The versatility of the fungal cytochrome P450 ... - Wiley Online Library

Molecular Microbiology (2011) 81(5), 1374–1389 䊏

doi:10.1111/j.1365-2958.2011.07772.x First published online 2 August 2011

The versatility of the fungal cytochrome P450 monooxygenase system is instrumental in xenobiotic detoxiﬁcation mmi_7772 1374..1389

Ljerka Lah,1,4,5* Barbara Podobnik,2 Metka Novak,1,5 Branka Korošec,1 Sabina Berne,3 Matjaž Vogelsang,1 Nada Kraševec,1 Neja Zupanec,1 Jure Stojan,3 Joerg Bohlmann5 and Radovan Komel1,3 1 National Institute of Chemistry, Hajdrihova 19, SI-1000 Ljubljana, Slovenia. 2 Lek Pharmaceuticals d.d., Verovškova 57, SI-1000 Ljubljana, Slovenia. 3 Institute of Biochemistry, Faculty of Medicine, University of Ljubljana, Vrazov trg 2, SI-1000 Ljubljana, Slovenia. 4 Department of Wood Science, Faculty of Forestry, University of British Columbia, 2424 Main Mall, Vancouver, BC V6T 1Z4, Canada. 5 Michael Smith Laboratories, University of British Columbia, Vancouver, BC V6T 1Z4, Canada.

Summary Cytochromes P450 (CYPs) catalyse diverse reactions and are key enzymes in fungal primary and secondary metabolism, and xenobiotic detoxification. CYP enzymatic properties and substrate specificity determine the reaction outcome. However, CYP-mediated reactions may also be influenced by their redox partners. Filamentous fungi with numerous CYPs often possess multiple microsomal redox partners, cytochrome P450 reductases (CPRs). In the plant pathogenic ascomycete Cochliobolus lunatus we recently identified two CPR paralogues, CPR1 and CPR2. Our objective was to functionally characterize two endogenous fungal cytochrome P450 systems and elucidate the putative physiological roles of CPR1 and CPR2. We reconstituted both CPRs with CYP53A15, or benzoate 4-hydroxylase from C. lunatus, which is crucial in the detoxification of phenolic plant defence compounds. Biochemical characterization using RP-HPLC shows that both redox partners support CYP activity, but with different product specificities. When reconstituted with CPR1, CYP53A15 converts benzoic acid to Accepted 6 July, 2011. *For correspondence. E-mail [email protected]; Te. (+386) 14760261; Fax (+386) 14760300.

© 2011 Blackwell Publishing Ltd

4-hydroxybenzoic acid, and 3-methoxybenzoic acid to 3-hydroxybenzoic acid. However, when the redox partner is CPR2, both substrates are converted to 3,4-dihydroxybenzoic acid. Deletion mutants and gene expression in mycelia grown on media with inhibitors indicate that CPR1 is important in primary metabolism, whereas CPR2 plays a role in xenobiotic detoxification.

Introduction Cytochromes P450 (CYPs) belong to a large superfamily of genes and enzymes that is present in all kingdoms of life. They appear to be most numerous and diverse in the ˇ rešnar genomes of plants and fungi (Nelson et al., 2008; C and Petricˇ, 2011). Some fungal genomes have over 150 CYPs, representing over 1% of all genes (Doddapaneni et al., 2005). These enzymes are crucial in primary and secondary metabolism pathways. They are often part of fungal biosynthetic gene clusters that process natural products like mycotoxins fumonisin and sterigmatocystin (Bräse et al., 2009). As well, CYPs like pisatin demethylase and benzoate para-hydroxylase act in pathways that degrade xenobiotic compounds (Matthews and Van Etten, 1983). Benzoate para-hydroxylase is a member of the relatively well-characterized fungal CYP53 family (Fujii et al., 1997; Van Den Brink et al., 2000; Faber et al., 2001; Matsuzaki and Wariishi, 2005; Matsuzaki et al., 2008). Enzymes from this family produce phenolic derivatives that are then channelled into the b-ketoadipate pathway for aromatic compound degradation (Harwood and Parales, 1996). We have shown that CYP53 is especially important in plant pathogenic fungi, because it detoxifies phenolic plant defence compounds such as benzoic acid and isoeugenol (Podobnik et al., 2008). Cytochromes P450 are haem-containing monooxygenases that catalyse a wide variety of reactions; e.g. hydroxylation, epoxidation, oxidation, reduction, deamination, dehalogenation, dealkylation, dehydrogenation and demethylation (e.g. Ortiz de Montellano and De Voss, 2005; Gillam and Hunter, 2007). They insert one atom of molecular oxygen into an organic substrate. To perform this reaction, they must activate molecular oxygen that

Fungal P450 systems in xenobiotic detoxification 1375

requires the sequential delivery of electrons to the haem cofactor in the CYP active site. The source of electrons are coenzymes NADH or NAD(P)H. The mechanisms and composition of cytochrome P450 electron transfer chains through which the electrons are channelled are highly diversified (Hannemann et al., 2007). The complex redox chemistry and substrate specificity reside in the active site of this enzyme (Sevrioukova and Poulos, 2010); however, the outcome of CYP-mediated reactions also seems to be influenced by its redox partners. The redox partner of eukaryotic microsomal CYPs (or class II CYPs) is the membrane-bound, diflavin cytochrome P450 reductase (CPR), which contains cofactors flavin adenine dinucleotide and flavin mononucleotide (FMN) (e.g. Laursen et al., 2011). In the CPR deletion mutant of the pathogenic ascomycete Fusarium fujikuroi, activities of CYPs involved in gibberellin biosynthesis resulted in changes in regioselectivity, reaction rate and alternate product formation (Malonek et al., 2004). These differences were caused by the CYP interaction with alternative redox partners: cytochrome b5 and NADH cytochrome b5 reductase (Troncoso et al., 2008). The same two redox partners were shown to support CYP51 activity in Saccharomyces cerevisiae (Lamb et al., 1999). Recently, we identified two CPR paralogues, CPR1 and CPR2, in the ascomycete Cochliobolus lunatus, and showed that other filamentous fungi with many cytochromes P450 also possess multiple CPRs (Lah et al., 2008). Because the C. lunatus genome has not yet been sequenced, the genomic context of cpr1 and cpr2 in this species is not known. The genome of the closest sequenced relative, Cochliobolus heterostrophus, only has the cpr1 homologue. Two of 11 other sequenced Dothideomycete genomes also possess the cpr2 orthologue (Rhytidhysteron rufulum and Dothistroma septosporum) (http://genome.jgi-psf.org/programs/fungi/ index.jsf). In these ascomycete CPR paralogues, only about 30% of amino acids are identical, most of which are part of cofactor binding domains (Fig. S1). The amino acid identity is surprisingly low for an enzyme well conserved across species. This raises questions about the functional significance of CPR paralogues in the fungus in terms of their effect on CYP activity and the activity of a particular CYP-CPR system in fungal metabolism. To the best of our knowledge, no study has yet compared the role of multiple CPRs within a fungal species. Here we functionally characterized two reconstituted endogenous fungal cytochrome P450 systems, and elucidated the putative physiological roles for the two redox partners, CPR1 and CPR2. We recombinantly expressed both CPR1 and CPR2 in E. coli, and reconstituted each with CYP53A15, the benzoate 4-hydroxylase from C. lunatus. We biochemically characterized both systems in terms of substrate and product specificity with reverse phase chromatography

(RP-HPLC). Using deletion mutants for the three redox partners, we characterized growth inhibition on media supplemented with toxic phenolic compounds. As well, we monitored gene expression of the redox partners in mycelia treated with phenolics. Finally, we studied the function of the two C. lunatus P450 systems heterologously expressed in yeast. By integrating the results of our in vitro and in vivo analyses we inferred the physiological role of each CPR. Moreover, we further described the function of CYP53A15, and the function of the respective endogenous fungal cytochrome P450 systems in xenobiotic detoxification.

Results Expression and activity of CPRs and CYP53A15 We attempted to biochemically characterize the effect of two different CPRs on the functions of a complete cytochrome P450 system of C. lunatus. Under inducible promoters, we recombinantly expressed C-terminally Histagged genes cpr1, cpr2, and the bph gene encoding CYP53A15. We optimized the expression of cpr1 and cpr2 genes in the E. coli strain C43(DE3), which was reported to improve synthesis of membrane proteins and overcome other toxic effects associated with overexpression (Miroux and Walker, 1996). Optimal gene expression occurred when the culture was induced at OD600 0.5–0.7 at 25°C for CPR1 and 30°C for CPR2, and then grown for 24 h. The protein expression levels were 625 nmol l-1 for CPR1 and 125 nmol l-1 for CPR2, which is within the range of values reported for fungal CPRs (Warrilow et al., 2002; Park et al., 2010). After we purified both CPRs with Ni-NTA affinity chromatography followed by size exclusion chromatography, we verified the homogeneity of both proteins via SDS-PAGE. The 80 kDa bands corresponded to CPR1 and CPR2 (Fig. S2A). We used a standard cytochrome c reduction assay to test the NADH- and nicotinamide adenine dinucleotide phosphate (NADPH)-dependent activity of the E. coli membrane fraction containing either CPR1 or CPR2, and the purified enzymes. Both CPRs reduced cytochrome c when NADPH was added, but not when the primary electron donor was NADH. CPR1, whether in membrane or purified, was more active in comparison with CPR2 (Table S1). The membrane fractions of E. coli expressing the empty plasmid pET17b were used as negative control and showed substantially lower cytochrome c reducing activity. The activities of both CPRs were higher in the membrane-bound than in the purified form. We therefore selected the membrane-bound CPRs for reconstitution and further analyses of the cytochrome P450 system. Using the classical Michaelis–Menten equation, we determined the kinetic parameters of CPR1 and CPR2 for reducing ferricyanide in the presence of a fixed 100 mM

© 2011 Blackwell Publishing Ltd, Molecular Microbiology, 81, 1374–1389

1376 L. Lah et al. 䊏

Fig. 1. Initial rates of ferricyanide reduction by 0.1 mM CPR1 and 0.05 mM CPR2 at different ferricyanide concentrations in the presence of fixed 0.1 mM NADPH concentration. In (A) and (C), the classical Michaelis–Menten equation was used to analyse data for CPR1 and CPR2 respectively. In (B) and (D), the same data were evaluated by the equation that includes cooperative effects of ligand binding for CPR1 and CPR2 respectively.

NADPH concentration and pH 7.6. CPR2 had higher affinities for the substrate as revealed by the KM value, whereas the activities (kcat values) of both CPRs are comparable (Fig. 1A and C). As a result, the catalytic efficiency, or the specificity constant (kcat/KM), was about three times higher for CPR2 (750 000 M-1 s-1) than for CPR1 (210 000 M-1 s-1). As well, these values for the Michaelis– Menten mechanism show that ferricyanide is a poor substrate for both fungal CPRs. We have also analysed the data with the equation that includes cooperative effects (Fig. 1B and D). We show that the theoretical Michaelis– Menten curve generated by using Eqn 1 (see Experimental procedures) fits the data satisfactorily, but that the curve generated by Eqn 2 reproduces them still better. We expressed the full-length bph gene for CYP53A15. Optimal expression was reached when the OD600 was 1.5–2.0 at the time of induction, and the culture then grown for 24 h at 30°C. The expression level for CYP53A15 was 3300 nmol l-1, which is relatively high (e.g. Wu et al., 2009). We purified (Fig. S2B) and spectrally characterized the enzyme following described protocols (Podobnik et al., 2008). The CO difference spectra of the purified CYP53A15 with a characteristic maximum at 450 nm confirmed expression of a functional enzyme (Fig. S3).

CPR1 and CPR2 induce CYP53A15 product specificity We found that both CPRs support the activity of the purified CYP53A15 in vitro. The redox partners were CPR1 and CYP53A15 in Reconstitution System 1 (RS1), and CPR2 and CYP53A15 in Reconstitution System 2 (RS2). We investigated substrate specificity and conversion capability of either RS1 or RS2 with two compounds: benzoic acid and 3-methoxybenzoic acid, using RP-HPLC. Both compounds were substrates for the two reconstituted systems. In RS1, benzoic acid was converted to 4-hydroxybenzoic acid in a single hydroxylation step, while 3-methoxybenzoic acid was demethylated to 3-hydroxybenzoic acid (Fig. 2A and B, respectively). The rate of product formation was higher when the substrate was benzoic acid than when the substrate was 3-methoxybenzoic acid (Fig. 3). However, in RS2, benzoic acid and 3-methoxybenzoic acid were both converted to the same product, 3,4dihydroxybenzoic acid (Fig. 2C and D). Benzoic acid was converted through two hydroxylation steps. The conversion of 3-methoxybenzoic acid involved demethylation and hydroxylation reactions. For RS2, the rates of product formation were substantially lower than for RS1 (Fig. 3). We performed a fluorometric assay based on oxygen consumption (Olry et al., 2007) to confirm the results of



Fig. 2. HPLC analysis of Reconstitution System 1 (RS1; CPR1 + CYP53A15) and Reconstitution System 2 (RS2; CPR1 + CYP53A15) products. (A) RS1, substrate: BA, product: 4-OH BA. (B) RS1, substrate: 3-MeO BA, product: 3-OH BA. (C) RS2, substrate: BA, product: 3,4-diOH BA. (D) RS2, substrate: 3-MeO BA, product: 3,4-diOH BA. (E) Chromatogram of standards (BA – benzoic acid, 3-OH BA – 3-hydroxybenzoic acid, 4-OH BA – 4-hydroxybenzoic acid, 3,4-diOH BA – 3,4-dihydroxybenzoic acid, 2-MeO BA – 2-methoxybenzoic acid, 3-MeO BA – 3-methoxybenzoic acid, 4-MeO BA – 4-methoxybenzoic acid, 4-OH-3-MeO BA – 4-hydroxy-3-methoxybenzoic acid).

HPLC analyses of RS1 and RS2, and screen other putative CYP53A15 substrates. We observed an increase in fluorescence due to substrate conversion with benzoic acid, 3-methoxybenzoic acid, isoeugenol and cinnamic acid for both RS (not shown). 4-hydroxycoumarin and 4-methoxybenzoic acid were substrates in RS1, but not RS2.

Growth morphology and inhibition of C. lunatus strains under varying culture conditions We constructed Dcpr1, Dcpr2 and Dbph deletion mutants to support the results obtained with our in vitro analyses with physiologically relevant in vivo analyses. In a previous study we found that benzoic acid and other phenolic



3-methoxybenzoic acid. Of all three substances, 3methoxybenzoic acid least inhibited the growth of the Dbph strain. Expression of bph and cpr2 genes are significantly induced with benzoic acid, and 3-methoxybenzoic acid and isoeguenol, respectively

Fig. 3. Time course of product formation in reaction catalysed by RS1 and RS2. Benzoic acid (BA) and 3-methoxybenzoic acid (3-MeO BA) were used as substrates (both 100 mM). RS and substrate combinations are shown in the legend. The product formed is indicated next to line (3-OH BA – 3-hydroxybenzoic acid, 4-OH BA – 4-hydroxybenzoic acid, 3,4-diOH BA – 3,4-dihydroxybenzoic acid).

compounds were CYP53A15 substrates, which inhibited fungal growth at higher concentrations (Podobnik et al., 2008). We investigated the function of both CPRs and CYP53A15 in vivo through: (i) growth inhibition studies of different strains, (ii) studies of gene expression following different treatment conditions and (iii) recombinant expression of cytochrome P450 system components in yeast. We examined the growth and hyphal morphology on malt extract medium (Malt extract agar – Blakeslee’s Formula, MBF). Mycelial growth and macroscopic appearance of the wild type (wt), and the Dbph, and Dcpr2 mutants were comparable on media that were not supplemented with substrates. The Dcpr1 gene deletion was not lethal; however, the deletion severely affected growth and resulted in growth initiation being delayed for 5 days. The Dcpr1 phenotype was characterized by melanized swollen hyphae with thickened cell walls. Because of its severely affected phenotype, we did not use the Dcpr1 deletion mutant in growth inhibition experiments. We monitored growth inhibition of C. lunatus strains on non-supplemented control MBF, MBF with added ethanol, and MBF supplemented with substrates dissolved in ethanol (1 mM benzoic acid, 3-methoxybenzoic acid or isoeugenol). Mycelial growth on non-supplemented and ethanol-supplemented media was comparable, so we used the latter as control in further experiments. Benzoic acid caused severe growth delay of Dbph. The growth of wt and Dcpr2 strains was affected to the same lesser extent (Fig. 4). Isoeugenol similarly affected growth of the Dbph strain; however, it also markedly reduced the growth of wt and Dcpr2 strains. All three strains showed a comparatively similar, albeit lesser reduction in growth on

We used quantitative PCR (QPCR) to assess transcriptional profiles of cpr1 and cpr2 genes, as well as the bph gene encoding CYP53A15 following different treatment conditions in the C. lunatus wild type (wt), and Dcpr1 and Dcpr2 deletion mutants. We monitored gene expression at four time points: 15, 45, 90 and 200 min. The results were analysed using REST software (Pfaffl et al., 2002). Overall, the temporal expression patterns of each of the three genes varied following treatment with 1 mM benzoic acid, 3-methoxybenzoic acid or isoeugenol, and were distinct in the three strains. In the wt strain, bph gene expression was highly induced by benzoic acid with over 260fold increase at 200 min (Fig. 5). In the Dcpr1 and Dcpr2 mutants, bph gene expression was induced 9- and 27-fold at 45 min and not appreciably at later times. When treated with 3-methoxybenzoic acid or isoeugenol, we observed increased expression of cpr2 in the wt strain at 90 min (Fig. 6). Notably, expression ratios of cpr2 (11.0 and 11.4, respectively) were higher than for cpr1 (1.6 and 1.7, respectively). The cpr1 gene is not induced by any of these compounds, if we consider a cut-off value

Fig. 4. Growth inhibition of C. lunatus wt, Dbph and Dcpr2 strains on media supplemented with benzoic acid, 3-methoxybenzoic acid or isoeugenol (all 1 mM), and growth on control MBF with ethanol (EtOH). The radial growth rate was measured daily for 1 week, or until the surface of the plate was homogeneously overgrown.



The yeast S. cerevisiae has a single CPR gene, NCP1. While deletion of NCP1 is not lethal, the mutant strain

grows poorly and has lowered ergosterol membrane content (Venkateswarlu et al., 1998; Tiedje et al., 2007). Both phenotypes are associated with the indirect role of the yeast CPR protein (Ncp1) in ergosterol biosynthesis. We produced the ncp1D deletion mutant for comparative gene complementation studies where we attempted to restore the function of the deleted gene with either CPR1 or CPR2 from C. lunatus. To further elucidate the possible in vivo role of CYP53A15, and both reconstitution systems, we expressed this CYP with C. lunatus redox partners in S. cerevisiae. All strains were grown on control media with dimethyl formamide (DMF) and media supplemented with 50 mM benzoic acid. This concentration was selected based on preliminary experiments where higher concentrations would inhibit growth to the extent of hindering comparative analyses. We used the wild-type S. cerevisiae as control. An additional positive control was the knock-out strain complemented with the native yeast CPR (Ncp1). Membrane ergosterol content of all control and transformed strains was measured using RP-HPLC. Expression of recombinant proteins was verified using SDS-PAGE (Fig. S4A and B), and Western blot analysis to enhance CPR1 visibility (Fig. S4C). The growth of strains and transformants was inhibited on benzoic acid-supplemented medium (Fig. 7). The inhibition, however, appeared to be general and not lower in particular strains complemented with either redox partner or entire reconstitution systems when compared with the control media. The reduced growth of the ncp1D deletion mutant was restored with yeast Ncp1, as well as with C. lunatus CPR2 and RS2. Complementation with CPR1

Fig. 6. Expression ratios of bph, cpr1 and cpr2 genes in the C. lunatus wild-type strain 90 min after treatment with 1 mM benzoic acid, isoeugenol or 3-methoxybenzoic acid, normalized to the reference gene (18S rRNA). Boxes represent the inter-quartile range. The horizontal line in the box represents median gene expression. Whiskers represent the minimum and maximum observations. Asterisks above boxes represent statistically significant results as calculated by the randomization test implemented in REST software.

Fig. 7. Complementation of ncp1D S. cerevisiae strains with components of RS. The yeast CPR deletion strain (ncp1D) was complemented with the S. cerevisiae native reductase Ncp1, CPR1, CPR2, CYP53A15, RS1 (CPR1 + CYP53A15) and RS2 (CPR2 + CYP53A15) and grown on medium containing 50 mM benzoic acid dissolved in DMF for 5 days. The wild-type strain was used as control. Undiluted yeast cells in the first row are followed by 10 ¥ serial dilutions.

Fig. 5. Temporal pattern of bph gene expression ratios in the C. lunatus wt, and Dcpr1 and Dcpr2 deletion strains following treatment with 1 mM benzoic acid, normalized to the expression of the reference gene (18S rRNA). Gene expression was measured at 15, 45, 90 and 200 min. All values are significant as calculated by the randomization test implemented in REST software.

of 2.0. In this same assay, bph gene expression was not significantly induced by either of these two substrates. Using the comparative CT method (Schmittgen and Livak, 2008), we have calculated that the fold expression of cpr2 is 10-2 to 10-3 relative to cpr1 in non-induced mycelia at the time points included in our study (Table S2). Comparison of both P450 reconstitution systems in S. cerevisiae



Fig. 8. Ergosterol levels in membrane fractions of yeast strains and transformants. Relative amounts of ergosterol were analysed in yeast membrane fractions grown to the stationary phase and given as percentage of dry weight of biomass. Yeast CPR deletion strain (ncp1D) was complemented with S. cerevisiae native reductase Ncp1, CPR1, CPR2, CYP53A15, RS1 (CPR1 + CYP53A15) and RS2 (CPR2 + CYP53A15); wt was used as control. Ergosterol was quantified by RP-HPLC as described in Experimental procedures. All values are means from two independent experiments with mean deviations as indicated by the error bars.

or CYP53A15 only slightly improved growth of the ncp1D deletion mutant, while the growth of RS1-transformed strains was even more affected than ncp1D. Ergosterol membrane contents of the ncp1D strain and the CPR1transformant were comparable and about one-third of that of the wt (Fig. 8). CPR2 and RS2 suppressed the ncp1D ergosterol phenotype. Relatively high levels of ergosterol, comparable with the wt, were measured in the CYP53A15- and RS1-transformed strains, which grew poorly on control and benzoic acid-supplemented media. It appears that the role of the transformed CYP53A15 in yeast differs from its role in C. lunatus, in terms of substrate utilization and the putative metabolic pathways context. We therefore tested a number of compounds involved in pathways of phenylalanine metabolism, specifically phenylalanine, phenylpyruvate, phenylacetaldehyde, phenylacetate, phenylethanol and benzyl alcohol. Of these compounds, only phenylacetaldehyde proved to be a substrate for CYP53A15 reconstituted with CPR1 as the redox partner when tested with the fluorometric assay of CYP activity based on oxygen consumption (Fig. S5).

Discussion CPR determines CYP53A15 product specificity In the work presented here, we show that the product formed by a given CYP from a specific substrate can vary with the nature of the CPR of the microsomal P450 system that participates in the reaction. When coupled

with CPR1, CYP53A15 converts benzoic acid to the expected 4-hydroxybenzoic acid in a single hydroxylation reaction. Similarly, 3-methoxybenzoic acid is demethylated in a single step to 3-hydroxybenzoic acid (Faber et al., 2001; Podobnik et al., 2008). However, when coupled with CPR2, CYP53A15 converts both of these substrates into the same product in a two-step reaction. This product, 3,4-dihydroxybenzoic acid, or protocatechuate, is the phenolic derivative that enters the b-ketoadipate pathway, which is the central pathway for aromatic compound degradation – and benzoic acid detoxification (Harwood and Parales, 1996). This result is of critical importance for biological systems that express more than one functional CPR gene, as it diversifies the metabolic capacity of modular P450 systems. The step in the pathway that converts 4-hydroxybenzoic acid to protocatechuate is catalysed by 4-hydroxybenzoate-3-monooxygenase, an enzyme that has been extensively studied in several bacteria (Ballou et al., 2005; Palfey and McDonald, 2010). This enzyme is not a cytochrome P450, but belongs to a family of flavin-dependent monooxygenases, which, like CYPs, insert a single atom of molecular oxygen into the substrate and catalyse oxygenation reactions like hydroxylation and epoxidation (Van Berkel et al., 2006). Flavin-dependent monooxygenases use a purely organic cofactor for oxygenation reactions and not a transition metal like the haem in cytochromes P450. These enzymes are relatively abundant in prokaryotes. However, reports on flavin-dependent monooxygenases from fungi are scarce (Kalin et al., 1992; Eppink et al., 2000; Torres Pazmino et al., 2010). Early work on Aspergillus niger identified 3-hydroxybenzoate-4hydroxylase that hydroxylated 3-hydroxybenzoic acid to protocatechuate, but did not convert 4-hydroxybenzoic acid (Premkumar et al., 1969). The gene encoding this enzyme has not been identified. Therefore, in addition to a hypothetical flavin-dependent monooxygenase catalysing selected steps leading to the formation of protocatechuate, CYP53A15 and the possible combinations of redox partners may be involved in more than one reaction in the pathway of aromatic compound degradation. Electrostatic binding interactions between basic residues of the cytochrome P450 and clusters of negatively charged acidic residues of CPR are important in the formation and stabilization of P450 redox partner complexes (Paine et al., 2005). Recently, crystallographic and NMR data suggested that CPR exists in equilibrium between physiologically relevant open and closed conformations that differ in the orientation of the FMN-binding domain (Aigrain et al., 2009; Ellis et al., 2009; Hamdane et al., 2009). Electron transfer to the cytochrome P450, or other acceptor, is possible when the reductase is in the open conformation and exposes a new interface on the surface of the FMN-binding domain. Evidence for cooperative



Fig. 9. The model of interaction of cytochrome P450 reductase (CPR; yellow) and CYP53A15 (black). The FMN-binding domain of CPR in open conformation interacts with the CYP53A15 that brings the FMN cofactor in close proximity to the CYP haem cofactor and facilitates electron transfer. (NADPH, nicotinamide adenine dinucleotide phosphate; FAD, flavin adenine dinucleotide; FMN, flavin mononucleotide).

substrate binding that we calculated from the rates of ferricyanide reduction agrees with these extensive conformational changes during the CPR catalytic cycle. Multiple redox partner docking sites were identified on the surface of the FMN-binding domain of CPR in helices B, C and F (Shen and Kasper, 1995; Ellis et al., 2009; Jang et al., 2010). Sequence alignment of mammalian and both C. lunatus CPRs reveals that acidic residues involved in redox partner interaction are fairly conserved between human CPR and CPR1 (Fig. S6). Notably, in CPR2 we find several instances where acidic residues are replaced by polar basic (Arg), non-polar basic (Ser) and non-polar hydrophobic (Ala) amino acids. Elimination of these negative charges can alter the mode of interaction between CPR and electron acceptors (Jang et al., 2010), which could also be another explanation for the lower activity of RS2. Product specificity of the two CPRs can partially be explained by visualizing the 3D complex between CPR1 and CYP53A15 (Fig. 9). Although all the structures were modelled, they help explain the nature of interactions between the two proteins. Molecular dynamics simulations of CYP53A15 show that although the enzyme’s haem group is well anchored, it is highly flexible within the active site. The root mean square of the haem movements

during the interval of 2.5 ns fluctuates between 0.25 and 0.8 A from its starting position (Fig. S7). It is therefore very likely that the FMN-binding domain of each CPR specifically affects these movements in complex with the CYP, and consequently the activity of each system. We investigated the interaction between CPR1 and CYP53A15 by subjecting the two facing structures to the RosettaDock server that predicted several protein complexes (http:// rosettadock.graylab.jhu.edu/). From the predicted lowenergy docking structures, we chose the one where the shortest distance between the FMN of CPR1 and the haem of CYP53A15. In the final 2 ns of the CPT-EWALD dynamic simulation, the distance between FMN and the haem iron fluctuated between 8.0 and 10.2 A. Consistent with the above, Sevrioukova and Poulos (2010) reported that close contact between P450cam and its redox partner putidaredoxin (Pdx) upon binding enables active site restructuring via haem displacement that couples electron transfer to substrate hydroxylation. Given that the FMN-binding domains of CPR1 and CPR2, share only 32% amino acid identity, it is reasonable to hypothesize that these two domains will modulate the haem group’s movements differently when CYP53A15 is complexed with either CPR. Such interactions could result in different product specificities, as the same substrate



would assume a slightly altered position in the active site permitting for example, hydroxylation reactions at particular sites of the molecule. The two CPRs do not greatly affect substrate specificity of CYP53A15, as the enzyme’s activity was successfully reconstituted with both redox partners, albeit with different rates of product formation from the same substrates. CYP53A15 reconstituted with CPR1 converted benzoic acid more readily than 3-methoxybenzoic acid. When reconstituted with CPR2, conversion of 3-methoxybenzoic acid was faster than that of benzoic acid. The rates of product formation from both substrates were faster with RS1 compared with RS2. CPR1 and CPR2 putatively affect the specificity of CYP53A15’s detoxification activity In addition to the differential role endogenous CPRs of C. lunatus play in CYP53A15 product specificity, their physiological function differs as well. Our in vivo experiments that include deletion mutant studies and gene expression data support the putative function of CPR1 in primary metabolism. Combined with the results of in vitro experiments, we suspect that this reductase is the physiological partner of CYP53A15 when the substrate is benzoic acid. The role of CPR2 and its putative function in specialized metabolism is more elusive and seems to be utilized in the metabolism of specific xenobiotic compounds. While the deletion of cpr1 is not lethal, the growth of the mutant is severely delayed. This phenotype indicates an important role of CPR1 in fungal primary metabolism, specifically as the electron donor to three enzymes in the evolutionary conserved pathway of ergosterol biosynthesis, sterol 22-desaturase (CYP61) and sterol 14ademethylase (CYP51) and squalene epoxidase (e.g. Tiedje et al., 2007). In fungi, CYP51 is essential and the central drug target of azole-based inhibitors (Kelly et al., 2001). Increased sensitivity to ketoconazole has been observed in the ncp1D S. cerevisiae deletion strain (Sutter and Loper, 1989). Unexpectedly, in the Dcpr1 mutant, CPR2 apparently does not complement the deletion of its paralogue to function as the alternative redox partner. Similar phenotypes have been observed when the homologue of cpr1 was deleted in the fungi Botrytis cinerea and Giberella fujikuroi (Malonek et al., 2004; Siewers et al., 2004). In the latter case, the alternative redox partners were identified to be cytochrome b5 (CYB5) and cytochrome b5 reductase (CBR) (Troncoso et al., 2008). As we have observed, the deletion of the native CPR gene in S. cerevisiae, NCP1, causes poor growth. The double deletion of NCP1 and the gene encoding CBR, CBR1, is lethal (Tiedje et al., 2007). It has also been shown that

CYB5 and CBR support CYP51 activity in vitro (Lamb et al., 1999). We therefore conclude that in C. lunatus CPR1 delivers electrons to enzymes in the ergosterol biosynthesis pathway. In the mutant, its role is probably replaced by CYB5 and CBR, but not by CPR2. Gene expression data show that cpr2 was relatively highly induced by isoeugenol and 3-methoxybenzoic acid, whereas expression of cpr1 was not induced by either of these substrates. Surprisingly, as it does not agree with literature data, expression of neither reductase was induced by benzoic acid (Van Den Brink et al., 2000; Matsuzaki and Wariishi, 2005). Comparison of gene expression of cpr2 relative to cpr1 shows that the fold expression of cpr2 is 10-2 to 10-3 relative to cpr1. The cpr1 gene therefore appears to be constitutively expressed (less inducible) whereas the expression of its more inducible paralogue in non-treated mycelia is lower. The severe growth inhibition of the Dbph mutant on benzoic acid and the high expression of the coding gene (over 260-fold) indicate that benzoic acid is the main substrate of CYP53. The preferred redox partner in benzoic acid elimination seems to be CPR1, as RS1 has the highest rates of product formation. As well, the benzoic acid-mediated growth inhibition of the Dcpr2 mutant is similar to the wild-type strain that suggests that CPR1, present in both strains, delivers electrons to CYP53A15. Growth of the wild type is most affected on media supplemented with isoeugenol, and to a lesser extent by 3-methoxybenzoic acid. The inhibition on these two supplemented media is similar in the Dcpr2 and Dbph mutants. As noted above, isoeugenol and 3methoxybenzoic acid are also the substrates that highly induce cpr2, and do not induce cpr1 gene expression. CPR2 may be the particular CYP53A15 redox partner when substrates are other phenolic compounds particularly those with a 3-methoxy substituent on the aromatic ring. Both RS1 and RS2 convert these phenolic compounds in vitro; however, it appears that these CYP systems (RS1 and RS2 in the wt and RS1 in the Dcpr2 mutant) are not as efficient in eliminating these two inhibitory compounds in vivo. As well, either the product of the reaction, or products of its downstream processing could result in the formation of a toxic intermediate. In the case of 3-methoxybenzoic acid as substrate, the reaction product is protocatechuate. Accumulation of this compound, or its di-oxygenated derivative carboxymuconate was shown to be toxic in bacteria (Parke et al., 2000). When isoeugenol is the substrate, the product of the RS2 is not known. To validate hypotheses of the toxicity of particular intermediates, we would need a better understanding of the sequential enzymatic steps in the pathway of aromatic compound degradation in C. lunatus. The putatively deleterious function of RS2 could also explain the increased synergistic



inhibitory effects of benzoic acid with phenolic compounds on the growth of C. lunatus reported previously (Podobnik et al., 2008). If benzoic acid is efficiently eliminated by RS1, the formation of RS2 would lower the amount of available CYP53A15 and perhaps simultaneously produce a toxic intermediate. While we have identified CPR1 as the preferred redox partner of CYP51 and CYP53A15, the CYPs that prefer CPR2 have yet to be identified. An indication could be the genomic context of these genes in genomes of species of the same taxonomic group. In two Dothideomycete species that possess the cpr2 homologue, a CYP gene of unknown function is in close proximity to the cpr2 gene. In Aspergillus nidulans, this gene belongs to the CYP630 family (Kelly et al., 2009). The cpr1 gene is present in all (12) analysed Dothideomycete genomes and has no neighbouring CYP genes. CYP53A15 has multiple functions in primary and secondary metabolism CYP53 enzymes in fungi use benzoic acid as the substrate to produce 4-hydroxybenzoic acid and are essential in detoxifying aromatic compounds (Fukuda et al., 1996; Faber et al., 2001; Fraser et al., 2002; Matsuzaki and Wariishi, 2005; Podobnik et al., 2008). CYP53s can have other substrates, and these can vary between fungal species. For example, 3-methoxybenzoic is demethylated to 3-hydroxybenzoic acid in C. lunatus and A. niger, but not in Phanerochaete chrysosporium (Faber et al., 2001; Matsuzaki and Wariishi, 2005). Although reports on substrate specificity can vary due to differences in experimental set-up and substrates tested, substrate structural requirements are generally rather strict. Mono-substitutions on the phenyl ring are allowed only at certain positions with specific groups, while the carboxyl group positions the substrate into the active site so that a spectral shift is induced which helps to identify substrates spectrophotometrically (Faber et al., 2001; Podobnik et al., 2008). With our flourometric assay, we identified additional compounds without the carboxyl group, like phenylacetaldehyde and cinnamic acid, as CYP53A15 substrates. A broader than expected spectrum of possible substrates in turn extends the putative physiological roles of CYP53A15. To study the CYP system in vivo, we expressed the redox partners, as well as the entire monooxygenase system in the S. cerevisiae with its native CPR gene (NCP1) knocked out. We followed growth on control and benzoic acid-supplemented media. As well, we have analysed the ergosterol content of transformants. As noted above, the ncp1D strain grows poorly. The wild-type growth is recovered after complementation with the native reductase, or C. lunatus CPR2 or RS2. Perhaps CPR2 better complements the deletion because of putatively

closer evolutionary relationships between the CPR2 family of Pezizomycotina and the CPR family of ascomycetous yeasts (Saccharomycotina) (Lah et al., 2008). In contrast, the complementation with CPR1 did not restore the wild-type growth. In these transformants, the measured levels of membrane ergosterol content agree with the observed phenotype, where poor growth is associated with lower ergosterol levels caused by defects in its biosynthetic pathway (Venkateswarlu et al., 1998). Complementation of the ncp1D strain with CYP53A15, or RS1 from C. lunatus resulted in no improvement of growth and almost no growth respectively. However, the ergosterol levels in these transformants with poor growth on solid media or liquid culture were relatively high. It has been observed that although ergosterol levels in the ncp1D mutant are about 25% of those in the wt, this was still the predominant sterol found in membranes (Venkateswarlu et al., 1998). Despite the fact that a strong correlation exists between ergosterol content and fungal dry mass, the amount of ergosterol in fungal tissue is not constant (Parsi and Gorecki, 2006; Silva et al., 2010; and references therein). It depends among others on the fungal species, developmental stage and growth conditions (growth media, pH and temperature). We have shown in vitro that phenylacetaldehyde, an intermediate of phenylalanine metabolism, can be a substrate of CYP53A15. As well, it has been reported that bph gene expression was induced by L-phenylalanine (Fujii et al., 1997). Based on our results and relevant literature reports, it is therefore possible that CYP53A15 (when not complemented with CPR2) assumes a function in yeast metabolism that does not impede ergosterol biosynthesis but still retards growth. CYP53A15 putatively associates with the alternative redox partners, CYB5 and CBR, as has been observed in fungi, insects and plants (e.g. Brankova et al., 2007; Murataliev et al., 2008; Troncoso et al., 2008). In double transformants, RS1 and RS2, it may associate with CPR1 and CPR2, respectively, to mediate different reaction outcomes. These outcomes, in turn, may cause the observed differences in phenotypes of transformed strains.

Experimental procedures Strains and growth conditions The filamentous fungus C. lunatus (m118) originated from the strain collection of the Friedrich Schiller University of Jena, Germany. Currently, it can be obtained from Belgian coordinated collections of micro-organisms (BCCM/MUCL) as strain MUCL 38696. It was cultured at 28°C in malt extract medium (MEM, Pivovarna Union, Slovenia) as described previously (Plemenitaš et al., 1988). Fungal mycelia, harvested by filtration, were frozen in liquid nitrogen, powder-ground and stored at -80°C for further applications.



For QPCR experiments, fungal mycelia were treated with the following substrates: benzoic acid, 3-methoxybenzoic acid and isoeugenol (all chemicals from Sigma-Aldrich, Germany). Briefly, 800 mg (wet weight) of mycelia was suspended in 10 ml phosphate buffer saline (PBS) (0.75 mM Na3PO4, 0.21 mM EDTA, 0.04 mM reduced glutathione; pH 5. 5). Substrates, dissolved in DMF, were added at 1 mM final concentration. As control, DMF alone was added to PBS. Mycelia were cultured on a rotary shaker at 110 r.p.m. and 28°C for 15, 45, 90 and 200 min. The yeast S. cerevisiae strains used in this study are described in Table S3. Yeast strains were routinely cultivated at 30°C on standard rich medium (YPD) (yeast extract/ peptone/dextrose) and on synthetic drop-out medium containing 2% glucose (SD) or 2% galactose (SG) (YNB: 0.67% w/v yeast nitrogen base, 2% w/v dextrose or galactose) without uracil, leucin, or both for selective pressure to maintain plasmid stability. Media were solidified with 2% w/v bacto-agar. S. cerevisiae cells were transformed with selected plasmids by a one-step method (Chen et al., 1992). For yeast plasmid propagation, the E. coli DH5a strain was used as bacterial host and grown in Luria–Bertani medium supplemented with 100 mg ml-1 ampicilin. C. lunatus genes were expressed in the E. coli C43(DE3) strain according to the method of Miroux and Walker (1996).

Nucleic acid manipulation The full-length nucleotide sequence of CYP53A15 (EU597483) was retrieved from the C. lunatus genomic library as described previously (Podobnik et al., 2008). Fulllength nucleotide sequences of CPR1 (EU111681) and CPR2 (EU111680) were obtained from C. lunatus DNA as described (Lah et al., 2008). Total RNA was isolated from frozen fungal mycelia with SV Total RNA Isolation System (Promega, USA) according to manufacturer’s instructions. RNA quality and quantity were assessed on the Bioanalyzer 2100 (Agilent Technologies, USA).

Construction of Dcpr1, Dcpr2 and Dbph deletion mutants in C. lunatus The bph deletion cassette with hygromycin resistance (hph) was constructed and transformed into C. lunatus protoplasts as described in Podobnik et al. (2008). Construction of cpr1 and cpr2 deletion cassettes followed the same protocol. C. lunatus deletion constructs are shown in Fig. S8. Primers and plasmids used are listed in Table S4. Transformants were selected on minimal medium agar plates with 1.2 M sorbitol and 100 mg ml-1 hygromycin (InvivoGen, France). Deletions of bph, cpr1 and cpr2 were confirmed with Southern blot and PCR.

Disruption of the NCP1 gene in yeast Disruption of the NCP1 gene encoding native yeast CPR was achieved using a short flanking homology method (Wach et al., 1998). The disruption cassette with the kanMX marker and flanking homology regions to target the NCP1 gene was

prepared by PCR with NCP1-kanMX-S1 and NCP1kanMX-S2 primers (Table S3). Plasmid pFA6a-KanMX4 was used as template. The PCR products were purified and transformed into yeast according to the method of Gietz and Woods (2002). Single colony yeast transformants were picked from YPD plates containing 200 mg l-1 geneticin (Sigma-Aldrich) after 3 days incubation at 30°C. Successful replacement of the NCP1 gene with kanMX cassette in geneticin-resistant transformants was verified by colony PCR (Ling et al., 1995).

Expression of cpr1, cpr2 and bph genes Expression of cpr1, cpr2 and bph genes in E. coli. The fulllength bph gene construct encoding the C-terminally His6-tagged CYP53A15 protein, was cloned into the pCWori+ plasmid and transformed into E. coli C43(DE3) cells. Bacteria were cultured in TB medium containing trace elements (listed in Table S5), 1 mM vitamin B1 (thiamin), and 100 mg ml-1 ampicillin at 37°C, with vigorous shaking until OD600 of 2.0 was reached. IPTG (1 mM final concentration) and daminolevulinic acid (1 mM final concentration) were then added to induce protein expression. Cells were grown for an additional 24 h at 30°C. For expression of C-terminally His4-tagged cpr1 and cpr2 genes, synthetic genes were ordered from GenScript (USA), optimized for expression in E. coli (sequence information available upon request). Bacterial cells were transformed with plasmid pET17b carrying cpr1 or cpr2. Single recombinant colonies were picked from LBA plates and inoculated into TB media supplemented with trace elements (see above). Bacteria were grown at 37°C and 170 r.p.m. until OD600 of 0.5–0.7 was reached. After addition of IPTG (1 mM) and riboflavin (10 mg ml-1), protein expression was induced for 24 h. CPR1 expression was achieved at 25°C and 160 r. p.m., while CPR2 was produced at 30°C and 160 r.p.m. Protein expression was confirmed by SDS-PAGE analysis of non-induced and induced bacterial culture samples. Expression of cpr1, cpr2 and bph genes in S. cerevisiae. The YepGAL1-CYP53A15 construct was prepared by yeast co-transformation of the bph gene amplified from pCW1-Wt vector with a C-terminal His4-tag, and BamHI-linearized YepGAL1-GFP1 target vector (kindly provided by Dr Mateja Novak Štagoj). Successful cloning was scored in Leu+ colonies. Cochliobolus lunatus cpr1 and cpr2 genes were amplified from cDNA and cloned into yeast expression vectors by in vivo recombination as described by Oldenburg et al. (1997). In brief, flanking regions for homologous recombination into the pYES2 target plasmid were added by PCR using 60-mer oligonucleotide primers containing a C-terminal His6-tag. Each PCR product was co-transformed in yeast together with XbaI/HindIII digested target plasmid (Gietz and Woods, 2002). Single colony Ura+ recombinats were picked from SD media lacking uracil (SD Ura-), inoculated in SD Ura- medium and grown overnight at 30°C and 170 r.p.m. Cells were then pelleted by centrifugation (13 400 g, 15 s), washed with sterile water and resuspended in SG Ura- medium, followed by 16 h incubation at 30°C and 170 r.p.m. until OD600 of 0.4–1 was reached.



To verify correct gene insertion, plasmid DNA was rescued from yeast colonies as described (Hoffman and Winston, 1987) and sequenced at Eurofins MWG Operon (Germany). Transformed strains were verified with PCR. Primer combinations are listed in Table S3. To monitor levels of expressed recombinant proteins in transformed strains with SDS-PAGE analysis and Western blot, yeast cells were harvested by centrifugation (1500 g, 5 min, 4°C), washed with sterile water, pelleted and stored at -80°C.

Purification of recombinant CPR1, CPR2 and CYP53A15 proteins Isolation of spheroplasts and membranes from E. coli. Bacterial cells were harvested by centrifugation (6000 g, 15 min) and resuspended in 50 mM K-phosphate (pH 7.4)/0.5 M NaCl/20% glycerol/0.1 mM DTT/0.1 mM PMSF. To obtain spheroplasts, 0.3 mg ml-1 lysozyme was added, and the bacterial suspension was gently shaken and incubated on ice for 45 min. Spheroplasts were recovered by centrifugation (5000 g, 15 min, 4°C) and resuspended in icecold 50 mM K-phosphate (pH 7.4)/20% glycerol/0.1 mM DTT/0.1 mM PMSF/250 U ml-1 benzonase. After homogenization and sonication (both carried out on ice), the lysate was centrifuged (12 000 g, 45 min, 4°C). The supernatant was then ultracentrifuged at 100 000 g and 4°C for 1 h. The pelleted membranes were resuspended in 50 mM K-phosphate (pH 7.4)/0.1 mM DTT/0.1 mM PMSF, aliquoted and stored at -80°C. Membrane solubilization and Ni-NTA affinity chromatography. Membranes were diluted (1:5, v/v) in 20 mM Na-phosphate (pH 7.4)/0.5 M NaCl/20% glycerol/0.2 mM DTT/0.1 mM PMSF/1 ¥ Complete, EDTA-free Protease Inhibitor Cocktail (Roche, Switzerland)/10 mM imidazole and solubilized with 2% Triton X-100 at 4°C for 1 h. The solution was ultracentrifuged (100 000 g, 45 min, 4°C) to remove insoluble material. The supernatant containing His-tagged proteins was purified by affinity chromatography on Ni2+ Sepharose High Performance (GE Healthcare, USA) resin. Bound proteins were eluted with 20 mM Na-phosphate (pH 7.0)/1.5 M NaCl/20% glycerol/500 mM imidazole/0.2 mM DTT/1¥ Complete EDTAfree Protease Inhibitor Cocktail (Roche). Fractions containing desired proteins were pooled and dialysed against 20 mM Na-phosphate (pH 7.4)/0.5 M NaCl/20% glycerol/0.1 mM EDTA. Samples were aliquoted and stored at -80°C. Purity was assessed by SDS-PAGE. Total protein was measured by the Bradford method (Bradford, 1976).

Measurement of CYP53A15 and CPR activity The quantity of active CYP53A15 was determined by CO-reduced difference spectra (Omura and Sato, 1964). Briefly, the sample was divided into a reference and sample quartz cuvette and gassed with CO for 60 s. Baseline from 350–500 nm was recorded on Agilent 8453 spectrophotometer at 25°C. A few grains of sodium dithionite were then added to the sample cuvette, while the same amount of buffer (20 mM Na-phosphate (pH 7.4)/0.5 M NaCl/20% glycerol/0.1 mM EDTA) was added to the reference cuvette.

CO difference spectra were measured several times within 15 min. CYP53A15 concentration was calculated using the equation based on the Beer-Lambert law: C = A/(∈L), where A is the difference in absorbance between oxidized and reduced CYP, L is the pathway length (1 cm), and ∈ is the extinction coefficient (185 mM–1 cm–1). The activity of membrane-bound and purified CPRs was assayed using Cytochrome c Reductase (NADPH) Assay Kit (Sigma-Aldrich). In the presence of NADPH, the reduction of cytochrome c by CPR changes the absorption spectrum of cytochrome c and an increase in absorbance at 550 nm is observed. One enzyme unit in this assay is defined as amount of enzyme that reduces 1.0 mmol of oxidized cytochrome c in the presence of 100 mM NADPH per minute at pH 7.8 and 25°C. CPR activity was measured spectrophotometrically for 3 min at 25°C and 550 nm. Cytochrome c reductase from rabbit liver (included in kit) was used as positive control. Membrane fractions of E. coli expressing the empty plasmid (pET17b) were used as negative control. Samples incubated without NADPH were used as blanks. Enzyme activity was calculated from:

ΔA550 min × D × 1.1 U = ml 21.1×Ve

and

A550 = ΔAsample − ΔAblank

where: D – dilution factor of enzyme sample, Ve – sample enzyme volume (ml), 21.1 – extinction coefficient (mM-1 cm-1) for reduced cytochrome c, and 1.1 – reaction volume (ml). Cytochrome c reducing activity of CPR1 and CPR2 was also assayed with 100 mM NADH as the primary electron donor.

Kinetic measurements To test the catalytic power of the two CPRs we followed the reduction of ferricyanide in the presence of saturating 100 mM NADPH on a UV-2450 Shimadzu spectrofotometer at 420 nm. Spectrophotometric assays were performed at room temperature using standard quartz cuvettes in a total reaction volume of 1 ml. All chemicals were purchased from Sigma-Aldrich. The extinction coefficient of 3360 mol-1 cm-1 was determined from known ferricyanide concentrations. From measured progress curves we determined initial rates at ferricyanide concentrations between 7.5 mM and 0.1 mM using 0.1 mM CPR1, and 0.05 mM CPR2 in 100 mM potassium phosphate (pH 7.6). We evaluated the obtained initial rate data using the classical Michaelis–Menten equation, and modified it to determine putative cooperativity:

v=

k cat ⋅ [E ]o ⋅ [S ] K M + [S ]

k cat ⋅ [E ]o ⋅ [S ] n K M + [S ]

n

and v =

where: v – reaction rate, E – enzyme concentration, and S – substrate concentration. In both equations KM and kcat represent the Michaelis and the catalytic constant, respectively, while n in the second equation stands for the Hill coefficient, representing the type and degree of cooperativity.

CYP53A15 reconstitution systems with CPRs, HPLC analysis and reaction kinetics Reconstitution of the CYP53A15 activity was tested with CPR1 and CPR2, and substrates as described (Podobnik



et al., 2008). For RS1, 2 ml of 4.5 mM CYP53A15, 2 ml of the membrane fraction of 0.25 mM CPR1 and 136 ml of 1% 10 mg ml-1 DLPC were prepared. For RS2, we combined 1 ml of 4.5 mM CYP53A15, 10 ml of the membrane fraction of 0.025 mM CPR2 and 550 ml of 1% 10 mg ml-1 DLPC. All experiments were carried out in 50 mM HEPES (pH 7.4 for CPR1 and pH 8.2 for CPR2) containing 15% glycerol and 0.1 mM EDTA. Buffer pH was adjusted to the theoretical isoelectric points of CPR1 and CPR2, 5.2 and 7.6 respectively. The reaction was started by adding 0.1 M NADPH and the substrate (100 mM benzoic acid or 3-methoxybenzoic acid). Organic product was extracted using acetonitrile (ACN). Reactions were stopped after 5, 10, 25, 45 and 90 min. 40 ml of sample was added to 60 ml of ACN. After centrifugation (16 000 g, 10 min), the supernatant was analysed by HPLC. CYP53A15 activity was measured by monitoring product formation on the Waters Alliance HPLC system, using an YMC-ODS chromatographic column using a gradient program with a two solvent system (solvent A: 5% w/v ACN, 0.1% w/v H2SO4; solvent B: 90% w/v ACN, 0.1% w/v H2SO4). Solvent A (100%) was used as the mobile phase for the initial 10 min, then lowered to 60% (10–12 min), then changed back to 100% of solvent A for the final 12 min. The flow rate was 1 ml per min, and the injection volume was 20 ml. The signals were detected at 215 nm and 250 nm. Standard phenolic acids were prepared in a solvent consisting of ACN : water (1:1; w/v).

Fluorometric CYP53A15 activity assay based on oxygen consumption To determine CYP53A15 activity in reconstitutions with either CPR, and in the presence of different substrates, a fluorometric assay was performed according to Olry et al. (2007). Both reconstitution systems were prepared as described above. For comparative studies of the functioning of RS1 and RS2, we tested benzoic acid, 3-hydroxybenzoic acid, 3-methoxybenzoic acid, 4-methoxybenzoic acid, isoeugenol, cinnamic acid, coumarin, 4-hydroxycoumarin and scopoletin as substrates. To establish which compounds were also substrates of CYP53A15 reconstituted with CPR1, we tested isoeugenol, phenylacetaldehyde, phenalalanine, phenylethanol, phenylacetate, phenylpyruvate and benzyl alcohol. All substrate concentrations were 300 mM. The assay uses standard BD Oxygen Biosensor System 96-well plates (BD Biosciences, Germany). Oxygen consumption due to substrate oxygenation was measured at 485 nm (lexc) on a FLUOstar Galaxy microplate reader (BMG Labtechnologies GmbH, Germany). The NADPH concentration was 500 mM. 3 mM Glucose-6-phosphate and 0.4 U/well of glucose-6-phosphate dehydrogenase were added to maintain saturating NADPH concentration and prevent CPR inhibition by oxidized NADPH. Fluorescence at 615 nm was recorded every min for 2 h.

Growth inhibition of C. lunatus strains We examined inhibitory activity of different substrates on C. lunatus growth. For temporal studies of growth inhibition, minimal medium agar plates were supplemented with

benzoic acid, 3-methoxybenzoic acid and isoeugenol (1 mM final concentration) and inoculated centrally with mycelial discs (6 mm diameter) obtained from a 7-day-old mycelial culture of C. lunatus wt, Dcpr2 and Dbph strains. As growth of the Dcpr1 strain was severely reduced, this mutant was not used in the experiment as a comparative analysis would not be possible. In all experiments, media with ethanol added were used as control. Mycelial growth at 28°C was monitored daily for 7 days or until the surface of the plate was homogeneously overgrown. Experiments were performed in triplicates.

Drop test assay in S. cerevisiae To test the growth of various yeast control strains and transformants (wt, ncp1D, and ncp1D with the native yeast CPR, Ncp1) and the ncp1D strain expressing C. lunatus proteins (CPR1, CPR2, CYP53A15, CPR1 + CYP53A15, CPR2 + CYP53A15) in the presence of benzoic acid, yeast cells were grown overnight in liquid SD Ura- Leu- medium. The plasmid pYeDP8 containing the NCP1 ORF was kindly provided by Dr Louise Agrain. Empty vectors pYES2 or Yep181 were used for restoration of uracil or leucine prototrophy, where necessary. Cultures were then diluted 1:1000 into SGal Ura- Leu- medium to induce the expression of recombinant genes under the GAL1 promoter, and cultivated to log phase. The OD600 of the cultures was adjusted to about 0.2. Ten-fold serial dilutions of the cell suspensions were prepared and spotted on solid SGal Ura- Leu- medium containing benzoic acid (50 mM). Growth at 30°C was monitored for 3–4 days.

Ergosterol quantification Ergosterol levels in all transformed strains (see above) were quantified following the protocol described in ArthingtonSkaggs et al. (1999). Briefly, single colony transformants from SD Ura- Leu- media were inoculated in 50 ml of SD Ura- and grown overnight at 30°C and 170 r.p.m. The stationary-phase cells were then harvested by centrifugation (1330 g, 5 min), washed with distilled water and dried in a DNA Speed Vac DNA110 (Savant Instruments, USA) for approximately 2 h. The dried pellet was then weighed. Three millilitres of 25% alcoholic potassium hydroxide solution (25 g of KOH and 35 ml of sterile distilled water, brought to 100 ml with 100% ethanol), were added to each pellet and vortexed for 1 min. Cell suspensions were transferred to a 12 ml sterile borosilicate glass vial, incubated in a 85°C water bath for 1 h and left to cool to room temperature. Sterols were extracted by adding a mixture of 1 ml sterile distilled water and 3 ml of n-heptane, then vortexed for 3 min. The heptane layer was transferred to a clean glass tube and stored at -20°C for 24 h. The heptane was left to evaporate in the Speed Vac for approximately 30 min. The sample was dissolved in 1 ml 99.8% ethanol and ultrasonicated for 10 min. Ergosterol and dehydroergosterol membrane content was analysed on the Waters Alliance HPLC system using an YMC-ODS chromatographic column and isocratic elution with a two solvent system [solvent A: 5% w/v ACN, 0.05% w/v trifluoroacetic acid (TFA); solvent B: 95% w/v ACN, 0.05% w/v TFA] for 30 min. The flow rate was 1.5 ml per min and the



injection volume was 20 ml. The signals were detected at 230 nm and 280 nm. External standards for 0.5 mM ergosterol and 0.5 mM dehydroergosterol were prepared in a solvent consisting of ACN : water (1:1; w/v).

QPCR The TaqMan (FAM MGB) ready-to-use probes and primers mixtures, specific for C. lunatus bph, cpr1 and cpr2 genes, were ordered from Custom TaqMan Gene Expression Assays Service (Applied Biosystems, USA). 18S rRNA (Eukaryotic 18S rRNA Endogenous Control (FAM MGB Probe, NonPrimer Limited), Applied Biosystems, USA) was used as the reference gene. The QPCR experiments were run on the LightCycler 480 Real-Time PCR System (Roche) using the TaqMan Universal PCR Master Mix (Applied Biosystems, USA) according to the recommendations of the Applied Biosystems’ Assays-by-Design Service. All experiments were performed in triplicates. Gene expression was analysed using the REST 2009 software tool where the expression ratio results of investigated transcripts are tested for significance by a randomization test (Pfaffl et al., 2002). The expression levels of cpr1 and cpr2 genes under normal growth conditions (i.e. non-induced mycelia) at four time points (15, 45, 90 and 200 min) were compared with the comparative CT method (Schmittgen and Livak, 2008). The equation, typically used to calculate fold change in gene expression between two samples, was used to investigate the fold change in expression of cpr2 relative to cpr1 at a particular time point:

Fold change = 2− ΔΔCT = [(CT cpr 1− CT 18sRNA ) − (CT cpr 2 − CT 18sRNA )].

Modelling and docking of CPR1 and CYP53A15 The FMN-binding domain of CPR1 was modelled using the structure of human-yeast chimeric CPR (PDB ID 3FJO) as the template as described previously (Podobnik et al., 2008). Briefly, the two sequences were aligned, residues mutated accordingly with the Whatif molecular modelling tool and the gaps manipulated manually (Vriend, 1990). After structure optimization, a 1 ns constant pressure and temperature (CPT) dynamic simulation (300 K, 1 bar, time step 1 fs) invoking the EWALD summation for calculating the electrostatic interactions was run. The last frame was optimized and used as an input for protein docking together with the homology model of CYP53A15 (PMDB ID: PM0075149) (http:// rosettadock.graylab.jhu.edu/viewjob?id=4294) (Lyskov and Gray, 2008). Initial orientation of the two structures was chosen on the basis of the binding site searching algorithm, implemented in ProBis (http://probis.cmm.ki.si/) (Konc and Janežicˇ, 2010). A structure was selected from the docking run (job#4294-protein-3.pdb) and used for further modelling where the prosthetic groups FMN and haem were added, and another 2 ns CPT-EWALD dynamic simulation was performed (40 536 atoms, 300 K, 1 bar, time step 1 fs). A 3D image was completed by adding the flavin adenine dinucleotide and NADPH-binding domains from the 3FJO structure and visualized using LIGHT software.

Acknowledgments The authors wish to thank Drs Colette Breuil and Gordon Robertson for helpful discussions and critical reading of the manuscript, and Jelka Lenarcic for expert technical assistance in the molecular biology lab. The work was supported by the Research Programme Grant No. P1-0104 from the Slovenian Research Agency.

References Aigrain, L., Pompon, D., Morera, S., and Truan, G. (2009) Structure of the open conformation of a functional chimeric NADPH cytochrome P450 reductase. EMBO Rep 10: 742– 747. Arthington-Skaggs, B.A., Jradi, H., Desai, T., and Morrison, C.J. (1999) Quantitation of ergosterol content: novel method for determination of fluconazole susceptibility of Candida albicans. J Clin Microbiol 37: 3332–3337. Ballou, D.P., Entsch, B., and Cole, L.J. (2005) Dynamics involved in catalysis by single-component and twocomponent flavin-dependent aromatic hydroxylases. Biochem Biophys Res Commun 338: 590–598. Bradford, M.M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248– 254. Brankova, L., Ivanov, S., and Alexieva, V. (2007) The induction of microsomal NADPH:cytochrome P450 ansssd NADH:cytochrome b5 reductases by long-term salt treatment of cotton (Gossypium hirsutum L.) and bean (Phaseolus vulgaris L.) plants. Plant Physiol Biochem 45: 691–695. Bräse, S., Encinas, A., Keck, J., and Nising, C.F. (2009) Chemistry and biology of mycotoxins and related fungal metabolites. Chem Rev 109: 3903–3990. Chen, D.-C., Yang, B.-C., and Kuo, T.-T. (1992) One-step transformation of yeast in stationary phase. Curr Genet 21: 83–84. ˇ rešnar, B., and Petricˇ, Š. (2011) Cytochrome P450 enzymes C in the fungal kingdom. Biochim Biophys Acta 1814: 29–35. Doddapaneni, H., Chakraborty, R., and Yadav, J.S. (2005) Genome-wide structural and evolutionary analysis of the P450 monooxygenase genes (P450ome) in the white rot fungus Phanerochaete chrysosporium: evidence for gene duplications and extensive gene clustering. BMC Genomics 6: 92. Ellis, J., Gutierrez, A., Barsukov, I.L., Huang, W.-C., Grossmann, J.G., and Roberts, G.C.K. (2009) Domain motion in cytochrome P450 reductase: conformational equilibria revealed by NMR and small-angle X-ray scattering. J Biol Chem 284: 36628–36637. Eppink, M.H.M., Cammaart, E., Van Wassenaar, D., Middelhoven, W.J., and Van Berkel, W.J.H. (2000) Purification and properties of hydroquinone hydroxylase, a FADdependent monooxygenase involved in the catabolism of 4-hydroxybenzoate in Candida parapsilosis CBS604. Eur J Biochem 267: 6832–6840. Faber, B.W., Van Gorcom, R.F.M., and Duine, J.A. (2001) Purification and characterization of benzoate-parahydroxylase, a cytochrome P450 (CYP53A1), from Aspergillus niger. Arch Biochem Biophys 394: 245–254.



Fraser, J.A., Davis, M.A., and Hynes, M.J. (2002) The genes gmdA, encoding an amidase, and bzuA, encoding a cytochrome P450, are required for benzamide utilization in Aspergillus nidulans. Fungal Genet Biol 35: 135–146. Fujii, T., Nakamura, K., Shibuya, K., Tanase, S., Gotoh, O., Ogawa, T., and Fukuda, H. (1997) Structural characterization of the gene and corresponding cDNA for the cytochrome P450rm from Rhodotorula minuta which catalyzes formation of isobutene and 4-hydroxylation of benzoate. Mol Gen Genet 256: 115–120. Fukuda, H., Nakamura, K., Sukita, E., Ogawa, T., and Fujii, T. (1996) Cytochrome P450rm from Rhodotorula minuta Catalyzes 4-Hydroxylation of Benzoate. J Biochem 119: 314–318. Gietz, D.R., and Woods, R.A. (2002) Transformation of yeast by lithium acetate/single-stranded carrier DNA/ polyethylene glycol method. In Guide to Yeast Genetics and Molecular and Cell Biology – Part B Methods in Enzymology. Guthrie, C., and Fink, G.R. (eds). London: Academic Press, pp. 87–96. Gillam, E.M.J., and Hunter, D.J.B. (2007) Chemical defense and exploitation. Biotransformation of xenobiotics by cytochrome P450 enzymes. In Metal Ions in Life Sciences. Volume 3: The Ubiquitous Roles of Cytochrome P450 Proteins. Sigel, A., Sigel, H., and Sigel, R.K.O. (eds). West Sussex: Wiley and Sons, pp. 477–560. Hamdane, D., Xia, C., Im, S.-C., Zhang, H., Kim, J.-J.P., and Waskell, L. (2009) Structure and function of an NADPHcytochrome P450 oxidoreductase in an open conformation capable of reducing cytochrome P450. J Biol Chem 284: 11374–11384. Hannemann, F., Bichet, A., Ewen, K.M., and Bernhardt, R. (2007) Cytochrome P450 systems – biological variations of electron transport chains. Biochim Biophys Acta 1770: 330–344. Harwood, C.S., and Parales, R.E. (1996) The b-ketoadipate pathway and the biology of self-identity. Annu Rev Microbiol 50: 553–590. Hoffman, C.S., and Winston, F. (1987) A ten-minute DNA preparation from yeast efficiently releases autonomous plasmids for transformaion of Escherichia coli. Gene 57: 267–272. Jang, H.-H., Jamakhandi, A.P., Sullivan, S.Z., Yun, C.-H., Hollenberg, P.F., and Miller, G.P. (2010) Beta sheet 2-alpha helix C loop of cytochrome P450 reductase serves as a docking site for redox partners. Biochim Biophys Acta 1804: 1285–1293. Kalin, M., Neujahr, H.Y., Weissmahr, R.N., Sejlitz, T., Johl, R., Fiechter, A., and Reiser, J. (1992) Phenol hydroxylase from Trichosporon cutaneum: gene cloning, sequence analysis, and functional expression in Escherichia coli. J Bacteriol 174: 7112–7120. Kelly, D.E., Kraševec, N., Mullins, J., and Nelson, D.R. (2009) The CYPome (Cytochrome P450 complement) of Aspergillus nidulans. Fungal Genet Biol 46 (Suppl. 1): S53–S61. Kelly, S.L., Lamb, D.C., Cannieux, M., Greetham, D., Jackson, C.J., Marczylo, T., et al. (2001) An old activity in the cytochrome P450 superfamily (CYP51) and a new story of drugs and resistance. Biochem Soc Trans 29: 122–128. Konc, J., and Janežicˇ, D. (2010) ProBiS algorithm for detec-

tion of structurally similar protein binding sites by local structural alignment. Bioinformatics 26: 1160–1168. Lah, L., Kraševec, N., Trontelj, P., and Komel, R. (2008) High diversity and complex evolution of fungal cytochrome P450 reductase: cytochrome P450 systems. Fungal Genet Biol 45: 446–458. Lamb, D.C., Kelly, D.E., Manning, N.J., Kaderbhai, M.A., and Kelly, S.L. (1999) Biodiversity of the P450 catalytic cycle: yeast cytochrome b5/NADH cytochrome b5 reductase complex efficiently drives the entire sterol 14-demethylation (CYP51) reaction. FEBS Lett 462: 283–288. Laursen, T., Jensen, K., and Moller, B.L. (2011) Conformational changes of the NADPH-dependent cytochrome P450 reductase in the course of electron transfer to cytochromes P450. Biochim Biophys Acta 1814: 132–138. Ling, M., Merante, F., and Robinson, B.H. (1995) A rapid and reliable DNA preparation method for screening a large number of yeast clones by Polymerase Chain Reaction. Nucl Acid Res 23: 4924–4925. Lyskov, S., and Gray, J.J. (2008) The RosettaDock server for local protein–protein docking. Nucl Acid Res 36: W233– W238. Malonek, S., Rojas, M.C., Hedden, P., Gaskin, P., Hopkins, P., and Tudzynski, B. (2004) The NADPH-cytochrome P450 reductase gene from Gibberella fujikuroi is essential for gibberellin biosynthesis. J Biol Chem 279: 25075– 25084. Matsuzaki, F., and Wariishi, H. (2005) Molecular characterization of cytochrome P450 catalyzing hydroxylation of benzoates from the white-rot fungus Phanerochaete chrysosporium. Biochem Biophys Res Commun 334: 1184– 1190. Matsuzaki, F., Shimizu, M., and Wariishi, H. (2008) Proteomic and metabolomic analyses of the white-rot fungus Phanerochaete chrysosporium exposed to exogenous benzoic Acid. J Proteome Res 7: 2342–2350. Matthews, D.E., and Van Etten, H.D. (1983) Detoxification of the phytoalexin pisatin by a fungal cytochrome P-450. Arch Biochem Biophys 224: 494–505. Miroux, B., and Walker, J.E. (1996) Over-production of proteins in Escherichia coli: mutant hosts that allow synthesis of some membrane proteins and globular proteins at high levels. J Mol Biol 260: 289–298. Murataliev, M.B., Guzov, V.M., Walker, F.A., and Feyereisen, R. (2008) P450 reductase and cytochrome b5 interactions with cytochrome P450: effects on house fly CYP6A1 catalysis. Insect Biochem Mol Biol 38: 1008–1015. Nelson, D., Ming, R., Alam, M., and Schuler, M. (2008) Comparison of cytochrome P450 genes from six plant genomes. Tropical Plant Biol 1: 216–235. Oldenburg, K.R., Vo, K.T., Michaelis, S., and Paddon, C. (1997) Recombination-mediated PCR-directed plasmid construction in vivo in yeast. Nucleic Acids Res 25: 451– 452. Olry, A., Schneider-Belhaddad, F., Heintz, D., and WerckReichhart, D. (2007) A medium-throughput screening assay to determine catalytic activities of oxygen-consuming enzymes: a new tool for functional characterization of cytochrome P450 and other oxygenases. Plant J 51: 331–340. Omura, T., and Sato, R. (1964) The carbon monoxide-binding pigment of liver microsomes. J Mol Biol 239: 2370–2378.



Ortiz de Montellano, P.R., and De Voss, J.J. (2005) Substrate oxidation by cytochrome P450 enzymes. In Cytochrome P-450: Structure, Mechanism and Biochemistry. Ortiz de Montellano, P.R. (ed.). 3rd edn. New York: Kluwer Academic/Plenum Publishers, pp. 183–246. Paine, M.J.I., Scrutton, N.S., Munro, A.W., Gutierrez, A., Roberts, G.C.K., and Wolf, C.R. (2005) Electron transfer partners of cytochrome P450. In Cytochrome P450: Structure, Mechanism, and Biochemistry. Ortiz De Montellano, P. (ed.). New York: Kluwer Academic/Plenum Publishers, pp. 115–138. Palfey, B.A., and McDonald, C.A. (2010) Control of catalysis in flavin-dependent monooxygenases. Arch Biochem Biophys 493: 26–36. Park, H.G., Lim, Y.R., Eun, C.Y., Han, S., Han, J.S., Cho, K.S., et al. (2010) Candida albicans NADPH-P450 reductase: expression, purification, and characterization of recombinant protein. Biochem Biophys Res Commun 396: 534–538. Parke, D., D’Argenio, D.A., and Ornston, L.N. (2000) Bacteria are not what they eat: that is why they are so diverse. J Bacteriol 182: 257–263. Parsi, Z., and Gorecki, T. (2006) Determination of ergosterol as an indicator of fungal biomass in various samples using non-discriminating flash pyrolysis. J Chromatogr A 1130: 145–150. Pfaffl, M.W., Horgan, G.W., and Dempfle, L. (2002) Relative expression software tool (REST©) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucl Acid Res 30: e36. Plemenitaš, A., Žakelj-Mavricˇ, M., and Komel, R. (1988) Hydroxysteroid dehydrogenase of Cochliobolus lunatus. J Steroid Biochem 29: 371–372. Podobnik, B., Stojan, J., Lah, L., Kraševec, N., Seliškar, M., Lanišnik Rižner, T., et al. (2008) CYP53A15 of Cochliobolus lunatus, a target for natural antifungal compounds. J Med Chem 51: 3480–3486. Premkumar, R., Rao, P.V.S., Sreeleela, N., and Vaidyanathan, C. (1969) m-Hydroxybenzoic acid 4-hydroxylase from Aspergillus niger. Biochem Cell Biol 47: 825–827. Schmittgen, T.D., and Livak, K.J. (2008) Analyzing real-time PCR data by the comparative CT method. Nat Protocols 3: 1101–1108. Sevrioukova, I.F., and Poulos, T.L. (2010) Structural biology of redox partner interactions in P450cam monooxygenase: a fresh look at an old system. Arch Biochem Biophys 507: 66–74. Shen, A.L., and Kasper, C.B. (1995) Role of acidic residues in the interaction of NADPH-cytochrome P450 oxidoreductase with cytochrome P450 and cytochrome c. J Biol Chem 270: 27475–27480. Siewers, V., Smedsgaard, J., and Tudzynski, P. (2004) The P450 monooxygenase BcABA1 is essential for abscisic acid biosynthesis in Botrytis cinerea. Appl Environ Microbiol 70: 3868–3876. Silva, R.R., Corso, C.R., and Matheus, D.R. (2010) Effect of

culture conditions on the biomass determination by ergosterol of Lentinus crinitus and Psilocybe castanella. World J Microbiol Biotechnol 26: 841–846. Sutter, T.R., and Loper, J.C. (1989) Disruption of the Saccharomyces cerevlsiae gene for NADPH-cytochrome P450 reductase causes increased sensitivity to ketoconazole. Biochem Biophys Res Commun 160: 1257–1266. Tiedje, C., Holland, D.G., Just, U., and Höfken, T. (2007) Proteins involved in sterol synthesis interact with Ste20 and regulate cell polarity. J Cell Sci 120: 3613–3624. Torres Pazmino, D.E., Winkler, M., Glieder, A., and Fraaije, M.W. (2010) Monooxygenases as biocatalysts: classification, mechanistic aspects and biotechnological applications. J Biotechnol 146: 9–24. Troncoso, C., Carcamo, J., Hedden, P., Tudzynski, B., and Cecilia Rojas, M. (2008) Influence of electron transport proteins on the reactions catalyzed by Fusarium fujikuroi gibberellin monooxygenases. Phytochemistry 69: 672– 683. Van Berkel, W., Kamerbeek, N.M., and Fraaije, M.W. (2006) Flavoprotein monooxygenases, a diverse class of oxidative biocatalysts. J Biotechnol 124: 670–689. Van Den Brink, J.M., Punt, P.J., Van Gorcom, R.F.M., and Van Den Hondel, C.A.M.J.J. (2000) Regulation of expression of the Aspergillus niger benzoate para-hydroxylase cytochrome P450 system. Mol Gen Genet 263: 601–609. Venkateswarlu, K., Lamb, D.C., Kelly, D.E., Manning, N.J., and Kelly, S.L. (1998) The N-terminal membrane domain of yeast NADPH-cytochrome P450 (CYP) oxidoreductase is not required for catalytic activity in sterol biosynthesis or in reconstitution of CYP activity. J Biol Chem 273: 4492–4496. Vriend, G. (1990) WHAT IF: A molecular modeling and drug design program. J Mol Graph 8: 52–56. Wach, A., Brachat, A., Rebischung, C., Steiner, S., Pokorni, K., Heesen, S., and Philippsen, P. (1998) 5 PCR-Based Gene Targeting in Saccharomyces cerevisiae. In Yeast Gene Analysis – Methods in Microbiology. Brown, A.J., and Tuite, M. (eds). London: Academic Press, pp. 67–81. Warrilow, A.G., Lamb, D.C., Kelly, D.E., and Kelly, S.L. (2002) Phanerochaete chrysosporium NADPH-cytochrome P450 reductase kinetic mechanism. Biochem Biophys Res Commun 299: 189–195. Wu, Z.L., Qiao, J., Zhang, Z.G., Guengerich, F.P., Liu, Y., and Pei, X.Q. (2009) Enhanced bacterial expression of several mammalian cytochrome P450s by codon optimization and chaperone coexpression. Biotechnol Lett 31: 1589–1593.

Supporting information Additional supporting information may be found in the online version of this article. Please note: Wiley-Blackwell are not responsible for the content or functionality of any supporting materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.
