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Department of Botany and Plant Pathology, 2082 Cordley Hall, Oregon State University, Corvallis, OR 97331-2902, USA
Correspondence
Daniel J. Arp
arpd{at}science.oregonstate.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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80 % of the hydroxylations occurring at the terminal carbon atom.
A supplementary table summarizing the purification of sBMO components is available with the online version of this paper.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Ashraf et al., 1994). The best characterized of these enzymes are soluble methane monooxygenase (sMMO) (Kopp & Lippard, 2002; Lipscomb, 1994; Wallar & Lipscomb, 1996) and the integral membrane monooxygenase, AlkB (Shanklin et al., 1997; van Beilen et al., 2005). Regiospecificity is not an issue for methane as it is the only physiologically relevant substrate of sMMO. However, AlkB is highly regiospecific for hydroxylation at the terminal carbon atom of C5–C16 alkanes. Recently, a family of cytoplasmic cytochrome P450 enzymes (the CYP153 family) has been identified and a few members from Mycobacterium sp. HXN-1500 and Alcanivorax borkumensis characterized (Funhoff et al., 2006; van Beilen et al., 2006). Studies of these enzyme systems constitute the bulk of the knowledge of the first step in alkane metabolism in prokaryotes.
Growth on C2–C4 gaseous alkanes is largely attributed to the Corynebacterium-Nocardia-Mycobacterium-Rhodococcus complex of Gram-positive bacteria (Ashraf et al., 1994; McLee et al., 1972; Perry, 1980). An exception to this association is the Gram-negative β-proteobacterium ‘Pseudomonas butanovora’ (ATCC 43655). Although originally identified as a pseudomonad, 16S rDNA analysis of ‘P. butanovora’ has shown that phylogenetically it is a member of the Rhodocyclus group, with the closest relatives belonging to the genus Thauera (Anzai et al., 2000). ‘P. butanovora’ was isolated from activated sludge sampled from an oil refining plant and was capable of aerobic growth on the linear alkanes C2–C9 (Takahashi et al., 1980). The same enzyme system was used to oxidize both the gaseous and liquid alkanes (Hamamura et al., 1999; Sluis et al., 2002). The downstream metabolism, in the case of butane, was determined to be via the terminal alcohol 1-butanol, with subsequent oxidations leading to butyraldehyde and butyrate. Presumably butyrate was shunted to the tricarboxylic acid cycle via the β-oxidation pathway (Arp, 1999). Previous work, through partial enzyme purification and insertional mutagenesis, established the genetic identity of a soluble three-component diiron monooxygenase, termed soluble butane monooxygenase (sBMO) and confirmed the role of sBMO in alkane oxidation (Sluis et al., 2002). That study also demonstrated the similarities of the bmo operon structure and the deduced amino acid sequence identities of the bmo genes to sMMO operons and gene products of the methanotrophs Methylosinus trichosporium OB3b and Methylococcus capsulatus Bath. Several detailed in vivo studies have focused on cometabolism of environmentally important pollutants by ‘P. butanovora’ for the purpose of bioremediation (Doughty et al., 2005; Halsey et al., 2005; Hamamura et al., 1997), the regulation of the bmo operon (Doughty et al., 2006; Sayavedra-Soto et al., 2001, 2005) and altering the regiospecificity of sBMO by site-directed mutagenesis of the hydroxylase 
-subunit (Halsey et al., 2006).
This study presents the first report of the purification of all the components of a soluble three-component diiron monooxygenase, with physiologically relevant activity, that was responsible for the oxidation of physiologically relevant C2–C9 alkanes as sources of carbon and energy. Physical characteristics and the roles for each of the subunits in catalysis are described. Product distribution studies revealed the strong bias that sBMO exhibits for terminal hydroxylation of C3–C6 linear and branched alkanes. Comparisons were made between the properties of sBMO and those of other soluble multicomponent diiron monooxygenases, and the implication of these findings for the catalytic mechanism of sBMO are discussed.
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METHODS |
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Bacterial cultivation.
‘P. butanovora’ (ATCC 43655) was cultivated at 30 °C in liquid medium consisting of 36 mM KH2PO4, 14 mM K2HPO4, 2 mM MgSO4.7H2O, 0.4 mM CaCl2.2H2O, 30 mM KNO3, 2 µM Na2MoO4.2H2O, 1 ml trace element solution (Wiegant & deBont, 1980), 10 ml vitamin solution (Lidstrom, 1988) and 500 µM FeSO4.7H2O at pH 7.2. Butane was supplied as an overpressure 7 % (v/v) in the headspace. Batch cultures were grown in a sealed 20 l glass carboy at 30 °C. Medium (15 l) was inoculated with 2 l overnight culture. The culture was constantly stirred and sparged with butane added as an overpressure to 7–10 % (v/v) via a vacuum-pressure pump with a 125 l Nalgene carboy serving as headspace. Cells were harvested (OD600 1.0–1.5) by centrifugation (7000 g for 30 min at 10 °C), suspended in 25 mM PIPES (pH 7.2), and centrifuged again. The cell paste was frozen in liquid N2 and stored at –70 °C. A yield of 2–3 g cells (l culture)–1 was typical.
Preparation of cell-free extract.
Approximately 60 g ‘P. butanovora’ cell paste was suspended 1 : 1 (w/v) in 25 mM PIPES (pH 7.2), 1 mM DTT, 5 % (v/v)glycerol and 200 µM Fe(NH4)2(SO4)2.6H2O (buffer A). The cells were disrupted by sonication by the method of Fox et al. (1990). The sonicated suspension was centrifuged at 10 000 g for 15 min at 4 °C. The supernatant was removed by careful decanting, diluted to 200 ml with buffer A and centrifuged at 150 000 g for 2 h at 4 °C. The supernatant was decanted, adjusted to pH 7.2 and designated the cell-free extract.
Separation of soluble butane monooxygenase components.
Unless otherwise noted, the following purification steps for each of the components were carried out at 4 °C. ‘P. butanovora’ cell-free extract was loaded onto a DEAE-Sepharose FF column (5.2x15 cm) equilibrated with buffer A at a linear flow rate of 28 cm h–1. The column was washed with two column volumes of buffer A, followed by a 1.2 l gradient of 0–0.4 M KCl in buffer A at a linear flow rate of 14 cm h–1. Fractions containing the monooxygenase components eluted in the following order and KCl concentrations, BMOH and BMOB (0.23–0.27 M) and BMOR (0.34–0.39 M).
Purification of BMOH.
Fractions containing BMOH and BMOB were pooled, brought to 25 % saturation with solid (NH4)2SO4 and stirred at 20 °C for 1 h. The solution was centrifuged at 10 000 g for 30 min and the supernatant decanted. Solid (NH4)2SO4 was added to the supernatant to 75 % saturation and stirred as above. After centrifugation at 10 000 g for 30 min, the pellet was suspended in a minimal volume of 25 mM PIPES (pH 7.2), 1 mM DTT, 5 % (v/v) glycerol, 100 µM Fe(NH4)2(SO4)2.6H2O and 1 M KCl (buffer B). This fractionated solution was loaded onto a Sephacryl S-300 HR column (3.2x90 cm) equilibrated with buffer B at a linear flow rate of 15 cm h–1. Fractions containing BMOH were pooled, concentrated and dialysed in 25 mM PIPES (pH 7.2), 1 mM DTT, 5 % (v/v) glycerol and 100 µM Fe(NH4)2(SO4)2.6H2O via ultrafiltration, frozen in liquid N2 and stored at –70 °C.
Purification of BMOB.
Pooled fractions containing BMOB from the Sephacryl S-300 HR column were concentrated and dialysed via ultrafiltration in 25 mM PIPES (pH 7.2) and 0.2 M KCl. The concentrated protein solution was applied to a Q Sepharose FF column (2.2x11 cm) equilibrated in the same buffer at a linear flow rate of 30 cm h–1. The small component was eluted with a 0.2 l gradient from 0.2 to 0.35 M KCl at a linear flow rate of 15 cm h–1. Fractions containing BMOB were pooled, concentrated and dialysed in 25 mM PIPES (pH 7.2) and 0.15 M KCl via ultrafiltration. This solution was applied to a Sephadex 75 FF column (1.6x25 cm) equilibrated in the same buffer at a linear flow rate of 15 cm h–1. BMOB fractions were pooled, concentrated and dialysed via ultrafiltration in 25 mM PIPES (pH 7.2), frozen in liquid N2 and stored at –70 °C.
Purification of BMOR.
BMOR fractions were pooled and diluted with two volumes of 25 mM PIPES (pH 7.2) and 5 mM sodium thioglycollate. This solution was applied to a Q Sepharose FF column (2.2x11 cm) equilibrated in the above buffer at a linear flow rate of 30 cm h–1. BMOR was eluted from the column with a 0.2 l gradient from 0.25–0.4 M KCl at a linear flow rate of 15 cm h–1. Fractions containing BMOR were pooled, concentrated via ultrafiltration and applied to a Sephadex 75 FF column (1.6x25 cm). The column was equilibrated in 25 mM PIPES (pH 7.2), 0.15 M KCl and 5 mM sodium thioglycollate and developed at a linear flow rate of 15 cm h–1. Fractions containing reductase were pooled, concentrated and dialysed via ultrafiltration in 25 mM PIPES (pH 7.2) containing 5 mM sodium thioglycollate, frozen in liquid N2 and stored at –70 °C.
NADH-coupled enzyme assays.
Soluble butane monooxygenase activity was assayed by following the epoxidation of ethene to ethene oxide by gas chromatography (Hamamura et al., 1997). BMOH (1 µM), BMOB (0.1–4 µM) and BMOR (0.2–8 µM) were placed in 25 mM PIPES (pH 7.2) in a total volume of 1 ml in a 10 ml reaction vial and sealed with a butyl rubber septum. Catalase, when present, was added to 4000 units per reaction. BMOR, when necessary, was added to 10 or 100 mM H2O2 in the above buffer and incubated at 30 °C for 30 min with shaking prior to addition of catalase and other components. Ethene (2 ml) was added as an overpressure to the headspace and allowed to equilibrate at 30 °C for 5 min. The reaction was started by the addition of NADH to 5 mM. Ethene oxide formation was analysed from 0.1 ml reaction vial headspace. Reactions to assess alcohol production contained optimized concentrations of monooxygenase components in 0.5 ml total volume. Gaseous alkanes were added as an overpressure to the headspace (3 ml). Liquid alkanes were added as neat liquid (5–10 µl). After addition of alkane the reaction was placed at 30 °C for 5 min and initiated by addition of NADH to 5 mM. The reaction was allowed to proceed for 5 min. The formation of alcohols was analysed by gas chromatography on a 0.2x180 cm 0.2 % Carbowax 1500 on Graphpac GC 80/100 column by injection of 1 µl of the liquid phase.
H2O2-coupled enzyme assays.
BMOH (200 µM) was placed in a 10 ml reaction vial in 25 mM PIPES, pH 7.2, and sealed with a butyl rubber septum. The headspace was subjected to evacuation and replacement with N2 for 3 cycles. Ethene (3 ml) was added to the headspace as an overpressure and allowed to equilibrate at 30 °C for 5 min. The reaction was initiated by the addition of H2O2. Ethene oxide was analysed by GC as described above and H2O2 was measured by the method of Zhang & Lipscomb (2006). Assays of recoverable activity were carried out as described above for NADH-coupled ethene oxidation with optimal concentration of BMOB and BMOR in the presence of catalase.
Other methods.
Denaturing SDS-PAGE was performed as described by Laemmli (1970). Protein concentrations were determined by the method of Bradford (1976) as modified by Nelson et al. (1982). Optical spectra were recorded with a Beckman DU 640 spectrophotometer. Sedimentation equilibrium experiments were carried out with a Beckman Optima XL-A analytical ultracentrifuge equipped with a Beckman An-60 Ti rotor. The reference and sample cell assemblies were monitored at a wavelength of 280 nm with a rotor speed of 22 000 r.p.m. at 20 °C. N-terminal amino acid sequence analysis was performed by Edman degradation with an Applied Biosystems 492 Procise Protein Sequencer coupled with a model 140C analyser by the Protein Facility at Iowa State University. Iron content was determined spectrophotometrically by the method of Percival (1991). H2O2 reduction was assayed with a Clark-style electrode measuring the evolution of oxygen as previously described (Arp, 1999).
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RESULTS |
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). Purified sBMO components are shown by denaturing SDS-PAGE in Fig. 1 and the UV-visible spectra are shown in Fig. 2. A table summarizing the purification of sBMO components is available as supplementary data with the online version of this paper.
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), and 40.2 kDa by analytical ultracentrifugation. Absorption maxima observed in the UV–visible spectrum were 270, 340, 400 and 458 nm (Fig. 2
) and are representative of a flavin and a [2Fe–2S] cluster as has been observed for the reductases of other multiple component monooxygenase systems (Colby & Dalton, 1979; Newman & Wackett, 1995; Patel, 1987). As purified, BMOR showed an A270/458 ratio of 3.1 and an A458/340 ratio of 1.4. These values were also consistent with other highly purified flavo-iron sulfur proteins (Fox et al., 1989). BMOR displayed a specific activity of 115 µmol min–1 (mg protein)–1 when catalysing the reduction of 2,6-dichlorophenol indophenol with a 20-fold purification of this activity from the cell-free extract.
BMOB.
Previous work on this system reported the requirement of a third component in addition to BMOR and BMOH. This third component, presumably BMOB, was found to elute ahead of BMOH during anion-exchange chromatography, but no peptide was found to be enriched in fractions containing this component (Sluis et al., 2002). In this work, BMOB was found to co-elute with BMOH and could be separated by including 1 M KCl during the gel-filtration step of the purification. The identity of BMOB was verified by Edman degradation analysis. Denaturing SDS-PAGE of BMOB revealed an apparent molecular mass of 22 kDa, an unexpected result considering the predicted mass of 15.1 kDa from the deduced amino acid sequence (Sluis et al., 2002). No optical absorbance maxima were observed above 300 nm (Fig. 2), a feature shared by all characterized small components of multicomponent monooxygenase systems (Fox et al., 1989; Green & Dalton, 1985; Newman & Wackett, 1995).
BMOH.
The migration of BMOH during gel filtrations indicated a molecular mass between 200 and 260 kDa and, by denaturing SDS-PAGE (Fig. 1), it consisted of three proteins in stoichiometric amounts exhibiting apparent molecular masses of 54, 43 and 25 kDa as previously observed (Sluis et al., 2002). This result suggests that, like other multicomponent monooxygenase systems, the holoenzyme has an (β
)2 quaternary structure and is consistent with the deduced amino acid sequences of the three subunits of the butane monooxygenase operon. BMOH showed an absorption maximum at 280 nm with no other significant features above 300 nm. This spectrum indicated no contamination by haem or iron–sulfur cluster prosthetic groups of other proteins (e.g. BMOR) in the purified preparations. BMOH was judged pure by denaturing SDS-PAGE and the absence of UV–visible spectral characteristics above 300 nm (Fig. 2). Iron content analysis of BMOH preparations, determined colorimetrically, revealed between 2.8 and 3.6 mol iron (mol BMOH)–1. This iron content translates to 1.4–1.8 diiron clusters per holoenzyme. BMOH specific activities for the epoxidation of ethene in the presence of catalase (see below) were 500–600 nmol min–1 (mg protein)–1.
Catalytic properties
Hydroxylation of substrates under multiple turnover conditions coupled to NADH oxidation required only the BMOR and BMOH components. Maximal activity was reached at a ratio of BMOR to BMOH of 2 : 1 (mol/mol). The addition of BMOB to assays containing BMOR and BMOH resulted in modest increases in rate, with a maximum 20 % increase in activity achieved between a ratio of 1.5–2 (mol/mol) BMOB to BMOH. The ratio of BMOB to BMOH that allowed for maximal activity varied between preparations of BMOH. Higher ratios of BMOB resulted in a decrease in the rate of substrate oxidation (data not shown). This characteristic was similar to other small components of multicomponent monooxygenase systems, where rate inhibition occurs when the small component to hydroxylase ratio exceeds 2 (Cadieux et al., 2002; Fox et al., 1989; Green & Dalton, 1985).
The rate of the reaction and yield for ethene oxidation were dependent on the presence of catalase in the reaction. Table 1 shows the results of NADH-coupled ethene oxidation, where NADH (462 µM) was limiting in the reaction for the purpose of assessing yield. In the absence of catalase, the coupling of NADH to ethene oxidation was 40–50 % lower, with the majority of the yield being produced in the first 3 min, and resulted in complete inactivation of ethene oxide production. The addition of catalase resulted in a twofold increase in the rate and 2.5–3-fold increase in the yield for ethene oxidation. The uncoupling of steady-state NADH turnover was shown to produce H2O2 during the hydroxylation of methane by sMMO (Zhang & Lipscomb, 2006). For sBMO, this was also the case in the presence of butane. H2O2 was produced by the reaction of NADH oxidation to the reduction of O2 catalysed by BMOR with a yield of 185±9 µM H2O2. When BMOH was present the amount of H2O2 at the end of the reaction was 16±2 µM, approximately 10 % of that with BMOR alone. Incubation of BMOR with 10 or 100 mM H2O2 for 30 min resulted in ethene oxide production rates that were 74 and 46 %, respectively, of those for BMOR incubated without addition of H2O2 but did not result in complete inactivation of ethene oxide production as shown in Table 1. Treatment of BMOR with 10 mM H2O2 and 5 mM NADH also resulted in decreased ethene oxide production rates but not complete inactivation of this activity. Addition of catalase and fresh BMOR to a reaction without catalase (Table 1) did not result in the restoration of ethene oxide production and showed that BMOH was completely inactivated during turnover in the absence of catalase. The catalase activity of BMOH (see below) made the determination of H2O2 yield difficult and thus the values shown should be viewed as a minimum. The presence of BMOB in the reaction had no statistically measurable effect on the yield of H2O2 generated (19±2 µM).
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; Fox et al., 1989; Green & Dalton, 1985; Newman & Wackett, 1995). BMOR and BMOH alone account for 80 % of the activity and 90 % product yield during NADH-coupled multiple turnover in the presence of catalase.
H2O2-dependent BMOH activities
The hydroxylase subunits of sMMO and toluene 2-monooxygenase (T2MO) of Burkholderia cepacia G4 were shown to form reactive oxygenating species from H2O2 resulting in multiple substrate turnover in the absence of O2 (Andersson et al., 1991; Froland et al., 1992; Newman & Wackett, 1995). BMOH was also capable of H2O2-dependent substrate turnover, also known as the peroxide shunt reaction. The results demonstrate the dependence of H2O2 concentration on the production of ethene oxide and the loss of recoverable BMOH activity in the initial absence of O2 (Table 2). Reactions containing only BMOR, BMOB, or acetylene-inactivated BMOH did not result in the formation of ethene oxide and no detectable loss of activity was observed when BMOH was incubated without H2O2. The product yield of the peroxidation reaction was three orders of magnitude lower compared to sMMO. A H2O2 utilization to product ratio (100 mM H2O2 reaction) of approximately 3000 : 1 was observed. As the concentration of H2O2 was increased, the recoverable activity of BMOH decreased, but did not result in total inactivation. At the end of the reaction, bubbles were visibly present in the reaction vial, suggesting the evolution of O2 from H2O2. Experiments confirming that BMOH was catalysing the dismutation of H2O2 were carried out utilizing an O2 electrode. BMOH was observed to follow Michaelis–Menten kinetics for the catalase reaction. The following kinetic parameters were deduced from a Hanes plot of the data: Km(app)=110 mM, Vmax=5500 nmol O2 min–1 (mg hydroxylase)–1, kcat=46 s–1 and kcat/Km=417 s–1 (M)–1.
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; Chang et al., 1999; Fox et al., 1991; Shinohara et al., 1998). The amino acid sequence of BMOB contains one cysteine residue at position 86. The possibility of disulfide bridge formation between two monomers of BMOB was examined by non-reducing conditions during denaturing SDS-PAGE (Fig. 3). As purified, BMOB migrated at apparent molecular masses of 44 and 22 kDa. Upon reduction with 10 % 2-mercaptoethanol in the sample buffer, BMOB displayed a molecular mass of 22 kDa. Reduction of BMOB with equimolar DTT (prior to addition of non-reducing SDS-PAGE sample buffer) produced the same results as shown (Fig. 3). Samples of reduced BMOB by both methods behaved similarly during SDS-PAGE regardless of heat treatment of the sample before separation. These results showed that a disulfide bridge was formed between two BMOB molecules and the cysteine residue at position 86 was responsible for the dimerization. Reduced BMOB did not show any demonstrable differences in enzymic analyses from the as-purified preparation of BMOB.
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). The associated error in the product determinations was ±5 % of the values shown. Remarkable similarity existed in the oxidations of the linear alkanes, where 80 % or more of the product was observed to be hydroxylated at the terminal carbon. Propane and butane showed the same distributions, with the secondary alcohols, 2-propanol and 2-butanol, respectively, constituting the remaining observed product. The remaining products of pentane oxidation were the secondary alcohols 2-pentanol and 3-pentanol. The small amount of 3-pentanol was not reported in the oxidation of pentane by sMMO (Froland et al., 1992). The addition of the hydroxyl radical scavenger mannitol (50 mM) had no effect on the production of 3-pentanol or the product distribution of pentane oxidation in general. Unlike the sMMO systems, where the addition of MMOB drastically shifts the distribution for most alkanes towards primary hydroxylation (Froland et al., 1992), there was no significant difference in the shift of products observed when BMOB was present in the reaction.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). The in vitro activity for ethene oxidation for the complete monooxygenase system was at least four times the typical in vivo activity. The physical characteristics of the purified enzyme components of sBMO were similar to those of other soluble diiron three-component monooxygenases of this class of diiron monooxygenase. A summary of the reactions possible by sBMO components and multiple component combinations is presented in Table 4.
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10-fold increase in product yield, depending on the substrate being oxidized (Cadieux et al., 2002; Fox et al., 1989; Newman & Wackett, 1995; Wallar & Lipscomb, 1996). BMOH and BMOR were also strongly biased for the terminal hydroxylation of C3–C6 linear and branched alkanes, with no statistically significant difference in the product distributions when BMOB was present in the reaction. This result was in contrast to the sMMO small component protein, MMOB. In the absence of MMOB, the oxidation of propane and butane catalysed by the sMMO system was biased for the production of secondary alcohols. When MMOB was included in the reaction, a strong shift in the product distribution toward the primary alcohols was observed (Froland et al., 1992). For the longer linear alkanes, pentane and hexane, sMMO displayed a shift toward primary hydroxylation as the chain length increased and was independent of MMOB. The branched alkane isobutane showed a small shift towards primary hydroxylation with MMOB while with isopentane the shift was more pronounced (Froland et al., 1992).
In the sMMO system, MMOB concentrations <5 % of MMOH concentrations were responsible for the full effect of MMOB on product distributions during NADH-coupled substrate oxidation (Andersson et al., 1991; Fox et al., 1989; Froland et al., 1992). Small amounts of contaminating BMOB could explain why there were not large effects on the product distributions for sBMO. Because BMOB was not found in denaturing SDS-PAGE of BMOH preparations, this explanation was not likely for sBMO. Given that there were no dramatic effects attributed to BMOB in the oxidation of substrates, it was imperative to question whether BMOB was interacting with BMOH. The identity of BMOB, given the aberrant migration during denaturing SDS-PAGE, was not in question due to the positive identification by Edman degradation analysis. The copurification of BMOB and BMOH during anion-exchange chromotography and the requirement of high ionic strength needed to separate these two components during gel filtration implied a close association. Dimerization of BMOB due to disulfide-bridge formation between cysteine residues at position 86 was confirmed. Reduction of BMOB prior to inclusion in steady-state ethene turnover experiments displayed the same catalytic properties as the purified preparation. MMOB proteins from the methanotrophs Methylosinus trichosporium OB3b, Methylococcus capsulatus Bath and Methylocystis sp. strain M are also capable of dimer formation under certain conditions (Brandstetter et al., 1999; Chang et al., 1999; Fox et al., 1991; Shinohara et al., 1998). Replacing BMOB in NADH-coupled steady-state ethene turnover reactions with MMOB from Methylosinus trichosporium OB3b had similar modest effects on ethene turnover (data not shown). The G113N mutant strain of ‘P. butanovora’ contains an amino acid substitution in the -subunit of BMOH, which is in the area adjacent to or contributing to formation of the active site (Halsey et al., 2006). In vivo, ‘P. butanovora’ G113N displayed an altered regiospecificity from the wild-type. When butane was supplied as substrate, G113N preferentially hydroxylated the secondary carbon atom, with 2-butanol comprising 92 % of the product observed. If BMOB had a role in shifting hydroxylation towards the primary carbon, like MMOB in the sMMO system, such a large shift towards secondary carbon hydroxylation by G113N would not be expected and a more even distribution of products would be observed. The above discussion suggests the unique properties associated with MMOB in the sMMO system have evolved specifically for methane oxidation (Brazeau et al., 2003). Given the rate of ethene oxidation by BMOH and BMOR in the absence of BMOB, this study suggests that the role of the effector component, BMOB, may be unnecessary for the in vivo oxidation of C2–C6 aliphatic alkanes.
The inactivation of sBMO during steady-state turnover differed from the sMMO system, where inactivation occurred only in the absence of substrate (Fox et al., 1989). During NADH-coupled turnover, sBMO progressively lost activity, ending in total inactivation, and the presence of substrate provided no protection to this loss. The addition of catalase to the reaction led to a constant rate of product accumulation, which suggested that BMOH and/or BMOR was rapidly inactivated by small amounts of H2O2 produced by the uncoupled oxidation of NADH by BMOR in the presence of butane. Pretreatment of BMOR with H2O2 (with or without NADH) led to a reduction in the rate of ethene oxide production but not complete inactivation of ethene oxide production. Adding catalase and fresh BMOR to an in vitro reaction resulted in complete inactivation of ethene oxide production (Table 1, without catalase). Inactivation of ethene oxide production by H2O2 was observed to occur to BMOR and BMOH, though in the case of BMOH inactivation was complete. This observation was not unprecedented for a diiron monooxygenase hydroxylase. The related enzyme system, phenol hydroxylase of Pseudomonas sp. strain CF600, was also inactivated in the presence of phenol during NADH-coupled substrate turnover (Cadieux et al., 2002). In contrast, inactivation was not seen in the H2O2 shunt reaction, where BMOH in the presence of 10 mM H2O2 retained nearly full activity at the end of the assay. Similarly, sMMO and T2MO retained 80 % and 50 %, respectively, of the initial activity after reacting with substrate and 10 mM H2O2 for 30 min (Andersson et al., 1991; Newman & Wackett, 1995). Assuming that sBMO follows a similar catalytic cycle as sMMO (Wallar & Lipscomb, 1996), BMOH would be reduced to the diferrous state during NADH-coupled substrate turnover before interacting with O2. The peroxide shunt mechanism bypasses the diferrous state as H2O2 serves as the source of both electrons and O2. It seemed reasonable to conclude that the rapid inactivation seen during NADH-coupled ethene turnover was dependent on interaction of the diferrous state of BMOH with H2O2. This reasoning was bolstered due to the protection afforded by the presence of catalase in the reaction.
This study is the first report of the purification of the enzyme system sBMO from the C2–C9 aliphatic alkane oxidizing bacterium ‘P. butanovora’. The reconstituted sBMO complex has been characterized by assessment of NADH coupling to ethene oxidation, H2O2 shunt chemistry and product distributions of physiologically relevant linear and branched alkanes. The efficient hydroxylation of ethene and the preferential production of primary alcohols required for downstream catabolism of ‘P. butanovora’ was shown to be independent of the small component protein, BMOB. Future experiments are focused on complexation of BMOH and BMOB and methane oxidation utilizing the site-directed mutants of the -subunit of the hydroxylase (Halsey et al., 2006).
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ACKNOWLEDGEMENTS |
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Edited by: J. A. Vorholt
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Received 6 December 2006;
revised 19 January 2007;
accepted 15 February 2007.
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