Enzymatic Synergism in the Synthesis of β-Keto Esters

Article abstract of DOI:10.1002/ejoc.201500676

Reaction of alcohols with ethyl and tert-butyl acetoacetate catalyzed by a combination of commercially available enzymes is shown to be a convenient method for the preparation of a range of acetoacetic acid derivatives. Systematic studies proved that the combination of two or more enzymes enhances the yield of the reaction. Application of the selected enzyme mixture for enzymatic transesterification of various β-keto esters provided the respective products in excellent yields up to 96% and quantitative within 24 and 48 hours, respectively. The presented methodology is simple and mild, and can be used to prepare acetoacetates from primary and secondary alcohols. Application of a selected enzyme mixture for enzyme-catalyzed esterification of various carboxylic acids provided the respective β-hydroxy esters in excellent yields up to 96% and quantitative within 24 and 48 hours, respectively.

Full text of DOI:10.1002/ejoc.201500676

FULL PAPER
DOI: 10.1002/ejoc.201500676
Enzymatic Synergism in the Synthesis of β-Keto Esters
[a]
[a]
[a]
[a]
´
Catalina Wisniewska, Dominik Koszelewski, Małgorzata Zysk, Szymon Kłossowski,
[a]
[a]
[a]
˙
Anna Zadło, Anna Brodzka, and Ryszard Ostaszewski*
˛
Keywords: Synthetic methods / Biotransformations / Enzyme catalysis / Transesterification / Alcohols / Esters
Reaction of alcohols with ethyl and tert-butyl acetoacetate
lected enzyme mixture for enzymatic transesterification of
catalyzed by a combination of commercially available en- various β-keto esters provided the respective products in ex-
zymes is shown to be a convenient method for the prepara- cellent yields up to 96% and quantitative within 24 and 48
tion of a range of acetoacetic acid derivatives. Systematic
studies proved that the combination of two or more enzymes
enhances the yield of the reaction. Application of the se-
hours, respectively. The presented methodology is simple
and mild, and can be used to prepare acetoacetates from pri-
mary and secondary alcohols.
chemical protocols have been explored to provide routes to
such compounds. There are a few procedures for the synthe-
sis of β-keto esters based on transesterification reaction of
ethyl or methyl acetoacetate with alcohols and, in general,
these reactions are catalyzed by protic or Lewis acids.[9]
Some of the drawbacks of homogeneous transesterification
include troublesome isolation of the product and large vol-
umes of waste generated during workup, and the catalysts
are not always recyclable. Furthermore, selectivity is often
low, resulting in the formation of unwanted mixtures of
products. More recently, various catalysts have been re-
ported to effect transesterification. Most of these methods
are not general and are equilibrium-driven reactions in
which the use of an excess of one of the reactants is manda-
tory to obtain the product in good yield. Transesterification
of β-keto esters is catalyzed by 4-(dimethylamino)pyridine
(DMAP) in good yield, but the application of this catalyst
is limited because of its toxicity and high price, and because
of the requirement for high temperature.[10] Furthermore,
the majority of protic acid catalysts are highly corrosive
and, hence, not environmentally friendly. These observa-
tions have led to the development of various heterogeneous
analogues for the transesterification of methyl and ethyl
acetoacetates in particular. The methods that are commonly
used employ titanium tetra-alkoxides, tin complexes or in-
dium iodide.[11] A few other homogeneous[12] and hetero-
geneous catalysts are also known to be effective for this
transformation.[13]
Introduction
β-Keto esters constitute a main class of organic building
blocks, and they are used for the efficient synthesis of a
range of complex natural products such as kermesic and
carminic acid esters, whic are commonly used as food-col-
oring additives.[1] The importance of β-keto esters stems
from the facile ability of such compounds to form bonds at
all four of the carbon atoms that make up their structural
unit; these consist of two different electrophilic carbonyl
carbon atoms and two nucleophilic carbon atoms, which
can react selectively under appropriate conditions. β-Keto
esters are commonly used in the pharmaceutical, agrochem-
ical, chemical, and polymer industries. They are suitable
substrates for the preparation of pyridazines,[2] function-
alized ketenimines,[3] aminopyrazole derivatives, and other
heterocycles.[4] Additionally, they have been successfully ap-
plied in cycloaddition reactions of halogenated quinines.[5]
Among the above organic compounds, β-keto esters are of
particular interest because their chelate complexes with
metals have found application as multifunctional additives
in various lubricating compositions, including those based
on oils of biological origin, as potential antiviral drugs, and
as catalysts for various stereoselective reactions.[6] Addition-
ally, β-keto esters are substrates for the Biginelli[7] reaction,
providing 3,4-dihydropyrimidin-2(1H)-ones, which show
antibacterial, and antiviral activities and they can be used
as a calcium channel blockers, mitotic kinesine inhibitors,
and adrenergic receptor antagonists.[8]
Depres and co-workers reported transesterification with-
out a catalyst with propargyl alcohols, but the reaction time
was very long (up to 12 days).[14]
In view of their medicinal importance, synthetic studies
on β-keto esters have attracted considerable interest. Several
Unfortunately, all the mentioned methods require high
temperatures.[15] In addition to this, the majority of cata-
lysts, especially metal complexes are toxic or expensive and
the products require complex purification procedures, so
they cannot be used in the pharmaceutical and cosmetic
[a] Institute of Organic Chemistry PAS,
Kasprzaka 44/52, 01-224 Warsaw, Poland
E-mail: ryszard.ostaszewski@icho.edu.pl
http://ww2.icho.edu.pl/z20/
Supporting information for this article is available on the
WWW under http://dx.doi.org/10.1002/ejoc.201500676.
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Enzymatic Synergism in the Synthesis of β-Keto Esters
industry. Another synthetic method to access β-keto esters synergy in degrading plant polymers by glucosidases and
is an acid-catalyzed reaction of aldehyde and diazo ester.[16] xylanases in aqueous medium,[26] this was, to our knowl-
A significant inconvenience of this method, however, is that edge, the first evidence of the cooperation between multiple
compounds containing a diazo group tend to be explosive, enzymes in organic solvents. Our previous work on enzyme-
unstable and toxic. For the synthesis of β-keto esters that catalyzed esterification of carboxylic acids prompted us to
are to be used in the pharmaceutical or food industry, regu- explore the application of the developed methodology in a
lations require that the procedures provide products with- consecutive way, using β-keto esters as the substrates. The
out inorganic and organic impurities. Especially in the aim of the presented work was to find a green alternative
pharmaceutical industry, for which pharmacopoeia limits to traditional chemical reactions, without use of high tem-
of heavy metal contaminations are below 5 ppm, most of perature or low pressure, by employing enzymes as cata-
the classical chemical methods with metal catalysts are un- lysts. The development of a method that allows transesteri-
acceptable. The development of economic and environmen- fication under neutral conditions should heighten the syn-
tally responsible methodologies for β-keto ester interconver- thetic potential of the reaction.
sion remains a challenge, because conventional processes
often lead to significant amounts of wastes and/or are per-
Results and Discussion
formed under harsh reaction conditions. We thus turn our
attention to enzymatic catalysis, because this approach al-
lows pure products to be obtained in high yields under mild
conditions, thereby fulfilling the general principles of green
chemistry.[17]
As was shown previously, a mixture of enzymes can cata-
lyze the esterification of carboxylic acids and is much more
efficient than the use of a single enzyme. Given that the
products were obtained with good to excellent yields, we
decided to use this method for the synthesis of β-acetoacet-
ates as another class of compounds.
Screening of biocatalysts was performed with 32 com-
mercially available enzymes in the model reaction con-
ducted in toluene at 40 °C. 2-Phenylethanol (2a) was chosen
for initial optimization study and reacted with ethyl aceto-
acetate for 24 hours to give 2-phenylethyl acetoacetate (3a)
(Scheme 1).
Jeromin and Welsch reported the synthesis of acetoacet-
ate esters catalyzed by lipase, applying diketene as the acyl
donor. Although this study was informative, it was narrow
in scope.[18] Typically, β-keto esters are prepared by decom-
posing highly reactive and unstable diketene with various
alcohols.[19] Although this methodology is atom economic,
the corrosiveness and handling difficulties of diketene make
it less attractive practically. The lachrymatory properties of
diketene together with concerns regarding its toxicity and
shipping have resulted in a need for alternative acetoacetyl-
ation technologies. One such “diketene-free” approach that
was described by Clemens, Hyatt, and Kato involves the
thermal reaction of 2,2,6-trimethyl-4H-dioxin-4-one with
nucleophiles to produce acetoacetic acid derivatives in good
yield.[20]
To our knowledge, there are only a few published pro-
cedures involving a single enzyme as a catalyst for the trans-
esterification of β-keto esters; unfortunately, no straightfor-
ward and general approach has been developed.
Janda and co-workers reported an elegant enzymatic
protocol for the transesterification of β-keto esters cata-
lyzed by immobilized Candida antarctica lipase B (CAL-B)
performed at 40 °C. However, although the products were
obtained with high yields, the reaction must be conducted
under low pressure (10 Torr), which required special equip-
ment.[21] Catalytic activity of Novozym 435 was also evalu-
ated in the systems involving nonactivated acyl donors, and
enhanced using microwave irradiation.[22] One of the major
drawbacks of this relatively new technology remains the
cost of the equipment. Furthermore, the microwave field is
usually nonuniform and localized superheating occurs.[23]
Finally, biocatalytic transesterification of β-keto esters in
ionic liquid (ILs) has been reported, but the sensitivity and
instability of the catalysts remain a serious problem.[24]
Scheme 1. Transesterification of ethyl/tert-butyl β-keto esters using
a range of alcohols.
To speed up the screening process, 32 enzymes were di-
vided into eight reaction mixtures (vials); each group con-
We recently reported the utilization of an enzyme mix- sisted of four different enzymes. It was observed that only
ture for the enzyme-catalyzed esterification of a range of one set – containing lipases from Candida antarctica (Novo-
carboxylic acids, providing the respective esters in excellent zym 435), Rhizopus niveus, lipoprotein lipase from Pseu-
yields.[25] Although there are a few examples of enzymatic domonas sp. and protease from papaine – catalyzed the
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R. Ostaszewski et al.
FULL PAPER
model reaction efficiently. An excess of starting β-keto ester ine the catalytic effect of the enzymes used, thermally deac-
1a (R1 = CH3, R2 = Et) was required to achieve high pro- tivated Candida antarctica present in Novozym 435 was ap-
ductivity in 24 hours (Table 1, entries 1–4). Given that ethyl plied together with native lipoprotein lipase from Pseudom-
acetoacetate is relatively inexpensive, it was used in fourfold onas sp. and vice versa; however, under these conditions,
excess in further experiments. The yield of isolated product only traces of product 3a were obtained.
3a was 92% (entry 4). The order of enzyme addition had
Moreover, the enzyme mixture could be reused without
no impact on the final result. Ester 3a was not observed in loss of activity by first washing it with ethyl ether and then
other groups. When the reaction was performed under re- drying in a dessicator. The preparation could be employed
duced pressure[21] no significant change in reaction effi- in more than five 48 hour cycles: the mixture of enzymes
ciency was observed, and the product 3a was obtained in retained 80% of the initial activity after five repetitive uses
87% yield.
in transesterification of ethyl acetoacetate with 2a.
Furthermore, this strategy could be used to create a β-
keto ester library. Both primary and secondary alcohols
were appropriate substrates for the enzyme mixture, as was
allylic alcohol (Table 2). These results then directed our at-
tention to possible donors of acetoacetate group. Experi-
ments with 2,2,6-trimethyl-4H-1,3-dioxine-4-one and tert-
butyl acetoacetate were performed; however, only with tert-
butyl acetoacetate was the corresponding product 3a ob-
tained with high isolated yield (88%; entry 2). The reaction
with 2,2,6-trimethyl-4H-1,3-dioxine-4-one was not cata-
lyzed by enzymes, which is probably because the acetyl-
ketene intermediate in transesterification can only be gener-
ated thermally.[27]
Table 1. Transesterification of ethyl acetoacetate (1a; R1 = Me, R2
= Et) with 2-phenylethanol (2a) catalyzed by combinations of en-
zymes.[a]
Entry
1 (equiv.)
Enzyme
Yield [%][b]
1
2
3
4
5
6
7
8
1
2
3
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
A, B, C, D
A, B, C, D
A, B, C, D
A, B, C, D
A
63
76
85
92
Ͻ 1
Ͻ 1
Ͻ 1
Ͻ 1
Ͻ 1
Ͻ 1
48
Ͻ 1
31
Ͻ 1
Ͻ 1
Ͻ 5
22
B
C
D
A, B
A, C
A, D
B, C
B, D
9
10
11
12
13
14
15
16
17
18
Table 2. Transesterification of acetoacetates with a range of
alcohols.[a]
C, D
Entry R1
R2
Alcohol 2 Yield [%][b] Yield [%][c]
A, B, C
A, B, D
A, C, D
B, C, D
1
2
3
4
5
6
7
8
CH3
Et
tBu
Et
tBu
Et
tBu
Et
tBu
Et
tBu
Et
tBu
Et
tBu
2a
2a
2b
2b
2c
2c
2d
2d
2e
2e
2f
92
88
80
76
96
72
67
86
Ͼ 99
Ͼ 99
98
96
Ͼ 99
94
89
Ͼ 99
83
Ͼ 99
72
Ͻ 5
92
CH3
CH3
CH3
CH3
CH3
CH3
CH3
CH3
CH3
CH3
CH3
CH3
CH3
Ͻ 5
[a] For a typical procedure, see the Exp. Section; A: Novozym 435,
B: protease from from Carica papaya, C: lipase from Rhizopus
niveus, D: lipoprotein lipase from Pseudomonas sp. [b] Isolated yield
after 24 h.
9
65
To find out which enzyme catalyzes the model reaction,
a series of experiments with each enzyme alone was per-
formed (Table 1). For each enzyme separately the yield of
isolated product was lower than 1% (entries 5–8), which is
consistent with results obtained in esterification of carb-
oxylic acids.[25] To identify the catalytic effect of enzyme
pairs, a second set of experiments was performed. Accord-
ing to the data included in Table 1, under standard condi-
tions, product 3a was obtained for some of these two-en-
zyme mixtures (entries 9–14). Two combinations of two en-
zymes gave the product in substantially higher yield (up to
10
11
12
13
14
15
16
84
30[d]
Ͻ 1
74
2f
2g
2g
2a
2b
Ͻ 1
78
84
Ͻ 5
Ͼ 99
Ͼ 99
4-MeOC6H4 Et
4-MeOC6H4 Et
[a] For a typical procedure, see the experimental section; 4 equiv.
of 1 was used. [b] Isolated yield after 24 h. [c] Isolated yield after
48 h. [d] Double acetylated product.
The results of the transesterification with various
48%; entries 11 and 13). However, this yield was still much alcohols are summarized in Table 2. In the majority of
lower in respect to the yield obtained with the four-enzyme cases, the products were obtained with excellent yields. For
mixture. Therefore, a further set of experiments was per- 2-phenylethanol (2a) and benzyl alcohol (2b) the corre-
formed with mixtures of three different enzymes (entries sponding products 3a and 3b were obtained with similar
15–18). Only, one combination of enzymes successfully pro- yields using ethyl or tert-butyl acetoacetate (entries 1–4). In
vided product 3a in 22% yield (entry 17). Two other enzyme case of chiral racemic alcohol 2c, slightly higher yield was
mixtures provided the product in less than 5% yield (entries obtained by using ethyl ester (entries 5 and 6). For alcohols
16 and 18). The presence of two enzymes: Novozym 435 with functional groups in the phenyl ring (–OCH3, –NO2)
and lipoprotein lipase from Pseudomonas sp. seem to be the yields of products 3d and 3e were higher for tert-butyl
crucial for productivity enhancement. Changing immobi- acetoacetate (entries 7–10). For cinnamyl alcohol (2g) and
lized Novozym 435 to native lipase from Candida antarctica 2-benzyl-1,3-propanediol (2f) the products 3f and 3g were
led to a significant drop in yield to less than 5%. To exam- obtained with good yield only with ethyl acetoacetate (en-
5434
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Enzymatic Synergism in the Synthesis of β-Keto Esters
tries 11–14). In the reaction of 2-benzyl-1,3-propanediol widespread application in academic and industrial fields.
(2f) with ethyl acetoacetate, 2-benzyl-1,3-propanediol di- Further research to widen the scope of this enhanced proto-
acetoacetate (3f) was obtained exclusively in 30% yield. col is in progress.
When the reaction time was extended to 48 h, the corre-
sponding products 3 were obtained with quantitative yields
(Table 2). To diversify the scope of the products that can be
obtained and to establish the generality of the developed
Experimental Section
General Experimental Methods: All the chemicals were obtained
from commercial sources. The solvents were of analytical grade.
NMR spectra were recorded in CDCl3 with TMS as an internal
standard using 200 MHz spectrometers. The chemical shifts are re-
ported in ppm (δ scale) and the coupling constants (J) are given in
Hertz (Hz). Enzymes: CAL-B (Novozym 435) {lipase acrylic resin
form Candida antarctica, Novozymes, 10 PLU/mg (PLU: propyl
laurate)}; protease from Carica papaya, SIGMA {lyophilized pow-
der, 12 U/mg (U: N-benzoyl-l-arginine ethyl ester)}; lipase from
Rhizopus niveus, Sigma {lyophilized powder, 1.5 U/mg (U: olive
procedure, ethyl 3-(4-methoxyphenyl)-3-oxopropionate was
applied to the enzymatic reaction, leading to the formation
of the corresponding esters 3h and 3i with quantitative
yields in 48 h (entries 15 and 16).
The enzymatic transesterification of tert-butyl acetoacet-
ate has, to our knowledge, not been reported. The applica-
tion of esterases or lipases for the hydrolysis of esters of
tertiary alcohols is hampered by the fact that most of the
commercially available enzymes do not accept tertiary
alcohols as substrates.[28] Only a limited group of enzymes oil)}; lipoprotein lipase from Pseudomonas sp., Sigma {lyophilized
powder, 50,000 U/mg (U: p-nitrophenyl butyrate)}. All enzymes
were stored in compliance with providers’ recommendations.
possessing a special active site structure are able to hy-
drolyze esters of tertiary alcohols.[29] The results obtained
in this study turned our attention to the mechanism of en-
zymatic transesterification of tert-butyl acetoacetate, which
is the subject of ongoing studies.
General Transesterification Procedure: Enzymatic reactions were
performed in a vortex (Heidolph Promax 1020) equipped with in-
cubator (Heidolph Inkubator 1000). All reactions were monitored
by TLC on Merck silica gel Plates 60 F254, detector UV/Vis, mo-
bile phase: hexanes/ethyl acetate (9:1, v/v).
Finally, we applied this transformation concept on a pre-
parative scale. Thus, 160 mg of benzyl alcohol (2b) was
transformed into ester 3i with complete conversion, within
48 h, and with 98% yield of the isolated product. It should
be noted that ester 3i is an important prodrug moiety.[30]
The mixture of enzymes appeared to be an excellent cata-
lyst system not only for the esterification of carboxylic
acids,[25] but also for the transesterification of acetoacetates,
and its use enables the synthesis of β-keto esters. It is clear
from Table 2 that the conversion from ethyl/tert-butyl keto
esters into higher homologues appears to be efficient and
practical through this procedure. It should be pointed out
that transesterification of β-keto esters with unsaturated
alcohols is rather difficult because it is offset by facile decar-
boxylation rearrangement; however, by using this method,
β-keto esters underwent such reactions smoothly.[31] The
present procedure is quite general for a wide range of struc-
turally varied alcohols. Neither high temperature, low pres-
sure, nor toxic catalysts are required in the synthesis The
procedure is very simple and provides the products with
quantitative yields. We believe our methodology will be of
ample use considering its mild conditions and simple pro-
cedure.
Experimental Setup: A solution of alcohol (0.5 mmol), ethyl or tert-
butyl acetoacetate (0.5, 1.0, 1.5 or 2 mmol) and 10 mg of each en-
zyme in toluene (3 mL) was placed in a 10 mL vial. The reaction
mixture was agitated at 200 rpm for 24 or 48 h at 40 °C. Enzymes
were removed by filtration, and toluene and remaining ethyl aceto-
acetate were removed under reduced pressure. The product was
purified by silica gel column chromatography (ethyl acetate/hex-
anes). The spectroscopic data were in accordance with reported
data.
Seven Other Sets of Enzymes Used in Screening:
1. Lipase from Candida rugosa (Sigma–Aldrich), lipase type II from
porcine pancreas (Sigma–Aldrich), cellulase from Aspergillus niger
(Sigma–Aldrich), acylase I from Aspergillus. melleus (Fluka).
2. Lipase from Candida lipolytica (Sigma–Aldrich), lipase from
Candida antarctica (Sigma–Aldrich), lipase from wheat germ, Am-
ano protease PS (Sigma–Aldrich).
3. Cellulase type VI from Trichoderma viride (Sigma–Aldrich), lyso-
zyme from chicken egg, pectinase from Rhizopus sp., Driselase Ba-
sidiomycetes sp. (Sigma–Aldrich).
4. Lipoprotein lipase from Pseudomonas sp. (Sigma–Aldrich), lipase
from Rhizopus niveus (Sigma–Aldrich) papain crude powder
(Merck), Novozym 435 (Novo Nordisk).
5. Amano lipase AK from Pseudomonas fluorescens (Sigma–Ald-
rich), lipase from Pseudomonas cepacia, lipase from hog pancreas
(Fluka), Amano lipase PS from Burkholderia cepacia (Sigma–Ald-
rich).
Conclusions
We have developed and demonstrated a viable new and
highly efficient procedure for the synthesis of a non-com-
mercial β-keto ester through transesterification in quantita-
tive yields. Noteworthy merits of this protocol are the sim-
ple operation, mild reaction conditions, and easy work-up
procedure, and no hazardous catalysts, or corrosive or toxic
solvents are required.
6. Lipase from Mucor javanicus (Sigma–Aldrich), lipase from Rhi-
zopus arrhizus (Sigma–Aldrich), lipase from Mucor miehei (Sigma–
Aldrich), lipase from Pseudomonas fluorescens (Sigma–Aldrich).
7. Lipase from Chromobacterium viscosum (Sigma–Aldrich), prote-
ase from Aspergillus oryzae (Sigma–Aldrich), lipase from Penicil-
lium roqueforti (Sigma–Aldrich), protease from Bacillus amylolique-
We consider that this is a more practical method than
the existing methodologies and expect that it should find faciens (Sigma–Aldrich).
Eur. J. Org. Chem. 2015, 5432–5437
© 2015 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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5435

R. Ostaszewski et al.
Synthesis of Benzyl 3-(4-Methoxyphenyl)-3-oxopropionate (3i): To a 191.2 ppm. C17H16O4 (284.31): calcd. C 71.82, H 5.67; found C
FULL PAPER
solution of benzyl alcohol (160 mg, 1.5 mmol) and ethyl acetoacet-
ate (780 mg, 6 mmol) and 25 mg of each enzyme in toluene (10 mL)
was placed in a 20 mL vial. The reaction mixture was agitated at
200 rpm for 48 h at 40 °C. Enzymes were removed by filtration and
toluene was evaporated under reduced pressure. The crude product
was purified by silica gel column chromatography (ethyl acetate/
hexanes) providing ester 3i (418 mg, 98%) as a colorless oil. The
spectroscopic data were in accordance with reported data.
71.78, H 5.55. HRMS (ESI): m/z calcd. for C17H16O4Na [M +
Na]+ 307.0946; found 307.0948.
Acknowledgments
This project was supported by Polpharma Starogard Gdan´ski. This
work was partially supported by the project “Biotransformations
for pharmaceutical and cosmetics industry” grant number
POIG.01.03.01-00-158/09-01, project number 2013/11/B/ST5/02199
and by the European Union (EU) in the frame of the COST Action
CM1303.
Analysis
2-Phenylethyl Acetoacetate (3a):[32] 1H NMR (CDCl3, 200 MHz): δ
= 2.20 (s, 3 H), 2.96 (t, J = 7.0 Hz, 2 H), 3.43 (s, 2 H), 4.37 (t, J =
7.0 Hz, 2 H), 7.22–7.31 (m, 5 H) ppm. 13C NMR (CDCl3,
50 MHz): δ = 26.5, 31.4, 46.5, 62.2, 123.1, 124.9, 125.3, 133.82,
167.1, 200,1 ppm.
Benzyl Acetoacetate (3b):[33] 1H NMR (CDCl3, 200 MHz): δ = 2.24
(s, 3 H), 3.50 (s, 2 H), 5.18 (s, 2 H), 7.35–7.37 (m, 5 H) ppm. 13C
NMR (CDCl3, 50 MHz): δ = 32.5, 71.6, 54.4, 132.6, 135.4, 136.3,
145.5, 175.2, 211.3 ppm.
1-Phenylethyl Acetoacetate (3c):[34] 1H NMR (CDCl3, 200 MHz): δ
= 1.57 (d, J = 6.4 Hz, 3 H), 2.23 (s, 3 H), 3.46 (s, 2 H); 5.93 (q, J
= 6.4 Hz, 1 H), 7.26–7.36 (m, 5 H) ppm. 13C NMR (CDCl3,
50 MHz): δ = 22.1, 30.3, 50.6, 73.7, 126.3, 128.3, 128.8, 139.0,
164.5, 198.6 ppm.
[1] a) S. J. Bingham, J. H. P. Tyman, J. Chem. Soc. Perkin Trans. 1
1997, 3637; b) G. Roberge, P. Brassard, J. Chem. Soc. Perkin
Trans. 1 1978, 1041–1046; c) V. Andrushko, N. Andrushko,
Stereoselective Synthesis of Drugs and Natural Products, John
Wiley & Sons, 2013, p. 1363; d) M. Marigo, K. Juhl, K. A.
Jørgensen, Angew. Chem. Int. Ed. 2003, 42, 1367–1369; Angew.
Chem. 2003, 115, 1405; e) I. Shinkai, The Structure, Reactions,
Synthesis and Uses of Heterocyclic Compounds, in: Comprehen-
sive Heterocyclic Chemistry II, vol. 3 (Ed.: I. Shinkai), Perga-
mon, Oxford, UK, 1996, p. 1–75.
[2] O. A. Attanasi, P. Filippone, C. Fiorucci, E. Foresti, F. Man-
tellini, J. Org. Chem. 1998, 63, 9880–9887.
[3] I. Yavari, F. Nourmohammadian, J. Chem. Res. 2000, 218–219.
2-(4-Nitrophenyl)ethyl Acetoacetate (3d):[35] 1H NMR (CDCl3, [4] a) F. Mathey, F. Robin, F. Mercier, M. Spagnol, PCT Int.
Appl., WO 98 00,375; Chem. Abstr. 1998, 128, 127603; b) K.
Niigata, T. Kimura, S. Hayashibe, H. Shikama, T. Takasu, E.
Hirasaki, Jpn. Kokai Tokkyo Koho, 1994, JP 06,298,737;
Chem. Abstr. 1995, 122, 13318; c) S. J. Binghama, J. H. P. Tym-
ana, Org. Prep. Proced. Int. 2001, 33, 357–361; d) R. V. Raga-
van, K. M. Kumar, V. Vijayakumar, S. Sarveswari, S. Ramaiah,
A. Anbarasu, S. Karthikeyan, P. Giridharan, N. S. Kumari,
Org. Med. Chem. Lett. 2013, 3, 6.
400 MHz): δ = 2.23 (s, 3 H), 3.08 (t, J = 6.4 Hz, 2 H), 3.45 (s, 2
H), 4.41 (t, J = 6.4 Hz, 2 H), 6.88 (d, J = 9.2 Hz, 2 H), 7.30 (d, J
= 9.2 Hz, 2 H) ppm. 13C NMR (CDCl3, 100 MHz): δ = 30.5, 34.9,
50.1, 64.9, 124.0, 129.9, 145.5, 167.1, 200.4 ppm.
4-Metoxybenzyl Acetoacetate (3e):[36] 1H NMR (CDCl3, 400 MHz):
δ = 2.23 (s, 3 H), 3.47 (s, 2 H), 3.81 (s, 3 H), 5.12 (s, 2 H), 6.88 (d,
J = 8.8 Hz, 2 H), 7.30 (d, J = 8.8 Hz, 2 H) ppm. 13C NMR (CDCl3,
50 MHz): δ = 30.4, 50.4, 55.5, 67.2, 114.2, 127.6, 130.5, 154.3,
168.5, 200.0 ppm.
2-Benzyl-1,3-propanediol Diacetoacetate (3f): 1H NMR (CDCl3,
400 MHz): δ = 2.28 (s, 6 H), 2.32–2.39 (m, 1 H), 2.69 (d, J = 7.6 Hz,
2 H), 3.48 (s, 4 H), 4.11–4.21 (m, 4 H), 7.16–7.30 (m, 5 H) ppm.
13C NMR (CDCl3, 100 MHz): δ = 30.4, 34.5, 39.3, 50.1, 64.5,
126.7, 128.8, 129.2, 138.5, 167.1, 200.6 ppm. C18H22O6 (334.37):
calcd. C 64.66, H 6.63; found C 64.63, H 6.67. HRMS (ESI): m/z
calcd. for C18H22O6Na [M + Na]+ 357.1314; found 357.1316.
Cinnamyl Acetoacetate (3g):[37] 1H NMR (CDCl3, 400 MHz): δ =
2.28 (s, 3 H), 3.50 (s, 2 H), 4.79 (dd, J = 1.2, 5.2 Hz, 2 H), 6.24–
6.31 (m, 1 H), 6.66 (d, J = 8 Hz, 1 H), 7.26–7.4 (m, 5 H) ppm. 13C
NMR (CDCl3, 100 MHz): δ = 30.4, 50.2, 66.1, 122.5, 126.8, 128.4,
128.8, 135.1, 136.1, 167.1, 200.6 ppm.
[5] M. Couturier, P. Brassard, Synthesis 1994, 703.
[6] a) L. I. Koval, V. I. Dzyuba, O. L. Ilnitska, V. I. Pekhnyo, Tet-
rahedron Lett. 2008, 49, 1645–1647; b) D. W. Hutchinson, Anti-
viral Res. 1985, 5, 193–205; c) E. Tramontano, F. Esposito, R.
Badas, R. Di Santo, R. Costi, P. La Colla, Antiviral Res. 2005,
65, 117–124.
[7] P. Biginelli, Gazz. Chim. Ital. 1893, 23, 360–416.
[8] a) H. Y. K. Kaan, V. Ulaganathan, O. Rath, H. Prokopcova,
D. Dallinger, C. O. Kappe, F. Kozielski, J. Med. Chem. 2010,
53, 5676; b) C. O. Kappe, Eur. J. Med. Chem. 2000, 35, 1043–
1052; c) M. Butters, C. D. Davies, M. C. Elliott, Org. Biomol.
Chem. 2009, 7, 5001–5009; d) J. S. S. Sandhu, ARKIVOC
(Gainesville, FL, U.S.) 2012, 1, 66–133.
[9] a) G. C. M. Kondaiah, L. A. Reddy, K. S. Babu, V. M. Gurav,
K. G. Huge, R. Bandichhor, P. P. Reddy, A. Bhattacharya,
R. V. Anand, Tetrahedron Lett. 2008, 49, 106–109; b) C. E.
Rehberg, C. H. Fisher, J. Am. Chem. Soc. 1944, 66, 1203; c)
C. E. Rehberg, W. A. Faucette, C. H. Fisher, J. Am. Chem. Soc.
1944, 66, 1723; d) C. E. Rehberg, Organic Synthesis Wiley, New
York, 1955, vol. 3, pp 146.
[10] a) F. T. Douglass, J. C. Amedio Jr., Y. K. Pate, J. Org. Chem.
1985, 50, 3619–3622; b) J. C. Gilbert, T. A. Kelly, J. Org. Chem.
1988, 53, 449–450.
[11] a) C.-T. Chen, J.-H. Kuo, C.-H. Ku, S.-S. Weng, C.-Y. Liu,
J. Org. Chem. 2005, 70, 1328–1339; b) J. S. Witzeman, W. D.
Nottingham, J. Org. Chem. 1991, 56, 1713–1718.
[12] a) P. Ilankumaran, J. G. Verkade, J. Org. Chem. 1999, 64, 3086;
b) K. Wakasugi, T. Misaki, K. Yamada, Y. Tanabe, Tetrahedron
Lett. 2000, 41, 5249.
[13] a) S. P. Chavan, K. Pasupathy, S. Shengule, V. Shinde, R. An-
and, ARKIVOC (Gainesville, FL, U.S.) 2005, 13, 162–168; b)
S. P. Chavan, P. K. Zubaidha, S. W. Dantale, A. Keshavaraja,
A. V. Ramaswamy, T. Ravindranathan, Tetrahedron Lett. 1996,
1
2-Phenylethyl 3-(4-Methoxyphenyl)-3-oxopropionate (3h): H NMR
(CDCl3, 200 MHz): δ = 2.93 (t, J = 7.2 Hz, 2 H), 3.86 (s, 3 H),
3.92 (s, 2 H), 4.36 (t, J = 7.2 Hz, 2 H), 6.91 (d, J = 9.0 Hz, 2 H),
7.18–7.32 (m, 5 H), 7.88 (d, J = 9.0 Hz, 2 H) ppm. 13C NMR
(CDCl3, 50 MHz): δ = 35.3, 46.2, 55.9, 66.2, 114.2, 127.0, 128.2,
128.9, 129.3, 129.5, 131.3, 137.9, 164.4, 168.1, 191.2 ppm.
C18H18O4 (298.34): calcd. C 72.47, H 6.08; found C 72.37, H 6.27.
HRMS (ESI): m/z calcd. for C18H18O4Na [M + Na]+ 321.1103;
found 321.1104.
Benzyl 3-(4-Methoxyphenyl)-3-oxopropionate (3i): 1H NMR
(CDCl3, 200 MHz): δ = 3.85 (s, 3 H), 3.97 (s, 2 H), 5.17 (s, 2 H),
6.90 (d, J = 8.8 Hz, 2 H), 7.31–7.37 (m, 5 H), 7.89 (d, J = 8.8 Hz,
2 H) ppm. 13C NMR (CDCl3, 50 MHz): δ = 46.2, 56.0, 67.6, 114.2,
127.4, 128.0, 128.7, 128.8, 129.1, 129.5, 131.3, 135.8, 164.4, 168.1,
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© 2015 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Eur. J. Org. Chem. 2015, 5432–5437

Enzymatic Synergism in the Synthesis of β-Keto Esters
37, 233; c) B. S. Balaji, M. Sasidharan, R. Kumar, B. Chanda,
Chem. Commun. 1996, 707; d) B. S. Balaji, B. M. Chanda, Tet-
rahedron 1998, 54, 13237; e) D. Ponde, V. H. Deshpande, V. J.
Bulbule, A. Sudalai, A. S. Gajare, J. Org. Chem. 1998, 63, 1058.
Ch. Mottet, O. Hamelin, G. Garavel, J. P. Depres, A. E.
Greene, J. Org. Chem. 1999, 64, 1380.
L. I. Koval, V. I. Dzyuba, O. L. Ilnitska, V. I. Pekhnyo, Tetrahe-
dron Lett. 2008, 49, 1645–1647.
C. R. Holmquist, E. J. Roskamp, J. Org. Chem. 1989, 54, 3258–
3260.
[26] a) M. Wang, G.-L. Lai, Y. Nie, S. Geng, L. Liu, B. Zhu, Z.
Shi, X.-L. Wu, Front. Microbiol. 6:509; DOI: 10.3389/
fmicb.2015.00509; b) M. Qi, H.-S. Jun, C. W. Forsberg, Appl.
Environ. Microbiol. 2007, 73, 6098–6105; c) J. Kopecˇny, A. G.
ˆ
Williams, Folia Microbiol. 1988, 33, 208–212; d) J. P. L. F.
Cairo, L. C. Oliveira, C. A. Uchima, T. M. Alvarez, A. P. S. Ci-
tadini, J. Cota, F. C. Leonardo, A. M. Costa-Leonardo, M. F.
Carazzolle, F. F. Costa, G. A. G. Pereira, F. M. Squina, Insect
Biochem. Mol. Biol. 2013, 43, 970–981; e) K. L. Kadam, E. C.
Rydholm, J. D. McMillan, Biotechnol. Prog. 2004, 20, 698–705;
f) L. Ji, J. Yang, H. Fan, Y. Yang, B. Li, X. Yu, N. Zhu, H.
Yuan, Biotechnol. Biofuels 2014, 7, 1–12.
[14]
[15]
[16]
[17]
[18]
M. Gupta, S. Paul, R. Gupta, Curr. Sci. 2010, 99, 1341–1360.
[27] A. D’Annibale, A. Pesce, S. Resta, C. Trogolo, Tetrahedron
Lett. 1996, 37, 7429–7432.
[28] R. Kourist, U. T. Bornscheuer, Appl. Microbiol. Biotechnol.
2011, 91, 505–517.
[29] E. Henke, J. Pleiss, U. T. Bornscheuer, Angew. Chem. Int. Ed.
2002, 41, 3211–3213; Angew. Chem. 2002, 114, 3338.
[30] C.-Q. Sun, P. T. W. Cheng, J. Stevenson, T. Dejneka, B. Brown,
T. C. Wang, J. A. Robl, M. A. Poss, Tetrahedron Lett. 2002, 43,
1161–1164.
[31] a) M. F. Carrol, J. Chem. Soc. 1940, 704–706; b) W. Kimel,
A. C. Cope, J. Am. Chem. Soc. 1943, 65, 1992–1998.
[32] S. Magens, M. Ertelt, A. Jatsch, B. Plietker, Org. Lett. 2008,
10, 53–56.
[33] S. Tayyari, F. Naghavi, S. Pojhan, R. W. McClurg, R. E. Sam-
melson, J. Mol. Struct. 2011, 987, 241–254.
[34] a) T. Hudlicky, T. Tsunoda, K. G. Gadamasetti, J. A. Murry,
G. E. Keck, J. Org. Chem. 1991, 56, 3619–3623; b) A. Nudel-
man, R. Kelner, N. Broida, H. E. Gottlieb, Synthesis 1989, 5,
387–388.
a) K. Suginaka, Y. Hayashi, Y. Yamamoto, Tetrahedron: Asym-
metry 1996, 7, 1153–1158; b) G. E. Jeromin, V. Welsch, Tetrahe-
dron Lett. 1995, 36, 6663.
a) S. R. Wilson, M. F. Price, J. Org. Chem. 1984, 49, 722; b) Y.
Tezuka, T. Ogura, S. Kawaguchi, Bull. Chem. Soc. Jpn. 1969,
42, 443.
[19]
[20] a) R. J. Clemene, J. A. Hyatt, J. Org. Chem. 1985, 50, 2431; b)
M. Sato, N. Kanuma, T. Kato, Chem. Pharm. Bull. 1982, 30,
1315; c) M. Sato, H. Ogaeawara, E. Yohizumi, T. Kato, Chem.
Pharm. Bull. 1983, 31, 1902; d) R. J. Clemens, J. S. Witzeman,
J. Am. Chem. Soc. 1989, 111, 2186.
[21] A. Cordova, K. D. Janda, J. Org. Chem. 2001, 66, 1906–1909.
[22] M. Rissoa, M. Mazzini, S. Kroger, P. Saenz-Mendez, G.
Seoane, D. Gamenara, Green Chem. Lett. Rev. 2012, 5, 539–
543.
[23] M. Nüchter, B. Ondruschka, W. Bonrath, A. Gum, Green
Chem. 2004, 6, 128–141.
[24] a) K. W. Kim, B. Song, M. Y. Choi, M. J. Kim, Org. Lett. 2001,
3, 1507; b) J. L. Kaar, A. M. Jesionowski, J. A. Berberich, R.
Moulton, A. J. Russell, J. Am. Chem. Soc. 2003, 125, 4125; c)
M. Persson, U. T. Bornscheuer, J. Mol. Catal. B 2003, 22, 21.
[25] M. Zysk, A. Zadlo, A. Brodzka, C. Wisniewska, R. Ostaszew-
ski, J. Mol. Catal. B 2014, 102, 225–229.
[35] N. Iqbal, E. E. Knaus, Arch. Pharm. 1996, 329, 23–26.
[36] T. Ogawa, A. Nakazato, K. Tsuchida, K. Hatayama, Chem.
Pharm. Bull. 1993, 41, 108–116.
[37] Y. Takekawa, K. Shishido, J. Org. Chem. 2001, 66, 8490–8503.
Received: May 27, 2015
Published Online: July 22, 2015
Eur. J. Org. Chem. 2015, 5432–5437
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