Cyanobacterial aldehyde deformylase oxygenation of aldehydes yields n - 1 aldehydes and alcohols in addition to alkanes

Article abstract of DOI:10.1021/cs400484m

Aldehyde-deformylating oxygenase (ADO) catalyzes O₂-dependent release of the terminal carbon of a biological substrate, octadecanal, to yield formate and heptadecane in a reaction that requires external reducing equivalents. We show here that ADO also catalyzes incorporation of an oxygen atom from O₂ into the alkane product to yield alcohol and aldehyde products. Oxygenation of the alkane product is much more pronounced with C _9-10 aldehyde substrates so that use of nonanal as the substrate yields similar amounts of octane, octanal, and octanol products. When using doubly labeled [1,2-¹³C]octanal as the substrate, the heptane, heptanal, and heptanol products each contained a single ¹³C-label in the C-1 carbons atoms. The only one-carbon product identified was formate. [¹⁸O]O₂ incorporation studies demonstrated formation of [¹⁸O]alcohol product, but rapid solvent exchange prevented similar determination for the aldehyde product. Addition of [1-¹³C]nonanol with decanal as the substrate at the outset of the reaction resulted in formation of [1-¹³C]nonanal. No ¹³C-product was formed in the absence of decanal. ADO contains an oxygen-bridged dinuclear iron cluster. The observation of alcohol and aldehyde products derived from the initially formed alkane product suggests a reactive species similar to that formed by methane monooxygenase (MMO) and other members of the bacterial multicomponent monooxygenase family. Accordingly, characterization by EPR and Moessbauer spectroscopies shows that the electronic structure of the ADO cluster is similar, but not identical, to that of the MMO hydroxylase component. In particular, the two irons of ADO reside in nearly identical environments in both the oxidized and fully reduced states, whereas those of MMOH show distinct differences. These favorable characteristics of the iron sites allow, for the first time for any biological system, a comprehensive determination of the spin Hamiltonian parameters describing the electronic state of the diferrous cluster. The nature of the diiron cluster and the newly recognized products from ADO catalysis hold implications for the mechanism of C-C bond cleavage.

Full text of DOI:10.1021/cs400484m

Research Article
pubs.acs.org/acscatalysis
Cyanobacterial Aldehyde Deformylase Oxygenation of Aldehydes
Yields n − 1 Aldehydes and Alcohols in Addition to Alkanes
Kelly G. Aukema,† Thomas M. Makris,‡,⊥ Sebastian A. Stoian,§,∥ Jack E. Richman,† Eckard Munck,§
̈
John D. Lipscomb,‡ and Lawrence P. Wackett*,†,‡
†BioTechnology Institute University of Minnesota, St. Paul, Minnesota 55108, United States
‡Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota, Minneapolis, Minnesota 55455, United
States
§Department of Chemistry, Carnegie Mellon University, Pittsburgh, Pennsylvania 15213, United States
S
* Supporting Information
ABSTRACT: Aldehyde-deformylating oxygenase (ADO) catalyzes O2-depend-
ent release of the terminal carbon of a biological substrate, octadecanal, to yield
formate and heptadecane in a reaction that requires external reducing equivalents.
We show here that ADO also catalyzes incorporation of an oxygen atom from O2
into the alkane product to yield alcohol and aldehyde products. Oxygenation of the alkane product is much more pronounced
with C9−10 aldehyde substrates so that use of nonanal as the substrate yields similar amounts of octane, octanal, and octanol
products. When using doubly labeled [1,2-13C]octanal as the substrate, the heptane, heptanal, and heptanol products each
contained a single 13C-label in the C-1 carbons atoms. The only one-carbon product identified was formate. [18O]O2
incorporation studies demonstrated formation of [18O]alcohol product, but rapid solvent exchange prevented similar
determination for the aldehyde product. Addition of [1-13C]nonanol with decanal as the substrate at the outset of the reaction
resulted in formation of [1-13C]nonanal. No 13C-product was formed in the absence of decanal. ADO contains an oxygen-
bridged dinuclear iron cluster. The observation of alcohol and aldehyde products derived from the initially formed alkane product
suggests a reactive species similar to that formed by methane monooxygenase (MMO) and other members of the bacterial
multicomponent monooxygenase family. Accordingly, characterization by EPR and Mossbauer spectroscopies shows that the
̈
electronic structure of the ADO cluster is similar, but not identical, to that of the MMO hydroxylase component. In particular,
the two irons of ADO reside in nearly identical environments in both the oxidized and fully reduced states, whereas those of
MMOH show distinct differences. These favorable characteristics of the iron sites allow, for the first time for any biological
system, a comprehensive determination of the spin Hamiltonian parameters describing the electronic state of the diferrous
cluster. The nature of the diiron cluster and the newly recognized products from ADO catalysis hold implications for the
mechanism of C−C bond cleavage.
KEYWORDS: biofuel, nonheme diiron enzyme, oxygenase, 13C NMR, GC/MS, Prochlorococcus marinus, EPR, Mossbauer
̈
second oxygen atom derived from atmospheric dioxygen.13,14
1. INTRODUCTION
In light of this, the hydrocarbon-generating enzyme has been
Bacterial synthesis of diesel-length alkanes and alkenes expands
given the name aldehyde-deformylating oxygenase (ADO).15
the biofuel inventory beyond alcohols and esters1−4 by making
a source of renewable, drop-in, liquid fuels.5−10 Recently, a two-
The amino acid sequence of ADO reveals similarity to a
number of enzymes that utilize a dinuclear iron cluster to
gene cluster required for alkane/alkene biosynthesis was
identified in cyanobacteria.11 The hydrocarbons formed had
activate O2 for oxygenation or oxidation reactions including
ribonucleotide reductase (RNR-R2), methane monooxygenase
an odd chain length (C13, C15, and C17), indicating they were
(MMO), a tRNA-modifying enzyme, MiaE, Δ9-fatty acid
biosynthesized by removal of the C-1 carbon of even-chain-
length fatty acyl groups. In this biosynthetic pathway, the fatty
desaturase and toluene monooxygenases.16−25 In addition, the
X-ray crystal structure of ADO shows that it has a dimetal
acyl group is reduced by an acyl-ACP reductase, a member of
center with coordination similar to that found for these
the short-chain dehydrogenase family, to produce the
enzymes.11 Each of these enzymes employs an oxygen-
corresponding aldehyde. The second enzyme in the pathway,
activation strategy whereby the reduced enzyme binds dioxygen
and subsequently carries out an oxidation reaction. In this
context, it is interesting that the ADO reaction is redox-neutral
a dinuclear iron-cluster-containing enzyme, removes the
aldehydic carbon to produce a hydrocarbon. Initially, the
displaced single-carbon product was proposed to be carbon
monoxide, on the basis of previous reports on similar reactions
observed in whole cells and cell extracts of plants and
animals.11,12 More recently, formate was shown to be the
coproduct of the hydrocarbon-generating enzyme, with the
Received: June 27, 2013
Revised: August 15, 2013
Published: August 16, 2013
© 2013 American Chemical Society
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overall. In fact, this led to a proposal that ADO does not require
dioxygen and that the oxygen atom in the formate product
derives from water.26 However, a reexamination of the data
showed that an oxygenolytic mechanism is operative.15,27
The mechanism of O2 activation and product formation of
ADO remains unknown. Mechanisms involving FeIIIFeIII-
superoxo,28 FeIIIFeIII-μ-peroxo,29,30 and high-valence FeIIIFeIV-
μ-oxo31 and FeIVFeIV-bis-μ-oxo32−34 species have been
proposed for other oxygen-activating enzymes in the dinuclear
iron cluster family. These species are known to catalyze
numerous types of reactions, such as hydrocarbon oxygenation,
alcohol and aldehyde oxidation, epoxidation, and desaturation,
as well as radical formation.32,34
Here, we have characterized the metal cluster of ADO for
comparison with those found in other oxygen-activating diiron-
cluster-containing enzymes. A study of the products formed
during reaction of ADO with both a biological substrate
octadecanal and shorter hydrocarbon aldehydes was conducted.
Surprisingly, it is found that, in addition to the well
documented formate and (n − 1) alkane products, ADO
catalyzes formation of products in which oxygen from O2 is
incorporated to form (n − 1) alcohols and aldehydes. These
newly recognized products give insight into the nature of the
ADO reaction and the reactive species generated during
catalysis.
a helium gas flow of 1.75 mL/min, and 250 °C injection port.
The sample was split at the column outlet between a flame
ionization detector, HP 7890A (Hewlett-Packard, Palo Alto,
CA), and a mass spectrometer, HP 5975C (GC/MS/FID). For
octanal, nonanal, decanal, and undecanal, the oven was held at
45 °C for 5 min, ramped to 60 °C at 5°/min, then ramped to
320 °C at 30°/min. For dodecanal, the oven temperature was
held at 60 °C for 5 min, ramped to 150 °C at 10°/min, then
ramped to 320 °C at 20°/min. For octadecanal, the oven was
ramped from 100 to 320 °C at 10°/min. All programs included
a 5 min hold at 320 °C. Concentrations of products were
determined using the equation of the best fit line for the plot of
MS peak area for aldehyde, alkane, or alcohol, ranging from 5
to 250 μM. Peaks were identified by comparison of retention
times, and mass spectra, to authentic standards.
2.4. ADO Reaction in [18O]-O2. After reaction mixtures
were degassed with alternating argon and vacuum on a Schlenk
line to remove 16O2, they were exposed to [18O]O2 under slight
positive pressure. The reactions were started with degassed
NADH and allowed to proceed for 10 min. Degassed alcohol
dehydrogenase (6 μg/mL) was added, and the reactions were
incubated for an additional 10 min before extraction,
bis(trimethylsilyl)acetamide derivatization (1% final in organic
layer), and GC/MS/FID analysis, as described above. The
abundance of the oxygen-containing m+/z ions at 187−191 was
determined.
2.5. [1-13C]-1-Nonanol Addition to Decanal ADO
Reaction. [1-13C]-1-Nonanol was synthesized from [1-13C]-
nonanoic acid with boranetetrahydrofuran using the methods of
Kende and Fludzinski.35 1-Nonanol or [1-13C]-1-nonanol was
added to a standard ADO reaction to 0.5 mM or 1.5 mM with
200 μM decanal and PMS/NADH. Percent 13C incorporation
was calculated as percent (m+ + 1)/z signal of the total signal
from a fragmentation ion pair (m+/z + (m+ + 1)/z). The
reported values are an average of three measurements, each for
the three major electron impact fragment ions that retain the
labeled carbon.36
2. EXPERIMENTAL SECTION
2.1. Chemicals and Reagents. All reagents, solvents,
standards, and enzymes were purchased from Sigma-Aldrich
with the following exceptions: heptadecanal and octadecanal
(TCI America, Portland, OR), nonanal (ThermoFisher
Scientific, Waltham, MA), nicotinamide adenine dinucleotide
(NADH) and nicotinamide adenine dinucleotide phosphate
(NADPH) (EMD Millipore, Billerica, MA), and [13C]-
formaldehyde (Cambridge Isotope Laboratories, Andover,
MA). Methods for syntheses of 13C-labeled aldehydes are
presented in the Supporting Information (SI).
2.2. Aldehyde-Deformylating Oxygenase Purification.
Prochlorococcus marinus ADO (NP_895059) was expressed in
BL21(DE3) with an N-terminal histidine tag from an
Escherichia coli-codon-optimized gene (DNA2.0, Menlo Park,
CA) in pET28b. To increase iron content of ADO, cells were
grown in terrific broth and induced at 25 °C with 0.2 mM
IPTG (University of Minnesota Biotechnology Resource
Center production batch WAC110911). ADO was purified in
25 mM HEPES, pH 7.5, by ion exchange chromatography on a
DEAE column using a linear gradient of 0−400 mM NaCl.
Purified protein was desalted by dialysis against 25 mM
HEPES, pH 7.5. The percent iron incorporation varied with
each purification from 40% to 80%.
2.3. ADO Reactions and GC/MS/FID Analysis. Reactions
were typically carried out in 300 μL reaction volumes in 2 mL
GC vials at room temperature for 30 min unless otherwise
noted. Reactants included 30 μM ADO, 500 μM aldehyde
substrate in ethyl acetate (1% v/v final), 100 mM HEPES pH
7.4, 75 μM phenazine methyosulfate (PMS), and 2 mM
NADH. When noted, spinach ferredoxin (100 μg/mL),
ferredoxin reductase (0.1 U/mL), and NADPH (2 mM) were
substituted for PMS and NADH. Dodecane standard was
added to 45 μM, and reactions were extracted with 300 μL of
methyl tert-butyl ether (MTBE). The organic layer was assayed
using a gas chromatograph (GC) with an HP-1 ms column
(100% dimethylsiloxane capillary; 30 m × 250 μm × 0.25 μm),
2.6. 13C NMR. Using a 7001 Megatron 700 MHz Bruker
NMR, spectra were collected using standard parameters for
protein-decoupled 13C NMR data with 24 000 transients. The
ADO reaction consisted of 3 mM [1-13C]nonanal, 3 mM ADO,
50 mM sodium phosphate pH 7.4, 0.1 mg/mL spinach
ferredoxin, 0.1 U/mL ferredoxin reductase, 2 mM NADPH, 1%
dimethyl sulfoxide (DMSO), and 10% D2O.
̈
2.7. Mossbauer and EPR Methods. Methods for
Mossbauer and EPR spectroscopies are presented in the SI.
̈
3. RESULTS
3.1. Novel Products of ADO Reaction Consistent with
an Oxygenase Reaction. ADO reaction mixtures containing
octadecanal and a reducing system (NADH/phenazine) were
extracted and analyzed by liquid GC/MS. Heptadecane, an
alkane one carbon shorter than the substrate, was observed, as
previously reported.11,13,37 In addition to heptadecane, we also
observed heptadecanal that accounted for less than 3% of the
heptadecane yield. Heptadecane and heptadecanal peaks on the
GC/MS chromatogram were validated by comparison of the
retention time and mass spectral fragmentation pattern with
those of authentic standards. Incomplete control reactions
without enzyme or without NADH lacked both this new
aldehyde product and the previously observed alkane product.
The production of heptadecanal indicated that ADO enzyme
was capable of incorporating O2 into a methylene carbon, and
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thus, the reaction mixture was also tested for the presence of an
alcohol product (heptadecanol). The high boiling point of the
long-chain heptadecanol complicates GC analysis, so reactions
with shorter-chain aldehyde substrates were examined. Rather
than sampling the reaction headspace, which would favor the
detection of volatile alkane products, liquid GC/MS was again
used to analyze the organic layer of extracted ADO reactions.
Using nonanal as the substrate, octane, octanal, and octanol
were observed (Figure 1). Peak identification was confirmed by
comparison of retention time and mass spectra to authentic
standards. Furthermore, a small amount of heptane was also
observed, and the time course of heptanal formation is
consistent with octanal’s being a precursor (SI Figure S1).
ADO was reacted with aldehydes of varied chain lengths to
determine the effect of chain length on product formation. The
formation of the additional products is a general feature of the
ADO reaction. However, the relative yields of aldehyde and
alcohol products varied significantly with the chain length of
the aldehyde substrate (Table 1 and SI Figure S2). In a separate
experiment, replicate reactions of 30 μM ADO with octanal or
nonanal (500 μM) and NADH/phenazine were quenched at 2
min. Products were determined by GC/MS (Table 1). A
marked decrease in relative yields of aldehyde and alcohol
products were observed as the alkyl chain length was decreased
from C9 to C8. Investigation of a range of aldehydes revealed
that relative yields of the one-carbon-shorter alcohol and
aldehyde products was optimum with nonanal and decanal and
decreased with shorter and longer alkyl chains.
Because the NADH/phenazine/O2 turnover system could
afford the production of reactive oxygen species, control
reactions were performed in which spinach ferredoxin, spinach
ferredoxin reductase, and NADPH were substituted for the
NADH/phenazine reduction system. The additional aldehyde
and alcohol products were still observed, suggesting that they
represent enzyme catalyzed products (SI Figure S3).
Furthermore, in the absence of NAD(P)H or enzyme, no
products are observed with either the ferredoxin or phenazine
systems.
Further experiments were conducted to confirm that the
observed novel products were not contaminants. First, the
incubation of aldehyde substrates with an increasing concen-
tration of enzyme yielded an increased rate of production of all
three one-carbon-shortened productsalkane, alcohol, and
aldehydeas well as the two-carbon-shorter alkane. Second, in
light of the recent finding that catalase greatly increases ADO
reaction turnover by removing the inhibitory H2O2 produced
by the inefficient coupling of the reduction system to the
enzyme,38 a control ADO reaction in the presence of catalase
(1 mg/mL) was also analyzed. The one-carbon-shortened
alcohol and aldehyde products could be detected in the
presence of catalase. Last, to definitely confirm that the
observed products were derived from the substrate aldehyde,
[1,2-13C]octanal was synthesized. The purity was analyzed by
NMR (SI Figure S4). The double-labeled aldehyde was used as
an ADO substrate. Upon reaction, products resulting from
removal of the [13C]-labeled aldehydic carbon from the double-
labeled aldehyde substrate would retain one 13C atom,
distinguishing products from any unlabeled contaminating
aldehydes or alcohols. In a complete reaction mixture with
[1,2-13C]octanal, the heptane, heptanal, and 1-heptanol
products could be detected by GC/MS, as expected (Figures
2 and SI S5). MS analysis was consistent with the presence of a
single 13C label on the new C1 of the products, revealing that
Figure 1. (A) GC/flame ionization detector (FID) chromatogram of
ADO reaction with nonanal. (B) Mass spectra of octanal and octanol
from GC/MS analysis (black) compared with mass spectra from the
National Institute of Standards and Technology (NIST) database
(red). (C) Time dependence of product formation in ADO reaction
with nonanal.
the oxygenated products are derived from [1,2-13C]octanal.
Electron ionization mass spectrometry (EI-MS) produced the
signature fragmentation pattern for the aldehyde and alcohol
products, yielding ions that result from dehydration [m − 18],
elimination of ethylene [m − 28], and dehydration with
terminal demethylation [m − 33].36
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Table 1. Alkyl-Products of 30 μM ADO Reactions
substrate
alkane
aldehyde
<5
alcohol
<5
a
octanal
16 10
16
nonanal
9
17
5
10
7
a
Product concentrations (μM) of 2 min reactions with NADH/
phenazine in triplicate standard deviation.
3.2. Single-Carbon Reaction Coproducts. In light of
these prominent oxygenated reaction products with C8−C10
aldehydes, we considered that other one-carbon products might
also form.
Since nonanal produced high relative yields of the alcohol
and aldehyde products (Table 1), experiments were conducted
using [1-13C]nonanal and 13C NMR to analyze coproducts.
The phenazine-reducing system was shown to cause minor
oxidation of formaldehyde to formate in preliminary experi-
ments, and thus, the ferredoxin-reducing system was used in
these experiments. Following the reaction, [13C]formate was
readily apparent in the 13C NMR spectrum (Figure 3). Another
single carbon compound, [13C]bicarbonate, appeared at
substoichiometric levels over the course of a long overnight
acquisition. The low level of bicarbonate detected is consistent
with its arising from a partial oxidation of formic acid.
Additional peaks were identified as [1-13C]-1-nonanol and
[1-13C]nonanoic acid, both detected in the aldehyde prepara-
tion (SI Figure S6), and DMSO, a reaction additive. An
observed peak at 60.81 ppm was not identified.
Figure 3. 13C NMR of ADO reaction mixture with [1-13C]nonanal.
production of other products. [13C]Formaldehyde was not
identified using an HPLC−MS assay for formaldehyde
derivatized with Nash reagent39 that was shown with standards
to readily detect 30 μM formaldehyde (SI Table S1). Carbon
monoxide was looked for in ADO reaction mixtures incubated
in sealed cuvettes. After reacting, reduced myoglobin was
injected into each cuvette, and the visible spectrum was
analyzed for the characteristic absorbance of myoglobin−CO at
425 nm. On the basis of the minimal spectral change, we can
infer that CO, if formed, would account for <1% of the
coproduct formed. (SI Figure S7). Other experiments using
[1-13C]octanal as the substrate yielded [13C]formate, as
expected.
3.3. Oxygen Incorporation into Alcohol and Aldehyde
Products. Before conducting experiments with [18O]O2, we
first confirmed that the formation of all of the reaction products
depended on the presence of oxygen. While oxygen levels can
be reduced to less than 10 ppm in an anaerobic chamber, if the
ADO has a low Km for oxygen, as has been suggested, some
Notably, formaldehyde, carbon monoxide, and methanol
were not detected by 13C NMR. The 13C NMR chemical shifts
of [13C]formaldehyde, [13C]methanol, and [13C]carbon mon-
oxide are 84.5, 51.6, and 181.3 ppm, respectively, as reported by
the University of Wisconsin Biological Magnetic Resonance
Data Bank. Further experiments mitigated against the
Figure 2. GC/MS spectra of heptanal (A, B) and 1-heptanol (C, D) products of ADO reaction with unlabeled octanal (A, C) and [1,2-13C]octanal
(B,D). EI-MS fragmentation patterns are inset with the mass of major fragments indicated.
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ADO activity would still be expected, even after degassing in an
anaerobic chamber.15,27,38 To further deplete oxygen, reactions
were run in an anaerobic chamber following preincubation with
homoprotocatechuate 2,3 dioxygenase from Brevibacterium
fuscum40 and homoprotocatechuate (HPCA) as an oxygen
scavenging system. Under these conditions, using dodecanal as
a substrate, the amount of undecane detected was comparable
to control reactions lacking NADH (SI Figure S8).
Accordingly, no undecanal or undecanol could be detected in
these reaction mixtures. These data are consistent with recent
findings and strongly support a strict oxygen requirement for
the ADO reaction, including the novel product species
identified in the present study.38
To follow oxygen-18 incorporation during the ADO reaction,
a reaction with ADO and nonanal was conducted under an
[18O]O2 atmosphere at pH 7.4. The reaction mixture at 10 min
was treated with an excess of equine alcohol dehydrogenase and
NADH to reduce any remaining aldehyde and prevent further
exchange. Any octanol thus produced, in addition to the ADO
product octanol, was derivatized with bis(trimethylsilyl)-
acetamide and analyzed by GC/MS (Figure 4A). The
the aldehyde 18O with solvent during the 10 min incubation
period, consistent with recent exchange measurements by Li et
al.15 and essentially complete incorporation of 18O from
[18O]O2 into the product octanol.
3.4. Reaction of ADO with Product Alcohol. The
observation of 18O-labeled oxygen incorporation into the
alcohol product of the ADO reaction with nonanal solidified
the identification of ADO as an oxygenase. Diiron oxygenases
such as MMO that form hydroxylated products are often able
to both react with hydrocarbons to form alcohol adducts and
also oxidize the alcohol products to aldehydes.41 In the case of
ADO, addition of nonane or nonanol to the complete reaction
mixture in the absence of decanal failed to yield any oxygenated
or oxidized products. However, after addition of excess [1-13C]-
1-nonanol to an ADO reaction with decanal, [1-13C]nonanal
was observed by GC/MS (Figure 5). (As expected, unlabeled
Figure 5. 13C incorporation into nonanal product. 13C-Labeled (black)
and unlabeled nonanol (gray) were added at 0.5 and 1.5 mM to a
complete ADO reaction with 200 μM decanal. Error bars indicate
standard deviation of a triplicate analysis.
nonane product was observed, whereas the nonanol product
detection was obscured by [1-13C]-1-nonanol addition.) The
percent incorporation of 13C from the [1-13C]-1-nonanol was
calculated to be 14 and 26% for 0.5 mM and 1.5 mM added
[1-13C]-1-nonanol, respectively, after correction for natural
abundance 13C in the [12C]-1-nonanol control. At 1.5 mM, the
added nonanol exceeded the solubility limit in water (∼1 mM);
therefore, the 2-fold increase in percent label incorporation
from 0.5 to 1.5 mM added [1-13C]-1-nonanol is consistent with
a 2-fold increase in soluble nonanol.
Although it appears that aldehyde products can react with
ADO to undergo deformylation with formation of a (n − 1)
alkane, we could find no evidence for enzyme-dependent
oxidation of the product aldehydes to form acid adducts. It is
also clear from the results shown in Figure 5 that the decanal
substrate does not prevent formation of the aldehyde product
from the 13C-labeled product alcohol. Finally, the total yield of
(n − 1) products is not increased by addition of 13C-labeled
product alcohol.
Figure 4. (A) ADO reaction scheme with [18O]O2 using alcohol
dehydrogenase (ADH) to convert octanal to octanol, followed by
derivatization with bis(trimethylsilyl)acetamide. The major ion of
trimethylsilyloctanol is shown. (B) Relative abundance of the major
ion (187) and isotopes of trimethylsilyloctanol. Error bars indicate
standard deviation of triplicate analysis.
derivatized alcohol product contained substantial levels of 18O
(Figure 4B) in the major m+/z ion (187). The major ion
corresponds to the loss of one methyl group from the
trimethylsilyl adduct. The mass spectrum derived from the
sample exposed to 16O2 showed the expected ratio of n, n + 1,
and n + 2 m+/z ions owing to the natural abundance of 13C
atoms (Figure 4B). The mass spectrum of the reaction product
obtained under [18O]O2 clearly exhibits an n + 2 ion at an m+/z
of 189 with a concomitant shift of the isotopic pattern by two
mass units. It was calculated from triplicate data that the oxygen
incorporation into the products from [18O]O2 was 36% 12%.
On the basis of the product ratios previously determined
(Table 1), octanol accounted for 37% of the oxygenated
products. Thus, the data suggested nearly complete exchange of
̈
3.5. Mossbauer and EPR Characterization of ADO. The
observation of alcohol and aldehyde products that apparently
derive from an intermediate alkane product suggests that the
reactive diiron cluster of ADO is capable of catalyzing types of
chemistry other than deformylation of aldehydes. Alkane
hydroxylation with incorporation of oxygen from [18O]O2 is
the hallmark reaction of the diiron-containing soluble form of
MMO and related enzymes from the bacterial multicomponent
monooxygenase (BMM) family. Past studies have employed
EPR, Mossbauer, and other spectroscopies to provide detailed
̈
characterizations of BMM family enzymes.42−51 Mossbauer
̈
spectroscopy, perhaps more than any other single technique,
can furnish detailed information about the electronic structure
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of the diferrous state of diiron(II) proteins. Spectra recorded in
applied magnetic fields are rich in information, and this
information can be used to assess the electronic structure and
reactivity of the protein under study and compare it with
corresponding properties of related proteins. The spectra of
proteins such as MMO hydroxylase component (MMOH)42,52
and RNR-R253 are immensely complex, and comprehensive fits
environment. For diferric MMOH, we have found ΔEQ = 1.05
mm/s and δ = 0.51 mm/s (identical sites) for one
preparation;52 ΔEQ1 = 1.16 mm/s, ΔEQ2 = 0.87 mm/s, and
δ(1) = δ(2) ≈ 0.51 mm/s for a two-site fit in a subsequent
preparation.
3.5.2. Diferrous ADO. After anaerobic addition of sodium
dithionite to the sample of Figure 6, a Mossbauer spectrum
̈
of the Mossbauer spectra, yielding a set of spin Hamiltonian
(Figure 7, top) was observed with ΔEQ = 3.10(4) mm/s and δ
̈
parameters, have not yet been reported. Fortunately, the
spectra of diferrous ADO, while still complex, have some
encouraging features (reflecting good structural homogeneity)
which allowed us to obtain the first complete set of spin
Hamiltonian parameters. The ADO data analysis, presented
below, has yielded insights that may allow us to fit the data of
diiron enzymes such as MMOH and RNR-R2 in the future,
thereby providing a detailed comparison of the electronic
structure of the diiron centers in these proteins.
3.5.1. Diferric ADO. Figure 6 top shows a Mossbauer
̈
spectrum of “as isolated” ADO recorded at 4.2 K in the absence
Figure 6. 4.2 K Mossbauer spectra of diferric ADO recorded in zero
̈
Figure 7. Variable field, variable temperature Mossbauer spectra of
̈
field (top) and in a field of 5.0 T applied parallel to the observed γ
radiation (bottom). The red lines are spectral simulations for a species
with S = 0 assuming two equivalent Fe sites with ΔEQ = 1.27 mm/s, η
= 0.41, and δ = 0.52 mm/s. Sign of ΔEQ was determined from fitting
the 5.0 T spectrum.
diferrous ADO (conditions indicated in figure). Spectra were recorded
in magnetic fields applied parallel to the observed γ radiation. The red
lines represent spectral simulations based on eqs 1−4 using the
parameters listed in Table 2. The spectra were simulated using the
2Spin routine of WMOSS. Additional spectra are shown in SI Figure
S9.
of an applied magnetic field (B = 0). The spectrum consists of a
doublet that is slightly asymmetric with a quadrupole splitting
ΔEQ = 1.27 mm/s and isomer shift δ = 0.52 mm/s. It is
possible to represent the spectrum equally well as a
superposition of two doublets with ΔEQ(1) = 1.30 mm/s and
ΔEQ(2) = 1.24 mm/s that may differ in δ by 0.01 mm/s, using
Lorentzian lines of 0.31 mm/s full width at half-maximum, but
we doubt whether such a representation is more meaningful
than the one for one doublet (comment: the sample was quite
concentrated. It had an iron concentration of ∼3.5 mM, which
leads to some line-broadening due to thickness effects). An
applied field B = 5.0 T (Figure 6, bottom) elicits a pattern
characteristic of a system with a diamagnetic (S = 0) ground
state, as is typically observed for antiferromagnetically coupled
diiron clusters with high-spin FeIII sites. The observation of one
doublet, or two differing slightly in ΔEQ, suggests that both iron
sites of ADO have a fairly similar, possibly identical, ligand
= 1.30(2) mm/s in zero magnetic field. These parameters are
characteristic of high-spin FeII sites, and they are close to those
reported for other diiron(II) enzymes, such as MMOH, toluene
4-monooxygenase, and RNR-R2.25,32,42,54 The observation of a
single doublet suggests that for the diferrous state, just as for
the diferric one, the two FeII sites are quite similar. Mossbauer
̈
spectra of high-spin FeII complexes recorded in variable applied
magnetic fields yield intricate spectral patterns from which
detailed fine structure and hyperfine structure parameters can
be extracted. These spectra, however, depend on numerous
parameters (as many as 20 for a diiron protein, even if all
tensors share the same principal axes frame), and because of
this complication, a complete set of parameters has not yet
been reported for any diiron(II) protein. However, well-
resolved spectra, rich in detail, of reduced ADO allowed us to
obtain such a set of parameters.
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The interactions relevant for a combined Mossbauer and
̈
EPR study of diferrous ADO are the zero-field splittings (ZFS),
electronic Zeeman, magnetic hyperfine, nuclear Zeeman, and
quadrupole interactions of the two Fe sites. We also require a
̂
̂
̂
̂
̂
̂
term, Hexch = JS1·S2, that describes exchange interactions
between the two sites with electronic spin S1 = S2 = 2. For
ADO, as for MMOH,42 the exchange coupling is quite small,
namely |J| < 0.5 cm−1, and ferromagnetic (J < 0); see below. In
the ensuing sections, it will be useful to employ a spin
Hamiltonian that can conveniently be evaluated and discussed
in the weak coupling limit, that is, under conditions for which |J|
≪ zero-field splittings.
̂
̂
̂
H = He + Hhf
(1)
⎧
⎩
⎡
̂
z,i
2
̂
̂
̂
̂
⎨
He = JS1·S2 + ∑ βS ·g ·B + D⎢S − 2
̃
i
i
i
⎣
Figure 8. X-band EPR spectra of diferrous ADO. The red traces are
simulations based on eq 2 using D1,2 = −6.6 cm−1, E/D1,2 = 0.21, σ(E/
D)1,2 = 0.0176; gx1,2 = gy1,2 = 2.0, gz1,2 = 2.15, Jx,y,z = −0.253 cm−1, and
packet width = 10 mT. Experimental conditions: T = 3 K; 9.408 GHz
in parallel mode and 9.646 GHz in transverse mode; 1 mW microwave
power, modulation 1 mT. For comments on spin−spin dipolar
interactions, see the SI. The insert shows the lowest four spin levels of
the coupled diferrous state.
i=1,2
⎫
⎬
⎭
⎤
Ei
D
2
2
̂
̂
+
(Sx,i − Sy,i)⎥
⎦
i
(2)
(3)
(4)
̂
̂
̂
̂
̂
i
Hhf
=
{S ·A ·I + H (i) − g β B·I }
∑
i
i
i
Q
n
n
i=1,2
eQVZZ,i
⎡
⎤
2
15
4
2
2
Y ,i
̂
̂
̂ ̂
+ η(IX,i − I )
HQ(i) =
I
−
⎢
⎣
⎥
⎦
Z,i
gap Δ (for B = 0). For the doublet considered here, the
resonance condition can be written as described in Hendrich et
al.,43
12
where the index i = 1,2 labels the two sites. Furthermore, in eq
2, D1, E1 and D2, E2 are the tetragonal and rhombic zero-field
splitting parameters of site 1 and 2, respectively, and g1 and g2
are the electronic g-tensors. A1 and A2 are the 57Fe magnetic
hyperfine tensors and HQ(i = 1,2) describes the interactions of
the nuclear quadrupole moment Q with the electric field
gradient (EFG) tensor (principal components VXX, VYY, and
2
hν = Δ + (geff βB cos α)2
(5)
̂
where ν is the microwave frequency, α is the angle between B
and the molecular z axis, and geff ≈ 4(g1z + g2z) = 8gz ≈ 16 with
gz = (g1z + g2z)/2. Briefly, the EPR-active doublet reflects the
following situation. The ZFS parameters D1 and D2 of the two
iron sites of reduced ADO are negative, indicating that for each
V
ZZ); η = (VXX − VYY)/VZZ is the asymmetry parameter. Our
analysis indicates that the ZFS and g and A tensors of both Fe
sites share a common principal axis system (x, y, z) and that the
principal axes of the two EFGs (X, Y, Z) are rotated with
respect to (x, y, z). While all parameters of eqs 1−4 are
site the ground state is a spin doublet of m1 = 2 or m2 =
2
parentage, where m1 and m2 are the magnetic quantum
numbers. In Figure 8, which shows the lowest four spin levels,
as well as in SI Figure S10, these states are labeled |2 ⟩. In the
absence of exchange interactions, a second-order perturbation
treatment yields for the spin doublet of each site a splitting Δi =
3(E/D)i2Di.
accessible by Mossbauer spectroscopy, only eq 2 is required for
the EPR results. To simplify the presentation of a rather
complex exchange-coupled system, we state at the outset a
̈
result obtained after a series of Mossbauer simulations: namely,
̈
the parameters of the two sites of ADO differ only in a minor
way, and within our resolution, the tensors of both sites have
the same spatial orientation. For simplicity, we will use a double
label when the quantities are the same for both sites, e.g. (E/
D)1,2 stands for (E/D)1 = (E/D)2.
For a mononuclear FeII, such quasidegenerate doublets
frequently yield a Δm = 0 EPR transition near geff = 4g(1z),(2z)
≈
8; however, in the case of weak exchange coupling between the
two sites, the two |2 ⟩ ground doublets combine to produce
two new doublets, which we labeled as dEPR and dEPR‑silent in
Figures 8 and SI S10. For ferromagnetic coupling, dEPR is the
ground doublet; its two levels are split by Δ ≈ Δ1Δ2/8|J| (or Δ
≈ Δ1,2/8|J| for equivalent sites).43 The expression for Δ is
approximate and serves to guide the reader; for our analysis, we
have used the exact solutions to eqs 1−4. This doublet is EPR-
active at the X-band, whereas dEPR‑silent, at an energy of ε ≈ 8J, is
essentially diamagnetic and EPR-silent. Diferrous ADO has a
very small J value. Theoretically, the sign of J can be determined
by studying the temperature dependence of the EPR signal
below ≈6 K. However, such measurements are difficult to
perform with the widely available EPR flow cryostats. The sign
of J can be unambiguously determined from the low
Figure 8 shows low-temperature X-band EPR spectra of
diferrous ADO recorded at T = 3 K with a bimodal cavity
operated in parallel (bottom) and transverse (top) modes. A
prominent signal near g = 16 is seen in both modes. A
transverse mode spectrum for diferrous ADO with a resonance
at g = 13 was previously reported by Marsh and co-workers;37
however, spectral simulations were not presented. Since the
spectra of Figure 8 are similar to those analyzed by Hendrich
and co-workers for reduced MMOH,43 we mention here only
the principal features of the electronic system as described by
eq 2. SI Figure S10, which leans in Figure 4 of Hendrich et al.,43
will aid the reader in the following discussion; the insert in
Figure 8 separately depicts the lowest four spin levels.
temperature Mossbauer spectra: we demonstrate by means of
̈
The observed EPR signal originates from transitions between
the levels of a quasidegenerate ground doublet with an energy
the spectra of SI Figure S9 that the coupling is ferromagnetic
and that dEPR is the ground state.
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Table 2. Fine Structure and Hyperfine Structure Parameters of Diferrous ADO Derived from the Analysis of the Mossbauer
̈
Spectra Using Eq 1−4
a
b
c
d
d
J
(cm−1
)
D
(cm−1
)
E/D
gx,y,z
Ax,y,z/gnβn (T)
δ (mm/s)
ΔEQ (mm/s)
η
−0.2
−6.6(10)
0.21(3)
2.00, 2.00, 2.15
−25(5), −22(3), −6.5/-8.0
1.30(2)
3.10(4)
0.7(2)
a
Spin−spin dipolar interactions between the two FeII spins may be comparable with the exchange term. Only the z-components of these interactions
b
matter (see SI). The quoted J may thus be Jeff,z = Jz + Jzspin‑dip . We estimate an uncertainty for Jz,eff of 0.1 cm−1. The parameters obtained for the
two Fe sites are essentially the same; thus, D stands for D1 = D2, etc. The two sites have different values for Az, indicated by the double entries. The
c
d
local electric field gradient tensors (EFG) are rotated relative to the ZFS by αEFG = 0°, βEFG = 20(5)° ,and γEFG = 40(10)°; the Euler angles (αEFG
,
βEFG, γEFG) rotate the principal axis frame of the EFG, (X, Y, Z), into the (x, y, z).
As argued in the SI, the z components of the 57Fe A tensors
listed in Table 2 and the theoretical curves (red lines) of
Figures 7 and SI S8. The simulations are not perfect, but given
the complexity of the system, the simulations shown in Figures
7 and SI S8 represent the data quite well over a wide range of
temperature and applied fields.
obtained from Mossbauer spectroscopy can be used to
̈
constrain g1z and g2z to 2.0 ≤ gz1,2 ≤ 2.15, a condition which
allowed us to determine Δ of eq 5 by simulating the EPR
spectra of Figure 8. A good representation of the EPR line
shape was obtained by assuming that the rhombicity parameters
(E/D)1,2 have a Gaussian distribution around a mean (E/D)1,2
= 0.21 with σ(E/D)1,2 = 0.017. The red lines in Figure 8 are
(SpinCount) simulations based on the parameters listed in the
caption. The simulation for the coupled system was obtained
for Δ = 0.31 cm−1 and g1z = g2z = 2.15. Note that the EPR
signal depends on Δ (and geff) and only indirectly on (E/D)i, Di
4. DISCUSSION
It is shown here that ADO can carry out reactions other than
deformylation of long-chain fatty aldehydes. This observation
was facilitated by using liquid rather than head space GC/MS,
which afforded the detection of low volatility compounds, and
by investigating aldehydes shorter than the C18 physiological
substrate. Importantly, it is shown that all reactions of ADO
require O2 and that oxygen from [18O]O2 is incorporated into
the (n − 1) alcohol products reported here. This suggests that
the diiron cluster can activate oxygen and promote reaction
chemistry similar to that observed for MMO and other diiron
cluster-containing oxygenases. This is consistent with the
formation of a high-valence intermediate at some point in the
ADO reaction cycle. The relevance of such a species to the
deformylation and oxygenation reactions is discussed here.
4.1. Correlation of Substrate Chain Length and
Alternative Product Formation. It is shown above that
the relative yields of (n − 1) alkane, alcohol, and aldehyde
products vary with the length of the substrate fatty aldehyde.
The peak production of the (n − 1) aldehyde and alcohol
products relative to alkane occurs with C9−C10-length aldehyde
substrates. The C9−C10 substrate length corresponds to the
position of a ∼ 90° bend in the substrate binding pocket of
ADO relative to the diiron cluster as revealed by the X-ray
crystal structure of the diferric enzyme (PDB 2OC5)11 (Figure
9). Without the additional anchoring gained from binding on
both sides of the bend, the (n − 1) carbon of the shorter
substrates may be able to shift more easily into position to react
at the diiron cluster following release of the terminal carbon as
formate, as further discussed below. Such a shift would facilitate
the types of reactions commonly observed for MMO and
similar bacterial multicomponent dinuclear-iron-cluster-con-
taining monooxygenases (BMM family). This suggests a
mechanism that accounts for both deformylation and the
formation of (n − 1) alcohol and aldehyde products.
and J; the latter quantities can be obtained from Mossbauer
̈
spectroscopy.42
We have recorded Mossbauer spectra of diferrous ADO
̈
between 4.2 and 160 K in applied magnetic fields up to 8.0 T.
Here, we list some relevant observations; detailed comments
can be found in the SI. The presence of an EPR-active ground
doublet, dEPR, with a small splitting,55 implies that the 4.2 K
Mossbauer spectra exhibit paramagnetic hyperfine structure,
̈
even in moderate applied fields (mixing of the two levels, |2up⟩
and |2down⟩, by the applied magnetic field is proportional to
gzβB/Δ). Indeed, small applied fields B induce in ADO sizable
magnetic hyperfine fields, Bint,z(i) at the 57Fe nuclei (i = 1, 2)
along the molecular z direction (see the 0.3 T spectra of SI
̂
Figure S12), given by Bint,z(i) = −⟨Szi⟩Azi/gnβn. The expectation
down
value ⟨Sz,i⟩ = ⟨2 |Sz,i|2down⟩ of the electron spin operator,
̂
̂
determined by eq 2, reaches saturation for B ≈ 1.5 T (SI Figure
S11), assuming the value ⟨Sz⟩ ≈ −2 for the ⟨2down⟩, the only
̂
level significantly populated at 4.2 K for B ≥ 1 T. Furthermore,
for B > 1.5 T, the system is essentially decoupled, that is, at
these fields for the spectral simulations of the 4.2 K spectra, one
can set J = 0 and treat the spectra as if they were originating
from two independent FeII sites.
Our simulations for B > 2.0 T yield Az1/gnβn = −6.5 T and
Az2/gnβn = −8.0 T. Az1 and Az2 are the only parameters for
which we found noticeable differences between the two sites.
Between T = 25 and 50 K, in the regime of fast relaxation of the
electronic spins, the spectra are fairly sensitive to the zero-field
splitting parameters D1,2; our simulations suggest that −7 cm−1
< D1,2 < −5 cm−1. Above 100 K, the magnetic hyperfine field
̂
int,i = −⟨Si⟩th·Ai/gnβn is only marginally dependent on the ZFS
B
4.2. A Proposal for the Mechanism of ADO and the
Formation of (n − 1) Products. Members of the BMM
family normally react poorly with aldehyde substrates, and
these reactions result in oxidation rather than deformylation. In
contrast, some cytochrome P450s, such as sterol 14R-
demethylase (CYP51) from Mycobacterium tuberculosis react
with aldehyde adducts of steroids, resulting in deformylation
and desaturation of the steroid.56−58 Computational studies
have suggested that the CYP51 reaction is initiated by attack of
the heme FeIII-peroxo intermediate at the aldehydic carbon to
form an Fe-peroxohemiacetal intermediate. In the protein
̂
̂
parameters (⟨Si⟩th is the expectation value of S averaged over all
thermally accessible spin levels).55 Under these conditions, we
established the sign of ΔEQ, η, and a good estimate for the A
tensor components. This is discussed in greater detail in the SI.
Finally, and significantly, the 0.3 T Mossbauer spectra of SI
̈
Figure S12 show that the EPR-active spin doublet with splitting
Δ is the ground state; hence, J < 0. With this information, and
taking the EPR constraint Δ ≈ 0.31 cm−1 into account, we have
simulated the spectra for the whole data set by least-squares
fitting groups of spectra. These efforts yielded the parameters
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Figure 10. Proposed mechanism for ADO reaction. Observed
products are boxed with gray background.
to aldehydes.41 If ADO forms a Q-like intermediate as a
consequence of the deformylation reaction, then it might form
the (n − 1) alcohol and aldehyde products we observe here by
the mechanism shown in Figure 10. The initially formed (n −
1) hydrocarbon could either be released or remain in the active
site and shift to react with the putative Q-like intermediate via a
hydrogen atom abstraction and rebound mechanism similar to
that described for MMO. Release of the resulting alcohol,
followed by rebinding to the fraction of ADO that initially
released hydrocarbon, could result in (n − 1) aldehyde
formation because this fraction would retain a Q-like
intermediate.
Several predictions can be made on the basis of the
mechanism shown in Figure 10. First, the (n − 1) alcohol
product should be 13C-labeled at (the new) C1 when the
substrate aldehyde is doubly labeled at C1 and C2 because it is
formed without intermediate release from the enzyme. Second,
the release and rebinding of the intermediate alcohol suggests
that addition of 13C-labeled alcohol to a reaction started with
unlabeled substrate aldehyde would result in incorporation of
label into the (n − 1) aldehyde. Third, the total product and, in
particular, the ratio of the (n − 1) alkane and aldehyde
products would not be altered by addition of exogenous alcohol
because the maximum aldehyde yield is determined by the
fraction of enzyme that initially releases (n − 1) alkane. Fourth,
the ratio of alkane released versus bound is likely to depend
strongly on the nature of the initial aldehyde substrate.
Consequently, the distribution of (n − 1) products should
differ from substrate to substrate. Each of these predictions has
been shown here to be correct.
Figure 9. The crystal structure of ADO (PDB 2OC5)11 showing a C18
fatty acid bound to the cluster. The binding orientation of the fatty
acid is presumably that of the aldehyde substrate and reveals a sharp
bend after carbon 10 (the first 10 carbons prior to the bend are shown
in green). The bend is enforced by the labeled residues, which are
shown with van der Waals radii superimposed.
environment, this initiates facile heterolytic C−C bond
cleavage58 to yield a substrate carbanion and an Fe-
peroxoformate adduct. Heterolytic O−O bond cleavage
would yield formate and the FeIV-oxo heme π cation radical
compound I of CYP51, but this reaction is apparently coupled
with one-electron transfer from the substrate carbanion to yield
compound II. Compound II can, in turn, accept a second
electron from the substrate to yield the desaturated product.
Very similar chemical steps can be proposed for the initial
deformylation reaction carried out by ADO (Figure 10). A
peroxo intermediate isoelectic to that formed in P450 is
observed for most members of the BMM family.29,32,34,59,60
One electron from each iron of the diferrous cluster is
transferred to O2 as the adduct forms. Attack of this species on
the substrate aldehyde and subsequent C−C bond cleavage
could proceed as described for CYP51. However, the additional
activation energy required to abstract an electron from the
carbanion located on the primary carbon of the alkane versus
the secondary carbon of the steroidal ring suggests that electron
transfer coincident with O−O bond cleavage is unlikely. The
product of direct expulsion of formate would be an FeIVFeIV-
oxo species isoelectronic with compound I of P450. Such a
species has been trapped and characterized as compound Q of
the MMOH catalytic cycle.32,34,61
One potential difficulty with the mechanism is the possible
binding of substrate aldehyde to the fraction of ADO with an
active Q-like cluster following release of the alkane. This might
compete with the low concentration of diffusible alcohol
product to form an (n − 1) carboxylic acid product, but no
enzyme-dependent acid formation is observed.
The mechanism we propose suggests that ADO can utilize
the diiron cluster to carry out three substantially different types
of chemistry: namely, attack by a peroxo adduct leading to
MMOH compound Q is known to carry out many types of
reactions, including desaturation.32 However, desaturation has
not been described for the terminal C−C bond of saturated
straight-chain hydrocarbon substrates. In contrast, two well
characterized reactions of compound Q are hydrocarbon
hydroxylation at the primary carbon and oxidation of alcohols
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deformylation, hydrogen atom abstraction followed by rebound
to give alkane hydroxylation, and two-electron oxidation of an
alcohol. The latter two reactions are well-known reactions for
MMO, but the first reaction has not been observed. This
suggests that the reactive diiron cluster of ADO, while similar
to that of MMOH, must have specific differences that tune it to
aldehyde deformylation. We do not yet know what these
difference are, but we wish to make two comments. The integer
spin EPR signals of diferrous ADO and MMOH are quite
similar. This similarity reflects two properties of the electronic
states: First, in both proteins, the two FeII have negative zero-
field splittings, which yield local ground states derived from the
m1 = 2 and m2= 2 spin levels of the two local sites. Second,
both proteins have diiron(II) centers coupled by weak
ferromagnetic exchange interactions. Although the two clusters
look deceptively similar from the perspective of EPR, the
equivalent of the FeIVFeIV cluster remaining after the
deformylation reaction must be returned to the FeIIIFeIII
state. The mechanism proposed here requires only 2 electrons
from NADH because the formal FeIVFeIV species is utilized for
mixture of hydroxylation and oxidation reactions in which the
electrons are derived from the substrates. Unfortunately, it is
not currently possible to accurately account for NADH (or
other reducing equivalent) utilization because the authentic
biological reductase has not been isolated.
CONCLUSIONS
■
We show here that ADO is a significantly more versatile catalyst
than previously recognized. It is apparently possible to activate
the enzyme for alkane hydroxylation by reacting the enzyme
with an intermediate chain aldehyde. The C−H bond
dissociation energy of the substrate alkane for the hydroxylation
reaction is ∼98 kcal/mol, placing ADO among the most potent
biological oxidants. This strongly supports the formation of a
compound Q-like intermediate as a key feature of the ADO
mechanism and opens the possibility of adapting the enzyme
for a wide range of chemistry involving abundant hydrocarbons.
Mossbauer data reveal substantial differences. Thus, diferrous
̈
ADO has (essentially) equivalent FeII sites in structurally well-
defined environments, in contrast to diferrous MMOH, which
has inequivalent Fe sites, as witnessed by the observations that
the sites have very distinct ΔEQ values (ΔEQ(1) ≈ 3.2 mm/s
and ΔEQ(2) ≈ 2.4 mm/s)44 and different magnetic hyperfine
tensors (see Figure 13 of ref 42). ΔEQ(1) of MMOH is well-
defined (in all preparations), yielding a quadrupole doublet
with sharp lines. In contrast, ΔEQ(2) in all preparations is
ASSOCIATED CONTENT
■
S
* Supporting Information
Methods for HPLC−MS detection of formate and form-
aldehyde, CO detection, syntheses of 13C-labeled compounds,
distributed with a variance σΔE = 0.3 mm/s if a Gaussian
Q
EPR and Mossbauer; time course of heptane production; GC/
̈
distribution about the average 2.4 mm/s is assumed. This
distribution reflects a structural heterogeneity that is not yet
understood. However, the successful analysis of the ADO
spectra gives us confidence that we may obtain, finally, a good
set of spin Hamiltonian parameters of this important protein.
The small value obtained for the exchange coupling constant
of diferrous ADO (J ≈ −0.2 cm−1) suggests, on the basis of the
reported studies of MMOH and RNR-R2,31,32,42 that the
ferrous sites are bridged by two carboxylates with unknown
bridging modes (μ-(η1, η2) and/or μ-1,3 coordination).62 The
bearing of these differences on the distinct reactivities of the
two enzymes is not clear and will require further study.
4.3. Comparison with Previous Mechanistic Studies.
Previous studies of ADO from two different cyanobacteria have
shown that (i) oxygen from [18O]O2 is incorporated into the
product formate, (ii) hydrogen is retained by the terminal
carbon, and (iii) hydrogen from solvent is incorporated into the
alkane product.14,26 Each of these findings is consistent with the
mechanism proposed in Figure 10. Recently, it has been shown
that an alternative octadecanal substrate with an additional
cyclopropyl group at C3−C4 is converted to a ring-open
terminal alkene product characteristic of intermediate radical
formation.63 It was suggested that one way this could occur is
through homolytic C−C bond cleavage. The alternative
presented in Figure 10 is that the substrate radical is formed
as in the MMO reaction by hydrogen atom abstraction from
the cyclopropyl alkane. Delocalization and ring-opening would
then rapidly occur, but in an MMO-type reaction, rebound of
the hydroxyl radical would be expected to yield octadecanol.
Instead, enzyme inactivation and formation of a covalent
adduct occurred, presumably by trapping of the radical by a
group in the enzyme active site. The mechanisms that have
been proposed by others for ADO on the basis of previous
studies require the input of four electrons from NADH or
another source for each cycle because the FeIIIFeIII cluster must
be reduced to the FeIIFeII state to bind O2, and then the
FID chromatograms of ADO reaction with ferredoxin/
ferredoxin reductase; 13C NMR analysis of synthesized
[1-13C]nonanal and [1,2-13C]octanal; MS of 13C-heptane
product of ADO reaction with [1,2-13C]octanal, HPLC−MS
analysis of derivatized [13C]-formaldehyde; investigation of CO
production in ADO reaction, O2-depleted ADO reaction;
comments on EPR and Mossbauer data analysis, including
̈
Mossbauer spectra. This material is available free of charge via
̈
the Internet at http://pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author
*E-mail: wacke003@umn.edu.
■
Present Addresses
⊥Department of Chemistry and Biochemistry, University of
South Carolina, 631 Sumter Street, Columbia, SC 29208.
∥National High Magnetic Field Laboratory, Florida State
University, 1800 E. Paul Dirac Drive, Tallahassee, FL 32310-
3706.
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
This work is supported by NIH Grant GM100943 to J.D.L.,
NSF grants CHE-1012485 and CHE-1305111 to E.M., and
University of Minnesota Initiative for Renewable Energy and
the Environment Grant to J.D.L. Funding for NMR
instrumentation was provided by the Office of the Vice
President for Research, the Medical School, the College of
Biological Science, NIH, NSF, and the Minnesota Medical
Foundation. The authors are thankful for the contributions of
the following people: Fred Schendel (fermentation, University
of Minnesota Biotech Resource Center), Tom Krick (HPLC−
MS, University of Minnesota Center for Mass Spectroscopy
and Proteomics), Todd Rappe (13C NMR, University of
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ACS Catalysis
Research Article
(35) Kende, A. S.; Fludzinski, P. Org. Synth. 1990, Coll. Vol. 7, 221.
(36) Liedtke, R. J.; Djerassi, C. J. Am. Chem. Soc. 1969, 91, 6814.
(37) Das, D.; Eser, B. E.; Han, J.; Sciore, A.; Marsh, E. N. Angew.
Chem., Int. Ed. 2011, 50, 7148.
(38) Andre, C.; Kim, S. W.; Yu, X.-H.; Shanklin, J. Proc. Natl. Acad.
Sci. U.S.A. 2013, 110, 3191.
Minnesota Biomedical NMR Resource Center), and Jennifer
Seffernick and Anna Komor for helpful discussion.
REFERENCES
■
(1) Lam, M. K.; Lee, K. T. Biotech. Adv. 2012, 30, 673.
(2) Jones, C. S.; Mayfield, S. P. Curr. Opin. Biotechnol. 2012, 23, 346.
(3) Malcata, F. X. Trends Biotechnol. 2011, 29, 542.
(4) Kung, Y.; Runguphan, W.; Keasling, J. ACS Synth. Biol. 2012, 1,
498.
(39) Nash, T. Biochem. J. 1953, 55, 416.
(40) Miller, M. A.; Lipscomb, J. D. J. Biol. Chem. 1996, 271, 5524.
(41) Stirling, D. I.; Colby, J.; Dalton, H. Biochem. J. 1979, 177, 361.
(42) Fox, B. G.; Hendrich, M. P.; Surerus, K. K.; Andersson, K. K.;
(5) Berry, D. A.; Robertson, D. E.; Skraly, F. A.; Green, B. D.; Ridley,
C. P.; Kosuri, S.; Reppas, N. B.; Sholl, M.; Afeyan, N. B. Joule
Biotechnologies, Inc., USA, 2009, WO2009111513A1.
(6) Schirmer, A.; Rude, M.; Brubaker, S. LS9, Inc., USA, 2010,
US20100249470A1.
(7) Berry, D. A.; Robertson, D. E.; Skraly, F. A.; Green, B. D.; Ridley,
C. P.; Kosuri, S.; Reppas, N. B.; Sholl, M.; Afeyan, N. B. Joule
Unlimited, Inc., USA, 2011, US20110262975A1.
(8) Franklin, S.; Somanchi, A.; Wee, J.; Rudenko, G.; Moseley, J.;
Rakitsky, W.; Zhao, X.; Bhat, R. Solazyme, Inc., USA, 2012,
WO2012106560A1.
(9) Lee, G. J.; Haliburton, J. R.; Hu, Z.; Schirmer, A. W. LS9, Inc.,
USA, 2013, WO2013039563A1.
(10) Shanklin, J.; Andre, C. Brookhaven Science Associates, LLC,
USA, 2013, WO2013032891A1.
(11) Schirmer, A.; Rude, M. A.; Li, X.; Popova, E.; del Cardayre, S. B.
Science 2010, 329, 559.
(12) Dennis, M.; Kolattukudy, P. E. Proc. Natl. Acad. Sci. U.S.A. 1992,
Froland, W. A.; Lipscomb, J. D.; Munck, E. J. Am. Chem. Soc. 1993,
̈
115, 3688.
(43) Hendrich, M. P.; Munck, E.; Fox, B. G.; Lipscomb, J. D. J. Am.
̈
Chem. Soc. 1990, 112, 5861.
(44) Banerjee, R.; Meier, K. K.; Munck, E.; Lipscomb, J. D.
̈
Biochemistry 2013, 52, 4331.
(45) Liu, K. E.; Valentine, A. M.; Wang, D.; Huynh, B. H.;
Edmondson, D. E.; Salifoglou, A.; Lippard, S. J. J. Am. Chem. Soc. 1995,
117, 10174.
(46) Shanklin, J.; Achim, C.; Schmidt, H.; Fox, B. G.; Munck, E. Proc.
̈
Natl. Acad. Sci. U.S.A. 1997, 94, 2981.
(47) Shu, L.; Nesheim, J. C.; Kauffmann, K.; Munck, E.; Lipscomb, J.
̈
D.; Que, L. Science 1997, 275, 515.
(48) Shu, L.; Broadwater, J.; Achim, C.; Fox, B.; Munck, E.; Que, L. J.
̈
Biol. Inorg. Chem. 1998, 3, 392.
(49) Cadieux, E.; Vrajmasu, V.; Achim, C.; Powlowski, J.; Munck, E.
̈
Biochemistry 2002, 41, 10680.
(50) Murray, L. J.; García-Serres, R.; Naik, S.; Huynh, B. H.; Lippard,
S. J. J. Am. Chem. Soc. 2006, 128, 7458.
89, 5306.
(13) Warui, D. M.; Li, N.; Nørgaard, H.; Krebs, C.; Bollinger, J. M.;
Booker, S. J. J. Am. Chem. Soc. 2011, 133, 3316.
(14) Li, N.; Nørgaard, H.; Warui, D. M.; Booker, S. J.; Krebs, C.;
Bollinger, J. M. J. Am. Chem. Soc. 2011, 133, 6158.
(15) Li, N.; Chang, W. C.; Warui, D. M.; Booker, S. J.; Krebs, C.;
Bollinger, J. M. Biochemistry 2012, 51, 7908.
(51) Makris, T. M.; Chakrabarti, M.; Munck, E.; Lipscomb, J. D. Proc.
̈
Natl. Acad. Sci. U.S.A. 2010, 107, 15391.
(52) Fox, B. G.; Surerus, K. K.; Munck, E.; Lipscomb, J. D. J. Biol.
̈
Chem. 1988, 263, 10553.
(53) Juarez-Garcia, C. H., Carnegie Mellon University, 1990.
(54) Lynch, J. B.; Juarez-Garcia, C.; Munck, E.; Que, L., Jr. J. Biol.
̈
(16) Shanklin, J.; Guy, J. E.; Mishra, G.; Lindqvist, Y. J. Biol. Chem.
2009, 284, 18559.
Chem. 1989, 264, 8091.
(55) Munck, E.; Surerus, K. K.; Hendrich, M. P. Methods Enzymol.
̈
(17) Friedle, S.; Reisner, E.; Lippard, S. J. Chem. Soc. Rev. 2010, 39,
2768.
(18) Nordlund, P.; Eklund, H. J. Mol. Biol. 1993, 232, 123.
(19) Rosenzweig, A. C.; Frederick, C. A.; Lippard, S. J.; Nordlund, P.
Nature 1993, 366, 537.
(20) Elango, N.; Radhakrishnan, R.; Froland, W.; Wallar, B.; Earhart,
C.; Lipscomb, J.; Ohlendorf, D. Protein Sci. 1997, 6, 556.
(21) Mathevon, C.; Pierrel, F.; Oddou, J.; Garcia-Serres, R.; Blonclin,
G.; Latour, J.; Menage, S.; Gambarelli, S.; Fontecave, M.; Atta, M. Proc.
Natl. Acad. Sci. U.S.A. 2007, 104, 13295.
(22) Lindqvist, Y.; Huang, W.; Schneider, G.; Shanklin, J. EMBO J.
1996, 15, 4081.
(23) Newman, L. M.; Wackett, L. P. Biochemistry 1995, 34, 14066.
(24) Sazinsky, M. H.; Bard, J.; Di Donato, A.; Lippard, S. J. J. Biol.
Chem. 2004, 279, 30600.
1993, 227, 463.
(56) Akhtar, M.; Calder, M. R.; Corina, D. L.; Wright, J. N. Biochem.
J. 1982, 201, 569.
(57) Vaz, A. D.; Pernecky, S. J.; Raner, G. M.; Coon, M. J. Proc. Natl.
Acad. Sci. U. S. A. 1996, 93, 4644.
(58) Sen, K.; Hackett, J. C. J. Am. Chem. Soc. 2010, 132, 10293.
(59) Bailey, L. J.; Fox, B. G. Biochemistry 2009, 48, 8932.
(60) Korboukh, V. K.; Li, N.; Barr, E. W.; Bollinger, J. M., Jr.; Krebs,
C. J. Am. Chem. Soc. 2009, 131, 13608.
(61) Lee, S. K.; Fox, B. G.; Froland, W. A.; Lipscomb, J. D.; Munck,
̈
E. J. Am. Chem. Soc. 1993, 115, 6450.
́
(62) Wei, P.-p.; Skulan, A. J.; Mitic, N.; Yang, Y.-S.; Saleh, L.;
Bollinger, J. M.; Solomon, E. I. J. Am. Chem. Soc. 2004, 126, 3777.
(63) Paul, B.; Das, D.; Ellington, B.; Marsh, E. N. J. Am. Chem. Soc.
2013, 135, 5234.
(25) Pikus, J. D.; Studts, J. M.; Achim, C.; Kauffmann, K. E.; Munck,
̈
E.; Steffan, R. J.; McClay, K.; Fox, B. G. Biochemistry 1996, 35, 9106.
(26) Eser, B. E.; Das, D.; Han, J.; Jones, P. R.; Marsh, E. N.
Biochemistry 2011, 50, 10743.
(27) Eser, B. E.; Das, D.; Han, J.; Jones, P. R.; Marsh, E. N.
Biochemistry 2012, 51, 5703.
(28) Bollinger, J. M., Jr.; Diao, Y.; Matthews, M. L.; Xing, G.; Krebs,
C. J. Chem. Soc., Dalton Trans. 2009, 905.
(29) Song, W. J.; Behan, R. K.; Naik, S. G.; Huynh, B. H.; Lippard, S.
J. J. Am. Chem. Soc. 2009, 131, 6074.
(30) Tinberg, C. E.; Song, W. J.; Izzo, V.; Lippard, S. J. Biochemistry
2011, 50, 1788.
(31) Burdi, D.; Willems, J.-P.; Riggs-Gelasco, P.; Antholine, W. E.;
Stubbe, J.; Hoffman, B. M. J. Am. Chem. Soc. 1998, 120, 12910.
(32) Wallar, B. J.; Lipscomb, J. D. Chem. Rev. 1996, 96, 2625.
(33) Kovaleva, E. G.; Neibergall, M. B.; Chakrabarty, S.; Lipscomb, J.
D. Acc. Chem. Res. 2007, 40, 475.
(34) Tinberg, C. E.; Lippard, S. J. Acc. Chem. Res. 2011, 44, 280.
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dx.doi.org/10.1021/cs400484m | ACS Catal. 2013, 3, 2228−2238