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Transcript of Investigation of a biofunctional polymeric coating deposited onto silicon microcantilevers
www.elsevier.com/locate/apsusc
Applied Surface Science 253 (2007) 4226–4231
Investigation of a biofunctional polymeric coating deposited
onto silicon microcantilevers
P. Bergese a,*, E. Bontempi a, M. Chiari b, P. Colombi a, F. Damin b,L.E. Depero a, G. Oliviero a, G. Pirri b, M. Zucca a
a INSTM and Laboratono di Chimica per le Tecnologie, Universita di Brescia, via Branze, 38, 25123 Brescia, Italyb Istituto di Chimica del Riconoscimento Molecolare, CNR, via Mario Bianco, 9, 20131 Milano, Italy
Received 3 August 2006; received in revised form 14 September 2006; accepted 14 September 2006
Available online 1 November 2006
Abstract
The paper deals with an appealing route to activate silicon microcantilevers (90, 110 and 130 mm long, 35 mm wide and 2 mm thick) for specific
binding of biochemical species. The method consists in coating the underivatized microcantilevers with a biofunctional copolymer (based on N,N-
dimethylacrylamide bearing silanating moieties) that was developed for low-density microarray assays on microscope glass slides. Coating
deposition was obtained by dip-coating and its microstructure investigated by analyzing the resonance frequency values of bare and coated
microcantilevers, by SEM and SFM imaging, SFM tip-scratch tests and XRR experiments. Results indicate that the coating is 2.5 nm thick and has
a density of 1.22 g/cm3. The coating surface is nanostructured, displaying nanoblobs, which are from few up to 20 nm wide and, on average, 1.6 nm
high. The diameter of the biggest nanoblobs is of the same order of magnitude of the gyration radius of the copolymer chains, suggesting that
nanoblobs may identify individual macromolecules.
# 2006 Elsevier B.V. All rights reserved.
PACS : 68.37.-d; 87.80.-y
Keywords: Coating; Polymers; Microcantilevers; Biofunctionalization
1. Introduction
Application of microcantilevers (MCs) to biosensing
promises to be a breakthrough in biochemistry, life science
and medicine [1–3]. Several experiments have been success-
fully performed in this direction, revealing DNA hybridization
(including detection of single nucleotide polymorphisms,
SNPs) [4–7] and detecting proteins and antibodies [8–11],
single virus particles [12–14] or bacteria [15,16].
MC biosensor working principle is simple: the MC is
functionalized with a probe that can selectively bind the target
biochemical species. Adsorption and binding site interactions
of the targets change the mechanical response of the MC system
(because of the mass increase and/or the surface stress
generated by changes in Gibbs free energy), providing the
transduction/sensing mechanism.
* Corresponding author. Tel.: +39 0303715802; fax: +39 0303702448.
E-mail address: [email protected] (P. Bergese).
0169-4332/$ – see front matter # 2006 Elsevier B.V. All rights reserved.
doi:10.1016/j.apsusc.2006.09.036
Probes immobilization can be obtained through direct
adsorption onto the MC surface [13,14,17] or through specific
functionalization methods. Indeed, accessibility, stability and
efficiency of surface functional groups are of crucial
importance and may constitute decisive feasibility drawbacks
for applications of MCs in biosensing experiments.
The use of thiolated molecules on gold coated MCs is the
most common functionalization method for DNA [4–7], protein
and antibodies [8–11]. Bacteria, viruses [18,19] and in some
cases proteins [20] have been immobilized by activation
techniques based on organosilanization. An alternative
approach to gold coating and organosilanization chemistry is
to coat the underivatized MCs with appropriate functional
polymers. This approach has found wide application in MC
chemical sensing [21–24]. On the contrary, to the best of our
knowledge, it was adopted only once in MC biosensing by
Gunter et al., who used a poly-ethylen-oxide (PEO) coating to
immobilize vaccinia polyclonal antibodies [25].
In the present paper the possibility of activating silicon MCs
by a co-polymeric thin coating based on N,N-dimethylacry-
P. Bergese et al. / Applied Surface Science 253 (2007) 4226–4231 4227
lamide bearing silanating moieties is assessed. This polymer
was developed for DNA and protein microarray assays on
microscope glass slides [26,27]. As showed in the previous
paper [26], the deposition of a thin layer of this polymer
represents a fast, inexpensive and robust method to covalently
bind amino-modified DNA probe strands on silicon dioxide
surfaces with high density. In addition, the film is stable in
aqueous buffers containing various additives, even at water
boiling temperature and, due to its hydrophilic nature and high
homogeneity, it minimizes non-specific interactions. This
feature is highly desirable for surface recognition bioassays,
for which interaction specificity is often an issue. Finally, the
macromolecular chains of the polymer act as spacer arms
keeping the probe molecules away from the surface more
effectively than self assembled monolayers (SAM) obtained by
organosilanization.
The deposition of a thin film of polymer onto the
underivatized MCs (90, 110 and 130 mm long, 35 mm wide
and 2 mm thick) was obtained by dip coating according to the
procedure previously devised for glass slides [26]. It is
worthwhile to notice that, because of the MC size, detailed
insight into the coating microstructure resulted a challenging
task. This goal was achieved by integrating the analysis of the
variation of the resonance frequency of the coated MCs with
respect to the bare MCs with the pieces of information obtained
from the images of scanning electron and scanning force
microscopies (SEM and SFM), SFM tip-scratch tests and from
X-ray reflectivity (XRR) data.
2. Experimental
The ter-polymer (DMA-co-NAS-co-MAPS) contains N,N-
dimethylacrylamide (DMA), N-acryloyloxysuccinimide (NAS)
and 3-(trimethoxysilyl)propyl methacrylate (MAPS), respec-
tively at 97%, 2% and 1% total monomer moles. The
procedures used to prepare and characterize the polymer are
reported in Ref. [26]. Hereafter it will be referred as (DMA-
NAS-MAPS).
We used commercial silicon MC arrays made of three MCs
(NSC12/Tipless/No Al by MikroMasch, Tallinn, Estonia). Each
array consists of three MCs of different lengths, namely 90, 110
and 130 mm, and of the same width and thickness: 35 and 2 mm,
respectively. Resonance frequency measurements were per-
formed in air, at room temperature using a scanning probe
microscope (SPM) head with an integrated laser lever readout
(Jeol, Tokyo, Japan). The Jeol SPM apparatus, equipped with
silicon tips by NT-MDT, Russia, was also used for SFM
experiments. Scanning electron microscopy (SEM) was
performed with a EVO 40 Microscope by LEO.
Polymer coating experiments were organized in two
separate batches. The coating procedure consisted of three
steps. Firstly the arrays were washed with acetone (Carlo Erba,
Milan, Italy) and after drying, they were immersed for 30 min
in a solution of the polymer (1% w/v in a water solution of
ammonium sulphate at 20% saturation level). Finally, they were
accurately washed with demineralized water and dried under
vacuum at 80 8C for 30 min.
The polymer deposition procedure previously assessed
[26,27] requires a pretreatment with HCl 1 M and NaOH
1 M. Such pretreatment is intended to activate the surface
silanols (a thin layer of native silicon dioxide is present on
the silicon surfaces of the MCs), maximizing the number of
hydrogen bonds with the polymer carbonyl groups and
ensuring the physisorption of a higher quantity of polymer.
However, the alkaline pretreatment resulted too aggressive
and changed the mechanical properties of MCs. This was
witnessed by an increase of the MC resonance frequency in
the order of tenth of kHz. In view of this, the MCs were just
carefully washed with acetone to eliminate superficial
contaminations.
In order to analyze the polymeric layer with X-ray
reflectivity (XRR), a silicon wafer slide (approximately
1.5 cm large and 2.5 cm long) was coated following the same
procedure and using the same polymer solution of the second
MCs batch. This allowed to obtain the best matching between
the MCs and the wafer coatings, as it was verified by SFM
analysis. XRR data were collected with a Bruker D8 Advance
reflectometer equipped with a Gobel mirror. The X-ray beam
(Cu Ka radiation, l = 0.154 nm) was collimated by proper slits
in order to set the cross section to 600 mm � 1.5 cm. All XRR
profiles were analyzed with the REFSIM software [28] in order
to extract information on the layer density, thickness and
roughness [29].
3. Results and discussion
The DMA-NAS-MAPS coating on the MCs was imaged by
SEM (Fig. 1) and quantitatively investigated by analyzing the
negative shift of the MC resonance frequency due to the mass of
the adsorbed DMA-NAS-MAPS layer (Fig. 2).
The SEM images of a DMA-NAS-MAPS coated MC
collected by secondary electrons and backscattered electrons
are reported in Fig. 1a and b, respectively. The secondary
electron image, which probes the surface morphology,
evidences that the polymeric layer is continuous, presenting
micron size corrugations localized far from the MC base.
These features disappear in the backscattered electron image.
Since backscattered electrons mainly probe the surface
chemical composition, this result indicates that the men-
tioned features are due to inhomogeneities of the polymeric
layer.
In order to evaluate the deposited mass of DMA-NAS-
MAPS, we used the general equation for the resonance
frequency ( f) of an oscillating cantilever in presence of fluid
damping and added mass [18,30]:
f ¼ 0:325
ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiK
mþ mf þ Dm;
r(1)
where K is the cantilever spring constant, m the cantilever mass,
mf the mass of the fluid film (mainly coming from ambient
moisture) adherent to the cantilever surface during oscillation
and Dm is the added mass. The constant 0.325 is a form factor
that accounts for the cantilever geometry. Eq. (1) is derived
P. Bergese et al. / Applied Surface Science 253 (2007) 4226–42314228
Fig. 1. (a) Secondary and (b) backscattered electrons SEM images of a DMA-
NAS-MAPS coated MC.
assuming that the added mass has no effect on the cantilever
mechanical properties. In general such assumption is valid
when the coating is much thinner than the cantilever or for
localized masses.
The DMA-NAS-MAPS coating on MCs resulted mainly
uniform, as showed by SEM imaging, and less than 5 nm thick
(see later in this section). Eq. (1) thus applies to the coated MCs
and can be simplified further. Actually both Dm and mf are
much smaller than m and can be neglected. Dm� m since the
coating is more than two orders of magnitude thinner than the
Fig. 2. Oscillation frequency profiles of a MC of the second batch before (solid
line) and after (dashed line) the coating with DMA-NAS-MAPS. The resonance
frequency shift, Df, between the two maxima is evidenced; according to Eq. (5),
it corresponds to a deposited mass per surface unit, Dms, of 6.7 fg/mm2.
MC, while the condition mf� m is ensured by the quality
factor of the frequency measurements (Fig. 2), which resulted
higher than 100 for all measurements. By these considerations,
Eq. (1) becomes:
f ¼ 0:325
ffiffiffiffiK
m
r: (2)
The equation that links the resonance frequency shift Df to
the mass variation (or added mass) Dm can be obtained by
differentiating Eq. (2):
Dm ¼ �2D f
fm ¼ �2
D f
frlwt; (3)
where l, w, t and r are the cantilever length, width, thickness,
and mass density, respectively. In the calculations we used the
nominal values of l and w, as they revealed at SEM check to be
reliable. On the contrary, the nominal value of t could not be
used as it resulted affected by a 30% uncertainty. Therefore t
was expressed through the Young modulus, E, using the rela-
tion:
t ¼ fl2
0:162
ffiffiffiffir
E
r; (4)
and taking into account that for the used silicon MCs
E = 169 GPa. In order to compare the coating of MCs of
different sizes it is convenient to use deposited masses per
surface unit (fg/mm2). Thus, by merging Eqs. (3) and (4), the
Fig. 3. Representative AFM image (non-contact mode) collected at the base of
a DMA-NAS-MAPS coated MC and height distribution curve. The vertical line
indicates the mean of the distribution.
P. Bergese et al. / Applied Surface Science 253 (2007) 4226–4231 4229
Fig. 4. SFM tip-scratch test (non-contact AFM image) at the base of a DMA-
NAS-MAPS coated MC. The reported line profile has been taken along the
white line.
Fig. 5. Representative AFM image (non-contact mode) collected on the DMA-
NAS-MAPS coated silicon wafer and height distribution curve. The vertical line
indicates the mean of the distribution.
operative equation we used in calculations reads:
Dms ¼ �12:35D f rl3w
S
ffiffiffiffir
E
r; (5)
where S = 2(lw + lt) + wt.
As described in Section 2, the experiments were organized in
two separate batches. The values of the deposited polymer mass
per surface unit, Dms, were calculated for each MC through
equation (5) and by using the experimental Df values (Fig. 2).
Finally, the mean value of Dms for each batch, Dm, was
obtained and the errors, intended as the standard deviations of
the means, calculated. Dm of the two batches resulted 10 � 2
and 7 � 1 fg/mm2, respectively. These values indicate fairly
good batch-to-batch reproducibility.
In order to gain insight into the microstructure of the DMA-
NAS-MAPS layer, scanning force microscopy (SFM) analysis
were performed on coated MCs. The coated MCs were scanned
in non-contact mode at their base, since it is a region in which
possible artifacts due to resonance interactions between the
observed MC and the SFM probe MC are prevented. A
representative SFM morphology image collected in non-
contact mode at the base of a coated MC and the related height
distribution curve are reported in Fig. 3. The coating surface
presents a nanostructure characterized by nanoblobs, which are
from few up to 20 nm wide with a mean height, according to the
height distribution curve, of 1.6 nm. The full width half
maximum (FWHM) of the height distribution curve is 1.5 nm.
It is worthwhile to note that the diameter of the biggest
nanoblobs is of the same order of magnitude of the gyration
radius of the DMA-NAS-MAPS polymer chains [31] suggest-
ing that each nanoblob may identify a single macromolecule.
We measured the thickness of the DMA-NAS-MAPS
coating by SFM tip-scratch tests [24,32,33]. The technique
consists in scratching the coating with the SFM tip. It is crucial
to set the normal force applied on the tip to a value high enough
to completely remove the polymeric layer but low enough to
avoid significant tip or silicon substrate damaging. After
scratching, the interested region is imaged in a non-contact
mode with the same tip used for scratching. Finally, the scratch
depth, i.e. the layer thickness, is evaluated by image analysis. A
representative scratch image of a scratched MC coating and a
related line profile (taken along the white line reported in the
image) are shown in Fig. 4. The line profile shows that the layer
thickness is about 2.5 nm. The surface area that was not
scratched is characterized by height oscillations of about 1 nm,
in agreement with the nanoblobs mean height evaluated by
SFM imaging. The spherical aggregates (hundreds of nm wide)
localized along the scratches are made of the polymer, which
was removed away from the scratches.
The average density of the DMA-NAS-MAPS coating was
evaluated by X-ray reflectivity (XRR) [29]. The experiment
was performed on a silicon wafer, which was coated by
P. Bergese et al. / Applied Surface Science 253 (2007) 4226–42314230
Fig. 6. SFM tip-scratch test (non-contact AFM image) collected on the DMA-
NAS-MAPS coated silicon wafer. The reported line profile has been taken along
the white line.
Table 1
XRR simulation parameters for the DMA-NAS-MAPS coated silicon wafer
Density
(g/cm3)
Thickness
(nm)
Roughness
(nm)
Wafer (Si) 2.32 1 <0.01
Layer 1 (SiO2) 2.98 1.52 0.12
Layer 2 (DMA-NAS-MAPS) 1.22 3.02 0.56
following the same procedure applied to MCs (see Experi-
mental section for details). The microstructural similarity
between MC and wafer coatings was verified through SFM by
repeating on the coated wafer the analysis performed on coated
MCs. The results, shown in Figs. 5 and 6, confirm a good
Fig. 7. Experimental and simulated XRR profiles of the DMA-NAS-MAPS
coated silicon wafer. The simulation parameters are reported in Table 1.
matching, and evidence some interesting fine differences. With
respect to MCs the DMA-NAS-MAPS forms on the wafer a
slightly thinner layer (about 2 nm) which presents a surface
characterized by less high nanoblobs (mean height of 1.3 nm)
with a narrower height distribution (FWHM = 0.6 nm), e.g.
lower surface roughness, intended as the FWHM of the height
distribution curve. These differences may be attributed to a
different adsorption efficiency of the surfaces due to the
differences of the native silicon oxide layers on the wafer and
MCs, respectively. Further investigation on the influence of the
silicon oxide layer on the DMA-NAS-MAPS adsorption is
ongoing.
The experimental and simulated XRR profiles and the
simulation parameters are reported in Fig. 7 and Table 1,
respectively. The polymeric layer resulted to be 3 nm thick,
with a surface roughness of 0.6 nm and a density of 1.22 g/cm3.
By keeping in mind that XRR overestimates the thickness of
layers of high roughness and that roughness values coincide
[29] it follows that XRR and SFM data are consistent.
The mass of adsorbed DMA-NAS-MAPS (on MCs) per
surface unit, Dms, may be exploited to assess the results
obtained from the different characterization techniques. Dms
can be calculated from the thickness t evaluated through SFM
tip-scratch tests and the density, rp, obtained by XRR,
according to the relation Dms = trp. It results Dms = 3 fg/
mm2, which is lower than the value obtained by analyzing the
MC resonance frequency, Dms ¼ 7� 1 fg=mm2. This differ-
ence was expected. Actually, Dms is related to uniform areas,
while Dms is an overall average that also accounts for the
coating inhomogeneities (Fig. 1). In view of this argument the
two values can be considered in fairly good agreement,
indicating that characterization based on MC resonance
frequency analysis is reliable for a coarse and quick (i.e.
routine) check of the coating.
Finally, by using Dms ¼ 7� 1 fg=mm2 and taking into
account that DMA-NAS-MAPS has an average molecular mass
of 1.5–105 amu [26] it follows that the coating layer (of the
second batch MCs) consists of (2.8 � 0.4) � 104 molecules/
mm2.
4. Conclusions
In this work we presented a method to activate silicon MCs
by a polymeric thin coating based on N,N-dimethylacrylamide
bearing silanating moieties [N,N-dimethylacrylamide (DMA),
N-acryloyloxysuccinimide (NAS) and 3-(trimethoxysilyl)pro-
pyl methacrylate (MAPS)]. The focus was placed on coating
characterization, which was achieved by analysis of the
P. Bergese et al. / Applied Surface Science 253 (2007) 4226–4231 4231
resonance frequency of the bare and coated MCs, SEM and
SFM imaging, SFM tip-scratch tests and XRR data. These
deeply different techniques provided complementary and
consistent results, which allowed to obtain a complete
microstructural picture of the coating.
MCs were coated with DMA-NAS-MAPS in two separate
batches by a dip-coating procedure, and a fairly good batch-to-
batch reproducibility was found. The coating is 2.5 nm thick
and has a density of 1.22 g/cm3. Its surface displays a
discontinuous nanostructure determined by nanoblobs, which
are from few up to 20 nm wide and, on average, 1.6 nm high.
From the average mass of adsorbed polymer per surface unit it
is possible to estimate that the coating consists of
(2.8 � 0.4) � 104 molecules/mm2. The diameter of the biggest
nanoblobs is of the same order of magnitude of the gyration
radius of the copolymer chains, suggesting that nanoblobs may
identify individual macromolecules.
This manuscript discusses the first issues encountered in
making innovative microcantilever-based (bio)sensors: suc-
cessful coating of microcantilevers with a high performance
functional layer and microstructural assessment of the coating,
which poses unique challenges due to the microscopic
dimensions of the microcantilevers. This latter aspect is often
neglected in the literature concerning microcantilever-based
sensors, even if it is mandatory for achieving interpretation of
sensing results, which goes beyond the straight ON/OFF
information.
Experiments to test the DMA-NAS-MAPS coated MCs as
biosensors are underway.
Acknowledgements
The authors acknowledge the funding of the Italian Ministry
of Education, University and Research through the project
PRIN 2005 ‘‘Nano-biosensors with micro-electro-mechanic
technology for genomic and proteomic analysis.’’ They also
wish to thank Alessandro Rivetti for SEM imaging.
References
[1] H.P. Lang, M. Hegner, C. Gerber, Nanotechnology 13 (2002) R29.
[2] R. Bashir, Adv. Drug Deliver. Rev. 56 (2004) 1565.
[3] K.K. Jain, Clin. Chim. Acta 358 (2005) 37.
[4] J. Fritz, M.K. Baller, H.P. Lang, H. Rothuizen, P. Vettiger, E. Meyer, H.J.
Guntherodt, C. Gerber, J.K. Gimzewski, Science 288 (2000) 316.
[5] R. Mukhopadhyay, M. Lorentzen, J. Kjems, F. Besenbacher, Langmuir 21
(2005) 8400.
[6] K.M. Hansen, H.F. Ji, G.H. Wu, R. Datar, R. Cote, A. Majumdar, T.
Thundat, Anal. Chem. 73 (2001) 1567.
[7] R. McKendry, J. Zhang, Y. Arntz, T. Strunz, M. Hegner, H.P. Lang, M.K.
Bailer, U. Certa, E. Meyer, H.J. Guntherodt, C. Gerber, Proc. Natl. Acad.
Sci. U.S.A. 99 (2002) 9783.
[8] G.H. Wu, R.H. Datar, K.M. Hansen, T. Thundat, R.J. Cote, A. Majumdar,
Nat. Biotech. 19 (2001) 856.
[9] P. Dutta, C.A. Tipple, N.V. Lavrik, P.G. Datskos, O. Hofstetter, M.J.
Sepaniak, Anal. Chem. 75 (2003) 2342.
[10] Y. Arntz, J.D. Seelig, H.P. Lang, J. Zhang, P. Hunziker, J.P. Ramseyer, E.
Meyer, M. Hegner, C. Gerber, Nanotechnology 14 (2003) 86.
[11] C. Grogan, R. Raiteri, G.M. O’Connor, T.J. Glynn, V. Cunningham, M.
Kane, M. Charlton, D. Leech, Biosens. Bioelectron. 17 (2002) 201.
[12] B. Ilic, Y. Yang, H.G. Craighead, Appl. Phys. Lett. 85 (2004) 2604.
[13] B. Ilic, D. Czaplewski, M. Zalalutdinov, H.G. Craighead, P. Neuzil, C.
Campagnolo, C. Batt, J. Vac. Sci. Technol. B 19 (2001) 2825.
[14] A. Gupta, D. Akin, R. Bashir, Appl. Phys. Lett. 84 (2004) 2004.
[15] K.Y. Gfeller, N. Nugaeva, M. Hegner, Biosens. Bioelectron. 21 (2005)
528.
[16] B.L. Weeks, J. Camarero, A. Noy, A.E. Miller, L. Stanker, J.J. Yoreo,
Scanning 25 (2003) 297.
[17] R. Marie, S. Schmid, A. Johansson, L.E. Ejsing, M. Nordstrom, D.
Hafliger, C.B.V. Christensen, A. Boisen, M. Dufva, Biosens. Bioelectron.
21 (2006) 1327.
[18] J. Zhang, H.F. Ji, Anal. Sci. 20 (2004) 585.
[19] N. Nugaeva, K.Y. Gfeller, Backmann, N. Lang, H.P. Duggelin, M. Hegner,
Biosens. Biolectron. 21 (2006) 849.
[20] G.A. Campbell, R. Mutharasan, Biosens. Bioelectron. 21 (2005) 597.
[21] F.M. Battiston, J.P. Ramseyer, H.P. Lang, M.K. Bailer, C. Gerber, J.K.
Gimzewski, E. Meyer, H.J. Guntherodt, Sens. Actuators B 77 (2001)
122.
[22] T.A. Betts, C.A. Tipple, M.J. Sepaniak, P.G. Datskos, Anal. Chim. Acta
422 (2000) 89.
[23] M. Maute, S. Raible, F.E. Prins, D.P. Kern, H. Ulmer, U. Weimar, W.
Gopel, Sens. Actuat. B 58 (1999) 505.
[24] G.G. Bumbu, G. Kircher, M. Wolkenhauer, R. Berger, J.S. Gutmann,
Macromol. Chem. Phys. 205 (2004) 1713.
[25] R.L. Gunter, W.G. Delinger, K. Manygoats, A. Kooser, T.L. Porter, Sens.
Actuators A: Phys. 107 (2003) 219.
[26] G. Pirri, F. Damin, M. Chiari, E. Bontempi, L.E. Depero, Anal. Chem. 76
(2004) 1352.
[27] M. Cretich, G. Pirri, F. Damin, I. Solinas, M. Chiari, Anal. Biochem. 332
(2004) 67.
[28] REFSIM Software, Bruker, 1998.
[29] E. Bontempi, Recent Research Development in Chemical Physics, Trans-
world Research Network, 2004, p. 461.
[30] A.N. Cleland, Foundations of Nanomechanics, Springer, 2003.
[31] E.A. Doherty, K.D. Berglund, B.A. Buchholz, I.V. Kourkine, T.M. Przy-
bycien, R.D. Tilton, A.E. Barron, Electrophoresis 23 (2002) 2766.
[32] C. Ton-That, A.G. Shard, R.H. Bradley, Langmuir 16 (2000) 2281.
[33] F. Anariba, S.H. DuVall, R.L. McCreery, Anal. Chem. 75 (2003) 3844.