Novel Gold-Capped Nanopillars Imprinted on a Polymer Film for

Jun 1, 2012 - Figure 3. SEM images of nanopillars, which were nanoimprinted on COP substrate. .... After the optimizing step, different concentrations...
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Novel Gold-Capped Nanopillars Imprinted on a Polymer Film for Highly Sensitive Plasmonic Biosensing Masato Saito,*,† Akito Kitamura,† Mizuho Murahashi,† Keiichiro Yamanaka,‡ Le Quynh Hoa,† Yoshinori Yamaguchi,‡ and Eiichi Tamiya†,‡ †

Department of Applied Physics, Graduate School of Engineering, Osaka University, 2-1 Yamadaoka, Suita, Osaka, Japan 565-0871 Photonics Advanced Research Center (PARC), Graduate School of Engineering, Osaka University, 2-1 Yamadaoka, Suita, Osaka, Japan 565-0871

‡

S Supporting Information *

ABSTRACT: Herein, a nanoporous alumina was fabricated to use as a mold in transforming nanopillar structures onto a thin film polymer by thermal nanoimprint lithography (NIL). The size of the pores was successfully controlled by varying the applied voltages and etching time. These nanoporous structures were transferred to the Cyclo-olefin polymer (COP) film surface from the porous mold by a thermal nanoimprinting process. A plasmonic substrate was fabricated by sputtering a thin layer of gold onto this nanopillar polymer structure, and the refractive index response in a variety of media was evaluated. Finally, the biosensing capacity of this novel plasmonic substrate was verified by analysis of Human immunoglobulin and achieved a minimum detection limit of 1.0 ng/ mL. With the advantages of mass production with consistent reproducibility stemming from the nanoimprint fabrication process, our gold-capped polymeric pillars are ready for the transition from academic interest into commercialization systems for practical use in diagnostic applications.

N

alumina (PAA), has a highly dense honeycomb nanostructure on the oxide surface, making it an attractive potential replacement. Thorough investigation of AAO undertaken by H. Masuda revealed a crucial relationship between the pore size, depth, structure, and so forth of synthesized AAO and anodizing conditions, further demonstrating its applicability as an antireflection surface and as a mold for polymeric photonic crystal imprinting fabrication.10,11 Thus, we propose AAO as a nanostructure array mold for NIL to fabricate the biosensing substrates to supplement limitations from the NIL mold fabrication process. We believe that the low cost, large area, size tuning capability, and ease of the anodizing process may enhance the production and marketability of NIL technology. Recently, a biosensing substrate with localized surface plasmon resonance (LSPR) for biosensing application was fabricated by immobilizing gold nanoparticles (Au-NPs) on the surface of an AAO layer formed on an aluminum (Al) substrate.12,13 Biosensing with LSPR has the merit of label-free measurement with a highly sensitive signal at long wavelengths. It is widely known that the collective charge density oscillations of nanoparticles are defined as localized surface plasmon resonance. LSPR, in turn, enhances an immediate change in the interfacial refractive index (RI) of the surrounding medium;

anoimprint lithography (NIL) has increasingly been recognized as a key manufacturing nanotechnology that plays a critical role in the future commercialization of nanodevices. In 1995, nanoimprinting lithography was proposed by S. Y. Chou and his students.1,2 Using NIL, one can create a nanometer pattern with sub-10 nm resolution in feature size and with critical dimension control3 at low cost, high throughput, large area, and excellent reproducibility.4 As a result, NIL has been extended to a variety of devices, such as single electron transistors,5 highly integrated magnetic memory disks,6 and optical devices.7,8 One of the outstanding advantages of NIL is that it can be applied in flexible electronics, e.g., fabrication of nanostructured devices on polymer film substrates, which is not possible with conventional lithography techniques. Thus, point-of-care testing (POCT) applications (e.g., biosensors, biochips) might be one of the most promising fields that benefits from this advanced technique. By controlling the polymer nature, we can expect the desired optical functionality in a polymer film that has a nanostructure with a pitch close to the optical wavelength.9 Although NIL has great potential as a high throughput nanofabrication technique, the mold for imprinting is fabricated through time-consuming electron beam lithography (EBL) followed by dry etching. Furthermore, the mold is expensive, is difficult to scale up, and has low throughput, thus limiting the technical applications of NIL. Meanwhile, self-ordered anodic alumina oxide (AAO), also referred to as porous anodic © 2012 American Chemical Society

Received: October 11, 2011 Accepted: June 1, 2012 Published: June 1, 2012 5494

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thus, the signal is greatly affected by the attachment of biomolecules. The excitation of LSPR by light at an incident wavelength, where resonance occurs, results in the appearance of intense surface plasmon (SP) absorption bands. The intensity and position of the SP absorption bands are characteristic to the type of nanomaterial, the diameter of the nanoparticles, and their distribution. Thus, for the development of practical LSPR biosensors, a reliable mass nanofabrication method for noble-metal nanoparticles is one of the key challenges. Generally, such nanoparticles are fabricated in colloidal form by the reduction of silver or gold ion solutions. This fabrication method feasibly produces noble-metal nanoparticles with low size dispersion.14−17 In all of our previous studies,12,13 we utilized presynthesized Au-NPs (50 nm in diameter, British Biocell, U.K.) to directly deposit on AAO substrates. However, by that method, Au-NPs could not be uniformly distributed on AAO substrates. It was observed that gold nanoparticles are trapped on top of many of the honeycomb-hole structures, but not all. Understanding that this limitation strongly inhibits the future commercialization process of Au-based biosensing devices, we now aim at fabricating uniformly distributed Au-NPs on a plasmon substrate that can enhance the signal-to-noise ratio by reducing the nonspecific binding area via densifying the measurement spots. Herein we report the fabrication of a plasmon polymeric nanopillar-structured substrate using NIL with a porous AAO mold. The nanopillar pattern was successfully transferred from the porous structure of the AAO mold to the polymer substrate during the NIL process. After that, to create a plasmon phenomenon, Au was deposited by sputtering to form a gold “cap” on top of each polymer pillar (hereon abbreviated as “Aucapped nanopillar”). We then investigated the optical properties of the synthesized plasmon chip and evaluated the biomolecular interactions on the Au-capped polymer nanopillar. A noteworthy achievement of our study is the mass production of a high density gold-based plasmon flexible chip for biosensing applications, paving the way forward to the commercialization of low-cost, high performance biosensors.

etching solution for 5 min. This fabricated mold will be denoted by PA-mold40. The second type of mold, which has larger pore size and pitch, was fabricated at a constant voltage of 80 V for 60 min at the first anodizing step, followed by a second anodizing at 60 V for 48 s and 12.5 min etching time. This fabricated mold will be denoted by PA-mold80. All samples were carefully rinsed with ultrapure water and underwent ultrasonication at each step, and finally dried completely by pure N2 gas. The morphologies of PA-mold40 and 80 were imaged using a commercial atomic force microscopy (AFM) unit (SPA400SPI4000, Seiko Instruments Inc.) equipped with a calibrated 20 μm xy-scan and 10 μm z-scan range PZT-scanner. All AFM images were taken in dynamic force mode (DFM mode, i.e. tapping mode) at optimal force. A silicon cantilever (OMCLAC160TS, OLYMPUS), which has a spring constant of 42 N/ m and a frequency resonance of 300 kHz, was also used for imaging in air at room temperature. Fabrication of Au-Capped Nanopillar Structured Polymer Film. Thermal nanoimprinting was performed with X-300H (SCIVAX Corp.). First, the PA molds were immersed in a mold release agent overnight (1% fluorocarbon compound (OPTOOL, DAIKIN Industries Ltd.) diluted by perfluorohexane (Demnumsolvent, DAIKIN Industries Ltd.). After setting the PA-mold and Cyclo-olefin polymer (COP) film onto the stage, a pressure of 0.83 MPa was applied for 1 min at 100 °C. Temperature was then increased to 160 °C (the glass-transition temperature of COP) while the pressure was maintained at 0.83 MPa. Next, the pressure was increased to 2 MPa while maintaining the temperature of 160 °C. After 10 min, the temperature was dropped to 100 °C and the pressure was released. Transformed nanopillar structures from PA-mold40 and PA-mold80 are denoted by NP40 and NP80, respectively. To estimate the reproducibility of nanopillar substrates, 20 nanopillar substrates were transferred from one PA-mold. The Au-capped nanopillar structure was formed by the deposition of between 24 and 96 nm of Au on the imprinted COP surface by sputtering (ACS4000, ULVAC) for 1 to 4 cycles at 10 min per cycle. To improve the wettability between the polymer substrate and the solution, the substrate surface was modified to be hydrophilic by an oxygen plasma treatment (PDC200, Yamato Scientific Co. Ltd.), at 75 mL/min O2 gas flow and 75 W for 10 s. Characterization of Au-Capped Nanopillar Structured Polymer Film. The Au-capped nanopillar structure on the COP surface, which was treated with osmium by an osumium coater (Neoc-ST, Meiwafosis Co. Ltd.), was imaged using a scanning electron microscope (SEM) (DB 235, FEI Company). SEM images were analyzed to obtain the distribution of nanopillar diameters based on an ellipsoidal fitting. The boundary of each nanopillar was determined by the segmentation of the SEM images at a certain threshold intensity, and the binarized SEM images were rendered to group the individual nanopillars. The x−y coordinates for the boundary of each nanopillar were assigned on the basis of the pixel coordinates of binarized individual images. An ellipse approximation was performed to set the x−y coordinate for the boundary of each nanopillar. The major axes (a) and minor axes (b) were obtained for every individual nanopillar using the least-squares approximation method. The least-squares approximation for the ellipsoidal fitting was performed on the basis of the following equation:

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MATERIALS AND METHODS Fabrication of Porous Alumina Mold. It is widely known that the pore size and pitch can be controlled by changing the anodizing condition. However, in our study, to investigate the size effect of NIL molds on the performance of a plasmon substrate, we fabricated two representative types of porous alumina (PA) molds: small and large pore size and pitch. These molds were fabricated by a two-step anodizing method that we developed.18 The first anodizing step was conducted under a constant voltage of 40 V in a 0.3 M aqueous oxalic acid solution for 60 min to generate an aluminum oxide layer on the mirrorlike polished aluminum plate. The electrolyte solution was vigorously stirred during the process to maintain a steady temperature and electrolyte concentration. The anodic temperature was kept at 273 K. The generated aluminum oxide layer was removed by immersing the substrate in an aqueous solution containing phosphoric acid (1.16%, w/v) and chromic acid (5%, w/v) at 333−353 K. After removing the aluminum oxide layer, a concave pattern was obtained on the surface of the aluminum plate. The second anodizing step for generating a PA layer on the concave pattern was carried out at a constant voltage of 40 V at 273 K. To increase pore size, the aluminum plate with the PA layer was immersed in a 0.23 M H3PO4 5495

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of SAM, followed by washing with phosphate buffered saline (PBS, pH 7.4) and drying with pure N2 gas. A 100 μg/mL concentration of goat anti-human-immunoglobulin G (IgG, BETHYL Laboratories, Inc.) in PBS was added to the substrate and incubated for 30 min followed by washing with PBS containing 0.05% Tween-20 (PBST) and drying. After the antibody was immobilized on the surface of Au-capped nanopillar substrate, a solution of 1% bovine serum albumin (BSA, Sigma-Aldrich Inc.) in PBS was used to block the nonspecific adsorption, followed by thorough washing with PBST and drying at room temperature. Finally, different concentrations of antigen Human IgG (Sigma-Aldrich) solutions (∼0 to 100 μg/mL) were introduced onto the system for 30 min, followed by washing with PBST and drying, and the optical characteristics were subsequently analyzed. The absorption spectrum of each step was measured in air. A 1 μg/ mL concentration of a C-reactive protein (CRP, Sigma-Aldrich) was used as the negative control sample.

major axis(a) = ((X 0 sin θ + Y0 sin θ )2 − E cos2 θ − {(X 0 sin θ − Y0 cos θ)2 − E sin 2 θ } 1

sin 2 θ − B cos2 θ ⎞ 2 ⎟ cos 2 θ − B sin 2 θ ⎠

(1)

minor axis(b) = ((X0 sin θ − Y0 cos θ )2 − E sin 2 θ − {(X 0 cos θ + Y0 sin θ)2 − E cos2 θ} 1

cos2 θ − B sin 2 θ ⎞ 2 ⎟ sin 2 θ − B cos2 θ ⎠

(2)

where:

( 1 −A B )

tan−1 θ=

■

(3)

2

X0 =

AD − 2BC 4B − A2

Y0 =

AC − 2D 4B − A2

RESULTS AND DISCUSSION Nanopillar Structure Formation Using Thermal Nanoimprinting. The fabrication of the Au-capped nanopillar polymer plasmon substrate is schematically illustrated in Figure 1. In this study, we utilized our two step anodizing method to

(4)

(5)

and ⎛∑ X 2 Y 2 i i ⎜ ⎜ i ⎜ 3 ⎛ A ⎞ ⎜∑ XiYi ⎜ ⎟ ⎜ i ⎜B ⎟ ⎜ 2 ⎜C ⎟ = ⎜∑ Xi Yi ⎜ ⎟ ⎜ i ⎜⎜ D ⎟⎟ ⎜ 2 ⎝ E ⎠ ⎜∑ XiYi ⎜ i ⎜ ⎜∑ XiYi ⎝ i

∑ XiYi3 ∑ Xi2Yi ∑ XiYi2 ∑ XiYi ⎞⎟

−1

i

∑

i

Y i4

i

∑ XiYi2 i

∑ Yi3 i

∑ Yi2

⎛− ∑ X 3Y ⎞ i i ⎟ ⎜ ⎜ i ⎟ ⎜ − X 2Y 2 ⎟ ∑ i i ⎟ ⎜ ⎜ i ⎟ ⎜ 3 ⎟ − X ∑ i ⎜ ⎟ ⎜ i ⎟ ⎜ 2 ⎟ ⎜− ∑ Xi Yi ⎟ ⎜ i ⎟ ⎜ 2 ⎟ ⎜ − ∑ Xi ⎟ ⎝ i ⎠

i

∑

i

XiYi2

i

∑ Xi2 i

∑ i

∑ XiYi i

∑ XiYi ∑ Yi2 i

∑ Xi i

i

∑ Yi i

⎟ ⎟ ⎟ i ⎟ X ∑ i ⎟⎟ i ⎟ ⎟ ∑ Yi ⎟ i ⎟ ⎟ ∑1 ⎟ ⎠ i i

Yi3

∑ Yi2

Figure 1. The schematic illustration of the experimental steps for the fabrication of LSPR substrate by nanoimprinting. (a) Fabrication of porous alumina mold. (b) Thermal nanoimprinting with high temperature and pressure to the COP resin. (c) Formation of the nanopillar structure by releasing from the alumina porous mold. (d) Au-capped nanopillar structure for the LSPR substrate. (6)

fabricate appropriate nanohole structured molds for the imprinting step. The key parameters for being an optimum imprinting mold are the pore size and pitch because they later define the diameter and the height, respectively, of the nanopillars on COP. The pore size and pitch of PA, in turn, can be controlled by varying applied anodizing voltage.19 By applying a high anodizing voltage, we succeed in obtaining a suitable pitch for our porous structure, but the pore diameter was not large enough to transform the pillared structure onto a polymer film when used as the imprinting mold. Furthermore, as depicted in Figure 1, the size of the gold caps is directly determined by the diameter of the nanopillars. Thus, if the pore size of PA mold is small, it creates thin nanopillars, and goldcapped thin nanopillars cannot produce a significant absorbance signal (data not shown). The reason is that the Au-covered area on the bottom layer of the substrate was larger

The absorbance spectrum was measured using UV-3600 (Shimadzu). The Au-capped nanopillar substrate was characterized under several different conditions, i.e., air, water, 1 M glucose, ethylene glycol, and glycerol. To demonstrate the plasmon biosensing applicability of our successfully fabricated Au-capped nanopillar substrate, an antibody−antigen reaction measurement was performed. The Au-capped nanopillar substrate was first interacted with 10 μL of 1 mM 10carboxyl-1-decanethiol (C10, Dojindo Inc., Japan) in 20 mL of ethanol (99.95%, Wako, Japan) and to form the self-assembled monolayer (SAM) after 30 min. Next, SAM functionalization was carried out by introducing 10 μL of 0.1 M Nhydroxysuccinimide (NHS, Dojindo Inc., Japan) and 0.4 M 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide.HCl (EDC, Dojindo Inc., Japan) for 30 min to activate the carboxyl groups 5496

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than that of the capped area on the top of the pillars. Thus, the absorbance signal from the Au-capped nanopillars would be overlapped by that of the bottomed layer of Au. To solve this problem, we etched the porous alumina substrate using phosphoric acid to expand the inner diameter to the desired value. Thus, we were able to successfully control pore size by varying the etching time as shown in Figure SI1. Figure 2 shows AFM images of fabricated porous alumina mold surfaces. During the fabrication process, we optimized

Figure 3. SEM images of nanopillars, which were nanoimprinted on COP substrate. (a) Nanopillar structure NP40 was transformed from the PA-mold40, which was made by 40 V anodizing and H3PO4 etching for 5 min. (b) Nanopillar structure NP80 was transformed from the PA-mold80, which was made by 80 V anodizing and 12.5 min H3PO4 etching.

Figure 2. AFM images of porous alumina (PA) mold. (a) PA-mold40 was formed by the second anodizing for 4 min and H3PO4 etching for 7.5 min. (b) PA-mold80 was formed by the second anodizing for 48 s and H3PO4 etching for 12.5 min. Line (*) and (**) in the AFM images indicate lineprofiles. Porous pitch of PAmold40 and PAmold80 were about 116.7 and 152.0 nm each.

etching time to obtain a pitch of less than 200 nm so that the nanopillars derived from the corresponding PA mold were highly dense and “standable” for capping with gold in following sputtering step. Figure SI2 clearly demonstrates the collapse of thin nanopillars at heights of 277 and 1000 nm. From the line profiles on the AFM images, the porous pitch of PA-mold40 is approximately 116.7 nm. In the case of PA-mold80, the porous pitch is about 152.0 nm, indicating successful size control achieved by applying a higher voltage than that of PA-mold40. Structural Characterization of Au-Capped Nanopillar Polymer Substrate. To transform the nanopillar structure onto the COP film from the PA-molds, we performed a thermal nanoimprinting at optimized condition of 160 °C and 2 MPa. Figure 3 shows SEM images of the nanopillar structures after coating with a 50 nm layer of Au via sputtering. The inset of each image clearly displays the highly dense and independently standing pillars, each with a gold “cap”. Transformed nanopillar structures from PA-mold40 and PA-mold80 are denoted by NP40 and NP80, respectively. It was observed from Figure 3A,B that the density and size of the gold caps on NP40 are lower than those of NP80 due to the original structures of PAmold40 and 80, respectively. To quantitatively determine the surface area and the distribution of gold for each case, we performed an ellipsoidal fitting as described in the Materials and Methods section and Figure 4. Major axis a and minor axis b were obtained from the ellipsoid fitting as shown in Figure 4a. The diameter of the nanopillar was measured from (a + b)/2 and distributed at 10.0 nm intervals (Figure 4b). The diameter distribution of NP80

Figure 4. Diameter distribution of Au-capped nanopillar structures (b). After the ellipsoidal fitting, the major axis a and minor axis b were obtained (a). The diameter of each nanopillar was calculated by (a + b)/2.

was centered around 60.0−69.9 nm, which is approximately double than that of NP40 (30.0−39.9 nm). These results indicated that the nanopillar size was successfully controlled and transformed from alumina porous molds’ size via the anodizing and imprinting steps. Furthermore, the fraction of gold-covered area over the whole geometric area was easily calculated from the axis after the ellipsoidal fitting (Figure 4a). 5497

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silica core and Au-capped shell, and observed the same redshift phenomenon with increasing silica size at a constant Au thickness.23 There was also a similar redshift of LSPR peaks with incremental increasing of the Au nanodisk diameter.22 Thus, it suggested that our Au-capped nanopillar structure exhibits plasmon properties similar to a Au nanodisk22 rather than a Au nanosphere whose LSPR peak positions are reported to be independent of their dimensions. Furthermore, to confirm the ability of our nanopillar substrate as a plasmonic sensor we evaluated the sensitivity of LSPR to the refractive index of various surrounding environments using these Au-capped NP40 and NP80 structures. Five surrounding environments with different refractive indexes were investigated: air (n = 1.0), water (n = 1.33), 1 M glucose (n = 1.35), ethylene glycol (n = 1.43), and glycerol (n = 1.47). The results showed that the LSPR peak positions of both nanopillar substrates red-shifted and their peak intensity linearly increased with the increase of the surroundings’ refractive index. Moreover, a bigger core-sized nanopillar resulted in a greater increase, as clearly shown in Figure 6C. On the basis of the slope of the fitted lines, the Au-capped NP80 exhibits higher sensitivity compared with that of NP40 due to its larger nanopillar diameter.24 From the obtained data, it is evident that the highly sensitive capacity of LSPR substrates was achieved by tuning the dimensions of the Au-capped nanopillar nanostructures. LSPR Au-Capped Nanopillar Biosensor with an IgG/ anti-IgG Reaction. The prospect of highly sensitive Aucapped nanopillar structures encouraged us to investigate their biosensing capacity with an IgG−anti-IgG interaction model (Figure 7). The Au-capped nanopillar substrate was first interacted with C10 to form SAM. Next, the NHS-EDCactivated SAM was used for antibody immobilization. After the antibody was immobilized on the surface of the Au-capped nanopillar substrate, BSA was used to effectively block nonspecific adsorption,25−28 followed by thorough washing with PBST. At this stage, one of the most important parameters was BSA concentration because it strongly effected the differentiation between positive and negative samples. For our own designed Au-capped nanopillar detection system, various BSA concentrations (∼0.01%−1%) were studied to reveal the most optimized value. It is predictable that the detection capacity decreases when the BSA concentration is lower than 0.1% because there is not enough BSA to cover all nonspecific adsorption sites, leading to poor differentiation between the positive and negative samples. On the other hand, when the BSA concentration is as high as 1%, the error bar is too large in comparison with the measured value. We attributed this severely scattered data to the changes in refractive index of the medium surrounding substrate surface. Due to the overloading of BSA, the PBS was not able to wash away the excess BSA completely, leading to unstable data for each measurement due to the errors resulting from the different refractive index values of PBS and BSA. We concluded that 0.1% BSA is the most optimized value for our Au-capped nanopillar detecting system. After the optimizing step, different concentrations of IgG solutions (∼0 to 100 μg/mL) were introduced onto the antiIgG immobilized Au-capped nanopillar system, and their optical characteristics were subsequently evaluated to examine detection limit by making a standard curve. As shown in Figure 8, the change in absorbance strength was recorded depending on the concentration of the antigen IgG. The wide detection

This fraction was 54.3% in the case of NP40, while it was enlarged and reached 67.0% in the case of NP80. This Au coverage was dramatically improved compared to our previous report,20 in which Au nanoparticles covered only 32.1% of the silicon oxide surface (in terms of a volume fraction of 21.4%). Fujiwara et al. have reported that the increasing absorption intensity is related to the increase of Au nanoparticle density.21 Thus, the enhancement of gold density not only helps to miniaturize the structure of the plasmonic biosensor, which is extremely important for application as point-of-care devices, but also contributes to improving the signal-to-noise ratio of the whole testing system. These results therefore have direct implications on the development of commercialized LSPR sensors by reducing size and enhancing sensitivity. Effect of Au Thickness on Optical Characteristics of the Au-Capped Nanopillars. The optical properties of Aucapped nanopillar structures and the effect of Au-cap thickness were studied by examining the absorbance spectra. To fabricate different Au-cap thicknesses, we sputtered Au on the NP80 layer-by-layer (1−4 times) to obtain thicknesses from 24 to 96 nm (the average Au-cap thickness after one sputtering cycle, or 10 min, was 24 nm). As experimentally shown in Figure 5, with

Figure 5. Feature absorbance spectra of “Au cap” structured NP80 with various Au thickness (a) and the corresponding peak positions calculated from the repeated experiments (b).

pillar size held constant, as the Au-cap thickness increases a blue-shift of the peak position can be clearly observed. The peak position of 24 nm-thick Au-cap was initially observed at 637.7 nm. After the second sputtering to obtain a 48 nm in thickness, the peak was positioned at a shorter wavelength (576.7 nm). Our results are in agreement with the computational model of Zheng et al. in which they observed the dimension-dependence LSPR characteristics of a Au nanodisk.22 These results also suggest that the LSPR peak could be red-shifted by reducing the thickness of the Au-cap via tuning the sputtering time. Effect of Nanopillar Size and Surrounding Refractive Index on Optical Characteristics of the Au-Capped Nanopillar. Unlike the above-mentioned thickness-dependent LSPR characteristic of Au-capped nanopillars, the opposite trend was observed upon changing the pillar size. In comparison with Au-capped NP40, the Au-capped NP80 displayed red-shifted LSPR peaks. For instance, the peak position in air of Au-capped NP40 was observed at 556.5 nm (Figure 6a), while it shifted to 575 nm in the case of Au-capped NP80 (Figure 6b). This strongly confirmed the dimensiondependent optical characteristic of Au-capped nanopillars, since not only thickness but also the diameter of the Au-cap directly influences the peak positions. In our previous studies, we reported a similar type of core−shell nanoparticle which had a 5498

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Figure 6. Absorption spectrum of Au-capped nanopillars of (a) NP40 and (b) NP80, when they were surrounded by air (n = 1.0), water (n = 1.33), 1 M glucose (n = 1.35), ethylene glycol (n = 1.43), and glycerol (n = 1.47). (c) The RI value dependence of peak shifts for NP40 (●) and NP80 (■).

Figure 8. Calibration curve for the IgG−anti IgG measurement using Au-capped nanopillar LSPR based-chip. N is 1 μg/mL of CRP as a negative control.

Figure 7. Optimization of BSA concentration as blocking agent for IgG−anti IgG detection.

range of our nanochip was linear up to 100 μg/mL, and the detection limit was down to 1.0 ng/mL, which is equal to 6.7 pM. For similar IgG reactions, Mayer et al.14 previously reported the detection limit of their Au-nanorod sensor was only about 1 nM. Thus, our novel Au-capped nanopillar flexible polymer chip evidently demonstrated a highly sensitive sensing capability along with its low-cost and the ease of its massproduction fabrication process.

sensitivity, further optimization steps for our device on the parameters of densities of the alumina porous mold and imprinted nanopillars are necessary. Although there is much more optimization work to be done, the label-free sensing Aucapped nanopillar structure proposed in this work is already quite suitable toward industrial and commercial application. Future work will involve dynamic range characterization, multispot array, kinetic constant calculations, and combination with micro-TAS device estimations of our biosensor for other biomolecules, such as various types of antigen−antibody interactions and DNA hybridizations.

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CONCLUSIONS In conclusion, we have developed a novel Au-capped nanopillar plasmonic biosensor based on the combination of porous alumina fabrication and thermal nanoimprinting techniques. The unique structure of our Au-capped nanopillar substrate resulted in enhanced sensitivity and binding capacities of biomolecules onto the gold surface. To achieve maximum

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ASSOCIATED CONTENT

S Supporting Information *

Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org. 5499

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AUTHOR INFORMATION

Corresponding Author

*Phone: +81-6-6879-4087. Fax: +81-6-6879-7840. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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REFERENCES

(1) Chou, S. Y.; Krauss, P. R.; Renstrom, P. J. Appl. Phys. Lett. 1995, 67, 3114−3116. (2) Chou, S. Y.; Krauss, P. R.; Renstrom, P. J. J. Vac. Sci. Technol., B 1996, 14, 4129−4133. (3) Buyukserin, F.; Aryal, M.; Gao, J.; Hu, W. Small 2009, 5, 1632− 1634. (4) Chou, S. Y.; Krauss, P. R.; Zhang, W.; Guo, L.; Zhuang, L. J. Vac. Sci. Technol., B 1997, 15, 2897−2904. (5) Fabrication and characterization of room temperature silicon single electron memory. Guo, L.; Leobandung, E.; Zhuang, L.; Chou, S. Y. J. Vac. Sci. Technol., B 1997, 15, 2840−2843. (6) Wu, W.; Cui, B.; Sun, X.-Y.; Zhang, W.; Zhuang, L.; Kong, L.; Chou, S. Y. J. Vac. Sci. Technol., B 1998, 16, 3825−3829. (7) Wang, J.; Schablitsky, S.; Yu, Z.; Wu, W.; Chou, S. Y. J. Vac. Sci. Technol., B 1999, 17, 2957−2960. (8) Li, M.; Wang, J.; Zhuang, L.; Chou, S. Y. Appl. Phys. Lett. 2000, 76, 673−675. (9) Yokoo, A.; Wada, K.; Kimerling, L. C. Jpn. J. Appl. Phys. 2007, 46 (9B), 6395−6397. (10) Yanagishita, T.; Nishio, K.; Masuda, H. Jpn. J. Appl. Phys. 2010, 49, 065202. (11) Yanagishita, T.; Nishio, K.; Masuda, H. J. Vac. Sci. Technol., B 2010, 28, 398−400. (12) Hiep, H. M.; Yoshikawa, H.; Tamiya, E. Anal. Chem. 2010, 82, 1221−1227. (13) Hiep, H. M.; Yoshikawa, H.; Taniyama, S.; Hoa, L. Q.; Kondoh, K.; Saito, M.; Tamiya, E. Jpn. J. Appl. Phys. 2010, 49, 06GM02. (14) Mayer, K. M.; Lee, S.; Liao, H.; Rostro, B. C.; Fuentes, A.; Scully, P. T.; Nehl, C. L.; Hafner, J. H. ACS Nano 2008, 2, 687−692. (15) Nath, N.; Chilkoti, A. Anal. Chem. 2002, 74, 504−509. (16) Nath, N.; Chilkoti, A. Anal. Chem. 2004, 76, 5370−5378. (17) Frederix, F.; Friedt, J.-M.; Choi, K.-H.; Laureyn, W.; Campitelli, A.; Mondelaers, D.; Maes, G.; Borghs, G. Anal. Chem. 2003, 75, 6894− 6900. (18) Kim, D. K.; Kerman, K.; Saito, M.; Sathuluri, R. R.; Endo, T.; Yamamura, S.; Kwon, Y. S.; Tamiya, E. Anal. Chem. 2007, 79 (5), 1855−1864. (19) Kim, D. K.; Kerman, K.; Ha, M. H.; Saito, M.; Yamamura, S.; Takamura, Y.; Kwon, Y. S.; Tamiya, E. Anal. Biochem. 2008, 379 (1), 1−7. (20) Hiep, H. M.; Yoshikawa, H.; Saito, M.; Tamiya, E. ACS Nano 2009, 3, 446−452. (21) Fujiwara, K.; Kasaya, H.; Ogawa, N. Anal. Sci. 2009, 25, 241− 248. (22) Zheng, Y. B.; Juluri, B. K.; Mao, X.; Walker, T. R.; Huang, T. J. J. Appl. Phys. 2008, 103, 014308. (23) Ha, M. H.; Endo, T.; Saito, M.; Chikae, M.; Kim, D. K.; Yamamura, S.; Takamura, Y.; Tamiya, E. Anal. Chem. 2008, 80 (6), 1859−1864. (24) Sekhon, J. S.; Verma, S. S. Plasmonics 2011, 6, 311−317. (25) Lahav, M.; Vaskevich, A.; Rubinstein, I. Langmuir 2004, 20, 7365−7367. (26) Huang, T. T.; Sturgis, J.; Gomez, R.; Geng, T.; Bashir, R.; Bhunia, A. K.; Robinson, J. P.; Ladisch, M. R. Biotechnol. Bioeng. 2003, 81, 618−624. (27) Peterfi, Z.; Kocsis, B. J. Immunoassay 2000, 21, 341−354. (28) Paulsson, M.; Kober, M.; Freij-Larsson, C.; Sollenwerk, M.; Wesslen, B.; Ljungh, A. Biomaterials 1993, 14, 845−853.

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