Adrenaline profiling of lipases and esterases with 1,2-diol and carbohydrate acetates

Article abstract of DOI:10.1016/j.tet.2003.11.059

The adrenaline test for enzymes is a general back-titration procedure to detect 1,2-diols, 1,2-aminoalcohols and α-hydroxyketones reaction products of enzyme catalysis by colorimetry. The method was used to profile a series of esterases and lipases for their esterolytic activity on a series of carbohydrate and polyol acetates. Substrates were prepared by peracetylation and used for parallel microtiter-plate analysis of enzyme activities. This method can be used to achieve a rapid and automated characterization of a set of enzymes during HTS screening.

Full text of DOI:10.1016/j.tet.2003.11.059

Tetrahedron 60 (2004) 703–710
Adrenaline profiling of lipases and esterases with 1,2-diol and
carbohydrate acetates
a
Denis Wahler, Olivier Boujard, Fabrice Lefevre and Jean-Louis Reymond
a
a
b
,
,
*
*
`
a
´
´
Proteus SA, 70 allee Graham Bell, 30000 Nimes, France
bDepartment of Chemistry and Biochemistry, University of Bern, Freiestrasse 3, 3012 Bern, Switzerland
Received 8 August 2003; revised 10 November 2003; accepted 19 November 2003
Abstract—The adrenaline test for enzymes is a general back-titration procedure to detect 1,2-diols, 1,2-aminoalcohols and
a-hydroxyketones reaction products of enzyme catalysis by colorimetry. The method was used to profile a series of esterases and lipases for
their esterolytic activity on a series of carbohydrate and polyol acetates. Substrates were prepared by peracetylation and used for parallel
microtiter-plate analysis of enzyme activities. This method can be used to achieve a rapid and automated characterization of a set of enzymes
during HTS screening.
q 2003 Elsevier Ltd. All rights reserved.
1. Introduction
even for laboratories not familiar with organic synthesis.
Herein we report that acetate derivatives of commercially
available carbohydrates and diols provide a readily
available substrate family for profiling lipases and esterases
based on the chromogenic adrenaline test for enzymes.
Enzyme assays suitable for high-throughput screening are
essential for the exploration of biological diversity in search
for novel enzymes,1 and the use of biocatalysis in industrial
processes is rapidly growing.2 The vast majority of
processes are based on hydrolytic enzymes, in particular
lipases and esterases, which are essential tools in fine
chemical synthesis. Several thousand different lipases and
esterases have been described, yet there is almost no
information available in the literature that allows comparing
these different enzymes in terms of their reactivities and
selectivities. We have described recently a general pro-
cedure for generating activity fingerprints of hydrolytic
enzymes using arrays of fluorogenic and chromogenic
substrates.3 The method was based on an indirect release
strategy including an oxidation step with NaIO4, followed
by a b-elimination leading to a fluorescent product, such
that various enzyme classes could be assayed including
alcohol dehydrogenases,4 aldolases,5 proteases,6 and
lipases.7 The diversity of reactivity patterns observed even
between closely related enzymes suggested that a large body
of differential reactivity information can indeed be estab-
lished by such procedures.8 Nevertheless each of the
substrates used in this study had to be individually
synthesized by asymmetric multistep synthesis, and was
available only in limited quantities. We wanted to explore
the possibility of using more readily available substrates
such that the method would become more broadly available
2. Results and discussion
The adrenaline test is a back-titration procedure which
measures the concentration of periodate-sensitive reaction
products formed from periodate-resistant substrates by an
enzymatic transformation.9 The test solution is treated with
a measured amount of sodium periodate, which rapidly
reacts with oxidizable functional groups presents, in parti-
cular 1,2-diols, 1,2-aminoalcohols or a-hydroxyketones
released from the hydrolysis of ester, amide, phosphate or
epoxide precursors used as substrates for the corresponding
enzymes. The unreacted periodate reagent is then revealed
by addition of adrenaline 1, which undergoes an instan-
taneous oxidation with periodate to give adrenochrome 2, a
cationic orthoquinone dye with a red absorption maximum
in the visible spectrum (Scheme 1). We had demonstrated
that this colorimetric procedure provided off-the-shelf
endpoint assays for lipases using vegetal oils as substrates,
phytases using phytic acid as substrate, and epoxide
hydrolases using epoxides as substrates.
In the perspective of fine chemical synthesis, kinetic
resolution of chiral alcohols by enantioselective lipases is
one of the most popular and reliable methods for producing
enantiomerically pure products. The preferred reactions are
the formation or hydrolysis of acetate esters of these
Keywords: Adrenaline; Esterases; Biocatalysis.
*
Corresponding authors. Tel.: þ33-46-6706463; fax: þ33-46-6706460;
e-mail addresses: flefevre@proteus.fr; jean-louis.reymond@ioc.unibe.ch
0040–4020/$ - see front matter q 2003 Elsevier Ltd. All rights reserved.
doi:10.1016/j.tet.2003.11.059

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D. Wahler et al. / Tetrahedron 60 (2004) 703–710
Table 1. Detection of free polyols by back-titration of sodium periodate
with adrenaline. EC50 is the product concentration that inhibits 50% of the
color reaction. Data calculated from OD decrease at 490 nm by increasing
polyol molar concentration, in the presence of 1 mM NaIO4 in 50 mM
aqueous borate buffer pH 8.0 for 1 h at 37 8C, followed by reaction with
1.5 mM of 1 for 5 min at 26 8C
Alcohol
Acetates
EC50 (mM)
D-Mannitol
D-Glucose
D-Lactose
D-Sorbitol
D-Mannose
D-Maltose
D-Galactose
D-Fructose
D-Xylose
D-Fucose
L-Arabinose
D-Arabinose
D-Ribose
D,L-Glyceraldehyde
2,3-Butanediol
Glycerol
1,2-Dodecanediol
Ethyleneglycol
1,2-Propanediol
1,2-Butanediol
Cis-1,2-cyclopentanediol
Cis-1,2-cyclohexanediol
1,2-Octanediol
(1R,2R)-trans-1,2-cyclohexanediol
3,3-Dimethyl-1,2-butanediol
(S)-(2)-1,1-Diphenyl-1,2-propanediol
Pinacol
37
28
31
32
24
36
34
29
25
34
35
30
23
13
10
8, 9
7
0.09
0.10
0.10
0.11
0.12
0.12
0.13
0.13
0.14
0.15
0.15
0.16
0.18
0.26
0.52
0.55
0.65
0.74
0.74
0.74
0.74
0.78
0.79
0.81
1.00
1.35
2.24
2.24
2.24
Scheme 1. Principle of the adrenaline test for esterolysis of 1,2-diol
acetates.
3
4
5
18
19
6
21
11
22
12
26
27
alcohols. However many enzymes show only very poor
reactivities towards acetates, and a preliminary screening
against such substrates is a necessary first step for assessing
the value of any enzyme collection in view of such
applications. It was shown recently that a multi-enzyme
detection kit for acetic acid can be used to test such
reactivities.10
D-Sucrose
D-Trehalose
We envisioned that our adrenaline test for enzymes would
provide an interesting opportunity for profiling such
reactivities with an array of 1,2-diol and carbohydrate
acetates. Indeed these substrates could be easily prepared by
peracetylation of the parent polyols. The resulting substrate
family was structurally diverse, such that the resulting
reactivity profiles would be useful for selecting enzymes on
the basis of their reactivity. Peracetate derivatives were
prepared by exhaustive acetylation with acetic anhydride in
pyridine. The products were purified simply by liquid
extraction and their structure confirmed by NMR-analysis.
The series was extended with a few commercially available
di- and triacetates to give a total of 35 different acetates
(Fig. 1).
carbohydrates undergo fivefold oxidation with periodate.
Sucrose, trehalose, and pinacol gave EC50 values in the
2 mM range, probably because their 1,2-diol functional
groups only react very slowly with periodate. This
calibration showed that all substrates were in principle
suited to generate a colorimetric signal upon complete
deacetylation. In general however, any partial deacetylation
liberating at least one 1,2-diol functionality would lead to a
decrease in the colorimetric signal in the adrenaline test.
A series of 33 lipases and esterases was assayed with the
peracetate substrate array shown in Figure 1, comprising 21
commercial lipases and 12 proprietary enzymes derived
´
from culture collections at Proteus (Table 2). These
The polyol substrates and their acetate derivatives were
conditioned as 10 mM stock solutions in water/acetonitrile
mixtures for the assay procedure. First, the reactivity of the
compounds against periodate in the adrenaline assay was
calibrated by testing serial dilutions of the products against
periodate (1 mM) followed by addition of 1 (1.5 mM). The
ester derivatives showed either full signal or only a weak
decrease of adrenochrome absorption with the adrenaline
test, in agreement with the absence of any free 1,2-diol
functionality in these derivatives. By contrast, the corre-
sponding polyols showed the expected reactivity, and the
results are shown in Table 1. The adrenaline procedure
efficiently detected most polyol in the expected range of
100 mM,EC50,750 mM based on the expected stoichio-
metry. Free hexitols and hexoses were detected with the
highest sensitivity in agreement with the fact that these
proprietary enzymes have been obtained during HTS
screening programs dedicated to the discovery of thermo-
philic esterases and lipases that can be used in the industrial
synthesis of fine chemical compounds at high temperature.
These enzymes, originating from strains isolated in either
deep sea hydrothermal vents, oil fields or hot terrestrial soil,
were characterized for their acyl chain length specificity
(ranging from C2 to C18).
The enzymes were incubated under a standard microtiter-
plate assay procedure consisting in incubating each of the 35
substrates at 1 mM in 100 mL of 50 mM aqueous borate
buffer pH 8.0, 5% DMF cosolvent, with 1 mg/mL enzyme
and 1 mM NaIO4 at 37 8C for 1 h, followed by treatment
with 1 for 5 min. The red color was quantitated at 490 nm
with a microtiter-plate reader. The relative signal intensity

D. Wahler et al. / Tetrahedron 60 (2004) 703–710
705
Figure 1. Array of polyol acetates used for adrenaline profiling.
was then corrected by subtracting the blank measure-
ment, which concerned mainly the acetylated fructose
derivative 29. The net signal thus obtained was then
used to generate activity patterns in grayscale, which is
a convenient display form for data analysis. The results
are shown in Figure 2.
Remarkably, the reactivity with the enzymes did not parallel
that of the blank data, showing that the specific enzyme
reactivities reflected selective interactions with the enzymes
rather than non-specific acceleration of the chemical
hydrolysis by the enzymes. Most enzymes showed the
strongest signal with acetylated carbohydrates in agreement
with the fact that these derivatives release the highest
number of periodate-sensitive 1,2-diols. All enzymes tested
showed individually differentiated reactivity patterns. These
patterns were reproducible and could thus be considered as
showing the intrinsic selectivity pattern of each enzyme.
The broad array of patterns observed illustrates the
enormous diversity in reactivities available in esterolytic
enzymes, despite of the fact that all of them are mostly
naturally selected to hydrolyze triglycerides. There was no
detectable association of the reactivity pattern observed
with the association of the enzyme to a lipase or esterase
type reactivity, in agreement with the fact that this
classification is problematic and sometimes grounded in
historical reasons.
The reactivity patterns obtained were strongly dependent
on the enzyme used. The reactivities observed should be
compared with the blank data observed without enzyme,
as well as with the values generated by reaction in the
presence of bovine serum albumin, which may be
considered as a measure of the non-specific reactivity of
the substrates. These blank values show an unusually high
reactivity of peracetylated fructose 29 and mannitol
hexaacetate 37 and may be associated with a spontaneous
partial deacetylation or intrinsic chemical unstability of
such substrates. The intrinsic reactivity of the substrates
was tested by treatment at higher temperatures under
conditions where complete solubility of all substrates was
ensured (Fig. 3). Most carbohydrate derivatives were
labile under these conditions, while simple aliphatic diols
not containing further oxygenated substituents were
clearly the most stable. The higher reactivity of carbo-
hydrate acetates compared to less substituted alcohol
esters might be related to their higher polarity as well as
to the lower pKa of their hydroxyl groups, which are
acidified by inductive effects of the neighbouring oxygen
atoms. Indeed the simple effect of stoichiometry (Table 1)
is not sufficient to explain the reactivity difference
between the two ester classes.
As for the proprietary enzymes, PE6 clearly resembled the
famous CAL-B enzyme often used for kinetic resolutions.
Remarkably, PE3 and PE4 displayed a high reactivity on the
hindered secondary ester 22 which was not found in any of
the other enzymes tested. PE7 was highlighted by a strong
and highly enantioselective reaction with the (R,R)-
enantiomer of trans-cyclohexanediol 21. This selectivity
was clearly more pronounced than that of ACE and PLE2 on
the 20/21 enantiomeric pair. These interesting reactivities
are under further investigation.

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D. Wahler et al. / Tetrahedron 60 (2004) 703–710
Table 2. Enzymes tested using the acetylated substrates array
Code
Enzymea
Specific activityb
(U/mg)
Specific periodate activity on 23c
(mU/mg)
Specific periodate activity on 9d
(mU/mg)
PE1
PE2
PE3
PE4
PE5
PE6
PE7
PE8
Proteus esterase number 1
Proteus esterase number 2
Proteus esterase number 3
Proteus esterase number 4
Proteus esterase number 5
Proteus esterase number 6
Proteus esterase number 7
Proteus esterase number 8
Proteus esterase number 9
Proteus esterase number 10
Proteus esterase number 11
Proteus esterase number 12
Electrophorus electricus AChE (F-01022)
Aspergillus lipase (F-84205)
Bacillus sp. esterase (F-46062)
Bacillus stearothermophilus esterase (F-46051)
Bacillus thermoglucosidasius esterase (F-46054)
Candida antarctica lipase (F-62299)
Candida antarctica lipase B
Candida cylindracea lipase (F-62316)
Candida cylindracea lipase (F-62302)
Candida lipolytica esterase (F-46056)
Hog kidney acylase I (F-01821)
Horse liver esterase (F-46069)
Hog pancreatic lipase (F-62300)
Mucor miehei lipase (F-62298)
Pseudomonas cepacia lipase (F-62309)
Pseudomonas fluorescens lipase (F-62312)
Pig liver esterase (S-E3019)
–
–
–
–
–
–
–
–
–
–
–
–
5.5
3.4
31.5
6.8
5.3
5.5
3.7
9.8
0.1
0.1
2.4
5.5
14.2
14.4
0.7
10.5
10.3
8.0
,0.1
0.3
8.2
1.0
5.2
5.4
0.4
0.7
0.1
0.7
,0.1
0.1
14.2
1.4
3.2
2.1
2.4
,0.1
14.7
4.8
3.7
3.5
3.5
1.8
,0.1
1.4
1.1
PE9
PE10
PE11
PE12
ACE
ASL
BAE
BSE
BTE
CAL
CAL B
CCL 1
CCL 2
CLE
HKA
HLE
HPL
MML
PCL
850e
0.5f
0.1g
0.4h
0.1g
3i
5.6
2i
13.1
12.1
13.8
14.7
2.8
14.7
1.8
9.0
10.8
13.9
14.8
7.9
13.9
13.1
20i
0.1j
16.5k
0.7l
25m
1i
50i
36i
15m
130g
10n
1.5o
10p
PFL
4.8
PLE 1
PLE 2
RAL
RNL
WGL
,0.1
14.8
0.9
2.6
0.8
Pig liver esterase (F-46058)
Rhizopus arrhizus lipase (F-62305)
Rhizopus niveus lipase (F-62310)
Wheat germ lipase (S-L3001)
a
Product reference from Fluka (F) or Sigma (S).
Activity given by the provider in units per milligram of powder; definition given in footnote.
b
c
Activity given in milliunits per milligram of powder of commercial samples, or per milligram of protein of proprietary enzyme samples; 1 U corresponds to
the amount of enzyme which liberates 1 mmol diol per minute at pH 8.0 and 37 8C, peracetylated D-ribose 23 as substrate.
Activity given in milliunits per milligram of powder of commercial samples, or per milligram of protein of proprietary enzyme samples; 1 U corresponds to
the amount of enzyme which liberates 1 mmol diol per minute at pH 8.0 and 37 8C, triacetin 9 as substrate.
1 U corresponds to the amount of enzyme which hydrolyzes 1 mmol acetylcholine per minute at pH 8.0 and 37 8C.
1 U corresponds to the amount of enzyme which liberates 1 mmol acetic acid per minute at pH 7.4 and 40 8C (triacetin as substrate).
1 U corresponds to the amount of enzyme which hydrolyzes 1 mmol of ethyl valerate per minute at pH 8.0 and 25 8C.
d
e
f
g
h
i
1 U corresponds to the amount of enzyme which releases 1 mmol 4-nitrophenol per minute at pH 7.0 and 65 8C (4-nitrophenyl-n-caproate as substrate).
1 U corresponds to the amount of enzyme which liberates 1 mmol oleic acid per minute at pH 8.0 and 40 8C (triolein as substrate).
1 U corresponds to the amount of enzyme which hydrolyzes 1 mmol of a-methyl-DL-phenylalanine-O-methylester per minute at pH 7.5 and 25 8C.
1 U corresponds to the amount of enzyme which hydrolyzes 1 mmol of N-acetyl-L-methionine per minute at pH 7.0 and 25 8C.
1 U will hydrolyze 1 mmol of ethyl butyrate to butyric acid and ethanol per minute at pH 8.0 and 25 8C.
j
k
l
m 1 U corresponds to the amount of enzyme which liberates 1 mmol fatty acid from triglycerides per minute at pH 8.0 and 37 8C (olive oil as substrate).
n
1 U corresponds to the amount of enzyme which liberates 1 mmol butyric acid per minute at pH 8.0 and 40 8C (tributyrin as substrate).
1 U corresponds to the amount of enzyme which liberates 1 mmol fatty acid from a triglyceride per minute at pH 7.7 and 40 8C (olive oil as substrate).
1 U will hydrolyze 1 mequiv. of fatty acid from a triglyceride in 1 h at pH 7.7 and 37 8C.
o
p
3. Conclusion
ditions, while lowering the level of false positive must be
used.11 At the end of the discovery process, novel enzymes
are validated for their intrinsic biochemical properties. This
step is generally cumbersome and time consuming.
Esterases and lipases are involved in industrial processes for
the synthesis of regioisomers of polyfunctional compounds,
resolutions of secondary alcohols and asymmetric synthesis
from prochiral or meso-compounds. The commercially
available enzymes are not robust and efficient enough to
investigate the feasibility of carrying out industrially
interesting syntheses, leading to increasing requests from
industry for novel enzymes. During the discovery processes
of such enzymes (which include microbial screening, HTS
gene isolation, and recombinant expression), the key point is
to be compatible with the timelines of the industry. To fit
with these timelines, robust and versatile HTS screening
tests that mimic the final industrial substrates and con-
The experiments above address this issue and demonstrate
that enzyme reactivity patterns can be readily obtained
using the adrenaline test and acetate esters of commercially
available diols and carbohydrates. Such reactivity profiling
can be readily established in any laboratory since the
synthetic chemistry involved is very limited and the
measurement method only uses inexpensive and simple
reagents. This biochemical fingerprint can be automated
using usual liquid handling systems. Comparison of
reactivity patterns of novel enzymes should help early

D. Wahler et al. / Tetrahedron 60 (2004) 703–710
707
Figure 2. Activity of esterases and lipases towards polyacetates 3–37 as determined by the adrenaline test. Each grayscale square corresponds to one ester
substrate according to the layout in Figure 1, and shows the percent reduction of OD490 from white (0%, no activity) to black (100%, maximum activity), after
correction from blank values. Conditions: (1) Enzyme in 50 mM aq. borate pH 8.0, 1 mM NaIO4, 1 mM substrate, 60 min at 37 8C; (2) 1.5 mM 1, 5 min, 26 8C.
Key for enzyme samples are given in Table 2. Commercial enzyme samples were tested at 1 mg/ml, proprietary esterases and lipases samples were used at one
tenth of crude enzyme extracts filtered through size-exclusion chromatography columns.
validation of discoveries and guide research efforts towards
those new enzymes displaying the most interesting and
novel reactivity patterns. In that respect, the fact that the
proprietary enzymes tested here all display patterns that
differ strongly from standard esterases and lipases highlights
the validity of pursuing biodiversity mining as a source for
new enzymes with novel selectivity properties.
4. Experimental
4.1. General
NMR spectra were recorded on a Bruker Avance 300 (1H:
300 MHz, 13C: 75 MHz) with a Bruker auto-sampler
B-ACS 120. Deuterated chloroform with the chemical shifts
1H: 7.26 ppm, 13C: 77.7 ppm, or deuterated methanol with
the chemical shifts 1H: 3.31, 4.79 ppm, 13C: 49.0 ppm were
used as solvents. All chemical shifts were expressed in ppm
according to the reference solvent peaks. Coupling con-
stants (J) are given in Hertz (Hz). Atoms of first acetate
Figure 3. Intrinsic hydrolytic lability of acetate substrates without enzyme.
Each grayscale square corresponds to one ester substrate according to the
layout in Figure 1, and shows the percent reduction of OD490 from white
(0%, no degradation) to black (100%, total consumption of sodium
periodate). Conditions: (1) 1 mM substrate in 50 mM aq. borate pH 8.0,
1 mM NaIO4, 60 min at the indicated temperature; (2) 1.5 mM adrenaline,
5 min, 26 8C.
0
residue were00quoted with , atoms of the second acetate
residue with , etc. The reactions were monitored by thin
layer chromatography (TLC) using plastic sheets with silica

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D. Wahler et al. / Tetrahedron 60 (2004) 703–710
gel 60 F254 (Merck). The melting points were determined
with a Bu¨chi Type S melting point apparatus and are
uncorrected.
NMR (300 MHz): d¼0.86 (t, 3H, J¼6.6 Hz, C8H3), 1.22
(m, 8H, C4H2, C5H2, C6H2, C7H2), 1.54 (m, 2H, C3H2),
2.04 (s, 3H, C20H3), 2.05 (s, 3H, C200H3), 4.01 (dd, 1H,
J¼11.8, 6.6 Hz, C1HH), 4.21 (dd, 1H, J¼12.0, 3.0 Hz,
C1HH), 5.05 (m, 1H, C2H1). 13C NMR (75 MHz): d¼14.7
(C8), 21.4 (C20), 21.7 (C200), 23.2 (C7), 25.7 (C6), 29.7 (C5),
31.3 (C4), 32.3 (C3), 65.8 (C1), 72.2 (C2), 171.3 (CvO0),
171.4 (CvO00).
Pyridine as the solvent for synthesis was purified and dried
before use. Polyols were obtained from Sigma-Aldrich-
Fluka or Lancaster. Diacetin, triacetin and ethyleneglycol
diacetate, as commercial polyacetate substrates, were
obtained from Fluka.
1
4.3.4. 1,2-Dodecanediol diacetate 7.15 Yellow syrup. H
4.2. General procedure for the preparation of
compounds 4–7, 10–11, 13, 15, 17–21, 24–25, 28, 30,
and 32–35
NMR (300 MHz): d¼0.83 (t, 3H, J¼6.4 Hz, C12H3), 1.20
(m, 16H, C4H2, C5H2, C6H2, C7H2, C8H2, C9H2, C10H2,
C11H2), 1.51 (m, 2H, C3H2), 2.01 (s, 3H, C20H3), 2.02 (s,
3H, C200H3), 3.97 (dd, 1H, J¼11.8, 6.6 Hz, C1HH), 4.17
(dd, 1H, J¼11.9, 3.2 Hz, C1HH), 5.01 (m, 1H, C2H1). 13C
NMR (75 MHz): d¼14.7 (C12), 21.3 (C20), 21.6 (C200), 23.3
(C11), 25.7 (C10), 29.8 (C9), 29.9 (C8), 30.0 (C7), 30.1
(C6), 30.2 (C5), 31.3 (C4), 3020 .5 (C3), 65.8 (C1), 72.3 (C2),
171.6 (CvO0), 171.7 (CvO ).
To an ice-bath cooled, stirred mixture of polyol in dry
pyridine were added 2 molar equiv. of acetic anhydride per
alcohol function. The reaction mixture was stirred over-
night, the temperature coming back slowly to room
temperature. The reaction was stopped by dilution with
water and ethylacetate, and the mixture washed with water
and brine. After separation and drying over magnesium
sulfate, the organic phase was concentrated to dryness.
Coevaporation under vacuum with pyridine and toluene of
the residue afforded pure polyacetate. Products were
characterized and validated by NMR analysis (proton and
carbon), and their homogeneity verified by TLC. Yields
obtained ranged from 83 to 99%.
4.3.5. 2,3-Butanediol diacetate 10.16 Yellow syrup. 1H
NMR (300 MHz): d¼1.19 (m, 3H, C1H3), 1.23 (m, 3H,
C4H3), 2.04 (s, 3H, C20H3), 2.06 (s, 3H, C200H3), 4.98 (m,
2H, C2H1, C3H1). 13C NMR (75 MHz): d¼8.3 (C1), 9.3
(C4), 14.4 (C20), 14.5 (C200), 64.6 (C2), 64.7 (C3), 163.7
(CvO0, CvO00).
4.3.6. 3,3-Dimethyl-1,2-butanediol diacetate 11. Yellow
1
4.3. General procedure for the preparation of
compounds 12, 22 and 29
syrup. H NMR (300 MHz): d¼0.94 (m, 9H, C4H3, C5H3,
C6H3), 2.01 (s, 3H, C20H3), 2.07 (s, 3H, C200H3), 3.98 (dd,
1H, J¼11.7, 8.9 Hz, C1HH), 4.37 (dd, 1H, J¼11.7, 2.4 Hz,
C1HH), 4.93 (dd, 1H, J¼8.9, 2.4 Hz, C2H1). 13C NMR
(75 MHz): d¼21.5 (C20), 21.6 (C200), 26.6 (C4, C5, C6),
34.2 (C3), 64.2 (C1), 67.4 (C2), 171.3 (CvO0), 171.6
(CvO00).
To an ice-bath cooled, stirred mixture of polyol in dry
pyridine were added 2 molar equiv. of acetic anhydride and
0.1 molar equiv. of 4-dimethylamino-pyridine per alcohol
function. The reaction mixture was stirred overnight, the
temperature coming back slowly to room temperature. The
reaction was stopped by dilution with water and ethyl-
acetate, and the mixture washed with water and brine. After
separation and drying over magnesium sulfate, the organic
phase was concentrated to dryness. Coevaporation under
vacuum with pyridine and toluene of the residue afforded
pure polyacetate. Products were characterized and validated
by NMR analysis (proton and carbon), and their homo-
geneity verified by TLC. Yields obtained ranged from 54 to
78%.
4.3.7. Pinacol diacetate 12.17 Yellow syrup. 1H NMR
(300 MHz): d¼1.20 (m, 12H, C1H3, C4H3, C5H3, C6H3,),
2.05 (m, 6H, C20, C200H3). 13C NMR (75 MHz): d¼22.4
(C1, C4, C5, C6), 25.6 (C20, C200), 75.3 (C2, C3), 172.1
(CvO0, CvO00).
4.3.8. D,L-Glyceraldehyde diacetate 13.18 White solid0. Mp
1
142–144 8C. H NMR (300 MHz): d¼2.08 (s, 3H, C2 H3),
2.14 (s, 3H, C200H3), 4.15 (dd, 1H, J¼12.4, 3.0 Hz, C3HH),
4.27 (dd, 1H, J¼12.5, 4.7 Hz, C3HH), 5.76 (d, 1H,
J¼7.9 Hz, C2H1). 13C NMR (75 MHz): d¼21.4 (C20),
21.6 (C200), 62.3 (C3), 75.0 (C2), 169.3 (CvO0), 171.2
(CvO00).
4.3.1. 1,2-Propanediol diacetate 4.12 Yellow syrup. 1H
NMR (300 MHz): d¼1.23 (d, 3H, J¼6.4 Hz, C3H3), 2.04 (s,
3H, C20H3), 2.05 (s, 3H, C200H3), 4.02 (dd, 1H, J¼11.8,
6.6 Hz, C1HH), 4.15 (dd, 1H, J¼11.8, 3.6 Hz, C1HH), 5.11
(m, 1H, C2H1). 13C NMR (75 MHz): d¼17.1 (C3), 210.4
(C20), 21.8 (C200), 66.7 (C1), 68.9 (C2), 171.1 (CvO ),
171.4 (CvO00).
4.3.9. (R)-(2)-Dihydro-5-(hydroxymethyl)-2(3H)-fura-
1
none acetate 15.19 Yellow syrup. H NMR (300 MHz):
d¼2.10 (s, 3H, C20H3), 2.35 (m, 2H, C3H2), 2.58 (m, 2H,
C2H2), 4.14 (dd, 1H, J¼12.2, 5.6 Hz, C5HH), 4.30 (dd, 1H,
J¼12.2, 3.2 Hz, C5HH), 4.73 (m, 1H, C4H1). 13C NMR
(75 MHz): d¼21.4 (C200), 24.6 (C3), 28.8 (C2), 66.0 (C5),
77.9 (C4), 171.2 (CvO ), 177.1 (CvO).
4.3.2. 1,2-Butanediol diacetate 5.13 Yellow syrup. 1H
NMR (300 MHz): d¼0.91 (t, 3H, J¼7.4 Hz, C4H3), 1.60
(m, 2H, C3H2), 2.04 (s, 3H, C20H3), 2.05 (s, 3H, C200H3),
4.03 (dd, 1H, J¼11.8, 6.6 Hz, C1HH), 4.21 (dd, 1H,
J¼11.8, 3.4 Hz, C1HH), 4.99 (m, 1H, C2H1). 13C NMR
(75 MHz): d¼10.1 (C4), 21.4 (C20), 21.7 (C200), 24.4 (C3),
65.4 (C1), 73.4 (C2), 171.2 (CvO0), 171.4 (CvO00).
4.3.10. (S)-(1)-Dihydro-5-(hydroxymethyl)-2(3H)-fura-
1
none acetate 17.19 Yellow syrup. H NMR (300 MHz):
d¼2.09 (s, 3H, C20H3), 2.34 (m, 2H, C3H2), 2.59 (m, 2H,
C2H2), 4.13 (dd, 1H, J¼12.2, 5.6 Hz, C5HH), 4.30 (dd, 1H,
J¼12.2, 3.2 Hz, C5HH), 4.73 (m, 1H, C4H1). 13C NMR
4.3.3. 1,2-Octanediol diacetate 6.14 Yellow syrup. 1H

D. Wahler et al. / Tetrahedron 60 (2004) 703–710
709
1
(75 MHz): d¼21.4 (C200), 24.6 (C3), 28.8 (C2), 66.0 (C5),
77.9 (C4), 171.2 (CvO ), 177.1 (CvO).
4.3.19. D-Fructose pentaacetate 29.28 0Yellow syrup. H
NMR (300 MHz): d¼2.04 (s, 3H, C2 H3), 2.06 (s, 3H,
C200H3), 2.08 (s, 3H, C2000H3), 2.09 (s, 3H, C20000H3), 2.10 (s,
3H, C200000H3), 4.16 (m, 2H, C5H2,), 4.75 (m, 2H, C6H2),
5.38 (m, 3H, C2H1, C3H1, C4H1). 13C NMR (75 MHz):
d¼21.3 (C20, C200, C2000, C20000, C200000), 62.3 (C5), 63.6 (C2),
64.1 (0C0 3), 67.0200 (C4), 68.8 (C6), 103.2 (C1), 170.2 (CvO0,
CvO , CvO , CvO0000, CvO00000).
4.3.11. cis-1,2-Cyclopentanediol diacetate 18.20 Yellow
syrup. 1H NMR (300 MHz): d¼1.80 (m, 12H, C3H2, C4H2,
C5H2, C20, C200H3), 5.12 (m, 2H, C1H1, C2H1). 13C NMR
(75 MHz): d¼19.8 (C4), 21.6 (C200,0 C200), 28.8 (C3, C5),
74.8 (C1, C2), 171.1 (CvO0, CvO ).
1
4.3.12. cis-1,2-Cyclohexanediol diacetate 19.21 Yellow
syrup. 1H NMR (300 MHz): d¼1.70 (m, 14H, C3H2, C4H2,
C5H2, C6H2, C20H3, C200H3,), 5.00 (m, 2H, C1H1, C2H1).
13C NMR (75 MHz): d¼21.8 (C20, C200), 22.3 (0C0 4, C5), 28.3
(C3, C6), 71.6 (C1, C2), 171.1 (CvO0, CvO ).
4.3.20. D-Arabinose tetraacetate 30.260 Yellow syrup. H
NMR (300 MHz): d¼2.02 (s, 3H, C2 H3), 2.06 (s, 3H,
C200H3), 2.12 (s, 3H, C2000H3), 2.15 (s, 3H, C20000H3), 4.10 (m,
2H, C5H2), 5.34 (m, 3H, C2H1, C3H1, C4H1), 6.34 (d, 1H,
J¼2.9 Hz, C1H1). 13C NMR00(0075 MHz): d¼21.2 (C20), 21.4
(C200), 21.6 (C2000), 21.7 (C2 ), 63.4 (C5), 67.3 (C2), 67.7
(C3), 69.1 (C4), 90.9 (C1), 169.8 (CvO0), 170.6 (CvO00),
170.8 (CvO000), 171.0 (CvO0000).
4.3.13. (1S,2S)-trans-1,2-Cyclohexanediol diacetate 20.22
Yellow syrup. 1H NMR (300 MHz):00 d¼1.70 (m, 14H,
C3H2, C4H2, C5H2, C6H2, C20H3, C2 H3,), 4.78 (0m0 , 2H,
C1H1, C2H1). 13C NMR (75 MHz): d¼21.8 (C20, C2 ), 24.1
(C4, C5), 30.8 (C3, C6), 74.4 (C1, C2), 171.1 (CvO0,
CvO00).
4.3.21. D-Sorbitol hexaacetate 32.29 White solid. Mp 87–
89 8C. 1H NMR (300 MHz): d¼2.05 (s, 3H, C20H3), 2.06 (s,
3H, C200H3), 2.07 (s, 3H, C2000H3), 2.08 (s, 3H, C20000H3), 2.09
(s, 3H, C200000H3), 2.13 (s, 3H, C2000000H3), 4.01 (dd, 1H,
J¼12.1, 6.2 Hz, C1HH), 4.12 (dd, 1H, J¼12.1, 5.2 Hz,
C6HH), 4.24 (dd, 1H, J¼12.1, 3.7 Hz, C1HH), 4.36 (dd, 1H,
J¼12.1, 4.3 Hz, C6HH), 5.05 (m, 1H, C2H1), 5.24 (m, 1H,
C5H1), 5.42 (m, 2H, C3H1, C4H1). 13C NMR (75 MHz):
d¼21.2 (C20, C200, C2000, C20000, C200000, C2000000), 62.1 (C1), 62.5
(C6), 69.0 (C2), 69.200(C5), 69.3 (C3), 70.0 (C4), 170.4
(CvO0), 170.5 (CvO ), 170.6 (CvO000), 170.7 (CvO0000),
171.1 (CvO00000), 171.2 (CvO000000).
4.3.14. (1R,2R)-trans-1,2-Cyclohexanediol diacetate 21.23
Yellow syrup. 1H NMR (300 MHz):00 d¼1.70 (m, 14H,
C3H2, C4H2, C5H2, C6H2, C20H3, C2 H3,), 4.78 (0m0 , 2H,
C1H1, C2H1). 13C NMR (75 MHz): d¼21.8 (C20, C2 ), 24.0
(C4, C5), 30.8 (C3, C6), 74.4 (C1, C2), 171.3 (CvO0,
CvO00).
4.3.15. (S)-(2)-1,1-Diphenyl-1,2-propanediol 2-acetate
1
22.24 White solid. Mp 137–139 8C. H NMR (300 MHz):
d¼1.09 (d, 3H, J¼6.4 Hz, C3H3), 1.87 (s, 3H, C20H3), 5.91
(q, 1H, J¼6.4 Hz, C2H1), 7.28 (m, 10H, CArom). 13C NMR
(75 MHz): d¼15.1 (C3), 21.8 (C20), 74.5 (C2), 80.2 (C1),
126.2, 126.5, 127.7, 128.9, 129.0, 143.8, 146.1 (CArom),
170.8 (CvO0).
4.3.22. D-Galactose pentaacetate 33.300 Yellow syrup. H
1
NMR (300 MHz): d¼1.98 (s, 3H, C2 H3), 1.99 (s, 3H,
C200H3), 2.02 (s, 3H, C2000H3), 2.03 (s, 3H, C20000H3), 2.14 (s,
3H, C200000H3), 4.07 (m, 2H, C5H1, C6HH), 4.32 (dd, 1H,
J¼12.1, 5.6 Hz, C6HH), 5.31 (m, 2H, C2H1, C4H1), 5.48
(m, 1H, C3H1), 6.35 (m, 1H, C1H1). 13C NMR (75 MHz):
d¼21.3 (C20, C200, C2000, C20000, C200000), 61.9 (C6), 67.1 (C4),
68.0 (C2), 68.1 (C3), 69.4 (C5), 90.3 (C1), 169.5 (CvO0),
170.0 (CvO00), 170.5 (CvO000), 170.8 (CvO0000), 171.0
(CvO00000).
1
4.3.16. D-Mannose pentaacetate 24.250 Yellow syrup. H
NMR (300 MHz): d¼1.99 (s, 3H, C2 H3), 2.03 (s, 3H,
C200H3), 2.07 (s, 3H, C2000H3), 2.15 (s, 3H, C20000H3), 2.16 (s,
3H, C200000H3), 4.09 (m, 2H, C5H1, C6HH), 4.26 (dd, 1H,
J¼12.2, 4.7 Hz, C6HH), 5.28 (m, 3H, C2H1, C3H1, C4H1),
6.06 (m, 1H, C1H1). 13C NMR (75 MHz): d¼21.3 (C20,
C200, C2000, C20000, C200000), 62.7 (C6), 66.2 (C4), 69.0 (C2), 69.4
(C3), 71.2 (C5), 91.2 (C1), 168.7 (CvO0), 170.1 (CvO00),
170.3 (CvO000), 170.6 (CvO0000), 171.2 (CvO00000).
4.3.23. D-Fucose tetraacetate 34.31 Yellow syrup. 1H NMR
(300 MHz): d¼1.13 (d, 3H, J¼6.4 Hz, C6H3), 1.98 (s, 3H,
C20H3), 1.99 (s, 3H, C200H3), 2.16 (s, 3H, C2000H3), 2.17 (s,
3H, C20000H3), 4.25 (m, 1H, C3H1), 5.31 (m, 3H, C2H1,
C4H1, C5H1), 6.31 (d, 1H, J¼2.8 Hz, C1H1). 13C NMR
(75 MHz): d¼21.3 (C6), 21.5 (C20, C200, C2000, C20000), 63.4
(C5), 66.8 (C2), 66.9 (C3), 68.0 (C4), 91.6 (C1), 169.4
(CvO0), 170.1 (CvO00), 170.4 (CvO000), 170.6 (CvO0000).
4.3.17. D-Xylose tetraacetate 25.26 Yellow syrup. 1H NMR
(300 MHz): d¼2.02 (s, 3H, C20H3), 2.04 (s, 3H, C200H3),
2.05 (s, 3H, C2000H3), 2.17 (s, 3H, C20000H3), 3.71 (dd, 1H,
J¼11.0 Hz, C5HH), 3.93 (dd, 1H, J¼11.0, 5.9 Hz, C5HH),
5.02 (m, 2H, C2H1, C4H1), 5.46 (dd, 1H, J¼9.5 Hz, C3H1),
5.47 (m, 1H, C3H1), 6.25 (d, 1H, J¼3.7 Hz, C1H1). 13C
NMR (75 MHz): d¼21.2 (C20), 21.3 (C200), 21.4 (C2000), 21.5
(C20000), 61.3 (C5), 69.300(C2, C4), 70.0 (C3), 89.9 (C1), 169.7
(CvO0), 170.4 (CvO ), 170.5 (CvO000), 170.8 (CvO0000).
1
4.3.24. L-Arabinose tetraacetate 35.320 Yellow syrup. H
NMR (300 MHz): d¼2.01 (s, 3H, C2 H3), 2.02 (s, 3H,
C200H3), 2.12 (s, 3H, C2000H3), 2.14 (s, 3H, C20000H3), 4.05 (m,
2H, C5H2), 5.30 (m, 3H, C2H1, C3H1, C4H1), 6.32 (d, 1H,
J¼2.6 Hz, C1H1). 13C NMR (75 MHz): d¼21.3 (C20, C200,
C2000, C20000), 63.4 (C5), 67.3 (C2), 67.7 (C3), 69.1 (C4), 90.8
(C1), 169.8 (CvO0), 170.5 (CvO00), 170.8 (CvO000), 170.9
(CvO0000).
4.3.18. D-Glucose pentaacetate 28.27 White solid.0 Mp
1
100–102 8C. H NMR (300 MHz): d¼2.01 (s, 3H, C2 H3),
2.04 (s, 3H, C200H3), 2.05 (s, 3H, C2000H3), 2.06 (s, 3H,
C20000H3), 2.09 (s, 3H, C20000H3), 4.09 (m, 2H, C5H1, C6HH),
4.26 (dd, 1H, J¼12.5, 4.1 Hz, C6HH), 5.11 (m, 2H, C2H1,
C4H1), 5.41 (m, 1H, C3H1), 6.32 (d, 1H, J¼3.7 Hz, C1H1).
4.4. Enzyme measurements
Substrates were diluted from 10 mM stock solutions in

710
D. Wahler et al. / Tetrahedron 60 (2004) 703–710
8. Reymond, J.-L.; Wahler, D. ChemBioChem 2002, 3, 701–708.
9. (a) Wahler, D.; Reymond, J.-L. Angew. Chem. Int. Ed. 2002,
41, 1229–1232. (b) . Reymond, J. -L.; Wahler, D. Patent
WO03046211, 2003-06-05.
water/acetonitrile mixtures. Enzymes were diluted from
10 mg mL21 stock solutions in aqueous borate buffer
(50 mM, pH 8.0). NaIO4 was added as a freshly prepared
10 mM stock solution in water. Adrenaline (as. HCl salt)
was added as a 15 mM stock solution in water. Assays
(0.1 mL) were conducted in individual wells of 96-well flat-
bottom half-area polystyrene microtiter-plates (Costar) as
described in the figure legends. The OD was recorded using
a Spectramax 190 Microplate Spectrophotometer (Molecu-
lar Devices, l¼490 nm).
10. Baumann, M.; Stu¨rmer, R.; Bornscheuer, U. T. Angew. Chem.
Int. Ed. 2001, 40, 4201–4204, Angew. Chem. 2001, 113,
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11. Reymond, J. -L.; Wahler, D.; Badalasssi, F.; Nguyen, H. K.
Patent WO0192563, 2001-12-07.
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Acknowledgements
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We are grateful to Professor B. K. C. Patel for some of the
strains isolation, to Dr. G. Ravot and V. Duvivier who took
part in the discovery and cloning of the proprietary esterases
and lipases, to Dr. O. Favre-Bulle and D. Buteux who have
achieved the fermentation of recombinant strains producing
the proprietary esterases/lipases, and to the other members
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´
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This was supported in part by the COST program, Action
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