New cholic acid derivatives: Biocatalytic synthesis and molecular docking study

Article abstract of DOI:10.1016/j.steroids.2015.12.014

A series of cholic acid derivatives was synthesized by enzyme catalysis. Eleven acetyl and ester derivatives of cholic acid, eight of them new compounds, were obtained through regioselective lipase-catalyzed reactions in very good to excellent yield. The influence of various reaction parameters in the enzymatic esterification, acetylation and alcoholysis reactions, such as enzyme source, alcohol or acylating agent: substrate ratio, enzyme: substrate ratio, solvent and temperature, was studied. Moreover, in order to shed light to cholic acid behavior in the enzymatic reactions, molecular docking of the lipase with cholic acid and some derivatives was carried out.

Full text of DOI:10.1016/j.steroids.2015.12.014

Accepted Manuscript
New cholic acid derivatives: biocatalytic synthesis and molecular docking study
M. Antonela Zígolo, Guadalupe García Liñares, Alicia Baldessari
PII:
S0039-128X(15)00312-8
http://dx.doi.org/10.1016/j.steroids.2015.12.014
STE 7887
DOI:
Reference:
Steroids
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Revised Date:
Accepted Date:
24 September 2015
7 December 2015
19 December 2015
Please cite this article as: Antonela Zígolo, M., Liñares, G.G., Baldessari, A., New cholic acid derivatives:
biocatalytic synthesis and molecular docking study, Steroids (2015), doi: http://dx.doi.org/10.1016/j.steroids.
2015.12.014
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New cholic acid derivatives: biocatalytic synthesis and
molecular docking study
M. Antonela Zígolo, Guadalupe García Liñares and Alicia Baldessari∗
Laboratorio de Biocatálisis. Departamento de Química Orgánica y UMYMFOR,
Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria,
Pabellón 2, piso 3, C1428EGA Buenos Aires, Argentina
∗Corresponding author: Alicia Baldessari. Phone: 5411 4576 3300 ext. 262; e-mail: alib@qo.fcen.uba.ar
1

Abstract
A series of cholic acid derivatives was synthesized by enzyme catalysis. Eleven acetyl and
ester derivatives of cholic acid, eight of them new compounds, were obtained through
regioselective lipase-catalyzed reactions in very good to excellent yield. The influence of
various reaction parameters in the enzymatic esterification, acetylation and alcoholysis
reactions, such as enzyme source, alcohol or acylating agent: substrate ratio, enzyme:
substrate ratio, solvent and temperature, was studied. Moreover, in order to shed light to
cholic acid behavior in the enzymatic reactions, molecular docking of the lipase with
cholic acid and some derivatives was carried out.
Keywords: cholic acid; lipase-catalyzed; molecular docking
2

1. Introduction
Enzymes are highly active and selective biocatalysts that turned out to be highly suitable
for the synthesis of many complex organic compounds [1,2]. The application of enzymes
for synthesizing specific compounds and the development of chemoenzymatic multi-step
processes are key research topics nowadays. Among the enzymes exploited for synthetic
purposes, lipases are very attractive due to their special properties: stability, recycling,
broad substrate tolerance and non-aqueous media activity. Due to their stereo-. chemo-
and regioselective nature, lipases have been employed for the preparation of compounds,
which are not easily obtainable by chemical methods. Steroids have been suitable
substrates for enzymatic transformations. In particular, the regioselective synthesis of
steroid derivatives containing multiple hydroxyl groups has been undertaken through
lipase-catalyzed acylation or deacylation procedures [3]. These mild and selective
interconversions of functional groups are more accessible than the standard chemical
approaches, because they avoid the tedious protection and deprotection steps [4].
Bile acids, widely distributed in nature as oxygenated metabolites of steroids, have been
the subject of numerous pharmacological studies [5,6]. Their amphiphilic character
generally increases cell membrane permeability [3]. For this reason their analogues are
widely used in drug formulations as excipients (intestinal absorption enhancers,
promoters, etc.) that can influence gastrointestinal solubility, and absorption of drugs [7].
O
OH
OH
HO
OH
1
Cholic acid (CA, 1) is one of the most important human bile acids and has attracted
significant attention primarily due to its wide availability and relatively low cost.
Synthetic analogues and derivatives of cholic acid have been widely explored in different
scientific areas such as combinatorial and supramolecular chemistry [8], syntheses of
various receptors [9], as antimalarials and antiproliferatives, etc. [10].
As other bile acids, CA exhibits two different faces: a hydrophobic face (β) containing
angular methyl groups and a hydrophilic face (α) with hydroxyl groups. This structure is
responsible for their amphipathic nature and, consequently, in water self-associates to
form multimers above a critical concentration, where hydrogen bonding plays a key role.
Because of its amphipathic properties and ability to form micelles, cholic acid has been
used as a building block in supramolecular chemistry to transport ion and polar molecules
across the membrane [11].
Unlike other bile acids, CA is the only one that has three hydroxyl groups at the C3, C7
and C12 positions facing the concave side of the molecule. This feature gives higher water
3

solubility and lower detergent power than the other bile acids [12]. Sometimes, the
potential use of bile acid is difficult because of their low solubility in both hydrophilic and
hydrophobic media. In order to find an increase in lipophilicity and enhance permeability
features, we have recently reported the synthesis of derivatives of chenodeoxycholic
(CDA), deoxycholic (DA) and lithocholic (LA) acids [13].
In the present work we describe an enzymatic strategy for the synthesis of a series of new
derivatives of cholic acid, obtained by lipase-catalyzed esterification, acetylation and
alcoholysis reactions. Due to its singular structure containing three hydroxyl groups, in the
enzymatic reactions CA showed a different behavior from that observed for the three bile
acids previously studied, containing one (LA) or two hydroxyl groups (CDA and DA). In
fact, we set about examining the particular reaction conditions to obtain CA derivatives.
This paper reports the results of our investigation in detail. Moreover, with the aim to
rationalize the selective behavior of lipases with CA as substrate, molecular modeling
studies were applied.
2. Experimental
2.1. General
Chemicals and solvents were purchased from Merck Argentina and Sigma-Aldrich de
Argentina and used without further purification. Lipase from Candida rugosa (CRL) (905
U/mg solid) and type II crude form porcine pancreas lipase ((PPL) (190 U/mg protein)
were purchased from Sigma Chemical Co.; Candida antarctica lipase B (CAL B):
Novozym 435 (7400 PLU/g) and Lipozyme RM 1M (LIP) (7800 U/g) were generous gifts
of Novozymes Spain and Carica papaya lipase (CPL) is the remaining solid fraction of
papaya latex, after wash off of proteases using distilled water was a generous gift of Dr.
Georgina Sandoval (CIATEJ). All enzymes were used "straight from the bottle".
Enzymatic reactions were carried out on Innova 4000 digital incubator shaker, New
Brunswick Scientific Co. at the corresponding temperature and 200 rpm. E/S: enzyme
amount in mg/substrate amount in mg. To monitor the reaction progress aliquots were
withdrawn and analyzed by TLC performed on commercial 0.2 mm aluminum-coated
silica gel plates (F254) and visualized by 254 nm UV or immersion in an aqueous solution
of (NH4)6Mo7O24.4H2O (0.04 M), Ce(SO4)2 (0.003 M) in concentrated H2SO4 (10%). %
Conversion was determined by analytical reverse-phase HPLC employing a Phenomenex
Phenogel column 5µM 10E5A, 300 x 7.8 mm and eluting with MeOH:H2O 80:20 at 1.00
mL/min. Melting points were measured in a Fisher Johns apparatus and are uncorrected.
Optical rotation values were measured in a CHCl3 solution with a Perkin Elmer-343
1
automatic digital polarimeter at 25 °C. H NMR and 13C NMR spectra were recorded at
room temperature in CDCl3 as solvent using a Bruker AM-500 NMR instrument operating
1
1
at 500.14 MHz and 125.76 MHz for H and 13C respectively. The H NMR spectra are
referenced with respect to the residual CHCl3 proton of the solvent CDCl3 at δ = 7.26
ppm. Coupling constants are reported in Hertz (Hz). 13C NMR spectra were fully
decoupled and are referenced to the middle peak of the solvent CDCl3 at δ = 77.0 ppm.
4

Splitting patterns are designated as: s, singlet; d, doublet; t, triplet; q, quadruplet; qn,
quintet; dd, double doublet, etc. IR spectra were recorded with a Nicolet Magna 550
spectrometer. High Resolution Mass Spectrometry was recorded with Thermo Scientific
EM/DSQ II – DIP. The results were within 0.02% of the theoretical values.
2.2. Enzymatic esterification. General procedure
Lipozyme (1g) was added to a solution of cholic acid (100 mg) in DIPE (10 mL) and the
corresponding alcohol (20 eq). The suspension was shaken (200 rpm) at 40 °C and the
progress of the reaction monitored by TLC/HPLC. Once the reaction was finished, the
enzyme was filtered off and the solvent evaporated under reduced pressure. The residue
was purified by column chromatography on silicagel employing mixtures hexane:ethyl
acetate as eluent (6:4-2:8). Reuse experiments: the filtered and washed enzyme was used
in the next enzymatic esterification under the same reaction conditions.
2.3. Enzymatic acetylation. General procedure
CAL B (1 g) was added to a solution of ethyl cholate (100 mg) in hexane (10 mL) and
ethyl acetate (0.25 mL). The suspension was shaken (200 rpm) at 40 °C and the progress
of the reaction monitored by TLC/HPLC. Once the reaction was finished, the enzyme was
filtered off and the solvent evaporated under reduced pressure. The residue was purified
by column chromatography on silicagel employing mixtures hexane:ethyl acetate as eluent
(7:3-1:1). Reuse experiments: the filtered and washed enzyme was used in the next
enzymatic acetylation under the same reaction conditions.
2.4. Enzymatic alcoholysis. General procedure
To a solution of the peracetylated cholic acid (100 mg) in 10 ml of the indicated solvent
containing 5 mol equiv of ethanol, CAL B (1 g) was added. The suspension was shaken
(200 rpm) at 40 °C and the progress of the reaction was monitored by TLC. After
indicated time, the enzyme was filtered off, the solvent was evaporated, and the crude
residue purified by column chromatography on silica gel employing mixtures hexane:ethyl
acetate as eluent (8:2-1:1).
2.5. Chemical peracetylation
In a typical procedure, a mixture of cholic acid (400 mg, 1 mmol), acetic anhydride (2 mL,
20 mmol), pyridine (2 mL) and DMAP (10 mg) was stirred at room temperature for 4 hs.
The progress of the reaction was monitored by TLC. After completion of the reaction, the
mixture was partitioned between saturated solution ammonium chloride (20 mL) and
methylene chloride (20 mL). The aqueous phase was extracted with methylene chloride (3
x 10 mL). The combined organic layers were washed with saturated solution of sodium
chloride (5 x 20 mL), dried (MgSO4), and the solvent was evaporated. The residue was
purified by column chromatography (silica gel) employing mixtures of hexane/EtOAc as
eluent (8:2–1:1).
5

Ethyl 3α,7α,12α−trihydroxy-5β-cholanate (2a): Yield 85 %; white solid, mp 160–161
°C; [α]D25 + 22.2° (c 1.0, CHCl3). 1H NMR (CDCl3, 500 MHz) δ 4.12 (2H, q, J = 6.7 Hz,
-OCH2CH3), 3.96 (1H, t, J = 2.9 Hz, H-12), 3.83 (1H, dd, J = 2.5, 3.0 Hz, H-7), 3.43 (1H,
tt, J = 11.2, 4.1 Hz, H-3), 2.35 (1H, m, H-23b), 2.22 (1H, m, H-23a), 1.24 (3H, t, J = 6.8
Hz, -OCH2CH3), 0.98 (3H, d, J = 6.4 Hz, H-21), 0.88 (3H, s, H-19), 0.67 (3H, s, H-18);
13C NMR (CDCl3, 500 MHz) δ 174.4 (C-24), 73.1 (C-12), 71.9 (C-7), 68.4 (C-3), 60.2 (-
OCH2CH3), 30.9 (C-23), 22.4 (C-19), 17.3 (C-21), 14.2 (-OCH2CH3), 12.5 (C-18).
HRMS: [M + Na]+ Calcd. C26H44NaO5 459.3086 Found: C26H44NaO5 459.3090.
Propyl 3α,7α,12α−trihydroxy-5β-cholanate (2b): Yield 70 %; white solid, mp 120–121
°C; [α]D25 + 25.3° (c 1.0, CHCl3). 1H NMR (CDCl3, 500 MHz) δ 4.02 (2H, t, J = 6.7 Hz, -
OCH2CH2CH3), 3.97 (1H, t, J = 3.0 Hz, H-12), 3.84 (1H, dd, J = 2.5, 3.0 Hz, H-7), 3.45
(1H, tt, J = 10.8, 4.0 Hz, H-3), 2.35 (1H, m, H-23b), 2.23 (1H, m, H-23a), 1.64 (2H, m, -
OCH2CH2CH3), 0.98 (3H, d, J = 6.2 Hz, H-21), 0.95 (3H, t, J = 7.0 Hz, -OCH2CH2CH3),
0.88 (3H, s, H-19), 0.68 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz) δ 174.4 (C-24), 73.0
(C-12), 71.9 (C-7), 68.4 (C-3), 65.9 (-OCH2CH2CH3), 30.9 (C-23), 22.5 (C-19), 17.3 (C-
21), 12.5 (C-18), 10.4 (-OCH2CH2CH3). HRMS: [M + Na]+ Calcd. C27H46NaO5 473.3243
Found: C27H46NaO5 473.3239.
Butyl 3α,7α,12α−trihydroxy-5β-cholanate (2c): Yield 66 %; white solid, mp 163–164
°C; [α]D25 + 23.8° (c 1.0, CHCl3). 1H NMR (CDCl3, 500 MHz) δ 4.06 (2H, t, J = 6.7 Hz, -
OCH2CH2CH2CH3), 3.98 (1H, t, J = 2.9 Hz, H-12), 3.85 (1H, dd, J = 2.5, 3.0 Hz, H-7),
3.45 (1H, tt, J = 10.8, 4.0 Hz, H-3), 2.35 (1H, m, H-23b), 2.23 (1H, m, H-23a), 1.61 (2H,
m, -OCH2CH2CH2CH3), 0.98 (3H, d, J = 6.2 Hz, H-21), 0.92 (3H, t, J = 7.0 Hz, -
OCH2CH2CH2CH3), 0.89 (3H, s, H-19), 0.69 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz)
δ 174.4 (C-24), 72.9 (C-12), 72.0 (C-7), 68.4 (C-3), 64.2 (-OCH2CH2CH2CH3), 30.9 (C-
23), 22.5 (C-19), 17.3 (C-21), 13.7 (-OCH2CH2CH2CH3), 12.5 (C-18). HRMS: [M + Na]+
Calcd. C28H48NaO5 487.3399 Found: C28H48NaO5 487.3392.
Dodecyl 3α,7α,12α−trihydroxy-5β-cholanate (2d): Yield 55 %; white solid, mp 71–72
°C; [α]D25 + 26.2° (c 1.0, CHCl3). 1H NMR (CDCl3, 500 MHz) δ 4.05 (2H, t, J = 6.7 Hz, -
OCH2(CH2)10CH3), 3.98 (1H, t, J = 2.8 Hz, H-12), 3.85 (1H, dd, J = 2.5, 3.0 Hz, H-7),
3.45 (1H, tt, J = 10.9, 4.2 Hz, H-3), 2.35 (1H, m, H-23b), 2.23 (1H, m, H-23a), 1.59 (2H,
m, -OCH2CH2(CH2)9CH3), 0.98 (3H, d, J = 6.2 Hz, H-21), 0.89 (3H, s, H-19), 0.88 (3H, t,
J = 7.0 Hz, -OCH2(CH2)10CH3), 0.68 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz) δ 174.4
(C-24), 73.0 (C-12), 71.9 (C-7), 68.4 (C-3), 64.5 (-OCH2(CH2)10CH3), 30.9 (C-23), 22.5
(C-19), 17.3 (C-21), 14.1 (-OCH2(CH2)10CH3), 12.5 (C-18). HRMS: [M + Na]+ Calcd.
C36H64NaO5 599.4651 Found: C36H64NaO5 599.4647.
Tetradecyl 3α,7α,12α−trihydroxy-5β-cholanate (2e): Yield 54 %; white solid, mp 65-
25
1
66 °C; [α]D + 30.1° (c 1.0, CHCl3). H NMR (CDCl3, 500 MHz) δ 4.05 (2H, t, J = 6.7
Hz, -OCH2(CH2)12CH3), 3.98 (1H, t, J = 2.9 Hz, H-12), 3.85 (1H, dd, J = 2.4, 3.0 Hz, H-
6

7), 3.45 (1H, tt, J = 11.0, 4.0 Hz, H-3), 2.35 (1H, m, H-23b), 2.22 (1H, m, H-23a), 1.59
(2H, m, -OCH2CH2(CH2)11CH3), 0.98 (3H, d, J = 6.1 Hz, H-21), 0.89 (3H, s, H-19), 0.87
(3H, t, J = 7.0 Hz, -OCH2(CH2)12CH3), 0.68 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz) δ
174.4 (C-24), 73.0 (C-12), 71.9 (C-7), 68.4 (C-3), 64.5 (-OCH2(CH2)12CH3), 30.9 (C-23),
22.5 (C-19), 17.3 (C-21), 14.1 (-OCH2(CH2)12CH3), 12.5 (C-18). HRMS: [M + Na]+
Calcd. C38H68NaO5 627.4964 Found: C38H68NaO5 627.4968.
Hexadecyl 3α,7α,12α−trihydroxy-5β-cholanate (2f): Yield 58 %; white solid, mp 74-75
°C; [α]D25 + 29.3° (c 1.0, CHCl3). 1H NMR (CDCl3, 500 MHz) δ 4.05 (2H, t, J = 6.7 Hz, -
OCH2(CH2)14CH3), 3.98 (1H, t, J = 2.9 Hz, H-12), 3.84 (1H, dd, J = 2.4, 3.0 Hz, H-7),
3.45 (1H, tt, J = 10.9, 4.0 Hz, H-3), 2.36 (1H, m, H-23b), 2.22 (1H, m, H-23a), 1.60 (2H,
m, -OCH2CH2(CH2)13CH3), 0.98 (3H, d, J = 6.1 Hz, H-21), 0.89 (3H, s, H-19), 0.88 (3H, t,
J = 7.0 Hz, -OCH2(CH2)14CH3), 0.68 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz) δ 174.4
(C-24), 73.0 (C-12), 71.9 (C-7), 68.4 (C-3), 64.5 (-OCH2(CH2)14CH3), 30.9 (C-23), 22.5
(C-19), 17.3 (C-21), 14.1 (-OCH2(CH2)14CH3), 12.5 (C-18). HRMS: [M + Na]+ Calcd.
C40H72NaO5 655.5278 Found: C40H72NaO5 655.5280.
Octadecyl 3α,7α,12α−trihydroxy-5β-cholanate (2g): Yield 51 %; white solid, mp 81-82
°C; [α]D25 + 30.6° (c 1.0, CHCl3). 1H NMR (CDCl3, 500 MHz) δ 4.05 (2H, t, J = 6.8 Hz, -
OCH2(CH2)16CH3), 3.98 (1H, t, J = 3.0 Hz, H-12), 3.84 (1H, dd, J = 2.5, 3.0 Hz, H-7),
3.45 (1H, tt, J = 11.0, 4.1 Hz, H-3), 2.35 (1H, m, H-23b), 2.23 (1H, m, H-23a), 1.60 (2H,
m, -OCH2CH2(CH2)15CH3), 0.98 (3H, d, J = 6.0 Hz, H-21), 0.89 (3H, s, H-19), 0.88 (3H, t,
J = 7.0 Hz, -OCH2(CH2)16CH3), 0.68 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz) δ 174.4
(C-24), 73.0 (C-12), 71.9 (C-7), 68.4 (C-3), 64.5 (-OCH2(CH2)16CH3), 30.9 (C-23), 22.5
(C-19), 17.3 (C-21), 14.1 (-OCH2(CH2)16CH3), 12.5 (C-18). HRMS: [M + Na]+ Calcd.
C42H76NaO5 683.5591 Found: C42H76NaO5 683.5590.
Ethyl 3α-acetoxy-7α,12α−dihydroxy-5β-cholanate (3a): Yield 87 %; white solid, mp
25
1
95–96 °C; [α]D + 24.5° (c 1.0, CHCl3). H NMR (CDCl3, 500 MHz) δ 4.58 (1H, tt, J =
11.2, 4.2 Hz, H-3), 4.12 (2H, q, J = 6.7 Hz, -OCH2CH3), 3.99 (1H, t, J = 2.9 Hz, H-12),
3.85 (1H, dd, J = 2.5, 3.0 Hz, H-7), 2.35 (1H, m, H-23b), 2.22 (1H, m, H-23a), 1.25 (3H, t,
J = 6.7 Hz, -OCH2CH3), 0.98 (3H, d, J = 6.5 Hz, H-21), 0.90 (3H, s, H-19), 0.70 (3H, s, H-
18); 13C NMR (CDCl3, 500 MHz) δ 174.2 (C-24), 170.8 (COCH3), 74.3 (C-12), 72.7 (C-
7), 68.2 (C-3), 60.2 (-OCH2CH3), 30.9 (C-23), 22.5 (C-19), 21.5 (CH3CO-), 17.3 (C-21),
14.2 (-OCH2CH3), 12.6 (C-18). HRMS: [M + Na]+ Calcd. C28H46NaO6 501.3192 Found:
C28H46NaO6 501.3195.
3α,7α,12α-triacetoxy-5β-cholanic acid (4a): Yield 90 %; white solid, mp 69.5–70 °C;
25
1
[α]D + 26.9° (c 1.0, CHCl3). H NMR (CDCl3, 500 MHz) δ 5.08 (1H, t, J = 3.0 Hz, H-
12), 4.90 (1H, dd, J = 3.0, 2.5 Hz, H-7), 4.57 (1H, tt, J = 11.2, 4.3 Hz, H-3), 2.32 (1H, m,
H-23b), 2.22 (1H, m, H-23a), 2.14 (3H, s, COCH3), 2.09 (3H, s, COCH3), 2.05 (3H, s,
COCH3), 0.91 (3H, s, H-19), 0.81 (3H, d, J = 6.5 Hz, H-21), 0.73 (3H, s, H-18); 13C NMR
7

(CDCl3, 500 MHz) δ 176.6 (C-24), 170.5 (2 x COCH3), 170.4 (COCH3), 75.4 (C-12),
74.1 (C-7), 70.7 (C-3), 30.8 (C-23), 22.6 (C-19), 21.6 (CH3CO-), 21.5 (CH3CO-), 21.4
(CH3CO-), 17.5(C-21), 12.2 (C-18). HRMS: [M + Na]+ Calcd. C30H46NaO8 557.3090
Found: C30H46NaO8 557.3086.
Ethyl 3α,7α,12α-triacetoxy-5β-cholanate (4b): Yield 92 %; white solid, mp 69.5–70 °C;
25
1
[α]D + 28.0° (c 1.0, CHCl3). H NMR (CDCl3, 500 MHz) δ 5.08 (1H, t, J = 3.0 Hz, H-
12), 4.90 (1H, dd, J = 3.0, 2.5 Hz, H-7), 4.57 (1H, tt, J = 11.2, 4.3 Hz, H-3), 4.11 (2H, q, J
= 6.7 Hz, -OCH2CH3), 2.32(1H, m, H-23b), 2.20 (1H, m, H-23a), 2.13 (3H, s, COCH3),
2.08 (3H, s, COCH3), 2.04 (3H, s, COCH3), 1.24 (3H, t, J = 6.7 Hz, -OCH2CH3), 0.91 (3H,
s, H-19), 0.81 (3H, d, J = 6.5 Hz, H-21), 0.73 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz)
δ 174.1 (C-24), 170.6 (COCH3), 170.5 (COCH3), 170.4 (COCH3), 75.4 (C-12), 74.1 (C-7),
70.7 (C-3), 60.2 (-OCH2CH3), 30.8 (C-23), 22.6 (C-19), 21.6 (CH3CO-), 21.5 (CH3CO-),
21.4 (CH3CO-), 17.5(C-21), 12.2 (C-18). HRMS: [M + Na]+ Calcd. C32H50NaO8 585.3403
Found: C32H50NaO8 585.3408.
Ethyl 7α,12α−diacetoxy-3α-hydroxy-5β-cholanate (5b): Yield 70 %; white solid, mp
25
1
75–76 °C; [α]D + 28.0° (c 1.0, CHCl3). H NMR (CDCl3, 500 MHz) δ 5.08 (1H, t, J =
3.0 Hz, H-12), 4.90 (1H, dd, J = 3.0, 2.5 Hz, H-7), 4.11 (2H, q, J = 6.7 Hz, -OCH2CH3),
3.64 (1H, tt, J = 11.0, 4.1 Hz, H-3), 2.32 (1H, m, H-23b), 2.20 (1H, m, H-23a), 2.13 (3H,
s, COCH3), 2.08 (3H, s, COCH3), 1.25 (3H, t, J = 6.7 Hz, -OCH2CH3), 0.90 (3H, s, H-19),
0.81 (3H, d, J = 6.5 Hz, H-21), 0.72 (3H, s, H-18); 13C NMR (CDCl3, 500 MHz) δ 174.1
(C-24), 170.7 (COCH3), 170.6 (COCH3), 75.4 (C-12), 71.7 (C-7), 70.8 (C-3), 60.3 (-
OCH2CH3), 30.8 (C-23), 22.6 (C-19), 21.6 (CH3CO-), 21.5 (CH3CO-), 17.5(C-21), 12.2
(C-18). HRMS: [M + Na]+ Calcd. C30H48NaO7 543.3298 Found: C30H48NaO7 543.3301.
2.6. Modecular modeling
CAL
B
structure
was
downloaded
from
RCSB
Protein
DataBank
(http://www.rcsb.org/pdb/), PDB code for CAL B is 1TCA. All substrates were
minimized using semiempirical AM1 method with the algorithm Polak-Ribiere in
Hyperchem. The docking calculations were carried out with Autodock 4.2 program [14].
The Autodock 4.2 method was applied considering a pair of rotatable bonds for cholic
acid and all the protein were defined it was considered stiff. For the location and extent of
the 3D area, the search space is defined by specifying a center, the number of points in
each dimension, and points between spaces to focus the search space in the active site of
the enzyme, it was taken as grid box center the coordinates x, y, z of a water molecule
close to Ser105
3. Results and Discussion
In this section we are going to describe the results obtained by applying lipases as
catalysts in three reactions: esterification, acetylation and alcoholysis.
8

3.1. Enzymatic Esterification
According to literature, only methyl and ethyl esters of cholic acid have been described so
far. CA methyl ester was prepared using methanol and acids such as hydrochloric acid,
methanesulfonic or p-toluensulfonic acid [15-17]. Ethyl ester was obtained by treatment
with absolute ethanol and fuming sulfuric acid (20-30% SO3) [18].
With the aim to get a mild procedure applied also to longer chain alcohols, we carried out
the esterification of CA with different alcohols and lipases as catalysts (Scheme 1).
Scheme 1. Enzymatic synthesis of esters of CA (1).
In order to optimize the reaction conditions we performed several experiments such as
lipase screening and variation of the reaction parameters such as solvent, temperature,
enzyme: substrate ratio (E/S) and alcohol: acid ratio (Nu/S).
Initially we examined the catalytic activity of five lipases, four of them commercially
available, in the esterification reaction of 1 with ethanol: Candida rugosa lipase (CRL),
Candida antarctica lipase B (CAL B), Lipozyme, lipase from the fungus Rhizomucor
miehei (LIP), porcine pancreas lipase (PPL) and Carica papaya lipase (CPL). Some
satistactory results are shown in Table 1. Among the enzymes tested, only LIP and CAL B
were active enough toward the esterification of CA. In terms of conversion the former was
the best while with CRL, PPL and CPL and in the absence of biocatalyst no product was
obtained.
Although several studies have already been reported on the effect of organic solvents on
lipase-catalyzed reactions [19], it is still difficult to predict the solvent effect. Therefore
the practical way is to select an appropriate solvent through screening experiments. We
tested hexane, diisopropyl ether (DIPE) and acetonitrile as low, medium and high polar
solvents respectively. In addition, the reaction without co-solvent using ethanol both as
nucleophile and solvent was also performed.
9

Table 1. Optimization of reaction parameters for lipase-catalyzed esterification of CA
with ethanol.
E/Sa
Entry Lipase
Solvent
T (°C)
Nu/S
t (h)
Conversion
Lipase and solvent
CAL B
CAL B
CAL B
CAL B
LIP
Ethanol
Hexane
DIPE
40
40
40
40
40
40
40
40
10
10
10
10
10
10
10
10
20
20
20
20
20
20
20
20
96
18
43
40
-
1
2
3
4
5
6
7
8
96
96
Acetonitrile
Ethanol
Hexane
DIPE
> 96
72
60
50
85
10
LIP
72
LIP
48
LIP
Acetonitrile
> 96
E/S
9
LIP
LIP
LIP
LIP
DIPE
DIPE
DIPE
DIPE
40
40
40
40
50
20
5
20
20
20
20
48
48
48
48
87
84
50
22
10
11
12
1
Nu/S
13
LIP
LIP
LIP
DIPE
DIPE
DIPE
40
40
40
10
10
10
500
100
10
48
48
48
88
85
76
14
15
Temperature
LIP
DIPE
DIPE
30
55
10
10
20
20
96
48
-
16
LIP
50
17
__________________________________________________________________________________________________________________________________
a E/S: enzyme amount in mg/substrate amount in mg.
Among the tested solvents, the lipase from Rhizomucor miehei (LIP) showed the highest
activity in DIPE, affording 2a at 85% conversion in 48 h of reaction (Table 1, entry 7).
This lipase was also active in hexane and without co-solvent, in neat ethanol, but to a
lesser extent (Table 1, entries 5 and 6). CAL B showed lower activity: about 40%
conversion in hexane and DIPE after 96 h. (Table 1, entries 2 and 3).
Regarding the influence of temperature on the enzymatic esterification, the reaction was
tested at 30 °C, 40 °C and 55 °C using an enzyme:substrate ratio E/S: 10 and
ethanol:substrate ratio: 20. The dependence of the reaction performance with temperature
was interesting because no reaction was observed at 30 °C and yield was only 50% at 55
10

°C affording 2a with a mixture of products. Therefore 40 °C was selected as the optimum
temperature.
To further optimize the reaction efficiency, the influence of the enzyme:substrate ratio in
the enzymatic esterification was evaluated at 48 h, using ethanol as nucleophile at Nu/S
ratio of 20, DIPE as solvent at 40 °C and variable amounts of LIP (Table 1, entries 9 -12).
From the results, it can be observed that an E/S ratio of 50, 20 and 10 (Table 1, entries 9,
10 and 7) are very similar. A conversion slightly lower was observed at E/S ratio of 10 and
this was the ratio of choice, considering the convenience of using the lowest amount of
enzyme. Anyway, although E/S = 10 could seem very high for a catalyst, in general the
amount of lipase in immobilized enzymes is very low. For example the ratio lipase:support
is 30 mg of lipase/g support in the case of CAL B [20]. Moreover, due the immobilization,
LIP can be reused. In this particular case it was observed that it retained 80 % activity
after three reaction cycles.
Finally, the influence of the Nu/S ratio on esterification yield was evaluated at 48 h using
LIP (E/S = 10) in DIPE at 40 °C, reaction time 48 h and variable amounts of ethanol.
From the results (Table 1, entries 13-16), it can be observed that there is not an important
dependence of the reaction yield with the ratio Nu/S ratio. Then, Nu/S: 20 was the ratio of
choice.
Considering the previously mentioned experiments, the following standard conditions for
the enzymatic esterification of cholic acid with ethanol were chosen: LIP as biocatalyst,
temperature: 40 °C, E/S: 10, Nu/S: 20 and DIPE as solvent (Table 1, entry 7).
After the establishment of the optimal conditions, the enzymatic strategy in the
esterification of 1 with a variety of alcohols was explored (Table 2).
Table 2. Lipase-catalyzed obtention of cholic acid esters
Entry
Alcohol
Product
Yield (%)
Ethanol
85
70
66
55
54
58
51
1
2
3
4
5
6
7
2a
2b
2c
2d
2e
2f
n-Propanol
n-Butanol
n-Dodecanol
n-Tetradecanol
n-Hexadecanol
n-Octadecanol
2g
Reaction conditions: Enzyme: LIP; Solvent: DIPE; Temperature: 40 °C; E/S: 10;
Nu/S: 20; t: 48 h.
Alkyl esters containing linear alkyl chains from two to eighteen carbon atoms were
synthesized in moderate to very good yield (50-80 %). It was observed that yield
decreased as the alkyl chain length increased. Every product was completely characterized
by spectroscopic methods and six of them (2b-2g) were not previously described in
11

literature. The advantage of the enzymatic procedure becomes more relevant in the case of
longer-chain derivatives, where the use of excess of alcohol in the traditional synthetic
approach is not convenient, in terms of product isolation and economy.
3.2. Enzymatic acetylation
At our best knowledge, the preparation of monoacetyl derivatives of cholic acid was not
reported so far. Some acyloxy derivatives of cholic acid, such as butanoyl, decanoyl- and
hexadecanoyloxy CA were prepared through protection and deprotection steps [21]. In a
previous work, peracetylation of methyl ester CA was carried out in acetic anhydride,
pyridine and DMAP at room temperature under microwave irradiation [16]. In the same
report, it was also described the obtention of methyl 3α-hydroxy-7α,12α-diacetoxy-5β-
cholanate by deacetylation using methanesulfonic acid in methanol. Moreover, a one-pot
esterification of carboxyl group and 3α-acetylation of CA was carried out using the
appropriate esters (ethyl-, propyl- and butyl acetate) in p-toluensulfonic acid and water at
reflux. Under these conditions the reaction was not selective and the monoacetyl
derivative was obtained in a mixture with ethyl 3α,7α-diacetoxy-12α-hydroxy-5β-
cholanate [22].
Recently, we have reported the synthesis of 3α-acetyl derivatives of chenodeoxycholic
and deoxycholic acids using CAL B as biocatalyst, ethyl acetate as acylation agent in
hexane at 55 °C. Under the same reaction conditions, with lithocholic acid as substrate, the
lipase catalyzed both the acetylation and esterification reactions affording ethyl
lithocholate 3-acetate as the only product in one step [13].
The presence of the three hydroxyl groups in CA makes it an interesting model for
studying the selectivity of the enzymatic acetylation. Therefore, we applied the enzymatic
procedure, studying a variety of biocatalysts and reaction conditions in the acetylation of
CA. Unfortunately no reaction was observed in any case, even when using activated
acylating agents such as vinyl or isopropenyl acetate. From these results and considering
the work previously reported, it is obvious that it is not possible to acetylate CA in a
regioselective way, chemically or enzymatically.
After this result, we decided to explore the reaction conditions for the lipase-catalyzed
acetylation of CA ethyl ester (2a) (Scheme 2). As usual, we conducted a screening varying
different parameters: lipases, solvents, temperature and E/S and acylating agent A/S ratios.
The results can be observed in Table 3.
Scheme 2. Enzymatic acetylation of ethyl cholate (2a).
12

In the absence of biocatalyst no product was obtained. In this reaction CAL B was the
only active enzyme giving the best results in hexane. The lipase was highly regioselective
affording ethyl 3-acetyl cholate (3a) at 90 % conversion in 96 h of reaction as the only
acetylation product (Table 3, entry 2). CAL B was also active in DIPE but to a lesser
extent (Table 3, entry 3) and working without co-solvent with ethyl acetate as acylating
agent and solvent, 3a conversion decreased to 70 % (Table 3, entry 1).
Table 3. Optimization of reaction parameters for lipase-catalyzed acetylation of ethyl
cholate (2a).
E/Sa
Entry Lipase
Solvent
Solvent
T (°C)
A/S
t (h) Conversion
(%)
CAL B
CAL B
CAL B
Ethyl acetate 40
10
10
10
-
96
70
90
65
1
2
Hexane
40
10
96
DIPE
40
10
96
3
E/S
CAL B
CAL B
CAL B
Hexane
Hexane
Hexane
40
40
40
5
2
1
10
10
10
96
96
96
76
54
35
4
5
6
A/S
7
CAL B
CAL B
CAL B
Hexane
Hexane
Hexane
40
40
40
10
10
10
20
5
96
96
96
90
80
61
8
1
9
Temperature
CAL B Hexane
CAL B Hexane
30
55
10
10
10
10
96
96
-
10
11
90
______________________________________________________________________
Lipases: LIP, PPL and CPL: inactive.
a E/S: enzyme amount in mg/substrate amount in mg.
Taking into account these studies, we have chosen as optimal conditions for the enzymatic
acetylation of CA ethyl ester: CAL B as biocatalyst, ethyl acetate as acylating agent,
hexane as solvent, temperature: 40 °C, E/S and A/S: 10.
Finally, in order to analyze the influence of the ester chain length on enzyme reactivity,
we applied the optimal conditions to the acetylation of the lauryl ester of CA (2d). Under
these conditions, no reaction was observed.
13

It is interesting to note that, despite the differences in structure and water solubility, in the
enzymatic acetylation of bile acid ethyl esters, CAL B showed the same regioselective
behavior for ethyl esters of CA, chenodeoxycholic and deoxycholic acid, affording the
monoacetyl derivative in the 3α position in every case [13].
3.3. Enzymatic alcoholysis
Since enzymatic acetylation was not a suitable way to get mono- or diacetyl derivatives of
cholic acid, we tried to carry out the enzymatic alcoholysis of peracetylated derivatives
both CA and its ethyl ester. Because CAL B was the only active enzyme in the acetylation
reaction, it was the enzyme of choice.
The substrates for the enzymatic alcoholysis 4a and 4b, were obtained in 90 % and 92%
yield respectively, by treatment of 1 and 2a with acetic anhydride and pyridine at room
temperature during 16 h.
Scheme 3. Chemical synthesis and enzymatic alcoholysis of peracetylated CA (4a) and
its ethyl ester (4b).
At this point, we attempted to carry out the enzymatic alcoholysis of 4a and 4b using
ethanol as nucleophile and DIPE or hexane as solvents (Table 4). The best results were
achieved when employing ethanol as nucleophile and hexane as solvent.
To our surprise under the above mentioned conditions, the alcoholysis of peracetylated
cholic acid (4a) afforded 3,7,12-triacetyl ethyl ester (4b) showing that deacetylation did
not take place in any of the three acyl positions. The lipase catalyzed the esterification of
the carboxyl group with ethanol instead of performing the nucleophilic attack to acetate
groups. On the other hand, with 4b as substrate, alcoholysis occurred at 3-position while
keeping unaltered the other two (7 and 12). Therefore, ethyl 7,12-diacetyl cholate (5b) was
obtained, using CAL B as catalyst (E/S:10), ethanol as nucleophile (Nu/S:1.2) and hexane
as solvent at 40 °C, after 72 h of reaction. It is remarkable that, at longer periods of time,
some deacetylation product afforded from 4b obtained from enzymatic alcoholysis of 4a.
14

Table 4. CAL B-catalyzed alcoholysis of 3α,7α,12α -triacetyl cholic acid (4a) and ethyl-
3α,7α,12α -triacetyl cholate (4b).
Conversion (%)a
Entry Substrate Solvent
Nu
Nu/S t (h)
4b
5b
Hexane
Hexane
DIPE
EtOH
EtOH
EtOH
EtOH
EtOH
EtOH
BuOH
OctOH
1.2
5
72
48
72
72
72
72
72
72
95
85
10
30
1
2
3
4
5
6
7
8
4a
4a
4a
4a
4b
4b
4b
4b
1.2
5
DIPE
Hexane
Hexane
DIPE
1.2
5
50
74
10
15
5
DIPE
5
______________________________________________________________________________________
Experimental conditions: Enzyme: CAL B; Temperature: 40 °C; E/S: 10.
3.4. Molecular Modeling
From the experimental results it is clear that only CAL B was able to perform the
acetylation of CA ethyl ester within reasonable reaction time and at mild reaction
conditions. It was also active in the esterification of CA with ethanol, although to a lesser
extent than Lipozyme. According to the mechanism proposed for lipases, the acyl-enzyme
intermediate is first formed by reaction with a serine residue in the active site [23].
Therefore, both steric and electronic properties of the substrate are likely to influence the
outcome of CAL B-catalyzed reactions.
To shed light this influence in the case of CA, we applied molecular modeling. This
approach helps to understand the structural relation between the enzymatic pocket and the
characteristic steroidal framework. Then, molecular docking studies were performed on
cholic acid with CAL B to evaluate the ability of the catalytic pocket to accommodate this
substrate. The aim of this part of the work was to explain the lack of reactivity of CA in
the enzymatic acetylation in comparison with the good performance showed when the
ethyl ester was used as substrate.
The analysis of the results led to the selection of a possible conformation for CA, using the
interaction with the amino acids of the catalytic site (Asp187-His224-Ser105) and the
population of the clusters as selection criteria. Docking analysis showed a majority cluster,
in which the substrate interacts with the active site (Ser105) of the enzyme through the
carboxyl moiety showing a distance of 2.48 Å (Table 5, Figure 1a). This optimal distance
could explain that the enzymatic esterification reaction of CA is favored.
15

Figure 1. Docking of CALB with cholic acid: interaction of substrate with Ser 105
through: a) the carboxyl moiety; b) hydroxyl gropus.
In addition, a less representative but considerable cluster, in which CA tries to enter the
active site of the lipase by the end of the hydroxyl groups, was also observed. This
entrance was unsuccessful as it can be assumed from Figure 1b. It can be observed that the
distance from Ser105 to the hydroxyl groups in C-3 (10.07 Å), C-7 (11.02 Å) and C-12
(12.41 Å) (Table 5). This confirms the lack of reactivity of CA towards the enzymatic
acetylation.
In the case of CAL B-catalyzed acetylation of CA ethyl ester, it could be assumed that a
decrease in the polarity of the substrate by esterification favors the income by the end of
OH groups in CA. As it could be expected, OH on C-3 is the only one located at an
optimal binding distance to the catalytic serine (2.07 Å) (Figure 2a).
Figure 2. Docking of CALB with ethyl ester CA (a) and lauryl ester CA (b).
a)
16

The negative result in the acetylation for 2d, the lauryl ester of CA, was also clarified by
molecular modeling. It was noted that this substrate cannot be properly accommodated in
the hydrophobic pocket of the enzyme. As the alkyl chain is bent, the OH groups are too
far to be able to interact with the catalytic triad. The nearest hydroxyl group is that on C-
12, with a distance of 9.24 Å to the catalytic serine for the most representative conformer
(Figure 2b).
In a previous work we performed molecular docking studies on chenodeoxycholic acid
(CDCA), deoxycholic acid (DCA) and lithocholic acid (LA) with CAL B [13]. Table 5
shows the data corresponding to the distance COOH-Ser105 and OH-Ser105 for these
three bile acids, CA and some derivatives.
Table 5. Distance in representative conformers of bile acids.
Bile
acid
Yield
(%)
Esterification
COOH-Ser distance (Å)
2.48
1.83
1.94
1.61
43a
99
89
93
CA
CDCA
DCA
LA
Acetylation
OH position
OH-Ser distance (Å)
3
7
12
3
3
3
7
3
12
3
10.07
11.02
12.41
2.07
9.24
1.94
4.38
1.95
4.73
1.95
CA
-
87
-
90
-
Et-CA
La-CA
CDCA
CDCA
DCA
99
-
DCA
LA
75b
a Using CAL B, with Lipozyme under the optimal conditions the product was obtained
in 80% yield.
b Ethyl ester LA-3-AcO, one-pot acetylation and esterification [13].
17

The comparison of these results, which are in accordance with the lipase activity and
regioselectivity experimentally observed, allowed us to draw some conclusions.
The four bile acids showed low values of the distance COOH-Ser105. They could be
esterified by lipase catalysis affording the ethyl ester, in excellent yield for CDCA, DCA
and LA. CA yield was poor with CAL B, but increased to 80% with Lipozyme.
For the four bile acids, acetylation was regioselective and only the 3-AcO was obtained.
Docking showed that the main favorable conformations correspond to those in which the
hydroxyl group on carbon 3 points toward the catalytic triad. It seems that the hydrophilic
α face in CA, containing the three hydroxyls and the carboxyl group, prevents it from
entering the active site by the end of any of the three hydroxyl groups, therefore no
product of acetylation from CA was obtained. The ethyl group in the ethyl ester of CA
decreases the polarity of the molecule allowing the interaction with the active site and
affording the product in very good yield.
In most cases, there is a correlation between the distance showed by the bile acids and
their derivatives with the activity and regioselectivity displayed by the lipase (Table 5).
The molecular modeling study confirmed the experimental results showing that CA
behavior in the enzymatic reactions is completely different from the other three bile acids
(CDCA, DCA and LA) previously studied.
4. Conclusions
In this work, we describe the application of enzymes to the synthesis of derivatives of
cholic acid. Scheme 4 shows the performed reactions and the obtained products.
The influence of various reaction parameters on the results was analyzed. Among the
tested enzymes, Rhizomucor miehei lipase gave the best results in esterification reactions,
affording alkyl esters as the only reaction products, six of them being new compounds.
It was not possible to obtain acetylation products from CA by any of the lipases tested.
According to literature, only the triacetylated derivative was obtained through chemical
procedures. However, ethyl ester CA 2a was a suitable substrate and Candida antarctica
B lipase was the catalyst of choice allowing us to obtain the ethyl 3α-acetyl cholate 3a in
excellent yield (87%). This derivative was chemically obtained in 75 % yield [15]. The
reaction was highly regioselective, neither 7-OH nor 12-OH was transformed. Under the
same reaction conditions the lauryl ester of cholic acid not was acetylated.
By lipase-catalyzed alcoholysis of the peracetylated cholic acid the corresponding ethyl
ester was obtained; no products of deacetylation were observed. Instead, by alcoholysis of
peracetylated ethyl ester of cholic acid it was possible to obtain a ethyl 7α,12α−diacetoxy-
3α-hydroxy-5β-cholanate as a new product.
In summary, the enzymatic catalysis offers a good alternative to synthesize a variety of
esters and mono- and diacetyl derivatives from cholic acid, under mild reaction conditions.
18

O
OH
OR
HO
OH
Alkyl-CA
2b-g: CH3(CH2)n n = 2,3,11,13,15,17
LIP
ROH / DIPE
40 ºC
O
OAc
O
OH
OH
OH
Ac2O / Py
DMAP r.t.
AcO
OAc
HO
OH
CA (1)
3,7,12-triAcO-CA (4a)
CAL B
EtOH / hexane
LIP
EtOH / DIPE EtOH / hexane
40 ºC 40 ºC
CAL B
40 ºC
O
O
OAc
OH
OCH2CH3
OCH2CH3
Ac2O / Py
DMAP r.t.
AcO
OAc
HO
OH
Ethyl CA (2a)
Ethyl 3,7,12-triAcOCA (4b)
CAL B
EtAcO / hexane
40 ºC
CAL B
EtOH / hexane
40 ºC
O
O
OAc
OH
OCH2CH3
OCH2CH3
HO
OAc
Ethyl 7,12-diAcOCA (5b)
AcO
OH
Ethyl 3-AcO-CA (3a)
Scheme 4. Reactions and products from CA.
The lipase-catalyzed procedure uses a series of alcohols as reagent for esterification,
ethanol for alcoholysis and ethyl acetate as acylating agent. Moreover, the reaction is
simple, it is performed at low temperature and the products are isolated by simple
filtration and solvent evaporation. The lipase is biodegradable and consequently more
compatible to the environment than the chemical catalysts. In addition, because the
enzyme is insoluble in the reaction medium, it is easily removed by filtration and can be
19

reused. In the esterification reaction LIP retained 80 % activity after three reaction cycles
and in the acetylation reaction, CAL B retained 88 % activity after five reaction cycles.
In addition, molecular modeling was applied to rationalize the selective behavior of CAL
B in the acetylation reactions and helped to explain the lack of reactivity of CA through
the particular interaction between the lipase active site and this bile acid. A comparison
between the behavior of cholic acid and chenodeoxycholic, deoxycholic and lithocholic
acid is also presented.
Supporting Information (see footnote on the first page of this article): Spectral data for
compounds 2a-g, 3a, 4a-b and 5b, associated with this article are supplied.
Acknowledgments
We thank UBA X010, UBACYT 20020100100304, CONICET PIP 112-200801-
00801/09, ANPCyT PICT 2011-00595 and Bunge y Born Foundation for partial financial
support. A.B. and G.G.L. are Research Members of CONICET (Argentina).
References
[1] Buchholz K., Kasche V., Bornscheuer UT. Biocatalysis and Enzyme Technology;
Wiley–VCH: Weinheim, 2005.
[2] Faber K. Biotransformations in Organic Chemistry, 6th ed.; Heidelberg: Springer
Verlag; 2011.
[3] (a) Baldessari A., Maier MS, Gros EG. Enzymatic deacetylation of steroids bearing
labile functions. Tetrahedron Lett. 1995; 36: 4349‒352; (b) Baldessari A.,
Bruttomeso, AC., Gros EG. Lipase-Catalysed Regioselective Deacetylation of
Androstane Derivatives. Helv. Chim. Acta 1996; 79: 999‒1004; (c) Brutomesso AC.,
Baldessari A. Lipase-catalysed deacetylation of androstane and pregnane derivatives:
influence of ring D substitution J. Mol. Catal. B: Enzym. 2004; 29: 149–53; (d)
Brutomesso AC., Tiscornia A., Baldessari A. Lipase-catalyzed preparation of
biologically active esters of dehydroepiandrosterone. Biocatal. Biotrans. 2004; 22:
215‒20; (e) Rustoy EM., Ruiz Arias IE., Baldessari A. Regioselective enzymatic
synthesis of estradiol 17-fatty acid esters (RL-1620KP). Arkivoc 2005; xii: 175–88;
(f) Monsalve LN., Machado Rada MY., Ghini AA., Baldessari A. An efficient
enzymatic preparation of 20-pregnane succinates: chemoenzymatic synthesis of
20beta-hemisuccinyloxy-5alfa-H-pregnan-3-one. Tetrahedron 2008; 64: 1721–30; (g)
Quintana PG., Baldessari A. Lipase-catalyzed regioselective preparation of fatty acid
esters of hydrocortisone. Steroids 2009; 74: 1007–14; (h) Quintana PG., Guillén, M.,
Marciello M., Palomo JM., Valero F., Baldessari A. Immobilized heterologous
Rhizopus oryzae lipase as an efficient catalyst in the acetylation of cortexolone. Eur. J.
Org. Chem. 2012; 4306-12; (i) Baldessari A., Iglesias LE. In Lipases and
Phospholipases, Methods and Protocols, Sandoval, G., Ed; Humana Press: New York,
2012, pp. 457-70; (j) Quintana PG., Camet A., Marciello, M. Valero F., Palomo JM.,
Baldessari A. Enzyme-catalyzed preparation of chenodeoxycholic esters by an
immobilized heterologous Rhizopus oryzae lipase. J. Mol. Catal. B: Enzym. 2015;
118: 36-42.
[4] Whitthall J., Sutton PW. In Practical Methods for Biocatalysis and Biotransformations;
John Wiley & Sons Ltd.: New York, 2012.
20

[5] Setchell KDR., Kritchevsky D., Nair PP. The Bile Acids: Chemistry, Physiology, and
Metabolism, Methods and Applications Vol.4, Springer Verlag: New York, 2011.
[6] Hofmann AF., Hagey LR. Living History of Lipids: Key discoveries in bile acid
chemistry and biology and their clinical applications: history of the last eight decades.
J. Lipid Res. 2014; 55: 1553–95.
[7] Coufalová L., Mrózek L., Rárová L., Plaček L., Opatřilová R., Dohnal J., Králová K.,
Paleta O., Král V., Drašar P., Jampílek J. New propanoyloxy derivatives of 5β-
cholan-24-oic acid as drug absorption modifiers. Steroids 2013; 78: 435–53.
[8] a)Pellicciari R., Gioiello A., Macchiarulo C., Thomas E., Rosatelli B., Natalini R.,
Sardella M., Pruzanski A., Roda E., Pastorini K., Schoonjans J., Auwerx J. Discovery
of 6alpha-ethyl-23(S)-methylcholic acid (S-EMCA, INT-777) as a potent and
selective agonist for the TGR5 receptor, a novel target for obesity. J. Med. Chem.
2009; 52: 7958–61. b) Secondo F., Carrea G., De Amici M., di Ventimiglia S.J.,
Dordick J. A combinatorial biocatalysis approach to an array of cholic acid
derivatives. Biotechnol. Bioeng. 2003; 81: 391-396.
[9] Suryanarayana V., Surendranatha Reddy O., Hari Babu B., Anuradha V. Facile
Synthesis of 1-(2-Morpholin-4-YL) 3, 5-Bis (2, 4, 6-Try SubstitutedArylidene)-
Piperidin-4-Ones. Res. J. Pharm. Biol. Chem. Sci. 2014; 5: 1188–94.
[10] Mukhopadhyay S., Maitra U. Chemistry and biology of bile acids. Curr. Sci. 2004;
12: 1666–83.
[11] Brotherhood PR., Davis AP. Steroid-based anion receptors and transporters. Chem.
Soc. Rev. 2010; 39: 3633–47.
[12] Hofmann AF., Roda A. Physicochemical properties of bile acids and their
relationship to biological properties: an overview of the problem. J. Lipid Res. 1984;
25: 1477–89.
[13] García Liñares G., Zigolo MA., Simonetti L., Longhi SA., Baldessari A. Enzymatic
synthesis of bile acid derivatives and biological evaluation against Trypanosoma
cruzi. Bioorg. Med. Chem. 2015; 23: 4804–14.
[14] Morris GM., Goodsell DS., Halliday RS., Huey R., Hart WE., Belew RK., Olson, AJ.
Automated Docking Using a Lamarckian Genetic Algorithm and and Empirical
Binding Free Energy Function. J. Comp. Chem. 1998; 19: 1639–62.
[15] Hu X., Zhang Z., Zhang X., Li Z., Zhu XX. Selective acylation of cholic acid
derivatives with multiple methacrylate groups. Steroids 2005; 70: 531–37.
[16] Kuhajda K., Kevresan S., Kandrac J., Fawcett JP., Mikov M. Chemical and metabolic
transformations of selected bile acids. Eur. J. Drug Metab. Pharm. 2006; 31: 179–235.
[17] Ibrahim-Oualia M., Rocheblaveb L. First synthesis of thia steroids from cholic acid.
Steroids 2010; 75: 701–9.
[18] (a) Cortese F. The Synthesis of Conjugated Bile Acids. IV, The Bondi and Mueller
Procedure. J. Am. Chem. Soc. 1937; 59: 2532–34; (b) Rasras AJM., Al-Tel TH., Al-
Aboudi AF., Al-Qawasmeh RA. Synthesis and antimicrobial activity of cholic acid
hydrazone analogues. Eur. J. Med. Chem. 2010; 45: 2307–13.
[19] Riva, S. Organic Synthesis with Enzymes in Non-Aqueous Media; Carrea G.; Riva
S., Eds.; Wiley–VCH: Weinheim, 2008.
[20] Cabrera, Z. Fernández-Lorente, G., Fernandez-Lafuente, R., Palomo, JM., Guisan.
JM. Novozym 435 displays very different selectivity compared to lipase from
Candida antarctica B adsorbed on other hydrophobic supports J. Mol. Cat. B: Enzym.
2009; 57: 171-176.
[21] Mrózek L., Dvořáková L., Mandelová Z., Rárová L., Řezáčová A., Plaček L.,
Opatřilová R., Dohnal J., Paleta O., Král V., Drašar P., Jampilek J. Investigation of
21

new acyloxy derivatives of cholic acid and their esters as drug absorption modifiers.
Steroids 2011; 76: 1082–97.
[22] Kuhada K., Kandraĉ J., Cirin-Novta V., Miljovic D. One-pot esterification and
selective 3α-acetylation of cholic and deoxycholic acid. Collect. Czech. Chem.
Commun. 1996; 61: 1073–76.
[23] Paiva AL, Balcão VM., Malcata F.X. Kinetics and mechanisms of reactions catalyzed
by immobilized lipases. Enzyme Microb. Technol., 2000; 27: 187-204.
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Graphical Abstract
O
OH
OH
HO
OH
Cholic acid
Lipase
23

Highlights
•
A series of cholic acid esters and mono- and diacetyl derivatives was prepared by
an enzymatic approach.
• The influence of various parameters on the enzymatic reactions was studied to
achieve high regioselectivity in the lipase catalysis.
• The enzymatic procedure is performed under mild reaction conditions compatible
to the environment.
• Molecular docking was applied to explain cholic acid behavior under enzymatic
conditions.
24