surface functionalization chemistries on highly sensitive silica-based sensor chips
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Surface functionalization chemistries on highly sensitive silica-based sensorchips†
Subash C. B. Gopinath,*a Koichi Awazu,a Makoto Fujimaki,*a Kazufumi Shimizu,b Wataru Mizutanic
and Kiyomi Tsukagoshic
Received 5th December 2011, Accepted 6th May 2012
DOI: 10.1039/c2an35159e
The surfaces of silica-based sensor chips, designed for evanescent-field-coupled waveguide-mode
sensors, were functionalized using various surface chemistries. The immobilization of molecular entities
on the functionalized silica surfaces was monitored using various microscopic techniques (scanning
electron, fluorescence, and atomic force microscopies). Further, gold nanoparticle-based signal
enhancement analyses were performed with protein conjugation on different functionalized surfaces
using a waveguide-mode sensor. Based on these analyses, the sensor surfaces modified with
glutaraldehyde (Glu) and carbonyldiimidazole were found to be good for molecules of different sizes.
In addition, it can be inferred that the Glu-modified surface may be suitable for small molecules with
diameters around 5 nm owing to its surface roughness. The modified surface with carbonyldiimidazole
is suitable for the direct immobilization of larger molecules especially for biomolecular assemblies
without intermediate chemical modifications.
Introduction
Surface modification is an approach for modifying the surface of
a material by physical, chemical, or biological means to produce
characteristics that are different from those of the original
surface. These surface modifications are highly dependent on
various factors such as roughness, hydrophilicity, surface charge,
surface energy, biocompatibility, and reactivity. On the other
hand, surface functionalization is a process of introducing
chemical functional groups on the surface to capture the mole-
cules to be analyzed, with a view to several applications, in
various fields involving biochemical, biophysical, biomedical and
molecular biological reactions for specific assays. High-density
microarrays on a solid glass support, known as protein chips or
DNA chips, are highly dependent on chemical modification in
order to accommodate higher numbers of molecules, and they
are powerful tools for gene discovery, expression, and mapping
analyses.1–4 The development of analytical devices requires the
formatting of various substrates in order to couple them with
the molecules of interest for various read-out formats of the
aElectronics and Photonics Research Institute, National Institute ofAdvanced Industrial Science and Technology (AIST), Central 5, 1-1-1Higashi, Tsukuba, Ibaraki, 305-8565, Japan. E-mail:[email protected]; [email protected] Research Center for Genome and Infectious Diseases, NihonUniversity School of Medicine, 30-1 Oyaguchikami-chou, Itabashi-ku,Tokyo 173-8610, JapancNanosystem Research Institute, AIST, Central 4, 1-1-1 Higashi, Tsukuba,Ibaraki, 305-8562, Japan
† Electronic supplementary information (ESI) available. See DOI:10.1039/c2an35159e
3520 | Analyst, 2012, 137, 3520–3527
molecular assembly or application area, such as surface plasmon
resonance, quartz crystal microbalance, microscopic methods,
cantilever-based methods, affinity chromatography, and elec-
trochemical and fluorescence methods.5
Among various metallic and non-metallic substrates, silica-
derived substrates are considered as one of the most versatile
materials with the potential for several chemical modifications.
They have been used to develop relatively cheap sensor surfaces
for analytical systems.6–13 Silica-based substrates provide a vari-
ation in the degree of packing densities, thicknesses, and
morphologies on the deposited surfaces.14,15 Silica-based nano-
particles are potential candidates for the development of nano-
scale composites with optical and chemical properties on a single
structure.16 In order to attach biomolecules to the silica surfaces,
various surface modifications have been proposed in previous
studies, including the attachment of modified biotin through
amino-coupling,15,17,18 a biotin–streptavidin–biotin sandwich on
an amino surface,10,19 the use of N-(2-trifluoroethane-
sulfonatoethyl)-N-(methyl)-triethoxysilylpropyl-3-amine-linked
oligonucleotides for duplex formation,6,10 the attachment of
proteins to amino couplings through glutaraldehyde (Glu)
molecules,11,19,20 an antibody–protein–antibody sandwich on
amino-coupled Glu,11 and thiol-coupling.8,19 By using a similar
strategy, aptamers have been attached to silica-based nano-
particles for cancer cell detection.21 Amino-coupling based
attachments on the silica surface have been quite common for
several years, and it is recognized that the high-density immo-
bilization of proteins on silica or other surfaces is necessary to
allow the use of low sample volumes.3,21,22 Parameters such as
chemistry, incubation time, reaction temperature, homogeneity,
This journal is ª The Royal Society of Chemistry 2012
Fig. 1 Spectral readout system of the EFC-WM sensor. The light from
the tungsten halogen lamp was guided to a collimator lens and the
collimated light was irradiated to a prism. The monolithic sensing plate
placed on the bottom of a prism was illuminated and a spectrum of the
reflected light was observed by a spectrophotometer. The prism was made
of SiO2 glass, and the bottom angle of the prism was 38�.
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spacing between molecules, conformation of captures, and
buffering condition influence the attachment of biomolecules.23
Further, improper immobilization may lead to loss of protein
activity partially or completely with wrong orientation of mole-
cules and structural deformation. Therefore, the choice of
immobilization chemistry is important before attaching the
ligand and analyte.3 Availing of these surface properties,
biosensor developments were pursued to analyze various sizes of
biomolecules and biomolecule–nanoparticle conjugates. Among
various proteins and oligonucleotides, there will be size varia-
tions and whole cell analyses will have larger molecules, neces-
sary to immobilize on the sensing surface with the suitable linker.
Recent interest in the sensor developments is to create a system to
analyze and screen small molecules, especially for the drug-
discovery processes. Previous studies carried out with different
surface modifications are mainly focused to study the property of
molecular interactions.7,10,11,13,15,24 Chemical and physical
properties of size-dependent nanoparticles for the sensitive
detections were overviewed by Wittenberg and Haynes.25
However, there are lacunae in the study of surface modifica-
tions for the immobilization of molecules based on molecular
sizes. To fill these lacunae, the present study investigates the use
of different surface functionalization chemistry, using carbonyl-
diimidazole (CDI), glycidoxypropyltrimethoxysilane (GOPTS),
and Glu for molecular size selection. All these molecules were
intended to be used with the sensor chip surfaces that are
routinely used in our laboratory to monitor various biomolecular
interactions, for an evanescent-field-coupled waveguide-mode
sensor (EFC-WM sensor) operating on the Kretschmann
principle.10,11,13,15,17–19,26
Experimental
Chemicals and biomolecules
3-(Triethoxysilyl)propan-1-amine (3APT) was purchased from
Sigma-Aldrich (Tokyo, Japan). N,N0-Carbonyldiimidazole
(CDI) and Glu were procured from Wako Chemicals (Osaka,
Japan); 3-glycidoxypropyltrimethoxysilane (GOPTS), from
Shin-Etsu, Silicon Chemicals, Japan; Alexa Fluor 555-labeled
goat anti-rabbit IgG (H+L), from Invitrogen, USA; gold nano-
particle (GNP) conjugated streptavidin (5 nm of 3 O.D. and
20 nm of 4 O.D.), from BBInternational, UK; 40 nm GNP–
streptavidin conjugates (15 O.D.), from BioAssay Works, MD,
USA; and anti-HA serum for H1N1 (Brisbane/59/2007) from
Denka Seiken, Tokyo, Japan.
Setup of the EFC-WM sensor
The EFC-WM sensor utilizes a sensing plate having a multilayer
structure consisting of a dielectric waveguide, a high-refractive
index layer, and a glass substrate.18 The sensing plate illuminated
under the Kretschmann configuration operates as a sensor that is
capable of detecting modifications in the dielectric environment
near the waveguide surface by measuring change in reflectivity.
In the present research, we applied a spectral readout system to
the EFC-WM sensor, which had been reported as a compact
optical system of SPR sensors.27 The optical setup is shown in
Fig. 1. In the system, a tungsten halogen lamp was used as a light
source. The light from the lamp was guided to a collimator lens
This journal is ª The Royal Society of Chemistry 2012
and the collimated light was irradiated to a prism, where the
incident angle was parallel to the bottom face of the prism. Then,
the monolithic sensing plate placed on the bottom of a prism is
illuminated and a spectrum of the reflected light is observed by
a spectrophotometer. The prism was made of SiO2 glass, and the
bottom angle of the prism was 38�. As the sensing plate,
a monolithic sensing plate that we previously developed was
applied.18 The monolithic sensing plate consists of a SiO2 glass
substrate, a single crystalline Si layer, and a thermally grown
SiO2 waveguide. In the present experiment, the thicknesses of the
SiO2 glass waveguide and the single crystalline Si layer were set to
be 45 and 360 nm, respectively. The optical system was designed
to show a dip in reflectance at around 520 nm, which corresponds
to the optical absorption of GNPs. If GNPs are attached on the
waveguide surface, the dip will be deepened by the optical
absorption of the GNPs.15
Reactions for the functionalization of the silica surface
Three different chemical strategies (CDI, GOPTS and Glu) were
employed for attaching the biomolecules to the silica-based
sensor chips. Before performing the chemical modifications, the
chips were treated with alkali solution for 30 min, washed
thoroughly with water, and dried to attach OH groups to the
surface of the silica. Free hydroxide ions were removed by
washing several times with phosphate buffer (pH 7.4). For the
CDI modification, the alkali-treated sensor chip was soaked in
a 0.5 M solution of CDI in dioxane. The reaction was carried out
at 37 �C for 2 h, after which the chip was rinsed with acetone
followed by distilled water and dried. For GOPTS modification,
GOPTS reactions on the sensor surface were performed by
immersion in anhydrous toluene containing 2% GOPTS for 4 h
at 50 �C, followed by washing with toluene and water to remove
adsorbed silane. For the Glu modification, silane coupling using
3APT on the chip followed by immobilization with Glu was
performed as previously described.10,19 In brief, the alkali
modified surface of the waveguide substrate was further modified
by immersion in a 0.5% (v/v) ethanolic solution of 3APT for 24 h.
The 3APT reacted with surface hydroxyl groups of the SiO2
waveguide to give an amine group-functionalized substrate that
was rinsed with ethanol and dried in a stream of nitrogen gas.
The amine-functionalized SiO2 surface was treated with a 2.5%
solution of aqueous Glu for 3 h at room temperature to form an
aldehyde-activated surface. After incubation, the surface was
washed thoroughly with the phosphate buffer. Unless otherwise
stated, surface modified glass substrates were stored in glass Petri
dishes until further analysis.
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On each surface modified sensing plate, experiments were
performed with 1 optical density of the GNP–streptavidin
conjugate, as this concentration did not give non-specificity on
the unmodified surfaces under optimal room temperature. The
dilutions for 1 optical density were made from the original stock
with phosphate buffer (pH 7.4). Reactions were performed for 20
min on the shell of the waveguide sensor and after each step the
sensing surface was thoroughly washed three times with 300 ml of
phosphate buffer.
Microscopic analyses
All microscopic observations were performed with 1 optical
density of GNP–streptavidin conjugates, as described above. The
GNP–streptavidin conjugate attached surfaces were examined
using field emission scanning electron microscopy (SEM; JEOL,
JSM-6340F). The surface functionalized with different chemis-
tries was observed by atomic force microscopy (AFM; Digital
Instruments Nanoscope). The AFM measurements were carried
out with a 1 mm scan size at a scan rate of 0.5003 Hz. Fluores-
cence images of Alexa Fluor 555-labeled goat anti-rabbit IgG
(H+L) attached surfaces were visualized using a BIOREVO
Keyence BZ-9000 instrument. 50 nM of a fluorescent-labeled
antibody was attached on each chemically modified surface and
incubated for 30 min at room temperature, washed, and dried
before observations were carried out.
Results and discussion
The sensor chips used for surface functionalization are correctly
designed for the EFC-WM sensors, and are stable against
chemical and physical changes. The principles and design of the
waveguide sensor system that we used were similar to that of
a surface plasmon resonance (SPR) system, the only difference
being that the mode used for measurement was not a surface
mode but a waveguide mode. The advantage of the waveguide
sensor over the SPR sensor is that higher sensitivity is easily
obtained by perforating the waveguide layer and/or by using dyes
or metal nanoparticles. In the case of the SPR sensor, the
wavelength of an incident light is restricted by the material used
in order to induce the surface plasmon, whereas there is no such
restriction in the case of the waveguide sensor. In addition,
amorphous SiO2, one of the most popular materials of the
waveguide layer of the waveguide sensor, is physically and
chemically more stable than typical materials (Au or Ag)
generally used for SPR.10,15,18,19 Moreover, silica materials are
inexpensive and have a relatively homogeneous chemical surface,
which offers the added advantage of easier scanning for visual-
ization of the spots for chemical and biochemical analyses.6,28
Silicates and silicon oxide surfaces require surface modifications
before attachment of molecules of interest.5 Attempts were made
to control the surface chemistry of these chips because surface
control is crucial for efficient immobilization of proteins in the
preparation of micro-nanodevices. There are two in vogue
approaches to the surface immobilization and patterning of solid
surfaces, namely, in situ surface synthesis and preparation of
prefabricated molecules.29Generally, in order to retain biological
activities on the sensor-surfaces, modifications of physico-
chemical and chemical properties are the pertinent options.
3522 | Analyst, 2012, 137, 3520–3527
Surface functionalization chemistry on waveguide sensor chips
The sensor surface was activated by alkali treatment and the
resulting surface has Si–OH groups. On these Si–OH surfaces,
the functionalization chemistry was incorporated with three
independent appropriate surface modifications, namely CDI,
GOPTS and Glu. CDI is an organic compound that forms
a white crystalline solid. It is often used for the coupling of amino
acids for peptide synthesis and as a reagent in organic synthesis.
CDI is very often used for coupling carboxylic acids with
aliphatic or aromatic amines to form amides for both small- and
large-scale applications.30 It can be dissolved in the solvent and
directly attached to the silica surfaces without the need for an
intermediate step, to produce silane residues,9 and thus, modifi-
cation with this reagent reduces the number of experimental
steps. A CDI functionalized surface will possess an imidazole
carbamate functionality and it can form a stable carbamate
linkage with amine groups. The epoxy-silane in GOPTS allows
for the formation of a covalent linkage to tertiary amine groups
on amino acids.20 Epoxy activated surfaces are considered to be
a universal type of support, on which nucleophilic and electro-
philic groups can be attached with high efficiency.31However, the
reaction time for this covalent reaction with proteins is higher
than that of the other modifications discussed;3,6 also, GOPTS
polymerizes easily in the presence of water to form multilayer
assemblies.32 This compound was dissolved in anhydrous toluene
to create an epoxylated surface and it forms a secondary amine
bond upon reacting with amines. GOPTS is stable at neutral pH,
wettable and reactive with several nucleophilic groups to form
strong bonds with protein.3 Glu needs an additional modification
step to immobilize the molecules on the sensor chip by first
modifying the chip with silane groups and then attaching Glu. A
glass substrate derived from organosilanes can be functionalized
for –OH, –NH2, –SH, –COOH, –CHO and so on, to capture the
subsequent molecules of interest (Todt and Blohm, 2009). In this
study, 3-(triethoxysilyl)propan-1-amine (in 95% ethanol) is used,
as this compound is considered as one of the potent functiona-
lization chemistries for glass substrates that can introduce reac-
tive amino groups on the surface. This surface activated with
amines was reacted with aldehyde groups of Glu. Glu is
a homobifunctional cross-linker, frequently used in biochemistry
applications as an amine-reactive group, and the oligomeric state
of proteins can be examined through this application. Glu has
two aldehyde groups at the ends and both ends can be coupled
with amine groups to form a Schiff’s base. As reported in the
literature, the Glu coupling reaction on chip was performed at
a pH of 7.4, a slightly alkaline environment, as a lower pH than
this would reduce the efficiency of the coupling reaction.33
Attachment of gold nanoparticle conjugated-streptavidin on
sensor chips: SEM and waveguide sensor analyses
Nanomaterials can facilitate signal transduction just like fluo-
rophores or electroactive tags and be useful for imaging studies.
GNP is the ideal nanomaterial commonly used in the sensor
developments, has unique characteristics, such as easy to disperse
in water, compatible with surface functionalization to link
biomolecules and can be tailored with the desired nano-sizes.34,35
In this study, the activity of the modified waveguide sensing
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surfaces was evaluated using GNPs of three different sizes. As
mentioned above, the surface chemistry modifications were
carried out using CDI, GOPTS, and Glu as modifiers (Fig. 2a).
Then, GNP–streptavidin conjugates were immobilized on the
modified waveguide-mode sensor chips, using GNPs of three
different sizes (Fig. 2b). Streptavidin is a tetravalent protein
molecule with a diameter of approximately 5 nm.10 Upon coating
the 40 nm sized gold nanoparticles with this molecule, the
conjugates may have a total diameter of approximately 50 nm,
whereas the 20 and 5 nm GNPs will produce conjugates with
sizes of 30 and 10 or 15 nm, respectively (Fig. 2b). Streptavidin
was chosen as the model protein for conjugation with GNP
because it finds widespread use in molecular biology owing to its
extraordinarily strong affinity for the vitamin biotin. The disso-
ciation constant (Kd) of the biotin–streptavidin complex is of the
order of 4 � 10�14 mol l�1, ranking it as one of the strongest
known non-covalent interactions in molecular, immunological
and cellular assays.36
SEM observation of the 40 nm GNP–streptavidin conjugates
showed clear attachment on all three chemically modified silica
surfaces used in the present study (Fig. 3a–c). As mentioned,
streptavidin has a size of about 5 nm, and the size of the GNP
conjugates using a 40 nm GNP is increased to roughly 50 nm
upon conjugation, in a non-aggregation situation (Fig. 2b). It is
hard to predict the number of streptavidin molecules attached on
GNP. To minimize the formation of aggregated forms of GNP,
the usage of NaCl was avoided. The addition of salt (NaCl) will
cap the repulsion among the unmodified negatively charged
GNPs, inducing the aggregation of these particles and failure to
stabilize.37,38 As mentioned in our previous report, the waveguide
system detects less than one GNP adsorbed per square
micrometer. According to our theoretical calculation based on
the experimental data the reflectance is expected to decrease by
0.01 (reflectivity in the vertical direction) by the adsorption of
0.75 GNP per square micrometer.15 When sensing surfaces are
analyzed with the EFC-WM sensor before and after attaching
the GNP–streptavidin conjugates, prominent changes in the dip
of the resonance were observed in the cases of CDI and Glu
Fig. 2 (a) Schematic surface functionalization on the sensor chip. CDI,
GOPTS, and Glu modifications are shown. (b) Diagrammatic represen-
tation of the sizes of the GNP–streptavidin conjugates. Three different
sizes (40, 20, and 5 nm) of GNP were used. The size of streptavidin is
considered to be 5 nm. The expected total sizes of GNP-conjugated
streptavidin are shown.
This journal is ª The Royal Society of Chemistry 2012
modification (Fig. 3d and f). In both cases, the reflection
changes in the dip were with values of 18.7 and 21.4, respectively.
In contrast, epoxy activation by surface modification with
GOPTS produced a reflection change with nearly half the value
(11.7) of those observed in the other two cases, indicating
stronger molecular attachments with CDI and Glu molecules
(Fig. 3d–f).
Further evaluation was performed by similar studies on all
three surfaces using 20 nm GNPs–streptavidin conjugates. These
GNP conjugates may have an estimated size of approximately
30 nm. Direct observation with SEM using the three chemically
modified surfaces shows the attachment of a larger number of
molecules of this size on the CDI modified surfaces than on the
GOPTS and Glu modified surfaces (Fig. 4a–c). Confirmatory
evidence was provided in the observation of the reflectance
changes using a EFC-WM sensor, similar to that presented
above. The observations with the sensor also support the
conclusion that the CDI-modified surface exhibited preferential
attachment of these particles, as more prominent reflectance
changes in the resonance dip were observed in the case of the
CDI modified surface, compared with the other two. With CDI-
modification the reflectance changes were about 18.7 which is
similar to 40 nm GNP–streptavidin attachment, whereas in the
case of Glu the reflectance change was roughly one fourth of the
reflectance observed for the CDI case. In the case of GOPTS
modification, no significant changes were observed in the dip in
the resonance (Fig. 4d–f).
Because the two studies presented above indicated clear
molecular-size discrimination, similar analyses were extended to
include 5 nmGNPs conjugated with streptavidin. The 5 nmGNP
can accommodate fewer streptavidin molecules, potentially
accommodating one or two streptavidin molecules per particle
on either side, to give conjugates with a size of 10 nm or 15 nm in
total (Fig. 2b). The 5 nm GNP was too small for the SEM system
that we used and we could not observe it using the system.
Waveguide-mode reflectance analysis of the 5 nm GNP–strep-
tavidin conjugates showed a reflectance change of 7.7 in the case
of the Glu-modified surface, which is higher than the value
observed for this surface using the 20 nm GNP–streptavidin
conjugates. The reflectance differences (with 20 nm and 5 nm
GNP–streptavidin) might be attributed to molecular accommo-
dation fitness on the Glu-modified surface in the case of the 5 nm
conjugates, relative to the 20 nm GNP–streptavidin conjugates.
The reflectance difference is, thus, due to a higher molecular
density of the 5 nm molecules attached to the Glu-immobilized
surface and a relatively lower density in the 20 nm case. However,
in the case of CDI-modification, there was a marked decrease in
the changes of the resonance dip, due to fewer molecular
attachments of the 5 nm conjugates on this surface compared to
the case of the 20 nm GNP–streptavidin conjugates (Fig. 5a and
b). As observed in the case of the 20 nm GNP–streptavidin
conjugates, there were no visible changes with the 5 nm GNP–
streptavidin conjugates for the GOPTS modifications (Fig. 5c).
The comprehensive analyses with three sizes of GNP–streptavi-
din conjugates and three chemical modifications are shown in
Fig. 5d. The non-specific attachments of all of the GNPs used in
this study were monitored on chemically un-modified surfaces,
and no significant non-specific attachments were observed (ESI,
Fig. 1a–c†).
Analyst, 2012, 137, 3520–3527 | 3523
Fig. 3 SEM images of 40 nmGNP–streptavidin conjugates on the surfaces modified with (a) CDI, (b) GOPTS and (c) Glu. Magnified SEM images are
shown as figure inset. Reflectivity measurements were carried out with the EFC-WM sensor using these surfaces on which the 40 nm GNP–streptavidin
conjugate was immobilized. (d), (e), and (f) illustrate the immobilization of the particles on CDI, GOPTS and Glu, respectively. The black and the red
curves indicate the spectra before and after the immobilization, respectively. Higher changes in reflectivity were observed with CDI and Glu, and lower
one was observed with GOPTS. Vertical arrows on the figures indicate the direction of the changes.
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Based on this observation, it can be deduced that particles
having larger sizes, i.e., �50 nm, are best suited for the surfaces
chemically modified with CDI and Glu, from which it can also be
extrapolated that these surfaces might be well suited for the
larger biomolecules such as nanoparticles and viruses. In one of
our recent studies to observe the interactions of antibody and
whole viral particles, a Glu-modified surface was used and good
sensitivity was achieved.19 In addition, the results indicate that
there is a significant dependence of the molecular attachments on
these surfaces, upon the molecular sizes below 50 nm. This might
be attributed to the high chemical molecular crowdedness in the
case of Glu modification, thereby restricting molecular sizes
around 30 nm, as the surface roughness forms ridges. Ultimately,
Fig. 4 SEM images of 20 nmGNP–streptavidin conjugates on the surfaces m
shown as figure inset. The particle immobilized surfaces were monitored by th
the particles on CDI, GOPTS and Glu, respectively. The black and the red cur
Vertical arrows on the figures indicate the direction of the changes.
3524 | Analyst, 2012, 137, 3520–3527
there was the possibility for the formation of ordered, arranged
molecular attachments on the CDI modified surfaces, but in the
case of 50 nm particle sizes used in the above study, the reflec-
tance was high due to larger particle sizes. These studies clearly
illustrate that the fit on the modified surfaces is based on the
chemical molecular crowding and molecular accommodation
operates within size-limitations.
Fluorescence microscopy analyses
Based on the above studies, there were expectations that there
would be differences in the surfaces of the sensors with the
various modifications. Consequently, the studies were further
odified with (a) CDI, (b) GOPTS, and (c) Glu. Magnified SEM images are
e waveguide-mode sensor. (d), (e), and (f) illustrate the immobilization of
ves indicate the spectra before and after the immobilization, respectively.
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Fig. 5 Reflectivity measurements with the EFC-WM sensor using the
surfaces modified with (a) CDI, (b) GOPTS and (c) Glu, on which the
5 nm GNP–streptavidin conjugate was immobilized. The black and
the red curves indicate the spectra before and after the immobilization,
respectively. The changes in reflectivity observed by the immobilization
of the three GNP–streptavidin conjugates with different sizes on the three
chemical modifications are summarized in (d). The error bars indicate the
averages of three values.
Fig. 6 Fluorescence microscopy images with fluorescent (Alexa Fluor
555) labeled antibody immobilized on the surfaces modified with (a) CDI,
(b) GOPTS, (c) Glu, and (d) the unmodified surface.
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extended to other microscopic approaches. Fluorescence
microscopy analyses were performed on the three chemically
modified surfaces using Alexa Fluor 555 labeled goat anti-rabbit
IgG. Alexa Fluor is a family of fluorescent dyes, a unique dye
obtained from Invitrogen, typically used for fluorescence
microscopy observations. The excitation and emission spectra of
Alexa Fluor dyes covers the visible spectrum and extends into the
infrared; and these dyes are generally more stable, brighter, and
less pH-sensitive than other common dyes. The results of the
fluorescence microscopy studies showed the presence of fewer
antibody molecules on the GOPTS-modified surface compared
with the CDI-modified surface (Fig. 6a and b). Several tens of
molecules could be clearly observed on the CDI surface. In the
case of the Glu-modified surface, a totally different behavior was
observed compared with the CDI and GOPTS surfaces, in which
there was a dense image as seen in the fluorescence microscopy
studies (Fig. 6c). The immobilizations of the fluorescent-labeled
antibody on the unmodified surfaces were also visualized as
blanks (Fig. 6d). The high-background caused by Glu has the
potential for creating cross-linkages between two protein mole-
cules via homobifunctional aldehyde groups leading to undefined
complex formation on the immobilized surface. Glu is also
reported to polymerize in aqueous solution to give a number of
molecular structures, which may cause immobilization of
a branched polyaldehyde rather than the native bifunctional
molecule.9 Another possibility could be the higher number of
molecular attachments of the amine and the Glu functionalized
surface as reported by Goddard and Erickson,9 resulting in
conjugation of Glu with higher hybridization densities. This clear
discrimination on the Glu-coated surface evidenced by the
indistinct image exemplified the correlation with the GNP–
streptavidin studies regarding the molecular size requirement for
attachment. Usually antibodies have a molecular weight in the
region of 150 000 and they may be similar in size to the 5 nm
This journal is ª The Royal Society of Chemistry 2012
GNP–streptavidin conjugated particles. In both cases, there
might be plenty of molecular attachments. Further, the fluores-
cence microscopy analyses allow us to conclude that Glu-modi-
fication may allow for the attachment of several proteins, as they
are the right-sized molecules. Even though our studies indicate
that the GOPTS-modified surface cannot accommodate many
molecules, the report of Goddard and Erickson9 indicates that
GOPTS and CDI behave similarly in their studies. As proposed
before, because GOPTS requires a longer reaction time, the
reaction efficiency may be lower than that of the other two
chemistries used.6
Validation of surface roughness by AFM on the CDI and Glu
surfaces
The immobilization of proteins via chemical reactions of the
amino acid side-chains are generally considered as random due
to the presence of different residues on different proteins, thereby
forming a heterogenic population on the sensor chip. Attach-
ments of the proteins through chemical bonds may be guided by
ordered processes to give proper orientation.3 However, this is
dependent on the chemical processes involved in the modification
procedure. The AFM approach was chosen for direct visualiza-
tion of the surface ordered-ness on the present sensor chip.
Based on the above experiments using protein conjugated
GNP and un-conjugated protein molecules, it was concluded
that there is a drastic change in the surface with both CDI and
Glu. To prove this, we monitored and measured the surface
roughness of both CDI and Glu modified surfaces after immo-
bilization of these chemical compounds. In the case of CDI, the
modification proceeds by a single step; therefore, the surface
roughness was observed directly for CDI using AFM, whereas in
the case of Glu the modification involves two steps, and surface
observations were made after each step. When the 3APT-coupled
surface, which is first step for Glu modification, was analyzed,
slight changes were observed in the roughness of the surface
(Fig. 7a). The surface roughness was increased drastically upon
Analyst, 2012, 137, 3520–3527 | 3525
Fig. 7 AFM images of the surfaces modified with (a) 3-APT, (b) Glu, and (c) CDI. The scanned images are also given at the bottom of each figure.
Graphical representation of surface roughness is shown in (d). The root-mean-square (RMS) roughnesses were measured at three different positions. (e)
An AFM image of antibodies attached on the CDI-modified surface.
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immobilization of the Glu molecules on the 3APT-coupled
surface (Fig. 7b), whereas, in the case of CDI, the roughness was
not as high compared with the Glu modified surface (Fig. 7c).
The overall comparison of these three surface roughnesses indi-
cates that they were different in each case as shown in Fig. 7d.
Antibody attachment on the CDI surface was monitored using
anti-Brisbane influenza viral anti-serum. The antibody was
directly attached to the CDI surface, and the antibody molecules
were clearly observed on the surface (Fig. 7e). This clear
discrimination between the attachment on the Glu and CDI
surface based on roughness might be attributed to crowdedness
and orientation of the molecular arrangements. In the case of
Glu, there are two aldehyde groups at the ends, and there is,
therefore, a higher probability for formation of a randomly
arranged chemical surface which will lead to a surface with more
roughness. Due to the rough nature of the Glu-immobilized
surfaces, there will be more surface area to receive lower
molecular sized molecules. Diagrammatic illustrations repre-
senting the molecular ordered-ness with different-sized molecules
on different surface chemistries are shown in the ESI, Fig. 2†.
Conclusion
Based on these results, it is clear that even though there is
a higher probability for more protein attachments on the Glu-
modified surface, it can be inferred that the CDI surface might be
the best choice for larger molecules (particles and viruses) as well
as for various molecular interactive analyses or biomolecular
assembly; and this surface can make direct linkage with the
molecules. However, for direct observation of proteins or protein
interactions with lower-sized molecules such as drugs, Glu might
be helpful and ideal for molecules with restricted sizes. The
3526 | Analyst, 2012, 137, 3520–3527
protein biochips with these chemical modifications will be useful
as diagnostic tools for proteomic analyses to obtain information
about the protein functions and interactions. Further, the
nanoparticle-conjugated proteins on these chemically modified
surfaces will enable the development of surfaces for nano-
biosensors, and nano-alignment using the particles can be carried
out on these sensor chip surfaces for nano-patterning purposes.
Acknowledgements
This study was supported by a grant from the Industrial Tech-
nology Research Program, 2009, of the New Energy and
Industrial Technology Development Organization (NEDO),
Japan. Part of this work was conducted at the AIST Nano-
Processing Facility, which is supported by the ‘‘Nanotechnology
Support Project’’ of the Ministry of Education, Culture, Sports,
Science and Technology (MEXT), Japan. The authors would
also like to thank the Advanced Functional Materials Research
Center of Shin-Etsu Chemical Co., Ltd. for supplying the
samples.
References
1 E. Defrancq, A. Hoang, F. Vinet and P. Dumy, Bioorg. Med. Chem.Lett., 2003, 13, 2683–2686.
2 F. Fixe, M. Dufva, P. Telleman and C. B. V. Christensen, NucleicAcids Res., 2004, 32(1), e9.
3 F. Rusmini, Z. Zhong and J. Feijen, Biomacromolecules, 2007, 8,1775–1789.
4 T. Sch€uler, A. Nykytenko, A. Csaki, R. M€oller, W. Fritzsche andJ. Popp, Anal. Bioanal. Chem., 2009, 395, 1097–1105.
5 S. Balamurugan, A. Obubuafo, S. A. Soper and D. A. Spivak, Anal.Bioanal. Chem., 2008, 390, 1009–1021.
6 P. Kumar, J. Choithani and K. C. Gupta, Nucleic Acids Res., 2004,32, e80.
This journal is ª The Royal Society of Chemistry 2012
Dow
nloa
ded
by C
orne
ll U
nive
rsity
on
05 J
uly
2012
Publ
ishe
d on
08
May
201
2 on
http
://pu
bs.r
sc.o
rg |
doi:1
0.10
39/C
2AN
3515
9E
View Online
7 G. Rong, A. Najmaie, J. E. Sipe and S. M. Weiss, Biosens.Bioelectron., 2008, 23, 1572–1576.
8 D. Goncalves, D. M. F. Prazeres, V. Chu and J. P. Conde, Biosens.Bioelectron., 2008, 24, 545–551.
9 J. M. Goddard and D. Erickson, Anal. Bioanal. Chem., 2009, 394,469–479.
10 S. C. B. Gopinath, K. Awazu, M. Fujimaki, K. Sugimoto, Y. Ohki,T. Komatsubara, J. Tominaga, K. C. Gupta and P. K. R. Kumar,Anal. Chem., 2008, 80, 6602–6609.
11 S. C. B. Gopinath, K. Awazu, M. Fujimaki, K. Sugimoto, Y. Ohki,T. Komatsubara, J. Tominaga and P. K. R. Kumar, Anal. Bioanal.Chem., 2009, 394, 481–488.
12 S. C. B. Gopinath, R. Kumaresan, K. Awazu, M. Fujimaki,M. Mizuhata, J. Tominaga and P. K. R. Kumar, Anal. Bioanal.Chem., 2010, 398, 751–758.
13 S. C. B. Gopinath, K. Awazu, M. Fujimaki and P. K. R. Kumar,Sens. Actuators, B, 2011, 155, 239–244.
14 J. Sabate, M. A. Anderson, H. Kikkawa, Q. Xu, S. Cervera-Marchand C. G. Hill, Jr, J. Catal., 1992, 134, 36–46.
15 M. Fujimaki, K. Nomura, K. Sato, T. Kato, S. C. B. Gopinath,X. Wang, K. Awazu, T. Komatsubara and Y. Ohki, Opt. Express,2010, 18, 15732–15740.
16 L. Wang, W. Zhao and W. Tan, Nano Res., 2008, 1, 99–115.17 K. Awazu, C. Rockstuhl, M. Fujimaki, N. Fukuda, J. Tominaga,
T. Komatsubara, T. Ikeda and Y. Ohki, Opt. Express, 2007, 15,2592–2597.
18 M. Fujimaki, C. Rockstuhl, X. Wang, K. Awazu, J. Tominaga,Y. Koganezawa, Y. Ohki and T. Komatsubara, Opt. Express, 2008,16, 6408–6416.
19 S. C. B. Gopinath, K. Awazu and M. Fujimaki, Anal. Methods, 2010,2, 1880–1884.
20 E. W. Olle, J. Messamore, M. P. Deogracias, S. D. McClintock,T. D. Anderson and K. J. Johnson, Exp. Mol. Pathol., 2005, 79,206–209.
21 C. D. Medley, S. Bamrungsap, W. Tan and J. E. Smith, Anal. Chem.,2011, 83, 727–734.
This journal is ª The Royal Society of Chemistry 2012
22 J. B. Lamture, K. L. Beattie, B. E. Burke, M. D. Eggers, D. J. Ehrlich,R. Fowler, M. A. Hollis, B. B. Kosicki, R. K. Reich, S. R. Smith,R. S. Varma and M. E. Hogan, Nucleic Acids Res., 1994, 22, 2121–2125.
23 S. Todt and D. H. Blohm, in DNA Microarrays for BiomedicalResearch: Methods and Protocols, Humana Press, 2009, vol. 529,pp. 81–100.
24 S. Devanathan, M. C. Walker, Z. Salamon and G. Tollin, J. Pharm.Biomed. Anal., 2004, 36, 711–719.
25 N. J. Wittenberg and C. L. Haynes,Wiley Interdiscip. Rev.: Nanomed.Nanobiotechnol., 2009, 1, 237–254.
26 E. Kretschmann, Z. Physik, 1971, 241, 313–337.27 O. R. Bolduc, L. S. Live and J. Masson, Talanta, 2009, 77, 1680–1687.28 E. LeProust, H. Zhang, P. Yu, X. Zhou and X. Gao, Nucleic Acids
Res., 2001, 29, 2171–2180.29 N. Dendane, A. Hoang, E. Defrancq, F. Vinet and P. Dumy, Bioorg.
Med. Chem. Lett., 2008, 18, 2540–2543.30 E. K. Woodman, J. G. K. Chaffey, P. A. Hopes, D. R. J. Hose and
J. P. Gilday, Org. Process Res. Dev., 2009, 13, 106–113.31 S. Mahajan, P. Kumar and K. C. Gupta, Bioconjugate Chem., 2006,
17, 1184–1189.32 A. K. Y. Wong and U. J. Krull, Anal. Bioanal. Chem., 2005, 383, 187–
200.33 D. Jamin, J. Demers, I. Shulman, H. T. Lam and R. Momparler,
Blood, 1986, 67, 993–996.34 V. K. Upadhyayula, Anal. Chim. Acta, 2012, 715, 1–18.35 B. S. Guirgis, C. S�a E Cunha, I. Gomes, M. Cavadas, I. Silva,
G. Doria, G. L. Blatch, P. V. Baptista, E. Pereira, H. M. Azzazy,M. M. Mota, M. Prudencio and R. Franco, Anal. Bioanal. Chem.,2012, 402, 1019–1027.
36 A. Holmberg, A. Blomstergren, O. Nord, M. Lukacs, J. Lundebergand M. Uhlen, Electrophoresis, 2005, 26, 501–510.
37 K. Song, M. Cho, H. Jo, K. Min, S. H. Jeon, T. Kim, M. S. Han,J. K. Ku and C. Ban, Anal. Biochem., 2011, 415, 175–181.
38 L. Li, B. Li, Y. Qi and J. Jin, Anal. Bioanal. Chem., 2009, 393, 2051–2057.
Analyst, 2012, 137, 3520–3527 | 3527