Carbon nanotubes as activating tyrosinase supports for the selective synthesis of catechols

Article abstract of DOI:10.1021/cs400856e

A series of redox catalysts based on the immobilization of tyrosinase on multiwalled carbon nanotubes has been prepared by applying the layer-by-layer principle. The oxidized nanotubes (ox-MWCNTs) were treated with poly(diallyl dimethylammonium chloride) (PDDA) and tyrosinase to yield ox-MWCNTs/PDDA/ tyrosinase I. Catalysts II and III have been prepared by increasing the number of layers of PDDA and enzyme, while IV was obtained by co-immobilization of tyrosinase with bovine serum albumin (ox-MWCNTs/PDDA/BSA-tyrosinase). Attempts to covalently bind tyrosinase provided weakly active systems. The coating of the enzyme based on the simple layer-by-layer principle has afforded catalysts I-III, with a range of activity from 21 units/mg (multilayer, II) to 66 units/mg (monolayer, I), the best system being catalyst IV (80 units/mg). The novel catalysts were fully characterized by scanning electron microscopy and atomic force microscopy, showing increased activity with respect to that of the native enzyme. These catalysts were used in the selective synthesis of catechols by oxidation of meta- and para-substituted phenols in an organic solvent (CH ₂Cl₂) as the reaction medium. It is worth noting that immobilized tyrosinase was able to catalyze the oxidation of very hindered phenol derivatives that are slightly reactive with the native enzyme. The increased reactivity can be ascribed to a stabilization of the immobilized tyrosinase. The novel catalysts I and IV retained their activity for five subsequent reactions, showing a higher stability in organic solvent than under traditional buffer conditions.

Full text of DOI:10.1021/cs400856e

Research Article
pubs.acs.org/acscatalysis
Carbon Nanotubes as Activating Tyrosinase Supports for the
Selective Synthesis of Catechols
Fabiana Subrizi,† Marcello Crucianelli,*,† Valentina Grossi,† Maurizio Passacantando,† Lorenzo Pesci,‡
and Raffaele Saladino*,‡
†Department of Physical and Chemical Sciences, University of L’Aquila, Via Vetoio, I-67100 Coppito, AQ, Italy
‡Department of Ecology and Biology, University of Tuscia, Largo dell’Universita,
̀
01100 Viterbo, VT, Italy
S
* Supporting Information
ABSTRACT: A series of redox catalysts based on the immobilization of tyrosinase
on multiwalled carbon nanotubes has been prepared by applying the layer-by-layer
principle. The oxidized nanotubes (ox-MWCNTs) were treated with poly(diallyl
dimethylammonium chloride) (PDDA) and tyrosinase to yield ox-MWCNTs/
PDDA/tyrosinase I. Catalysts II and III have been prepared by increasing the
number of layers of PDDA and enzyme, while IV was obtained by co-immobilization
of tyrosinase with bovine serum albumin (ox-MWCNTs/PDDA/BSA-tyrosinase).
Attempts to covalently bind tyrosinase provided weakly active systems. The coating
of the enzyme based on the simple layer-by-layer principle has afforded catalysts I−
III, with a range of activity from 21 units/mg (multilayer, II) to 66 units/mg
(monolayer, I), the best system being catalyst IV (80 units/mg). The novel catalysts
were fully characterized by scanning electron microscopy and atomic force microscopy, showing increased activity with respect to
that of the native enzyme. These catalysts were used in the selective synthesis of catechols by oxidation of meta- and para-
substituted phenols in an organic solvent (CH2Cl2) as the reaction medium. It is worth noting that immobilized tyrosinase was
able to catalyze the oxidation of very hindered phenol derivatives that are slightly reactive with the native enzyme. The increased
reactivity can be ascribed to a stabilization of the immobilized tyrosinase. The novel catalysts I and IV retained their activity for
five subsequent reactions, showing a higher stability in organic solvent than under traditional buffer conditions.
KEYWORDS: tyrosinase, carbon nanotubes, catechols, biocatalyst, layer by layer, supported enzyme
catalysts by immobilization of tyrosinase on the epoxy resin
Eupergit C250L.5 These catalysts provided high catalytic
efficiency in the synthesis of catechols and DOPA peptides,
showing greater storage life, pH stability, and reusability when
coated by the layer-by-layer (LbL) procedure. However, some
disadvantages have been highlighted, such as the long duration of
loading procedures (incubation time of 24 h), the low enzyme
loading, and the use of covalent attachment that, although it
improves stability and reduces the extent of leaching, could
severely damage the structural integrity of the protein. In this
study, the choice of carbon nanotubes (CNTs) as a supporting
material for tyrosinase was made to improve the previously
mentioned approach. These fascinating nanomaterials offer a
high surface area for enzyme loading, with the establishment of
hydrophobic or electrostatic interaction for a better assembly
between the enzyme and support, as well as biocompatibility and
mechanical resistance.6 In the past, most of the efforts toward the
development of tyrosinase supported on CNTs have been
focused on the exploitation of their electronic properties and
electrode kinetics,7 with the aim of designing biosensors and
INTRODUCTION
■
Biological conversions using enzymes, being highly specific,
commonly require milder reaction conditions than classical
chemical catalysis. The poor stability of enzymes, the easy
inactivation caused by solvent or mechanical stress, and the high
cost of purification are sometimes the main drawbacks of this
methodology. To improve their performance and economic
viability, enzymes are generally immobilized onto a support.
Immobilization can improve the activity and stability, allowing
better handling and usage under unconventional operative
conditions.1 The increase in stability can be achieved via
multipoint or multisubunit immobilization, generation of a
favorable environment, and prevention of intermolecular
phenomena.1 Increased activity is somehow more complex and
unexpected, except in certain cases like multimeric proteins,
allosteric proteins, and enzymes that suffer conformational
changes, or by ensuring prevention of inhibition−inactivation
processes.1 In addition, immobilization leads to an easier
separation of the enzyme from the reaction mixture. Tyrosinase
is a copper enzyme that catalyzes the hydroxylation of
monophenols to o-diphenols and o-quinones using dioxygen
(O2) as the primary oxidant.2 This enzyme has been widely
studied because of its relevance in several applied areas.3,4 In a
recent work, we described the preparation of heterogeneous
Received: September 26, 2013
Revised: January 19, 2014
Published: January 27, 2014
© 2014 American Chemical Society
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Figure 1. Multishell LbL strategy.
Scheme 1. Synthesis of Catalyst I As Depicted in a Schematic Representation of the Adsorption of Tyrosinase (Tyr) on Ox-
a
MWCNTs/PDDA
a
Tyrosinase (negatively charged) is attracted by PDDA (positively charged).
biofuel cells, particularly useful in the detection of phenols8 or
sulfites.9 On the other hand, their application for synthetic
purposes has been largely understudied. Here we describe the
preparation and characterization of novel catalysts based on the
immobilization of tyrosinase on multiwalled carbon nanotubes
(MWCNTs) by the LbL method, focusing our attention on their
use as efficient biocatalysts for the synthesis of catechols. This
method is based on the consecutive deposition of alternatively
charged polyelectrolytes onto the active species (Figure 1).10
The LbL method is an effective tool for the stabilization of
enzymes because polyelectrolyte films can protect proteins from
high-molecular mass denaturing agents.11a−d
The strategy based upon the formation of a covalent bond by
the diimide-activated amidation and the glutaraldehyde-assisted
co-immobilization of tyrosinase and bovine serum albumin
(BSA) was also exploited, in this work, to anchor tyrosinase on
carbon nanotubes. Glutaraldehyde is one of the most widely used
reagents in the design of biocatalysts, acting as a useful cross-
linker and a versatile tool for enzyme immobilization, by reacting
with amino groups of enzymes.11e This process may further
stabilize multimeric enzymes (like tyrosinase, in this case) by
preventing subunit dissociation. Moreover, it can be reasonably
assumed that BSA will react via glutaraldehyde with the enzyme,
giving a higher stability. Indeed, BSA has been frequently used as
a proteic feeder to obtain cross-linked enzyme aggregates
(CLEAs).11f Catalysts conceived in this way, by combining the
unique role of CNTs in supporting the immobilized enzyme and
the advantages of enzymatic catalysis in phenol oxidation, would
increase the efficiency of the synthetic process with a reduced
economic impact.
immobilization. Hence, before the loading steps, the pristine
nanotubes were oxidized under acidic conditions, with the aim of
introducing a negative charge onto their surface. The acidic
treatment allowed the removal of impurities (residual con-
tamination and amorphous carbons) and the introduction of
carboxyl and hydroxyl groups at the ends or sidewall defects of
the structure. After prolonged sonication at room temperature
with a 3:1 mixture of concentrated sulfuric and nitric acids, the as
obtained nanotubes were solubilized in water, and the increase in
their solubility was verified by UV−vis spectroscopy (maximal
absorbance at 500 nm).6,12 A linear relationship between
absorbance and the concentration of oxidized nanotubes (ox-
MWCNTs) was observed: the solutions were stable for more
than 1 week (see Figure S1 of the Supporting Information).12
The ox-MWCNTs were successively treated with a positively
charged polyelectrolyte by the LbL technique to facilitate the
loading of tyrosinase, negatively charged at the operative pH
(7.0; isoelectric point of 4.7−5.0).13 Commercially available
poly(diallyl dimethylammonium chloride) (PDDA) was used as
polycation, and the coating was applied.14,15 Briefly, ox-
MWCNTs were treated with PDDA in 0.5 M NaCl, and the
suspension was subjected to orbital shaking to yield ox-
MWCNTs/PDDA. The 0.5 M NaCl solution was used to
optimize the ionic strength of the mixture, helping the dispersion
of nanotubes to facilitate the deposition of the polyelectrolyte.
Before the deposition of the next layer, the excess PDDA was
removed by centrifugation−redispersion cycles. The absence of
residual PDDA in the solution is critical to the adsorption of the
following layer because a precipitate would form upon mixing
́
with the enzyme. Ziołkowska et al. recently reported a simple
method for the determination of the PDDA concentration in
solution based on the Bradford procedure.16 To ensure the
complete removal of the polyelectrolyte, UV−vis spectra of each
supernatant with Coomassie Brilliant Blue (CBB) were recorded
until no evidence of the CBB−PDDA complex was detected.
This method, being quantitative, also permits the determination
of the amount of PDDA in the solution and, hence, the efficiency
of PDDA deposition. Monitoring the absorbance at 595 nm, we
RESULTS AND DISCUSSION
■
Preparation of the CNT Support and Optimization of
the Immobilization Procedures. Studies were conducted
with MWCNTs as supports because of their lower costs and
greater commercial availability. MWCNTs show low dispersi-
bility in aqueous solutions, also after prolonged sonication, and
this leads to minimization of the available surface for protein
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found the amount of PDDA adsorbed per milligram of
nanotubes was 0.573 mg. In the preparation of catalyst I (ox-
MWCNTs/PDDA/tyrosinase), the optimal weight ratio be-
tween the enzyme and the support was investigated. The ox-
MWCNTs/PDDA (1.0 mg) system was treated with different
amounts of tyrosinase (0.02−2.00 mg) in sodium phosphate
buffer (PBS, 0.1 M, pH 7.0), and the effectiveness of the
immobilization was evaluated by analyzing the residual activity in
the waste waters (Scheme 1).
Tyrosinase (Tyr) activity was determined by the dopachrome
assay17 following the oxidation of L-tyrosine at 475 nm; the
enzyme unit is defined in terms of the increase in absorbance of
10−3 unit/min at 25 °C in 0.1 M phosphate buffer (pH 7.0). The
activity of immobilized tyrosinase was expressed as the activity
unit per milligram of support (eq 1), where Ux is the activity
(units) of the immobilized enzyme assayed by the dopachrome
method.
to increase it. The low activity at high enzyme concentrations
(Table 1, entry 1) can reasonably be attributed to close packing
effects. On the other hand, the possibility that a high enzyme
concentration may affect the structure of the polymeric bed,
producing a negative effect on the activity, cannot be completely
ruled out. The activity yield exhibited a similar behavior.18 Note
that the unsupported enzyme may be successfully immobilized in
following batches.
Next, experiments were performed using a support:enzyme
ratio of 5:1. These optimized conditions would ensure maximal
efficiency of the catalyst with minimal enzyme waste.
Subsequently, we applied the LbL technique to increase the
number of layers of PDDA and enzyme. In particular, catalysts II
(Figure 2A) and III (Figure 2B) were prepared by addition of
PDDA and PDDA/Tyr layers, respectively.
These catalysts showed lower activity (21 and 43 units/mg,
respectively) than catalyst I. Thus, the efficiency of the catalysts is
not influenced by the number of layers. A different pattern was
observed when the Tyr/PDDA multilayer system was pro-
gressively assembled on a quartz slide.3g The lower activity
observed for catalysts II and III compared to that of I requires a
brief comment. In the case of II, the addition of the outer layer of
PDDA can affect the diffusion of the substrate, inhibiting the
interaction with the enzyme. On the other hand, in catalyst III,
the outer layer is similar to I. In this latter case, it is possible to
suggest that the second layer of tyrosinase is partially dissolved in
PDDA, even if the possibility of an active role of nanotubes
(which are redox active and now more distant from the surface of
the catalyst) cannot be completely ruled out. The chemical
immobilization of tyrosinase on ox-MWCNTs was also
performed. The use of a covalent bond to anchor enzymes to
carbon nanotubes is well-documented, and we tried to follow this
method to increase the stability of the system. The
immobilization was achieved using a two-step process through
the diimide-activated amidation and employing the optimal
support:enzyme ratio. Briefly, ox-MWCNTs were activated by
N-ethyl-N′-(3-dimethylaminopropyl)carbodiimide hydrochlor-
ide (EDC) in the presence of N-hydroxysuccinimide (NHS),
followed by enzyme coupling.9,19 Despite the high value of
protein immobilization (immobilization yield of 52%), a low
activity (9.0 units/mg) was obtained: probably, in this case,
chemical immobilization had a negative effect on enzyme
conformation. With the aim of further improving the efficiency
of catalyst I, we also investigated the glutaraldehyde-assisted co-
immobilization of tyrosinase and BSA.20 The immobilization
efficiency is dependent upon the number of enzyme molecules
loaded onto a given surface area. In such situations, given the
large surface area of nanotubes, the enzyme strives for the
greatest surface coverage, leading to conformational changes that
activity (units/mg) = U /W
x
support
(1)
Immobilization and activity yields were evaluated as reported
in eqs 2 and 3, respectively.
activity yield (%) = [U /(U − U)] × 100
(2)
(3)
x
a
r
immobilization yield (%) = [(U − U)/U ] × 100
a
r
a
where Ua is the total activity (units) of the enzyme added to the
solution and Ur is the residual activity (units) evaluated by the
dopachrome method in the washing solutions. As reported in
Table 1, the immobilization yield increased with an increase in
a
Table 1. Activity Parameters for Catalyst I
support/Tyr
(mg/mg)
immobilization
yield (%)
activity yield
(%)
activity
(units/mg)
b
entry
1
2
3
4
5
6
7
0.5
1
16
17
46
47
70
73
74
<1
11
59
66
44
23
10
3
11
5
27−34
19−24
13
10
15
25
50
13
6
a
All experiments were conducted in triplicate. Average errors were
∼0.1% for immobilization yield and 0.1−0.2% for activity yield and
b
activity. The activity of native tyrosinase, evaluated by the
dopachrome method, was 2568 units/mg of enzyme (see ref 25).
the support:enzyme ratio while the activity reached a maximum
at a ratio of 5 (Table 1, entry 3). Thus, it appears that there is a
critical concentration value beyond which any further immobi-
lization of the enzyme decreases its activity instead of continuing
Figure 2. Representations of catalysts II (A) and III (B).
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Scheme 2. Synthesis of Catalyst IV As Depicted by a Schematic Representation of the BSA-Assisted Co-immobilization of
Tyrosinase (Tyr) on ox-MWCNTs
could influence its activity. This effect can be avoided by the
addition of an inert protein that reduces the amount of surface
area available to the enzyme. BSA, with an isoelectric point close
to that of tyrosinase, was the protein selected for this purpose.21
Glutaraldehyde (GA) was used as the cross-linking agent for the
formation of a three-dimensional network “to confine” the
enzyme as shown for the catalyst ox-MWCNTs/PDDA/BSA-
tyrosinase IV (Scheme 2).
Briefly, Tyr and BSA were immobilized on PDDA-coated
MWCNTs in the presence of GA following the general
procedure reported by Kim et al. for the preparation of Tyr-
based sensitive electrochemical immunosensors.22a In particular,
different amounts of BSA were studied, maintaining the
MWCNTs:Tyr ratio of 5:1 (milligrams) already selected for
catalyst I, and the optimal value of GA concentration [0.5−1.0%
(v/v)] previously described in the literature.3h The immobiliza-
tion yield increased when the BSA:Tyr ratio increased to 3:1 and
then decreased, suggesting this last value was the optimal one
(Table 2, entries 1−4). The activity parameters showed a similar
behavior. The 3:1 BSA:Tyr ratio had been previously reported as
the optimal value in the preparation of biosensors based on the
immobilization of Tyr on MWCNTs in a glassy carbon
electrode.22b
The activity and immobilization parameters varied slightly
close to the optimal value (see, for example, entries 2 and 4 vs
entry 3) in accordance with data previously reported for similar
systems.20 The treatment with GA before the removal of the
unbounded proteins allowed the increase in enzyme loading by
cross-linking additional tyrosinase and BSA without affecting the
activity. Catalyst IV was more active than previous systems (80
units/mg), showing a 67% immobilization yield. Among the
novel catalysts, I and IV (the most active) were selected for
successive characterizations and applications.
Scanning Electron Microscopy (SEM) and Atomic Force
Microscopy (AFM). The topography and morphology of
catalysts I and IV were examined by SEM and AFM to
demonstrate the effective polyelectrolyte deposition and enzyme
immobilization. The analyses were performed on well-dispersed
and separated MWCNTs supported on a silicon substrate (Si).
Figure 3 shows the SEM images and the corresponding AFM
images23 of the selected areas of the different samples. MWCNTs
well dispersed and separated on Si are shown in Figure 3a. The
corresponding AFM image (Figure 3b) shows the detail of a
single nanotube immobilized on Si; the white areas, which are
higher than the substrate, are drag from the AFM tip during the
scanning. The SEM image of ox-MWCNT/PDDA (Figure 3c)
showed a cluster of functionalized nanotubes. Only at the cluster
edge is it possible to find individual functionalized nanotubes.
The AFM morphological study (Figure 3d) allowed the
evaluation of the cluster height that is ∼1 μm.
a
Table 2. Activity Parameters for Catalyst IV
immobilization yield
(%)
activity yield
(%)
activity
(units/mg)
b
entry BSA:Tyr
1
2
3
4
1:1
2:1
3:1
4:1
56
60
67
63
38
40
47
42
68
71
80
73
The SEM analysis (Figure 3e) of catalyst I showed that the
nanotubes with the immobilized enzyme were more dispersed,
broken, and shorter than the original MWCNTs. The AFM
analysis (Figure 3f) showed the remarkable roughness of the
external structure that completely covers the nanotube.
For catalyst IV, we obtained a good dispersion of nanotubes
onto Si, but after sonication, they appeared to be broken (Figure
3g). The AFM image (Figure 3h) shows small clusters of
a
b
Support:Tyr ratio of 5:1 (milligrams). Final concentration of GA of
0.5−1.0% (v/v). Note that changes in both activity and immobilization
parameters in the range of used GA concentrations [0.5−1.0% (v/v)]
were not observed. In accordance with ref 22a, treatment with GA
after the removal of the unbound Tyr inhibited the performance of
catalyst IV (49% of immobilization yield, 36% of activity yield).
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Figure 3. SEM and AFM images of MWCNTs (a and b), ox-MWCNTs/PDDA (c and d), catalyst I (e and f), and catalyst IV (g and h) deposited on
silicon substrates.
nanotubes, and individual functionalized nanotubes are clearly
visible.
Figure 4 reports the high-resolution AFM images of individual
MWCNTs and the related image profiles for selected directions
(line in Figure 4a). The nanotube surface appears to be smooth,
and the maximal height (h) and width at half-height (d),
evaluated by a profile analysis (Figure 4c), were 39.0 0.5 and 53
2 nm, respectively. The width of the nanotube has been
estimated at half-height and not at the bottom to reduce the size
of errors resulting from effects of tip convolution.
High-resolution AFM images of ox-MWCNT/PDDA at the
same scale (300 nm × 300 nm) of the previous sample are
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Figure 4. High-resolution AFM images for MWCNTs: (a) bidimensional and (b) tridimensional analyses. The line in panel a indicates the section that
was further analyzed for the image profile (c).
Figure 5. High-resolution AFM images for ox-MWCNT/PDDA: (a) bidimensional and (b) tridimensional analyses. The line in panel a indicates the
section that was further analyzed for the image profile (c).
Figure 6. High-resolution AFM images for catalyst I: (a) bidimensional and (b) tridimensional analyses. Lines I−III in panel a indicate the sections that
were further analyzed for the image profiles (c).
reported in Figure 5. Although the surface is smooth, h and d
dimensions of the system are doubled, confirming that an
were 60.5 0.5 and 90 2 nm, respectively (Figure 5c). The
external shell covers the nanotube.
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Figure 7. High-resolution AFM images for catalyst IV: (a) bidimensional and (b) tridimensional analyses. Lines I and II in panel a indicate the sections
that were further analyzed for the image profiles (c).
Figure 8. Activity of catalysts I and IV vs the number of runs, in buffer and in organic solvent.
The AFM analysis of catalyst I (Figure 6) showed the
remarkable roughness and the presence of external “lumps” that
can be attributed to tyrosinase, which completely covers the
nanotube. To estimate the size of both the nanotube and the
lumps, three different image profiles were performed (Figure 6c).
From profile II, the maximal h of the whole structure is 199.0
0.5 nm and the d is 259 2 nm. From profiles I and III, we
obtained the lump d that ranges from 62 2 to 80 2 nm.
The AFM image (Figure 6b) shows small clusters where
individual functionalized nanotubes are clearly visible. Finally,
high-resolution AFM images of catalyst IV (Figure 7) show a
lower roughness and fewer lumps than catalyst I. The smoother
surface can be explained by suggesting that BSA occupies
interstitial sites between tyrosinase molecules. Maximal heights
and widths of 46.0 0.5 and 101 2 nm, respectively, for profile
I and 70.0 0.5 and 126 2 nm, respectively, for profile II
(Figure 7c) were obtained.
Reusability Properties and Storage Stability. The
reusability properties of catalysts I and IV were evaluated in
both buffer and organic solvent, performing five consecutive
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oxidations of L-tyrosine in 0.1 M PBS (pH 7.0) (Figure 8, red
line) and p-cresol in CH2Cl2 and buffer (Figure 8, blue line),
respectively. Several advantages can be obtained using enzymes
in organic solvents, such as the enhanced solubility of
hydrophobic substrates, reduced side reactions, enhancement
of catalyst performance, and simple recovery from the reaction
mixture.24 Previously published data from our laboratory showed
While the native enzyme and catalyst IV showed a slight
decrease of activity in the first 7 days, thereby reaching a constant
value of ∼70% of residual activity, catalyst I retained the full
activity for up to 20 days. With respect to the storage stability of
IV, Bakir et al.27 described a loss of activity, for cross-linked
immobilized tyrosinase in the presence of BSA, in the range of
10−20% after 20 days (depending on the purity of the enzyme).
These data are comparable with the results observed, in our case,
for catalyst IV. Thus, the maintenance of the catalytic activity for
successive reactions shown above, along with data on their
storage stability, confirms the excellent general behavior in terms
of the stability of the prepared catalysts I and IV.
Kinetic Analyses. Kinetic parameters for native tyrosinase
and selected catalyst I were evaluated in CH2Cl2 and PBS, using
p-cresol as the substrate. The initial reaction rates were measured
at substrate concentrations ranging from 1.0 to 20 mM at 25 °C.
Because catechol dehydrogenation is reported to follow
Michaelis−Menten kinetics, benzoquinone formation was
measured at 389 nm. Kinetic constants were evaluated by
using different linear regression equations [Lineweaver−Burk,
Eadie−Hofstee, and Hanes (Figures S4−S6, respectively, of the
Supporting Information) and a nonlinear regression plot (Figure
S7 of the Supporting Information). The Hanes plot (Figure S6 of
the Supporting Information) provided excellent correlation
values without the necessity of discarding any point. From a
comparison between the linear regression and nonlinear
regression plot analyses, we can observe how the Hanes (mainly)
and Eadie−Hofstee (to a lesser extent) plots, respectively, gave
the best estimations while the Lineweaver−Burk plot has
afforded under- or overestimated values (compare Figures S4−
S6 of the Supporting Information).
that CH2Cl2 plays a benign role in the synthesis of catechols.25
A
minimal amount of phosphate buffer was also used to disperse
the catalysts, because water is fundamental for the correct folding
of the enzyme and consequently for its proper functionality. The
oxidations were followed at 475 nm (in buffer) or 389 nm (in
CH2Cl2 and buffer), and after the absorbance plateau had been
reached, the immobilized enzyme was recovered by simple
centrifugation, washed, and reused with a fresh substrate. Data
referring to tyrosinase immobilized on Eupergit (Tyro/E-LbL)
are shown in Figure 8 as ref 25. As a general trend, the reusability
of catalysts I and IV was higher in organic solvent than in buffer.
The organic medium strongly improved the stability and
efficiency of the catalysts, and only a minimal loss of activity
was observed during the first five runs, for catalysts I and IV.
Irrespective of experimental conditions, catalyst IV was more
reactive than I. This result may be due to a more rigid enzyme
structure in IV that can resist conformational changes that could
affect activity.26 The T50 values for I and IV in buffer (defined as
the run number at which the catalyst activity is reduced to 50%)
were found to be 3 and 4, respectively [T50 was calculated by
linear regression of the percentage of activity vs run plot (see
Figures S2 and S3 of the Supporting Information)]. In organic
solvent, the T50 values of I and IV were 8 and 12, respectively.
The novel catalysts exhibited better performance with respect to
the Tyro/E-LbL system.25
The reusability experiments showed different behaviors of
catalysts I and IV in the different solvents. In particular, the lower
T50 values for catalysts I and IV in buffer can be justified by
invoking the stronger tendency of the o-quinones (formed in the
oxidation of o-diphenols) to undergo a sequence of non-
enzymatic reactions, in this medium, leading to synthetic melanin
as a final product. These insoluble polymers can precipitate and
be strongly adsorbed on the functionalized nanotubes that
cannot be reused. This phenomenon is particularly evident in
buffer, where the polymerization process is favored.3g
The kinetic parameters (Km, Vmax, and Kcat) for the native
enzyme and catalyst I are listed in Table 3.
Table 3. Kinetic Parameters for Native Tyrosinase (Tyr) and
a
Catalyst I
Vmax × 104 (ΔAbs
min−1 μg of
Km (mM)
enzyme−1
)
Kcat (min−1
)
plot
Tyr catalyst I Tyr catalyst I
Tyr
catalyst I
Lineweaver−
3.92
7.68
49.3
56.1
1310
1683
Burk
Hanes
2.52
2.82
2.9
9.91
8.12
43
44
44
63
58
63
1287
1330
1318
1891
1732
1882
Further, the stability of the new systems, namely, native
tyrosinase and catalysts I and IV, was evaluated by storing the
catalysts in PBS at 4 °C for 20 days. The activity was measured at
specific times at room temperature by the oxidation of p-cresol in
the mixed organic medium (PBS and CH2Cl2) (Figure 9).
Eadie−Hofstee
nonlinear
regression
10
a
Kcat is defined as vmax (Δabsorbance per minute)/[Tyr] (micromoles
per milliliter). All experiments were conducted in triplicate using free
and immobilized tyrosinase. Average errors in kinetic parameters were
2−4% for Km and 1−3% for Vmax
.
As a general trend, the immobilized enzyme (catalyst I)
showed the highest Km, probably because of the enzyme
conformational changes and mass transfer limitations (Table
3). Irrespective of the regression plot used, catalyst I showed a
higher rate and a higher turnover number than the native
enzyme, thus suggesting the stabilizing effect of the nanotubes.28
Experimental values were also fit using nonlinear regression
procedures like GraphPad Prism6: this method permits us to
obtain standard deviations of the kinetic parameters and to
evaluate the reliability of the previously reported regression plots
(Figure S7 of the Supporting Information).
Figure 9. Storage stability of native tyrosinase and catalysts I and IV at 4
°C.
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a
Scheme 3. Oxidation of Phenols
a
(i) Tyro-based catalysts, O2; (ii) CH2Cl2 and buffer; (iii) phosphate buffer.
Table 4. Oxidation of Phenols 1−5 with Native Tyrosinase and Catalysts I and IV in Buffer or CH2Cl2 with PBS
a
d
entry
catalyst
solvent
substrate
product(s)
1a
conversion (%)
yield (%)
b
c
1
tyrosinase
buffer/AA
1
1
1
1
1
1
2
2
2
3
3
3
4
4
4
5
5
5
98
99
99
93
96
98
55
80
85
9
13
b
c
2
I
buffer/AA
1a
6
b
c
3
IV
buffer/AA
1a
6
4
tyrosinase
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
CH2Cl2/buffer
1a [1b] {1c}
54 [4] {14}
5
I
1a [1b] {1c}
78 [8] {9}
6
IV
1a [1b] {1c}
82 [4] {3}
7
tyrosinase
2a [2b]
2a
42 [13]
80
8
I
9
IV
2a
85
10
11
12
13
14
15
16
17
18
tyrosinase
3a
<4
I
3a
22
25
8
22
IV
3a
24
tyrosinase
4a
<5
I
4a
21
34
42
35
39
21
IV
4a
34
tyrosinase
1a [1b]
1a
35 [7]
33
I
IV
1a
39
a
Reaction conditions: native or immobilized tyrosinase (240 units) in PBS (275 μL), 0.05 mmol of substrate, CH2Cl2 (2.5 mL) at 25 °C. All
b
c
d
experiments were conducted in triplicate. Solvent of 0.1 M phosphate buffer (pH 7.0). AA, ascorbic acid. The average error in the yield was 0.3%.
Oxidation of Phenols. The synthetic versatility of catalysts I
and IV was evaluated in the oxidation of a panel of selected
phenols (1−5 in Scheme 3) to the corresponding catechol
derivatives. Catechols are characterized by several biological
activities and are well recognized as antioxidant compounds.29
The potential of tyrosinase in the synthesis of catechols has
received a great degree of attention because they are difficult to
obtain under environmentally friendly conditions by means of
traditional chemical procedures.4d,30 The application of
tyrosinase in buffer is partially limited because of the formation
of reactive o-quinones that can covalently bind to the enzyme or
polymerize producing brown pigments. For this reason, the
oxidation of p-cresol 1 was initially performed in the presence of
ascorbic acid (AA) that acts as an internal reducing agent.5 As a
general procedure, phenol, native or immobilized tyrosinase
(240 units), and AA (1.5 equiv) in a buffer (PBS, 0.1 M, pH 7.0)
solution (5.0 mL) were vigorously stirred at room temperature,
under a dioxygen atmosphere (O2), according to a procedure
previously reported by our laboratory.25 Under these conditions,
catalysts I and IV behaved like the native enzyme, showing
excellent substrate conversions but only low yields in the desired
catechol 1a (Table 4, entries 1−3).
The low mass balance value observed for the transformation
was probably caused by the formation of products of further
oxidation and/or of polymeric material not isolated under our
experimental conditions. Better results were obtained in organic
medium. The substrate in CH2Cl2 was treated, at 25 °C, with
native or immobilized tyrosinase (240 units) in the presence of
the appropriate amount of PBS, thus ensuring the minimal
amount of hydration water for the proper functioning of the
enzyme. Under these experimental conditions, catalysts I and IV
were more active than the native enzyme, affording 1a in high
yield and conversion of substrate (Table 4, entries 4−6). Dimers
1b and 1c, characterized by the formation of the C−C bond
between two phenol units, were also detected in very small
amounts (Scheme 3 and Table 4, entries 4−6). The blank test
performed under conditions similar to those described above
showed the absence of any type of reactivity when the support
ox-MWCNT was used, without the enzyme, for the oxidation of
p-cresol 1, thus confirming its neutral role. The effectiveness of I
and IV in CH2Cl2 and buffer was further confirmed with more
hindered para-substituted phenols 2−4. Again, in the oxidation
of 4-ethyl phenol 2, catalysts I and IV were more reactive than
native tyrosinase, catechol 2a being detected in high yields and
conversions of substrate (Table 4, entries 7−9). It is noteworthy
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that no dimers were produced with immobilized tyrosinase in
entries 8 and 9. Catalysts I and IV were highly active also in the
oxidation of hindered 4-s-butyl phenol 3 and 4-tert-butyl phenol
4, because these substrates were not converted by native
tyrosinase under the experimental conditions tested. Catechols
3a and 4a were obtained in 22−24 and 21−34% yields,
respectively (Table 4, entries 10−15). Previous studies showed
the possibility of slightly oxidizing 3 and 4 with native tyrosinase
in CH2Cl2 and buffer using only 600−1400 units of the
enzyme.19 As a general trend, the efficiency of catalysts decreased
with an increase in the steric hindrance of the para substituent.
These results are in accordance with previously reported data on
the low reactivity of bulky phenols with tyrosinase in organic
medium, because of the reduced conformational flexibility of the
enzyme in a hydrophobic solvent.31 Catalysts I and IV were also
active in the oxidation of 3-methyl phenol (m-cresol) 5 as a
selected example of a meta-substituted substrate, and 1a was
obtained in 33 and 39% yields, respectively [conversion of
substrate in the range of 35−39% (Table 4, entries 16−18)].
Irrespective of the experimental conditions, IV was more reactive
than I, confirming the positive role of BSA in the immobilization
of tyrosinase. From a comparative point of view, if we refer to
catechol 1a as the main reaction product, the yields obtained with
catalysts I and IV are comparable and, in some cases, slightly
larger than those previously obtained by us with the reference
catalyst (Tyro/E-LbL),25 in spite of the remarkable fact that in
catalysts I and IV a smaller amount of enzyme has been
immobilized per weight of support.
the immobilized tyrosinase. The greater efficiency of the two
biocatalysts probably depends on several factors, including the
stabilization of the enzyme by nanotubes and the poor formation
of polyphenols as side products that can precipitate on the
nanotube inhibiting the tyrosinase activity.3g Furthermore, a
possible synergic effect of the support in the oxidation cannot be
completely ruled out. Recently, Zhong and co-workers showed
that oxidized carbon nanotubes are redox active species that can
cooperate during the oxidation.28 They could be themselves the
primary oxidant and also could accelerate the reaction of O2 by
energetically favorable absorption processes on the surface (ΔH
= −16.3 kcal/mol).32 These effects were directly related to an
increase in peroxidase activity using nanotubes as mediators of
oxidation. The possibility of a similar effect in the case of
tyrosinase can in part justify the improved reactivity of our
catalysts in the oxidation of sterically congested substrates. The
active redox role of nanotubes can also explain the observed
decrease in activity of catalysts II and III with an increase in the
number of layers with respect to I, leading to the hypothesis of a
reduction of the synergic effect with an increase in the distance of
the enzyme from the support. The high activities of catalysts I
and IV open new stimulating entries for novel oxidation
processes of hindered phenols based on tyrosinase.
EXPERIMENTAL SECTION
■
Reagents and Materials. Multiwalled carbon nanotubes
(MWCNTs), mushroom tyrosinase from Agaricus bisporus (T-
3824, multimeric enzyme with H2L2-type structure),33 BSA,
dichloromethane (CH2Cl2), ethyl acetate (EtOAc), hexane
(Hex), sodium dithionite (Na2S2O4), anhydrous sodium sulfate
(Na2SO4), dodecane, pyridine, hexamethyldisilazane (HMDS),
trimethylchlorosilane (TMCS), glutaraldehyde (GA), PDDA, L-
tyrosine, and phenols 1−5 were purchased from Sigma-Aldrich.
All spectrophotometric measurements were taken with a Varian
Cary50 UV−vis spectrophotometer equipped with a Peltier
thermostated single-cell holder. Dichloromethane was dried on
anhydrous sodium sulfate prior to use. All experiments were
conducted in triplicate using native and immobilized tyrosinase
in the dichloromethane/buffer system and in an aqueous
medium. Sodium phosphate buffer (PBS, 0.1 M, pH 7.0) is a
solution containing NaH2PO4 and Na2HPO4 in Milli-Q water;
no NaCl is present. This buffer solution was used at pH 7.0 and
0.1 M unless differently specified.
CONCLUSIONS
■
The activity and efficiency of tyrosinase immobilized on carbon
nanotubes vary significantly depending on the applied
immobilization procedure. The coating of the enzyme based
on the LbL principle has afforded the best catalysts since the
deposition of the first layer of enzyme. The activity of catalyst I
reached a maximum at a support:tyrosinase ratio of 5, and it was
found to decrease in the presence of a high enzyme
concentration, probably because of close packing effects. The
deposition of the second PDDA layer decreased the activity,
although it started to increase again with a further increase in the
number of enzyme layers. In the glutaraldehyde-assisted
immobilization procedure, the presence of BSA as protein
spacer leads to the more active catalyst IV, confirming the
favorable role of the protein to avoid steric effects and
conformational changes in tyrosinase. In accordance with the
literature, the chemical linkage was the less efficient procedure.
The new catalysts were more active than previously described
heterogeneous systems based on the immobilization of
tyrosinase on epoxy resin Eupergit or on His entrapment in a
matrix of chitosan.3h From a kinetic point of view, the
immobilized enzyme showed higher rates and turnover numbers
than native tyrosinase. With regard to concerns about stability,
catalysts I and IV retained their activity for more subsequent
reactions, showing a higher stability in organic solvent than in
buffer. The catalytic performance in the oxidation of phenols to
catechols deserves special attention. In fact, catalysts I and IV
were more efficient than the native enzyme in affording catechols
1a−4a in high yield and conversion of substrate. The oxidation in
organic medium was the only synthetically useful procedure,
because in buffer the products of overoxidation were obtained
even in the presence of ascorbic acid as an internal reducing
agent. This was particularly evident with bulky substituents, such
as 4-s-butyl phenol 3 and 4-tert-butyl phenol 4, only oxidized by
Preparation of Catalysts I−IV. Multiwalled carbon nano-
tubes (>95% pure, inner diameter of 5−10 nm, outer diameter of
10−30 nm, and length of 0.5−50 μm) were treated with a
concentrated sulfuric acid (98%)/nitric acid (65%) mixture [3:1
(v/v)] in a sonication bath for 4 h. The resulting solution was
diluted with deionized water and extensively washed by
consecutive sonication−centrifugation steps until the pH
became neutral. The oxidized nanotubes (ox-MWCNTs) were
lyophilized and stored at room temperature as a dry powder.
Coating of MWCNTs with a PDDA Solution. PDDA (2.0 mg)
and ox-MWCNTs (1.0 mg) were dispersed in 0.5 M NaCl (2.0
mL) and subjected to sonication for 5 min to give a stable black
suspension. After being shaken at 170 rpm for 20 min, the
mixture was centrifuged (6000 rpm for 20 min), and the
composite was washed with Milli-Q water to remove the residual
polyelectrolyte. The absence of a polyelectrolyte in the solution
was confirmed by the Bradford method.16,34
Preparation of Catalyst I.35 Tyrosinase was adsorbed on the
first polycation layer by incubating the PDDA-coated ox-
MWCNTs (1.0 mg) with different amounts of the enzyme
819
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(ranging from 2.0 to 0.02 mg) in PBS (0.1 M, pH 7.0) in an
orbital shaker at room temperature (40 min at 150 rpm). After
the mixture had been shaken, the excess enzyme was removed by
centrifugation (6000 rpm × 20 min) and the supernatant was
used for immobilization and activity yield calculations. The
complex was washed several times with PBS to ensure the
complete removal of the unbound enzyme. The absence of
tyrosinase in the washing waters was confirmed by both the
activity assay and the Bradford method.
Preparation of Catalysts II and III. To reduce the possibility
of tyrosinase desorption, a final layer of PDDA (catalyst II) and a
further layer of tyrosinase (catalyst III) were added on I
following the reported general procedure.
using commercial silicon tips (frequency range of 51−94 kHz)
scanned by means of a Veeco Nanoman closed loop XY head. For
the SEM and AFM analysis, different solutions of the samples
were prepared. In particular, MWCNT and ox-MWCNTs/
PDDA samples were dispersed in pure ethanol and catalysts I
and IV in Milli-Q water, to prevent enzyme denaturation. All the
solutions were sonicated for 5 min at room temperature, and a
drop of each solution was taken and deposited onto silicon
substrates (Si). At the end, the obtained samples deposited on Si
were subjected to annealing at 50 °C to quickly evaporate the
solvent (ethanol or water).
Determination of the Kinetic Constants. The catalytic
properties of native and immobilized tyrosinase were determined
in the organic solvent by measuring the initial rates of the
reaction with the substrate at 25 °C. The reactions were
conducted by using different concentrations of p-cresol 1,
ranging from 2 to 18 mM. The appropriate amount of 1 was
dissolved in DCM (2.5 mL), and free or immobilized tyrosinase
in the optimal aqueous amount (275 μL of PBS) was added. The
reaction mixture was stirred for 30 min. Sampling was performed
every 6.0 min, the absorbance at 389 nm (due to o-quinone)
measured, and the sample returned to the flask as rapidly as
possible. One unit of enzyme activity was defined as the increase
in absorbance of 0.001 at 389 nm and 25 °C in CH2Cl2 and 0.1 M
PBS (pH 7.0). Km and Vmax values were calculated by plotting
data in Lineweaver−Burk, Hanes, Eadie−Hofstee, and nonlinear
regression plots. All experiments were conducted in triplicate
using native and immobilized tyrosinase.
Storage Stability. The storage stability of native and
immobilized tyrosinase was estimated by measuring enzyme
activity at specific time intervals (0−20 days) and at room
temperature in organic medium. The catalysts were stored in
PBS (2.0 mg/mL) at 4 °C, and at each time point, an appropriate
amount was added to a solution of p-cresol 1 (18 mM) in CH2Cl2
(2.5 mL) containing the opportune amount of buffer (275 μL).
The initial rates of the reaction were measured by means of the
same procedure described for the determination of the kinetic
constants. Each sample was assessed in triplicate, and the
tyrosinase activity was expressed as the relative percentage
activity with respect to that at time zero.
Enzyme Recycling. The reusability of catalysts I and IV in
aqueous medium was determined by recording the time-
dependent increase in absorbance at 475 nm (dopachrome
formation) during L-tyrosine oxidation. The catalyst was then
recovered by centrifugation, washed to remove the substrate, and
reused again. The reusability of catalysts I and IV in the organic
solvent was determined by measuring the initial rates of the
reaction with the substrate at 25 °C. Briefly, 5.0 mg of p-cresol 1,
1.0 mg of I or IV, 275 μL of buffer, and 2.5 mL of CH2Cl2 were
placed in a flask at 25 °C under O2. The reaction mixture was
vigorously shaken, and the initial rates of the reaction were
measured by means of the same procedure described for the
determination of the kinetic constants. After 30 min, the catalyst
was recovered in the aqueous solution by phase separation,
centrifuged, washed to remove the substrate, and reused again.
For each run, the tyrosinase activity was expressed as the relative
percentage activity with respect to that of the first run.
Preparation of Catalyst IV.22 Tyrosinase (60 μL, 3.4 mg/
mL) and BSA (300 μL, 2.0 mg/mL) were added to a suspension
of the PDDA-coated MWCNTs (1.0 mg) in PBS (280 μL). After
the mixture had been shaken for 30 min, 2.5% glutaraldehyde
(GA) was added to reach a final volume of 800 μL and the
mixture was shaken again at room temperature for 30 min and
then at 4 °C overnight. Subsequently, the black suspension was
centrifuged (6000 rpm × 20 min) to remove the supernatant
containing the excess enzyme and GA. The supernatant was used
both for immobilization and activity yield calculations. The
catalyst was finally treated with 1.5 mL of 0.1 M Tris-HCl buffer
(pH 7.2) while being shaken for 1 h at 4 °C (to cap unreacted
aldehyde groups) and centrifuged and the supernatant removed.
The conjugate sample was washed several times with PBS to
ensure the complete removal of the unbound enzyme, and the
absence of tyrosinase, in the washing waters, was confirmed by
both the activity assay and the Bradford method.
Chemical Attachment of Tyrosinase. To a suspension of
nanotubes (1 mg/mL) in 2-(N-morpholino)ethanesulfonic acid
(MES) buffer (50 mM, pH 6.2) was added N-hydroxysuccini-
mide (NHS) (1 mL, 400 mM in MES buffer). After sonication
for 30 min, 1 mL of fresh N-ethyl-N′-(3-dimethylaminopropyl)-
carbodiimide hydrochloride (EDC) in MES buffer (20 mM) was
added quickly, and the mixture was stirred at room temperature
for 30 min. The activated nanotube solution was then centrifuged
at 6000 rpm for 20 min and washed three times with the buffer to
remove excess EDC and NHS. A solution containing tyrosinase
(0.2 mg in PBS) was added to the activated nanotubes, and the
dispersion was shaken at 200 rpm during the conjugation of the
enzyme. Unbound tyrosinase was removed by consecutive
centrifugation and washing steps.
Determination of the Activity of the Native and
Immobilized Enzyme. The activity of the native and
immobilized enzyme was determined by measuring the
tyrosinase-catalyzed oxidation of L-tyrosine (L-Tyr). The
reaction was started by adding the substrate to the solution
containing the appropriate amounts of native enzyme or catalysts
I−IV in PBS under magnetic stirring. The initial rates were
measured as linear increases in the optical density at 475 nm, due
to dopachrome formation. One unit of enzyme activity was
defined as the increase in absorbance of 0.001 per minute at pH 7
and 25 °C in a 3.0 mL reaction cuvette containing 0.83 mM L-
tyrosine and 67 mM PBS (pH 7.0). Spectrophotometric data
were analyzed with Cary WinUV. All experiments were
conducted in triplicate using free and immobilized tyrosinase.
SEM and AFM Characterization of Catalysts. The surface
morphology of the samples has been studied by SEM analysis,
making use of a Zeiss LEO 1530 apparatus equipped with a field
emission electron gun, while AFM was conducted with a Digital
Dimension D5000 instrument with a Nanoscope IV controller,
Oxidation of Phenols. The oxidation of phenols 1−5 in the
biphasic CH2Cl2/buffer system was performed following a
procedure previously optimized by our group.25 Briefly, to a
solution of the substrate (0.05 mmol) in CH2Cl2 (2.5 mL) was
added the opportune amount of native or immobilized tyrosinase
(240 units) in PBS (275 μL), and the mixture was stirred at room
820
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6307. (d) Zhou, Z.; Hartmann, M. Chem. Soc. Rev. 2013, 42, 3894−
3912. (e) Verma, M. L.; Barrow, C. J.; Puri, M. Appl. Microbiol.
Biotechnol. 2013, 97, 23−39. (f) Hwang, E. T.; Gu, M. B. Eng. Life Sci.
2013, 13, 49−61. (g) Rajkumar, K.; Palla, S.; Paladugu, A.; Reddy, E. R.;
Reddy, K. R. Int. Res. J. Pharm. 2013, 4, 36−44. (h) Tran, D. N.; Balkus,
K. J., Jr. ACS Catal. 2011, 1, 956−968. (i) Garcia-Galan, C.; Berenguer-
Murcia, A.; Fernandez-Lafuente, R.; Rodrigues, R. C. Adv. Synth. Catal.
2011, 353, 2885−2904.
temperature under O2. Reactions were monitored by thin layer
chromatography (TLC). After 24 h, the two phases were
separated by decantation, and in the case of immobilized
tyrosinase, the catalyst was recovered by centrifugation of the
aqueous layer. The organic fraction was concentrated with a
rotary evaporator and taken up with a solution of sodium
dithionite in THF and H2O [1:1 (v/v)]. The mixture was stirred
at room temperature for 5 min to allow the complete reduction of
benzoquinones to catechols and extracted twice with ethyl
acetate. The collected organic extracts were dried over anhydrous
sodium sulfate, filtered, and concentrated under vacuum. For gas
chromatography and mass spectrometry (GC−MS), the residue
was treated with anhydrous pyridine (kept over NaOH pellets)
and a 2:1 HMDS/TMCS mixture under vigorous stirring at
room temperature for 30 min and then allowed to stand for 5
min.36 Dodecane (1 μL) was used as an internal standard. The
oxidation of 1 in buffer was performed following a previously
reported protocol.5 In a general procedure, a solution of 1 (0.05
mmol), ascorbic acid (1.5 equiv), and the appropriate amount of
native or immobilized tyrosinase (240 units) in 0.1 M PBS (pH
7.0, 5.0 mL) were stirred at 25 °C under O2. The reactions were
monitored by TLC. After the disappearance of the substrate, the
mixture was acidified with a solution of 1.0 M HCl and extracted
twice with EtOAc. In the case of immobilized tyrosinase, the
biocatalyst was first removed from the reaction mixture by
centrifugation. The collected organic extracts were washed with a
saturated solution of NaCl, dried over anhydrous Na2SO4,
filtered, and concentrated. The obtained colored residue was
analyzed by GC−MS as previously reported. All experiments
were conducted in triplicate.
(2) (a) Seo, S.-Y.; Sharma, V. K.; Sharma, N. J. Agric. Food Chem. 2003,
51, 2837−2853. (b) Halaouli, S.; Asther, M.; Sigoillot, J.-C.; Hamdi, M.;
Lomascolo, A. J. Appl. Microbiol. 2006, 100, 219−232.
(3) (a) Duran, N.; Rosa, M. A.; D’Annibale, A.; Gianfreda, L. Enzyme
Microb. Technol. 2002, 31, 907−931. (b) Munjal, N.; Sawhney, S. K.
Enzyme Microb. Technol. 2002, 30, 613−619. (c) Sanz, V. C.; Mena, M.
L.; Gonzalez-Cortes, A.; Yanez-Sedeno, P.; Pingarron, J. M. Anal. Chim.
Acta 2005, 528, 1−8. (d) Arslan, A.; Kiralp, S.; Toppare, L.; Yagci, Y. Int.
J. Biol. Macromol. 2005, 35, 163−167. (e) Jaafar, A.; Musa, A.; Nadarajah,
K.; Lee, Y. H.; Hamidah, S. Sens. Actuators, B 2006, 114, 604−609.
(f) Zejli, H.; Hidalgo-Hidalgo de Cisneros, J. L.; Naranjo-Rodriguez, I.;
Liu, B.; Temsamani, K. R.; Marty, J. L. Anal. Chim. Acta 2008, 612, 198−
203. (g) Fiorentino, D.; Gallone, A.; Fiocco, D.; Palazzo, G.; Mallardi, A.
Biosens. Bioelectron. 2010, 25, 2033−2037. (h) Dincȩ r, A.; Becerik, S.;
Aydemir, T. Int. J. Biol. Macromol. 2012, 50, 815−820.
(4) (a) Thalmann, C. R.; Lotzbeyer, T. Eur. Food Res. Technol. 2002,
̈
214, 276−281. (b) Chen, T.; Embree, H. D.; Wu, L. Q.; Payne, G. F.
Biopolymers 2002, 64, 292−302. (c) Huang, T. H.; Kuwana, T.;
Warsinke, A. Biosens. Bioelectron. 2002, 17, 1107−1113. (d) Anghileri,
A.; Lantto, R.; Kruus, K.; Arosio, C.; Freddi, G. J. Biotechnol. 2007, 127,
508−519. (e) Marìn-Zamora, M. E.; Rojas-Melgarejo, F.; García-
́
Canovas, F.; García-Ruiz, P. A. J. Biotechnol. 2009, 139, 163−168.
(f) Tuncagil, S.; Kayahan, S. K.; Bayramoglu, G.; Arica, M. Y.; Toppare,
L. J. Mol. Catal. B: Enzym. 2009, 58, 187−193. (g) Hanifah, S. A.; Heng,
L. Y.; Ahmad, M. Anal. Sci. 2009, 25, 779−784. (h) Yin, H.; Zhou, Y.; Xu,
J.; Ai, S.; Cui, L.; Zhu, L. Anal. Chim. Acta 2010, 659, 144−150.
(i) Donato, L.; Algieri, C.; Miriello, V.; Mazzei, R.; Clarizia, G.; Giorno,
L. J. Membr. Sci. 2012, 407, 86−92.
ASSOCIATED CONTENT
* Supporting Information
■
S
(5) Guazzaroni, M.; Crestini, C.; Saladino, R. Bioorg. Med. Chem. 2012,
20, 157−166.
(6) Feng, W.; Ji, P. Biotechnol. Adv. 2011, 29, 889−895.
(7) Apetrei, I. M.; Rodriguez-Mendez, M. L.; Apetrei, C.; de Saja, J. A.
Sens. Actuators, B 2013, 177, 138−144.
Relationship diagram between absorbance and the concentration
of oxidized carbon nanotubes (ox-MWCNTs), plots of activity
(percent) of catalysts I and IV versus run in buffer and in organic
medium, linear regression equations and plots of Lineweaver−
Burk, Eadie−Hofstee, and Hanes analyses for native tyrosinase
and catalyst I, nonlinear regression plot for native tyrosinase and
catalyst I, and data for the identification and characterization of
the oxidation products. This material is available free of charge
via the Internet at http://pubs.acs.org.
(8) (a) Kong, L.; Huang, S.; Yue, Z.; Peng, B.; Li, M.; Zhang, J.
Microchim. Acta 2009, 165, 203−209. (b) Yina, H.; Zhoua, Y.; Xua, J.;
Aia, S.; Cuia, L.; Zhub, L. Anal. Chim. Acta 2010, 659, 144−150.
(c) Hashemnia, S.; Khayatzadeh, S.; Hashemnia, M. J. Solid State
Electrochem. 2012, 16, 473−479.
(9) Romao Sartori, E.; Campanha Vicentinia, F.; Fatibello-Filho, O.
̃
̃
AUTHOR INFORMATION
Corresponding Authors
*E-mail: marcello.crucianelli@univaq.it. Fax: (+39) 0862-
433753.
Talanta 2011, 87, 235−242.
■
(10) (a) Decher, G.; Schmitt, J. Prog. Colloid Polym. Sci. 1992, 89, 160−
164. (b) Decher, G. Science 1997, 277, 1232−1237.
(11) (a) Mateo, C.; Fernandes, B.; Van Rantwijk, F.; Stolz, A.; Sheldon,
R. A. J. Mol. Catal. B: Enzym. 2006, 38, 154−157. (b) Wilson, L.; Illanes,
*E-mail: saladino@unitus.it. Fax: (+39) 0761-357242.
Notes
The authors declare no competing financial interest.
A.; Abian
Biomacromolecules 2004, 5, 852−857. (c) Abian
C.; Fernandez-Lorente, G.; Palomo, J. M.; Fernan
Guisan, J. M.; Re, D.; Tam, A.; Daminatti, M. J. Mol. Catal. B: Enzym.
2002, 19−20, 295−303. (d) Abian, O.; Mateo, C.; Fernandez-Lorente,
G.; Palomo, J. M.; Fernandez-Lafuente, R.; Guisan, J. M. Biocatal.
́
, O.; Pessela, B. C. C.; Fernan
́
dez-Lafuente, R.; Guisan
, O.; Wilson, L.; Mateo,
dez-Lafuente, R.;
́
, J. M.
́
́
́
́
́
́
ACKNOWLEDGMENTS
■
́
́
F.S. thanks the University of L’Aquila for a research grant. The
Italian Space Agency (ISA) is greatly acknowledged for its
financial support.
Biotransform. 2001, 19, 489−503. (e) Barbosa, O.; Ortiz, C.; Berenguer-
Murcia, A.; Torres, R.; Rodrigues, R. C.; Fernandez-Lafuente, R. RSC
Adv. 2014, 4, 1583−1600. (f) Shah, S.; Sharma, A.; Gupta, M. N. Anal.
Biochem. 2006, 351, 207−213.
REFERENCES
(12) Asuri, P.; Karajanagi, S. S.; Sellitto, E.; Kim, D.-Y.; Kane, R. S.;
Dordick, J. S. Biotechnol. Bioeng. 2006, 95, 804−811.
(13) Robb, D. A.; Gutteridge, S. Phytochemistry 1981, 20, 1481−1485.
(14) (a) Iamsamai, C.; Soottitantawat, A.; Ruktanonchai, U.;
Hannongbua, S.; Dubas, S. T. Carbon 2011, 49, 2039−2045. (b) Lee,
J.; Park, S.-J.; Moon, Y.-K.; Kim, S.-H.; Koh, K. Korean J. Chem. Eng.
2009, 26, 1790−1794.
■
(1) For selected recent reviews on this general topic, see: (a) Brena, B.;
Gonzal
́
ez-Pombo, P.; Batista-Viera, F. Methods Mol. Biol. 2013, 1051,
15−31. (b) Barbosa, O.; Torres, R.; Ortiz, C.; Berenguer-Murcia, A.;
Rodrigues, R. C.; Fernandez-Lafuente, R. Biomacromolecules 2013, 14,
2433−2462. (c) Rodrigues, R. C.; Ortiz, C.; Berenguer-Murcia, A.;
́
Torres, R.; Fernandez-Lafuente, R. Chem. Soc. Rev. 2013, 42, 6290−
821
dx.doi.org/10.1021/cs400856e | ACS Catal. 2014, 4, 810−822

ACS Catalysis
Research Article
(15) Xiang, Y.; Zhang, Y.; Jiang, B.; Chai, Y.; Yuan, R. Sens. Actuators, B
2011, 155, 317−322.
(16) Zioł
303−305.
́
kowska, D.; Shyichuk, A.; Zelazko, K. Polimery 2012, 57,
(17) Masamoto, Y.; Iida, S.; Kubo, M. Planta Med. 1980, 40, 361−365.
(18) (a) Martın, M. T.; Plou, F. J.; Alcalde, M.; Ballesteros, A. J. Mol.
Catal. B: Enzym. 2003, 21, 299−308. (b) Knezevic, Z.; Milosavic, N.;
Bezbradica, D.; Jakovljevic, Z.; Prodanovic, R. Biochem. Eng. J. 2006, 30,
269−278.
(19) Gao, Y.; Kyratzis, I. Bioconjugate Chem. 2008, 19, 1945−1950.
(20) Yu, C.-M.; Yen, M.-J.; Chen, L.-C. Biosens. Bioelectron. 2010, 25,
2515−2521.
(21) Shah, S.; Solanki, K.; Gupta, M. N. Chem. Cent. J. 2007, 1, 30−36.
(22) (a) Piao, Y.; Jin, Z.; Lee, D.; Lee, H.-J.; Na, H.-B.; Hyeon, T.; Oh,
M.-K.; Kim, J.; Kim, H.-S. Biosens. Bioelectron. 2011, 26, 3192−3199.
(b) Ren, J.; Kang, T.-F.; Xue, R.; Ge, C.-N.; Cheng, S.-Y. Microchim. Acta
2011, 174, 303−309.
(23) Horcas, I.; Fernan
Gomez-Herrero, J.; Baro, A. M. Rev. Sci. Instrum. 2007, 78, 013705/1−
013705/8.
́ ́
dez, R.; Gomez-Rodríguez, J. M.; Colchero, J.;
́
(24) (a) Kazandjiand, R. A.; Klibanov, M. J. Am. Chem. Soc. 1985, 107,
5448−5450. (b) Kermasha, S.; Tse, M. J. Chem. Technol. Biotechnol.
2000, 75, 475−483. (c) Kermasha, S.; Bao, H.; Bisakowski, B.; Yaylayan,
V. J. Mol. Catal. B: Enzym. 2001, 11, 929−938. (d) Kermasha, S.; Bao,
H.; Bisakowski, B. J. Mol. Catal. B: Enzym. 2002, 19, 335−345.
(25) Guazzaroni, M.; Pasqualini, M.; Botta, G.; Saladino, R.
ChemCatChem 2012, 4, 89−99.
(26) Torchilin, V. P.; Maksimenko, A. V.; Smirnov, V. N.; Berezin, I. V.;
Klibanov, A. M.; Martinek, K. Biochim. Biophys. Acta 1978, 522, 277−
283.
(27) Aytar, B. S.; Bakir, U. Process Biochem. (Oxford, U.K.) 2008, 43,
125−131.
(28) (a) Ren, L.; Zhong, W. Environ. Sci. Technol. 2010, 44, 6954−
6958. (b) Ren, L.; Yan, D.; Zhong, W. Carbon 2012, 50, 1303−1310.
(29) (a) Zheng, L. T.; Ryu, G. M.; Kwon, B. M.; Lee, W. H.; Suk, K.
Eur. J. Pharmacol. 2008, 588, 106−113. (b) Nolan, L. C.; O’Connor, K.
E. Biotechnol. Lett. 2008, 30, 1879−1891. (c) Shimizu, T.; Nakanishi, Y.;
Nakahara, M.; Wada, N.; Moro-Oka, Y.; Hirano, T.; Konishi, T.;
Matsugo, S. J. Clin. Biochem. Nutr. 2010, 47, 181−190.
(30) (a) Carvalho, G. M. J.; Alves, T. L. M.; Freire, D. M. G. Appl.
Biochem. Biotechnol. 2000, 84, 791−800. (b) Ullrich, R.; Hofrichter, M.
Cell. Mol. Life Sci. 2007, 64, 271−293. (c) Otavio de Faria, R.; Rotuno-
Moure, V.; Lopes deAlmeida, M. A.; Krieger, N.; Mitchell, D. A. Food
Technol. Biotechnol. 2007, 45, 287−294.
(31) Burton, S. G.; Duncan, J. R.; Kaye, P. T.; Rose, P. D. Biotechnol.
Bioeng. 1993, 42, 938−944.
(32) Da Silva, A. M.; Junqueira, G. M. A.; Anconi, C. P. A.; Dos Santos,
H. F. J. Phys. Chem. C 2009, 113, 10079−10084.
(33) Ismaya, W. T.; Rozeboom, H. J.; Schurink, M.; Boeriu, C. G.;
Wichers, H.; Dijkstra, B. W. Acta Crystallogr. 2011, F67, 575−578.
(34) (a) Bradford, M. M. Anal. Biochem. 1976, 72, 248. (b) Sedmak, J.
J.; Grossberg, S. E. Anal. Biochem. 1977, 79, 544−552.
(35) Bi, S.; Zhou, H.; Zhang, S. Biosens. Bioelectron. 2009, 24, 2961−
2966.
(36) (a) Sweely, C. C.; Bentlry, R.; Makita, M.; Wells, W. W. J. Am.
Chem. Soc. 1963, 85, 2497−2507. (b) Rodríguez-Lop
Fenoll, L.; Penalver, M. J.; García-Ruíz, P. A.; Varon, V.; Martínez-Ortíz,
F.; García-Canovas, F.; Tudela, J. Biochim. Biophys. Acta 2001, 1548,
238−256.
́ ́
ez, J. N.; Gomez-
́
́
822
dx.doi.org/10.1021/cs400856e | ACS Catal. 2014, 4, 810−822