Mechanism of flavin transfer and oxygen activation by the two-component flavoenzyme styrene monooxygenase

ABB Archives of Biochemistry and Biophysics 442 (2005) 102–116 www.elsevier.com/locate/yabbi

Mechanism of flavin transfer and oxygen activation by the two-component flavoenzyme styrene monooxygenase Auric Kantz, Franklin Chin, Nagamani Nallamothu, Tim Nguyen, George T. Gassner * Department of Chemistry and Biochemistry, San Francisco State University, San Francisco, CA 94132-4163, USA Received 21 June 2005, and in revised form 21 July 2005 Available online 18 August 2005

Abstract Styrene monooxygenase (SMO) from Pseudomonas putida S12 is a two-component flavoenzyme composed of the NADH-specific flavin reductase, SMOB, and FAD-specific styrene epoxidase, SMOA. Here, we report the cloning, and expression of native and histidine-tagged versions of SMOA and SMOB and studies of the flavin transfer and styrene oxygenation reactions. In the reductive half-reaction, SMOB catalyzes the two-electron reduction of FAD with a turnover number of 3200 s
1. Single turnover studies of the reaction of reduced SMOA with substrates indicate the formation of a stable oxygen intermediate with the absorbance characteristics of a flavin hydroperoxide. Based on the results of numerical simulations of the steady-state mechanism of SMO, we find that the observed coupling of NADH and styrene oxidation can be best explained by a model, which includes both the direct transfer and passive diffusion of reduced FAD from SMOB to SMOA.
2005 Elsevier Inc. All rights reserved. Keywords: Electron transfer; Flavin monooxygenase; Epoxidation; Enzyme intermediate; Redox; Pre-steady-state kinetics; Transient kinetics; Styrene; Reductase; Singlet oxygen

Pyridine nucleotide-dependent oxygenases have evolved in prokaryotes as a diverse group of enzymes existing in both soluble and membrane-bound forms. Structurally, they range from self-contained single-component enzymes to complex systems including separate reductase, electron-transfer, regulatory, and oxygenase activities. These enzymes engage heme, non-heme iron, copper, or flavin-dependent active sites in their substrate oxygenation reactions [1–5]. Flavin-dependent monooxygenases catalyze reactions ranging from the genesis of bioluminescence to the catabolic oxidation of hydrocarbons [6,7]. Single-component flavoenzymes house both flavin reduction and substrate oxygenation activities in the same contiguous peptide and have been shown to proceed through flavin *

Corresponding author. Fax: +1 415 338 2384. E-mail address: [email protected] (G.T. Gassner).

0003-9861/$ - see front matter
2005 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2005.07.020

C-4a hydroperoxide and hydroxide intermediates in the substrate oxygenation reaction [7]. Similar intermediates have been reported and proposed for the closely related two-component systems [8–12]. Two distinct classes of two-component flavoenzymes have been identified. The first of these, exemplified by 4hydroxyphenylacetate-3-monooxygenase from Pseudomonas putida, is composed of a separate oxygenase and regulatory protein required for the efficient coupling of pyridine nucleotide oxidation and substrate oxygenation activities [9,13]. More commonly encountered twocomponent flavoenzymes are composed of distinct flavin reductase and monooxygenase activities segregated on separate peptide units [6,10–12,14–16]. The spatial resolution of flavin reduction and substrate oxygenation activities in the two-component flavin monooxygenases requires either a direct or a diffusive transfer of reduced flavin from the reductase

A. Kantz et al. / Archives of Biochemistry and Biophysics 442 (2005) 102–116

to the monooxygenase. Reductase components of these enzymes have been distinguished based on differences in their mechanisms of interaction with flavin substrates [17]. Analysis of the steady-state mechanisms of bacterial luciferase and alkanesulfonate monooxygenase suggests that these systems operate through a mechanism of direct transfer of reduced flavin [16,18,19]. Two-component styrene monooxygenases (SMOs)1 from Pseudomonas are similarly composed of highly conserved flavin reductase and epoxidase components. It has been suggested that in these enzymes and closely related systems including 4-hydroxyphenylacetate-3-monooxygenase isolated from Escherichia coli W flavin transfer from the reductase to oxygenase component is purely diffusive [12,15,20]. In support of this mechanism it has been shown that the monooxygenase component of SMO is capable of substrate epoxidation in the absence of reductase when provided with an alternate source of reduced FAD [21]. However, rapid reactions of reduced FAD with oxygen and oxidized FAD are likely to make the diffusive mode flavin transfer quite inefficient [7]. Substrate epoxidation is a common reaction of both the heme and non-heme iron-containing enzymes [22– 25], but unusual for flavoenzyme monooxygenases. Studies of the oxygen reaction of SMO may provide insight into flavin-based epoxidation mechanism of this enzyme and other flavoenzyme epoxidases such as squalene and zeaxanthin monooxygenases [26,27]. Here, we report the cloning expression of native and N-terminally histidine-tagged versions of the styrene monooxygenase reductase and epoxidase components of P. putida S12. Kinetic data recording the time-dependent evolution and spectral characteristics of oxygen intermediates are presented. To further explore the mode of flavin transfer in this system, the observed efficiency of coupling NADH oxidation to styrene consumption over a range of enzyme component ratios and FAD concentrations is compared with the results of simulations of direct and diffusive flavin-transfer mechanisms.

Materials and methods Cell growth, protein, and DNA purification from native P. putida S12 Pseudomonas putida S12 was purchased from the ATCC [28]. This strain of cells grows poorly in culture with styrene as the sole source of carbon and energy and expression from the styrene operon is subdued by catabolite repression by alternate carbon sources found in enriched media [29]. The styrene operon of P. putida S12 is not catabolite-repressed by phenyl acetic acid [30], 1 Abbreviations used: SMO, styrene monooxygenase; PMSF, phenylmethylsulfonyl fluoride; EDTA, ethylenediaminetetraacetic acid.

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but the cells grow poorly in liquid culture when limited to this substrate. To circumvent these limitations, cells were initially grown in Trypticase-Soy Broth supplemented with 0.1% phenyl acetic acid in a 10 L New Brunswick BioFlow 2000 Fermenter. Approximately 130 g of cell paste was recovered by centrifugation following a 16-h growth period at 33 C. The cell paste was then resuspended in minimal media containing phenylacetic acid as the only carbon source. After a half hour equilibration time, the styrene operon was induced by the addition of styrene to the cell culture to a final concentration of 1 mM. Following a 1-h induction period, approximately 130– 150 g of cell paste was recovered. Cells were resuspended in 20 mM phosphate buffer (pH 7) containing 1 mM of each phenylmethylsulfonyl fluoride (PMSF) and ethylenediaminetetraacetic acid (EDTA) and disrupted by sonication for a total time of 6 min, while maintaining the solution temperature below 10 C. Particulate cell debris was removed from the soluble protein fraction by centrifugation for 45 min at 18,000 rpm in an SS34 rotor at 4 C. Ultimately native styrene monooxygenase components were partially purified by anion exchange, dye-ligand, and reverse-phase chromatography. Cloning design of expression vectors and sequencing DNA was isolated from P. putida S12 after sub-culturing on agar plates supplemented with 0.1% phenyl acetic acid and 1 mM indole. Indigo producing cells were recovered by centrifugation and subjected to alkaline lysis and genomic DNA extraction by using a Qiagen Genomic Tip protocol. Primers were designed based on the reported sequences of styA and styB from Pseudomonas fluorescens [31] and purchased from Operon. The forward and reverse primers used for the amplification of styA were 5 0 -CCATATGAAAAAGCGTAT CGGTATT-GTTGGTG-3 0 and 5 0 -CCTTAAGTCAG GCCGCGATAGTGGGTGC-3 0 , respectively. Forward and reverse primers for the amplification of styB were 5 0 -CCATATGACGTTAAAAAAAGATATGGCGGT GG-3 0 and 5 0 -CCTTAAGTTAATTCAGCGGCAA CGGGTTAC-3 0 . In each case, the primers were designed to introduce an NdeI site at the 5 0 -end of the amplified styA and styB genes. The styA and styB genes were amplified from genomic DNA by 30 cycles of PCR with Pfu-Turbo polymerase. PCR products were recovered after gel purification by using Q-Biogene silica gel spin-columns. The purified PCR products were 5 0 -phosphorylated by treatment with T4 polynucleotide kinase and inserted by blunt-end ligation with T4 ligase into the EcoRV site of pZErO-2 vector purchased from Invitrogen. E. coli TOP-10 cells were transformed and positive clones were selected. pZErO-2 vectors containing styA and styB were recovered by the Qiagen mini prep

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plasmid purification protocol. The styA and styB genes were then ligated into the NdeI and EcoRI restriction sites in the multiple cloning site of pET-28 and pET-29 expression vectors from Novagen. This work yielded vectors pET-29SMOA and pET-29SMOB designed for the synthesis of native SMOA and SMOB and vectors pET-28NSMOA and pET-28NSMOB for the synthesis of thrombin cleavable, N-terminally histidine-tagged SMOA and SMOB. Sequencing primers were designed and used to verify the styA and styB gene sequences in the recombinant expression vectors. It was determined that several mutations were introduced due to differences in the P. fluorescens and S12 DNA sequences. However, the encoded amino acid sequences of SMOA and SMOB were unaffected due to degeneracy of the genetic code translation. The sequence of styA from P. putida S12 was previously reported (GenBank Accession No. Y13349). Our results confirm this sequence. The SMOB sequence was determined to be identical to that of styB isolated from Pseudomonas sp. VLB [32]. Expression and purification of recombinant proteins Native and N-terminally histidine-tagged forms of SMOA were expressed in E. coli BL21(DE3) cells. Typically, a yield of 30 mg of protein per liter was recovered after a 1-h induction period in growth medium supplemented with 30 lg mL
1 of ampicillin and 1 mM IPTG at 37 C. A large quantity of indigo dye accumulates when these proteins are expressed in LB medium and for this reason SMOA was typically expressed in M-9 medium. Native and N-terminally histidine-tagged SMOB was expressed from E. coli BL21 (DE3) under the same conditions. SDS–PAGE assay of soluble and insoluble cell fractions recovered after expression indicated that whereas SMOA and N-SMOA are over-expressed as purely soluble protein, the over-expressed SMOB and N-SMOB primarily form inclusion bodies. Only about 1% of the SMOB expressed remains in the soluble fraction, and we were unsuccessful in our efforts to increase the fraction of protein expressed in soluble form by inducing the cells to express protein at lower temperatures. It was possible to recover soluble, active SMOB from inclusion bodies by denaturation in 8 M urea containing 5 mM DTT as previously reported [15]. N-terminally histidine-tagged SMOA and SMOB were recovered from 4–10 L cell growths in M-9 medium. Cell pellets were resuspended and sonicated in Ni– NTA equilibration buffer composed of 50 mM sodium phosphate, 10 mM imidazole, 300 mM sodium chloride, 1 mM PMSF, and 1 mM EDTA. Immediately after centrifugation, soluble extracts containing N-His-SMOA and N-His-SMOB were pumped onto a 1.5 · 5 cm Omnifit column containing freshly charged Sigma HisSelect resin at a flow rate of 3 mL min
1. The column was then washed with one column volume of equilibra-

tion buffer followed by a linear 50 mL gradient of imidazole from 10 to 250 mM. Protein fractions were pooled based on UV absorbance and SDS–PAGE assay and stored at
80 C after addition of dithiothreitol to a final concentration of 1 mM and glycerol to 50%. Protein concentration measurements and thiol titration Molecular weights and extinction coefficients for SMO components were estimated by using the webbased application Prot Param [33]. Protein concentrations were calculated by using these values and by the Pierce BCA assay with BSA as a protein standard. Accessible thiols were titrated with 5 0 ,5 0 -dithionitrobenzoic acid with dithiothreitol as a thiol standard. Steady-state reaction mechanism of SMOB The steady-state reaction of native SMOB with an array of NADH and FAD concentrations was monitored by stopped-flow absorbance spectroscopy. In these studies, apo enzyme at a concentration of 10 nM after mixing was reacted with defined substrate mixtures in air-saturated 20 mM phosphate buffer, pH 7, containing 5% glycerol and 1 mM dithiothreitol. A stopped-flow spectrophotometer equipped with an Ocean Optics USB-2000 diode array spectrophotometer was used for mixing and data collection. Absorbance changes corresponding to NADH reduction were monitored at 340 nm, while the FAD oxidation state was monitored at 450 nm. Initial rates were calculated from linear fits through the first 5–10 s of absorbance data recording the oxidation of NADH to NAD+. No significant reduction of FAD was detected over this time period due to the rapid reoxidation of FADH by molecular oxygen. Three to five replicate reactions were analyzed for each experimental condition. The reaction rate data corresponding to each experimental NADH and FAD concentrations were entered into a single data array and fit globally along each substrate concentration axis to the function describing a sequential bimolecular model (Eq. (1)), in which the apparent Vmax and KM values are given by Eqs. (2) and (3). v¼

V app max ½NADH ; K app M þ ½NADH

ð1Þ

V app max ¼

V max ½FAD ; K FAD þ ½FAD M

ð2Þ

K app M ¼

NADH K FAD þ K NADH M KS M . K FAD þ ½FAD M

ð3Þ

Data points analyzed in these fits were weighted based on standard deviation from the mean experimental values by using the program GraphPad Prism 4 (GraphPad Software). This global-fitting approach provides clear

A. Kantz et al. / Archives of Biochemistry and Biophysics 442 (2005) 102–116

advantages over alternate methods that rely on data manipulation to generate linear plots, sequential, independent fitting of data sets, and construction of secondary kinetic plots [34]. Pre-steady-state kinetic analysis of SMOA Equal concentrations of N-terminally histidinetagged SMOA and oxidized FAD were combined and made anaerobic in a tonometer equipped with a titration port and 1 cm pathlength quartz cuvette by repeatedly evacuating and back filling with purified nitrogen gas on a vacuum Schlink line. The SMOA–FAD complex was reduced by incremental addition of dithionite from a gas-tight Hamilton syringe attached to the tonometer through a ground glass joint. Stock dithionite solutions were prepared by addition of solid dithionite to an anoxic buffer solution prepared by exhaustive bubbling with nitrogen and confirmed by measuring the absorbance at 314 nm using an extinction coefficient of 8000 M
1 cm
1 [35]. The extent of flavin reduction was recorded by monitoring the absorbance changes at 450 and 314 nm. Reductive titrations were halted just prior to complete FAD reduction or just after the first detectable increase in absorbance 314 nm was observed corresponding to the accumulation of 5–10 lM excess dithionite. Reduced enzyme solutions were transferred to a double-mixing stopped-flow spectrophotometer designed and constructed at San Francisco State University. In test reactions we measured a 3.2 ms dead time for this instrument, which is adequate for the studies described in this paper. The plumbing throughout the instrument is PEAK plastic excepting the drive syringes which are 2.5 mL Hamilton syringes. The heart of stopped flow is a mixer and 20 lL flow cell equipped with fiber optic cables that allow absorbance measurements to be made across a 1 or 0.35 cm optical path length. The plumbing and flow cell are completely contained in a thermostated water bath. During anaerobic studies, the water bath was continuously sparged with industrialgrade nitrogen. Under these conditions, flavin solutions contained in the plumbing and flow cell of the instrument remained fully reduced over the course of several hours. This instrument was used in single-mixing mode to rapidly mix equal volumes reduced SMOA with aerobic buffer solutions containing oxygen and styrene or benzene. The flow cell was illuminated with a 75 W Xenon lamp and absorbance changes were monitored by a USB2000 diode array spectrophotometer interfaced with the stopped-flow spectrophotometer through solarization-resistant fused silica optical fibers purchased from Polymicro Technologies. Each processed absorbance spectrum represents the average of seven raw spectra each recorded at a frequency of 333 s
1. Noise deriving from small fluctuations in xenon lamp intensity was diminished by reference monitoring. The total effec-

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tive 21 ms exposure time was short enough to resolve the fastest reactions detected in our studies and long enough to provide a good signal-to-noise ratio. Kinetic data were fit to functions consisting of sums of 3–4 exponentials (Eq. (4)) by using the program KaleidaGraph (Synergy Software). Aðk;tÞ ¼

i X

an e
kn t þ const.

ð4Þ

n¼1

Observed rate constants obtained in this way were used to estimate the time-dependent changes in concentrations in the sequential transformation of reactants to intermediate species and products by using the KINSIM program [36,37]. The resulting concentration tables were then referenced to find the best-fitting extinction coefficients corresponding to each intermediate spectrum by using the table function built into the KaleidaGraph program. Efficiency of coupling NADH and styrene oxidation reactions of SMO The steady-state kinetic reaction of the two-component styrene monooxygenase system was recorded by using a Ocean Optics DT-1000 deuterium lamp and USB2000 diode array spectrophotometer interfaced with the stopped-flow instrument described above. The stopped-flow instrument allowed precise 1:1 mixing and measurement of initial rates and the control of temperature, styrene, and oxygen concentration. Steady-state kinetics were monitored simultaneously at 450 nm where only oxidized FAD has
1 significant absorbance ðEM cm
1 Þ, at 450 ¼ 11; 300 M
1 M 340 nm where both FAD ðE340 ¼ 4680 M cm
1 Þ and
1 NADH ðEM cm
1 Þ absorb significantly, 340 ¼ 6220 M
1 and at 245 nm where FAD ðEM cm
1 Þ, 245 ¼ 18; 434 M +
1 M
1 NADH ðE245 ¼ 10; 300 M cm Þ, NAD ðEM 245 ¼
1
1 M
1 12; 576 M cm Þ, and styrene ðE245 ¼ 8880 M cm
1 Þ all absorb. Under conditions of aerobic steady-state turnover, the absorbance at 450 nm remains constant at a value corresponding to the initial concentration of oxidized FAD included in the assay. For this reason, it was possible to calculate reaction rate and coupling efficiency by considering absorbance changes corresponding only to the depletion of styrene and the transformation of NADH to NAD+ at 245 and 340 nm. Data collected at higher styrene concentrations were recorded along the short (0.35 cm) path length of the stopped-flow flow cell to allow the full dynamic range of UV-absorbing reactant concentrations to be studied. The rate of NADH oxidation was calculated by dividing the time-dependent absorbance change at 340 nm by the extinction coefficient for NADH at this wavelength. The rate of styrene oxidation was calculated from the time-dependent absorbance changes at 245 and 340 nm with Eq. (5). The coupling effi-

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ciency was determined by calculating the ratio of the NADH and styrene oxidation rates.
NADH  e245
eNAD D½Styrene DA340 245 ¼ Styrene NADH  Dt Dt e245 e340 b


1 eStyrene b 245



DA245 . Dt

ð5Þ

Plots of styrene and NADH oxidation rate data were analyzed by non-linear regression analysis and by numerical simulation according to the models described in the text. The relatively small molar absorbtivity change associated with styrene oxidation and the technical difficulty of accurately preparing low-concentration styrene solutions set the limit of detection by this method at 5–10 lM. Oxidation–reduction potential measurements The redox potentials of free- and enzyme-bound FAD were calculated from anaerobic titration with sodium dithionite as the reductant. These studies were conducted at 25 C in solutions buffered at pH 7 with 20 mM MOPSO. Reactions typically included 20–50 lM concentration FAD and one of a series of solution potential indicators [38]. These studies were performed in a specially designed quartz cuvette and made anaerobic by alternately drawing a vacuum and back flushing with wet, anaerobic nitrogen. Absorbance changes were recorded during the titration at wavelengths corresponding to absorbance peaks of the oxidized indicator and FAD. The absorbance changes recorded at each point in the titration were transformed into concentrations by using Eqs. (6) and (7) in which the D represents the difference between the oxidized and reduced value of each parameter. Each calculated concentration was corrected for dilution by multiplying by the ratio of the final volume after each addition of dithionite divided by the initial volume prior to the addition of dithionite. ½FADred  ¼

½Dyered  ¼

Dye DAk2 DeDye k1
DAk1 Dek2 Dye FAD DeDye
DeFAD k1 Dek2 k1 Dek2

DAk1 DeFAD
DAk2 DeFAD k2 k1 Dye FAD DeDye
DeFAD k1 Dek2 k1 Dek2

.

;

ð6Þ

ð7Þ

The solution potential was calculated at each point in the titration by entering the calculated dye concentrations into the Nernst equation. Results Protein expression and purification More than 95% of the total native and N-terminally tagged versions of SMOB are expressed in the form of

insoluble inclusion bodies. Native SMOB we recovered by resolubilization from inclusion bodies had very similar catalytic activity compared with the soluble fraction of N-terminally tagged SMOB recovered by nickel– NTA affinity chromatography. Enzymes were stored in 50% glycerol at
80 C in a concentration of 1–5 lM. Native and N-terminally tagged SMOB tended to aggregate and lose activity when concentrated beyond this level centrifugally or by reverse osmosis. Dilute solutions of the native and engineered versions of SMOB were stabilized in assay buffers containing 5% glycerol and 1 mM dithiothreitol for use in initial rate measurements. In the presence of phosphate buffer, SMOA precipitates and for this reason experiments including SMOA were buffered with MOPSO. Native and N-terminally histidine-tagged versions of SMOA were found to be soluble and amenable to concentration. The engineered version of SMOA behaves very similarly to the native protein in kinetic assays (Table 1), and since it was significantly easier to isolate the histidine-tagged protein, we elected to use it rather than the native protein in the studies described in this paper. A picture of a gel showing purified native and histidine-tagged proteins is shown in Fig. 1. Steady-state mechanism of SMOB The pH profile of SMOB reacting with NADH and FAD suggests an ideal operating pH between 6 and 7. By fitting the pH profile between pH 5 and 8 it was possible to calculate the value of macroscopic pKa values of 4.4 ± 0.2, 7.6 ± 0.1, and 8.2 ± 0.2 associated with the Vmax and 6.1 ± 1.2, 7.9 ± 3.3, and 8.9 ± 1.9 associated with KM (Fig. 2). In these studies, 20 nM SMOB was combined with 120 lM FAD in 5 mM MOPSO buffer at pH 7 containing 5% glycerol and 1 mM DTT. The enzyme–FAD complex was then rapidly mixed in the stopped flow with each of a series of solutions containing a 100 mM pH defining buffers including citrate, MES, POPSO, CHES, CAPS, and various concentrations of NADH. The exact pH values after mixing were measured experimentally. Since these data were recorded at only one FAD concentration, the pKa values correspond only to apparent values of Vmax and KM and cannot be assigned to individual steady-state parameters. Product inhibition studies were used to characterize the sequence of substrate binding and product release in the reaction of SMOB with NADH and FAD (Fig. 3A). These results suggest that NAD+ acts as a competitive inhibitor of NADH and mixed inhibitor of FAD in its interaction with SMOB. This inhibition pattern is consistent with the sequential bimolecular reaction mechanism in which NADH serves as the leading substrate as previously reported for the reaction of FMN with SMOB from the VLB strain [15].

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Table 1 Comparison of native and N-terminally histidine-tagged SMOA SMOA

(Vmax/KM)app (s
1)a

(Vmax/KM)app (s
1)b

[NADH]/[Styrene]c

[NADH]/[Styrene]d

Native N-His

0.07 ± 0.03 0.05 ± 0.01

0.02 ± 0.01 0.05 ± 0.03

0.9 ± 0.1 1.1 ± 0.1

6.7 ± 2.7 4.2 ± 0.9

Steady-state reactions included 10 nM native SMOB 0.5 lM native or N-terminally histidine-tagged SMOA and 50 lM NADH. aKinetic parameters from reactions which included 10 lM FAD and styrene as the varied substrate. bKinetic parameters from reactions which included 20 lM styrene and FAD as the varied substrate. Coupling of NADH consumption to styrene oxidation in reactions including 10 lM FAD and 100 lM cstyrene or 20 lM styrene and 100 lM dFAD.

Global fits through initial rate data from reactions of SMOB with various concentrations of FAD and NADH are plotted in Fig. 3B. The best-fitting steady-state kinetic parameters and their experimental uncertainties were: kcat = 3240 ± 650 lM s
1, KM (NADH) = 101 ± 53 lM, Ki (NADH) = 114 ± 41 lM, KM (FAD) = 86 ± 29 lM. Steady-state analysis of SMOA

Fig. 1. SDS–PAGE 4–20% acrylamide gradient gel of purified SMO components. The contents of each lane are as follows: Lane 1, molecular weight markers; lane 2, native SMOB after concentration and purification through a BioRad Uno-Q anion exchange column; lane 3, N-terminally histidine-tagged SMOB after resolution by Ni– NTA chromatography; lane 4, molecular weight markers; lane 5, native SMOA after partial purification by anion exchange chromatography; and lane 6, N-terminally histidine-tagged SMOA after purification by Ni–NTA chromatography.

A plot of reaction velocity of SMO as a function of styrene concentration is presented in Fig. 4A. SMOA concentration was 50-fold greater than SMOB and the flavin concentration was maintained at a low level so as to maximize the efficiency of coupling NADH oxidation to styrene consumption. Under these conditions, NADH/styrene ratio was found to decrease from approximately 1.7 in the low range of styrene concentration to 1.0 in the presence of saturating styrene. Best-fitting parameters gave an apparent KM for styrene of 4.6 ± 0.7 lM and Vmax of 200 ± 5 nM s
1. Benzene was determined to be a competitive inhibitor of styrene. Fig. 4B shows a fit of initial velocity data from the reaction of SMOA with styrene over a range of benzene concentrations. These data were fit according to the equation for simple substrate inhibition to calculate an estimate of the equilibrium dissociation constant for benzene. In this way, a dissociation constant of 173 ± 10 lM was determined. Pre-steady-state investigation of SMOA

Fig. 2. pH profile of the reaction of native SMOB with NADH and FAD. Apparent values of Vmax (s) and KM (h) determined at 25 C and pH values of 5.2, 5.6, 6.2, 6.7, 7.2, 7.6, 7.9, 8.3, and 9.2 were fit as described in the text. Solid lines represent the best fits through these data. Dashed line represents the Vmax/KM ratio calculated as a function of pH.

Reaction of reduced SMOA with oxygen was investigated by single-mixing stopped-flow experiments. In these studies, solutions containing equimolar concentrations of SMOA and FAD were first titrated in an anaerobic cuvette and then transferred to the stopped flow where they were rapidly mixed with aerobic buffer solutions. Titrations performed in the presence of a small amount of residual oxygen resulted in the formation of a stable intermediate spectrum with an absorbance maximum centered between 370 and 380 nm, as might be expected for a flavin hydroperoxide [39–41]. Under anaerobic conditions, this intermediate remained stable and showed no evidence of reduction by dithionite. Under aerobic conditions, the intermediate was observed to decompose. The rate of

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Fig. 3. Steady-state mechanism of SMOB reacting with NADH and FAD. (A) Product inhibition studies with NAD included as inhibitor of substrates NADH (h) and FAD (s). (B) Global fit of the bi-sequential model through initial velocity data from the reaction of 10 nM SMOB with NADH concentrations ranging from 20 to 350 lM and FAD concentrations ranging from 1 to 100 lM in 20 mM phosphate buffer at pH 7 and 25 C.

Fig. 4. Steady-state reaction of SMO with styrene and inhibition by benzene. Reaction of 0.5 lM N-terminally histidine-tagged SMOA and 10 nM native SMOB with 10 lM FAD, 50 lM NADH, and 270 lM oxygen, and styrene in 20 mM aerobic MOPSO buffer pH 7 at 25 C. (A) Michaelis– Menten fit through initial rate data from reactions, which included styrene at concentrations ranging from 5 to 100 lM. (B) Competitive inhibition fit through data from reactions, which included 15.6 lM styrene and benzene concentrations ranging from 20 to 500 lM.

decomposition was accelerated by the addition of styrene and inhibited by the addition of benzene. Kinetic data recorded at 314, 390, and 450 nm are shown in Figs. 5A–C. These data illustrate the different reaction kinetics, which occur in the oxidative reaction of SMOA with air in the presence or absence of styrene or benzene. Solid lines passing through the data in each plot represent the best three or four exponential fits through these data and the best-fitting rate constants corresponding to each fit are listed in Table 2. Extinction coefficients corresponding to the initial, intermediate, and final spectra involved in each reaction are given in Figs. 5D–F. Measurement of the Redox potential of FAD associated with SMOA Titration of N-terminally histidine-tagged SMOA in the presence of FAD and anthraquinone-2-sulfonate

resulted in the sequential reduction first of enzymebound FAD and then the indicator. It was clear from this result that flavin binding to the enzyme results in a significant positive shift in midpoint potential. However, the disparity in the anthraquinone-2-sulfonate and bound flavin potentials was too large to allow accurate determination of the bound FAD potential. Analysis of data from titrations in which anthraquinone-(1,5)disulfonate (Em7 =
174 mV) and indigo-(5,7)-disulfonate (Em7 =
125 mV) were used as solution potential indicators allowed us to establish the midpoint potential of bound FAD to be
149 ± 0.4 mV. Studies with a fourth indicator, indigotrisulfonate (Em7 =
81 mV), resulted in the complete reduction of the indicator prior to flavin reduction. Absorbance spectra resulting from the reduction of FAD bound to N-terminally histidine-tagged SMOA in the presence of indigodisulfonate are presented in Fig. 6A. Fits through the data used to calculate the potential of bound FAD and free FAD

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Fig. 5. Single turnover kinetics of the reaction of N-terminally histidine-tagged SMOA with oxygen. Reduced 25–30 lM N-terminally histidinetagged SMOA was reacted with 130 lM oxygen in 50% air-saturated buffer (d) and with mixtures of 130 lM oxygen and 500 lM styrene (s) or benzene (n). Absorbance data superimposed with exponential fits are shown at (A) 314 nm (B), 390 nm, and (C) 450 nm. Extinction coefficients corresponding to oxidized (m) and reduced (d) FAD–enzyme complexes and the first (s), second (j), and third (n) intermediates detected in the reaction of N-terminally histidine-tagged SMOA with (D) oxygen, (E) oxygen and styrene, and (F) oxygen and benzene.

Table 2 Best-fitting values of rate constants in the oxygen reaction of SMOA Reaction

k1obs (s
1)

k2obs (s
1) k3obs (s
1)

Air only 4.7 ± 0.3 1.1 ± 0.1 Air + benzene 5.0 ± 1.2 0.8 ± 0.2 Air + styrene (15 ± 4.3)a 1.0 ± 0.2

k4obs (s
1)

0.30 ± 0.01 0.082 ± 0.004 0.032 ± 0.001 0.006 ± 0.01 0.02 ± 0.01 —

Absorbance data were fit to sums according to Eq. (4). a This value may be significantly influenced by the 21 ms time constant associated with data averaging.

are shown in Fig. 6B. The oxidation–reduction potential of free FAD was estimated at
212 ± 0.5 mV by analyzing the results of a reductive titration conducted in the presence of the solution potential indicator, anthraquinone-2-sulfonate, at 25 C in 20 mM phosphate buffer (pH 7). There was no significant perturbation of the midpoint potentials of either FMN or riboflavin when they were reduced in the presence of N-terminally histidine-tagged SMOA. The ratio of the electron equilibrium dissociation constants calculated from the free FAD and SMOA-bound FAD equilibrium midpoint potentials indicates that N-terminally histidine-tagged SMOA binds reduced FAD 137 times more tightly than oxidized FAD.

Mechanism of reduced FAD transfer from SMOB to SMOA The oxidation rate of reduced FAD in aerobic solution is quite rapid and dependent on both the concentration of oxygen and oxidized FAD present in the reaction. Kinetic traces from reactions of free reduced FAD with air-saturated buffer containing oxidized FAD are shown overlaid with kinetic simulations of this reaction in Fig. 7. The rate constants and model of reduced flavin oxidation used in these simulations were based on the known rate constants and kinetic mechanism previously reported for this reaction [7]. Clearly the presence of oxygen and oxidized flavin will interfere with the efficiency of electron-transfer from SMOB to SMOA if this transfer is mediated by diffusion of reduced flavin. The direct transfer of reduced flavin from SMOB to the active site of SMOA through the formation of transient complex of SMOA and SMOB during catalysis would eliminate interference caused by oxygen and oxidized FAD with the efficient transfer of electrons in SMO. To better resolve the nature of reduced flavin transfer in this system, we measured the coupling of NADH oxidation to styrene oxide produc-

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Fig. 6. Equilibrium midpoint potential of FAD bound to N-terminally histidine-tagged SMOA. (A) Absorbance spectra from the dithionitemediated reductive titration of 70 lM N-terminally histidine-tagged SMOA in complex with 34.7 lM FAD at 25 C. The reaction was conducted in 20 mM MOPSO buffer, pH 7, containing 200 lM styrene and 35 lM indigodisulfonate. (B) Plots of extent of reduction as a function of solution potential used to verify equilibrium midpoint potentials of free FAD (d) and FAD bound to N-terminally histidine-tagged SMOA (s).

enzyme complex and the data were fit to a model describing uncompetitive substrate inhibition (Eq. (8)). Based on this fit, KM(app) (FAD) = 15 ± 8 lM and the inhibition constant for oxidized FAD is 37 ± 22 lM. v¼

Fig. 7. Reaction of free FAD with oxygen. Absorbance changes at 450 nm corresponding to reactions of 6.3 lM reduced FAD (lower trace) with 123 lM oxygen and 5.5 lM oxidized FAD and 8.3 lM reduced FAD (upper trace) with 123 lM oxygen and 3.5 lM oxidized FAD in 20 mM phosphate buffer, pH 7, at 23 C. Solid lines passing through the data represent simulations of each reaction.

tion as a function of FAD concentration. Results of these studies are shown in Fig. 8. The initial velocity data describing the rate of NADH oxidation and styrene oxidation were recorded simultaneously as described in Materials and methods. It is clear from inspection of these data and the NADH/styrene coupling ratio that the efficiency of reduced FAD transfer decreases with increasing FAD concentration. Apparent Vmax and KM values for NADH reacting with SMOB of 0.29 ± 0.1 lM s
1 and 4.8 ± 0.5 lM were calculated from a Michaelis–Menten fit through the initial rate data corresponding to the oxidation of NADH. The styrene oxidation kinetics were fit in two ways. In the first approach, SMOB and SMOA were treated as an

V maxðappÞ ½FADox   . ox  K MðappÞ þ ½FADox  1 þ ½FAD Ki

ð8Þ

In the second approach to fitting these data, reduced FAD transfer was considered to occur exclusively by flavin diffusion. In this mode of operation, reduced flavin is a substrate of SMOA and in order to calculate the apparent KM of reduced FAD for SMOA it was necessary to estimate the concentration of reduced FAD contributing to each measurement of initial rate of styrene oxidation. This was done by inputting the apparent Vmax and KM values for the reaction of SMOB with NADH into the integrated steady-state rate expression (Eq. (9)).   K NADH ½NADHo ½NADHo
½NADHt MðappÞ ln þ V maxðappÞ ½NADHt V maxðappÞ ¼ t.

ð9Þ

The average concentration of reduced FAD generated over a 10-s time interval was determined in this way for each of the reactions of oxidized FAD with SMOB. The NADH/styrene coupling ratio calculated for each reaction was then used to correct for the fraction of reduced FAD consumed prior to interaction with SMOA. In this way, we obtained estimates of the reduced FAD concentration present in each of the initial rate determinations for the reaction of SMOA with styrene. The program GraphPad Prism 4 was then used to fit the styrene concentration dependence data in Fig. 8 to a simple product inhibition model (Eq. (10)).

A. Kantz et al. / Archives of Biochemistry and Biophysics 442 (2005) 102–116

Fig. 8. Coupling of NADH and styrene oxidation as a function of FAD concentration. Initial rates of NADH (h) and styrene (s) oxidation superimposed with fits providing the best agreement with the data as described in the text. Simulations of the experimentally determined NADH/styrene coupling ratio (j) according to Model A (dashed line) and Model B (dotted line) described in the text.

v¼

V ½FAD   maxðappÞ  red . ½FADox  K MðappÞ 1 þ K i þ ½FADred 

ð10Þ

In these fits, reduced FAD was treated as substrate and oxidized FAD as competitive inhibitor. The best estimates of the fit parameters were KM(app) (reduced FAD) = 1.3 ± 0.3 lM and Ki (oxidized FAD) = 96 ± 68 lM. The equilibrium dissociation constant of reduced FAD is calculated from the Ki and the observed shift in equilibrium midpoint potential of FAD upon binding to SMOA was found to be 0.7 lM, which is in close agreement with apparent KM for reduced FAD. The coupling of NADH use to styrene oxidation was further studied as a function of SMOA/SMOB ratio. In this investigation, the concentration of SMOA was varied at fixed SMOB, FAD, NADH, and oxygen concentration to yield SMOA:SMOB ratios varying from 1:1 to 500:1. The observed initial rates of NADH and styrene oxidation are displayed in Fig. 8. A fit through the styrene oxidation data in accordance with the Michaelis–Menten equation provides apparent values of Vmax and KM of 0.095 ± 0.007 lM s
1 and 0.68 ± 0.16 lM. Two models describing the dependence of SMO reaction on FAD concentration and enzyme ratio were investigated by numerical simulation (Scheme 1). Model A characterized by reactions 1–3 and 5 in Scheme 1 represents flavin transfer from SMOB to SMOA by free diffusion. Model B is identical to Model A except that it also includes Scheme 1 reaction number 4. This model represents a mechanism in which both direct and diffusive flavin transfer are significant features of catalysis. As illustrated, each model includes the synthesis of reduced FAD by SMOB and the parallel pathways of fla-

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vin reoxidation by either collision and reaction of freely diffusing reduced flavin with molecular oxygen or the association of reduced flavin with SMOA followed its reoxidation by the styrene epoxidation. Each model also includes substrate inhibition of SMOA by oxidized FAD in solution. Parameters used in modeling these reactions were as follows: the rate constants used to calculate the rate of the reaction of reduced flavin with oxygen and oxidized flavin were based on the mechanism and values reported in the literature [7]. The complex set of reactions involved in the reoxidation of free FAD is represented by k* in step 5 of Scheme 1. The relative amounts of superoxide and hydrogen peroxide generated in this reaction depend on the initial concentrations of oxygen, oxidized and reduced flavin. The rate of reduced flavin production by SMOB was based on our values of the apparent Vmax and KM of FAD corresponding to the concentrations of oxidized FAD, NADH, and oxygen present in the reaction. The rate of styrene oxidation by free SMOA was modeled by using the calculated Ki of oxidized FAD and apparent Vmax and KM values of reduced FAD were estimated from fits through the flavin concentration dependence of the reaction rate. The results from simulations of the FAD and SMOA concentration dependence data according to Model A are compared with the data in Fig. 8. The model correctly predicts the general trends of increased substrate inhibition of SMOA and decreased efficiency of coupling NADH consumption to styrene oxidation as a function of increasing FAD concentration. Model A also correctly predicts the experimentally observed trend of increased coupling efficiency with increasing SMOA/ SMOB ratio. Although Model A correctly predicts the

Fig. 9. Coupling of NADH and styrene oxidation as a function of Nterminally histidine-tagged SMOA/SMOB ratio. (A) Initial rates of NADH (h) and styrene (s) oxidation catalyzed by various ratios of SMOB and N-terminally histidine-tagged SMOA plotted together with (d) the NADH/styrene coupling ratio. Simulations of the experimentally determined NADH/styrene coupling ratio according to Model A (dashed line) and Model B (dotted line).

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Scheme 1.

general trends observed in the experimental data, the estimates of coupling efficiency calculated from the simulation are inconsistent with the experimentally observed values. At high FAD concentrations, the reaction is more than two orders of magnitude more coupled than predicted by the simulation of the SMO reaction by Model A (Fig. 9). At low SMOA/SMOB ratios, the experimentally observed coupling of NADH to styrene oxidation is about an order of magnitude greater than that calculated from the Model A simulation (Fig. 8). In Model B, the channeling of FAD from SMOB to SMOA through the formation of a catalytic complex and the direct transfer of reduced flavin from SMOB to SMOA is predicted to decrease the fraction of reduced flavin released in solution and the rate of free-flavin oxidation by reaction with molecular oxygen. To study this model, it was necessary to estimate the magnitude of the dissociation equilibrium constant of the complex formed by SMOA and SMOB and the catalytic rate of styrene oxidation by this complex. The sensitivity of the coupling efficiency to the magnitude of equilibrium dissociation constant of the complex formed between SMOA and SMOB during catalysis was estimated by comparing the experimental data to the results of a series of simulations in which this value was varied in decades from 10 nM to 10 lM. A similar approach was taken in estimating the catalytic rate of styrene epoxidation by the SMOA–SMOB complex. Based on this evaluation, parameters giving the best agreement with the experimentally determined data were an SMOA–SMOB dissociation constant of 100 nM and observed catalytic rate constant of 10 s
1 for the styrene oxidation reaction. The results of this Model B simulation provided in Figs. 8 and 9 are in better agreement with the experimental data than those derived from the Model A. In

particular, the coupling efficiency predicted by Model B is very close to the observed coupling efficiency of the SMO system over the full range of FAD and enzyme concentrations investigated.

Discussion The primary sequences of the flavin-dependent styrene monooxygenases are highly conserved [31,32,42,43]. The epoxidase components isolated from Pseudomonas S12 and VLB strains differ by only two C-terminal residues and the flavin reductases of these systems have identical primary sequences [32]. Steady-state kinetic analysis of the reductase component from the VLB system was previously reported with FMN as substrate [15]. We find a different set of best-fitting kinetic parameters in our analysis of the reaction mechanism with FAD as the flavin substrate but are otherwise in agreement regarding the sequential nature of the reduction reaction mechanism. The N-terminally histidine-tagged versions of SMOB and SMOA are more convenient to purify than the native enzymes and appear to provide excellent functional models for catalysis by the native enzyme components. As previously reported for the flavin reductase from the VLB system, we find that native SMOB expresses predominantly in the form of inclusion bodies, but this protein can be recovered in fully active form [15]. The N-terminally histidine-tagged version of SMOB similarly expresses primarily as insoluble protein and neither decreasing the temperature nor the concentration of inducer during expression significantly improves the partitioning of SMOB into the soluble phase. However, we were able to recover soluble highly purified N-terminally histidine-tagged SMOB constructs by Niaffinity chromatography. The steady-state reactions of

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native or N-terminally histidine-tagged SMOB with substrates and native or N-terminally tagged SMOA are very similar. Based on homology modeling through the 3dPSSM server [44], the closest structural homology match for SMOB was an FMN-containing flavin reductase from Thermus thermophilis (Tahir Tahirov, unpublished results). Other closely related structures are those of FMN-dependent ferric ion and flavin reductases [45,46]. Homology modeling and model building studies of the reductase of phenol hydroxylase reductase from Bacillus thermoglucosidasius suggest that this enzyme may also be a close structural relative of SMOB [14]. SMOA was most closely matched to p-hydroxybenzoate hydroxylase and D-amino acid oxidase by structural homology modeling [47,48]. For this reason, we believe that the single-component flavin monooxygenases can provide valuable structural and mechanistic insight into our studies of the oxidative half-reaction of SMO. Reductive titrations of the SMOA–FAD complex in the presence of 10–20 lM oxygen result in the formation of an extremely stable flavin intermediate. This species has an absorbance spectrum maximizing between 375 and 380 nm, which is consistent with that of a C4a-flavin hydroperoxide. Interestingly, the intensity of this spectrum was not diminished by the addition of excess dithionite. Upon mixing with styrene or excess oxygen this species rapidly returns to that of oxidized flavin. No oxygen intermediates were stabilized when the SMOA– FAD complex was titrated with dithionite in the presence of 10–20 lM oxygen and 200 lM styrene. In this case, the equilibrium absorbance spectra recorded were representative of mixtures of oxidized and two-electron reduced FAD. In our rapid mixing studies of the reaction of SMOA with oxygen, we observe the formation of the putative flavin hydroperoxide intermediate at an observed rate constant of 4.7 s
1 and calculate an apparent extinction coefficient of 11 mM
1 cm
1 at 380 nm for this species. In the presence of oxygen, this intermediate is transformed at an observed rate constant of 1 s
1 to a second species with diminished extinction coefficient of 8 mM
1 cm
1. Based on the difference spectrum resulting from subtraction of the spectrum of the first intermediate from that of the second, we conclude that approximately 20% of the bound FAD is reoxidized in this phase of the reaction. Approximately 90% of the bound FAD returns to the oxidized form in a third step with an observed rate of 0.3 s
1. Enzyme-bound FAD is fully reoxidized in a final step with an observed rate constant of 0.08 s
1. In the presence of the substrate analog benzene, the first intermediate again has an absorbance maximum in the 375–380 nm region and this species forms and decays with observed rate constants, which are very similar

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to those found to provide the best fit in the reaction of SMOA with oxygen in the absence of benzene. In this case, the first observed intermediate is spectrally less intense, having an extinction coefficient of only 8 mM
1 cm
1 at 380 nm. The enzyme returns to oxidized flavin with observed rate constants that are an order of magnitude less than those observed in the absence of benzene. The rate of hydrolysis of the C4a-hydroxyflavin has been shown to be sensitive to active site occupancy by substrate and products [49,50]. It seems likely that the slow reoxidation of SMOA in the presence of benzene may similarly block the dissociation of hydrogen peroxide and regeneration of oxidized flavin in the active site of this enzyme. When reduced SMOA is reacted with styrene, we observe the rapid formation of an intermediate with an apparent molar extinction of 8 mM
1 cm
1 and absorbance maximum centered at 370 nm. This intermediate forms at an observed rate of 15 s
1 then decays to yield an oxidized FAD spectrum in a biphasic reaction. Each decay phase results in the formation of oxidized flavin. The first phase, which accounts for 90% of the increase at 450 nm, occurs with an observed rate constant of 0.95 s
1. A final increase in absorbance at 450 nm, which returns the fully oxidized FAD spectrum, occurs at 0.02 s
1. In this experiment, the rate of styrene oxidation may have been limited by the reaction of oxygen with the reduced FAD–SMOA. In this case, the C4a-hydroperoxide intermediate would not accumulate and the first observed intermediate may be a C4a-hydroxyflavin. The present data suggest the reactive intermediates employed in the oxidative half-reaction of SMOA are similar to those previously identified in related flavoenzyme monooxygenases as C4a-peroxy and hydroxyflavins. However, many details of this reaction remain to be resolved. It is possible that the reactive species is actually a flavin peroxide as has been established in the case of cyclohexanone monooxygenase [51]. Alternatively, the flavin hydroperoxide may collapse to form a reactive oxaziridine intermediate which then catalyzes the observed epoxidation [52] (Scheme 2). Our titration data indicate that the equilibrium midpoint potential of SMOA-bound FAD is shifted positively by 63 mV. This implies that the SMOA binds reduced FAD approximately 137 times more tightly than oxidized FAD (Scheme 3). Based on the best estimate of the FAD inhibition constant, we calculate the binding constant of reduced FAD to SMOA is in the range of 500 nM. We presently have no data identifying the structural basis of this interaction, but the calculated change in binding energy
36 kJ mol
1 is comparable with that which would be gained through the formation of a single hydrogen bond [53]. There was no evidence for the formation of a stable oxygen intermediate when either riboflavin or FMN was substituted for FAD. Moreover, the midpoint

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Scheme 2.

Scheme 3.

potentials of these FAD analogues were unaffected by SMOA based on reductive titrations, which included the solution potential indicators anthraquinone-2-sulfonate and indigodisulfonate. Our work supports the previous finding for the VLB system that whereas riboflavin, FMN, and FAD each function as reductase substrates only FAD is an effective substrate for the monooxygenase component. It was previously shown that the hydroxylase components of SMO (VLB) and the closely related 4-hydroxyphenylacetate-3-monooxygenase system are able to catalyze substrate oxygenation in the absence of their reductase components [12,21]. It was further shown through gel filtration and analytical ultracentrifugation studies that the reductase and monooxygenase components of these systems do not form stable complexes at equilibrium [15,20]. Additional studies of the 4-hydroxyphenylacetate-3monooxygenase system were conducted to test for evidence of a reductase–hydroxylase interaction during catalysis. In this work, the apparent KM of the reductase for FAD was measured and found to be uninfluenced by the hydroxylase component [20]. A significant increase or decrease in the apparent KM value would have provided evidence in support of a direct reductase–hydroxylase interaction. However, this apparent KM value depends on a ratio of microscopic rate constants, which may increase, decrease, or not be significantly affected by protein–protein interactions during catalysis, and the absence of a significant change in this value does not preclude the formation of a transient reductase–hydroxylase complex during catalysis.

In our studies of this problem in the SMO system, we developed a real-time assay, which allows us to simultaneously monitor the steady-state rates of styrene and NADH consumption. We find that within experimental uncertainty NADH and styrene are oxidized in a 1:1 ratio only when the SMOA:SMOB ratio is large and when FAD concentrations are kept low. Lower SMOA:SMOB ratios and elevated FAD concentrations are found to decrease the efficiency of this reaction. To seek evidence for the formation of a transient flavin-transfer complex during catalysis by the SMO system, we compared our experimental data with the results of numerical simulation of models, which included or excluded such a complex. Parameter estimates for these simulations were derived from both our experimental data and from rate constants reported in the literature for the reactions of free reduced flavin with oxygen. Based on this comparison, we find the observed steady-state coupling of NADH and styrene oxidation to be in best agreement with a reduced flavin-transfer mechanism in which SMOA and SMOB form a transient complex during catalysis. In this way, the reduced flavin is protected from counterproductive reactions with free oxygen and oxidized flavin in solution. Comparison of the data with numerical simulations indicates that whereas the diffusion of free reduced flavin from SMOB to SMOA is a significant mechanism of electron delivery in the SMO system it does not account for the observed reaction coupling efficiency over the broad range of FAD concentrations and SMOA/SMOB ratios investigated. Flavin dynamics have been shown to be critical feature in the catalysis of flavoenzyme monooxygenases [54]. In the two-component enzymes, this dynamic reaches an extreme in which the flavin is physically transferred from the reductase to hydroxylase. Since neither FMN nor riboflavin is able to substitute for FAD in the styrene epoxidation reaction, we propose that the AMP functionality represents a critical structural motif in the recognition and binding of FAD to SMOA during catalysis. Reduction of SMOA-bound FAD may be coupled to the movement of the isoalloxazine ring system from a solvent-exposed position to a sequestered position within the active site of the protein. This change of environment is consistent with both the observed positive shift in midpoint potential and the ability of the FAD to form a stable C4a-hydroperoxide. The isoalloxazine ring of the oxidized FAD may be sufficiently accessible such that SMOB can provide it with electrons in the reductive half-reaction while the AMP portion of FAD is still associated with SMOA. On a similar note, the AMP moiety of FAD may remain solvent exposed and accessible to SMOA during binding and reduction of the isoalloxazine ring of free-FAD by SMOB. Based on the present information we suggest that in the most efficient mode of catalysis by the

A. Kantz et al. / Archives of Biochemistry and Biophysics 442 (2005) 102–116

Scheme 4.

SMO system, SMOB may form a transient flavin-transfer complex with SMOA in which the AMP moiety is associated with FAD and isoalloxazine ring is associated with SMOB (Scheme 4). Release of a tethered isoalloxazine ring from the active site of SMOB would ensure its efficient transfer to the oxygenation active site of SMOA and minimize interaction with molecular oxygen and oxidized FAD in solution.

Acknowledgments We thank the UCSF DNA-sequencing core facility for assistance in the verification of our cloned gene sequences, Tom Franco in the San Francisco State University College of Science and Engineering Machine Shop for his help in constructing our stopped-flow instrument, Drs. Ishan Erden (San Francisco State University), David Ballou and Bruce Palfey (University of Michigan), and Tahir Tahirov (RIKEN Institute) for helpful discussions, NIH S06 GM52588 and for funding this research, and Robert Yen in the SFSU Mass Spectrometry Core Facility supported by NIH P20 MD000262, NCMHD for his help in verifying the molecular weight of SMOB.

References [1] L.M. Cunane, Z.W. Chen, R.C. Durley, J.D. Barton, F.S. Mathews, Biochem. Soc. Trans. 27 (1999) 179–184. [2] A.C. Stainthorpe, V. Lees, G.P. Salmond, H. Dalton, J.C. Murrell, Gene 91 (1990) 27–34. [3] E.L. Neidle, C. Hartnett, L.N. Ornston, A. Bairoch, M. Rekik, S. Harayama, J. Bacteriol. 173 (1991) 5385–5395. [4] B. Galan, E. Diaz, J.L. Garcia, Environ. Microbiol. 2 (2000) 687– 694. [5] O. Pylypenko, I. Schlichting, Annu. Rev. Biochem. 73 (2004) 991– 1018. [6] S.C. Tu, H.I. Mager, Photochem. Photobiol. 62 (1995) 615–624. [7] V. Massey, J. Biol. Chem. 269 (1994) 22459–22462. [8] B. Lei, Q. Ding, S.C. Tu, Biochemistry 43 (2004) 15975–15982. [9] U. Arunachalam, V. Massey, S.M. Miller, J. Biol. Chem. 269 (1994) 150–155.

115

[10] M.R. Gisi, L. Xun, J.Bacteriol. 185 (2003) 2786–2792. [11] P. Chaiyen, C. Suadee, P. Wilairat, Eur. J. Biochem. 268 (2001) 5550–5561. [12] B. Galan, E. Diaz, M.A. Prieto, J.L. Garcia, J. Bacteriol. 182 (2000) 627–636. [13] U. Arunachalam, V. Massey, C.S. Vaidyanathan, J. Biol. Chem. 267 (1992) 25848–25855. [14] U. Kirchner, A.H. Westphal, R. Muller, W.J. van Berkel, J. Biol. Chem. 278 (2003) 47545–47553. [15] K. Otto, K. Hofstetter, M. Rothlisberger, B. Witholt, A. Schmid, J. Bacteriol. 186 (2004) 5292–5302. [16] B. Gao, H.R. Ellis, Biochem. Biophys. Res. Commun. 331 (2005) 1137–1145. [17] F. Fieschi, V. Niviere, C. Frier, J.L. Decout, M. Fontecave, J. Biol. Chem. 270 (1995) 30392–30400. [18] C.E. Jeffers, J.C. Nichols, S.C. Tu, Biochemistry 42 (2003) 529– 534. [19] B. Lei, S.C. Tu, Biochemistry 37 (1998) 14623–14629. [20] T.M. Louie, X.S. Xie, L. Xun, Biochemistry 42 (2003) 7509–7517. [21] F. Hollmann, P.C. Lin, B. Witholt, A. Schmid, J. Am. Chem. Soc. 125 (2003) 8209–8217. [22] J.T. Groves, K.M. Fish, G.E. Avaria-Neisser, M. Imachi, R.L. Kuczkowski, Prog. Clin. Biol. Res. 274 (1988) 509–524. [23] P.C. Wilkins, H. Dalton, C.J. Samuel, J. Green, Eur. J. Biochem. 226 (1994) 555–560. [24] J. Green, H. Dalton, Biochem. J. 236 (1986) 155–162. [25] P. Liu, A. Liu, F. Yan, M.D. Wolfe, J.D. Lipscomb, H.W. Liu, Biochemistry 42 (2003) 11577–11586. [26] K. Buch, H. Stransky, A. Hager, FEBS Lett. 376 (1995) 45–48. [27] T.D. Porter, J. Biochem. Mol. Toxicol. 16 (2002) 311–316. [28] S. Hartmans, M.J. van der Werf, J.A. de Bont, Appl. Environ. Microbiol. 56 (1990) 1347–1351. [29] K. OÕConnor, C.M. Buckley, S. Hartmans, A.D. Dobson, Appl. Environ. Microbiol. 61 (1995) 544–548. [30] F.J. Weber, L.P. Ooijkaas, R.M. Schemen, S Hartmans, J.A.M. De Bont, Appl. Environ. Microbiol. 59 (1993) 3502–3504. [31] F. Beltrametti, A.M. Marconi, G. Bestetti, C. Colombo, E. Galli, M. Ruzzi, E. Zennaro, Appl. Environ. Microbiol. 63 (1997) 2232– 2239. [32] S. Panke, B. Witholt, A. Schmid, M.G. Wubbolts, Appl. Environ. Microbiol. 64 (1998) 2032–2043. [33] C. Hoogland, E. Gasteiger, A. Gattiker, S. Duvaud, M.R. Wilkins, R.D. Appel, A. Bairoch, Protein Identification and Analysis Tools on the ExPASy Server, Humana Press, Totowa, NJ, 2005. [34] H. Motulsky, A. Christopoulos, Fitting Models to Biological Data Using Linear and Nonlinear Regression, Oxford University Press, New York, 2004. [35] D.O. Lambeth, G. Palmer, J. Biol. Chem. 248 (1973) 6095–6103. [36] D.H. Wachsstock, T.D. Pollard, Biophys. J. 67 (1994) 1260–1273. [37] B.A. Barshop, R.F. Wrenn, C. Frieden, Anal. Biochem. 130 (1983) 134–145. [38] W.M. Clark, Oxidation–Reduction Potentials of Organic Systems, Williams and Wilkins, Baltimore, MD, 1960. [39] J.W. Hastings, C. Balny, C. Le Peuch, P. Douzou, Proc. Natl. Acad. Sci. USA 70 (1973) 3468–3472. [40] B. Entsch, D.P. Ballou, V. Massey, J. Biol. Chem. 251 (1976) 2550–2563. [41] K.C. Jones, D.P. Ballou, J. Biol. Chem. 261 (1986) 2553–2559. [42] A. Velasco, S. Alonso, J.L. Garcia, J.L. Garcia, J. Perera, E. Diaz, J. Bacteriol. 180 (1998) 1063–1071. [43] N.D. OÕLeary, K.E. OÕConnor, W. Duetz, A.D.W. Dobson, Microbiology 147 (2001) 973–979. [44] L.A. Kelley, R.M. MacCallum, M.J. Sternberg, J. Mol. Biol. 299 (2000) 499–520. [45] J.J. Tanner, B. Lei, S.C. Tu, K.L. Krause, Biochemistry 35 (1996) 13531–13539.

116

A. Kantz et al. / Archives of Biochemistry and Biophysics 442 (2005) 102–116

[46] H.J. Chiu, E. Johnson, I. Schroder, D.C. Rees, Structure (Camb.) 9 (2001) 311–319. [47] H.A. Schreuder, J.M. van der Laan, W.G. Hol, J. Drenth, J. Mol. Biol. 199 (1988) 637–648. [48] H. Mizutani, I. Miyahara, K. Hirotsu, Y. Nishina, K. Shiga, C. Setoyama, R. Miura, J. Biochem. (Tokyo) 120 (1996) 14–17. [49] M.G. Taylor, V. Massey, J. Biol. Chem. 265 (1990) 13687–13694. [50] M.G. Taylor, V. Massey, J. Biol. Chem. 266 (1991) 8291–8301.

[51] D. Sheng, D.P. Ballou, V. Massey, Biochemistry 40 (2001) 11156– 11567. [52] W. Adam, C.R. Saha-Moller, P.A. Ganeshpure, Chem. Rev. 101 (2001) 3499–3548. [53] T.W. Martin, Z.S. Derewenda, Nat. Struct. Biol. 6 (1999) 403– 406. [54] B. Entsch, L.J. Cole, D.P. Ballou, Arch. Biochem. Biophys. 433 (2005) 297–311.