Comprehensive study of bioanalytical platforms: Xanthine oxidase

Article abstract of DOI:10.1021/ac051676l

A comprehensive study of a general bioanalytical platform for biosensor applications is presented using xanthine oxidase (XnOx) as a case study within the framework of developing approaches of broad applicability. In this context, emphasis is placed on amperometric biosensors based on XnOx, which has been immobilized by covalent binding to gold electrodes modified with dithiobis-N-succinimidyl propionate. The immobilized XnOx layers have been characterized using atomic force microscopy under liquid conditions and quartz crystal microbalance techniques. In addition, spatially resolved mapping of enzymatic activity has been carried out using scanning electrochemical microscopy. Redox dyes of phenothiazine derivatives, specifically, thionine and methylene blue, have been found to work well as electron acceptors for reduced XnOx. The kinetic parameters and equilibrium constants of the mediated enzymatic oxidation of xanthine in the presence of the above-mentioned redox dyes have been calculated. The response of the enzymatic electrode to varying xanthine concentrations has been obtained in the presence of thionine or methylene blue as redox mediator in solution. Under these conditions, xanthine could be determined amperometrically at +0.2 V versus SSCE.

Full text of DOI:10.1021/ac051676l

Anal. Chem. 2006, 78, 530-537
Comprehensive Study of Bioanalytical Platforms:
Xanthine Oxidase
E. Casero,† A. Martinez G. de Quesada,† J. Jin,‡ M. C. Quintana,† F. Pariente,† H. D. Abrun˜a,‡
L. Va´ zquez,§ and E. Lorenzo*,†
Departamento de Qu´ımica Anal´ıtica y Ana´lisis Instrumental, Universidad Auto´noma de Madrid, Campus de Cantoblanco,
28049 Madrid, Spain, Instituto de Ciencia de Materiales de Madrid (CSIC), C/Sor Juana Ine´s de la Cruz, no. 3. 28049
Madrid, Spain, and Department of Chemistry and Chemical Biology, Baker Laboratory, Cornell University,
Ithaca, New York 14853-1301
immobilization process, using techniques that allow quantification
as well as visualization of the immobilized enzymes, would
represent a great advantage.
A comprehensive study of a general bioanalytical platform
for biosensor applications is presented using xanthine
oxidase (XnOx) as a case study within the framework of
developing approaches of broad applicability. In this
context, emphasis is placed on amperometric biosensors
based on XnOx, which has been immobilized by covalent
binding to gold electrodes modified with dithiobis-N-
succinimidyl propionate. The immobilized XnOx layers
have been characterized using atomic force microscopy
under liquid conditions and quartz crystal microbalance
techniques. In addition, spatially resolved mapping of
enzymatic activity has been carried out using scanning
electrochemical microscopy. Redox dyes of phenothiazine
derivatives, specifically, thionine and methylene blue,
have been found to work well as electron acceptors for
reduced XnOx. The kinetic parameters and equilibrium
constants of the mediated enzymatic oxidation of xanthine
in the presence of the above-mentioned redox dyes have
been calculated. The response of the enzymatic electrode
to varying xanthine concentrations has been obtained in
the presence of thionine or methylene blue as redox
mediator in solution. Under these conditions, xanthine
could be determined amperometrically at +0.2 V versus
SSCE.
Among the techniques that have been employed to determine
the mass deposited on a surface, the quartz crystal microbalance
(QCM) has been shown to be most useful. In addition, this
technique can be used not only for quantification but also to obtain
kinetic information. Atomic force microscopy (AFM) has achieved
particular relevance as a surface characterization technique, which
provides morphological and mechanical information at the na-
nometer level. Since the development of tapping operation mode
under aqueous environment,2 AFM has been able to provide
morphological data of protein deposits without damaging the
sample surface.3 In addition, AFM provides very valuable informa-
tion on protein-protein and protein-surface interactions, through
force spectroscopy measurements4 and on surface-induced con-
formational changes of proteins.5
The assessment of the enzymatic activity is also very important
for the optimization of enzyme immobilization and functional
parameters of sensors.6 Imaging of local activity of redox enzymes
often requires the availability of electrochemically detectable
compounds indicative of the biocatalytic reaction. In addition, the
use of redox mediators with fast interfacial kinetics can enhance
the sensitivity and reproducibility of the determinations.7 In this
sense, scanning electrochemical microscopy (SECM) has provided
a very sensitive and versatile tool for determining the local activity
of immobilized enzymes.8 In the case of oxidases, mapping of
enzymatic activity can be achieved either by detecting the
enzymatic reaction product (hydrogen peroxide) or by following
changes in the steady-state current of a mediator as a consequence
of the enzymatic reaction. Among the main advantages of the
SECM detection, one can highlight its suitability to detect
molecules in very small volumes9 and that it does not require an
For the past decade, highly specific analytical devices have
been developed by coupling immobilized redox enzymes with
electrochemical sensors.1 In comparison with other analytical
techniques, such sensors possess simplicity of operation, high
sensitivity, and selectivity. In addition, the immobilization of an
enzyme has the advantage that a small amount of enzyme can be
used for a large number of analytical determinations. Therefore,
there is a great deal of continued interest in the development of
enzyme immobilization methods that give rise to robust and
successful biosensors. A crucial step in these studies is to know
both the amount of immobilized enzyme and its conformation/
distribution on the surface. Thus, the characterization of the
(2) Putman, C. A.; Van der Werf, K. O.; De Grooth, B. G.; Van Hulst, N. F.;
Greve, J. Appl. Phys. Lett. 1994, 64, 2454-2456.
(3) Forbes, J. G.; Jin, A. J.; Wang, K. Langmuir 2001, 17, 3067-3075.
(4) Feldman, K.; Ha¨hner, G.; Spencer, N. D.; Harder, P.; Grunze, M. J. Am.
Chem. Soc. 1999, 121, 10134-10141.
* To whom correspondence should be addressed. E-mail: encarnacion.lorenzo@
uam.es.
(5) Moulin, A. M.; O’Shea, S. J.; Badley, R. A.; Doyle, P.; Welland, M. E.
Langmuir 1999, 15, 8776-8779.
† Universidad Auto´noma de Madrid.
(6) Csoka, B.; Kova´cs, B.; Nagy, G. Electroanalysis 2003, 15, 1335-1342.
(7) Pierce, D. T.; Unwin, P. R.; Bard, A. J. Anal. Chem. 1992, 64, 1795-1804.
(8) Wittstock, G. Fresenius J. Anal. Chem. 2001, 370, 303-315.
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‡ Cornell University.
§ Instituto de Ciencia de Materiales de Madrid.
(1) Wang, J. Anal. Chem. 1995, 67, 487R-492R.
530 Analytical Chemistry, Vol. 78, No. 2, January 15, 2006
10.1021/ac051676l CCC: $33.50 © 2006 American Chemical Society
Published on Web 11/30/2005

incubation time period since the products of the enzymatic
reaction are detected directly in proximity to the target.10 In
addition, the biorecognition reactions can be localized in very small
areas by micropattering techniques,11,12 which not only decrease
reagent consumption but also provide the basis for multianalyte
assays.
The characterization studies described above are preliminary
and essential steps in the design of biosensors based on (redox)
enzymes. In this sense, the goal of this work was to carry out a
comprehensive characterization of a general bioanalytical platform
for biosensor applications using xanthine oxidase (xanthine
oxygen oxidoreductase; XnOx) as a case of study with the purpose
of developing approaches of broad applicability.
phenazine and phenothiazine derivatives, quinones/hydroquino-
nes and related materials are often used. Recently, some redox-
mediated XnOx-based sensors have been developed.23-25 In these
previous studies, ferrocene and its derivatives as well as prussian
blue have been employed as redox mediators either in solution23,24
or incorporated in the carbon paste electrodes.25 Detection limits
in the micromolar regime were reported for both theophylline
and hypoxanthine employing well-known immobilization schemes,
such as enzyme retention behind a dialysis membrane, enzyme
cross-linked with glutaraldhehyde on a porous Nylon membrane,
or incorporation into electropolymerized pyrrole films. However,
the drawbacks of the use of such enzyme immobilization schemes
are well known and documented.
In the present work, we have focused our attention on XnOx-
based biosensor applications using artificial redox mediators.
Specifically, phenothiazine dyes thionin and methylene blue as
well as hydroxymethylferrocene have been employed as electron-
transfer mediators in solution.
XnOx is a complex metalloprotein involved in purine catabo-
lism, oxidizing hypoxanthine to xanthine and xanthine to uric acid,
with the concomitant reduction of oxygen.13-15 In addition, the
enzyme catalyzes the hydroxylation of a large number of nitrogen-
containing heterocyclic compounds and the oxidation of many
aldehydes.
XnOx-modified electrodes have been frequently used for the
determination of biological purines, in particular, hypoxanthine,16-18
a major metabolite in the degradation of adenine nucleotide, which
is found to accumulate in fish and beef. As a result, the level of
hypoxanthine is generally used in the food industry as an index
of freshness. Most of these reported sensors are generally based
on the determination of either consumed oxygen or enzymatic
reaction products, i.e., peroxide19,20 or uric acid.21
Electrical communication/connection between redox proteins
and the electrode surfaces provides a general means of enhancing
the activity of the redox-active biocatalyst.22 Direct contact between
the protein’s redox center and the electrode interface is, however,
generally ineffective due to the insulation of the active site by the
protein matrix. Various methodologies have been employed to
enhance the interaction between redox proteins and electrodes.
One of the more common approaches involves the use of redox
mediators in solution. In addition, mediated oxidase-based bio-
sensors frequently operate at much lower potentials than those
based on the determination of an enzymatic reaction product, i.e.,
peroxide. In solution, electron-transfer mediators operate by a
diffusional route facilitating electrical contact between the en-
zyme’s redox center and the electrode surface. In the case of
oxidoreductases, it has been shown that both, one- and two-
electron, mediators can be employed. Typical of the first would
be ferrocenyl derivatives and hexacyanoferrates. In the latter case,
As we have previously mentioned, the aim of this work is to
describe bioanalytical sensor platforms based on oxidoreductase
enzymes. In this sense, the work involves (1) the study of different
artificial redox mediators that can be used as electron acceptors
for XnOx and evaluation of their equilibrium constants and kinetics
parameters, (2) the immobilization of XnOx by covalent binding
to gold electrodes modified with dithiobis-N-succinimidyl propi-
onate (DTSP) as a general reagent for a covalent enzyme
immobilization, (3) the characterization of the resulting enzyme
layer by QCM and AFM, (4) the assessment of the enzymatic
activity of immobilized XnOx by SECM, and (5) the study of the
enzyme electrode response to xanthine using thionin and meth-
ylene blue as solution redox mediators.
EXPERIMENTAL SECTION
Materials. XnOx (EC1.1.3.22) from buttermilk was com-
mercially available from Sigma Chemical Co. (St. Louis, MO) and
used without further purification. Stock solutions were prepared
by dilution and stored at 4 °C. Under these conditions, the
enzymatic activity remains stable for several weeks. Xanthine,
thionin, methylene blue, hydroxymethylferrocene, 3,3′-dithiodipro-
pionic acid di(N-succinimidyl ester) (DTSP), and dimethyl sul-
foxide were obtained from Aldrich Chemical Co. (Milwaukee, WI)
and were used as received. All other chemicals were of at least
reagent grade quality and were used as received. Sodium
phosphate (Merck) was employed for the preparation of buffer
solutions (0.1 M, pH 6.5). Water was purified with a Millipore
Milli-Q-System. All solutions were prepared just prior to use.
Apparatus and Procedures. Spectrophotometric Mea-
surements. Changes in the absorbance were measured for 0.1
mM solutions of the oxidized form of thionin or methylene blue
after addition of xanthine to a 0.1 M pH 6.5 phosphate buffer
solution containing 0.2 unit of XnOx. Absorbance measurements
were carried out at 25 °C using a Shimadzu UV-1700 spectropho-
tometer and a quartz cell having a light path length of 1 cm. Before
adding the xanthine solution, the phosphate buffer solution was
deoxygenated by bubbling nitrogen for 20 min.
(10) Shiku, H.; Matsue, T.; Uchida, I. Anal. Chem. 1996, 68, 1276-1278.
(11) Turyan, I.; Matsue, T.; Mandler, Anal. Chem. 2000, 72, 3431-3435.
(12) Wilhelm, T.; Wittstock, G. Langmuir 2002, 18, 9485-9493.
(13) Harris, C. M.; Massey, V. J. Biol. Chem. 1997, 272, 28335-28341.
(14) Krenitsky, T. A.; Neil, S. M.; Elion, G. B.; Hitchings, G. H. Arch. Biochem.
Biophys. 1972, 150, 585-599.
(15) Rosemeyer, H.; Seela, F. Eur. J. Biochem. 1983, 134, 513-515.
(16) Lanqun, M.; Fang, X.; Qi, X.; Litong, J. Anal. Biochem. 2001, 292, 94-101.
(17) Cayuela, G.; Pen˜a, N.; Reviejo, A. J.; Pingarro´n, J. M. Analyst 1998, 123,
371-377.
(18) McKenna, K.; Brajter-Toth, A. Anal. Chem. 1987, 59, 954-958.
(19) Hu, S.; Xu, C.; Luo, J.; Cui, D. Anal. Chim. Acta 2000, 412, 55-61.
(20) Rehak, M.; Snejdarkova, M.; Otto, M. Biosens. Bioelectron. 1994, 9, 337-
341.
(21) Gonzalez, E.; Pariente, F.; Lorenzo, E.; Herna´ndez, L. Anal. Chim. Acta
1991, 242, 267-273.
(22) Barlett, P. N.; Tebbut, P.; Whitaker, R. G. Prog. React. Kinet. 1991, 16,
55-60.
(23) Liu, Y.; Nie, L.; Tao, W.; Yao, S. Electroanalysis 2004, 16, 1271-1277.
(24) Luong, J. H. T.; Thatipamala, R. Anal. Chim. Acta 1996, 319, 325-333.
(25) Stredansky, M.; Pizzariello, A.; Miertus, S.; Svorc, J. Anal. Biochem. 2000,
285, 225-229.
Analytical Chemistry, Vol. 78, No. 2, January 15, 2006 531

Quartz Crystal Microbalance Measurements. AT-cut quartz
crystals (5.0 MHz) of 25-mm diameter with Au electrodes
deposited over a Ti adhesion layer (Maxtek Inc., Santa Fe Springs,
CA) were used for QCM measurements. An asymmetric keyhole
electrode arrangement was used, in which the circular electrode
geometrical areas were 1.370 (front side) and 0.317 cm2 (back-
side). The electrode surfaces were overtone polished. Prior to use,
the quartz crystals were cleaned by immersion in piranha solution,
H2SO4/H2O2 (3:1 v/v). Caution: Piranha solution is extremely
reactive. They were subsequently rinsed with water and dried in
air. The quartz crystal resonator was set in a probe (TPS-550,
Maxtek) made of Teflon in which the oscillator circuit was
included, and the quartz crystal was held vertically. The probe
was connected to a cell by a homemade Teflon joint, which was
immersed in water-jacketed beaker thermostated at the assay
temperature (25.0 ( 0.1 °C) with a thermostatic bath (Digital
Temperature Controller Haake F6). The frequency was measured
with a plating monitor (PM-740, Maxtek Inc.) and simultaneously
recorded by a personal computer.
Atomic Force Microscopy Measurements. Supports used
for protein attachment consisted of glass substrates (1.1 × 1.1
cm) covered with a chromium adhesion layer (1-4 nm thick) on
to which a gold layer (200-300 nm thick) was deposited
(Arrandee, Germany). Prior to use, the gold surfaces were
annealed for 2 min in a gas flame in order to obtain Au (111) flat
terraces. The atomic force microscope was a Nanoscope IIIa
(Veeco, Santa Barbara, CA) used with a D scanner (maximal scan
range of ∼14 µm). Measurements were carried out, under liquid
environment, with Si3N4 cantilevers (nominal radius of 20 nm and
spring constant of 0.38 N/m) from Veeco. The samples were
imaged with different cantilevers in order to ensure that the
imaged structures were not due to tip artifacts. The substrate was
first imaged in buffer solution to ensure that the surface was flat
and clean before carrying out the protein immobilization step. Both
sample and cantilever were located within a Plexiglas fluid cell
into which ∼50 µL of buffer was added.
saturated calomel electrode (SSCE) without taking into account
the liquid junction. All solutions were deaerated with nitrogen gas
before use, and the gas flow was kept over the solution during
experiments. Cronoamperometric measurements were carried out
by poising the enzyme electrode at a potential of +0.2 V, where
oxidation of the mediator is assured. After a steady-state back-
ground current baseline was obtained, aliquots (typically 100 µL)
of a xanthine stock solution (typically 5 mM) were added. After
the mixture was stirred for 30 s and allowed 2 min for equilibra-
tion, the steady-state current in the unstirred solution was
recorded.
SECM Enzyme Assay Procedure. SECM measurements
were carried out using CH Instrument model 900B equipment.
The setup is formed by an electrochemical cell located on an XYZ
positioning stage. The distance between the tip and the sample
was controlled using a stepper motor (coarse approach) combined
with a piezoelectric micropositioning system (fine approach). The
three- or four-electrode cell was formed by a Ag/AgCl reference
electrode, a platinum wire auxiliary electrode, a substrate (highly
ordered pyrolytic graphite), and a Pt ultramicroelectrode with a
10-µm radius, usually termed the tip or probe electrode. Substrates
were previously modified with the enzyme and subsequently
mounted in the cell. Measurements were performed in the
feedback mode (probe potential E ) + 0.4 V, sample at open
circuit, lateral scan speed vt ) 15 µm s-1) in pH 6.5, 0.1 M
phosphate buffer + 0.1 M KCl containing 1 mM hydroxymethyl-
ferrocene and 3 mM xanthine. Previously, the assay solution was
thoroughly deoxygenated in the SECM cell.
RESULTS AND DISCUSSION
Spectrophotometric Study of Redox Mediators. Evaluation
of Equilibrium Constants and Kinetic Parameters. Based on
previous studies on artificial electron acceptors for oxidase
enzymes, which, like XnOx have an FAD active redox center,28
we have examined the ability of several phenothiazine dyes, with
emphasis on thionin and methylene blue, to accept electrons from
the reduced XnOx. To carry out these studies, the absorption
spectra of 0.1 mM solutions of the dye in pH 6.5, 0.1 M phosphate
buffer containing 0.2 unit of XnOx were recorded (data not
shown). Upon addition of xanthine (0.1 mM), a dramatic decrease
in absorbance with reaction time was observed. In the case of
thionin, the absorption peak at 600 nm decreased rapidly, and
after 30 s, the spectrum was that of the known reduced form of
thionin. However, a qualitatively similar behavior was observed
for methylene blue except the final spectrum was reached after
∼60 s. These observations were consistent with the well-known
ping-pong mechanism:
For morphological imaging of the different protein deposits
and substrates, we employed tapping mode AFM microscopy. In
this mode, the cantilever is oscillated at a frequency close to its
resonant frequency, f0, in the ∼8.5-10.5 kHz range with an
amplitude A0 (∼15 nm) above the surface. As the cantilever-
sample distance decreases, the probe interaction with the sample
surface increases, leading to a decrease of the cantilever ampli-
tude, A (A < A0). Thus, the lower the A/A0 ratio, the higher the
average applied force on the sample.26 Accordingly, one can
measure, at a fixed spot, the change of A with the relative tip-
sample distance. These curves, when compared with those
obtained with the same tip on a hard substrate, can provide an
estimation of the indentation induced by the tip on the imaged
material.27
Enzyme Electrode Response. The enzyme electrode was
immersed in a stirred pH 6.5, 0.1 M phosphate buffer solution in
a three-compartment electrochemical cell with standard taper
joints so that all compartments could be hermetically sealed with
Teflon adapters. A large-area coiled platinum wire was employed
as a counter electrode. All potentials are reported against a sodium-
XnOx (ox) + S T XnOx-S T XnOx (red) + P (1)
XnOx (red) + Medox T XnOx-Med T
XnOx (ox) + Medred (2)
From such experiments we have obtained the time course of
changes in the concentration of the oxidized form of the dye after
addition of increasing concentrations of xanthine ranging from
(26) Garc´ıa, R.; Pe´rez, R. Surf. Sci. Rep. 2002, 47, 197-301.
(27) Kopp-Marsaudon, S.; Lecle`re, Ph.; Dubourg, F.; Lazzaroni, R.; Aime´, J. P.
Langmuir 2000, 16, 8432-8437.
(28) Nakaminami, T.; Kuwabata, S.; Yoneyama, H. Anal. Chem. 1997, 69, 2367-
2372.
532 Analytical Chemistry, Vol. 78, No. 2, January 15, 2006

Figure 1. (A) Time course of changes in the concentration of electron acceptors (thionin and methylene blue) in oxidized forms after addition
of 0.4 µM xanthine in pH 6.5, 0.1 M phosphate buffer solution containing 0.2 unit of XnOx and 0.6 µM thionin or 0.5 µM methylene blue in initial
concentration. (B) Lineweaver-Burk plots obtained from data presented in (A).
XnOx, are smaller for methylene blue than for thionin. Since
thionin and methylene blue have very similar E0′ values (-0.21
and -0.22 V in 0.02 M pH 7.0 phosphate buffer solution,
respectively) one would not, a priori, anticipate any differences
in the ability of the mediators to accept electrons from XnOx.
Thus, these results suggest that other effects, such as size or
hydrophobicity of the redox mediators, may also play an important
role in determining their reactivity. Although largely speculative
in our part, the differences might arise from the enhanced ability
of thionin to engage in hydrogen bonding.
Table 1. Equilibrium Constants and Kinetic Parameters
of XnOx
νmax (µmol min-1
)
KM (µM)
Keq
τ (s)
thionin
methylene Blue
125
55
0.325
0.077
6800
454
18
25
0.1 to 1.2 µM and a concentration of thionin and methylene blue
of 0.6 and 0.5 µM, respectively. Figure 1A shows representative
plots for both mediators at a xanthine concentration of 0.4 µM.
As can be seen, the dye concentrations decrease with time. Given
that the product of the enzymatic reaction is uric acid, the
following equilibrium applies:
A similar effect has been previously reported by Nakaminami
et al.28 in the case of cholesterol oxidase and several phenazines
and phenothiazine derivatives, including thionin and methylene
blue.
XnOx
Development of Xanthine Bioanalytical Platforms. One of
the objectives of this investigation was the development of
bioanalytical platforms based on XnOx. For this purpose, in a first
step, we attempted the immobilization of XnOx on modified gold
electrode surfaces. As we have previously described,32 gold
electrodes can be modified with a monolayer of DTSP through
the disulfide group, so that terminal succinimidyl groups allow
further covalent immobilization of enzymes or other materials
including XnOx. To characterize the XnOx immobilization process
on DTSP-modified gold substrates, we have employed several
techniques as described below.
To determine the amount of XnOx immobilized on the DTSP-
modified gold electrode, QCM studies were carried out. QCM
measurements allow simultaneous acquisition of kinetic informa-
tion about the binding process, as well the surface coverage of
the resulting XnOx layer. The immobilization process was carried
out, as described above, in a homemade cell coupled to the QCM
probe containing the DTSP-modified Au quartz resonator in
contact with 10 mL of buffer solution. After the temperature and
the frequency were stabilized, an aliquot of XnOx stock solution
was added to the cell in order to obtain a final concentration of
protein of 1.8 µM. As can be seen in Figure 2A, upon addition of
xanthine + Med
7
8 uric acid + Medred
(3)
ox
Keq
where Keq is the equilibrium constant. The values of Keq were
calculated for both thionin and methylene blue by determining,
from the absorbance measurements, the concentrations of Medox
and Medred once equilibrium has been reached. The values
obtained are presented in Table 1. This table also includes kinetic
parameters of the maximum rate of xanthine oxidation (Vmax) and
the Michaelis constant for xanthine (KM) evaluated from the
corresponding Lineweaver-Burk plots (see Figure 1B) of v-1
versus Cs-1, where Cs is the concentration of xanthine.29,30 The
time employed to achieve equilibrium (τ) is also given in Table
1. This parameter is defined as the reaction time required to
consume (1/n)∆CMedox, where n is a natural number and ∆CMedox
is the difference between the initial concentration of the oxidized
form of the mediator (Medox) and the corresponding value at
equilibrium.31
As can be observed in Table 1, both Vmax and Keq, which reflect
the ability of the two redox mediators to accept electrons from
(29) Segel, I. H. Enzyme Kinetics; Wiley-Interscience: New York, 1993; Chapter
9-I.
(30) Dixon, M.; Webb, E. C. Enzymes; Longman: London, 1979; Chapter 4.
(31) Atkins, P. W. Physical Chemistry; Oxford University Press: New York, 1990;
Chapter 26.
(32) Darder, M.; Takada, K.; Pariente, F.; Lorenzo, E.; Abrun˜a, H. D. Anal. Chem.
2000, 72, 3784-3792.
Analytical Chemistry, Vol. 78, No. 2, January 15, 2006 533

Figure 3. Scheme of XnOx. The most exposed lysine groups are
marked in pink.
Figure 2. Time dependence of the frequency changes of a DTSP-
gold substrate resonator (A) and a bare gold substrate (B) in pH 6.5,
0.1 M phosphate buffer solution upon addition of XnOx to a final
concentration of 1.8 µM.
Table 2. First-Order Rate Constant (k), Frequency
Change (∆Fmax), Mass of Xanthine Oxidase (m), and
Surface Coverage (Γ) for the Immobilization Process of
Xanthine Oxidase on a Gold Surface and on a
DTSP-Gold Substrate
XnOx, the frequency decreased slowly during 40 min until a steady
state was reached.
Assuming that the frequency decrease (∆F) is only due to the
change in mass arising from the adsorption of the enzyme, one
can calculate the amount and surface coverage, Γ, of the im-
mobilized XnOx by using the Sauerbrey equation33
k
∆Fmax
m
Γ × 1012
(min-1
)
(Hz)
(ng cm-2
)
(mol cm-2
)
XnOx-DTSP-Au
XnOx-Au
0.1
1.0
-23.90
-39.01
423
690
1.4
2.3
can be seen in Figure 2B, upon addition of XnOx, a rapid decrease
in frequency was observed during the first 2 min and afterward a
steady state was reached. These observations indicate that the
enzyme adsorbs to bare gold surfaces. The shapes of the
frequency time plots for bare and DTSP-modified surfaces are
clearly different as can be ascertained from Figure 2 with the
decay being faster for the bare gold surface. In addition, the
surface coverages for bare gold surfaces were consistently larger
than for DTSP-modified surfaces (Table 2). This could reflect
differences in packing arising from the interaction of XnOx with
the bare and DTSP-modified surfaces.
As mentioned above, the shape of the frequency-time profile
can be employed to study the kinetics of adsorption. The process
can be controlled by either transport (diffusion) or by kinetics
(activation controlled), which predict time dependencies of t1/2
and exp(t), respectively. Fits to both transport and kinetically
controlled models were carried out, and the latter gave consis-
tently better results. Assuming that the immobilization process
is kinetically controlled, the data were fit (Figure 2A and B solid
∆m ) -Cf∆F
(4)
where ∆m is the mass change (ng cm-2) and Cf (17.7 ng Hz-1
cm-2) is a proportionality constant for the 5.0-MHz crystals used
in this study. Assuming a molecular mass of 290.000 Da for
XnOx,34 a surface coverage value of ∼1.4 × 10-12 mol cm-2 was
obtained. It should be noted that, due to the asymmetric shape
of the protein (see Figure 3), the theoretical surface coverage
estimated from a monolayer of XnOx according to its molecular
size would be in the range of 1.2 × 10-12-2.7 × 10-12 mol cm-2
depending on orientation and packing in the monolayer.
As a comparison to the above measurements, QCM experi-
ments were also performed to study the direct adsorption of XnOx
onto a bare gold quartz crystal resonator under the same
conditions as described above for DTSP-modified resonators. As
(33) Sauerbrey, G. Z. Phys. 1959, 155, 206-222.
(34) Enroth, C.; Eger, B. T.; Okamoto, K.; Nishino, T.; Nishino, T.; Pai, E. F.
Proc. Natl. Acad. Sci. USA 2000, 97, 10723-10728.
534 Analytical Chemistry, Vol. 78, No. 2, January 15, 2006

line) to a first-order kinetics equation: ∆F ) ∆Fmax (1 - e-kt),
where ∆F is the frequency change (in Hz), ∆Fmax is the frequency
change between the initial and the final steady-state values, and
k is the first-order rate constant (expressed in min-1). The values
of ∆Fmax and k obtained from the fits are summarized in Table 2.
It is evident from the different values obtained for the constant
(k) that the frequency decreases more slowly when XnOx is
immobilized via covalent binding than through direct adsorption.
The difference in the kinetics of adsorption and in the final surface
coverage may be explained based on potentially different mech-
anisms involved in the immobilization of XnOx on DTSP-modified
and bare gold surfaces. On bare gold, the adsorption of the
enzyme ostensibly takes place with a random distribution of
protein adsorption geometries. However, when the immobilization
of XnOx takes place through DTSP, the process involves a
nucleophilic attack of the primary amino groups present in the
enzyme to the terminal N-succinimidyl esters exposed to the
solution in the monolayer. Therefore, in this case, the enzyme
immobilization will occur only for particular orientations (see
Figure 3) in which the amino groups are exposed to the substrate.
Thus, one would anticipate slower kinetics than in the case of
direct enzyme adsorption, which occurs in a nonspecific fashion.
To further characterize the nature of the resulting XnOx layers
obtained either by direct adsorption or covalently bonded through
DTSP, and to establish the differences between them, we have
carried out AFM experiments that provide morphological informa-
tion that complement the results obtained by QCM. The advantage
of AFM is that it allows characterization of the enzyme monolayer
at the local (i.e., nanoscale) level.
Figure 4. The 800 nm × 800 nm tapping mode AFM image taken
under buffer conditions of XnOx layer adsorbed on a DTSP-modified
gold substrate (A) and a bare gold substrate (B).
Panels A and B in Figures 4 show 800 nm × 800 nm images
of a DTSP-modified gold substrate and a bare gold substrate after
10 min of adsorption of XnOx. From Figure 4, it is evident that
the surface coverage obtained for the DTSP-modified surface is
smaller than that obtained for the bare gold surface. In addition,
closer inspection of Figure 4B reveals brighter spots, which
correspond to XnOx molecules adsorbed on top of an apparently
homogeneous layer in which globular structures are also ob-
served. These results are qualitatively consistent with the surface
coverage calculated from QCM plots (see Figure 2). Note that
the typical diameter of the globular structures obtained by AFM
is larger than the reported dimensions of XnOx.34 This is likely
due to tip convolution effects since the tip radius is similar to the
enzyme size. These effects become more evident for DTSP-
modified surfaces because the enzyme molecules appear isolated
on top of the flat substrate. In contrast, for bare gold, due to the
monolayer enzyme packing, the measured enzyme size is smaller.
In this case, the tip starts to image the next enzyme molecule
before reaching the underlying substrate because the proteins
are very close to each other. This relatively high enzyme packing
precludes a more detailed analysis of the height of the enzyme
structures. However, such analysis can be carried out for DTSP-
modified surfaces since the isolated adsorption of the enzymes
on the flat substrate allows us to determine their height. Note
that the highlighted protein in Figure 4B, which lies clearly
isolated on top of the protein layer, shows sizes similar to those
observed in Figure 4A. The corresponding height histogram
obtained from Figure 4A is shown in Figure 5. A relatively wide
height distribution is observed, ranging from 5 to 24 nm. The
Figure 5. Height histogram obtained from Figure 4A.
largest values (>17 nm), which correspond to 25% of the
observations, are likely related to aggregated structures since the
protein dimensions are about 17, 7, and 9 nm. The rest of the
features, with heights in the 5-17-nm range, represent 75% of
the observations, displaying a maximum for heights in the 14-
15-nm range. These values, taking into account the likely deforma-
tion of the enzyme when imaged by AFM, are in agreement with
the protein dimensions. In fact, the maximum at 14-15 nm
suggests a predominant geometry of adsorption with the longest
protein axis perpendicular to the substrate.
The DTSP-modified surfaces allow further analysis of the
distribution of the enzyme on the flat substrate and the protein
deformation induced by the tip during AFM imaging. First, from
the AFM image, one can obtain its autocovariance (Figure 6A).
This figure shows an outer diffuse bright circle (relative maxi-
mum) containing both a black ring (minimum) and a bright central
Analytical Chemistry, Vol. 78, No. 2, January 15, 2006 535

Figure 7. Approach AFM amplitude versus distance curves ob-
tained on XnOx-Au and XnOx-DTSP-Au monolayers and Au
surfaces in a liquid environment with the same cantilever. The dotted
vertical line indicates the tip-sample surface first contact point.
can be regenerated by the immobilized enzyme so that the
feedback current provides a direct measurement of enzymatic
activity. For systems with high catalytic rates, the SECM current
will clearly show a positive feedback response, whereas for a slow
process, a negative feedback response will be observed. We have
carried out analogous studies with immobilized XnOx using
hydroxymethylferrocene as redox mediator. Although thionin and
methylene blue are two-electron mediators, it is well known that
XnOx can also be catalyzed by one-electron mediator such as
hydroxymethylferrocene. We have employed hydroxymethylfer-
rocene as mediator rather than thionin or methylene blue because
of the propensity of the latter two to strongly adsorb. Such
adsorption could affect the measured response in an unpredictable
way, and thus, we used hydroxymethylferrocene as mediator even
though we recognize that it is not the most efficient.
Figure 8B shows the activity of an XnOx layer over an area 50
× 50 µm2 with hydroxymethylferrocene acting as mediator after
addition of xanthine to the solution. As can be observed, the tip
current measured by the SECM exhibits clear positive feedback
behavior. In contrast, no positive feedback was observed in the
absence of xanthine. From the difference between both images,
the spatially resolved enzymatic activity can be assessed. The
variations in feedback current, which would correspond to
variations in enzymatic activity, can be understood in terms of
the spatial distribution of the enzyme and its activity. As mentioned
in the Experimental Section, we employed direct adsorption for
enzyme immobilization. This method often gives rise to a
nonuniform spatial distribution of enzymes on surfaces, especially
in the monolayer regime, as was the case in this study. In addition,
it is often found that immobilized enzyme layers have nonuniform
activity, reflecting variations in the specifics of adsorption (and
conformation) for particular locations. Thus, such spatially inho-
mogeneous enzymatic activity would, in fact, be anticipated, as
we indeed observed.
Figure 6. (A) Autocovariance of the AFM image shown in Figure
4A. (B) Profile measured along the dotted line.
spot (global maximum). In Figure 6B is shown a profile of the
autocovariance in which a central absolute maximum flanked by
two smaller maximums is observed. The distance between the
central maximum and the smaller maximums corresponds to the
average distance between next protein neighbors. In our case,
this value is in the 70-95-nm range, which is larger than that
corresponding to that defined by two adjacent enzyme molecules
(15-35 nm). These results support that during the adsorption
process the XnOx molecules do not tend to adsorb close each
other, but rather leave a relatively large gap between them.
As we have mentioned previously, both isolated protein
structures and flat substrate regions are observed in Figure 4A.
Thus, we can obtain amplitude versus relative tip-surface distance
curves (Figure 7) on both regions with the same tip. From Figure
7, it is clear that the slope of the curve obtained on XnOx
structures is smaller than that corresponding to the DTSP-Au
substrate. Also, the slope of the curve obtained on DTSP-Au
substrates is smaller than that of the bare Au substrates. This
result implies that the protein is deformed by the tip action to a
larger extent than the DTSP layer. Furthermore, this deformation
can be quantified by measuring the horizontal shift between both
curves for a given A/A0 ratio. Thus, for A/A0 ) 0.75, the
deformation obtained is close to 5 nm, whereas for A/A0 ) 0.92,
it is close to 2-3 nm. It should be noted that for standard AFM
topographical imaging we employ A/A0 ratios in the 0.9-0.93
range.
Mapping of Local Enzyme Activity by Scanning Electro-
chemical Microscopy. To assess the reactivity of immobilized
XnOx and to map its reactive sites, SECM experiments, using
hydroxymethylferrocene as redox mediator, were carried out.
Previous SECM studies based on oxidoreductases, especially
glucose oxidase,7 have shown that in the feedback mode, the
redox mediator, whose active form is generated at the SECM tip,
Enzyme Electrode Response. In our configuration, the
immobilized XnOx oxidizes xanthine to uric acid in the presence
of thionin/methylene blue, which act as acceptors of electrons
generated in the enzymatic reaction and are transformed to their
reduced form. The mediator, in turn, diffuses to the electrode,
where it is reoxidized back to its oxidized form. The electrode
536 Analytical Chemistry, Vol. 78, No. 2, January 15, 2006

Figure 8. SECM surface plot image (50 × 50 µm2) of immobilized XnOx. Image was taken with a platinum microelectrode tip (10 µm, rate 15
µm s-1). Positive feedback with hydroxymethylferrocene mediator (1 mM) at ET ) +0.4 V vs Ag/AgCl in pH 6.5, 0.1 M phosphate buffer solution
in the absence (A) and in the presence of 3 mM xanthine (B).
acts as a secondary acceptor of electrons able to regenerate the
redox mediator used in the enzymatic reaction. This sequence of
redox events represents a catalytic process. Thus, the magnitude
of this catalytic current can be employed as the analytical signal
in the determination of the substrate (xanthine) concentration.
The steady-state current response obtained at +0.2 V for either
thionin or methylene blue was plotted as a function of the bulk
concentration of xanthine in solution. The response was linear to
xanthine concentration over the range of 5.0 × 10-5-8.0 × 10-4
M for both redox mediators. In the case of thionin, the response
(y ) 0.002 + 130x; r ) 0.99) exhibited a good sensitivity (130
nA/mM). We estimate a limit of detection of ∼30 µM. Methylene
blue (y ) 0.001 + 28x) exhibited a low sensitivity (28 nA/mM).
The high sensitivity exhibited by thionin compared to that
obtained with methylene blue agrees well with the results obtained
from the spectrophotometric studies reported above. Concerning
the stability of immobilized XnOx layer, it was found that the
biosensor response decreases ∼30% of its initial value after one
assay and then it remains constant for more than one month.
value of 1.4 × 10-12 mol cm-2. This value represents ∼50% of a
compact monolayer, assuming an orientation with the longest
enzyme axis perpendicular to the substrate. From QCM studies,
we have also been able to determine that the kinetics of absorption
is activation controlled with a value of 0.1 min-1 for the rate
constant. AFM measurements have provided morphological
characterizations that complement the results obtained by QCM.
Moreover, the AFM studies provided data concerning the protein
distribution on the substrate and the tip-induced deformation
during AFM imaging. AFM measurements showed that on DTSP-
modified surfaces XnOx molecules appear to adsorb in clusters
or domain separated by 70-95-nm gaps. Thionin and methylene
blue have been used as electron acceptors for reduced XnOx, and
the kinetic parameters and equilibrium constants of the mediated
enzymatic oxidation of xanthine have been calculated. Electro-
chemical and scanning electrochemical microscopy assays indicate
that the immobilized enzyme retains its enzymatic activity after
immobilization. Finally, a xanthine biosensor was developed
employing thionin.
ACKNOWLEDGMENT
CONCLUSIONS
This work was supported by the Ministerio de Ciencia y
A comprehensive study of a general bioanalytical platform for
Tecnolog´ıa of Spain through BQU2001-0163 and by the Univer-
sidad Auto´noma de Madrid through Proyecto Emergente 2005.
biosensor applications has been described. In particular, emphasis
has been placed on amperometric biosensors based on XnOx,
which was immobilized by covalent binding to gold electrodes
modified with DTSP. This material provides a general way of
covalent enzyme immobilization. The amount of immobilized
enzyme has been estimated from QCM measurements giving a
Received for review September 20, 2005. Accepted
November 1, 2005.
AC051676L
Analytical Chemistry, Vol. 78, No. 2, January 15, 2006 537