Báo cáo khoa học: Adventitious reactions of alkene monooxygenase reveal common reaction pathways and component interactions among bacterial hydrocarbon oxygenases ppt

Adventitious reactions of alkene monooxygenase reveal
common reaction pathways and component interactions
among bacterial hydrocarbon oxygenases
William L. J. Fosdike
1
, Thomas J. Smith
1,2
and Howard Dalton
1
1 Department of Biological Sciences, University of Warwick, Coventry, UK
2 Biomedical Research Centre, Sheffield Hallam University, UK
Alkene monooxygenase (AMO) (EC 1.14.13.69), of
Rhodococcus rhodochrous (formerly Nocardia corallina)
B-276 belongs to a family of soluble multicomponent
oxygenases that possess a binuclear iron active centre
[1–5]. This group of enzymes includes AMOs from two
other bacterial sources [6–8], the soluble methane
monooxygenases (sMMOs) (EC 1.14.13.25) produced
by certain methanotrophic bacteria [9] and a range of
oxygenases that perform aromatic ring hydroxylations
with a variety of specificities [3,10]. All enzymes for
which data are available have been found to comprise
at least three components: (1) multisubunit binuclear
iron active centre-containing terminal oxygenase,
where oxygen activation and substrate oxygenation
occur; (2) an NAD(P)H-dependent reductase that sup-
plies electrons to the active centre of the terminal
Keywords
alkene monooxygenase; alkyne; component
interactions; peroxide shunt reaction;
turnover-dependent inhibition

Correspondence
H. Dalton, Department of Biological
Sciences, University of Warwick, Coventry
CV4 7AL, UK
Fax: +44 24 76523568
Tel: +44 24 76523552
E-mail:
Website: />frame.asp?id=4;
/>bmrc/tomsmith.htm
(Received 24 January 2005, revised
14 March 2005, accepted 21 March 2005)
doi:10.1111/j.1742-4658.2005.04675.x
Alkene monooxygenase (AMO) from Rhodococcus rhodochrous (formerly
Nocardia corallina) B-276 belongs to a family of multicomponent nonheme
binuclear iron-centre oxygenases that includes the soluble methane mono-
oxygenases (sMMOs) found in some methane-oxidizing bacteria. The
enzymes catalyse the insertion of oxygen into organic substrates (mostly
hydrocarbons) at the expense of O
2
and NAD(P)H. AMO is remarkable in
its ability to oxidize low molecular-mass alkenes to their corresponding
epoxides with high enantiomeric excess. sMMO and other well-character-
ized homologues of AMO exhibit two adventitious activities: (1) turnover-
dependent inhibition by alkynes and (2) activation by hydrogen peroxide in
lieu of oxygen and NAD(P)H (the peroxide shunt reaction). Previous stud-
ies of the AMO had failed to detect these activities and opened the possi-
bility that the mechanism of AMO might be fundamentally different from
that of its homologues. Thanks to improvements in the protocols for culti-
vation of R. rhodochrous B-276 and purification and assay of AMO, it has
been possible to detect and characterize turnover-dependent inhibition of

AMO by propyne and ethyne and activation of the enzyme by hydrogen
peroxide. These results indicate a similar mechanism to that found in
sMMO and also, unexpectedly, that the enantiomeric excess of the chiral
epoxypropane product is significantly reduced during the peroxide shunt
reaction. Inhibition of the oxygen⁄ NADH-activated reaction, but not the
peroxide shunt, by covalent modification of positively charged groups
revealed an additional similarity to sMMO and may indicate very similar
patterns of intersubunit interactions and ⁄ or electron transfer in both
enzyme complexes.
Abbreviations
AMO, alkene monooxygenase; sMMO, soluble methane monooxygenase; sulfo-NHS-acetate, sulfosuccinimidyl acetate.
FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS 2661
oxygenase; and (3) a small component, known as the
coupling or gating protein, which is also required for
full activity [3–5]. The terminal oxygenase components
of soluble methane monooxygenases [9,11], aromatic
monooxygenases [10] and the AMO of Xanthobacter
sp. strain Py2 [12] all have an (abc)
2
quaternary struc-
ture, whereas the AMOs from R. rhodochrous B-276
[13] and Mycobacterium sp. [8] lack the c subunit. In
addition, the aromatic monooxygenases and AMOs
from Xanthobacter Py2 and Mycobacterium sp. also
possess a Rieske iron-sulfur protein that appears to
pass electrons between the reductase and terminal
oxygenase [12,14–16].
The binuclear iron-centre monooxygenases are char-
acterized by close biochemical and, apparently, struc-
tural similarities. There is complete conservation of

the protein ligands to the binuclear iron site (four glu-
tamyl and two histidyl residues) and all catalyse
similar hydrocarbon monooxygenation reactions at the
expense of NADH and dioxygen [3–5]. The enzymes
show a gamut of substrate ranges and regio- and
stereo-selectivities. For instance, the substrate range of
AMO from R. rhodochrous B-276 is restricted almost
exclusively to alkenes [17], whereas sMMOs have a
remarkably wide range of substrates that includes alka-
nes, alkenes and aromatic compounds [18]. Consistent
with their different enzymatic activities, the terminal
oxygenase components of AMOs are termed epoxygen-
ases as their products are epoxides; the equivalent
components of the other enzymes add hydroxyl groups
to their natural substrates and are therefore known as
hydroxylases. There are also important differences in
the enantiopurity of products obtained from oxygen-
ation of alkene substrates: AMO from R. rhodochrous
B-276 epoxygenates propene to R-epoxypropane
with high enantiomeric excess (83%) [1,13], whereas
sMMO produces a nearly racemic mixture of products
with the same substrate (S.E. Slade, T.J. Smith and
H. Dalton, unpublished observations).
In addition to these oxygenation reactions, several
well characterized binuclear iron-centre monooxygen-
ases have been found to exhibit two adventitious reac-
tions: turnover-dependent inhibition by alkynes and
the so-called peroxide shunt reaction. Terminal alkynes
such as ethyne have been shown to act as suicide sub-
strates of sMMO [19], soluble butane monooxygenase

[20], several aromatic monooxygenases [21,22] and the
AMO from Xanthobacter sp. Py2 [12,23], presumably
by oxygenation to ketenes that then covalently modify
and inactivate the enzymes [19]. Irreversible inhibition
of the heme active-site monooxygenases of the cyto-
chrome P450 family, which are not related to the binu-
clear iron centre-containing enzymes, may also proceed
via similar ketene intermediates [24]. The peroxide
shunt reaction allows the terminal oxygenase compo-
nent to perform oxygenation reactions in the absence
of other protein components and NAD(P)H, if the
oxidant is hydrogen peroxide rather than dioxygen.
The peroxide shunt has been observed in sMMO
[25,26] and toluene 2-monooxygenase [27], as well as
unrelated monooxygenases of the cytochrome P450
family [28]. In sMMO it has been shown that the
whole-complex (i.e. NADH-dependent) and peroxide-
activated activities can be functionally separated by
covalently modifying positively charged groups on the
hydroxylase, which inhibits the whole-complex reaction
but not the peroxide shunt [29].
Neither of these adventitious activities has to our
knowledge previously been reported in the rhodococcal
AMO. The rhodococcal AMO was previously found to
be inhibited by propyne, but the inhibition appeared
to be competitive because K
m
for propene oxygen-
ation was increased by the inhibitor but V
max

was
unchanged [1]. This, together with the lack of any pub-
lished account of the peroxide shunt reaction in the
rhodococcal AMO, opened the possibility of funda-
mental differences in reaction mechanism and ⁄ or sub-
strate selectivity between AMO and its homologues. In
addition to their possible mechanistic significance,
these inferences had potential implications for com-
mercial application of the enzyme. An AMO that was
not destroyed by alkynes would be tolerant of alkyne
contamination of the alkene feedstock, whereas one
that did not exhibit the peroxide shunt reaction could
not be economically employed in a cell-free system
without a means for regeneration of NADH. In the
context of a whole-cell biocatalyst that would bypass
problems with coenzyme regeneration, we began the
present study of AMO by investigating the effect of
alkynes on AMO-dependent growth of R. rhodochrous
B-276. Subsequently, by refining the purification, inhi-
bition and activity assay protocols we have been able
to produce large amounts of pure high-activity AMO
and to perform a more thorough investigation of its
interaction with alkynes and hydrogen peroxide than
has previously been possible.
Results
Alkynes strongly inhibit growth of
R. rhodochrous B-276 on propene
The effect of alkynes on whole-cell systems expressing
AMO was investigated by monitoring the growth of
flask cultures of R. rhodochrous B-276 in the presence

of a range of alkynes (Fig. 1). A fivefold molar excess
Adventitious reactions of alkene monooxygenase W. L. J. Fosdike et al.
2662 FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS
of propene over the alkyne was used as the carbon and
energy source to ensure that cell growth was dependent
on AMO. The observed complete inhibition of growth
in the presence of propyne and but-1-yne was qualita-
tively a much more severe effect than was expected from
the relatively low level (£ 70%) of competitive inhibition
that the previous data implied [1] and suggested either
that these alkenes were inhibiting AMO to a greater
extent than had been observed in the purified enzyme
experiments or that some other essential metabolic pro-
cess was inhibited by alkynes. Even more remarkably,
ethyne (which barely inhibited purified AMO in the
previous study [1]), caused significantly delayed growth
and reduced growth rate relative to the control.
Preincubation of AMO with alkyne and NADH
allowed turnover-dependent inhibition to be
observed
In order to explain the unexpectedly large effect of
alkynes on the growth of AMO-expressing R. rhodo-
chrous B-276 cells on propene, the effect of alkynes
on the activity of purified AMO was re-examined.
Previously, the inhibition properties of AMO were
investigated in reactions in which propene and the
alkyne were present simultaneously and were added
before NADH, which is essential for turnover of
the AMO complex [1,13]. Hence, although the effect
of propyne in causing an increase in K

m
with respect
to propene without significant change in V
max
was
consistent with reversible competitive inhibition, turn-
over-dependent and -independent events could not
be distinguished. Consequently, the nature of the
inhibition could not be unambiguously assigned. To
resolve this uncertainty, purified AMO components
and NADH were preincubated aerobically in the pres-
ence of ethyne or propyne (5% v ⁄ v in the headspace)
and subsequent assay of the propene oxygenation
activity after removal of the alkyne clearly demonstra-
ted irreversible inhibition at 80% (Table 1). It was
clear that this inhibition was predominantly turnover-
dependent because omission of NADH during the pre-
incubation phase completely abolished it. Conversely,
when a much higher concentration of ethyne (50%
v ⁄ v, corresponding to an increase in calculated liquid-
phase concentration from 1.8 mm to 18 mm) and pro-
pene were added at the same time, before NADH, no
inhibition was observed during a 10-min assay. A sim-
ilar assay using propyne (35% v ⁄ v, corresponding to
an increase in calculated liquid-phase concentration
from 3.1 mm to 22 mm) in place of ethyne resulted
in only a 50% reduction in epoxide formation over a
10-min assay, relative to the control in which nitrogen
was substituted for the alkyne. The fact that the pres-
ence of propene protects against inhibition by the alky-

nes is consistent with the alkynes’ acting as suicide
substrates which compete for the same active site on
the enzyme as the natural substrate propene.
Residual AMO activity was measured as a function
of the time between the start of the reaction (addition
of NADH) and removal of the alkyne by flushing with
nitrogen. First-order decay of enzyme activity was
observed, from which first order decay constants for
the inactivation of the enzyme by propyne and ethyne
under these conditions could be calculated (Fig. 2). It
is likely that the linear part of the graph in Fig. 2 does
not cross the ordinate at the position corresponding to
the uninhibited enzyme activity because the measured
reaction times do not include the time taken to remove
the alkyne during the flushing process and are there-
fore uniformly underestimated. The positive deviation
of the latest data points from the extrapolated linear
Fig. 1. Effect of alkynes on the growth of R. rhodochrous B-276
using propene as the growth substrate. Cultures were incubated
aerobically in the presence of propene plus nitrogen (solid symbols,
solid line), ethyne (solid symbols, dotted line), propyne (open sym-
bols, solid line) and but-1-yne (open symbols, dotted line).
Table 1. Turnover-dependent inhibition of AMO by alkynes.
Preincubation before assay
a
Percent activity
b
None 100 ± 10
Ethyne 96 ± 12
Propyne 85 ± 8

Ethyne + NADH 18 ± 4
Propyne + NADH 21 ± 2
a
Reactions contained 8 lM of epoxygenase and 12 lM each of
reductase and GST-coupling protein fusion in a total volume of
100 lL; the headspace concentration of alkyne was 11% (v ⁄ v) and
the reaction was preincubated for 10 min before removal of the
alkyne and assaying with propene as the substrate, in the presence
of O
2
and NADH, as described in the Experimental procedures.
b
Specific activities are given as percentages of the uninhibited
activity of 192 nmolÆmin
)1
Æ(mg of epoxygenase)
)1
.
W. L. J. Fosdike et al. Adventitious reactions of alkene monooxygenase
FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS 2663
portion possibly reflects the exhaustion of alkyne later
in the reaction, if a number of turnovers are required
per inactivation event.
Improved epoxygenase preparations allowed
detection of the AMO-catalysed peroxide shunt
reaction
Investigation of the effect of hydrogen peroxide on
AMO was facilitated by the improved protocols for
growth of R. rhodochrous B-276 and purification of the
binuclear iron site-containing epoxygenase component

described in the Experimental procedures. Use of pro-
pene as the sole carbon and energy source ensured a
high level of AMO expression by making cell growth
dependent on AMO. The subsequent purification pro-
tocol yielded > 95% pure epoxygenase with a specific
activity of at least 145 nmolÆmin
)1
Æ[mg of protein]
)1
,
which was more than twice as active as previous pre-
parations produced from cells grown in rich medium
[1,13]. In addition, the use of smaller assay reaction
volumes (0.1 mL rather than 0.5 mL used previously)
enabled extensive investigations to be carried out at
high epoxygenase concentrations. Thanks to these
improvements, the peroxide shunt reaction was found
to be detectable and easily quantifiable at 40 lm
epoxygenase after only 3 min reaction time, using
500 mm hydrogen peroxide and propene at 37% (v ⁄ v)
in the headspace gas. The specific activity of the
hydrogen peroxide-activated reaction under these con-
ditions was 32 nmolÆmin
)1
Æ[mg of epoxygenase]
)1
, i.e.
22% of the NADH-dependent reaction catalysed by
the whole AMO complex. The amount of epoxypro-
pane produced was unchanged when the reaction was

purged of oxygen by flushing with oxygen-free nitro-
gen for 5 min before addition of the hydrogen per-
oxide, showing that the reaction did not require
molecular oxygen. Formation of product was depend-
ent on the presence of hydrogen peroxide and, more
importantly, active epoxygenase. Omission or prior
heat-denaturation (100 °C, 5 min) of the epoxygenase
completely abolished formation of epoxypropane
under otherwise identical conditions.
The rate of epoxypropane formation via the peroxide
shunt reaction was linear with epoxygenase concentra-
tion between 20 and 60 lm (data not shown). When a
reaction time of 3 min was used, the reaction rate was
linear with hydrogen peroxide concentration up to
1.0 m, the highest concentration tested, suggesting that
the K
m
for hydrogen peroxide was considerably greater
than 1 m. This contrasts with the lower value of K
m
for
hydrogen peroxide of 66 mm estimated from experi-
ments with the corresponding hydroxylase component
of sMMO [25]. When longer reaction times were used,
the average rate did not increase beyond about 500 mm
hydrogen peroxide (Fig. 3), probably because of pro-
gressive inactivation of the enzyme by higher concentra-
tions of hydrogen peroxide in a manner that was also
observed with sMMO [25]. The AMO epoxygenase-
catalysed peroxide shunt reaction showed moderate

inhibition by the coupling protein component of
AMO (29 ± 8% inhibition relative to an activity of
30 ± 2 nmolÆmin
)1
Æ[mg of epoxygenase]
)1
at a coupling
protein ⁄ epoxygenase molar ratio of 3 : 1), again similar
to the result obtained with sMMO [25].
Fig. 2. Kinetics of inactivation of AMO by (A) propyne and (B)
ethyne. Alkynes were added to the headspace at 6.25% (v ⁄ v), which
was calculated to give equilibrium aqueous-phase concentrations
of propyne and ethyne of 3.9 and 2.2 m
M, respectively. Residual
AMO activity was measured after removal of the alkyne using pro-
pene as the substrate and are derived from single-timepoint meas-
urements of the product after 10 min reaction time; 100% activity
corresponded to 153 nmolÆmin
)1
Æ(mg of epoxygenase)
)1
. Error bars
show standard deviation from three experiments. First order decay
constants during the exponential decay periods were 0.083 min
)1
and 0.13 min
)1
for propyne and ethyne, respectively. The zero-time
measurement is derived from enzyme that was not exposed to the
alkyne.

Adventitious reactions of alkene monooxygenase W. L. J. Fosdike et al.
2664 FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS
The peroxide shunt reaction in AMO showed low
stereoselectivity even in the presence of the
coupling protein
When epoxypropane produced via the whole-complex
reaction was subjected to chiral analysis, the R-enantio-
mer predominated with an enantiomeric excess >80%,
consistent with the 83% previously observed [1]. In
contrast, epoxypropane produced by the AMO epoxy-
genase via the peroxide shunt reaction, whilst still
showing a predominance of the R-enantiomer, had an
enantiomeric excess of only 25%.
In sMMO from Methylosinus trichosporium OB3b
[30], it has been observed that the protein B compo-
nent, equivalent to the coupling protein of AMO,
influences the regioselectivity of the enzyme. If the
coupling protein influenced product distribution in
AMO also, it was possible that the low stereoselecti-
vity observed via the peroxide shunt was due to the
absence of the coupling protein. It was found that the
presence of the coupling protein (up to a threefold
molar excess relative to the epoxygenase) had no signi-
ficant effect on the chiral composition of the epoxypro-
pane product in the peroxide shunt reaction, although
the possibility that the coupling protein was damaged
by the high concentration of hydrogen peroxide used
in this experiment cannot be excluded.
Modification of positively charged residues on
the surface of the epoxygenase allowed

functional separation of the whole-complex
and peroxide shunt activities
As treatment of the hydroxylase component of
sMMO (equivalent to the epoxygenase of AMO)
with reagents specific for positively charged moieties
completely inhibited the whole-complex reaction but
not the peroxide shunt [29], covalent modification of
AMO afforded an additional test of its biochemical
similarity to sMMO. When the epoxygenase of
AMO was reacted with the primary amine-specific
reagent sulfo-NHS-acetate, the whole complex reac-
tion was abolished whilst the activity via the per-
oxide shunt was unaffected, as in the sMMO system.
The control experiment described in the Experimental
procedures confirmed that the specific inactivation of
the whole-complex reaction was due solely to the
effect of the sulfo-NHS-acetate on the epoxygenase
component and not due to the effect of any carried
over reagent during the assay reaction. A similar
functional separation of the peroxide shunt and
whole-complex reactions was observed by using the
arginine-specific reagent p-hydroxyphenylglyoxal, which
did not significantly inhibit the peroxide shunt reac-
tion but resulted in progressive inactivation of the
whole-complex reaction (data not shown). These
results suggested that accessible positively charged
residues were necessary for interactions between the
enzyme components but not per se for substrate oxy-
genation at the active site. Consistent with the hypo-
thesis that protein–protein interactions between the

AMO components require positive charges on the
epoxygenase, chemical modification of the coupling
protein with sulfo-NHS-acetate or p-hydroxyphenyl-
glyoxal had no effect on its activity in the whole-
complex propene oxygenation reaction (data not
shown).
Discussion
The observations that the rhodococcal AMO, like
other binuclear iron-centre monooxygenases, exhibits
turnover-dependent inhibition by alkynes and can be
activated by hydrogen peroxide support a unified
mechanism for oxygenation reactions catalysed by this
family of enzymes. The results of Gallagher et al. [1],
where it was observed using partially purified AMO
that propyne increased the apparent K
m
with respect
to propene but left V
max
unchanged, can now be
reinterpreted as showing that competition between the
substrate propene and the inhibitor propyne not only
alleviates inhibition at high substrate concentration by
preventing propyne from blocking the active site but in
so doing also protects the enzyme from the irreversible
inhibition that would result from turnover of the
alkyne. The relatively small amount of inhibition of
growth and AMO activity seen when ethyne was pre-
sent at the same time as propene are consistent with
the conclusion that the two-carbon ethyne competes

Fig. 3. Kinetics of peroxygenation of propene catalysed by the
AMO epoxygenase. Activity was measured by quantifying epoxy-
propane formation as described in the Experimental procedures,
using 40 l
M epoxygenase and reaction times of 3 min (solid line),
10 min (dotted line) and 15 min (dashed line). Error bars show
standard deviations from three experiments.
W. L. J. Fosdike et al. Adventitious reactions of alkene monooxygenase
FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS 2665
less well with propene for the active site than the
three-carbon propyne. The approximately twofold dif-
ference in solubility of the two alkynes would not be
expected to account for the gross difference in inacti-
vation that was observed in competition with the pro-
pene substrate.
As it is now clear that the adventitious activities of
AMO are broadly similar to those of other binuclear
iron-centre monooxygenases, the range of activities
exhibited by this family of enzymes can be explained
by differences in substrate binding and the ability of
the highly oxygenating species at the active site to acti-
vate high energy C–H bonds such as those in benzene,
methane and other alkanes. The fact that AMO can be
activated via hydrogen peroxide is consistent with the
presence of a binuclear iron–peroxo intermediate sim-
ilar to that found during the catalytic cycle of sMMO
[31,32]. Whether, as previously proposed [33], AMO is
unable to oxidize methane because it lacks the charac-
teristic and probably diferryl (Fe
IV

2
) intermediate Q
observed in sMMO must await future studies.
Whatever the structural differences that underlie the
difference in active-site reactivity of the binuclear iron-
centre monooxygenases, it appears to be relatively
subtle as far as the immediate environment of the
binuclear iron site is concerned. Not only are the ligat-
ing residues perfectly conserved throughout the group
[3] but also the recent structure of the hydroxylase
component of toluene ⁄ o-xylene monooxygenase has
indicated that the only appreciable difference between
the active-centre ligation environments of this enzyme
and sMMO, which share only 24% identity in their
a-subunits, are relatively minor differences in active
site hydrogen bonding patterns and the coordinating
nitrogen atom of one histidine ligand [34].
The results obtained with the reagents specific for
positively charged groups reveal an additional level
of similarity between AMO and sMMO that allows
functional separation of the whole-complex and per-
oxide shunt reaction in both enzyme systems. In
both systems, dioxygen activation and ⁄ or functional
interactions between the enzyme components require
accessible positively charged moieties on the terminal
oxygenase, whereas the actual process of oxygen
insertion into the substrate does not. There may be
at least two positively charged moieties, perhaps
unidentified conserved residues, that are independ-
ently necessary for electron transfer and⁄ or intersub-

unit interactions in each enzyme because primary
amine- and arginine-specific reagents have similar
effects in both systems.
The diminished enantiomeric excess of epoxypro-
pane production observed when AMO was activated
via the peroxide shunt reaction is reminiscent of the
altered substrate specificity and product distribution
seen in the peroxide shunt reaction in sMMO
[25,30]. In sMMO from Methylosinus trichosporium
OB3b, the coupling protein component has been
shown to influence the regioselectivity of oxygenation
reactions [30] and mutagenesis and modelling studies
have suggested that the coupling protein interacts
directly with the hydroxylase active site and controls
substrate entry [35]. The binding site for the coup-
ling protein of AMO may be similarly positioned on
the epoxygenase. However, unless the coupling pro-
tein is damaged by the high concentration of hydro-
gen peroxide used, presence of the coupling protein
is clearly not the sole determinant of enantioselective
catalysis in AMO because addition of the coupling
protein to the AMO epoxygenase-catalysed peroxide
shunt reaction reduced the overall reaction rate but
did not increase the enantiomeric excess of the prod-
uct. Whilst the operation of the peroxide shunt
reaction in AMO presents an opportunity for devel-
opment of an AMO-based biocatalyst that is inde-
pendent of reduced coenzyme, the low product
enantiomeric excess obtained via the peroxide shunt
presents an important new question about the origin

of enantioselectivity in this enzyme and a challenge
to novel applications of AMO for chiral synthesis.
Experimental procedures
Bacterial strains and growth conditions
R. rhodochrous B-276 was cultivated at 30 °C in nitrate min-
imal salts liquid medium or on nitrate minimal salts agar
[36], using propene as the sole carbon and energy source.
Batch cultures for analysis of the effect of alkynes on growth
were performed in 1-L cultures in 2-L conical flasks (Quick-
fit, Fisher, Loughborough, UK), sealed with Subaseal
Ò
rub-
ber seals (W. Freeman, Barnsley, UK). After inoculation,
600 mL of headspace gas was removed and replaced by
500 mL of propene, plus 100 mL of the appropriate alkyne
or nitrogen, as stated for each experiment. Growth was
monitored over a 5-day incubation at 30 °C with shaking
(180 r.p.m.) by measuring the OD
600
of samples removed
through the seal with a hypodermic syringe. Large-scale con-
tinuous growth of R. rhodochrous B-276 for purification of
the AMO epoxygenase and reductase component was
achieved by using a 2000 Series fermentor (LH Engineering,
Stoke Poges, UK) with a working volume of 4 L and a dilu-
tion rate of 0.035 h
)1
. The culture was gassed with a 1 : 10
(v ⁄ v) propene ⁄ air mixture at a flow rate of 1 LÆh
À1

, agitated
at an imepellor speed of 450 r.p.m. and maintained at
pH 7.0. The Escherichia coli strain used to produce the
Adventitious reactions of alkene monooxygenase W. L. J. Fosdike et al.
2666 FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS
recombinant glutathione S-transferase (GST)-coupling pro-
tein fusion was cultivated as described previously [37].
Purification of the AMO components
All protein purification procedures were conducted at
0–4 °C. For purification of the AMO epoxygenase (i.e. ter-
minal oxygenase) and reductase components, cells from
20 L of propene-grown R. rhodochrous B-276 culture at an
OD
600
of 12–15 were harvested at 3000 g using a Westfalia
continuous centrifuge (Northvale, NJ, USA), washed by
centrifugation (10 000 g, 10 min) and resuspension in
10 mm MgSO
4
,5%(v⁄ v) glycerol, 25 mm Mops pH 7.5
and resuspended in a minimal volume of the same buffer
containing benzamidine (1 mm), dithiothreitol (1 mm) and
a total glycerol concentration of 15% (v ⁄ v) (buffer B).
Deoxyribonuclease I (Sigma, Gillingham, UK) was added
to 20 lgÆmL
)1
and the cells were broken by passing twice
through a high-pressure cell disrupter (172 MPa; Constant
Systems, Warwick, UK), after which the cell-free extract
was centrifuged (48 000 g, 1 h) and the supernatant (the

soluble extract) was removed.
The soluble extract was applied to a DE52 anion
exchange column (Whatman, Maidstone, UK; 5 cm ·
15 cm), previously equilibrated with buffer B. After wash-
ing with buffer B, the column was eluted with a step gradi-
ent of 150 and 250 mm NaCl in buffer B. The reductase
was purified from the 250-mm NaCl fraction as described
previously [13]. The 150-mm NaCl fraction was concentra-
ted to a volume of 200 mL using a 900 cm
2
surface-area
spiral-wound ultrafiltration cartridge (Amicon, Stonehouse,
UK) with a molecular size cut-off of 10 kDa. MgSO
4
was
added to a final concentration of 0.5 m and then the
epoxygenase-containing solution was applied to a Phenyl
Sepharose high-performance column (Amersham-Pharma-
cia, Little Chalfont, UK; 2.6 · 12 cm) previously equili-
brated with buffer D (25 mm Mops pH 7.5 plus
benzamidine and dithiothreitol, each at 1 mm) containing
0.5 m MgSO
4
. Proteins were eluted with a linear gradient
of 0.5–0 m MgSO
4
in buffer D. Fractions containing
epoxygenase activity, which eluted at 0 mm MgSO
4
, were

adjusted to an MgSO
4
concentration of 10 mm and concen-
trated using a 30 kDa molecular size cut-off membrane
(Amicon). The phenyl Sepharose column was washed with
0.7% (w ⁄ v) sodium cholate after use to remove residual
protein and prevent loss of epoxygenase yield during subse-
quent preparations. The concentrated epoxygenase was
applied to a Mono Q anion exchange column (Amersham-
Pharmacia; 1 · 10 cm) that had been equilibrated with
25 mm Mops pH 7.5 containing 15% (v ⁄ v) glycerol and
10 mm MgSO
4
and then proteins were eluted with a lin-
ear gradient of 0–400 mm NaCl in the same buffer. The
pure epoxygenase eluted at 300 mm NaCl.
The coupling protein used throughout this study, which
was a recombinant GST-coupling protein fusion that is
fully active with the GST tag attached, was produced in
Escherichia coli and purified by affinity chromatography
[37].
AMO assays and inhibition studies
AMO assays and alkyne inhibition reactions were per-
formed in 100-lL reaction volumes in 1.7-mL crimp-seal
glass vials. For measurement of activity via the whole-
complex AMO reaction, the three AMO components were
mixed with 25 mm Mops pH 7.5 containing 10 mm MgSO
4
to give 8 lm epoxygenase, 12 lm reductase and 12 lm
coupling protein. The vial was then sealed and 0.7 mL of

the headspace gas was removed and replaced with 0.7 mL
of propene, after which the vial was preincubated at 30 °C
for 30 s. The reaction was initiated by adding NADH (to
100 lm) and the vial was shaken (180 r.p.m.) at 30 °C for a
further 3 min, unless otherwise stated, before the epoxypro-
pane formed was quantified by gas chromatography of
0.5-mL gas phase samples, as described previously [1].
Activity assays of the individual AMO components during
purification were performed in the presence of an excess of
the other two components. Peroxide shunt assays were per-
formed in a similar manner to the whole-complex reactions
except that reductase and (except where stated otherwise)
the coupling protein were omitted and the reaction was
started by adding hydrogen peroxide instead of NADH.
The reaction time and concentrations of protein compo-
nents and hydrogen peroxide are stated for each experi-
ment.
Inhibition of the AMO complex by alkynes was
achieved as follows. Epoxygenase (8 lm), reductase
(12 lm) and coupling protein (12 lm) were mixed and the
reaction vial was sealed. Headspace gas was removed and
replaced with an equal volume of the alkyne, to give the
alkyne concentration stated for each experiment. Concen-
trations of alkynes in the aqueous phase were calculated
using Henry’s law and Henry’s constants of 0.068 mÆatm
)1
[38] and 0.037 mÆatm
)1
( />$sander/res/henry.html), respectively, for propyne and eth-
yne. The vial was preincubated with shaking at 30 °C for

30 s and then, except where stated otherwise, NADH was
added to 100 lm. Turnover-dependent inhibition was then
allowed to proceed under the same incubation conditions,
for 10 min unless stated otherwise. Unreacted alkyne was
removed by flushing with nitrogen for 5 min; the vial was
then flushed with air and the remaining AMO activity
was assayed by replacing 0.7 mL of the headspace gas
with 0.7 mL of propene and incubating at 30 °C for a fur-
ther 10 min, before removing 0.5 mL of the headspace gas
for quantitation of the epoxypropane product by gas chro-
matography as above. Residual propene oxidation activity
was measured after 10 min reaction with the alkyne, at
a range of concentrations, in the presence of NADH
(100 lm). The decay of enzyme activity as a function of
W. L. J. Fosdike et al. Adventitious reactions of alkene monooxygenase
FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS 2667
time was analysed by plotting the natural logarithm of the
residual activity against time, to yield the apparent first-
order decay constant, k_app, from the slope of the linear
portion of the graph [39].
Chiral analysis
Reactions for analysis of chiral composition were scaled up
to 1-mL volume and incubated at 30 °C until the epoxypro-
pane concentration was about 2 mm. Epoxypropane was
extracted using 200–300 lL of diethyl ether, which was
dried using molecular sieve and analysed by means of capil-
lary GC using a Phillips 4500 GC apparatus fitted with a
Chiraldex (20 m · 0.25 mm) a-cyclodextrin trifluoroacetate
column (Advanced Separation Technologies, Whippany,
NJ, USA). The injection volume was 1 lL, the split ratio

was 100 : 1 and the carrier gas (nitrogen) flow rate was
1.3 mLÆmin
)1
. The injector and flame ionization detector
were maintained at 150 °C and the column temperature
was 30 °C.
Modification of positively charged groups
Covalent modification of the AMO epoxygenase and coup-
ling protein was effected using protocols based on those
used with sMMO [29], as follows. Accessible primary
amine groups were modified by reacting the epoxygenase or
coupling protein (10 mgÆmL
)1
) with a 13-fold molar excess
of sulfosuccinimidyl acetate (sulfo-NHS-acetate; Pierce,
Rockford, IL, USA) at room temperature for 30 min. Un-
reacted sulfo-NHS-acetate was removed by buffer exchange
via three cycles of ultrafiltration [using a Microcon centrifu-
gal concentrator (Amicon)] and dilution. In the case of the
epoxygenase, a 30 kDa cut-off membrane was used and
each time the sample was diluted with a volume equal to
the original sample volume of 25 mm Mops pH 7.5 con-
taining 15% (v ⁄ v) glycerol. In the case of the coupling pro-
tein, the membrane had a 10 kDa cut-off and glycerol was
omitted from the dilution buffer. The reaction between the
epoxygenase and sulfo-NHS-acetate abolished the activity
of the epoxygenase in the AMO whole-complex reaction
and so a control was performed to confirm that the
observed inhibition was due to reaction of the sulfo-NHS-
acetate with the epoxygenase and not due to reaction of

carried over reagent with the other AMO components.
Here, the epoxygenase was not added to the reaction until
after the sulfo-NHS-acetate had been removed by buffer
exchange. No inhibition of the reaction was observed, con-
firming that the buffer exchange procedure was effective in
removing the sulfo-NHS-acetate and showing that inhibi-
tion required contact between the epoxygenase and sulfo-
NHS-acetate. Modification of accessible arginyl side-chain
guanidinium groups was effected by reacting the epoxyge-
nase or coupling protein (10 mgÆmL
)1
) with p-hydroxyphe-
nylglyoxal (12 mm) for 20 min at room temperature, after
which the reaction was quenched by adding arginine to
60 mm.
References
1 Gallagher SC, Cammack R & Dalton H (1997) Alkene
monooxygenase from Nocardia corallina B-276 is a
member of the class of dinuclear iron proteins capable
of stereospecific epoxygenation reactions. Eur J Biochem
247, 635–641.
2 Saeki H & Furuhashi K (1994) Cloning and characteri-
sation of the Nocardia corallina B-276 gene cluster
encoding alkene monooxygenase. J Ferment Bioeng 78,
399–406.
3 Leahy JG, Batchelor PJ & Morcomb SM (2003) Evolu-
tion of the soluble diiron monooxygenases. FEMS
Microbiol Rev 27, 449–479.
4 Kotani T, Yamamoto T, Yurimoto H, Sakai Y & Kato
N (2003) Propane monooxygenase and NAD

+
-depend-
ent secondary alcohol dehydrogenase in propane
metabolism Gordonia sp. strain TY-5. J Bacteriol 185,
7120–7128.
5 Sluis MK, Sayavedra-Soto LA & Arp DJ (2002) Mole-
cular analysis of the soluble butane monooxygenase
from ‘Pseudomonas butanovora’. Microbiology 148,
3617–3629.
6 Zhou NY, Jenkins A, Chion CKNCK & Leak DJ
(1999) The alkene monooxygenase from Xanthobacter
strain Py2 is closely related to aromatic monooxy-
genases and catalyzes aromatic monohydroxylation of
benzene, toluene, and phenol. Appl Environ Microbiol
65, 1589–1595.
7 Hartmans S, Weber FJ, Somhorst DPN & de Bont
JAM (1991) Alkene monooxygenase from Mycobacter-
ium – a multicomponent enzyme. J Gen Microbiol 137,
2555–2560.
8 Coleman NV & Spain JC (2003) Distribution of the
coenzyme M pathway of epoxide metabolism among
ethene- and vinyl chloride-degrading Mycobacterium
strains. Appl Environ Microbiol 69, 6041–6046.
9 Murrell JC, Gilbert B & McDonald IR (2000) Molecu-
lar biology and regulation of methane monooxygenase.
Arch Microbiol 173, 325–332.
10 Johnson GR & Olsen RH (1995) Nucleotide-sequence
analysis of genes encoding a toluene benzene-2-monooxy-
genase from Pseudomonas sp. strain JS150. Appl Environ
Microbiol 61, 3336–3346.

11 Lipscomb JD (1994) Biochemistry of the soluble
methane monooxygenase. Annu Rev Microbiol 48,
371–399.
12 Small FJ & Ensign SA (1997) Alkene monooxygenase
from Xanthobacter strain Py2 – purification and charac-
terization of a four-component system central to the
bacterial metabolism of aliphatic alkenes. J Biol Chem
272, 24913–24920.
Adventitious reactions of alkene monooxygenase W. L. J. Fosdike et al.
2668 FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS
13 Miura A & Dalton H (1995) Purification and characteri-
sation of the alkene monooxygenase from Nocardia
corallina B-276. Biosci Biotechn Biochem 59, 853–859.
14 Pikus JD, Studts JM, Achim C, Kauffmann KE, Mu
¨
nck
E, Steffan RJ, McClay K & Fox BG (1996) Recombi-
nant toluene 4-monooxygenase: catalytic and Mo
¨
ss-
bauer studies of the purified diiron and Rieske
components of a four-protein complex. Biochemistry 35,
9106–9119.
15 Byrne AM, Kukor JJ & Olsen RH (1995) Sequence analy-
sis of the gene cluster encoding toluene-3-monooxygenase
from Pseudomonas pickettii PKO1. Gene 154, 65–70.
16 Bertoni G, Martino M, Galli E & Barbieri P (1998)
Analysis of the gene cluster encoding toluene ⁄ o-xylene
monooxygenase from Pseudomonas stutzeri OX1. Appl
Environ Microbiol 64, 3626–3632.

17 Furuhashi K (1992) Biological routes to optically active
epoxides. In Chirality in Industry (Collins, A, Sheldrake,
G & Crosby, J, eds), pp. 167–186. John Wiley & Sons,
London.
18 Smith TJ & Dalton H (2004) Biocatalysis by methane
monooxygenase and its implications for the petroleum
industry. In Petroleum Biotechnology, Developments and
Perspectives (Vazquez-Duhalt, R & Quintero-Ramirez,
R, eds). Elsevier, Amsterdam.
19 Prior SD & Dalton H (1985) Acetylene as a suicide sub-
strate and active site probe for methane monooxygenase
from Methylococcus capsulatus (Bath). FEMS Microbiol
Lett 29, 105–109.
20 Hamamura N, Storfa RT, Semprini L & Arp DJ (1999)
Diversity in butane monooxygenases among butane-
grown bacteria. Appl Environ Microbiol 65, 4586–4593.
21 Yeager CM, Bottomley PJ, Arp DJ & Hyman MR
(1999) Inactivation of toluene 2-monooxygenase in Bur-
kholderia cepacia G4 by alkynes. Appl Environ Microbiol
65, 632–639.
22 Keener WK, Watwood ME, Schaller KD, Walton MR,
Partin JK, Smith WA & Clingenpeel SR (2001) Use of
selective inhibitors and chromogenic substrates to differ-
entiate bacteria based on toluene oxygenase activity.
J Microbiol Methods 46, 171–185.
23 Ensign SA, Hyman MR & Arp DJ (1992) Cometabolic
degradation of chlorinated alkenes by alkene monooxy-
genase in a propylene-grown Xanthobacter strain. Appl
Environ Microbiol 58, 3038–3046.
24 Foroozeh M, Primrose G, Guo Z, Bell LC, Alworth WL

& Guengerich FP (1997) Aryl acetylenes as mechanism-
based inhibitors of cytochrome P450-dependent mono-
oxygenase enzymes. Chem Res Toxicol 10, 91–102.
25 Jiang Y, Wilkins PC & Dalton H (1993) Activation of
the hydroxylase of sMMO from Methylococcus capsula-
tus (Bath) by hydrogen peroxide. Biochim Biophys Acta
1163, 105–112.
26 Andersson KK, Froland WA, Lee S-K & Lipscomb JD
(1991) Dioxygen independent oxygenation of
hydrocarbons by methane monooxygenase hydroxylase
component. New J Chem 15, 411–415.
27 Newman LM & Wackett LP (1995) Purification and
characterization of toluene 2-monooxygenase from Bur-
kholderia cepacia G4. Biochemistry 34, 14066–14076.
28 White RE & Coon MJ (1980) Oxygen activation by
cytochrome P-450. Annu Rev Biochem 49, 315–356.
29 Balendra S, Lesieur. C, Smith. TJ & Dalton H (2002)
Positively charged amino acids are essential for electron
transfer and protein–protein interactions in the soluble
methane monooxygenase complex from Methylococcus
capsulatus (Bath). Biochemistry 41, 2571–2579.
30 Froland WA, Andersson KK, Lee SK, Liu Y &
Lipscomb JD (1992) Methane monooxygenase compo-
nent B and the reductase alter the regioselectivity of the
hydroxylase component-catalysed reactions. J Biol Chem
267, 17588–17597.
31 Lee S-K, Nesheim JC & Lipscomb JD (1993) Transient
intermediates of the methane monooxygenase catalytic
cycle. J Biol Chem 268, 21569–21577.
32 Liu KE, Valentine AM, Qiu D, Edmondson DE, Apple-

man EH, Spiro TG & Lippard SJ (1995) Characterisa-
tion of a diiron (III) peroxo intermediate in the catalytic
cycle of methane monooxygenase hydroxylase from
Methylococcus capsulatus (Bath). J Am Chem Soc 117,
4997–4998.
33 Valentine AM, Stahl SS & Lippard SJ (1999) Mechanis-
tic studies of reduced methane monooxygenase hydroxy-
lase with dioxygen and substrates. J Am Chem Soc 121,
3876–3887.
34 Sazinski MH, Bard J, Di Donato A & Lippard SJ
(2004) Crystal structure of the toluene ⁄ o-xylene mono-
oxygenase hydroxylase from Pseudomonas stutzeri OX1.
J Biol Chem 279, 30600–30610.
35 Brazeau B, Wallar BJ & Lipscomb JD (2003) Effector
proteins from P450
cam
and methane monooxygenase:
lessons in tuning nature’s powerful reagents. Biochim
Biophys Res Commun 312, 143–148.
36 Dalton H & Whittenbury R (1976) The acetylene reduc-
tion technique as an assay for nitrogenase activity in the
methane oxidizing bacterium Methylococcus capsulatus
(Bath). Arch Microbiol 109, 147–151.
37 Smith TJ, Lloyd JS, Gallagher SC, Fosdike WLJ,
Murrell JC & Dalton H (1999) Heterologous expression
of alkene monooxygenase from Rhodococcus rhodo-
chrous B-276. Eur J Biochem 260, 446–452.
38 Simpson LB & Lovell FP (1962) Solubility of methyl,
ethyl, and vinyl acetylene in several solvents. J Chem
Eng Data 7, 498–500.

39 Cornish-Bowden A (1995) Fundamentals of Enzyme
Kinetics, 2nd edn. Portland Press, London.
W. L. J. Fosdike et al. Adventitious reactions of alkene monooxygenase
FEBS Journal 272 (2005) 2661–2669 ª 2005 FEBS 2669