Mechanistic insights from reaction of α-oxiranyl-aldehydes with cyanobacterial aldehyde deformylating oxygenase

Article abstract of DOI:10.1021/cb400772q

The biosynthesis of long-chain aliphatic hydrocarbons, which are derived from fatty acids, is widespread in Nature. The last step in this pathway involves the decarbonylation of fatty aldehydes to the corresponding alkanes or alkenes. In cyanobacteria, this is catalyzed by an aldehyde deformylating oxygenase. We have investigated the mechanism of this enzyme using substrates bearing an oxirane ring adjacent to the aldehyde carbon. The enzyme catalyzed the deformylation of these substrates to produce the corresponding oxiranes. Performing the reaction in D₂O allowed the facial selectivity of proton addition to be examined by ¹H NMR spectroscopy. The proton is delivered with equal probability to either face of the oxirane ring, indicating the formation of an oxiranyl radical intermediate that is free to rotate during the reaction. Unexpectedly, the enzyme also catalyzes a side reaction in which oxiranyl-aldehydes undergo tandem deformylation to furnish alkanes two carbons shorter. We present evidence that this involves the rearrangement of the intermediate oxiranyl radical formed in the first step, resulting in aldehyde that is further deformylated in a second step. These observations provide support for a radical mechanism for deformylation and, furthermore, allow the lifetime of the radical intermediate to be estimated based on prior measurements of rate constants for the rearrangement of oxiranyl radicals.

Full text of DOI:10.1021/cb400772q

Articles
pubs.acs.org/acschemicalbiology
Mechanistic Insights from Reaction of α‑Oxiranyl-Aldehydes with
Cyanobacterial Aldehyde Deformylating Oxygenase
Debasis Das,† Benjamin Ellington,‡ Bishwajit Paul,† and E. Neil G. Marsh†,‡,*
†Department of Chemistry, University of Michigan, Ann Arbor, Michigan 48109, United States
‡Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, United States
S
* Supporting Information
ABSTRACT: The biosynthesis of long-chain aliphatic hydrocarbons,
which are derived from fatty acids, is widespread in Nature. The last
step in this pathway involves the decarbonylation of fatty aldehydes to
the corresponding alkanes or alkenes. In cyanobacteria, this is catalyzed
by an aldehyde deformylating oxygenase. We have investigated the
mechanism of this enzyme using substrates bearing an oxirane ring
adjacent to the aldehyde carbon. The enzyme catalyzed the
deformylation of these substrates to produce the corresponding
oxiranes. Performing the reaction in D2O allowed the facial selectivity
1
of proton addition to be examined by H NMR spectroscopy. The
proton is delivered with equal probability to either face of the oxirane
ring, indicating the formation of an oxiranyl radical intermediate that is
free to rotate during the reaction. Unexpectedly, the enzyme also catalyzes a side reaction in which oxiranyl-aldehydes undergo
tandem deformylation to furnish alkanes two carbons shorter. We present evidence that this involves the rearrangement of the
intermediate oxiranyl radical formed in the first step, resulting in aldehyde that is further deformylated in a second step. These
observations provide support for a radical mechanism for deformylation and, furthermore, allow the lifetime of the radical
intermediate to be estimated based on prior measurements of rate constants for the rearrangement of oxiranyl radicals.
hydroxylase superfamily; in this case, the aldehyde carbon is
ong-chain aliphatic hydrocarbon waxes are synthesized by
La wide variety of organisms, including plants, insects, and released as CO.9,13,18 In cyanobacteria, the enzyme is somewhat
1
2
other animals3 and microbes.4,5 They serve important functions
surprising, a small, soluble protein that contains a “2-histidine,
such as water-proofing the feathers of waterfowl,3 preventing
4-carboxylate” non-heme di-iron cofactor similar to class 1
desiccation of plant leaves and stems,6 acting as contact
ribonucleotide reductase, methane monooxygenase, and
ferritin;4,19 in this enzyme the aldehyde carbon is converted
to formate.20,21 For all three types of decarbonylases, their
reactions represent highly unusual variations on the canonical
oxidation reactions catalyzed by other members of their
respective families, and their mechanisms remain poorly
understood.
pheromones in insects,7 and energy storage in green algae.8
These hydrocarbons are derived from various fatty acid
biosynthesis pathways that may involve elongation and
desaturation but conclude in two common steps that first
convert the fatty-acyl-CoA ester to the corresponding
aldehyde9−12 and then remove the aldehyde carbon to form
the final hydrocarbon product.13 This latter reaction is
catalyzed by a group of enzymes collectively known as aldehyde
decarbonylases.
The increasing interest in developing “next generation”
biofuels, those that can effectively function as “drop-in”
replacements for gasoline, diesel, and jet fuel, has spurred
renewed attention toward enzymes involved in hydrocarbon
biosynthesis.4,14 The mechanisms of the enzymes are also of
considerable interest because, as in the case of the decarbon-
ylases, they catalyze unusual and chemically difficult reactions.15
It has recently become apparent that there are at least three
mechanistically distinct types of aldehyde decarbonylases.16
The decarbonylase in insects has been shown to be a P450
enzyme, CYP4G1, and the aldehyde carbon is released as
CO2.2,17 In plants (and most likely in green algae), the enzyme
is an integral membrane protein belonging to the fatty acid
Our studies have focused on the cyanobacterial enzyme,
aldehyde deformylating oxygenase, cADO (also referred to as
cyanobacterial aldehyde decarbonylase in earlier reports20−24).
The enzyme has been shown to be iron-dependent,21,25 to
24,26,27
require O2
and an auxiliary reducing system for
activity,4,20,21 and to be inhibited by hydrogen peroxide.26
The aldehyde hydrogen is retained in formate, whereas the
proton in the alkane product derives from the solvent (Figure
1A).20,21 During the reaction, one atom of O2 is incorporated
into formate, requiring a mechanism in which O2 is completely
reduced to the oxidation state of water to accommodate the
stoichiometry of the reaction.24 In this respect, the reaction is
Received: October 8, 2013
Accepted: December 6, 2013
Published: December 6, 2013
© 2013 American Chemical Society
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Figure 1. (A) Deformylation reaction catalyzed by cADO. (B) Proposed mechanism of cADO involving homolytic cleavage of the C1−C2 bond of
aldehyde by di-iron peroxo species. (C) A recently proposed mechanism for deformylation involving heterolytic cleavage of the C1−C2 bond.
unlike that of other iron-dependent oxygenases that catalyze
the net oxidation of their substrates. A mechanism that
accommodates these observations is shown in Figure 1B.24
Stopped-flow UV-visible spectroscopy and rapid quench
Mossbauer spectroscopy have recently provided evidence to
support the formation of an FeIII/FeIII peroxide (or
peroxyhemiacetal) species in cADO.25 This species was
relatively stable, t1/2 ≈ 400 s at 5 °C, but once additional
electrons in the form of reduced O-methoxy-phenazine
methosulfate were added, it rapidly decayed, in accord with
the mechanism shown in Figure 1B. Evidence for a radical
mechanism for C−C bond scission comes from the reaction of
cADO with a β,γ-substituted cyclopropyl aldehyde designed to
function as a radical clock. Ring-opening of the cyclopropyl ring
was observed, consistent with homolytic cleavage of the formyl
group.23 We also note that recent experimental observations of
oxidative products arising from the reaction of cADO with
medium-chain aldehydes have led to a mechanistic proposal
involving heterolytic C−C bond scission (Figure 1C).28 We
discuss this proposal in more detail later.
modification is therefore limited by steric constraints. However,
examination of the structure suggested that aldehydes
containing three-membered rings adjacent to the aldehyde
carbon could be accommodated with minimal perturbation of
the structure. We therefore synthesized as analogues of
dodecanal and octadecanal, trans-3-nonyloxirane-2-carbalde-
hyde, 1, and trans-3-pentadecanyloxirane-2-carbaldehyde, 2,
which bear an oxirane ring adjacent to the aldehyde carbon
(Scheme 1). The compounds were synthesized using standard
Scheme 1. Structures of trans-3-Nonyloxirane-2-
carbaldehyde, 1, and trans-3-Pentadecanyloxirane-2-
carbaldehyde, 2, Used in These Studies
literature procedures, as described in the Methods section. We
reasoned that introducing the oxiranyl functionality at the site
of C−C bond scission should provide insights into the
mechanism of deformylation by altering the stability of
intermediates. Furthermore, the introduction of the three-
membered oxirane ring adjacent to the aldehyde carbon would
allow the stereochemistry of proton transfer to be investigated.
Reaction of 1 and 2 with cADO. Initially we examined the
activity of 1 under both air-saturated and microaerobic
conditions and compared its activity with that of dodecanal.
Typical assays contained 10 μM cADO, 300 μM substrate, and
an auxiliary reducing system comprising NADH, 2 mM, as the
reductant and PMS, 100 μM, as the electron mediator, as
described previously.23 Under fully aerobic conditions, very
little activity was observed with either 1 or dodecanal, most
likely because the nonenzymatic reaction of O2 with reduced
Alternate substrates that incorporate functionality that can
perturb the stability of putative reaction intermediates or
facilitate stereochemical analysis of the reaction have proved to
be useful tools to diagnose enzyme mechanisms. In this report,
we have investigated the reaction of cADO with aldehydes
bearing an oxirane ring adjacent to the aldehyde carbon, which
has allowed us to probe the mechanism of the key carbon−
carbon bond-breaking step in the reaction.
RESULTS AND DISCUSSION
■
Choice of Substrate Analogues. The substrate-binding
site of cADO comprises a narrow hydrophobic channel that
terminates at the metal center.19 The scope for substrate
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PMS depleted the reducing system before significant turnover
could occur. All subsequent experiments were therefore
performed microaerobically, as described previously. Under
microaerobic conditions, cADO catalyzed the conversion 1 to
2-nonyloxirane and dodecanal to undecane at approximately
similar rates (apparent kcat = 0.016 0.001 and 0.01 0.001
min−1, respectively). In both cases, the reaction was linear for
several hours during which time about three turnovers occurred
(Figure 2). Formate was formed as the coproduct, as expected.
recent study examined the relative rates of reaction of aliphatic
aldehydes with chain lengths ranging from 18 to 4 carbons.29
Interestingly, it was found that the enzyme is more active with
either long-chain (C18−C14) or short-chain (C9−C5)
aldehydes whereas medium chain aldehydes, including
dodecanal, were turned over considerably more slowly.
Therefore, we were curious whether faster rates of turnover
could be obtained by increasing the chain length of the alkyl-
oxiranyl-carbaldehyde. We examined the activity of 2 with
cADO and compared it with the “fast” substrate octadecanal.
We found that 2 was converted to 2-pentadecanyloxirane and
formate with apparent kcat = 0.029 0.001 min−1, that is, about
twice as fast as 1 was converted to 2-nonyloxirane. However,
this is ∼4-fold slower than the turnover of octadecanal for
which apparent kcat = 0.11 0.01 min−1.22,29
We were curious whether the slow turnover of 1 and 2 might
be due to these compounds inactivating the enzyme, as we
previously observed mechanism-based inactivation, resulting in
covalent modification of the enzyme, in the reaction of cADO
with a β,γ-substituted cyclopropyl aldehyde designed to
function as a radical clock.23 However, LC-ESI-MS analysis of
the enzyme after reaction with the oxiranyl aldehydes
established that neither 1 nor 2 covalently modified the
enzyme (Figure S8, Supporting Information). Moreover, the
time course of the reaction does not show evidence for time-
dependent inactivation of the enzyme. The slow reactions of 1
and 2 with cADO therefore appear to be intrinsic to their
chemical functionality.
■
Figure 2. Comparison of the rates of deformylation of 1 ( ) and
dodecanal ( ) by cADO.
●
The sluggish nature of the cADO reaction has been noted
previously in investigations by various laboratories.20,21,26
A
1
Figure 3. Facial selectivity of proton addition to 2-nonyloxirane. H NMR spectra of the oxirane ring protons Ha, Hb, and Hc are shown. (A) An
authentic standard of racemic 2-nonyloxirane (for clarity, the structure of the (R)-enantiomer is drawn); (B) products of the reaction of 1 with
cADO in H2O; (C) products of the reaction of 1 with cADO in D2O. In each case integrations are relative to Ha. Peak identified by ∗ in spectrum C
is a contaminant that contributes slightly to the integration of Hc.
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Facial Selectivity of Proton Transfer. Previous studies
have established that the proton in the product alkane derives
from the solvent in the cADO-catalyzed reaction;20,21 this is in
contrast to the decarbonylation reactions catalyzed by the
insect and plant enzymes in which the aldehyde hydrogen is
transferred to the alkane.13,30 However, the stereochemistry of
this step has not been investigated for any of these enzymes.
We took advantage of the oxirane ring generated by the
reaction of 1 with cADO to examine the facial selectivity of
proton transfer. Reactions were set up containing 40 μM
cADO, 400 μM 1, 2 mM NADH, and 100 μM PMS in 10 mM
potassium phosphate buffer, pH/pD 7.2, in either H2O or D2O.
After 2 h incubation at 37 °C, the products of the reaction,
together with unreacted substrate, were extracted with CDCl3
1
and dried, and their H NMR spectra were recorded.
The oxirane protons (Figure 3A) are clearly separated from
other resonances and comprise a broad multiplet due to Ha, δ =
2.89 ppm (J1 = 3.22, J2 = 5.83 Hz), an overlapping doublet-of-
doublets due to Hb, δ = 2.73 ppm (J1 = 3.90, J2 = 5.08 Hz), and
a doublet-of-doublets due to Hc, δ = 2.45 ppm (J1 = 2.75 Hz, J2
= 5.07 Hz). For the reaction performed in H2O, integration of
Ha, Hb, and Hc reveals, as expected, equal intensities for all
three protons (Figure 3B). For the reaction performed in D2O,
however, both Hb and Hc are almost equally reduced in
intensity to 0.67 and 0.74, respectively, relative to Ha (the
slightly higher integration for Hc reflects the presence of an
overlapping contaminant peak, Figure 3C). This indicates that
the deuteron can be transferred with equal probability to either
face of the oxirane ring. It is evident that the intensities of Hb
and Hc are not reduced to the theoretical value of 0.50. We
attribute this to residual protons in the D2O buffer, which are
estimated to comprise ∼5% of the solvent. Proton incorpo-
ration is most likely enhanced by a solvent kinetic isotope
effect.
Figure 4. Formation of (n − 2)-alkanes from 1 and 2 by cADO. (A)
GC-MS chromatograph of the products of reaction of 1 with cADO.
(B) GC-MS traces of the products of reaction of 2 with cADO; in this
case small amounts of enzymatically derived heptadecanal are resolved
in the chromatograph. Inset, comparison of the rates of formation of 2-
pentadecyloxirane ( ) and hexadecane ( ) from 2. Peaks identified
by ∗ and ∗∗ are contaminants.
■
●
This observation provides evidence for the existence of an
intermediate species, most likely the 3-nonyloxiran-2-yl radical
that is able to undergo rapid rotation about the C−C bond to
the alkyl group before delivery of the solvent-derived proton.
Oxiranyl radicals are known to be pyramidal at the carbon
center,31 indicating little or no delocalization of the radical onto
the oxygen, and undergo rapid interconversion between cis and
trans forms. For unsubstituted oxiranyl radicals, the rate of
interconversion is especially fast, ∼107 s−1 at −110 °C.32 Thus,
the observed stereochemical scrambling of deuterium indicates
that interconversion between cis and trans radicals occurs much
faster than proton delivery to form the product.
Evidence for Rearrangement of Oxiranyl Radical
Intermediates. During the course of our investigations, we
consistently noticed small amounts of decane (Figure 4A) and
hexadecane (Figure 4B) in the products of the reaction of 1 and
2, respectively, with cADO. Further investigations established
that the appearance of these products was linearly dependent
on enzyme concentration and that they were formed in direct
proportion to the major 2-alkyloxirane products (Figure 4B,
inset). Furthermore, the appearance of these (n − 2)-alkanes
was dependent on the presence of all the components in the
assay, including the substrates (Figure S9 and S10, Supporting
Information). These observations suggested that they were
derived from reaction of the oxiranyl-aldehydes with the
enzyme.
ment of 2-nonyloxirane and 2-pentadecanyloxirane to un-
decanal and heptadecanal respectively, which would then
undergo deformylation to decane and hexadecane. However,
no alkanes were formed when either 2-nonyloxirane or 2-
pentadecanyloxirane were incubated with the diferric enzyme
alone. Neither was rearrangement of these compounds
observed when they were incubated with the enzyme with
the other components of the assay for prolonged periods.
These observations suggest that the (n − 2)-alkanes most
likely arise through partitioning of an intermediate formed in
the deformylation of 1 and 2 by cADO between two reaction
pathways. To explain the formation of the (n − 2)-alkanes, we
considered a variant of the deformylation mechanism in which
after homolytic cleavage of the Cα−CO bond to form the 3-
alkyloxiran-2-yl radical and formate, ring-opening of this radical
occurs to generate the Cα radical of the (n−1)-aldehyde.
Quenching of this radical would thus generate the (n − 1)-
aldehyde that could subsequently undergo deformylation
(Figure 5). Ring-opening reactions of alkyloxiranyl radicals
are well documented in the literature.32,34
This mechanism predicts that (n − 1)-aldehydes should be
formed as intermediates. Close examination of the gas
chromatograph for the reaction of 2 with cADO revealed the
presence of a minor peak at 10.46 min that eluted just before 2-
pentadecanyloxirane (Figure 4B). The intensity of the peak
increased with time during the course of the reaction and was
dependent upon all the components of the assay being present.
The mass spectrum of the compound matched that of
Oxirane rings can be rearranged to carbonyl compounds by
Lewis acid catalysts.33 We therefore considered the possibility
that the diferric form of cADO might catalyze the rearrange-
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Figure 5. Mechanism for the conversion of oxiranyl aldehydes to Cn−1
oxiranes and Cn−2 alkanes involving a branched pathway that arises
through the slow rearrangement of an oxiranyl radical intermediate.
Figure 6. GC-MS analysis of hexadecane (m/z = 226.2) produced
from reaction of 2 with cADO (A) in H2O buffer and (B) in D2O
buffer. The molecular ion for hexadecane produced in D2O buffer is
shifted by 2 mass units to m/z = 228.2.
heptadecanal, and the peak coeluted with an authentic standard
of heptadecanal (Figure S11, Supporting Information). It was
similarly possible to detect the formation of undecanal in the
reaction products formed through the reaction with cADO with
1, although in this case it was necessary to modify the
chromatography conditions to separate undecanal from 2-
nonyloxirane (Figure S12, Supporting Information).
A further prediction of the mechanism is that the alkanes
derived from 1 or 2 should incorporate two protons from the
solvent during the course of the reaction. To evaluate this
prediction, the reaction of 2 in D2O was investigated. cADO
was reacted with 2 for 2 h under the usual conditions in assay
buffer in which the D2O content was ∼97%. The products of
the reaction were extracted and analyzed by GC-MS. The
molecular ion for hexadecane was clearly visible and shifted by
2 mass units to m/z = 228.2 from the expected value of m/z =
226.2 for unlabeled material (Figure 6). A smaller peak at m/z
= 227.2 corresponding to monodeuterated heptadecane was
also present, which may be explained by incomplete
deuteration of the solvent combined with a solvent isotope
effect.
rates that allow them to be used as slow radical clocks. The rate
constant for rearrangement of the 3-methyloxiran-3-yl radical to
the corresponding acetonyl radical has been measured as 3.1 ×
104 s−1 at 25 °C.35 The rate constant for the rearrangement of
the 3-methyloxiran-2-yl radical to the corresponding propanal-
derived radical, which serves as a better reference for the
reaction of oxiranyl aldehydes with cADO, has not been
measured directly but is estimated to occur about an order of
magnitude more slowly.32 From the observed partitioning
ratios between alkyloxirane and (n − 2)-alkane, the products
derived from the reaction of 1 and 2 with cADO, we estimate
that the rate constant for this proton + electron transfer step
(there are no residues in the active site that might serve as
hydrogen atom donors, for example, cysteine or tyrosine, but
whether this is formally a proton-coupled electron transfer is
currently unclear) is ∼104 s−1 at 25 °C and is unlikely to be
faster than 105 s−1, using the rate constant for rearrangement of
the 3-methyloxiran-3-yl radical as an upper limit.
The relatively fast rate constant for this step is indicative of
electron transfer directly from a site on the protein, rather than
directly from the external reducing system. The most likely
source of the electron is the di-iron metal center. This step
could conceivably involve either transient oxidation of the
diferric center to generate a mixed valent FeIII−FeIV species
followed by reduction back to FeIII or initial reduction of the
diferric center to generate a mixed valent FeIII−FeII species,
followed by electron transfer to regenerate the diferric enzyme.
Recently, an interesting observation has been made that
cADO catalyzes the oxidation of alkanes derived from
deformylation of C9 and C10 aldehydes to the corresponding
(n − 1)-alcohols and -aldehydes, albeit extremely slowly.28
However, enzyme was unable to oxidize alkanes or alcohols in
the absence of aldehydes. To accommodate these findings, a
mechanism was proposed in which the deformylation step
occurred by a heterolytic mechanism to generate a reactive
FeIV-oxo species, akin to that generated in P450 reactions, that
was responsible for the subsequent oxidative chemistry. We
note that oxiranes can be rearranged to aldehydes by strong
hindered bases,36,37 so in principle, a mechanism involving
Mechanistic Implications. To investigate the nature of
C−C bond cleavage step following initial formation of the
metal peroxide species, we previously examined the reaction of
cADO with an aldehyde bearing a strategically placed
cyclopropyl group that could act as a radical clock.23 This
substrate partitioned between two pathways when reacted with
cADO. In the productive pathway, ring-opening of the
cyclopropyl group occurred to produce 1-octadecene, providing
support for a radical mechanism for C−C bond cleavage. In a
nonproductive reaction, alkylation of the protein occurred after
deformylation, resulting in inactivation of the enzyme.
Compounds that generate cyclopropylcarbinyl radicals can
be used to measure the lifetimes of radical intermediates when
they are of similar duration to the well-characterized ring-
opening reactions so that product partitioning ratios can be
measured. In this case because only the ring-opened product
was observed, all that could be inferred was that the lifetime of
the intermediate radical species was in excess of 10 ns.23
However, oxiranyl radicals, although not as extensively studied,
also undergo ring-opening rearrangements but at much slower
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7.2, containing 100 mM KCl and 10% glycerol under microaerobic
conditions, employing phenazine methosulfate (PMS) and NADH as
the auxiliary reducing system as described previously.23 Aldehyde
substrates were made up as a 10 mM stock solution in DMSO. A
typical assay contained 10 μM cADO, 20 μM ferrous ammonium
sulfate, 300 μM aldehyde substrate, 100 μM PMS, and 2 mM NADH
in a total volume of 500 μL. Assays were shaken at 37 °C at 200 rpm.
Reactions were quenched by addition of 500 μL of ethyl acetate and
vigorous vortexing, followed by centrifugation at 14000 rpm for 30
min to separate the organic phase. The ethyl acetate layer was
collected, and 8 μL of sample was subjected to GC-MS analysis as
described previously.
heterolytic C−C bond cleavage and the formation of an
oxiranyl carbanion could be operating. However, we consider
that this is less likely for the following reasons.
The heterolytic mechanism predicts that it should be possible
to form up to 1 equiv of product under the assay conditions
resulting from the initial reaction of the apoenzyme, Fe(II), and
O2 with aldehyde. However, we found no evidence for any
turnover unless PMS and NADH were included.21,23 The
reaction of cADO with a cyclopropyl aldehyde radical clock
substrate, discussed above, provides strong support for a radical
mechanism. It was suggested that ring-opening could be a
secondary reaction arising from reaction of the cyclo-
propylalkane product with the FeIV-oxo species to generate a
cyclopropylcarbinyl radical in a P450-like manner.28 But the
products from such a reaction should bear a hydroxyl group; in
practice the product was found to be (unoxidized) octadecene.
Lastly, spectroscopic evidence has recently been published
supporting the formation of a diferric peroxide (peroxyhemia-
cetal) intermediate in the reaction of cADO, as mentioned
above.25 This species was stable until the addition of the
reductant necessary to initiate homolytic bond cleavage; in
contrast a heterolytic mechanism would imply that it could
undergo spontaneous conversion to the putative FeIV-oxo
species.
In conclusion, our studies demonstrate that oxiranyl-
aldehydes are substrates for cADO that undergo turnover at
rates that are comparable to the corresponding unfunctional-
ized aldehydes. The unexpected observation that these oxiranyl-
aldehydes are processed by cADO to generate both (n − 1)-
oxiranes and (n − 2)-alkanes provides support for the
formation of a relatively long-lived radical in the reaction.
This is supported by stereochemical analysis demonstrating that
the proton is delivered with equal probability to either face of
the oxirane ring, indicative of an intermediate that is free to
rotate during the reaction. From this work and the results of
other studies, it is becoming clear that cADO catalyzes a wide
range of reactions that are initiated by activated oxygen species.
Further studies are needed to both clarify the physiological role
of the enzyme and better understand the mechanisms of the
unusual and chemically challenging reactions catalyzed by this
enzyme.
Formate Determination. Formate was determined to be the
coproduct of reaction of 1 and 2 with cADO by reaction with 2-
nitrophenylhydrazine, as described previously.21
Deuterium Incorporation Experiments. To investigate deute-
rium incorporation into alkane products, assays were performed in 100
mM HEPES buffer containing 100 mM KCl in 99.9% D2O, pD 7.2.
Substrates were made up as 10 mM stock solutions in 99.9% DMSO-
d6. cADO was added as a concentrated stock solution in
nondeuterated buffer. The final H2O concentration in the reaction
mixture did not exceed 3%. The enzyme was incubated in the buffer
for 1 h prior to initiating the reaction by addition of substrate. Assays
were shaken at 37 °C for 2 h at 200 rpm. Products were extracted and
analyzed as described above.
Preparation of Samples for NMR. Assays were performed as
described above except that phosphate buffer was substituted for
HEPES buffer, which otherwise interfered with the NMR spectra.
Assays were carried out either in 10 mM potassium phosphate, pH 7.2,
containing 50 mM KCl in H2O or in 10 mM potassium phosphate, pD
7.2, containing 50 mM KCl in D2O (99.9%). Aldehyde solutions were
made up as a stock solution in DMSO or DMSO-d6 for the respective
experiments. A typical assay contained 40 μM Np cADO, 80 μM
ferrous ammonium sulfate, 100 μM PMS, 2 mM NADH, and 400 μM
substrate in a total volume of 500 μL. For assays performed in
deuterated buffer, the final H2O concentration was ∼5% after adding
all the assay components. Ten identical 500 μL reactions were set up
in each buffer and shaken at 37 °C at 200 rpm for 2 h. The reaction
mixtures were sequentially extracted with a total volume of 1 mL of
CDCl3 (99.9%). The CDCl3 layers were washed with D2O, dried over
1
sodium sulfate, and filtered before analysis by H NMR.
ASSOCIATED CONTENT
■
S
* Supporting Information
Procedures for the synthesis and characterization of the various
oxiranyl compounds described in the manuscript and GC-MS
chromatographs of enzymatic reaction products and authentic
standards referred to in the main text. This material is available
free of charge via the Internet at http://pubs.acs.org.
METHODS
■
Materials. Phenazine methosulfate (PMS) and ferrous ammonium
sulfate were from Sigma-Aldrich. NADH, 2-nitrophenylhydrazine, and
1-ethyl-3-(3-dimethylaminopropyl) carbodiimide were obtained from
Acros Organics. D2O (99.9%) and DMSO-d6 (99.9%) were from
Cambridge Isotope Laboratories, Inc. All other reagents were of the
purest grade commercially available and used without further
purification.
AUTHOR INFORMATION
■
Corresponding Author
Synthesis of Oxirane Aldehydes. The synthesis of trans-3-
nonyloxirane-2-carbaldehyde, 1, was accomplished by standard
methods starting from commercially available (E)-dodec-2-en-1-ol.
trans-3-Pentadecanyloxirane-2-carbaldehyde, 2 (Figure 2), was synthe-
sized by standard methods utilizing the Horner−Wittig reaction of
hexadecanal with ethyl-2-(diethyloxyphosphoryl) acetate to obtain the
corresponding α,β-unsaturated carboxylic acid ethyl ester that was
subsequently elaborated to 1 and 2.23,38 Authentic standards of
nonyloxirane and pentadecanyloxirane were synthesized by epox-
idation of 1-undecene and 1-pentadecene using metachloroperbenzoic
acid.38 Full details of the synthetic procedures are included as
Supporting Information.
*E-mail: nmarsh@umich.edu.
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
We thank M. Waugh and F. Lin for helpful discussions in the
preparation of this manuscript. We thank N. Theddu for
asistance with synthesis of substrate analogues. This research
was supported in part by National Science Foundation, Grants
CHE 1152055 and CBET 1336636, the National Institutes of
Health, Grant GM 093088, and the European Union, Grant
FP-7 256808.
Enzyme Assay. The purification of recombinant Nostoc
punctiforme cADO from Escherichia coli was performed as described
previously.23 Assays were performed in 100 mM HEPES buffer, pH
575
dx.doi.org/10.1021/cb400772q | ACS Chem. Biol. 2014, 9, 570−577

ACS Chemical Biology
Articles
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