Lipase and Laccase Encapsulated on Zeolite Imidazolate Framework: Enzyme Activity and Stability from Voltammetric Measurements

Article abstract of DOI:10.1002/cctc.201801293

Abstract: Lipase (Pseudomonas fluorescens) and laccase (Trametates versicolor) were encapsulated on two zeolite imidazolate framework, ZIF-8 and ZIF-zni, materials using a one-pot synthesis-immobilization method in aqueous solution at room temperature. Th

Full text of DOI:10.1002/cctc.201801293

Accepted Article
Title: Lipase and Laccase Encapsulated on Zeolite Imidazolate
Framework: Enzyme Activity and Stability from Voltammetric
Measurements
Authors: Maryam Naseri, Federica Pitzalis, Cristina Carucci, Luca
Medda, Lida Fotouhi, Edmond Magner, and Andrea Salis
This manuscript has been accepted after peer review and appears as an
Accepted Article online prior to editing, proofing, and formal publication
of the final Version of Record (VoR). This work is currently citable by
using the Digital Object Identifier (DOI) given below. The VoR will be
published online in Early View as soon as possible and may be different
to this Accepted Article as a result of editing. Readers should obtain
the VoR from the journal website shown below when it is published
to ensure accuracy of information. The authors are responsible for the
content of this Accepted Article.
To be cited as: ChemCatChem 10.1002/cctc.201801293
Link to VoR: http://dx.doi.org/10.1002/cctc.201801293
A Journal of
www.chemcatchem.org

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
Lipase and Laccase Encapsulated on Zeolite Imidazolate
Framework: Enzyme Activity and Stability from Voltammetric
Measurements
Maryam Naseri,[a],[b] Federica Pitzalis, Cristina Carucci, Luca Medda, Lida Fotouhi, Edmond
[a]
[c]
[a]
[b]
Magner,[c] and Andrea Salis*[a]
Abstract: Lipase (Pseudomonas fluorescens) and laccase
Trametates versicolor) were encapsulated on two zeolite
Although lipases and laccases offer excellent activities, their use
is limited by some practical problems. In aqueous solution, the
activity of free laccase is very sensitive to the reaction medium
making it difficult to recover and reuse the enzyme without
(
imidazolate framework, ZIF-8 and ZIF-zni, materials using a one-pot
synthesis-immobilization method in aqueous solution at room
temperature. The synthesized immobilized biocatalysts (Lip@ZIF-8,
Lip@ZIF-zni, Lac@ZIF-8, and Lac@ZIF-zni) were characterized by
X-ray diffraction, scanning electron microscopy, Fourier transform
infrared spectroscopy, and thermogravimetric analysis. The
enzymatic activities of the four immobilized biocatalysts were
characterized via the electrochemical detection of the substrates, p-
nitrophenyl butyrate and 2,2-azinobis-3-ethylbenzthiazoline-6-
[15]
incurring additional losses due to the low operational stability.
In contrast, lipases, are quite robust enzymes which retain their
activity even in organic media. Nevertheless, lipases are
insoluble in such media and the low amount of water necessary
to keep the lipase in its active form can promote the formation of
large enzyme particle aggregates, which display very low
catalytic performance.[
1]
-
1
sulfonic acid. For Lip@ZIF-8 the specific activity was 91.9 U mg
-
1
Enzyme immobilization on solid supports can overcome these
and 123.1 U mg for Lip@ZIF-zni, while for Lac@ZIF-8 and
drawbacks.[
16]
In comparison to free enzymes, immobilized
-
1
-1
Lac@ZIF-zni, the activity was 51
U mg and 163 U mg ,
respectively, confirming that laccase retains a higher level of activity
when immobilized onto ZIF-zni than on ZIF-8. Lac@ZIF-8 was the
most stable system on storage (15 days at 5˚C), retaining 94% of
initial activity, while Lip@ZIF-zni biocatalyst had the optimal level of
reusability, retaining 40% of initial activity after five reaction cycles.
enzymes are more resistant and more robust to harsh
environmental conditions.[ For example, a laccase (Pleurotus
sajor-caju) immobilized on SBA-15 mesoporous silica could be
reused for several reaction cycles in the oxidation of phenolic
compounds.[ In non-aqueous media lipases immobilized on
supports of high specific surface area display high catalytic
efficiencies. Such types of materials allow the effective
dispersion of lipase molecules in a high surface area, with
retention of catalytic activity. Immobilized enzymes can be
prepared via different approaches including physical adsorption
or covalent binding to a support, entrapment, or cross-linking.[
The performance of the resulting immobilized biocatalyst may be
affected by the immobilization procedure as well as by the
nature of the solid support. A wide range of materials, e.g.
17]
7]
1. Introduction
16]
Lipases (EC 3.1.1.3; triacylglycerol hydrolases) are
enzymes that catalyze the hydrolysis of triacylglycerols. Since
triacylglycerols are poorly soluble in water, the hydrolytic
reaction normally occurs at the oil–water interface where lipases
mesoporous
silica,[
18]
magnetic
nanomaterials,
[19]
[
1]
are adsorbed. Lipases can also operate in non-aqueous
microspheres,[ and microchips have recently been reported
20]
[21]
media,[2] - i.e. organic solvents, ionic liquids, solvent free -
catalyzing esterification and transesterification (alcoholysis,
as enzyme supports.
[
1]
Metal organic frameworks (MOFs) are hybrid materials
which can be synthesized by combining metal ions or metal-
containing clusters (connectors) with anionic polyatomic organic
ligands (linkers) through coordination bonds.[ MOFs generally
offer unique characteristics such as high surface area, large
internal pore volumes, and uniform-structured cavities. They
have attracted considerable attention for potential applications,
acidolysis, interesterification) reactions. For these reasons
lipases are used for a wide range of purposes including
[
3]
[4]
[5]
pharmaceutical applications, food modification, biodiesel,
22]
and biopolymer production.[6]
Laccases
(E.C.
1.10.3.2,
p-diphenol:dioxygen
oxidoreductase) are multi-copper containing enzymes that
[
7]
catalyze the oxidation of a variety of phenolic compounds,
[
23]
[24]
[25]
[28]
diamines and aromatic amines.[8] Since laccases possess a wide
range of substrate specificity they can be used for applications
especially for gas storage,
catalysis,
separation,
biomedicine,[
26]
sensing,
[27]
and enzyme immobilization.
[
9]
[10]
[11]
Zeolitic imidazolate frameworks (ZIFs) are a class of MOFs in
such as bleaching of denim and paper,
decolorization,
2
+
2+
2+
[
12]
which tetrahedral metal centers (i.e. M = Zn , Co , and Cd )
are linked by imidazole (Im) or functionalized imidazole (RIm)
ligands.[ By combining metal ions with either Im or RIm, a
large number of crystalline ZIFs have been prepared which have
high thermal and chemical stability.[ ZIF-8, comprised of zinc
and 2-methyl imidazole (MeIm), is widely used. Recent works
and in bioremediation reactions such as the biodegradation of
industrial dyes[
wastewaters and contaminated soils.
13]
or the removal of phenolic pollutants in
29]
[
7-8, 14]
30]
[
a]
M. Naseri, Dr. F. Pitzalis, Dr. L. Medda, and Prof. A. Salis
Department of Chemical and Geological Sciences
University of Cagliari-CSGI and CNBS, Cittadella Universitaria,
S.S. 554 bivio Sestu, 09042 Monserrato, Cagliari, Italy.
E-mail: asalis@unica.it
have shown the potential of ZIF-8 for use as membranes for gas
permeation, capture, and separation,[
31]
in electric energy
storage devices,[ catalysis, and for water desalination.
32]
[29]
[33]
A
range of lipases (Burkholderia cepacia, Rhizomucor miehei,
Alcaligenes sp., Candida rugosa, Aspergillus niger, etc.)[ have
been immobilized on ZIF-8, while only one report to date on the
34]
[
[
b]
c]
M. Naseri and Prof. L. Fotouhi
Department of Chemistry
University of Alzahra
Teheran, Iran
immobilization of a laccase (Trametes hirsuta)[ . ZIF-zni, has a
15]
[29]
tetrahedral structure comprised of zinc and imidazole
and has
C. Carucci and Prof. E. Magner
not been examined extensively. However, ZIF-zni has very good
mechanical properties and is considered to be the most stable
Department of Chemical Sciences
Bernal Institute, University of Limerick
Limerick, Ireland
[35]
ZIF material. To date ZIF-zni has not been used as a support
for enzyme immobilization.
Supporting information for this article is given via a link at the end of
the document.
The activity of enzymatic reactions is quantified by
measuring the initial rate of the disappearance/formation of the
substrate/product. For lipases several assays
- based on
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
titrimetry, spectroscopy (photometry, fluorimetry, infrared),
2. Results and Discussion
chromatography,
radioactivity,
interfacial
tensiometry,
nephelometry, turbidimetry, conductimetry, immunochemistry,
and microscopy – have been reported.[36] Spectrophotometric or
chromatographic (HPLC) methods are used to measure the
activity of laccases.[36b] UV-Vis spectroscopy is the most widely
used method. For example, lipase activity can be determined by
measuring the rate of hydrolysis of p-nitrophenyl esters with
different chain-length fatty acids. The release of p-nitrophenol is
detected spectrophotometrically at 410 nm.[37] Similarly, laccase
activity is commonly determined through UV-Vis spectroscopy
using 2,2′-azinobis(3-ethylbenzthiazoline-6-sulfonic acid) (ABTS)
as substrate.[ Laccase catalyzes the oxidation of ABTS to a
green–blue colored radical cation ABTS , the formation of which
can be followed at 436 nm. Spectrophotometric activity assays
are suitable when free enzymes are tested. With immobilized
enzymes, spectrophotometric measurements can be difficult due
to the presence of immobilized enzyme particles. For this
reason, the development of alternative methods to measure the
activity of immobilized enzymes is of value. This is particularly
important for laccases, where the activity is predominantly
measured using spectrophotometric methods, but also for
lipases when used for biosensing.[34a]
2
.1. Samples characterization
The synthesized ZIF-8 and ZIF-zni materials and the lipase
Lip@ZIF-8, Lip@ZIF-zni) and laccase (Lac@ZIF-8, Lac@ZIF-
(
zni) immobilized biocatalysts were characterized using X-ray
diffraction (Figure 1). The immobilization of laccase and lipase
on ZIF-8 resulted in slight modifications to the XRD pattern when
compared to that of ZIF-8. Even though the peaks occur at the
same position for all ZIF-8 samples, a peak splitting occurred
with Lip@ZIF-8 at 18° and 27° which is likely due to the
immobilization process. The diffractogram of pure ZIF-8 has a
different pattern to that of the simulated ZIF-8 pattern reported in
38]
+
•
[40]
2+
the literature.
This may be due to the concentration of Zn
ions and 2-MIm which could play key roles in the morphology
and XRD patterns of the material. Previous studies have
reported different XRD patterns for ZIF-8 materials prepared
using different 2-MIm concentrations.
samples obtained using a Zn:2-MIm molar ratio ranging from 20
to 100
:60, 1:80, 1:100) had a crystalline ZIF-8 SOD like structure,
whereas mixed phases associated with the formation of
Zn(OH) , Zn(OH)NO , and Zn (OH) (NO (H O) were obtained
at lower ratios (Zn:2-MIm ratio 1:20). We recently observed
different XRD patterns for ZIFs samples obtained with 1:40 and
:4 Zn :2-MIM ratios.
at Zn:2-MIm 1:40 molar ratio, the optimal catalytic performance
of Lipase@ZIF-8 biocatalyst was observed at a low Zn:2-MIm
[34c, 41]
XRD patterns of ZIF
[41b]
demonstrated that high molar Zn:2-MIm ratios (1:40,
1
2
3
5
8
3
)
2
2
2
In this work, lipase (Pseudomonas fluorescens) (Lip) and
laccase (Trametates versicolor) (Lac) were encapsulated in ZIF-
[41b]
8
and ZIF-zni - through a one-pot synthesis-immobilization
2+
[41c]
1
Although a pure ZIF-8 phase resulted
method performed in aqueous solution at room temperature - to
obtain Lip@ZIF-8, Lip@ZIF-zni, Lac@ZIF-8, and Lac@ZIF-zni
biocatalysts (Scheme 1). In comparison to the mulit-step and
(
=1:4) molar ratio. Despite the fact that the structure of the latter
long synthesis of ordered mesoporous silica,[18a,
39]
, the
sample was not similar to that of ZIF-8, the material with a Zn:2-
MIm ratio of 1:4 was chosen as the optimal material due to the
higher levels of activity and stability. Figure 1B shows the XRD
pattern of ZIF-zni. In contrast to ZIF-8 (Figure 1A), the XRD
pattern is similar to that previously reported for ZIF-zni
structure.
preparation of ZIF-based biocatalysts is simple and rapid.
Indeed, the one-pot synthesis/immobilization of enzyme@ZIF-8
can be performed in 12 hours, and that of enzyme@ZIF-zni in
30 min. The synthesized enzyme@ZIF biocatalysts were
characterized using X-ray diffraction (XRD), scanning electron
microscopy (SEM), Fourier transform infrared (FTIR)
spectroscopy, and thermogravimetric analysis (TGA). The
activity and the stability of the biocatalysts were examined using
electrochemical assays.
[29, 40a]
The XRD patterns of Lac@ZIF-zni and Lip@ZIF-
zni samples did not show any significant differences when
compared to that of ZIF-zni.
Figure 1. XRD patterns of A) ZIF-8 and enzyme@ZIF-8 and B) ZIF-zni and
enzyme@ZIF-zni samples.
Figure 2 shows the thermogravimetric analysis of ZIF and
enzyme@ZIF samples. Mass loss data in the different
temperature ranges (< 230°C; 230-500°C; > 500°C) are listed in
Table 1. Both ZIF-8 and ZIF-zni samples displayed almost no
appreciable mass loss up to 500°C. The mass loss below 230°C
observed for enzyme@ZIF samples is likely due to the loss of
water and unreacted reagents.[ Small mass losses for the
ZIF-8 (4.73%) and ZIF-zni (2.50%) were observed in the range
42]
230-500°C. In the same range (230-500°C) higher mass losses
were reported for all enzyme@ZIF samples, that is: 14.26%
(Lac@ZIF-8), 33.82 % (Lip@ZIF-8), 3.26 % (Lac@ZIF-zni), and
10.28 % (Lip@ZIF-zni). This is ascribed to the thermal
Scheme 1. Schematic representation of the synthesis of the enzyme@ZIF
biocatalysts.
decomposition of the encapsulated enzymes and other organic
compounds (i.e. carbohydrates) present in the commercial
enzymatic preparations.[
43]
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
We ascribe these discrepancies to the presence of other
materials in the enzyme preparations, and to a different behavior
to the thermal treatment between ZIF-8 and ZIF-zni which may
have different thermal stability in the temperature range 230-
500°C. However, for both methods, the enzyme loading
decreased in the sequence: Lip@ZIF-8 > Lac@ZIF-8 and
Lip@ZIF-zni > Lip@ZIF-8. The encapsulation efficiency (EE),
the amount of immobilized enzyme relative to the total enzyme
amount in the immobilizing solution, is shown in Figure 3B.
Laccase was fully immobilized in the ZIF-8 material with an EE
of 100%, whereas an EE of 84.3% was obtained for ZIF-zni. The
encapsulation of lipase was less efficient, with EE values of
Figure 2. TGA curves of A) enzyme@ZIF-8 and B) enzyme@ZIF-zni samples.
Table 1. Mass loss (%) of ZIF and enzyme@ZIF samples and enzyme loading
30.0% and 33.8% for Lip@ZIF-8 and Lip@ZIF-zni, respectively.
Hence, the two enzymes behaved differently, that is,
immobilized lipases gave high loadings and low EE, whereas
immobilized laccases resulted low loadings and high EE.
enzyme@ZIF
(L )
Sample
Mass loss (%)
T < 230˚C
0.68
Mass loss (%)
230˚C < T< 500˚C
4.73
Mass loss (%)
T > 500˚C
57.44
L
enzyme@ZIF
(mg g )
-
1
ZIF-8
-
2.3 Electrochemical determination of lipase activity
Lac@ZIF-8
Lip@ZIF-8
ZIF-zni
66.02
40.70
2.46
14.26
33.82
2.43
54.51
61.16
58.7
24.6
60.2
-
The catalytic activities of the immobilized lipases, Lip@ZIF-8
and Lip@ZIF-zni, were measured by cyclic voltammetry. To
[45]
detect the oxidation of p-nitrophenol (eq. 2)
Figure 4A
displays cyclic voltammograms of PNP at different
concentrations in 0.1 M acetate buffer at pH 5. As shown in
Figure 4B, a good linear trend between the oxidation current and
the concentration of PNP, over the range 0.045-0.675 mM, was
observed. To measure the catalytic activity of Lip@ZIF-8 and
Lip@ZIF-zni samples, the cyclic voltammograms were recorded
at various times after adding the biocatalyst to the p-nitrophenyl
butyrate solution. A peak around 0.9 V appeared during the
Lac@ZIF-zni
Lip@ZIF-zni
47.50
29.32
3.26
29.07
50.39
16.3
48.8
10.28
Finally, the mass loss measured above 500°C is likely due to
the decomposition of the ZIF structures.[42] Additional
characterization details, e.g. scanning electron microscopy
Figure S1 and S2) and Fourier transform infrared spectroscopy
Figure S3), are reported in supporting information. In a recent
work Patil and Yadav investigated the encapsulation of
Trametes hirsute laccase on ZIF-8.[15] They used FTIR
spectroscopy to study the conformational changes of the
enzyme after the immobilization. A similar study was not
possible with the commercially available lower purity enzymatic
preparations used here.
anodic scan, corresponding to the oxidation of PNP (Figure
(
(
45]
4
C).[ The catalytic activities of the free lipase, Lip@ZIF-8, and
Lip@ZIF-zni are shown in Figure 5. The specific activities
-
1
-1
-1
-1
(
μmolPNP min mgenzyme = U mg ) were 110.4 U mg (free
-1
-1
lipase), 91.9 U mg (Lip@ZIF-8), and 123.1 U mg (Lip@ZIF-
zni).
2.2. Determination of enzyme loading of immobilized
biocatalysts
Enzymatic loadings of the enzyme@ZIF samples, quantified by
mean of the Bradford assay,[44] are shown in Figure 3. Lip@ZIF
samples showed higher enzyme loadings than Lac@ZIF: 60.2
mg g (Lip@ZIF-8), 24.6 mg g (Lac@ZIF-8), 48.8 mg g-
(
-
1
-1
1
-1
Lip@ZIF-zni), and 16.3 mg g (Lac@ZIF-zni). When the TGA
data (Table 1), particularly the mass loss (%) in the range 230-
50°C, was compared with the enzyme loading data (Figure 3A),
a mass loss of 33.82% (338.2 mg g , Lip@ZIF-8) was obtained
corresponding to an enzyme loading 5.6 times higher than that
obtained by the Bradford assay (60.2 mg g ). A similar
discrepancy was observed for Lac@ZIF-8 (5.8 times), whereas
the loading was only 2 fold higher for the Lip@Zif-zni and
Lac@ZIF-zni samples.
3
-1
-1
Figure 4. Determination of lipase activity through cyclic voltammetry. A) cyclic
voltammograms at different p-nitrophenol concentrations. B) calibration curve
2
of p-nitrophenol. (I
p
(µA) = 13.02 [PNP](mM) + 1.41 and R = 0.994) C) Activity
of lipase AK@ZIF-8: cyclic voltammograms at different reaction times through
cyclic voltammetry. D) Activity of lipase AK@ZIF-8: concentration of p-
nitrophenol versus time.
Figure 3. A) Protein loading and B) encapsulation efficiency of Lip@ZIF-8/zni
and Lac@ZIF-8/zni biocatalysts.
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
Although enzyme immobilization can sometimes result in
decreased catalytic activity,[28b] the obtained specific activities of
the immobilized lipases are comparable to that of the free
lipase.The highest specific activity was obtained for Lip@ZIF-zni
conformational changes due to unfavorable enzyme-support
interactions. The results indicate the lipase is better adapted
[49]
to the ZIF structures than the laccase. Moreover, the structure of
ZIF appears to play an important role. Indeed, although the
enzyme loading of Lac@ZIF-8 is about 1.5 times higher than
that of the Lac@ZIF-zni, the catalytic activity of Lac@ZIF-8 is
about 1.6 lower than that of the Lac@ZIF-zni. The activity of
laccase was preserved to a more significant extent in Lac@ZIF-
zni likely because of more favorable interactions between the
enzyme and the ZIF-zni in comparison to ZIF-8. The enzymatic
activities of Lac@ZIF-8/zni samples were also measured using
the spectrophotometric ABTS assay (Figure S6 supporting
information file).[
(
Figure 5). While Lip@ZIF-zni possesses a lower enzyme
loading than Lip@ZIF-8, the ZIF-zni matrix is more favorable
than ZIF-8 in retaining lipase activity. The enzymatic activity of
Lip@ZIF-zni sample was also measured spectrophotometrically
through the p-nitrophenyl butyrate assay (Figure S5 Supporting
[36a]
-1
Information file).
The specific activity was 114±21U mg
(
Lip@ZIF-zni) which is comparable with that obtained by cyclic
-1
voltammetry (123±25 U mg ). This result confirms the validity of
the electrochemical method to measure the activity of lipase.
38]
Recently Ma et al. used a similar electrochemically based lipase
ZIF-8.[
34a]
assay to successfully determine the concentration of
the pesticide methyl parathion.
Typically, lipases in oil in water emulsions undergo a
structural change in the presence of hydrophobic interfaces at
the oil:water interface. An amphiphilic polypeptide chain called
the ‘lid’, that in the closed conformation covers the active site,
moves allowing the substrate to bind to the catalytic triad in the
active site. As lipase is still active after encapsulation in ZIF, it is
clear that the conformational change required for enzyme
activation can still occur.
Figure 6. Determination of laccase activity through DPV. A) Differential pulse
Figure 5. Specific activity of free lipase (Pseudomonas fluorescens),
Lip@ZIF-8 and Lip@ZIF-zni biocatalysts measured by cyclic voltammetry.
voltammograms at different ABTS concentrations. B) calibration curve of ABTS
2
for investigation of activity of laccase. (I
p
(µA) = 5.40 [ABTS](mM) + 0.37 and R =
0.999) C) Activity of LAC@ZIF-8: voltammograms at different reaction times
through DPV. D) Activity of LAC@ZIF-8: concentration of ABTS versus time.
2.4 Electrochemical determination of laccase activity
The catalytic activity of the immobilized laccases was measured
by following the decrease in the concentration of ABTS using
differential pulse voltammetry (DPV). DPV was chosen because
of its high sensitivity. Fabbrini et al. reported that laccases
+
oxidize ABTS to a radical-cation (ABTS ˙) and to a dication
2
+ [46]
(
ABTS ).
In oyther reports, only the radical monocation is
[
47]
[48]
formed. Gonzalez et al. reported that laccase catalyzes the
formation of a stable monocation (ABTS ˙) with no evidence of
+
2
+
dication (ABTS ) formation. Figure 6A shows the DPV
responses of different concentrations of ABTS in 0.1 M acetate
buffer solution (pH 5). A good linear relationship, was obtained,
in the range of 0.1-0.6 mM (Figure 6B).. As shown in Figure 6C,
the anodic current decreased with time after addition of
Lac@ZIF-8 or Lac@ZIF-zni The catalytic activity of Lac@ZIF
samples was then estimated by calculating the initial rate of
decrease in the concentration of ABTS. The specific activities of
free laccase, Lac@ZIF-8, and Lac@ZIF-zni samples obtained
through DPV are shown in Figure 7. Specific activities were
Figure 7. Specific activities of free laccase (Trametates versicolor), Lac@ZIF-
8, and Lac@ZIF-zni biocatalysts measured by differential pulse voltammetry.
Specific activities were 163±23 U mg-1 and 51±3U mg-1 for
Lac@ZIF-zni and Lac@ZIF-8, respectively. Although the specific
activities obtained using the spectrophotometric assay are
higher than those obtained using the DPV method, both
methods confirm that immobilization of laccase on ZIF-ini yields
a more active enzyme than on ZIF-8.
-1
-1
7
4±3 U mg (free laccase), 9.1±0.5 U mg (Lac@ZIF-8), and
-1
14.6±1 U mg (Lac@ZIF-zni).
In contrast with the results obtained for Lip@ZIF, the catalytic
activities of immobilized laccases were lower than that of the
free laccase. The activities of most enzymes are negatively
2.5 Determination of enzyme@ZIF sample stability
affected by the immobilization procedure as
a result of
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
Soluble enzymes are generally immobilized for industrial scale
applications, where many cycles of high yield processes are
required.[
Ouyang et al.[51] showed that laccase immobilized on Cu– MOF
retained a 48% of activity after the seventh reaction cycle. Our
biocatalysts were less stable to recycling but more stable to
storage than the reported Lac@Cu-MOF. Since our and Ouyang
et al. laccases were of different origin and the used MOFs were
also different, it is difficult to recognize the cause of the
observed activity loss.
50]
3
Conclusions
In summary, we described a rapid and facile approach to
immobilize lipase from Pseudomonas fluorescens and laccase
from Trametates versicolor onto two kinds of zeolitic imidazolate
frameworks through an in-situ encapsulation in aqueous solution
and at room temperature. Enzyme@ZIF-8 synthesis was carried
out in 12 hours whereas that of enzyme@ZIF-zni took only 30
min, significantly faster than the time required to prepare other
immobilized biocatalysts. The structures of the synthesized
enzyme@ZIF systems were characterized using XRD, TGA, and
FTIR. The enzymatic activities of the free enzymes and
enzyme@ZIF were measured using voltammetric methods. For
Lip@ZIF-zni, the specific activity obtained by voltammetry was
comparable with that obtained by UV-Vis spectrophotometry. On
the contrary, the specific activities of Lac@ZIF-8 and Lac@ZIF-
zni obtained through DPV were lower than those obtained
spectrophotometrically. Nonetheless, both methods confirm that
ZIF-zni is a better enzyme support than ZIF-8. Furthermore,
Lip@ZIF-zni, Lac@ZIF-8, and Lac@ZIF-zni had good stability
with retention of activity after 15 days storage at 4 ˚C. Finally,
the operational stability to recycling of the Lip@ZIF-zni,
Lac@ZIF-8, and Lac@ZIF-zni biocatalysts was evaluated
confirming that lipases are generally more robust enzymes than
laccases. To the best of our knowledge this the first work where
the dense and highly stable ZIF-zni structure was used for
enzyme immobilization. The electrochemical activity assays
used here are a very promising alternative to the UV-Vis
spectrophotometric assay and may find application in the
development of electrochemical biosensors.
Figure 8. Determination of A) storage stability and B) recycling stability
of Lip@ZIF-zni biocatalyst measured by cyclic voltammetric PNP assay.
In this regard, the stability of immobilized enzymes to storage
and to reuse needs to be evaluated. Figure 8 shows the storage
stability and the recycling of Lip@ZIF-zni sample. This
biocatalyst was chosen since it had the highest enzyme loading
and specific activity. The biocatalyst was stored at 4 ˚C for 15
days, with retention of 70.3% of activity (Figure 8A). The storage
conditions did not significantly affect the relative activity of the
biocatalyst. The operational stability to recycling was also
investigated. The Lip@ZIF-zni biocatalyst retained 40% of its
initial activity at the fifth reaction cycle (Figure 8B). We recently
investigated the long-term stability and recycling of
Pseudomonas fluorescens lipase immobilized on ZIF-8.[41c] After
15 days storage, the immobilized lipase retained 99% of its initial
activity. Similarly to the present case, also Lip@ZIF-8 could be
recycled five times with about 40% of activity retention at the
[41c]
fifth reaction cycle.
Although Lip@ZIF-zni was less stable to
storage than Lip@ZIF-8, both samples showed practically the
same stability to recycling. Figure 9 shows the storage and the
recycling stabilities of Lac@ZIF-8 and Lac@ZIF-zni biocatalysts
determined through DPV. The biocatalysts were stored at 4 ˚C
for 15 days. Activity retentions of 94% and 76% were obtained
for Lac@ZIF-8 and Lac@ZIF-zni, respectively (Figure 9A). The
storage conditions did not significantly affect the relative activity
of the biocatalysts. Hence, although the immobilization
procedure had a negative effect on activity in comparison with
the free enzyme, it permits to obtain a good storage stability.
Ouyang et al.[ investigated the long-term stability and recycling
of laccase from white rot fungi immobilized on Cu–MOF. They
found that only 19% of the initial immobilized laccase activity
was retained after storage at 4 ˚C for 3 weeks. The operational
stability to recycling of the two immobilized biocatalysts was then
evaluated (Figure 9B). The Lac@ZIF-8 biocatalyst retained only
Experimental Section
51]
Chemicals
-1
Laccase from Trametates versicolor (1.07 U mg ), 2,2’-azinobis-
3-ethylbenzothiazoline-6-sulfonate) diammonium salt (≥ 98%),
(
p-nitrophenyl butyrate (≥ 98%), p-nitrophenol (≥ 99%), zinc
nitrate hexahydrate (98%), 2-methylimidazole (2-MIm, 99%),
imidazole (≥ 99%), Bradford reagent, 2-propanol were
purchased from Sigma-Aldrich (Milan, Italy). Lipase from
Pseudomonas fluorescens was purchased from Amano
Enzymes Inc (Japan) and was used without any further
purification.
12.6% of its initial activity at the fifth reaction cycle. Instead,
Lac@ZIF-zni biocatalyst displayed a better performance, as it
could be reused five times with activity retention of 25% at the
fifth reaction cycle (Figure 9B).
Preparation of ZIF-8 and enzyme@ZIF-8 biocatalysts
The synthesis of enzyme@ZIF-8 composites was carried out
according with previously described methods.[
52]
Typically, a
mass of 50 mg of enzyme powder was dissolved in 2 mL of
buffer acetate 0.1 M (pH 5) and 60 mg of 2-methylimidazole
were then dissolved in 5 mL of milli-Q water. A mass of 66 mg
3 2 2
Zn(NO ) ×6H O was dissolved in 5 mL of milli-Q water and
mixed with the enzymatic and 2-methylimidazole solutions. The
resulting solution was incubated at room temperature for 12 h,
then a white precipitate was collected by centrifugation at 4500
Figure 9. Determination of A) storage stability of Lac@ZIF-8/zni; B) stabiltiy to
recycling of Lac@ZIF-8/zni.
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
rpm and washed three times with milli-Q water. Finally, the white
powder was dried at room temperature and stored at 4 °C. The
synthsis of the pure ZIF-8 was carried out in the same way
except for the fact that no enzyme was added.
and 100 µL of 2-propanol were added to a volume of 50 μL of
free lipase solution or a defined amount of immobilized lipase.
Lipase catalyzes the hydrolysis of p-nitrophenyl butyrate to p-
nitrophenol (PNP):
푙푖푝푎푠푒
Preparation of ZIF-zni and enzyme@ZIF-zni biocatalysts
p-nitrophenyl butyrate → ꢀp-nitrophenol + butyric acid
(2)
The synthesis of enzyme@ZIF-zni composites was carried out
[
52]
according previously described methods.
2
Briefly, a volume of
mL of an enzymatic solution (10 mg mL ) in acetate buffer (0.1
M, pH 5) was added to 10 mL of an imidazole (0.851g) solution
in Milli-Q water, and 1 mL of a Zn(NO solution (0.31 M). The
The formation of PNP as a function of time was followed by
reading the CV peak current of its anodic oxidation.
activity is defined as the amount of PNP formed per minute (U =
µmolPNP min ). The enzymatic activities of free Trametates
versicolor laccase, Lac@ZIF-8, and Lac@ZIF-zni were obtained
by differential pulse voltammetry. A volume of 4.9 mL of 0.1 M
acetate buffer (pH 5), and 100 µL of ABTS 0.02 M were added
to 50 μLvolume of free laccase solution or a weighed amount of
immobilized laccase. According to eq. 3, laccase catalyzes the
-1
[45]
Lipase
)
2
-1
3
obtained mixture was stirred at room temperature for 30 min,
and then a white solid was collected by centrifugation at 4500
rpm and washed three times with Milli-Q water. Finally, the
obtained sample was dried at room temperature and stored at
4°C. The pure ZIF-zni synthesis was carried out in the same way
except for the fact that no enzyme was added.
+
oxidation of ABTS to ABTS ˙. The concentration of ABTS as a
function of time was followed measuring the DPV peak current
for the anodic oxidation of ABTS.
Characterization of ZIF and enzyme@ZIF samples
X-ray diffraction was performed through
PAnalytical diffractometer using Cu-Kα radiation. The
diffractograms were obtained using a 2θ step size of 0.04° from
a X’PERT Pro
푙푎푐푐푎푠푒
ABTS → ꢀABTS+˙
(3)
4
to 45° and accumulation time 20 s. Thermogravimetric
Laccase activity is defined as the amount of ABTS
transformed per minute (µmolABTS min ).
analysis measurements were performed in TGA 4000 Perkin
-1
Elmer from 30 °C to 900 °C with a ramp rate of 10°C min-1 under
Specific activities of the biocatalysts are defined as the
−
1
continous flow of nitrogen gas at flow rate = 20 mL min .
Scanning electron microscopy studies at various magnifications
were carried out in a HITACHI SU-70 at 10 kV.
-
1
-1
activity per mass (mg) of enzyme (U mgenzyme = µmol min
-1
mgenzyme ). The mass of enzyme in mg (menzyme
immobilized preparation is calculated by the formula:
)
in the
Determination of enzyme loading
menzyme = menzyme@ZIF x Lenzyme@ZIF
(4)
The loading of enzyme@ZIF biocatalysts was performed by
measuring the enzyme concentrations of the initial, final and
washing solutions. Although not always correct,[43] here we
assume that the commercially available enzymatic (lipase and
laccase) preparations contain only the selected enzyme and no
other inactive enzymes. The concentrations of enzyme in the
where, menzyme@ZIF is the mass of the immobilized biocatalyst (g)
used for the activity assay, and Lenzyme@ZIF is the enzyme loading
as defined above. Blank reactions carried out with ZIF-8 and
ZIF-zni samples showed that, in the absence of enzymes, the
supports were not catalytically active.
Storage and recycling stabilities were investigated by means
of the electrochemical methods as described above. For storage
stabilities measurements, the wet biocatalysts were stored at 4
different
solutions
were
obtained
through
using BSA as the
UV-Vis
[
44]
spectrophotometry by the Bradford assay,
standard. the enzyme loading (Lenzyme@ZIF = mgenzyme
was determined by the following equation:
-1
enzyme@ZIF
g )
˚
C and dried under vacuum the day before the activity
(
[E]f−[퐸]i)V−[퐸]w[V]w
푚
Lenzyme@ZIF
=
(1)
measurements. For recycling measurements, the biocatalyst
was recovered by centrifugation (4500 rpm for 2 min) and
washed with acetate buffer (0.1 M, pH 5) before the addition of
the fresh substrate solution. The change of loading due to
enzyme leaching after each reuse was tested through the
Where, [E]
concentrations (mg mL ), respectively; V is the volume of the
enzyme solution (mL); [E] and [V] are the enzyme
f
and [E]
i
-
are the final and initial enzyme
1
w
w
th
Bradford assay. A negligible leaching was obtained after the 5
concentration and the volume of the washing solution; m is the
mass of enzyme@ZIF biocatalyst (g).
reuse suggesting that activity decrease was due to enzyme
inactivation. All activity and stability tests were performed in
triplicate at a minimum.
Determination of enzymatic activities through voltammetric
techniques
An Autolab Potentiostat/Galvanostat was used for the
electrochemical measurements. The electrochemical cell was
constituted by a glassy carbon electrode (diameter 2 mm), an
Ag/AgCl electrode and a platinum wire as working, reference,
and counter electrodes, respectively. Convectional conditions of
the solutions were achieved by means of magnetic stirring. The
Acknowledgements
FIR 2018, Fondazione di Sardegna/Regione Autonoma della
Sardegna (CUP F72F16003070002), FFABR 2017 (MIUR) are
acknowledged for financial support. MN would like to express
their deep gratitude to Alzahra University for supporting this
study. FP is thankful to Agenzia delle Dogane e dei Monopoli
glassy carbon electrode surface was pretreated with
mechanical polishing step (0.5 µm alumina slurry on micro cloth
pads) followed by sonication in different solutions (HNO 30%,
a
(
Italy) for the financial support during her PhD. We thank the
3
anonymous reviewers, whose comments have helped to
improve the presentation of our work.
ethanol and pure water) for 2 min. The electrode was then dried
at room temperature.
The enzymatic activity of the free Pseudomonas fluorescens
lipase, Lip@ZIF-8, and Lip@ZIF-zni immobilized biocatalysts
were measured by cyclic voltammetry. A solution of 5 mL of 0.1
M acetate buffer (pH 5), 100 µL of p-nitrophenyl butyrate 0.02 M,
Conflict of interest
The authors declare no conflict of interest.
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
[
[
26]
27]
S. Wuttke, A. Zimpel, T. Bein, S. Braig, K. Stoiber, A.
Vollmar, D. Müller, K. Haastert‐Talini, J. Schaeske, M.
Stiesch, Adv. Healthc. Mater. 2017, 6.
a) I. Stassen, B. Bueken, H. Reinsch, J. Oudenhoven, D.
Wouters, J. Hajek, V. Van Speybroeck, N. Stock, P.
Vereecken, R. Van Schaijk, Chem. Sci. 2016, 7, 5827-
5832; b) X.-L. Zhao, D. Tian, Q. Gao, H.-W. Sun, J. Xu,
X.-H. Bu, Dalton Trans. 2016, 45, 1040-1046.
Keywords: • Enzyme activity • Laccase • Lipase • Voltammetry •
Zeolite imidazolate framework
[
[
1]
2]
P. Adlercreutz, Chem. Soc. Rev. 2013, 42, 6406-6436.
C. D. Anobom, A. S. Pinheiro, R. A. De-Andrade, E. C.
Aguieiras, G. C. Andrade, M. V. Moura, R. V. Almeida, D.
M. Freire, BioMed Res. Int. 2014, 2014.
[
[
[
[
3]
4]
5]
6]
V. Gotor-Fernández, R. Brieva, V. Gotor, J. Mol. Catal. B
Enzym. 2006, 40, 111-120.
R. Aravindan, P. Anbumathi, T. Viruthagiri, Indian J.
Biotechnol. 2007, 6, 141-158.
A. Salis, M. F. Casula, M. S. Bhattacharyya, M. Pinna, V.
Solinas, M. Monduzzi, ChemCatChem 2010, 2, 322-329.
G. Sandoval, I. Rivera, K. A. Barrera‐Rivera, A. Martínez‐
Richa, in Macromolecular symposia, Vol. 289, Wiley
Online Library, 2010, pp. 135-139.
A. Salis, M. Pisano, M. Monduzzi, V. Solinas, E. Sanjust, J.
Mol. Catal. B Enzym. 2009, 58, 175-180.
J. Subramanian, T. Ramesh, M. Kalaiselvam, Int. J.
Pharm. Biol. Arch. 2014, 5, 8-16.
[28]
a) E. Gkaniatsou, C. Sicard, R. Ricoux, J.-P. Mahy, N.
Steunou, C. Serre, Mater Horiz. 2017, 4, 55-63; b) V.
Gascón, C. Carucci, M. B. Jimenez, R. M. Blanco, M.
Sánchez‐Sánchez, E. Magner, ChemCatChem 2017, 9,
1182-1186.
T. Hikov, C. A. Schröder, J. Cravillon, M. Wiebcke, K.
Huber, Phys. Chem. Chem.Phys. 2012, 14, 511-521.
A. Phan, C. J. Doonan, F. J. Uribe-Romo, C. B. Knobler,
M. O’keeffe, O. M. Yaghi, Acc. Chem. Res. 2010, 43, 58-
67.
a) A. Knebel, C. Zhou, A. Huang, J. Zhang, L. Kustov, J.
Caro, Chem. Eng. Technol. 2018, 41, 224-234; b) Z. Lai,
Curr. Opin. Chem. Eng. 2018, 20, 78-85; c) X. Gong, Y.
Wang, T. Kuang, ACS Sustain. Chem. Eng. 2017, 5,
11204-11214.
[29]
[30]
[
[
[
[
[
[
[
[
7]
[31]
8]
9]
E. Forgacs, T. Cserhati, G. Oros, Environ. Int. 2004, 30,
9
53-971.
A. P. Virk, P. Sharma, N. Capalash, Biotechnol. Prog.
012, 28, 21-32.
10]
11]
12]
13]
14]
[32]
[33]
[34]
X. Xu, H. Wang, J. Liu, H. Yan, J. Mater. Sci. Mater.
Electron. 2017, 28, 7532-7543.
Z. Hu, Y. Chen, J. Jiang, J. Chem. Phys. 2011, 134,
134705.
2
E. Grassi, P. Scodeller, N. Filiel, R. Carballo, L. Levin, Int.
Biodeterior. Biodegradation 2011, 65, 635-643.
M. Kuddus, B. Joseph, P. W. Ramteke, Biocatal. Agric.
Biotechnol. 2013, 2, 333-338.
a) B. Ma, L.-Z. Cheong, X. Weng, C.-P. Tan, C. Shen,
Electrochim. Acta 2018, 283, 509-516; b) S. S. Nadar, V.
K. Rathod, Enzyme Microb. Technol. 2018, 108, 11-20; c)
K. Liang, R. Ricco, C. M. Doherty, M. J. Styles, S. Bell, N.
Kirby, S. Mudie, D. Haylock, A. J. Hill, C. J. Doonan, Nat.
Commun. 2015, 6, 7240; d) J. Zhao, D. Yang, J. Shi, J. Li,
S. Zhang, Y. Wu, Z. Jiang, Ind. Eng. Chem. Res. 2018; e)
H. He, H. Han, H. Shi, Y. Tian, F. Sun, Y. Song, Q. Li, G.
Zhu, ACS Appl. Mater. Interfaces 2016, 8, 24517-24524; f)
M. Adnan, K. Li, L. Xu, Y. Yan, Catal. 2018, 8, 96; g) L.-Z.
Cheong, Y. Wei, H. Wang, Z. Wang, X. Su, C. Shen, J.
Nanopart. Res. 2017, 19, 280; h) M. Adnan, K. Li, J. Wang,
L. Xu, Y. Yan, Int. J. Mol. Sci.2018, 19, 1424.
P. Zucca, G. Cocco, F. Sollai, E. Sanjust, Biocatal. 2015, 1,
8
2-108.
R. Chandra, P. Chowdhary, Environ. Sci. Process. Impact
015, 17, 326-342.
2
[
[
15]
16]
P. D. Patil, G. D. Yadav, Chem. Select 2018, 3, 4669-4675.
R. A. Sheldon, S. van Pelt, Chem. Soc. Rev. 2013, 42,
6223-6235.
[
[
17]
18]
R. Ahmad, M. Sardar, Biochem. Anal. Biochem. 2015, 4, 1.
a) E. Magner, Chem Soc. Rev. 2013, 42, 6213-6222; b) Z.
Zhou, M. Hartmann, Chem. Soc. Rev. 2013, 42, 3894-
3912; c) A. Salis, L. Medda, F. Cugia, M. Monduzzi,
Colloids Surf., B 2016, 137, 77-90; d) F. Pitzalis, M.
Monduzzi, A. Salis, Microporous Mesoporous Mater. 2017,
[35]
[36]
T. D. Bennett, J. C. Tan, S. A. Moggach, R. Galvelis, C.
Mellot‐Draznieks, B. A. Reisner, A. Thirumurugan, D. R.
Allan, A. K. Cheetham, Chem. Eur. J. 2010, 16, 10684-
10690.
a) F. Beisson, A. Tiss, C. Rivière, R. Verger, Eur. J. Lipid
Sci. Technol. 2000, 102, 133-153; b) A. Li, Y. Zhu, L. Xu,
W. Zhu, X. Tian, Afr. J. Biochem.Res. 2008, 2, 181-183; c)
M. Stoytcheva, G. Montero, R. Zlatev, J. A Leon, V.
Gochev, Curr. Anal.Chem. 2012, 8, 400-407.
2
41, 145-154.
a) Y. Cao, L. Wen, F. Svec, T. Tan, Y. Lv, Chem. Eng. J.
016, 286, 272-281; b) H. Vaghari, H. Jafarizadeh-Malmiri,
[
19]
2
M. Mohammadlou, A. Berenjian, N. Anarjan, N. Jafari, S.
Nasiri, Biotechnol. Lett. 2016, 38, 223-233.
a) G. Peng, X. Hou, B. Liu, H. Chen, R. Luo, RSC Adv.
[
[
[
[
20]
21]
22]
23]
2016, 6, 91431-91439; b) H. Liu, K. Guo, C. Duan, X.
Chen, Z. Zhu, Mater. Sci. Eng. C 2016, 64, 243-248.
a) A. Kecskemeti, A. Gaspar, Talanta 2017, 166, 275-283;
b) Y. Tang, S. Liu, R. Pi, Z. Cheng, RSC Adv. 2016, 6,
[37]
[38]
R. Gupta, P. Rathi, N. Gupta, S. Bradoo, Biotechnol. Appl.
Biochem. 2003, 37, 63-71.
M. Terrón, M. López-Fernández, J. Carbajo, H. Junca, A.
Téllez, S. Yagüe, A. Arana-Cuenca, T. González, A.
González, Biochim. 2004, 86, 519-522.
20867-20875.
a) J. L. Rowsell, O. M. Yaghi, Microporous Mesoporous
Mater. 2004, 73, 3-14; b) J. Lei, R. Qian, P. Ling, L. Cui, H. [39]
Ju, TrAC Trends Anal. Chem. 2014, 58, 71-78.
a) D. DeSantis, J. A. Mason, B. D. James, C. Houchins, J.
R. Long, M. Veenstra, Energy Fuels 2017, 31, 2024-2032;
b) D. De, T. K. Pal, S. Neogi, S. Senthilkumar, D. Das, S.
S. Gupta, P. K. Bharadwaj, Chem. Eur. J. 2016, 22, 3387-
M. Hartmann, X. Kostrov, Chem. Soc. Rev. 2013, 42,
6277-6289.
[40]
a) D. W. Lewis, A. R. Ruiz-Salvador, A. Gómez, L. M.
Rodriguez-Albelo, F.-X. Coudert, B. Slater, A. K.
Cheetham, C. Mellot-Draznieks, CrystEngComm 2009, 11,
2272-2276; b) D. Fairen-Jimenez, S. Moggach, M.
Wharmby, P. Wright, S. Parsons, T. Duren, J. Am. Chem.
Soc. 2011, 133, 8900-8902.
a) Q. Shi, Z. Chen, Z. Song, J. Li, J. Dong, Angew. Chem.
Int. Ed 2011, 50, 672-675; b) K. Kida, M. Okita, K. Fujita, S.
Tanaka, Y. Miyake, CrystEngComm 2013, 15, 1794-1801;
c) F. Pitzalis, C. Carucci, M. Naseri, L. Fotouhi, E. Magner,
A. Salis, ChemCatChem 2018, 10, 1578-1585.
Y. Pan, Y. Liu, G. Zeng, L. Zhao, Z. Lai, Chem.Commun.
2011, 47, 2071-2073.
3396.
[
[
24]
25]
a) Q. Yang, Q. Xu, S. H. Yu, H. L. Jiang, Angew. Chem.
Int. Ed. 2016, 55, 3685-3689; b) P.-Z. Li, X.-J. Wang, J.
Liu, J. S. Lim, R. Zou, Y. Zhao, J. Am. Chem. Soc. 2016,
[41]
138, 2142-2145.
a) H.-M. Wen, B. Li, H. Wang, R. Krishna, B. Chen,
Chem.l Commun. 2016, 52, 1166-1169; b) H. Liu, Y. He, J.
Jiao, D. Bai, D. l. Chen, R. Krishna, B. Chen, Chem. Eur. J. [42]
016, 22, 14988-14997.
2
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
[
[
[
[
[
[
43]
44]
45]
46]
47]
48]
M. A. Bjurlin, S. Bloomer, M. J. Haas, J. Am. Oil
Chem.Soc. 2001, 78, 153-160.
D. Steri, M. Monduzzi, A. Salis, Microporous Mesoporous
Mater. 2013, 170, 164-172.
A. Arvinte, M. Mahosenaho, M. Pinteala, A.-M. Sesay, V.
Virtanen, Microchim. Acta 2011, 174, 337-343.
M. Fabbrini, C. Galli, P. Gentili, J. Mol. Catal. B Enzym.
2
002, 18, 169-171.
C. Johannes, A. Majcherczyk, J. Biotechnol. 2000, 78,
93-199.
1
M. Solıs-Oba, V. M. Ugalde-Saldıvar, I. González, G.
Viniegra-González, J. Electroanal. Chem. 2005, 579, 59-
66.
[
[
49]
50]
Y. Zhang, J. Ge, Z. Liu, ACS Catal. 2015, 5, 4503-4513.
C. Mateo, J. M. Palomo, G. Fernandez-Lorente, J. M.
Guisan, R. Fernandez-Lafuente, Enzym. Microb.Technol.
2007, 40, 1451-1463.
[
[
51]
52]
Z. Zhong, S. Pang, Y. Wu, S. Jiang, J. Ouyang, J. Chem.
Technol. Biotechnol. 2017, 92, 1841-1847.
a) X. Wu, J. Ge, C. Yang, M. Hou, Z. Liu, Chem.Commun.
2015, 51, 13408-13411; b) F. Lyu, Y. Zhang, R. N. Zare, J.
Ge, Z. Liu, Nano Lett. 2014, 14, 5761-5765.
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201801293
FULL PAPER
FULL PAPER
Lipase
and
laccase
were
M. Naseri, F. Pitzalis, C Carucci, L.
Medda, L. Fotouhi, E. Magner, and A.
Salis*
encapsulated on ZIF-8 and ZIF-zni
metal organic framework materials
using
a
one-pot
synthesis-
immobilization method in aqueous
solution at room temperature (see
figure). The synthesized enzyme@ZIF
samples were characterized by XRD,
FT-IR, and TGA. Enzymatic activities
and stabilities of enzyme@ZIF
samples were obtained through
voltammetric methods.
Page ∎– Page ∎
Lipase and Laccase Encapsulated on
Zeolite
Imidazolate
Framework:
Enzyme Activity and Stability from
Voltammetric Measurements
This article is protected by copyright. All rights reserved.