Dioxygenase-catalysed cis-dihydroxylation of meta-substituted phenols to yield cyclohexenone cis-diol and derived enantiopure cis-triol metabolites

Article abstract of DOI:10.1039/c0ob00894j

cis-Dihydroxylation of meta-substituted phenol (m-phenol) substrates, to yield the corresponding cyclohexenone cis-diol metabolites, was catalysed by arene dioxygenases present in mutant and recombinant bacterial strains. The presence of cyclohexenone cis-diol metabolites and several of their cyclohexene and cyclohexane cis-triol derivatives was detected by LC-TOFMS analysis and confirmed by NMR spectroscopy. Structural and stereochemical analyses of chiral ketodiol bioproducts, was carried out using NMR and CD spectroscopy and stereochemical correlation methods. The formation of enantiopure cyclohexenone cis-diol metabolites is discussed in the context of postulated binding interactions of the m-phenol substrates at the active site of toluene dioxygenase (TDO). The Royal Society of Chemistry 2011.

Full text of DOI:10.1039/c0ob00894j

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PAPER
Dioxygenase-catalysed cis-dihydroxylation of meta-substituted phenols to
yield cyclohexenone cis-diol and derived enantiopure cis-triol metabolites†
Derek R. Boyd,*a Narain D. Sharma,a Paul J. Stevenson,a Marine Blain,b,c Colin McRoberts,b
John T. G. Hamilton,b,d Jose´ M. Argudo,d Harpinder Mundi,d Leonid A. Kulakovd and Christopher C. R. Allend
Received 18th October 2010, Accepted 16th November 2010
DOI: 10.1039/c0ob00894j
cis-Dihydroxylation of meta-substituted phenol (m-phenol) substrates, to yield the corresponding
cyclohexenone cis-diol metabolites, was catalysed by arene dioxygenases present in mutant and
recombinant bacterial strains. The presence of cyclohexenone cis-diol metabolites and several of their
cyclohexene and cyclohexane cis-triol derivatives was detected by LC-TOFMS analysis and confirmed
by NMR spectroscopy. Structural and stereochemical analyses of chiral ketodiol bioproducts, was
carried out using NMR and CD spectroscopy and stereochemical correlation methods. The formation
of enantiopure cyclohexenone cis-diol metabolites is discussed in the context of postulated binding
interactions of the m-phenol substrates at the active site of toluene dioxygenase (TDO).
and bacteria. A range of enzymes may be involved including heme
Introduction
and non-heme monooxygenases, dioxygenases, phenol oxidases
Enzyme-catalysed mono- and poly-hydroxylations of aromatic
and peroxidases.1a,1b Over the past two decades, many reviews
rings are ubiquitous in nature, occurring in animals, plants, fungi
published on this topic indicate that stereoselective bacterial
dioxygenase (DO)-catalysed hydroxylations are among the most
aSchool of Chemistry and Chemical Engineering, Queen’s University Belfast,
widely studied types of arene biotransformations.2a–2n The role
Belfast, UK BT9 5AG. E-mail: dr.boyd@qub.ac.uk; Fax: (+44) (0)28
of bacterial dioxygenases in the oxidation of monosubstituted
90975418; Tel: +44 (0)28 90975419
bAgri-food and Biosiences Institute for Northern Ireland, Belfast, UK BT9
5PX
(A, R¢ = H, R π H) and disubstituted benzenes (A, R and
R¢ π H) and polycyclic arenes, to yield the corresponding cis-
dihydrodiols (B, Scheme 1) is of particular interest because of
their numerous applications as chiral precursors in the synthesis of
enantiopure natural and unnatural products.2a–2n cis-Dihydrodiols
(B) are generally formed as initial intermediates which may
cEnsicaen, Boulevard du Marechal Juin, 14050 Caen, Cedex, France
dSchool of Biological Sciences, Queen’s University Belfast, Belfast, UK BT9
5AG
† Electronic supplementary information (ESI) available. See DOI:
10.1039/c0ob00894j
Scheme 1 Aromatic hydroxylation and mineralization pathways via toluene dioxygenase (TDO), cis-diol dehydrogenase (DD), catechol dioxygenase
(CDO), ene reductase (ERED) and carbonyl reductase (CRED) biocatalysis.
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undergo spontaneous dehydration to yield ortho- (D, R¢ = H)
or m-phenols (E, R¢ = H) or cis-diol dehydrogenase (DD)-
catalysed dehydrogenation to yield catechols (C, R¢ = H). The
latter bioproducts can also be produced via arene dioxygenase-
catalysed oxidation of phenols (D or E, R¢ = H) prior to a ring-
opening process catalysed by an extradiol catechol dioxygenase
(CDO, Scheme 1) or an intradiol CDO.
As phenols are often found to be the initial metabolites isolated
from both arenes A and their cis-dihydrodiol derivatives B, e.g.
ortho-phenols (A → B → D, R¢ = H), m-phenols (A → B → E,
R¢ = H), their further metabolism is of considerable importance in
relation to the environment. Based on earlier literature reports, on
the bacterial metabolism of a range of phenol types, e.g. D or E,3a–3h
it might be inferred that further aromatic hydroxylation, to yield
achiral catechols, e.g. C, was the major or sole metabolic pathway
(Scheme 1). However, studies have indicated that while catechols
C are the dominant bioproducts, a further range of water soluble
metabolites, including ketocarboxylic acids F, hydroxycarboxylic
acids G and picolinic acids H, may also be formed during the
mineralization process of phenols.
Following the pioneering work of Gibson et al.,4 initially using
mutant strains of the soil bacterium Pseudomonas putida, and later
Escherichia coli recombinant strains containing TDO, it became
possible to intercept, characterise and utilize the cis-dihydrodiol
metabolites B. Recent biotransformation studies, using whole
cells of the P. putida UV4 mutant strain, showed that the TDO-
catalysed cis-dihydroxylation was more efficient for monosubsti-
tuted (A, R π H, R¢ = H) compared with disubstituted benzenes
(A, R and R¢ π H). meta-Disubstituted benzene substrates in
particular were generally much more resistant to this type of
oxidation (the meta effect).5a Where possible, the latter type
of meta-substituted benzene substrates were found to undergo
alternative types of TDO-catalysed oxidation according to the
nature of substituents e.g. alkylaryl sulfide sulfoxidation (A, R =
S-Alkyl, meta-R¢ π H),5b alkene dihydroxylation, (A, R = vinyl,
meta-R¢ π H)5c or benzylic hydroxylation (A, R = CH2-Alkyl, meta-
R¢ π H).5a
It has been postulated that bacterial dioxygenase-catalysed cis-
dihydroxylation of phenols (Scheme 2), for example 1e, at the 1,2-
bond to give a transient triol 2e,3f or 1d at the 4,5-bond to yield
the enol cis-diol 4d,3j followed in each case by their spontaneous
dehydration, could in principle account for the formation of the
corresponding catechol 3e (from 2e) or hydroquinone 5d (from
4d).
To our knowledge, no direct evidence of either of these two
types of phenolic cis-dihydrodiols, 2 or 4, has been reported in the
literature. However, a recent preliminary communication of this
study,6 showed that a P. putida UV4 mediated biotransformation
of ortho-, e.g. 1b, and m-phenols, e.g. 1c or 1f, and disubstituted
phenols, e.g. 1d, could yield, in addition to the expected catechols
3b–3d and 3f, enantiopure cyclohexenone cis-diols 6bR–dR and
6fS, as isolable metabolites, albeit in generally modest yields. Since
these compounds are the more stable tautomeric forms of the
initial cis-dihydrodiols 4bR–dR, and 4fS, they have provided the first
direct evidence that phenols, in common with other mono- and
di-substituted benzene substrates, undergo stereoselective TDO-
catalysed cis-dihydroxylation. Tentative evidence of this type of
hydroxylation also occurring at the 5,6-bond of phenols, e.g. 1d,
was found when enantiopure dimeric bioproducts, e.g. 8d, were
obtained.6 The pathway presumably involves rapid dehydration
of the transient phenolic cis-diol metabolite, e.g. 7d, followed
by spontaneous cycloaddition between two molecules of the
Scheme 2 Dioxygenase-catalysed oxidation of phenols 1 to yield catechols 3, hydroquinones 5, cyclohexenone cis-diols 6S (or 6R) and cycloadducts 8.
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Table 1 Relative proportions of cyclohexenone cis-diols 6, triols 12 and carboxylic acids 15 found by LC-TOFMS analysisa and catechols 3 from NMR
analysisb
Phenol (R)
cis-Diol
Relative ratioa
Triol
Relative ratioa
Acid
Relative ratiob
Catechol
Relative ratioa
c
c
1c (Me)
1f (I)
1g (Et)
1h (iPr)
1i (tBu)
1j (CF3)
1k (Ph)
6cS
6fS
6gS
6hS
6iS
++
++++
++
+++
++++
++
12c
12f
12g
12h
12i
15c
15f
15g
15h
15i
3c
3f
3g
3h
3i
+++
++
+
—
d
d
++
d
+
+
+
d
++
c
6jS
12je
12k
+++
15j
3j
+
f
d
d
6kS
++
15k
+
3k
a LC-TOFMS based on peak areas; b NMR; c Not detected; d Trace; e Triol 11j also found (+); f cis-Dihydrodiol 17 also found (+)
resulting diene to yield a diketone, e.g. 8d. This result, and those
obtained from earlier studies,3f,3j,6 where bacterial metabolism of
specific phenols 1 yielded the corresponding achiral catechols
3, cyclohexenone cis-diols 6S and cycloadducts 8, could all be
accounted for by alternative regioselective cis-dihydroxylation
pathways (Scheme 2). Thus, the postulated TDO-catalysed oxi-
dation of phenols 1 occurring at the 1,2-bond3f (e.g. on compound
1d to give catechol 3d via intermediate 2d), at the 4,5-bond (to give
keto cis-diol 6dR via intermediate 4dR)6 and at the 5,6-bond (to
give bis-ketol 8d via intermediate 7d),6 contrasts with the almost
exclusive regioselectivity of TDO-catalysed cis-dihydroxylation at
the 2,3-bond of monosubstituted benzene substrates (A → B, R¢ =
H, Scheme 1).
Following on from our earlier Communication,6 this more
comprehensive study was undertaken in order to establish if:
(i) (a) the new cyclohexenone cis-diol family of metabolites
6S (R¢ = H) could be formed using a wider range of m-phenol
substrates 1 (R¢ = H) and (b) any second generation chiral products
could be derived from these initial metabolites.
(ii) (a) TDO was the enzyme responsible for catalysing the
production of cyclohexenone cis-diols 6S (R¢ = H) or other m-
phenol metabolites by employing a recombinant strain containing
TDO and (b) these bioproducts could be formed using alternative
dioxygenase enzymes.
(iii) the preferential TDO-catalysed cis-dihydroxylation of m-
phenols 1 (R¢ = H), to form enantiopure cyclohexenone cis-diols
6S (R¢ = H), which contrasts with the behaviour of most other
ortho- and para-phenols and other non-phenolic meta-substituted
benzene substrates, could be rationalised in terms of binding
interactions of these substrates within the TDO active site.
Scheme 1). Dehydration of a small proportion of this diol during
biotransformation yielded phenol 1d (Scheme 2). Further TDO-
catalysed cis-dihydroxylation of phenol 1d then gave enol 4dR
which preferentially exists as the keto tautomer, i.e. cyclohexenone
cis-diol 6dR (Scheme 2).6 When phenol 1d was added as substrate
to whole cells of P. putida UV4, a higher yield of cis-diol 6dR
was obtained, compared with that found using para-xylene as
substrate.6 Preliminary studies have shown that alternative para-
disubstituted phenolic substrates, e.g. 1d (R¢ = Cl or Et and R =
Me, or R¢ = Me and R = F), for P. putida UV4, also yield
the corresponding keto cis-diols 6R. The structures and absolute
configurations of keto cis-diols 6R (R = Me, R¢ = Cl or Et and
R = F, R¢ = Me), each containing three chiral centres, derived
from the corresponding disubstituted phenols 1d, will be discussed
elsewhere.
When the m-phenols 1c (and 1f) were previously examined as
substrates for P. putida UV4,6 it was found that the corresponding
cyclohexenone cis-diols, 6cS (and 6fS) were formed as the major
metabolites (15–70% isolated yield) along with the expected cate-
chols 3c (and 3f) as minor bioproducts. ortho-Cresol 1b also proved
to be a substrate for P. putida UV4 producing mainly catechol 3
(R = Me, R¢ = H) and a low yield of cyclohexenone cis-diol 6bR (ca.:
1%).6 When para-cresol was used as substrate, the corresponding 4-
methylcatechol was the only identifiable bioproduct formed. Thus,
the biotransformation of these ortho- and para-substituted phenols
strongly favoured the formation of catechols. As m-phenols 1c
and 1f yielded enantiopure cyclohexenone cis-diols, preferen-
tially, a wider range of m-phenols was examined to extend the
study.
Following the earlier report of cyclohexenone cis-diol formation
from meta-cresol 1c (R = Me, R¢ = H),6 three further alkyl-
substituted m-phenols 1g (R = Et, R¢ = H), 1h (R = iso-Pr, R¢ =
H) and 1i (R = tert-Bu, R¢ = H) were initially found to yield
the corresponding cyclohexenone cis-diols (6gS–iS) as bioproducts
(>5% isolated yield, Scheme 3 and Table 1) accompanied by
minor or trace amounts of the corresponding known catechols
3g–i which were not isolated. NMR spectroscopic analysis was
used to identify the structures of the new keto cis-diols 6gS–iS. It
is noteworthy that the relative yields of these cyclohexenone cis-
diols varied between experiments but generally diminished with
decreasing substituent size (tert-Bu > iso-Pr > Et > Me, Table 1).
The variable isolated yields of both cyclohexenone cis-diols 6gS–
iS, and catechols 3g–i formed during repeated biotransformations
were attributed to further metabolism of these bioproducts.
Results and discussion
(a) Dioxygenase-catalysed cis-dihydroxylation of m-phenols 1c,
1f–k to form cyclohexenone cis-diols 6cS, 6fS–kS, their carbonyl
reductase-catalysed ketone reductions to yield triols 12fS–kS and
other metabolites
The first member of the new cyclohexenone cis-diol family 6dR
(Scheme 2) was isolated from a serendipitous biotransformation
of p-xylene (A, R = R¢ = Me, Scheme 1) using a constitutive mutant
strain (UV4) of P. putida, a source of TDO.6 The initial step in
the biosynthetic sequence involved the TDO-catalysed formation
of the corresponding p-xylene cis-dihydrodiol (B, R = R¢ = Me,
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Scheme 3 Dioxygenase-catalysed oxidation of phenols 1c, 1f–k, to yield cyclohexenone cis-diols 6cS , 6fS –kS , cyclohexene cis-triols 12fS –kS , cyclohexane
cis-triols 11a and 11j, catechols 3c, 3f–k, hydroxycarboxylic acid 15g–i,15k, and picolinic acid 16i.
The enantiopurity of each of the metabolites 6gS–iS was
established as ≥98% ee by formation of chiral boronate deriva-
tives, using (R)- and (S)-2-(1-methoxymethyl)benzene boronic
acid (MEBBA) and 1H-NMR analysis as previously reported
selected as substrate since it possesses a readily replaceable iodine
atom. In addition to the major product, cyclohexenone cis-diol
6fS, bioproducts 10a6 and 11a6 and triol 12fS were also isolated
as minor metabolites. Compounds 10a and 11a were assumed to
be formed via the cyclohexanone intermediate 9a which could
only be detected by LC-TOF/MS. The metabolite 12fS was fully
characterized using NMR spectroscopy and other methods. The
absolute configuration shown in Scheme 3 was determined by
stereochemical correlation via hydrogenolysis/hydrogenation to
yield triol 11a of established absolute configuration. LC-TOF/MS
analysis of the crude culture medium resulting from biotransfor-
mation of phenols 1g–k with P. putida UV4 detected the presence
of the corresponding triol metabolites 12g–k. Only the triol
diastereoisomers 12fS, 12iS and 12jS, were obtained in sufficient
quantity and purity for complete characterization. The relative
cis stereochemistry of the three chiral centres in each of the triols
12fS, 12iS and 12jS, was determined by NMR spectroscopy. The
pseudo axial proton on C3 exhibited a strong geminal coupling
and two strong diaxial couplings, thereby establishing that the
hydroxyl groups on C2 and C4 were diequatorial and, hence,
confirmed the relative stereochemistry. It is thus presumed that all
the triol metabolites, 12g–i, are likely to possess similar relative and
absolute configurations. Chemoenzymatic synthesis studies, using
the corresponding, more readily available, cis-dihydrodiols (B, R¢ =
H, Scheme 1) are in progress and should provide adequate samples
of all the triols for unequivocal confirmation of their structures
and stereochemistry. It was assumed that each of the minor
6
for the cyclohexenone cis-diols 6cS or 6fS and for standard
arene cis-dihydrodiols.7a–c The absolute configuration of the iodo-
substituted cyclohexenone cis-diol 6fS had earlier been assigned
as (4S,5S) on the basis of X-ray crystallographic analysis and
was stereochemically correlated with the (4R,5S) configuration of
the parent cyclohexenone cis-diol 6aS following hydrogenolysis.6
Recent studies, employing both experimental and calculated
circular dichroism spectra, have shown that cyclohexenone cis-
diols 6aS, 6cS, 6gS–iS all have similar bisignate CD spectra, with
positive Cotton effects at shorter wavelengths and negative Cotton
effects at longer wavelengths, indicative of identical absolute
(4R,5S) configurations.8
Initial analysis of the bioproducts in the crude aqueous culture
medium obtained from P. putida UV4 biotransformations of
m-phenols 1c, 1f–k, based on the HPLC/ESI-TOF/MS (LC-
TOF/MS) method, provided further evidence of the alkyl-
substituted cyclohexenone cis-diols 6cS, 6fS–kS, being formed as
the most significant metabolites prior to extraction (Table 1).
Although this method was not found to be suitable for the
detection of catechols 3, which were only identified after extraction
and NMR analysis, it resulted in the detection of several other
minor metabolites (Scheme 3). This prompted a large-scale
biotransformation of the meta-substituted phenol 1f, which was
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triol metabolites, 12fS–kS, was a second generation metabolite
derived from the corresponding enantiopure cyclohexenone cis-
diols 6fS–iS. This premise was confirmed when compounds 6fS
and 6iS, were used as substrates for P. putida UV4 and yielded the
corresponding triols 12fS and 12iS as the major metabolites; it also
confirmed that a carbonyl reductase (CRED) was present in this
organism.
In addition to the cyclohexenone cis-diols, 6gS–kS, and triols,
12gS–kS, obtained from biotransformation of the phenols 1g–k
and 1k in P. putida UV4 cultures, LC-TOF/MS analysis revealed
a third family of bioproducts. These were tentatively identified as
carboxylic acids, 15g–i and 15k, based on the parent ion data ([M +
H]+, [M + Na]+,[M + NH4]+, [M + K]+ or [MH - H2O]+) and their
relative retention times (Table 1). Although compounds 15g–i and
15k, (Scheme 3), appeared to be new metabolites, bioproduct 15h
and several other carboxylic acids (15, R = n-Pr, n-Bu and iso-Bu)
had been observed as metabolites of the parent alkylbenzenes
using wild-type Pseudomonas strains.9a–9d Carboxylic acid 15k
was not isolated as a bacterial metabolite from biphenyl, but its
immediate metabolic precursor 14k was identified.9d In our studies
only carboxylic acid 15i was obtained in sufficient quantity for full
structure determination by spectroscopic methods.
It was assumed that each of the carboxylic acids 15g–i and
15k resulted from ring-opening of the corresponding catechol
metabolites 3g–i and 3k. An extradiol catechol dioxygenase, CDO,
appeared to be responsible for this oxidative ring-opening process
leading to the undetected conjugated ketocarboxylic acids 13g–i
and 13k, via the corresponding lactone intermediates.10a,b Earlier
work9a–d had shown that the next step involved an ene reductase
(ERED)-catalysed hydrogenation of an alkene group to give the
corresponding non-conjugated diketocarboxylic acids; a similar
metabolic sequence is proposed for the formation of diketocar-
boxylic acids 14g–i and 14k. This ERED-catalysed hydrogenation
was similar to the conversion of cis-diol 6aS into cyclohexane
cis-triol 11a. CRED-catalysed ketone reduction, similar to the
conversion of keto cis-diols 6 into triols 12, occurred in the
final biosynthetic step to give the carboxylic acids 15g–i and
15k. Further metabolism of cyclohexenone cis-diols, 6gS–iS to
give triols, 12g–i, catechols 3g–i, 3k and then carboxylic acids,
15g–i and 15k, may explain the relatively low but variable yields
obtained. The presence of these carboxylic acids also provides
indirect evidence of catechols 3g, 3h, 3i as precursors of value, since
the LC-TOF/MS analysis protocol used in his study was found to
be unsuitable for the detection of catechols in general. This study,
has extended the known range of carboxylic acid metabolites (15,
R = Et, n-Pr, iso-Pr, n-Bu, iso-Bu, tert-Bu, Ph)9a–d derived from the
corresponding catechols 3.
An authentic sample of picolinic acid derivative 16i, available
from independent studies allied to LC-TOF/MS analysis, was
used to confirm the presence of trace amounts of this acid, as
a further metabolite of catechol 3i (Scheme 3). Earlier work on
the biotransformation of a series of monosubstituted benzenes,
e.g. diphenylacetylene,11a and biphenyl (A, R = Ph, R¢ = H),11b
involving the sequence: (i) TDO-catalysed cis-dihydroxylation (to
give cis-dihydrodiols B, R = Ph, R¢ = H), (ii) DD-catalysed
dehydrogenation (to form catechols, C, R = Ph, R¢ = H), (iii)
CDO-catalysed synthesis (to yield ketocarboxylic acids, F, R =
Ph, R¢ = H) and (iv) non-enzymatic cyclization as the final step to
yield picolinic acids (H, R = Ph, R¢ = H, Scheme 1).
The presence of extradiol catechol dioxygenase (CDO), ene
reductase (ERED), and carbonyl reductase (CRED) and other
enzymes in whole cells of P. putida UV4, would be expected to
significantly reduce the isolated yields of both catechols C and
derived metabolites, e.g. G and H (Scheme 1). Thus, the relative
proportions of these metabolites, as determined by LC-TOFMS or
NMR analysis, were often present only in trace amounts (Table 1).
Earlier experiments had shown that m-phenols bearing bulky
substituents, e.g. tert-Bu or I, gave better yields of cyclohexenone
cis-diols. Phenols 1j and 1k, having large CF3 and Ph groups,
were therefore selected for further study as substrates for P. putida
UV4. 3-Trifluoromethylphenol 1j, on biotransformation, gave
the expected enantiopure cyclohexenone cis-diol 6jS but only as
a minor metabolite after column chromatography (4% yield).
A second more polar fraction proved to be a mixture of two
triol metabolites (16% yield) that was then separated by multi-
elution preparative layer chromatography (PLC) with MeOH–
CHCl3 (1 : 5) into triols 12jS and triol 11j; the structure, relative
and absolute stereochemistry of each triol was assigned by NMR
spectroscopy. The proportion of the major triol 12jS, relative
to keto cis-diol 6jS, was much higher than that observed for
other phenol substrates (Table 1). This observation suggests that
a ketone bearing a vinylogous CF3 group [-CH C(CF3)-] in
metabolite 6jS is more readily reduced by a CRED from P. putida
UV4 compared with the other substituted vinyl ketones (6cS, 6fS–
iS).
The metabolic pathway (1 → 6 → 12) shown in Scheme 3 is an
example of a tandem redox biotransformation i.e. a TDO/CRED-
catalysed cis-dihydroxylation/ketone reduction of phenols 1. An
earlier biotransformation of a,a,a-trifluoromethyl acetophenone
(CF3COPh) using P. putida UV4 resulted in the reverse redox tan-
dem sequence, i.e. a CRED/TDO-catalysed ketone reduction/cis-
dihydroxylation, indicative of the facile reduction of the ketone
bearing a CF3 group.12 Only trace amounts of catechol 3j, but no
evidence of the carboxylic acid 15j, were detected by LC-TOF/MS
(Table 1).
3-Phenylphenol 1k, proved to be among the most interesting of
the m-phenol substrates; it provided the TDO enzyme present in
P. putida UV4 with the option of catalysing cis-dihydroxylation
of either the monosubstituted phenyl ring or the disubstituted
m-phenol ring. m-Disubstituted benzenes are normally much
poorer substrates for TDO.5a Thus, we anticipated preferential
cis-dihydroxylation of the monosubstituted ring in compound
1k to give cis-dihydrodiol 17. Surprisingly, more cyclohexenone
cis-diol 6kS was formed relative to metabolite 17 in a readily
separable mixture (Scheme 4, Table 1). Triol 12k was only
detected by LC-TOFMS as a very minor metabolite (<1%).
This observation suggests that a substrate with a m-phenol ring
presents a more attractive cis-dihydroxylation site for TDO than
its monosubstituted phenyl group. 1H-NMR analysis of MEBBA
derivatives was used to determine the ee values of cis-diols 6kS and
17 and, as anticipated, they were both enantiopure (≥98% ee).
The absolute configuration of cis-dihydrodiol 17 was deter-
mined by comparison of its CD spectrum with that of the cis-
dihydrodiol of biphenyl 18. Due to the presence of the additional
phenyl chromophore in compound 6kS, determination of the
absolute configuration by comparison of its CD spectrum with
those of cyclohexenone cis-diols 6aS, 6cS, 6f–6iS, was considered
to be less reliable. However, as part of a separate study designed
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Scheme 4 Dioxygenase-catalysed oxidation of phenol 1k to yield cis-dihydrodiol 17 and cyclohexenone cis-diol 6kS with absolute configuration
assignment by stereochemical correlation with cis-dihydrodiol 18.
to provide synthetic routes to keto cis-diols, e.g. 6kS, and triols,
e.g. 12k, from the corresponding arene cis-dihydrodiols (B, R¢ =
H, Scheme 1), it was possible to provide unequivocal confirmation
of its absolute configuration via stereochemical correlation with
the biphenyl cis-dihydrodiol metabolite 18 (unpublished data).
As shown in Table 1, other very minor metabolites from 3-
hydroxybiphenyl 1k, tentatively identified, included catechol 3k
(detected by NMR spectroscopy), and the derived carboxylic acid
metabolite 15k (detected by LC-TOF/MS).
It is noteworthy that biotransformation of the six m-phenols
1f–k in P. putida UV4 generally yielded the corresponding cyclo-
hexenone cis-diols 6fS–kS, as major metabolites and triols, 12f–k,
as minor bioproducts based on LC-TOF/MS analyses (Table 1).
No evidence of the corresponding triol 12cS or carboxylic acid
metabolites 15c or 15j was observed.
(ii) other types of arene dioxygenases can also catalyse this cis-
dihydroxylation process
(iii) the strong preference shown by TDO for m-phenols could be
rationalized in terms of favourable substrate binding interactions
at the active site
As part of the study, a new E. coli recombinant strain bearing
the TDO genes was constructed. The X-ray crystal structure of
TDO derived from P. putida F1 has recently been reported14 and
a cloned TDO obtained from P. putida UV4 has now been shown
to be identical to the P. putida F1 enzyme. A plasmid, pCL-4t,
containing all four tod-genes from P. putida UV4, was constructed
using a pBAD-ET vector. E. coli TOP10 cells were transformed
with the pCL-4t plasmid to produce an E. coli TOP10 (pCL-4t)
clone. The complete sequence of the cloned P. putida UV4 TDO
ISPTOL large and small subunits, is exactly equivalent to the tod C1
and C2 components respectively from P. putida F1.15
The relatively low but variable yields of the initial cyclohexenone
cis-diol bioproducts, 6fS–kS, obtained could be accounted for by
several factors. These include:
(i) the toxicity of phenolic substrates which are often used as
antimicrobial agents
The E. coli TOP10 (pCL-4t) recombinant strain, hereafter
described as E. coli CL-4t, was tested with the m-phenols 1c, 1f–
k as substrates. In each case the corresponding cyclohexenone
cis-diol, 6cS–kS, was found to be present using LC-TOF/MS
analysis. The isolated yields of cyclohexenone cis-diols obtained
using the clone, were however, consistently lower than those found
using P. putida UV4 under the reaction conditions used. We
speculate that this may reflect the increased toxicity of the phenolic
substrates 1c, 1f–k against the E. coli host cell. In addition, the
(ii) the competition from alternative metabolism routes of
phenols, particularly formation of catechols 3
(iii) the inhibition effect of catechols 3 on the toluene dioxyge-
nase enzyme13
(iv) the CDO-catalysed ring opening of catechols 3 to yield
ketocarboxylic 15 and picolinic acids 16 prior to mineralization
(v) the ERED-catalysed hydrogenations of cyclohexenone cis-
diols 6 to yield cyclohexanone cis-diols 9 and CRED-catalysed
ketone reductions of both cyclohexenone cis-diols 6 and cyclohex-
anone cis-diols 9 to yield triols 12 and 11.
The CRED-catalysed reduction of the keto group to yield
cyclohexene triols 12 and ERED/CRED-catalysed hydrogena-
tion/reduction to yield cyclohexane cis-triols 11 have been iden-
tified as further steps in the biodegradation sequence of the new
family of cyclohexenone cis-diol metabolites 6. The possibility
that cis-diol dehydrogenases, already known to be present in
the environment, can also biodegrade this new family of phenol
metabolites remains to be investigated.
13
known inhibition effect of some catechols on TDO, possibly
including catechols 3c, 3f–k, would be much stronger in E. coli CL-
4t compared with P. putida UV4, since they would not be readily
removed in the absence of the extradiol catechol dioxygenase
enzyme. Significantly, this result confirms unequivocally that the
same TDO enzyme, known to be present in P. putida F1 and now
P. putida UV4, was responsible for the formation of cyclohexenone
cis-diols 6. In contrast to the earlier biotransformations of the m-
phenols, 1f–k, using P. putida UV4, no evidence of the formation
of triol 12 or carboxylic acid 15 was found using E. coli CL-4t. This
was consistent with the CRED, responsible for ketone reductions,
and the ring opening CDO enzyme in P. putida UV4 both being
absent in the recombinant strain.
The question of what other types of dioxygenase enzyme could
catalyse the cis-dihydroxylation of phenol 1, to yield the cis-diols 6
or 17, was addressed using four dioxygenase-containing bacterial
strains (Table 2). These were P. putida UV4 (a constitutive mutant
source of TDO), E. coli CL-4t (a recombinant source of TDO),
P. putida 9816/11 (an inducible mutant source of naphthalene
dioxygenase, NDO) and Sphingomonas yanoikuyae B8/36 (an
inducible mutant source of biphenyl dioxygenase, BPDO). 3-
Hydroxybiphenyl 1k was used as substrate since earlier studies
(b) Enzymes responsible for the production of cyclohexenone
cis-diols 6 from the corresponding m-phenols 1
Preliminary results obtained during the biotransformation of m-
phenols 1 needed further work to establish if:
(i) TDO is the enzyme responsible for the formation of the new
family of cyclohexenone cis-diols 6 in P. putida UV4
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Table 2 Relative ratiosa of metabolites 6kS and 17 obtained via biotrans-
members of this new family of metabolites. When the large number
of anthropogenic or natural product phenols, and phenolic
metabolites resulting from direct aromatic hydroxylation,1 dehy-
dration of cis-dihydrodiols,18a or isomerisation of arene oxides,18b
are combined with the diverse range of bacterial strains containing
arene dihydroxylating dioxygenases, present in the environment, it
is anticipated that a much larger family of cyclohexenone cis-diols
will be identified in the future.
In light of the recently available X-ray crystal structure of
TDO,14 it was timely to re-examine the earlier observation that
meta-disubstituted benzene substrates were generally found to
be poorer substrates for TDO compared with mono-substituted,
ortho-disubstituted and para-disubstituted benzenes (meta-effect).
A representation of the natural substrate, toluene, within the active
site is shown in Fig. 1. This image has been generated from the
published X-ray crystal structure of the enzyme14 associated with
the toluene substrate, first described in the same reference, using
the PDB database Ligand Explorer (see www.pdb.org; database
file code is 3EN1). Coordination of the substrate in the active site
is a result of many hydrophobic interactions between the arene
and active site amino acid residues. In Fig. 1 only interactions
formation using different dioxygenase enzymes
Dioxygenase
Cyclohexenone-cis diol 6kS
cis-Dihydrodiol 17
TDO (UV4)
TDO (CL-4t)
NDO
61
63
48
29b
39
37
52
71b
BPDO
a Based on peak areas using LC-TOF/MS analysis. b The corresponding
triol 12k was also observed as a minor metabolite.
had shown that biaryls, e.g. biphenyl16 or phenyl pyridines,17 were
the only types of monocyclic arenes acceptable to each of these
dioxygenases (Table 2).
LC-TOF/MS analysis showed that in each case a mixture of the
cis-diols 17 and 6kS was formed (Table 2). Whilst the protonated
molecular ion, (M + H)+ at m/z 205 was readily detected for
the cyclohexenone cis-diol 6kS, the corresponding ion for the
dihydrodiol 17 was not. The sodiated molecular ion, (M + Na)+,
at m/z 227 was strong for both metabolites 6kS and 17, and in all
cases this allowed the relative ratios to be estimated from their peak
areas (calibrated using pure samples of diols 6kS and 17 of known
concentration) in the crude aqueous culture medium (Table 2).
It should be emphasised that, while cis-diols 6kS and 17 were
detected as metabolites using all three enzymes (TDO, NDO and
BPDO), the strongest preference (61–63%) for cis-dihydroxylation
of the phenolic ring was found with TDO. Thus, the relative yields
of cyclohexenone cis-diol 6kS, produced by individual strains
under different culture conditions, decreased in the sequence
TDO > NDO > BPDO.
˚
with calculated bond lengths below 1.7 A are shown.
The binding site consists of an ellipse-shaped substrate pocket
lined by 17 mainly hydrophobic residues. The ortho and meta
carbon atoms on one side of the bound toluene substrate are
proximate to the mononuclear non-heme iron atom. The toluene
structure is held within the active site by a series of Van der
Waals and hydrophobic interactions, including possibly edge-to-
face bonding, to the surrounding amino acids bearing non-polar
phenyl or alkyl groups, e.g. Phe 366, Phe 216, Ile 32. This spatial
arrangement is fully consistent with the regio- and stereo-selective
delivery of two Fe-bound oxygen atoms at the 2,3-bond and to
only one face of the benzene ring. Of particular note, in the
context of this study, is the proximity of the positively charged
imidazole ring of His 311 to the meta position of the toluene
In the preliminary communication6 we reported that the TDO
enzyme present in P. putida UV4 was responsible for the formation
of these previously undetected but stable intermediate cyclo-
hexenone cis-diol metabolites 6. This study has now demonstrated
that other bacterial strains and dioxygenases, can yield more
Fig. 1 Toluene bound at the active site of TDO showing all its hydrophobic interactions with amino acid residues with bond distances calculated below
˚
3.7 A.
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substrate furthest from the dihydroxylation site. The basicity of
the imidazole ring would then result in an attractive interaction
with an acidic phenolic OH group at a meta position and similarly
a repulsive interaction when non-hydroxylic meta-alkyl groups are
present.
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. CD spectra were recorded in
spectroscopic grade acetonitrile using a JASCO J-720 instrument.
A PerkinElmer 341 polarimeter was used for optical rotation
Precedents for histidine-ligand hydrogen bonding/electrostatic
interactions have also been revealed through analysis of X-
ray crystal structures of the active sites for other enzyme-
phenolic ligand complexes. These include (i) a catechol dioxy-
genase with pyrogallol,19a (ii) a toluene monooxygenase with 4-
bromophenol,19b (iii) a para-hydroxyphenyl acetate hydroxylase
with the natural substrate,19c and (iv) 1,2-dihydroxynaphthalene
dioxygenase with 4-methylcatechol.19d A role for histidine residues
in coordinating phenolic substrates in TDO, may provide an
explanation for this preference for meta-disubstituted, and to a
lesser extent ortho-disubstituted6 benzene substrates bearing a
phenolic hydroxyl group, and for the production of cyclohexenone
cis-diols 6.
While X-ray crystallography shows that there are many simi-
larities between the active sites of TDO, BPDO and NDO there
are also significant differences. Thus the histidine (His 311) in
TDO is replaced by an aspartic acid (Asp 204) in BPDO and
by an asparagine (Asn 297) in NDO. The stronger attractive H-
bonding interaction between the m-phenolic OH group and the
proximate imidazole ring of His 311 in TDO could explain the
higher proportion (62%) of cyclohexenone cis-diol 6kS found.
These attractive interactions would be weaker with proximate Asn
297 and Asp 204 amino acids and are consistent with the lower
proportions (48% and 29% respectively, Table 1) of keto cis-diol
6kS observed.
◦
([a]D) measurements (ca. 20 C, 10-1 deg cm2 g-1). 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. m-Phenol substrates 1c, 1g–k,
were purchased from Aldrich and used as received. Authentic
samples of catechol bioproducts derived from the corresponding
cis-dihydrodiols and two cyclohexenone cis-diols 6cS and 6fS were
available from earlier studies.6,20
Liquid chromatography-time of flight mass spectrometry (LC-
TOF/MS) 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.
Conclusion
TDO-catalysed biotransformation of the m-phenols 1c, 1f–k,
using P. putida UV4, yielded the enantiopure cyclohexenone cis-
diols 6cS, 6fS–kS, which were structurally and stereochemically
characterized. CRED-catalysed reductions of the cyclohexenone
cis-diols, 6fS–jS, to yield the corresponding cyclohexene cis-
triols, 12f–j, and ERED/CRED-catalysed reductions to give
cyclohexane cis-triols, 11a and 11j, appeared to be further steps
in the biodegradation of phenols by P. putida UV4. Other
phenol metabolites identified by LC-TOF/MS analysis include
the corresponding catechols 3, a-hydroxyacids 15 and picolinic
acids 16.
The ability of three different dioxygenases, TDO, NDO and
BPDO, to catalyse the cis-dihydroxylation of both the phenyl and
phenol rings in 3-hydroxybiphenyl 1k has been established. The
role of the TDO enzyme in the production of cyclohexenone cis-
diols 6 has been confirmed by the use of a new E. coli recombinant
strain over-expressing this enzyme. The ability of TDO to accept
meta-substituted phenols as substrates, in preference to ortho- and
para-phenols, may be rationalized interms oftheir optimalbinding
and catalysis within the active site, facilitated by binding between
the phenolic OH group and a proximate histidine amino acid.
Construction of recombinant biocatalyst strain
A derivative of the arabinose PBAD promoter vector pBAD24 was
used.21 This vector, pBADET (obtained from Dr V Ksenzenko),
has a translation start codon within a unique NdeI restriction
site. Primers were designed for cloning of TDO genes from
Pseudomonas putida F1 (Accession number for the DNA sequence
used is J04996). Forward primer (611–636 nt): 5¢-GAG AAG
CAT ATG AAT CAG ACC GAC ACA TC-3¢ and Reverse
primer (4199–4229 nt): 5¢-GCG AAT TCG CCT TCA AGT
CTC AGC TTA GGT C-3¢ NdeI restriction site was introduced
into the 5¢- part of the forward primer and EcoRI site into the
reverse primer (sites are underlined). A DNA fragment amplified
with these primers included all four TDO genes: todC1, todC2,
todA and todB. PCR amplification was performed with PfuTurbo
polymerase obtained from Stratagene. Reaction conditions were
as follows: 95 ◦C for 3 min and then 32 cycles of 95 ◦C (30 s), 55
◦C (30 s) and 72 ◦C (4.5 min). Fragments were purified using PCR
DNA and a Gel Band Purification kit (Amersham), and digested
with the corresponding enzymes. The PBAD-ET DNA fragment
was digested with both NdeI and EcoRI restriction enzymes prior
to cloning. Ligated DNA was transformed into E. coli TOP10
cells (Invitrogen). Transformation mixtures were plated onto 2YT
media containing ampicillin (100 mg ml-1) and arabinose (0.02%).
Colonies were analysed after incubation at 30 ◦C for 24–30 h.
Experimental
1H and 13C NMR spectra were recorded on Bruker Avance 400,
DPX-300 and DRX-500 instruments. Chemical shifts (d) are
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The E. coli TOP10 clones containing TDO genes were initially
selected by their ability to convert indole to indigo and then the
activity of TDO was confirmed in biotransformation experiments.
The pCL-4t plasmid, was constructed with the complete TDO
sequence from P. putida UV4. The cloned genes were under a tight
regulation by the PBAD promoter (Fig. 2) that allowed efficient
modulation of its expression by addition of an inducer (arabinose)
to cultivation media. E. coli TOP10 cells containing this plasmid
were used in all subsequent biotransformation experiments.
triol 11a have already been reported6 and the fourth, iodo-1,2,3-
triol 12fS, isolated after PLC, was a new bioproduct.
(1S,2S,4R)-6-Iodo-cyclohex-5-ene-1,2,3-triol
12fS. White
crystals (ca. 2% yield), m.p. 102 ◦C (EtOAc); Rf (0.48, EtOAc);
[a]D -60 (c 0.85, MeOH); LC-TOF/MS: Retention time 7.08 min;
m/z obs. 256.9681 calcd. 256.9669 for [M + H]+; m/z obs.
278.9480 calcd. 278.9489 for [M + Na]; HRMS (EI): Found: [M -
H2O]+ 237.9505 requires C6H7O2I 237.9491; 1H-NMR (500 MHz,
CD3OD) d 6.49 (1H, td, J = 2.2, 1.1 Hz, 5-H), 4.13 (2H, m, 1-H,
4-H), 3.69 (1H, ddt, J = 12.4, 5.4, 3.8 Hz, 2-H), 2.04 (1H, m, J =
12.3 Hz, 3-H), 1.75 (1H, dddd, J = 12.1, 12.1, 9.9, 2.2 Hz, 3-Ha);
13C-NMR (125 MHz, CD3OD) d 146.2, 101.7, 77.1, 69.8, 68.6,
35.1; ee ≥ 98%.
Fig. 2 Construction of the TDO biocatalyst. Genes encoding subunits
of the F1/UV4 TDO are shown as black arrows. The transcription of the
system is determined by a PBAD promoter and indicated by the white arrow.
(iii) Biotransformation of 3-ethylphenol 1g. After separation
and purification by PLC, enonediol 6gS was isolated as the major
product with two other metabolites, 12g and 15g, in trace amounts.
General procedure
(4R,5S)-4,5-Dihydroxy-3-ethyl-cyclohex-2-enone 6gS. White
silky needles (ca. 60% yield), m.p. 77–78 ◦C (Et2O–hexane); Rf
(0.24, EtOAc–hexane, 3 : 1); [a]D -103 (c 0.94, MeOH); LC-
TOF/MS: Retention time 11.4 min; m/z obs. 157.0856 calcd.
157.0859 for [M + H]+; m/z obs. 179.0678 calcd. 179.0679 for [M +
Na].+; HRMS (EI): Found 156.0797 requires C8H12O3 156.0786;
1H-NMR (500 MHz, CDCl3) d 5.91 (1H, s, 2-H), 4.35 (1H, m,
4-H), 4.24 (1H, m, 5-H), 3.671 (2H, m, 2 x OH), 2.74 (1H, dd, J =
7.3, 16.4 Hz, 6-H), 2.57 (1H, dd, J = 3.3, 16.4 Hz, 6¢¢-H), 2.43 (2H,
m, J = 7.0 Hz, CH2Me), 1.14 (3H, t, J = 7.0 Hz, CH2Me); 13C-
NMR (125 MHz, CDCl3) d 198.3, 165.1, 125.5, 69.6, 69.1, 42.4,
27.4, 11.5; IR (KBr) nmax/cm-1 1664 (a,b unsaturated ketone); ee ≥
98%.
Small scale (0.2–4.00 g) shake flask biotransformations, with the
whole cells of P. putida UV4 (TDO), P. putida 9816/11 (NDO),
S. yanoikuyae B8/36 (BPDO) and the E. coli CL-4t recombinant
strains were performed using methods described earlier22–24 for
non-phenolic aromatic substrates. The biotransformation condi-
tions were not optimised for the phenol substrates used in this
study and isolated yields (5–70%) of cyclohexenone cis-diols, 6fS–
jS, varied between different strains and repeat biotransformations.
The aqueous culture medium, obtained after the biotransfor-
mation, was concentrated under reduced pressure at ca. 40 ◦C,
the concentrate repeatedly extracted (EtOAc), and the extract
concentrated under reduced pressure to give the crude mixture
of bioproducts. 1H-NMR spectra of the bioproduct mixtures
were routinely recorded, before further purification. The bio-
products were separated either by flash column chromatography
and/or PLC. The enantiomeric excess (ee) values of metabolites
6cS, 6fS–kS, were indirectly estimated from NMR spectroscopic
analysis of their boronate derivatives. The (-)-(S)- and (+)-(R)-
2-(1-methoxyethyl)benzene boronic acids were synthesised and
used according to the literature method.7a–7c Phenol metabolism
also yielded the corresponding known 2,3-catechols, (1H-NMR
analysis) but these were not investigated further.
6-Ethylcyclohex-5-ene-1,2,3-triol 12g. LC-TOF/MS: Reten-
tion time 11.98 min; m/z obs. 181.08373 calcd. 181.08352 for [M +
Na].+
2- Hydroxyl-6-oxooctanoic acid 15g. LC-TOF/MS: Retention
time 14.42 min; m/z obs. 197.0782 calcd. 197.0783 for [M + Na]+,
m/z obs. 213.0519 calcd. 213.0524 for [M + K]+, 157.0859 calcd.
157.0859 for [MH - H2O].+
(iv) Biotransformation of 3-iso-propylphenol 1h. The crude
mixture of bioproducts yielded enonediol 6hS as the major product
(after PLC) and the following two metabolites, 12h and 15h, in
trace amounts.
Biotransformations of phenols and enone cis-diols using P. putida
UV4
(4R,5S)-4,5-Dihydroxy-3-iso-propylcyclohex-2-enone
6hS.
(i) Biotransformation of m-cresol 1c. Chromatographic sep-
aration and purification from the crude mixture of bioproducts
from substrate 1c, yielded enone cis-diol 6cS (ca. 13% yield) as
reported earlier.6
◦
White crystals (ca. 65% yield), m.p. 39–40 C; Rf (0.24, EtOAc–
hexane, 3 : 1); [a]D -79 (c 1.00, MeOH); LC-TOF/MS: Retention
time 13.62 min; m/z obs. 171.1022 calcd. 171.1016 for [M + H]+;
m/z obs. 193.0841 calcd. 193.0835 for [M + Na].+; HRMS (EI):
Found 170.0943 requires C9H14O3 170.0943; 1H-NMR (400 MHz,
CDCl3) d 5.92 (1H, s, 2-H), 4.42 (1H, dd, J = 3.4, 3.4 Hz, 4-H),
4.28 (1H, dq, J = 7.9, 4.0 Hz, 5-H), 3.50 (1H, d, J = 3.5 Hz, OH),
3.43 (1H, d, J = 3.4 Hz, OH), 2.78 (1H, m, -CHMe2), 2.74 (1H,
m, J = 17.0 Hz, 6-H), 2.57 (1H, dd, J = 3.8, 17.0 Hz, 6¢¢-H), 1.16
(3H, d, J = 6.7 Hz, Me), 1.10 (3H, d, J = 7.0 Hz, Me); 13C-NMR
(100 MHz, CDCl3) d 198.5, 168.8 124.5, 69.6, 68.3, 42.1, 32.4,
21.7, 20.2; IR (KBr) nmax/cm-1 1666 (a,b unsaturated ketone);
ee ≥ 98%.
(4R,5S)-4,5-Dihydroxy-3-methyl-cyclohex-2-enone 6cS. [a]D
-115 (c 1.0, MeOH); LC-TOF/MS: Retention time 4.89 min; m/z
obs. 143.06977 calcd. 143.07027 for [M + H]+; m/z obs. 165.05221
calcd. 165.05222 for [M + Na].+ Spectroscopic data was identical
to that reported.6
(ii) Biotransformation of 3-iodophenol 1f. The crude mix-
ture of bioproducts yielded four compounds. Three of these,
viz, (4R,5S)-4,5-dihydroxy-3-iodocyclohex-2-enone 6fS, (4R)-
hydroxycyclohexenone 10a and (1R,2S,4S)-cyclohexane-1,2,4-
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6-iso-Propylcyclohex-5-ene-1,2,3-triol 12h. LC-TOF/MS: re-
tention time 14.25 min; m/z obs. 195.0982 calcd. 195.0992 for
[M + Na].+
(4R,5S)-4,5-Dihydroxy-3-trifluromethylcyclohex-2-enone 6jS.
Light yellow viscous oil (ca. 4% yield); Rf (0.31, MeOH–CHCl3,
7 : 93); [a]D -65 (c 1.00, MeOH); LC-TOF/MS: Retention time
10.58 min. m/z obs. 197.0425 calcd. 197.0420 for [M + H]+;
m/z obs. 219.0246 calcd. 219.0240 for [M + Na].+; HRMS
(EI): Found: 196.0344 requires C7H7O3F3 196.0347; 1H-NMR
(400 MHz, CD3OD) d 6.41 (1H, s, 2-H), 4.58 (1H, d, J = 3.3 Hz,
4-H), 4.28 (1H, ddd, J = 9.6, 4.1, 3.3 Hz, 5-H), 2.83 (1H, dd,
J = 9.6, 16.8 Hz, 6-H), 2.66 (1H, dd, J = 4.1, 16.8 Hz, 6¢¢-H);
13C-NMR (100 MHz, CD3OD) d 198.7, 146.9 (q, J = 30.8 Hz),
130.9 (q, J = 5.2 Hz), 124.4 (q, J = 272.4 Hz, CF3), 69.7, 65.9,
42.7; IR(liquid film) nmax/cm-1 1694 (a,b unsaturated ketone);
ee ≥ 98%.
2- Hydroxyl-7-methyl-6-oxooctanoic acid 15h. LC-TOF/MS:
Retention time 15.37 min; m/z obs. 189.1125 calcd. 189.1121 for
[M + H]+, m/z obs. 211.0933 calcd. 211.0941 for [M + Na]+, m/z
obs. 206.1390 calcd. 206.1387 for [M + NH4]+, m/z obs. 227.0680
calcd. 227.0676 for [M + K]+, 171.1018 calcd. 171.1016 for [MH -
H2O]+.
(v) Biotransformation of 3-tert-butylphenol 1i. The crude mix-
ture of bioproducts yielded enonediol 6iS as the major product and
its corresponding triol 12iS and a-hydroxyacid 15i as the minor
bioproducts. All proved to be separable using PLC.
(1R,2S,4R)-6-Trifluoromethylcyclohex-5-ene-1,2,4-triol
12jS.
◦
White crystals (ca. 6% yield), m.p. 144–45 C (EtOAc); Rf (0.46,
MeOH–CHCl3, 1 : 4); [a]D -29 (c 0.68, MeOH); LC-TOF/MS:
Retention time 9.21 min; m/z obs. 221.0383 calcd. 221.0396 for
[M + Na]+; m/z obs. 216.0831 calcd. 216.0842 for [M + NH4].+;
HRMS (EI): Found: [M - H2O]+ 180.0405 requires C7H7O2F3
180.0398; 1H-NMR (500 MHz, CD3OD) d 6.47 (1H, s, 5-H), 4.32
(1H, m, J = 12.3, 2.6 Hz, 4-H), 4.05 (1H, d, J = 3.6 Hz, 1-H),
3.61 (1H, ddd, J = 3.6, 12.3, 3.6 Hz, 2-H), 2.09 (1H, m, J = 3.6,
10.3, 2.6 Hz, 3-H), 1.68 (1H, ddd, J = 10.3, 12.3, 12.3 Hz, 3¢-H);
13C-NMR (100 MHz, CD3OD) d 139.4 (q, J = 5.3 Hz), 130.6 (q,
J = 29.2 Hz), 125.4 (q, J = 270.1 Hz, CF3), 68.4, 66.8, 64.7, 34.5;
ee ≥ 98%.
(4R,5S)-4,5-Dihydroxy-3-tert-butylcyclohex-2-enone
6iS.
Colourless crystals (ca. 68% yield), m.p. 57–58 ◦C (acetone–
hexane); Rf (0.36, EtOAc–hexane, 3 : 1); ee ≥ 98%; [a]D -9 (c
0.8, MeOH); LC-TOF/MS: Retention time 15.41 min; m/z obs.
185.1164 calcd. 185.1172 for [M + H];+ m/z obs. 207.0992 calcd.
207.0992 for [M + Na].+; HRMS (EI): Found 184.1096 requires
1
C10H16O3 184.1099; H-NMR (400 MHz, CDCl3) d 6.02 (1H, s,
2-H), 4.49 (1H, m, 4-H), 4.00 (1H, m, 5-H), 3.11 (1H, m, OH),
2.96 (1H, m, OH), 2.77 (1H, dd, J = 11.4, 16.3 Hz, 6-H), 2.60
(1H, dd, J = 5.3, 16.3 Hz, 6¢-H), 1.21 (9H, s, CMe3); 13C-NMR
(100 MHz, CDCl3) d 199.0, 168.6, 126.0, 69.1, 66.3, 40.4, 36.4,
28.9; IR (KBr) nmax/cm-1 1651 (a,b unsaturated ketone); ee ≥
98%.
(1R,2S,4S,6R)-6-Trifluoromethylcyclohexane-1,2,4-triol
11j.
Colourless viscous liquid (ca. 10% yield); Rf (0.35, MeOH–CHCl3,
1 : 4); [a]D -3.0 (c 1.0, MeOH); LC-TOF/MS: Retention time
5.73 min. m/z obs. 201.0743 calcd. 201.0733 for [M + H]+;
m/z obs. 223.0565 calcd. 223.0553 for [M + Na]+; m/z obs.
(1R,2S,4R)-6-tert-Butylcyclohe◦x-5-ene-1,2,4-triol 12iS. White
crystals (ca. 5% yield), m.p. 107 C (CH2Cl2); Rf (0.57, MeOH–
CHCl3, 1 : 4); [a]D -21 (c 1.25, MeOH); LC-TOF/MS: Retention
time 15.78 min; m/z obs. 209.1148 calcd. 209.1148 for [M + Na]+;
m/z obs. 204.1590 calcd. 204.1594 For [M + NH4]+; HRMS (EI):
1
218.1006 calcd. 218.0999 for [M + NH4]+; H-NMR (400 MHz,
CD3OD) d 3.93 (1H, m, 1-H), 3.54 (1H, tt, J = 11.7, 3.8 Hz, 4-H),
3.41 (1H, ddd, J = 12.0, 4.4, 2.8, 2-H), 2.18 (1, m, 6-H), 1.82
(1H, m, 3-He), 1.72 (1H, m, 5-He), 1.59 (2H, q, 12.0, 3-Ha and
5-Ha); 13C-NMR (100 MHz, CD3OD) d 128.6 (q, J = 278.1 Hz,
CF3), 70.7, 67.8, 67.3 (q, J = 2.7 Hz), 43.4 (q, 26.4), 37.9, 28.5
(q, J = 2.1 Hz).
Found: [M - H2O]+ 168.1161 requires C10H16O2 168.1150; H-
1
NMR (400 MHz, CD3OD) d 5.57 (1H, dd, J = 2.9, 2.9 Hz, 5-H),
4.08 (1H, m, J = 10.2, 2.9, 2.9 Hz, 4-H), 4.05 (1H, dd, J = 6.7,
2.9 Hz, 1-H), 3.11 (1H, ddd, J = 12.2, 2.9, 2.9 Hz, 2-H), 1.90 (1H,
m, J = 10.2, 2.9 Hz, 3-He), 1.68 (1H, ddd, J = 12.2, 12.2, 10.2 Hz,
3¢-Ha); 13C-NMR (100 MHz, CD3OD) d 148.3, 127.8, 70.0, 67.9,
67.3, 35.8, 34.6, 30.3; ee ≥ 98%.
(vii) Biotransformation of 3-phenylphenol 1k. PLC separation
of the mixture of bioproducts yielded enonediol 6kS as the major
compound, dihydrodiol 17 as the minor metabolite and two
metabolites, 12k and 15k, in trace amounts.
2-Hydroxy-7,7-dimethyl-6-oxooctanoic acid 15i. Light yellow
viscous oil (< 1% yield); Rf (0.26, MeOH–CHCl3, 1 : 3); [a]D +8.5
(c 0.41, MeOH); MS (ES-) Found: 202.1128 requires C10H17O4
201.1127; LC-TOF/MS: Retention time 16.98 min; m/z obs.
203.12808 calcd. 203.12779 for [M + H]+, m/z obs. 220.1543 calcd.
220.1543 for [M + NH4]+, m/z obs. 225.1091 calcd. 225.1097 for
[M + Na]+, m/z obs. 241.0840 calcd. 241.0837 for [M + K]+,
(4R,5S)-4,5-Dihydroxy-3-phenylcyclohex-2-enone 6kS. White
crystals (ca. 28% yield), m.p. 83–84 ◦C (CHCl3); Rf (0.40, EtOAc–
hexane, 3 : 1); [a]D -20 (c 0.74, MeOH); LC-TOF/MS: Retention
time 15.52 min; m/z obs. 205.0853 calcd. 205.08592 for [M + H]+;
m/z obs. 227.0672 calcd. 227.0679 for [M + Na].+; HRMS (EI):
Found 204.0788 requires C12H12O3 204.0786; 1H-NMR (500 MHz,
CDCl3) d 7.65 (2H, m, Ar-H), 7.46 (3H, m, Ar-H), 6.42 (1H, s,
2-H), 4.92 (1H, m, 4-H), 4.33 (1H, m, 5-H), 2.88 (1H, dd, J =
9.8, 16.3 Hz, 6-H), 2.72 (2H, m, J = 4.4, 16.3 Hz, 6¢-H and OH),
2.55 (1H, d, J = 5.8, OH); 13C-NMR (100 MHz, CDCl3) d 197.4,
156.1, 136.5, 130.5, 129.1, 127.0, 126.8, 68.7, 68.0, 41.1; IR (KBr)
1
m/z obs. 185.1168 calcd. 185.1172 for [MH - H2O]; H-NMR
(400 MHz, CD3OD) d 3.94 (1H, dd, J = 3.8, 7.1 Hz, 2-H), 2.62
(2H, t, J = 7.3 Hz, 5-H), 1.68 (4H, m, 2 ¥ 3-H, 2 ¥ 4-H), 1.17 (9H, s,
-CMe3); 13C-NMR (100 MHz, CD3OD) d 218.7, 181.6, 73.4, 45.2,
37.6, 35.8, 26.7, 21.0.
Biotransformation of 3-trifluoromethylphenol 1j. The mixture
of bioproducts was separated by PLC and was found to contain
enonediol 6jS as the minor product and the corresponding triols
12jS and 11j as the major metabolites.
n
max/cm-1 1648 (a,b unsaturated ketone); ee ≥ 98%.
(1S,2R)-1,2-Dihydroxy-3-(3¢-hydroxyphenyl)cyclohexa-3,5-
diene 17. White crystals (ca. 10% yield); m.p. 137–38 ◦C
1488 | Org. Biomol. Chem., 2011, 9, 1479–1490
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(EtOAc–hexane); Rf (0.38, EtOAc–hexane, 3 : 1); [a]D +222
(c 1.09, MeOH); LC-TOF/MS: Retention time 15.90 min; m/z
obs. 227.0672 calcd. 227.0679 for [M + Na].+; MS (ES-) Found:
203.0697 requires C12H11O3 203.0708; 1H-NMR (400 MHz,
CD3OD) d 7.05 (1H, dd, J = 7.8, 7.8 Hz, 5¢-H), 6.95 (1H, m,,
J = 7.8, 2.2, 1.0 Hz, 4¢-H), 6.91 (1H, dd, J = 2.2, 1.0 Hz, 2¢-
H), 6.59 (1H, ddd, J = 7.8, 2.2, 1.0 Hz, 6¢-H), 6.27 (1H, d,
J = 6.0 Hz, 4-H), 5.96 (1H, ddd, J = 9.5, 6.0, 2.9 Hz, 5-H),
5.73 (1H, m, J = 9.5, 6.0 Hz, 6-H), 4.41 (1H, m, J = 6.0 Hz,
1-H), 4.26 (1H, d, J = 6.0 Hz, 2-H); 13C-NMR (100 MHz,
CD3OD) d 158.7, 142.2, 139.5, 132.3, 130.5, 125.0, 122.8, 118.0,
115.6, 113.3, 72.7, 69.9; CD (MeCN) l 208 nm (De -11.37), l
212.6 nm (De -9.14), l 229.8 nm (De -13.57), l 320 nm (De -6.34);
ee ≥ 98%.
Acta, 1999, 32, 35; (h) D. T. Gibson and R. E. Parales, Curr. Opin.
Biotechnol., 2000, 11, 236; (i) D. R. Boyd, N. D. Sharma and C. C. R.
Allen, Curr. Opin. Biotechnol., 2001, 12, 564; (j) R. A. Johnson, Org.
Reactions, 2004, 63, 117; (k) D. R. Boyd and T. D. H. Bugg, Org. Biomol.
Chem., 2006, 4, 181; (l) K. A. B. Austin, M. Matveenko, T. A. Reekie
and M. G. Banwell, Chem. Aust., 2008, 75, 3; (m) T. Hudlicky and J. W.
Reed, Synlett, 2009, 685; (n) T. Hudlicky and J. W. Reed, Chem. Soc.
Rev., 2009, 38, 3117.
3 (a) R. C. Bayly, S. Dagley and D. T. Gibson, Biochem. J., 1966, 101,
293; (b) D. T. Gibson, V. Mahadevan and J. F. Davey, J. Bacteriol.,
1974, 119, 930; (c) J. A. Buswell, J. Bacteriol., 1975, 124, 1399; (d) J. C.
Spain and D. T. Gibson, Appl. Environ. Microbiol., 1988, 54, 1077;
(e) F. K. Higson and D. D. Focht, Appl. Environ. Microbiol., 1989,
55, 946; (f) J. C. Spain, G. J. Zylstra, C. K. Blake and D. T. Gibson,
Appl. and Environ. Microbiol., 1989, 55, 2648; (g) C. Hinteregger, R.
Leitner, M. Loidl, A. Ferschl and F. Streichsbier, Appl. Microbiol.
Biotechnol., 1992, 37, 252; (h) G. Bestetti, E. Galli, B. Leoni, F.
Pelizzoni and G. Sello, Appl. Microbiol. Biotechnol., 1992, 37, 260;
(i) K. Lee, FEMS Microbiol. Lett., 2006, 255, 316; (j) D. Kim, J. S.
Lee, K. Y. Choi, Y-S. Kim, J. N. Choi, S-K. Kim, J-C. Chae, G.
Zylstra, C. H. Lee and E. Kim, Enzyme Microb. Technol., 2007, 41,
221.
6-Phenylcyclohex-5-ene-1,2,3-triol 12k. LC-TOF/MS: Reten-
tion time 15.16 min; m/z obs. 229.0838 calcd. 229.0835 for [M +
Na].+
4 D. T. Gibson, J. R. Koch and R. E. Kallio, Biochemistry, 1968, 7,
2653.
2-Hydroxyl-6-phenyl-6-oxohexanoic acid 15k. LC-TOF/MS:
Retention time 19.70 min; m/z obs. 223.0972 calcd. 223.0965 for
[M + H]+, m/z obs. 245.0793 calcd. 245.0784 for [M + Na].+
5 (a) D. R. Boyd, N. D. Sharma, N. I. Bowers, H. Dalton, M. D. Garrett,
J. S. Harrison and G. N. Sheldrake, Org. Biomol. Chem., 2006, 4, 3343;
(b) D. R. Boyd, N. D. Sharma, A. King, B. Byrne, S. A. Haughey, M. A.
Kennedy and C. C. R. Allen, Org. Biomol. Chem., 2004, 2, 2530–2537;
(c) D. R. Boyd, N. D. Sharma, I. N. Brannigan, M. R. Groocock, J. F.
Malone, G. McConville and C. C. R. Allen, Adv. Synth. Catal., 2005,
347, 1081.
6 D. R. Boyd, N. D. Sharma, C. C. R. Allen and J. F. Malone, Chem.
Commun., 2009, 3633.
7 (a) J. Gawronski, M. Kwit, D. R. Boyd, N. D. Sharma, J. F. Malone
and A. Drake, J. Am. Chem. Soc., 2005, 127, 4308; (b) D. R. Boyd,
N. D. Sharma, G. P. Coen, P. Gray, J. F. Malone and J. Gawronski,
Chem.–Eur. J., 2007, 13, 5804; (c) M. Kwit, N. D. Sharma, D. R. Boyd
and J. Gawronski, Chirality, 2008, 20, 609.
(viii)
Biotransformation
of
(4R,5S)-4,5-dihydroxy-3-
iodocyclohex-2-enone 6fS. The mixture of bioproducts obtained
from substrate 6fS (0.100 g), was separated by a combination
of column chromatography (hexane → EtOAc → 10% MeOH
in EtOAc) and PLC (EtOAc–hexane, 1 : 1) to give the reported
metabolites (+)-(R)-4-hydroxycyclohex-2-enone 10a (0.011 g)6,
(-)-(1R,2S,4S)-cyclohexane-1,2,4-triol 11a (0.015 g)6 and (-)-
(1S,2S,4R)-6-iodo-cyclohex-5-ene-1,2,3-triol 12fS (0.018 g)
having identical properties to those obtained using phenol 1f as
substrate.
8 M. Kwit, J. Gawronski, D. R. Boyd, N. D. Sharma and M. Kaik, Org.
Biomol.. Chem., 2010, 8, 5635.
9 (a) Y. Jigami, T. Omori and Y. Minoda, Agric. Biol. Chem., 1974, 38,
1757; (b) Y. Jigami, T. Omori and Y. Minoda, Agric. Biol. Chem., 1975,
39, 1781; (c) Y. Jigami, Y. Kawasaki, T. Omori and Y. Minoda, Appl.
Environ. Microbiol., 1979, 38, 783; (d) T. Omori and H. Ishigooka,Y.
Minoda, Agric. Biol. Chem., 1988, 52, 503.
10 (a) T. D. H. Bugg, Tetrahedron, 2003, 59, 7075; (b) T. D. H. Bugg and
S. Ramaswamy, Curr. Opin. Chem. Biol., 2008, 12, 134.
11 (a) J. C. Spain, S. F. Nishino, B. Witholt, L-S. Tan and W. A. Duetz,
Appl. Environ. Microbiol., 2003, 69, 4037; (b) K. Shindo, A. Osawa, R.
Nakamura, Y. Kagiyama, S. Sakuda, Y. Shizuri, K. Furukawa and N.
Misawa, J. Am. Chem. Soc., 2004, 126, 15043.
12 D. R. Boyd, N. D. Sharma, V. Ljubez, J. F. Malone and C. C. R. Allen,
J. Chem. Technol. Biotechnol., 2007, 82, 1072.
(ix) Biotransformation of (4R,5S)-4,5-dihydroxy-3-tert-
butylcyclohex-2-enone 6iS. The only compound obtained after
PLC of the crude mixture from substrate 6iS (0.030 g) was
identified as (-)-(1R,2S,4R)-6-tert-butylcyclohex-5-ene-1,2,3-triol
12iS (0.02 g). This product was indistinguishable from the sample
produced earlier using the phenol 1i.
Acknowledgements
We thank Professors Tim Bugg and Jim Spain for helpful
discussions, Stewart Floyd for assistance with the LC-TOF/MS
analyses and Peter Gray for microbiological support. Financial
support was provided by the Department for Education and
Learning NI (HM) and the European Social Fund (JMA).
13 G. K. Robinson, G. M. Stevens, H. Dalton and P. J. Geary, Biocatal.
Biotransform., 1992, 6, 81.
14 R. Friemann, K. Lee, E. N. Brown, D. T. Gibson, H. Eklund and S.
Ramaswamy, Acta. Cryst., 2009, D65, 24.
15 G. J. Zylstra and D. T. Gibson, J. Biol. Chem., 1989, 264,
14940.
16 R. E. Parales, S. M. Resnick, C. L. Yu, D. R. Boyd, N. D. Sharma and
D. T. Gibson, J. Bacteriol., 2000, 184, 5495.
17 D. R. Boyd, N. D. Sharma, G. P. Coen, F. Hempenstall, V. Ljubez, J. F.
Malone, C. C. R. Allen and J. T. G. Hamilton, Org. Biomol. Chem.,
2008, 6, 3957.
18 (a) D. R. Boyd, J. Blacker, B. Byrne, H. Dalton, M. V. Hand, S. Kelly,
R. A. MoreO’Ferrall, S. N. Rao, N. D. Sharma, G. N. Sheldrake
and H. Dalton, J. Chem. Soc., Chem. Commun., 1994, 313; (b) D. R.
Boyd, N. D. Sharma, J. S. Harrison, J. F. Malone, W. C. McRoberts,
J. T. G Hamilton and D. B. Harper, Org. Biomol. Chem., 2008, 6,
1251.
19 (a) I. Matera, M. Ferraroni, M. Kolomytseva, L. Golovlova, A.
Scozzafava and F. Briganti, J. Struct. Biol., 2010, 170, 548 PDB
database file code 3HHX;; (b) M. H. Sazinsky, J. Bard,A. Di Donato
and S. J. Lippard, J. Biol. Chem., 2004, 279, 30600 PDB database file
code 1TOS; (c) A. Alfieri, F. Fersini, N. Ruangchan, M. Prongjit, P.
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