Performance of different immobilized lipases in the syntheses of short- and long-chain carboxylic acid esters by esterification reactions in organic media

Article abstract of DOI:10.3390/molecules23040766

Short-chain alkyl esters and sugar esters are widely used in the food, pharmaceutical and cosmetic industries due to their flavor and emulsifying characteristics, respectively. Both compounds can be synthesized via biocatalysis using lipases. This work aims to compare the performance of commercial lipases covalently attached to dry acrylic beads functionalized with oxirane groups (lipases from Candida antarctica type B—IMMCALB-T2-350, Pseudomonas fluorescens—IMMAPF-T2-150, and Thermomyces lanuginosus—IMMTLL-T2-150) and a home-made biocatalyst (lipase from Pseudomonas fluorescens adsorbed onto silica coated with octyl groups, named PFL-octyl-silica) in the syntheses of short- and long-chain carboxylic acid esters. Esters with flavor properties were synthetized by esterification of acetic and butyl acids with several alcohols (e.g., ethanol, 1-butanol, 1-hexanol, and isoamyl alcohol), and sugar esters were synthetized by esterification of oleic and lauric acids with fructose and lactose. All biocatalysts showed similar performance in the syntheses of short-chain alkyl esters, with conversions ranging from 88.9 to 98.4%. However, in the syntheses of sugar esters the performance of PFL-octyl-silica was almost always lower than the commercial IMMCALB-T2-350, whose conversion was up to 96% in the synthesis of fructose oleate. Both biocatalysts showed high operational stability in organic media, thus having great potential for biotransformations.

Full text of DOI:10.3390/molecules23040766

molecules
Article
Performance of Different Immobilized Lipases in the
Syntheses of Short- and Long-Chain Carboxylic Acid
Esters by Esterification Reactions in Organic Media
Lionete Nunes de Lima 1, Adriano Aguiar Mendes 2 ID , Roberto Fernandez-Lafuente 3,
*
,
ID
Paulo Waldir Tardioli 1,
*
and Raquel de Lima Camargo Giordano 1
ID
1
Graduate Program in Chemical Engineering, Department of Chemical Engineering,
Federal University of São Carlos, 13565-905 São Carlos, SP, Brazil; lionetenunes@yahoo.com.br (L.N.d.L.);
raquel@ufscar.br (R.d.L.C.G.)
Institute of Chemistry, Federal University of Alfenas, 37130-001 Alfenas, MG, Brazil;
adriano.mendes@unifal-mg.edu.br
Department of Biocatalysts, ICP-CSIC, Campus UAM-CSIC, 28049 Madrid, Spain
Correspondence: rfl@icp.csic.es (R.F.-L.); pwtardioli@ufscar.br (P.W.T.);
Tel.: +34-91-595-4941 (R.F.-L.); +55-16-3351-9362 (P.W.T.)
2
3
*
Academic Editor: Derek J. McPhee
ꢀꢁꢂꢀꢃꢄꢅꢆꢇ
ꢀꢁꢂꢃꢄꢅꢆ
Received: 23 February 2018; Accepted: 24 March 2018; Published: 27 March 2018
Abstract: Short-chain alkyl esters and sugar esters are widely used in the food, pharmaceutical and
cosmetic industries due to their flavor and emulsifying characteristics, respectively. Both compounds
can be synthesized via biocatalysis using lipases. This work aims to compare the performance
of commercial lipases covalently attached to dry acrylic beads functionalized with oxirane groups
(lipases from Candida antarctica type B—IMMCALB-T2-350, Pseudomonas fluorescens—IMMAPF-T2-150,
and Thermomyces lanuginosus—IMMTLL-T2-150) and a home-made biocatalyst (lipase from
Pseudomonas fluorescens adsorbed onto silica coated with octyl groups, named PFL-octyl-silica)
in the syntheses of short- and long-chain carboxylic acid esters. Esters with flavor properties
were synthetized by esterification of acetic and butyl acids with several alcohols (e.g., ethanol,
1-butanol, 1-hexanol, and isoamyl alcohol), and sugar esters were synthetized by esterification of
oleic and lauric acids with fructose and lactose. All biocatalysts showed similar performance in
the syntheses of short-chain alkyl esters, with conversions ranging from 88.9 to 98.4%. However,
in the syntheses of sugar esters the performance of PFL-octyl-silica was almost always lower than the
commercial IMMCALB-T2-350, whose conversion was up to 96% in the synthesis of fructose oleate.
Both biocatalysts showed high operational stability in organic media, thus having great potential
for biotransformations.
Keywords: immobilized lipases; esterification; flavor esters; sugar esters
1. Introduction
Many short-chain alkyl esters exhibit flavors similar to those of common fruits [1,2]. Consequently,
they are widely used in pharmaceutical, cosmetics and food industries. Even though aromatic esters
might be obtained from natural sources, an alternative to the extraction of these compounds is
their chemical synthesis by esterification reactions. These can be catalyzed by inorganic catalysts or
biocatalysts [
and present low selectivity, making the downstream process complex [
the reaction conditions are milder and the reactions are more selective [4,5
3
,4
]. Inorganic catalysts require high temperatures, hence requiring high energy costs,
]. When using biocatalysts,
]. In addition, esters
4
Molecules 2018, 23, 766; doi:10.3390/molecules23040766
www.mdpi.com/journal/molecules

Molecules 2018, 23, 766
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synthesized through biocatalysis may be considered closed to “identical to natural” [
labeled as green products.
4,6
,7] and thus
Another ester family of great interest is that composed by the sugar fatty acid esters, which
are non-ionic surfactants [ ]. These substances are biodegradable, with a varied variety of uses in
8
many different economic sectors, e.g., the cosmetic, food, oral-care, detergent and pharmaceutical
industries [4,9,10].
The industrial synthesis of sugar fatty acid esters (e.g., long- or medium-chain fatty acids
of glucose, fructose or sucrose) is carried out by transesterification of fatty acid methyl esters
(more expensive than the◦fatty acids) using alkaline or metallic catalysts. The reaction requires
high temperatures (>100 C) and the use of reduced pressure in order to remove the methanol
released from the activated acyl donor to prevent competition with the hydroxyl groups of the
carbohydrates [
dimethylformamide (DMF), pyridine or dimethylsulfoxide (DMSO) that are toxic solvents. This implies
high energy costs, the production of undesired by-products and low selectivity [11 15]. Moreover,
the most challenging features of this process are the control of the degree of esterification and the
position of acylation [ 16], and both are difficult to achieve using alkaline catalysis. Therefore,
enzymatic synthesis of sugar esters might be a useful alternative, which is already used in industrial
scale for some applications [ ]. In these reactions, in order to dissolve the carbohydrate, the use of very
8]. In addition, the chemical catalysts are mostly alkaline and the reaction often uses
–
8,
4
polar solvents is required. The commonly used organic solvents, such as DMF or DMSO, are able to
dissolve saccharides, but they usually have very negative effects for enzymes and are not compatible
for many applications of carbohydrate-derived products [7,17,18]. Hence, the solvent must reconcile
carbohydrate solubility and low polarity to reduce the damage of the enzyme. In addition, the solvent
must not present reactive hydroxyl groups able to compete with the sugars, carboxylic groups or
esters bonds, which could be substrates of the enzyme and reduce the product yield and/or produce
side-products. In general, tertiary alcohols meet these requirements, as lipases do not recognize
them as substrates [19
ionic liquids [ 22 24]. Within the biocatalysts which can be used in the synthesis of esters, the most
common ones are lipases [ 14 18 25 35]. Lipases can catalyze esterification reactions, provided
that the reaction occurs at low water content [36]. Lipases present a peculiar catalytic mechanism,
the so-called interfacial activation [37 38]. According to this mechanism, in homogenous aqueous
–21]. Other examples of solvents that might be used in such reactions are
7, –
3,4, , , –
,
media, the lipase molecules are in a conformational equilibrium between a closed form, with the active
center inaccessible for the reaction media due to the presence of a polypeptide chain called lid that
block it, and the open form, where the lid is displaced and exposes the lipase active center to the
medium [39
–
41]. In the presence of any hydrophobic surface, e.g., a hydrophobic support, another
42 43], the open form of the lipase becomes
open form of the lipase or a hydrophobic protein [40
,
,
adsorbed via the hydrophobic area formed by the surrounding of the active center and the internal
side of the lid, shifting the conformational equilibrium and fixing the open form of the lipase stabilized
versus the surface of the support [44–46].
Esterification is a thermodynamically controlled process, where the yields are only defined by the
thermodynamics of the process [21,47], the catalyst only defines the velocity and the possibility of the
process [48]. To improve the yields, the thermodynamic equilibrium must be shifted. Reducing the
water content (even eliminating the produced water) is the usual way to improve the yields. In these
esterification reactions, the use of ultrasound irradiation to prevent the formation of water layers on
the immobilized enzyme [49
water in the enzyme surroundings) [51
water [ 32 53 54], have all proven to be very useful to shift the equilibrium and prevent biocatalyst
,
50], the use of very hydrophobic supports (also to avoid the gathering of
,52] and the use of molecular sieves to eliminate the produced
7
, , ,
inactivation. In the case of sugars, the situation is more complex, as the catalyst may define the position
or positions that are acylated, therefore the biocatalyst may also define in a certain sense the yields of
the target product. The possibility of acyl migration must be considered, as this may produce the final
modification of positions different to the ones modified by the enzyme [51,55,56].

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In this work, lipase from Pseudomonas fluorescens (PFL), Candida antarctica type B (CALB) and
Thermomyces lanuginosus (TLL) in one of their commercial immobilized forms (covalently attached to
an acrylic resin functionalized with oxirane groups) were used as biocatalysts in the syntheses of esters
with flavor and surfactant properties. These lipases, specially CALB, have been frequently employed
in the synthesis of esters in nonaqueous medium [4,7,8,10,57,58]. PFL immobilized on silica particles
coated with octyl groups (octyl-silica) was also used. Hydrophobically coated supports immobilize
the lipases very efficiently and enable the one step immobilization, purification and stabilization of the
open form of the lipase, due to the interfacial activation of the lipases on the hydrophobic surface of
the support [39,44,48]. Among the several pre-existing supports, silica has been reported as a good
choice for enzyme immobilization due to some properties, such as thermal and mechanical stabilities,
high surface area, high resistance to organic solvents and microbial degradation and nontoxicity [59].
Biocatalysts prepared by immobilization of several lipases onto ocytl-silica has been successfully used
in biotransformation reactions in organic media [26,28,59–61]. The preparation of octyl-silica and the
immobilization of PFL on this support were previously reported by our group and evaluated in the
synthesis of ethyl fatty acid esters [60] and fructose fatty acid ester [28]. In this new research effort,
we intend to analyze if the immobilization protocol may affect the final properties of immobilized
lipases in esterification reactions involving substrates as different as sugars, short alcohols and short or
long chain carboxylic acids.
2. Results and Discussion
2.1. Synthesis of Aroma Esters
Table 1 shows the conversions observed in the syntheses of short-chain carboxylic acid esters
by esterification catalyzed by home-made PFL-octyl-silica and commercial immobilized lipases at
37 ◦C after 24 h of reaction. All biocatalysts showed excellent performance in the synthesis of flavor
esters, yielding conversions ranging from 80.3 to 98.4% after 24 h reaction. Similar results have been
reported for other reaction systems (reaction time, enzyme and organic solvent) [4]. PFL-octyl-silica,
the home made preparation, offered yields over 95% for all alcohols. IMMTLL-T2-150 offered the
worst performance when hexanol was the substrate, only giving 80% after 24 h, and just 90% when
isoamyl alcohol and ethanol were used. IMMAPF-T2-150 only offered yields under 90% using ethanol,
the higher yields were obtained using butanol (with both, acetic and butyric acid). IMMCALB-T2-350
gave the worst results using ethanol (just over 90%). In an esterification, yields should be identical,
thus the difference in yields should be related to some kinetic problem that can slow down the reaction
rate in some instances. PFL-octyl-silica seems to be the most suitable biocatalyst for these esterifications
among the assayed ones.
◦
Table 1. Conversions of syntheses of short-chain carboxylic acid esters at 37 C for 24 h, catalyzed by
home-made PFL-octyl-silica and commercial immobilized lipases. Reactant concentrations: 0.1 M in
heptane for acids and alcohols, and 200 U/g of acid (esterification activity) 20 g/L molecular sieves for
all biocatalysts.
Reactants and Products
Conversions (%) by Biocatalyst
Acyl
donor
FEMA flavor
profile 1
apple, banana,
pungent
apple, banana,
grass, herb, pear
Apple, banana,
glue, pear
Acyl acceptor
Ester
PFL-octyl-silica
96.8 ± 0.1
98.0 ± 2.9
96.9 ± 0.2
98.4 ± 2.3
96.7 ± 0.1
IMMTLL-T2-150
97.0 ± 0.0
IMMAPF-T2-150
93.8 ± 0.4
IMMCALB-T2-350
96.8 ± 0.1
Acetic acid
Acetic acid
Acetic acid
1-butanol
Butyl acetate
Hexyl acetate
Isoamyl acetate
Butyl butyrate
Ethyl acetate
1-hexanol
80.3 ± 1.5
96.9 ± 0.1
97.5 ± 3.5
isoamyl
alcohol
91.5 ± 0.0
93.7 ± 0.2
95.0 ± 2.4
Butyric
acid
1-butanol
floral
93.5 ± 0.0
96.9 ± 0.1
96.6 ± 0.1
Aromatic,
brandy, grape
Acetic acid
1
ethanol
90.8 ± 0.2
88.9 ± 1.9
90.5 ± 0.3
Flavor & Extract Manufactures Association (FEMA), available in https://www.femaflavor.org/flavor-library.

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2.2. Operational Stability of the Different Biocatalyst in the Synthesis of Butyl Butyrate
The operational stabilities of all biocatalysts were evaluated in the syntheses of butyl butyrate at
37 ◦C for reactions cycles of 24 h (Figure 1). All biocatalysts maintained maximum conversion after
eight reaction cycles, except the commercial IMMAPF-T2-150, whose performance in the synthesis of
butyl butyrate was greatly reduced to approximately 60% conversion after seven cycles. This contrasts
with the very good behavior of PFL-octyl-silica, that suggests a great improvement on the enzyme
performance upon immobilization on the hydrophobic support prepared in this work. This could be
related to the lower adsorption of water, acids and alcohol in the enzyme environment [51,55,62].
◦
Figure 1. Reaction conversions as function of the number of cycles of butyl butyrate synthesis at 37 C
for 24 h cycles, using 0.1 M in heptane for acids and alcohols, and 200 U/g of acid (esterification activity)
and 20 g/L molecular sieves for all biocatalysts. IMMTLL-T2-150 (dark gray column), IMMAPF-T2-150
(gray column), IMMCALB-T2-350 (black column), PFL-octyl-silica (hatched column)-100% was the
conversions of the first reaction cycle. All assays were performed in duplicate and the standard
deviations were very closed to zero, with only a few closed to 2%.
2.3. Reaction Course of Butyl Butyrate Synthesis by PFL-Octyl-Silica and IMMCALB-T2-350 Biocatalysts
The home-made biocatalyst PFL-octyl-silica and the commercial IMMCALB-T2-350 were chosen
to evaluate the reaction course in the conversion of butyl butyrate synthesis. In this set of experiments,
the reaction progress was monitored by consumption of both, 1-butanol by gas chromatography and
butyric acid by titration. Figure 2 (PFL-octyl-silica catalyzed synthesis) and Figure 3 (IMMCALB-T2-350
catalyzed synthesis) shows that over 90% conversion is achieved within only 2 h of reaction. Although
both preparations were very active, IMMCALB-T2-350 was slightly more active in this reaction. Besides,
it can be noticed that the conversion calculated from data of titration and gas chromatography agree
reasonably well throughout the reaction. Thus, monitoring the reaction by acid titration (particularly
when stoichiometric ratio is used) is a simpler and faster method to be used routinely.

Molecules 2018, 23, 766
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◦
Figure 2. Time-conversion profiles of synthesis of butyl butyrate at 37 C catalyzed by PFL-octyl-silica
in heptane, using 0.1 M for acids and alcohols, 200 U/g of acid (esterification activity) and 20 g/L
molecular sieves. The conversions were assessed by consumption of 1-butanol by gas chromatography
(squares) and butyric acid by titration (triangles).
Figure 3.
Time-conversion profiles of synthesis of butyl butyrate at 37 ◦C catalyzed by
IMMCALB-T2-350, using 0.1 M in heptane for acids and alcohols, 200 U/g of acid (esterification
activity) and 20 g/L molecular sieves. The conversions were assessed by consumption of 1-butanol by
gas chromatography (squares) and butyric acid by titration (triangles).
2.4. Syntheses of Sugar Esters
2.4.1. Syntheses of Fructose Oleate—Stoichiometric Ratio
Figure 4 shows reactions courses of syntheses of fructose oleate catalyzed by IMMCALB-T2-350
and PFL-octyl-silica, using tert-butyl alcohol or tert-amyl alcohol as solvents, which are two of the most
frequently employed solvents for lipase-catalyzed carbohydrate ester syntheses [32]. These solvents
were chosen because they are sufficiently hydrophobic (logP of 0.80 and 1.4, respectively) to prevent
lipase inactivation, but sufficiently hydrophilic to dissolve the carbohydrates [7,32,47]. The highest
conversion (79%) was obtained with IMMCALB-T2-350 as biocatalyst and tert-butyl alcohol as solvent.
This conversion was reduced to 54% when tert-amyl alcohol was used as solvent using the same

Molecules 2018, 23, 766
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biocatalyst. On the other hand, using PFL-octyl-silica, similar conversions (around 60%) were obtained
with both solvents.
It has been reported that interactions between organic solvent molecules and essential water for
enzymatic activity in the environment of the enzyme could cause enzyme deactivation, particularly
when using more hydrophilic solvents. The more hydrophobic the organic solvent is (higher logP
value) the more active and stable the enzyme behaves [32,35,48]. On the other hand, the carbohydrate
capability of absorbing water could alter the essential water for the enzyme activity, reducing
ester conversion [35]. In these very complex systems, composed by organic solvent, organic acid,
the hydrophilic carbohydrate (logP =
−
2.23, according to the PubChem Data Base), molecular sieves
and different support surfaces, it is difficult to explain the behavior of the results with a unique
phenomenon. In fact, when using immobilized lipases, the adsorption of hydrophilic substrates
(e.g., carboxylic acid and alcohol), water molecules, or reaction products on the support is one of the
most relevant cause of biocatalyst inactivation [21].
Neta et al. [63] also reported conversions of 70% and 74% in the syntheses of fructose oleate
(stoichiometric ratio of fructose and oleic acid in ethanol as solvent) catalyzed by lipase from
Candida antarctica B immobilized on chitosan and commercial lipase B from Candida antarctica,
respectively. However, the authors did not report the selectivity of the reaction, since ethanol can
also be a substrate in the esterification reaction. Vescovi et al. [27] reported conversions between
◦
84% and 88% in the synthesis of fructose oleate at 55 C after 6 h of reaction catalyzed by Novozym 435
(immobilized CALB supplied by Novozymes Latin America Ltda., Araucária, PR, Brazil) and CALB
immobilized onto silica bi-functionalized with octyl and glutaraldehyde moieties, using fructose and
oleic acid (molar ratio of 1:5) in tert-butyl alcohol. Compared to PFL immobilized onto octyl-silica,
Vescovi et al. [
28] reported conversion around 40% in the synthesis of fructose oleate at 35 ◦C after
12 h of reaction using fructose and oleic acid (molar ratio of 1:5) in tert-butyl alcohol in absence of
water. However, when 1% (v/v) water was added to the reaction medium, around 90% conversion
was achieved. These examples show that the reaction conditions (temperature, time reaction, molar
ratio of substrates, organic solvent, absence/presence of water, etc.) need to be optimized for each
specific biocatalyst.
In our case, PFL-octyl-silica was better catalyst than IMMCALB-T2-350 using tert-amyl alcohol as
solvent, but it was worse using tert-butyl alcohol.
◦
Figure 4. Oleic acid conversion as a function of time for syntheses of fructose oleate at 45 C, 250 rpm
stirring, oleic acid and fructose initial concentrations of 25 mM, using IMMCALB-T2-350 (
ꢀ
) and
PFL-octyl-silica (
#
) in the presence of tert-butyl alcohol and IMMCALB-T2-350 (
ꢁ
) and PFL-octyl-silica
( ) in the presence of tert-amyl alcohol. For all assays 0.5 g of biocatalyst and 1.5 g of molecular sieves
were used.

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2.4.2. Syntheses of Lactose Oleate—Stoichiometric Ratio
Figure 5 shows the profiles of the reactions courses of the synthesis of lactose oleate catalyzed
by IMMCALB-T2-350 and PFL-octyl-silica, using tert-butyl alcohol and tert-amyl alcohol as solvents.
The maximum conversions were around 40% after 72 h reaction using both solvents and biocatalysts.
Lactose is a more hydrophilic carbohydrate (logP =
−
5.03, from the PubChem Data Base) than fructose
(logP = 2.23). This higher hydrophilicity of lactose could easily create a layer close to the biocatalyst
−
stripping away essential water for catalytic activity of the enzyme and/or impairing the access of oleic
acid to the enzyme active site. Besides, different affinities of lipases for different acyl acceptors in the
syntheses of sugar esters have also been reported [64]. Supposedly, more bulky carbohydrate could
exhibits more steric hindrances to the formation of the enzyme-substrate complex.
However, high conversion for syntheses of lactose fatty acid esters has been described.
Zaidan et al. [65] reported conversions above 70% in the acylation of lactose with capric acid (molar
ratio of 2:1) at 55 ◦C for 48 h, using acetone as solvent and lipase from Candida rugosa immobilized
on mica with amino groups. Neta et al. [63] also reported conversions above 80% in the synthesis
of lactose oleate (40 ◦C, 72 h reaction, stoichiometric ratio, initial reagent concentration of 83.3 mM)
catalyzed by lipase B from Candida antarctica immobilized on chitosan and acrylic resin (commercial
Novozym 435). However, ethanol was used as solvent and the authors did not report whether ethyl
oleate was produced as by-product.
In our case, both solvents and biocatalyst yielded similar yields after 72 h of reaction under the
chosen conditions. Curiously, in this reaction IMMCALB-T2-350 performance was not significantly
dependent on the solvent used. That is, the binomial solvent/enzyme may change when the substrate
changes, showing the complexity of the optimization of these processes.
◦
Figure 5. Oleic acid conversion as a function of time for syntheses of lactose oleate at 45 C, 250 rpm
stirring, oleic acid and lactose initial concentrations of 50 mM, using IMMCALB-T2-350 (
PFL-octyl-silica ( ) in the presence of tert-butyl alcohol and IMMCALB-T2-350 ( ) and PFL-octyl-silica
) in the presence of tert-amyl alcohol. For all assays 0.5 g of biocatalyst and 1.5 g of molecular sieves
ꢁ) and
ꢀ
(
#
were used.
2.4.3. Syntheses of Fructose and Lactose Laurate—Stoichiometric Ratio
Figures 6 and 7 show the reaction courses of the syntheses of fructose and lactose laurates.
Conversions of around 80% were achieved in the acylation of fructose catalyzed by IMMCALB-T2-350
when tert-amyl alcohol was used as solvent (Figure 6). Using tert-butyl alcohol as solvent,

Molecules 2018, 23, 766
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the conversion dropped to 40%. Using PFL-octyl-silica, again there was a lack of effect of the solvent.
As pointed above, IMMCALB-T2-350 showed to be more influenced by the solvent.
For the synthesis of lactose laurate (Figure 7), conversions of 40% were achieved using both
solvents and IMMCALB-T2-350, while PFL-octyl-silica yields decreased to 30%. This lower enzyme
activity versus lactose was observed in the syntheses of lactose oleates (Figure 5). As pointed above,
the carbohydrate hydrophilicity and/or the affinity of the lipase seem to be related to the lower
conversion using lactose as acyl acceptor. In this case, the effect of the solvent was not relevant for
any biocatalyst assayed, suggesting very complex relations and interferences involving, the support,
the enzyme and the substrate, as pointed above.
Figure 6. Lauric acid conversion as a function of time for syntheses of fructose laurate at 45 ◦C, 72 h,
250 rpm, oleic acid and fructose initial concentrations of 25 mM, using IMMCALB-T2-350 (
PFL-octyl-silica ( ) in the presence of tert-butyl alcohol and IMMCALB-T2-350 ( ) and PFL-octyl-silica
) in the presence of tert-amyl alcohol. For all assays 0.5 g of biocatalyst and 1.5 g of molecular sieves
ꢁ) and
ꢀ
(
#
were used.
Figure 7. Lauric acid conversion as a function of time for syntheses of lactose laurate at 45 ◦C, 72 h,
250 rpm stirring, oleic acid and lactose initial concentrations of 50 mM, using IMMCALB-T2-350 ( ) and
PFL-octyl-silica ( ) in the presence of tert-butyl alcohol and IMMCALB-T2-350 ( ) and PFL-octyl-silica
) in the presence of tert-amyl alcohol. For all assays 0.5 g of derivative and 1.5 g of molecular sieves
ꢁ
ꢀ
(
#
were used.

Molecules 2018, 23, 766
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2.4.4. Effect of the Fatty Acid:Carbohydrate Molar Ratio in the Synthesis of Oleate Fructose Esters
Oleic acid and tert-butyl alcohol were chosen for an analysis of the influence of oleic acid:fructose
molar ratio on the oleic acid conversion using different biocatalysts. Figure 8 shows the reaction
courses for the syntheses of fructose oleate at different concentrations of oleic acid and fructose. It can
be observed that the reaction conversion using IMMCALB-T2-350 as biocatalyst yielded 96% when a
oleic acid:fructose molar ratio 1:2 (25/50 mM) was used. On the other hand, using PFL-octyl-silica
the reaction conversion reached only 50% at the same concentrations of substrates, slightly lower
than the one obtained when the oleic acid:fructose molar ratio was 1:1 (25/25 mM), i.e., around 60%
conversion. In other words, excess fructose favored the reaction catalyzed by IMMCALB-T2-350,
but slightly disfavored the reaction catalyzed by PFL-octyl-silica. The chemical features of the supports
are very different and can explain these results. IMMCALB-T2-350 is prepared by covalent linkage
of CALB onto an polyacrylic resin activated with oxirane groups (ImmobeadTM IB-350, product
catalog from ChiralVision B.V., Leiden, The Netherlands), while PFL-octyl-silica is prepared by
hydrophobic adsorption of PFL onto silica surface modified with octyl groups [60]. It has been
described that lipase immobilized on hydrophobic supports may be released from the support at
presence of organic cosolvents, high temperatures, [66] or in the presence of high concentrations of some
substrates-products with surfactant properties [67–69]. In the case of PFL-octyl-silica, the reversible
nature of the linkage enzyme-support in the presence of the substrate oleic acid and a product with
detergent properties (fructose oleate) could desorb the enzyme from the support causing enzyme
inactivation, thus reducing the reaction conversion. In fact, as it will be show below, the operational
stability of the biocatalyst PFL-octyl-silica was lower than IMMCALB-T2-350 in successive batches of
syntheses of fructose oleate.
When excess of oleic acid was used (molar ratio oleic acid:fructose of 2:1) the maximum yield
regarding the acid should be 50% if the reaction stopped in monoester. However, reaction conversions
higher than 50% were achieved using both biocatalysts, suggesting that some diesters were being
produced during the reaction. This is relatively simple to explain manly considering that acyl
migrations may liberate the hydroxyl group that is modified by the enzyme [51,55,56].
Figure 8. Oleic acid conversion as a function of time for syntheses of fructose oleate in tert-butyl alcohol
as solvent at 45 ◦C, 72 h reaction and 250 rpm stirring. Molar ratio oleic acid/fructose of 1:2 (oleic
acid initial concentration 25 mM, fructose initial concentration 50 mM) using IMMCALB-T2-350 (ꢁ)
and PFL-octyl-silica (
fructose initial concentration 25 mM) using IMMCALB-T2-350 (
assays 0.5 g of derivative and 1.5 g of molecular sieves were used.
#
). Molar ratio oleic acid/fructose of 2:1 (oleic acid initial concentration 50 mM,
ꢀ
) and PFL-octyl-silica ( ). For all
The effect of the temperature on the conversion of syntheses of fructose oleate was evaluated in
the molar ratio oleic acid:fructose of 1:2 (25 mM of oleic acid and 50 mM of fructose) for syntheses

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catalyzed by IMMCALB-T2-350, and stoichiometric ratio (25 mM of fructose and oleic acid) for
syntheses catalyzed by PFL-octyl-silica. Figure 9 shows the reaction courses for syntheses of fructose
oleate carried out at 45 ◦C and 55 ◦C.
Figure 9. Oleic acid conversion as function of time for syntheses of fructose oleate using tert-butyl
alcohol as solvent, 72 h reaction, 250 rpm stirring, oleic acid initial concentration 25 mM, fructose initial
concentration 25 mM, using PFL-octyl-silica at 45 ◦C (
#
), PFL-octyl-silica at 55 ◦C ( ) and oleic acid
◦
initial concentration 25 mM, fructose initial concentration 50 mM, using IMMCALB-T2-350 at 45 C (
ꢀ
)
◦
and 55 C (ꢁ). For all assays 0.5 g of derivative and 1.5 g molecular sieves were used.
At 55 ◦C the conversion reached 96% within 24 h using IMMCALB-T2-350 as biocatalyst, while
using PFL-octyl-silica the conversion was slightly reduced from 57 (after 72 h at 45 ◦C) to 46% (after
72 h at 55 ◦C), although at shorter times the reaction progressed more rapidly at 55 ◦C. These results
could be related to the lower stability of the PFL-octyl-silica due to enzyme desorption from the
support surface, followed by enzyme inactivation.
2.4.5. Effect of the Biocatalyst Mass
Figure 10 shows the oleic acid consumption during the syntheses of fructose oleate with different
derivative masses (0.25, 0.5 and 1.0 g) using initial oleic acid and fructose concentrations of 25 mM
when utilizing PFL-octyl-silica as biocatalyst, and oleic acid and fructose initial concentrations of 25 and
50 mM, respectively, when employing IMMCALB-T2-350 as catalyst, using tert-butanol as solvent.
◦
Figure 10. Oleic acid conversion as a function of time on the syntheses of fructose oleate at 45 C,
250 rpm stirring, fructose initial concentration 25 mM, oleic acid initial concentration 25 mM using
0.25 g (
#
), 0.5 g ( ) and 1.0 g ( ) of PFL-octyl-silica and oleic acid initial concentration 25 mM, fructose
), 0.5 g ( ) and 1.0 g ( ) of IMMCALB-T2-350, using
initial concentration 50 mM using 0.25 g (
tert-butyl alcohol as solvent and 1.5 g molecular sieves for all synthesis reactions.
ꢁ
♦
ꢀ

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For the reactions catalyzed by IMMCALB-T2-350 the behavior was typical of enzyme catalyzed
reactions: the initial rate increased with increasing the mass of biocatalyst from 0.25 to 0.5 g,
but reaching similar conversions (90 to 96%) for 72 h reaction. When using a higher mass of biocatalyst
(1.0 g), the reaction rate did not significantly change, probably because the biocatalyst particles
aggregated and produced merged particles with higher diffusional limitations, more water retention,
etc. In the reactions catalyzed by PFL-octyl-silica, the reaction rate increased when the biocatalyst mass
increased from 0.25 to 0.5 g, but decreased when the mass was increased from 0.5 to 1.0 g, very likely
due to the same problem.
2.4.6. Operational Stability of the Biocatalysts in the Syntheses of Fructose Oleate
Figure 11 shows the operational stability of the biocatalysts PFL-octyl-silica and IMMCALB-T2-350
◦
in successive batches of 48 h in◦the synthesis of fructose oleate at 45 C. The performance of both
biocatalysts was reduced at 45 C, but more accentuated for PFL-octyl-silica: the conversion was
reduced to 50% after the fifth reaction cycle, while using IMMCALB-T2-350 the conversion was only
reduced to 80%. It may be assumed that enzyme desorption from the octyl-silica support occurs in the
presence of products with detergent properties (oleic acid and fructose oleate) [67–69].
Figure 11. Relative conversion as a function of the number of recycles for syntheses of fructose oleate at
◦
45 C, cycles of 48 h reaction, 250 rpm stirring, 25 mM of initial concentrations of fructose and oleic acid
using PFL-octyl-silica (
50 mM using IMMCALB-T2-350 (
of biocatalyst and 1.5 g molecular sieves. The conversion for the first cycle was considered as 100%.
ꢁ) and oleic acid initial concentration of 25 mM, fructose initial concentration of
#
). All assays were carried out in tert-butyl alcohol as solvent, 0.5 g
3. Materials and Methods
3.1. Materials
Lipase from Pseudomonas fluorescens (PFL) was acquired from Sigma-Aldrich Co. (St. Louis, MO,
USA). Commercial immobilized lipases from Candida antarctica type B (CALB, IMMCALB-T2-350),
Thermomyces lanuginosus (Lipolase, IMMTLL-T2-150) and Pseudomonas fluorescens (Amano AK,
IMMAPF-T2-150) were acquired from ChiralVision (Leiden, The Netherlands). All these commercial
lipases are covalently attached to dry acrylic beads (ImmobeadTM IB-150 or IB-350—crosslinked
copolymer of methacrylate exhibiting oxirane groups—particle size 150–200 micrometer or
300–700 micrometer, respectively) Product catalog (ChiralVision B.V., Leiden, The Netherlands).
Macroporous silica (Immobead S60S, particle size of 60–200 micrometer) was attained from

Molecules 2018, 23, 766
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ChiralVision (Supplementary material). Molecular sieve UOP type 3 Å (rod, size 1/16 in.) was
purposed by Sigma-Aldrich Co. 2-Methyl-2-butanol (tert-amyl alcohol), 1-hexanol and 1-butanol were
purchased from Sigma-Aldrich Co. Ethanol (99.7%) was purchased from JT Baker (Phillipsburg, NJ,
USA). 2-Methyl-2-propanol (tert-butyl alcohol), lactose and lauric acid were purchased from Vetec
(Rio de Janeiro, Brazil). Heptane, isoamyl alcohol, fructose and oleic acid were purchased from Synth
(São Paulo, Brazil). Acetic acid was purchased from Qhemis (Sao Paulo, Brazil).
3.2. Production of Octyl-Silica Support
The chemical modification of silica with octyl groups was performed according to Tani and
Suzuki [70]. Firstly, silica particles were incubated in 0.1 M hydrochloric acid and washed with
◦
distilled water until the pH was neutral. Afterwards, the particles were dried at 200 C for 5 h.
One gram of dried silica was maintained unde◦r reflux with 20 mL of a mixture of toluene and
octyltriethoxysilane (volume ratio of 10:1) at 85 C. After 3 h, the suspension was filtered and the
support was thoroughly washed with toluene, methanol, and distilled water. Finally, the support
was dried at room temperature for 24 h. The activated support, named octyl-silica, was used for the
immobilization of PFL.
3.3. Immobilization of PFL on Octyl-Silica
Octyl-silica particles were suspended in an enzymatic solution prepared in 5 mM sodium
phosphate (pH 7.0) in a support:enzymatic solution ratio of 1:19 (w/v). Two derivatives were prepared:
a low-loaded derivative (5 mg protein/g support) and a high-loaded derivative (40 mg protein/g
support), although the loading capability of this support◦is around 168 mg protein/g support [60].
The suspension was gently stirred in an orbital shaker at 25 C for 24 h [39]. PFL-octyl-silica biocatalyst
was filtered and washed with Milli-Q water. In order to monitor the enzyme immobilization, enzyme
hydrolytic activity versus emulsified olive oil was assessed in the supernatant solution and in a control
enzyme solution incubated under the same conditions, but without support. The immobilization
parameters (immobilization yield and recovered activity) were calculated as previously described [60].
3.4. Synthesis of Flavor Esters
The syntheses of flavor esters were performed according to Abbas and Comeau [3] in closed glass
flasks. Briefly, 10 mL of a solution of short-chain acids (acetic or butyric acid) and alcohols (1-butanol,
1-hexanol, ethanol or isoamyl alcohol) were prepared in heptane to a final concentration of 0.1 M
◦
(for both acid and alcohol). The esterification reactions were performed at 37 C under 250 rpm stirring
in the presence of molecular sieves (20 g/L). A load of 200 U/g of acid (in terms of esterification
activity as describe below) was used for all biocatalysts: home-made PFL-octyl-silica (low-loaded
derivative, i.e., 5 mg protein/g support) and commercial immobilized lipases (IMMCALB-T2-350,
IMMTLL-T2-150 and IMMAPF-T2-150).
The reaction conversion (Equation (1)) was assessed from the results of acid consumption by
titration with 0.025 M KOH. Samples of 0.5 mL of the reaction medium were takenat regular time
periods and diluted with 10 mL of a solution of acetone:ethanol (1:1 v/v) containing phenolphthalein
as the pH indicator. All assays were performed in triplicate:
molt=0 − molt=t
C(%) =
× 100
(1)
molt=0
where C(%) is the reaction conversion, molt is the initial number of moles of fatty acid and molt=t is
=0
the number of moles of fatty acid at time t.
In order to evaluate whether the titration was suitable to monitor the reaction, the synthesis
of butyl butyrate was also followed by gas chromatography. Samples of 1
µL were injected in a
HP5890 Gas Chromatograph (Hewlett-Packard Company, Palo Alto, CA, USA) equipped with a

Molecules 2018, 23, 766
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flame ionization detector, using nitrogen as carrier gas and a crossbond acid deactivated Carbowax
polyethelene glycol column (15 m 0.32 mm 0.45 m, Restek Corporation, (Bellefonte, PA, USA)).
×
×
In this case, the reaction conversion was calculated based on the 1-butanol consumption (in this case,
all experiments were carried out using molar ratio butyric acid/1-butanol of 1:1).
3.5. Synthesis of Sugar Esters
The syntheses of sugar esters were carried out according to Paula et al. [71] in closed glass flasks.
Briefly, 10 mL of a solution of fructose (25 or 50 mM) or lactose (50 mM) were prepared by dissolving the
carbohydrates in organic solvents (tert-butyl alcohol or tert-amyl alcohol). Afterwards, oleic or lauric
acids were added to final concentrations of 25 or 50 mM. The reactions were carried out in the presence
of 1.5 g of molecular sieves (150 g/L), in order to reduce and control the water activity in the reaction
medium. Adachi and Kobayashi [47] reported an increase in the equilibrium conversion for lauroyl
mannose synthesis adding to the system until 100 g/L of molecular sieve 3A. In this work, we adopt
a concentration of 150 g/L to guarantee water removal from the reaction system. The biocatalysts
used in these reactions were PFL-octyl-silica (high-loaded, i.e., 40 mg protein/g support) and the
commercial immobilized lipase from Candida antarctica (IMMCALB-T2-350), using biocatalyst masses
of 0.25, 0.5 and 1.0 g. The reactions were carried out in triplicate at 45 or 55 ◦C for 72 h under 250 rpm
stirring. The reaction conversion was monitored by titration with 0.01 M KOH solution.
3.6. Operational Stability of the Biocatalysts
The operational stability of the biocatalysts was studied in the syntheses of butyl butyrate and
fructose oleate, as described below.
3.6.1. Flavor Ester (Butyl Butyrate)
PFL-octyl-silica, IMMCALB-T2-350, IMMTLL-T2-150 and IMMAPF-T2-150 were used in eight
24 h-cycles of synthesis of butyl butyrate at 37 ◦C under 250 rpm stirring, using heptane as solvent,
acid and alcohol concentrations of 0.1 M and 20 g/L molecular sieve. The reaction volume and the
enzyme load were 10 mL and 200 U/g butyric acid (in terms of esterification activity as described
below), respectively. After each cycle, the biocatalysts were washed with heptane before being used in
a new reaction cycle. The conversion of the reaction after 24 h was calculated from the consumption of
acid, monitored by titration. All assays were performed in duplicate.
3.6.2. Sugar Ester (Fructose Oleate)
PFL-octyl-silica and IMMCALB-T2-350 were employed in five 48 h-cycles of synthesis of fructose
oleate employing 0.5 g of biocatalyst, 10 mL of tert-butyl alcohol as solvent, and 1.5 g of molecular
◦
sieves at 45 C under 250 rpm stirring. The initial molar ratio fructose/oleic acid was 25:25 (mM)
for PFL-octyl silica and 25:50 (mM) for IMMCALB-T2-350. After each 48 h-batch, the biocatalysts
were washed with tert-butanol and used in a new esterification. The conversion of the reaction
was calculated from the consumption of acid, monitored by titration. All assays were carried out
in triplicate.
3.7. Esterification Activity Assay
The esterification activity of the biocatalysts was measured at 37 ◦C under 250 rpm stirring in
closed glass flasks, using 1-butanol and butyric acid (both at 0.1 M) in heptane (10 mL reaction volume)
as acceptor and donor of acyl group, respectively [71]. Samples were taken from the reaction medium
at the defined times (up to 40 min) for quantification of butyric acid by titration with 0.0105 M KOH
solution. One unit of esterification activity was defined as the consumption of 1
per minute under the reaction conditions.
µmol of butyric acid

Molecules 2018, 23, 766
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4. Conclusions
Lipase from Pseudomonas fluorescens immobilized on silica modified with octyl groups
(PFL-octyl-silica biocatalyst) presented similar performance and operational stability compared to
the covalent commercial preparation of the same lipase or other lipases in the ester syntheses with
flavor properties. On the other hand, different interactions between biocatalysts/medium/substrate
might have caused different catalytic behavior of the enzyme during sugar ester synthesis. Such
differences might be related either to the nature of the enzymes or interactions between substrates
and the biocatalyst itself (support and/or enzyme). In the syntheses of sugar fatty acid esters,
PFL-octyl-silica reached lower conversion and showed lower operational stability compared to the
CALB IM biocatalyst.
Supplementary Materials: The following are available online.
Acknowledgments: This work was supported by the São Paulo Research Foundation (FAPESP) (grant numbers
#2011/23194-0, #2016/10636-8), Brazilian National Research Council—CNPq (Lionete Nunes de Lima’ doctorate
fellowship), and Spanish MINECO (Project CTQ2017-86170-R). The help and comments from Ángel Berenguer
(Instituto de Materiales, Universidad de Alicante) are gratefully recognized.
Author Contributions: Lionete Nunes de Lima performed the experiments; Adriano Aguiar Mendes
designed the experiments of synthesis of short-chain carboxylic esters; Raquel de Lima Camargo Giordano,
Roberto Fernandez-Lafuente and Paulo Waldir Tardioli designed the experiments and wrote the paper.
Conflicts of Interest: The authors declare no conflict of interest.
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