Reductive biotransformation of nitroalkenes via nitroso-intermediates to oxazetes catalyzed by xenobiotic reductase A (XenA)

Article abstract of DOI:10.1039/c0ob01216e

A novel reductive biotransformation pathway for β,β-disubstituted nitroalkenes catalyzed by flavoproteins from the Old Yellow Enzyme (OYE) family was elucidated. It was shown to proceed via enzymatic reduction of the nitro-moiety to furnish the corresponding nitroso-alkene, which underwent spontaneous (non-enzymatic) electrocyclization to form highly strained 1,2-oxazete derivatives. At elevated temperatures the latter lost HCN via a retro-[2 + 2]-cycloaddition to form the corresponding ketones. This pathway was particularly dominant using xenobiotic reductase A, while pentaerythritol tetranitrate-reductase predominantly catalyzed the biodegradation via the Nef-pathway.

Full text of DOI:10.1039/c0ob01216e

View Online / Journal Homepage / Table of Contents for this issue
Organic &
Biomolecular
Chemistry
Dynamic Article Links
Cite this: Org. Biomol. Chem., 2011, 9, 3364
www.rsc.org/obc
PAPER
Reductive biotransformation of nitroalkenes via nitroso-intermediates to
oxazetes catalyzed by xenobiotic reductase A (XenA)†
a
a
b
a
c
Katharina Durchschein, Walter M. F. Fabian, Peter Macheroux, Klaus Zangger, Gregor Trimmel and
Kurt Faber*
a
Received 22nd November 2010, Accepted 21st February 2011
DOI: 10.1039/c0ob01216e
A novel reductive biotransformation pathway for b,b-disubstituted nitroalkenes catalyzed by
flavoproteins from the Old Yellow Enzyme (OYE) family was elucidated. It was shown to proceed via
enzymatic reduction of the nitro-moiety to furnish the corresponding nitroso-alkene, which underwent
spontaneous (non-enzymatic) electrocyclization to form highly strained 1,2-oxazete derivatives. At
elevated temperatures the latter lost HCN via a retro-[2 + 2]-cycloaddition to form the corresponding
ketones. This pathway was particularly dominant using xenobiotic reductase A, while pentaerythritol
tetranitrate-reductase predominantly catalyzed the biodegradation via the Nef-pathway.
11,14
Introduction
furnished the corresponding amines in low yields.
Aiming
at the production of a-chiral primary amines in nonracemic
form, we have recently investigated the bioreduction of aliphatic
sec-nitro-compounds using oxygen-stable flavin-reductases from
the OYE family. In contrast to our expectations, OYEs did
not furnish amines, but reductively degraded aliphatic sec-nitro-
compounds to the corresponding carbonyl-compounds via a
cascade of enzymatic (nitro- and oxime)-reductions followed by
The biodegradation of aromatic nitro-compounds predominantly
proceeds through a three-step bioreduction of the nitro-group
via the nitroso- and hydroxylamine-intermediates to furnish the
corresponding aniline or (especially for di- and tri-nitroarenes) via
reduction of the electron-deficient aromatic system to form the so-
1
called ‘Meisenheimer-complex’, which is stabilized via elimination
2
3
of nitrite. This metabolic activity is widespread among bacteria,
spontaneous isomerisation/hydrolysis steps, which overall consti-
4
5
fungi, plants, and yeasts. The enzymes responsible for these
reductive biotransformations are flavoproteins from the Old
Yellow Enzyme (OYE) family. In the context of these studies,
15–18
tutes a biocatalytic equivalent to the Nef-reaction
(Scheme 1).
During the Nef-pathway, the corresponding nitroso- and imine-
6
intermediates could not be directly observed due to their inherent
7
xenobiotic reductases XenA and XenB from Pseudomonas sp.
15,16
instability.
However, the corresponding oxime-intermediate
and pentaerythritol tetranitrate (PETN) reductase from Enter-
could be unambiguously identified using (E/Z)-1-nitro-2-phenyl-
8
obacter cloacae have been aptly denoted as ‘nitroreductases’
15
1
-propene (1) as substrate (Scheme 2).
for their ability to catalyze the reductive biodegradation of
explosives such as PETN, trinitrotoluene (TNT) and nitroglyc-
erin. It was recently reported that nitroreductase NRSal from
Salmonella typhimurium constitutes the first single isolated enzyme
Results and discussion
Much to our surprise, the bioreduction of nitroalkene (E/Z)-1
using xenobiotic reductase A (XenA) furnished the expected Nef-
reaction products (nitroalkane 1a, oxime 1b and aldehyde 1c) only
in minor amounts (9% in total, Scheme 2, Table 1). In contrast,
a new compound was detected by TLC and GC formed in 73%
yield. The latter was sufficiently stable to allow chromatographic
9
catalyzing the reduction of nitrobenzene to aniline accompa-
nied by non-enzymatic (chemical) condensation of nitroso- and
hydroxylamino-intermediates to yield azoxybenzene as a side
9,10
product.
In contrast to the bioreduction of aromatic nitro-
compounds, much less is known about nitroaliphatics. So far,
the bioreduction of aliphatic nitro-compounds appears to be
purification on silica gel. High resolution mass spectrometry
11–13
mainly associated with facultative anaerobic organisms,
which
+
(
ESI) revealed a composition of C
9
H
10NO ([M+H] m/z calcd.
1
13
for 148.0762, found 148.0758). Analysis of DEPT-135, H-, C-
NMR and two-dimensional HSQC and HMBC NMR spectra
revealed oxazete 1e as product. Multiplicities obtained from a
DEPT-135 (CDCl ) showed signals for d 28.08 (CH ), 73.46 (C ),
a
Department of Chemistry, Organic & Bioorganic Chemistry, University of
Graz, Heinrichstrasse 28, A-8010, Graz, Austria. E-mail: Kurt.Faber@Uni-
Graz.at; Fax: +43-316-380-9840; Tel: +43-316-380-5332
b
Institute of Biochemistry, Petersgasse 12, Graz University of Technology,
3
3
q
A-8010, Graz, Austria
1
25.06 (CH), 127.61 (CH), 128.54 (CH), 143.93 (C
q
) and 155.24
c
Institute of Chemistry and Technology of Materials, Stremayrgasse 16, Graz
1
13
(
CH). Using 2D H- C-HSQC and HMBC NMR spectra, the
University of Technology, 8010, Graz, Austria
proton signals between 7.25 ppm and 7.51 ppm and the carbon
signals at 125.1, 127.6, 128.5 and 143.9 ppm could be assigned to
†
1
Electronic supplementary information (ESI) available. See DOI:
0.1039/c0ob01216e
3
364 | Org. Biomol. Chem., 2011, 9, 3364–3369
This journal is © The Royal Society of Chemistry 2011

View Online
Scheme 1 Reductive biotransformation of nitroalkenes of OYEs via the Nef-pathway.
Scheme 2 Reductive biotransformation of nitroalkene (E/Z)-1 using XenA.
Table 1 Bioreduction of (E/Z)-1 using various OYEs in presence of NAD(P)H-recycling systems (Scheme 2)
Conversion [%]
a
b
Entry
Enzyme
XenA
Conditions
Nitroalkane 1a
Oxime 1b
Aldehyde 1c
Oxazete 1e
Acetophenone 1f
1
2
3
4
5
6
7
8
9
NADH
2
3
2
4
6
6
14
24
22
21
38
49
49
48
66
0
5
4
7
15
1
4
4
4
1
12
15
21
26
32
8
2
2
2
2
1
1
1
2
2
3
1
1
2
2
1
0
51
52
23
50
19
25
19
15
12
27
7
8
9
12
7
5
22
21
18
27
11
8
12
16
16
16
5
NADPH
+
NAD /GDH/Glu
NADP /GDH/Glu
+
Mor-R
NADH
+
NAD /GDH/Glu
NerA
OPR3
NADH
NADH
NADPH
+
1
1
1
1
1
1
1
0
1
2
3
4
5
6
NADP /GDH/Glu
PETN
NADH
NADPH
6
3
4
8
+
NAD /GDH/Glu
NADP /GDH/Glu
+
XenB
YhdA
NADH
NADH
2
9
a
XenA, XenB = xenobiotic reductase A and B, NerA = glycerol trinitrate reductase, MorR = morphinone reductase, OPR3 = 12-oxophytodienoic acid
b
reductase, PETN-R = pentaerythritol tetranitrate reductase, YhdA = flavin-dependent quinone reductase. Standard conditions: substrate (E/Z)-1 (1 eq,
1
U), glucose (2 eq, 20 mM); reaction time 24 h, shaking (120 rmp), 30 C.
+
+
0 mM), Tris-buffer (50 mM, pH 7.5), NADH or NADPH (1.5 eq, 15 mM); cofactor recycling: NAD or NADP (100 mM), glucose dehydrogenase (10
◦
a mono-substituted phenyl group, with the quaternary carbon at
signals at 73.5 ppm and 155.2 ppm. A shift of 73.5 ppm
for a quaternary carbon indicates a single bond to oxygen and the
CH group at 155.2 ppm has to be a sp -hybrized carbon bound
1
43.9 ppm being connected to the methyl protons at 1.74 ppm
2
within three bonds according to the HMBC. This spectrum
also showed correlations from the methyl protons to the carbon
to a heteroatom, most likely nitrogen. Therefore, the methyl
This journal is © The Royal Society of Chemistry 2011
Org. Biomol. Chem., 2011, 9, 3364–3369 | 3365

View Online
group can only be bound directly to the quaternary carbon
at 73.5 ppm, with the aromatic carbon at 143.9 ppm and the
signal at 155.3 ppm being attached to the one at 73.5 ppm. Since
no exchangeable proton was found and according to the sum
formula, there is a single bond between oxygen and nitrogen
leading to the proposed oxazete structure 1e. Characteristic
(E)-trans-1d → (E)-cis-1d and (Z)-trans-1d → (Z)-cis-1d occurs
-
1
via a barrier of approximately 10 kcal mol . In contrast to these
rotational barriers, which are slightly larger in aqueous solution,
the barrier for the cyclization is significantly lowered by solvent
-
1
effects by ca. 6–7 kcal mol . Moreover, the four-membered oxazete
1e is preferentially stabilized by solvation, resulting in the greater
-
1
-1
absorptions for C N bonds (1555, 1602 and 1644 cm ) but no
indications for C O or nitro-groups were found in the IR.
In order to rationalize the unexpected formation of this highly
strained heterocycle, the following pathway, which was supported
stability of the product of ca. 2.5 kcal mol compared to the most
stable (E)-trans-nitrosoalkene 1d. According to these calculations,
the more unstable (Z)-cis-isomer of 1d underwent cyclization to
furnish 1e.
19–24
by theoretical calculations
based on relative Gibbs free energies
Although oxazete 1e proved to be reasonably stable at ambient
temperatures, it underwent retro-[2 + 2]-cycloaddition at elevated
temperatures forming acetophenone 1f and HCN, which was
proven by differential scanning calorimetry (DSC) measurements,
in the gas phase and in aqueous solution (SM8 bulk solvation
25,26
model ) was elucidated (Fig. 1): in the first step, the nitro-
moiety of (E/Z)-1 was enzymatically reduced by a hydride
delivered via the flavin cofactor to furnish the corresponding
◦
which showed the highest HCN formation within a range of 120 C
27
◦
nitroso-alkene (E/Z)-1d. The (Z)-isomer of the latter underwent
to 160 C, indicated by a broad sigmoidal shape of the DSC (Fig.
electrocyclization yielding the highly strained four-membered
heterocycle 1e.
S2, ESI†).
28
Since oxazete 1e was racemic as demonstrated by GC using a
chiral stationary phase and CD measurements (Fig. S1, ESI†),
we concluded that the electrocyclization step is of a spontaneous
nature and does not require the involvement of the flavoprotein.
In order to elucidate whether this oxazete formation is of a
general nature or was an isolated phenomenon associated to
XenA, nitroalkene 1 was tested with a broad range of OYEs using
NADH or NADPH as cofactor, as well as in presence of the
corresponding cofactor recycling systems. The data presented in
Table 1 reveal that the type of pathway strongly depends on the
nature of the enzyme:
(i) Using XenA, the oxazete-pathway was clearly dominant
as indicated by the formation of oxazete 1e and its degrada-
tion product acetophenone 1f in up to 77% in total (entries
1
–4). Morphinone reductase showed a similar, but somewhat
less pronounced trend (entries 5 and 6, 1e + 1f up to 33%).
With both enzymes, oxime/aldehyde formation yielding 1b and
1
c (as an indicator of the Nef-pathway) constituted only a
Fig. 1 Energy-diagram calculated for the cyclization of nitroso-alkene 1d
to oxazete 1e in H O (TS = transition state).
side-activity.
2
(
ii) Glycerol trinitrate reductase NerA and 12-oxophytodienoic
acid reductase OPR3 showed both remarkable oxazete formation
(entries 7 and 10) going hand in hand with pronounced ene-
reductase activity as indicated by the C C-bond reduction of
nitroalkene 1 forming nitroalkane 1a in up to 21% (entry 10).
Oxime formation (1b) was somewhat suppressed.
An alternative mechanism for oxazete formation could be
envisaged via conjugate addition of H O across the C C bond of
nitrosoalkene 1d, thereby forming a benzylic tert-alcohol species
2
which would undergo subsequent cyclisation with concomitant
formation of water. Since the latter mechanism would involve
(iii) Pentaerythritol tetranitrate reductase showed only weak
oxazete formation (1e up to 12%), but dominant alkene reduction
(1a up to 49%) and remarkably high oxime formation (1b up to
32%, entry 14).
(iv) XenB predominantly proved to behave as an ene-reductase
(1a 66%, entry 15), and the quinone reductase YhdA was almost
inactive (entry 16).
Overall, there was no dramatic impact on the product distribu-
tion depending on the type of cofactor (NADH or NADPH) or
on the respective cofactor recycling system employed. All blank
experiments in the absence of protein and/or cofactor showed a
conversion below the level of detectability (<3%).
In order to map the substrate tolerance for oxazete formation,
a range of nitroalkenes 2–6 which are structurally related to
substrate 1 was studied (Scheme 3).
-
nucleophilic attack of [OH ] at the (sterically demanding) C
b
of the nitrosoalkene, it was not considered as most plausible.
,2-Oxazetes are known to occur as unstable intermediates in
1
29
aza-Claisen rearrangements and they have been postulated as
intermediates during the formation of carbonyl-compounds and
nitriles from vinyl-radicals and NO. In addition, they are known
to occur as reactive intermediates in thermal and photochemical
reactions of a,b-unsaturated nitro-compounds. Their stability
can be significantly enhanced by sterically demanding substituents,
which allow their synthesis via intramolecular cyclization of a,b-
30
31
32
unsaturated nitroso-compounds or by MCPBA-oxidation of
oximes.
33
According to our calculations, the (E)-configuration of nitroso-
alkene 1d is more stable than the (Z)-isomer, especially for
the s-cis-conformation of the nitroso-group (DG (E/Z)-cis 1d =
The product distribution derived from the bioreduction of ni-
troalkenes 2–6 shows that oxazete formation is strongly dependent
-
1
1
.8 kcal mol ) (Fig. 1). Interconversion between the two rotamers,
3
366 | Org. Biomol. Chem., 2011, 9, 3364–3369
This journal is © The Royal Society of Chemistry 2011

View Online
Scheme 3 Bioreduction of substituted nitroalkenes 2–6 via the Nef-pathway versus oxazete formation.
on the substrate structure (Table 2). 1-Nitrocyclohexene (2) was
mainly reduced at the C C-bond yielding nitrocyclohexane (2a),
the Nef-product cyclohexanone (2c) was formed in minor amounts
with XenA (27% 5e + 5f), while C C-bond reduction remained
low (OPR3 11% 5a). Finally, a close analog of the original test
substrate 1 bearing a 2-naphthyl group (6) was solely converted
via the oxazete pathway, again XenA being best (41% 6e+6f),
no C C-bond reduction was detected. Both oxazete derivatives
5e + 5f, 6e + 6f were characterised by GC-MS using compound
1e as a lead. Overall, the corresponding oximes 2b–6b were not
detectable. From these results it can be concluded, that oxazete
formation only takes place with b,b-disubstituted nitroalkenes (1,
5, 6) which – in line with the absence of an enantiomeric excess of 1e
– supports the hypothesis that it is of a non-enzymatic nature and
proceeds via spontaneous electrocyclization of the corresponding
nitrosoalkenes.
(
up to 12%) using OPR3. The corresponding oxazete pathway
yielding 2e and/or 2f was not detectable. The open-chain substrate
gave similar results, i.e. mainly nitroalkane 3a (XenB 96% 3a)
3
and Nef-degradation product 3c (OPR3 15% 3c) were formed.
Both of the latter substrates share two a,b-substituents at the
C
C-bond. In contrast, all enzymes solely gave the C C-bond
reduction product 4a with the b-monosubstituted nitroalkene 4
in up to 67% conversion (XenB 67% 4a). However, two b,b-
substituents in substrate 5 opened the oxazete pathway, forming
5e and its degradation product 5f, this was most pronounced
Table 2 Product distribution during bioreduction of nitroalkenes 2–6
a
Substrate
Enzyme
Products [%]
2
XenA
XenB
NerA
OPR3
XenA
XenB
NerA
OPR3
XenA
XenB
NerA
OPR3
XenA
XenB
NerA
OPR3
XenA
XenB
NerA
OPR3
2a
3a
4a
5a
6a
97
99
90
85
7
96
88
84
16
67
<1
13
5
2b
3b
4b
5b
6b
<1
<1
<1
<1
<1
2c
3c
4c
5c
6c
3
<1
9
12
<1
3
11
15
<1
2e
3e
4e
5e
6e
<1
<1
<1
2f
3f
4f
5f
6f
<1
<1
<1
3
4
5
6
<1
<1
18
n.d.
3
9
n.d.
3
n.d.
8
11
<1
3
8
13
<1
<1
<1
28
<1
15
31
a
◦
Standard conditions: substrate 2–6 (1 eq, 10 mM), Tris-buffer (50 mM, pH 7.5), NADH (1.5 eq, 15 mM), reaction time 24 h, shaking (120 rmp), 30 C;
organic cosolvents were used to solubilize the following substrates: 20% v:v DMSO substrate 3; tBuOMe (10% v:v) substrate 4; tBuOMe (15% v:v)
substrate 6; XenA, XenB = xenobiotic reductase A and B, NerA = glycerol trinitrate reductase, OPR3 = 12-oxophytodienoic acid reductase; n.d. = not
determined.
This journal is © The Royal Society of Chemistry 2011
Org. Biomol. Chem., 2011, 9, 3364–3369 | 3367

View Online
Conclusions
and 120 rpm and after 24 h products were extracted with EtOAc
(
2 ¥ 0.8 ml). The combined organic phases were dried (Na
2
SO )
4
The reductive biotransformation of b,b-disubstituted nitroalkenes
catalyzed by flavoproteins from the OYE family was shown
to proceed not only via the recently elucidated Nef-reaction,
but also via enzymatic reduction of the nitro-moiety instead
of C C-reduction to furnish the correponding nitroso-alkene,
which spontaneously underwent electrocyclization to form highly
strained 1,2-oxazete derivatives in racemic form. The latter were
sufficiently stable to allow their structural characterization; at ele-
vated temperatures they lost HCN via a retro-[2 + 2]-cycloaddition
to form the corresponding ketones. The type of bioreduction
pathway, i.e. Nef-pathway versus oxazete formation, depended not
only on the substitutional pattern of the nitroalkene, but also on
the type of flavoprotein: whereas PETN-reductase predominantly
catalyzed the Nef-pathway, xenobiotic reductase A was strong in
oxazete formation.
and the resulting samples were analyzed on achiral GC or GC-
MS after TLC control. For substrates of low solubility in Tris-
HCl buffer cosolvents were used, 4: 10% v:v t-butyl-methyl-ether
(
TBME); 3: 20% v:v DMSO; 6: 15% v:v TBME.
Synthesis and isolation of oxazetes
-Methyl-4-phenyl-4H-1,2-oxazete (1e). The general proce-
4
dure was up-scaled 60-fold using substrate (E/Z)-1. After the
biotransformation all samples were pooled and extracted twice
with 50 ml ethyl acetate. Removal of the solvent under reduced
pressure gave an oily residue which was purified via flash chro-
matography on silica (ethyl acetate : petroleum ether 1 : 3) to yield
2
5 mg of pure 4-methyl-4-phenyl-4H-1,2-oxazete 1e. TLC: R
f
0.30
(silica gel, ethyl acetate : petroleum ether 1 : 3, molybdate blue);
GC-MS (EI): m/z 32, 43, 53, 63, 77, 91, 105, 122, 132, 148;
+
HRMS (1.250.000 resolution, ESI) (M + H) = m/z calcd. for
Experimental section
-
1
C
1
9
H
10NO 148.0762, found 148.0758; IR (cm ): 1374, 1447, 1494,
1
555, 1602, 1644, 2930, 2982. H-NMR (300 MHz, CDCl
3
): d 1.74
Cloning, expression and purification of XenA, XenB and NerA
(
7
3H, s), 7.25–7.35 (1H, t, J = 7.3 Hz), 7.35–7.44 (2H, t, J = 7.3 Hz),
.46–7.51 (2H, d, J = 7.2 Hz), 7.68 (1H, br); C-NMR (75 MHz,
(
GTN)
1
3
Xenobiotic reductase A from Pseudomonas putida (XenA), xeno-
biotic reductase B from Pseudomonas putida JLR11 (XenB),
and glycerol trinitrate reductase from Agrobacterium radiobacter
CDCl
3
): d 28.1, 73.5, 125.1, 127.6, 128.5, 143.9, 155.2.
4
-Ethyl-4-phenyl-4H-1,2-oxazete (5e). TLC: R 0.70 (silica,
f
ethyl acetate : petroleum ether 1 : 3, molybdate blue); GC-MS (EI):
m/z 32, 44, 51, 72, 77, 105, 134, 149, (161); 5f m/z 32, 39, 44, 51,
(
NerA) were cloned, expressed and purified as follows. The DNA
sequences were retrieved from the NCBI Genebank (accession files
Q9R9V9 for XenA, Q9RPM1 for XenB and O31246 for NerA)
and synthesized with a C-terminal hexahistidine tag. The genes
were cloned into pET21a and transformed into the expression
strain E. coli BL 21 (DE3). Heterologous expression of the genes
in E. coli BL21 (DE3) was performed as follows: 100 ml starter
LBAmp cultures were inoculated with aliquots from a frozen stock
55, 65, 77, 91, 104, 115, 119, 120. The data (GC, GC-MS, TLC)
corresponded to 1e.
4
-Methyl-4-(naphthalen-2-yl)-4H-1,2-oxazete (6e). GC-MS
EI): m/z 32, 43, 57, 77, 127, 155, 170, 180 (197); 6f m/z 32, 43, 56,
3, 77, 101, 127, 155, 170. The data (GC, GC-MS) corresponded
(
6
to 1e.
◦
culture and grown overnight at 37 C. Each starter culture was
used to inoculate a 700 ml culture, which was grown to an OD600
of 0.6–0.8. Gene expression was induced with 0.2 mM IPTG (final
concentration) and protein expression was performed for 4 h at
Acknowledgements
This project was performed within the DK ‘Molecular Enzymol-
ogy’ and financial support from the Austrian Science Fund FWF
◦
◦
3
7
C. After centrifugation, cell pellets were frozen at -80
or used directly for the protein purification. The pellets were
resuspended in lysis buffer (50 mM NaH PO , 300 mM NaCl
C
(Vienna, project W9) is gratefully acknowledged. Cordial thanks
2
4
go to Harald Koefeler (Center for Medical Research ZMF, CF
Mass Spectrometry - Lipidomics, Graz) for HRMS-measurements
and to Neil C. Bruce (York) for providing plasmids of PETN- and
morphinone reductase.
and 20 mM imidazole) and sonicated on ice (t = 5 min, pulse 1 s,
pause 2 s, amplitude 40%) with addition of FMN (ca. 3 mg). After
◦
centrifugation (30 min, 18 000 rpm, 4 C), the supernatant was
2
+
subjected to protein purification on a Ni -NTA column according
34
to standard protocol.
References
General procedure for the bioreduction of substrates
1 J. C. Spain, Annu. Rev. Microbiol., 1995, 49, 523–555; F. D. Marvin-
Sikkema and J. A. M. de Bont, Appl. Microbiol. Biotechnol., 1994, 42,
4
99–507.
An aliquot of enzyme [OPR3 isoenzyme of 12-oxophytodienoate
reductase from Lycopersicon esculentum, PETN-reductase from
E. cloacae PB2 (PETN-Red), morphinone reductase (Mor-Red),
flavin-oxidase (YhdA), xenobiotic reductases (XenA, XenB) and
2
C. E. French, S. J. Rosser, G. J. Davies, S. Nicklin and N. C. Bruce,
Nat. Biotechnol., 1999, 17, 491–494; J. W. Pak, K. L. Knoke, D. R.
Noguera, B. G. Fox and G. H. Chambliss, Appl. Environ. Microbiol.,
2000, 66, 4742–4750; M. M. Gonz a´ lez-P e´ rez, P. van Dillewijn, R.-M.
Wittich and J. L. Ramos, Environ. Microbiol., 2007, 9, 1535–1540.
Z. C. Symons and N. C. Bruce, Nat. Prod. Rep., 2006, 23, 845–850.
J. L. Ramos, M. Mar Gonz a´ l e´ z-Perez, A. Caballero and P. van
Dillewijn, Curr. Opin. Biotechnol., 2005, 16, 275–281.
-
1
glycerol trinitrate reductase (NerA), protein content ~100 mg ml ],
was added to a Tris-HCl buffer solution (0.8 ml, 50 mM, pH 7.5)
containing the substrate (10 mM) and the cofactor NADH
or NADPH (15 mM). Alternatively, the oxidized form of the
3
4
5 C. L. Davey, L. W. Powell, N. J. Turner and A. Wells, Tetrahedron
Lett., 1994, 35, 7867–7870; J. A. Blackie, N. J. Turner and A. S.
Wells, Tetrahedron Lett., 1997, 38, 3043–3046; A. Navarro-Ocana, L. F.
Olgu ´ı n, H. Luna, M. Jim e´ nez-Estrada and E. B a´ rzana, J. Chem. Soc.,
Perkin Trans. 1, 2001, 2754–2756.
+
+
cofactor NAD or NADP (100 mM) was used in combination
with a recycling enzyme (glucose dehydrogenase, 10 U) and the
cosubstrate (glucose, 20 mM). The mixture was shaken at 30 C
◦
3
368 | Org. Biomol. Chem., 2011, 9, 3364–3369
This journal is © The Royal Society of Chemistry 2011

View Online
6
7
R. E. Williams, D. A. Rathbone, N. S. Scrutton and N. C. Bruce, Appl.
Environ. Microbiol., 2004, 70, 3566–3574.
study, see: J. U. Nef, Justus Liebigs Ann. Chem., 1894, 280, 263–342;
H. W. Pinnick, Org. React., 1990, 38, 655–792; R. Ballini and M. Petrini,
Tetrahedron, 2004, 60, 1017–1047; L. Petrus, M. Petrusov a´ , D.-P. Pham-
Huu, E. Lattov a´ , B. Pribulov a´ and J. Turjan, Monatsh. Chem., 2002,
133, 383–392.
D. S. Blehert, B. G. Fox and G. H. Chambliss, J. Bacteriol., 1999, 181,
6
254–6263; J. J. Griese, R. P. Jakob, S. Schwarzinger and H. Dobbek,
J. Mol. Biol., 2006, 361, 140–152.
8
H. S. Toogood, A. Fryszkowska, V. Hare, K. Fisher, A. Roujeinikova,
D. Leys, J. M. Gardiner, G. M. Stephens and N. S. Scrutton, Adv.
Synth. Catal., 2008, 350, 2789–2803; H. Nivinskas, J. Sarlauskas, Z.
Anusevicius, H. S. Toogood, N. S. Scrutton and N. C e´ nas, FEBS J.,
19 S. Arulmozhiraja and P. Kolandaivel, J. Mol. Struct., 1998, 429, 165–
173.
20 S. Arulmozhiraja and P. Kolandaivel, Mol. Phys., 1997, 90, 55–62.
21 O. Kikuchi, Tetrahedron Lett., 1981, 22, 859–862.
22 P. Gerbaux, N. Dechamps, R. Flammang, P. C. Nam, M. T. Nguyen,
F. Djazi, F. Berruyer and G. Bouchoux, J. Phys. Chem. A, 2008, 112,
5418–5428.
2
008, 275, 6192–6203.
Y. Yanto, M. Hall and A. S. Bommarius, Org. Biomol. Chem., 2010, 8,
826–1832.
9
1
1
0 V. Hoeller, D. Wegricht, I. Yuranov, L. Kiwi-Minsker and A. Renken,
Chem. Eng. Technol., 2000, 23, 251–255; R.-M. Wittich, A. Haidour, P.
van Dillewijn and J.-L. Ramos, Environ. Sci. Technol., 2008, 42, 734–
23 T. Sakaizumi, A. Usami, H. Satoh, O. Ohashi and I. Yamaguchi, J. Mol.
Spectrosc., 1994, 164, 536–549.
24 T. Sakaizumi, M. Nishikawa, A. Usami, H. Satoh and O. Ohashi,
J. Anal. Appl. Pyrolysis, 1995, 34, 219–227.
25 C. J. Cramer and D. G. Truhlar, Acc. Chem. Res., 2008, 41, 760–768.
26 A. V. Marenich, R. M. Olson, C. P. Kelly, C. J. Cramer and D. G.
Truhlar, J. Chem. Theory Comput., 2007, 3, 2011–2033.
27 Nitrosoalkenes are very unstable and have been identified by microwave
spectroscopy, see ref. 23.
28 For a related nitroso-Diels–Alder cyclization, see: B. Yang, P. A. Miller,
U. M o¨ llmann and M. J. Miller, Org. Lett., 2009, 11, 2828–2831; for a
related peri-cyclization, see: C. Dolka, K. van Hecke, L. van Meervelt,
P. G. Tsoungas, E. V. van der Eycken and G. Varvounis, Org. Lett.,
2009, 11, 2964–2967; for a related cyclization of nitroalkenes yielding
1,2-oxazet-N-oxides, see: A. Berndt, Angew. Chem. Int. Ed., 1968, 7,
637–638.
29 N. Coskun and N. Arikan, Turk. J. Chem., 2003, 27, 15–20.
30 A. G. Sherwood and H. E. Gunning, J. Am. Chem. Soc., 1963, 85,
3506–3508.
31 T. H. Kinstle and J. G. Stam, J. Org. Chem., 1970, 35, 1771–1774; J. T.
Pinhey and E. Rizzardo, Tetrahedron Lett., 1973, 41, 4057–4058; J. M.
Surzur, C. Dupuy, M. P. Bertrand and R. Nouguier, J. Org. Chem.,
1972, 37, 2782–2784.
32 K. Wieser and A. Berndt, Angew. Chem. Int. Ed., 1975, 14, 69–70.
33 H. G. Corkins, L. Storace and E. R. Osgood, Tetrahedron Lett., 1980,
21, 2025–2028.
7
39.
1
1
1 L. Angermaier and H. Simon, Hoppe-Seyler s´ Zeitschrift f u¨ r physiolo-
gische Chemie, 1983, 364, 961–975.
2 The bioreduction of oximes by baker’s yeast was reported to yield
amines in modest ee and yield, see: D. E. Gibbs and D. Barnes,
Tetrahedron Lett., 1990, 31, 5555–5558.
1
1
3 H. Braun, F. P. Schmidtchen, A. Schneider and H. Simon, Tetrahedron,
1
991, 47, 3329–3334.
4 A. Mori, I. Ishiyama, H. Akita, K. Suzuki, T. Mitsuoka and T. Oishi,
Xenobiotica, 1990, 20, 629–634; A. Mori, I. Ishiyama, H. Akita, K.
Suzuki, T. Mitsuoka and T. Oishi, Chem. Pharm. Bull., 1990, 38, 3449–
3
451.
1
1
1
1
5 K. Durchschein, B. Ferreira-Silva, S. Wallner, P. Macheroux, W.
Kroutil, S. M. Glueck and K. Faber, Green Chem., 2010, 12, 616–
6
19.
6 D. H. Aue and D. Thomas, J. Org. Chem., 1974, 39, 3855–3862; M.
Witanowski, L. Stefaniak, H. Januszewski, S. Szymanski and G. A.
Webb, Tetrahedron, 1973, 29, 2833–2836.
7 S. Trivic, V. Leskovac, D. Pericin and G. W. Winston, Biotechnol. Lett.,
2
002, 24, 807–812; B. Ferreira-Silva, I. Lavandera, A. Kern, K. Faber
and W. Kroutil, Tetrahedron, 2010, 66, 3410–3414.
8 Although the Nef-reaction was originally described as the (redox-
neutral) hydrolysis of prim- and sec-nitro-alkenes to yield the corre-
sponding carbonyl-compound and N
2
O, oxidative variants are known,
34 K. Kitzing, T. B. Fitzpatrick, C. Wilken, J. Sawa, G. P. Bourenkov, P.
Macheroux and T. Clausen, J. Biol. Chem., 2005, 280, 27904–27913.
which nicely parallel the enzymatic reductive equivalent shown in this
This journal is © The Royal Society of Chemistry 2011
Org. Biomol. Chem., 2011, 9, 3364–3369 | 3369