Direct electrochemistry of human and rat NADPH cytochrome P450 reductase

Electrochemistry Communications 8 (2006) 1845–1849 www.elsevier.com/locate/elecom

Direct electrochemistry of human and rat NADPH cytochrome P450 reductase Alka Shukla a, Elizabeth M.J. Gillam a, Paul V. Bernhardt a

b,*

Centre for Metals in Biology, School of Biomedical Sciences, University of Queensland, Brisbane 4072, Australia b Centre for Metals in Biology, Department of Chemistry, University of Queensland, Brisbane 4072, Australia Received 19 July 2006; received in revised form 10 August 2006; accepted 10 August 2006 Available online 15 September 2006

Abstract The diflavo-protein NADPH cytochrome P450 reductase (CPR) is the key electron transfer partner for all drug metabolizing cytochrome P450 enzymes in humans. The protein delivers, consecutively, two electrons to the heme active site of the P450 in a carefully orchestrated process which ultimately leads to the generation of a high valent oxo-heme moiety. Despite its central role in P450 function, no direct electrochemical investigation of the purified protein has been reported. Here we report the first voltammetric study of purified human CPR where responses from both the FMN and FAD cofactors have been identified using both cyclic and square wave voltammetry. For human CPR redox responses at 2 and 278 mV (with a ratio of 1e:3e) vs NHE were seen at pH 7.9 while the potentials for rat CPR at pH 8.0 were 20 and 254 mV. All redox responses exhibit a pH dependence of approximately 59 mV/pH unit consistent with proton coupled electron transfer reactions of equal stoichiometry. Ó 2006 Elsevier B.V. All rights reserved. Keywords: Cytochrome P450 reductase; Protein; Voltammetry

1. Introduction Cytochrome P450s (P450s) are heme containing enzymes that play a major role in the metabolism of xenobiotics [1]. Most P450s catalyze a 2-electron oxidation reaction of an organic substrate coupled to O-transfer (Eq. (1)). Dioxygen and the substrate (RH) bind at the heme active site and electrons from NADPH act to cleave the O–O bond leaving a high valent oxo-heme moiety that is capable of O-insertion into an otherwise inert C–H bond [1]. NADPH + RH + O2 + Hþ ! NADPþ + ROH + H2 O ð1Þ Delivery of the two electrons from NADPH to the P450 must be carefully orchestrated in order to avoid so-called uncoupling i.e., the futile catalytic reduction of dioxygen to superoxide, hydrogen peroxide or water without sub*

Corresponding author. Tel.: +61 7 3365 4266; fax: +61 7 3365 4299. E-mail address: [email protected] (P.V. Bernhardt).

1388-2481/$ - see front matter Ó 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.elecom.2006.08.020

strate turnover. Microsomal P450s require the NADPH cytochrome P450 reductase (CPR) as a relay, which accepts electrons from NADPH and transfers them sequentially to the P450. CPR is a rare example of a mammalian protein containing both flavin adenine dinucleotide (FAD) and flavin mononucleotide (FMN) cofactors within a single polypeptide chain [2]. The FMN domain has strong resemblance to bacterial flavodoxins whereas the FAD domain is related to ferredoxin NADP+-reductase. It has been proposed that CPR and other FMN/FAD containing proteins have evolved as a fusion of two ancestral genes [3]. CPRs from a number of eukaryotic organisms (including humans) are highly homologous [4]. Crystal structures of wild type and mutant forms of CPR from rat liver have been solved [5,6]. These structures reveal that NADP+ binds at the FAD site. Electrons are passed sequentially to the adjacent ˚ away) and then on to any of a numFMN cofactor (5 A ber of P450s; a single form of CPR serves all microsomal P450s [1,2].

1846

A. Shukla et al. / Electrochemistry Communications 8 (2006) 1845–1849

To facilitate isolation and solution studies, the hydrophobic amino terminal domain of CPR may be cleaved by treating the enzyme with pancreatic steapsin or trypsin, releasing the soluble C-terminal 72-kDa hydrophilic domain. This soluble CPR contains both the FAD and FMN cofactors and is catalytically active with many electron acceptors (such as cytochrome c), but it does not reduce cytochromes P450 in its modified form [7]. Potentiometric studies on human [8] and rabbit liver [9] CPR have been reported, where the redox potentials of the FAD and FMN cofactors were determined. Redox properties of the closely related methionine synthase reductase (MSR) [10], nitric oxide synthase (NOS) [11] and human novel reductase 1 (NR1) [12] have also been determined recently by potentiometric titrations. A feature of CPR, and its related di-flavoproteins, is a capacity to stabilize single-electron reduced (‘semiquinone’) forms of both its FAD and FMN cofactors. This is in contrast to the free FMN and FAD cofactors which undergo obligate cooperative 2-electron transfer reactions. In the case of CPR, four redox couples (FADox/FADsq, FADsq/FADred, FMNox/FMNsq and FMNsq/FMNred) may be resolved potentiometrically. In human and rabbit CPR the FMNox/FMNsq couple is found at a potential almost 200 mV positive of the other three redox couples. Indeed the singly reduced form of CPR (FMNsq:FADox) is the resting state of the enzyme. Given its importance in P450 catalysis, it is perhaps surprising that there has been very little work on the electrochemistry of any CPR. Recently a report of the cyclic voltammetry of insect cell microsomes containing recombinant human P450s 1A2 and 3A4 (comprising a mixture of CPR, lipid, cytochromes P450 and other membrane proteins) has appeared [13]. A single low potential voltammetric response (250 mV vs NHE, pH 7.4) was reported, which was assigned to that of CPR and not due to any P450s present or other proteins in the mixture. A similar response was seen in the voltammetry of isolated rabbit CPR. No mention was made of the absent high potential FMNox/FMNsq response of CPR, which apparently was not observed [13]. Herein we report the first direct electrochemical study of isolated recombinant human CPR and a parallel study with recombinant rat CPR. We have identified both low and high potential responses that for the first time unequivocally show that CPR may undergo electrochemically driven redox reactions. This is a first important step towards the construction of direct electrochemically driven, reconstituted CPR/P450 systems that may lead toward a protocol for rapid screening of P450 metabolites on the basis of electrochemical detection. 2. Materials and methods 2.1. Materials 2 0 AMP (2 0 adenosine monophosphate), FMN and adenosine were obtained from Sigma Chemical Company. Cel-

lulose ester dialysis membrane (MWCO: 8000) was obtained from Spectrum Laboratories (Los Angeles, CA, USA). Aprotonin was obtained from Boehringer Mannheim Australia Pty Ltd. (Castle Hill, NSW, Australia). Bactotryptone, yeast extract and bactopeptone were purchased from Difco (Detroit, MI, USA), Ampicillin, chloramphenicol and isopropyl b-D-thiogalatopyranoside (IPTG) were obtained from Progen Industries Pty Ltd. (Darra, Qld, Australia). All other reagents were obtained from local suppliers at the highest quality commercially available. 2.2. Protein expression and purification Human and rat CPR expression was performed using the expression construct pCW/hNPR and pOR263 in DH5aF’IQe strain of E. coli (Invitrogen, Life Technologies, Carlsbad, USA) cotransformed with the chaperone expression plasmid pGro7 [14] using the method of Inoue et al. [15]. The same procedure was used for the expression and purification of human and rat CPR. The rat CPR was purified by Rebecca Wunsch. Expression was done according to established procedures [16]. Spheroplasts were resuspended in 50 mM Tris/HCl buffer pH 8.0, containing 5 mM EDTA, 20% glycerol and protease inhibitors (0.1 mM PMSF, 10 lg/mL aprotinin and 2 lM leupeptin), then sonicated in salt/ice bath, and centrifuged at 10,000g for 40 min at 4 °C. The resultant supernatant was centrifuged at 105,000g for 65 min at 4 °C to collect membranes which were then solubilized in 120 mL 100 mM Tris/HCl pH 7.7, containing 1 mM EDTA, 20% glycerol, 0.1% Tergitol, 0.6% CHAPS, 1 lM FMN, 1 mM DTT, 0.1 mM PMSF, and 10 lg/ml aprotinin with gentle stirring at 4 °C for 2–3 h. The solubilized membrane fraction was centrifuged at 4 °C for 65 min at 105,000g. The supernatant was adjusted to 0.3 M KCl and then loaded onto an ADP Sepharose column pre-equilibrated with 50 mM potassium phosphate buffer pH 7.7, containing 300 mM KCl, 20% w/v glycerol, 0.1 mM EDTA, 0.35% w/v cholate, 1 lM FMN, 0.2 mM DTT, and 0.4 mM PMSF. The column was washed with 100 mL 50 mM potassium phosphate buffer pH 7.7, containing 20% w/v glycerol, 0.35% cholate, 5 mM adenosine, 1 lM FMN, 0.2 mM DTT, 0.4 mM PMSF. The column was eluted with 50 mM potassium phosphate buffer pH 7.7, containing 20% glycerol, 0.1 mM EDTA, 0.35% cholate, 1 lM FMN, 5 mM 2 0 AMP, and 0.4 mM PMSF and 2–5 mL colored fractions were collected. The reductase was dialyzed against two changes (1 L and 12 h each) of 10 mM potassium phosphate buffer pH 7.7, containing 20% w/v glycerol, 0.35% w/v cholate, 1 lM FMN, and 0.4 mM PMSF. The post-dialysis sample was loaded onto hydroxyapatite column equilibrated with 10 mM potassium phosphate buffer pH 7.7, containing 20% w/v glycerol, 0.35% w/v cholate, 0.1 mM DTT, 0.1 mM PMSF and washed with 250 mL 10 mM potassium phosphate buffer pH 7.7,

A. Shukla et al. / Electrochemistry Communications 8 (2006) 1845–1849

containing 20% w/v glycerol, 0.1 mM DTT, and 0.1 mM PMSF. Elution was performed with 300 mM potassium phosphate buffer pH 7.7, 20% w/v glycerol, 0.1 mM EDTA, and 0.1 mM PMSF, and 1 mL fractions containing reductase were pooled and dialyzed against 2–4 changes of 1 L each of 10 mM Tris acetate pH 7.4, containing 1 mM EDTA, and 20% w/v glycerol. The concentration of human (6.9 lM) and rat CPR (51 lM) was determined from its activity towards cytochrome c using a specific activity of 3200 nmol cytochrome c reduced/ min/nmol [17].

1847

400 nA

SWV (offset)

200 nA

CV (background subtracted)

2.3. Electrochemistry -400

Cyclic and square wave voltammetry were performed on a BAS 100B/W analyzer employing an edge-plane pyrolytic graphite (EPG) electrode (modified as described below), a Pt wire counter and Ag/AgCl (3 M KCl) reference electrode (Eo +196 mV vs NHE). All measurements were carried out in a glove box (Belle Technology) with oxygen level below 10 ppm and at 25 °C. The electrochemical cell contained 0.4 mL of 50 mM of bis-tris propane and 2amino-2-methylpropane hydrochloride buffer (adjusted to the desired pH with AcOH or NaOH) in addition to 10 mM NaCl as electrolyte. The EPG electrode was cleaned by cleaving several 2 lm layers from the surface using a microtome then the electrode was thoroughly rinsed with water, sonicated twice and allowed to dry at room temperature. Purified human or rat CPR (5 lL) was mixed with 5 lL of an aqueous didodecyldimethylammonium bromide (DDAB) solution (5 mM) and then cast onto the working electrode surface and the film was allowed to set overnight at 4 °C. Baseline subtractions were performed with either the program UTILS [18], which was kindly provided by the author, or the BAS100W software (vers. 3.2). 3. Results and discussion Cyclic voltammetry of both purified recombinant human and rat CPR confined to an edge-plane pyrolytic graphite electrode within a didodecyldimethylammonium bromide (DDAB) surfactant film yielded two well separated pairs of anodic and cathodic redox responses. The voltammogram for human CPR in Fig. 1 at pH 7.9 (red curve) has the background capacitive current removed for clarity. The square wave voltammogram at the same pH is also shown (Fig. 1 blue curve) where peaks at 2 and 278 mV vs NHE are resolved. The anodic and cathodic current maxima exhibit a linear dependence on sweep rate which identifies the responses as due to a surface adsorbed molecule. The higher potential anodic and cathodic peaks have a width of 110 mV at half height and are consistent with a single-electron reaction (theoretically 90/n mV at 25 °C) [19]. Integration of the peak (after correcting for charging current) gives a charge of 1.6 nC (840 fmol of electroactive protein). The lower

-300

-200

-100

0

100

E (mV vs NHE) Fig. 1. Background subtracted cyclic voltammogram of human CPR (red curve, bottom). Sweep rate 20 mV s1, pH 7.9. Square wave voltammogram also shown (blue curve, top) square wave amplitude 5 mV, frequency 8 Hz. Note different current scales.

potential response has a peak width at half height of 70 mV and as such is consistent with a convoluted multi electron wave. Integration of this peak yields a charge of 4.6 nC; ca. 3 times that of the higher potential response. Similar cyclic voltammetry results were obtained for rat CPR. Integration of the voltammetric peaks gave, once more, a ratio of 3:1 for the charge passed in the low and high potential peaks and an electroactive coverage of 570 fmol rat CPR. Most importantly, we observed high potential redox responses (ca. 150–200 mV higher than the response at ca. 0 V in Fig. 1) that were due to surface oxidation processes on the EPG electrode as described by Evans and Kuwana [20]. These responses fall well outside the potential range shown in Fig. 1 are of no consequence to the present study. For both human and rat CPR pH dependences of about 59 mV/pH unit were identified for their high (1 electron) and low potential (3 electron) responses (Fig. 2), which is consistent with an overall stoichiometry of 1e/1 H+ (high potential) and 3e/3H+ (low potential) transferred during the sweep within the range 6 < pH < 9. Redox potentiometric studies of the FMN-dependent flavodoxins [21–24] have found that the high potential FMNox/FMNsq couple typically exhibits a dependence of 59 mV/pH unit with no pKa i.e., the FMNsq form is protonated at all accessible pH values. The lower potential couple of flavodoxins often exhibits a pH dependence below about pH 7, where the FMNred form binds an additional proton. There is greater variation in the redox potentials of FAD-dependent proteins such as the ferredoxin reductases with some being able to stabilize a semiquinone [25] while others do not [26]. The redox potentials of human CPR as determined by redox potentiometry at pH 7.0 [8] were 66 mV (FMNox/ FMNsq), 269 mV (FMNsq/FMNred), 283 mV (FADox/ FADsq) and 382 mV (FADsq/FADred). Our voltammetric

1848

A. Shukla et al. / Electrochemistry Communications 8 (2006) 1845–1849

200

Em(mV vs NHE)

100 0 -100 -200

Human CPR (DDAB film) Rat CPR (DDAB film) Human CPR (in solution) DDAB film (no protein)

(a)

abs. (arbitrary scale)

E1 human CPR E2 human CPR E1 rat CPR E2 rat CPR

(b)

(c)

-300 (d)

-400 6

0

7

8

9

pH

400

450

500

550

600

650

wavelength (nm) Fig. 2. pH dependence of the redox potentials of human (circles and squares) and rat (triangles) CPR as determined by square wave voltammetry (square wave amplitude 5 mV, frequency 8 Hz). The high potential (E1) and low potential (E2) responses are indicated.

data at the same pH are +58 mV (FMNox/FMNsq) and 228 mV for the three remaining couples. The pH dependences of the redox potentials of human or rat CPR have not been reported before. Regardless of pH, we were unable to resolve the lower potential 3-electron response into more than one component. The anodic shift in redox potentials seen here relative to those reported from solution potentiometric redox titrations may be a consequence of the surfactant used in immobilizing the protein to the electrode surface. We [27] and others [28] have noted similar anodic shifts in the voltammetry of P450s relative to their potentials determined in solution using redox potentiometry. Voltammetry of the individual protein-free FAD and FMN cofactors was conducted under identical experimental conditions as those employed for the CPR holoprotein. Single, reversible, 2-electron voltammetric waves were seen at 280 mV (pH 8) for both FAD and FMN. This is consistent with previous studies of these compounds [29–33]. Coincidentally these waves fall in a similar region as the low potential couple of both human and rat CPR. However, the free flavins lack the high potential, single electron wave of both CPR proteins as they cannot benefit from specific hydrogen bonding interactions within the protein that stabilize the semiquinone form of the FMN cofactor in particular at close to neutral pH [34]. In order to examine whether any major structural changes might have occurred upon immobilizing the protein in a DDAB surfactant film, we measured the electronic spectra of both human and rat CPR within a DDAB film cast onto a glass slide. The results are shown in Fig. 3 in comparison with the solution spectrum of human CPR (Fig. 3c), which is essentially indistinguishable from that of rat CPR in solution (not shown). The spectrum of the blank DDAB film is also given for comparison (Fig. 3d).

Fig. 3. Visible spectra of (a) human CPR in a DDAB film (cast on a glass slide); (b) rat CPR in a DDAB film (cast on a glass slide); (c) human CPR in solution and (d) a DDAB film cast on a glass slide with no protein. The absorption scale of each spectrum has been normalised to correct for variations in protein concentrations and film thicknesses and also offset for clarity.

The most prominent electronic maxima apparent in the solution spectrum of human CPR emerge at 380 nm, 450 nm and a very broad peak around 600 nm. The two higher energy maxima are associated with the fully oxidized flavin chromophore (FMN or FAD) while the low energy maximum is from the flavin semiquinone [8]. The human CPR sample used here is evidently already in a partially reduced form (Fig. 3c). The DDAB film spectra of human (Fig. 3a) and rat (Fig. 3b) CPR exhibit the same features as the solution spectrum although the quality of the spectra are affected by a sloping baseline (opaque films) and inherently small optical density changes due to the thin films (short path lengths). Nevertheless, there is no evidence to suggest that the protein has been altered by immobilization in the DDAB film. 4. Conclusions Having established that direct electron transfer is possible between either human or rat CPR and a working electrode, future studies will focus on using these CPRs as an electron bridge between the electrode and human P450s under turnover conditions i.e., in air and in the presence of substrate. It is our hope that the use of a natural electron partner will lead to tighter coupling of the electrochemically driven catalytic reaction and greater overall efficiency than seen in previous work using artificial mediators or no electron relay to the heme active site. Acknowledgments We wish to acknowledge the help of Dr. Pavel Soucek in the purification of human reductase and of Rebecca

A. Shukla et al. / Electrochemistry Communications 8 (2006) 1845–1849

Wunsch in purifying the rat reductase. Financial support of the Australian Research Council and an Australia Asia Award for India (to AS) is gratefully acknowledged. References [1] F.P. Guengerich, in: P.R. Ortiz de Montellano (Ed.), Cytochrome P450: Structure Mechanism and Biochemistry, Plenum Press, New York, 2005. [2] A. Gutierrez, A. Grunau, M. Paine, A.W. Munro, C.R. Wolf, G.C.K. Roberts, N.S. Scrutton, Biochem. Soc. Trans. 31 (2003) 497–501. [3] T.D. Porter, C.B. Kasper, Biochemistry 25 (1986) 1682–1687. [4] A.L. Shen, C.B. Kasper, Handbook of Experimental Pharmacology 105 (1993) 35–59. [5] M. Wang, D.L. Roberts, R. Paschke, T.M. Shea, B.S.S. Masters, J.J.P. Kim, Proc. Natl. Acad. Sci. USA 94 (1997) 8411–8416. [6] P.A. Hubbard, A.L. Shen, R. Paschke, C.B. Kasper, J.-J.P. Kim, J. Biol. Chem. 276 (2001) 29163–29170. [7] I.F. Sevrioukova, J.A. Peterson, Biochimie 77 (1995) 562–572. [8] A.W. Munro, M.A. Noble, L. Robledo, S.N. Daff, S.K. Chapman, Biochemistry 40 (2001) 1956–1963. [9] T. Iyanagi, N. Makino, H.S. Mason, Biochemistry 13 (1974) 1701– 1710. [10] K.R. Wolthers, J. Basran, A.W. Munro, N.S. Scrutton, Biochemistry 42 (2003) 3911–3920. [11] P.E. Garnaud, M. Koetsier, T.W.B. Ost, S. Daff, Biochemistry 43 (2004) 11035–11044. [12] R.D. Finn, J. Basran, O. Roitel, C.R. Wolf, A.W. Munro, M.J.I. Paine, N.S. Scrutton, Eur. J. Biochem. 270 (2003) 1164–1175. [13] N. Sultana, J.B. Schenkman, J.F. Rusling, J. Am. Chem. Soc. 127 (2005) 13460–13461. [14] K. Nishihara, M. Kanemori, M. Kitagawa, H. Yanagi, T. Yura, Appl. Environ. Microbiol. 64 (1998) 1694–1699.

1849

[15] H. Inoue, H. Nojima, H. Okayama, Gene 96 (1990) 23–28. [16] H.K. Crewe, L.M. Notley, R.M. Wunsch, M.S. Lennard, E.M.J. Gillam, Drug Metab. Dispos. 30 (2002) 869–874. [17] Y. Yasukochi, B.S.S. Masters, J. Biol. Chem. 251 (1976) 5337–5344. [18] H.A. Heering, UTILS, Utilities for Data Analysis, University of Delft, The Netherlands, 2001. [19] A.J. Bard, L.R. Faulkner, Electrochemical Methods: Fundamentals and Applications, 2nd ed., Wiley, New York, 2001. [20] J.F. Evans, T. Kuwana, Anal. Chem. 49 (1977) 1632–1635. [21] S.G. Mayhew, G.P. Foust, V. Massey, J. Biol. Chem. 244 (1969) 803– 810. [22] M. Dubourdieu, J. Le Gall, V. Favaudon, Biochim. Biophys. Acta 376 (1975) 519–532. [23] G.A. Sykes, L.J. Rogers, Biochem. Soc. Trans. 10 (1982) 414–415. [24] G.A. Sykes, L.J. Rogers, Biochem. J. 217 (1984) 845–850. [25] J.J. Pueyo, C. Gomez-Moreno, S.G. Mayhew, Eur. J. Biochem. 202 (1991) 1065–1071. [26] K.J. McLean, N.S. Scrutton, A.W. Munro, Biochem. J. 372 (2003) 317–327. [27] A. Shukla, E.M. Gillam, D.J. Mitchell, P.V. Bernhardt, Electrochem. Commun. 7 (2005) 437–442. [28] Z. Zhang, A.-E.F. Nassar, Z. Lu, J.B. Schenkman, J.F. Rusling, J. Chem. Soc., Faraday Trans. 93 (1997) 1769–1774. [29] R.D. Braun, J. Electrochem. Soc. 124 (1977) 1342–1347. [30] O.S. Ksenzhek, S.A. Petrova, Bioelectrochem. Bioenerg. 11 (1983) 105–127. [31] V.I. Birss, H. Elzanowska, R.A. Turner, Can. J. Chem. 66 (1988) 86– 96. [32] L.T. Kubota, L. Gorton, A. Roddick-Lanzilotta, A.J. McQuillan, Bioelectrochem. Bioenerg. 47 (1998) 39–46. [33] R. Garjonyte, A. Malinauskas, L. Gorton, Bioelectrochemistry 61 (2003) 39–49. [34] D.M. Hoover, C.L. Drennan, A.L. Metzger, C. Osborne, C.H. Weber, K.A. Pattridge, M.L. Ludwig, J. Mol. Biol. 294 (1999) 725–743.