U.S. patent application number 16/245376 was filed with the patent office on 2019-12-05 for method for the topographically-selective passivation of micro- and nanoscale devices.
The applicant listed for this patent is University of Rochester. Invention is credited to Mark A. Lifson, Benjamin L. Miller, Dhruba Jyoti Basu Roy.
Application Number | 20190369092 16/245376 |
Document ID | / |
Family ID | 52428011 |
Filed Date | 2019-12-05 |
United States Patent
Application |
20190369092 |
Kind Code |
A1 |
Miller; Benjamin L. ; et
al. |
December 5, 2019 |
METHOD FOR THE TOPOGRAPHICALLY-SELECTIVE PASSIVATION OF MICRO- AND
NANOSCALE DEVICES
Abstract
Disclosed is a method of preparing a biosensor that involves
providing a substrate including a surface having a topographical
pattern formed at one or more sites on or in the surface, coating
the substrate with a solution including hydrogel particles, wherein
the hydrogel particles self-assemble on the surface to mask the
surface except at the one or more sites, and binding one or more
capture molecules to the one or more sites to form the biosensor.
Systems that include the biosensor, as well as methods of using the
biosensor, are also disclosed.
Inventors: |
Miller; Benjamin L.;
(Penfield, NY) ; Lifson; Mark A.; (Rochester,
NY) ; Roy; Dhruba Jyoti Basu; (Rochester,
NY) |
|
Applicant: |
Name |
City |
State |
Country |
Type |
University of Rochester |
Rochester |
NY |
US |
|
|
Family ID: |
52428011 |
Appl. No.: |
16/245376 |
Filed: |
January 11, 2019 |
Related U.S. Patent Documents
|
|
|
|
|
|
Application
Number |
Filing Date |
Patent Number |
|
|
14452141 |
Aug 5, 2014 |
10215753 |
|
|
16245376 |
|
|
|
|
61862374 |
Aug 5, 2013 |
|
|
|
Current U.S.
Class: |
1/1 |
Current CPC
Class: |
G01N 2021/757 20130101;
G01N 2021/7776 20130101; G01N 33/54373 20130101; G01N 2021/7783
20130101; G01N 2021/7789 20130101; G01N 2021/7773 20130101; G01N
33/54393 20130101; G01N 21/77 20130101 |
International
Class: |
G01N 33/543 20060101
G01N033/543; G01N 21/77 20060101 G01N021/77 |
Goverment Interests
[0002] This invention was made with government support under 5R01
AI080770 awarded by the National Institutes of Health. The
government has certain rights in the invention.
Claims
1-12. (canceled)
13. A biosensor prepared according to a process comprising the
steps of providing a substrate comprising a surface having a
topographical pattern formed at one or more sites on or in the
surface; coating the substrate with a solution comprising hydrogel
particles, wherein the hydrogel particles self-assemble on the
surface to mask the surface except at the one or more sites; and
binding one or more capture molecules to the one or more sites to
form the biosensor.
14. A biosensor comprising: a substrate comprising a surface having
a topographical pattern formed at one or more sites on or in the
surface; and one or more capture molecules bound to the one or more
sites to form the biosensor, the one or more capture molecules
binding specifically to a target molecule under suitable
conditions, wherein at least 80 percent of one or more types of
capture molecules bound to the surface of the substrate are bound
at the one or more sites.
15. The biosensor according to claim 14, wherein at least 90
percent of one or more types of capture molecules bound to the
surface of the substrate are bound at the one or more sites.
16. The biosensor according to claim 14, wherein the biosensor
exhibits at least an order of magnitude improvement in the limit of
detection of the target molecule compared to a biosensor having the
one or more capture molecules indiscriminately bound across the
entire surface of the substrate.
17. The biosensor according to claim 14, wherein the substrate
comprises a 2D photonic crystal array, a ring resonator, a toroidal
microcavity, waveguide, a photonic bandgap fiber, a Bragg
reflector, a diffraction grating, a plasmonic waveguide, or a
nanoplasmonic pore.
18. The biosensor according to claim 14, wherein the substrate
comprises an inlet for coupling light into, onto, or across the
topographical pattern and an outlet for coupling light that passes
from, through, or past the topographical pattern.
19. A method of detecting the presence of a target molecule in a
sample comprising: providing a biosensor according to claim 14;
exposing a sample to the one or more sites on the biosensor
surface; and detecting a change in an optical property of the
biosensor surface at the one or more sites following said
exposing.
20. A system comprising: a biosensor according to claim 14; a light
source coupled to the biosensor to pass light into or across the
topographical pattern; a detector coupled to the biosensor to
detect light passing from or across the topographical pattern.
21. A method of quantifying the amount of a biological target
present in a sample comprising: providing a biosensor according to
claim 14; exposing a sample to the one or more sites on the
biosensor surface; and detecting a change in an optical property of
the biosensor surface at the one or more sites following said
exposing, wherein the amount of biological target is quantifiable
based on the extent of the change of the optical property.
22. A method of detecting the presence of a target molecule in a
sample comprising: providing a biosensor according to claim 13;
exposing a sample to the one or more sites on the biosensor
surface; and detecting a change in an optical property of the
biosensor surface at the one or more sites following said
exposing.
23. A system comprising: a biosensor according to claim 13; a light
source coupled to the biosensor to pass light into or across the
topographical pattern; a detector coupled to the biosensor to
detect light passing from or across the topographical pattern.
24. A method of quantifying the amount of a biological target
present in a sample comprising: providing a biosensor according to
claim 13; exposing a sample to the one or more sites on the
biosensor surface; and detecting a change in an optical property of
the biosensor surface at the one or more sites following said
exposing, wherein the amount of biological target is quantifiable
based on the extent of the change of the optical property.
Description
[0001] This application is a continuation of U.S. patent
application Ser. No. 14/452,141 filed Aug. 5, 2014, which claims
the benefit of U.S. Provisional Patent Application Ser. No.
61/862,374, filed Aug. 5, 2013, which is hereby incorporated by
reference in its entirety.
TECHNOLOGICAL FIELD
[0003] This application relates to a method for the
topographically-selective passivation of micro- and nanoscale
devices, the resulting devices formed thereby, and biological
sensors containing the same.
BACKGROUND
[0004] The detection of biomedically significant molecules with
high-sensitivity nanoscale optical sensors has been the focus of
major development efforts by many research groups worldwide (Fan et
al., Anal. Chim. Acta 620:8-26 (2008)). Novel structures resulting
from these efforts, including ring- and whispering-gallery
resonators (Chao et al., Appl. Phys. Lett. 83:1527-1529 (2003);
Armani et al., Science 317:783-787 (2007); Barrios et al., Opt.
Lett. 32:3080-3082 (2007)), waveguides (Heideman et al., Sens.
Actuators 10:209-217 (1993); Goddard et al., Analyst 119:583-588
(1994); Salamon et al., Biophys. J. 80:1557-1567 (2001)), and
photonic crystals (Vollmer et al., Appl. Phys. Lett. 80:4057-4059
(2002); Krioukov et al., Opt. Lett. 27:1504-1506 (2002)), operate
by resolving minute changes in refractive index that occur when a
target molecule or virus interacts with the device. While all of
these devices have remarkable theoretical sensitivities, their
observed limits of detection ("LoD") under real-world conditions
are often unsatisfactory (Fan et al., Anal. Chim. Acta 620:8-26
(2008); Sheehan et al., Nano Lett. 5:803-807 (2005).
[0005] The LoD of a biosensor is dependent not only on the
sensitivity of the transduction mechanism, but also on the
biomolecular thermodynamics of the immobilized probe and the target
analyte in solution (Lambeck, Meas. Sci. Technol. 17:R93-R116
(2006); Kusnezow et al., Mol. Cell. Proteomics 5:1681-1696 (2006)).
In addition to presenting unique challenges for analyte mass
transport, nanoscale sensors require careful functionalization with
capture molecules (for example, antibodies) since the active
sensing region is orders of magnitude smaller than the overall
device. If the placement of capture molecules (probes) onto the
surface is indiscriminate and both the sensing and non-sensing
regions are functionalized (Sapsford et al., Anal. Chem.
73:5518-5524 (2001); Choi et al., Anal. Biochem. 405:1-10 (2010)),
the target loss to the non-sensing regions may become substantial
enough to disturb the bulk concentration of target. This can lead
to a lower fraction of material being bound to the sensing area,
and a higher (worse) LoD (Ekins et al., Clin. Chem. 37:1955-1967
(1991); Ekins, Clin. Chem. 44:2015-2030 (1998); Parpia et al.,
Anal. Biochem. 401:1-(2010)). Conventional passivation techniques
(Taylor et al., Nucleic Acids Res. 31:e87 (2003)) involving
incubation with proteins (e.g. bovine serum albumin) or synthetic
blocking chemicals cannot be used to avoid this issue, since they
would result in equal application to the non-sensing and sensing
areas of nanoscale devices. A common top-down approach to this
problem has been to shrink the size of the probe droplet in
manufacturing to closely overlay only the active sensing region
(McKendry et al., Proc. Natl. Acad. Sci. U.S.A 99:9783-9788 (2002);
Lee et al., Nano Lett. 4:1869-1872 (2004)). However, there are
considerable challenges with alignment and uniform dispensing on
such a small scale. Others have exploited material differences
within a nanoscale biosensor. For example, Fuez et al. showed
material-selective surface chemistry that selectively bound a
blocking agent to inactive titanium dioxide surfaces of a plasmonic
nanostructure leaving the gold sensing region to bind biomolecules
(Feuz et al., ACS Nano 4:2167-2177 (2010)). Since incorporation of
different materials into the device is not always feasible,
alternative strategies are clearly needed.
[0006] The devices and methods disclosed herein are directed to
overcoming these and other deficiencies in the art.
SUMMARY
[0007] A first aspect relates to a method of preparing a biosensor
that includes providing a substrate including a surface having a
topographical pattern formed at one or more sites on or in the
surface, coating the substrate with a solution comprising hydrogel
particles, wherein the hydrogel particles self-assemble on the
surface to mask the surface except at the one or more sites, and
binding one or more capture molecules to the one or more sites to
form the biosensor.
[0008] A second aspect relates to a biosensor prepared according to
the method described above.
[0009] A third aspect relates to a biosensor that includes a
substrate including a surface having a topographical pattern formed
at one or more sites on or in the surface, and one or more capture
molecules bound to the one or more sites to form the biosensor, the
one or more capture molecules binding specifically to a target
molecule under suitable conditions, wherein at least 80 percent of
one or more types of capture molecules bound to the surface of the
substrate are bound at the one or more sites.
[0010] A fourth aspect relates to a method of detecting the
presence of a target molecule in a sample that includes providing a
biosensor as described above, exposing a sample to the one or more
sites on the biosensor surface, and detecting a change in an
optical property of the biosensor at the one or more sites
following the exposing.
[0011] A fifth aspect relates to a system that includes a biosensor
as described above, a light source coupled to the biosensor to pass
light into or across the topographical pattern, and a detector
coupled to the biosensor to detect light passing from or across the
topographical pattern.
[0012] A sixth aspect relates to a method of quantifying the amount
of a biological target present in a sample that includes providing
a biosensor as described above, exposing a sample to the one or
more sites on the biosensor surface, and detecting a change in an
optical property of biosensor surface at the one or more sites
following the exposing, wherein the amount of biological target is
quantifiable based on the extent of the change of the optical
property.
[0013] Nanoscale biosensors have remarkable theoretical
sensitivities, but often suffer from sub-optimal limits of
detection in practice. This is in part because the sensing area of
nanoscale sensors is orders of magnitude smaller than the total
device substrate. Current strategies to immobilize probes (capture
molecules) functionalize both the sensing and non-sensing regions,
leading to target depletion and diminished limits of detection. The
difference in topography between these regions on nanoscale
biosensors offers a way to selectively address only the sensing
area. A bottom-up, topographically selective approach employing
self-assembled hydrogel nanoparticles as a mask to preferentially
bind target to the active sensing region of a biosensor has been
developed using a photonic crystal (PhC) as a proof of concept.
This led to over one order of magnitude improvement in the limit of
detection for the device, in agreement with finite element
simulations. Since the sensing elements in many nanoscale sensors
are topographically distinct, as in the PhC biosensor, this
approach should be widely applicable.
BRIEF DESCRIPTION OF THE DRAWINGS
[0014] FIG. 1A illustrates a detection system that includes a
biosensor, light source, and detector which operate via
illumination and detection through an ambient medium (e.g., air or
aqueous). FIG. 1B illustrates a detection system that includes a
biosensor, light source, and detector which operate via light
coupled into and out of the biosensor via waveguides.
[0015] FIGS. 2A-B show geometry and meshing of a droplet with
"active" (sensing) and "inactive" (non-sensing) areas for finite
element simulations. The surface mesh in FIG. 2A is plotted on a
10.sup.-4 m scale, while the mesh in FIG. 2B is a cross-section of
the geometry volume shown on a scale of 0.5 mm.
[0016] FIG. 3A shows simulated dose-response curves at 100%
antibody localization ("ABL") (red solid line), 95% ABL (green
dot-dash line), and 0% ABL (blue dashed line). FIG. 3B highlights
the low-concentration regime of the dose response, plotted on a
log/log scale.
[0017] FIG. 4A is a scanning electron microscope ("SEM") image of a
2D PhC device (10 .mu.m.times.7 .mu.m) with an array of wells
(radius=111 nm) etched in a triangular lattice. A row of wells was
removed to create awl waveguide. A defect (radius=73 nm) (arrow)
disrupts the periodicity, giving rise to an absorption peak. FIG.
4B is a low-pass filtered transmission spectra for air (n.about.1)
and water (n.about.1.32) show a peak shift to longer wavelengths
upon hydration, demonstrating the RI sensitivity of the 2D PhC.
[0018] FIG. 5A shows fluorescence image taken at 10.times.
magnification (2 sec. exposure) of 0.1 .mu.m r-IgG incubated on a
protein reactive (glutaraldehyde) substrate which was partially
covered with a Poly(N-isopropyl acrylamide) ("PNIPAM") nanoparticle
("PNP") mask. FIG. 5B shows relative fluorescence intensities of
the PNP masked surface for different concentrations of r-IgG was
calculated by normalizing the ratio of the fluorescence intensities
of the masked to the unmasked regions. Error bars for each
condition are calculated from the standard deviations of the
fluorescence intensities. See experimental section for detailed
formulae.
[0019] FIGS. 6A-B show SEM images of HSQ (5A) and embedded PMMA
(6B) 2D PhCs dipcoated with PNPs. The SEM platform was tilted 20
degrees in order to optimize contrast between the particles and the
substrate. Scale bars represent 2 .mu.m.
[0020] FIGS. 7A-B show post-processed experimental peak shifts for
two representative 2D-PhC sensors (7A) which were incubated with
capture antibody and set to zero red-shift (black dashed line);
peak-shifts after incubation with 5 nm target without ABL (red
dotted line) and with ABL (solid blue line). FIG. 7B shows the
dose-response without ABL (red bar) and with ABL (blue bar); values
that were statistically significant (unpaired 2-tailed student
t-test, p-value<0.05) were marked with an asterisk (*). Error
bars were calculated by taking the square root of the sum of the
control standard deviation squared plus the experimental standard
deviation squared.
DETAILED DESCRIPTION
[0021] According to one aspect a method of preparing a biosensor is
disclosed herein. The method includes providing a substrate
including a surface having a topographical pattern formed at one or
more sites on or in the surface, coating the substrate with a
solution including hydrogel particles, wherein the hydrogel
particles self-assemble on the surface to mask the surface except
at the one or more sites (where the topographical pattern is
formed), and binding one or more capture molecules to the substrate
or a material forming the topographical pattern at the one or more
sites to form the biosensor.
[0022] It should be appreciated by those of ordinary skill in the
art that any of a variety of substrates can be employed in the
present invention. Preferably, the substrate has a topographical
pattern that comprises a plurality of pits, pores, or troughs
formed in the substrate, a raised structure formed on the
substrate, or any combination of two or more such features. In one
embodiment, the topographical pattern comprises a structural
feature formed on the substrate. In another embodiment, the
topographical pattern comprises a structural feature formed in the
substrate (i.e., below a surface of the substrate). It should also
be appreciated that the hydrogel particles assemble on the surface
of the substrate in the regions around the topographical
pattern.
[0023] Exemplary substrates include, but are not limited to, a 2D
photonic crystal array, a ring resonator, a toroidal microcavity,
waveguide, a photonic bandgap fiber, a Bragg reflector, a
diffraction grating, a plasmonic waveguide, glass-supported gold
disk pairs, or a nanoplasmonic pore.
[0024] Substrates can be formed using any of a variety of
materials. Exemplary materials include, without limitation, silicon
such as crystalline silicon, amorphous silicon, or single crystal
silicon, oxide glasses such as silicon dioxide, and polymers such
as polystyrene. The materials can also include metal
(nano)particles or coatings applied to one or more surfaces of the
substrates.
[0025] The 2D photonic crystal array may have any suitable
arrangement of pores formed in a substrate. One example of a 2D
photonic crystal array is described in U.S. Patent Application
Publication No. 2010/0279886 to Fauchet et al., the disclosure of
which is incorporated herein by reference in its entirety. Photonic
crystals (or crystal arrays) are an attractive sensing platform
because they provide strong light confinement. These crystals can
be designed to localize the electric field in the low refractive
index region (e.g., air pores), which makes the sensors extremely
sensitive to a small refractive index change produced by the
capture of a targeted bio-molecule on the pore walls. In certain
embodiments, the hydrogel particles are predominantly or completely
localized to the surface of the photonic crystal array at sites
other than where the pores reside. In this case, the hydrogel
particles effectively mask the outer surface of the photonic
crystal array, but leave the pore structures formed in the
substrate exposed and accessible for subsequent binding of capture
molecules to the interior pore surfaces.
[0026] The ring resonator may have any suitable arrangement of ring
features and working waveguide surfaces, including single or
multiple ring resonator constructions. One example of a ring
resonator detector is described in PCT Publication WO2013053459,
the disclosure of which is incorporated herein by reference in its
entirety. A substrate of this type is very sensitive as a surface
of the ring is scanned by an evanescent field of a light wave
propagating within the ring. Currently, ring resonators are used to
perform measurements with a selectively working absorber surface,
which is labeled with one or more capture molecules and therefore
plays an important role for an adequate specificity of the sensor.
The capture of a targeted bio-molecule at the working surface cause
an optical ring circumference to vary. Thus, an effective
refractive index of the ring resonator changes upon capture of the
targeted bio-molecule such that wavelengths of resonant modes are
shifted. The detection of the shift into a coupled detection
waveguide can indicate presence of the bio-molecule. When utilized
in the devices and methods described herein, the hydrogel particles
are used to mask regions of the ring resonator substrate other than
where the working absorber surface resides.
[0027] Ultrahigh-Q silica toroidal microcavities are particularly
attractive for use in applications of biomolecular sensing because
they can be fabricated on a chip. The toroidal microcavity can have
any desired configuration, e.g., ring, ellipsoidal, or polygonal
configurations. In one approach, an SiO.sub.2 disk cavity can be
fabricated on a silicon wafer by, e.g., thermal dioxidation,
photolithography, and SiO.sub.2 etching. The dioxide layer can be
on the micron or submicron level. Next, the silicon sacrificial
layer is undercut to form a Si post. With a combination of
isotropic and anisotropic etching, a silicon post can be obtained
and then the SiO.sub.2 is exposed with a laser suitable to transfer
the shape of the silicon post to the SiO.sub.2 and form a smooth
toroidal cavity of the desired configuration. As an alternative to
SiO.sub.2, other oxide glasses can be used to form the toroidal
microcavity. The toroidal microcavity may have any suitable
arrangement between the microcavity and working waveguide surfaces,
including single or multiple microcavity constructions. Toroidal
microcavities are useful to increase the distance between adjacent
resonance wavelengths. One suitable structure of the microcavity
sensor is illustrated in U.S. Application Publ. No. US20090097031
A1 to Armani et al., the disclosure of which is incorporated herein
by reference in its entirety. One example for use of toroidal
microcavities in a biosensor is described in U.S. Patent
Publication No. 20090093375 to Arnold et al., the disclosure of
which is incorporated herein by reference in its entirety. When
utilized in the devices and methods described herein, the hydrogel
particles are used to mask regions of the toroidal microcavity
substrate other than where the microcavity is formed. This can
facilitate coupling of the capture molecules to the microcavity
surface.
[0028] A waveguide is a structure which guides optical waves by
total internal reflection (TIR). When a light beam traveling in a
waveguide is totally internally reflected at the interface between
the waveguide and an adjacent medium having a lower refractive
index, a portion of the electromagnetic field of the TIR light
penetrates shallowly into the adjacent medium. The use of
waveguides in the design of biosensors has been described in
numerous publications including U.S. Pat. No. 5,814,565 to Reichert
et al., the disclosure of which is incorporated herein by reference
in its entirety. The waveguide can be fabricated on a substrate
surface, in which case the hydrogel particles are used to mask
regions of the substrate other than where the waveguide is
formed.
[0029] Alternatively, a waveguide can be formed within a recessed
region of the substrate so as to form trenches on either side of
the waveguide. With this configuration, hydrogel particles mask
regions on either side of the trenches leaving the waveguide
exposed.
[0030] Photonic bandgap structures allow light within certain
well-defined wavelength bands to be guided without a total internal
reflection mechanism. Photonic band gap structures are configured
so as to confine and guide light through resonant reflections, and
do not depend on total internal reflections. Accordingly, much
greater flexibility is allowed in the design and construction of
such structures.
[0031] Photonic band gap structures may be fabricated by machining
blocks of dielectric material, although other methods of
fabricating photonic band gap structures may involve the mechanical
drilling or machining of holes or cavities in solid blocks of a
dielectric material. Another method may involve the use of chemical
removal, such as reactive ion etching, to fabricate holes or
cavities in solid blocks of dielectric material. Alternatively,
photonic band gap structures may be fabricated by stacking a
collection of dielectric elements in a desired pattern. Where the
photonic band gap structure is formed in a block of solid material
by forming holes or cavities, the hydrogel particles are used to
mask regions of the substrate other than where the holes or
cavities are formed. Where the photonic band gap structure is
formed in a block of solid material by forming holes or cavities,
the hydrogel particles are used to mask regions of the substrate
other than where the holes or cavities are formed.
[0032] A Bragg reflector is a sensor element utilizing more than
one layer of materials with varying refractive indexes that result
in detection of a reflectivity shift having one or more sharply
defined luminescent peaks. A biosensor comprising a Bragg reflector
is described in U.S. Pat. No. 7,226,733 to Chan et al., the
disclosure of which is incorporated herein by reference in its
entirety. The periodicity and design of the upper and lower Bragg
reflectors can have any suitable configuration. When used with
macroporous or mesoporous Bragg structures, it is possible to
confine capture molecule location to the pores of the Bragg
structures. Confinement to the pores rather that the outer surface
of the Bragg structure can be achieved by masking the outer
surfaces with the hydrogel particles prior to capture molecule
coupling.
[0033] A diffraction grating operates at a fixed wavelength and
detection angle by exploiting the variation in diffraction
efficiency that occurs due to the presence of a chemical or
biological species on a diffraction grating. Any of a variety of
suitable diffraction grating structures (channel depth, width, and
spacing) can be employed. In traditional diffraction-based
biosensors, chemical or biological species are selectively adsorbed
onto the top surface of a diffraction grating, giving rise to an
increase in the diffraction efficiency proportional to the change
in the grating thickness. One exemplary diffraction grating based
sensor is described in U.S. Pat. No. 8,349,617 to Weiss et al., the
disclosure of which is incorporated herein by reference in its
entirety. In addition, surface-plasmon enhancement can be enhanced
by nanoparticles (Wark et al., "Nanoparticle-Enhanced Diffraction
Gratings for Ultrasensitive Surface Plasmon Biosensing," Anal.
Chem. 79:6697-6701 (2007), the disclosure of which is incorporated
herein by reference in its entirety). In the devices and methods
described herein, the hydrogel particles can be used to mask
regions of the device that lack diffraction grating structures.
[0034] A plasmonic waveguide involves excitations which do not
exhibit the disadvantages associated with using light sources to
determine a specific binding event. These surface plasmon
polaritons or plasmonic mode excitations, i.e., electromagnetic
excitations at a metal-dielectric interface, may be guided using
structures that are much smaller than the wavelength of photons of
the same frequency. Any of a variety of SPR-biosensor structures
can be utilized in forming a biosensor. These structures can be
provided with any of a variety of topographical structures on the
sensing surface. One exemplary plasmonic waveguide is described in
U.S. Pat. No. 6,373,577 to Brauer et al., the disclosure of which
is incorporated herein by reference in its entirety. Where the
SPR-based sensor structure includes grating patterns, or holes or
cavities in which capture molecules are intended to reside, the
hydrogel particles are used to mask the outer surface of the
substrate other than where the grating, holes or cavities are
formed. Where the SPR-based sensor structure includes raised
structures intended to contain the capture molecules, the hydrogel
particles are used to mask regions of the substrate other than
where the raised structures are formed.
[0035] Glass-supported gold nanostructure dimers include a pair of
the nanostructures supported on a glass substrate and separated
from one another by a nanogap on the order of about 1 to about 10
nm. The gold nanostructures include, without limitation, disks,
bowties, nanorods, and rings, which can have any suitable
dimension. Light passes through the nanogap, and polarization of
longitudinal and transverse polarizations are obtained. As the gold
dimers approach each other, an exponential red shift and slight
blue shift of coupled bonding mode is obtained for these
polarizations. Near-field coupling in gold dimers causes an
exponential increase in sensitivity to refractive index of
surrounding medium with decreasing the gap distance. Thus, capture
of the target molecule at the gap interface will induce a change in
the refractive index and a shift in the coupled bonding modes.
Exemplary gold dimers are described in Tsai et al., "Plasmonic
Coupling in Gold Nanoring Dimers: Observation of Coupled Bonding
Mode," Nano Lett. 12(3):1648-54 (2012); Tanaka et al., "Nanoscale
Interference Patterns of Gap-mode Multipolar Plasmonic Fields,"
Scientific Reports 2(764):doi:10.1038/srep00764 (2012); and Ye et
al., "Plasmonic Behaviors of Gold Dimers Perturbed by a Single
Nanoparticle in the Gap," Nanoscale 4(22):7205-11 (2012), the
disclosures of which are incorporated herein by reference in their
entirety. Because the dimer paired nanostructures are supported on
the surface of a substrate, hydrogel particles can be used to mask
regions of the substrate so as to confine capture molecule binding
to only the raised nanostructure pairs.
[0036] Nanoplasmonic pores have the advantage of exhibiting unique
optical transmission characteristics at resonant wavelengths. Any
sensor structure comprising nanoplasmonic pores can be used in the
present invention. The nanopores are formed in a submicron membrane
including a metal film (e.g., gold, silver, platinum). The
nanopores can be dimensioned to facilitate maximal response in
consideration of the target molecule, but typically the nanopores
are on the order of less than 250 nm, preferably less than 150 nm
in diameter. Capture molecules bound within the nanopore features
allow for specific binding of the target molecule within the
nanopore structures. By monitoring the temporal variation in the
plasmon resonance of the structure, flow-through nanoplasmonic
sensing of specific biorecognition events (i.e., detection of the
target molecule) can be achieved quickly in a low-volume flow
through device. Because hydrogel particles can be used to mask the
upper and lower surfaces of the membrane, it is possible to confine
capture molecule binding to within the nanopore structures.
Exemplary nanoplasmonic biosensors are disclosed in U.S. Patent
Publication No. 20120218550 to O'Mahony; and Jonsson et al.,
"Locally Functionalized Short-range Ordered Nanoplasmonic Pores for
Bioanalytical Sensing," Anal. Chem. 82(5):2087-94 (2010), the
disclosures of which are incorporated herein by reference in their
entirety.
[0037] The hydrogel particles can be solid hydrogel polymers or
they can be hybrid particles, e.g., a hydrogel coating that
surrounds a metal or polymer core (Kim and Lee, "Hydrogel-Coated
Gold Nanoparticles," Polymeric Materials: Science and Engineering
90: 637-638 (2004); Dingenouts et. al., "Observation of the Volume
Transition in Thermosensitive Core-Shell Latex Particles by
Small-Angle X-Ray Scattering," Macromolecules 31: 8912-8917 (1998),
the disclosures of which are incorporated herein by reference in
their entirety. Hydrogel particles can be formed as small units of
crosslinked monomers.
[0038] The hydrogel particle diameter selected for use in the
invention will depend, in part, on the nature of the topographical
features of the substrate and, thus, the nature of masking desired.
The hydrogel particles are preferably, though not necessarily,
submicron in diameter. In certain embodiments, the average size of
the hydrogel particles is between about 100 nm to about 900 nm, or
about 200 nm to about 800 nm in diameter.
[0039] The hydrogel particles can be formed of any suitable
hydrogel material. Exemplary hydrogel materials include, without
limitation, poly-N-isopropylacrylimide (PNIPAM), PNIPAM
copolymerized with allyl-iminodiacetic acid, PNIPAM grafted with
polyethylene glycol-succinic acid, hydroxypropyl cellulose, or a
pullulan acetate/sulfonamide conjugate. Solutions containing the
hydrogel particles can be formed using aqueous solutions, including
mild buffer solutions, and water. The hydrogel particles can be
present in the solution at any suitable concentration that allows
for hydrogel particle coverage of the substrate regions devoid of
the topographical features. By way of example, the concentration
can range from about 10.sup.8 to about 10.sup.13 particles per
milliliter, with a solids weight percentage between about 0.001% to
about 1%. This is exemplary, and deviations from these ranges are
contemplated. The optimum concentration for uniform coverage
depends on extrinsic and intrinsic factors such as the
characteristics of the substrate and its topographical features,
and the size and area covered by the droplet. For instance, when
using a droplet in the picoliter regime, a higher concentration of
particles is desirable (e.g., about 10.sup.11 to about 10'.sup.3
particles per milliliter).
[0040] After selecting the design and substrate for the biosensor
device, the structural features of the biosensor device are formed
at the one or more sites, and then the coating is applied to the
substrate. Coating of the hydrogel particles onto the substrate can
be performed using techniques well known in the art. Exemplary
coating techniques include, but are not limited to, spraying,
spotting, depositing, dip-coating, spin-coating, evaporative
lithography, and evaporative deposition of the solution onto the
substrate.
[0041] Once the hydrogel mask is applied to the substrate, capture
molecules can be applied to the substrate whereby the capture
molecules bind to or associate with unmasked regions of the
substrate, i.e., where the topographical pattern resides.
[0042] As used herein, a "capture molecule" is any molecule that is
capable of binding to an analyte (i.e. capturing it). Suitable
capture molecules include, without limitation, a protein or
polypeptide, a nucleic acid molecule, or an organic small molecule
probe. It is desirable that the capture molecule binds specifically
to the analyte of interest.
[0043] Exemplary small molecules include, without limitation:
avidin, peptido-mimetic compounds, and vancomycin. One class of
peptido-mimetic compounds is disclosed in U.S. patent application
Ser. No. 09/568,403 to Miller et al., filed May 10, 2000, the
disclosure of which is incorporated herein by reference in its
entirety. A preferred peptido-mimetic compound which binds to
lipopolysaccharide is a tetratryptophan ter-cyclopentane as
disclosed in the above-noted application to Miller et al. Other
peptidomimetic compounds can also be employed.
[0044] Exemplary polypeptides include, without limitation, a
receptor for cell surface molecule or fragment thereof; a lipid A
receptor; an antibody or fragment thereof; peptide monobodies of
the type disclosed in U.S. patent application Ser. No. 09/096,749
to Koide, filed Jun. 12, 1998, and U.S. patent application Ser. No.
10/006,760 to Koide, filed Nov. 19, 2001, the disclosures of which
are incorporated herein by reference in their entirety; a
lipopolysacchardide-binding polypeptide; a peptidoglycan-binding
polypeptide; a carbohydrate-binding polypeptide; a
phosphate-binding polypeptide; a nucleic acid-binding polypeptide;
and polypeptides which bind organic warfare agents such as tabun,
sarin, soman, GF, VX, mustard agents, botulinum toxin,
Staphylococcus entertoxin B, and saitotoxin.
[0045] Exemplary nucleic acid molecules can be DNA, RNA, or
modified nucleic acids that include 2' or 5'-modified sugars,
modified nucleotide bases, or peptide-nucleic acids. The nucleic
acids can be any length which is suitable to provide specificity
for the intended target. Typically, nucleic acids which do not
contain modified nucleotides will be at least about 12 to about 100
nucleotides in length. For nucleic acids which contain modified
bases, oligonucleotides should be at least about 7 nucleotides in
length, up to about 100 nucleotides in length. Nucleic acid capture
molecules can be used for Watson-Crick base-pairing with a
complementary or partially complementary target nucleic acid
molecule depending on the conditions employed (i.e., low
stringency, moderate stringency, or high stringency).
Alternatively, nucleic acid aptamer molecules can be used for
specific binding to other target molecules, typically proteins,
carbohydrates, lipids, etc.
[0046] The available strategies for attaching the one or more
capture molecules include, without limitation, covalently bonding a
capture molecule to the surface of the substrate, ionically
associating the capture molecule with the surface of the substrate,
adsorbing the capture molecule onto the surface of the substrate,
or the like. Such association can also include covalently or
noncovalently attaching the capture molecule to another moiety (of
a coupling agent), which in turn is covalently or non-covalently
attached to the surface of the substrate.
[0047] In one embodiment, prior to coating the substrate with the
hydrogel particles, the substrate surface is treated with a
reactant that promotes covalent binding of the one or more capture
molecules. This can be achieved by providing a coupling agent
precursor and then covalently or non-covalently binding the
coupling agent precursor to the surface of the substrate. Once the
substrate has been primed with the coupling agent, the capture
molecule is exposed to the primed surface under conditions
effective to covalently bind to the coupling agent. The binding of
the capture molecule to the substrate is carried out under
conditions which are effective to allow the one or more
target-binding groups thereon to remain available for binding to
the target molecule. Suitable coupling agent precursors include,
without limitation, silanes functionalized with an epoxide group, a
thiol, or an alkenyl.
[0048] In a further embodiment, any of the reactant that remains
unbound by a capture molecule following the binding is blocked. The
blocking agent can be structurally similar to the capture molecules
except that they lack a target-binding group or the blocking agents
can simply be simple end-capping agents. By way of example, an
amino alkyl ester (e.g., glycine methyl ester, glycine ethyl ester,
3-alanine methyl ester, etc.) blocking agent can be introduced to
an epoxide-functionalized substrate.
[0049] In one embodiment, the one or more capture molecules are
directly bound to the substrate surface.
[0050] In another embodiment, the one or more capture molecules are
attached to the substrate surface via a linker molecule, and the
one or more capture molecules and the linker molecule have an
affinity-based, non-covalent interaction. For example, the capture
molecule and the linker molecule can interact via a
streptavidin/biotin interaction or a Protein A/G-immunoglobulin
interaction.
[0051] Binding of the one or more capture molecules to the one or
more sites on the substrate will be carried out in accordance with
the selected capture molecules and approaches for attaching the
capture molecules to the substrate. In general, this is achieved by
exposing the one or more sites to a solution comprising a capture
molecule under conditions effective to allow the capture molecule
to bind to the site exposed to the solution. This exposure step can
be performed using methods well known in the art including, without
limitation, printing, spraying, spotting, or depositing the
solution onto the substrate or flowing the solution over the
substrate at the one or more sites.
[0052] After binding of the capture molecules, it is desirable to
wash any reactant from the substrate surface and/or blocking any
reactant that remains unbound by a capture molecule. This will
minimize the likelihood that the target molecule or macromolecular
structure can be bound non-specifically to the substrate.
[0053] In accordance with one embodiment, two or more sites and two
or more capture molecules may be present, with each capture
molecule being used at a different site. Thus, the different sites
can be prepared using different attachment chemistries (i.e.,
mutually exclusive) such that a single solution may be used to
attach different capture molecules are different sites.
Alternatively, the exposure steps can be carried in parallel at
distinct sites on the substrate.
[0054] In one embodiment, after attaching/binding of the one or
more capture molecules the hydrogel particles are removed from the
substrate surface. Removal is carried out before exposing the
biosensor to a sample for detection of a target molecule. Removal
of the hydrogel particles can be achieved by washing the surface of
the biosensor with distilled water or a buffer solution.
Additionally or alternatively, the hydrogels particles can be
removed with sonication. In certain embodiments, even after
removing hydrogel particles from the surface, it is desirable for
the reasons noted above to wash any reactant from the substrate
surface and/or block any reactant that remains unbound by a capture
molecule.
[0055] In an alternative embodiment, the hydrogel particles are
allowed to remain on the substrate surface. Removal is carried out
before exposing the biosensor to a sample for detection of a target
molecule.
[0056] Another aspect relates to a biosensor. The biosensor
includes a substrate including a surface having a topographical
pattern formed at one or more sites on or in the surface. The
substrate can have any of the constructions described above. One or
more capture molecules are bound to the one or more sites to form
the biosensor, and the one or more capture molecules bind
specifically to a target molecule under suitable conditions. The
biosensor is unique in that at least 80 percent of one or more
types of capture molecules bound to the surface of the substrate
are bound at the one or more sites. In other words, less than 20
percent of the one or more types of capture molecules is bound to a
region on the surface of the substrate where the capture molecules
are not intended.
[0057] In one embodiment, at least 85 percent of one or more types
of capture molecules bound to the surface of the substrate are
bound at the one or more sites. In other words, less than 15
percent of the one or more types of capture molecules is bound to a
region on the surface of the substrate where capture molecules are
not intended.
[0058] In another embodiment, at least 90 percent of one or more
types of capture molecules bound to the surface of the substrate
are bound at the one or more sites. In other words, less than 10
percent of the one or more types of capture molecules is bound to a
region on the surface of the substrate where capture molecules are
not intended.
[0059] In a further embodiment, at least 95 percent of one or more
types of capture molecules bound to the surface of the substrate
are bound at the one or more sites. In other words, less than 5
percent of the one or more types of capture molecules is bound to a
region on the surface of the substrate where capture molecules are
not intended.
[0060] In another embodiment, the biosensor exhibits at least an
order of magnitude improvement in the limit of detection of the
target molecule compared to a biosensor having the one or more
capture molecules indiscriminately bound across the entire surface
of the substrate.
[0061] In certain types of biosensors, the biosensor has an inlet
for coupling light into, onto, or across the topographical pattern
and an outlet for coupling light that passes from, through, or past
the topographical pattern.
[0062] The biosensor may also be present in a microfluidic device
and exposing the biosensor to the sample involves flowing the
sample over the one or more sites on the biosensor surface.
[0063] A further aspect relates to a system that includes a
biosensor of the type described above, a light source coupled to
the biosensor to pass light into or across the topographical
pattern, and a detector coupled to the biosensor to detect light
passing from or across the topographical pattern.
[0064] FIG. 1A illustrates one embodiment of the system, where
light is directed onto the biosensor substrate through an ambient
medium (e.g., air or water) and detected from the biosensor
substrate through the ambient medium. The system 10 includes the
light source 12, the biosensor 14, and the detector 16.
[0065] FIG. 1B illustrates another embodiment of the system, where
light is coupled into the biosensor via a waveguide and detected
from the biosensor via a waveguide. The system 20 includes the
light source 22, waveguide 23, biosensor 24, waveguide 25, and
detector 26.
[0066] The system embodiments illustrated in FIG. 1A-B are
exemplary, and are capable of modification to accommodate different
biosensor substrates (i.e., different sensing platforms) of the
type described above.
[0067] The light source functions as a source of illumination and
may be, for example, an argon, cadmium, helium, or nitrogen laser
and accompanying optics positioned to illuminate the biosensor and
the detector. The detector is positioned to capture
photoluminescent emissions from the biosensor and to detect changes
in photoluminescent emissions from the biosensor. Exemplary
detectors include, without limitation, a charge coupled display,
spectrophotometer, photodiode array, photomultiplier tube array, or
active pixel sensor array.
[0068] The system optionally includes a polarizer positioned
between the light source and the biosensor. The system also
optionally includes a filter positioned between the light source
and the biosensor or, alternatively, between the biosensor and the
detector. In FIG. 1A, the optional polarizer 17 and filter 18 are
shown.
[0069] The system may include two or more of the biosensors, each
of the biosensors being coupled to the light source and the
detector.
[0070] A further aspect also relates to a method of detecting the
presence of a target molecule in a sample that involves providing a
biosensor of the type described above, exposing a sample to the one
or more sites on the biosensor surface, and detecting a change in
an optical property of biosensor surface at the one or more sites
following said exposing.
[0071] Samples which can be examined include blood, water, a
suspension of solids (e.g., food particles, soil particles, etc.)
in an aqueous solution, or a cell suspension from a clinical
isolate (such as a tissue homogenate from a mammalian or other
patient), cell free extracts, and similar types of suspensions or
solutions.
[0072] According to this aspect, target molecules may include,
without limitation, proteins (including without limitation enzymes,
antibodies or fragments thereof), glycoproteins, peptidoglycans,
carbohydrates, lipoproteins, a lipoteichoic acid, lipid A,
phosphates, nucleic acids which are expressed by certain pathogens
(e.g., bacteria, viruses, multicellular fungi, yeasts, protozoans,
multicellular parasites, etc.), or organic compounds such as
naturally occurring toxins or organic warfare agents, etc. These
target molecules can be detected from any source, including food
samples, water samples, homogenized tissue from organisms, etc.
Moreover, the biological sensor can also be used effectively to
detect multiple layers of biomolecular interactions, termed
"cascade sensing." Thus, a target, once bound, becomes a probe for
a secondary target. This can involve detection of small molecule
recognition events that take place relatively far from the
substrate's surface.
[0073] Presence of the target molecule in the sample will dictate
the change in optical property. The specific optical property that
is modified will vary depending upon the particular structure used
for the biosensor, but generally includes any one or more of
transmission peak wavelength shift, absorption peak wavelength
shift, or refractive index change. To determine whether a change in
optical property has occurred, a baseline optical measurement is
made prior to exposure to a sample. After exposure to the sample, a
second optical measurement is made and the first and second
measurements are compared. Typically any change will depend on the
size of the target to be recognized and its concentration within
the sample. In another embodiment, the topographical pattern is
illuminated with light before and after exposing the sample to the
one or more sites on the biosensor surface.
[0074] A still further aspect relates to a method of quantifying
the amount of a biological target present in a sample. This method
includes providing a biosensor of the type described above,
exposing a sample to the one or more sites on the biosensor
surface, and detecting a change in an optical property of biosensor
surface at the one or more sites following said exposing, wherein
the amount of biological target is quantifiable based on the extent
of the change of the optical property.
[0075] To quantify the amount of biological target present in a
sample, the light source and the detector can both be present in a
spectrometer. A computer with an appropriate microprocessor can be
coupled to the detector to receive data from the spectrometer and
analyze all the data to compare the optical properties before and
after exposure of the biosensor to a target molecule.
EXAMPLES
[0076] The following examples are provided to illustrate
embodiments of the disclosed methods and devices, but they are by
no means intended to limit their scope.
Materials and Methods for Examples 1-5
[0077] Photonic Crystal Design.
[0078] The PhC design used in Example 1-5 has been described before
(Pal et al., Biosens. Bioelectron. 26:4024-4031 (2011), the
disclosure of which is incorporated herein by reference in its
entirety). Briefly, the 2D PhC slab structure consists of a
25.times.26 array of air wells in a triangular lattice pattern with
row of wells removed from the center creating a w1 waveguide (line
defect). A nanocavity was created by modifying the radius of a
single air well adjacent to the waveguide (point defect).
[0079] Device Fabrication.
[0080] A p-type silicon-on-insulator (SOI) wafer (<100>) with
a 450 nm silicon device layer on top of 1 .mu.m thick buried
silicon oxide (BOX) was used as the starting substrate for the
PhCs. For fabrication with PMMA, a 130 nm oxide hard mask was
thermally grown on the Si layer via wet oxidization.
Polymethylmethacrylate (PMMA) was used as an e-beam resist and a
JEOL JBX-9300FS system was used to write the PhC patterns. The
pattern was developed and dry etched using argon assisted CHF.sub.3
gas in a reactive-ion-etcher to transfer the oxide hard mask,
followed by a gas etch with CF.sub.4 and BCl.sub.3 to etch the Si
device layer. The individual PhC devices were cleaved with a
diamond scribe to create smooth waveguide facets to facilitate
light coupling. For fabrication with HSQ, the native oxide layer of
the SOI substrate was stripped using a buffered oxide etch (6:1
hydrofluoric acid/ammonium fluoride). Hydrogen silsesquioxane (HSQ)
was used as an e-beam resist and a JEOL JBX-9300FS system was used
to write the PhC patterns. After exposure, the pattern was
developed and transferred using a CF.sub.4 and BCl.sub.3 gas etch.
The individual PhC devices were cleaved with a diamond scribe to
create smooth waveguide facets to facilitate light coupling.
[0081] Finite Element Modeling. All solutions were generated using
COMSOL Multiphysics (v.4.2a). Bulk diffusion was modeled using the
Transport of Diluted Species module. Surface reactions were modeled
using General Form Boundary PDEs. Optical Set-up: A tunable laser
(Hewlett Packard, model 8168F, output power: -7 to 7 dBm) operating
within the wavelength range of 1440-1590 nm (wavelength resolution
of 0.02 nm) was used to scan and optically probe the 2D PhC device.
A polarization controller was used to excite the TE modes and light
was coupled through tapered ridge waveguides into the PhC device
using a tapered lensed fiber (Nanonics, Israel). The transmitted
optical power was measured using an indium gallium arsenide
(InGaAs) photodiode detector (Teledyne Judson Technologies, PA,
USA).
[0082] Nanoparticle Synthesis.
[0083] Poly(N-isopropylacrylamide) microgels were prepared via free
radical precipitation polymerization. The monomers
N-isopropylacrylamide (0.76 g) and bis-acrylamide (BIS) (0.013 g)
were dissolved in double distilled water (ddH.sub.2O)(50 mL) inside
of a 3-neck flask (500 mL). The solution was then mixed with
aqueous 1% sodium dodecyl sulphate (SDS) (0.34 mL). The flask
containing the solution was equipped with a nitrogen line, overhead
stirrer, and gas outlet. The solution was bubbled with nitrogen for
45 minutes to remove dissolved oxygen. The mixture was heated to
60.degree. C. Ammonium persulfate (0.0166 g) was dissolved in
ddH.sub.2O (0.5 mL) and injected into the flask to start the
reaction. The reaction proceeded for 5 hours in an inert atmosphere
at a constant stir rate of 200 RPM. After 10 minute the solution
became visibly turbid, which was indicative of particle formation.
At 5 hours, the flask was removed from heat opened to ambient
oxygen while maintaining a constant stir-rate for 15 minutes. The
solution was filtered through a 1.2 .mu.m cut-off syringe filter
(Millipore). The purified solution was used as is.
[0084] Nanoparticle Dipcoating.
[0085] Both the flat silicon dioxide and nanostructured SOI chips
were dipcoated using a syringe pump (Yale Apparatus YA-12) which
was modified to hold a pair of tweezers and mounted vertically. The
chips were dipped into a 1:100 v/v dilution of PNIPAM particles in
ddH.sub.2O at a speed of 50 .mu.m per minute until the chip was
submerged in solution to approximately half the height of the chip
(0.2-0.5 cm, depending on the chip length). The pump was stopped
for 30 seconds to allow the chip to equilibrate with the solution
before being pulled out at a rate of 100 .mu.m per minute. Once the
chips had cleared the water line, the edges of the chip were
inspected to ensure they were completely dry before removing
them.
[0086] Aminosilane-Glutaraldehyde Surface Functionalization.
[0087] To generate a protein-reactive surface, both planar silicon
oxide and SOI chips were functionalized with glutaraldehyde (GA) as
per the following protocol. First, the chips were carefully cleaned
in piranha solution (3:1 (v/v) conc. sulfuric acid to 30% hydrogen
peroxide for 30 minutes, followed by ddH.sub.2O rinse and dried
under a stream of nitrogen gas. Next, the chips were incubated with
a 1% (v/v) solution of (3-aminopropyl)dimethylethoxysilane in
anhydrous toluene for 20 minutes on an orbital shaker. The chips
were then repeatedly washed with anhydrous toluene, dried under a
stream of nitrogen and baked at 110.degree. C. for 30 minutes.
After the chips had cooled to room temperature (approximately 5
minutes), a solution of 1.25% (v/v) GA in modified PBS buffer
(MPBS: 10 mM NaH.sub.2PO.sub.4, 10 mM Na.sub.2HPO.sub.4, 150 mM
NaCl at pH 7.2) was poured over them, and the chips were left in
this solution on a shaker for 60 minutes. Afterwards, they were
washed with MPBS and ddH.sub.2O and dried under a nitrogen
stream.
[0088] Antibody Localization on Planar Silicon Oxide Chips.
[0089] Planar square silicon oxide chips (1 cm.times.1 cm) were
washed in piranha solution and functionalized with GA as described
above. The chips were carefully dipcoated with PNIPAM nanoparticles
such that half of each chip was passivated while the other half
remained protein reactive. Next, the chips were incubated with
three different concentrations of rhodamine-labelled IgG (0.1
.mu.M, 0.5 .mu.M and 1 .mu.M) for 60 minutes. Lastly, the chips
were washed with MPBS-ET (MPBS buffer with 3 mM
Ethylenediaminetetraacetic acid and 0.05% (v/v) Tween-20) for 30
minutes, rinsed with ddH.sub.2O and imaged with
epifluorescence.
[0090] Epifluorescence Microscopy.
[0091] The fluorescent intensity of the passivated and
un-passivated areas of the planar silicon oxide chips was evaluated
with an Olympus-BX60 microscope with a Qicam FAST-1394 (Qimaging)
active cooled CCD camera. A silicon dioxide chip with no
fluorophore was used as a control to measure the background
fluorescence. The exposure time was kept constant at 2 seconds.
Data analysis was performed with ImageJ (NIH). The relative
fluorescence intensity (RFI) was calculated as:
RFI = 100 * ( 1 - ( F M _ - F C _ ) ( F U _ - F C _ ) )
##EQU00001##
where F.sub.U, F.sub.M, and F.sub.C are the unmasked, masked, and
control average fluorescence intensities. The error bars for each
condition were computed as the relative fluorescence intensity
multiplied by the square root of the sum of the squares of the
normalized standard deviations (standard deviation/mean) of the
masked and unmasked portions of the chip, and a control chip with
no fluorophore:
SD RFI = RFI * ( SD F U F U _ ) 2 + ( SD F M F M _ ) 2 + ( SD F C F
C _ ) 2 ##EQU00002##
where SD.sub.RFI is the calculated error, SD.sub.F.sub.U,
SD.sub.F.sub.M, and SD.sub.F.sub.C are the standard deviations of
unmasked, masked, and control fluorescence intensities
respectively.
[0092] Antibody Localization on PhC Chips. Newly fabricated PhC
chips were thermally oxidized in a furnace with an oxygen gas
stream at 900.degree. C. for 15 minutes. The chips were then
functionalized with aminosilane-glutaraldehyde chemistry. Next, the
PhC chips were dipcoated with PNIPAM particles at a rate of 100
.mu.m sec.sup.-1. After passivation, a 10 .mu.L droplet of 0.1
.mu.M IgG in MPBS buffer was placed on the chip covering the PhC
sensing region. The chips were then placed in a humidity chamber
for 1 hour (no evaporation appeared to occur), allowing the
human-IgG to covalently bind to the exposed GA groups via
amine-aldehyde coupling chemistry (Schiff base formation). After
antibody immobilization, the remaining aldehyde groups were blocked
by incubating the chips in BSA (10 .mu.M) solution in HBS buffer
(20 mM2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid
(HEPES), 150 mM NaCl, at pH 7.2) for 1 hour. The chips were then
washed with MPBS, dried under nitrogen stream and incubated with a
10 .mu.L droplet of the target solution (anti-IgG) at increasing
concentrations diluted in MPBS buffer for a period of 1 hour.
Finally, the chips were washed with MPBS-ET for 30 min on an
orbital shaker, rinsed with ddH.sub.2O and dried under a stream of
nitrogen. The chips were then subjected to optical
characterization.
[0093] Peak Fitting and Data Analysis.
[0094] After spectra collection, each data set was filtered using
Origin (OriginLab, Northampton, Mass.). A fast Fourier transform
low-pass filter with a frequency cut-off of 0.5 was used to remove
high frequency noise. The peaks were then fit to the Lorentz
equation with transmission intensity (a.u.) plotted as a function
of wavelength (nm):
y = y 0 + 2 A .pi. * w 4 ( x - x c ) 2 + w 2 ##EQU00003##
The fitted value of x.sub.c (represented in units of wavelength)
was used as the location of the minimum absorption for the
data-set.
Example 1--Numerical Simulations
[0095] Whether antibody localization to active areas of a nanoscale
device will enhance the limit of detection was tested through
simulation using finite element methods. The geometry used was a
section of a sphere, representing a water droplet with a contact
angle of 55 degrees and a volume of approximately 10 .mu.L (FIG.
2A). The flat region of the sectioned sphere was composed of a 70
.mu.m.sup.2 area circle (active region) surrounded by a circular
contact surface with a 4.5 mm diameter (inactive region). Due to
the large size difference between the two areas, a much finer
volume (tetrahedral) mesh was required near the active site, which
was located at the center of the flat region (FIG. 2B).
[0096] The surface reaction was modeled as immunoglobulin gamma
(IgG) binding to anti-immunoglobulin gamma (anti-IgG) with 1:1
binding stoichiometry. The surface density of antibody sites was
assumed to be a monolayer with a value of 1.2.times.10.sup.12
antibodies cm.sup.-2. The k.sub.on and k.sub.off values were 250
m.sup.3s.sup.-1 mol.sup.-1 and 0.0003 s.sup.-1, respectively. The
diffusion coefficient for IgG was modeled as 5.times.10.sup.-11
m.sup.2 s.sup.-1. The following diffusion equation was solved for
the entire domain:
.differential. c .differential. t + .gradient. ( - D .gradient. c )
= 0 ( 1 ) ##EQU00004##
where c and D are the bulk analyte concentration (mol m.sup.-3) and
diffusion coefficient of the target (m.sup.2 sec.sup.-1),
respectively. Boundary conditions representing the flux balance
between the surface and bulk concentrations of target species were
set for the active (sensing) and inactive areas:
n(D.gradient.c)=R.sub.c.sub.s=-(k.sub.on*c*(e*.theta..sub.max-c.sub.s)-k-
.sub.off*c.sub.s) (2)
[0097] where n is a unit vector normal to the reaction surfaces,
R.sub.cs is the inward flux of the target into the bulk (can be
either positive or negative), k.sub.on and k.sub.off are the
kinetic on (m.sup.3 sec.sup.-1 mol.sup.-1) and kinetic off
(s.sup.-1) rate constants for a target and capture-molecule pair,
c.sub.s is the surface concentration of bound target (mol
m.sup.-2), .theta..sub.max is the maximum surface concentration of
available binding sites (mol m.sup.-2) which was explicitly set at
the active and inactive regions, and e is an efficiency factor for
antibody localization (dimensionless). The active boundary,
representing the "nanoscale sensor" was modeled with an efficiency
factor of 1 (e=1) implying that the active region had the maximal
surface density of antibodies. The efficiency factor on the
inactive boundaries was set to one of three values: e=0 for perfect
antibody localization (ABL) (no antibodies at the inactive region),
e=0.05 for 95% ABL (surface density of antibody at the inactive
region was 5% of the value at the active region), and e=1 for no
ABL (the inactive area had the same surface density of antibodies
as the active area).
[0098] Dose-response curves (FIG. 3A-B) were generated by plotting
fractional occupancy of target,
( c s .theta. max ) ##EQU00005##
at the active region for different concentrations of analyte after
24 hours of simulated incubation. The results show a marked
difference in the location and shape of the dose-response curve for
perfect ABL (e=0), partial ABL (e=0.05), and no ABL (e=1). An
assumption of perfect ABL produces a dose-response curve whose
midpoint lies at the IgG/anti-IgG equilibrium constant, as expected
(at 50% surface coverage, the concentration value matches the
expected equilibrium constant,
K D = k off k on = 1.2 nM ) . ##EQU00006##
Simulations suggest that even a 95% ABL provides a 10-fold higher
surface coverage at the active sensing area relative to the case
with no ABL (FIG. 3A). Thus, these simulations predict that ABL
with a nanoscale sensor will generate a significantly higher signal
compared to a sensor with no ABL at the same analyte concentration,
thereby improving the limit of detection.
Example 2-2D Photonic Crystal Biosensor Operation
[0099] To test these predictions experimentally, a 2-dimensional
photonic crystal (2D PhC) biosensor previously employed to detect
proteins (Pal et al., Biosens. Bioelectron. 44:229-234 (2013),
which is hereby incorporated by reference in its entirety) and
virus-like particles (Pelton et al., Colloids Surf 20:247-256
(1986), the disclosure of which is incorporated herein by reference
in its entirety) was used. The active sensing area of this device
is approximately 7 .mu.m by 10 .mu.m and contains of 509
cylindrical wells in silicon with a diameter of 220 nm and a single
cylindrical well 150 nm in diameter, with all wells etched to
depths of .about.400 nm. A w1 waveguide allows propagation of
guided modes within the photonic band-gap (PBG) of the crystal.
Light is confined within the embedded silicon (Si) layer by total
internal reflection from the encasing silicon oxide (SiO.sub.2),
which has a lower refractive index. Breaking the translational
symmetry by modifying the radius of a well to create a defect (FIG.
4A), gives rise to a localized mode within the PBG. Thus, the w1
waveguide photonic crystal allows light transmission at all
frequencies except at the resonant defect nano-cavity wavelength,
resulting in a characteristic sharp dip in its transmission
spectrum. The electric field is strongly confined in the defect at
resonance. Analyte binding causes the local refractive index to
change resulting in a red-shift of the resonant wavelength due to
the strong light-matter interaction within the defect. FIG. 4B
depicts the red shifts observed due to the refractive index change
when a 2D PhC sensor is analyzed in air (n.about.1.0) followed by
water (n.about.1.32 at 1550 nm wavelength).
Example 3--Antibody Surface Coverage Evaluation and Localization
with PNIPAM Nanoparticle Mask
[0100] PNIPAM is versatile and can be synthesized as nanoparticles
(Andersson et al., J. Polym. Sci., Part B: Polym. Phys.
44:3305-3314 (2006), the disclosure of which is incorporated herein
by reference in its entirety) with control over their size
(Blackburn et al., Colloid Polym. Sci. 286:563-569 (2008); Kratz et
al., Colloids Surf, 170:137-149 (2000), the disclosures of which
are incorporated herein by reference in their entirety) and charge
(Hoare et al., Macromolecules 37:2544-2550 (2004); Karg et al.,
Langmuir 24:6300-6306 (2008); Tsuji et al., Langmuir 21:8439-8442
(2005), the disclosures of which are incorporated herein by
reference in their entirety). Several groups have demonstrated that
PNIPAM nanoparticles (PNPs) form self-assembled well packed
monolayers (Kawaguchi et al., Colloid Polym. Sci. 270:53-57 (1992);
Pelton et al., Colloids Surf 20:247-256 (1986), the disclosures of
which are incorporated herein by reference in their entirety).
[0101] While continuous PNIPAM films have previously been used in
surface blocking, a first step was to assess the ability of PNP
masks to function in this capacity. This was accomplished using
flat silicon dioxide-on-silicon chips, first made protein reactive
by aminosilane-gluteraldehyde chemistry followed by dipcoating 315
nm diameter PNIPAM particles on half of the substrate. The entire
chip was then exposed to a solution of rhodamine labelled IgG
(r-IgG) concentrations and rinsed. The fluorescence intensities of
the PNIPAM-masked and unmasked portions of the chip were determined
by epifluorescence microscopy, and were visibly higher on the
unmasked regions of the substrate (FIG. 5A). The relative
fluorescence intensities of the masked regions were found to be 95%
lower than the unmasked regions at 0.1 .mu.M r-IgG (FIG. 5B).
Higher concentrations had decreased relative fluorescence
intensities; however, the nanoparticle masks could still localize
antibody effectively within the concentration range likely to be
used for sensor functionalization.
Example 4--Nanoparticle Assembly on 2D PhC Structures
[0102] PNPs were deposited on a PhC chip via dip coating. The PNPs
preferentially assembled around lithographed features, and did not
settle on top of the wells. This behavior was reproducible and
consistent on both "extruded" and "embedded" versions of the
sensor, fabricated via negative (Hydrogen silsesquioxane (HSQ)) and
positive (Polymethylmethacrylate (PMMA)) tone resists respectively
with e-beam lithography (FIGS. 6A-B).
Example 5--Enhanced Limit of Detection Via Topographically
Selective Passivation
[0103] To test the effect of nanoparticle-mediated blocking of
non-active portions of the sensor surface during functionalization,
2D PhC chips were functionalized with IgG (probe) and used to
detect anti-IgG (target) at various concentrations. The red-shift
for each concentration was the difference in the wavelength of the
minimum absorption before and after a chip was incubated at that
particular anti-IgG concentration, subtracted from the red-shift of
an identically functionalized control chip which was not exposed to
anti-IgG. Two sets of experiments were run: one in which the chips
were unmodified, and another in which the chips were dip-coated
with PNIPAM particles prior to probe (IgG) functionalization. Each
data point was tested with at least 3 devices. Normalizing the
control shift value to zero allowed for a comparison of the
relative shift (normalized red-shift) for chips with and without
ABL (FIG. 7A). Neither chip had statistically significant shifts
from the background with a target concentration of 50 pM. However,
the antibody-localized (ABL) chip showed an increasing signal at
all higher concentrations of target (FIG. 7B), while it was
necessary to test a 10-fold higher concentration of anti-IgG on the
non-ABL chip than the highest tested concentration examined on the
ABL sensor to see a measurable signal. The lowest detectable
concentration of anti-IgG was found to be 0.5 nM with ABL,
demonstrating that the LoD of the sensor was improved by at least
one order of magnitude.
Discussion of Examples 1-5
[0104] To improve the limit of detection of nanoscale biosensors,
non-productive loss of target to inactive regions needs to be
minimized. Precisely aligning top-down target delivery systems to
nanoscale features on a substrate is challenging and requires
expensive instrumentation. As an alternative, a fast and
inexpensive bottom-up technique was developed based on coating with
a monolayer of PNPs. It was demonstrated that these PNPs assemble
around topographically distinct features of 2D PhCs, leaving the
active sensor area free for immobilization of capture antibodies.
Consistent with FEM calculations, this provided over an order of
magnitude improvement in the lowest limit of detection for the
sensor. It is believed that this strategy will work equally well
with other nanoscale sensors, since most have topographical
features distinguishing the active sensing area from the remainder
of the device.
[0105] It will be appreciated that variants of the above-disclosed
and other features and functions, or alternatives thereof, may be
combined into many other different systems or applications. Various
presently unforeseen or unanticipated alternatives, modifications,
variations, or improvements therein may be subsequently made by
those skilled in the art which are also intended to be encompassed
by the following claims.
* * * * *