Bacterial dioxygenase- and monooxygenase-catalysed sulfoxidation of benzo[b]thiophenes

Article abstract of DOI:10.1039/c1ob06678a

Asymmetric heteroatom oxidation of benzo[b]thiophenes to yield the corresponding sulfoxides was catalysed by toluene dioxygenase (TDO), naphthalene dioxygenase (NDO) and styrene monooxygenase (SMO) enzymes present in P. putida mutant and E. coli recombinant whole cells. TDO-catalysed oxidation yielded the relatively unstable benzo[b]thiophene sulfoxide; its dimerization, followed by dehydrogenation, resulted in the isolation of stable tetracyclic sulfoxides as minor products with cis-dihydrodiols being the dominant metabolites. SMO mainly catalysed the formation of enantioenriched benzo[b]thiophene sulfoxide and 2-methyl benzo[b]thiophene sulfoxides which racemized at ambient temperature. The barriers to pyramidal sulfur inversion of 2- and 3-methyl benzo[b]thiophene sulfoxide metabolites, obtained using TDO and NDO as biocatalysts, were found to be ca.: 25-27 kcal mol^-1. The absolute configurations of the benzo[b]thiophene sulfoxides were determined by ECD spectroscopy, X-ray crystallography and stereochemical correlation. A site-directed mutant E. coli strain containing an engineered form of NDO, was found to change the regioselectivity toward preferential oxidation of the thiophene ring rather than the benzene ring.

Full text of DOI:10.1039/c1ob06678a

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PAPER
Bacterial dioxygenase- and monooxygenase-catalysed sulfoxidation of
benzo[b]thiophenes†
Derek R. Boyd,*a Narain D. Sharma,a Brian McMurray,a Simon A. Haughey,a Christopher C. R. Allen,b
John T. G. Hamilton,b,c W. Colin McRoberts,c Rory A. More O’Ferrall,d Jasmina Nikodinovic-Runic,e
Lydie A. Coulombele and Kevin E. O’Connor*e
Received 3rd October 2011, Accepted 26th October 2011
DOI: 10.1039/c1ob06678a
Asymmetric heteroatom oxidation of benzo[b]thiophenes to yield the corresponding sulfoxides was
catalysed by toluene dioxygenase (TDO), naphthalene dioxygenase (NDO) and styrene monooxygenase
(SMO) enzymes present in P. putida mutant and E. coli recombinant whole cells. TDO-catalysed
oxidation yielded the relatively unstable benzo[b]thiophene sulfoxide; its dimerization, followed by
dehydrogenation, resulted in the isolation of stable tetracyclic sulfoxides as minor products with
cis-dihydrodiols being the dominant metabolites. SMO mainly catalysed the formation of
enantioenriched benzo[b]thiophene sulfoxide and 2-methyl benzo[b]thiophene sulfoxides which
racemized at ambient temperature. The barriers to pyramidal sulfur inversion of 2- and 3-methyl
benzo[b]thiophene sulfoxide metabolites, obtained using TDO and NDO as biocatalysts, were found to
be ca.: 25–27 kcal mol-1. The absolute configurations of the benzo[b]thiophene sulfoxides were
determined by ECD spectroscopy, X-ray crystallography and stereochemical correlation. A
site-directed mutant E. coli strain containing an engineered form of NDO, was found to change the
regioselectivity toward preferential oxidation of the thiophene ring rather than the benzene ring.
Introduction
Monooxygenase- and dioxygenase-catalysed oxidations of the
isosteric substrates indene 1,1a–c,2a–f benzo[b]thiophene (B[b]T) 2,3a–l
and substituted derivatives, have previously been studied in these
and other laboratories. The stereoselective oxidation product from
indene 1, was the (1S,2R)-epoxide 3 (up to 97% ee, Scheme 1) when
using SMO from a Pseudomonas putida strain (CA-3) and derived
E. coli recombinant strains.1a–c
Earlier studies 2a–f have also shown that biocatalytic asymmetric
cis-dihydroxylation of the alkene bond in substrate 1, using
different P. putida strains, can give the corresponding (1S,2R)-
dihydrodiol 4cis with variable degrees of stereoselectivity (20–
Scheme 1 Isolated and potential products resulting from enzyme–
>98% ee) depending on the type of dioxygenase used. Rhodococ-
catalysed oxidation of the five-membered rings in indene
1 and
benzo[b]thiophene 2.
aSchool of Chemistry and Chemical Engineering, Queen’s University Belfast,
Belfast, UK, BT9 5AG. E-mail: dr.boyd@qub.ac.uk; Tel: +44 (0) 28
90975419
cus strains have also been reported to yield epoxide 3, trans-
indandiol 4trans and cis-indandiol 4cis among bioproducts obtained
via oxygenase-catalysed oxidation of indene 1.2e,2f In addition,
asymmetric benzylic hydroxylation of indene 1 to yield (1R)-
indenol 5 of variable enantiopurity (up to >98% ee) was also
observed using different P. putida dioxygenases (Scheme 1).2a–d
Thus epoxide 3, cis-diol 4cis, and benzylic alcohol 5, can each be
formed by oxygenase-catalysed asymmetric oxidation of indene 1
with high ee values (≥97%) according to the type of oxygenase
enzyme selected.
bSchool of Biological Sciences, Queen’s University Belfast, Belfast, UK, BT9
5AG
cAgri-food and Biosiences Institute for Northern Ireland, Belfast, UK,
BT95PX
dSchool of Chemistry and Chemical Biology, University College Dublin,
Belfield, Dublin, 4, Ireland
eSchool of Biomolecular and Biomedical Sciences and Centre for Synthesis
and Chemical Biology, University College Dublin, Belfield, Dublin, 4,
Ireland. E-mail: kevin.oconnor@ucd.ie
† Electronic supplementary information (ESI) available. See DOI:
10.1039/c1ob06678a
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The prevalence of B[b]T 2 and derivatives in the environment,
e.g. in crude oil, coal and their combustion products, had prompted
earlier studies of their bacterial biotransformation.3a–l However,
with the exception of our preliminary reports from this study,
obtained with B[b]T substrates and TDO from Pseudomonas
putida UV4,3e,i,j most of the earlier studies3a–d,3f–h on prokaryotic
metabolism of B[b]T 2 and methyl substituted benzo[b]thiophene
(MB[b]T) derivatives were conducted on a relatively small scale.
After extraction (EtOAc) of aqueous culture medium from bio-
transformations and concentration of the extract, the bioproducts
were generally detected using GC-MS or GC-FTIR analysis of
the crude mixture.3a–d,3f–h The GC analytical method resulted in
partial decomposition of some metabolites, e.g. B[b]T sulfoxides
(B[b]T-1-oxides), within the injection port,3d and precluded the full
structural and stereochemical characterization (ee and absolute
configuration) of individual metabolites from the mixture of
bioproducts.
Scheme 2 Oxygenase-catalysed oxidative pathways of monocyclic thio-
phenes (R and R¢ = H, alkyl or aryl groups and halogen atoms).
The major objectives of this more comprehensive study follow-
ing from our earlier communications3e,i,j are: (i) to fully charac-
terize our previously reported and newly isolated TDO-catalysed
sulfoxidation products from B[b]T 2 and MB[b]T substrates, (ii) to
compare the range and stereochemistry of bioproducts obtained
using these substrates with both dioxygenase (TDO, NDO) and
monooxygenase (SMO) enzymes and (iii) to determine the ee
values and absolute configurations of chiral sulfoxide metabolites
from three B[b]T substrates and to study their thermally-induced
racemization.
sulfoxides C followed by dimerization to yield disulfoxides F and,
after chemical deoxygenation, monosulfoxide adducts G.7 The elu-
sive monosulfoxides C were also found to form sulfoxide–thioether
adducts H with thiols. The isolation of trans-hydroxythioether
metabolites J, has been postulated to result from the CYP-450
monooxygenase-catalysed formation of unstable thiophene arene
oxide metabolites I followed by nucleophilic attack of thiols (e.g.
glutathione).7 Due to their instability, arene oxide derivatives of
monocyclic thiophenes have not yet been isolated or detected.4
The metabolic steps shown in Scheme 2 could, in principle, be
applicable to mono- and bi-cyclic thiophenes. Preliminary commu-
nications of this study showed that the major biotransformation
pathway of benzo[b]thiophene 2 using P. putida UV4, resulted in
TDO-catalysed cis-dihydroxylation of the thiophene ring to yield
cis-diol 7cis, which equilibrated with trans-diol 7trans (Scheme 3).3e,i,j
The initial biotransformation of B[b]T 2 was carried out over an
extended period (24 h) followed by column chromatography of
the crude mixture of bioproducts; it yielded mainly dihydrodiols
7cis/7trans and phenols 14 and 15 as minor bioproducts (Scheme
3) with little evidence of sulfoxidation products. It was assumed
that phenol 14 and 15 were derived from dehydration of the
carbocyclic cis-dihydrodiol 13 which was much less stable than the
heterocyclic dihydrodiols 7cis/7trans. When the biotransformation
was repeated on a larger scale using a shorter biotransformation
period (7 h), the heterocyclic ring dihydrodiols 7cis/7trans were
again the dominant metabolites along with phenols 14 and 15.
However, additional products, including the less stable carbocyclic
cis-dihydrodiol 13 and the tetracyclic sulfoxidation products, 10–
12 were also isolated.
Results and discussion
The biocatalytic oxidation products 3–5, derived earlier from
indene 1, were of sufficient stability to be isolated and fully char-
acterized structurally and stereochemically.1a–c,2a–f This contrasted
with our preliminary observations from the oxygenase-catalysed
oxidation of the isosteric substrate 2.3e,3i,3j Thus it was found that:
(i) the initially formed minor sulfoxide metabolite 8, when obtained
using TDO, was too unstable to be detected and isolated, (ii)
the major cis-diol metabolite 7 formed using TDO was prone to
spontaneous equilibration with the trans isomer in CDCl3,3e (iii)
the potential arene oxide metabolite 6, using SMO enzyme (cf.
the indene epoxide metabolite 3) was not detected. Arene oxide 6
has only been characterized by 1H-NMR analysis (THF-d8) of the
chemically synthesised crude sample which rapidly decomposed
to an isomeric ketone during attempted purification.4,5
TDO-catalysed oxidation (TO) of monocyclic thiophenes A,
using P. putida UV4 whole cells (Scheme 2), has recently been
found to occur at both the sulfur atom (sulfoxidation) to
yield the unstable monosulfoxides C and at the 2,3-bond (cis-
dihydroxylation) to give cis-diols B as initial metabolites.6 Under
acidic conditions the equilibrating mixture of cis-diols B and trans-
diols D was also found to dehydrate yielding hydroxythiophenes
E. Due to the spontaneous dimerization of the transient mono-
cyclic thiophene monosulfoxides C, and their disproportionation
products, thiophenes A and sulfones K, and disulfoxide adducts
F were isolated as metabolites. A sulfoxide reductase-catalysed
deoxygenation of stable disulfoxides F also gave the corresponding
monosulfoxides G.5
A comparison of the relative yields of bioproducts, obtained
from biotransformation of B[b]T, 2-MB[b]T and 3-MB[b]T
with several dioxygenases (TDO, NDO1, NDO2, NDO3) and
a monooxygenase enzyme (SMO), that demonstrated a strong
preference for the sulfoxidation pathway is shown in Table 1. As
this study is primarily focused on oxygenase-catalysed heteroatom
oxidation and the structure, stereochemistry and reactivity of
the resulting sulfoxides, only the relative proportions of other
metabolites have been shown (Table 1).
CYP-450 monooxygenase-catalysed sulfoxidation (MO) of
monocyclic thiophenes A was again found to yield the unstable
A discussion of the full structural characterization, enan-
tiopurity, absolute configuration and synthetic applications of
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Scheme 3 Dioxygenase-catalysed oxidation of B[b]T 2.
Table 1 Relative yields of bioproducts obtained using substrates 2, 17 and 22
Bioproducts detected (Relative % yield)
Enz.
B[b]T
B[b]TSOa
BC/TDb
BCDc
Other bioproducts
TDOd
NDO1
NDO2
NDO3
SMOh
TDOd
NDO1
NDO2
NDO3
SMOh
TDOd
NDO1
NDO2
NDO3
SMOh
2
7 (63)
7 (85)
13 (10)
10 (5), 11 (5), 12 (2), 14 (8), 15 (7)
e
f
2
11 (15)
2
10 (50), 11 (50)
11 (15)
g
2
7 (85)
2
8 (>95)
18 (7)
18 (10)
18 (90)
18 (40)
18 (100)
17
17
17
17
17
22
22
22
22
22
19 (10)
20 (83)
20 (90)
e
f
21 (10)
g
19 (60)
24 (10)
24 (100)
e
f
23 (90)
23 (39)
26 (33), 27 (28)
g
23 (100)
i
a B[b]TSO (B[b]T sulfoxides). b BC/TD (B[b]T) heterocyclic ring cis or trans diols). c BCD (B[b]T carbocyclic cis-diols). d P. putida UV4. e P. putida 9816/11.
f P. putida NCIMB 8859. g E. coli F352V. h E.coli BL21 (DE3)-pRSET-styAB. i No sulfoxide formed.
cis-dihydrodiols derived from both the carbocyclic and hetero-
cyclic ring of benzo[b]thiophenes isolated during this work, and
from the corresponding benzo[b]furan (B[b]F) substrates, will be
presented in a later publication.7
The biotransformation of B[b]T 2 in P. putida UV4 resulted
in a strong preference for TDO-catalysed oxidation to occur
within the electron-rich thiophene ring of the substrate with cis-
hydroxylation and sulfoxidation accounting for ca.: 75% of the
identified metabolites (7cis 10cis, 10trans, 11, 12).
The equilibrating cis/trans diol mixture 7cis/ 7trans was isolated
as the major product (63% relative yield). However, the stable
tetracyclic derivatives 10–12, resulting from dimerization of the
initial transient B[b]T sulfoxide (8 → 9), accounted for a minor, but
significant, proportion (12%) of the isolated metabolites (Table 1).
Although the initially formed sulfoxide 8 was not detected during
the earlier biotransformations of B[b]T 2, recent studies using
LC-TOFMS analysis showed it to be an identifiable metabolite
formed during the early stages of the biotransformation. However
it could not be isolated from among the mixture of metabolites
(8–15, Scheme 3).
An early attempt to synthesise an authentic sample of sulfoxide
8 by MCPBA oxidation of B[b]T 2 gave mainly the B[b]T sulfone
(B[b]T-1,1-dioxide) 16 (60% isolated yield) along with small
amounts (<1%) of each of the tetracyclic sulfoxides 10cis, 10trans
and 11 and benzo[b]naphtho[2,1-d]thiophene 12. Formation of the
last product suggests that a degree of spontaneous deoxygenation
had occurred, possibly via disproportionation. A later attempt to
obtain the unstable sulfoxide 8, under milder conditions, using
dimethyldioxirane as oxidant in acetone-d6 solvent at ambient
1
temperature, provided direct H-NMR evidence of its formation
in solution.
The biosynthetic sequence involved in the formation of the tetra-
cyclic metabolites 10–12 from B[b]T 2 has not yet been rigorously
established. However, the earlier formation and dimerization of
a wide range of monocyclic thiophene sulfoxides, using P. putida
UV4, to give a series of stable racemic disulfoxides (compound
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F, R = H and R¢ = H; R¢ = H and R = Me, Et, Cl, Br, Ph,
Scheme 2), whose structure and stereochemistry was confirmed by
X-ray crystallography and NMR spectroscopy, provide a credible
precedent.6
Although disulfoxide 9 was not isolated, a fully deoxygenated
derivative has recently been detected by GC-MS analysis following
a whole cell biotransformation of B[b]T 2 using a Sphingomonas
strain (XLDN2-5).3l Our inability to detect disulfoxide 9 as a
metabolite of B[b]T 2, or as a product from chemical oxidation,
may be due to the spontaneous formation of the stereoisomeric
substrates.10a,10c However, the wild-type strain cannot accumulate
cis-dihydrodiols due to the presence of a cis-diol dehydrogenase
which catalyses their further biotransformation into catechols. In
contrast with TDO, NDO can only catalyse a very limited range
of arene cis-dihydroxylations e.g. mainly biaryls and polycyclic
arenes.11a–c A site-directed mutant strain, E. coli F352V, containing
the same NDO gene (NDO3) as P. putida 9816/4, but with the
phenylalanine amino acid, Phe 352, at the active site replaced
by valine,11a was also used in the biotransformation of B[b]T
substrates. Earlier studies had shown that the regioselectivity of
cis-dihydroxylation found using NDO1 from P. putida 9816/11 on
biaryl substrates (e.g. biphenyl and 2-phenyl pyridine), polycyclic
arenes (e.g. naphthalene, phenanthrene), and polycyclic azaarenes
(e.g. benzo[h]quinoline and phenanthridine) could be redirected
in some cases, when E. coli F352V (NDO3) was used.11a–c Evi-
dence of a similar change in regioselectivity was sought during
NDO3-catalysed sulfoxidation or cis-dihydroxylation of polycyclic
thiaarenes when using B[b]T and MB[b]T substrates.
The oxidation of B[b]T 2 using P. putida 9816/11 (NDO1), P.
putida NCIMB 8859 (NDO2) and E. coli F352V (NDO3) only
showed evidence of oxidation within the electron-rich thiophene
ring giving bioproducts 7, 10 or 11 (Table 1). As anticipated, no
cis-dihydrodiol products were observed using the wild-type strain
(P. putida NCIMB 8859) and only the tetracyclic adducts, 10 and
11 derived from B[b]T 2 were detected, albeit in very low isolated
yield (<5%). The heterocyclic dihydrodiol 7cis/7trans was formed as
the major product (85%) along with the tetracyclic sulfoxide 11
(15%) using both NDO2 and NDO3 (Table 1). As expected from
earlier cis-dihydroxylations, using E. coli F352V (NDO3),11a–c the
isolated yields were much lower compared with those obtained
using P. putida 9816/11 (NDO1).
tetracyclic metabolites 10cis and 10trans
.
The extrusion of sulfur monoxide from tetracyclic disulfoxide
9 to yield 10cis and 10trans would result in the reformation of a
benzene ring while a similar reaction of a bicyclic disulfoxide (e.g.
F, R and R¢ = H, Scheme 2) formed from a monocyclic thiophene
sulfoxide (e.g. C, R and R¢ = H) would produce a thiophene ring.
The larger increase in resonance energy resulting from formation
of the tetracyclic monosulfoxides 10cis and 10trans compared with
the bicyclic monosulfoxides (e.g. G, R and R¢ = H, Scheme 2) may
be an important factor in the instability of disulfoxide 9.
In our preliminary communication,3j the possibility of a cy-
cloaddition reaction occurring between the substrate B[b]T 2 and
the B[b]T sulfoxide metabolite 8 was discussed. However, in light
of more recent results showing that: (i) P. putida UV4 contains
a sulfoxide reductase enzyme with an ability to catalyse the
mono-deoxygenation of monocyclic thiophene sulfoxide dimers
(F, Scheme 2) 8 and (ii) both benzo[b]naphtho[2,1-d]thiophene 12
and its monosulfoxide 11 can be formed as minor products from
MCPBA oxidation of B[b]T 2 followed by abiotic deoxygenation,
the metabolic sequence 8 → 9 → 10cis/10trans → 11 → 12, shown
in Scheme 3, is proposed.
In common with the disulfoxides, formed by dimerization of
monocyclic thiophene sulfoxides (e.g. compound F → G, R and
R¢ = H, R = H and R¢ = Me, Scheme 2),8 the trans isomer 10trans
was formed in preference (3 : 1) to the cis diastereoisomer 10cis. The
separated tetracyclic sulfoxide metabolites 10trans ([a]D +65) and 11
([a]D -13), whose absolute configurations were not determined,
had very low enantiopurity values (8% ee and 3% ee respectively
by chiral stationary phase HPLC analysis).
Following earlier bacterial metabolism studies of B[b]T 2, GC-
MS analysis showed the presence of both sulfoxide 8 and sulfone
16.3g,3h,3l A similar GC-MS approach was initially employed in the
current study using the recombinant strain E.coli BL21 (DE3)-
pRSET-styAB, a source of styrene monooxygenase (SMO).12a–c
This method again showed the presence of B[b]T-1-oxide 8 as
the major metabolite (>95%), and B[b]T-1,1-dioxide 16 as a
minor product (<5%). This metabolic profile was confirmed by
reverse phase LC-TOFMS analysis (Table 1). The relatively pure
sample of B[b]T sulfoxide 8 thus obtained, enabled a full structure
characterization to be carried out.
(-)-Benzo[b]naphtho[2,1-d]thiophene-7-oxide 11 appeared to
be configurationally stable, i.e. showed no evidence of spontaneous
racemization at ambient temperature. This would be expected for
a sulfoxide inversion barrier (DG‡) greater than 23 kcal mol-1 and
was in accord with the calculated value of DG‡ = 32.3 kcal mol-1
for dibenzo[b]thiophene sulfoxide.9a The low enantiopurity values
(<10% ee) observed for the tetracyclic sulfoxides 10trans and 11
could result from several metabolic sequences. These include: (i)
an initial enzyme-catalysed asymmetric sulfoxidation (2 → 8),
(ii) a sulfoxide reductase-catalysed kinetic resolution involving a
mono-deoxygenation step (8 → 2 or 11 → 12), (iii) a combination
of these pathways allied to partial racemization of sulfoxide 2.
Whole cells of P. putida UV4, a source of TDO, are known
to catalyse a wide range of aromatic dihydroxylations and
sulfoxidations.3e,3i,3j,10a–c The relative yields of the B[b]T metabolites
obtained using three bacterial strains each containing naph-
thalene dioxygenase (NDO) are shown in Table 1. P. putida
9816/11(NDO1, an inducible mutant strain) and P. putida NCIMB
8859 (NDO2, a wild-type strain), each contain NDO and can catal-
yse both vicinal dihydroxylations and sulfoxidations of aromatic
Sulfone 16 could have been formed from sulfoxide 8 by: (a)
decomposition of B[b]T sulfoxide 8 in the GC injection port (as
reported for 2-MB[b]T sulfoxide 18,3d or (b) a disproportionation
reaction (with B[b]T 2 as the other product) during the work up
procedure involving extraction, concentration and purification.
We anticipated that it would be quite difficult to isolate and
determine either the enantiopurity or absolute configuration of
sulfoxide 8 due to its thermal instability and possible spontaneous
racemisation, as predicted from an earlier calculation of its barrier
o
to inversion (DG‡ ca.: 24 kcal mol-1 at 25 C).9a To circumvent
these potential difficulties, the cooled (ca.: 4 ◦C) aqueous culture
medium, containing the bioproducts from a biotransformation
of B[b]T 2, using SMO [whole cells of E. coli BL21 (DE3)-
pRSET-styAB], was concentrated at ice bath temperature and the
concentrate quickly extracted (EtOAc) to minimize the exposure
of bioproducts at ambient temperature. The solvent was removed
at a similar temperature, the residue dried at 0 ◦C under high
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Scheme 4 Dioxygenase-catalysed oxidation of B[b]T 17 and 22.
vacuum and stored below -20 ◦C. The 1H-NMR spectrum of the
crude bioproduct indicated the presence of sulfoxide 8 as the major
metabolite (ca.: 97%) and sulfone 16 as the minor product (ca.:
2%) with trace amounts of substrate 2. After recording the optical
rotation (sign and magnitude) of a portion of the crude product in
CHCl3 solution, the sample was carefully recovered and purified
by PLC (EtOAc–hexane, 1 : 1). The purified, dried and weighed
sample was then used to calculate the optical rotation ([a]D -235,
CHCl3) of sulfoxide 8. The [a]D value of the purified sulfoxide 8
recorded may not be the maximum possible. It registered a rapid
decrease and after keeping the solution at room temperature for
a few hours the sample was found to have an extremely low value
([a]D -3). 1H- and 13C-NMR spectral data of this sample, recorded
every hour, confirmed the purity of sulfoxide 8 (≥97%), indicating
that a rapid racemisation without significant decomposition had
occurred.
When the sample of B[b]T sulfoxide 8 was allowed to remain
at ambient temperature for several days, it was again found to
have decomposed into a mixture of achiral disproportionation and
racemic dimerization products. This observation suggests that the
enantiomeric excess found in bioproducts 10trans ([a]D +65) and 11
([a]D -13), obtained using P. putida UV4, is consistent with either
an asymmetric synthesis and partial racemization of sulfoxide 8
and/or a sulfoxide reductase-catalysed kinetic resolution of the
racemized dimer 9.
The effects of methyl substituent location around the B[b]T
ring, on the regioselectivity of oxygenase-catalysed oxidation,
were also investigated using substrates 17 and 22 (Scheme 4,
Table 1). Both TDO- and NDO1-catalysed oxidations of 2-
methylbenzo[b]thiophene (2-MB[b]T) 17 showed a marked pref-
erence for cis-dihydroxylation of the carbocyclic ring to yield cis-
dihydrodiol 20 (83% and 90% relative yields respectively, Table 1).
This result was consistent with a 2-methyl substituent inhibiting
attack in the thiophene ring at either the sulfur atom or the substi-
tuted 2,3-bond. While the heterocyclic cis-diol 19 was also found
as a minor metabolite using TDO (10%), it was not observed using
NDO1 from P. putida 9816/11. Sulfoxide 18 was only obtained as a
minor metabolite (<10%) from these TDO- and NDO1-catalysed
oxidations. Conversely, oxidation catalysed by NDO2 from the
wild-type strain, P. putida NCIMB 8859, gave no evidence of cis-
dihydroxylation and, as expected, yielded mainly the sulfoxide 18
(90%) and a minor achiral metabolite (10%) that was tentatively
assigned as 3-hydroxy-5-methyl-thiophene-2-carboxaldehyde 21,
based on 1H-NMR and MS data. Further evidence in support of
this assignment was available from the similar characteristics of 3-
hydroxythiophene-2-carboxaldehyde, a metabolite isolated from
the bacterial biotransformation of the parent B[b]T 2.3f It was
assumed that bioproduct 21 originated from the initially formed
but undetected carbocyclic cis-dihydrodiol 20 and ring opening of
the derived catechol.
Significantly, when NDO3 was used as biocatalyst, a marked
change in regioselectivity, compared with NDO1, was observed.
A comparison of relative yields showed that oxidation occurred
exclusively within the thiophene ring to give cis-diol 19 (60%)
and 2-MB[b]T sulfoxide 18 (40%) using NDO3, while oxidation
catalysed by NDO1 occurred mainly in the carbocyclic ring to give
cis-dihydrodiol 20 (90%).
The sulfoxide metabolite 18 proved to be relatively stable
in comparison to the parent sulfoxide 8 but was only found
as a minor metabolite from TDO (7%), or NDO1-catalysed
oxidation (10%, Table 1). To separate sulfoxide 18 from the
cis-dihydrodiol metabolites 19 and 20 derived from 2-MB[b]T
17, flash column chromatography was used. Although sulfoxide
18 had been detected earlier as a P. putida metabolite, using
GC-MS and GC-FTIR analysis,3c it had not been isolated and
structurally/stereochemically assigned prior to the preliminary
report of this study.3i,j The recrystallized sulfoxide 18 isolated as
a minor metabolite using TDO, proved to be a single enantiomer
1
([a]D -476, CHCl3) based on H-NMR analysis in the presence
of (+)-(S)-1-(9-anthryl)-2,2,2-trifluoroethanol. The (1R) absolute
configuration, established for (-)-2-MB[b]T sulfoxide 18 by X-
ray crystallography in our earlier communication,3i showed a
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characteristic electronic circular dichroism (ECD) spectrum. Sul-
foxide 18, obtained in higher relative (90%) and isolated yields
(26%), using P. putida NCIMB 8859 (NDO2), had the opposite
(1S) configuration ([a]D +267, CHCl3), and a lower ee value
(56%).
It was not possible to obtain an accurate value for the barrier
to racemization of B[b]T sulfoxide 8, due to its reduced configura-
tional and thermal stability. However, concomitant observations
1
using polarimetry and H-NMR spectroscopy allowed a crude
estimate from the faster rate of racemization (t1/2 value of ca.
180 min at 25 ◦C, k = 6.4 ¥ 10-6 s -1). This was entirely consistent
with the calculated lower barrier to racemization DG‡ 23.9 kcal
In contrast to the results obtained using dioxygenase enzymes,
2-MB[b]T 17 was converted in quantitative yield to the cor-
responding sulfoxide 18 using SMO [E. coli BL21(DE3)]. For
preparative purposes, the SMO-catalysed oxidation appeared to be
the most promising enzymatic route but stereochemical analysis of
the isolated metabolite 18 indicated that, stereoselectivity during
sulfoxidation of 2-MB[b]T 17 was relatively low ([a] D +15, CHCl3;
3% ee). Although chemically stable, sulfoxide 18, in common with
sulfoxide 8, was found to have limited configurational stability.
Thus, a significant degree of racemisation (pyramidal sulfur
inversion) of sulfoxides 8 and 18 was found to occur over a period
of 24 h at 25 ◦C in CHCl3 solution with the former racemizing
faster. Kinetic studies of the thermal racemization of enantiopure
(-)-(1R)- 2-MB[b]T sulfoxide 18 in CHCl3 solution were conducted
using a thermostatically controlled polarimeter cell. The rate
constants (k) for racemization at 25 ◦C and 50 ◦C were 2.45 ¥
10-6 and 7.11 ¥ 10-5 s-1 respectively, giving similar values for the
free energy of activation (DG‡ = 25.1 kcal mol-1) and activation
energy (Ea = 25.7 kcal mol-1).
The biotransformation of 3-MB[b]T 22 using either TDO or
SMO did not produce the expected sulfoxide metabolite 23 (Table
1). However, the other bacterial strains (NDO1-NDO3) yielded
sulfoxide 23 as the major or sole identified metabolite. NDO1
present in the mutant strain P. putida 9816/11 yielded mainly
sulfoxide 23 (90%) and a small proportion of the heterocyclic
ring cis-dihydrodiol 24 (10%). When the wild type strain P. putida
NCIMB 8859 was used, as anticipated, no cis-diols were detected
but the sulfoxide 23 (39%, ([a]D -181, CHCl3; 41% ee) was
obtained as one of the three major bioproducts. The inversion
barrier (DG‡) for thermal racemisation of 3-MB[b]T sulfoxide 23
was found to be 26.4 kcal mol-1.
Utilization of the E. coli F352V strain (NDO3) with 3-MB[b]T
substrate 22 resulted in the detection of sulfoxide 23 as the sole
isolated metabolite but the very low isolated yield obtained using
NDO3 precluded any rigorous stereochemical analysis. Earlier
reports showed how regioselectivity, during NDO-catalysed car-
bocyclic arene cis-dihydroxylation, could be redirected towards
alternative C C bonds, when using NDO3.11a–c The present study
now indicates that the NDO3 enzyme, present in the E. coli F352V
site directed mutant strain, catalyses preferential oxidation of the
heterocyclic ring (and the sulfur atom in particular) rather than
the carbocyclic ring when using MB[b]T substrates 17 and 22.
The ECD spectra of sulfoxides (-)-18 and (-)-23 obtained
using TDO were very similar and thus confirmed that the [1R]
absolute configuration was common to both. The other two
achiral bioproducts, 26 and 27, resulting from NDO-catalysed
oxidation of the methyl group had been reported as bioproducts
from bacterial metabolism of 3-MB[b]T 22.3h On account of its
thermal and configurational instability, only a qualitative ECD
spectrum of (-)-sulfoxide 8 metabolite (using SMO) was recorded
in a cooled MeCN solution. The ECD spectrum showed similar
Cotton effects to those found for (-)-(1R)-sulfoxides 18 and (-)-
23. Based on this observation (-)-B[b]T sulfoxide 8 was tentatively
assigned a similar (1R) absolute configuration.
mol -1
)
9a compared with those found experimentally for sulfoxides
-
18 (k = 2.45 ¥ 10-6 s 1, DG‡ = 25.1 kcal mol -1) and 23 (DG‡ =
26.4 kcal mol-1).
An earlier value for the barrier to pyramidal inversion of 2,5-di-
o
tert-octylthiophene sulfoxide, obtained experimentally at 25 C
by NMR spectroscopy, (DG‡ = 14.8 kcal mol -1),9b compared
favourably with that obtained by semiempirical calculation of the
inversion barrier for 2,5-dimethylthiophene (DH‡ = 13.3 kcal mol
-1).9c These9b and later ab initio calculations9a led to the prediction
that the barriers to pyramidal inversion for sulfoxides 8 (DH‡ =
9a
-1 9c
23.9 kcal mol-1) and 18 (DH‡ = 19.4 kcal mol
) would be
significantly higher than those calculated for thiophene sulfoxide
(DH‡ = 11.2 kcal mol -1 9a as a result of 2,3-annelation. The higher
inversion barriers (DG‡) obtained experimentally for sulfoxides 8
(< 25 kcal mol -1), 18 (25.1 kcal mol -1) and 23 (26.4 kcal mol -1
)
)
are in general accord with the calculated values. As the normal
barriers for racemisation and pyramidal inversion of sulfoxides
are generally in the range DG‡ = 37–42 kcal mol,-1 the calculated
and experimental values obtained for monocyclic and bicyclic
thiophene sulfoxides are evidently much lower. These markedly
reduced pyramidal inversion barriers for thiophene sulfoxides
have been ascribed to maximal cyclic p-conjugation and aromatic
stabilization of the planar transition state during inversion.11b
Conclusion
The biotransformations of B[b]T substrates 2, 17 and 22 us-
ing TDO, NDO and SMO enzymes were found to yield the
corresponding chiral sulfoxides 8, 18, 23 and derived products.
The unstable B[b]T sulfoxide 8 from B[b]T 2 was isolated and
characterized following a biotransformation using SMO. Dimer-
ization of sulfoxide 8 followed by sulfoxide reductase-catalysed
deoxygenation is proposed to account for the formation of the
tetracyclic metabolites 10–12 using TDO.
The regioselectivity observed during biocatalysed oxidation of
B[b]T substrates 2, 17, 22, examined using the oxygenase enzymes
TDO, NDO1, NDO2, NDO3 and SMO, was found to be dependent
on the substituent pattern and enzyme type. The site-directed
mutant strain E. coli F352V (NDO3) generally showed a strong
preference (85–100%) for oxidation of the thiophene ring over the
benzene ring compared to TDO and NDO1.
The first example of an enantiopure thiophene sulfoxide,
metabolite (-)-18, along with enantioenriched sulfoxides (-)-8 and
(-)-23, were found to racemize at ambient temperature. Absolute
configurations of the (-)-B[b]T sulfoxides were assigned as (1R)
on the basis of ECD spectroscopy (8, 18 and 23) and X-ray
crystallography (18).
The barriers to pyramidal inversion were determined for the
(1R)-sulfoxides (-)-18 and (-)-23 (DG‡ = 25–27 kcal mol.-1)
and compared with previously calculated barriers. The inversion
barrier for (-)-sulfoxide 8 appeared to be slightly lower (DG‡ <
25 kcal mol.-1).
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(Na2SO4). The crude product obtained after evaporating the
solvent was separated into two fractions by flash chromatography
(CHCl3). The earlier major fraction on evaporation gave sulfone
16 and the minor fraction on PLC (Et2O : pentane, 1 : 1) further
separated into four minor products 10trans, 10cis, 11 and 12.
Experimental
1H and 13C NMR spectra were recorded on Bruker Avance 400,
DPX-300 and DRX-500 instruments. Chemical shifts (d) are
reported in ppm relative to SiMe4 and coupling constants (J) are
given in Hz. Mass spectra were run at 70 eV, on a VG Autospec
Mass Spectrometer, using a heated inlet system. Accurate molec-
ular weights were determined by the peak matching method, with
perfluorokerosene as the standard. Flash column chromatography
and preparative layer chromatography (PLC) were performed
on Merck Kieselgel type 60 (250–400 mesh) and PF254/366 plates
respectively. Merck Kieselgel type 60F254 analytical plates were
employed for TLC. The benzo[b]thiophene substrates (2, 17,
22), were available from commercial sources and from earlier
studies.3e,i,j,5,9a A PerkinElmer 341 polarimeter was used for optical
rotation ([a]D) measurements at ambient temperature (23–25 ◦C).
Enantiomeric excess values were determined by chiral stationary
phase HPLC using Chiralcel OJ or OD-H columns (method
Benzo[b]thiophene-1,1-dioxide 1614a. White crystalline solid
◦
(7.63 g, 60%); m.p. 141–143 C (from MeOH), (lit.14a m.p. 142–
143 ◦C); 1H-NMR (300 MHz, CDCl3) d 6.73 (1 H, d, J3,2 6.9, 3-H),
7.23 (1 H, d, J2,3 6.8, 2-H), 7.37 (1 H, dd, J4,5 7.8, J4,6 1.8, 4-H),
7.53–7.58 (2 H, m, 6-H), 7.73 (1 H, dd, J7,6 7.6, J7,5 1.3, 7-H); m/z
∑
166 (M+ , 52%), 137 (100).
Benzo[b]naphtho[2,1-d]thiophene 123g,14b. White solid (0.018
g); m.p. 98–100 ◦C (MeOH), (lit.14b m.p. 102–103 ◦C); Rf 0.95; 1H-
NMR (300 MHz, CDCl3) d 7.48–7.53 (1 H, m, Ar–H), 7.56–7.63
(2 H, m, Ar–H), 7.72–7.77 (1 H, m, Ar–H), 7.88–7.94 (2 H, m,
Ar–H), 8.00–8.04 (2 H, m, Ar–H), 8.87 (1 H, d, J5,6 8.2, 5-H), 9.02
∑
(1 H, d, J6,5 8.3, 6-H); m/z 234 (M+ , 100%).
trans-6a,11b-Dihydronaphtho[2,1-d]benzo[b]thiophene-7-
oxide 10trans.3l. White solid (0.045 g); m.p. 155–156 ◦C
(EtOAc/pentane); Rf 0.42; (Found: C, 76.0; H 4.7; S, 12.5
C16H12OS requires C, 76.2, H 4.8; S, 12.7%); 1H-NMR (500 MHz,
CDCl3) d 4.51 (1 H, d, J11b,6a 7.2, 11b-H) 4.74–4.76 (1 H, m, 6a-H),
6.23 (1 H, dd, J6,5 9.9, J6,6a 3.0, 6-H), 6.76 (1 H, dd, J5,6 9.9, J5,6a
2.2, 5-H), 7.13–7.17 (2 H, m, Ar–H), 7.27–7.30 (3 H, m, Ar–H),
7.39–7.46 (2 H, m, Ar–H), 7.79 (1 H, d, J8,9 7.4, 8-H); 13C-NMR
(125 MHz, CDCl3) d 44.64, 63.97, 117.91, 126.54, 126.70, 128.55,
128.67, 128.69, 128.74, 129.49, 132.00, 132.44, 132.64, 132.78,
1
A), by H-NMR analysis in the presence of the chiral resolving
reagent (+)-(S)-1-(9-anthryl)-2,2,2-trifluoroethanol (method B) or
by comparison of optical rotation values (method C). Absolute
configurations were determined using ECD spectroscopy and
spectra were recorded using spectroscopic grade acetonitrile and
a JASCO J-720 instrument.
Liquid chromatography-time of flight mass spectrometry (LC-
TOFMS) analyses were conducted using an Agilent 1100 series
HPLC coupled to an Agilent 6510 Q-TOF (Agilent Technologies,
USA). Separation was performed using a reverse phase column
(Agilent Eclipse Plus C18, 5 mm, 150 ¥ 2.1 mm) together with the
corresponding guard column (5 mm, 12.5 ¥ 2.1 mm). The mobile
phase consisted of 95% methanol containing 0.1% formic acid
in channel A, and 5% methanol containing 0.1% formic acid in
channel B. The system was programmed to perform an analysis
cycle consisting of 100% B for 1 min, followed by gradient elution
from 100% to 5% B over a 14 min period, hold at 5% B for 10
min, return to initial conditions over 2 min and then hold these
conditions for a further 8 min. The flow rate was 0.20 ml min-1
and the injection volume was 5 ml. MS experiments were carried
out using ESI in positive ion mode with the capillary voltage set
at 4.0 kV. The desolvation gas was nitrogen set at a flow rate of 8
L min-1 and maintained at a temperature of 350 ◦C.
Shake flask and fermenter scale (50 L) biotransformations were
carried out using whole cells of P. putida UV4 (TDO), P. putida
NCIMB 8859 (NDO1), P. putida 9816/11 (NDO2), and E. coli
F352V (NDO3) using methods described earlier.10a–c,13a–c SMO-
catalysed sulfoxidations were conducted using cell suspensions
of the E.coli BL21(DE3)-pRSET-styAB strain expressing styrene
monooxygenase from Pseudomonas putida CA-3 as previously
described,1c using B[b]T 2 and 2-MB[b]T 17 as substrates.
∑
142.25, 143.58; m/z 252 (M+ , 40%), 235 (100).
Benzo[b]naphtho[2,1-d]thiophene-7-oxide 1114c. White crys-
◦
talline solid (0.026 g); m.p. 142–144 C (Et2O/pentane); (lit.14c
◦
1
147–148 C); Rf 0.40; H-NMR (300 MHz, CDCl3) d 7.50–7.55
(1 H, m, Ar–H), 7.61–7.72 (3 H, m, Ar–H), 7.96–8.08 (4 H, m,
Ar–H), 8.45 (1 H, d, J5,6 8.2, 5-H), 8.74 (1 H, d, J6,5 8.3, 6-H); m/z
∑
250 (M+ , 14%), 234 (100).
cis-6a,11b-Dihydronaphtho[2,1-d]benzo[b]thiophene-7-oxide
10cis.3l. White solid (0.015 g); m.p. 156–157 ◦C (EtOAc/pentane);
Rf 0.36; (Found: M+, 252.0609 C16H12OS requires 252.0593);
1H-NMR (300 MHz, CDCl3) d 4.60–4.63 (1 H, m, 6a-H), 5.24
(1 H, d, J11b,6a 6.7, 11b-H) 5.87 (1 H, dd, J6,5 9.7, J6,6a 2.4, 6-H),
6.47 (1 H, dd, J5,6 9.8, J5,6a 2.5, 5-H), 7.08–7.14 (2 H, m, Ar–H),
7.30–7.46 (5 H, m, Ar–H), 7.86 (1 H, d, J8,9 6.9, 8-H); 13C-NMR
(125 MHz, CDCl3) d 45.84, 69.16, 119.20, 126.59, 126.73, 128.08,
128.28, 128.53, 128.71, 129.13, 131.15, 132.62, 132.45, 132.60,
∑
143.25, 143.63; m/z 252 (M+ , 48%), 235 (100).
(b) Using dimethyl dioxirane (DMD).
A
sample of
benzo[b]thiophene 2 (ca. 0.1 g) was treated, in an ice cold
bath, with an excess of pre-dried d6-acetone solution of DMD
(0.085 M), prepared using the literature method.17 The reaction
mixture was stirred, monitored by TLC (EtOAc : hexane, 1 : 1),
and was allowed to warm to room temperature. When most of
the starting compound had been consumed, the products of the
reaction mixture were examined, in situ, by NMR spectroscopy
and showed mainly the presence of racemic benzo[b]thiophene-
1-oxide 8. Removal of the solvent under vacuum and storage of
crude product at ambient temperature resulted in a mixture of
products.
Oxidation of benzo[b]thiophene 2
(a) Using m-chloroperoxybenzoic acid (MCPBA). To
a
stirred solution of benzo[b]thiophene 2 (10.0 g, 77 mmol) in
methylene chloride (200 ml) maintained at 0 ◦C was added, in small
portions, MCPBA (16.8 g, 81 mmol). The mixture was stirred
overnight at ambient temperature. The chlorobenzoic acid was
filtered from the reaction mixture, the filtrate washed successively
with Na2SO3 solution, NaHCO3 solution, water and then dried
( )-Benzo[b]thiophene-1-oxide 8. 1H-NMR (300 MHz,
CD3COCD3) d 7.30–7.35 (2 H, m, 5-H, 7-H), 7.38 (1 H, d, J3,2
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5.5, 3-H), 7.60 (1 H, d, J2,3 5.4, 2-H), 7.82–7.85 (1 H, m, 6-H),
7.90–7.93 (1H, m, 4-H); 13C-NMR (125 MHz, CD3COCD3) d
122.98, 124.21, 124.53, 124.78, 124.85, 127.20, 140.26, 140.45.
Data also given for (-)-(R)-sulfoxide 8 in section (c, iii).
benzo[b]thiophene-1,1-dioxide 16 (ca. 2%) and trace amounts of
benz[b]thiophene 2.
(-)-(1R)-Benzo[b]thiophene-1-oxide 8. Light yellow oil (purified
by PLC); (0.078 g, 70%); Rf 0.28 (EtOAc–hexane, 1 : 1); (Found
∑
M+ 150.0146. C8H6SO requires 150.0139); [a]D -235 (c 1.25,
(c) Oxygenase-catalysed sulfoxidations of benzo[b]thiophenes
2, 17, 22.
1
CHCl3); H-NMR (500 MHz, CDCl3) d 7.08 (1 H, d, J 6.3, 3-
H), 7.22 (1 H, d, J 6.3, 2-H), 7.46–7.54 (3 H, m, 5-H, 6-H and
7-H), 7.92 (1H, d, J 7.3 4-H); 13C-NMR (125 MHz, CDCl3) d
125.42, 126.64, 129.46, 132.45, 135.27, 137.54, 137.93, 145.67.
(iv) Biotransformation of 2-methylbenzo[b]thiophene 17 with
P. putida UV4. Substrate 17 (6.0 g) was metabolised with
P. putida UV4 and the bioproducts were extracted (EtOAc)
from the aqueous phase. The crude mixture of products ob-
tained after removal of EtOAc was purified by flash chro-
matography (75% Et2O in pentane → Et2O) to afford (+)-2-
methyl-cis-2,3-dihydroxy-2,3-dihydro-benzo[b]thiophene 19 (0.2
g, 3%, Rf 0.4, 50% Et2O/pentane), (+)-2-methyl-cis-4,5-dihydroxy-
4,5-dihydrobenzo[b]thiophene 20 (1.8 g, 25%, Rf 0.23, 50%,
(i) Biotransformation of benzo[b]thiophene 2 with P. putida
UV4. A biotransformation with P. putida UV4 and sub-
strate 2 (13 g, 100 mmol) followed by flash column chro-
matography (50% Et2O in pentane) yielded pure samples
of benzo[b]naphtho[2,1-d]thiophene 12 (0.180 g, Rf 0.95), 4-
hydroxy-benzo[b]thiophene 15 (0.520 g, Rf 0.88), 5-hydroxy-
benzo[b]thiophene 14 (0.705 g, Rf 0.85), cis/trans-2,3-dihydroxy-
2,3-dihydrobenzo[b]thiophene 7 (4.9 g, Rf 0.50),7 and cis-4,5-
dihydroxy-4,5-dihydrobenzo[b]thiophene 13 (0.785 g, Rf 0.20).7
Further purification of unresolved fractions by multiple elution
PLC (50% Et2O in pentane) yielded three other bioproducts
10trans, 10cis, and 11 which were identified by direct compar-
ison with authentic samples synthesised earlier by chemical
oxidation. Metabolites 5-hydroxybenzo[b]thiophene 14 and 4-
hydroxybenzo[b]thiophene 15 were found to have similar prop-
erties to those reported in the literature.14c Full spectroscopic and
other characterization data for compounds 14, 15 and 7 will be
given in the separate report.7 The samples of 10trans, 10cisand 11
showed identical spectra to the racemic products obtained by
chemical oxidation in Section (a).
7
Et2O/pentane) and (-)-2-methylbenzo[b] thiophene-1-oxide 18
(0.15 g, 2%).
(-)-(1R)-2-Methylbenzo[b]thiophene-1-oxide 18.15 white crys-
talline solid (0.057 g, 51%); Rf 0.24 (CHCl3); m.p. 80–82 ◦C (from
CHCl3/hexane), (lit.15 m.p. 80–81 ◦C); [a]D -476 (c 0.41, CHCl3);
>99% ee (Method B); nmax (KBr) 1060 cm-1 (S O); 1H-NMR (300
MHz, CDCl3) d 2.36 (3 H, d, J 1.5, Me), 6.78 (1 H, q, J 1.5, 3-H),
7.32–7.46 (3 H, m, 5-H, 6-H and 7-H), 7.82 (1 H, d, J 7.0, 4-H);
∑
m/z 164 (M+ , 96%), 147 (70), 121 (100); ECD: 332 nm (De -7.0),
(+)-trans-6a,11b-Dihydronaphtho[2,1-d]benzo[b]thiophene-7-
oxide 10trans. (0.245 g); Rf 0.42; [a]D +65 (c 0.47, EtOH); 8% ee
(Method A, Chiralcel OJ, 20% EtOH/hexane).
( )-cis-6a,11b-Dihydronaphtho[2,1-d]benzo[b]thiophene-7-oxide
10cis. (0.080 g); Rf 0.36.
(-)-Benzo[b]naphtho[2,1-d]thiophene-7-oxide 11. (0.390 g); Rf
0.40; [a]D -13 (c 0.52, CHCl3); 3% ee (Method A, Chiralcel OJ,
20% EtOH/hexane).
301 nm (De +5.8), 270 nm (De -4.5), 224 nm (De -35.0).
(v) Biotransformation of 2-methylbenzo[b]thiophene 17 using
P. putida 8859. Substrate 17 (0.2 g) on biotransformation using
P. putida 8859 gave a mixture of two metabolites. Separa-
tion by PLC (1% MeOH in CHCl3) afforded of 3-hydroxy-5-
methylthiophene-2-carboxaldehyde 21 and the more polar (+)-
(1S)-2-methylbenzo[b]thiophene-1-oxide 18.
3-Hydroxy-5-methylthiophene-2-carboxaldehyde 21. (0.006 g,
3%); Rf 0.57; nmax (neat) 1610 cm-1 (C O) and 3330 cm-1 (OH);
1H-NMR (300 MHz, CDCl3) d 2.51 (3 H, s, Me), 6.5 (1 H, s, 4-H),
(ii) Biotransformation of benzo[b]thiophene 2 with NDO.
Biotransformation of substrate 2 was carried out using P. putida
9816/11 (NDO1), P. putida NCIMB 8859 (NDO2) and E. coli
F352V (NDO3) in two litre shake flasks for comparative study
under standard conditions (ca. 0.4 g substrate/500 ml culture
medium). The metabolites 7, 10 and 11 present in the crude
aqueous extract were identified by LC-TOFMS. The aqueous
biotransformed material was extracted with EtOAc and the crude
mixture of bioproducts obtained after removal of EtOAc was
separated by PLC. The separated bioproducts 7, 10 and 11 were
identified by direct comparison with authentic samples (Table 1).
∑
9.47 (1 H, s, CHO); m/z 142 (M+ , 87%), 141 (100).
(+)-(1S)-2-Methylbenzo[b]thiophene-1-oxide 18. (0.057 g, 26%);
[a]D +267 (c 0.81, CHCl3); 56% ee (methods B and C).
(vi) Biotransformation of 2-methylbenzo[b]thiophene 17 with
SMO [E.coli BL21(DE3)-pRSET-styAB]. Substrate 17 (0.02
g) was metabolized using SMO. The NMR spectrum of the
crude bioproduct showed the presence of metabolite 18 only
and a trace amount of starting substrate. Purification by PLC
(EtOAc : hexane, 1 : 1) furnished sulfoxide 18 (0.019 g, 94%); [a]D
+15 (c 0.8, CHCl3); 3% ee (method C).
(vii) Biotransformation of 3-methylbenzo[b]thiophene 22 with
P. putida UV4. Substrate 22 (7.5 g) when biotransformed using
P. putida UV4 yielded a single bioproduct, (+)-3-methyl-(4R,5S)-
cis-4,5-dihydroxy-4,5-dihydrobenzo[b]thiophene 25 (4.1 g, 45%)
whose characterization will be reported elsewhere.7
(viii) Biotransformation of 3-methylbenzo[b]thiophene 22
with P. putida NCIMB 8859. Substrate 22 (0.3 g) yielded
a crude mixture of three metabolites on biotransformation
with P. putida NCIMB 8859. Separation by PLC (Et2O)
gave 3-hydroxymethylbenzo[b]thiophene 26, benzo[b]thiophene-
3-carboxylic acid 27 and (-)-(R)-2-methylbenzo[b]thiophene-1-
oxide 23.
(iii) Biotransformation of benzo[b]thiophene 2 with SMO
[E.coli BL21(DE3)-pRSET-styAB]. Substrate 2 (ca. 0.1 g) in
500 ml of cell suspension of E.coli BL21(DE3)-pRSET-styABO
(5 g l-1) in 50 mM phosphate buffer (pH 7.4) containing 1% (w/v)
glucose was biotransformed as previously described.1c The cellular
debris was removed by centrifugation and the clear aqueous por-
tion was stored in the fridge. LC-TOFMS analysis of the aqueous
medium showed benzo[b]thiophene sulfoxide 8 to be the dominant
metabolite (>95%) with only traces of benzo[b]thiophene-1,1-
dioxide 16 (<5%) detected. A small sample of the cooled aqueous
portion was then extracted (EtOAc) and the extract analysed by
GC-MS. It showed the presence of benzo[b]thiophene-1-oxide 8
as the major product (ca. 97%) along with minor amounts of
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3-Hydroxymethylbenzo[b]thiophene 26.3h (0.02 g, 6%); Rf 0.64;
J. Chromatogr., A, 1992, 591, 362; (e) D. R. Boyd, N. D. Sharma, R.
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4 D. R. Boyd and N. D. Sharma, Chem. Soc. Rev., 1996, 289.
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6 P. M. Dansette, G. Bertho and D. Mansuy, Biochem. Biophys. Res.
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∑
nmax (neat) 3350 cm-1 (OH); (Found M+ 164.0292. C9H8OS
1
requires 164.0296); H-NMR (300 MHz, CDCl3) d 4.91 (2 H,
m, CH2) 7.35–7.42 (3 H, m, Ar–H), 7.82–7.87 (2 H, m, Ar–H);
m/z 164 (M+, 100%), 147 (83), 135 (84).
Benzo[b]thiophene-3-carboxylic acid 27.3h (0.021 g, 5%); Rf
0.38 (Et2O); nmax (neat) 1674 cm-1 (C O); (Found M+ 178.0087
C9H6O2S requires 178.0088); 1H-NMR (300 MHz, CDCl3) d 7.40–
7.53 (2 H, m, Ar–H), 7.89 (1 H, d, J 7.9, Ar–H), 8.55 (1 H, s, 2-H),
∑
8.62 (1 H, d, J 8.1, Ar–H); m/z 178 (M+ , 100%), 161 (58).
(-)-(1R)-3-Methylbenzo[b]thiophene-1-oxide 23. (0.023 g, 8%);
Rf 0.18 (CHCl3); m.p. 75–77 ◦C (from CHCl3/hexane), (lit.15 m.p.
77 ◦C); [a]D -181 (c 0.8, CHCl3); 41% ee (method B); nmax (KBr)
1050 cm-1 (S O); 1H-NMR (300 MHz, CDCl3) d 2.28 (3 H, d, J
1.5, Me), 6.77 (1 H, q, J 1.4, 2-H), 7.43–7.59 (3 H, m, Ar–H), 7.90
∑
(1 H, d, J 7.4, Ar–H); m/z 164 (M+ , 30%), 147 (22), 135 (100);
7 D. R. Boyd, N. D. Sharma, I. N. Brannigan, T. A. Evans, S. A. Haughey,
B. T. McMurray, J. F. Malone and C. C. R. Allen, Org. Biomol. Chem.,
2011, manuscript in preparation.
ECD: 325 nm (De -8.0), 296 nm (De +3.8), 267 nm (De -1.8), 225
nm (De -26.2).
8 D. R. Boyd, N. D. Sharma, A. King, S. D. Shepherd, C. C. R. Allen, R.
Holt, H. Luckarift and H. Dalton, Org. Biomol. Chem., 2004, 2, 554.
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1, 66.
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B. Byrne, M. V. Hand, J. F. Malone, G. N. Sheldrake, J. Blacker and
H. Dalton, J. Chem. Soc., Perkin Trans. 1, 1998, 1935; (c) K. Lee, J. M.
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9.
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D. T. Gibson, J. Bacteriol., 2000, 182, 1641; (b) R. E. Parales, S. M.
Resnick, C. L. Yu, D. R. Boyd, N. D. Sharma and D. T. Gibson, J.
Bacteriol., 2000, 182, 5495; (c) D. R. Boyd, N. D. Sharma, G. P. Coen,
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Hamilton, Org. Biomol. Chem., 2008, 6, 3957.
Acknowledgements
We thank Science Foundation Ireland for postdoctoral funding
(JR-N, LAC, NDS; Grant no. 04/IN3/B581), and DENI for
a postgraduate studentship (SAH) and a CASE award (BTM).
We also gratefully acknowledge valuable preliminary biotrans-
formation results provided by Friedrich Schmitt and Margarethe
Plotkowiak (University of Applied Sciences, Mannheim, Ger-
many) and LC-TOFMS data from Stewart Floyd (AFBI). The E.
coli F352V strain was kindly supplied by Professors D. T. Gibson
and R. E. Parales (University of Iowa).
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