specific substrate conversion and electrochemical signal transduction can be expected in the next generation of electronic biochips.

Acknowledgments The funding from the Fond der Chemische Industrie, European Community (Intellisens QLK-3-CT2000-01481) and German Ministry of Education and Research (BMBF 3i308B) is greatly appreciated.

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