Covalent immobilization of organic solvent tolerant lipase on aluminum oxide pellets and its potential application in esterification reaction

Article abstract of DOI:10.1016/j.molcatb.2012.10.002

This study was carried out to covalently immobilize the partially purified lipase from Bacillus sp. DVL2 on glutaraldehyde-activated aluminum oxide pellets and subsequently use the immobilized enzyme for esterification of oleic acid and ethanol. The immobilization process parameters were optimized through response surface methodology. Under optimized conditions, maximum immobilization yield of the enzyme was 78.20%. The immobilized lipase could be reused for 5 consecutive cycles without any loss of enzyme activity. However, the enzyme showed 75% residual activity after 10th cycle. The stability of the immobilized enzyme towards pH, temperature and organic solvents was enhanced as compared with its soluble counterpart. Higher values of half-lives, D-values, enthalpy and free energy change after covalent immobilization implied an improvement in enzyme stability. The K_m and V_max values of the enzyme were enhanced after immobilization. The immobilized lipase was found to be most stable in DMSO followed by toluene, hexane and xylene, exhibiting more than 90% residual activity in these solvents after 24 h of incubation. The immobilized lipase was more efficient in catalyzing the esterification between oleic acid and ethanol in hexane. The formation of ethyl oleate was confirmed by TLC and ¹H NMR spectroscopy.

Full text of DOI:10.1016/j.molcatb.2012.10.002

Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
Contents lists available at SciVerse ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Covalent immobilization of organic solvent tolerant lipase on aluminum oxide
pellets and its potential application in esterification reaction
Davender Kumara, Sushil Nagara, Indu Bhushanc, Lalit Kumara, Rajinder Parshadb,
Vijay Kumar Guptaa,∗
a Department of Biochemistry, Kurukshetra University, Kurukshetra 136119, India
b Indian Institute of Integrative Medicine, Jammu 180001, India
c Shri Mata Vaishno Devi University, Katra, Jammu, India
a r t i c l e i n f o
a b s t r a c t
Article history:
This study was carried out to covalently immobilize the partially purified lipase from Bacillus sp. DVL2
on glutaraldehyde-activated aluminum oxide pellets and subsequently use the immobilized enzyme
for esterification of oleic acid and ethanol. The immobilization process parameters were optimized
through response surface methodology. Under optimized conditions, maximum immobilization yield
of the enzyme was 78.20%. The immobilized lipase could be reused for 5 consecutive cycles without any
loss of enzyme activity. However, the enzyme showed 75% residual activity after 10th cycle. The stability
of the immobilized enzyme towards pH, temperature and organic solvents was enhanced as compared
with its soluble counterpart. Higher values of half-lives, D-values, enthalpy and free energy change after
covalent immobilization implied an improvement in enzyme stability. The Km and Vmax values of the
enzyme were enhanced after immobilization. The immobilized lipase was found to be most stable in
DMSO followed by toluene, hexane and xylene, exhibiting more than 90% residual activity in these sol-
vents after 24 h of incubation. The immobilized lipase was more efficient in catalyzing the esterification
between oleic acid and ethanol in hexane. The formation of ethyl oleate was confirmed by TLC and 1H
NMR spectroscopy.
Received 30 March 2012
Received in revised form
21 September 2012
Accepted 5 October 2012
Available online 13 October 2012
Keywords:
Covalent immobilization
Response surface methodology
Lipase
Bacillus sp. DVL2
Aluminum oxide pellets
© 2012 Elsevier B.V. All rights reserved.
1. Introduction
activities, the high yields possible, ease of genetic manipulation,
regular supply due to absence of seasonal fluctuations and easy
Lipase (triacylglycerol acylhydrolase, EC 3.1.1.3) belongs to a
group of enzymes whose biological function is to catalyze the
hydrolysis of triacylglycerols into diacylglycerols, monoacylglyc-
erols, free fatty acids (FFA) and glycerol at an oil–water interface
[1]. In addition to their natural function of hydrolyzing carboxylic
ester bonds, lipases can catalyze esterification, interesterification,
and transesterification reactions in non-aqueous media [2–4].
These reactions usually proceed with high chemo-, regio- and/or
enantioselectivity, making lipases an important group of biocata-
lysts [5]. Lipases are highly diversified in enzymatic properties and
substrate specificity. Lipases have emerged as key enzymes which
find usage in food, paper, textile and detergent industries, waste
water treatment, production of fine chemicals, pharmaceuticals,
cosmetics and so on [6–10]. Though lipases are widely distributed
among microorganisms, animals and plants yet, microbial lipases
are receiving much more attention in industry mainly because
of the availability of a wide range of hydrolytic and synthetic
cultivation of microbes on inexpensive media [11–13].
Lipases have been used as biocatalyst in esterifica-
tion/transesterification reactions [14]. The biosynthesis of esters
is of commercial interest because of the increasing popularity
and demand for natural products amongst consumer. A range of
fatty acid esters is now being produced commercially using the
immobilized lipase in non-aqueous solvents. Ethyl oleate esters
have been reported to possess applications in several industrial
fields such as food, aromatics, cosmetics, detergents, flavors
and pharmaceuticals. They may also be used as plasticizers and
lubricants, biological additives and hydraulic fluids [15].
The application of lipase has been deterred by the high cost
of enzymes, contamination of products with residual protein and
their low storage, operational, and thermal stabilities. Immobi-
lization of lipase might be the best solution to overcome these
limitations. Immobilization of lipases on water insoluble carriers,
helps in their economic reuse and in the development of continu-
ous bioprocesses. Immobilization also facilitates the separation of
products and improves lipase properties such as thermostability
and activity in non-aqueous media. The choice of appropriate
support as well as method of immobilization is essential to make
∗
Corresponding author. Tel.: +91 9896045695.
E-mail address: vkgupta59@rediffmail.com (V.K. Gupta).
1381-1177/$ – see front matter © 2012 Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.molcatb.2012.10.002

52
D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
industrially important biocatalyst. Accordingly, various immobi-
lization methods for lipase have been developed including physical
adsorption, ionic binding, covalent attachment, entrapment and
microencapsulation [16–23]. Although physical adsorption on
solid supports is simple, inexpensive and easy to perform yet it
suffers from the limitation of enzyme leaching from the support.
Encapsulation is a mild and especially useful method for enzymes
which are easily deactivated, for instance by covalent immobiliza-
tion. Covalent immobilization, which involves multipoint covalent
attachment of an enzyme to a carrier either directly or through
a spacer, has the advantage of preventing enzyme leaching. The
immobilization of lipase has been carried out on different supports
including both hydrophobic (e.g. octyl-agarose, sol–gels and
octyl-sephabeads) and hydrophilic (e.g. celite, alumina, silica,
agarose and nylon-6) [17,24]. Nanostructured materials such as
carbon nano tubes [25] and electrospun nanofibres [26] were
also used as supports for immobilization. Recently, lipases have
been immobilized on poly (allyl glycidyl ether-co-ethylene glycol
dimethacrylate) polymer microsphere [27], hydrophobic/strong
cation-exchange functional silica particles [28] and magnetic silica
nanocomposite particles [29,30].
2.3. Assay of free lipase
The activity of free lipase was determined spectrophotometri-
cally using p-nitrophenyl palmitate (p-NPP) as substrate according
to the method of Nawani et al. [32] with some modifications. The
reaction mixture containing 0.3 mL of 0.05 M phosphate buffer (pH
8.0), 0.1 mL of 0.8 mM p-NPP and 0.1 mL of lipase was incubated at
37 ◦C for 10 min. The reaction was then terminated by adding 1 mL
ethanol. A control was run simultaneously, which contained the
same contents but the reaction was terminated prior to addition
of the enzyme. Absorbance of the resulting yellow colored product
was measured at 410 nm in a spectrophotometer. One International
Unit (IU) of lipase activity was defined as the amount of enzyme cat-
alyzing the release of 1 ␮mol of p-nitrophenol per min from p-NPP
under the standard assay conditions.
2.4. Estimation of protein content
Protein content in different enzyme preparations was estimated
by Lowry’s method [33].
The yield of the immobilized enzyme may be enhanced through
process optimization by using statistical approach, which is
easy, less laborious, quick and more accurate. Response surface
methodology (RSM), a collection of mathematical and statistical
techniques, is widely used to determine the effect of several vari-
ables at a time to optimize the biotechnological processes [31].
In this study, the strain Bacillus sp. DVL2 used for lipase pro-
duction was isolated in our laboratory. The lipase produced by this
strain was optimally active at 37 ◦C and pH 8.0. It showed stability
in various organic solvents. This lipase was covalently immobi-
lized on glutaraldehyde-activated aluminum oxide pellets and the
immobilized enzyme was examined for its properties. The above
support was selected for immobilization due to its stability at high
pH and temperature. As per our knowledge, no report is available
for the immobilization of lipase on aluminum oxide pellets through
covalent attachment. The immobilization process parameters were
statistically optimized to maximize the immobilization yield. The
potential of the immobilized lipase in esterification of oleic acid
and ethanol to ethyl oleate was also investigated.
2.5. Partial purification of enzyme
All the steps of enzyme purification were performed at 4 ◦C.
2.5.1. Ammonium sulphate precipitation
To 500 ml of culture supernatant, ammonium sulphate was
added (20–70% saturation) and the protein was allowed to pre-
cipitate by keeping it overnight in a refrigerator. The protein
precipitates were collected by centrifugation at 12,000 × g for
20 min at 4 ◦C and dissolved in a small volume of 0.05 M phosphate
buffer (pH 8.0).
2.5.2. Hydrophobic interaction chromatography
The concentrated ammonium sulphate fraction was applied on
to a column of Phenyl-sepharose CL-4B (Sigma) pre-equilibrated
with 1.0 M ammonium sulphate dissolved in 0.05 M phosphate
buffer (pH 7.0). The bound lipase was eluted by applying a negative
linear gradient of 1.0–0 M ammonium sulphate in 0.05 M phos-
phate buffer (pH 7.0). The fractions were collected and analyzed
for lipase activity and protein content. The fractions containing
high activity of lipase were pooled and concentrated using a 10 kDa
cut-off membrane.
2. Materials and methods
Aluminum oxide pellets (3 mm) and p-Nitrophenyl palmitate
(p-NPP) were purchased from Sigma–Aldrich Chemical Company
(St. Louis, USA) and all the other chemicals were of analytical grade.
2.6. Immobilization of lipase on aluminum oxide pellets
2.1. Microorganism
Aluminum oxide pellets were employed as support for immo-
bilization of partially purified lipase through covalent attachment.
The pellets were activated by dipping in 2% aqueous glutaralde-
hyde solution for 1 h at room temperature followed by addition
of the enzyme. The coupling of enzyme with the glutaraldehyde-
activated pellets was allowed to occur at room temperature for
90 min. The pellets were washed with distilled water at each step
to remove the excess of glutaraldehyde and the unbound enzyme.
The enzyme activity was determined in the supernatant as well
as in the enzyme bound pellets. Immobilization yield (IY %) was
calculated in the following manner:
The microorganism Bacillus sp. DVL2 was isolated from dairy
industry waste collected from Karnal district of Haryana (India) and
identified by morphological, biochemical and16 S rRNA sequenc-
ing. The 16S rDNA sequence of 1406 bp was generated and
submitted to NCBI Gene Bank depository under the Accession No.
JX846495.
2.2. Lipase production
The culture (isolated Bacillus sp. DVL2) was grown under opti-
mal conditions for lipase production. The basal production medium
containing (g/L) peptone 5.0, yeast extract 5.0 and olive oil 1.0 was
autoclaved at 1.05 kg/cm2 for 20 min, inoculated with 3.0% (v/v)
of 18 h old inoculum and incubated in an Orbital shaker at 30 ◦C
under agitation of 200 rpm for 24 h. The contents of the flasks were
then centrifuged at 10,000 × g for 20 min at 4 ◦C to remove the cells.
The clear supernatant containing the lipase was used for its partial
purification and subsequent immobilization.
Immobilization yield(%)
Total activity recovered on pellets
Total activity offered for immobilization
=
× 100
The observed activity of the immobilized enzyme refers to
the activity of the enzyme immobilized on aluminum oxide
pellets whereas total activity immobilized on pellets refers to

D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
53
Table 1
Experimental range of each selected variable for lipase immobilization.
2.11. Effect of temperature on free and the immobilized lipase
The temperature optima of soluble and the immobilized lipase
were investigated by determining the enzyme activity at different
temperatures (25–45 ◦C). The relative activity (%) at each temper-
ature was calculated with reference to the enzyme activity at the
optimum temperature (which was taken as 100%). The activation
energy (Ea) of catalysis for free and the immobilized lipase was
determined from the slope of the Arrhenius plot [log V (logarithm
of % relative activity) versus reciprocal of absolute temperature in
Kelvin (1000/T)], using the following equation:
Factor Name
Unit
Low level High
(−) level (+)
Mean SD
(0)
A
B
C
D
No. of beads
Glutaraldehyde
Enzyme dose
numeric
(%, v/v)
mL
4.0
8.0
6.0
1.79
2.0
6.0
4.0
0.04
1.79
0.02
0.02
0.06
Coupling period min
30.0
120.0
75.5
39.21
the difference in enzyme activity offered for immobilization and
activity recovered in the supernatant.
Ea
Slope =
R
2.7. Assay of the immobilized lipase
Thermal stability of soluble and the immobilized lipase was
studied by pre-incubating the enzyme at various temperatures
(25–60 ◦C) for 1 h followed by determination of activity using
standard assay at the optimum temperature. The residual activ-
ity (%) was calculated relative to the maximum enzyme activity
taken as 100%. Thermal stability of the immobilized lipase was also
investigated by pre-incubating it at temperatures ranging from 30
to 60 ◦C for different periods (30, 60, 90 and 120 min). The resid-
ual activity was calculated by taking the enzyme activity at 0 min
incubation as 100%.
Activity of the immobilized lipase was assayed by adding 0.1 mL
of 0.8 mM p-NPP and 0.4 mL of 0.05 M phosphate buffer (pH 8.0) to
the enzyme bound aluminum oxide pellets in an eppendorf tube.
After incubating the reaction mixture for 10 min at 37 ◦C, the pellets
were removed by transferring the contents of the reaction mix-
ture to another eppendorf tube. The reaction was then stopped
by adding 1 mL ethanol and intensity of the resulting yellow col-
ored product was read at 410 nm. The pellets were washed with
phosphate buffer and reused.
The first order thermal deactivation rate constants (kd) were
determined from the regression plot of log relative activity (%)
versus time (min). The activation energy (Ed) for lipase denatur-
ation was determined by a plot of log denaturation rate constants
(ln kd) versus reciprocal of the absolute temperature (K) using the
following equation:
2.8. Statistical optimization of immobilization
To examine the cumulative effect of immobilization parame-
ters response surface methodology was employed using a statistical
software package Design Expert 7.1.2, Stat-Ease, Inc. A 24 full fac-
torial central composite design (CCD) with 16 trials for factorial
design, 8 trials for axial point and 6 replicate trials at the central
point, leading to a set of 30 experiments was designed. The range
and levels of variables (−␣, −1, 0, 1, +␣) are presented in Table 1
and the experimental design is shown in Table 2. All the variables
were taken at a central value represented by “0”. The response value
from each experiment of CCD was the average of triplicates.
Ed
Slope =
R
The half-lives (t1/2) and D-values (decimal reduction time or
time required to pre-incubate the enzyme at a given temperature
to maintain 10% residual activity) of the immobilized lipase at each
temperature were determined from the following relationships:
ln
kd
2
2.9. Operational stability of the immobilized lipase
t1/2
=
To evaluate the recycling stability of the immobilized lipase,
enzyme bound pellets were incubated with buffer and p-NPP for
10 min at 37 ◦C followed by termination of the reaction according to
standard assay conditions. The bound enzyme was repeatedly used
to hydrolyze p-NPP up to 15 batch reactions. The residual activity
(%) was determined as:
ln 10
kd
D-value =
The changes in enthalpy (ꢀH◦, kJ mol−1), free energy (ꢀG◦,
kJ mol−1) and entropy (ꢀS◦, J mol−1 K−1) for thermal denaturation
of lipase were determined using the following equations:
enzyme activity in nth cycle
Residual activity =
× 100
ꢀH◦ = Ed − RT
enzyme activity in 1st cycle
ꢀ
ꢁ
kdh
kBT
ꢀG◦ = −RT ln
2.10. Effect of pH on free and the immobilized lipase
The effect of pH on soluble and the immobilized lipase was
studied by determining the enzyme activity using different buffers
(0.05 M) viz. sodium citrate (pH 4.0–6.0), sodium phosphate (pH
7.0–9.0) and glycine–NaOH (pH 10.0–11.0). The residual activity (%)
at each pH was calculated with reference to the enzyme activity at
the optimum pH (taken as 100%).
The pH stability of soluble as well as the immobilized lipase
was investigated by pre-incubating the enzyme in the above-
mentioned buffers of different pH for 2 h followed by measurement
of its activity under standard assay conditions. The residual activity
(%) at each pre-incubating pH was calculated relative to the max-
imum enzyme activity (taken as 100%). The results of the effect of
pH on the immobilized enzyme were compared with those of the
free enzyme.
ꢀH◦ − ꢀG◦
ꢀS◦
=
T
where T is the corresponding absolute temperature (K), R is
the gas constant (8.314 J mol−1 K−1),
h is the Planck’s con-
stant (11.04 × 10−36 J min), and kB is the Boltzmann constant
(1.38 × 10−23 J K−1).
The z-value (temperature rise necessary to reduce D-value by
one logarithmic cycle) was calculated from the slope of graph
between log D versus temperature (◦C) using the following equa-
tion:
1
Slope =
z

54
D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
Table 2
Experimental design (CCD) for lipase immobilization using RSM.
Sr. No.
A: no. of pellets
B: glutaraldehyde (%, v/v)
C: enzyme dose (mL)
D: coupling period (min)
% immobilization
1
2
3
4
5
6
7
8
4
8
4
8
4
8
4
8
4
8
4
8
4
8
4
8
2
10
6
6
6
6
6
6
6
2
2
6
6
2
2
6
6
2
2
6
6
2
2
6
6
4
4
0
8
4
4
4
4
4
0.02
0.02
0.02
0.02
0.06
0.06
0.06
0.06
0.02
0.02
0.02
0.02
0.06
0.06
0.06
0.06
0.04
0.04
0.04
0.04
0
30
30
30
30
30
30
30
30
120
120
120
120
120
120
120
120
75
75
75
75
75
26.60
36.32
32.30
40.17
18.58
40.86
21.30
46.01
39.70
55.02
51.44
62.30
28.10
46.10
30.20
59.20
16.20
78.20
19.30
31.20
15.40
29.14
18.10
40.79
60.72
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25–30
0.08
0.04
0.04
0.04
75
0
165
75
heating on hot plate. 1H NMR spectra were recorded with BUCKER
500 MHz NMR instrument. Chemical data for protons are reported
in parts per million (ppm, scale) downfield from tetramethylsilane
(TMS) and are referenced to the residual proton in the NMR solvent
(CDCl3: ı 7.26).
2.12. Effect of organic solvents on the activity of the immobilized
enzyme
To study the effect of organic solvents on the immobilized
lipase, the enzyme bound aluminum oxide beads were treated
with organic solvents viz. isopropanol, xylene, methanol, ethanol,
dimethylsulphoxide (DMSO), toluene, hexane and acetonitrile to
a final concentration of 25% (v/v) for 12, 24 and 36 h. The resid-
ual activity of the immobilized lipase was then determined under
standard assay conditions. The enzyme activity of the sample with-
out adding any organic solvent was taken as control (100%). The
stability of the immobilized lipase in various organic solvents was
also compared with that of free enzyme after an incubation of 24 h.
3. Results and discussion
The lipase produced by the isolate Bacillus sp. DVL2 under
submerged fermentation was partially purified through ammo-
nium sulphate fractionation (20–70%) and Hydrophobic Interaction
Chromatography on Phenyl-sepharose CL-4B column. The partially
purified enzyme (specific activity 1.5 IU/mg protein) was covalently
immobilized on glutaraldehyde-activated aluminum oxide pellets.
This support is preferred for immobilization as these pellets are
mechanically tough and resistant to high pH and temperature. In
addition, the conformation of pellets was maintained during mod-
erate shaking for esterification reaction. The parameters of the
immobilization process were statistically optimized using RSM to
enhance yield of the immobilized enzyme.
2.13. Determination of kinetic parameters
The apparent Michaelis–Menten constant (Km) and maximum
velocity (Vmax) of free and the immobilized enzyme were calculated
using the Lineweaver–Burk plot. The substrate concentration used
for enzyme assay ranged from 0.17 to 2.6 mM.
2.14. Application of the immobilized lipase in esterification of
ethanol with oleic acid
3.1. Optimization of lipase immobilization through RSM
Four process parameters, i.e. the number of pellets, glutaralde-
hyde concentration, volume of lipase and coupling time were
optimized using RSM. Tables 1 and 2 represent the range of com-
ponents and experimental design for RSM, respectively. Analysis
of the results obtained from RSM showed that immobilization
yield (IY) varied greatly from 15.4 to 78.2% (Table 2). These results
encouraged the use of statistical optimization of process parame-
ters. The responses of the CCD design were fitted with a polynomial
quadratic equation (Eq. (1)). The overall polynomial equation for
the immobilization (Y) of lipase was
Partially purified lipase immobilized on aluminum oxide pel-
lets was used as biocatalyst for the esterification of oleic acid and
ethanol in 1:1 (v/v) ratio in hexane. The reaction was carried out at
37 ◦C with shaking at 100 rpm for 4, 8, 12, 16, 20 and 24 h with heat
inactivated free enzyme as control. The ester content was quanti-
fied by using the alkalimetric method of titrating unreacted acid
with 0.1 N NaOH using phenolphthalein as an indicator. The per-
centage conversion in ester synthesis was based on the amount of
acid consumed [34]. Reusability of the immobilized lipase in the
above esterification reaction was also tested.
Y = +60.19 + 10.91 × A + 3.14 × B − 1.08 × C + 7.25 × D + 0.45
× A × B + 3.14 × A × C + 0.54 × A × D − 0.34 × B × C + 1.05
× B × D − 2.26 × C × D − 1.65 × A2 − 7.14 × B2 − 7.88
2.15. TLC and 1H NMR
Analytical TLC was performed using pre-coated silica gel 60
F254 MERCK TLC plates (20 cm × 20 cm). The spots were visualized
by immersing the plates in 10% H2SO4 in ethanol followed by
× C2 − 7.08 × D2
(1)

D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
55
Fig. 1. Response surface plots showing cumulative effect of two variables on the immobilization of lipase while keeping other variables at “0” level; (a) interaction between
factors A and B, (b) between factor C and A, (c) between factors C and B, (d) between factors D and B and (e) perturbation graph showing the effect of variables on immobilization
yield (where A: number of beads; B: glutaraldehyde concentration; C: enzyme dose and D: incubation period).
where A is the number of pellets, B is glutaraldehyde concentration,
C is enzyme dose and D is coupling time.
Statistical significance of the model equation was evaluated by
the F-test for analysis of variance (ANOVA), which showed that the
regression was statistically significant (Table 3). The model F value
for IY was 7.62 as shown by Fisher’s test, which indicated that the
model was significant. The value of “Prob > F” less than 0.05 was
desirable for a significant model. The P > F value of the model was
<0.0002; therefore, the model terms were also significant. It implied
that the model was statistically significant for the lipase immobi-
lization. There was only a 0.01% chance that a “Model F-Value” was
not significant which could occur due to noise. The determination

56
D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
Table 3
Analysis of variance (ANOVA) for response surface quadratic model for lipase immobilization.
Term
F-value
P-value (Prob > F)
Value
Model
A: no. of beads
7.62
39.34
3.27
0.39
0.0002
<0.0001
0.0907
0.5423
–
–
–
–
–
0.88
0.76
0.30
Significant
B: glutaraldehyde
C: enzyme dose
D: incubation period
R2
16.18
0.0011
–
–
–
–
–
–
–
–
–
–
–
–
–
–
–
–
Adjusted R2
Predicted R2
Adequate precision
Standard deviation
Mean
Coefficient of variation %
Predicted residual sum of squares (PRESS)
9.93
8.52
41.56
20.50
6172.09
coefficient (R2) of the model was 0.88 which showed the aptness
of the model. The “Adeq Precision” measures the signal to noise
ratio (a ratio of greater than 4 was desirable) and ratio of 9.93 of
the model indicated an adequate signal. Thus, the model was sig-
nificant for the process. A lower value of coefficient of variation (CV
20.50%) showed that the experiments conducted were precise and
reliable.
enzyme retained nearly 75% and 25% of its activity. So, the immo-
bilized enzyme could apparently be reused for 10 cycles without
substantial loss of activity. The decrease in enzyme activity during
repeated use might be due to inactivation of enzyme as suggested
earlier [35].
3.3. Effect of pH
The response plots (3D and contour curves) were analyzed to
determine interaction among the variables and the optimum value
of each component for maximum IY. Fig. 1a shows the cumulative
effect of two factors viz. number of beads and glutaraldehyde con-
centration (%) on IY at central level of the remaining two factors. It
showed clearly that IY increased proportionately when number of
beads was increased from −1 to +1 level at 0 level of glutaraldehyde.
The IY declined when glutaraldehyde concentration was increased
or decreased beyond the central level. Fig. 1b reveals the variation
of IY with incubation period and enzyme dose. The IY enhanced
with increase in incubation period and enzyme dose initially. Max-
imum IY was obtained close to mid of 0 and +1 level of incubation
period at 0 level of enzyme dose. The effect of incubation period
and number of beads on IY is shown in Fig. 1c. Incubation period at
mid of 0 and +1 level resulted in maximum IY. However, number
of beads resulted in maximum IY at +1 level. Both the factors viz.
incubation period and number of beads showed strong interaction
effect as indicated by a more elliptical contour plot. Fig. 1d analy-
sis reveals that the maximum immobilization yield was obtained
when both enzyme dose and glutaraldehyde concentration were
taken near ‘0ꢀ level. Umbrella shaped curve indicates strong interac-
tion between factors close to ‘0ꢀ level. Any change in concentration
of both the factors from the central level leads to decrease in IY.
As glutaraldehyde is the cross linking agent, so its concentration is
directly proportional to the cross linking of enzyme. The simulta-
neous effect of all the variables on the lipase immobilization yield
can be seen in the perturbation graph (Fig. 1e). The plot reveals
that the immobilization process is most sensitive to the number of
pellets while least influenced by the enzyme dose.
The pH optima of free and the immobilized lipase are shown in
Fig. 3. It was evident from the data that the free enzyme showed a
broad pH optimum (6.0–9.0) with maximum at pH 8.0. The immobi-
lized lipase also showed broad pH optimum in the range of 7.0–9.0
but it exhibited slightly higher activity at acidic and alkaline pH
values as compared to its soluble counterpart. It could be due to
effect of the microenvironment of the immobilized enzyme.
The pH stability data shown in Fig. 4 revealed that both free
and the immobilized lipase exhibited substantial stability in the
pH range of 6.0–10.0 upon pre-incubation for 2 h but the immobi-
lized form showed better stability at extreme pH values. The soluble
enzyme was completely inactive at pH 4.0 and exhibited 12.4%
activity at pH 11.0 while the immobilized enzyme showed 25% and
46.2% activity at pH 4.0 and 11.0, respectively. Ortega et al. [36]
also reported enhanced stability of lipase at acidic and alkaline pH
upon immobilization. The increase in stability upon immobiliza-
tion might be due to multipoint covalent linking between lipase
and support, which prevents lipase denaturation in acid or alkaline
environments as suggested by Yang et al. [37]. Similar results were
obtained with the immobilized Candida rugosa lipase on chitosan
by Hung et al. [38].
All the RSM graphs were analyzed in the similar way to deter-
mine the optimum value of each factor for maximum IY. The
maximum IY of 78.20% was obtained when 10 aluminum oxide
pellets were treated with 4% glutaraldehyde and subsequent incu-
bation with 0.04 mL of enzyme for 75 min.
3.2. Operational stability of the immobilized enzyme
Immobilized enzymes are preferred as they can be recycled,
resulting in lower production costs. The operational stability of the
lipase immobilized on glutaraldehyde-activated aluminum oxide
pellets was investigated for 15 consecutive cycles. There was no
loss in enzyme activity up to 5 consecutive cycles but thereafter,
a decrease was observed (Fig. 2). After 10th and 15th cycles, the
Fig. 2. The operational stability of covalently the immobilized lipase on
glutaraldehyde-activated aluminum oxide pellets for 15 cycles.

D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
57
after 1 h incubation at 60 ◦C while the immobilized lipase retained
25% of its activity (Fig. 5b). So, the immobilized lipase was
found to have better thermo-stability as compared to its soluble
form. It could be due to the formation of intermolecular cova-
lent linkages between the enzyme and support which impart
rigidity to the structure of the enzyme molecule [26,32]. Similar
to our finding, Dosanjh and Kaur [40], observed that temper-
ature stability of the immobilized enzyme was enhanced from
50 to 60 ◦C in comparison with the soluble enzyme from Bacil-
lus sp. Increased thermo-stability of lipase upon immobilization
was also reported by other researchers [26,36]. The restricted
interaction between the immobilized enzyme molecules could
also be responsible for retaining the enzyme activity at higher
temperatures [41].
The investigation of thermal stability of the immobilized
enzyme at different temperatures (30–60 ◦C) with respect to time
period revealed that the enzyme was nearly fully stable up to 40 ◦C
for 60 min but gradually lost its activity upon incubation at higher
temperatures (Fig. 5c). The immobilized enzyme showed 45.2, 22.1
and 12.0% residual activity on incubation for 120 min at 45, 50 and
55 ◦C, respectively.
Fig. 3. Effect of pH on the activity of free (solid circle) and the immobilized (hollow
circle) lipase. The lipase activity was measured at different pH values using buffers
viz. sodium citrate (pH 4.0–6.0), sodium phosphate (pH 7.0–9.0) and glycine–NaOH
(pH 10.0–11.0) at 0.05 M each under standard assay conditions.
The plots of log (% residual activity) versus time were linear indi-
cating the first order kinetics of immobilized enzyme (Fig. 5d). To
determine the thermodynamic parameters for thermal stability,
the activation energy (Ed) for thermal denaturation was deter-
mined from slope of the plot between ln kd and reciprocal of
the absolute temperature (Fig. 5d inset). The Ed for the immobi-
lized enzyme (178.67 kJ mol−1) was higher in comparison to free
enzyme (166.08 kJ mol−1). The higher value of Ed meant more
energy was required to denature the bound enzyme. These obser-
vations implied that the immobilized enzyme was more stable and
resistant to heat denaturation. At higher temperatures, the rate of
enzyme inactivation was greater as indicated by lower D-values
[42].
The thermo stability parameters of free and the immobilized
lipase are summarized in Table 4. The half-lives and D-values of
lipase prolonged remarkably at all temperatures after covalent
attachment indicating better thermo stability of the immobilized
lipase. The values of ꢀH◦ were higher for the immobilized lipase
as compared to free lipase (Table 4). At 30 ◦C, ꢀH◦ of free lipase
was 163.54 kJ mol−1 while that of the immobilized enzyme was
176.12 kJ mol−1 indicating the requirement of more energy for
thermal denaturation of the immobilized enzyme. The thermal
denaturation of enzymes is accompanied by the disruption of
non-covalent linkages, including hydrophobic interactions, with
concomitant increase in the enthalpy of activation [43]. The ꢀG◦
for thermal unfolding at 30 ◦C was 110.16 kJ mol−1 for free enzyme
while after immobilization it was 114.85 kJ mol−1. A higher value of
free energy of thermal denaturation indicated the resistance of the
immobilized enzyme towards thermal unfolding at higher temper-
atures which meant more stability of the immobilized enzyme. The
ꢀS◦ values were also enhanced after immobilization (Table 4). This
could result from thermal agitations and conformational changes.
In accordance with the results of this study, immobilization caused
significant increase in both entropy and enthalpy changes com-
pared to the native enzyme.
3.4. Effect of temperature on free and the immobilized lipase
The effect of varying temperature from 25 to 45 ◦C revealed that
the activity of free lipase increased up to 30 ◦C and declined on
further raising the temperature while the immobilized lipase dis-
played maximum activity at 37 ◦C (Fig. 5a). It was inferred from
these results that the temperature optima for catalysis of the reac-
tion by free and the immobilized enzymes were 30 and 37 ◦C,
respectively. So, there was displacement of optimum temperature
of the enzyme from 30 to 37 ◦C after immobilization. Nawani et al.
[32] also observed a shift in the optimum temperature after the
immobilization of lipase.
Activation energy for the immobilized lipase was calculated
from the slope of the Arrhenius plot (Fig. 5a inset). The regres-
sion equation for Arrhenius plot of free and the immobilized
lipase was y = −0.600x + 0.751 and y = −0.758x + 1.138, respec-
tively. The activation energy calculated for the immobilized lipase
was 6.3 kJ mol−1, which was higher than that of free enzyme
(5.0 kJ mol−1). The strong affinity between the lipase and alu-
minum oxide pellets might cause a higher energy barrier of the
first deactivation step. Similar results were reported during the
immobilization of lipase by other researchers [7,39].
On comparing the thermal stability of free and the immobi-
lized lipase it was found that free lipase was completely inactive
The z-values of free and the immobilized lipases were 12.2 and
11.1 ◦C, respectively (Fig. 5e). The higher z-value of the immobi-
lized enzyme implied that it was more sensitive to increase in
temperature rather than its duration as compared to its soluble
counterpart.
3.5. Effect of organic solvents
Fig. 4. Investigation of pH stability of free (solid circle) and the immobilized (hollow
circle) lipase. Free and the immobilized lipases were pre-incubated separately in the
buffers (mentioned in Fig. 3) for 2 h followed by determination of residual activity
at the optimum temperature under standard assay conditions.
The immobilized lipase showed better stability than free
enzyme in organic solvents (Fig. 6a). However, the extent of

58
D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
Fig. 5. Effect of temperature on free (solid circle) and the immobilized (hollow circle) lipase. (a) Temperature optima, Inset is the Arrhenius plot to calculate activation energy
(Ea) of catalysis; (b) comparison of thermal stability of free and the immobilized lipase; (c) thermal stability of the immobilized lipase with respect to time; (d) first order
thermal deactivation of the immobilized lipase, Inset is the Arrhenius plot to calculate activation energy (Ed) for denaturation and (e) temperature dependence of the decimal
reduction of free and the immobilized lipase to calculate z-values.
stability varied from solvent to solvent. The enhancement in the
stability of the immobilized lipase might be due to the increase in
rigidity of the enzyme upon covalent attachment of the enzyme
on aluminum oxide pellets. Increase in solvent stability after the
immobilization on synthetic beaded macroporous copolymers has
earlier been reported [44].
The effect of organic solvent on the activity of the immobi-
lized lipase with respect to time of incubation (12, 24 and 36 h)

D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
59
Table 4
Kinetic and thermodynamic parameters for thermal inactivation of free and immobilized lipase.
Thermodynamic entity
Temperature
30 ◦
C
35 ◦
C
40 ◦
C
45 ◦
C
50 ◦
C
55 ◦
C
60 ◦
C
F
I
0.00005
0.00001
0.00008
0.00004
0.00200
0.00100
0.00500
0.00200
0.01000
0.00500
0.01700
0.00700
0.01500
0.01500
kd (min−1
t1/2 (min)
)
F
I
13,862.94
115,524.53
8664.34
17,328.68
346.57
693.15
138.63
346.57
69.32
138.63
40.77
99.02
46.21
46.21
F
I
46,051.70
383,764.18
28,782.31
57,564.63
1151.29
2302.59
460.52
1151.29
230.26
460.52
135.45
328.94
153.51
153.51
D-value (min)
F
I
166.48
179.06
166.46
179.04
166.44
179.02
166.43
179.01
166.42
179.00
166.42
179.00
166.42
179.00
ꢀH◦(kJ mol−1
ꢀG◦(kJ mol−1
)
F
I
109.51
114.85
110.16
111.93
103.61
105.41
102.88
105.31
102.68
104.54
102.87
104.29
102.82
103.62
)
F
I
188.02
211.91
182.82
217.90
200.73
235.15
199.83
231.77
197.34
230.53
193.76
224.74
184.96
222.74
ꢀS◦(J mol−1 K−1
)
F and I represent free and the immobilized lipase, respectively.
was also investigated and the results are shown in Fig. 6b. The
immobilized lipase was found to be 100% stable in xylene, ethanol,
methanol, toluene and hexane after 12 h of incubation. The stability
in these solvents though declined after 24 h incubation as com-
pared to 12 h yet the relative activity was greater than 90%, except
methanol (80%). After 36 h of incubation, the bound enzyme was
most stable in DMSO followed by toluene, hexane and xylene with
residual activity of 93.1, 81.8, 73.4 and 71.9%, respectively. Further,
isopropanol and acetonitrile inactivated the immobilized lipase to
some extent as indicated by low residual activity, i.e. 69.5% and
45.8% after 12 h, respectively. The greater stability of Bacillus sp.
DVL 2 lipase in water immiscible solvents is in accordance with
earlier findings [18,21]. Most lipases exhibit a lid of hydrophobic
amino acids over the active site. In the presence of hydrophobic
organic solvents such as toluene, the lid is opened up and more
substrate gets accessible to active site resulting in higher enzyme
activity than the control [45,46]. The effect of organic solvents may
also vary among different lipases [47,48].
3.6. Determination of kinetic parameters
The Kinetic parameters were calculated using the
Lineweaver–Burk plot. The Km was 0.94 and 1.3 mM while
Vmax was 99 and 108 ␮mol/min for free and the immobilized
lipase, respectively (Fig. 7). An increase in Km upon immobilization
might be due to limited diffusion of the substrate to the active site
of the enzyme. The increase in Vmax upon immobilization might be
due to enhanced stability of the covalently bound enzyme.
3.7. Esterification
Enzymatic methods of ester synthesis are more effective in
non-aqueous media. In this study, the potential of free and the
immobilized lipases were compared in esterification of oleic acid
with ethanol to form ethyl oleate in hexane (Fig. 8a). The immo-
bilized lipase showed maximum 63% conversion of oleic acid into
ethyl oleate in 16 h whereas free lipase showed maximum 60% con-
version of oleic acid into ethyl oleate in 24 h. The ester formation by
the immobilized enzyme was also confirmed by TLC chromatogram
and 1H NMR spectra. Analysis of 1H NMR spectrum of ethyl oleate
is shown below:
Fig. 6. (a) Comparison of stability of free (solid circle) and the immobilized (hollow
circle) lipase in different organic solvents each viz. isopropanol, xylene, methanol,
ethanol, dimethylsulphoxide (DMSO), toluene, hexane and acetonitrile at a final
concentration of 25% (v/v) after 24 h incubation. (b) Effect of the above mentioned
organic solvents on the activity of the covalently immobilized lipase with respect to
time of incubation.
1H NMR (500 MHz, CDCl3): ı 5.38–5.30 (m, 2H), 4.14–4.10 (q, 2H,
J = 7.1 Hz), 2.30–2.27 (t, 2H, J = 7.5 Hz), 2.05–2.00 (m, 4H), 1.62–1.58
(m, 6H), 1.35–1.24 (m, 19H), 0.89–0.86 (t, 3H, J = 7.0 Hz).
Since the immobilized lipase enhanced the conversion of acid
into ester by 3% and reduced esterification time by 8 h, it was more

60
D. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 87 (2013) 51–61
efficient in esterification than free lipase. This might be due to
greater stability of the covalently immobilized enzyme in organic
solvents. These results are similar to those of Kanwar et al. [49] who
observed 58% conversion of oleic acid and ethanol into ethyl oleate
in n-nonane using synthetic hydrogel-bound lipase of Bacillus coag-
ulans.
The investigation of reusability of the immobilized lipase in the
above esterification reaction for seven cycles revealed a gradual
decrease in esterification efficiency after 2 cycles, retaining about
50, 31 and 12% of the initial activity after 5th, 6th and 7th cycle
respectively (Fig. 8b) which could be due to several reasons includ-
ing the inhibitory effect of water released during esterification
[50] and prolonged exposure to the organic solvent. In agreement
with these findings, repeated use of C. rugosa lipase covalently
immobilized on chitin for butyl butyrate synthesis lost almost 83%
of its initial activity in just 5 s [51]. Similarly, reuse of immobi-
lized Candida antarctica lipase-B by adsorption on Lewatit for three
consecutive esterification of lauric acid and isopropanol showed
40% reduction in percent relative conversion after each cycle [52].
They attributed it to enzyme deactivation and protein desorption.
Another report [53] revealed linear decay in percentage of residual
activity of cross linked C. antarctica lipase-B on chitosan in ethyl
oleate synthesis and the enzyme was completely deactivated after
four cycles. They related this deactivation to a combined effect
of deactivation/desorption of lipase and repeated manipulation
operations. It is likely that recyclability of the immobilized lipase
depends on either the nature of support or the enzyme source as
suggested earlier [52].
Fig. 7. Lineweaver–Burk plot for estimation of kinetic parameters for free (solid
circle) and the immobilized (hollow circle) lipase. The substrate concentration used
to generate this plot ranged from 0.17 to 2.6 mM.
4. Conclusion
In this study Bacillus sp. DVL2 lipase was covalently immobi-
lized on glutaraldehyde activated aluminum oxide pellets, which
are mechanically tough and resistant to high pH and temperature.
This is apparently the first report for the immobilization of lipase on
aluminum oxide pellets through covalent attachment. The immo-
bilized lipase showed maximum immobilization yield of 78.20%
under RSM optimized conditions and exhibited operational stabil-
ity for 5 consecutive cycles without any loss of enzyme activity.
The stability of the immobilized enzyme to pH, temperature and
organic solvents was enhanced as compared to free lipase. Thermo-
dynamic studies also indicated an improvement in enzyme stability
after immobilization. The efficiency of the immobilized lipase in
esterification of oleic acid and ethanol was better as it increased
the conversion of acid to ester by 3% and reduced the time of
esterification by 8 h as compared to free enzyme, which showed
60% conversion of acid to ester in 24 h. Reuse of the immobilized
enzyme for seven cycles in esterification reaction showed a gradual
decrease in esterification efficiency after 2 cycles, retaining about
50, 31 and 12% of the initial activity after 5th, 6th and 7th cycle
respectively.
Appendix A. Supplementary data
Supplementary data associated with this article can be
found, in the online version, at http://dx.doi.org/10.1016/
j.molcatb.2012.10.002.
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