Metallic Nanohole Arrays on Fluoropolymer Substrates as Small Label

Aug 19, 2008 - and Plasmonic Systems, 33 Oxford Street Cambridge, Massachusetts 02138,. The Charles Stark Draper Laboratory, 555 Technology Square, ...
0 downloads 0 Views 1MB Size
Download PDF
NANO LETTERS

Metallic Nanohole Arrays on Fluoropolymer Substrates as Small Label-Free Real-Time Bioprobes

2008 Vol. 8, No. 9 2718-2724

Jiun-Chan Yang,†,‡ Jin Ji,† James M. Hogle,*,†,‡ and Dale N. Larson*,†,‡,§ Department of Biochemistry and Molecular Pharmacology, HarVard Medical School, 240 Longwood AVenue, Boston, Massachusetts 02115, HarVard Center for Microfluidic and Plasmonic Systems, 33 Oxford Street Cambridge, Massachusetts 02138, The Charles Stark Draper Laboratory, 555 Technology Square, Cambridge, Massachusetts 02139 Received April 11, 2008; Revised Manuscript Received July 3, 2008

ABSTRACT We describe a nanoplasmonic probing platform that exploits small-dimension (e20 µm2) ordered arrays of subwavelength holes for multiplexed, high spatial resolution, and real-time analysis on biorecognition events. Nanohole arrays are perforated on a super smooth gold surface (roughness rms < 2.7 Å) attached on a fluoropolymer (FEP) substrate fabricated by a replica technique. The smooth surface of gold provides a superb environment for fabricating nanometer features and uniform immobilization of biomolecules. The refractive index matching between FEP and biological solutions contributes to ∼20% improvement on the sensing performance. Spectral studies on a series of small-dimension nanohole arrays from 1 µm2 to 20 µm2 indicate that the plasmonic sensing sensitivity improves as the gold-solution contact area increases. Our results also demonstrate that nanohole arrays with a dimension as small as 1 µm2 can be used to effectively detect biomolecular binding events and analyze the binding kinetics. The future scientific opportunities opened by this nanohole platform include highly multiplexed analysis of ligand interactions with membrane proteins on high quality supported lipid bilayers.

Nanoplasmonic sensing is an emerging technique that utilizes the interaction of light with surface plasmons, collective oscillations of free electrons in the conduction band, in metallic nanostructures to transduce chemical bindings to signal for remote optical readout. With receptive biological macromolecules immobilized on nanostructured metal surfaces, whose length scale is close to the wavelength of light, the binding of target biomolecules leads to an increase of the local refractive index, which is then transduced into the change of spectral features observed at the far-field. The greatest attraction of nanoplasmonic sensing is it does not require extrinsic fluorescent or radioactive labeling (i.e., “label-free”), which would potentially change the binding properties of biomolecules and cause other significant problems like background binding and autofluorescence.1 The small sensing volume resulting from short electromagnetic field decay length leads to high sensitivity for detecting biomolecular binding events in local environments. Significant improvement in nanofabrication and nanomaterials synthesis in the past decade has allowed a variety of * Address correspondence to [email protected] and [email protected]. † Harvard Medical School. ‡ Harvard Center for Microfluidic and Plasmonic Systems. § The Charles Stark Draper Laboratory. 10.1021/nl801043t CCC: $40.75 Published on Web 08/19/2008

 2008 American Chemical Society

nanostructures to be utilized as nanoplasmonic sensors. A wide range of nanostructure shapes such as triangles, cubes, rods, and rings 2,3 have been reported to exhibit high sensitivity based on localized surface plasmon resonance (LSPR) or coupled plasmonic resonance. The discovery of extraordinary light transmission through metallic subwavelength-hole arrays,4 with an efficiency that is orders of magnitude greater than that predicted by classic optical theory, has created another exciting opportunity for nanoplasmonic sensing. Surface plasmon polaritons (SPPs) are generally proposed to explain this phenomenon observed on periodically arrayed nanoholes, although some works have claimed that other mechanisms like diffraction and LSPR also play important roles.5-7 Several studies exploiting periodically arrayed nanoholes fabricated by focused ion beam (FIB),8-12 ebeam lithography,13 and soft lithography14,15 have demonstrated their excellent sensitivity when used as label-free chemical sensing devices. Label-free plasmonic sensing is traditionally performed on prisms (Kretschmann geometry) or grating couplers, on which biomolecules are immobilized on a continuous metal film and the angular distribution or the reflected intensity is measured.16 Those real-time techniques have been successful in performing binding affinity analysis of proteins or small molecules, and commercial instruments are available.1 Compared with

conventional coupling techniques, the nanohole geometry has a much smaller probing area and can be fabricated in high spatial density. The optical transmission configuration enables multiplexed detection and offers better opportunities to develop low-cost devices with integrated microfluidics for rapid bioanalytical measurements. Nanohole plasmonic sensing devices that have been reported to have excellent detection limits are mostly based on wavelength interrogation with relatively low sampling rates on infinite or quasi-infinite nanohole arrays. High spatial density multiplexed real-time measurements are difficult to achieve in those designs. For screening binding affinity of thousands of proteins in parallel, small-dimension array nanoholes are of more interest because they offer high packing density of sensing elements and can be integrated with compact microfluidic devices that require less analyte volume. Small-dimension nanohole arrays have been reported to exhibit strikingly different transmission features from infinite arrays15 because the contribution from the edge becomes significant.17 Their refractive-index sensing properties have not been extensively studied. Another key point in developing a nanohole sensing system is the interface between the nanostructure surface and the chemical or biological systems. For lipid-membranemediated reactions, especially those associated with membrane proteins or membrane fusion processes, a smooth surface is greatly desired to help building a high-quality single lipid-bilayer that incorporates protein receptors and improving experimental reproducibility.18,19 Visualization of chemisorbed proteins and molecules can also be achieved by using scanning probe techniques (AFM/STM) on atomically smooth surfaces.19,20 On the other hand, from the view of nanofabrication, smoother surface will be beneficial on improving the quality of fabricated features, especially when they are in nanometer sizes. Here, we present a nanoplasmonic sensing platform which exploits small-dimension ordered arrays of nanoholes (e4.5 µm × 4.5 µm) for multiplexed (30 arrays in a 48 µm × 58 µm area) and real-time (time resolution e1s) analysis on biorecognition events. The measuring platform was built on a conventional inverted microscope with regular bright-field optics. Intensity interrogation was performed instead of commonly used wavelength interrogation to achieve multiplex and real-time sensing capability. Nanohole arrays were perforated on super smooth gold surfaces fabricated by replica techniques. We chose fluorinated ethylene propylene copolymer (FEP) as the replica substrate because it is chemically inert, thermoplastic, transparent in the visible region, and has a refractive index (1.341 at λ ) 590 nm) close to biological solutions. It has been reported that, when the refractive indices on both sides of the gold film match, the intensity of extraordinary transmission through nanoholes would be strongly enhanced because of the overlap of transmission maxima from two interfaces and the higher lifetime of a coupled state from strong coupling of SP modes on each side of the nanohole arrays.21-23 Nano Lett., Vol. 8, No. 9, 2008

Meanwhile, although the change in local dielectric environments normally causes more shift of the spectral features in the near-infrared and infrared range than in the visible range,10,14 we used visible light as the sensing wavelength to prevent potential near-infrared adsorption of water and many biological species. Moreover, shorter excitation wavelengths generate surface plasmon wave with shorter propagation lengths in the X-Y plane, leading to higher possible packing density and hence higher throughput on multiplexed biosensing. The substrate for nanoholes was prepared by the templatestripped-gold (TSG) method18,19 to obtain a super smooth surface for biomolecule immobilization. FEP was used to replace epoxy resins used in the regular TSG method (see Supporting Information). The resulting TSG substrate has a 220 nm thick gold film on FEP with the smooth surface facing up (Figure 1a). The surface roughness is ∼1.2 nm (peak to peak, rms ) 2.7 Å; see Supporting Information). 220 nm thickness was used to effectively block the direct transmitted light. For comparison, a regular gold substrate was prepared by thermally depositing 8 nm chromium and 220 nm gold on a cleaned glass slide. FEI DB235 focused ion beam system with 30 kV acceleration voltage and 10 pA ion current was applied to mill nanoholes arrays on the gold film. Ion image magnification was calibrated before each milling. After immobilizing biomolecules, a PDMS block with a microfluidic channel (volume ∼ 3 µL) was mounted on the nanohole film. A syringe pump was used to deliver solutions through this channel at a 30 µL/min flow rate. Transmission spectra and real-time sensing responses were collected using an inverted microscope with bright-field accessories (Nikon eclipse TE300, Figure 1b). For multiplexed spectral acquisition, a multispectral illumination source and an intensity imaging CCD were used instead of a polychromatic spectrometer.24 Nonpolarized white light from a 75 W xenon lamp was passed through a double monochromator (Jobin-Yvon SPEX 1680B) and a condenser lens. Divergence of light (5°∼39°) can be adjusted using the diaphragm on the condenser lens. The light coming out from the nanohole arrays was collected with a 20× objective (NA ) 0.5) and then focused on an uncooled CCD (QImage, Intensified Retiga). The spectra were corrected by the lamp transmission profile through the glass substrate. The bandpass of the monochromator was set to 1 nm for acquiring transmission spectra. An image processing program was created to extract intensity value of individual nanohole arrays from CCD images (see Supporting Information). A series of nanohole patterns with different nanohole diameters (d), periodicity (P, hole-to-hole distance), and numbers was fabricated on TSG substrates. Figure 1c is the CCD image of one of the nanohole array patterns we studied. This pattern contains thirty 9 × 9 nanohole arrays spaced by 11 µm. Since the calculated SPP propagation length on gold along the X-Y direction is less than 3 µm for excitation wavelengths shorter than 630 nm,16 11 µm spacing ensures sufficient separation between individual arrays to avoid interference, while allowing a maximum number of nanohole arrays in the CCD field of view. The hole diameter was 2719

Figure 1. (a) Fluoropolymer-based nanohole array device with the super smooth gold surface facing up, fabricated by the template-stripped method. (b) Schematic of the experimental setup. (c) CCD image of 30 sets of nanohole arrays at λ ) 540 nm. (d) SEM micrography of a 9 × 9 nanohole array on the TSG substrate. (e) SEM cross section image of a fluoropolymer-based nanohole array device. Title angle ) 53°.

changed from 130 to 180 nm, and the D value (hole periodicity/diameter) was varied from 1.67 to 3.00. Figure 1d,e shows SEM micrographs of the top view and cross section of a nanohole array. A protruded ring of gold approximately 160 nm in height was found underneath the hole. It is mainly due to Ga ion bombardment on soft FEP since this phenomenon was not observed on normal gold substrates. We also found that it was relatively easier to fabricate high-quality nanoholes (Figure 1d inset) on TGS substrate by FIB than on fresh thermally deposited gold surface (surface roughness ∼10 nm). The optical setup and design of nanohole arrays allow collecting transmission spectra of all thirty small nanohole arrays simultaneously. This feature enables fast screening of small nanohole array patterns to determine the sensing sensitivity at different wavelengths. The black curve in Figure 2a represents a typical transmission spectrum of one of the nanohole arrays (d ) 180 nm, D ) 2.33, TSG substrates) in Figure 1c when flowing DI water and the red curve represents the spectrum upon switching flowing media from DI water (RI ) 1.3331) to 21% NaCl (RI ) 1.3702). Bulk plasmon resonance peak around λ ) 510 nm was found in almost all spectra and did not shift upon the change of flowing media. The wavelength shift sensitivity is 323 nm/RIU for λ ∼ 600 nm. The blue curve in Figure 2b represents the differential intensity (∆T/T in percentage) between two spectra in Figure 2a. Two maxima (562 and 618 nm) and one minima (588 nm) are found in the visible range. These wavelengths 2720

(denoted as S wavelengths in the following context) will be useful for single-wavelength sensing by intensity interrogation. We compared the spectra collected with different collimation conditions. Larger light divergence (39°) normally excites a wider distribution of wavevectors and leads to lower sensitivity than smaller light divergence (5°). However, the signal-to-noise ratio (S/N) with 5° divergence is much worse than 39° divergence because of lower photon flux. We hence used 39° divergence (NA ) 0.34) for the rest of the experiment. From analyzing the transmission spectral features from a series of nanohole arrays, we discovered that the hole diameters only slightly affect the transmission maxima, and the periodicity is the dominant factor. Larger diameters lead to higher transmission. For the same periodicity, the transmission intensity from a d ) 180 nm array (9 × 9, P ) 360) is around 7 times that of a d ) 130 nm array at the resonance peak. This observation is consistent with previous reports using large nanohole arrays and collimated/polarized light sources.4,25-27 Figure 2c shows the relation between S wavelengths (plotted if higher than 5% intensity change) and hole periodicity from 30 different small nanohole arrays in Figure 1c from TSG and regular gold substrates. Maxima and minima of differential intensity form four different linear relations (lines 1∼4) in the visible region. Two different nanohole substrates (glass and FEP) exhibit the same linear relations of maxima and minima of differential intensity, although peak positions in transmission spectra differ due Nano Lett., Vol. 8, No. 9, 2008

Figure 2. (a) Typical transmission spectra of a nanohole array (d ) 180 nm, P ) 419 nm) on the TSG substrate. Black plot: in DI water. Red plot: in 21% NaCl solution. (b) Intensity difference (∆T/T) between two plots in Figure 2a. (c) The relation between hole periodicity (P) and the wavelengths of maxima/minima in differential intensity from all 30 sets of nanohole arrays. The hole diameter labels are removed for clarity. Symbols: (O) minima from TSG; (b) minima from regular gold; (]) maxima from TSG; ([) maxima from regular gold. Lines 1 and 2: minima. Lines 3 and 4: maxima. (d) Bubble plots of line 1 and 2 in Figure 2c.

to the different interfaces between gold and substrates. This observation confirms that the refractive-index sensing occurs exclusively on the gold-solution interface. For periodicity 450 nm, the few minima and maxima at the right-bottom corner of Figure 2c indicate another set of trends extending to the largeperiodicity region. No maxima or minima of differential intensity were found on the left-hand side of Line 1 in the visible region. Each line in Figure 2c represents a red-shift edge of transmission peaks. For a small refractive index change (e.g., 0.0371 in this figure), only the shift of a single edge of transmission peaks was observed in most spectra collected. A full theoretical analysis of the nature of transmission features is beyond the scope of this article. Nevertheless, Figure 2c is helpful in determining nanohole periodicity when using a monochromatic excitation source that has better intensity and stability than white light sources. The magnitude of intensity change of lines 1 and 2 was plotted in Figure 2d in the bubble plot format. A strong trend can be observed that shorter periodicity mostly leads to weaker intensity change for both lines, that is, poorer sensing performance. For the same diameter, a long-periodicity array always has more intensity change than the short-periodicity one. On average, line 2 has slightly better sensing performance than line 1, especially for P < 400 nm. Lines 3 and 4 have the same trend as lines 1 and 2 (data not shown for clarity). The extraordinary transmission from nanohole arrays on TSG substrates is typically more intense than on normal gold substrates by 10∼20% and the intensity change (sensing performance) is on average also about 20% better. The fact that longer periodicity leads to higher sensitivity implies that a larger nanohole array (for the same nanohole number) is a better sensor. This suggests the interaction area between gold and flowing solutions substantially affects the sensor sensitivity. We also investigated the effect of the number of nanoholes on sensitivity. In Figure 3a,b, the number of nanohole arrays was changed from 9 (3 × 3) to Nano Lett., Vol. 8, No. 9, 2008

196 (14 × 14) on TSG substrates. The transmission intensity was proportional to the number of nanoholes (Figure 3a). The fwhm (full-width at half maximum) was found to be smaller when increasing the number of nanoholes, which is consistent with a previous report.5 For intensity interrogation used in this work, small fwhm (sharper resonance peaks) will be preferable for obtaining higher sensing sensitivity. According to this result, we suggest that, when comparing the sensitivity of different nanohole sensing systems based on intensity interrogation or wavelength interrogation, the size of nanohole arrays should be taken into consideration. For real-time sensing measurements, the monochromator was set to the wavelength around the S wavelengths of target nanohole arrays in Figure 2c. The bandpass was increased to 4 nm to improve the S/N ratio. More than 30 real-time sensing signals could be collected simultaneously. Figure 3c,d shows the real-time response from a 9 × 9 and a 3 × 3 array (with different d and P). When the refractive index varied from 1.3331 and 1.3702, both arrays exhibited a linear relation between intensity changes and the refractive index (insets in Figure 3c,d). For the 9 × 9 array, the highest intensity change was observed as a 37.5% increase for ∆n ) 0.0371 (DI water to 21% NaCl) at λ ) 590 nm; for the best 3 × 3 array, it was only 7.5% at best. Less intense transmission from the 3 × 3 array also leads to a low S/N ratio. Hence, we observe an inevitable tradeoff between the sensitivity and the size of the nanohole arrays. Nevertheless, 1010%/RIU sensitivity for the 9 × 9 array is fairly close to the regular grating coupler-based SPR sensors operated in the intensity interrogation mode,16 with the sensor size of nanohole arrays only 1/2000 of that used in commercial instruments (typical spot size ∼200 µm in diameter). Although the TSG surface offers better opportunity for more uniform immobilization of biomolecules, it also has a smaller surface area than the regular gold for immobilization (3% less, estimated by AFM). In order to demonstrate the capability of detecting biorecognition events, Glutathione S-Transferase (GST) and anti-GST antibody binding, a high 2721

Figure 3. (a) Transmission intensity at λ ) 568 nm (max. intensity) of nanohole arrays from 3 × 3 to 14 × 14 (d ) 150 nm, P ) 450 nm) on TSG substrates. (b) Full-width at half-maximum (fwhm) of nanohole arrays from 3 × 3 to 14 × 14. (c) Response to surface refractive index change from a 9 × 9 nanohole array (d ) 150 nm, P ) 450 nm), λ ) 590 nm. (d) Response to surface refractive index change from a 3 × 3 nanohole array (d ) 180 nm, P ) 419 nm), λ ) 590 nm. No noise reduction or smoothing algorithms were applied to c,d.

Figure 4. (a) Sensing response upon injecting 615 nM anti-GST and subsequently the regenerator. (b) Schematic illustration of surface chemistry on nanohole arrays (drawing not to scale). (c) Sensing response (black plot) of a 9 × 9 nanohole array (d ) 200 nm, P ) 540 nm) upon injecting three different concentrations of anti-GST. Red plot: fitting with a bimolecular reaction model. (d) Sensing response (black plot) of a 5 × 5 nanohole array (d ) 180 nm, P ) 419 nm) upon injecting 615 nM anti-GST and subsequently the regenerator. (e) Sensing response (black plot) of a 3 × 3 nanohole array (d ) 180 nm, P ) 419 nm) upon injecting 615 nM anti-GST and subsequently the regenerator. Measurements were all performed at λ ) 590 nm with bandpass ∼4 nm. No noise reduction or smoothing algorithms were applied. Signal change direction was inverted to meet the requirement of the fitting software.

affinity interaction that has been extensively used in Western blots and dot blots to detect GST-fusion proteins, was selected in this work (Figure 4b). We immobilized GST and used anti-GST as the target molecule since anti-GST is a bigger protein (MW ∼ 150 kD) that will induce more change in the local dielectric environment. The TSG surface was first functionalized with a thiol terminated with a primary amine. The GST proteins were then immobilized onto the gold surface with a cross-linker that covalently attached to the amino terminus of GST and thiol (see Supporting Information for details). Different concentrations of anti-GST 2722

antibodies prepared in PBS 7.4 buffer were introduced to the flow cell through a sample injector. After the binding event, 10 mM glycine (pH 2.2) was used to dissociate GST and anti-GST complex to regenerate the surface. In these experiments, we functionalized all of the nanohole arrays on the same substrate to demonstrate the sensing capability. For a practical multiplexed sensing, commercial AFM-based nanospotters28 may be used to functionalize individual arrays with different biomolecules. Figure 4a shows that the transmission intensity from a nanohole array immobilized with GST changed significantly Nano Lett., Vol. 8, No. 9, 2008

after 615 nM anti-GST was introduced. Subsequent stripping with 10 mM glycine (pH 2.2) successfully removed antiGST from the gold surface, and the signal baseline came back to the previous level. The reaction was fully reversible for at least 10 cycles. In Figure 4c, three different concentrations of anti-GST were injected in a random order. The red curves represent the best fit of the binding responses to a simple bimolecular reaction model (A + B T AB)29 using BIAevaluation (GE Health). The dissociation affinity constant (KD) of this specific assay was determined by commercially available BIACORE 3000 (GE Health) as 2.97 nM, which is typical for a high affinity binding. Although immobilized GST could bind with more than one anti-GST and the bimolecular model may not be the best model to describe this reaction pathway, the calculated association constant (ka) from nanohole arrays was 2.1 × 104 M-1 S-1, fairly close to 3.7 × 104 M-1 S-1 determined by BIACORE 3000 with dissociation and association constants fit separately. The dissociation constant (1.1 × 10-4 S-1, determined by BIACORE 3000) was too small to be accurately verified in this nanohole system because of the high affinity of this assay and the baseline drift. For smaller nanohole arrays (Figure 4d,e), although the S/N ratio was not as good as the 9 × 9 array, similar association constants (2.1 × 104 M-1 S-1 for 5 × 5 and 2.6 × 104 M-1 S-1 for 3 × 3) still can be determined by the fitting software. The detection limit (2σ) is around 10 nM for the 9 × 9 array. It should be mentioned that the measuring setup was designed to acquire multiple transmission spectra so highly intense and stable laser sources were not used. Low detection limits, though also highly desirable, are not the highest priority in this work. Although TSG substrates show better sensing performance than the regular thermally evaporated gold substrates, the transmission enhancement and performance improvement are far less than we expected from literature.21,23 The possible reasons are that the refractive index of FEP still does not exactly match the biological solutions, and there might be a small amount of air trapped between gold and FEP during the template-stripping process. Optimization of the TSG process with FEP is in progress. The protruded gold ring in the FEP layer caused by ion bombardment might have also played an important role. The sharp edge could contribute to some localized surface plasmon characters,9 which have been proven to markedly affect the transmission intensity. In summary, we present a plasmonic sensing platform that exploits small-dimension ordered arrays of nanoholes perforated on a super smooth gold surface that adhered on fluoropolymer substrate. It is capable of performing multiplexed and real-time binding affinity analysis on biorecognition events. The spectral study on 1∼20 µm2 nanohole arrays indicates that higher gold-solution contact area results in higher plasmonic sensing sensitivity. There is an inevitable tradeoff between the sensitivity and the size of the periodically arrayed nanoholes. Nevertheless, our result demonstrated that nanohole arrays with a dimension as small as 1 µm2 could be used to detect biomolecular binding events effectively and analyze the binding kinetics. TSG on FEP substrates provides a smooth environment for higher-quality Nano Lett., Vol. 8, No. 9, 2008

nanofabrication and more uniform immobilization. The refractive index matching between FEP and water solutions contributes to ∼20% improvement on the sensing performance. This nanohole approach presents a sophisticated platform for studies ranging from highly multiplexed protein binding screening to viral fusion and transmembrane protein analysis that require a high-quality supported lipid membrane. Acknowledgment. Funding for this work was providing by DARPA/MTO N/MEMS S&T Fundamentals Program and two NIH Grants, 5R01HG003828-04 and 5R21EB00433302. We thank Garland O’Connell for his instrumentation support and helpful discussion, Dane Stebbings and John Slusarz for making optical accessories. This work was performed in part at the Center for Nanoscale Systems (CNS), a member of the National Nanotechnology Infrastructure Network (NNIN), which is supported by the National Science Foundation under NSF Award No. ECS0335765. CNS is part of the Faculty of Arts and Sciences at Harvard University. Supporting Information Available: Detailed experimental procedures of TSG on FEP substrates, the GST/anti-GST assay, and image processing. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Cooper, M. A. Nat. ReV. Drug DiscoVery 2002, 1, 515–528. (2) Willets, K. A.; Van Duyne, R. P. Annu. ReV. Phys. Chem. 2007, 58, 267–297. (3) Stewart, M. E.; Anderton, C. R.; Thompson, L. B.; Maria, J.; Gray, S. K.; Rogers, J. A.; Nuzzo, R. G. Chem. ReV. 2008, 108, 494–521. (4) Ebbesen, T. W.; Lezec, H. J.; Ghaemi, H. F.; Thio, T.; Wolff, P. A. Nature 1998, 391, 667–669. (5) Lezec, H. J.; Thio, T. Opt. Express 2004, 12, 3629–3651. (6) Cao, Q.; Lalanne, P. Phys. ReV. Lett. 2002, 88. (7) van der Molen, K. L.; Klein Koerkamp, K. J.; Enoch, S.; Segerink, F. B.; van Hulst, N. F.; Kuipers, L. Phys. ReV. B 2005, 72. (8) De Leebeeck, A.; Kumar, L. K. S.; de Lange, V.; Sinton, D.; Gordon, R.; Brolo, A. G. Anal. Chem. 2007, 79, 4094–4100. (9) Lesuffleur, A.; Im, H.; Lindquist, N. C.; Oh, S. H. Appl. Phys. Lett. 2007, 90. (10) Tetz, K. A.; Pang, L.; Fainman, Y. Opt. Lett. 2006, 31, 1528–1530. (11) Brolo, A. G.; Gordon, R.; Leathem, B.; Kavanagh, K. L. Langmuir 2004, 20, 4813–4815. (12) Stark, P. R. H.; Halleck, A. E.; Larson, D. N. Methods 2005, 37, 37– 47. (13) Ji, J.; O’Connell, J. G.; Carter, D. J. D.; Larson, D. N. Anal. Chem. 2008, 80, 2491–2498. (14) Stewart, M. E.; Mack, N. H.; Malyarchuk, V.; Soares, J.; Lee, T. W.; Gray, S. K.; Nuzzo, R. G.; Rogers, J. A. Proc. Nat. Acad. Sci. U.S.A. 2006, 103, 17143–17148. (15) Henzie, J.; Lee, M. H.; Odom, T. W. Nat. Nanotechnol. 2007, 2, 549– 554. (16) Homola, J.; Yee, S. S.; Gauglitz, G. Sens. Actuators, B 1999, 54, 3– 15. (17) Genet, C.; Ebbesen, T. W. Nature 2007, 445, 39–46. (18) Naumann, R.; Schiller, S. M.; Giess, F.; Grohe, B.; Hartman, K. B.; Karcher, I.; Koper, I.; Lubben, J.; Vasilev, K.; Knoll, W. Langmuir 2003, 19, 5435–5443. (19) Jeuken, L. J. C.; Connell, S. D.; Henderson, P. J. F.; Gennis, R. B.; Evans, S. D.; Bushby, R. J. J. Am. Chem. Soc. 2006, 128, 1711–1716. (20) Jeuken, L. J. C.; Connell, S. D.; Nurnabi, M.; O’Reilly, J.; Henderson, P. J. F.; Evans, S. D.; Bushby, R. J. Langmuir 2005, 21, 1481–1488. (21) Dintinger, J.; Degiron, A.; Ebbesen, T. W. MRS Bull. 2005, 30, 381– 384. (22) Martin-Moreno, L.; Garcia-Vidal, F. J.; Lezec, H. J.; Pellerin, K. M.; Thio, T.; Pendry, J. B.; Ebbesen, T. W. Phys. ReV. Lett. 2001, 86, 1114–1117. 2723

(23) Krishnan, A.; Thio, T.; Kima, T. J.; Lezec, H. J.; Ebbesen, T. W.; Wolff, P. A.; Pendry, J.; Martin-Moreno, L.; Garcia-Vidal, F. J. Opt. Commun. 2001, 200, 1–7. (24) Liu, G. L.; Doll, J. C.; Lee, L. P. Opt. Express 2005, 13, 8520–8525. (25) van der Molen, K. L.; Segerink, F. B.; van Hulst, N. F.; Kuipers, L. Appl. Phys. Lett. 2004, 85, 4316–4318. (26) Wang, G. P.; Yi, Y. X.; Wang, B. J. Phys.: Condens. Matter 2003, 15, 8147–8156.

2724

(27) Thio, T.; Ghaemi, H. F.; Lezec, H. J.; Wolff, P. A.; Ebbesen, T. W. J. Opt. Soc. Am. B: Opt. Phys. 1999, 16, 1743–1748. (28) For example, Bioforce Nanosciences, www.bioforcenano.com. (29) Quinn, J. G.; O’Kennedy, R. Anal. Biochem. 2001, 290, 36–46.

NL801043T

Nano Lett., Vol. 8, No. 9, 2008