Highly selective purification of three lipases from Geotrichum candidum 4013 and their characterization and biotechnological applications

Article abstract of DOI:10.1016/j.molcatb.2013.09.012

Simple purification steps were used to purify two lipases (Lip1 and Lip2) and one novel lipase (Lip3) produced by Geotrichum candidum 4013. The octyl-Sepharose at low ionic strength was used to adsorbe Lip2 and novel Lip3, by addition of octadecyl-Sepabea

Full text of DOI:10.1016/j.molcatb.2013.09.012

Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Highly selective purification of three lipases from Geotrichum
candidum 4013 and their characterization and biotechnological
applications
Jana Brabcováa,b,c, Zuzana Demianováa,e,f, Jirˇí Vondrásˇeka, Michal Jágrd,
Marie Zarevúckaa,∗, Jose M. Palomob,∗∗
a Institute of Organic Chemistry and Biochemistry AS CR, 166 10 Prague 6, Flemingovo n. 2, Czech Republic
b Departamento de Biocatálisis, Instituto de Catálisis (CSIC) Campus UAM Cantoblanco, 28049 Madrid, Spain
c Institute of Chemical Technology Prague, 160 28 Prague 6, Czech Republic
d Institute of Physiology, Academy of Science of the Czech Republic, Vídenˇská 1083, 142 20 Prague 4, Czech Republic
e Research Institute of Molecular Pathology (IMP), Dr. Bohr-Gasse 7, 1030 Vienna, Austria
f
IMBA Institute of Molecular Biotechnology of the Austrian Academy of Sciences, Dr. Bohr-Gasse 3, 1030 Vienna, Austria
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 26 July 2013
Received in revised form 28 August 2013
Accepted 9 September 2013
Available online 27 September 2013
Simple purification steps were used to purify two lipases (Lip1 and Lip2) and one novel lipase (Lip3) pro-
duced by Geotrichum candidum 4013. The octyl-Sepharose at low ionic strength was used to adsorbe Lip2
and novel Lip3, by addition of octadecyl-Sepabeads the Lip1 was also adsorbed from the crude mixture.
Desorption of proteins with a Triton X-100 gradient or 1% (v/v) Triton X-100 permitted to fully purify
Lip1, Lip2 and Lip3 lipases.
The identification of Lip1 and Lip2 was confirmed by using mass spectrometry approach and their
characterization by molecular modeling approach. CD spectra and fluorescence showed differences in
the secondary and tertiary structure; the apparent difference was the presence of 4 extra loops in Lip2
in contrast to Lip1 structure. The Edman degradation together with mass spectrometry revealed the
presence of unique peptide AVGGGATLPEK in the novel lipase Lip3. The catalytic properties of the lipases
were tested through hydrolysis of p-nitrophenyl esters, triacylglycerols and peracetylated thymidine
esters. The enantioselectivity potential of purified lipases was studied for the hydrolyses of trans-2-(4-
methoxybenzyl)-1-cyclohexyl acetate, where Lip3 was the only active lipase with an excellent selectivity
(eep > 99%, E = 372). The all three purified lipases from G. candidum 4013 have shown promising catalytic
properties with biotechnological application.
Keywords:
Lipases
Hydrophobic supports
Substrate specifity
Geotrichum candidum
© 2013 Elsevier B.V. All rights reserved.
1. Introduction
considerably in their positional specificity to the hydrolysis of
triacylglycerols (TAGs, distinction between sn-1 and sn-3 position
Lipases (glycerol ester hydrolases, EC 3.1.1.3) catalyze the
hydrolysis of ester linkages in lipids, which result in release of the
constituent alcohol and acid moieties. Lipases act at the interface
between a hydrophobic substrate phase, and a hydrophilic phase
in which the enzyme is dissolved. These enzymes can differ
on the TAGs) and stereospecificity (ability to resolve racemic
mixtures) [1–3]. Some lipases show a very high versatility in
catalyzing the reactions like aminolysis, alcoholysis, esterification
and transesterification [4–7]. Therefore, lipases are widely used
in industrial applications for biosynthesis, in oil processing, pro-
duction of surfactants and preparation of enantiomerically pure
compounds [4,5,8].
However, most of the applications of lipases have been devel-
oped using commercially available lipases, and these in many
instances are not pure preparations, containing many impurities,
in some cases with activity against the target compound. This
presents a problem as the properties of each lipase may be different,
exhibiting different activity, stability or selectivity, or even being
affected in a very different way by the experimental conditions.
In these cases, the obtained results are the average of the whole
mixture, making the optimization or even the understanding of the
Abbreviations: c, conversion; E, enantiomeric ratio; eep, enantiomeric excess
of the product; ees, enantiomeric excess of the substrate; FOA, free oleic acid; MD,
molecular dynamics; MOE, modeling enviroment; MTPA, (R)-(+)-3,3,3-trifluoro-2-
methoxy-2-phenylpropanoic acid; MUF-butyrate, methylumbelliferone butyrate;
pNPB, p-nitrophenyl butyrate; pNPE, p-nitrophenyl ester; PDB, Protein Data Bank;
PLB, protein loading buffer; TAG, triacylglycerol; TO, trioleoylglycerol.
∗
Corresponding author. Tel.: +420 220183281.
Corresponding author. Tel.: +34 91 5854768; fax: +34 91 5854760.
E-mail addresses: zarevucka@uochb.cas.cz (M. Zarevúcka),
∗∗
josempalomo@icp.csic.es (J.M. Palomo).
1381-1177/$ – see front matter © 2013 Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.molcatb.2013.09.012

J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
63
Scheme 1. Cascade purification of lipases from Geotrichum candidum 4013.
processes very difficult. Moreover, the results may change between
2.2. Preparation of inoculum and lipases production
the enzyme batches. Therefore, pure lipase preparations are needed
to have full control and understanding of the processes [9].
High selective purification techniques are very useful. Among
various methods, the interfacial activation of lipases on hydropho-
bic supports has been described as a highly efficient methodology
to separate and purify lipases from a crude protein mixture, and
even from several isoenzymes [10,11]. This strategy permits the
one-pot purification by a solid-phase strategy obtaining excellent
final protein purification yield.
Geotrichum candidum has been described as a producer of
several lipases [3,12–21]. Most of the studies based on the identifi-
cation and characterization of lipases produced by yeast Geotrichum
sp. have reported the presence of two lipases with molecular mass
58–66 kDa [22–24] or from 50 to 62 kDa [25,26] depending on
the individual. Treatment with endo-␤-N-acetylglucosaminidase
indicated that these enzymes are glycosylated [26]. The homology
among the different lipases was high although they exhibited dif-
ferent substrate specificity [25]. Only a few studies have mentioned
the existence of other lipases [21,27], for example Gopinath et al.
obtained a pure lipase of 32 kDa by SDS-PAGE [21].
Generally, lipases have been used during hydrolysis of tria-
cylglycerols with various degree of specificity [19,26], especially
in the production of specific compounds from fats and oils
due to their specificity for unsaturated fatty acids production
[28].
Herein, we describe for the first time the highly selective purifi-
cation of three different lipases from a G. candidum strain 4013 and
their identification, characterization and biotechnological applica-
tions. The recent biocatalytic investigation of G. candidum 4013
showed promising catalytic properties of its crude lipases [3,15].
This study gives us answer concerning relationship structure and
catalytic abilities of the isolated lipases.
Preparation of inoculum and lipases production has been
already described by Zarevúcka et al. [3]. For solid culture, G. can-
didum strain 4013 was inoculated from a culture slant. A growing
solid medium was used with following composition per liter: 30 g of
glucose, 10 g of corn steep, 0.5 g of MgSO4 •7H2O, 1 g of KH2PO4, 1 g
of K2HPO4, 2 g of NaNO3, 0.5 g KCl, 0.02 g of FeSO4•7H2O and 15 g of
agar. The tubes were incubated at 25 ◦C for 3 days and conserved at
4 ◦C. Liquid culture was prepared as solid medium without addition
of agar. The medium was inoculated with the cells originally grow-
ing on solid medium and incubated with shaking at 30 ◦C for 24 h.
The lipases production was stimulated by induction medium [pep-
tone (50 g/L), glucose (10 g/L), MgSO4•7H2O (1 g/L), NaNO3 (1 g/L]
with olive oil as inductor (1%, v/v) at 30 ◦C for 48 h [3]. The cell-free
medium was used as a source of extracellular lipases.
2.3. Lipases cascade purification
1 g of octyl-Sepharose (or octadecyl-Sepabeads) was added to
75 mL of culture media (containing 45 mg of proteins) and pH was
set to 7.0. The mixture was continuously shaken for 24 h at 25 ◦C.
Periodically, the activity of suspension and supernatant was mon-
itored using the p-nitrophenyl butyrate (pNPB) assay described
below. The support with adsorbed proteins was washed by dis-
tilled water (10 times with 75 mL) and proteins were desorbed by
using different concentrations of detergent Triton X-100 in 10 mM
(0–1%, v/v) sodium phosphate buffer (pH 7.0, Scheme 1, Table 1).
2.4. Immobilization of lipases on CNBr-activated Sepharose
CNBr-activated SepharoseTM 4B (GE Healthcare) was sus-
pended in acidic aqueous solution (pH 2–3) for 30 min. Then the
support (0.5 g) was added to lipase solution in 10 mM phosphate
buffer (pH 7). The mixture was shaken at 25 ◦C for 2 h. The super-
natant was removed by filtration and the immobilized lipases on
CNBr-activated Sepharose were treated by 10 mL of blocking solu-
tion (1 M ethanolamine hydrochloride, pH 8) for 1 h 30 min under
shaking at 25 ◦C. Finally the mixture was filtered and support was
washed 10 times by distilled water.
2. Experimental
2.1. Materials
Octyl-Sepharose and CNBr-activated Sepharose were purchased
from GE Healthcare. Octadecyl-Sepabeads was kindly donated by
Resindion Srl., Triton X-100, p-nitrophenyl esters (pNPE), peracety-
lated thymidine, methylumbelliferone butyrate (MUF-butyrate)
and Fast Blue RR were obtained from Sigma. Coomassie brilliant
blue R250 and Protein Test Mixture of molecular mass standards
were obtained from Serva and [14C] trioleoylglycerol from New
England Nuclear, Perkin-Elmer (catalog no. NEC-674, 3.7 MBq/mL).
Other reagents and solvents were of analytical or HPLC grade.
Racemic trans-2-(4-methoxybenzyl)cyclohexanol was synthesized
and kindly donated by Prof. Zdeneˇk Wimmer from Institute of
Chemical Technology Prague, Czech Republic.
2.5. SDS-PAGE and zymogram analysis
Support with immobilized enzymes (50 mg) was suspended in
70 ␮L protein loading buffer (PLB; pH 6.8 containing 0.125 mM Tris-
base, bromophenol blue 10% (v/v) 2-mercaptoethanol, 40% glycerol,
4% SDS), the fractions after were diluted (1:1) with PLB and boiled
at 95 ◦C for 5 min.
SDS-PAGE was performed according to Laemmli [29], using
5% and 12% polyacrylamide for the stacking and resolving gels,
respectively. Broad range molecular mass standards (Protein Test

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J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
Table 1
Summary of Geotrichum candidum 4013 lipases purification.
Purification step
Total activity (U)
Protein (mg)
Specific activity (U/g)
Purification factor
Crude
Lip1
Lip2
0.03
0.22
0.14
0.06
0.6
0.2
0.5
1.1
45.9
1110.0
282.0
55.5
1
24.2
6.1
1.2
Lip3
Mixture from Serva) were used for mass determinations. Gels
were stained with Coomassie brilliant blue R250 (Serva). Non-
denaturing SDS-PAGE was performed using SDS-PAGE gel and
under non-reduced condition (protein samples were not treated
with 2-mercaptoethanol and with high heat before electrophore-
sis). Both electrophoresis were carried out at 180 mV at 4 ◦C [30].
For the zymogram analysis, SDS molecules were removed from
the SDS-PAGE gel by washing for 20 min with 20% 2-propanol
and twice for 10 min with distilled water at the 25 ◦C tempera-
ture [31]. The gel was briefly washed in 50 mM Tris–HCl pH 8.0,
and covered by a solution of 100 ␮M methylumbelliferone butyrate
(MUF-butyrate) in the same buffer [32]. Active bands became visi-
ble in a short time after UV illumination.
the amount of radioactivity recovered in the assay. The final reac-
tion mixture (175 ␮L) contained: 125 mM [14C]trioleoylglycerol, 1%
(w/v) fatty acid-free bovine serum albumin, 25 mM Tris–HCl, pH
7.5 at 0 ◦C, and 25 mM nonradiolabeled competing substrate (the
total TAG concentration in the assay was 150 mM, with 83.3% of
radiolabeled trioleylglycerol and 16.6% of nonradioactive TAG).
2.8. Biotransformation experiments
The immobilized lipases (300 mg) on CNBr-activated
SepharoseTM 4B were added to 2 mL of 0.5 mM substrate
peracetylated thymidine in acetate buffer, pH 5. The progress of
the reactions was analyzed by HPLC. The enzymatic resolution of
trans-2-(4-methoxybenzyl)cyclohexyl acetate (20 mg; 0.09 mmol)
was dispersed in 50 mM phosphate buffer (1.7 mL; pH = 6.5),
and the free enzyme (released from support, 2U) was added.
The reaction mixture was stirred for 72 h at 25 ◦C. Work-up of
the reaction mixture consisted of filtering the enzyme from the
reaction mixture. The rest of the mixture was extracted with
diethylether, dried over sodium sulfate, then the solvent was
evaporated and finally the products were separated by preparative
TLC and analyzed by HPLC.
2.6. Hydrolysis of p-nitrophenyl esters
Hydrolysis of p-nitrophenyl esters was performed by measur-
ing the increase in the absorbance at 348 nm (isosbestic point)
produced by the release of p-nitrophenol during the hydrolysis.
To start the reaction, 40 ␮L of lipase solution or suspension were
added to 2.5 mL of 0.4 mM p-nitrophenyl butyrate (pNPB) in 25 mM
sodium phosphate buffer pH 7.0. The unit of lipase activity was
defined as the amount of enzyme necessary to release 1 ␮mol of p-
nitrophenol per minute (U) under the conditions described above.
The concentration of proteins in crude extract was determined
spectrophotometrically at 595 nm using bovine serum albumin as
a standard protein [33].
The substrate specificity of the enzymes was determined
by the hydrolysis of p-nitrophenyl esters of different C-chain
lengths (p-nitrophenyl butyrate (C4), p-nitrophenyl caprylate (C8),
p-nitrophenyl decanoate (C10), p-nitrophenyl laurate (C12), p-
nitrophenyl palmitate (C16), p-nitrophenyl stearate (C18)).
2.9. Mass spectroscopy and structures modeling
In-gel digestion of Lip1 and Lip2, the specific bands were excised
from the gel and digested with trypsin (0.4 ␮g). The resulting tryp-
tic peptides were acidified with 30 ␮L of 0.1% formic acid than
10 ␮L of the mixture was separated using the HTC-PAL autosam-
pler containing the Rheos 2000 pumps and nano-LC system with a
PepSwift Monolithic Trap Column PS-DVB, 200 ␮m x 5 mm couple
to a PepMap C18, 75 ␮m x 150 mm x 3 ␮m (LC Packings, USA) using
the flow-rate 200–300 ␮L/min. The mobile phases consisted of 0.1%
formic acid (A); and 0.1% formic acid in 100% acetonitrile (B). A
four-step linear gradient of 2–15% B in 35 min, 15–25% B in 15 min,
25–40% B in 20 min, 40–95% B in 15 min and 100% B for 10 min
was used throughout this study. The LC system was online cou-
pled to the LTQ-XL ORBITRAP mass spectrometer (Thermo Fisher
Scientific). The peptide ions were detected in a survey MS scan
from 350 to 2000 amu with resolution of 100,000 and top three
precursor ions selected for subsequent MS/MS scans with resolu-
tion of 7500 and normalization collision energy 35 V. All MS/MS
spectra were searched using Sequest algorithm [36]. The collected
MS spectra were searched against Saccharomycetales-uniprot pro-
tein database containing canonical and isoform sequence data
(version released February, 2012) by the Bioworks Browser 3.3.1
SP1 (Thermo Fisher Scientific, USA) using following criteria: the
mass tolerance for peptide identification was 5 ppm, variable mod-
ifications were carbamidomethyl modification of cysteine and
methionine oxidation.
2.7. 14C labeled trioleoylglycerol activity assay and hydrolysis of
various TAGs
14C labeled trioleoylglycerol (isotope was present at
the carboxyl carbon of the oleic acid esterified to each
of the three hydroxyl group of glycerol) was used as
a
substrate, and activity was measured as radioactivity recov-
ered in free fatty acid after exposure to lipase [34,35].
Sample solution (50 ␮L) was added to 175 ␮L of the final
reaction mixture buffer (150 mM [14C] trioleylglycerol, 1% (w/v)
fatty acid-free bovine serum albumin, 50 mM Tris–HCl, pH 8). The
reactions were shaken 5 h at 25 ◦C and terminated by the addition
of mixture [chloroform:methanol:benzene (1:2.4:2, v/v), 1 mL)]
to extract free fatty acids. The pH was set to 11.5 by addition of
0.1 M NaOH (40 ␮L) and aqueous and organic solvent phases were
separated by centrifugation at 1000 × g for 10 min. The upper
aqueous phase was taken for determination of radioactivity on
Tri-Carb 2900TR liquid scintillation spectrometer (Perkin-Elmer).
Lipase activity was expressed in units of pmoles [14C] oleic acid
released per min/mg protein.
In-gel digestion of Lip3, the protein bands were excised from
the gel, and further processed as described in Shevchenko et al.
[37]. The resulting tryptic peptides were extracted from the gel
pieces. The nano-LC apparatus used for protein digests analysis
was a Proxeon Easy-nLC (Proxeon) coupled to a maXis Q-TOF
(quadrupole – time of flight) mass spectrometer with ultrahigh res-
olution (Bruker Daltonics) by nanoelectrosprayer. The nLC–MS/MS
The substrate specificity of the enzyme was inferred from the
effects of the addition of non-radiolabeled competing substrates
(trimyristoylglycerol, tripalmitoylglycerol, tristeroylglycerol). Sub-
strates more effective than trioleoylglycerol are expected to reduce

J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
65
instruments were controlled with the software packages HyStar
3.2 and micrOTOF-control 3.0. The data were collected and manip-
ulated with the software packages ProteinScape 3.0, DataAnalysis
4.0, and BioTools 3.2 (Bruker Daltonics).
The peptide mixture (5 ␮L) was injected into a NS-AC-11-
C18 Biosphere C18 column (particle size: 5 ␮m, pore size: 12 nm,
length: 150 mm, inner diameter: 75 ␮m), with a NS-MP-10 Bio-
sphere C18 pre-column (particle size: 5 ␮m, pore size: 12 nm,
length: 20 mm, inner diameter: 100 ␮m), both manufactured by
NanoSeparations (Nieuwkoop).
The separation of peptides was achieved via a linear gradient
between mobile phase A (water) and B (acetonitrile), both contain-
ing 0.1% (v/v) formic acid. A four step linear gradient of 5–30% in
70 min, 30–50% in 10 min, 50–100% in 8 min and 100% B for 2 min
was used (the flow rate was 250 ␮L/min at 25 ◦C).
methanol/water (4:1, v/v) as the mobile phase at 0.3 mL/min. The
eluate was monitored at 220, 254 and 275 nm while the UV spectra
were run from 200 to 300 nm.
The products of hydrolysis of peracetylated thymidine were
analyzed by HPLC (column Kromasil C18, the flow 1 mL/min, UV
detection at 260 nm). The mobile phase was composed of medium
A: 10 mM ammonium dihydrogen phosphate (pH 4.2, 90%): ace-
tonitrile (10%) and medium B: acetonitrile (90%): distilled water
(10%) under conditions of the analysis: 0–6 min 100% A, 6–14 min
65% A and 35% B, 14–18 min 100% A. The retention times were: per-
acetylated thymidine – 2.1 min, monodeprotected 3ꢀOH – 7.6 min,
monodeprotected 5ꢀOH – 8.4 min, acetylated thymidine – 13.3 min.
2.12. Calculation of enantiomeric excess and E value
Mass spectra processing: Data were processed using Protein-
Scape and BioTools software. Trypsin was chosen as the enzyme
parameter. One missed cleavage was allowed, and an initial pep-
tide mass tolerance of 15.0 ppm was used for MS and 0.03 Da for
MS/MS analysis. Cysteines were assumed to be carbamidomethy-
lated, and methionine was allowed to be oxidated. All these possible
modifications were set to be variable. Monoisotopic peptide charge
was set to 1+, 2+ and 3+. Only significant hits (Mascot score ≥80),
as defined by the MASCOT probability analysis (P < 0.05) were
accepted.
For the construction of the 3D models of both lipases we
used as a template a publicly available crystal structure of the
LIP2 from G. candidum identifier 1THG from Protein Data Bank
(PDB, http://www.rcsb.org/pdb/home/home.do). To assemble the
alignment the modeled sequences of the Lip1 and Lip2 was uti-
lized CLUSTALW2 server at the EBI (http://www.ebi.ac.uk/Tools/
msa/clustalw2/). The aligned Lip1 with 1THG and Lip2 with the
1THG were inputs into modeling environment (MOE). The homol-
ogy module of the MOE program was used for the modeling of the
Lip1 and the Lip2 structures [38]. The final models were subject of
geometry optimization in parm99 force field to adjust local minima
geometry for given model.
To calculate the enantiomeric excess values (eep and ees), the
ratio of the corresponding enantiomers was determined by an
analysis of diastereoisomeric derivatives of the respective chiral
compounds. The synthesized convenient diastereoisomeric deriva-
tives were the diastereoisomeric esters of the chiral alcohols
with the respective pure enantiomers of (R)-(+)-3,3,3-trifluoro-2-
methoxy-2-phenylpropanoic acid (MTPA, Mosher acid). The second
method consisted of the application of a chiral stationary phase of
a cyclodextrin type to separate enantiomers of the chiral alcohols
using chiral HPLC analysis. A combination of both methods gives
two independent analytical methods for assigning the absolute
configuration of the major enantiomers of chiral alcohols [39–42].
The absolute configurations of the major enantiomers of the MTPA
esters were assigned by measuring their 1H and 19F NMR spectra.
Three key parameters allow an efficient evaluation of an enzyme-
mediated resolution process: the optical purity of the product,
expressed as enantiomeric excess of the chiral product (eep) or that
of the deracemized substrate (ees), the extent of conversion of the
racemic substrate c (c = ees/(ees + eep)), and the enantiomeric ratio
E (E = ln[1 − c(1 + eep)]/ln[1 − c(1 − eep)]).
The higher is the E value, the higher is the value of the cor-
responding enzyme-mediated process in a selection procedure
aiming to use the optimum enzymic system for the production of
a chiral compound with requested absolute configuration.
2.10. Circular dichroism and fluorescence spectroscopy
Circular dichroism spectra of the three pure lipases were
recorded in a Chirascan spectropolarimeter (Applied Photophysics)
at 25 1 ◦C. Far-UV CD spectra were measured at wavelengths
between 190 and 250 nm in a 1 mm path-length cuvette, with 2 ␮M
protein solutions in distilled water on a Varian Cary Eclipse Fluo-
rescence Spectrophotometer (Agilent Technologies). The intrinsic
tryptophan fluorescence was monitored using an excitation wave-
length of 280 nm, with excitation and emission bandwidths of
5 nm, and recording fluorescence emission spectra between 300
and 400 nm.
3. Results and discussion
3.1. Purification and characterization of extracellular lipases
produced by G. candidum strain 4013
SDS-PAGE analysis of the crude mixture of extracellular proteins
produced by yeast G. candidum strain 4013 showed a high num-
ber of bands (Fig. 1). Non-denaturing electrophoresis, revealed the
presence of at least three enzymes with esterase activity (Fig. 1,
Scheme 1). In order to separate pure lipases from crude mix-
ture a selective purification strategy based on domino purification
by using matrixes with different hybrophobicity was developed
(Fig. 2). The molecular mass were calculated from SDS-PAGE gels
using GelAnalyzer version 2012a – 70.5 kDa lipase for Lip1, 75.5 kDa
for Lip2 and 43 kDa for Lip3.
More than 76% of the active protein was adsorbed on the support
(octyl-Sepharose) during 17 h. Further addition of support did not
improve the immobilization yield.
The supernatant after immobilization showed hydrolytic activ-
ity toward pNPB, which confirmed the presence of lipases or
esterases. The immobilization on hydrophobic support is a method
how to selectively adsorb lipases with presence of the esterases.
However, sometimes a more hydrophobic material is needed
[10] due to various nature of lipases. Matrixes with different
hydrophobic degree (butyl-Sepharose, phenyl-Sepharose, Lewatit,
2.11. Analytical methods
Characterization of the products of biotranformations (1–6) was
performed by the 1H NMR and the 19F NMR spectra were recorded
on a Varian UNITY 500 spectrometer (in a FT mode) at 499.8
and 470.3 MHz, respectively, in deuteriochloroform using either
tetramethylsilane (ı 0.0) as the internal reference or hexafluo-
robenzene as external reference (ı – 162.9). Preparative column
chromatography was performed on a silica gel type 60 (parti-
cle size 0.04–0.063 mm). TLC was performed on aluminum sheets
precoated with silica gel 60. Analytical HPLC was carried out on
a Thermoseparation Products instrument equipped with a Con-
staMetric 4100 Bio pump and a SpectroMonitor 5000 UV DAD.
Analysis of the chiral products was performed on a chiral Nucle-
odex ␤-OH column (150 × 4 mm; Macherey-Nagel, Germany) using

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J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
Finally, these pure lipases were analyzed by zymogram analy-
sis for lipolytic activity toward MUF-butyrate in order to confirm
the previous reference (Fig. 3A). Far-UV CD shoved the differences
on the secondary structure for the different lipases (Fig. 3B). A CD
spectrum of Lip2 differs from spectra of Lip1 and Lip3. According
to the far-CD spectrum, the content of ␣-helix secondary structure
strongly increased in comparison to Lip1. In the fluorescence spec-
trum there was a displacement of the maximum by l = 11 nm (from
l = 303 to 314 nm, Fig. 3C) suggesting a different enzyme tertiary
structure.
Till now, all studies dealing with identification and characteri-
zation of lipases produced by yeast Geotrichum sp. have reported
the presence of only two lipases with molecular mass 58–66 kDa
[22–24]. Shimada et al. isolated the lipase encoding cDNA clone
of G. candidum ATCC 34614 and found presence of two genes on
the chromosome, confirming two forms of lipase (lipase I and II)
obtained after purification procedures [22]. The isoenzymes had
similar amino acid composition but different terminal sequences
not found in the predicted primary structure. This confirmed two
different lipase genes and excluded the possibility of the forma-
tion of observed multiple forms by proteolytic digestion [22,23].
Sidebottom et al. purified and determined the substrate speci-
ficity of four lipases from two strains of G. candidum (ATCC 34614
– lipases I and II, CMICC 335426 lipases A and B). The puri-
fied enzymes were monomeric and had similar molecular masses
(from 50 to 62 kDa) and pI (from 4.5 to 4.7). Treatment with
endo-␤-N-acetylglucosaminidase caused a reduction in molecu-
lar mass of approximately 4.5 kDa for all four lipases, indicating
that these enzymes are glycosylated, it was also observed that
the lipases are related and they share common antigenic determi-
nants [25,26]. Although, the homology among the different lipases
was very high (84% between lipase A and B, and 97% between
lipase B and I), they exhibited a different substrate specificity [43].
Bertolini et al. have found lipase genes located in four strains
(ATCC 34614, NRCC 205002, NRRLY-552 and NRRLY-553) of G. can-
didum [25]. Each strain contained two lipase genes which were
closely related to lipase I and II of G. candidum ATCC 34614.
Most of the substitutions in amino acid composition were located
mainly on the protein surface and some were present in struc-
tural features possibly involved in determining substrate specificity
[25].
Fig. 1. Zymogram analysis of crude mixture of extracellular proteins produced by
yeast Geotrichum candidum strain 4013: (A) the gel stained with Coomassie Brilliant
Blue R-250 and (B) detection of lipolytic activity by using MUF-butyrate.
Fig. 2. SDS-PAGE analysis of desorption of the immobilized lipases on hydrophobic
supports: (A) proteins in supernatant after desorption from octadecyl-Sepabeads
by incubation with 1% Triton X-100 and (B) desorption from octyl-Sepharose by
incubation in increasing Triton X-100 concentration (v/v).
In a few studies, it has been mentioned existence of lipases with
molecular mass around 40 kDa. Gopinath et al. purified the lipase
to homogeneity with the apparent molecular mass of the purified
enzyme 32 kDa by SDS-PAGE [21]. In the study of Cai et al. two
new cold-adapted lipases (Lipase-A and Lipase-B) of mesophilic
Geotrichum sp. SYBC WU-3 were purified. The molecular mass of
Lipase-A and Lipase-B were determined to be approximately 41.1
and 35.8 kDa, respectively by SDS-PAGE. The two lipases showed
hydrolysis efficiency to various p-nitrophenyl esters, but they were
more active toward shorter p-nitrophenyl esters (C2 and C4) [27].
In the last years have been found several lipases produced by yeast
Geotrichum sp., as is shown in Table 2.
octadecyl-Sepabeads) were tested. The best results were found
using octadecyl-Sepabeads, 90% of the enzymatic activity of the
supernatant was disappeared and two bands were observed on the
SDS-PAGE of the adsorbed proteins, a main band corresponding to
Lip1 with traces of Lip2 (Fig. 2).
The simple method to purify the Lip1 consisted of 45 mg of
the crude protein mixture at the pH 7 to 1 g of octyl-Sepharose
with incubation overnight. The lipases Lip2 and Lip3 were adsorbed
on the support (76% of the pNPB activity), while Lip1 was free
in supernatant. Then, the supernatant was incubated with 1 g of
octadecyl-Sepabeads and 21.6% (90% yield at the beginning) of
the remaining activity was incorporated to the support after 8 h.
Therefore by these cascade stages of immobilization, a single band
corresponding to Lip1 was obtained. The pure lipase was fully des-
orbed from the support in an active form by incubation of 1 g of
immobilized Lip1 with 10 mL solution of 1% (v/v) of Triton X-100
for 1 h (Fig. 2A).
3.2. Structure sequence determination of Lip1 and Lip2 by mass
spectrometry
Gel bands were manually excised and subjected to tryptic
digestion and analyzed by mass spectrometer [37]. This mass spec-
trometric peptide mapping was compared with the theoretical
maps of known proteins. Substitutions in amino acid compositions
of identified peptides found in both lipases are marked in bold in
Table 3. The grey color shows new peptide sequences obtained
accurately in individual samples.
To separate the other two lipases adsorbed on the octyl-
Sepharose, the support was incubated in a growing concentration
of Triton X-100 from 0 to 1% (v/v) in 10 mM phosphate buffer (pH
7). The supernatants were analyzed by SDS-PAGE (Fig. 2 B). Lip3 was
desorbed from the support by using only 0.4% of detergent, with-
out any trace of Lip2, which was much more strongly adsorbed and
required 0.8% detergent to be eluted.

J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
67
Fig. 3. Characterization of the pure lipases. (A) zymogram analysis: the gels stained with Coomassie Brilliant Blue R-250 (1) and analyzed by for lipolytic activity using
MUF-butyrate (2), (B) Far CD spectra, and (C) fluorescence spectra. Lip1 (violet squares), Lip2 (red rhombus), Lip3 (blue triangles). (For interpretation of references to color in
this figure legend, the reader is referred to the web version of this article.)
3.3. Molecular model structure of Lip1 and Lip2
Lip2 and LIP 2 respectively. The amino acids of the active sites (Ser,
Glu and His) and oxyanion hole residue placement were carefully
adjusted. The missing residues in both lipases were 1:1 substituted
by the amino acids from the original UniProt sequences of the LIP1
and the LIP2 (Uniprot ids P17573 and P22394). The reconstructed
Lip2 full length sequence was longer than its Lip1 counterpart.
Available crystal structure of the LIP2 from G. candidum (PDB
code 1THG) was used as a template for the construction of the 3D
models of both lipases as described in Section 2.
The first apparent difference is the presence of 4 extra loops in
Lip2 in contrast to Lip1 where the extra residues of Lip2 are accom-
modated. It is interesting that the provided CLUSTALW2 alignment
also placed 2 extra loops into the Lip1 comparing to the Lip2. This
means that both sequenced lipases were of different length and
The sequencing of two forms of the experimentally isolated
lipases from G. candidum 4013 – Lip1 and Lip2 was incomplete.
Bioinformatic analysis on lipases from different organisms and
sources were used to reconstruct the sequences of both isolated
forms. The analysis of LIP1 and LIP2 from G. candidum (Uniprot
ids P17573 and P22394 – 84.014%), Candida rugosa (P20261 and
P32946 – 78.689%), Candida albicans (Q5APE4 and Q5APG1 –
58.849%) provided two important features of the above listed
molecules: (i) Lipase 1 and Lipase 2 from these organisms have
usually almost the same length; (ii) Both forms of lipases share
high level of sequence identity. Full sequence reconstruction was
based on the pair-wise local alignment of the Lip1 and LIP 1 and the
Table 2
Lipases found in database uniprot for Geotrichum sp. (http://www.uniprot.org).
Entry
Protein names
Organism
Length
P17573
P79066
P22394
A0FLP5
I7C458
Q1ZZV0
Q12614
E6Y1H2
Q0MVP3
Q12616
D0F1S1
E6Y1H1
Q2PPA8
Lipase 1 (EC 3.1.1.3) (GCL I) (Lipase I)
Lipase 1 (EC 3.1.1.3) (Lipase I) (TFL I)
Lipase 2 (EC 3.1.1.3) (GCL II) (Lipase II)
Lipase (fragment)
Geotrichum candidum
Geotrichum fermentans
Geotrichum candidum
Geotrichum sp. DTQ-26.3
Geotrichum candidum
Geotrichum candidum
Geotrichum candidum
Geotrichum candidum
Geotrichum candidum
Geotrichum candidum
Geotrichum candidum
Geotrichum candidum
Geotrichum candidum
563
563
563
544
563
544
544
544
563
563
563
544
563
Lipase (EC 3.1.1.3)
Lipase I (EC 3.1.1.3) (fragment)
Triacylglycerol lipase (EC 3.1.1.3) (fragment)
Lipase 2 (fragment)
Lipase
Triacylglycerol lipase (EC 3.1.1.3)
Lipase
Lipase 1 (fragment)
Lipase

68
J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
Table 3
The mass spectrometric peptide mapping of Lip1 and Lip2 produced by Geotrichum candidum 4013.
Peptide
DeltaM (ppm)
P (pep)
Score XC
Lip1
-.VML ATK.-
0.91742
−0.24839
−0.12622
0.95009
0.84519
0.93690
0.87985
1.28143
1.20320
1.07589
1.22910
0.67155
0.24676
0.78292
0.75039
−0.48839
−0.11921
0.75848
0.86426
0.97426
1.19236
0.97586
0.78615
1.28143
1.16264
1.51852
0.90522
1.06702
0.24676
1.83274
1.27435
0.53087
0.46164
1.30004
0.67906
1.27262
1.81474
8.84E−01
2.06E−02
5.73E−03
7.99E−07
2.34E−04
1,03E−04
4.42E−06
9,69E−01
1.21E−08
4.80E−01
1.55E−02
5.99E−01
4.27E−01
2.59E−01
2.31E−03
6.27E−02
3.86E−01
1.58E−06
5.36E−06
7.54E−07
5.45E−10
4.27E−05
5.27E−05
9.99E−01
2.55E−07
1.16E−03
3.73E−03
3.70E−03
6.59E−01
4.80E−04
2.68E−06
2.47E−06
4.53E−04
2.86E−07
1.07E−04
2.48E−07
6.78E−12
0.22
1.18
1.75
3.93
1.95
2.41
3.84
0.47
4.14
0.94
1.06
1.39
0.88
1.13
2.00
0.56
1.35
5.15
4.65
4.09
5.27
2.01
3.90
0.47
3.19
4.56
1.52
2.28
0.88
3.22
2.41
2.75
1.79
4.49
2.28
3.86
5.79
N
K.HPQPFTGSYQGLK.A
K.SSDVLH AQNSYDLK.D
S
R. AAI
I
LFQSPR.R
FTDL
K.GLEWVSDNIANFGGDPDK.V
K.GIPFADPP DLR.F
VG
R.TGILNALTPQFK.R
-.VMIFGESAGAMSVAHQLIAYGGDNTYNGK.-
K.ANDFS ACMQLDPGN
LLDK.V
AIS
S
R.YFISFANHHDPNVGTNL .Q
K
R.IEGISNFE DV
L
G.-
F
S
T
R.PDGNIIPDAAYELYR.S
K.DLFGLLPQFLGFGPR.P
R.TGPYGFLGGDAITAEGSTNAGLHDQR.K
K.ES EMGQPVVFVSINYR.T
VE
K.QWDMYTDAGK.E
Lip2
R.RVML ATK.D
S
R.FKHPQPFTGSYQGLK.A
K.SSSVLH AQNSYDLK.D
D
R. AAI
V
FQSPR.R
LSDM
R.KGLEWVSDNIANFGGDPDK.V
K.GIPFADPP DLR.F
LN
R.TGILNALTPQFK.R
K.VMIFGESAGAMSVAHQLIAYGGDNTYNGK.K
K.ANDFS ACMQLDPGN
LLDK.A
SLT
P
R.YFISFANHHDPNVGTNL
.E
LQWDQYTDEGK
R.IEGISNFE DV
L
G.-
Y
T
N
R.PDGNIIPDAAYELFR.S
K.DLFGLLPQFLGFGPR.P
R.TGPFGFLGGDAITAEGNTNAGLHDQR.K
K.ES MGQPVVFVSINYR.T
IN
R.GPLYDMAK.G
K.EMLEIHMTDNVMR.T
K.KLFHSAILQSGGPLPYHDSSSVGPDISYNR.F
K.GTVSMNEDCLYLNVFR.P
R.VLSLYPQTLSVGSPFR.T
K.WLQYIFYDASEASIDR.V
that the level of the sequence identity was not comparable with
the canonical LIP1 and LIP2 forms from G. candidum. Interestingly,
the rotameric states of the residues in the binding site are not the
same for Ser, Glu and His. The largest difference is the placement of
the Glu residue in Lip1 where it is oriented out of the triacylglycerol
binding site whereas in the Lip2 is oriented toward the binding cleft
(Fig. 4).
One of the physical–chemical features of both lipases is the
placement of the lid. The 3-helical segment covers the binding
cleft and it should possesses hydrophobic character on the side
oriented to the interior of the protein and making hydrophobic
contacts with aliphatic part of triacylglycerol ligand. As shown in
Fig. 4 the model succeeded in placement of hydrophobic residues
toward the protein interior and moreover the contact residues are
also of hydrophobic character. It is necessary to stress that the level
of conservation for the lid is relatively high.
The further attempt to characterize the models was a placement
of the naturally occurred ligand TAG into the binding cleft Lip1.
It was not perform the full molecular dynamics (MD) simulation
of the lid opening, ligand binding and the lid closing. The model
of the TAG was placed into the binding site by manual docking
utilizing dock module of the MOE), solvated and short MD simu-
lations was performed. Expected binding pattern was reached in
relatively short simulation time of 500 ps. The models could not
provide any additional information without reasonable dynamics
simulationmappingthenon-staticcharacteristics. Thefirstattempt

J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
69
Fig. 4. Model structures of two lipases from yeast Geotrichum candidum 4013: (A)
a
comparison of Lip1 and Lip2, the composition of
4
extra loops in Lip2
–
Loop1 (Val140–Asn152, VGPDISYNRESIN), Loop2 (Ile189–Ala209, IANFGGDPDKTGPFGFLGGDA), Loop3 (Asn514–His532, NIGPANSYLKYFISFANHH), Loop4 (Gln559–Ile570,
QTLSVGSDPFRI), (B) the active sites of lipases (Lip1 – Ser 257, Glu359, His468 Lipase 61 kDa – Ser 292, Glu 394, His 503), (C) the differences in composition of lid for Lip1 and
Lip2, (D) composition of aminoacids in extra loops of Lip2, Loop3 (in red color), Loop2 (in green color), and (E) manual docking for a placement of trioleoylglycerol into the
binding cleft Lip1. (For interpretation of references to color in this figure legend, the reader is referred to the web version of this article.)
to uncover the dynamics and binding properties of the model was a
short MD simulation of the trioleoylglycerol in binding in the bind-
ing cleft. From the simulation is apparent that the hydrophobic
character of the binding cavity can easily accommodate the tria-
cylglycerol ligand without dramatic geometry reorientation of the
binding side. Importantly, the correct placement of the polar part
of TAG between the residues of the active site confirmed sound
adjustment of the Lip1 and Lip2 models.
Probably the most interesting results of our modeling effort are
the found differences between Lip1 and Lip2 and their position in
the structure. A distinct environment provided by 4 extra loops of
the Lip2 makes possible to hypothesize that different specificity and
Fig. 5. Substrate specificities of Lip1 and Lip2 from Geotrichum candidum 4013 to p-nitrophenyl esters of different C-chain lengths: p-nitrophenyl butyrate (C4), p-nitrophenyl
caprylate (C8), p-nitrophenyl decanoate (C10), p-nitrophenyl laurate (C12), p-nitrophenyl palmitate (C16), p-nitrophenyl stearate (C18). Hydrolysis of p-nitrophenyl esters
was performed by measuring the increase in the absorbance at 348 nm (isosbestic point) produced by the release of p-nitrophenol in 25 mM sodium phosphate buffer of or
with pH 7 at 25 ◦C.

70
J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
Fig. 6. Effect of various species of triacylglycerols on rates of free oleic acid (FOA)
released from 14C-labeled trioleoylglycerol (TO) by Lip1 and Lip2 from Geotrichum
candidum 4013. 14C labeled trioleoylglycerol (isotope was present at the carboxyl
carbon of the oleic acid esterified to each of the three hydroxyl group of glycerol)
was used as a substrate, and activity was measured as radioactivity recovered in
free fatty acid after exposure to lipase.
different behavior of both lipases could be accounted to these four
loops. Most importantly the loop in proximity of the lid could serve
as a modulator and important regulatory segment of the lid dynam-
ics. The important regulatory role of the loop is also suggested in
recently published work of Rehm et al. [44].
Fig. 8. Substrate specificity of purified Lip3 from Geotrichum candidum 4013: (A)
p-nitrophenyl esters of different C-chain lengths: p-nitrophenyl butyrate (C4), p-
nitrophenyl caprylate (C8), p-nitrophenyl decanoate (C10), p-nitrophenyl laurate
(C12), p-nitrophenyl palmitate (C16), p-nitrophenyl stearate (C18), and (B) various
species of triacylglycerols on rates of free oleic acid released from 14C-labeled tri-
oleoylglycerol. Hydrolysis of p-nitrophenyl esters was performed by measuring the
increase in the absorbance at 348 nm (isosbestic point) produced by the release of
p-nitrophenol in 25 mM sodium phosphate buffer of or with pH 7 at 25 ◦C.
3.4. Substrate specificity of purified lipases Lip1 and Lip2
The specificity of lipases Lip1 and Lip2 was firstly studied in
the hydrolysis of p-nitrophenyl esters of different C-chain lengths
(Fig. 5). The Lip1 exhibited similar activity values for substrates from
C8 to C12, with the highest hydrolytic activity toward p-nitrophenyl
laurate (Fig. 5). Lip1 showed slight lipolytic activity toward C16 or
C18 substrates.
However, Lip2 proved a profile with a clearly preferential speci-
ficity toward shorter chain length substrates (esterase activity
rather than lipase activity) with a maximum activity value toward
p-nitrophenyl caprylate (C8). No activity of Lip2 was observed in
the hydrolysis of p-nitrophenyl stearate. These results are similar
to those with crude enzyme extracts, which were obtained in our
previous work [15].
allowed evaluating the relative substrate specificity comparing
with hydrolysis rate with nonradiolabeled trioleoylglycerol. The
increasing of lipase activity in the presence of competitive sub-
strate means that the substrate competes more effectively for the
enzyme than trioleoylglycerol and it would indicate either a true
stimulation of lipase activity. In the opposite case, when the release
of oleic acid is decreased, the enzyme hydrolyzes this substrate
preferentially than another. The substrate which has the great-
est effect on inhibition is the most readily hydrolyzed [35]. The
main difference in substrate specificity showed stimulation of tri-
oleoylglycerol hydrolysis in the presence of tripalmitoylglycerol
(Fig. 6). Tripalmitoylglycerol was a not substrate for Lip1 in compar-
ison to Lip2. Trimyristoylgycerol depressed the rates of hydrolysis
of trioleoylglycerol for both studied lipases. This indicated that
To understand the enzyme activity toward natural substrates
– TAGs, we focused on the influence of addition of a second
competitive substrate (TAG with different acyl chain length of
fatty acids: C14, C16, C18) to [14C] labeled trioleoylglycerol. This
Fig. 7. MS–MS spectrum of N-terminal peptide (AVVGGGATLPEK) of new lipase Lip3 isolated from Geotrichum candidum 4013.

J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
71
Table 4
Regioselective deprotection of peracetylated thymidine 1 catalyzed by lipases from Geotrichum candidum 4013 at pH 5.0.
O
O
O
NH
NH
NH
AcO
HO
AcO
N
O
N
O
N
O
O
O
O
+
H
H
H
OAc
OAc
OH
3
1
2
.
Biocatalyst
Activitya
Conversion (%)b
2b (%)
3b (%)
CNBr-Lip1
CNBr-Lip2
CNBr-Lip3
0.68
0.71
0.79
1.35
12.85
5.00
18.57
41.85
8.22
2.20
7.47
3.74
2.80
9.95
7.02
Octadecyl-Lip1
28.82
a
Units of ␮moles [14C] oleic acid released per min/g support.
Yield of monodeprotected product at 336 h.
b
Table 5
Kinetic resolution of trans-2-(4-methoxybenzyl)cyclohexyl acetate catalyzed by lipases from Geotrichum candidum 4013.
.
Biocatalyst
Conversion (%)
eep (%)
E
Lip1
Lip2
Lip3
0
0
38
–
–
>99
–
–
372
trimyristoylglycerol was preferentially hydrolyzed than trioleoyl-
glycerol.
trioleoylglycerol for all studied enzymes, indicating that it is more
preferentially hydrolyzed substrate than trioleoylglycerol.
3.5. Structure sequence determination of Lip3 by mass
spectrometry
3.7. Biotransformation experiments
The activity of the pure lipases immobilized on CNBr-activated
Sepharose was tested in the hydrolysis of compound 1 at pH 5.0
(Table 4). The specificity of the three lipases in pure form was
2–3 times higher than the crude mixture. The highest activity was
achieved for immobilized Lip1. Indeed when this lipase was immo-
bilized on octadecyl-Sepabedas (matrix for purification and where
the lipase fixes the open conformation) the activity of the enzyme
was 2-fold higher. Also the regioselectivity was different depending
on using lipases in crude or pure form. All the enzymes hydrolyzed
regioselectively the 6-OH (compound 2) and 3-OH (compound 3)
positions but with different specificity. Immobilized crude mix-
ture and pure Lip1 was more specific to produce 2. The adsorption
on octadecyl-Sepabeads increased the specificity of Lip1 for com-
pound 2 (two-fold) (Table 4). However, a slightly higher specificity
to produce compound 3 was observed for Lip2 and Lip3.
More interesting results were found in the evaluation of the
enantioselectivity of the lipases in the kinetic resolution of 4
(Table 5). Lip1 and Lip2 were unable to catalyze the desired hydrol-
ysis. However, Lip3 resolved the racemate 4, with an excellent
enantioselectivity (E = 372, eep > 99%) producing enantiomerically
pure alcohol (1R,2S)-5. This result shows that Lip3 is the only one
responsible for the selectivity observed previosly when the crude
mixture obtained from G. candidum strain 4013 was used in the
resolution of 4 [3].
The identity of the Lip3 could not be revealed by the first attempt.
The band was digested by trypsin [37] and analyzed by nLC–MS/MS.
The analysis showed several spectra probably possessing peptides
with unknown amino acid structure. Therefore, it was hypothesized
that this band belongs to the new original lipase.
For these reasons it was used the Edman degradation proce-
dure to reveal a part of the N-terminal amino acid sequence of the
enzyme. The first 15 amino acids of the new lipase Lip3 produced by
yeast G. candidum 4013 were determined as AVVGGGATLPEKLYG.
The most of this sequence was moreover confirmed by additional
searching of the MS/MS data was found MS spectra that matched
with the AVVGGGATLPEK peptide. This peptide was identified by
correlating measured mass spectra with the predicted spectra.
Fragmentation pattern of the MS/MS spectra was in good agree-
ment with the proposed sequence (Fig. 7). Using this approach, it
was confirmed the unique character of the new lipase.
3.6. Substrate specificity of the new Lip3
Substrate specificity of pure Lip3 toward different p-nitrophenyl
esters showed a slight different profile than for the other two
lipases (Fig. 5). A clear preference for C8 substrate was observed
(higher than for Lip2), and maintaining similar activity for sub-
strates from C12 to C18 (Fig. 8A).
The competitive substrates, tripalmitoylglycerol had higher
increasing effect on hydrolysis of 14C labeled trioleoylglycerol
using Lip3 Fig. 8B) as catalyst than that determined with the
other two lipases (Fig. 6). Only Lip3 showed the stimulation of
trioleoylglycerol hydrolysis in the presence of tristearaoylglycerol
(Fig. 8B). Trimyristoylglycerol depressed the rates of hydrolysis of
4. Conclusions
In this study, three different extracellular lipases from G.
candidum 4013 were simply and rapidly purified with a good
yields from a highly selective absorption supports. CD spectra

72
J. Brabcová et al. / Journal of Molecular Catalysis B: Enzymatic 98 (2013) 62–72
and fluorescence showed differences among their structures.
By MD simulations we presented the three-dimensional struc-
ture of Lip1 and Lip2. The new lipase Lip3 showed remarkable
ability in the hydrolytic kinetic resolution of racemic trans-2-(4-
methoxybenzyl)-1-cyclohexyl acetate, whereas lipases Lip1 and
Lip2 did not recognize the substrate.
[13] T. Jacobsen, J. Olsen, K. Allermann, Biotechnol. Lett. 12 (1990) 121–126.
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catal. Agric. Biotechnol. 1 (2012) 147–151.
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Enzymatic 61 (2009) 188–193.
In conclusion, it was discovered and described a new lipase from
yeast G. candidum 4013. The identity of the lipase was confirmed
with finding the unique peptide by two independent methods:
nLC–MS/MS and Edman degradation procedure. This peptide has
not been previously described for any yeast microorganism yet.
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Technol. 100 (2009) 2182–2188.
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Acknowledgments
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359–364.
This work has been sponsored by the Spanish National Research
Council (CSIC). Jana Brabcová is grateful to the Academy of Science
of the Czech Republic (project No. M200551203). Authors thank
to Zdeneˇk Vobu˚ rka by the N-terminal analysis experiments and
Prof. ZdeneˇkWimmer forkindlydonatingracemiccompoundtrans-
2-(4-methoxybenzyl)cyclohexyl acetate and for helping with the
analysis of chiral compounds.
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