Protein micro- and nanopatterning using aminosilanes with protein-resistant photolabile protecting groups

Article abstract of DOI:10.1021/ja1103662

An approach to the integration of nanolithography with synthetic chemical methodology is described, in which near-field optical techniques are used to selectively deprotect films formed by the adsorption of aminosilanes protected by modified 2-nitrophenylethoxycarbonyl (NPEOC) groups. The NPEOC groups are functionalized at the m- or p-position with either a tetraethyleneglycol or a heptaethylene glycol adduct. We describe the synthesis of these bioresistant aminosilanes and the characterization of the resulting photoreactive films. Photodeprotection by exposure to UV light (λ = 325 nm) yielded the amine with high efficiency, at a similar rate for all four adsorbates, and was complete after an exposure of 2.24 J cm^-2. Following photodeprotection, derivatization by trifluoroacetic anhydride was carried out with high efficiency. Micropatterned samples, formed using a mask, were derivatized with aldehyde-functionalized polymer nanoparticles and, following derivatization with biotin, were used to form patterns of avidin-coated polymer particles. Fluorescence microscopy and atomic force microscopy data demonstrated that the intact protecting groups conferred excellent resistance to nonspecific adsorption. Nanometer-scale patterns were created using scanning near-field photolithography and were derivatized with biotin. Subsequent conjugation with avidin-functionalized polymer nanoparticles yielded clear fluorescence images that indicated dense attachment to the nanostructures and excellent protein resistance on the surrounding surface. These simple photocleavable protecting group strategies, combined with the use of near-field exposure, offer excellent prospects for the control of surface reactivity at nanometer resolution in biological systems and offer promise for integrating the top-down and bottom-up molecular fabrication paradigms.

Full text of DOI:10.1021/ja1103662

ARTICLE
pubs.acs.org/JACS
Protein Micro- and Nanopatterning Using Aminosilanes with
Protein-Resistant Photolabile Protecting Groups
†
,||,#
§,||
†,‡
†,‡
,†
Shahrul A. Alang Ahmad, Lu Shin Wong, Ehtsham ul-Haq, Jamie K. Hobbs, Graham J. Leggett,*
,
§
and Jason Micklefield*
†
Department of Chemistry, University of Sheffield, Brook Hill, Sheffield S3 7HF, United Kingdom
‡
Department of Physics and Astronomy, University of Sheffield, Hicks Building, Hounsfield Road, Sheffield S3 7RH, United Kingdom
§
School of Chemistry & Manchester Interdisciplinary Biocentre, The University of Manchester, 131 Princess Street,
Manchester M1 7DN, United Kingdom
S Supporting Information
b
ABSTRACT: An approach to the integration of nanolithography
with synthetic chemical methodology is described, in which
near-field optical techniques are used to selectively deprotect
films formed by the adsorption of aminosilanes protected by
modified 2-nitrophenylethoxycarbonyl (NPEOC) groups. The
NPEOC groups are functionalized at the m- or p-position with
either a tetraethyleneglycol or a heptaethylene glycol adduct.
We describe the synthesis of these bioresistant aminosilanes and
the characterization of the resulting photoreactive films. Photodeprotection by exposure to UV light (λ = 325 nm) yielded the amine
-
2
with high efficiency, at a similar rate for all four adsorbates, and was complete after an exposure of 2.24 J cm . Following
photodeprotection, derivatization by trifluoroacetic anhydride was carried out with high efficiency. Micropatterned samples, formed
using a mask, were derivatized with aldehyde-functionalized polymer nanoparticles and, following derivatization with biotin, were
used to form patterns of avidin-coated polymer particles. Fluorescence microscopy and atomic force microscopy data demonstrated
that the intact protecting groups conferred excellent resistance to nonspecific adsorption. Nanometer-scale patterns were created
using scanning near-field photolithography and were derivatized with biotin. Subsequent conjugation with avidin-functionalized
polymer nanoparticles yielded clear fluorescence images that indicated dense attachment to the nanostructures and excellent protein
resistance on the surrounding surface. These simple photocleavable protecting group strategies, combined with the use of near-field
exposure, offer excellent prospects for the control of surface reactivity at nanometer resolution in biological systems and offer
promise for integrating the top-down and bottom-up molecular fabrication paradigms.
’
INTRODUCTION
framework, we require generalized methodologies that permit
the creation, on demand, of a range of surface molecular
architectures.
The integration of top-down (lithographic) with bottom-up
(synthetic chemical) approaches remains a central challenge in
Photochemical methods have a long-standing and well-established
place in synthetic chemical methodology. The development
nanofabrication. The majority of work on molecular nano-
patterning has utilized physical methods for the deposition
or removal
1
-5
19
6
-8
of light-directed chemical synthesis by Fodor et al. enabled
the translation of synthetic chemical methodology into micro-
technology, by exploiting photolithography, in combination with
the deployment of highly efficient photocleavable protecting
groups, to carry out spatially defined solid-phase synthesis. The
use of light for this purpose is appealing because it enables
of material, with subsequent elaboration of
structures formed this way being possible via a variety of routes
9
(for example, host-guest chemistry or interactions between
10
Ni-NTA and His-tagged proteins ). However, the development
of tools for the direct initiation of specific chemical transforma-
tions on nanometer length scales would add a new dimension to
the construction of nanostructured assemblies. The importance
“
reagent-free” deprotections and allows a high degree of spatial
registry. Such approaches have proved enormously valuable in
11,12
of this was recognized by Sagiv and co-workers,
who used a
20
oligonucleotide synthesis and form the basis for the solid-phase
potential difference applied between an AFM tip and a surface to
drive oxidative modification of silane films. Subsequently, a
number of approaches to the excitation of specific chemical
pathways have been reported, including methods based upon
electrochemistry, nanopipettes, and the use of tip-immobilized
enzymes, catalytically active metals,
fabrication of DNA chips by sequential photodeprotection-base
21-23
coupling steps.
Light-directed synthesis thus provides a model for the integra-
tion of top-down and bottom-up fabrication methods. The main
drawback is the limited resolution of conventional far-field
13
14
1
5
16,17
and metal nano-
1
8
particles to catalyze surface reactions. However, for the full
integration of synthetic chemical methodology into a top-down
Received: November 18, 2010
Published: February 8, 2011
r 2011 American Chemical Society
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ARTICLE
optical techniques. However, extensive work in the authors’
laboratories has demonstrated that near-field optical techniques
enable the translation of photochemical patterning to the nano-
nanofabricated features, and any undesired adsorption results in
a significant degradation in resolution and performance. Hence,
any strategy for the integration of top-down and bottom up
methodologies that is to be capable of use for biomolecular
nanofabrication must also address the important problem of
nonspecific adsorption. Ideally for the fabrication of materials
with integrated biomolecules, a surface is required that is
bioresistant and chemically passive, yet switchable to a surface
that is able to undergo further synthetic manipulation upon
photoirradiation.
2
4,25
meter scale,
because evanescent (near) fields are not subject
to the diffraction effects that limit the resolution of conventional
optical methods. In scanning near-field photolithography (SNP),
a UV laser coupled to a scanning near-field optical microscope is
used to initiate photochemical reactions. Using SNP, structures
as small as 9 nm have been fabricated in self-assembled mono-
26
layers (SAMs) of alkylthiolates on gold. The methodology has
subsequently been extended to a wide range of other materials
28,29
In the present work, we have addressed these complex
challenges by attaching a nitrophenylethoxycarbonyl protecting
group to aminopropyltriethoxysilane, because amine groups
(formed by photodeprotection) are a convenient basis for
subsequent solid-phase synthesis. We demonstrate the reactivity
of deprotected amines by coupling biotin using a succinimidyl
ester derivative, although the free amine offers a generic point
from which many other bioconjugation strategies are accessible.
To prevent nonspecific adsorption, we have functionalized the
27
including monolayers of alkylsiloxanes and phosphonic acids,
3
0,31
32
as well as films of polymers,
metallic nanoparticles. and
3
3
fullerenes. Recently, we described new approaches to the
parallelization of near-field lithography, with a device (the
“
Snomipede”) that uses parallel near-field exposure to execute
photodeprotection steps in organic films and can achieve a spatial
resolution of 70 nm with an array of probes immersed under
3
4
water.
62-66
To fully integrate top-down and bottom-up approaches, it is
necessary to develop generic protecting group strategies that
enable the photoswitchable or photoactivatable introduction
of a wide range of functional groups to surfaces. Nitrophenyl-
based protecting groups are very promising in this respect. The
phenyl ring with protein-resistant oligoethylene glycol chains.
For alkylthiolate SAMs, a minimum of three ethylene glycol
6
7
(
EG) units is required to ensure protein resistance. Because
siloxane films are known to exhibit lower order and a wider
distance between neighboring headgroups compared to alkylthiol
SAMs, we have compared protecting groups functionalized with
four and seven EG units, to ensure complete coverage of the
underlying surface. Further, because the effect of the OEG
attachment on the photochemistry of the nitrophenylethyloxy-
carbonyl group is unknown, we have additionally compared
molecules with OEG substituents located m- and p- to the nitro
group.
35
o-nitroveratryloxycarbonyl (NVOC) and R-methyl-o-nitro-
36-38
piperonyloxycarbonyl (MeNPOC)
groups are now well
established in synthetic chemistry and are suitable for the
protection of carboxy and hydroxy moieties, while the recently
introduced nitrophenylpropyloxycarbonyl (NPPOC) protecting
2
0,39
group
may be used to protect amines and yields an excep-
40
tionally high photodeprotection efficiency. MeNPOC caging
of biotinylated surfaces has been used in micrometer scale
41
biological patterning, while Nakayama et al. attached biotin
to a nitrophenyl photocleavable linker, enabling its selective
’
RESULTS AND DISCUSSION
42
removal from surfaces. In a different approach, Alonso et al.
synthesized an NVOC protected aminosilane, which, following
photodeprotection, was derivatized with tris-nitrilo triacetic acid
Synthesis of Photoreactive Silanes. The syntheses of all the
photoreactive silanes were completed following a similar route
(Scheme 1). Initially, monomethoxy oligoethylene glycols 1a,b
43
68,69
(tris-NTA), enabling the immobilization of His-tagged proteins.
were first tosylated
and then treated with the appropriate
Other promising avenues offering generic surface functionaliza-
methylnitrophenol isomers to give 2a-d in good yields. These
44,45
tion capability include the use of thiol-ene reactions
and
were then converted to the alcohols 3a-d with paraformalde-
46
t
70
photoactivated click reactions. These results highlight the
power of photochemistry for carrying out organic synthesis at
surfaces. However, none of these papers has demonstrated
molecular patterning with nanometer-scale spatial resolution.
In efforts to merge the versatility of this synthetic chemistry and
the spatial resolution of near-field photolithography, we recently
reported the synthesis of NPPOC-protected aminopropyl-
triethoxysilane and demonstrated that nanometer-scale patterns
may readily be generated by using a near-field probe to expose
hyde under KO Bu catalysis in DMSO. Unlike previous reports
40,47,71
with the NPPOC and related photolabile groups,
the use
of Triton B as the base did not yield any desired product in all
cases. For 3a-c, these were then coupled with 3-isocyanatopropyl-
47
triethoxysilane to give the desired products 4a-c. However, 4d
0
was found to be inseparable from the N,N -di(3-triethoxysilyl-
propyl)urea side product. This was instead prepared through the
intermediate formation of the 4-nitrophenyl carbonate followed
by reaction with 3-aminopropyltriethoxysilane in a one-pot two-
step sequence.
Film Formation and Characterization. Films were formed
from the nitrophenylethyoxycarbonyl-protected aminopropyl-
triethoxysilanes 4a-d, henceforth NPEOC-APTES. The advan-
cing water contact angles of all four films were in close agree-
ment, falling in the range 43.8-44.9° (Table 1). The differences
between the values obtained for the films were small as compared
to the experimental error, suggesting that the composition of the
surface presented by all four adsorbate films is similar.
Film thicknesses were measured using ellipsometry. Films
4
7
the adsorbates. These deprotected amine groups could sub-
sequently undergo a variety of chemical manipulations including
acetylations and imine formation.
Nanometer-scale biological structures offer the potential for
addressing a plethora of scientific problems, including the
development of ultrasensitive detection systems for biological
4
8-57
diagnostics
cellular attachment.
and fundamental studies of mechanisms of
5
,53,58-61
However, biological systems pre-
sent several important and difficult issues for nanofabrication
including, in particular, nonspecific adsorption (fouling). Pro-
teins are particularly challenging in this regard because they
adsorb strongly and irreversibly to most surfaces. At nanometer
length scales, biomolecules are on a comparable size scale to the
of tetraethyleneglycol NPEOC-APTES (EG NPEOC-APTES)
4
were ca. 6 nm thick, making them fractionally thicker than the
4
7
films of NPPOC-APTES reported in an earlier study (5.5 nm),
2
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Table 1. Contact Angle and Ellipsometry Data for Films of Photosenstive Silanes
protecting group
NPEOC (4b) m-EG
m-EG
4
NPEOC (from 4a)
p-EG
4
7
NPEOC (4c)
7
p-EG NPEOC (4d)
θ
a
/deg
44.9 ( 1.5
5.7 ( 0.30
44.5 ( 0.9
6.3 ( 0.3
43.1 ( 0.8
9.2 ( 1.1
43.8 ( 0.8
8.9 ( 0.5
ellipsometric thickness/nm
a
Scheme 1. Sy nthesis of Photoreactive Silanes
a
2
Reagents and conditions: (a) TsCl, NaOH, THF/H O, 3 h, 84% (1a)
or 70% (1b); (b) 4-methyl-3-nitrophenol or 3-methyl-4-nitrophenol,
K
2
CO
3
, acetone (2a-c) or DCM (2d), reflux, 6 h (2a and 2b) or 16 h
(
(
2c and 2d), 91% (2a) or 88% (2b) or 89% (2c) or 84% (2d);
c) paraformaldehyde, KO Bu (cat.), DMSO, 80 °C, 30 min, 35% (3a) or
t
34% (3b) or 25% (3c) or 30% (3d); (d) 3-isocyanatopropyltriethoxy-
silane, Et
3
N (cat.), DCM, reflux, 24 h, 63% (4a) or 62% (4b) or 65%
N, DCM, 6 h, then 3-amino-
(
4c); (e) 4-nitrophenol chloroformate, Et
3
propyltriethoxysilane, EtOH/DCM, 16 h, 30% (4d over two steps).
Figure 1. C 1s (top) and N 1s (bottom) photoelectron spectra for
EG NPEOC-APTES monolayers.
which were estimated to be ca. two molecular layers thick.
n
EG NPEOC-APTES 4a and 4b are longer than NPPOC-
4
APTES, by approximately the length of the EG unit, estimated
the carbon atoms in the EG tail and also the carbon next to the
nitrogen of the carbamate group. There is additionally a compo-
nent corresponding to the carbonyl carbon atom. In each case,
the ether peak was slightly larger than anticipated, based on the
molecular formula of the adsorbate, because of attenuation of the
photoelectrons emitted from the underlying carbon atoms.
The N 1s spectra exhibited two components at binding
energies of ca. 401 and 407 eV, attributed to the carbamate
and nitro group nitrogen atoms, respectively. On the basis of
consideration of the molecular formulas of the adsorbates, these
components would be expected to exhibit the same areas.
However, the nitro group component was smaller than the
carbamate component in all spectra. The samples were all stored
in the dark so photodeprotection was unlikely. Rather, the
observation was attributed to X-ray-induced modification of
the samples during analysis. To confirm that this was the case,
and that deprotection of adsorbates did not occur during storage,
4
to be ca. 1 nm. Hence, it may be concluded that the EG NPEOC-
4
APTES films were also close to bilayer thickness, on average.
Films of EG NPEOC-APTES 4c and 4d were thicker than films
7
of EG NPEOC-APTES by ca. 3 nm, consistent with the forma-
4
tion of a film of bilayer thickness, given the greater length of the
heptaethylene glycol adducts. These data are consistent with the
well-documented tendency of silanes to self-polymerize, and in
general to form more disordered films than comparable alkyl-
thiolates on gold. The film thicknesses reported here were highly
reproducible under the conditions used.
X-ray photoelectron spectra were acquired for all four films.
Figure 1 shows the C 1s and N 1s spectra. There are two
dominant peaks in each C 1s spectrum: the main hydrocarbon
peak at 285 eV, corresponding to the carbon atoms in the
aromatic ring, the carbon adjacent to it, and the propyl chain;
and a peak at ca. 1.5 eV higher binding energy corresponding to
2
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Table 2. Contact Angles of Films Formed from EG NPEOC-
n
APTES Following UV Exposure and Derivatization with
TFAA in THF
θ
a
/virgin
θ
a
/3 min
a
θ /5 min
protecting group
film
exposure
exposure
m-EG
4
NPEOC (from 4a)
46.3 ( 2.5
45.3 ( 2.1
44.0 ( 1.0
45.0 ( 1.0
68.3 ( 2.1
69.7 ( 1.2
66.5 ( 1.5
65.3 ( 3.5
92.0 ( 3.4
89.4 ( 2.1
87.5 ( 4.0
88.8 ( 2.5
p-EG
4
NPEOC (4b)
m-EG
7
NPEOC (4c)
p-EG
7
NPEOC (4d)
Figure 2. N 1s photoelectron spectra following exposure to UV light for
m-EG NPEOC-APTES.
7
Figure 3. Variation in the ratio of the areas of the nitro and carbamate/
amine components in the N 1s spectra of films formed from EG N-
n
PEOC-APTES.
samples were incubated with trifluoroacetic anhydride (TFAA),
and their contact angles were measured. The values obtained
were analytically identical to those of the films before incubation,
suggesting no derivatization had occurred and hence that no
deprotection of the adsorbates had occurred. In contrast, after
photodeprotection, significant changes were observed in the
contact angle following reaction with TFAA (see below). It has
been reported that nitrophenyl groups are reduced rapidly upon
Figure 4. Tapping mode AFM images, and representative line sections,
showing micropatterns formed by deprotection of films with different
protecting groups through a mask and subsequent derivatization with
aldehyde-functionalized polymer nanoparticles. (a) m-EG NPEOC;
4
(
b) m-EG NPEOC; (c) p-EG NPEOC; (d) p-EG NPEOC. Vertical
7 4 7
72
exposure to X-rays and electrons, and this has been used
scale range: 0-100 nm dark to light.
73
successfully as a means of patterning nitrophenyl SAMs. The
reduced area of the nitro components in the N 1s spectra is here
attributed to a similar process. A similar although much less
pronounced effect was also observed with the NPPOC-APTES
carbamate component for the unmodified adsorbate and the
amine component following deprotection) was measured as a
function of the UV exposure time (Figure 3). The ratio is less
than unity for the virgin materials, as described above. Further,
the limiting value reached at long exposures was not zero. Taken
together with the elipsometry data, which indicate that the
average thickness of the assembled films corresponded to two
layers of siloxanes, this result suggests that the top layer may be
47
that we previously reported.
Photodeprotection. Figure 2 shows the evolution of the N
s region of the XPS spectrum with UV exposure for m-
1
EG NPEOC-APTES. The nitro component was visible in the
7
spectrum of the virgin material but quickly declined following
exposure to UV light, and after 5 min exposure (1.86 J cm-2) the
intensity of this component was very much reduced.
The N1 s spectra of the other three adsorbates exhibited very
similar qualitative changes as a function of exposure. To quanti-
tatively compare the behavior of the four adsorbates, the ratio of
the nitro peak to the main peak (representing the sum of the
partially preventing the removal of the EG NPEOC groups in
n
underlying layer. For all four adsorbates, a limiting value was
-
2
reached after 6 min (2.24 J cm ), and the rate of change was
similar in all four cases.
To confirm that deprotection of the adsorbates was occurring,
samples were derivatized in two different ways. First, deprotected
2
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Figure 5. (a-d) Fluorescence micrographs showing micropatterns formed by attachment of NeutrAvidin particles to biotinylated micropatterns
formed in films with different protecting groups. (a) m-EG NPEOC; (b) p-EG NPEOC; (c) m-EG NPEOC; (d) p-EG NPEOC. (e-h) Tapping
mode topographical AFM images of micropatterns formed by attachment of NeutrAvidin to biotinylated, deprotected films. (e) m-EG NPEOC;
f) p-EG NPEOC; (g) m-EG NPEOC; (h) p-EG NPEOC. Cross sections along the dashed lines indicated are shown beneath (e)-(h).
4
4
7
7
4
(
4
7
7
films were immersed in a dilute solution of TFAA in tetrahy-
drofuran (THF). Reaction between TFAA and the deprotected
amine groups yielded an increase in the water contact angle of the
film due to the increased hydrophobicity of the trifluoroacetyl-
ated surfaces. Table 2 shows data for virgin film (i.e., not exposed
to UV light) and films that have been exposed to UV light for
was measured to be ca. 40 nm, consistent with the formation of a
monolayer of immobilized nanoparticles in those regions.
Protein Patterning. Deprotected EG NPEOC-APTES
n
films were functionalized by biotin molecules attached to
N-hydroxysuccinimidyl ester-terminated linkers (henceforth
NHS-biotin). NeutrAvidin particles (avidin-coated nanoparticles
fluorescently labeled with fluorescein isothiocyanate (FITC))
were coupled to the patterns via the specific biotin-avidin
recognition interaction, and the samples were imaged using
confocal fluorescence microscopy (Figure 5a-d). Adhesion of
the NeutrAvidin particles to the surface is dominated by the
interactions between the avidin coating and the surface; they thus
exhibit adsorption characteristics similar to those of Avidin, but
also possess a significant loading of dye (FITC) facilitating
imaging by fluorescence microscopy. Bright fluorescence was
observed from the exposed regions (squares) where the OEG-
functionalized protecting groups had been removed, and dark
contrast was observed where the protecting groups were intact.
Tapping mode AFM images (Figure 5e-h) confirmed that
nonspecific adsorption onto the unexposed regions was negli-
gible. The NeutrAvidin particles were sufficiently well resolved
that they could be readily observed where they did adsorb on the
3
and 5 min. The contact angles of the virgin films following
exposure to the TFAA solution were very similar to those shown
in Table 1, confirming that the protecting groups remained
intact, despite the smaller-than-expected size of the nitro com-
ponent in the N 1s spectra. After 3 min exposure to UV light
(
1.12 J cm- ), the contact angle rose substantially, to 65-68°,
2
-
2
and after 5 min exposure (1.86 J cm ), the contact angle
measured was very similar to that recorded for a control sample,
formed by adsorption of APTES onto a silicon dioxide surface.
47
These data indicate that photodeprotection of the films was
extensive, in agreement with the data in Figure 3.
Samples were also exposed to UV light through an electron
microscope grid, as a simple micrometer-scale photomask, and
treated with a solution of aldehyde-functionalized polymer nano-
particles(42nmdiameter). The aldehydefunctional groupson the
surfaces of the nanoparticles react with the amine groups produced
by photodeprotection of EG NPEOC-APTES films to form imine
regions covered with intact EG NPEOC protecting groups, but
n
n
bonds, providing topographical contrast and enabling the deriva-
tization to be confirmed by AFM (Figure 4). Clear topographical
contrast was evident, with nanoparticles observed to be closely
packed across the exposed areas (squares) and not present on the
masked regions where the NPEOC protecting groups remained
intact. The height of the nanoparticle films on the exposed regions
only a very small number of isolated particles were detected. In
contrast, on the deprotected, biotinylated regions, dense attach-
ment was observed, such that it was difficult to distinguish
individual particles. The mean height across the square, exposed
regions was 40 nm for all four films, equal to the diameter of the
NeutrAvidin particles, confirming that a monolayer formed.
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Nanofabrication. Nanofabrication was carried out by SNP.
Light from the HeCd laser was coupled to a SNOM probe, and
the probe was traced across the sample to deprotect lines of
adsorbates. Before adsorbing NeutrAvidin particles, friction force
ness are not likely to account for the friction contrast in figure 6. The
OEG chains are lengthy, and likely to present multiple modes for the
dissipation of energy in the sliding tip-sample interaction through
tip-induced conformational change. This would be expected to raise
the coefficient of friction, relative to the deprotected amines terminal
groups, and probably accounts for the darker contrast (i.e., smaller
friction forces) measured in the exposed regions.
74,75
microscopy (FFM)
fabricated by SNP. FFM yields contrast due to changes in
was first used to characterize the features
76-78
79,80
molecular composition
and packing.
Figure 6 shows
FFM images of patterns fabricated by SNP, in which the exposed
regions exhibit darker contrast (i.e., a smaller friction force) than the
unmodified regions of surface. The contact angles measured for the
OEG-functionalized surfaces were very similar to values obtained for
deprotected amines, indicating that differences in surface adhesive-
The line width in Figure 6 was 180 nm. In the present case,
commercially fabricated SNOM probes were used, and SEM
characterization of the tips revealed that the width of the aperture
was also ca. 180 nm, suggesting that the resolution is determined
by the aperture dimensions. On the basis of previous work, we
would thus expect smaller line widths to be feasible if probes with
smaller apertures were used. Reducing the aperture diameter
would also diminish the throughput of the probes, but the
81
reduction in throughput would be nonlinear. The reduction in
throughput could be compensated by increasing the intensity of
the laserbeam coupled to the probe, or decreasing the writing rate,
8
2,83
or, potentially, by the use of probes with a larger cone angle.
Figure 7a and b shows patterns fabricated by SNP following
derivatization with NHS-biotin, and then incubated in a solution
of NeutrAvidin particles. To assist in relocating the nanopatterns
after the ex situ bioconjugation steps, a micrometer-scale ex-
posure was first carried out using an electron microscope “finder”
grid to create patterns of larger features. These were visible in the
optical microscope of the SNOM instrument, enabling the probe
to be located with good precision adjacent to a particular registry
Figure 6. Friction force microscopy images of patterns formed in films
of m-EG
7
NPEOC by SNP. Vertical scale range: 0.0-0.5 mV dark to
light (photodetector deflection).
Figure 7. (a and b) Fluorescence micrographs showing NeutrAvidin nanopatterns formed by SNP adjacent to larger registry features formed using a
mask-based process. Protecting groups: (a) p-EG NPEOC; (b) p-EG NPEOC. (c) AlexaFluor 647 conjugated streptavidin nanopattern formed in a film
4
7
of of p-EG NPEOC-APTES. (d and e) Cross sections through the nanopatterns shown in (a) and (b), respectively. (f) Higher magnification image of
7
the nanopattern formed in (b). Scale bars: 50 μm in (a)-(c) and 20 μm in (f).
2
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feature. After exposure of a region of intact adsorbates by SNP,
the sample was removed, incubated in solutions of NHS-biotin
followed by NeutrAvidin, and transferred to a confocal micro-
scope for characterization. The microscale patterns are also
readily located by confocal microscopy after avidin nanoparticle
labeling, providing registry features to enable the nanopattern to
be relocated easily.
Figure 7a-c shows low magnification images. The optical
intensity of the nanolines was relatively low, because they were
only ca. 4 NeutrAvidin particles wide, but they are nevertheless
clearly visible even at this magnification. Figure 7c shows a
pattern, formed by attachment of AlexaFluor 647-labeled strep-
tavidin, demonstrating that the patterning method is equally
efficacious for protein molecules, as opposed to protein-coated
nanoparticles. Line sections through fluorescence images of the
features yielded line widths somewhat larger than the widths
obtained by FFM directly following SNP, due to the resolution
limit of the confocal microscope (Figure 7d and e). Neverthe-
less, the line sections demonstrate that sharp, well-defined
features have been formed. The full width at half-maximum
nanopatterns. The fluorescently labeled nanopatterns also exhib-
ited excellent fluorescence contrast between exposed and unmo-
dified regions. Overall, these data suggest that simple strategies
based on the use of a PEGylated photocleavable protecting group
offer great potential for the control of biological organization at
micrometer and nanometer length scales. More generally, when
integrated with near-field exposure methods, they hold great
promise for the integration of synthetic chemical methodologies
with nanofabrication, and for the ultimate unification of the top-
down and bottom-up fabrication paradigms.
’ EXPERIMENTAL SECTION
Materials. The tetraethylene and heptaethylene glycol monome-
thyl ethers 1a and b were purchased from ABCR (Karlsruhe, Germany)
and TCI Europe (Zwijndrecht, Belgium) respectively. The 3-isocyanato-
propyltriethoxysilane, trifluoroacetic anhydride (>99%), phosphate
buffered saline (PBS), and (þ)-biotin N-succinimidyl ester (>98%
TLC) were supplied by Sigma-Aldrich (Poole, UK). Both methylnitro-
phenol isomers and all other reagents were purchased from Fisher
Scientific (Loughborough, UK). The DMSO was dried over 3 Å mole-
cular sieves prior to use. HPLC grade absolute ethanol was used as
(fwhm) was 340 nm in Figure 7d,e. The lenses used had
numerical apertures of 1.3 and 1.4, meaning that the diffrac-
tion-limited resolution was ca. 229 and 213 nm, respectively.
Given that the minimum width of the lines in the fluorescence
image would be approximately equal to the actual line width plus
the width of the point spread function, a fwhm of 340 nm is
consistent with a protein line no larger than the features shown in
Figure 6. AFM images did not enable an accurate determination
of the feature size because of the crystallization of salt from the
buffer solution onto the surface.
Strong fluorescence contrast was observed between the lines
and the unexposed material on either side. The fluorescence
signal does not reach zero anywhere in the image because of a low
level of background intensity. However, given the strong signal in
Figure 7f from the nanolines, which are only a few particles wide,
one might expect any nonspecifically bound particles to be clearly
evident. The absence of such features, apart from a very small
number of isolated spots likely resulting from adventitious
deposition, suggests that the NPEOC-protected film efficiently
resists adsorption.
3
received. The Et N was dried over 4 Å molecular sieves for a minimum
of 72 h prior to use. Tetrahydrofuran (THF), dimethylformamide
(DMF), and toluene were supplied by a Grubbs dry solvent system.
Aldehyde-coated 42 nm diameter nanoparticles (white aldehyde/sulfate
polystyrene latex, 4% solids in suspension), NeutrAvidin conjugated
yellow-green fluorescent (505/515) nanoparticles (1% solids) and
AlexaFluor647-conjugated streptavidin were obtained from Invitrogen
Molecular Probes (Paisley, UK). Immersol 518 F fluorescence free
immersion oil and Citifluor glycerol/PBS solution mounting media were
purchased from Carl Zeiss and Citifluor Ltd., respectively. Silicon (100)
wafers were purchased from Goodfellow (Cambridge, UK).
Synthesis of Photolabile Silanes. Tosylation of both the
glycols 1a and b was effected with tosyl chloride according to previously
6
8,69
described procedures.
-Nitro-4-(2,5,8,11-tetraoxatridec-22-yloxy)toluene, 2a. 4-Methyl-
2
3
4
-nitrophenol (689 mg, 4.5 mmol) and tosylate derivative of 1a (1.5 g,
.14 mmol) were dissolved in acetone (8 mL) and finely powdered
K CO (691 mg, 5 mmol) added. The suspension was heated under
2
3
reflux at ca. 70 °C for 6 h, after which water (20 mL) was added and the
mixture was extracted with Et O (15 mL twice). The organic layer was
2
evaporated under reduced pressure, and the residue was purified by flash
column chromatography (Hexanes:EtOAc, 1:1 f 1:4) to yield the
’
CONCLUSIONS
desired 2a as a yellow oil (1.29 g, 3.77 mmol, 91%); R
f
0.31 (Hex:EtOAc,
We have synthesized four photosensitive silanes possessing
-
1
2 2
1:3); νmax(BaF )/cm 2875 (alkyl), 1529 (NO ), 1349 (alkyl), 1249
tetra- and hepta-ethylene glycol adducts at the m- or p- positions
relative to the NPEOC group. In all cases, the nitrophenylethox-
ycarbonyl groups exhibit excellent sensitivity to irradiation at 325
(
(
CH
alkyl), 1107 (ether); δ (400 MHz, CDCl ) 2.53 (3H, s, CH Ar), 3.38
H
3
3
3H, s, CH
CH
-Ar), 7.23 (1H, d, J 8.5, 6-Ar), 7.54 (1H, d, J 2.5, 3-Ar); δ
3
O), 3.51-3.78 (12H, m, CH
2
OCH
2
), 3.88 (2H, t, J 4.8,
-
2
2
2
OAr), 4.17 (2H, t, J 4.8, CH
2
OAr), 7.10 (1H, d,d, J 2.5 and 8.5,
(100 MHz,
nm, being extensively deprotected at an exposure of 2.24 J cm
5
C
to give an amine-functionalized surface, as revealed both by XPS
measurements and by contact angle data for samples derivatized
using TFAA. The XPS data show that for macroscopic UV
exposures, all four adsorbates yield similar rates of photodepro-
tection. The potential use of this simple photoprotecting group
strategy for carrying out surface modifications was demonstrated
through the observation of highly efficient reactions with trifluor-
oacetic anhydride, with aldehyde-functionalized polymer nano-
particles, and with NHS-biotin, which facilitated the subsequent
attachment of NeutrAvidin particles. All four adsorbates exhibited
excellent resistance to nonspecific adsorption, and micropatterns
yielded sharp fluorescence images with high contrast between
masked and exposed regions. Near-field exposure also yielded
photodeprotection of the adsorbates and facilitated fabrication of
CDCl ) 19.7 (ArCH ), 59.0 (CH O), 68.1, 69.5, 70.5, 70.6, 70.6, 70.9,
3
3
3
2 2
71.9 (all CH OCH ), 110.0 (3-Ar), 120.5 (5-Ar), 125.7 (1-Ar), 133.4
(
6-Ar), 149.34 (2-Ar), 157.3 (4-Ar); m/z (ESþ) 344 (100%, [M þ
þ þ þ
H] ), 366 (60%, [M þ Na] ); HRMS found 344.1723, [M þ H]
requires 344.1709, δ 4.1 ppm.
2-Nitro-5-(2,5,8,11-tetraoxatridec-22-yloxy)toluene, 2b. This com-
pound was prepared using the procedure identical to that of 2a except
using 3-methyl-4-nitrophenol to yield 2b as a brown oil (1257 mg,
-1
3
.66 mmol, 88%); R 0.25 (Hex:EtOAc, 1:3); ν (BaF )/cm 2877
f max 2
(alkyl), 1580 (aryl), 1511 (nitro), 1336 (alkyl), 1295 (alkyl), 1258
(alkyl), 1107 (ether); δH (400 MHz, CDCl ) 2.63 (3H, s, CH Ar),
3
3
3.38 (3H, s, CH O), 3.51-3.79 (12H, m, CH OCH ), 3.88 (2H, t,
3
2
2
J 4.5, OCH CH OAr), 4.20, 2H, t, J 4.5, CH OAr), 6.80-6.85 (2H, m, 4
2
2
2
C 3
and 6-Ar), 8.08 (2H, d, J 8.6, 3-Ar); δ (100 MHz, CDCl ) 21.6
2
755
dx.doi.org/10.1021/ja1103662 |J. Am. Chem. Soc. 2011, 133, 2749–2759

Journal of the American Chemical Society
ARTICLE
t
(
CH
3
Ar), 59.0 (CH
3
O), 68.0, 69.4, 70.5, 70.6, 70.6, 70.9, 71.9 (all
under an inert atmosphere. KO Bu (63 mg, 0.56 mmol) was added, and
the mixture was stirred under an inert atmosphere at 85 °C for 1 h. The
mixture was then neutralized with 0.05 M aqueous HCl (ca. 15 mL) and
extracted with EtOAc (10 mL thrice). The organic layer was evaporated
under reduced pressure and purified by flash column chromatography
(EtOAc:EtOH, 1:0 f 4:1) to give the desired product as a brown oil
CH OCH
1
2
2
), 112.3 (4-Ar), 118.1 (6-Ar), 127.5 (3-Ar), 137.0 (1-Ar),
þ
42.3 (2-Ar), 162.3 (5-Ar); m/z (ESþ) 344 (100%, [M þ H] ), 366
þ
þ
(
3
30%, [M þ Na] ); HRMS found 344.1715, [M þ H] requires
44.1709, δ 1.7 ppm.
4-(2,5,8,11,14,17,20-Heptaoxadocosan-22-yloxy)-2-nitrotoluene, 2c.
4
1
K
-Methyl-3-nitrophenol (612 mg, 4.0 mmol) and tosylate 1b (865 mg,
.75 mmol) were dissolved in acetone (10 mL), and finely powdered
(465 mg, 1.24 mmol, 34%); R
f
0.33 (EtOAc:EtOH, 9:1); νmax(BaF
cm 3448 (hydroxy), 2879 (alkyl), 1514 (nitro), 1338 (alkyl), 1258
(alkyl), 1084 (ether); δ (400 MHz, CDCl ) 1.84 (1H, s(br), OH), 3.21
(2H, t, J 6.2, ArCH ), 3.38 (3H, s, CH O), 3.50-3.78 (12H, m,
CH OCH ), 3.88 (2H, t, J 4.5, OCH CH OAr), 3.95 (2H, td, J 6.0
and 5.5, CH OH), 4.21 (2H, t, J 4.5, CH OAr), 6.86 (1H, dd, J 9.0 and
2.5, 4-Ar), 6.91 (1H, d, J 2.5, 6-Ar), 8.05 (1H, d, J 9.0, 3-Ar); δ (100
MHz, CDCl ) 37.5 (ArCH ), 59.5 (CH O), 63.0 (CH OH), 68.5, 69.8,
70.9, 71.0, 71.3, 72.3 (all CH OCH ), 113.4 (4-Ar), 118.7 (6-Ar), 128.2
2
)/
-
1
2
CO
3
(691 mg, 5.0 mmol) was added. The suspension was refluxed at
H
3
ca. 70 °C for 16 h, after which the solids were removedbyfiltration and the
solution was evaporated under reduced pressure. The residue was purified
by flash column chromatography (MeOH:DCM, 1:39 f 1:29) to yield
2
3
2
2
2
2
2
2
the desired product as a pale yellow oil (737 mg, 1.55 mmol, 89%); R
f
0.29
Ar), 3.38
), 3.59-3.74 (22H, m,
OAr), 4.16 (2H, t, J 4.8,
OCH CH OAr), 7.09 (1H, d,d, J 2.6 and 8.5, 5-Ar), 7.22 (1H, d, J 8.5,
C
(
(
MeOH:DCM, 1:19); δ
3H, s, CH O), 3.55 (2H, t, J 3.0, CH
CH O), 3.87 (2H, t, J 4.9, OCH
H
(500 MHz, CDCl
OCH
CH
3
) 2.52 (3H, s, CH
3
3
2
3
2
3
3
2
CH
2
2
2
OCH
2
2
2
2
(3-Ar), 137.6 (1-Ar), 142.9 (2-Ar), 162.7 (5-Ar); m/z (ESþ) 374 (10%,
þ þ
[M þ H] ), 396 (100%, [M þ Na] ); HRMS found 396.1649, [M þ
2
2
þ
6
5
1
1
-Ar), 7.53 (1H, d, J 2.6, 3-Ar); δ (125 MHz, CDCl ) 20.2 (ArCH ),
9.4 (CH O), 68.4, 69.9, 70.9, 70.9, 71.0, 71.0, 71.3, 72.9 (all CH OCH ),
10.3 (3-Ar), 120.9 (5-Ar), 126.1 (1-Ar), 133.8 (6-Ar), 149.6 (2-Ar),
57.7 (4-Ar); m/z (ESþ) 476 (20%, [M þ H] ), 494 (100%, [M þ
H] requires 396.1634, δ 3.8 ppm.
C
3
3
2-[4-(2,5,8,11,14,17,20-Heptaoxadocosan-22-yloxy)-2-nitrophenyl]-
3
2
2
ethanol, 3c. The nitrotoluene 2c (346 mg, 0.68 mmol) was mixed with
paraformaldehyde (21 mg, 0.68 mmol) under an inert atmosphere, and
DMSO (anhydrous, 2 mL) was added. KO Bu (7 mg, 0.06 mmol) was
added and then stirred under an inert atmosphere at 80 °C for 30 min.
The dark brown mixture was then quenched by the addition of 0.1 M
aqueous HCl (20 mL) and extracted with DCM four times (10 mL
2 4
each). The organic fractions were combined, dried with Na SO , and
evaporated under reduced pressure. The residual oil was then purified by
þ
t
þ
þ
þ
NH
4
] ), 498 (15%, [M þ Na] ); found 476.2471, [M þ H] requires
4
76.2471, δ 0 ppm.
-(2,5,8,11,14,17,20-Heptaoxadocosan-22-yloxy)-2-nitrotoluene, 2d.
-Methyl-4-nitrophenol (612 mg, 4.0 mmol) and tosylate derivative of 1b
1.5 g, 3.03 mmol) were dissolved in DCM (10 mL), and finely powdered
CO (553 mg, 4.0 mmol) was added. The mixture was heated under
5
3
(
K
2
3
flash column chromatography (MeOH:DCM, 1:49 f 1:9) to give the
desired product as a yellow liquid (86 mg, 0.17 mmol, 25%); R 0.10
reflux at ca. 50 °C for 16 h, after which the solids were removed by
filtration and the solution evaporated under reduced pressure. The
residual liquid was purified by flash column chromatography (MeOH:
DCM, 49:1 f 1:24) to yield the desired product as a bright yellow oil
f
(
MeOH:DCM, 1:19); δ
ArCH ), 3.35 (3H, s, CH
.58-3.75 (22H, m, CH OCH
and CH OH), 4.15 (2H, t, J 4.4, CH
-Ar), 7.29 (1H, d, J 8.8, 6-Ar), 7.45 (1H, d, J 2.8, 3-Ar); δ (100 MHz,
H
(400 MHz, CDCl
O), 3.50-3.57 (2H, m, CH
), 3.80-3.92 (4H, m, OCH CH
OAr), 7.11 (1H, dd, J 2.4 and 8.4,
3
) 3.07 (2H, t, J 6.4,
OCH ),
OAr
2
3
2
2
3
2
2
2
2
(
1.21 g, 2.55 mmol, 84%); R
CDCl ) 2.62 (3H, s, CH Ar), 3.37 (3H, s, CH
OCH CH O), 3.87 (2H, t, J 4.6, OCH CH
OCH CH OAr), 6.78-6.83 (2H, m, 4,6-Ar), 8.07 (1H, d, J 9.6, 3-Ar); δ
C
f
0.22 (MeOH: DCM, 1:19); δ
O), 3.50-3.76 (24H, m,
OAr), 4.18 (2H, t, J 4.6,
H
(400 MHz,
2
2
3
3
3
5
C
2
2
2
2
CDCl ) 35.5 (ArCH ), 59.1 (CH O), 62.7 (CH OH), 68.1, 69.5, 70.5,
3
2
3
2
2
2
7
0.6, 70.6, 70.9, 71.9 (all CH OCH ), 110.2 (3-Ar), 120.3 (5-Ar), 125.9
(100 MHz, CDCl ) 22.1 (ArCH ), 59.4 (CH O), 67.3, 69.8, 70.9, 70.9,
2 2
3
3
3
(
1-Ar), 133.6 (6-Ar), 149.9 (2-Ar), 157.7 (4-Ar); m/z (ESþ) 506 (35%,
71.0, 71.0, 71.3, 72.3 (all CH OCH ), 112.7 (4-Ar), 118.4 (6-Ar), 127.9
2 2
þ þ þ
[
M þ H] ), 523 (100%, [M þ NH
4
] ), 528 (40%, [M þ Na] );
(
3-Ar), 137.4 (1-Ar), 142.6 (2-Ar), 162.7 (5-Ar); m/z (ESþ) 476 (10%,
þ
þ þ þ
HRMS found 506.2578, [M þ H] requires 506.2601, δ 4.5 ppm.
[M þ H] ), 493 (100%, [M þ NH
4
] ), 498 (30%, [M þ Na] ); HRMS
found 476.2498, [M þ H] requires 476.2490, δ 1.6 ppm.
-[2-Nitro-4-(2,5,8,11-tetraoxatridec-22-yloxy)phenyl]ethanol, 3a.
þ
2
-[5-(2,5,8,11,14,17,20-Heptaoxadocosan-22-yloxy)-2-nitrophenyl]-
ethanol, 3d. This compound was prepared using the same procedure as
c except using the pegylated nitrotoluene 2d. The product was isolated
as a yellow oil (103 mg, 0.20 mmol, 30%); R 0.14 (MeOH:DCM, 1:19);
2
3
The pegylated nitrotoluene 2a (1.72 g, 5.0 mmol) and paraformalde-
hyde (158 mg, 5.0 mmol) were suspended in DMSO (anhydrous, 6 mL)
under an inert atmosphere. KO Bu (84 mg, 0.75 mmol) was added, and
the mixture was stirred under an inert atmosphere at 85 °C for 1 h. The
mixture was then neutralized with 0.05 M aqueous HCl (ca. 15 mL) and
extracted with EtOAc (10 mL thrice). The organic layer was evaporated
under reduced pressure and purified by flash column chromatography
f
t
δ (400 MHz, CDCl ) 2.52 (1H, s(br), OH), 3.20 (2H, t, J 6.2, ArCH ),
H
3
2
3.37 (3H, s, CH
3
O), 3.51-3.78 (24H, m, CH
2
OCH
2
), 3.87 (2H, t, J 4.6,
OCH
CH OAr), 6.86 (1H, dd, J 2.8 and 8.8, 4-Ar), 6.92 (1H, d, J 2.8, 6-Ar),
8.04 (1H, d, J 8.8, 3-Ar); δ (100 MHz, CDCl ) 37.0 (ArCH ), 59.0
(CH O), 62.4 (CH OH), 68.0, 69.4, 70.4, 70.5, 70.6, 70.9, 71.9 (all
CH OCH
2 2 2
CH OAr), 3.95 (2H, t, J 6.2, CH OH), 4.21 (2H, t, J 4.6,
2
C
3
2
(
(
EtOAc:EtOH, 1:0 f 4:1) to give the desired product as a yellow oil
654 mg, 1.75 mmol, 35%); R 0.39 (EtOAc:EtOH, 9:1); ν (BaF )/
3
2
), 113.0 (4-Ar), 118.3 (6-Ar), 127.8 (3-Ar), 137.3 (1-Ar),
f
max
2
2
2
-
1
þ
cm 3436 (hydroxy), 2877 (alkyl), 1530 (nitro), 1350 (alkyl), 1251
142.5 (2-Ar), 162.3 (5-Ar); m/z (ESþ) 374 (10%, [M þ H] ), 396
þ
þ
(alkyl), 1106 (ether); δ
H
(400 MHz, CDCl
3
) 1.69 (1H, s(br), OH), 3.10
(100%, [M þ Na] ); HRMS found 528.2420, [M þ Na] requires
(2H, t, J 6.0, ArCH ), 3.38 (3H, s, CH O), 3.53-3.78 (12H, m,
528.2415, δ 0.9 ppm.
2
3
CH
(
2
OCH
2
), 3.80-3.99 (4H, m, OCH
2
CH
2
OAr and CH
2
OH), 4.17
N-{2-[2-Nitro-4-(2,5,8,11-tetraoxatridec-22-yloxy)phenyl]ethoxy-
carbonyl}-3-aminopropyltriethoxysilane, 4a. The pegylated nitrophenyl
alcohol 3a (167 mg, 0.43 mmol) was dissolved in DCM (2 mL), and to this
was added 3-isocyanatopropyltriethoxysilane (129 mg, 0.52 mmol) fol-
0
2
2H, t, J 4.7, CH OAr), 7.13 (1H, dd, J 2.5 and 8.6, 5 -Ar), 7.31 (1H, d,
0
0
J 8.6, 6 -Ar), 7.48 (1H, d, J 2.5, 3 -Ar); δ
C
(100 MHz, CDCl
3
) 35.9
(
(
ArCH ), 59.5 (CH O), 63.2 (CH OH), 68.5, 69.9, 70.9, 71.0, 71.3, 72.3
all CH
2
3
2
OCH
2
), 110.6 (3-Ar), 120.7 (5-Ar), 126.2 (1-Ar), 134.0 (6-Ar),
lowed by Et N (5.5 μL, 0.04 mmol). The solution was heated under reflux
3
2
þ
1
50.3 (2-Ar), 158.2 (4-Ar); m/z (ESþ) 374 (15%, [M þ H] ), 396
for 16 h and then evaporated under reduced pressure. The residual liquid
was purified by flash column chromatography (Hex:EtOAc, 1:5 f 1:19) to
þ
þ
(
100%, [M þ Na] ); HRMS found 396.1647, [M þ H] requires
96.1634, δ 3.3 ppm.
-[2-Nitro-5-(2,5,8,11-tetraoxatridec-22-yloxy)phenyl]ethanol, 3b.
3
yield the desired silane 4a as a yellow oil (169 mg, 0.27 mmol, 63%); R
f
0.18
3
-1 -1
2
(Hex:EtOAc, 1:5); λmax(EtOH)/nm 218 (ε/dm mol cm 16 974),
-1
The pegylated nitrotoluene 2b (1.28 g, 3.7 mmol) and paraformalde-
265 (3398), 330 (1995); νmax(BaF
)/cm 3344 (carbamate), 2883
2
hyde (113 mg, 3.7 mmol) were suspended in DMSO (anhydrous, 6 mL)
(alkyl), 1721 (carbamate), 1531 (nitro), 1249 (silane), 1103, 1077 (ether
2
756
dx.doi.org/10.1021/ja1103662 |J. Am. Chem. Soc. 2011, 133, 2749–2759

Journal of the American Chemical Society
ARTICLE
or siloxane); δ
H
(400 MHz, CDCl
3
) 0.60 (2H, t, J 8.5, CH
2
Si), 1.22 (9H, t,
formate (30 mg, 149 μmol). The mixture was stirred in the dark, and
after 24 h, all the alcohol was observed to be consumed by TLC together
with the formation of a new product corresponding to the 4-nitrophenyl
J 6.5, CH CH ), 1.52-1.65 (2H, m, CH CH CH ), 3.10-3.23 (4H, m,
2
3
2
2
2
2 2 3 2
NHCH and ArCH ), 3.38 (3H, s, CH O), 3.52-3.57 (12H, m, CH O),
3.81 (6H, q, J 7.0, SiOCH ), 3.87 (2H, t, J 4.5, CH CH OAr), 4.17 (2H, t,
f
carbonate (R 0.40; MeOH:DCM, 1:19). EtOH (9 μL) was added, and
2
2
2
J 4.5, CH
1H, dd, J 8.0 and 1.7, 5-Ar), 7.27 (1H, d, J 8.0, 6-Ar), 7.46 (1H, d, J 1.7,
-Ar); δC (100 MHz, CDCl ) 8.0 (CH Si), 18.7 (CH CH ), 23.7
2
OAr), 4.27 (2H, t, J 4.5, CH
2
OCO), 4.87 (1H, s(br), NH), 7.11
the mixture was stirred for 10 min to quench the excess chloroformate.
3-Aminopropyltriethoxysilane (24 μL, 115 μmol) was added, and the
reaction was observed to be complete by TLC after a further 24 h of
stirring. The mixture was evaporated under reduced pressure, and the
oily residue was purified by flash column chromatography (MeOH:
DCM, 1:49 f 1:29) to yield the desired product (10 mg, 0.013 mmol,
(
3
3
2
2
3
(
CH
CH
1.0, 71.3, 72.3 (all OCH ), 110.6 (3-Ar), 120.6 (5-Ar), 125.6 (1-Ar), 133.7
2
CH
2
CH
2
), 32.4 (ArCH
2
), 43.8 (NHCH
2
), 58.9 (SiOCH
2
), 59.5
(
3
O), 64.6 (CH
2
OCO), 68.5 (CH
2
OAr), 69.9 (CH
2
CH OAr), 70.9,
2
7
2
(
6-Ar), 150.6 (2-Ar), 156.6 (CO), 158.3 (4-Ar); m/z (ESþ) 356 (100%,
30%); R
J 8.1, CH
CH CH CH
ArCH ), 3.34 (3H, s, CH
3.53-3.72 (22H, m, CH
(2H, t, J 4.7, CH CH OAr), 4.16 (2H, t, J 4.7, CH
J 6.5, CH OCO), 4.92 (1H, t(br), J 5.6, NH), 6.78-6.85 (2H, m, 4- and
6-Ar), 7.98 (1H, d, J 8.7, 3-Ar); δ (100 MHz, CDCl ) 7.6 (CH Si), 18.3
(CH CH ), 23.3 (CH CH CH ), 33.5 (ArCH ), 43.4 (NHCH ), 58.4
(SiOCH ), 59.0 (CH O), 63.9 (CH OCO), 68.1 (CH OAr), 69.4
(CH CH OAr), 70.5, 70.6, 70.6, 70.6, 70.9, 71.9 (all OCH ), 113.0
f
0.33 (MeOH:DCM, 1:19); δ
Si), 1.81 (9H, t, J 7.0, CH CH
), 3.11 (2H, dt, J 6.5 and 6.7, NHCH
O), 3.51 (2H, t, J 4.6, CH
OCH ), 3.77 (6H, q, J 7.0, SiOCH
OAr), 4.28 (2H, t,
H
(400 MHz, CDCl
), 1.61 (2H, tt, J 7.4 and 7.8,
), 3.22 (2H, t, J 6.5,
OCH CH O),
), 3.84
3
) 0.62 (2H, t,
þ
þ
[
[
M - H
2
O þ H] ), 643 (55%, [M þ Na] ); HRMS found 643.2897,
2
2
3
þ
M þ Na] requires 643.2874, δ 3.6 ppm.
2
2
2
2
N-{2-[2-Nitro-5-(2,5,8,11-tetraoxatridec-22-yloxy)phenyl]ethoxy-
2
3
3
2
2
carbonyl}-3-aminopropyltriethoxysilane, 4b. The alcohol 3b (251 mg,
.67 mmol) was dissolved in DCM (2 mL), and 3-isocyanatopropyl-
triethoxysilane (193 mg, 0.78 mmol) and Et N (5.5 μL, 0.04 mmol) were
2
2
2
0
2
2
2
3
2
added in succession. This mixture was heated under reflux for 16 h, the
solvent evaporatedunder reducedpressure, and the residual oil purifiedby
flash column chromatography (Hex:EtOAc, 1:5 f 1:19) to give the
desired 4b as an orange oil (259 mg, 0.42 mmol, 62%); R
EtOAc, 1:5); λ (EtOH)/nm 232 (ε/dm mol cm 7510), 303
C
3
2
2
3
2
2
2
2
2
2
3
2
2
f
0.15 (Hex:
2
2
2
3
-1 -1
(4-Ar), 118.1 (6-Ar), 127.6 (3-Ar), 136.4 (1-Ar), 142.7 (2-Ar), 156.3
max
-1
þ
(8258); νmax/cm 3340 (carbamate), 2885 (alkyl), 1722 (carbamate),
(CO), 162.2 (5-Ar); m/z (ESþ) 753 (10%, [M þ H] ), 770 (100%,
þ
þ
1
516 (nitro), 1259 (silane), 1104, 1082 (ether or siloxane); δH (400
[M þ NH
4
] ), 775 (10%, [M þ Na] ); HRMS found 775.3658, [M þ
þ
MHz, CDCl
1
3
CH
3
) 0.63 (2H, t, J 8.0, CH
2
Si), 1.23 (9H, t, J 7.0, CH
2
CH
3
),
),
Na] requires 775.3655, δ 0.4 ppm.
.55-1.65 (2H, m, CH CH CH ), 3.16 (2H, td, J 6.5 and 6.5, NHCH
2
2
2
2
Film Preparation, Patterning, and Functionalization. Solu-
tions (1 mM) of the isomers of NPEOC-APTES were prepared in dry
toluene and chemisorbed on clean silicon oxide surfaces using previously
.27 (2H, t, J 6.3, ArCH ), 3.38 (3H, s, CH O), 3.53-3.77 (12H, m,
2
3
2 2 2 2
O), 3.82 (6H, q, J 7.0, SiOCH ), 3.88 (2H, t, J 4.5, CH CH OAr),
4
7
4
.20 (2H, t, J 4.5, CH
s(br), NH), 6.81-6.92 (2H, m, 4- and 6-Ar), 8.02 (1H, d, J 9.0, 3-Ar); δC
100 MHz, CDCl ) 8.0 (CH Si), 18.7 (CH CH ), 23.7 (CH CH CH ),
3.9 (ArCH ), 43.8 (NHCH ), 58.9 (SiOCH ), 59.5 (CH O), 64.4
2 2
OAr), 4.32 (2H, t, J 4.5, CH OCO), 4.92 (1H,
published procedures. Photodeprotection and patterning were con-
ducted using a He-Cd laser (IK3202R-D, Kimmon, Tokyo, Japan),
emitting at a wavelength of 325 nm. The samples of approximately
(
3
3
2
2
3
2
2
2
2
1.0 cm were irradiated at a power of 11 mW with an area of irradiation of
2
2
2
3
2
2
(
7
1
H
CH OCO), 68.5 (CH OAr), 69.8 (CH CH OAr), 70.9, 71.0, 71.3,
1.77 cm . Masks consisting of 12.7 ꢀ 12.7 and 16.9 ꢀ 16.9 μm square
features were used for micrometer scale patterning. For SNP, the laser
was coupled to an AlphaSNOM scanning near-field optical microscope
2
2
2
2
2
2.3 (all OCH ), 113.3 (4-Ar), 118.5 (6-Ar), 128.0 (3-Ar), 136.8 (1-Ar),
45.6 (2-Ar), 156.6 (CO), 162.6 (5-Ar); m/z (ESþ) 356 (100%, [M -
þ
þ
O þ H] ), 643 (50%, [M þ Na] ); HRMS found 643.2897, [M þ
(WiTec, Ulm, Germany). WiTec AlphaSNOM cantilever-type probes
2
þ
-1
Na] requires 643.2874, δ 3.6 ppm.
N-{2-[2-Nitro-4-(2,5,8,11,14,17,20-heptaoxadocosan-22-yloxy)phenyl]-
were used (WiTec, Ulm, Germany). The writing rate was 1 μm s ,
meaning that it took ca. 50 s to write each of the lines in Figure 7.
ethoxycarbonyl}-3-aminopropyltriethoxysilane, 4c. The alcohol 3c
After UV exposure, the reaction of the surface amine groups and
TFAA was carried out in the presence of Et N. The samples were
immersed in a 20 mM solution of the reagent in anhydrous tetra-
hydrofuran, with 40 mM triethylamine in Schlenck tubes under a dry N
(
36 mg, 71 μmol) was dissolved in DCM (anhydrous, 3 mL), and
-isocyanatopropyltriethoxysilane (46 μL, 184 μmol) was added, fol-
lowed by Et N (2 μL, 14 μmol). The mixture was heated under reflux
3
3
3
2
(
ca. 60 °C) for 16 h. The mixture was evaporated under reduced
pressure, and the residue was purified by flash column chromatography
MeOH:DCM, 1:49 f 1:19) to yield the title product as a pale yellow
oil (35 mg, 46.5 μmol, 65%); R 0.29 (MeOH:DCM, 1:19); δ (400 MHz,
atmosphere. Following the reaction, the derivatized samples were rinsed
and sonicated in tetrahydrofuran for 10 min to remove any unreacted
(
reagents. Following cleaning, the samples were blown dry with N
to further analysis.
2
prior
f
H
CDCl ) 0.60 (2H, t, J 8.2, CH Si), 1.20 (9H, t, J 7.0, CH CH ), 1.59
Derivatization with aldehyde-functionalized nanoparticles was car-
ried out by coating the micropatterned surface in a suspension of
nanoparticles for 30 min (30 μL). The samples were then washed with
ethanol and dried with nitrogen.
3
2
2
3
(
2H, tt, J 7.6 and 7.6, CH
2
CH
2
CH
2
), 3.04-3.22 (4H, m, NHCH
2
and
O),
ArCH
2
), 3.36 (3H, s, CH
3
O), 3.53 (2H, t, J 4.6, CH
3
OCH CH
2
2
3
.55-7.73 (22H, m, CH OCH ), 3.79 (6H, q, J 6.9, SiOCH ), 3.85
2
2
2
(
2H, t, J 4.6, CH
2
CH
2
OAr), 4.15 (2H, t, J 4.6, CH
2
OAr), 4.25 (2H, t,
Biotin-terminated patterns were created on the amine surfaces
using a solution consisting of biotin N-hydroxysuccinimide ester (10
2
J 6.4, CH OCO), 4.88 (1H, t(br), J 5.6, NH), 7.09 (1H, dd, J 2.8 and
8
(
-1
.8, 5-Ar), 7.24 (1H, d, J 8.0, 6-Ar), 7.43 (1H, d, J 2.4, 3-Ar); δ
100 MHz, CDCl ) 7.6 (CH Si), 18.3 (CH CH ), 23.3 (CH CH
C
mg mL ) in anhydrous dimethylformamide (DMF). The exposed
films were cleaned, placed in a Schlenk tube, and purged with N
3
2
2
3
2
2
-
2
.
CH ), 32.0 (ArCH ), 43.4 (NHCH ), 58.5 (SiOCH ), 59.1 (CH O),
These samples were immersed in the biotin solution for 2 h, then
2
2
2
2
3
6
7
4.2 (CH
0.6, 70.6, 70.9, 71.9 (all OCH
2
OCO), 68.1 (CH
2
OAr), 69.5 (CH
2
CH
2
OAr), 70.5, 70.6,
removed and rinsed twice with DMF followed by ethanol, and finally
2
), 110.2 (3-Ar), 120.2 (5-Ar), 125.2
dried with N gas.
2
(
(
1-Ar), 133.4 (6-Ar), 150.1 (2-Ar), 156.2 (CO), 157.8 (4-Ar); m/z
To attach NeutrAvidin beads to the biotinylted regions, NeutrAvidin
bead suspension (10 μL) was mixed with phosphate buffered saline
(PBS) solution (1 mL), and the patterned substrates were incubated for
2 h with the diluted suspension. For Alexa Fluor 647-conjugated
streptavidin attachment, the same procedure was used, but in the final
þ
þ
ESþ) 771 (100%, [M þ NH
4
] , 776 (20%, [M þ Na] ); HRMS
þ
found 775.3645, [M þ Na] requires 775.3655, δ 1.3 ppm.
N-{2-[2-Nitro-4-(2,5,8,11,14,17,20-heptaoxadocosan-22-yloxy)-
phenyl]ethoxycarbonyl}-3-aminopropyltriethoxysilane, 4d. The alcohol
-
1
3
d (22 mg, 43 μmol) was dissolved in DCM (anhydrous, 100 μL), and
step a solution of the protein in PBS (10 μg mL ) was used in place of
Et N (21 μL, 100 μmol) was added followed by 4-nitrophenylchloro-
3
the NeutrAvidin suspension.
2
757
dx.doi.org/10.1021/ja1103662 |J. Am. Chem. Soc. 2011, 133, 2749–2759

Journal of the American Chemical Society
ARTICLE
Surface Characterization. Contact angle measurements were
made using a Rame-Hart model 100-00 water contact angle goniometer.
For each sample, measurements were obtained at five different points on
the surface of the samples, and the mean was calculated. The thicknesses
of the films were obtained using a Jobin-Yvon UVISEL ellipsometer
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5
incident angle of 70°, in the 200-850 nm spectral range. The Cauchy
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(
J. D.; Berenschot, E.; Hunter, C. N.; Huskens, J.; Subramaniam, V.; Otto,
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layer of native SiO and the NPEOC film.
2
Contact and tapping mode AFM imaging were performed using a
Digital Instruments Nanoscope IIIa Multimode atomic force micro-
scope (Veeco, Santa Barbara, CA). For contact mode imaging, silicon
nitride probes (Nanoprobes, Veeco) with average spring constants 0.06
(
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1
1
-
1
or 0.12 N m and nominal tip radii between 20 and 60 nm were used.
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(
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acquired at pass energies of 160 and 80 eV, respectively. The Casa XPS
software was used to analyze and curve-fit the spectra. The peaks were
corrected relative to the C 1s hydrocarbon signal at a binding energy of
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(
285.0 eV.
Fluorescence images were acquired with a LSM 510 Meta laser
scanning confocal microscope (Carl Zeiss, Welwyn Garden City, UK).
The samples were mounted in a glycerol/PBS-based anti fade solution
(
2
(
(
Citifluor AF1, Agar Scientific, UK) and observed with 40ꢀ and 63ꢀ oil
immersion objectives (numerical apertures of 1.30 and 1.40, respecti-
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(
(
2
(
’
ASSOCIATED CONTENT
(
Hinsberg, W. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 13555.
(23) Singh-Gasson, S.; Green, R. D.; Yue, Y.; Nelson, C.; Blattner, F.;
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S
Supporting Information. Full citation for ref 45. This
b
material is available free of charge via the Internet at http://pubs.acs.org.
(24) Sun, S.; Chong, K. S. L.; Leggett, G. J. J. Am. Chem. Soc. 2002,
1
24, 2414.
’
AUTHOR INFORMATION
(25) Sun, S.; Leggett, G. J. Nano Lett. 2004, 4, 1381.
Corresponding Author
graham.leggett@shef.ac.uk; jason.micklefield@manchester.ac.uk
(26) Montague, M.; Ducker, R. E.; Chong, K. S. L.; Manning, R. J.;
Rutten, F. J. M.; Davies, M. C.; Leggett, G. J. Langmuir 2007, 23, 7328.
(27) Sun, S.; Montague, M.; Critchley, K.; Chen, M.-S.; Dressick,
Present Addresses
Department of Chemistry, Faculty of Science, Universiti Putra
W. J.; Evans, S. D.; Leggett, G. J. Nano Lett. 2006, 6, 29.
(28) Sun, S.; Leggett, G. J. Nano Lett. 2007, 7, 3753.
#
Malaysia, 43400 UPM Serdang, Selangor, Malaysia.
(29) Tizazu, G.; Adawi, A.; Leggett, G. J.; Lidzey, D. G. Langmuir
2
009, 25, 10746.
30) Alang-Ahmad, S. A.; Hucknall, A.; Chilkoti, A.; Leggett, G. J.
Langmuir 2010, 26, 9937.
31) Hurley, C. R.; Ducker, R. E.; Leggett, G. J.; Ratner, B. D.
Langmuir 2010, 26, 10203.
32) Sun, S.; Mendes, P.; Critchley, K.; Diegoli, S.; Hanwell, M.;
Evans, S. D.; Leggett, G. J.; Preece, J. A.; Richardson, T. H. Nano Lett.
006, 6, 345.
33) Iqbal, P.; Sun, S.; Hanwell, M. D.; Attwood, D.; Leggett, G. J.;
Preece, J. A.; Richardson, T. H.; Tunnicliffe, D. J. Mater. Chem. 2008, 18,
016.
34) Haq, E. u.; Liu, Z.; Zhang, Y.; Ahmad, S. A. A.; Wong, L.-S.;
Author Contributions
These authors contributed equally.
(
(
’
ACKNOWLEDGMENT
(
We thank Research Councils UK (grant no. EP/C523857/1)
2
for financial support. S.A.A.A. thanks the Malaysian Government
for a research scholarship. L.S.W. gratefully acknowledges the
EPSRC for the award of an LSI Fellowship (EP/F042590/1).
(
2
(
’
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