Biocatalytic synthesis of chiral cyclic γ-oxoesters by sequential C-H hydroxylation, alcohol oxidation and alkene reduction

Article abstract of DOI:10.1039/c7gc02215h

A three-step biocatalytic procedure is described for the conversion of methyl and ethyl cyclopentene- and cyclohexenecarboxylates into both the enantiomers of the corresponding chiral 3-oxoesters, which are useful building blocks for the synthesis of active pharmaceutical ingredients. The allylic hydroxylation of the starting cycloalkenecarboxylates is carried out by using R. oryzae resting cells entrapped in alginate beads, in acetate buffer solution at 25 °C. The oxidation of the intermediate allylic alcohols to unsaturated ketones, performed by the laccase/TEMPO system, and the ene-reductase mediated hydrogenation of the alkene bond were carried out in the same reaction vessel in a sequential mode at 30 °C. Being entirely biocatalytic, our multistep procedure provides considerable advantages in terms of selectivity and environmental impact over reported chemical methods.

Full text of DOI:10.1039/c7gc02215h

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Page 1 of 28
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DOI: 10.1039/C7GC02215H
Biocatalytic synthesis of chiral cyclic γ-oxoesters by sequential C-H
hydroxylation, alcohol oxidation and alkene reduction
a,
a
b
Elisabetta Brenna,*,a,b Michele Crotti, Francesco G. Gatti, Daniela Monti, Fabio
§
Parmeggiani,a Andrea Pugliesea, Francesca Tentoria
a Politecnico di Milano, Dipartimento di Chimica, Materiali, Ingegneria Chimica, Via Mancinelli 7,
Iꢀ20131 Milano, Italy.
b
Istituto di Chimica del Riconoscimento Molecolare – CNR, Via M. Bianco 9, Iꢀ20131 Milano,
Italy.
§Authors are listed in alphabetical order. M. C. is the principal author of the publication.
1

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Abstract
A threeꢀstep biocatalytic procedure is described for the conversion of methyl and ethyl
cyclopenteneꢀ and cyclohexenecarboxylates into both the enantiomers of the corresponding chiral
3ꢀoxoesters, which are useful building blocks for the synthesis of active pharmaceutical ingredients.
The allylic hydroxylation of the starting cycloalkenecarboxylates is carried out by using R. oryzae
resting cells entrapped in alginate beads, in acetate buffer solution at 25°C. The oxidation of the
intermediate allylic alcohols to unsaturated ketones, performed by the laccase/TEMPO system, and
the eneꢀreductase mediated hydrogenation of the alkene bond were carried out in the same reaction
vessel in a sequential mode at 30°C. Being entirely biocatalytic, our multistep procedure provides
considerable advantages in terms of selectivity and environmental impact over reported chemical
methods.
2

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Introduction
The key role played by biocatalysis in improving the sustainability of fine and bulk chemicals
manufacturing is well recognized.1 The capability of enzymes to catalyze a remarkably wide range
of organic reactions with excellent chemoꢀ, regioꢀ, and stereoselectivity under mild conditions
(temperature, pH, pressure), limiting the amount of waste and byꢀproducts, is greatly appreciated,
especially in the pharmaceutical field.2 Furthermore, the combination of multiꢀenzymatic reactions
in a cascade sequence3 represents an attractive approach for the production of complex valuable
chemicals from simple precursors, taking advantage of the fact that, to date, the biocatalytic
counterparts of many classical organic reactions have been established and optimized to work under
similar and compatible operating conditions.4
For example, the oxyfunctionalization of allylic carbon atoms represents a very useful strategy for
converting readily available unsaturated hydrocarbons into high valueꢀadded allylic alcohols and
α,βꢀunsaturated ketones.5 Traditionally, the process has been performed by using stoichiometric
oxidants, e.g. chromium(VI)ꢀbased reagents, manganese dioxide, potassium permanganate or
selenium dioxide. During the last years, the thrust towards more environmentally benign and costꢀ
effective protocols has promoted the search and optimization of catalytic procedures using suitably
activated molecular oxygen or peroxides as oxidants.6 Biocatalysis offers advantageous strategies
for this reaction, based on the versatile oxidative capabilities of (monoꢀ and diꢀ) oxygenases,
peroxygenases and oxidases.7 Several examples of selective allylic CꢀH hydroxylation of steroids,
terpenes and terpenoids have been described in the literature using either wholeꢀcell
microorganisms or isolated enzymes.8
Recently, we envisaged the possibility to employ this reaction as the first step of a multiꢀenzymatic
procedure for the preparation of the enantiomers of cyclic ketoesters 1a,b and 2a,b (Scheme 1)
from readily available cycloalkenecarboxylates 3a,b and 4a,b. Herein, we report on the
development and optimization of a completely biocatalytic and environmentally benign system
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combining a twoꢀstep allylic biooxidation with the biohydrogenation of the oxidized products 5a,b
and 6a,b, for the preparation of both enantiomers of cyclic γꢀoxoesters 1a,b and 2a,b.
These compounds, along with their corresponding ketoacids, are exceptionally useful building
blocks in the synthesis of active pharmaceutical ingredients (APIs), because they make available
rigid cyclic cores bearing two functional handles for further manipulation. Such structures are often
incorporated into the skeleton of drug candidates, in order to investigate the effects due to limited
conformational freedom and reduced rotation around single bonds. Representative examples of
APIs built around such cores are reported in Figure 1.
Scheme 1. Synthetic plan to chiral cyclic ketoesters 1 and 2.
F
F
N
O
O
NH2
N
HOOC
H
N
NH
N
N
N
N
N
(a)
(b)
F
O
O
O
F
F
O
N
COOR
1
F
HO OH
P
R = Me, Et, H
O
Ar
O
O
OH
O
H2N
N
R3
O
OH
(c)
(d)
O
O
4
COOR
O
OR6
2
O
R4 R5
O
O
Figure 1. Structures of active pharmaceutical ingredients incorporating a 1,3ꢀdifunctionalised C5 or
C6 ring: (a) antagonists of the A2A receptor, employed for the treatment of neurological disorders;9
(b) inhibitors of acetylꢀCoA:diacylglycerol acyltransferase activity, employed for the treatment of
type II diabetes;10 (c) PPAR (peroxisome proliferatorꢀactivated receptor) ligands, employed for the
treatment of hyperlipidemia and diabetes;11 (d) leustroducsin B, a potent colonyꢀstimulating factor
inducer isolated from the culture broth of Streptomyces platensis SANK 60191.12
4

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Results and discussion
Known chemical approaches to enantiomerically enriched derivatives 1 and 2
To date, the chemical synthetic procedures that have been developed for enantiomerically enriched
derivatives 1 and 2 include classical resolution and a few enantioselective approaches using metal
or organocatalysts.
The fractional crystallization of diastereoisomeric salts was exploited for both cyclopentane and
cyclohexane derivatives. (R)-3ꢀoxocyclopentanecarboxylic acid was obtained (ee = 98%, 18%
yield) from the corresponding racemate after four crystallisations of the diastereoisomeric salts with
(–)ꢀbrucine from water.13 The (S)ꢀenantiomer had to be recovered from the mother liquors.14 The
crystallisation of (±)ꢀ3ꢀoxocyclohexanecarboxylic acid with (–)ꢀbrucine afforded the (R)ꢀ
enantiomer (23% yield), as well.15
The organocatalytic transfer hydrogenation of the C=C double bond was described only for
compound 5a, prepared by CrO3 oxidation of 3a. (S)ꢀ1a was synthesised (ee = 91%, 78% yield) by
treatment of 5a with tertꢀbutyl Hantzsch ester in diethyl ether solution in the presence of a chiral
amine as a catalyst.16
(R)ꢀ3ꢀoxocyclopentanecarboxylic acid (ee = 95%, 60% yield) and (S)ꢀ3ꢀoxocyclohexanecarboxylic
acid (ee = 93%, 71% yield) were also synthesized by enantioselective rhodiumꢀcatalyzed 1,4ꢀ
addition of lithium 2ꢀfuryltriolborates (prepared at –78°C by alkylation of B(OiPr)3 with
furyllithium followed by ester exchange with 1,1,1ꢀtris(hydroxymethyl)ethane) to the corresponding
easily available cycloalkenones, followed by oxidation of the furyl ring with ozone.17
Another enantioselective approach led to the synthesis of (R)ꢀ3ꢀoxocyclopentanecarboxylic acid (ee
> 99%, 58% yield) by nucleophilic addition of SAMPꢀderived formaldehyde hydrazone to
cyclopentenone promoted by TDSOTf. Ozone was employed to cleave the C=N bond and create the
aldehydic moiety, which was then submitted to Jones oxidation.18
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The enantioselective intramolecular hydroacylation of ethyl 2ꢀmethyleneꢀ5ꢀoxopentanoate
catalyzed by rhodium diphosphine complexes afforded (S)ꢀ1b with ee >99%. The starting substrate
was obtained by Hg2+ catalyzed vinylation of ethyl 2ꢀ(hydroxymethyl)acrylate followed by Claisen
rearrangement.19
The importance of chiral building blocks 1a,b and 2a,b in the field of medicinal chemistry, and the
lack of costꢀeffective, sustainable synthetic routes for these compounds at a preparative scale,
prompted us to investigate the combination of redox enzymes in a sequential procedure to achieve
the conversion of derivatives 3 and 4 into chiral γꢀoxoesters 1 and 2.
Allylic hydroxylation of methyl and ethyl cycloalkenecarboxylates
A survey of chemical literature on the allylic oxidations of esters 3 and 4 highlighted that chemical
reagents afford directly the corresponding unsaturated oxoꢀderivatives 5 and 6. The following
methods have been described: (i) stoichiometric chromium(VI) oxide, with acetic anhydride and
acetic acid in CH2Cl2 at 0ꢀ5°C;20,21 (ii) tꢀbutyl hydroperoxide as stoichiometric oxidant, in the
presence of either 20% Pd(OH)2/C (5 mol%)22 or dirhodium complexes23,24 as catalysts in CH2Cl2
solution. The only example of biocatalysed oxyfunctionalisation is the hydroxylation of derivatives
3a and 4a mediated by suitable mutants of monooxygenase P450ꢀBM3, which was studied by Reetz
et al. with the aim of preparing both enantiomers of allylic alcohols 7a and 8a.25 In a following
work,26 the same authors employed designer cells to achieve a multiꢀenzymatic cascade
transformation to convert substrate 4a into (S)ꢀ and (R)ꢀ2a, by using two different types of
engineered Escherichia coli cells: one producing P450ꢀBM3 mutants, for the regioselective
oxidation of 4a to 6a, and the other producing variants of the eneꢀreductase YqjM, suitable for the
(S)ꢀ or (R)ꢀselective hydrogenation of the alkene bond.
We chose methyl cyclopentenecarboxylate 3a as a model compound to investigate the oxidative
capability of a set of four filamentous fungi, Colletotrichum lini, Rhizopus oryzae, Aspergillus niger
6

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and Alternaria tenuis, which had been described to perform the allylic CꢀH hydroxylation of
steroids and terpenes.27ꢀ30 Screening experiments were carried out by incubating substrate 3a with
the fungal cell cultures, and monitoring the biotransformations by GC/MS analysis. After 5 days,
wellꢀdefined oxygenated metabolites could be identified only in the biotransformation of 3a with R.
oryzae, while the other strains afforded complex mixtures of products. Preliminary experiments
with R. oryzae were performed also on substrates 3b and 4a,b and the results are reported in Figure
2, Procedure A. The identification of the components of each reaction mixture was performed by
comparison with authentic samples prepared by chemical synthesis (see Supporting Information).
Interestingly, the biotransformation of compound 3a showed also the formation of the ethyl ester 3b
during the first day. This could be ascribed to a transesterification reaction promoted by the ethanol
produced by fermentation processes typical of the fungus growing on sugar medium even under
aerobic conditions.31 After five days (Figure 2a, Procedure A) the reaction mixture contained
cyclopentanemethanol 9 (14%), obtained by reduction of both the C=C double bond and the ester
moiety, and the two allylic alcohols 7a (41%) and 7b (30%). Neither the corresponding ketones 5a
and 5b, nor the other possible regioisomeric allylic alcohols could be detected by GC/MS among
the other components of the reaction mixture (unknown products 15%), not even in trace amounts.
When ethyl ester 3b was employed as a substrate (Figure 2b, Procedure A), after 5 days reaction
time, R. oryzae gave cyclopentanemethanol 9 (10%), allylic alcohol 7b (22%), and the
corresponding saturated derivative ethyl 3ꢀhydroxycyclopentaneꢀ1ꢀcarboxylate (45%), besides other
unidentified compounds (10%). Derivative 4a was hydroxylated very neatly by R. oryzae (Figure
2c, Procedure A), to give only allylic alcohol 8a (95 %), and 5% of cyclohexanemethanol (10).
Finally, the biotransformation of the ethyl ester 4b (Figure 2d, Procedure A) was found to be rather
slow, albeit quite selective: after 9 days a mixture containing 55% of the starting substrate and 45%
of the hydroxylated derivative 8b was recovered.
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After these positive preliminary results on the oxidative capability of R. oryzae towards
cycloalkenecarboxylates, we tried to improve the reaction conditions by entrapping the
microorganism in calcium alginate beads. The long mycelia of R. oryzae are known to form large
pellets or clumps in free cell cultivation, causing a severe mass transfer resistance. Immobilization
has been described to be a useful method to control the length of mycelia, facilitating oxygen mass
transfer.32 Thus, we studied the biocatalysed oxydation of 3a,b and 4a,b by using Caꢀalginate
entrapped R. oryzae. The spore suspension was mixed with Naꢀalginate and then added to a
sterilized solution of CaCl2, to give cell beads which were filtered, washed and preꢀcultured for 2
days at 25°C. A dense layer of mycelia was formed on the outer surface of the beads.32 In separate
experiments 3a,b and 4a,b, dissolved in DMSO, were added to the flasks containing the R. oryzae
beads suspended in the culture medium. The biotransformations were monitored by GC/MS and the
results are reported in Figure 2, Procedure B. The oxidation of 3a (Figure 2a, Procedure B) occurred
with a limited quantity of byꢀproducts (3% of 9, and 10% of unknown compounds), affording a
1:4.4 mixture of allylic alcohols 7a and 7b in 4 days reaction time. The reaction of ethyl ester 3b
(Figure 2b, Procedure B) was nearly complete in 2 days, affording compound 7b as the main
component (77%) of the reaction together with a small quantity of ethyl 3ꢀhydroxycyclopentaneꢀ1ꢀ
carboxylate (10%) and of unidentified products (11%). When derivative 4a was employed as the
starting compound (Figure 2c, Procedure B), allylic alcohol 8a was the main product (82%, 4 days),
but the formation of ethyl ester 8b (11%) was also observed. The hydroxylation of 4b to allyl
alcohol 8b was nearly complete (97%) in 3 days without formation of byꢀproducts (Figure 2d,
Procedure B).
Further experiments were performed by resuspending the beads in acetate buffer solution (pH 6.0),
in order to minimize the effects of extracellular enzymes and residual activities present in the
culture medium. The biotransformations were monitored by GC/MS and the results improved
considerably, as reported in Figure 2, Procedure C. For the biotransformation of 3a, methanol was
8

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used as a cosolvent instead of DMSO, in order to prevent the formation of ethyl ester 7b (Figure 2a,
procedure C): in 5 days the reaction afforded 95% of methyl ester 7a, along with 5% of unknown
products. The biotransformation of 3b (Figure 2b, Procedure C) occurred smoothly, and afforded
after 2 days the allylic alcohol 7b (78%) with a small quantity of the corresponding saturated
alcohol (4%, lower than that observed in culture medium). Under the same conditions, substrates 4a
and 4b were converted nearly quantitatively into allyl alcohols 8a (97%) and 8b (96%) in 3 days
(Figure 2c and 2d, Procedure C).
The allylic alcohols 7a,b and 8a,b were found to be enantiomerically enriched (Supporting
Information, Table S1): the enantiomeric excess values of the cyclohexene derivatives (60ꢀ86%)
were generally higher than those of the cyclopentene compounds (35ꢀ75%).
(a) 3a
(c) 4a
100
100
80
60
40
20
0
80
60
40
20
0
PAro(5ceddauyrse) A BP(r4ocdeadyusr)e BCP(5rodcaeydsu)re C
A (5 days) B (4 days) C (3 days)
Procedure A Procedure B Procedure C
(b) 3b
(d) 4b
100
100
80
60
40
20
0
80
60
40
20
0
other
products
other
products
A (5 days) B (2 days) C (2 days)
Procedure A Procedure B Procedure C
A (5 days) B (3 days) C (3 days)
Procedure A Procedure B Procedure C
Figure 2. Results of R. oryzae mediated biotransformations of compounds 3a (a), 3b (b), 4a (c), 4b
(d), according to the following experimental procedures. Procedure A: 5 mM substrate (DMSO as
cosolvent), R. oryzae whole cells, 25°C; Procedure B: 5 mM substrate (DMSO as cosolvent), R.
oryzae beads suspended in culture medium, 25°C; Procedure C: 5 mM substrate (MeOH or DMSO
as cosolvent), R. oryzae beads suspended in buffer pH 6.0, 25°C.
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In contrast with most of the reported chemical methods, the R. oryzae oxidation was demonstrated
to stop cleanly at the allylic alcohol stage without formation of unsaturated ketoesters 5 and 6.
While this may be a desirable chemoselectivity trait in different synthetic plans, a further oxidation
step had to be implemented to obtain 5 and 6 prior to the final bioreduction.
Biocatalysed oxidation of allylic alcohols 7a,bꢀ8a,b
Allylic alcohols 7a,bꢀ8a,b were converted into ketones 5a,bꢀ6a,b by using molecular oxygen as the
oxidant, in the presence of a laccase and a suitable chemical mediator. Laccases are blue
multicopper oxidases able to catalyze the fourꢀelectron reduction of molecular oxygen to water,
coupled with the concomitant oxidation of organic aromatic substrates.33 They are not effective
towards nonꢀphenolic derivatives, so to carry out the oxidation of primary and secondary alcohols
they require electron transfer mediators, such as 2,2’,6,6’ꢀtetramethylpiperidineꢀNꢀoxyl (TEMPO).34
The treatment of allylic alcohols 7a,bꢀ8a,b (5 g L–1, with DMSO as a cosolvent) with catalytic
–
TEMPO+ BF4 and laccase Amano M120 in acetate buffer (pH 5.0) under O2 atmosphere, afforded
quantitative oxidation to the corresponding ketones 5a,bꢀ6a,b over 24 hours at 30°C.
Enantioselective reduction of cyclic unsaturated ketoesters 5a,bꢀ6a,b
The enantioselective reduction of compounds 5a,bꢀ6a,b had been previously described in the
literature. Reetz et al.35 had reported that the reduction of methyl esters 5a and 6a mediated by the
eneꢀreductase YqjM afforded the (R)ꢀenantiomer of saturated ketoesters 1a and 2a very efficiently.
Moreover, while we were finalizing our experimental work on this topic, a paper by Faber et al. was
published,36 describing the enantioselective hydrogenations of substrates 5a,bꢀ6a,b (obtained by
traditional chemical oxidation with CrO3) mediated by a panel of known eneꢀreductases.
We submitted the unsaturated ketoesters 5a,bꢀ6a,b, prepared according to the twoꢀstep oxidation
protocol, to eneꢀreductase mediated hydrogenation, using as catalysts a panel of enzymes belonging
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to the Old Yellow Enzyme family: OYE1 (from Saccharomyces pastorianus), OYE2 and OYE3
(from S. cerevisiae), OYE2.6 (from Pichia stipitis), LeOPR1 (from Solanum lycopersicum), and
YqjM (from Bacillus subtilis). The recycling of the catalytic NADPH cofactor was performed with
glucose dehydrogenase (GDH from Bacillus megaterium), using glucose as a sacrificial cosubstrate.
The results of the preliminary screening are reported in Table 1.
In our hands, OYE1ꢀ3 and OYE2.6 invariably gave the same enantiomer of the reduced ketoesters
1a,bꢀ2a,b, while the reactions mediated by YqjM and LeOPR occurred with opposite
enantioselectivity.37 The absolute configuration of compounds 1a,b and 2a, recovered from OYE1
mediated reduction, was assigned to be (S) by comparison of the optical rotation data of these
samples with literature values (see Experimental Section). The configuration of compound 2b,
obtained by OYE1 mediated reduction of 2b, was established to be (S) by converting this sample
(94% ee) into the (+)ꢀ(S)ꢀenantiomer of the corresponding carboxylic acid.
Table 1. Eneꢀreductase mediated hydrogenationa of compounds 5a,bꢀ6a,b.
O
O
O
COOEt
COOMe
COOEt
6a
6b
5a
eec
5b
eec
Enzyme
OYE1
OYE2
cb
[
%]
[
%]
cb
[
%]
[
%]
cb
[
%]
eec
[
%]
cb
[
%]
eec
[%]
99
99
99
99
99
99
90 (S)
82 (S)
90 (S)
92 (S)
88 (R)
86 (R)
99
99
99
99
99
99
86 (S)
80 (S)
71 (S)
87 (S)
99 (R)
86 (R)
99
99
99
99
99
99
94 (S)
75 (S)
62 (S)
96 (S)
98 (R)
99 (R)
99
99
99
99
99
99
96 (S)
90 (S)
42 (S)
92 (S)
99 (R)
99 (R)
OYE3
OYE 2.6
LeOPR1
YqjM
a 5 mM substrate, 20 mM glucose, isolated OYE, GDH, NADP+, 1% DMSO, phosphate buffer pH 7.0, 30°C, 24 h;
b conversion calculated on the basis of GC analysis of the crude mixture after 24 h (isolation yields are reported in
c
the Experimental Section); enantiomeric excess calculated on the basis of GC analysis on a chiral stationary
phase (see Experimental Section).
The enantioselectivities of these cycloalkene bioreductions are in agreement with literature data for
structurally related βꢀalkylcycloalkenones. For instance, 3ꢀmethylcyclopentenone is reduced to the
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(S)ꢀenantiomer of the saturated ketone by OYE1ꢀ3 (>99% ee),38 while it is converted very poorly by
LeOPR1 and YqjM,39 and no information are available about its reduction with OYE2.6. Similarly,
3ꢀmethylcyclohexenone is converted into (S)ꢀ3ꢀmethylcyclohexanone by OYE1ꢀ340,38 and
OYE2.6,41 and it is scarcely accepted by LeOPR1 and YqjM.39 Furthermore, investigations
performed by using OYE142 and OYE2.641 highlighted that the reduction of 3ꢀalkylcyclohexenones
is very sensitive to steric hindrance of the substituent at the βꢀposition: conversions decreased by
increasing the size of the alkyl chain, and substrates with a substituent longer than a nꢀpropyl chain
were not accepted by the enzymes. In the transformation of the cyclic γꢀoxoesters of this work, the
presence of either a methyl or an ethyl ester group linked to the β olefinic carbon atom is not
detrimental for conversion and enantioselectivity of the reactions mediated by the six investigated
ERs.
Conversion of cycloalkenecarboxylates 3a,b-4a,b into (R)- and (S)-1a,b-2a,b
The two oxidative steps and the alkene reduction were combined to convert
cycloalkenecarboxylates 3a,bꢀ4a,b into (R)ꢀ and (S)ꢀ1a,bꢀ2a,b (Scheme 2). The CꢀH allylic
hydroxylation of derivatives 3a,bꢀ4a,b was performed according to Procedure C (30 g of R. oryzae
beads in 120 mL of acetate buffer pH 6.0, in 300 mL flasks), with 5 mM substrate concentration.
The reactions were monitored by GC/MS.
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Scheme 2. Conversion of cycloalkenecarboxylates 3a,bꢀ4a,b into (R)ꢀand (S)ꢀ1a,bꢀ2a,b.
The entrapment of R. oryzae in calcium alginate beads considerably simplified the reaction workꢀ
up, which consisted solely in the removal of the beads by filtration and extraction of the filtrate with
EtOAc. As discussed above, the use of a buffer solution instead of the fungal cultural medium
greatly decreased the amounts of subproducts and metabolites in the isolated allylic alcohols. In
particular, 7a and 8a,b were obtained in satisfactory purity and were submitted to the following step
without further treatment. Only in the case of ethyl ester 3b, the formation of 10% of saturated
ketone (S)ꢀ1b was observed and column chromatography purification of allylic alcohol 7b was thus
necessary.
The allylic alcohols 7a,bꢀ8a,b were converted directly by laccase/TEMPO oxidation into γꢀ
oxoesters 5a,bꢀ6a,b (total reaction volume 10 mL, substrate concentration 35 mM) affording
complete conversion. Since the reaction conditions for the laccase/TEMPO system and for the
OYE/NADP+/GDH/glucose system were found to be compatible, these intermediates γꢀoxoesters
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were not isolated and the final C=C reduction was carried out telescopically. Firstly, in order to
avoid the consumption of NADP+ by reaction with TEMPO+, after the oxidation the excess of
molecular oxygen dissolved in the reaction medium was removed by bubbling nitrogen, and
isopropanol was added to reduce the residual oxoammonium cation. Then, the suitable eneꢀ
reductases, together with NADP+ and the cofactor regenerating system (GDH/glucose), were added
to the reaction mixtures for the final reduction, which occurred quantitatively in all cases. Both the
enantiomers of cyclic γꢀoxoesters 1a,bꢀ2a,b were obtained in 47ꢀ68% overall isolation yields from
the starting cycloalkenecarboxylates, without any chromatographic separation (except for
intermediate 7b).
Conclusions
This enzymatic synthesis of enantiomerically enriched γꢀoxoesters 1 and 2 represents a convenient
alternative to the known chemical methods summarised above. In spite of the necessity of rather
long reaction times, the procedure shows several advantages. The starting materials 3a and 4a are
commercially available and less expensive than racemic 3ꢀoxocyclopentaneꢀ and 3ꢀ
oxocyclohexanecarboxylic acid. Metal catalysts or complex reagents/catalysts to be specifically
prepared are avoided, thanks to the use of enzymes from renewable sources. Atoms are inserted
sequentially and maintained within the molecular structure, and no removal of auxiliary fragments
or protecting groups is required.
The twoꢀstep oxidation procedure, using R. oryzae resting cells entrapped in alginate beads and the
laccase/TEMPO system, represents a sustainable enzymatic synthetic approach to unsaturated
cyclic oxoesters 5a,b-6a,b, showing lower environmental burden and higher efficiency than
classical chemical oxidations. The biohydroxylation step is characterised by high regioselectivity
with limited formation of sideꢀproducts, such as regioisomeric allyl alcohols and epoxides
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(commonly occurring in chemical oxidation procedures, and causing a decrease of the overall yield
of the desired products). The use of beads suspended in buffer solution makes the reaction workꢀup
much easier: the fungal catalyst is removed by filtration, and the allylic alcohols are isolated in high
purity (except for 7b, that required column chromatography) without further purification before the
following oxidation step, thus minimizing solvents/energy requirements of the process. The
possibility to combine the laccase/TEMPO oxidation with the alkene bioreduction in a sequential
cascade increases the value of the procedure. The availability of eneꢀreductases characterized by
broad substrate tolerance for cyclic γꢀoxoesters 4a,b-5a,b and showing high and opposite
enantioselectivity, enabled the preparation of both the enantiomers of the corresponding saturated
derivatives of satisfactory purity for API synthesis. Finally, the procedure is performed in water
(with minimal amounts of DMSO or methanol as cosolvents) and the nonꢀtoxic waste generated
during the whole process can be easily disposed of, in contrast with the traditional chemical
procedures.
Keywords
biocatalysis; enantioselectivity; oxidoreductases; eneꢀreductases; laccases
Experimental
General
GCꢀMS analyses were performed using a HPꢀ5MS column (30 m × 0.25 mm × 0.25 µm, Agilent).
The following temperature program was employed: 60°C (1 min) / 6°C min–1 / 150°C (1 min) /
12°C min–1 / 280°C (5 min). The enantiomeric excess values of compounds 1a,b-2a,b were
determined by GC analysis. Chiral GC analyses were performed on a MEGAꢀDEX DACꢀBeta
column (25 m × 0.25 mm × 0.25 ꢁm, Mega), carrier gas H2, constant flow 3.7 mL min–1, injector
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temperature 250°C, detector temperature 250°C with the following temperature programs: a)
compound 1a: 90°C / 0.7°C min–1 / 112°C / 90°C min–1 / 220°C (3 min), (S)ꢀ1a tR = 25.1 min; (R)ꢀ
1a tR = 26.7 min; b) compound 1b: 100°C (35 min) / 90°C min–1 / 220°C (3 min), (S)ꢀ1b tR = 23.7
min; (R)ꢀ1b tR = 26.2 min; c) compound 2a: 100°C / 0.3°C min–1 / 110°C / 90°C min–1 / 220°C (3
min), (R)ꢀ2a tR = 20.4 min; (S)ꢀ2a tR = 21.6 min; d) compound 2b: 95°C (45 min) / 90°C min–1
/
220°C (3 min), (R)ꢀ2b tR = 38.5 min; (S)ꢀ2b tR = 41.0 min. 1H and 13C NMR spectra were recorded
on a 400 or 500 MHz spectrometer, and the chemical shift scale was based on internal
tetramethylsilane. TLC analyses were performed on Merck Kieselgel 60 F254 plates. All the
chromatographic separations were carried out on silica gel columns.
Strains and plasmids
Colletotrichum lini CBS 112.21, Rhizopus oryzae Mucl 28423, Aspergillus niger (Microbiologia
Industriale Milano) and Alternaria tenuis (Microbiologia Agraria, Alimentare, Ecologica) were
obtained from the fungal collection of the Department of Food, Environmental and Nutritional
Science (DeFENS), University of Milan (Prof. F. Molinari and Dr. D. Romano). Laccase Amano
M120 was employed. Eneꢀreductases (OYE1ꢀ3, OYE2.6, and LeOPR1) and glucose dehydrogenase
(GDH) were overproduced in E. coli BL21(DE3) or E.coli RosettaTM(DE3) strains harbouring a
specific plasmid prepared as previously reported: pET30aꢀOYE1 from the original plasmid
provided by Neil C. Bruce,43 pET30aꢀOYE2 and pET30aꢀOYE3 from S. cerevisiae BY4741 and
pKTSꢀGDH from B. megaterium DSM509;44 pDJBxꢀOYE2.6 and pDJBxꢀLeOPR1 from the
original plasmids provided by Prof. Jon D. Stewart. As for YqjM, the original plasmid provided by
Prof. M. Hall was used directly as provided.36
Entrapment of R. oryzae
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Rhizopus oryzae CBS 112.07 was routinely maintained on MEA (malt extract agar) medium (malt
extract 20 g L–1, glucose 20 g L–1, peptone 2 g L–1, agar 20 g L–1). The fungal spores were
suspended in sterile water, mixed with a sterile Naꢀalginate solution (40 g L–1), and added dropwise
to a sterile solution of CaCl2 (20 g L–1) under mild stirring, to form cell beads with the average
diameter of 3.5 mm. The beads were filtered, and preꢀcultured in a flask (ca. 10 g of beads per 40
mL of MEA medium) at 25°C on a rotary shaker at 160 rpm for 2 days, to form a shell of mycelia
on the outer surface. For Procedure B, the beads were employed as a suspension in this culture
medium. For Procedure C the beads were recovered by filtration, washed with sterile water and
suspended in acetate buffer (20 mM, pH 6.0).
Overproduction of enzymes in E. coli BL21(DE3)
LB medium (5 mL) containing the appropriate antibiotic (50 ꢁg mL–1 kanamycin for pET30a, 100
ꢁg mL–1 ampicillin for pKTS, pDJBx and 100 µg mL ꢀ1 ampicillin and 30 µg mL ꢀ1 chloramphenicol
for YqjM) was inoculated with a single colony from a fresh plate and grown for 8 h at 37°C and
220 rpm. This starter culture was used to inoculate 500 mL LB medium (TB medium in the case of
YqjM). The latter culture was shaken at 37°C and 220 rpm until OD600 reached 0.4ꢀ0.5, then
enzyme expression was induced by the addition of 0.1 mM IPTG (50 ng mL–1 anhydrotetracycline
was also added in the case of the pKTSꢀGDH plasmid) at 30°C and 160 rpm.
In the case of OYE1ꢀ3, GDH and YqjM, after 5ꢀ6 h the cells were harvested by centrifugation
(5000 g, 20 min, 4°C), resuspended in 50 mL of lysis buffer (20 mM KPi buffer pH 7.0, 300 mM
NaCl, 10 mM imidazole) and disrupted by sonication (Omni Ruptor 250 ultrasonic homogeniser,
five sonication cycles, 15 s each, 50% duty). YqjM was employed as cellꢀfree extract, whereas for
OYE1ꢀ3 and GDH the cellꢀfree extract, after centrifugation (20000 g, 20 min, 4°C), was
chromatographed on IMAC stationary phase (NiꢀSepharose Fast Flow, GE Healthcare) with a
mobile phase composed of 20 mM KPi buffer pH 7.0, 300 mM NaCl and a 10ꢀ300 mM imidazole
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gradient. Protein elution was monitored at 280 nm, the fractions were collected according to the
chromatogram and dialysed twice against 1.0 L of 50 mM KPi buffer pH 7.0 (12 h each, 4°C) to
remove imidazole and salts. Purified protein aliquots were stored frozen at –80°C.
In the case of OYE2.6, and LeOPR1, produced as GSTꢀfusion proteins, cell pellets were
resuspended in cold sepharose binding buffer (PBS, 140 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4,
1.8 mM KH2PO4, pH 7.3) and lysed by sonication (Omni Ruptor 250 ultrasonic homogeniser, five
sonication cycles, 15 s each, 50% duty). The cellꢀfree extract was centrifuged (12000 rpm, 40 min,
4°C). The resulting supernatant was passed through Glutathione Sepharose 4 Fast Flow (GE
Healthcare), with PBS buffer as the mobile phase. Once the absorbance (280 nm) returned to a
baseline reading, the desired protein was eluted by adding a reduced glutathione (GSH) buffer
solution (10 mM γꢀLꢀglutamylꢀLꢀcysteinylglycine, 50 mM TrisꢀHCl, pH 8.0). Protein elution was
monitored at 280 nm and the fractions were collected according to the chromatogram. Purified
protein aliquots were stored frozen at –80°C.
General procedure for the multi-enzymatic conversion of 3a,b-4a,b into 1a,b-2a,b
Allylic hydroxylation of substrates 3a,bꢀ4a,b by R. oryzae (Procedure C)
Preꢀcultivated R. oryzae beads (corresponding to ca. 30 g of those prepared according to the
entrapment procedure) were suspended in acetate buffer (120 mL, 20 mM, pH 6.0) in a 300 mL
flask, and a solution of the substrate (1.2 mL, 500 mM) in MeOH (for compound 3a) or DMSO (for
compounds 3b-4a,b) was added. The flask was shaken at 160 rpm at 25°C, and the reaction was
monitored by GC/MS analysis. After consumption of the starting substrate, the beads were filtered,
and the aqueous solution was extracted with EtOAc. The organic phase was dried over anhydrous
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Na2SO4 and the residue was submitted to the following oxidation step without any further
purification.
Methyl 3-hydroxycyclopent-1-ene-1-carboxylate (7a)
From 3a (0.076 g, 0.60 mmol), according to Procedure C after 5 days, derivative 7a was obtained
(0.074 mg, 87%): 1H NMR (CDCl3, 400 MHz):45 δ = 6.70 (q, J = 2.1 Hz, 1H, CH=C), 4.97 (m, 1H,
CHOH), 3.76 (s, 3H, COOCH3), 2.78 ꢀ 2.66 (m, 1H, CHH), 2.56 ꢀ 2.32 (m, 2H, 2CHH), 1.85 – 1.73
(m, 1H, 2CHH); 13C NMR (CDCl3, 100.6 MHz): δ = 166.1, 143.0, 138.9, 77.2, 52.0, 33.2, 30.0;
GC/MS (EI) tR = 11.83 min: m/z (%) = 142 (M+, 5), 127 (53), 81 (100).
Ethyl 3-hydroxycyclopent-1-ene-1-carboxylate (7b)
From 3b (0.084 g, 0.60 mmol), according to Procedure C after 2 days, derivative 7b was recovered
after purification by column chromatography (0.061 g, 65%): 1H NMR (CDCl3, 400 MHz): δ = 6.69
(q, J = 2.1 Hz, 1H, CH=C), 4.97 (m, 1H, CHOH), 4.21 (q, J = 7.1 Hz, COOCH2CH3), 2.80 ꢀ 2.65
(m, 1H, CHH), 2.55ꢀ2.30 (m, 2H, 2CHH), 1.86ꢀ1.75 (m, 1H, 2CHH), 1.30 (t. J = 7.1 Hz,
COOCH2CH3; 13C NMR (CDCl3, 100.6 MHz): δ = 165.3, 142.9, 139.2, 77.3, 60.7, 33.7, 30.0, 14.3;
GC/MS (EI) tR = 13.54 min: m/z (%) = 127 (M+–29, 75), 111 (38), 83 (100).
Methyl 3-hydroxycyclohex-1-enecarboxylate (8a)
From 4a (0.084 g, 0.60 mmol), according to Procedure C after 3 days, derivative 8a was obtained
(0.082 g, 88%): 1H NMR (CDCl3, 400 MHz):25 δ = 6.87 (m, 1H, CH=), 4.35 (m, 1H, CHOH), 3.75
13
(s, 3H, COOCH3), 2.36ꢀ2.16 (m, 2H, 2CHH ), 2.00ꢀ1.50 (m, 4H, 2 CH2); C NMR (CDCl3, 100.6
MHz: 25 δ = 167.9, 140.3, 132.1, 65.8, 51.8, 31.0, 24.2, 19.2; GC/MS (EI) tR = 14.85 min: m/z (%) =
156 (M+, 12), 141 (71), 124 (65), 97 (100).
Ethyl 3-hydroxycyclohex-1-enecarboxylate (8b)
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From 4b (0.092 g, 0.60 mmol), according to Procedure C after 3 days, derivative 8b was obtained
1
(0.088 g, 86%): H NMR (CDCl3, 400 MHz): δ = 6.87 (q, J = 2.2 Hz, 1H, CH=C), 4.36 (m, 1H,
CHOH), 4.20 (q, J = 7.1 Hz, COOCH2CH3), 2.35ꢀ2.15 (m, 2H, 2CHH), 2.00ꢀ1.75 (m, 2H, 2CHH),
1.70ꢀ1.50 (m, 2H, 2CHH), 1.28 (t, J = 7.1 Hz, 3H, COOCH2CH3); 13C NMR (CDCl3, 100.6 MHz):
δ = 167.4, 139.5, 132.9, 66.1, 60.7, 31.4, 24.4, 19.2, 14.4; GC/MS (EI) tR = 16.44 min: m/z (%) =
170 (M+, 4), 141 (100), 124 (78), 114 (22), 97 (100).
Oneꢀpot sequential twoꢀstep conversion of allylic alcohols 7a,bꢀ8a,b into (S)ꢀ and (R)ꢀ1a,bꢀ2a,b
A solution of crude allylic alcohols 7a,bꢀ8a,b in DMSO (0.70 mL, 500 mM) was added to an
−
acetate buffer (10 mL, 50 mM, pH 5.0), containing TEMPO+BF4 (70 ꢀmol, 0.2 eq.) and laccase
Amano M120 (50 mg). After stirring at 30°C for 24 hours under O2 atmosphere to achieve complete
conversion into intermediate oxoesters 5a,bꢀ6a,b, iPrOH (100 uL) was added to quench the excess
of TEMPO+, and N2 was bubbled through the solution to remove dissolved oxygen. The required
OYE (1ꢀ2 mg) was then added, together with glucose (1.40 mmol, 4 eq.), NADP+ (35.0 ꢁmol, 0.1
eq.), and GDH (5 U mL−1). The mixture was incubated for 24 h in an orbital shaker (160 rpm,
30°C). The solution was extracted with EtOAc, centrifuging after each extraction (15000 g, 1.5
min), and the combined organic solutions were dried over anhydrous Na2SO4, to give either (S)ꢀ or
(R)ꢀ1a,bꢀ2a,b.
(S)-Methyl 3-oxocyclopentanecarboxylate ((S)-1a)
From 7a (0.050 g, 0.35 mmol), after TEMPO/laccase oxidation and OYE1ꢀmediated reduction,
compound (S)ꢀ1a (0.037g, 75%) was obtained: ee = 90% (chiral GC); [α]D = –36.2 (c 1.0, CHCl3)
[lit. ref. 46 [α]D = 37 (c 3.5, CHCl3, for (R)ꢀ1a with ee = 95%); 1H NMR (CDCl3, 400 MHz):36 δ =
3.70 (s, 3H, COOCH3), 3.11 (m, 1H, CHCOOCH3), 2.51ꢀ2.01 (m, 6H, 3CH2); 13C NMR (CDCl3,
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100.6 MHz):36 δ =216.5, 174.8, 52.2, 41.2, 40.9, 37.5, 26.6; GC/MS (EI) tR = 10.43 min: m/z (%) =
142 (M+, 13), 114 (75), 55 (100).
(R)-Methyl 3-oxocyclopentanecarboxylate ((R)-1a)
From 7a (0.050 g, 0.35 mmol), after TEMPO/laccase oxidation and LeOPR1ꢀmediated reduction,
compound (R)ꢀ1a (g, %) was obtained: ee = 88% (chiral GC); [α]D = +35.0 (c 1.0, CHCl3). 1H and
13C NMR spectra were in agreement with those of the (S)ꢀenantiomer.
(S)-Ethyl 3-oxocyclopentanecarboxylate ((S)-1b)
From 7b (0.054 g, 0.35 mmol), after TEMPO/laccase oxidation and OYE1ꢀmediated reduction,
compound (S)ꢀ1b (0.040 g, 73%) was obtained: ee = 86% (chiral GC); [α]D = –18.8 (c 1.5, MeOH)
[lit. ref. 47 [α]D = ꢀ22.2 (c 1.0, MeOH, for (S)ꢀ1b with ee = 99%); 1H NMR (CDCl3, 400 MHz):36 δ
= 4.18 (q, J = 7.1 Hz, COOCH2CH3), 3.10 (m, 1H, CHCOOCH2CH3), 2.55ꢀ2.09 (m, 3 CH2, 6H),
1.29 (t, J = 7.1 Hz, 3H, COOCH2CH3); 13C NMR (CDCl3, 100.6 MHz):36 δ = 216.6, 174.3, 61.0,
41.2, 41.0, 37.5, 26.7, 14.3; GC/MS (EI) tR = 12.18 min: m/z (%) = 156 (M+, 5), 128 (50), 111 (25),
100 (75), 55 (100).
(R)-Ethyl 3-oxocyclopentanecarboxylate ((R)-1b)
From 7b (0.058 g, 0.38 mmol), after TEMPO/laccase oxidation and LeOPR1ꢀmediated reduction,
compound (R)ꢀ1b (0.041 g, 70%) was obtained: ee = 99% (chiral GC); [α]D = +21.7 (c 1.0, MeOH).
1H and 13C NMR spectra were in agreement with those of the (S)ꢀenantiomer.
(S)-Methyl 3-oxocyclohexanecarboxylate ((S)-2a)
From 8a (0.052 g, 0.34 mmol), after TEMPO/laccase oxidation and OYE1ꢀmediated reduction,
compound (S)ꢀ2a (0.041 g, 77%) was obtained: ee = 94% (chiral GC); [α]D = +2.7 (c 2.0, EtOH)
1
[lit. ref. 35 [α]D = –3.2 (c 0.135 g mLꢀ1, EtOH, for (R)ꢀ1a with ee = 99%); H NMR (CDCl3, 400
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MHz):36 δ = 3.70 (s, 3H, COOCH3), 2.80 (m, 1H, CHCOOCH3), 2.55 (d, J = 7.9 Hz, 2H,
CH2CHCOO), 2.45 ꢀ2.25 (m, 2H, 2CHH), 2.20 ꢀ2.10 (m, 2H, 2CHH), 1.90 ꢀ 1.65 (m, 2H, 2CHH);
13C NMR (CDCl3, 100.6 MHz):36 δ = 209.3, 174.2, 52.2, 43.20, 43.19, 41.0, 27.8, 24.6; GC/MS
(EI) tR = 13.02 min: m/z (%) = 156 (M+, 21), 124 (11), 113 (32), 97 (100).
(R)-Methyl 3-oxocyclohexanecarboxylate ((R)-2a)
From 8a (0.054 g, 0.35 mmol), after TEMPO/laccase oxidation and LeOPR1ꢀmediated reduction,
compound (R)ꢀ2a (0.038 g, 69%) was obtained: ee = 98% (chiral GC); [α]D = –3.0 (c 1.5, MeOH).
1H and 13C NMR spectra were in agreement with those of the (S)ꢀenantiomer.
(S)-Ethyl 3-oxocyclohexanecarboxylate ((S)-2b)
From 8b (0.055 g, 0.33 mmol), after TEMPO/laccase oxidation and OYE1ꢀmediated reduction,
compound (+)ꢀ2b (0.042 g, 74%) was obtained: ee = 96% (chiral GC); [α]D = +2.7 (c 1.6, MeOH).
1H NMR (CDCl3, 400 MHz): δ = 4.15 (q, s, 2H, COOCH2CH3), 2.78 (m, 1H, CHCOOCH2CH3),
2.55 (d, J = 7.9 Hz, 2H, CH2CHCOO), 2.40ꢀ2.25 (m, 2H, 2CHH), 2.20ꢀ2.05 (m, 2H, 2CHH), 1.95 ꢀ
1.70 (m, 2H, 2CHH), 1.27 (t, J = 7.1 Hz, 3H); 13C NMR (CDCl3, 100.6 MHz): δ = 209.4, 173.8,
61.0, 43.3, 43.2, 41.0, 27.8, 24.6, 14.3; GC/MS (EI) tR = 14.70 min: m/z (%) = 170 (M+, 7), 142 (5),
128 (12), 125 (9), 113 (6), 102 (6), 97 (100).
Compound (+)ꢀ2b (0.035 g, 0.21 mmol) was dissolved in MeOH (2 mL) and treated with a solution
of LiOH in water (4 mL, 1 M). After stirring at room temperature for 6 h, the solution was
concentrated under reduced pressure, acidified with 10% HCl and extracted with EtOAc. The
solvent was removed in vacuo to afford (+)ꢀ(S)ꢀ3ꢀoxocylcohexanecarboxylic acid (0.026 g, 87%):
1
[α]D = +16.5 (c 1, MeOH) lit. ref. 48 [α]D = –17.6 (c 4, MeOH) for the (R)ꢀenantiomer; H NMR
(CDCl3, 400 MHz):49 δ = 2.92ꢀ2.80 (m, 1H, CHCOOH), 2.65ꢀ2.50 (m, 2H, CH2CHCOO), 2.45ꢀ2.25
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(m, 2H, 2CHH), 2.20ꢀ2.00 (m, 2H, 2CHH), 1.95ꢀ1.70 (m, 2H, 2CHH); GC/MS (EI) tR = 15.45 min:
m/z (%) = 142 (M+, 33), 114 (20), 97 (92), 55 (100).
(R)-Ethyl 3-oxocyclohexanecarboxylate ((R)-2b)
From 8b (0.060 g, 0.36 mmol), after TEMPO/laccase oxidation and LeOPR1ꢀmediated reduction,
compound (R)ꢀ2b (0.046 g, 75%) was obtained: ee = 99% (chiral GC); [α]D = ꢀ 2.9 (c 1.7, MeOH).
1H and 13C NMR spectra were in agreement with those of the (S)ꢀenantiomer.
Acknowledgement
This work was supported by Fondazione Cariplo (grant no. 2014ꢀ0568). Prof. F. Molinari, and Dr.
D. Romano (University of Milan, Department of Food, Environmental and Nutritional Science) are
kindly acknowledged for making available the fungal strains. Prof J. D. Stewart (University of
Florida) is acknowledged for the kind gift of OYE2.6 and LeOPR1 expression constructs, and Prof.
M. Hall (University of Graz) for that of YqjM. Mr. Davide De Simeis and Giacomo Toselli are
kindly acknowledged for helpful technical assistance.
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Graphical Abstract
The conversion of C5 and C6 cycloalkenecarboxylates into both enantiomers of the corresponding
cyclic γꢀoxoesters is performed at room temperature using three enzymes, two isolation steps, and
no environmentallyꢀunfriendly oxidants.
86-96% ee
Rhizopus oryzae
88-99% ee
n = 1,2
R = Me,Et
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Notes and References
1. T. Itoh, U. Hanefeld, Green Chem., 2017, 19, 331ꢀ332.
2. G. W. Huisman, S. J. Collier, Curr. Opin. Chem. Biol., 2013, 17, 284 ꢀ 292.
3. (a) S. P. France, L. J. Hepworth, N. J. Turner, S. L. Flitsch, ACS Catal. 2017, 7, 710ꢀ724. (b)
E. GarcíaꢀJuncedaa, I. Lavandera, D. Rother, J. H. Schrittwieser, J. Mol. Catal. B, 2015,
114, 1ꢀ6.
4. Organic Synthesis Using Biocatalysis, Eds. A. Goswami and J. D. Stewart, 2016, Elsevier.
5. E. J. Horn, B. R. Rosen, Y. Chen, J. Tang, K. Chen, M. D. Eastgate and P. S. Baran, Nature,
2016, 533, 77ꢀ81.
6. K. Chen, P. Zhang, Y. Wang, and H. Li, Green Chem., 2014, 16, 2344ꢀ2374, and references
therein.
7.
(a) F. Hollmann, I. W. C. E. Arends, K. Buehler, A. Schallmey and B. Bühler, Green
Chem., 2011, 13, 226ꢀ265; (b) E. Roduner, W. Kaim, B. Sarkar, V. B. Urlacher, J. Pleiss, R.
Gläser, W.ꢀD. Einicke, G. A. Sprenger, U. Beifuß, E. Klemm, C. Liebner, H. Hieronymus,
S.ꢀF. Hsu, B. Plietker, S. Laschat, ChemCatChem, 2013, 5, 82ꢀ112
8. V. B. Urlacher and M. Girhard, in Enzyme Catalysis in Organic Synthesis, ed. K. Drauz, H.
Gröger, and O. May, WileyꢀVCH, Weinheim, Third Edition, 2012, Vol.3, Chapter 30, 1225
ꢀ1267.
9. Astra Zeneca AB, WO 2009/024821 A2, 2009.
10. Merck Sharp & Dohme Corp., WO 2015/027431 A1, 2015.
11. SanofiꢀAventis Deutschland GMBH, WO 2006/018115 A1, 2006.
12. K. Shimada, Y. Kaburagi, T. Fukuyama, J. Am. Chem. Soc. 2003, 125, 4048ꢀ4049.
13. K. Ausmees, A. Selyutina, K. Kuett, K. Lippur, T. Pehk, M. Lopp, E. Zusinaite, A. Merits,
T. Kanger, Nucleosides, Nucleotides Nucleic Acids, 2011, 30, 897 – 907.
14. K. Toki, Bull. Chem. Soc. Jpn., 1958, 31, 333ꢀ334.
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15. R. D. Allan, G. A. R. Johnston, B. Twitchin, Aust. J. Chem., 2017, 34, 2231ꢀ22366.
16. J. B. Tuttle, S. G. Ouellet, D. W. C. MacMillan, J. Am. Chem. Soc. 2006, 128, 12662ꢀ12663
17. X.ꢀQ. Yu, T. Shirai, Y. Yamamoto, N. Miyaura, Chem. Asian J. 2011, 6, 932 – 937.
18. E. Díez, R. Fernández, C. Gasch, J. M. Lassaletta, J. M. Llera, E. MartínꢀZamora, J.
Vázquez, J. Org. Chem. 1997, 62, 5144ꢀ5155.
19. R. W. Barnhart, X. Wang, P. Noheda, S. H. Bergens, J. Whelan, B. Bosnich, J. Am. Chem.
Soc, 1994, 116, 1821ꢀ1830.
20. Y. Aye, S. G. Davies, A. C. Garner, P. M. Roberts, A. D. Smith, J. E. Thomson, Org.
Biomol. Chem., 2008, 6, 2195 – 2203.
21. M. K. Brown, T. L. May, C. A. Baxter, A. H. Hoveyda, Angew. Chem., Int. Ed., 2007, 46,
1097ꢀ1100.
22. J.ꢀQ. Yu, E. J. Corey, J. Am. Chem. Soc. 2003, 125, 3232ꢀ3233.
23. A. J. Catino, R. E. Forslund, M. P. Doyle, J. Am. Chem. Soc., 2004, 126, 13622ꢀ13623.
24. S. J. Na, B. Y. Lee, N. – N. Bui, S. Mho, H. – Y. Jang, J. Organomet. Chem., 2007, 692,
5523ꢀ5527.
25. R. Agudo, G.ꢀD. Roiban, M. T. Reetz, ChemBioChem 2012, 13, 1465ꢀ1473.
26. R. Agudo, M. T. Reetz, Chem. Commun., 2013, 49, 10914ꢀ10916.
27. A. Romano, D. Romano, E. Ragg, F. Costantino, R. Lenna, R. Gandolfi, F. Molinari,
Steroids, 2006, 71, 429ꢀ434.
28. S. Petrič, T. Hakki, R. Bernhardt, D. Zigon, B. Crešnar, J. Biotechnol. 2010, 150, 428ꢀ437.
29. I. A. Parshikov, J. B. Sutherland, Proc. Biochem. 2014, 49, 2086ꢀ2100.
30. M. Ye, G. Qu, H. Guo, D.Guo, Appl. Environ. Microbiol., 2004, 70, 3521ꢀ3527.
31. R. H. W. Maas, J. Springer, G. Eggink, R. A. Weusthuis, J. Ind. Microbiol. Biotechnol.,
2008, 35, 569–578.
32. J. Lin, M. Zhoub, X. Zhaob, S. Luoc, Y. Luc, Chem. Eng. Process. 2007, 46, 369ꢀ374.
26

Page 27 of 28
Green Chemistry
View Article Online
DOI: 10.1039/C7GC02215H
33. S. Riva, Trends Biotechnol. 2006, 24, 219ꢀ226.
34. M. Fabbrini, C. Galli, P. Gentili, D. Macchitella, Org. Lett. 2001, 42, 7551ꢀ7753.
35. D. J. Bougioukou, S. Kille, A. Taglieber, M. T. Reetz, Adv. Synth. Catal. 2009, 351, 3287 –
3305.
36. N. G. Turrini, R. C. Cioc, D. J. H. van der Niet, E. Ruijter, R. V. A. Orru, M. Hall, K. Faber,
Green Chem., 2017, 19, 511ꢀ518
37. In contrast with previously published data (ref. 28), no inversion of enantioselectivity was
observed with OYE1 and 3 with respect to OYE2.
38. M. Hall, C. Stueckler, B. Hauer, R. Stuermer, T. Friedrich, M. Breuer, W. Kroutil, K. Faber,
Eur. J. Org. Chem. 2008, 1511ꢀ1516.
39. M. Hall, C. Stueckler, H. Ehammer, E. Pointner, G. Oberdorfer, K. Gruber, B. Hauer, R.
Stuermer, W. Kroutil, P. Macheroux, K. Faber, Adv. Synth. Catal. 2008, 350, 411ꢀ418.
40. D. Monti, M. C. Forchin, M. Crotti, F. Parmeggiani, F. G. Gatti, E. Brenna, S. Riva
ChemCatChem 2015, 7, 3106ꢀ3109.
41. A. PattersonꢀOrazem, B. Sullivan, J. D. Stewart, Bioorg. Med. Chem. 2014, 22, 5628ꢀ5632
42. M.A. Swiderska, J.D. Stewart, J. Molec. Catal. B: Enzym. 2006, 42, 52ꢀ54.
43. R.E. Williams, D. A. Rathborne, N. S. Scrutton, N. C. Bruce, Appl. Environ. Microbiol.
2004, 70, 3566–3574.
44. M. Bechtold, E. Brenna, F.C. Femmer, F. G. Gatti, S. Panke, F. Parmeggiani, A. Sacchetti,
Org. Process Res. Dev. 2012, 16, 269–276.
45. Y. Aye, S. G. Davies, A. C. Garner, P. M. Roberts, A. D. Smith, J. E. Thomson, Org.
Biomol. Chem., 2008, 6, 2195ꢀ2203.
46. X.ꢀQ. Yu, T. Shirai, Y. Yamamoto, N. Miyaura, Chem. Asian J. 2011, 6, 932ꢀ937.
47. R. W. Barnhart, X. Wang, P. Noheda, S. H. Bergens, J. Whelan, B. Bosnich, J. Am. Chem.
Soc. 1994, 116, 1821ꢀ1830.
27

Green Chemistry
Page 28 of 28
View Article Online
DOI: 10.1039/C7GC02215H
48. R. D. Allan, G. A. R. Johnston, B. Twitchin, Aust. J. Chem., 1981, 34, 2231ꢀ2236.
49. A. P. Marchand, R. Sharma, U. R. Zope, J. Org. Chem. 1993, 58, 7
28