From the Max-Planck-Institut für marine
Mikrobiologie, Celsiusstrasse 1, 28359 Bremen, and
§ Mikrobiologie, Institut für Biologie II,
Universität Freiburg, Schänzlestrasse 1,
79104 Freiburg, Germany
Received for publication, February 22, 2001, and in revised form, April 5, 2001
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ABSTRACT |
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The initial enzyme of ethylbenzene
metabolism in denitrifying Azoarcus strain EbN1,
ethylbenzene dehydrogenase, was purified and characterized. The soluble
periplasmic enzyme is the first known enzyme oxidizing a nonactivated
hydrocarbon without molecular oxygen as cosubstrate. It is a novel
molybdenum/iron-sulfur/heme protein of 155 kDa, which consists of three
subunits (96, 43, and 23 kDa) in an Three bacterial species capable of anaerobic degradation of the
aromatic hydrocarbon ethylbenzene are known to date. All of these are
denitrifying bacteria that belong to the genus Azoarcus of
the structure. The N-terminal
amino acid sequence of the
subunit is similar to that of other
molybdenum proteins such as selenate reductase from the related species
Thauera selenatis. Ethylbenzene dehydrogenase is unique in
that it oxidizes the hydrocarbon ethylbenzene, a compound without
functional groups, to (S)-1-phenylethanol. Formation of the
product was evident by coupling to an enantiomer-specific (S)-1-phenylethanol dehydrogenase from the same organism.
The apparent Km of the enzyme for ethylbenzene is
very low at <2 µM. Oxygen does not affect ethylbenzene
dehydrogenase activity in extracts but inactivates the purified enzyme,
if the heme b cofactor is in the reduced state. A variant of
ethylbenzene dehydrogenase exhibiting significant activity also with
the homolog n-propylbenzene was detected in a related
Azoarcus strain (PbN1).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-proteobacteria. For one of these strains, Azoarcus
sp. EB-1, ethylbenzene is the only known hydrocarbon utilized as growth substrate (1). The other two strains utilize either ethylbenzene or an
alternative hydrocarbon compound, namely toluene (strain EbN1) or
n-propylbenzene (strain PbN1) (2). The proposed pathway of
anaerobic degradation of ethylbenzene by these bacteria is shown in
Fig. 1. It is initiated by a novel
biochemical reaction, namely an oxygen-independent oxidation of
ethylbenzene to (S)-1-phenylethanol. This intermediate is
then oxidized further to acetophenone by an alcohol dehydrogenase
(1-4). Activities of an ethylbenzene-oxidizing enzyme and an
enantio-specific (S)-1-phenylethanol dehydrogenase have been
reported in cell extracts of strain EB-1 (4), and a substrate-specific
(S)-1-phenylethanol dehydrogenase has been purified and
characterized from strain
EbN1.1 The intermediate
acetophenone is apparently degraded further by carboxylation to
benzoylacetate to yield benzoyl-CoA and acetyl-CoA eventually (Fig. 1;
for review, see Ref. 5). The catabolic pathway of
n-propylbenzene in strain PbN1 is supposed to be analogous to that of ethylbenzene, yielding benzoyl-CoA and propionyl-CoA as
intermediates (2). Toluene degradation in strain EbN1 proceeds via a
completely different pathway and involves the formation of
benzylsuccinate from toluene and fumarate as initial reaction (3, 6,
7).
View larger version (29K):
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Fig. 1.
Proposed anaerobic degradation pathway of
ethylbenzene in strain EbN1. Fc+, ferricenium; Fc,
ferrocene.
In this report, we analyze the biochemical properties of the first
enzyme of anaerobic ethylbenzene metabolism, ethylbenzene dehydrogenase. The enzyme was purified and shown to be a new
periplasmic molybdenum/iron-sulfur/heme protein that oxidizes
ethylbenzene stereospecifically to (S)-1-phenylethanol. We
also provide evidence that the same enzyme catalyzes anaerobic
oxidation of ethylbenzene and n-propylbenzene.
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EXPERIMENTAL PROCEDURES |
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Growth of Bacteria and Preparation of Cell Extracts--
Strain
EbN1 was isolated previously from an enrichment culture on ethylbenzene
by Rabus and Widdel (2). Growth of the bacteria in 1-2-liter scale
cultures was performed as described previously (2, 3). Cells were grown
by subsequent transfer for at least 30 generations on the same
substrate prior to harvesting for the described experiments. Harvesting
was performed anoxically while the cultures were in the exponential
growth phase. Fermenter cultures (200 liters) were set up as described
previously (3) and run in fed-batch mode with a growth-limiting and
exponentially increasing feeding rate of nitrate and discontinuous
supply of ethylbenzene. Growth rates of 0.015-0.025
h1 and cell yields of 200-300 g (wet
mass)/fermenter were usually obtained. Extract preparation was usually
performed aerobically. Cells (10 g, wet mass) were suspended in 10 ml
of water and passed through a French pressure cell at 137 megapascals.
Cell debris and membranes were removed by ultracentrifugation (1 h at
100,000 × g). Washed membrane fractions were prepared
from the supernatant of a 20,000 × g centrifugation
step, which was centrifuged at 100,000 × g for 1 h. The pellet was washed and resuspended in the same volume of basal
buffer (10 mM Tris-Cl, 1 mM MgCl2,
10% glycerol, pH 7.5). For anaerobic extract preparation, all
solutions were degassed and stored under nitrogen, and all handling
steps were performed in an anaerobic glove box as described earlier (3). Strain PbN1 (2) was grown in 2-liter bottles under the same
conditions as described for strain EbN1. The hydrocarbon substrates
were added to the cultures in an inert carrier phase (2,2,4,4,6,8,8-heptamethylnonane) containing 2% (v/v) ethylbenzene or
4% (v/v) n-propylbenzene. Shortest doubling times of
10.5 h on ethylbenzene and 12 h on n-propylbenzene
were recorded.
Enzyme Assays--
Ethylbenzene dehydrogenase was routinely
assayed in 100 mM Tris-Cl buffer (pH 7.5) containing 0.2 mM ferricenium hexafluorophosphate as electron acceptor.
Enzyme solution was added, and the reactions were started by adding
ethylbenzene or n-propylbenzene (final concentration, 100 µM) from saturated aqueous solutions, which contained 2 mM ethylbenzene (8) or 1 mM
n-propylbenzene (9). Decrease of absorption of the
ferricenium ion was followed at 290 nm ( = 9,000 M
1
cm
1). The tests were routinely performed
under aerobic conditions because identical activities were observed in
control tests under anaerobic conditions. To assess the pH optimum of
ethylbenzene dehydrogenase, the enzyme assay was also performed in
sodium phosphate buffers within a pH range of 6.0-8.0. Alternative
assays for ethylbenzene oxidation were set up with 0.1 mM
dichlorophenol indophenol as electron acceptor in the presence and
absence of the redox mediator phenazine methosulfate (0.05 mM). These tests were performed under anaerobic conditions
as described above and were monitored for dichlorophenol indophenol
reduction at 546 nm. Reversibility of the ethylbenzene dehydrogenase
reaction was tested under strictly anaerobic conditions in 100 mM Tris-Cl buffer (pH 7.5), containing 1 mM
methyl viologen and 0.5 mM dithionite. Oxidation of reduced methyl viologen was followed at 710 nm (
= 2,400 M
1
cm
1). After adding the enzyme, the reaction
was started by adding 1 mM (S)-1-phenylethanol.
The same buffer was also used to assess the purified enzyme for
possible selenate reductase or nitrate reductase activities. In these
cases, the reaction was started by the addition of 1 mM
respective electron acceptor. (S)-1-Phenylethanol dehydrogenase activity was assayed in 100 mM Tris-Cl buffer
(pH 7.5) containing 2 mM MgCl2, 0.5 mM NAD, and 1 mM (S)-1-phenylethanol and enzyme. Malate dehydrogenase activity was measured in 100 mM potassium phosphate buffer (100 mM, pH 7)
containing 0.25 mM NADH, 0.2 mM oxaloacetate,
and cell extract. Reduction of NAD+ or oxidation of NADH
was followed photometrically at 365 nm (
= 3.4 mM
1
cm
1).
Enzyme Purification--
All column chromatography steps were
performed in an anaerobic glove box with an FPLC System (Amersham
Pharmacia Biotech). Extract of ethylbenzene-grown cells of strain EbN1
(20 ml of a 100,000 × g supernatant) was applied to a
DEAE-Sepharose column (Amersham Pharmacia Biotech; 2.2-cm diameter,
50-ml volume), which had been equilibrated with basal buffer (2 mM Tris acetate buffer, pH 8.0, and 10% w/v glycerol). The
column was washed at a flow rate of 5 ml min1
for 2 column volumes and eluted with a gradient from 0 to 50 mM KCl in basal buffer over 500 ml. Fractions of 7 ml were
collected. Ethylbenzene dehydrogenase activity eluted in a volume of 80 ml between 40 and 50 mM KCl. A yield of 77% and an
enrichment factor of 20 were obtained after this step (see Table III).
The active fractions were applied on a ceramic hydroxyapatite column
(10 ml; Bio-Rad, Hercules, CA), which had been equilibrated with basal buffer. The column was washed with 2 volumes of basal buffer. A
gradient over 100 ml was then applied from 0 to 300 mM
potassium phosphate, and fractions of 5 ml were collected. Enzyme
activity eluted in a volume of 40 ml when 160-250 mM
potassium phosphate was applied. Active fractions were pooled, and
ethylbenzene dehydrogenase was concentrated by ammonium sulfate
precipitation under anaerobic conditions (60% saturation of ammonium sulfate).
Separation of Subcellular Compartments-- Cells of strain EbN1 were grown and harvested as described above. Spheroplasts were formed by using a modification of previous procedures (10, 11). Freshly harvested cells (0.8 g, wet mass) were resuspended in 64 ml of TS buffer (30 mM Tris-Cl, 30% sucrose, pH 8). EDTA (9 mM final concentration) and lysozyme (2.6 × 106 units) were added, and the suspension was incubated on ice for 120 min to produce spheroplasts. Periplasmic proteins were prepared by centrifugation of the spheroplast suspension for 30 min at 16,000 × g. Most of the periplasmic proteins were recovered in the supernatant, whereas the pellet contained the intact spheroplasts. These were washed in TS buffer, suspended in 25 ml of buffer (20 mM Tris-Cl, 10 mM MgCl2, 10% glycerol, pH 8) containing 10 mg DNase I and lysed by one passage through a French pressure cell. The membrane and soluble cytoplasmic fractions of the cell lysate were separated by centrifugation at 100,000 × g.
Other Methods--
Protein concentrations were determined
according to Lowry (12) or by the Coomassie dye binding test (12) with
bovine serum albumin as standard, and discontinuous
SDS-PAGE2 was performed in
15% (w/v) polyacrylamide gels according to standard procedures (12).
Molecular mass standards were phosphorylase b, bovine serum
albumin, ovalbumin, lactate dehydrogenase, carbonic anhydrase, trypsin
inhibitor, and lysozyme. Gels were analyzed by the
ImageMaster® one-dimensional software (Amersham Pharmacia
Biotech). UV-visible spectra were recorded with a 2 photometer
(PerkinElmer Life Sciences). Cytochrome c content in
subcellular fractions was analyzed and calculated as described (13).
The native molecular mass of ethylbenzene dehydrogenase was determined
by gel filtration on a calibrated Superdex 200 column (Amersham
Pharmacia Biotech) and by analysis of purified enzyme on native
polyacrylamide gels. Gels containing five different polyacrylamide
concentrations between 6 and 8% (w/v) were used, and ovalbumin and the
monomer, dimer, trimer, and tetramer bands of bovine serum albumin were
used as standards for a Ferguson plot (12). Photometric quantitation of
molybdenum, tungsten (14), iron (15), and inorganic sulfide (16) was performed by standard chemical techniques. Additionally, a simultaneous determination of 32 elements in purified enzyme was performed by
inductively coupled plasma optical emission spectroscopy (ICP-OES) using a Jarrel Ash Plasma Comp 750 instrument at the center of Complex
Carbohydrate Research, University of Georgia. For protein microsequencing, cell extract or purified enzyme was separated by
SDS-PAGE and blotted on a polyvinylidene difluoride membrane (Pro
Blott, Applied Biosystems, Weiterstadt, Germany) using a Semi-Phor
(model TE77) semidry blotting device as described in (17). Proteins on
polyvinylidene difluoride membrane were stained by Coomassie Blue
R-250. The proteins were subjected to Edman degradation microsequencing
(Procise 492 Sequencer, Applied Biosystems) with repetitive yields of
>96%. Ferricenium hexafluorophosphate was synthesized following a
published procedure (18); all other chemicals were from Fluka, Sigma
(Deisenhofen, Germany), or Merck (Darmstadt, Germany) and were of the
highest available purity.
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RESULTS |
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Ethylbenzene and n-Propylbenzene Dehydrogenase Activities in Strain
EbN1--
A photometric enzyme assay was developed for the first
enzyme of anaerobic ethylbenzene degradation of strain EbN1,
ethylbenzene dehydrogenase. The artificial electron acceptors
dichlorophenol indophenol or phenazine methosulfate were tested without
success for coupling to ethylbenzene oxidation. However, significant
activity of an ethylbenzene dehydrogenase was detected in extracts of
ethylbenzene-grown cells with the ferricenium cation as electron
acceptor. The assay was dependent on the amount of protein, and a pH
optimum of 7.0 was determined. Identical activities were obtained under
oxic and strictly anoxic conditions, indicating that molecular oxygen is not required for ethylbenzene oxidation. No decrease in activity was
recorded when extracts were adjusted to pH 5.5 or to pH 9.0 prior to
starting the enzyme assay. Ethylbenzene dehydrogenase activity was
detected exclusively in the soluble fraction after 100,000 × g centrifugation; no activity was found in washed membrane fractions. With untreated cell extracts, an ethylbenzene:ferricenium stoichiometry (ethylbenzene:electron ratio) of 1:3.9 was determined, indicating that ethylbenzene is oxidized to acetophenone in these assays. Assuming a stoichiometry of 1:4, the specific ethylbenzene oxidation rate in 100,000 × g supernatants was 22 ± 4 nmol min1 (mg of
protein)
1. This value is close to a
calculated minimum substrate degradation rate of 28 nmol
min
1 (mg of
protein)
1 required in growing cells at the
time of harvesting (doubling time 11 h). Very low ethylbenzene
dehydrogenase activity was recorded in extracts of cells grown on
(S)-, (R)-, or racemic 1-phenylethanol, or on
acetophenone, and none in benzoate-grown cells (Table
I). Clear activity was also observed when
ethylbenzene was replaced by n-propylbenzene in the assays
with ethylbenzene-grown cells, albeit only at about 15% of the
activity measured with ethylbenzene (Table I).
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Ethylbenzene and n-Propylbenzene Dehydrogenase Activities in Strain PbN1-- Cells of strain PbN1, which were grown in 2-liter cultures on ethylbenzene or n-propylbenzene, were also tested for dehydrogenase activities for ethylbenzene and n-propylbenzene. Ethylbenzene-grown cells of strain PbN1 contained two times higher specific activities of ethylbenzene dehydrogenase than observed in strain EbN1 (Table I). In extracts of strain PbN1, n-propylbenzene was oxidized at a rate of 62-66% of that measured with ethylbenzene. If strain PbN1 was grown on n-propylbenzene, the specific activity of the enzyme was slightly lower, but the ratio of the rates with n-propylbenzene or ethylbenzene as substrates was nearly the same as with ethylbenzene-grown cells (Table I). These findings suggest that ethylbenzene and n-propylbenzene are oxidized by a common enzyme. Specific activities for both substrates were sufficient to explain the observed growth rates of strain PbN1 on either hydrocarbon.
Protein Patterns of Cells of Strain EbN1 and PbN1 Grown on
Different Substrates--
Ethylbenzene-grown cells of strain EbN1
contained large amounts of several substrate-induced polypeptides,
which were lacking in cells grown on 1-phenylethanol, the next
intermediate of the metabolic pathway. The most prominent
ethylbenzene-induced polypeptides visible in cell extracts separated by
SDS-PAGE are shown in Fig. 2. Several
other substrate-induced polypeptides were also observed in cells grown
on 1-phenylethanol or acetophenone compared with benzoate-grown cells
(Fig. 2), indicating sequential induction of the catabolic enzymes of
anaerobic ethylbenzene metabolism. Extracts of ethylbenzene- or
n-propylbenzene-grown cells of strain PbN1 showed no obvious
difference in polypeptide patterns. Several prominent substrate-induced
polypeptides of identical sizes were observed in cells grown on either
hydrocarbon substrate compared with benzoate-grown cells (Fig. 2).
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Stoichiometry and Stereospecificity of Ethylbenzene Dehydrogenase-- Proteins in extracts of ethylbenzene-grown cells of Azoarcus sp. strain EbN1 were separated initially by chromatography on DEAE-Sepharose in 10 mM Tris-Cl buffer (pH 7.5). Under these conditions, 100% of ethylbenzene dehydrogenase activity was found in the flow-through fractions. (R)- and (S)-1-Phenylethanol dehydrogenases from the cell extract were retained on the column,1 thus allowing complete separation of the first two enzymes of ethylbenzene degradation. Using the separated enzymes, we determined whether the four-electron oxidation of ethylbenzene to acetophenone, as detected in cell extracts, is catalyzed by ethylbenzene dehydrogenase alone, or whether it depends on the subsequent (S)-1-phenylethanol dehydrogenase reaction. Enzyme assays with ethylbenzene dehydrogenase containing flow-through fractions, which are devoid of (S)-1-phenylethanol dehydrogenase, showed an ethylbenzene:ferricenium stoichiometry of 1:2.3. The stoichiometry changed to 1:4.1 when the tests were supplied with NAD+ and purified (S)-1-phenylethanol dehydrogenase1 and remained at 1:2.3 when the same experiment was performed in the absence of NAD+. Therefore, ethylbenzene dehydrogenase catalyzes a two-electron oxidation of ethylbenzene, and stereospecifically produces (S)-1-phenylethanol, which is subsequently oxidized to acetophenone by (S)-1-phenylethanol dehydrogenase in cell extract. The NADH generated by the alcohol dehydrogenase is apparently reoxidized by an NADH:acceptor oxidoreductase with ferricenium as electron acceptor.
Subcellular Localization of Ethylbenzene Dehydrogenase-- After generating spheroplasts from freshly harvested cells of ethylbenzene-grown strain EbN1 by lysozyme/EDTA treatment in isotonic Tris-Cl/sucrose buffer, 94% of the ethylbenzene dehydrogenase was released into the medium (Table II). The periplasmic marker protein cytochrome c was also largely released in these experiments, as determined from difference spectra (dithionite-reduced minus oxidized), whereas 93% of the cytoplasmic marker enzyme malate dehydrogenase was retained in the spheroplasts (Table II). Determination of (S)-1-phenylethanol dehydrogenase activities in the subcellular fractions showed that 20% of this enzyme was released, and about 80% was retained in the spheroplasts (Table II). This indicated that ethylbenzene oxidation to (S)-1-phenylethanol occurs in the periplasm, whereas further oxidation of (S)-1-phenylethanol to acetophenone occurs in the cytoplasm. This is corroborated by the stoichiometries of ethylbenzene oxidation versus ferricenium reduction in the different subcellular fractions. In cytoplasmic fractions, an ethylbenzene:ferricenium stoichiometry of 1:3.2 was determined, indicating coupling of the residual ethylbenzene dehydrogenase activity with (S)-1-phenylethanol dehydrogenase, whereas the periplasmic fractions showed an ethylbenzene:ferricenium stoichiometry of 1:1.9.
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Purification of Ethylbenzene Dehydrogenase from Strain EbN1-- Because ethylbenzene dehydrogenase did not bind to DEAE-Sepharose columns when 10 mM Tris-Cl buffer (pH 7.5) was used, the elution buffer was changed to a chloride-free 2 mM Tris acetate buffer (pH 7.5). Under these conditions, the enzyme bound to the DEAE-column and was then eluted from the column by a linear KCl gradient. Although enzyme activity in extracts was not affected by air, enriched enzyme from the first column quickly lost activity under oxic conditions. Therefore, purification of the enzyme was performed under anoxic conditions, which resulted in typical yields of >75% and an enrichment factor of 20 after the first column (Table II). Chromatography on ceramic hydroxyapatite (Bio-Rad) was performed as second purification step. The pooled enzyme eluted from the DEAE-Sepharose column was applied directly on this column and eluted by a linear potassium phosphate gradient (pH 7.5). A typical enrichment factor of 24 at a yield of 20% was obtained after this step. The enzyme was essentially pure after the second column, as shown by SDS-PAGE. A summary of the purification scheme is given in Table III.
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Molecular Properties of Ethylbenzene
Dehydrogenase--
Ethylbenzene dehydrogenase consists of three
subunits of 96, 43, and 23 kDa, as revealed by SDS-PAGE of the purified
enzyme (Fig. 3). The apparent native
molecular mass of the enzyme was determined as 155 ± 15 kDa by
gel filtration and Ferguson plot analysis of native polyacrylamide
gels. These values are consistent with an assumed composition
of the enzyme. The N-terminal amino acid sequences of the three
subunits were determined from enzyme that had been separated by
SDS-PAGE. The N terminus of the
subunit of purified ethylbenzene
dehydrogenase was blocked, whereas short sequences of the
and
subunits were obtained. However, a sequence of the
subunit was
obtained when the ethylbenzene-induced 96-kDa polypeptide was cut out
from blotted cell extracts and used for sequencing. The N-terminal
amino acid sequence of the
subunit (GTKAPGYASWEDIYRKEWKWDKVN) was
highly similar to that of other molybdoproteins such as selenate
reductase of Thauera selenatis (63% identity) or several
nitrate reductases (e.g. 63% identity with nitrate
reductase subunit 1 of Haloarcula marismortui). The
sequences obtained for the other subunits (
,
XGPXXYLRP; and
, XKAKRVPGGKELLLDL)
did not show significant matches with known proteins.
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Additionally, molar contents of molybdenum, iron, and acid-labile
sulfide were determined in purified ethylbenzene dehydrogenase. Molybdenum content in ethylbenzene dehydrogenase was determined as
0.9 ± 0.1 mol (mol of enzyme)1 by
colorimetric assay, and as 1.2 ± 0.1 mol (mol of
enzyme)
1 by ICP-OES. Iron was determined
chemically at a stoichiometry of 17 ± 2 mol (mol of
enzyme)
1 compared with 16 ± 1 mol of
iron (mol of enzyme)
1 determined by ICP-OES.
Finally, colorimetric analysis of acid-labile sulfide yielded 12 ± 4 mol S2
(mol of enzyme)
1.
These values are consistent with the presence of one molybdenum, four
[Fe4S4] clusters, and one heme in
ethylbenzene dehydrogenase, as reported previously for selenate
reductase of T. selenatis (19, 20). Other metals detected by
ICP-OES in significant amounts were magnesium and calcium (2.5 mol/mol
each), but no further transition metals or selenium were present in
purified enzyme.
Spectral Properties of Ethylbenzene Dehydrogenase--
UV-visible
spectroscopic analysis of purified ethylbenzene dehydrogenase showed a
complex spectrum. The spectrum of the purified enzyme showed a shoulder
around 400 nm and distinct absorption maxima at 424, 528, and 559 nm,
which indicated the presence of a reduced heme b cofactor. After
anaerobic oxidation of the enzyme by stoichiometric concentrations of
ferricenium hexafluorophosphate, the and
peaks of the
tentatively identified heme at 559 and 528 nm disappeared, and the
Soret band at 424 nm was shifted to 414 nm (Fig.
4). The difference spectrum of reduced
and oxidized enzyme was indeed indicative of the presence of a heme b
cofactor (Fig. 4). Addition of stoichiometric concentrations of
ethylbenzene to ferricenium-oxidized enzyme resulted in restoration of
the spectrum of reduced enzyme (Fig. 4). The spectra of the
substrate-reduced enzyme and the enzyme obtained from the final column
were identical, suggesting that ethylbenzene dehydrogenase was purified
in the completely reduced form. Treatment of the reduced enzyme with 0.2 mM dithionite did not result in further reduction of
the heme cofactor but resulted in further bleaching of the absorption
between 400 and 500 nm (Fig. 4). This is indicative of the presence of iron-sulfur clusters in ethylbenzene dehydrogenase. These clusters are
obviously not completely reduced by the substrate and need a strong
chemical reductant such as dithionite for being entirely reduced. The
heme content of the enzyme was determined as 0.95 mol/mol from
dithionite-reduced enzyme, using an
of 34.7 mM
1 cm
1
for the
band (21).
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Catalytic Properties of Ethylbenzene Dehydrogenase--
Purified
ethylbenzene dehydrogenase catalyzed the
ferricenium-dependent oxidation of ethylbenzene to
(S)-1-phenylethanol. The specific activity recorded for
purified enzyme corresponds to a very low catalytic number of 1.2 s1, a value similar to that recorded for
benzoyl-CoA reductase and phenylacetyl-CoA:acceptor oxidoreductase from
Thauera aromatica (22, 23). Ethylbenzene dehydrogenase
activity was already saturated at extremely low ethylbenzene
concentrations, which prevented the accurate determination of the
Km value for ethylbenzene. An approximation of <2
µM for the Km value was derived from
the residual substrate concentration under half-saturating turnover
conditions (for estimation, see Fig. 5).
About 15% of the activity recorded with ethylbenzene was obtained when
n-propylbenzene was used as substrate. The ratio of
activities with the two substrates did not change significantly between
cell extract and purified enzyme. Other aromatic substrates, such as toluene, p-cymene, phenylacetate,
(R,S)-1-phenylethanol or 2-phenylethanol, were
not oxidized by ethylbenzene dehydrogenase. The potential of
ethylbenzene dehydrogenase to catalyze the reversed reaction was tested
by an anaerobic enzyme assay with reduced methyl viologen as electron
donor and (S)-1-phenylethanol as starting substrate. No
reduction of (S)-1-phenylethanol to ethylbenzene was
detected by this test, suggesting that the reaction of ethylbenzene
dehydrogenase is irreversible under physiological conditions.
Ethylbenzene dehydrogenase also did not catalyze methyl
viologen-dependent reduction of selenate or nitrate,
despite the strong similarity of the N-terminal sequences of the
respective
subunits.
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Inhibition of Ethylbenzene Dehydrogenase--
Ethylbenzene
dehydrogenase was not inhibited in assays containing sodium azide or
sodium cyanide (1 mM each). Addition of cyanide to the
assay buffer caused a strong nonenzymatic background reaction, probably
by sequestering iron from the ferricenium into a cyanide complex, but
enzyme activities were still measurable reliably after starting with
ethylbenzene. Whereas ethylbenzene dehydrogenase activity in cell
extracts was not affected by aerobic extract preparation and incubation
in air for up to 12 h, purified enzyme, which was apparently in
the reduced state (see above), was inactivated irreversibly by
incubation in air with a half-life time of 7 min. This inactivation was
prevented efficiently by addition of the artificial electron acceptor
ferricenium hexafluorophosphate (1 mM) to the enzyme
preparations. Under these conditions, >90% of the enzyme activity was
still present after 2 h and >70% after a 6-h incubation in air.
Because ferricenium has been shown to convert the heme cofactor of
ethylbenzene dehydrogenase to the oxidized form, it can be concluded
that the reduced heme is most likely responsible for inactivation by
oxygen. The enzyme is probably held oxidized in cell extracts by
natural electron acceptors and acquires oxygen sensitivity when these
electron acceptors are removed during purification.
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DISCUSSION |
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The pathway of anaerobic ethylbenzene metabolism is initiated by two consecutive two-electron oxidation steps of ethylbenzene to (S)-1-phenylethanol and further to acetophenone (1-3). In this study, we analyzed the first enzyme of the predicted pathway, ethylbenzene dehydrogenase. To our knowledge, ethylbenzene dehydrogenase is the first described enzyme that catalyzes oxygen-independent hydroxylation of a hydrocarbon, namely, of an apolar compound without functional groups. Enantiomer specificity of ethylbenzene dehydrogenase was demonstrated by coupling the reaction to that of purified (S)-1-phenylethanol dehydrogenase. This matches the product reported previously for a different Azoarcus strain (4). Ethylbenzene dehydrogenase was induced specifically in cells grown anaerobically on ethylbenzene, and only very low activities were measured in cells grown on 1-phenylethanol or acetophenone. Three of the ethylbenzene-induced polypeptides described in this study showed molecular masses identical to those of the subunits of ethylbenzene dehydrogenase (96, 44, and 23 kDa).
Cells of the ethylbenzene-metabolizing strain EbN1 contained an enzyme exhibiting n-propylbenzene dehydrogenase activity at 15% of the specific activity measured with ethylbenzene. In contrast, cells of the ethylbenzene and n-propylbenzene-metabolizing strain PbN1 contained an enzyme exhibiting high activity with either ethylbenzene or n-propylbenzene. Similar ratios of activity with the two hydrocarbons in cells grown on either substrate strongly suggest that the same enzyme is used for metabolism of ethylbenzene and n-propylbenzene. This assumption is supported by the apparent identity of the polypeptide patterns of ethylbenzene an n-propylbenzene grown cells. Ethylbenzene dehydrogenase showed no activity with toluene as substrate, which is consistent with previous observations that strain EbN1 catabolizes toluene via a completely different pathway, namely, the addition of the methyl group to fumarate (5-7).
The present data suggest that ethylbenzene oxidation occurs in the periplasm, whereas the product, (S)-1-phenylethanol, is oxidized further in the cytoplasm, as evident from the use of NAD+ as electron acceptor. It is unknown presently whether and how 1-phenylethanol is transported into the cytoplasm. A passive diffusion of 1-phenylethanol as a hydrophobic compound via the cytoplasmic membrane appears principally possible. The advantage of a periplasmic location of ethylbenzene dehydrogenase for the organism is unknown. One may speculate that scavenging of the poorly water-soluble ethylbenzene is more effective outside of the cytoplasmic membrane. The low Km value (<2 µM) of ethylbenzene dehydrogenase supports the assumption that the capacity of the enzyme for effective substrate binding is an important factor in the metabolism of this hydrocarbon.
Even though the redox potential of the 1-phenylethanol/ethylbenzene
couple is around +0.03 V relative to standard hydrogen electrode
(estimated from thermodynamic data of other hypothetical alcohol/hydrocarbon couples), ethylbenzene oxidation was only observed
in this study with an electron acceptor of significantly higher redox
potential (ferricenium/ferrocene, E0 = +0.38 V).
One may assume that redox centers in the enzyme as well as the natural
electron acceptor have high redox potentials to achieve reasonable
oxidation rates with the relatively inert hydrocarbon substrate. Redox
centers of high midpoint potential in the enzyme may offer a possible
explanation for the observed irreversibility of ethylbenzene
dehydrogenation, even in tests with strongly reducing electron donors
(e.g. methyl viologen+/methyl viologen,
E0 = 0.446 V). Because ethylbenzene
dehydrogenase is a periplasmic enzyme, a possible natural acceptor in
the Azoarcus strains could be cytochrome c. The
properties of ethylbenzene dehydrogenase from strain EbN1 are in
contrast to the recently reported benzoquinone dependence of
ethylbenzene oxidation in strain EB-1. Reaction rates in this strain
were 3-fold lower than those reported here, and the enzyme was
membrane-associated and not induced in ethylbenzene-grown cells (4).
Apparently, there are different types of ethylbenzene dehydrogenases in
different strains of denitrifiers.
Ethylbenzene dehydrogenase is a new molybdenum/iron-sulfur/heme enzyme,
which is composed of three subunits. In analogy to other known
three-subunit molybdoenzymes, it may be assumed that the subunit
contains the molybdenum cofactor; the
subunit carries the Fe/S
clusters; and the
subunit, the heme cofactor. The enzyme, which is
most similar to ethylbenzene dehydrogenase with respect to subunit
composition, cofactor content and location, is presumably the recently
characterized selenate reductase from a closely related species,
T. selenatis (19, 20). However, ethylbenzene dehydrogenase
did not catalyze reduction of selenate or nitrate. Another recently
characterized molybdoprotein, phenylacetyl-CoA:acceptor oxidoreductase
from T. aromatica, catalyzes a similar dehydrogenation reaction with a polar aromatic substrate, but this enzyme is
membrane-bound, devoid of a heme cofactor, and apparently catalyzes the
four-electron oxidation of phenylacetyl-CoA to phenylglyoxylate, with
coupling to quinones and without release of intermediates (23).
A striking finding is the discrepancy between the stability of
ethylbenzene dehydrogenase under air in cell extracts and the fast
inactivation of the purified enzyme by oxygen. We showed that the
enzyme becomes relatively insensitive to air by anaerobic oxidation of
the heme b cofactor, suggesting that the fully reduced heme in the
enzyme may generate damaging oxygen metabolites. In cell extract,
ethylbenzene dehydrogenase is probably kept oxidized by interaction
with its natural electron acceptor and only becomes reduced when the
electron acceptor is removed during purification.
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ACKNOWLEDGEMENTS |
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We thank G. Fuchs (Universität Freiburg) and F. Widdel (MPI Bremen) for constant support and fruitful discussions. We also thank B. Auxier (University of Georgia, Athens) for ICP-OES analysis and H. Schägger (Universität Frankfurt) and Toplab, Martinsried, for N-terminal sequencing.
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FOOTNOTES |
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* This work was supported by the Deutsche Forschungsgemeinschaft and the Max-Planck-Gesellschaft.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ To whom correspondence should be addressed. Tel.: 49-761-203-2774; Fax: 49-761-203-2626; E-mail heiderj@uni-freiburg.de.
Published, JBC Papers in Press, April 9, 2001, DOI 10.1074/jbc.M101679200
1 O. Kniemeyer and J. Heider, submitted for publication.
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ABBREVIATIONS |
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The abbreviations used are: PAGE, polyacrylamide gel electrophoresis; ICP-OES, inductively coupled plasma optical emission spectroscopy.
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