Covalently immobilized lipase catalyzing high-yielding optimized geranyl butyrate synthesis in a batch and fluidized bed reactor

Article abstract of DOI:10.1016/j.molcatb.2011.11.009

Three commercially available polymers (Sepabeads EC-EP, Sepabeads EC-HA and Purolite A-109) were tested for potential application as supports for covalent immobilization of lipase from Candida rugosa by analyzing some critical properties of immobilized enzymes such as enzyme loading, activity and activity immobilization yield. Among them, lipase covalently immobilized on Sepabeads EC-EP via epoxy groups appeared to show the best performance in a standard hydrolytic reaction. Therefore, it was selected and assayed in the esterification of butyric acid and geraniol to produce geranyl butyrate, first in a batch system followed by continuous geranyl butyrate synthesis in a fluidized bed reactor, as one being potentially applicable for large-scale production. Based on statistical analysis, optimal conditions for the production of geranyl butyrate by selected, immobilized lipase in the batch system are recommended as: temperature at 25-30°C, water concentration at 3.6% (v/v) and acid/alcohol molar ratio at 2.5. A set of optimal conditions for the ester synthesis in a fluidized bed reactor system has also been determined, specifically, flow rate at 10 mL min^-1, temperature at 35°C, water concentration at 2% (v/v), substrate concentration at 0.1 M and acid/alcohol ratio at 2.0. Implementation of the optimized parameters in a batch system and in a fluidized bed reactor enabled production of target ester with high molar conversion, at > 99.9% for 48 h in the batch process, and 78.9% for 10 h in fluidized bed reactor. Although when assayed at their optimal conditions, lower molar conversion was achieved in the fluidized bed reactor system compared to the batch system, the volumetric productivity in fluidized bed reactor was more than five fold higher than that obtained in the batch system.

Full text of DOI:10.1016/j.molcatb.2011.11.009

Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
Contents lists available at SciVerse ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Covalently immobilized lipase catalyzing high-yielding optimized geranyl
butyrate synthesis in a batch and fluidized bed reactor
a
a
a
c
Jasmina J. Damnjanovic´ a,b,∗, Milena G. Zuzˇa , Jova K. Savanovic´ , Dejan I. Bezbradica , Dusˇan Z. Mijin ,
ˇ
ˇ
Nevenka Bosˇkovic´-Vragolovic´ d, Zorica D. Knezˇevic´-Jugovic´ a
a Department of Biochemical Engineering and Biotechnology, Faculty of Technology and Metallurgy, Karnegijeva 4, University of Belgrade, Belgrade, Serbia
b Laboratory of Molecular Biotechnology, Graduate School of Bioagricultural Sciences, Nagoya University, Nagoya, Japan
c Department of Organic Chemistry, Faculty of Technology and Metallurgy, Karnegijeva 4, University of Belgrade, Belgrade, Serbia
d Department of Chemical Engineering, Faculty of Technology and Metallurgy, Karnegijeva 4, University of Belgrade, 11000 Belgrade, Serbia
a r t i c l e i n f o
a b s t r a c t
Three commercially available polymers (Sepabeads® EC-EP, Sepabeads® EC-HA and Purolite® A-109)
were tested for potential application as supports for covalent immobilization of lipase from Candida
rugosa by analyzing some critical properties of immobilized enzymes such as enzyme loading, activity
and activity immobilization yield. Among them, lipase covalently immobilized on Sepabeads® EC-EP via
epoxy groups appeared to show the best performance in a standard hydrolytic reaction. Therefore, it was
selected and assayed in the esterification of butyric acid and geraniol to produce geranyl butyrate, first
in a batch system followed by continuous geranyl butyrate synthesis in a fluidized bed reactor, as one
being potentially applicable for large-scale production.
Based on statistical analysis, optimal conditions for the production of geranyl butyrate by selected,
immobilized lipase in the batch system are recommended as: temperature at 25–30 ◦C, water concentra-
tion at 3.6% (v/v) and acid/alcohol molar ratio at 2.5. A set of optimal conditions for the ester synthesis
in a fluidized bed reactor system has also been determined, specifically, flow rate at 10 mL min−1, tem-
perature at 35 ◦C, water concentration at 2% (v/v), substrate concentration at 0.1 M and acid/alcohol ratio
at 2.0. Implementation of the optimized parameters in a batch system and in a fluidized bed reactor
enabled production of target ester with high molar conversion, at > 99.9% for 48 h in the batch process,
and 78.9% for 10 h in fluidized bed reactor. Although when assayed at their optimal conditions, lower
molar conversion was achieved in the fluidized bed reactor system compared to the batch system, the
volumetric productivity in fluidized bed reactor was more than five fold higher than that obtained in the
batch system.
Article history:
Received 10 June 2011
Received in revised form 4 November 2011
Accepted 8 November 2011
Available online 17 November 2011
Keywords:
Geranyl butyrate
Covalent immobilization
Enzymatic esterification
Fluidized bed reactor
Optimization study
© 2011 Elsevier B.V. All rights reserved.
1. Introduction
and poor quality of the product [4,5]. Also, chemical processes
demand high costs for additional separation and purification steps
Candida rugosa lipase (CRL) is a versatile, robust, frequently
used lipase since it can be easily produced in lyophilized form and
efficiently prepared in large amounts. Reports have been made
describing successful CRL catalyzed synthesis of terpene esters
[1–3]. In industrial scale, aromatic terpene esters are currently syn-
thesized in a nonspecific chemical process obtaining low yields
[6]. Besides higher product yields, mild operating conditions, syn-
thesis of products that do not need further purification, enzymatic
reaction delivers “natural product”, in terms of its origin, which is
of a high importance, especially in the food industry [7].
However, industrial-scale synthesis using soluble enzymes is
economically unacceptable, since these enzymes lack reusability as
well as possibility of continuous type synthesis due to their low sta-
bility and complex separation techniques [8]. One way to address
these issues is enzyme immobilization.
Covalent immobilization provides formation of a very sta-
ble catalyst via multipoint covalent attachment by maintaining
enzyme’s active conformation and reducing denaturing effects
of environmental factors [9]. Among materials used for covalent
immobilization, Sepabeds® EC series show physical and chemical
stability, high protein binding capacity, low swelling tendency in
Abbreviations: CRL, Candida rugosa lipase; FBR, fluidized bed reactor; CCRD,
central composite rotatable design.
∗
Corresponding author at: Laboratory of Molecular Biotechnology, Graduate
School of Bioagricultural Sciences, Nagoya University, Furo-cho, Chikusa-ku, 464-
8601 Nagoya, Japan. Tel.: +81 90 9180 1389.
E-mail address: damnjanovic.jasmina@b.mbox.nagoya-u.ac.jp
(J.J. Damnjanovic´).
1381-1177/$ – see front matter © 2011 Elsevier B.V. All rights reserved.
doi:10.1016/j.molcatb.2011.11.009

J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
51
2. Experimental
high molar solutions and resistance to the microbial degradation,
making them suitable for application in industrial bioprocesses
[10–13]. Specifically, Sepabeads® EC-EP particles are shown to
efficiently stabilize penicillin G acylase providing hundreds-fold
2.1. Materials
more stable catalyst compared to the one attached to Eupergit®
C
Nonspecific lipase AY, Type VII, L 1754, in powdered form from
C. rugosa, lipase substrate (stabilized olive oil emulsion), Triton X-
100, bovine serum albumin (BSA) and terpene alcohol, geraniol
(98.0%) were purchased from Sigma–Aldrich Inc. St. Louis, MO, USA.
Butyric acid from Merck, Darmstadt, Germany was used as acid
substrate while isooctane (HPLC grade) from Arcos Organics, New
Jersey, USA, was used as organic solvent. Sepabeads® EC-EP and
Sepabeads® EC-HA were donated by Resindion S.R.L., Mitsubishi
Chemical Corporation, Milan, Italy. Purolite® A-109 was purchased
from Purolite International Ltd. (Llantrisant, United Kingdom). All
other chemicals were purchased from Merck, Darmstadt, Germany.
[14]. Hilterhaus et al. successfully immobilized three industrially
important enzymes, endoglucanase, benzoylformate decarboxy-
lase from Pseudomonas putida and lipase from Candida antarctica on
Sepabeads® EC-EP, EC-EA and EC-BU, producing stable and active
enzymes suitable for industrial application [15].
The selection of a proper reactor configuration is another
important aspect in designing industrial enzymatic synthe-
sis. Fluidized bed reactors (FBRs) are widely and successfully
used in many industrial processes, like aerobic fermentation
processes, catalytic reactions and biological waste-water treat-
ment, due to continuous operational mode as well as for
improved heat and mass transfer [16]. However, information
on their application for ester synthesis is currently rather
limited in the literature. The majority of studies concern-
ing ester synthesis focused on batch systems and packed bed
reactors [17], while ester synthesis in FBRs remains yet unex-
plored.
In studies conducted so far, geranyl butyrate was produced in
yields between 85 and 99.9% using free or immobilized lipases
[5,18–20]. Recently, it was shown that CRL immobilized on
Sepabeads® EC-EP can be employed as a robust biocatalyst in ester-
ification of geraniol with butyric acid in a low aqueous system
[21]. However, these studies have been generally performed in the
flasks with magnetic stirring or in the vials immersed in an orbital
shaker. Analysis of the effect of reactor configuration and hydro-
dynamic conditions on the reaction rate and enzyme stability was
not included in these contributions. To the best of our knowledge,
this report presents the first study of the parameters affecting the
synthesis of geranyl butyrate with the immobilized lipase in FBR
system.
Previous research of Saponjic et al. regarded application of CRL
immobilized on Sepabeads® EC-EP for amyl caprylate synthesis
in FBR [22]. The study revealed that both batch and continuous
synthesis of amyl caprylate can be accomplished in a high yield.
Continuous esterification had slightly improved kinetics since
90.2% yield was achieved within 14 h compared to the batch syn-
thesis where almost complete conversion was observed within 24 h
[22]. As a further research, we aimed to prove robustness and versa-
tility of immobilized CRL and efficiency of the reaction optimization
pathway by employing them for geranyl butyrate synthesis. Using a
different reaction model which includes acid substrate of three fold
higher polarity and synthesis of a higher polarity ester would pose
additional limitations to the ongoing esterification, since lipase
activity is known to be affected by changes in substrate partitioning
between organic phase and water layer surrounding the enzyme
[23,24].
2.2. Immobilization method
Immobilization of CRL on epoxy-Sepabeads® (EC-EP): Immobiliza-
tion of CRL on epoxy-activated support, Sepabeads® EC-EP, was
achieved by direct lipase coupling to the polymer via epoxy groups
in the presence of very high salt concentration (standard protocol
recommended by Resindion S.R.L., Mitsubishi Chemical Corpora-
tion). Immobilization was performed in 70 mL of 1.25 M potassium
phosphate buffer, pH = 8.0 at 22.5 ◦C with orbital mixing for 48 h.
Immobilized enzyme was then washed with water and buffer and
kept at 4 ◦C followed by drying in vacuo for 48 h, prior to the reac-
tion.
Immobilization of CRL on Sepabeads® EC-HA and Purolite® A-109:
The immobilization procedure on amino-supports consisted of two
steps: (1) oxidation of the lipase by sodium periodate; and (2) cou-
pling of the oxidized enzyme to the amino-supports.
Therefore, first step was the lipase oxidation by sodium
periodate following the methodology previously described [25].
According to this protocol, 1 mg mL−1 of crude enzyme solutions,
corresponding to 0.25 mg mL−1 of pure protein determined by the
Bradford method [26] were incubated in 5 mM solution of sodium
periodate in sodium acetate buffer, pH = 5.0, for 6 h in the dark at
4 ◦C. The reaction mixture was stirred occasionally and the reaction
was quenched with 10 mM ethylene glycol for 30 min. To remove
by-products, the oxidized lipase solution was then dialyzed against
50 mM sodium acetate buffer, pH = 5.0, for 18 h.
Polymers with amino groups (1.0 g) were incubated with 35 mL
of oxidized lipase solution in sodium acetate buffer at pH = 5.0 and
4 ◦C for 48 h. Afterwards, obtained biocatalysts were washed with
water and sodium phosphate buffer, pH = 7.0 and stored in the same
buffer at 4 ◦C until use. In previous research, it was reported that
Schiff’s bases formed between oxidized enzyme and aminated sup-
ports proved as very stable, therefore, additional reduction step was
not applied to the immobilized preparations [25,27]. The lipase oxi-
dation was checked using FT-IR spectrometry as described bellow.
Focus was set on designing an economical fluidized-bed-
immobilized-enzyme system for geranyl butyrate synthesis
employing an efficient and inexpensive commercial support
and immobilization method. In the first part of the study,
characteristics of CRL covalently immobilized on Sepabeads®
EC-EP have been compared to those obtained with other two
commercial supports. Some critical properties of immobilized
enzymes such as protein loading, activity, specific activity and
immobilization yield were considered. Selected highly active
immobilized lipase was used for statistical assessment of
relevant process conditions in the batch system and in fur-
ther study as a biocatalyst and constituent of the FBR. The
bioreactor hydrodynamic characteristics and the reaction condi-
tions have been investigated in order to improve the process
performance.
2.3. Immobilization parameters
Protein loading, Pg defined as the amount of the pure protein
coupled to the supports (mg of protein/g of the supports) is cal-
culated as a difference between protein amount (mg) added in
the immobilization process and the protein amount (mg) found
in the filtrate and wash-through after immobilization. The protein
loading efficiency, YP (%) was calculated according to Eq. (1):
C0V0 − (CfVf + CwVw
)
YP (%) =
(1)
C0V0
where C0 is the protein concentration of the initial immobilization
solution (mg mL−1); V0 its volume (mL); Cf the protein concentra-
tion in the filtrate (mg mL−1); Vf the filtrate volume (mL); Cw the

52
J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
protein concentration in washing solution (mg mL−1) and Vw its
volume (mL).
used in the calculation of enzyme mass (me) in samples. Calculation
is based on the following equation:
Specific activity of the immobilized enzyme, SAIE was defined
A
as the hydrolytic activity of the immobilized protein, A (IU g−1
)
me
=
Vs
(3)
ls
divided by the protein loading and expressed as IU mg−1 protein.
The activity immobilization yield, YA (%) was calculated by dividing
the specific activity of the immobilized lipase by specific activity of
the free lipase.
where Vs stands for sample volume, and
A
stands for the
absorbance. The final value of enzyme concentration is the average
of two values obtained on two wave lengths. Modified standard
curves in the presence of Triton X-100 or NaCl were used for the
desorption experiments with the detergent (0.5%, v/v), or NaCl
(0.50–1.25 M).
2.4. Desorption study
100 mg of the immobilized enzyme samples were suspended
in 10 mL of 10 mM sodium phosphate buffer at pH = 7.0 contain-
ing 0.5% (v/v) Triton X-100 or 0.50–1.25 M NaCl and incubated at
room temperature in orbital shaker (150 rpm). Samples were taken
from the supernatant after 60 min and the amount of lipase was
determined as described above. Desorption study from different
supports was performed for two values of protein loading in each
support. Desorption yield, YD (%) was defined as the amount of des-
orbed protein (calculated per 1 g of biocatalysts) divided by the
protein loading expressed in percentage [25].
2.6.2. Fourier transform infrared spectroscopy (FT-IR) analysis
Samples of free lipase and oxidized CRL were subjected to FT-
IR analysis and the spectra were obtained using a Bomem MB 100
FT-IR Spectrophotometer. The amount of 10 wt% of the sample was
mixed and ground with 100 wt% of potassium bromide and then
compressed into a pellet under a pressure of 11 t, for about a minute,
using Graseby Specac Model: 15.011. The Spectra were recorded in
the 400–4000 cm−1 wave number range.
2.6.3. Ester analysis
Ester synthesis was monitored by determination of the residual
acid content by titration against sodium hydroxide using phenolph-
thalein as an indicator and methanol as a quenching agent. After
48 h reaction time, 0.5% phenolphthalein in methanol was added in
the reaction mixture followed by titration with 0.1 M NaOH solu-
tion to determine the amount of residual butyric acid [30]. The
reported molar conversion was calculated as
2.5. Geranyl butyrate synthesis in a batch system
Syntheses were done in a screw capped flasks (100 mL) con-
taining precalculated proportions and concentrations of geraniol,
butyric acid and water in isooctane. 500 mg of immobilized lipase
was added to the preheated reaction mixture. Flasks were incu-
bated at variable temperatures in orbital shaker (150 rpm) for
48 h. The reaction time was selected based on preliminary results
of the batch runs showing that the reaction reached the equi-
librium after 48 h. Substrate concentration was fixed at 0.25 M.
Other reaction parameters varied according to experimental design
(Supplemental Table 1). Blank experiments were done under iden-
tical conditions.
Five-level-five-factor central composite rotatable design (CCRD)
was used which included 32 experiments consisting of 16 factorial,
10 axial, and 6 central points [28]. Experiments were run in random
order to avoid bias. Obtained data were fitted to a second-order
polynomial equation:
B0 − A0
Y (%) =
× 100
(4)
B0
where Y stands for molar conversion (%), B0 (mL) and A0 (mL) stand
for volume of 0.1 M NaOH spent to titrate reaction mixture before
and after synthesis, respectively.
The accuracy of this method was tested by determination
of ester concentration on Varian 3400 gas chromatograph as
described previously [22]. The ester yield (%) was defined as the
amount of geranyl butyrate produced from initial substrate in
defect (mol ester/mol initial substrate in defect × 100). The ester
yield (%) and molar conversion (%) were found to be in good agree-
ment.
5
5
4
5
ꢀ
ꢀ
ꢀꢀ
Y = ˇk0
+
ˇkiXi +
ˇkiiXi2
+
ˇkijXiXj
(2)
2.7. Lipase activity assay
i=1
i=1
i=1 j=i+1
Activities of immobilized enzymes were determined in the reac-
tion of olive oil hydrolysis by Sigma method as described previously
[25] and expressed in IU. This activity assay was carried out with
reaction mixtures containing 3 mL of Sigma lipase substrate, 1 mL
of Trizma buffer, and 3.5 mL of distilled water. 1 IU is defined as the
amount of enzyme required to produce 1 ␮mol of free fatty acid per
minute under the assay conditions (37 ◦C, pH 7.7).
where Y is the response (molar conversion in %), ˇk0, ˇki, ˇkii and ˇkij
are regression coefficients for intercept, linear, quadratic and inter-
action terms, respectively, and Xi and Xj are independent variables.
The values and statistical significances of the response function
coefficients were calculated using the method of least squares in
the MATLAB software (version 6.5, Release 13, The MathWorks,
Juc, Matick, MA, USA). Contour plots were obtained by using fitted
model. Ester synthesis was optimized and obtained response equa-
tion enabled prediction of molar conversion from known values of
the five main factors.
2.8. Geranyl butyrate synthesis in a fluidized bed reactor (FBR)
2.8.1. Hydrodynamic characteristic of the FBR system
Tracer response analysis was used to characterize and model
the flow through the reactor, according to previously described
methodology [31]. The system was disturbed by introducing a step
input change of the tracer (black ink) concentration and system
response was then monitored at the reactor output. Experimen-
tal setup is given in Fig. 1a. Effluent of the reactor was sampled
at timed intervals, and tracer concentration was determined spec-
trophotometrically at 400 nm. The experiments were performed in
duplicates. Obtained tracer concentration profiles were normalized
2.6. Analysis
2.6.1. Enzyme concentration assay
Enzyme concentration in samples was determined spectropho-
tometrically at 215 nm and 225 nm using CRL as a standard [29].
Calibration plots for enzyme were generated for both wave lengths
by plotting averaged absorbance against enzyme concentration and
presented by their linear regression formulas. Line slopes (ls) are

J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
53
In this study, Tanks-in-series model, presented by Eqs. (6) and (7)
was used to characterize non-ideal flow within the reactor:
ꢁ
ꢂ
2
N−1
¯
¯
(N(t/t))
t
(N(t/t))
¯
F = 1 − e(−N(t/t)) 1 + N
+
+ · · · +
(6)
(7)
¯
t
2!
(N − 1)!
NN
tN−1
¯
E =
e−(Nt/t)
N−1
¯
t
(N − 1)!
¯
where N is the number of stirred tanks, t is mean residence time in
the system, and t is time of sampling.
2.8.2. Enzymatic reactor setup
Fluidized bed reactor (FBR) was a cylindrical glass column
(10 mm i.d. × 136 mm length) packed with 2.0 g of immobilized
enzyme (3.75 cm g−1, density 1.13 g mL−1) and equipped with a
water jacket for temperature control. The reaction mixture (80 mL,
containing 2% (v/v) water) was fed upwards through the column
using a peristaltic pump (Behrotest® Labor-Schlauchpumpe PLP
66, Dusseldorf) at different flow rates. Prior to entry into the reac-
tor zone, the substrate mixture was preheated to a predetermined
temperature by thermostatic bath, while the identical tempera-
ture in the reactor zone was maintained constant by circulating
preheated water through the reactor jacket. Reaction was carried
out by recirculation of the reaction mixture through the FBR. Sam-
pling was conducted at specified times in the substrate reservoir.
Experimental set-up is shown in Fig. 1b.
For estimation of optimal flow rate and substrate concentration,
parameter P or volumetric productivity, was used. It describes effi-
ciency and feasibility of the process regarding the concentration of
the product formed and a reaction time. Calculation is presented
by the following equation:
[E]
t
P =
(8)
where [E] stands for ester concentration and t stands for the reac-
tion time needed for production of [E].
3. Results and discussion
3.1. Enzyme immobilization study
As reported in our previous studies, CRL was successfully immo-
bilized on epoxy activated supports [22,25]. This time, CRL was
covalently immobilized on epoxy-activated Sepabeads® EC-EP,
and two amino-supports, Sepabeads® EC-HA and Purolite® A-
109, in order to compare the immobilization performances of
obtained biocatalysts. The selected supports used in this work
are relatively inexpensive and readily available commercially. The
coupling of enzyme to epoxy-activated carrier, Sepabeads® EC-EP,
involved reactions between enzyme’s available amino or sulfhydryl
groups and active epoxy groups of the polymer in the presence
of high salt concentrations. In fact, the reactivity of epoxy groups
towards nucleophilic groups of the enzyme is very low under
mild experimental conditions (neutral pH, low ionic strength) [32].
On the other hand, successful immobilization could be achieved
by using hydrophobic epoxy supports such as Sepabeads® EC-
EP and performing the enzyme coupling in the presence of high
salt concentrations (e.g. in 1.25 M potassium phosphate buffer).
The immobilization is reported to follow a very interesting two
step mechanism: first, a rapid mild physical adsorption between
the enzyme and the support occurs followed by covalent reac-
tion between adsorbed protein and epoxy groups of the support
[32,33]. Immobilization of CRL on amino-supports, Sepabeads® EC-
HA and Purolite® A-109, could be achieved via lipase carbohydrate
moiety previously modified by periodate oxidation. The exact way
of enzyme attachment is not well known, but covalent coupling
Fig. 1. Experimental setup for flow pattern analysis (a) and for lipase catalyzed
esterification (b): (1) distilled water reservoir; (2) solution of labeled substance
(black ink); (3) peristaltic pump; (4) fluidized bed reactor; (5) reactor water jacket;
(6) cooling/heating water; (7) substrate reservoir; (8) reactor temperature control;
(9) sampling.
with respect to the inlet tracer concentration, and resulting normal-
ized C/C0 functions were plotted against time to give experimental F
curve. Experimental E curves (exit age distribution functions) were
then obtained by derivation, as follows:
dF
E =
(5)
dt

54
J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
between supports’ amino groups and enzyme carbohydrate moiety
was expected to be the most probable mechanism. The idea behind
the method was the formation of a Schiff’s base linkage between
carbonyl groups of the modified lipase and free amino groups on
the supports [34].
other functional groups covered oxidized lipase carbonyl bands
detectable in that range. However, it is clear that CRL and oxidized
CRL showed differences in the FT-IR spectra, more specifically in the
position and intensities of the primary hydroxyl group and carbonyl
group related bands, characterizing differences in the oxidation
states.
Immobilization data for three tested supports and covalent tech-
niques are summarized in Table 1, along with desorption data for
each immobilization system. Although the enzyme binding capac-
ity of supports often largely depends on the surface density of
functional groups and the nature of the coupling reactions, the
protein loading for all three tested supports was similar and quite
satisfactory. Protein loading increased with increasing protein con-
centration in the coupling solution reaching maximum values of
19.2, 16.1 and 20.0 mg g−1 dry supports for Sepabeads® EC-EP,
Sepabeads® EC-HA and Purolite® A-109, respectively. The slight
decrease in protein loading on Sepabeads® EC-EP at higher lipase
concentration than 0.75 mg mL−1 may be due to the fact that this
support has a finite number of binding sites and the amount of
lipase for immobilization was overloading in this instance. Reasons
affecting protein loading to decrease at higher protein concentra-
tions could also include effects of steric hindrances and diffusional
limitations, typical for concentrated enzyme solutions. Specific
activity of immobilized enzyme appeared to follow the oppo-
site trend, with maximal value reached at 1.98 IU mg−1 for the
Sepabeads® EC-EP sample having the lowest amount of protein
loading (6.9 mg g−1). Influences of diffusional limitations and steric
effects are notable in values of specific activities of CRL immobilized
on all three supports. As protein loading increased, these effects
were more pronounced and likely limited the amount of substrate
reaching active site, thus causing decrease in specific activity.
Successful immobilization on Sepabeads® EC-EP is probably the
consequence of high affinity of epoxy groups present on the poly-
mer surface towards nucleophilic groups of the enzyme, as well as
of simple, mild and effective immobilization procedure [13].
Immobilization outcome can also be influenced by type and
origin of the enzyme. In previous studies, Kunamneni et al. immobi-
lized recombinant laccase expressed in Aspergillus on Sepabeads®
EC-EP3 using enzyme/support ratio 100 mg g−1 and reported
enzyme loading efficiency to be 32.6% [11]. However, Hilterhaus
et al. immobilized C. antarctica lipase on Sepabeads® EC-EP using
enzyme/support ratio of 138.7 mg g−1 and immobilization process
succeeded with enzyme loading efficiency of 85% [15]. Differences
in enzyme loading efficiencies might be affected by the content
and availability of the groups reacting with the support in differ-
ent enzymes. Also, different degree of enzyme glycosilation may
influence enzyme-support interactions [11].
In order to obtain more information concerning the nature of
the lipase attachment to amino-supports, immobilized prepara-
tions were incubated in 10 mM sodium phosphate buffer, pH 7
containing 0.5% (v/v) Triton X-100 or 0.5–1.25 M NaCl. The oxida-
tion of CRL was also confirmed by FT-IR analysis (supplemental Fig.
1). The FT-IR spectrum of the lipase exhibited characteristic bands
of 1657.74 cm−1 (amide I), 1538.01 cm−1 (amide II) while the broad
absorption bands at 3400 cm−1 and 2934.52 cm−1 were apparently
caused by the symmetrical amine N–H vibration. The absorp-
tion bands at 1410.92 cm−1 (–CH symmetric bending vibration in
–CHOH–), 1122.38 cm−1 (–CO stretching vibration in –CH–O–CH)
and 889.442 cm−1 (–CN stretching vibration) could be characteris-
tic for enzyme’s carbohydrate moiety. It is evident that some major
changes occurred due to CRL oxidation by sodium periodate. The
band 1122.38 cm−1 shifted to 1127.59 cm−1 showing decreased
intensity, revealing major changes in primary hydroxyl group. In
addition, bands 1685.39 cm−1 (C O stretch vibration) appeared
indicating the appearance of free carbonyl groups. Two bands in
the range 2900–3010 cm−1 could be also related to free carbonyl
group ( C–H stretch) but they were moved to higher values since
The nature of the linkage between enzyme and amino-supports
was further highlighted using desorption study and the results are
presented in Table 1. It appeared that the lipase was tightly bound to
Sepabeads® EC-EP and was not removed by treatment with Triton
X-100 or NaCl (desorption yield was lower than 5%). This provided
evidence that most of the enzyme has been bound covalently on
this support rather than by hydrophobic or ionic interactions.
Lower than 5 and 10% of bound lipase was desorbed from
Sepabeads® EC-EP, and from other two types of amino-supports,
respectively, after incubation with 0.5% Triton X-100 for 60 min.
However, non-covalently bound lipase was easily lost after treat-
ment with salt solutions of high ionic strength, revealing that
the non-covalent binding was primary ionic. Overall, the results
emphasized that non-covalent binding of CRL occurred in addition
to covalent coupling of enzyme to amino-supports indicating the
importance of the removal of this non-covalently bound lipase after
covalent coupling.
In conclusion, among three supports and coupling chemistry
tested, lipase immobilized on Sepabeads® EC-EP is the immobi-
lization process of choice. This support is a unique material for
enzyme immobilization and the process of immobilization is sim-
ple, effective and very economical. The stable binding achieved
in this case is a useful characteristic for practical applications
of biocatalysts that would allow enzyme recycling with minimal
leaching.
Since immobilized enzyme obtained using protein concentra-
tion 0.75 mg mL−1 showed highest activity and protein loading, still
retaining high specific activity, it was selected for geranyl butyrate
synthesis optimization.
3.2. Optimization of geranyl butyrate synthesis in the batch
system by statistical approach
Reaction parameters were optimized using response surface
methodology based on 5-level-5-factor central composite rotat-
able design (CCRD) requiring 32 experiments. The variables studied
in the batch process of geranyl butyrate synthesis were: water
concentration (1.0–5.0%, v/v), temperature (25–45 ◦C), enzyme
concentration during immobilization, indicating enzyme loading
(2.0–4.0 g L−1), initial substrate molar ratio (acid/alcohol: from 1:2
to 5:2) and time of molecular sieves introduction for removal
of excess water (0–32 h). The range and levels of the variables
were chosen based on our previous study [21] where tempera-
ture was found to have a significant negative influence on ester
yield, masking the effects of all other tested variables. Thus, in
the new experimental design used in this paper, the range for
temperature was decreased (25–45 ◦C) while for other variables
remained the same. A higher temperature would not increase
molar conversion, but could cause denaturation of the immobilized
lipase.
The experimental data obtained for 32 experiments of the sta-
tistical design are presented in supplemental Table 2. Significances
of the factors were estimated based on the t-test and p-value statis-
tical parameters. The effects of temperature, water concentration
and temperature-substrate molar ratio interaction seem to be sig-
nificant (p < 0.05), as well as three quadratic coefficients (p < 0.05).
The fit of the model is verified by Fisher test (F) for 5% level of signif-
icance. Since the experimental F value (−3.31) was lower compared
to theoretical (4.58), it was concluded that the model is adequate
for description of this reaction system. After the introduction of

J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
55
Table 1
Comparison of the methods applied for covalent immobilization of CRL on three different supports using two methods.
Support
C0 (mg mL−1
)
Pg (mg g−1
)
YP (%)
A (IU g−1
)
SAIE (IU mg−1
)
YA (%)
YD (%), for 0.5%
Triton X-100
YD (%), for
0.50 M NaCl
YD (%), for
1.00 M NaCl
YD (%), for
1.25 M
NaCl
0.25
0.50
0.75
0.90
6.9
11.4
19.2
17.5
39.3
32.6
36.5
28.6
13.7
19.4
26.2
17.2
1.98
1.70
1.36
0.98
90.6
77.35
62.1
44.6
3.2
4.3
Minor
Minor
Minor
Minor
Minor
Sepabeads® EC-EP
Sepabeads® EC-HA
Minor
0.25
0.50
0.75
1.25
1.40
1.5
7.1
9.4
16.0
16.1
17.4
40.4
36.0
36.6
33.4
4.94
4.78
5.76
5.76
5.58
3.29
0.67
0.61
0.36
0.35
–
76.8
69.3
40.8
39.4
9.3
8.7
12.4
12.0
13.5
13.0
24.2
24.0
0.25
0.50
0.75
1.25
1.50
2.6
6.0
10.1
16.2
20.0
29.7
34.4
38.4
37.0
38.0
3.9
3.0
1.7
1.3
1.1
1.50
0.50
0.17
0.08
0.05
–
56.6
19.2
9.1
9.9
9.3
10.5
12.5
15.4
17.8
33.7
30.2
Purolite® A-109
6.3
C0, the protein concentration of the initial immobilization solution (mg mL−1); Pg, the protein loading (mg protein/g of the support); YP (%), the protein loading efficiency; A,
the hydrolytic activity of the immobilized lipase (IU g−1); SAIE, the specific activity of the immobilized enzyme lipase (IU mg−1); YA (%), the activity immobilization yield; YD
(%), the desorption yield.
calculated regression coefficients into second-order polynomial
equation, following model was obtained:
Y = 86.28 + 6.05X1 − 15.8X2 − 4.64X12
− 4.84X22 − 10.6X2X4 + 4.34X32
(9)
Contour plots obtained using the model are given in Fig. 2.
Temperature appears to have the most significant effect
on ester synthesis, although there are other factors, such as
water concentration, substrate molar ratio, enzyme content, and
temperature–substrate molar ratio interaction which should also
be considered.
Significant negative interactive effect of temperature and sub-
strate molar ratio, indicate that high molar conversion could
be expected at low temperatures and high acid/alcohol molar
ratios (Fig. 2a). Molar conversion increased as reaction tempera-
ture decreased, reaching maximum at 25–30 ◦C. Several previous
researches also indicated lower reaction temperatures as favorable
for production of butyric acid esters catalyzed by lipases. Shieh
et al. reported optimal temperature of 35 ◦C for geranyl butyrate
synthesis in transesterification reaction catalyzed with CRL [20].
Also, Hari Krishna et al. reported optimal temperature of 30 ◦C for
isoamyl butyrate synthesis in esterification catalyzed by Lipozyme
IM-20 [35]. Further on, it is apparent form Fig. 2a that the increase
in acid/alcohol molar ratio had a positive influence on ester synthe-
sis. Similar conclusions were made by Pereira et al., in the study of
butyl butyrate synthesis catalyzed by CRL immobilized on porous
chitosan grains. Optimal initial molar ratio of butanol/butyric acid
was 1:1.5 [36]. In esterification where butyric acid is present as
acyl donor, usually moderate acid excess favors synthesis from the
aspect of reaction equilibrium, thus having a beneficial effect on
molar conversion. Furthermore, excess acid in the esterification is
often necessary to prevent competition of two acyl donors, acid
and ester formed during the reaction [37]. Also, CRL shows affin-
ity towards small chain fatty acids, but high acid concentrations
eventually inhibit lipase activity. Polar butyric acid molecules tend
to concentrate in the water layer around the enzyme changing its
pH, and thus causing enzyme inactivation. Excess geraniol proved
to have an adverse effect on molar conversion, as also found by
Chatterjee et al. in the research regarding production of geranyl
capronate, geranyl caprylate, geranyl caprate and geranyl laurate
catalyzed by lipase from R. miehei [38].
Fig. 2. Contour plots for molar conversion as a function of temperature and initial
substrate molar ratio (a) and for the molar conversion as a function of tempera-
ture and water concentration (b). Reaction conditions: reaction time 48 h, limiting
substrate concentration 0.25 M.

56
J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
Fig. 4. Effect of temperature on geranyl butyrate synthesis in a FBR. Reaction
conditions: substrate concentration 0.1 M, substrate molar ratio 1:1, flow rate
3.70 mL min−1, water concentration 2% (v/v) and enzyme loading 76.7 mg g−1
.
Fig. 3. Application of Tanks-in-series model on experimental F and E curves. Flow
rate was set to 3.70 mL min−1. Simbols stand for experimental data while lines stand
for model predictions. In case of E curve, best fitting between experimental and
predicted data stand for N = 4 (according to criteria of lowest STD).
identical with residence time calculated using reactor volume and
flow rate, temp, and was 3.1 min, indicating absence of dead regions
within the reactor volume.
The effect of temperature and water concentration is presented
in Fig. 2b.
3.3.2. Optimization of esterification conditions in FBR
3.3.2.1. Effects of water content and temperature. Preliminary study
revealed increase in molar conversion as the initial water con-
centration in the reaction mixture rose from 0 to 2% (v/v) (data
not shown). However, a sharp decrease in molar conversion was
observed if water content further increased from 2 to 6% (v/v). Also,
the effect of temperature (30, 35 and 45 ◦C) on geranyl butyrate
production in FBR was investigated using fixed initial acid/alcohol
molar ratio at 1:1, substrate concentration at 0.1 M, and flow rate
at 3.70 mL min−1. Time course of molar conversion at different
temperatures is shown in Fig. 4. Although the highest conversion
was achieved at 35 ◦C, initial reaction rate was higher at 45 ◦C,
revealing that the biocatalyst was inactivated when subjected to
higher temperature for longer period under non-aqueous condi-
tions. Therefore, control of water and temperature at optimal values
(2%, v/v, 35 ◦C) in the system is important to optimize esterification
in FBR.
Ester production is represented by a contour plot with a max-
imum at a low temperature of around 25–30 ◦C for a water
concentration of 3.6% (v/v). Low water content is necessary to retain
enzyme catalytic activity by keeping the integrity of the molecule
in active conformation. However, exceeding the critical amount of
water, the thickness of water layer around the enzyme is increasing
causing increase in enzyme flexibility and eventually denaturation
[39]. Excess water would also favor the reverse hydrolytic reaction.
Data obtained by surface response analysis were used as refer-
ence to further optimize geranyl butyrate synthesis in a FBR.
3.3. Esterification in a FBR
The majority of studies concerning ester synthesis focused on
stirred tank and packed bed reactors. Fluidized-bed reactors could
present a number of advantages such as improved heat and mass
transfer, optimal liquid mixing and absence of plugging. Also, pres-
sure drop across the bed appears to be much lower than that in the
corresponding packed bed reactor. Thus, this study aimed to further
test fluidization properties of biocatalyst beads (CRL-Sepabeads®
EC-EP system, 150–300 ␮m) and their ability to catalyze geranyl
butyrate synthesis in a FBR.
3.3.2.2. Effect of substrate concentration. Enzymatic reactions,
especially esterifications, are often challenged by high substrate
concentrations due to the inhibitory effects of acid, alcohol or both
of the substrates. This influence was not part of the initial study
in the batch system, but may have a strong impact on the reaction
course. Therefore, we compared the influence of different substrate
concentrations, namely, 0.05 M, 0.1 M and 0.25 M on molar conver-
sion for stoichiometric amounts of reactants. Results are given in
Fig. 5a.
Increase in substrate concentration caused a decrease of molar
conversion. As the substrate concentration increases, inhibitory
effects of the substrates might become more pronounced. Observed
effect could also be partially attributed to intensified water forma-
tion in the presence of higher substrate concentrations. However,
efficient water removal in FBR still remains to be established. The
highest molar conversion (67.3%) was obtained for the lowest sub-
strate concentration 0.05 M, but the volumetric productivity (P
defined by Eq. (8)) was 38% lower than the corresponding value
obtained for the highest substrate concentration 0.25 M. Thus, to
make a correct comparison, the volumetric productivity has been
calculated for each substrate concentration. The highest P value
(7.1 mmol L−1 h−1) is obtained for the system with substrate con-
centration 0.1 M, which was selected as optimal. Lowest P value
(4.2 mmol L−1 h−1) is calculated for initial substrate concentration
0.05 M, since in this case substrate concentration is unnecessarily
3.3.1. Hydrodynamic characteristic of the FBR system
For continuous reactors, residence time distribution is an essen-
tial tool to assess fluid velocity profile, which can be used to describe
flow regime in the reactor. Since flow regime affects reactor per-
formance, its description may enable better process control. Flow
pattern studies conducted previously by Saponjic et al. revealed
nearly plug flow for the FBR studies of amyl caprylate synthesis,
using immobilized CRL in the similar FBR system [22]. Due to dif-
ferences in scale-up strategy, size of particle layer used to fill reactor
vessel for geranyl butyrate synthesis was around 40% higher com-
pared to the amount used for amyl caprylate synthesis. Thus, flow
pattern analysis was conducted once more.
Using Tanks-in-series model (Eqs. (6) and (7)), flow pattern in
this system can be described by the cascade of four ideal continuous
flow stirred tanks (Fig. 3). This number of tanks describes the flow
through tubular fluidized bed reactor as nearly plug flow with mod-
erately low deviations, due to a certain degree of axial dispersion.
Mean residence time, determined by the model, for N = 4, tmod, is

J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
57
Fig. 6. Effect of flow rate on geranyl butyrate synthesis in a FBR. Reaction conditions:
substrate molar ratio 1:1, substrate concentration 0.1 M, water concentration 2%
(v/v), enzyme loading 76.7 mg g−1 and temperature 35 ◦C.
It seems that the flow rate strongly influenced ester production
by immobilized CRL in FBR. According to P value, which was highest
at flow rate 10.0 mL min−1 reaching the value of 26.7 mmol L−1 h−1
,
this flow rate is selected as optimal. It appears that the esterifi-
cation reaction in FBR was mass transfer limited and thus can be
improved by increasing flow rate. At high flow rates, mass trans-
fer is enhanced and diffusional limitations reduced. These reasons
affect molar conversion and could be used to explain obtained
results. However, one adverse effect was also observed, specifi-
cally, increased flow rate led to a reduction of contact time between
the enzyme surface and substrates; the reaction was incomplete
causing reduction in maximal molar conversion [22,40]. This effect
seems to be predominant at lower flow rates. Thus, if flow rate
is set to 2.67 mL min−1, P value reaches only 6.1 mmol L−1 h−1
,
while at flow rate 3.70 mL min−1, P value is even less and reaches
5.8 mmol L−1 h−1. Results obtained by other authors suggest lower
flow rates and longer residence times as more productive due to
prolonged contact between enzyme and substrates [40,41]
Fig. 5. Effect of substrate concentration (a) and initial substrate molar ratio (b)
on geranyl butyrate synthesis in FBR. Reaction conditions: substrate molar
a
ratio 1:1, flow rate 3.70 mL min−1, water concentration 2% (v/v), enzyme loading
76.7 mg g−1, temperature 35 ◦C (a) and, limiting substrate concentration 0.1 M, flow
rate 3.70 mL min−1, water concentration 2% (v/v), enzyme loading 76.7 mg g−1 and
temperature 35 ◦C (b).
3.3.3. Operational stability of the immobilized CRL
Operational stability was analyzed in a semi-continuous man-
ner, with the biocatalyst regeneration after each esterification
cycle, as previously done in this research group [22]. Trend in rel-
ative molar conversions for eight successive cycles conducted in a
batch system is given in Fig. 7.
low, enzyme/substrate ratio very high, indicating that amount of
enzyme in the reactor was not yet used with its full catalytic
capacity. Therefore, this system was characterized as economically
unfeasible even though, from Fig. 5a, it produced highest molar
conversion (%).
Effect of substrate molar ratio on esterification in FBR (Fig. 5b)
followed the same trend as determined by statistical analysis for
batch conditions, providing maximal molar conversion of 78.9%
(P = 7.9 mmol L−1 h−1) obtained within 10 h by introducing excess
butyric acid.
During seven complete reaction cycles, immobilized lipase
retained high level of stability reflected in high relative molar
3.3.2.3. Effect of flow rate. In continuous type reactors, flow rate
proved to be significant for reactions catalyzed by immobilized
enzymes, since it has reciprocal relationship with residence time.
Usually, longer residence time enhances the reaction by enabling
longer enzyme–substrate contact, which can enhance conversion.
In this work, we used three different flow rates (residence
times): 2.67 mL min−1 (4.3 min), 3.70 mL min−1 (3.1 min) and
10.0 mL min−1 (1.1 min). Flow rate 2.67 mL min−1 is a minimal flow
rate that fluidized layer of immobilized enzyme particles, while
10.0 mL min−1 is a maximal flow rate which caused maximal layer
expansion without particle leakage. It was shown that biocatalyst
beads can be used to create stable expanded beds at all flow rates.
Determined relation between molar conversion and flow rates is
given in Fig. 6.
Fig. 7. Relative operational stability of lipase immobilized on Sepabeads® EC-
EP. Molar conversion is presented as a relative value referred to initial reaction
cycle. Reaction conditions: substrate concentration 0.25 M, substrate molar ratio
acid/alcohol 2:1, enzyme loading 76.7 mg g−1 and temperature 30 ◦C.

58
J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
conversion. By the end of the eighth cycle, approximately 50%
decrease in molar conversion was observed indicating loss of
operational stability (the operational enzyme half-life was 368 h).
Operational stability achieved in the batch system corresponds to
the results published previously using lipases in flavor ester syn-
thesis. Claon et al. reported five and 10 cycles operational stability
of the commercial enzyme preparations SP382 and SP435 (indus-
trially immobilized lipase from C. antarctica) used in a batch geranyl
acetate synthesis [5]. In a similar way, Yee et al. investigated oper-
ational stability of lipase from Pseudomonas sp. immobilized on
Duolite and PVP in the reaction of citronellyl butyrate and ger-
anyl capronate synthesis. The sudden drop in enzyme activity was
registered after the fifth cycle of synthesis [42].
Generally, stability might be affected by the method of catalyst
regeneration, immobilization mechanism or nature of substrates.
Immobilization mechanism would influence the strength of the
enzyme-support bond. In this sense, covalent immobilization pro-
vides strong enzyme-support bond preventing enzyme leakage
from the reactor vessel [16]. Immobilized enzyme regeneration
was done by thorough rinsing with the solvent followed by
overnight drying at room temperature in a dessicator containing
silica gel. This regeneration could be insufficient to completely
remove adsorbed polar substrate resulting in accumulation of acid
molecules on the carrier surface causing diffusional limitations
to the substrates migrating from the reaction medium towards
enzyme, as proposed by Marty et al. [43] and verified by Saponjic
et al. [22]. Therefore, highly effective catalyst regeneration remains
yet to be established.
but the productivity in FBR was more than 5-fold higher compared
to that obtained in the batch system.
Generally, reaction conditions in a batch reactor and FBR fol-
lowed the same trend with minor differences originating from
different reactor set-ups and consequently different transfer phe-
nomena. Both, however, enabled highly productive synthesis of
geranyl butyrate. Covalently immobilized CRL applied in FBR syn-
thesis proved to be robust and versatile way for aroma ester
production even in the medium posing additional limitations due to
substrate polarity and unfavorable substrate partitioning between
organic phase and microaqueous layer.
Results of the conducted study imply that this system has high
potential for further improvement and scale-up as well as for
application in continuous enzymatic synthesis of other esters or
products of enzymatic conversions. Besides of enzyme stabiliza-
tion achieved by immobilization, conditions in the reactor provide
improved mass transfer necessary for the reaction advancement.
Acknowledgements
The authors are grateful to Resindion S.R.L. (Mitsubishi Chemical
Corporation, Milan, Italy) for donation of Sepabeads® EC-EP and
Sepabeads® EC-HA carriers. Also, the authors are grateful to the
Ministry of Education and Science of the Republic of Serbia (Project
No. III 46010) for the financial support during this research which
was conducted at the Department of Biochemical Engineering and
Biotechnology, Faculty of Technology and Metallurgy, University of
Belgrade, Belgrade, from 2006 to 2009.
3.3.4. Comparison of the batch and FBR systems for geranyl
butyrate synthesis
Appendix A. Supplementary data
To compare batch system with FBR system, when assayed at
their optimal conditions, the volumetric productivity was calcu-
lated for each one. It was shown that in the batch system, a rather
high molar conversion >99.9% can be obtained. However, a draw-
back of this system was low reaction rate thus the conversion was
achieved after 48 h, corresponding to the volumetric productivity
of 5.2 mmol L−1 h−1. The kinetics in the FBR system seems to have a
better profile compared to batch system since the highest conver-
sion of 78.9% was achieved in 10 h, corresponding to the volumetric
productivity of 7.9 mmol L−1 h−1. At flow rate of 10 mL min−1, the
molar conversion of 53.3% may have been reached after 2 h reac-
tion (volumetric productivity 26.7 mmol L−1 h−1), indicating, as
expected, improved reaction kinetics compared to the batch sys-
tem. Observed molar conversion is possibly still susceptible to
further increase by implementation of effective control of water
concentration, and by better understanding of transfer phenomena
inside the reactor zone.
Supplementary data associated with this article can be found, in
the online version, at doi:10.1016/j.molcatb.2011.11.009.
References
[1] C. Akoh, G.-C. Lee, J.-F. Shaw, Lipids 39 (2004) 513–526.
[2] N. Gandhi, Journal of the American Oil Chemists Society 74 (1997) 621–634.
[3] D. Bezbradica, D. Mijin, S. Siler-Marinkovic, Z. Knezevic, Journal of Molecular
Catalysis B: Enzymatic 45 (2007) 97–101.
[4] M. Karra-Chaabouni, S. Pulvin, D. Touraud, D. Thomas, Biotechnology Letters
18 (1996) 1083–1088.
[5] P. Claon, C. Akoh, Journal of the American Oil Chemists Society 71 (1994)
575–578.
[6] Y. Ikeda, Y. Kurokawa, Journal of the American Oil Chemists Society 78 (2001)
1099–1103.
[7] B. Gillies, H. Yamazaki, D.W. Armstrong, Biotechnology Letters
709–714.
9 (1987)
[8] M.G. Carneiro-da-Cunha, J.M.S. Rocha, F.A.P. Garcia, M.H. Gil, Biotechnology
Techniques 13 (1999) 403–409.
[9] C. Mateo, J.M. Palomo, G. Fernandez-Lorente, J.M. Guisan, R. Fernandez-
Lafuente, Enzyme and Microbial Technology 40 (2007) 1451–1463.
[10] S.R.L. Resindion, Sepabeads® EC series-enzyme carriers (Instruction manual),
M.C. Corporation, Milan, Italy, 2006.
4. Conclusions
[11] A. Kunamneni, I. Ghazi, S. Camarero, A. Ballesteros, F.J. Plou, M. Alcalde, Process
Biochemistry 43 (2008) 169–178.
A process to obtain geranyl butyrate has been developed using
batch and FBR systems with CRL immobilized on Sepabeads® EC-
EP by covalent binding. Batch geranyl butyrate synthesis catalyzed
by immobilized CRL was efficiently optimized using response sur-
face methodology. Derived optimal conditions were temperature
of 25–30 ◦C, water concentration of 3.6% (v/v) and substrate molar
ratio acid/alcohol 2.5. Enzyme concentration and time of molecu-
lar sieves addition did not significantly influence molar conversion.
Preliminary results in repeated batch esterification showed an
acceptable stability of the immobilized enzyme.
[12] C. Mateo, V. Grazu, B.C. Pessela, T. Montes, J.M. Palomo, R. Torres, F. Lopez-
Gallego, R. Fernandez-Lafuente, J.M. Guisan, Biochemical Society Transactions
35 (2007) 1593–1601.
[13] C. Mateo, V. Grazu, J.M. Palomo, F. Lopez-Gallego, R. Fernandez-Lafuente, J.M.
Guisan, Nature Protocols 2 (2007) 1022–1033.
[14] C. Mateo, O. Abian, G. Fernández-Lorente, J. Pedroche, R. Fernández-Lafuente,
J.M. Guisan, Biotechnology Progress 18 (2002) 629–634.
[15] L. Hilterhaus, B. Minow, J. Müller, M. Berheide, H. Quitmann, M. Katzer, O. Thum,
G. Antranikian, A. Zeng, A. Liese, Bioprocess and Biosystems Engineering 31
(2008) 163–171.
[16] M. Chaplin, Enzyme Technology, Faculty of Engineering, Science and the Built
Environment, South Bank University, London, UK, 2004.
[17] S.-W. Chang, J.-F. Shaw, C.-K. Yang, C.-J. Shieh, Process Biochemistry 42 (2007)
1362–1366.
[18] F. Molinari, G. Marianelli, F. Aragozzini, Applied Microbiology and Biotechnol-
ogy 43 (1995) 967–973.
[19] S.S. Kanwar, S. Gehlot, M.L. Verma, R. Gupta, Y. Kumar, G.S. Chauhan, Journal of
Applied Polymer Science 110 (2008) 2681–2692.
Geranyl butyrate synthesis in the FBR was optimized at tem-
perature 35 ◦C, substrate concentration 0.1 M, acid/alcohol ratio 2.0
and flow rate 10.0 mL min−1. When assayed at their optimal condi-
tions, higher molar conversion was achieved in the batch system,

J.J. Damnjanovi´c et al. / Journal of Molecular Catalysis B: Enzymatic 75 (2012) 50–59
59
[20] C.-J. Shieh, C.C. Akoh, L.N. Yee, Biotechnology and Bioengineering 51 (1996)
371–374.
[33] J.B. Wheatley, D.E. Schmidt Jr., Journal of Chromatography
11–16.
[34] Z.D. Knezˇevic´-Jugovic´, D.I. Bezbradica, D.Z. Mijin, M.G. Antov, in: S.D. Minteer
(Ed.), Enzyme stabilization and immobilization: Methods and protocols,
Humana Press, New York, 2011, pp. 99–111.
[35] S. Krishna, B. Manohar, S. Divakar, N. Karanth, Journal of the American Oil
Chemists Society 76 (1999) 1483–1488.
[36] G.M.Z.E.B. Pereira, H.F. de Castro, Brazilian Journal of Chemical Engineering 20
(2003) 343–355.
A
644 (1993)
ˇ
ˇ
[21] Z.D. Knezˇevic´-Jugovic´, J.J. Damnjanovic´, D.I. Bezbradica, Z.D. Mijin, Chemical
Industry and Chemical Engineering Quarterly 14 (2008) 245–249.
[22] S. Saponjic, Z.D. Knezevic-Jugovic, D.I. Bezbradica, M.G. Zuza, O.A. Saied, N.
Boskovic-Vragolovic, D.Z. Mijin, Electronic Journal of Biotechnology 13 (2010).
[23] P.J. Halling, Enzyme and Microbial Technology 16 (1994) 178–206.
[24] R.H. Valivety, P.J. Halling, A.R. Macrae, Biochimica et Biophysica Acta (BBA):
Protein Structure and Molecular Enzymology 1118 (1992) 218–222.
[25] Z. Knezevic, N. Milosavic, D. Bezbradica, Z. Jakovljevic, R. Prodanovic, Biochem-
ical Engineering Journal 30 (2006) 269–278.
[37] E.B.P.H.F. de Castro, W.A. Anderson, Journal of the Brazilian Chemical Society 7
(1996) 219–224.
[26] B.M. Marion, Analytical Biochemistry 72 (1976) 248–254.
[27] R. Prodanovic´, S. Jovanovic´, Z. Vujcˇic´, Biotechnology Letters 23 (2001)
1171–1174.
[38] T. Chatterjee, D.K. Bhattacharyya, Biotechnology Letters 20 (1998) 865–868.
[39] G.D. Yadav, P.S. Lathi, Journal of Molecular Catalysis B: Enzymatic 27 (2004)
113–119.
[28] W.G. Hunter, G.E.P. Box, J.S. Hunter, Statistics for experimenters: An introduc-
tion to design, data analysis and model building, Wiley, New York, 1978.
[29] J.M. Walker, Protein Protocols Handbook, Humana Press Inc, New York, 1996.
[30] J. Nelson, Chemistry: The Central Science, Prentice-Hall, 1985.
[31] Z. Knezevic, G. Kukic, M. Vukovic, B. Bugarski, B. Obradovic, Process Biochem-
istry 39 (2004) 1377–1385.
[40] L.M. Esteban, d.M. Mun˜ío, A. Robles, E. Hita, M.J. Jiménez, P.A. González, B.
Camacho, E. Molina, Biochemical Engineering Journal 44 (2009) 271–279.
[41] A. Sánchez, F. Valero, J. Lafuente, C. Solà, Enzyme and Microbial Technology 27
(2000) 157–166.
[42] L. Yee, C. Akoh, R. Phillips, Journal of the American Oil Chemists Society 74
(1997) 255–260.
[32] J.M. Guisan, C. Mateo, G. Fernandez-Lorente, O. Abian, R. Fernandez-Lafuente,
Biomacromolecules 1 (2000) 739–745.
[43] A. Marty, V. Dossat, J.-S. Condoret, Biotechnology and Bioengineering 56 (1997)
232–237.