Department of Biochemistry and Biophysics, Laboratory for Nitrogen Fixation Research1, and Department of Botany and Plant Pathology2, Oregon State University, Cordley 2082, Corvallis, 97331-2902 OR, USA
Author for correspondence: Daniel J. Arp. Tel: +1 541 737 4214. Fax: +1 541 737 3573. e-mail: arpd{at}bcc.orst.edu
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ABSTRACT |
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Keywords: butane metabolism, 1-butanol dehydrogenase, quinohaemoprotein
Abbreviations: ADH, alcohol dehydrogenase; BDH, 1-butanol dehydrogenase; DCPIP, 2,6-dichlorophenolindophenol; EDH, ethanol dehydrogenase; NBT, nitro blue tetrazolium; PMS, phenazine methosulfate; PQQ, pyrroloquinoline quinone
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INTRODUCTION |
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Except for the Gram-negative bacteria P. butanovora and Pseudomonas sp. strain CRL71 (Hou et al., 1983 ), most butane-oxidizing bacteria are members of the Gram-positive CorynebacteriumNocardiaMycobacteriumRhodococcus complex (Ashraf et al., 1994
). The pathways for metabolism of gaseous n-alkanes (C2C4) by some of these organisms have been investigated by several techniques. In all cases, metabolism is initiated by a monooxygenase (Ashraf et al., 1994
; Hamamura et al., 1999
; Perry, 1980
; Stephens & Dalton, 1986
; Woods & Murrell, 1989
). Terminal oxidation of an alkane leads to the corresponding 1-alcohol, while subterminal oxidation produces the 2-alcohol. Evidence for either, and in certain organisms for both, terminal and subterminal oxidation has accumulated. For example, the terminal oxidation of butane was inferred by the induction of isocitrate lyase in Mycobacterium vaccae JOB5 grown on butane or butyrate. Furthermore, isocitrate lyase was not induced when cells were grown on butanone, a predicted intermediate in the subterminal oxidation of butane (Phillips & Perry, 1974
). In contrast, subterminal oxidation of propane was suggested by the accumulation of acetone in propane-grown cells (Vestal & Perry, 1969
). Moreover, an inducible 2-propanol dehydrogenase was purified from M. vaccae JOB5 grown on either propane or 2-propanol (Coleman & Perry, 1985
). Pseudomonas fluorescens NRRL-B1244 can utilize propane, 1-propanol and 2-propanol as growth substrates. Two soluble NAD+-linked alcohol dehydrogenases (ADHs) (one primary ADH and one secondary ADH) were detected in propane-grown cells, which suggested that propane is metabolized through both terminal and subterminal oxidation pathways (Ashraf & Murrell, 1992
). The presence of both terminal and subterminal propane oxidation pathways in Rhodococcus rhodochrous PNKb1 was suggested because mutants unable to utilize either 1-propanol or 2-propanol were also unable to use propane as a growth substrate, indicating that both alcohols are intermediates of propane metabolism (Ashraf & Murrell, 1992
). The terminal oxidation of butane by butane-grown Nocardia TB1 was indicated by the accumulation of butyrate in the presence of appropriate inhibitors (Van Ginkel et al., 1987
). These results indicate the diversity of butane and propane oxidation pathways, including the diversity of the ADHs involved in alkane metabolism.
The pathway of butane metabolism by butane-grown P. butanovora was recently determined to follow the terminal oxidation pathway, that is butane1-butanol
butyraldehyde
butyrate (Arp, 1999
). Each intermediate (1) accumulated in the presence of appropriate inhibitors, (2) supported cell growth, and (3) stimulated O2 consumption of butane-grown cells. Although no production of 2-butanol was detected, 2-butanol was consumed and stimulated O2 consumption by butane-grown P. butanovora. Beers (1988)
examined the expression of ADH in butane-grown P. butanovora. Three soluble ADHs (one primary ADH and two secondary ADHs) that required NAD+ as an electron acceptor were found. A membrane-bound, NAD(P)+-independent ADH was also described.
In this study, we focused on the characterization of the ADH(s) induced in P. butanovora in response to growth on butane. Although butane-grown cells can consume 1- and 2-butanol, it was not known if one ADH was oxidizing both substrates. Our goal was to determine the number and specificity of ADHs present in butane-grown P. butanovora, and to purify and characterize the inducible ADH primarily involved in the butane metabolism pathway. We report here on the purification and characterization of a quinohaemoprotein, type II ADH induced in butane-grown P. butanovora. This report is the first to describe a quinohaemoprotein produced from n-alkane-grown bacteria.
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METHODS |
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Bacterial strains and growth conditions.
P. butanovora (ATCC 43655) cultures were grown in 12 l bottles at 30 °C with vigorous recirculation of a gas mixture (150 l consisting of 10%, v/v, butane; 5%, v/v, CO2; 85%, v/v, air) through the bottle. The growth medium consisted of, per liter, 8·0 g (NH4)2HPO4, 1·9 g Na2HPO4 . 7H2O, 2·0 g KH2PO4, 0·5 g MgSO4 . 7H2O, 0·06 g CaCl2.2H2O, 0·05 g yeast extract, and 1 ml of the trace element solution described by Wiegant & de Bont (1980) . For growth on 1-butanol, 2-butanol or lactate, an appropriate amount of each growth substrate was added to the medium to make the final concentration 4 mM. Butane and CO2 were omitted from the gas mixture. Cells were harvested at mid- to late-exponential growth phase (
4548 h). Cells were harvested by centrifugation (JA-14; Beckman J2-21) (15 min at 11000 g, 4 °C), resuspended at a ration of 1:1 (w/v) in 25 mM Tris/HCl pH 8·0 and kept at -80 °C until used.
Preparation of soluble fraction and purification of 1-butanol dehydrogenase (BDH).
All procedures for preparation of the soluble fraction and further purification were performed at 4 °C unless stated otherwise. Unbroken cell suspensions were passed three times through a French pressure cell at 5562 MPa. Unbroken cells were removed by centrifugation at 11000 g for 15 min. The cell-free extract was centrifuged at 200000 g for 1 h (SW40; Beckman L8-70); the resulting supernatant was the soluble fraction and the sediment was collected as the membrane fraction.
BDH was purified from the soluble fraction prepared from butane-grown cells (see above) by the following steps. In each step, the active fraction(s) were selected primarily by activity stain of non-denaturing gels and then the 1-butanol-dependent phenazine methosulfate (PMS) reductase activity of each active fraction was quantified spectrophotometrically. All active fractions were pooled and dialysed against 25 mM Tris/HCl buffer pH 8·0 at 4 °C overnight before being applied to the next column. (i) Fractionation with ammonium sulfate. To the soluble fraction, ammonium sulfate (to 30% saturation) was gradually added. The resulting precipitate was removed by centrifugation at 14000 g for 30 min. Additional ammonium sulfate was added to the supernatant to reach 70% saturation. The resulting precipitate was collected by centrifugation and redissolved at a ratio of 1:2 (w/v) in 25 mM Tris/HCl buffer pH 8·0. (ii) Q-Sepharose FPLC. The dialysed 3070% ammonium sulfate fraction was applied to a Q-Sepharose anion exchange FPLC column (Millipore) (2·2x10 cm) which had been equilibrated with 25 mM Tris/HCl buffer pH 8·0. The proteins were eluted by a continuous gradient of 01 M NaCl in the same buffer. The ADH of interest, which has a pinkish-red colour, eluted at 200250 mM NaCl, while a secondary ADH eluted at approximately 600 mM NaCl. (iii) 4-Amino-1-butanol affinity chromatography. The dialysed Q-Sepharose fraction containing a primary ADH was applied to 4-amino-1-butanol substrate-analogue affinity column prepared as previously described (Beers, 1988 ; Lange & Vallee, 1976
). The proteins were eluted by a step gradient of 0·05, 0·075, 0·1, 0·2 and 1 M NaCl in 25 mM Tris/HCl pH 8·0. Most proteins did not bind to the column. The ADH of interest eluted at 0·05 M NaCl.
Enzyme assays.
All enzyme assays were performed at 25 °C. NAD+-independent ADH activity was routinely measured as PMS reductase activity. PMS reductase activity was measured spectrophotometrically by monitoring the reduction of 2,6-dichlorophenolindophenol (DCPIP) at 600 nm in a reaction mixture consisting of 25 mM Tris/HCl pH 8·0, 0·7 mM PMS, 0·1 mM DCPIP, 4 mM NH4Cl, 0·8 mM KCN and 1 mM 1-butanol in a total volume of 3 ml. The enzyme activity of NAD+-dependent ADH was measured by the increase in A340 due to NADH in a reaction mixture (total volume 3 ml) consisting of 25 mM Tris/HCl pH 8·0, 2 mM NAD+ and the enzyme solution. The endogenous rate of DCPIP or NAD+ reduction obtained without substrate was subtracted from the rate observed with substrate. The reaction mixture (2 ml) for ferricyanide reductase activity assay contained 25 mM Tris/HCl pH 8·0, 1 mM potassium ferricyanide, 1 mM 1-butanol and enzyme solution. The enzyme concentration was fixed at 50 nM. One unit of enzyme activity was defined as the amount of enzyme catalysing oxidation of 1 µmol substrate min-1 under the conditions described. Protein content was estimated by the bicinchonic acid protein assay reagent (Pierce) with bovine serum albumin as a standard (Smith et al., 1986 ).
Determination of 1-butanol oxidation and product formation by GC.
The concentration of substrate (alcohol or aldehyde) utilized and type of products (aldehyde or organic acid) formed were determined by GC (Shimadzu GC-8A column equipped with a flame-ionization detector and 60 cm long by 0·1 cm i.d. stainless steel column packed with Porapak Q resin). The column temperature was 150 °C for detection of 1-butanol and butyraldehyde, and 120 °C for detection of 2-butanol and butanone.
Analytical gel filtration.
The molecular mass of native protein was determined by gel filtration (1·6x30 cm column) with Sephacryl S-300 (Pharmacia). Elution was performed at a flow rate of 0·5 ml min-1 with 25 mM Tris/HCl buffer pH 8·0 containing 0·15 mM NaCl, and was monitored at A280. The following molecular mass marker proteins were used for calibration: catalase (Mr 240000), bovine serum albumin (Mr 66000), carbonic anhydrase (Mr 30000) and cytochrome c (Mr 13000).
Electrophoresis.
10% SDS-PAGE (Mighty Small SE245; Hoefer Scientific Instruments) (Laemmli, 1970 ; Woods & Murrell, 1989
) and Coomassie brilliant blue G250 staining were used to analyse the homogeneity of the purified enzyme.
Non-denaturing gel and activity staining.
Native protein was applied to non-denaturing gels prepared as above but without SDS. After PAGE, the gel was incubated for 5 min in the dark with a reaction mixture (50 ml) containing 0·01 g nitro blue tetrazolium (NBT), together with 0·7 mM PMS or 1 mM NAD+ alone or in combination. Either 1-butanol or 2-butanol was then added to a final concentration of 1 mM and incubated with gentle rocking for another 5 min to develop the colour. The reaction was stopped by rinsing the gel with water.
Kinetic measurements.
Kinetic parameters were calculated from the initial rates determined with the PMS reductase standard assay described above by varying the concentrations of substrates tested from 2 µM to 1 mM. The enzyme concentration was fixed at 50 nM.
Preparation of antibodies and immunoblotting.
Purified BDH (2x100 µg) well mixed 1:1 (v/v) with TitreMax adjuvant (Sigma) was injected into two New Zealand rabbits. Two weeks later, a booster injection of another 100 µg protein mixed with TitreMax was performed. Another two weeks later, blood was collected, then left at room temperature for 30 min and overnight at 4 °C. After centrifugation at 1000 g for 10 min to remove the erythrocytes, antisera were collected and stored at -80 °C. For immunoblotting analysis, the proteins in the SDS gel after PAGE were transferred electrophoretically with a semi-dry system (Ausubel et al., 1996 ) onto a PVDF membrane at 10 mA overnight. The membrane was washed with 15 ml Tris-buffered-saline (TBS) (20 mM Tris/HCl, 150 mM NaCl, pH 7·5) before blocking with 15 ml 1% BSA in TBST (TBS with 0·1% Tween 20) for 1 h. The membrane was then incubated with primary antiserum (at a dilution of 1:5000 in TBST) for 1 h. The membrane was washed twice with TBST before incubation with TBST containing secondary antibody [1 mg anti-rabbit IgG (Fc) ml-1; Promega] conjugated with alkaline phosphatase (at a dilution of 1:5000 in TBST) for 30 min. After the membrane had been washed with 20 ml TBST three times, the immunosorbent protein bands were visualized by the addition of 15 ml of Western blue solution (Promega). The colour was developed within 520 min. Pre-stained molecular mass marker proteins (Bio-Rad) were included.
Haem staining.
A lithium dodecylsulfate-PAGE gel was prepared and pre-run overnight (Delepelaire & Chua, 1979 ). Purified BDH was then applied to the gel and run at 5 mA. Haem staining was performed as described by Thomas et al. (1976)
.
Measurement of haem c content.
The amount of haem c in the purified BDH was measured from the reduced minus oxidized difference spectra of the pyridine haemochrome (Paul et al., 1953 ), using a millimolar absorption coefficient of 26·5 (550534 nm). The oxidized form of pyridine haemochrome was prepared by adding pyridine to 20% (v/v), NaOH to a final concentration of 0·2 M, and potassium ferricyanide to a final concentration of 3 mM, to the enzyme solution. To reduce the reaction mixture, 2 mg sodium dithionite was added to the completely oxidized pyridine haemochrome solution. Cytochrome c from horse heart (Boehringer Mannheim) was used to test the protocol and gave a value of 1·0 haem per protein molecule.
Measurement of pyrroloquinoline quinone (PQQ) content.
The PQQ content of purified BDH was determined by HPLC analysis as described by Duine et al. (1983) .
N-terminal and internal amino acid sequence analysis.
To separate the upper and lower bands of the purified BDH, a 13x13 cm SDS gel was used. Both upper and lower bands were then blotted onto PVDF membrane and the N-terminal amino acid sequences were determined directly. The internal amino acid sequence was obtained by digesting the purified BDH with endoproteinase Lys-C (Endo Lys-C) in a reaction mixture consisting of 0·2 M Tris pH 9·1, 20% acetonitrile. The enzyme was digested at 37 °C for 20 h, then DTT was added to a final concentration of 5 mM and incubated at 37 °C for another 20 h. Peptides in the digest were separated by HPLC using a Vydac C18 column with a Brownlee guard column. The mobile phase A was 0·1% trifluoroacetic acid (TFA) and the mobile phase B was 70% acetonitrile plus 0·1% TFA. Chromatography was performed with a gradient of from 1 to 65% phase B in 75 min. Peaks were monitored at 220 nm and collected for sequence analysis. The prominent fragment was selected for sequencing. The N-terminal and internal amino acid sequencing, including enzyme digestion, were determined by the Biotechnology Laboratory, Institute of Molecular Biology, University of Oregon.
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RESULTS |
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BDH is induced during growth of P. butanovora on butane
Growth-substrate-dependent differences in the induction of primary and secondary ADH activities in P. butanovora were apparent from the activity-stained non-denaturing gels (Fig. 1c, d
). To quantify these differences, BDH activity was determined by spectrophotometric assay in extracts of cells grown on butane, 1-butanol, 2-butanol or lactate. The highest activity was present in cells grown on butane [32·9±1·7 nmol min-1 (mg protein)-1]. The activities of extracts of cells grown on 1-butanol, 2-butanol or lactate were 10·6±0·5, 4·4±0·1 and 1·9±0·1 nmol min-1 (mg protein)-1, respectively. The activities determined in solution correlated with the intensities of BDH activity bands on native gels.
To further confirm the correlation of BDH activity with butane metabolism, the induction of BDH upon exposure of cells to butane was examined. Cells were grown in citrate (5 mM) because this substrate does not repress the induction of butane metabolism when butane is provided. In contrast, lactate and succinate induce catabolic repression in Pseudomonas (Duetz et al., 1994 ; Yuste et al., 1998
). When cells reached an OD600 of 0·6 (2 d), they were divided into three parts. One part remained in the citrate-containing medium, another part received 10% butane in the gas phase, and the third part was supplemented with 1-butanol (to 4 mM). Cells were incubated at 30 °C with shaking. After 6 and 12 h, samples of cells were harvested, washed and broken. Further investigation of the induction of BDH activity was performed by Western blot analysis. The results from the activity assay were consistent with the increases of cross-reacting band intensities in Western blot analysis. As shown in Fig. 2(a)
, the cross-reacting band of an inducible BDH was first seen after cells were incubated with 1-butanol for 6 h. After 12 h, clearly visible cross-reacting bands were present in cell-free extracts of 1-butanol-induced and butane-induced cells, but not in citrate-grown cells. The 1-butanol-PMS/DCPIP-dependent activity assay (Fig. 2b
) showed 22-fold and 30-fold increases in BDH activity after citrate-grown cells were induced by 1-butanol and butane for 12 h, respectively.
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Purification and physical properties of BDH
BDH activity was the predominant ADH activity in butane-grown cells. The activity was closely correlated with butane metabolism. Therefore, we focused on this enzyme for additional characterization. BDH from butane-grown P. butanovora was purified from the soluble fraction to near homogeneity by a six-step procedure (Table 1). Many contaminating proteins were removed by the first anion exchange (Q-Sepharose) column fractionation, which was followed by a butanol affinity column fractionation. Repeating these two column fractionations was sufficient to purify the BDH to near homogeneity. The 37-fold increase in specific activity indicated that BDH accounted for about 2·7% of soluble protein.
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The molecular mass of BDH, the primary ADH, was estimated to be 66 kDa by SDS-PAGE (Fig. 3) and 69 kDa by gel permeation chromatography (Sephacryl S-300), which indicates that BDH exists as a monomer. This characteristic is similar to other monomeric quinohaemoproteins with a range of molecular masses of 6973 kDa (Groen et al., 1986
; Hopper et al., 1991
; Toyama et al., 1995
; Zarnt et al., 1997
). Because of the sequence similarity to the quinohaemoprotein EDH, the PQQ content of BDH was examined. BDH contains an almost 1:1 molar ratio of PQQ [0·99 nmol (nmol protein)-1], which indicates that PQQ was not lost during the purification. It has been reported that PQQ is non-covalently bound to quinohaemoprotein-type enzymes and can be dissociated by dialysis overnight in Tris/HCl buffer pH 8·0 containing 2 mM EDTA (Zarnt et al., 1997
) or by heat treatment (Toyama et al., 1995
).
BDH exhibits a pinkish-red colour in solution and appears as a red band in non-denaturing gels. A haem-staining band corresponding to a molecular mass of 65 kDa (data not shown) suggested the presence of haem cofactor in this enzyme. BDH as isolated exhibited a typical spectrum of reduced c-type cytochrome with absorption peaks at 416 nm ( peak), 520 nm (ß peak) and 550 nm (
peak) (Fig. 4
). When BDH was oxidized with 200 µM potassium ferricyanide, the
and ß peaks disappeared and the
peak was shifted to 410 nm (Fig. 4
). This spectral change from the reduced form to oxidized form is characteristic of cytochrome c.
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Catalytic properties of BDH
In addition to the 1-butanol-dependent PMS/DCPIP activity of BDH, we also examined its ferricyanide reductase activity, which is one of the characteristics of quinohaemoproteins. The ferricyanide reductase activity exhibited by BDH from butane-grown P. butanovora was 0·62±0·2 µmol min-1 (mg protein)-1 when 1-butanol was used as substrate.
The effect of calcium ions on BDH catalytic activity was determined. The presence of Ca2+ ions facilitated the reconstitution of inactive apoenzyme (PQQ-free, but haem-c-containing) quinohaemoprotein from C. testosteroni and alkaloid-degrading Pseudomonas and increased the ADH catalytic activity (de Jong et al., 1995 ; Groen et al., 1986
; Zarnt et al., 1997
). Ca2+ fulfils a structural role upon reconstitution of apoenzyme with PQQ, facilitating the binding of PQQ to an apoenzyme and activating the bound cofactor (Groen et al., 1986
; Zarnt et al., 1997
). The activity of BDH from butane-grown P. butanovora was significantly increased when Ca2+ was added into the reaction mixture. BDH activity with 1-butanol was increased from 3·4±0·2 µmol min-1 (mg protein)-1 (without Ca2+) to 4·2±0·01 µmol min-1 (mg protein)-1 with 5 mM CaCl2 and 4·6±0·2 µmol min-1 (mg protein)-1 with 10 mM CaCl2. The maximal and saturated activity was reached at a Ca2+ concentration of 10 mM. Ca2+ was recently reported to have an important role in the catalytic mechanism of quinoprotein-type methanol dehydrogenase (Zheng & Bruice, 1997
). The possible roles of Ca2+ complexed with PQQ in methanol oxidation include: (i) modest reduction of the pKa of the substrate and facilitating the association of substrate to active site, (ii) polarizing the oxygen at the C-5 position of PQQ, and (iii) placing the reaction components in the right positions to react, therefore contributing to the formation of enzymesubstrate complex (Zheng & Bruice, 1997
).
BDH activity was increased approximately threefold when ammonium ions (as 4 mM NH4Cl) were present. A similar result was obtained in tetrahydrofurfuryl ADH from R. eutropha where the activity was increased 2·5-fold in the presence of ammonium sulfate (Zarnt et al., 1997 ).
The optimum pH for PMS/DCPIP-dependent BDH activity was 8·0. The temperature optimum was 60 °C, at which the specific activity was increased 9·6-fold relative to that at 25 °C. The temperature range of maximum stability of the enzyme was 2532 °C, within which the loss of BDH activity was insignificant after a 30 min incubation. In contrast, BDH activity was decreased 77% when incubated at 60 °C for the same period of time. BDH activity was stable for more than 6 months when stored in 25 mM Tris/HCl pH 8·0 at -80 °C.
BDH exhibited broad substrate specificity towards various primary alcohols, secondary alcohols, diols, aldehydes and aromatic alcohols (Table 3). Nonetheless, the highest activity was observed with 1-butanol, the butane oxidation product in the terminal oxidation pathway. Therefore, the activities obtained on other substrates were compared to the maximal activity obtained on 1-butanol. Besides 1-butanol, 1-propanol showed a relatively high activity (66%) among the primary alcohols tested. The relative activities among primary alcohols decreased with longer-chain alcohols. Little activity (12%) was detected with methanol, which is consistent with the observation that P. butanovora cannot grow with either methane or methanol as a substrate (Takahashi, 1980
). Among the secondary alcohols tested, BDH exhibited a marked preference towards 2-pentanol and the activity gradually decreased with longer-chain secondary alcohols. DCPIP was slowly reduced by BDH when 2-propanol or 2-butanol was added. In activity-stained native gels, BDH activity bands were also detected when 2-butanol was added (Fig. 1f
, g
). However, when 2-butanol consumption was determined directly by GC, no 2-butanol consumption was observed. Furthermore, no butanone formation was detected within the 60 min assay (data not shown). The low level of DCPIP reduction activity obtained upon addition of either 2-propanol or 2-butanol was probably due to the presence of contaminants that supported activity. BDH was active towards C4 diols when one of the hydroxyl groups was at the
-position. Benzyl alcohol and phenol were chosen as the representative aromatic alcohols and were slowly oxidized by BDH. BDH readily oxidized the three aldehydes we tested, with butyraldehyde showing the highest activity. In fact, only 1-butanol was oxidized more rapidly than butyraldehyde.
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DISCUSSION |
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We purified this soluble, NAD+-independent primary ADH from butane-grown P. butanovora and showed it to be a quinohaemoprotein. As pointed out by Toyama et al. (1995) , there have been no reports of PQQ-containing ADHs associated with alkane oxidation by bacteria. This work is the first report of a quinohaemoprotein induced from alkane-grown bacteria. Interestingly, this inducible BDH from butane-grown P. butanovora was found to be different from ADHs induced in other alkane-grown Pseudomonas or other short-, medium- or long-chain-alkane-oxidizing bacteria. ADHs induced in propane-grown R. rhodochrous PNKb1, M. vaccae JOB5 and P. fluorescens NRRL B-1244 were purified and characterized as NAD+-dependent secondary ADH(s) (Ashraf & Murrell, 1990
, 1992
; Coleman & Perry, 1985
). Heptane- and octane-grown Pseudomonas aeruginosa strain 473 contain soluble NAD(P)+-dependent ADH and at least two NAD(P)+-independent quinoprotein (without haem c) primary ADHs (Van Der Linden & Huybregtse, 1969
).
In addition to methanol dehydrogenase, other quinoprotein ADHs found in oxidative nonmethylotrophic bacteria have been classified into three groups (types IIII) according to their molecular properties, catalytic properties and localization (Matsushita et al., 1994 ). The molecular structure of type I ADH resembles that of methanol dehydrogenase, but it has very low affinity for methanol. Type II ADH is a soluble quinohaemoprotein. To date, type II ADHs have been purified from five bacteria: as the apoenzyme, lacking PQQ, from alkaloid-degrading Pseudomonas (Hopper et al., 1991
) and C. testosteroni (de Jong et al., 1995
; Groen et al., 1986
); and as the active holoenzyme from P. putida HK5 (Toyama et al., 1995
), Rhodopseudomonas acidophila M402 (Yamanaka & Tsuyuki, 1983
) and R. eutropha (Zarnt et al., 1997
). Type III ADH is a membrane-bound ADH found in the cytoplasmic membrane of acetic acid bacteria.
The characterization of purified BDH from butane-grown P. butanovora indicates that this enzyme is a type II NAD+-independent ADH or quinohaemoprotein (Goodwin & Anthony, 1998 ). This is also the first report of a type II NAD+-independent ADH synthesized by an Azoarcus-like bacterium. PQQ, a prosthetic group in quinohaemoprotein, is expected to participate directly in the oxidation of alcohols and aldehydes (Groen et al., 1986
). Apo-EDH, lacking PQQ but containing haem c, has no activity with alcohol or aldehyde substrates. The presence of haem c as a functional prosthetic group in BDH was indicated by the following results. (i) A red band, which also stained for ADH activity, was observed in non-denaturing gels. (ii) Protein corresponding to a molecular mass of 65 kDa in lithium dodecylsulfate-PAGE also stained positive for haem. (iii) The absorption spectrum of purified BDH, with peaks at 416, 520 and 550 nm, was a typical reduced cytochrome c spectrum that changed to an oxidized spectrum upon addition of ferricyanide, and back to a reduced spectrum upon addition of 1-butanol. (iv) A ferricyanide reductase activity was exhibited by BDH; quinoproteins, lacking haem c, such as methanol dehydrogenase, EDH from P. aeruginosa and P. putida, and soluble glucose dehydrogenase from Escherichia coli, do not react with ferricyanide, suggesting the oxidation of the enzyme by ferricyanide occurs at the haem c site (Dokter et al., 1986
; Geerlof et al., 1994
; Groen et al., 1984
). For a cell extract from 1-butanol-grown P. putida HK5, the ratio of 1-butanol-dependent PMS/DCPIP-reductase activity to ferricyanide-reductase activity was 2:1 (Toyama et al., 1995
). BDH from butane-grown P. butanovora as isolated expressed, with 1-butanol present, a specific activity ratio of PMS/DCPIP reductase activity to ferricyanide reductase activity of 7·8. This specific activity ratio is consistent with the fact that only one quarter of the BDH as isolated contained haem c.
The physical properties of BDH presented here, including high internal sequence similarity with C. testosteroni EDH, indicates that BDH belongs to the type II quinohaemoprotein group. However, the affinity of BDH for 1-butanol is much higher than that of C. testosteroni EDH for ethanol or of R. eutropha tetrahydrofurfuryl ADH for n-pentanol (Geerlof et al., 1994 ; Zarnt et al., 1997
). The Km of BDH towards 1-butanol (7±1 µM) is the lowest Km reported among other quinohaemoprotein ADHs obtained from ethanol-grown C. testosteroni (Km 2230±64 µM towards ethanol; Geerlof et al., 1994
) or 1-butanol grown P. putida HK5 (Km 105 µM towards 1-butanol; Toyama et al., 1995
). Coupled with much lower affinity for butyraldehyde (Km 535±13 µM), 1-butanol is clearly preferred over butyraldehyde by BDH. In contrast, C. testosteroni EDH has a higher affinity for the aldehyde (Km for acetaldehyde 669±22 µM), such that acetic acid is produced even in the presence of ethanol.
It was recently demonstrated that butane metabolism by P. butanovora was mainly via a terminal oxidation pathway (Arp, 1999 ). Up to 90% of the butane consumed was accounted for as 1-butanol. The results of this work are consistent with terminal oxidation. The predominant ADH activity in butane-grown cells is much more efficient towards 1-butanol than 2-butanol. Furthermore, the enzyme is expressed at the highest level observed in butane-grown cells. However, butane-grown cells also expressed a low level of 2-butanol-dependent ADH activity that was not present in 1-butanol-grown cells. This result may reflect a low level of subterminal butane oxidation involved in butane metabolism in P. butanovora. Studies of the genes encoding each of these enzymes, including expression experiments and gene disruptions, will be pursued to further characterize the role of each ADH in butane metabolism.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 22 August 2000;
revised 6 November 2000;
accepted 13 November 2000.