Biocatalytic oxidation by chloroperoxidase from Caldariomyces fumago in polymersome nanoreactors

Article abstract of DOI:10.1039/b911370c

The encapsulation of chloroperoxidase from Caldariomyces fumago (CPO) in block copolymer polymersomes is reported. Fluorescence and electron microscopy show that when the encapsulating conditions favour self-assembly of the block copolymer, the enzyme is incorporated with concentrations that are 50 times higher than the enzyme concentration before encapsulation. The oxidation of two substrates by the encapsulated enzyme was studied: i) pyrogallol, a common substrate used to assay CPO enzymatic activity and ii) thioanisole, of which the product, (R)-methyl phenyl sulfoxide, is an important pharmaceutical intermediate. The CPO-loaded polymersomes showed distinct reactivity towards these substrates. While the oxidation of pyrogallol was limited by diffusion of the substrate into the polymersome, the rate-limiting step for the oxidation of thioansiole was the turnover by the enzyme.

Full text of DOI:10.1039/b911370c

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www.rsc.org/obc | Organic & Biomolecular Chemistry
Biocatalytic oxidation by chloroperoxidase from Caldariomyces fumago
in polymersome nanoreactors†‡
H. M. de Hoog,a M. Nallani,a,b J. J. L. M. Cornelissen,*a,c A. E. Rowan,a R. J. M. Noltea and
I. W. C. E. Arends*d
Received 10th June 2009, Accepted 29th July 2009
First published as an Advance Article on the web 3rd September 2009
DOI: 10.1039/b911370c
The encapsulation of chloroperoxidase from Caldariomyces fumago (CPO) in block copolymer
polymersomes is reported. Fluorescence and electron microscopy show that when the encapsulating
conditions favour self-assembly of the block copolymer, the enzyme is incorporated with concentrations
that are 50 times higher than the enzyme concentration before encapsulation. The oxidation of two
substrates by the encapsulated enzyme was studied: i) pyrogallol, a common substrate used to assay
CPO enzymatic activity and ii) thioanisole, of which the product, (R)-methyl phenyl sulfoxide, is an
important pharmaceutical intermediate. The CPO-loaded polymersomes showed distinct reactivity
towards these substrates. While the oxidation of pyrogallol was limited by diffusion of the substrate into
the polymersome, the rate-limiting step for the oxidation of thioansiole was the turnover by the enzyme.
In recent decades we have seen the emergence of vesicular sys-
tems as scaffolds for the harboring of enzymes, of which the most
thoroughly studied systems are liposomes. Although progress in
Introduction
Biohybrid compounds that are able to form sub-micrometer (i.e.
nano-) stable capsules in an aqueous environment by self-assembly
the entrapping of enzymes has been made, liposomes generally
are of interest to the biomedical and physical fields from both
lack stability and disintegrate over time.7,8 In addition, the
a fundamental and applied perspective.1,2 When loaded with
permeability of the vesicle bilayers for externally added substrate
therapeutic proteins or enzymes, and made responsive to external
molecules is usually low.8 These shortcomings have been addressed
stimuli, these capsules may be used in diagnostics or targeted drug
by cross-linking the membrane and/or by the incorporation of
delivery. For diagnostics, the ideal system combines permeability
membrane channels.9 Recently, bottom-up engineered nano-sized
to analytes with shielding of its contents to the environment.3
capsule systems have emerged based on (amphiphilic) polymers, al-
Ideally, drugs or gene vectors loaded into nano-capsules are
lowing the tuning of the properties of the corresponding particles.10
protected from the biological surroundings, but should be readily
This has led to a diversity of functional nano-reactors containing
released once ingested by the target cell.4 Alternatively, one
enzymes.5,11-14 An interesting approach involves layer-by-layer
could envision self-assembled nano-capsules as mild immobi-
(LbL) assembly, by which oppositely charged polyelectrolytes are
lization agents for enzymes to perform environmentally benign
adsorbed on a sacrificial template.15,16 This technique allows the
biotransformations.5,6 Immobilization of biocatalysts is widely
incorporation of enzymes while an attractive characteristic is the
applied for the stabilization of the generally labile enzyme and
capability to modulate permeability of the thus-prepared capsule.17
facilitates removal of the catalyst from the reaction mixture. Often,
In addition, spatial confinement of different enzymes in the LbL-
immobilization is accompanied by activity loss as a result of either
constructs has been shown recently.18 Magnetic nanoparticles
diffusion limitation or the fact that the enzyme has now become a
constitute another emerging platform of enzyme immobilization,
heterogeneous instead of a homogeneous catalyst. Encapsulation
of which the main advantage is the ease by which the catalyst can
of enzymes into the nano-sized volumes of porous polymer
be removed from the reaction mixture.19
capsules diminishes the diffusion limitation while retaining the
A promising approach for the synthesis of nano-sized capsules
homogeneous nature of the system.
involves the direct self-assembly of amphiphilic block copolymers
into polymersomes.20,21 These structures are reminiscent of lipo-
somes but exhibit enhanced stability, and the possibility to encap-
sulate enzymes has been reported.22 As stability often comes with
reduced permeability, polymersomes have been equipped with
trans-membrane channels even though their membrane dimen-
sions exceed that of the natural cell membrane.23,24 Furthermore,
polymersomes have been exploited as a platform to covalently at-
tach enzymes.25-27 Interesting functional systems have been created
based on the triblock copolymer PMOXA-PDMS-PMOXA, rang-
ing from pH-switchable nano-reactors to nano-compartments for
the selective recovery of DNA.23,28 Unique in this aspect are poly-
mersomes assembled from polystyrene-b-poly(L-isocyanoalanine-
(2-thiophene-3-yl-ethyl)amide) (PS-PIAT), which are permeable
aInstitute for Molecules and Materials. Radboud University Nijmegen,
Heyendaalseweg 135, 6525 AJ, Nijmegen, The Netherlands
bInstitute of Materials Research & Engineering (IMRE), Research Link 3,
Singapore 117602
cLaboratory for Biomolecular Nanotechnology, MESA+ Institute, Univer-
sity of Twente, P.O. Box 217, 7500 AE, Enschede, the Netherlands. E-mail:
J.J.L.M.Cornelissen@tnw.utwente.nl; Fax: +3153 489 4645; Tel: +31 53
489 4380
dDept. of Biotechnology, Delft University of Technology, Julianalaan 136,
2628 DL, Delft, The Netherlands. E-mail: I.W.C.E.Arends@tudelft.nl
† Electronic supplementary information (ESI) available: Experimental
procedures and additional characterization. See DOI: 10.1039/b911370c
‡ This paper is part of an Organic & Biomolecular Chemistry web theme
issue on biocatalysis.
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to low-molecular weight organic molecules but at the same
time are able to retain enzymes.29,30 Furthermore, the positional
assembly of enzymes in either the membrane or the aqueous
inner compartment of PS-PIAT polymersomes has recently been
described.31
solution in THF (1 mg ml-1) to 5 volumes of an aqueous solution
containing enzymes (0.2–2 mg ml-1).31,37 During formation of
the polymersomes by self-assembly, statistically a fraction of the
enzymes is incorporated into the polymersomes. The solution is
left to equilibrate for 24 hours and the turbid solution is filtered, to
remove the non-encapsulated enzyme. Both the enzymatic activity
of the flow-throughs and the activity of the supernatant containing
the polymersomes is measured. Ideally, after sufficient washings
the activity of the flow-throughs reaches zero while the activity of
the supernatant is retained, demonstrating the incorporation of
the enzyme into the polymersomes.
For incorporation of CPO inside the polymersomes the proce-
dure was adjusted. First, the final volume percentage of THF was
lowered, because at the initial volume percentage (20% v/v) the
enzyme was irreversibly deactivated (Fig. S1†), as was observed by
determining its chlorinating activity in the presence of increasing
amounts of THF.38 It was found that at a concentration of 10%
THF the enzyme showed similar activity as the enzyme in aqueous
buffer. In order to have a concentration of PS-PIAT similar to
the previously used conditions for the successful encapsulation
of enzymes, the initial concentration of PS-PIAT in THF was
doubled.
Although the enzyme retained its activity in aqueous THF,
encapsulation in PS-PIAT polymersomes was not successful under
the conditions (pH = 3, 0.1 M phosphate/citric acid buffer)
used in a halogenation assay. After several washings with buffer
no chlorination activity was left in the flow-throughs or in the
supernatant, indicating that all active protein was washed out.
Upon examination of the unfiltered polymersomes by TEM,
spherical aggregates were still visible, ranging in diameter from
~500 nm (the size of the regular enyzme-containing polymersomes)
to aggregates of more than 10 mm (Fig. S2†). Clearly the
combination of enzyme and buffer has a noticeable effect on
the aggregation properties of the PS-PIAT and apparently affects
the encapsulation as well. This is also the case for previously
encapsulated enzymes like CALB, HRP and GOx, of which
the resulting polymersomes showed varying sizes and different
membrane characteristics.31,37 In addition, dispersions of empty
polymersomes precipitated after a few days, whereas co-aggregates
of PS-PIAT and proteins stayed intact for months.30
Enzymes are environmentally benign catalysts and well suited
for the synthesis of enantiopure synthons, which is of importance
to the fine-chemical and pharmaceutical industries.32 This fact
has inspired us to encapsulate industrially relevant enzymes inside
PS-PIAT polymersomes and study their activity. Conversion of
industrially interesting substrates in polymersomes has not been
demonstrated so far. Moreover, studies on nano-scale enzyme
reactors in aqueous environment generally are limited to substrates
selected on their desirable spectroscopic properties.3,6,11-14,17,33,34
Often, these substrates include fast-reacting and activated com-
pounds like p-nitrophenyl adducts, the structure of which relies
on the type of enzyme. Organic synthons do not exhibit these
favourable properties and generally react at different rates, de-
pending on solubility and affinity and, probably as a consequence
of this, have been studied in much less detail.
As an example of a potential biocatalyst useful to organic
synthesis, we decided to incorporate chloroperoxidase from Cal-
dariomyces fumago (CPO; EC 1.1.11.1) into the polymersomes.
CPO fills the niche between the cofactor-dependant cytochrome
P450 oxygenases and the hemeperoxidases, and is unique in its
ability to catalyze the enantioselective epoxidation of a variety of
alkenes.35 Its industrial application is, however, still hindered by the
rapid deactivation of the enzyme in the presence of oxidant. At this
moment the mechanism of deactivation is not well understood but
seems to result from degradation of the heme-group of the enzyme
or the oxidation of the axial cysteine ligand.36 Encapsulation of
CPO into polymersomes might have a stabilizing effect on the
limited operational stability of the enzyme. In this paper, we
report on a first step in this direction, i.e., by the functional
encapsulation of CPO into PS-PIAT polymersomes and the study
of the oxidation of two different substrates (Scheme 1). We found
that the oxidation of pyrogallol to purpurogallin, a reaction
commonly used to assay peroxidase activity, shows a different
kinetic behaviour than the oxidation of thioanisole to (R)-phenyl
methyl sulfoxide – a chiral pharmaceutical intermediate.
As encapsulation of PS-PIAT at the standard operating condi-
tions of CPO was not trivial, CPO was incorporated under the self-
assembly conditions reported for PS-PIAT (milli-Q; pH ~ 6). It was
observed that under these unbuffered conditions the enzyme did
not behave differently than at pH = 5 in 50 mM acetate buffer (Fig.
S3†). At pH 5 or above the halogenating activity of CPO is lost
but the enzyme is still active in the oxidation of various substrates,
which is our reaction of interest.35,39,40 The CPO-polymersome
(CP) activity was measured as the rate of oxidation of pyrogallol
to purpurogallin (Scheme 1).41 The oxidation activity apparently
was not affected by the presence of THF.
After preparation of the polymersomes and filtering the dis-
persion, it was found that the oxidation activity was retained in
the supernatant, indicating encapsulation of the enzyme inside the
polymersomes. Investigation of the aggregates by TEM revealed
the presence of spherical objects, which, in contrast to the
typical membrane-structure observed for empty polymersomes,
showed a uniform density. However, upon closer inspection the
membrane was clearly visible (Fig. S4†). Interestingly, functional
Scheme 1 Chloroperoxidase-catalyzed reactions studied in this work.
Results and discussion
Encapsulation of chloroperoxidase
In general, the incorporation of enzymes into PS-PIAT poly-
mersomes is achieved by addition of 1 volume of a PS-PIAT
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Table 1 Concentration of CPO, as determined by UV-vis, in the initial
polymersome preparation (before filtration) and in the filtered dispersion.
Also shown is the estimated local concentration of the enzyme in the
volume of the polymersomes
encapsulation only occurred at starting concentrations of CPO of
3 mM (0.1 mg ml-1) or higher. Based on the activity of the resulting
CPs in the oxidation of pyrogallol, the starting concentration
of CPO (i.e. the concentration of CPO before filtering) did not
seem to have an effect on the eventual activity of the aggregates
(Fig. 1). Varying the starting concentration of CPO between 7,
23 and 36 mM (0.3, 0.9 and 1.5 mg ml-1) resulted in an activity
between 0.4 abs min-1 and 0.6 abs min-1, and was lowest for
the CPs with a starting concentration of 23 mM. In contrast,
activity measurements with the free enzyme in a similar activity
range showed a linear relation between activity and enzyme
concentration (Fig. 1 inset). From a comparison with the data-
points in the graph in the inset, the apparent amount of enzyme in
the polymersomes can be calculated. For the CPs prepared from
a 23 mM CPO solution, an activity was found of 0.4 abs min-1.
This corresponds to a concentration of 0.1 mM, indicating an
encapsulation efficiency of 0.5%. Although this is a small number,
a low value may be expected when the estimated volume fraction of
the polymersomes is taken into account (0.1%), and it is assumed
that encapsulation is based solely on statistics.31 On the other hand,
the activity of the CPs was found to be considerably lower than
observed previously.31
Initial CPO concentration (mM)a
9 ( 4)
23
36
Mean concentration in filtered
solution (mM)a
0.05 ( 0.04) 0.9 ( 0.4) 2 ( 0.4)
Encapsulation (%)
0.6
4.0
0.8
5.9
2.0
Local concentration in CPs (mM)b 0.05
a Mean of 3 measurements. b Based on the assumption that the volume frac-
tion of the polymersomes is 0.0011 and encapsulation occurs statistically.31
Fig. 2 (Left) UV-Vis spectra of CPs containing different loadings of
CPO (solid line). The absorption spectrum of free CPO exhibiting the
characteristic absorption of the heme at l = 420 nm (dashed line) is shown,
as well as the absorption spectrum of empty polymersomes (dotted line).
(Right) Absorption spectra of a dispersion of PS-PIAT and CPO labelled
with Alexa-633 dye, before and after filtering off the non-encapsulated
enzyme. The initial concentration of CPO was 23 mM. The straight lines
underneath the absorption peaks were taken as the base-line for the
calculation of the ratio between the peaks.
To probe the location of the enzyme inside the polymersomes,
the enzyme was labeled with the fluorescent dye Alexa-633 and
encapsulated, after which the encapsulated system was studied
with confocal fluorescence microscopy (CFM, Fig. 3). Polymer-
somes containing the labeled enzyme were prepared following
Fig. 1 Enzymatic activity measured at l = 420 nm by the pyrogallol assay
(abs min-1) of CPs obtained by filtration of CPO/PS-PIAT dispersions with
increasing concentrations of enzyme (mM). The inset shows the dependence
of the activity on the concentration of the free enzyme (mM) in the same
activity range (abs min-1).
The observed results prompted us to investigate in more detail
the concentration and the location of CPO in the polymersomes.
Since the concentration of CPO can be determined from the
absorbance of the heme group at l = 400 nm, the concentration
of CPO inside the polymersomes was determined by UV-Vis
measurements on the aggregate solutions.42 The data showed
large absorption/scattering from the polymersomes, but also the
distinct absorption of the enzyme was visible (Fig. 2; left graph). To
estimate the concentrations of the CPO inside the polymersomes,
the absorbance of the empty polymersomes was subtracted from
the absorption of the CPs. These calculations showed that un-
like the outcome of the pyrogallol activity measurements, the
concentration of enzyme inside the polymersomes increased
linearly from 0.05 to 2 mM with increasing starting concentration
of enzyme (Table 1). Interestingly, the measured concentration of
the enzyme inside the polymersomes appeared to be considerably
higher than the concentration that was determined from the
activity tests.
Fig. 3 (Left; blue) Confocal scanning fluorescence microscope image
at lem = 505 nm of polymersomes loaded with Alexa-633 labeled CPO
(initial concentration 23 mM) showing fluorescence of the PS-PIAT itself.
(Right; red) Fluorescence image at lem = 650 nm of polymersomes
loaded with labeled CPO showing the emission from the labeled CPO.
The polymersome distribution on the substrate is the same at both
wavelengths, as is emphasized by the white squares. For fluorescence images
of non-labeled CPO encapsulated in the polymersomes at both emission
wavelengths; see Fig. S5†.
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exactly the same procedure as was used for the non-labeled
enzyme (enzyme concentration before filtration = 23 mM). The
fluorescence emission from the CPs containing the labeled CPO
and the non-labeled protein was then studied both at lem = 505 nm
(lex = 488 nm) and lem = 650 nm (lex = 633). At lem = 505 nm
for both systems emission was observed, originating from the
PS-PIAT polymer itself (as emission from the dye is absent below
l = 600 nm). When lex = 633 nm a different pattern was found, that
is, fluorescence emission was only observed for CPs with labeled
CPO. No signal was observed for the CPs containing non-labeled
enzyme. Furthermore when overlaying the fluorescence images at
lem = 505 nm and lem = 650 nm of CPs containing labeled CPO,
the same distribution pattern was observed (Fig. 3), confirming
that CPO was located inside the polymersomes.
The dye labeling also provided a means of determining the
concentration of CPO inside the polymersomes more accurately
(Fig. 2; right graph). The labeled CPO shows an intense absorption
at l = 633 nm, and this signal is virtually undisturbed by the
adsorption from the polymersomes compared to the signal of
the heme at l = 400 nm. The ratio in absorption between the
filtered CPs and the CPs before filtration gives the percentage
of encapsulation, which for a labeled CPO solution containing
an initial concentration of 23 mM enzyme gives an encapsulation
efficiency of 4%. Moreover, dye labeling experiments with initial
CPO concentrations of 9, 18, 27 to 36 mM showed a concurrent,
linear increase in CPO concentration after filtration, indicating
that the encapsulation efficiency was constant over the concen-
trations studied. These results correlated with the calculations
based on the UV-Vis measurements on the unlabeled protein
encapsulated in the polymersomes. The CPs prepared from 7 mM
CPO solutions give a slightly lower value, which might be the result
of disturbance of the heme-signal by the absorption/scattering
of the polymersomes. For the CPs prepared from 36 mM CPO
solutions, the enzyme concentration appears to be 50 times what
it would be if encapsulation is considered to be a purely statistical
process. Assuming that the volume fraction of the polymersomes
is 0.0011,31 this would lead to a concentration of ~0.04 mM. In
the present case, the local concentration of the enzyme inside
the polymersomes is estimated to be 2.0 mM (Table 1). This is
a considerable concentration, and may explain partly why for
CALB, HRP and GOx it was found that the encapsulated enzyme
was 100-fold more active than the free enzyme.31
valuable chiral auxiliaries, and the sulfoxyl moiety is frequently
found in bioactive natural products and synthetic drugs.43 The
enzymatic oxidation of this compound by CPO has been studied in
detail in the field of biocatalysis.44-47 Depending on the conditions
used the reaction affords the corresponding (R)-sulfoxide in high
ee (99%) and high yields. When the enzyme concentration is kept
sufficiently high, oxidation occurs very rapidly, thereby avoiding
the non-enzymatic oxidation of the substrate.
It turned out that sulfoxidation also proceeded readily inside
the CPs, showing that the structure of the polymersomes allowed
substrate and oxidant to reach the active site of the enzyme
and did not hamper the introduction of the oxygen into the
substrate. Furthermore, based on gas chromatography (GC), the
polymersome-anchored enzyme allowed the formation of the
desired sulfoxide in the same high enantioselectivity as observed
for the free enzyme. When the oxidation of thioanisole by the
CPs was followed over time, a subtle effect of the polymersome
structure on the kinetics was observed (Fig. 4). Although the
initial rates of the CPs and free CPO were of the same order of
magnitude, the free CPO at a concentration of 60 nM (comparable
apparent activity as the encapsulated enzyme) already showed
100% conversion of substrate after 2 minutes, whereas for the CPs
thioanisole conversion was only 39–65%. When the concentration
of free CPO was increased to 1 mM (concentration determined by
UV for the enzyme concentration inside the polymersomes), full
conversion was already observed after 40 seconds.
Fig. 4 Sulfoxidation of thioanisole in time by CPs obtained from filtration
of 7 (᭺), 23 (ꢀ) and 36 mM (ꢀ) dispersions of CPO and PS-PIAT. The
data reported are the average of three measurements. The curves obtained
with free CPO at 60 nM (dotted line) and 1 mM (dashed line) are also
shown.
The reduced activity of the CPs in the oxidation of pyrogallol,
and the non-linear variation of the activity of the CPs in the
oxidation of pyrogallol with increasing enzyme-loading, points
to diffusion limitation which is imposed by locating the enzyme
inside the polymersomes. Most likely this effect results from
the substrate having to cross the physical barrier of the block
copolymer membrane in order to reach the active site of the
enzyme (see later).
To study if the same effect was observed for the oxidation of
thioanisole as for the oxidation of pyrogallol (Fig. 1), CPs were
prepared with the same starting concentrations of CPO, e.g., 7,
23, and 36 mM (Fig. 5) and after 40 s the yields for sulfide
oxidation were determined. Within this short reaction time the
conversion is still in the linear phase and deactivation of CPO due
to hydrogen peroxide is not important.41 Based on the activity for
thioanisole sulfoxidation of the free enzyme the concentration
of the enzyme inside the polymersomes was 0.1 mM (enzyme
concentration before filtration = 23 mM), approaching the number
calculated for the pyrogallol oxidation. However, with respect to
this substrate a trend was now observed in which the initial rate
of the reaction, Vi, was lowest for the polymersomes containing
the lowest concentration of enzyme, and Vi was highest for the
polymersomes containing the highest enzyme concentration (see
also Fig. 4). Although this is in agreement with the data observed
Sulfoxidation inside CPs
Following the successful encapsulation of CPO in the polymer-
somes and the study of the oxidation of pyrogallol, we decided to
also study the conversion of a relevant model organic synthon. For
these investigations the sulfoxidation of thioanisole (methyl phenyl
sulfide) to (R)-methyl phenyl sulfoxide was chosen (Scheme 1).
Chiral sulfoxides have been used in asymmetric synthesis as
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Table 2 Kinetic parameters for sulfoxidation of thioanisole by CPs and
free CPO
CP
CPO (20 nM)a
CPO (300 nM)
KM
kcat
5
4
400
3
275
34b
a Activity determined by UV-Vis. b Assuming the enzyme concentration in
the polymersomes is that found by direct measurement of the concentration
by UV-Vis.
Fig. 5 Variation of the initial rate of the sulfoxidation of thioanisole of
CPs obtained from filtration of 7, 23 and 36 mM dispersions of CPO
and PS-PIAT. Upon the increase of enzyme concentration inside the
polymersomes, the activity increases as well, as is the case for the free
enzyme (inset).
for sulfoxidation by the free CPO, it is in contrast to the results
obtained with pyrogallol (compare Fig. 1 and 5).
To explain these observations, we consider that sulfoxidation
of thioanisole is a much slower process (kcat = 200 s-1)40 than
the oxidation of pyrogallol (kcat = 2500 s-1)48 and that the
kinetics of substrate conversion in the polymersome is a 2-step
process (Fig. 6). The first step, represented by the rate constant
Fig. 6 Model proposed to explain the different kinetics observed for
pyrogallol and thioanisole conversion by chloroperoxidase encapsulated
in CPs. Two steps can be discerned: 1) diffusion of the substrate into the
polymersome, 2) the conversion of the substrate by the enzyme.
k
transfer, involves the diffusion of the substrate molecules into the
before filtration = 23 mM)) than the concentration measured by
optical spectroscopy (~0.9 mM). For pyrogallol oxidation this
partial deactivation appears to be less relevant, because the rate-
limiting step is ktransfer. For thioanisole oxidation this translates to
a value for the turnover number (kcat = 34 s-1) that is an order of
magnitude lower than the value for the free enzyme (kcat = 400 s-1).
polymersome, which depends on the membrane properties. The
second step is the turnover by the enzyme and is represented by kcat,
assuming that Michaelis–Menten kinetics is obeyed. The former
rate constant, ktransfer, is a complex parameter and depends on the
concentration of the substrate both inside, [S]in, and outside, [S]out
,
the polymersomes (including such parameters as the local block
copolymer structure, the enzyme used, pH and polarity of the
solvent). Because of the high kcat of pyrogallol by the enzyme,
the rate-limiting step for the CPs, in this case, is ktransfer. For the
sulfoxide formation the rate-limiting step is probably turnover
by the enzyme, because kcat is much lower. In this case ktransfer
ꢀ kcat and as a result ktransfer does not seem to have an effect
on the conversion of the thioanisole. The encapsulation of the
enzymes into the polymersomes evidently introduces an additional
parameter to the enzyme kinetics, that is the ratio ktransfer/kcat. At
high ratios the substrate conversion by the enzyme is unrestricted
while at lower ratios the turnover frequency is dictated by the
polymersome properties. This property of the polymersomes is
important as it may be useful to tune reaction rates relative to
each other in cascade reactions.31
The observation that the sulfoxidation activity varied linearly
with the enzyme concentration inside the polymersomes suggests
that the kinetics can be described by the Michaelis–Menten
model. For peroxidases (being two-substrate enzymes) these
parameters can be determined when the H2O2 concentration is
kept at saturating concentrations (>5 mM) and the substrate
concentration is varied.49,50 Determining the kinetic parameters
indeed showed that the encapsulated enzyme displayed the same
saturation kinetics as the free enzyme and the KM for thioanisole
appeared to be similar (Table 2 and Fig. S6†). The comparable
KM suggests that [S]in = [S]out and that the kinetics in the case of
sulfoxidation with the CPs are determined by step 2 (Fig. 6).
For both the pyrogallol oxidation and the thioanisole oxidation
the concentration of CPO inside the polymersomes, determined
on the basis of activity, is lower (~0.1 mM, (enzyme concentration
This lower catalytic efficiency cannot be the result of the ktransfer
,
as it was concluded that the diffusion into the membrane is not
important for thioanisole.
The decrease in catalytic efficiency could either be caused by a
deactivation of the enzyme or by the fact that a significant fraction
of the enzyme present in the polymersomes is not accessible to
substrate molecules. In the latter case it can be calculated that
the activity observed only comes from ~10% of the encapsulated
enzyme molecules. We were able to discriminate between these
two possibilities by the observation that 43% of the enzymatic
activity could be released from the polymersomes by redispersion
of the CPs upon filtration in buffer, while 39% remained in the
polymersomes (Fig. S7†). When the activity of the filtrates and
the remaining activity in the polymersomes were tested for their
protein content similar percentages were found (Fig. S8†), showing
that the protein population as a whole was deactivated.
Conclusion
In this paper we have described the functional encapsulation of
chloroperoxidase (CPO) into PS-PIAT polymersomes. It is shown
that CPO can be encapsulated with an efficiency of 5% under
conditions that favour self-assembly of the PS-PIAT molecules.
In the oxidation of pyrogallol catalyzed by CPO, it was found
that the activity of the enzyme is 0.4% with respect to the activity
before encapsulation. Furthermore, it was observed that there is
no linear correlation between the activity of the CPO-containing
polymersomes (CPs) and the concentration of the enzyme in the
polymersomes, suggesting that the rate of conversion of pyrogallol
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is limited by the transfer of substrate molecules over the mem-
brane. In order to show the potential of nano-structured materials
for the immobilization of enzymes, we also studied the oxidation
of thioanisole to the corresponding (R)-sulfoxide, an important
organic synthon. Interestingly, this substrate was readily oxidized
with the same high enantioselectivity. The kinetic parameters for
the oxidation of thioanisole by the CPs suggest that for this reac-
tion the turnover by the enzyme is the rate-limiting factor. The dif-
ferent kinetic behaviours of the substrates is explained by a model
that takes into account the rate of diffusion into the polymersomes
and the rate of conversion of the substrate by the enzyme.
Acknowledgements
Victor I. Claessen (HFML) is acknowledged for help with
the CSLM measurements and Dr. M. Koepf for providing
the rhodamine-azide. The Research School NRSC-Catalysis, the
Royal Netherlands’ Academy for Arts and Sciences, and the
Chemical Council of the Netherlands Organization for Scientific
Research are acknowledged for financial support.
References
1 M. Antonietti and S. Fo¨rster, Adv. Mater., 2003, 15, 1323–1333.
2 D. M. Vriezema, M. C. Aragones, J. A. A. W. Elemans, J. J. L. M.
Cornelissen, A. E. Rowan and R. J. M. Nolte, Chem. Rev., 2005, 105,
1445–1489.
3 E. W. Stein, D. V. Volodkin, M. J. McShane and G.B. Sukhorukov,
Biomacromolecules, 2006, 7, 710.
Experimental section
UV analysis was performed on a Shimadzu UV-4201 SPC
spectrophotometer using an electronic stirrer (Rank Brothers) at
400 rpm at 21 ◦C. Chiral GC was performed on a Shi-
madzu GC 17A with AOC-201 automatic injector. Electron
microscopy including sample preparation was carried out as
reported previously.30 CFM was carried out with a set-up reported
previously.51 However, to discriminate between fluorescence emis-
sion from PS-PIAT (lem = 505 nm) and the fluorescent dye, the en-
zyme was labelled with Alexa-633 succinimidyl ester and purified
following the protocol supplied. The average degree of labelling
was 1.4 moles of Alexa-633 per mole of enzyme. Encapsulation
was performed under conditions as described in the text.
4 Y. Hu, T. Litwin, A. R. Nagaraja, B. Kwong, J. Katz, N. Watson and
D. J. Irvine, Nano Lett., 2007, 7, 3056.
5 J. Kim, J. W. Grate and P. Wang, Chem. Eng. Sci., 2006, 61, 1017.
6 J. Kim, H. F. Jia and P. Wang, Biotech. Adv., 2006, 24, 296.
7 P. Walde and S. Ichikawa, Biomol. Eng., 2001, 18, 143–177.
8 P. Walde and B. Marzetta, Biotechnol. Bioeng., 1998, 57, 216.
9 G. E. Lawson, F. Y. Lee and M.R.A. Singh, Adv. Funct. Mater., 2005,
15, 267–272.
10 H. Burt and K. Letchford, Eur. J. Pharm,. Biopharm., 2007, 65.
11 Y. Lvov, A. A. Antipov, A. Mamedov, H. Mohwald and G. B.
Sukhorukov, Nano Lett., 2001, 1, 125.
12 M. J. Boerakker, N. E. Botterhuis, P. H. H. Bomans, P. M. Frederik,
E. M. Meijer, R. J. M. Nolte and N.A. J. M. Sommerdijk, Chem. Eur. J.,
2006, 12, 6071.
CPO (EC 1.11.1.10) was bought from BioChemika as a dark-
brown suspension in 0.1M phosphate buffer, pH = 4. The suspen-
sion was used as received. The stated amount of units was 22731
U ml-1. UV showed the Rz value to be 0.7 (Fig. S9†), and the con-
centration of the CPO in the solution to be ~25 mg ml-1 (600 mM).
Halogenase activity was measured by the monochlorodimedon
(2-chloro-5,5-dimethyl-1,3-cyclohexanedione) assay, and peroxi-
dase activity was measured by the pyrogallol (1,2,3-trihydroxyben-
zene) assay.38,41 Peroxidase activity was always measured by UV
as the increase in absorbance per minute at l = 420 nm. The
reaction was initiated by the addition of 50 ml of the sample
solution to a cuvette containing 2.9 ml of Milli-Q solution
(pH = 6), pyrogallol (43 mM) and H2O2 (9 mM). As the assay
conditions were constant during the study, pyrogallol activity is
reported in AU min-1. For thioanisole oxidation typical reaction
volumes were 250 ml, comprising 100 ml of the CPO-containing
polymersomes, an additional amount of substrate (always taken
from an 18 mM thioanisole stock in milli-Q and extensively
dispersed before addition) and an 18 mM solution of H2O2. The
reaction was initiated by addition of the oxidant and shaken
on a Thermotwister Comfort (Quantifoil instruments) at 5 rpm
at 21 ◦C. Reaction times were 40 s. After the desired time the
reaction was quenched by the addition of Na2SO3. Complete
quenching of the oxidant was checked with peroxide test sticks
(Quantofix). The solution was extracted with 0.50 ml DCM
containing hexadecane as the internal standard. After drying over
a small column of anhydrous magnesium sulfate the phases were
separated by spinning down for 10 s and the solution was directly
analyzed for yield and ee by GC on a Chiraldex G-TA column
(50 m ¥ 0.25 mm ¥ 0.12 mm), running isothermally at 160 ◦C for
20 min (column flow: 0.9 ml min-1; split ratio: 1/60). Elution times
were 7.2 min for (R)-methyl phenyl sulfoxide and 11.1 min for
(S)-methyl phenyl sulfoxide.
13 J. Rubio, F. Tamimi Marino and E. Lopez, J. Biomed. Mater. Res., Part
B, 2007, 83, 145–152.
14 E. A. Simone, T. D. Dziubla, F. Colon-Gonzalez, D. E. Discher and
V. R. Muzykantov, Biomacromolecules, 2007, 8, 3914–3921.
15 G. Decher, Science, 1997, 277, 1232.
16 C. S. Peyratout and L. Dahne, Angew. Chem., Int. Ed., 2004, 43, 3762–
3783.
17 A. Yu, Z. Liang and F. Caruso, Chem. Mater., 2005, 17, 171.
18 O. Kreft, M. Prevot, H. Mo¨hwald and G. B. Sukhorukov, Angew.
Chem., Int. Ed., 2007, 46, 5605–5608.
19 A. Dyal, K. Loos, M. Noto, S. W. Chang, C. Spagnoli, K. V. Shafi, A.
Ulman, M. Cowman and R. A. Gross, J. Am. Chem. Soc., 2003, 125,
1684.
20 K. Kita-Tokarczyck, J. Grumelard, T. Haefele and W. Meier, Polymer,
2005, 46, 3540.
21 L. Zhang, K. Yu and A. Eisenberg, Science, 1996, 272, 1777.
22 B. M. Discher, Y. Y. Won, D. S. Ege, J. C. M. Lee, F. S. Bates, D. E.
Discher and D. A. Hammer, Science, 1999, 284, 1143.
23 P. Broz, S. Driamov, J. Ziegler, N. Ben-Haim, S. Marsch, W. Meier and
P. Hunziker, Nano Lett., 2006, 6, 2349.
24 H. J. Choi and C. D. Montemagno, Nano Lett., 2005, 5, 2538.
25 J. A. Opsteen, R. P. Brinkhuis, R. L. M. Teeuwen, D.W. P. M. Lo¨wik
and J.C.M. van Hest, Chem. Commun., 2007, 3136.
26 B. Li, A. L. Martin and E. R. Gillies, Chem. Commun., 2007, 5217.
27 S. F. M. Van Dongen, M. Nallani, S. Schoffelen, J. J. L. M. Cornelissen,
R. J. M. Nolte and J.C.M. van Hest, Macromol. Rapid Commun., 2008,
29, 321.
28 M. Nallani, S. Benito, O. Onaca, A. Graff, M. Lindemann, M.
Winterhalter, W. Meier and U. Schwaneberg, J. Biotechnol., 2006, 123,
50–59.
29 D. M. Vriezema, J. Hoogboom, K. Velonia, K. Takazawa, P. C. M.
Christianen, J. C. Maan, A. E. Rowan and R. J. M. Nolte, Angew.
Chem., Int. Ed., 2003, 42, 772.
30 H.-P. M. de Hoog, D. M. Vriezema, M. Nallani, S. Kuiper, J. J. L. M.
Cornelissen, A. E. Rowan and R. J. M. Nolte, Soft Matter, 2008, 4,
1003–1010.
31 D. M. Vriezema, P. M. L. Garcia, N. S. Oltra, N. S. Hatzakis, S. M.
Kuiper, R. J. M. Nolte, A. E. Rowan and J. C. M. van Hest, Angew.
Chem., Int. Ed., 2007, 46, 7378.
32 R. A. Sheldon, I. W. C. E. Arends and U. Hanefeld, Green Chemistry
and Catalysis, Wiley-VCH, 2007.
This journal is
The Royal Society of Chemistry 2009
Org. Biomol. Chem., 2009, 7, 4604–4610 | 4609
©

View Article Online
33 A. Kawamura, A. Harada, K. Kono and K. Kataoka, Bioconjugate
Chem., 2007, 18, 1555.
34 N. Bruns and J. C. Tiller, Nano Lett., 2005, 5, 45.
35 V. M. Dembitsky, Tetrahedron, 2003, 59, 4701–4720.
36 C. E. Grey, M. Hedstro¨m and P. Adlercreutz, ChemBioChem, 2007, 8,
1055–1062.
44 S. Kobayashi, M. Nakano and T. Goto, Biochem. Biophys. Res.
Commun., 1986, 135, 166–171.
45 S. Colonna, N. Gaggero, A. Manfredi, L. Casella, M. Gullotti, G.
Carrea and P. Pasta, Biochemistry, 1990, 29, 10465–10468.
46 S. Lutz, K. Vuorilehto and A. Liese, Biotechnol. Bioeng., 2007, 98,
525–534.
37 M. Nallani, H.-P. M. de Hoog, J. J. L. M. Cornelissen, A. R. A. Palmans,
J. C. M. van Hest and R. J. M. Nolte, Biomacromolecules, 2007, 8, 3723–
3728.
38 D. R. Morris and L. P. Hager, J. Biol. Chem., 1966, 241, 1763.
39 J. A. Thomas, D. R. Morris and L. P. Hager, J. Biol. Chem., 1970, 245,
3129.
47 N. Spreti, R. Germani, A. Incani and G. Savelli, Biotechnol. Prog.,
2004, 20, 96–101.
48 M. B. McCarthy and R. E. White, J. Biol. Chem., 1983, 258, 9153–
9158.
49 R. B. Bhatia, C. J. Brinker, A. K. Gupta and A. K. Singh, Chem. Mater.,
2000, 12, 2434–2441.
40 F. Van Rantwijk and R. A. Sheldon, Curr. Opin. Biotechnol., 2000, 11,
50 P. Toti, A. Petri, V. Pelaia, A. M. Osman, M. Paolini and C. Bauer,
Biophys. Chem., 2005, 114, 245–251.
554–564.
41 J. B. Park and D. S. Clark, Biotechnol. Bioeng., 2006, 93, 1190–1195.
42 P. F. Hallenberg and L. P. Hager, Methods Enzymol., 1978, 52, 521–529.
43 J. Gawronski, J. Grajewski, J. Drabowicz and M. Mikolajczyk, J. Org.
Chem., 2003, 68, 9821–9822.
51 M. Comellas-Aragone`s, H. Engelkamp, V. I. Claessen, N. A. J. M.
Sommerdijk, A. E. Rowan, P. C. M. Christianen, J. C. Maan, B. J. M.
Verduin, J. J. L. M. Cornelissen and R. J. M. Nolte, Nat. Nanotechnol.,
2007, 2, 635.
4610 | Org. Biomol. Chem., 2009, 7, 4604–4610
This journal is
The Royal Society of Chemistry 2009
©