New Monodisperse Magnetic Polymer Microspheres Biofunctionalized for Enzyme Catalysis and Bioaffinity Separations

Article abstract of DOI:10.1002/mabi.201100393

Magnetic macroporous PGMA and PHEMA microspheres containing carboxyl groups are synthesized by multi-step swelling and polymerization followed by precipitation of iron oxide inside the pores. The microspheres are characterized by SEM, IR spectroscopy, AAS

Full text of DOI:10.1002/mabi.201100393

Full Paper
New Monodisperse Magnetic Polymer
Microspheres Biofunctionalized for Enzyme
Catalysis and Bioaffinity Separations
Daniel Hor a´ k,* Jana Ku cˇ erov a´ , Lucie Koreck a´ , Barbora Jankovi cˇ ov a´ ,
Ji ˇr ´ı Palar cˇ ´ı k, Petr Mikul a´ sˇ ek, Zuzana B ´ı lkov a´
Magnetic macroporous PGMA and PHEMA microspheres containing carboxyl groups are
synthesized by multi-step swelling and polymerization followed by precipitation of iron
oxide inside the pores. The microspheres are characterized by SEM, IR spectroscopy, AAS, and
zeta-potential measurements. Their functional groups enable bioactive ligands of various
sizes and chemical structures to couple covalently. The applicability of these monodisperse
magnetic microspheres in biospecific catalysis and bioaffinity separation is confirmed by
coupling with the enzyme trypsin and huIgG. Trypsin-modified magnetic PGMA-COOH and
PHEMA-COOH microspheres are investi-
gated in terms of their enzyme activity,
operational and storage stability. The
presence of IgG molecules on micro-
spheres is confirmed.
[
1]
1. Introduction
cules (proteins, enzymes, antibodies, and nucleic acids),
[
2]
[3]
biospecific catalysis, immunomagnetic cell separation,
[
4]
Interest in superparamagnetic microspheres has been
rapidly increasing recently due to their potential in
biotechnologyandbiomedicine. Afterbiofunctionalization,
they find practical uses in immunoprecipitations, gentle
but highly efficient isolation and purification of biomole-
diagnosis and prognosis of malignant diseases, etc. Their
main advantages are easy manipulation, high capacity and
simple separation from complex heterogeneous biological
mixtures including blood, urine, foodstuffs, or tissues.
Conventional mechanical methods, such as filtration or
centrifugationormulti-stepcolumnseparationtechniques,
including size-exclusion chromatography, fractional pre-
cipitation, or ion exchange chromatography, can thus be
avoided. Moreover, magnetic microsphere-based bioaffi-
nity separation methods enable the effective isolation of
viable cells and labile molecules due to the lower
D. Hor a´ k
Institute of Macromolecular Chemistry, Academy of Sciences of
the Czech Republic, Prague, Czech Republic
E-mail: horak@imc.cas.cz
J. Ku ˇc erov a´ , L. Koreck a´ , B. Jankovi ˇc ov a´ , Z. B ´ı lkov a´
Department of Biological and Biochemical Sciences, Faculty of
Chemical Technology, University of Pardubice, Pardubice,
Czech Republic
[
5]
mechanical stress than the aforementioned methods.
Several methods are commonly used for the preparation
of magnetic polymer microspheres. Encapsulation of
magnetic cores with a polymer results in polydisperse
J. Palar ˇc ´ı k, P. Mikul a´ sˇ ek
Institute of Environmental and Chemical Engineering, Faculty of
Chemical Technology, University of Pardubice, Pardubice,
Czech Republic
[
6]
particles with irregular shapes,
while miniemulsion
[
7]
polymerization in the presence of iron oxide produces
Macromol. Biosci. 2012, 12, 647–655
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wileyonlinelibrary.com
DOI: 10.1002/mabi.201100393
647

D. Hor a´ k et al.
www.mbs-journal.de
[
8,9]
particles <500 nm, emulsion polymerization
particles
covalently attach suitable bioactive ligands and subse-
quently isolate the analyte of interest from the complex
mixture in the required purity. Due to the increased
stability of immobilized ligands, covalent bonds are
preferred for a wide range of applications. Hence, bioactive
ligands exemplified by the proteolytic enzyme trypsin and
human immunoglobulin G (huIgG) were immobilized on
magnetic PGMA and PHEMA microspheres containing
carboxyl groups (further denoted as PGMA-COOH and
PHEMA-COOH microspheres). Trypsin was selected for
the simple quantification of its proteolytic activity.
Human IgG served to simulate a biologically active high-
molecular-weight biocompound that was immobilized on
the magnetic microspheres to form an immunosorbent.
The amount of IgG immobilized on the microspheres
was quantified by standard bioanalytical methods.
[
10]
<
1 mm, dispersion polymerization
around 1 mm, and
suspension polymerization provides polydisperse particles
[
11]
measuring hundreds of micrometers.
An advantage of
the multi-step swelling and polymerization technique
[
12]
pioneered by Ugelstad et al.
and elaborated by
[
13,14]
others
is that it produces strictly monodisperse
magnetic particles larger than 1 mm, which are difficult
to produce by other techniques. This technique was
therefore for the first time adapted in this report on earlier
elaborated by us suspension polymerization of glycidyl
methacrylate (GMA) and 2-hydroxyethyl methacrylate
(
HEMA). Another novelty consists in utilization of a new
porogen (cyclohexyl acetate) and new stabilizers [2-
hydroxyethylcellulose and (hydroxypropyl)methylcellu-
lose] for preparation of macroporous poly(GMA) or poly-
(
HEMA) (PGMA and PHEMA, respectively) microspheres
and also in formation of magnetic iron oxide in the pores by
oxidation of Fe(OH)
with ammonia.
2
obtained by precipitation of neat FeCl
2
2. Experimental Section
2
.1. Materials
Despite the wide availability of commercial magnetic
carriers, they do not often fulfill all the criteria needed for
unique bioanalytical research. The main requirements laid
on magnetic microspheres include superparamagnetic
behavior, proper size, monodispersity, applicability, and
stability in various media (aqueous,
Monomers such as styrene (Kau cˇ uk Kralupy, Czech Republic),
HEMA (R o¨ hm, Darmstadt, Germany), GMA (Fluka, Buchs, Switzer-
land), and ethylene dimethacrylate (EDMA; Ugilor S. A., France)
were vacuum distilled. 2-[(Methoxycarbonyl)methoxy]ethyl
organic, and mixed) for
a
given
[
15–17]
usage.
The appropriate type of
functional groups on the surface enables
the covalent attachment of various bio-
molecules. The density of functional
groups is then one of the key parameters
of the microspheres. Even though the
number of various applications of mag-
netic microspheres is continually grow-
ing, there are still many areas where the
use of newly developed magnetic micro-
spheres with suitable features could be
beneficial, e.g., micrototal analysis sys-
[
18,19]
tems.
Microspheres need to meet
particularly specific requirements in
such developing fields as microfluidics.
Combining microfluidic devices with
magnetic microspheres is creating new
possibilities in innovative bioapplica-
[
19,20]
tions.
This is the reason why atten-
tion has been paid to the development of
new types of microspheres.
In order to be able to routinely utilize
magnetic microspheres, it is vital to
deeply characterize them, verify their
stability in various media during the
activation process as well as during
regular usage and confirm the ability to
Scheme 1. Synthesis of (a) MCMEMA and (b) HEMA-Ac.
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methacrylate (MCMEMA) was prepared from
ethylene glycol (300 mL) and chloroacetic acid
(
94.5 g) in the presence of NaOH (80 g), produ-
cing sodium hydroxyethoxyacetate, which
was then transformed (in the presence
2 4
of H SO and methanol) to the methyl ester
of hydroxyethoxyacetic acid and finally to
MCMEMA using methacrylic anhydride
(
(
Scheme 1a). 2-(Methacryloyl)oxyethyl acetate
HEMA-Ac; Scheme 1b) was obtained from
HEMA and acetic anhydride. Cyclohexyl acet-
ate was obtained from cyclohexanol and acetic
anhydride. Trypsin from bovine pancreas (EC
Scheme 2. Preparation of monodisperse magnetic macroporous polymer microspheres
by multi-step swelling and polymerization method and precipitation of iron oxide inside
their pores.
3
.4.22.2), bovine serum albumin (BSA), IgG
from human serum, benzamidine, 1-ethyl-3-
3-dimethylaminopropyl)carbodiimide (EDC),
N-a-benzoyl-D,L-arginine-4-nitroanilide (BApNA),
(
and 2-(N-morpholino)ethanesulfonic acid (MES) were obtained
1
amountofDBPandthenfour timeswithdoubletheamountofDBP.
The resulting PS latex contained 4.3 g DBP in 17.5 mL of dispersion.
Third, DBP-swollen PS particles were swollen with the mono-
mers, porogen, and initiator in a 30-mL reaction vessel. Briefly, a
DBP-swollen PS dispersion (2 mL) was swelled with an emulsion of
a solution of BPO (30 mg), GMA (1.5 g), MCMEMA (0.3 g), and EDMA
(1.2 g) in 0.1% SDS solution (7.5 mL) for 16 h with gentle stirring
(30 rpm). A mixture of cyclohexyl acetate (4 g) in 0.1% SDS solution
(10 mL) was then treated with ultrasound at 22 8C for 3 min to form
an emulsion and transferred to the above monomer-swollen PS
dispersion; swelling proceeded for 3 h under stirring (300 rpm).
Alternatively, HEMA-Ac replaced GMA in the swelling solution
when PHEMA-COOH microspheres were being prepared.
from Sigma-Aldrich (St. Louis, MO, USA). The Pierce BCA Protein
Assay Kit was produced by ThermoScientific (Rockford, IL, USA).
The sodium salt of N-hydroxysulfosuccinimide (sulfo-NHS),
FeCl
2
ꢀ 4H
2
O, 2-hydroxyethyl cellulose, sodium dodecylsulfate
(
SDS), benzoyl peroxide (BPO), and Methocel 90 HG [(hydroxypro-
pyl)methyl cellulose] were obtained from Fluka, sodium persulfate
was from Lachema (Brno, Czech Republic). Sodium azide was
produced by Chemapol (London, United Kingdom). The remaining
chemicals were supplied by Sigma-Aldrich, Lachema, or Penta
Chemicals (Chrudim, Czech Republic) and were of analytical
reagent grade. Ultrapure Q-water ultrafiltered with a Milli-Q
Gradient A10 system (Millipore, Molsheim, France) was used for
preparing solutions.
In the fourth step, a 2 wt% solution of 2-hydroxyethyl cellulose
(
2 mL), 2 wt% solution of Methocel 90 HG (2 mL), and a solution of
citric acid (30 mg) in water (0.5 mL) were mixed with the above
monomer-swollen PS latex and the polymerization proceeded at
2
.2. Synthesis of Monodisperse Polystyrene (PS) Seeds
7
0 8C for 16 h at stirring rate of 600 rpm under a carbon dioxide
PS seeds were obtained by the emulsifier-free emulsion polymer-
ization of styrene in a 150-mL reaction vessel equipped with an
anchor-type stirrer. In brief, sodium persulfate (44 mg) and
sodium carbonate (39 mg) were dissolved in water (90 mL) to form
the aqueous phase. The monomer phase (10 g styrene) was added,
the mixture stirred (300 rpm), and the temperature increased
to 80 8C. Polymerization proceeded for 20 h under a nitrogen
atmosphere. The resulting latex was separated by centrifugation
atmosphere. Theresultingmicrosphereswereremovedbyfiltering,
washed with 0.05 wt% Tween 20 solution, ethanol, toluene, and
ethanol (five times each), and finally transferred to water. In order
to change the methyl ester of MCMEMA to a carboxyl group, the
PGMA microspheres were separated by centrifugation and
2 4
hydrolyzed in 0.2 M H SO (50 mL) at 22 8C for 60 h. The micro-
spheres were then washed five times with water with ultrasonic
treatment. In order to yield PHEMA microspheres, the acetate of
poly[2-(methacryloyl)oxyethyl acetate] was hydrolyzed in 0.5 M
NaOH solution (30 mL) in the presence of Tween 20 at 60 8C for 16 h
withstirring(300 rpm);themicrosphereswerepurifiedbywashing
four times with water and twice with 1,4-dioxane.
(
4000 rpm) and thoroughly washed with a 0.25% aqueous solution
of SDS.
2
.3. Synthesis of Monodisperse Macroporous PGMA-
COOH and PHEMA-COOH Microspheres
2
.4. Precipitation of Iron Oxide Inside the
Macroporous PGMA-COOH and PHEMA-COOH microspheres were
synthesized by the modified multi-step swelling and polymeriza-
Macroporous Microspheres
[12]
tion method originally developed by Ugelstad et al.
(Scheme 2).
Macroporous PGMA-COOH or PHEMA-COOH microspheres (1 g)
First, PS latex (0.3 g) was dispersed in 0.25% SDS solution (1 mL) and
the mixture sonicated (4710 Series Ultrasonic homogenizer; Cole-
Parmer, Chicago, IL, USA) at 15 8C for 3 min. Second, the latex was
mixed with an emulsion of dibutyl phthalate (DBP; 0.4 g) in 0.25%
weredispersedin a solutionof FeCl
2 min with ultrasonic treatment, removed by filtering, washed
again with FeCl solution, and left to dry at 40 8C for 30 min. They
were then transferred to a 0.5 M NH OH solution (20 mL), the
2
ꢀ 4H
2
O (2 g) in water(10 mL)for
2
4
SDS(1.2 mL)and2%NaHCO
5 8C for 4 min. PS latex was swollen with DBP for 4 d with mild
stirring (30 rpm); swelling was repeated once more with the same
3
solutions(0.05 mL)undersonicationat
mixture stirred (100 rpm) in air at 22 8C for 3 h, the particles
separated using a magnet, washed several times with water
(100 mL), and mixed with 5 wt% sodium hypochlorite solution
1
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(
2 mL) with stirring (100 rpm) for 10 min. Finally, the microspheres
stored at 4–8 8C in 0.1 M phosphate buffer (pH ¼ 7.3) containing
were washed with water, ethanol, and water. Before trypsin
immobilization, the microspheres were stored in distilled water
3
0.05% NaN solution.
Trypsin activity was determined using low-molecular-weight
chromogenic substrate BApNA according to method modified from
3
with 0.1% NaN .
[
2]
the literature.
incubated in 0.1 M NH
N,N-dimethylformamide (0.02 mL) at room temperature for
0 min under mild stirring. The reaction was stopped by the
Soluble or immobilized trypsin (0.1 mL) was
4
HCO (0.88 mL) and 0.55 M BApNA in
3
2
.5. Characterization of Magnetic Microspheres
3
The microsphere morphology, size, and size distribution were
analyzed by scanning electron microscopy (SEM; JEOL JSM 6400,
Tokyo, Japan). The number-average diameter (Dn), weight-average
diameter (Dw), and uniformity (polydispersity index PDI ¼ Dw=Dn)
were calculated using Atlas software (Tescan Digital Microscopy
Imaging, Brno, Czech Republic) by counting at least 500 individual
particles from SEM microphotographs. The Dn and Dw can be
expressed as follows:
addition of acetic acid (30 wt%, 0.2 mL) and the absorbance was
measured at 405 nm.
2.7. Immobilization of huIgG on Magnetic PGMA-
COOH and PHEMA-COOH Microspheres
Magnetic PGMA-COOH and PHEMA-COOH microspheres (1 mg)
P
P
niDi
were washed five times with 0.1 M MES buffer (pH ¼ 5.0) and
Dn ¼
P
ꢁ1
ni
incubated with EDC (7.5 mg ꢀ mL of 0.1 M MES) for 10 min, the
excess of unreacted EDC was removed from the activated micro-
niD4i
niD3i
spheres by washing with 0.1 M MES buffer (2 ꢂ 1 mL) and huIgG
Dw ¼ P
ꢁ1
(0.2 mg ꢀ mL
of 0.1 M MES) was then added. The immobilization
mixture was incubated at 4–8 8C for 16 h. Subsequent washing was
identical to that described above. The amount of immobilized
huIgG was assessed by absorbance measurement at 260/280 nm
The microspheres were examined with a Paragon 1000 PC FTIR
spectrometer (Perkin-Elmer) with a SpecacMKIIGoldenGate Single
Reflection ATR System with a diamond crystal and a ray angle of
incidence of 458. The iron content was analyzed by atomic
absorption spectrometry (AAS Perkin-Elmer 3110) of an extract
from a sample obtained by treatment with 70% perchloric and 65%
nitricacid at100 8Cfor30 min. A 799GPTTitrinotitrator(Metrohm)
was used to evaluate the carboxyl group content of the micro-
(
(
Eppendorf BioPhotometer; Hamburg, Germany) and by BCA test
1
Pierce BCA Protein Assay Kit).
3
. Results and Discussion
spheres by titrating with 0.1 M NaOH.
The electrostatic stability of the microspheres (46 mg ꢀ mLꢁ1 of
3.1. Synthesis and Characterization of Magnetic
Macroporous Microspheres
0
.01–0.1 M phosphate buffer; pH ¼ 7.3) was investigated by zeta-
potential measurements using a ZetaPALS apparatus (Brookhaven
Instruments; New York, USA).
In this report, carboxyl-terminated microspheres were
preferred due to the availability of a range of standardized
andoptimizedprotocols forbioconjugating variousligands.
Carboxyl groups were introduced in the microspheres by
copolymerizing a relatively small amount (10 wt%) of
MCMEMA (Scheme 1a) in the feed, whichwas subsequently
hydrolyzed. The microspheres were based both on PGMA
and PHEMA. PGMA has the advantage of reactive oxirane
groups which can be potentially modified into any
functional groups required. In addition to PGMA-COOH
microspheres, PHEMA-COOH particles were fabricated as a
standard, because PHEMA is commonly used in a range of
2
.6. Immobilization of Trypsin on Magnetic PGMA-
COOH and PHEMA-COOH Microspheres
Three immobilization approaches were investigated for the
covalent attachment of trypsin to magnetic microspheres: a one-
stepprocedureusingthezero-lengthcrosslinkerEDCandsulfo-NHS
andaone-ortwo-stepprocedurewithneatEDCasacouplingagent.
In a typical one-step immobilization using EDC and sulfo-NHS,
magneticPGMA-COOHorPHEMA-COOH microspheres(1 mg) were
washed four times with 0.1 M phosphate buffer (pH ¼ 7.3),
magnetically separated and mixed with the following agents:
EDC (7.5 mg/0.2 mL), sulfo-NHS (1.25 mg/0.2 mL), and trypsin
[
21]
bioapplications
due to its biocompatibility, inertness,
[
22]
and low-protein adsorption.
Generally, PGMA micro-
(
4 mg/0.5 mL; 0.03 wt% benzamidine). This one-step method can
spheres of narrow size distribution can be synthesized by
a single-step swelling of PS template (obtained by the
dispersion polymerization) with GMA followed by the
also be carried out without the addition of sulfo-NHS. In a two-step
procedure, microspheres were first incubated in buffer with EDC
ꢁ1
(
7.5 mg ꢀ mL ) for 10 min, the supernatant with the excess of EDC
wasremovedandtrypsinsolution(4 mg/0.5 mL)withbenzamidine
0.03 wt%) was added to activated microspheres. For all the above-
[23]
polymerization.
In this report, however, magnetic PGMA-COOH and
PHEMA-COOH microspheres were prepared by a rather
complicated multi-step swelling and polymerization
method (Scheme 2). In contrast to the single-step method,
cyclohexylacetatecouldbeusedasaporogenthatdissolved
PS seeds. Removal of PS seeds during washing after
(
mentioned procedures the reaction proceeded in 0.1 M phosphate
buffer (1 mL of total volume; pH ¼ 7.3) at 23 8C for 3 h with
gentle stirring. The trypsin-immobilized magnetic microspheres
(further denoted as Tryp-PGMA-COOH or Tryp-PHEMA-COOH)
were washed ten times with 0.1 M phosphate buffer (pH ¼ 7.3) and
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Figure 1. SEM micrographs of (a) PS seeds, (b,c) PGMA-COOH, and (d,e) PHEMA-COOH non-magnetic (b and d) and magnetic
c and e) microspheres.
(
completion of the polymerization thus did not induce
formation of dents so typical for single-step swelling and
polymerization method. Moreover, multi-step procedure
produced monomer-swollen particles containing muchless
PS due to much higher swelling of the precursor seed than
single-step swelling and polymerization could provide. The
multi-step swelling and polymerization method thus
produces a highly monodisperse and better quality product
than other techniques. First, seeds 0.7 mm in size were
prepared by emulsifier-free emulsion polymerization
monomer (Scheme 1b), which was hydrolyzed to PHEMA
after the polymerization reached completion. An advan-
tage of the HEMA-Ac monomer, compared to HEMA, is its
insolubility in water, thus facilitating particle swelling and
suspension polymerization in an aqueous phase. Figure 1b
and d represents scanning electron micrographs (SEM) of
both PGMA-COOH and PHEMA-COOH microspheres pre-
pared by the multi-step swelling and polymerization
method. The morphology of the PGMA-COOH and
PHEMA-COOH particles was spherical; the particles were
monodisperse (PDI ¼ 1.04) and had diameters of 3.9 and
(Figure 1a). Second, the seeds were activated (pre-swelled)
with a highly water-insoluble compound (DBP) to enable
subsequent swelling with the monomers, initiator, and
porogen. In the third step, DBP-swollen PS seeds 1.5 mm in
diameter were swelled with the mixture of monomers,
initiator, and porogen. This was followed in the fourth step
by 2-hydroxyethyl cellulose- and (hydroxypropyl)methyl
cellulose-stabilized and BPO-initiated suspension polymer-
ization. In order to introduce carboxyl groups in PGMA and
PHEMAmicrospheres, aMCMEMAcomonomer(Scheme1a)
was incorporated in the polymerization feed. After
its hydrolysis, [2-(methacryloyloxy)ethoxy]acetic acid
1
2
0
8
6
4
1
(MOEAA) was obtained and 0.19 mmol of carboxyl groups
was determined per g of dry PHEMA-COOH microspheres
by titration with NaOH solution (Figure 2). This was less
than the amount of MCMEMA in the feed (0.5 mmol), which
is most likely due to the inaccessibility of some carboxyl
groups buried inside the highly crosslinked polymer bulk.
Nevertheless, the amount of carboxyl groups available for
futuremodifications was sufficient according toour results.
In the synthesis of PHEMA microspheres, HEMA-Ac was a
0
1
2
3
Volume (ml)
Figure 2. Titration of PHEMA-COOH microspheres (0.2 g) with
0.1 M NaOH.
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D. Hor a´ k et al.
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4
.4 mm, respectively. Their surface was rough, containing
the precipitation of iron oxide inside the microsphere pores
(Figure1cande).MagneticPGMA-COOHandPHEMA-COOH
microspheres contained 13.2 and 19.4 wt% Fe, respectively,
according to atomic absorption spectrometry (AAS).
This amount of Fe in the microspheres is sufficient to
respond quickly to a magnetic field as was confirmed
by earlier measurements of magnetic properties of
analogous PGMA microspheres (prepared by the
macropores formed by phase separation during the
polymerization in the presence of the porogen (cyclohexyl
acetate). Due to the presence of the crosslinking agent
(
EDMA), the specific surface area of PGMA-COOH and
2
ꢁ1
PHEMA-COOH particles amounted to 88.3 and 89.4 m ꢀ g
,
respectively, according to the dynamic desorption of
nitrogen. PGMA microspheres contained oxirane groups,
which made additional functionalization possible, result-
[
27]
dispersion polymerization) with a magnetometer.
The
[
24]
ing in a range of various reactive groups.
In this study,
advantage of maghemite over magnetite is its oxidation
stability. Due to its low solubility in water and ability
to form coordination complexes with carboxyl groups,
it was not released from the microspheres in aqueous
media.
oxirane groups were transformed into hydroxyl groups by
hydrolysis, thus increasing the hydrophilicity of the
product, which is important in reducing non-specific
protein adsorption. This was confirmed by attenuation
total Fourier-transform reflectance infrared (ATR-FTIR)
spectra of initial PGMA-COOH microspheres with the
3
.2. Biofunctionalization of Magnetic PGMA-COOH
ꢁ
1
typical transmission peak of oxirane groups at 910 cm
,
and PHEMA-COOH Microspheres
which disappeared after the hydrolysis (Figure 3). More-
over, the transmission peak in the region of OꢁH stretching
Trypsin was immobilized on magnetic PGMA-COOH and
PHEMA-COOH microspheres via EDC/sulfo-NHS chemistry.
The amount of immobilized trypsin was subsequently
measured by colorimetric assay with a chromogenic
substrate. This approach enables the amount of actually
active enzyme molecules to be quantified. Benzamidine is a
low-molecular-weight competitive inhibitor of trypsin,
which prevents the self-cleavage of trypsin during the
immobilization process. Its presence in the immobilization
mixture enhances the binding efficiency of trypsin. After
the binding procedure, benzamidine is removed by simple
washing.
ꢁ
1
vibration (ꢃ3400 cm ) was higher after hydrolysis,
signifying that the number of hydroxyl groups had
increased.
Both PGMA and PHEMA hydrogels are known to absorb
large amounts of water but remain both insoluble and
biologically, chemically, and mechanically stable, preser-
[
25]
ving their shape.
The equilibrium water uptake of
PGMA-COOH and PHEMA-COOH microspheres was 2.7 and
1
2
.8 mL ꢀ gꢁ , respectively, thus reflecting the hydrophilicity
of the particles.
To produce magnetic carriers, maghemite (g-Fe O ) was
2
3
prepared inside the pores of the macroporous microspheres
byatwo-stepoxidation ofFe(OH) with oxygen andsodium
hypochlorite (Scheme 2). First, magnetite (Fe ) was
For PGMA-COOH microspheres, the highest activity of
immobilized trypsin was achieved with a one-step
immobilization using EDC/sulfo-NHS at 23 8C for 3 h
(Figure 4). The same trend was observed for magnetic
PHEMA-COOH microspheres.
2
3 4
O
formed by the precipitation of a ferrous salt with
ammonium hydroxide and oxidation with oxygen. Second,
magnetite was then oxidized with sodium hypochlorite,
Figure 5 shows the effect of increasing amounts of
trypsin in the binding mixture on the amount of enzyme
[
26]
yielding chemically more stable g-Fe O (maghemite).
2
3
Neither the morphology nor size of the PGMA-COOH or
PHEMA-COOH microspheres substantially changed after
1000
100
9
8
7
6
5
0
0
0
0
5
00
-
1
9
10 cm
0
(
a)
6 h, 4° C
(b)
(c)
3 h, 23° C
1
0
4
000
3500
3000
2500
2000
1500
1000
500
Figure 4. Proteolytic activity of magnetic Tryp-PGMA-COOH
microspheres under various reaction conditions (time, tempera-
ture, and presence of EDC and sulfo-NHS). (a) One-, (b) two-step
protocol with EDC, and (c) one-step immobilization with EDC and
sulfo-NHS.
-1
Wavenumbers (cm )
Figure 3. ATR-FTIR spectra of PGMA-COOH microspheres (—)
before and ( ꢀ ꢀ ꢀ ) after hydrolysis with H SO
2
4
.
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1
500
000
tional and storage stability were therefore verified. The
activity of magnetic Tryp-PGMA-COOH and magnetic
Tryp-PHEMA-COOH microspheres was repeatedly deter-
mined in nine cycles within 1 d (0.1 mg aliquot, using the
sameoneeachtime).TheactivityofmagneticTryp-PHEMA-
COOH microspheres decreased only moderately to 60% in
the fifth cycle. The activity of magnetic Tryp-PGMA-COOH
microspheres decreased more considerably, reaching 39.6%
of theoriginal value in the fifth cycle. These results were not
what we expected; previous results showed that >50% of
the original activity was retained even after the 10th
1
5
00
0
2
3
4
5
Added trypsin (mg)
Tryp-PGMA-COOH
Tryp-PHEMA-COOH
[
2]
measurement. One of several reasons for the activity
decreasecouldbetheabsenceofBSAduringimmobilization
or the autoproteolytic behavior of trypsin. This issue still
needs to be studied in more detail. Storage stability tests,
i.e. determination of enzyme activity over one-week
periods (0.1 mg aliquot, a fresh one each time) gave
excellent results and confirmed the suitability of the
prepared biofunctionalized carriers for long-term storage
as well as their applicability. More than 70% of the trypsin
molecules immobilized on Tryp-microspheres were active
after seven weeks of storage, which fulfilled our expecta-
tions (Figure 6).
Figure 5. Determination of proper amount of trypsin in binding
mixture per mg of magnetic microspheres.
which was immobilized onto the microspheres and
simultaneously active. In both types of magnetic carriers,
a slight increase in activity was observed as a consequence
of increasing the trypsin content in the binding mixture.
The optimalamount was foundto be4 mgof trypsinpermg
of both magnetic microspheres. Immobilizing higher
amounts of enzyme (>5 mg) was not beneficial; the
immobilization efficiency slightly decreased (Figure 5).
The average enzyme activity of magnetic Tryp-PGMA-
COOH and Tryp-PHEMA-COOH microspheres determined
Accordingtotheliterature, zeta-potentialmeasurements
of free and enzyme-modified carriers correlated with
[
29]
from three repetitions was 1021 ꢄ 44 and 1418 ꢄ
enzyme binding efficiency.
Thus, we measured the
1
3
2 U ꢀ mgꢁ of microspheres, respectively.
zeta-potential of trypsin-free and trypsin-modified mag-
netic microspheres in various buffers at various molar
concentrations (0.01, 0.05, and 0.1 M). The absolute values of
zeta-potential of the microspheres in phosphate buffer
decreased after their biofunctionalization with trypsin
(Figure 7). A similar tendency was also observed for
measurements of the zeta-potential of magnetic Tryp-
PGMA-COOH and Tryp-PHEMA-COOH microspheres in
To verify that the enzyme molecules were specifically
adsorbed, they were washed with 1 M NaCl solution.
Enzyme activity then slightly decreased to 99.8 and
9
9.3% for magnetic Tryp-PGMA-COOH and Tryp-PHEMA-
COOH microspheres, respectively. The above-mentioned
reduction of trypsin activity is statistically insignificant;
hence, theresultsindicatethatmorethan99%ofthetrypsin
was coupled to the microspheres through covalent bonds.
Based on good experience with the simultaneous
co-immobilization of proteins and other ligands to micro-
spheres, and keeping in mind the aim of covering these
hydrophobic clusters, which are responsible for the non-
specific adsorption of biomolecules from complex biologi-
cal mixtures, the addition of inert protein – BSA – could
increase the quality of enzyme-biofunctionalized micro-
NH
4
HCO
3
buffer. The lower molar concentrations of buffer
120%
Tryp-PGMA-COOH
Tryp-PHEMA-COOH
8
0%
0%
[
28]
spheres.
BSA solution (33 wt%) was added to the
immobilization mixture 10 min after the start of the
immobilization. This was followed by a 16-h incubation
and subsequent washing with 0.1 M phosphate buffer
4
(
pH ¼ 7.3) containing 1 M NaCl for the elimination of
eventual non-specifically bound molecules. Magnetic
Tryp-PGMA-COOH microspheres achieved a trypsin acti-
vity comparable to the procedure without albumin
addition (difference <5%).
0%
0
1
2
3
4
5
6
7
Time (weeks)
Figure 6. Storage stability of magnetic Tryp-PGMA-COOH and
magnetic Tryp-PHEMA-COOH microspheres at 4 8C in 0.1 M phos-
phate buffer (pH ¼ 7.3) containing 0.03 wt% benzamidine (0.1 mg
aliquots).
Thechemicalaswellasbiologicalstabilityofthesenewly
developed biofunctionalized magnetic microspheres needs
to be confirmed before their routine application. Opera-
Macromol. Biosci. 2012, 12, 647–655
ß 2012 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
653
www.MaterialsViews.com

D. Hor a´ k et al.
www.mbs-journal.de
4
. Conclusion
-
-
-
-
15
25
35
45
Magnetic monodisperse PGMA-COOH and PHEMA-COOH
microsphereswere obtainedbythemulti-step swellingand
polymerization method followed by the precipitation of
iron oxide inside the pores of the particles. To verify the
applicability of the developed magnetic PGMA-COOH and
PHEMA-COOH microspheres for immobilizing biocom-
pounds, the proteolytic enzyme trypsin and human IgG
were used as model ligands. Trypsin was immobilized on
the microsphere surface by the well-known carbodiimide-
mediated one-step protocol with heterobifunctional cross-
linker EDC and sulfo-NHS agent, and strong bond creation
was confirmed. These newly developed biofunctionalized
magnetic microspheres demonstrated the possibility of
long-term storage without significant changes, which thus
indicates great potential for their successful use in
bioapplications. Also, IgG-bionfunctionalized microspheres
have been shown to be convenient carriers for, e.g., widely
used immunoprecipitations.
0
0.05
Concentration of PBS (mol.l )
0.1
-1
Neat PGMA-COOH
Neat PHEMA-COOH
Tryp-PGMA-COOH
Tryp-PHEMA-COOH
Figure 7. Zeta-potential of magnetic PGMA-COOH and magnetic
PHEMA-COOH microspheres before and after biofunctionaliza-
tion with trypsin in phosphate buffer (pH ¼ 7.3); activity
ꢁ1
ꢁ1
9
70 U ꢀ mg of Tryp-PGMA-COOH and 1417 U ꢀ mg of Tryp-
PHEMA-COOH.
Acknowledgements: Financial support of the Grant Agency of the
Czech Republic (203/09/0857 and P503/10/0664), the Ministry of
Education, Youth and Sports (no. MSMT 0021627502), and EU
were used, the higher absolute values of zeta-potential of
carriers were measured indicating better colloidal
stability and lower rate of aggregation due to the repulsion
among individual particles. Based on these results, the
buffers with a lower molar concentration are preferred for
this type of microspheres.
(
CaMiNEMS project no. 228980 and NaDiNe project no. 246513) is
gratefully acknowledged.
Received: October 6, 2011; Revised: November 29, 2011; Published
online: March 13, 2012; DOI: 10.1002/mabi.201100393
Regarding the potential use of microspheres as carriers of
antibodies in the field of immunoprecipitation, human IgG
was immobilized on both types of new magnetic micro-
Keywords: enzyme catalysis; glycidyl methacrylate; 2-hydro-
xyethyl methacrylate; magnetic microspheres; trypsin
[
6]
spheres. Based on earlier investigations, an optimized
two-step carbodiimide method was chosen for covalently
binding huIgG to the microspheres. Two different
approaches were used for determining the amount of
immobilized huIgG on the microspheres. The total amount
of immobilized huIgG was calculated as the difference
between the amount of IgG in the binding solution
before immobilization and the sum of IgG from all
solutions after immobilization, measured by UV spectro-
photometry at 260/280 nm. Additionally, the BCA test
with IgG-immobilized microspheres was done according
to the manufactures’ instructions. For both methods,
the amount of immobilized IgG was calculated for
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