Immobilization of Thermomyces lanuginosus lipase on ZnO nanoparticles: Mimicking the interfacial environment

Article abstract of DOI:10.1039/c5ra02249e

Thermomyces lanuginosus lipase (TL lipase) was immobilized covalently on ZnO nanoparticles (NPs) functionalized with small amino acid molecules, like glycine. Glutaraldehyde was used as a spacer between the ZnO/glycine Nps and the enzyme. This study is based on the observation that the favorable conformation of an enzyme (in which the catalytic lid is exposed to reactant molecules) can be obtained at the lipid/water interface and such an interfacial environment can be mimicked by properly designing the carrier used as the support for its immobilization. Glycine functionalized ZnO NPs were covalently bonded with glutaraldehyde and consequently TL lipase enzyme immobilization was carried out by a simple wet chemical method. The resulting assemblies were characterized by using techniques like XRD, UV absorption and photoluminescence spectroscopy. The particle size was determined by using Transmission Electron Microscopy (TEM). The immobilized TL lipase enzyme showed high activity for esterification of oleic acid (C-18) with methanol in an organic medium. The catalyst was recovered and reused several times without any significant loss of activity.

Full text of DOI:10.1039/c5ra02249e

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Immobilization of Thermomyces lanuginosus lipase
on ZnO nanoparticles: mimicking the interfacial
environment†
Cite this: RSC Adv., 2015, 5, 26291
Ekta Shah, Paramita Mahapatra, Ashutosh V. Bedekar and Hemant P. Soni*
Thermomyces lanuginosus lipase (TL lipase) was immobilized covalently on ZnO nanoparticles (NPs)
functionalized with small amino acid molecules, like glycine. Glutaraldehyde was used as a spacer
between the ZnO/glycine Nps and the enzyme. This study is based on the observation that the favorable
conformation of an enzyme (in which the catalytic lid is exposed to reactant molecules) can be obtained
at the lipid/water interface and such an interfacial environment can be mimicked by properly designing
the carrier used as the support for its immobilization. Glycine functionalized ZnO NPs were covalently
bonded with glutaraldehyde and consequently TL lipase enzyme immobilization was carried out by a
simple wet chemical method. The resulting assemblies were characterized by using techniques like XRD,
UV absorption and photoluminescence spectroscopy. The particle size was determined by using
Transmission Electron Microscopy (TEM). The immobilized TL lipase enzyme showed high activity for
esterification of oleic acid (C-18) with methanol in an organic medium. The catalyst was recovered and
reused several times without any significant loss of activity.
Received 5th February 2015
Accepted 5th March 2015
DOI: 10.1039/c5ra02249e
www.rsc.org/advances
heterogenized and the immobilized enzyme can be recovered
and recycled for several times maintaining the activity. This
1
. Introduction
With thousands of years of evolution nature has developed makes overall operation simple, efficient in both aqueous and
molecules called enzymes which can carry out specic non-aqueous medium, and of course, economically viable.
biochemical conversions in the constrained environment of a
Lipase (triacylglycerol ester hydrolase EC 3.1.1.3) is an
cell. Chemically, enzymes are long chain polypeptides folded in enzyme which catalyzes the hydrolysis of triacylglycerol to
such a way that unique reaction sites (pockets) are generated glycerol and fatty acids. It nds wide applications in diverse
according to a predened genetic program enabling them to act areas like dairy industry, specialty chemicals, organic synthesis
as catalysts. Highly selective and specic reactions occur at and manufacture of enantiomerically pure pharmaceuticals.
1
these reaction sites usually in a narrow range of temperatures Immobilization of lipase is carried out by various techniques
and pH values in an aqueous medium. From an industrial point such as (i) non-covalent adsorption on robust supports like
2,3
4,5
6
7
of view, enzymes can be manufactured or extracted from the polymeric beads, lms, natural kaolin clay, nanoparticles,
cells and utilized for large scale production of high purity etc. (ii) entrapment of enzyme in a polymeric gel or on
stereoisomers which would be difficult by conventional catalytic membranes by physical adsorption, inclusion or covalent
8
–11
processes. However, the major drawbacks are (i) denaturation of bonding
(iii) covalent attachment with the supports like
1
2
13
enzymes when the temperature or pH of the reaction are dras- polymers or nanoparticles and (iv) cross-linking of an
14
tically changed (ii) enzymes function mostly in an aqueous enzyme making it carrier free. However, each method has its
medium under homogeneous conditions (iii) enzymes are own drawbacks. For example, non-covalent adsorption of an
denatured sometimes and the reaction sites may be distorted/ enzyme may cause multilayers of unfavorable orientations on
blocked at the end of a single reaction cycle during the the support that hamper the activity. Cross-linking may cause
recovery process. Such problems can be solved by immobilizing inactive enzyme aggregates, deteriorating the catalytic efficiency
enzyme on a suitable support. By this way the reactions can be of preparation. Physical entrapment in gels or membranes also
requires the trapping of enzyme in active orientation in the
15
matrix which is laborious and demands a lot of experimenta-
1
6
Department of Chemistry, Faculty of Science, The Maharaja Sayajirao University of tion. This may restrict the natural movement of the enzyme
Baroda, Vadodara-390 002, Gujarat, India. E-mail: drhpsoni@yahoo.co.in; Tel:
diminishing the catalytic efficiency. The fact is that there is no
universal method available for immobilizing the enzyme
without restricting its activity. Suitable selection of a carrier,
reaction conditions and enzyme itself are the three major
+
91-0265-2795552
†
Electronic supplementary information (ESI) available: FTIR spectra of
17
1
13
immobilized enzyme at various stages, H NMR, C NMR and HPLC analysis of
the product methyl oleate. See DOI: 10.1039/c5ra02249e
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components governing the performance of the developed the growth. The synthesized ZnO NPs were used as support to
18
immobilized enzyme. Proper strategy based on these param- covalently immobilize TLL enzyme. It was observed that the
eters is required to develop a suitable biocatalyst for a chemical microenvironment in the vicinity of the enzyme in terms of
transformation.
lipophilic–lipophobic balance and exibility of the adopted
ZnO is an ionic solid and environmentally friendly semi- conformation as well as porosity of the support directly affect
1
8,33
conducting material with band gap energy of 3.37 eV at room the activity.
The coating of zwitterionic glycine molecules
19
temperature. It nds wide applications in solar cells, photo- functions in two ways. First, they stabilize ZnO NPs by adjusting
20
21
22
catalysis,
biosensors,
luminescent material
etc. Its the surface charge, making the surface hydrophilic. Secondly,
biocompatible, biodegradable and antimicrobial nature they provide sites for glutaraldehyde activation. The glycylimine
encourage researchers using it as a sensor for biotransforma- assembly acts as hydrophobic spacer between ZnO NPs and
23,24
tion.
In the present strategy ZnO/glycine/glutaraldehyde NPs enzyme. Consequently, the microenvironment in its vicinity is
have been developed as a catalyst akin to a-Al O , zeolites and tailored in such a way that the enzyme can be in its high activity
2
3
cationic clay sheets which could provide an inert support for state. The activity of the developed immobilized enzyme was
covalent binding of TLL enzyme and/or it may also enhance the evaluated for the esterication of oleic acid with methanol.
catalytic activity of lipase synergistically by maintaining hydro-
philic–hydrophobic balance.
Nowadays, biodiesel which is a mixture of methyl and ethyl
ester of fatty acids (mainly palmitic, linolenic, linoleic, and oleic
2
. Experimental
2.1 Materials
acids) is gaining an importance because of its comparatively Zinc acetate Zn(CH
COO)
was obtained from Loba chemicals,
CH COOH),
3
OH) were purchased from S.D. Fine Chemicals,
3
2
clean burning, renewability, biodegradability and superior India. Potassium hydroxide (KOH), glycine (NH
2
2
25
lubricating properties. Non-edible oils and fats, vegetable oils methanol (CH
and their rening by-products are good sources of biodiesel. The Mumbai, India. All the chemicals were AR grade and used
amount of free fatty acids (FFA) content and the storage stability without further purication. Freshly prepared aqueous
are some of the parameters affect the quality of the biodiesel. solutions were used for the synthesis of ZnS nanoparticles.
Oxidation of unsaturated esters in biodiesel occurs by contact Alkaline solution of Thermomyces lanuginosus lipase (TLL,
5
ꢀ1
with air and other pro-oxidizing conditions during long term activity-10 U g ) procured from Aumgene Chemicals Limited,
storage. It was observed that the extent of unsaturation in fatty India. Glutaraldehyde was purchased from Merck.
acid molecules (in FFA, mono-, di- or triglycerides) adversely
2
6,27
affect the storage stability.
diesel should be less than 1% for superior quality and storage
Ideally, the FFA content in bio- 2.2 Synthesis of ZnO nanoparticles
ZnO NPs were synthesized by a co-precipitation method.
Methanolic solutions of zinc acetate (40 mL, 0.03 mmol) and
glycine (20 mL, 0.04 mmol) were mixed. To this mixture, alco-
holic KOH solution (40 mL, 0.13 mmol) was added drop wise
and with constant stirring at 8000 rpm at room temperature.
28
stability. For the purpose, acid or base-catalyzed esterication of
FFA is carried out. The major problem associated with base
catalysis is that it reacts with FFA instead of catalyzing the reac-
tion and form soap like sticky mass. These decrease the bio-
diesel yield and create problems associated to the separation and
purication steps due to emulsions and partial dissolution of
Fatty Acid Methyl Esters (FAME) into the glycerol phase. Mineral
acid (e.g. H SO or HCl) catalyzed esterication of FFA is also
associated with the work-up problems. It is very difficult to tackle
the issues like recovery and recycling of the by-products, corro-
sion and other such environmental problems crop up during the
29
ꢁ
The resulting colloidal solution of ZnO NPs was ripened at 70 C
for three days. The solution was centrifuged, washed several
times with distilled water then with absolute alcohol to remove
any impurities and dried at 60 C under vacuum for three days.
30
2
4
ꢁ
2.3 Lipase immobilization
31
Immobilization of lipase on the surface of ZnO NPs was carried
out at room temperature. 200 mg of glycine functionalized ZnO
NPs were dispersed in 0.5 mL PBS (5 mmol) buffer at 7.7 pH.
Subsequently, glutaraldehyde (10 mL) was added and the mixture
was stirred for about 4–5 h. Then 0.5 mL native TL lipase
reaction. Solid acid and supported Lewis acid catalysis may be
preferred; however, the water produced as by-product severely
affects the efficiency (in terms of leaching or solubility) of the
catalyst and an environmental problem of discarding the catalyst
32
aer several cycles still persists. Catalysis involving the immo-
bilized lipase enzymes for the esterication of FFA may appear a
good choice. Careful selection of the support (in term of biode-
gradability and/or toxicity) for immobilization of the enzyme can
minimize the environmental issues. Nanomaterials have high
surface area to volume ratio and when coated with catalyst,
provide efficient catalysis with minute quantity of material which
can be reused for several cycles and regenerated by immobilizing
fresh enzyme on the surface instead of polluting the environment
by unrestricted waste disposal.
ꢀ
1
(1.675 mg mL , determined from protein assay) aqueous solu-
tion prepared in 0.5 mL PBS buffer (pH 7.7) was introduced. The
mixture was then gently but thoroughly stirred to ensure
complete mixing for further 4 h and then centrifuged at
8000 rpm. The solid collected was washed three times with 60 mL
(5 mmol, 20 mL for each washing) PBS solution (Scheme 1).
2.4 Protein estimation
Protein estimation was carried out by Lowry's method with
3
4
In the present study, we have attempted the wet chemical bovine serum albumin as standard. It was calculated that on
synthesis of ZnO NPs using glycine as capping agent to restrict 100 mg of ZnO, 32.6 mg of lipase was loaded.
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incubation respectively. One international unit of lipase activity
was dened as the amount of enzyme catalyzing the release of
1
mmol of p-NP per min from p-NPP under the standard assay
36
conditions.
ꢀ1
ꢀ1
Specific activity (U mg ) ¼ {enzyme activity (U mL )/
ꢀ
1
protein content (mg mL )}
(2)
2.6 Operational stability of immobilized lipase
2.6.1 Effects of pH on enzyme activity. Lipase activity was
measured in different buffers (0.05 M) of pH (5–12) viz. sodium
citrate (pH 4.0–6.0), sodium phosphate (pH 7.0–9.0) and
ꢁ
glycine–NaOH (pH 10.0–11.0) at 37 C by incubating the free
and immobilized lipase with p-NPP (maintaining other condi-
tions same as mentioned above) for 1 h. The lipase activity was
measured by the above standard assay method.
Scheme 1 Covalent immobilization of TL lipase enzyme on ZnO
nanoparticles.
35
2.6.2 Effect of temperature on enzyme activity. The
optimum temperature of lipase activity was determined by
carrying out enzymatic reactions at different temperature (55,
2
.5 Enzyme assay
ꢁ
65, 75, 85 and 95 C) at pH 7.4 by incubating the free and
Lipase activity was assayed using p-nitrophenylpalmitate
immobilized lipase with p-NPP for 1 h. Subsequently, NaOH
1 N, 1 mL) was added to stop further hydrolysis and then the
35
(p-NPP) as substrate. The basis of this assay protocol is the
(
colorimetric estimation of p-NP released as a result of enzymatic
hydrolysis of p-NPP at 410 nm. Enzyme solution (0.8375 mg/
lipase activity was determined by above mentioned assay
35
method.
50 mL) was added to 950 mL of the substrate solution consist-
2.6.3 Thermal stability of immobilized enzyme. Thermal
ing mixture of solution A (50 mL, 5.0 mM p-NPP in 2-propanol)
and solution B (900 mL, 100 mM potassium phosphate buffer
with pH 7.0, 0.4% Triton X-100 and 0.1% gum arabic), which
were freshly prepared before use. The reaction mixture was
stability of free and immobilized lipase was studied at 55, 65
ꢁ
and 75 C. The stability was studied by collecting the aliquot at
different time interval and then lipase activity was determined
35
by standard assay method. The stability of the support
ꢁ
incubated at 37 C for 10 min. The reaction was then terminated
ZnO/glycine/glutaraldehyde was also studied using differential
thermal analysis.
by adding 1 mL ethanol. The absorbance of the yellow color
product p-nitrophenol (p-NP) was measured in spectropho-
tometer at 410 nm. Enzyme activity and specic activities were
calculated as per the following formula,
2.7 Esterication reaction
Immobilized TLL on ZnO NPs was used as biocatalyst for
(1) esterication of oleic acid with methanol. For the reaction,
A ꢂ B
ꢀ
1
Lipase activity U mL
¼
C ꢂ D ꢂ E
100 mg of immobilized lipase was dispersed in 5 mL of petro-
where, A, B, C, D and E are mmol of p-NPA released, total volume leum ether at room temperature. To this solution, oleic acid
of the reaction mixture, volume used in spectrophotometric (5.95 mmol), methanol (2.97 mmol) and molecular sieves (1.6 g,
determination, volume of enzyme used in assay and time of 3 ꢀA ) were introduced. The temperature of the reaction then
Scheme 2 Esterification of oleic acid (C-18) with methanol in presence of TL lipase enzyme immobilized on ZnO nanoparticles.
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ꢁ
gradually raised to 55 C. Aer about 10 h of reux, further b cos q) and the FWHM (full width at half maximum) value
methanol (2.97 mmol) was added and the reaction was allowed corresponding to the major plane (101), was 40 nm. On
to continue for about 14 h. Progress of the reaction was moni- glutaraldehyde activation and covalent attachment of lipase on
tored on TLC plate. The purity of the product, aer the the ZnO/Gly NPs, the XRD peak positions almost remain same
1
13
completion of reaction, was determined by HPLC. For H,
C
with change in intensity. It can be observed (Fig. 1) that on
NMR, and FTIR spectral analysis the reaction mixture was loading enzyme, the intensity of (002) peak increases indicating
passed through column having silica as stationary phase and preferred unidirectional growth. The compactness and clear
ethyl acetate:petroleum ether as eluent (Scheme 2).
resolution of the peaks towards lower angles indicate properly
) d(ppm): ordered and compact enzyme layer on the surface of NPs.
1
2.7.1 Methyl oleate. H NMR: (400 MHz, CDCl
3
5
.37 (m, 2H), 3.69 (s, 3H), 2.33 (2H), 2.19–1.70 (m, 4H), 1.62 Further, the size and shape of ZnO/Gly NPs were studied by TEM
1
3
(m, 2H), 1.47–1.28 (m, 20H), 0.88 (t, 3H). C NMR (CDCl , 100 analysis. The TEM image of pristine ZnO/Gly NPs shows almost
3
MHz) d (ppm): 174.31, 129.98, 129.73, 51.42, 34.09, 31.90, 29.76, monodispersed spherical particles with an average size 9–16 nm
2
1
9.68, 29.52, 29.32, 29.15, 29.08, 27.21, 27.15, 24.94, 22.68, (Fig. 2a). On covalent binding of an enzyme, the size and shape
4.16.
of the NPs almost remains same (Fig. 2b). The surface
morphology of the pristine ZnO/Gly NPs was studied by FEG-
SEM analysis (Fig. 3). It can be seen from the Fig. 3 that the
surface of the NPs is highly porous. The pores and cracks are
formed due to agglomeration of NPs on the surface, indicating
that the surface is capable to show catalytic activity.
2
.8 Characterization of synthesized nanoparticles and
esterication products
X-ray powder diffraction (XRD) pattern of the ZnO/Gly NPs and
ZnO/Gly/glutaraldehyde/enzyme NPs were obtained from X-ray
powder diffractometer (Bruker D8 Advance) with Cu Ka radia-
tion, l ¼ 0.15418 nm. The nanoparticles were dispersed in
water, sonicated for 10 minutes, and then UV-visible absorption
spectra were recorded by means of Perkin-Elmer Lambda 35
UV-visible spectrophotometer. Photoluminescence (PL) spectra
were recorded on a Jasco FP-6300 using xenon lamp as the
excitation source at 260 nm. The size and shape of the nano-
particles were studied by means of Transmission Electron
Microscopy (TEM, Philips Tecnai 20). A BIC 90 plus (Broo-
khaven) equipped with 35.0 mW solid state laser operating at
3.2 Optical studies
The quality of the pristine ZnO/Gly NPs was further conrmed
using optical studies. As can be seen from Fig. 4 that ZnO/Gly
NPs absorb sharply at 371 nm which is blue shied when
compared with bulk ZnO (380 nm). This observation conrms
38
the quantum connement and nano regime. When suspended
in ultrapure water and excited at 200 nm, the ZnO/Gly lumi-
nesces at 425 nm. It can be observed from the Fig. 5 that virgin
enzyme uoresce at 470 nm and on covalent binding with
glutaraldehyde on the surface of ZnO/Gly NPs, it is blue shied
to 360 nm. Haldar et al. explained the phenomena of blue
shiing of lmax of enzyme on excitation at a suitable wavelength
due to unfavorable interactions of dipole moments of polar
solvent molecules with uorophores of enzyme under excited
state. This observation may lead to assume the conforma-
tional changes in enzyme and the exposure of catalytic hydro-
phobic triad (Asp–Hys–Ser) in aqueous medium on covalent
binding.
660 nm and an avalanche photodiode detector was used for the
measurement of surface charges in term of zeta potential (x). All
measurements were made at 25 C in deionized water. Differ-
ential scanning calorimetric (DSC) analysis of ZnO/glycine/
glutaraldehyde was also carried out using Mettler Toledo DSC
8
up. The heating rate was 10 C min from RT to 500 C in N
atmosphere. The FTIR (Perkin-Elmer, RX-FTIR) spectra of the
samples (biocatalyst and esterication products) were obtained
in the range of 400 to 4000 cm . The H NMR and C spectra
ꢁ
39
22. For the purpose, the material was heated inside a DSC set-
ꢁ
ꢀ1
ꢁ
2
ꢀ1
1
13
3.3 Surface charges
of the esterication products were recorded on a Bruker
Advance 400 MHz spectrometer using TMS as an internal Surface charges on the NPs were studied by zeta potential (x)
standard in CDCl . HPLC analysis was carried out by using measurement. It gives an idea about the electrostatic charge
Shimadzu LC-20AD having C18 column (250 nm ꢂ 4.5 m) and
methanol : H O (80 : 20 v/v) as mobile phase with a ow rate of
mL min for 30 minutes.
3
2
ꢀ1
1
3. Results and discussion
3.1 Compositional and morphological studies
The XRD patterns of the as-synthesized glycine functionalized
ZnO NPs (ZnO/Gly) are shown in Fig. 1a. The XRD patterns
manifest predominant diffraction peaks at 2q values 31.7, 34.3,
36.2, 47.4, 56.5, 62.7, 66.3, 67.8, 69.0, 74.4 and 76.8. These peaks
are well matched with standard JCPDS card no. 89-1397 corre-
sponding to ZnO with hexagonal wurtzite phase. The particle
size was calculated using Debye–Scherrer formula (D ¼ 0.9l/ enzyme immobilized ZnO nanoparticles.
Fig. 1 XRD patterns of (a) bulk ZnO nanoparticles (JCPDS card no.
9-1397) (b) glycine functionalized ZnO nanoparticles (c) TL lipase
8
37
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Fig. 4 UV absorption spectrum of ZnO/Gly nanoparticles.
ꢀ
During crystal growth, both OH ions and glycine act as
ꢀ
capping agents, however, OH ions are more prone to be
adsorbed on the surface of NPs while glycine makes coordina-
2
+
tion with the surface Zn ions. On glutaraldehyde activation
Fig. 2 TEM images of (A) ZnO/Gly nanoparticles (B) TL enzyme
immobilized ZnO/Gly nanoparticles.
stabilization during the growth of NPs. From Table 1 it can be
seen that pristine ZnO/Gly NPs possess negatively charged
surface with x value ꢀ17.15 mV and on glutaraldehyde activa-
tion and enzyme binding the surface charge decrease to
ꢀ
6.12 mV. The emergence of charge on the surface is directly
inuenced by the presence of counter ions in the medium and
40
mode of crystallization. The negative charge on pristine ZnO
NPs is due to presence of OH ions in the basic medium.
ꢀ
Fig. 5 Photoluminescence spectra of (A) ZnO/glycine nanoparticles
Fig. 3 FEG-SEM image of glycine functionalized ZnO nanoparticles.
(B) free TL enzyme and TL enzyme immobilized ZnO nanoparticles.
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Table 1 Surface charge analysis of as-synthesized nanoparticles
Charge, x
System
(in mV)
ZnO/glycine
ZnO/glycine/glutaraldehyde/TLL enzyme
ꢀ17.15
ꢀ6.12
and covalent binding of enzyme, as proposed earlier, the
conformational changes in enzyme takes place which increases
ꢀ
the hydrophobicity and expel the OH ions or polar water
molecules present in the vicinity. However, the polar molecules
could not be totally expelled due to surface coordinated
hydrophilic glycine molecules. This fact is expressed in term of
decrease in x value to ꢀ6.12.
ꢁ
Fig. 7 Effect of pH on hydrolysis of p-NPP at 40 C.
3
.4 FTIR study
two fundamental modes of vibrations of carboxylate ion in free
acetate form can be observed: one is COO asymmetric stretch-
To understand the interactions among the ligands and the
surface of ZnO NPs, FTIR spectroscopy is one of the best tools.
The mode of interaction of carboxylate ions present in an ionic
glycine can be conrmed from the vibrational spectroscopy
ꢀ
ing mode n (–COO–) and the other is symmetric stretching mode
as
n (–COO–). The fundamental frequencies of vibrations for free
s
ꢀ1
acetate ion are 1583 and 1422 cm for nas(–COO–) and n
s
(–COO–)
(FTIR). Fig. S1† shows FTIR spectra of native enzyme, ZnO/Gly
respectively. If carboxylate ligand coordinate with surface Zn(II) in
bidentate mode then nas(–COO–) decreases and n (–COO–)
increases from the normal modes in free state and vice versa in
and ZnO/Gly/glutaraldehyde/enzyme. In FTIR spectra of native
enzyme (Fig. S1a†), the peak near 1632 cm is amide I band
due to C]O stretching vibrations, the hump near 1510 cm is
amide II band due to N–H bending/C–H stretching vibration,
the amide III band at 1222 cm is due to N–H bending vibra-
tion. The band at 3430 cm is due to N–H bending and at 1404
cm is resulted from protein side chain COO .
the ways by which carboxylate ion of glycine (at pH 7.7,
isoelectric point 6.0) can coordinate to the surface Zn(II) ion
either as a unidentate ligand or as a chelating (bidentate
ꢀ
1
s
ꢀ
1
47
case of monodentate mode. On comparing D(nas(–COO–) ꢀ
0
0
0
0
n (–COO–)) with D (n (–COO–) ꢀ n (–COO–)), we found D > D ,
ꢀ
1
s
as
s
(
where D indicates difference in the absorption bands for free
ꢀ1
0
carboxylate ion and D indicates the same for metal bound
carboxylate ion) which suggests bidentate coordination of
glycine with Zn(II) ion on the surface of ZnO NPs. On glutaral-
dehyde activation and enzyme covalent binding, the broad
amide peak becomes sharp and blue shied compared to ZnO/
glycine due to high energy peptidal N–H stretching vibrations
ꢀ
1
ꢀ
41,42
Fig. 6 shows
4
3–46
fashion) ligand.
This can be detected on the basis of COO stretching vibra-
tion frequencies. On the basis of a normal coordinate treatment
ꢀ
ꢀ1
and at 1440 cm imine stretching vibrations indicating cova-
lent binding of TLL enzyme with glutaraldehyde on the surface
of ZnO NPs (Fig. S1C†).
It was observed from the previous studies that the catalytic
activity of the most of the lipase enzymes is due to Ser–His–Asp/
Fig. 6 Modes of interaction of glycine with the surface of the ZnO
nanoparticles (orange balls indicate ZnO nanoparticles, light blue for
carbon atoms, red for oxygen atoms, white for hydrogen atoms and
blue for nitrogen atom).
Fig. 8 Effect of temperature on hydrolysis of p-NPP at 7.4 pH.
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conformation of enzyme. However, this hydrophobic catalytic
triad can be made available to the substrate by forcing the
enzyme to change its conformation. It was proved that at lipid/
water interface the microenvironment around the enzyme
forces it to change its conformation in such a way that the active
sites (triad) expose on the surface and easily available to the
48–51
substrate.
It may be assumed that by proper tuning of the
environment around the bound enzyme its suitable conforma-
tion can be locked. The lipophilic–lipophobic balance in vicinity
of enzyme plays a key role to achieve the catalytically active
conformation. We have designed the enzyme based on the idea
that both hydrophilic (in term of zwitterionic glycine around the
solid ZnO NPs) and hydrophobic (in term of glutaraldehyde
chain) environment mimicking the lipid/water interface avail-
able to enzyme which stabilize the catalytically active confor-
mation of enzyme in hydrophobic (organic) medium.
3.5 Operational stability of lipase enzyme
3.5.1 Effect of pH. Fig. 7 shows the effect of pH on lipase
activity measured by spectrometric method. It can be observed
that the immobilized TLL enzyme shows always higher activity
than free form at the pH range 4–13. The free enzyme shows
very less activity at pH 4.0 which increases with rise in pH.
While immobilized lipase manifest constant increase in activity
with rise in pH. It can be observed that enzyme activity increases
ꢀ1
to 21 from 19 U mL on immobilization at pH 5 while at pH 12
ꢀ1
it is increases to 26 from 22.5 U mL of free counterpart. So it
can be concluded that both free and bound enzyme show
substantial stability at lower pH and it is increases with pH.
Higher enzyme activity can be achieved in basic medium.
Ortega et al. also reported enhanced stability of lipase at acidic
52
and alkaline pH upon immobilization. The increase in
stability upon immobilization might be due to multipoint
covalent linking between lipase and support, which prevents
36,53
lipase denaturation in acid or alkaline environments.
Similar results were obtained with the immobilized Candida
54
rugosa lipase on chitosan by Hung et al.
.5.2 Effect of temperature. It is a fact that all enzymes
3
denature and show degradation in activity at higher tempera-
tures. It can be observed from the Fig. 8 that the activity of lipase
increases three folds on immobilization and maximum activity
ꢁ
ꢁ
can be achieved at 55 C. At 65 C the immobilized enzyme
shows almost same activity which would have been shown by
ꢁ
free enzyme at 55 C i.e. on immobilization the operational
stability and activity of the TLL enzyme can be extended to
ꢁ
1
0 C. Industrial point of view, the immobilized enzyme shows
ꢁ
moderate activity upto 75 C. It is interesting to note that at
ꢁ
9
5 C, both free and immobilized enzyme show some activity on
hydrolysis of p-NPP. To explain this, a controlled experiment
using only glycine functionalized ZnO nanoparticles (without
any type of enzymes) was carried out. Similar kind of p-NPP
ꢁ
Fig. 9 Thermal stability of TL lipase at (A) 55, (B) 65 and (C) 75 C and
.4 pH.
7
ꢁ
hydrolysis activity was observed in the range of 90–95 C. Hence,
it can be inferred that this ‘residual’ activity is due to the
thermal effect and not to the enzyme activity.
Glu catalytic triad present inside the surface loop having
hydrophobic environment. This catalytic triad is not available to
the substrates in aqueous medium and under normal
3
.5.3 Thermal stability. Based on temperature studies, we
ꢁ
have selected 55, 65 and 75 C to monitor the thermal stability
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Fig. 10 Thermal analysis of ZnO/Gly/glutaraldehyde system.
of immobilized lipase (Fig. 9). Both type of enzymes exhibited a the rise in the amount of water as by-product in the reaction
similar trend; however, the immobilized lipase is more stable mixture. Generally, enzymes are involved to carry out
than that of free one. The half-life of the immobilized lipase is reactions/biotransformations in aqueous media. The enzyme
much longer than the free lipase at these three temperatures. activity is greatly hampered by shiing the reaction media
This study also support our argument that the immobilized from aqueous to non-aqueous. The other factors like resis-
lipase shows three folds increase in activity than its free coun- tance of the substrate to diffuse into the immobilized matrix
ꢁ
terpart at 55 C. It is obvious that with increase in temperature and resistance of the product to diffuse out (diffusional
the efficiency of both the enzymes decreases. We adopted a limitations), pH of the medium, exibility of the enzyme
rational approach to design immobilized lipase catalyst. Firstly, molecules etc. directly affect the efficiency of the enzymatic
5
0
ZnO NPs were functionalized by hydrophilic glycine and then reactions. The problem of diffusional limitation can be
hydrophobic glutaraldehyde activation was carried out before solved by inducing hydrophilic–hydrophobic environment
binding the enzyme.
on the surface of the support, the substrate (long chain fatty
The stability of resulting assembly (ZnO/Gly/glutaraldehyde) acid) can be attracted and oriented near the hydrophobic
was conrmed from thermal analysis (Fig. 10). Two major weigh chains of supported matrix such that there would be
losses were observed on TG curve. The rst loss (11.0%) maximum hydrophobic–hydrophobic interactions before
ꢁ
occurred at a temperature range between 50 to 110 C accom- undergoing the reactions in enzymatic lids and the products
ꢁ
panied by an endotherm at 104 C with a rate of decomposition formed can also be diffused by the same mechanism. The
ꢀ
1
0
4
.426 mg min then the curve becomes almost constant up to advantage of presence of hydrophilic area near the support is
ꢁ
ꢁ
00 C. The horizontal part of the curve between 150 to 400 C that the by-product water can be collected at the beginning
indicates stability of the support throughout the temperature and then entrapped in molecular sieves at certain level.
ꢁ
range. Aer this, 8.1% weigh loss at 458 C can be observed in Hence, the equilibrium can be shied continuously to
term of exotherm in DTA curve with maximum rate of decom- forward direction by ‘pseudo’ removal of water from the
ꢀ
1
position 0.84 mg min . This may be due to degradation of reaction mixture (Scheme 2). Hence, the supported enzyme
covalent bonding on the surface of ZnO NPs. This is a assembly can be viewed as ‘nano reactor’ to which the
straightforward conrmation of presence of hydrophilic– diffusion of reactants and products continuously takes place
hydrophobic environment on the surface.
with instant entrapment of by-product.
The maximum esterication yield (90%) is obtained in
ꢀ
1
presence of TLL enzyme (20 mg mL ) with oleic acid (C18) and
methanol in the molar ratio of 1 : 1 in petroleum ether as
solvent at 40 C. On HPLC analysis, the purity of the product
was found to be 84% for the rst cycle and 82.5 and 81% for the
next two consecutive cycles (Fig. S3†).
3
.6 Esterication reaction
ꢁ
Esterication of long chain fatty acid with alcohol is equi-
librium dominated reaction. The rate of reaction becomes
slow as it approaches towards the equilibrium and also with
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.7 Leaching study
RSC Advances
3
3 T. Boller, C. Meier and S. Menzler, Org. Process Res. Dev.,
2
002, 6, 509.
The stability and reusability of the immobilized enzyme is very
important. The leaching study of immobilized enzyme was
carried out using n-hexane as washing medium. The recycled
4
5
R. Dalla-Vechhia, D. Sebr ˜a o, M. G. Nascimento and V. Soldi,
Process Biochem., 2005, 40, 2677.
J. S. Crespo, N. Queiroz, M. G. Nascimento and V. Soldi,
Process Biochem., 2005, 40, 401.
6 M. B. A. Rahman, S. M. Tajudin, M. Z. Hussein,
R. N. Z. R. A. Rahman, A. B. Salleh and M. Basri, Appl. Clay
Sci., 2005, 29, 111.
M. I. Kim, H. O. Ham, S.-D. Oh, H. G. Park, H. N. Chang and
S.-H. Choi, J. Mol. Catal. B: Enzym., 2006, 39, 62.
55
immobilized TLL enzyme was used 4 consecutive cycles using
2
0 mL n-hexane for washing each cycle exhibiting almost same
th
activity as in the rst cycle, however, aer 4 cycle there was fall
in the activity and become almost 80% compared to the rst
cycle.
7
8
9
4
. Conclusion
N. Hilal, V. Kochkodan, R. Nigmatullin, V. Goncharuk and
L. Al-Khatib, J. Membr. Sci., 2006, 268, 198.
K. Nagayama, N. Yamasaki and M. Imai, Biochem. Eng. J.,
From this study it can be concluded that (1) microenvironment
surround the immobilized enzyme plays an important role for
activity and favorable enzymatic conformation can be obtained
by mimicking the environment existing at lipid/water interface
2
002, 12, 231.
0 J.-P. Chen and W.-S. Lin, Enzyme Microb. Technol., 2003, 32,
01.
1 M. Goto, C. Hatanaka and M. Goto, Biochem. Eng. J., 2005, 24,
1.
1
1
1
8
(2) diffusion limitation which is the major factor to restrict the
enzyme activity can be controlled by prior designing of the
biocatalyst (3) maximum catalytic activity and reusability can be
achieved for several cycles by proper covalent binding of the
biocatalyst with the solid support and (4) immobilized lipase
enzymes can serve as a ‘green catalyst’ for the production of
biodiesel. This study may provide a direction to future research
to achieve maximum catalytic activity from the enzyme by
changing its microenvironment e.g. using ‘spacer’ molecules
having different chain length or different amino acid molecules
as capping agent around the support altogether with optical
activity from the metal oxide support which also may be useful
in the area of developing bio-sensors for enzymatic reactions.
9
2 P. Ye, Z.-K. Xu, Z. G. Wang, J. Wu, H. T. Deng and P. Seta,
J. Mol. Catal. B: Enzym., 2005, 32, 115.
3 G. Jun, D. Lu, Z. Liu and Z. Liu, Biochem. Eng. J., 2009, 44, 53.
4 L. Cao, L. Langen and R. A. Sheldon, Curr. Opin. Biotechnol.,
1
1
2
003, 14, 387.
1
1
5 B. Al-Duri and Y. P. Yong, Biochem. Eng. J., 2000, 4, 207.
6 M. J. J. Litens, K. Q. Le, A. J. J. Straathof, J. A. Jongejan and
J. J. Heinen, Biocatal. Biotransform., 2001, 19, 1.
7 A. Dyal, K. Loos, M. Noto, S. W. Chang, C. Spangnoli,
K. V. Sha, A. Ulman, M. Cowman and R. A. Gross, J. Am.
Chem. Soc., 2003, 125, 1684.
1
1
1
8 L. Cao, Curr. Opin. Chem. Biol., 2005, 9, 217.
9 W. J. E. Beek, M. M. Wienk and J. Raj, Adv. Mater., 2004, 16,
Author contributions
1
009.
The manuscript was written through contributions of all
authors. All authors have given approval to the nal version of
the manuscript.
2
2
2
0 H. Zeng, W. Cai, P. Liu, X. Xu, H. Zhou, C. Klingshirn and
H. Kalt, ACS Nano, 2008, 2, 1661.
1 A. Umar, M. M. Rahman, S. H. Kim and Y. B. Hahn, Chem.
Commun., 2008, 166.
2 P. Dolcet, M. Casarin, C. Maccato, L. Bovo, G. Ischia,
S. Gialanella, F. Mancin, E. Tondello and S. J. Gross, Mater.
Chem., 2012, 22, 1620.
Abbreviations
TLL
FFA
Thermomyces lanuginosus lipase
Free fatty acids
23 X. Shi, W. Gu, B. Li, N. Chen, K. Zhao and Y. Xian, Microchim.
Acta, 2014, 181, 1.
NPs
Nanoparticles
HPLC
TLC
High performance liquid chromatography
Thin layer chromatography
24 M.-K. Liang, M. J. Lima, A. Sola-Rabada, M. J. Roe and
C. C. Perry, Chem. Mater., 2014, 26, 4119.
2
2
2
5 P. Yin, L. Chen, Z. Wang, R. Qu, X. Liu and S. Ren, Bioresour.
Technol., 2012, 110, 258.
6 E. Christensen and R. L. McCormick, Fuel Process. Technol.,
2014, 28, 339.
Acknowledgements
7 B. Abderrahim, M. Mercedes and A. Jos ´e , Fuel, 2007, 86,
We wish to thank DBT, New Delhi for Research Associateship
2596.
(RA) to one of us (PM).
28 V. G. Shashikant and R. Hiur, Bioresour. Technol., 2006, 97,
79.
3
2
3
9 M. J. Haas, Fuel Process. Technol., 2005, 86, 1087.
0 L. F. Bautista, G. Vicente, R. Rodr ´ı guez and M. Pacheco,
Biomass Bioenergy, 2009, 33, 862.
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