Article pubs.acs.org/ac
Electrochemical Surface Plasmon Resonance Measurement Based on Gold Nanohole Array Fabricated by Nanoimprinting Technique Kohei Nakamoto,†,‡ Ryoji Kurita,† and Osamu Niwa*,†,‡ †
National Institute of Advanced Industrial Science and Technology (AIST), Central 6, 1-1-1 Higashi, Tsukuba 305-8566, Japan Institute of Materials Science, Graduate School of Pure and Applied Sciences, University of Tsukuba, 1-1-1 Tennodai, Tsukuba 305-8573, Japan
‡
S Supporting Information *
ABSTRACT: In this paper, we describe our development of an electrochemical surface plasmon resonance (EC-SPR) measurement device based on a bottom-filled gold nanohole array. The polymer based gold nanohole array was fabricated with a UV nanoimprint technique and electron beam gold deposition. Direct reflection mode measurement was used to monitor the SPR dip in the reflection spectra. A cyclic voltammogram was also operated by using the standard three electrodes containing working electrode having a gold nanohole array and counter and reference electrodes. The gold nanohole array was modified with an osmium-poly(vinylpyridine)-wired horseradish peroxidase (Os-gel-HRP) film, and its redox state induced by the change in potential was monitored simultaneously. The redox state of the local film was obtained simply by scanning the sample substrate stage. The substrate modified with Os-gel-HRP film was incorporated in a microfluidic chip, and then the hydrogen peroxide was determined in terms of the redox change in the Os complex mediator from the slope of the SPR dip shift. The linear relation of hydrogen peroxide from 10 to 250 μM was successfully monitored, and a high conversion efficiency was realized.
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is that they offer the possibility of obtaining large amounts of optical information or local 2D imaging on a substrate simply by scanning the irradiation area. In regards to 2D EC-SPR using a conventional Kretschmann configuration, one of the authors has reported the measurement of enzymatic reactions by converting a substrate reaction into the redox change of an Ospolymer based mediator modified on the gold film.12 This type of measurement can image enzymatic activity by using 2D-SPR equipment13 combined with the control of the potential of the gold film surface, although an enzymatic reaction is difficult to measure with SPR due to the short lifetime of complex state between enzyme and the substrate, which are different from those of DNA hybridization and an immune reaction. In fact, one of the authors has also demonstrated the imaging of glucose detection by detecting the H2O2 reduction induced by HRP, which caused a redox state change in the Os-polymer film.14 However, 2D SPR imaging requires more complicated optical equipment such as a polarizer and a collimator to obtain a parallel light beam, a prism, and a 2D CCD device. Improved spatial resolution is also required by using a highresolution CCD device and precisely collimating the incident light. Therefore, the EC-SPR based multiple biosensing method
anostructured plasmonic materials have attracted much attention in the field of biological and chemical sensing.1 Gold or silver nanoparticles,2 nanorods,3 nanoshells,4 and periodic nanopost5 or nanohole arrays6 have been studied for localized surface plasmon resonance (LSPR) or extraordinary optical transmission (EOT)7 devices. Despite being less sensitive than surface plasmon resonance (SPR) sensors based on the Kretschmann configuration, these types of devices are expected to be used as portable diagnostic tools or chemical/biochemical sensors. This is because nanostructured plasmonic devices provide simpler optical instrumentation than the Kretschmann configuration, which requires a prism and complicated optical alignment. Van Duyne et al. fabricated an LSPR sensor that employed triangular silver nanoparticles with nanosphere lithography8 and detected low concentration biomolecules.9 Recently, these plasmonic structures have been combined with electrochemistry to study potential dependent changes in optical properties. For example, Leroux et al. investigated the optical properties of metallic nanoparticles by applying a potential through a surrounding thin conducting polymer layer.10 Hiep et al. also combined LSPR and electrochemical impedance techniques by fabricating core−shell (silica-gold) array nanostructures, and they used their approach to measure melittin.11 Although EC-SPR measurements have been studied using SPR equipment based on the Kretschmann configuration, one of the advantages of these nanostructured EC-SPR devices © 2012 American Chemical Society
Received: November 28, 2011 Accepted: January 29, 2012 Published: January 30, 2012 3187
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nanohole array as a working electrode and a reference and counter electrode were fabricated by the following method. As shown in Figure S1 in the Supporting information, several microliters of PUA prepolymer were poured into a glassy carbon (GC) mold with a circular post whose diameter, height, and periodicity were 300, 50, and 600 nm, respectively. GC molds fabricated with electron beam (EB) lithography and reactive ion etchings were obtained from NTT Advanced Technology (Atsugi, Japan). No further preparation was employed such as coating with a mold releasing agent. After positioning 188 μm thick poly(ethylene terephthalate) (PET) film with an area of 18 mm × 18 mm (Cosmo Shine A4300, Toyobo Co., Ltd.) as a backing film, we exposed the substrate to ultraviolet (UV) light (EXECURE 4000-D, HOYA; Saitama, Japan) of no less than 200 mJ (Figure S1b in the Supporting Information). UV light and ozone gas produced by UV light are harmful to humans. The UV exposure process should be performed in an isolated box that is able to shield the experimenter from UV light and that safely disposes of the ozone gas that is produced. Subsequently the PET film with the replicated nanohole array was removed from the GC mold and rinsed in isopropyl alcohol and distilled water (Figure S1c in the Supporting Information). A gold layer was selectively patterned on the substrates by employing an adhesive sheet (made of polyvinyl chloride, no. GD-60-23A, Denka Adtech, Japan) as a mask after cutting out three electrode patterns for the working electrode (patterned on the nanohole array area) (WE), counter electrode (CE), and reference electrode (RE). After depositing gold with a thickness of 100 nm (EB-680, Eiko Engineering, Tokyo, Japan), we removed the adhesive sheet from the substrate. Here an electrode modified with silver paste (Fujikura Kasei, Tokyo, Japan) was used as an RE. Os-gel-HRP film was formed by spotting 1 μL of a 1/2 dilute solution of Os-gel-HRP solution on the working electrode and allowing it to dry in a refrigerator. The refrigerator temperature was controlled at 4 °C, and the sample was dried for 10−12 h overnight. Apparatus and Measurement. Our measurement system is shown in Figure 1b. The optical system was constructed with a halogen cold light source, a spectrophotometer, and a fiber bundle (Ocean Optics). To measure the refractive index change induced by a redox change of the Os complex, a gold nanohole array modified with Os-gel-HRP, which also performed as a WE, was illuminated with a vertical incident light from a light source through an optical fiber. The spectrum of the light reflected from the patterned area was monitored with a spectrometer and recorded in a personal computer. Cyclic voltammograms (CVs) were obtained using an electrochemical analyzer (CHI instrument, model 802) that was operated by connecting three electrodes. These three electrodes were equipped with alligator clips and wired to the potentiostat. A linear arrangement of three electrodes was employed so that gold film could be embedded in a rectangular microfluidic channel with an area of 2 mm × 12 mm. The three electrodes were arranged in the order reference, working, and counter electrodes. The working and reference electrodes should be close to each other to avoid any IR drop in the microfluidics. For the simultaneous measurement of reflection spectra and CVs induced by the redox state change of the Os-gel-HRP, three electrodes in a stationary solution were surrounded with a PDMS sheet in which there was a square hole (3 mm × 10 mm) with a depth of 3 mm for voltammetric measurement. For the H2O2 measurement, with a PDMS sheet with a straight
could be utilized more conveniently if we could realize SPR imaging with a much simpler device. We have recently developed a polymer based plasmonic gold nanohole array and undertaken a systematic study in relation to the hole geometry for an immunoassay by utilizing a nanoimprint technique.15 This technique makes it possible to fabricate plasmonic devices over a large area more accurately and with a higher throughput than by arranging Au nanoparticles or employing colloidal lithography as frequently used in previous studies. By combining this gold nanohole array with electrochemistry, we are able to realize more convenient EC-SPR devices such as multiple enzyme sensors without fabricating an electrode array. Here we report a gold nanohole array based EC-SPR device incorporated into a microfluidic channel. The synchronized measurement of cyclic voltammetry and optical sensing on the gold nanohole array were employed to investigate the effect on the SPR dip shift in the reflection spectra. A compact sensing system is particularly advantageous for realizing a high throughput and integrating with portable and disposable sensing chips. The measurement of H2O2 was demonstrated by monitoring the rate of the SPR dip wavelength change due to the reduction of the Os complex via horseradish peroxidase (HRP). Localized change of Os-gel-HRP film could be observed solely by scanning the sample substrate stage.
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EXPERIMENTAL SECTION Chemicals and Materials. Dulbecco’s phosphate-buffered saline (PBS), pH 7.4, was purchased from Sigma Chemical Co. (St. Louis, MO). Osmium-poly(vinylpyridine)-wired horseradish peroxidase (Os-gel-HRP) was purchased from Bioanalytical Systems (West Lafayette, IN). Ascorbic acid (AA) was purchased from Wako (Osaka, Japan). H2O2 was purchased from Nacalai Tesque, Inc. (Kyoto, Japan). UV curable poly(urethane acrylate) (PUA) was obtained from Minuta Tech. (Seoul, South Korea). Poly(dimethylsiloxane) (PDMS) was purchased from Dow Corning Asia (Tokyo, Japan). All the chemicals were analytical grade and were used as received. Fabrication of Microfluidic EC-SPR Device. Figure 1a shows a photograph of our microfluidic EC-SPR device. A gold
Figure 1. (a) Photograph of EC-SPR chip. (Inset) SEM image of gold nanohole array. (b) Schematic illustration of measurement setup. A 2 mm × 2 mm gold nanohole array in the center was vertically illuminated with white light through a fiber bundle. Three electrodes, namely, a counter electrode (film A), a working electrode (film B), and a reference electrode (film C), were incorporated in a PDMS microfluidic channel that was 12 mm long, 2 mm wide, and 20 μm deep. The three electrodes were connected to a potentiostat. The SPR and EC results were displayed on a PC. 3188
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Figure 2. (a) Reflection spectra from a gold nanohole array without Os-gel-HRP film (black dots) and with the film when applying different potentials of 0.1 (red dots and dashes), 0.3 (green dashes), and 0.5 B (blue solid line) through CV measurement. (b) Cyclic voltammogram (red dots) and potential dependent surface plasmon resonance wavelength sensorgram (light blue solid line) in PBS buffer without a substrate (H2O2). The gold nanohole array substrate (working electrode) was modified with 0.5 μL of Os-gel-HRP.
microflow channel (2 mm wide and 20 μm deep), the substrate was covered with a three-electrode pattern. Two perfluoro(ethylene-propylene) plastic (FEP) tubes (BAS Inc., Tokyo, Japan) were connected to the inlet and outlet on the PDMS cover as shown in Figure 1b. A CMA 102 syringe pump (CMA Microdialysis, Stockholm, Sweden) was used to introduce the sample solutions onto the sensor surfaces incorporated in the flow channel in the suction mode. The sensor surface can be used repeatedly by reducing the Os-gel-HRP (Os3+ to Os2+) by introducing 100 μM AA. Characterization of Nanoimprinted Surface. A field emission scanning electron microscope (FE-SEM) (Hitachi S4800, Hitachi, Japan) was used to characterize the gold nanohole array. The diameter of each nanohole was calculated by using image processing software (Eizo-kun, Asahi Kasei Engineering, Shizuoka, Japan).
result of the potential change from 0 to 0.5 V. Kang et al. quantified the H2O2 concentration by monitoring the reflection intensity change by a conventional Kretschmann configuration based SPR sensor because it varies more than the SPR angle shift owing to the large spectrum change of the conducting film they used.16 However in our case, the λSPR shift occurred without changing the reflection intensity, which meant we could always evaluate the λSPR shift with a sharp peak. Thus we utilize the λSPR shift by detecting the substrate concentration in the experiment described below. Figure 2b shows the typical λSPR shift (light blue solid line) that occurs when scanning the potential from 0 to 0.5 V without H2O2 (HRP substrate). The Os-gel-HRP was in a fully reduced state at the starting potential of 0 V. The voltammogram (red dots) shows a narrow peak separation of 10 mV. This CV has an almost symmetric shape. Part of the reason for the slight asymmetry is as follows. First, the Os-gel-HRP film on the gold layer is much thicker than an ideal monolayer film in order to utilize the surface sensitive area of several hundred nanometers from the gold layer. Second, the Os-gel-HRP film contains the enzyme, HRP, which increases the peak separation.17 λSPR was blue-shifted as the surface potential increased as shown in Figure 2a,b. The λSPR blue shift was about 6.5 nm and was caused by Os complex oxidation, and then λSPR was red-shifted and returned to its original value (761 nm) when the Os complex was reduced again. When the Os-gelHRP film was not modified on the gold nanohole array, the λSPR was slightly positive shifted (blue solid line) about 0.5 nm and no oxidation peak was observed (red dashes) as shown in Figure S3 in the Supporting Information. The narrow red shift was caused by the formation of a chloride ion layer in the electric double layer on the gold surface and depended on the potential increase on the gold nanohole array. Similar results have also been reported by Sannomiya et al.18 Since the reflection spectra depend greatly on the thickness of the Os-gel-HRP, we obtained several reflection spectra from the Os-gel-HRP film coated gold nanohole array at different positions by moving the sample stage. This is because the Osgel-HRP film was formed with manual spotting and natural drying, and so the layer forms a ringlike structure. However, a thickness dependence study is very important if we are to select suitable conditions such as the detection limit (dip resolution) and measurable concentration range (response time). When the fiber bundle was scanned from the center to the edge position of the deposited film with a length of about 1 mm
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RESULTS AND DISCUSSION First, we observed the SPR dip (λSPR) shift in the reflection spectra and characterized the λSPR position and the shape induced by surface potential variation. A set of four reflection spectra obtained from a bottom-filled gold nanohole array with or without Os-gel-HRP is shown in Figure 2a. The first reflection spectrum (black dot) in Figure 2a was obtained from the bare gold nanohole array surface. We observed λSPR positioned at around 733 nm, and a narrow full width at halfmaximum (fwhm) of 20.9 nm. Two other shallow peaks were observed at 720 and 740 nm. Although these two dips must be related to one of the plasmonic modes based on the enhancement of the electromagnetic wave at the top and bottom of the gold shown in our previous report,15a we have not yet confirmed the respective origins of the dips. However, these shallow dips are of limited use when monitoring the dip wavelength shift for analytical applications. Thus we focused on the strong dip at 730 nm in the following experiment. The three other spectra in Figure 2a were obtained from the bottom-filled gold nanohole substrate by modifying Os-gelHRP film at different surface potentials of 0.1 V (blue solid line), 0.3 V (green dashed line), and 0.5 V (red dash dotted line). These three dips were broader than the SPR dip obtained from the bare gold nanohole array substrate due to variations in the film thickness. On the other hand, the broadness of the dip and the reflection intensity of the three spectra were almost unchanged even after the Os redox reaction had occurred as a 3189
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(Figure 3a), the λSPR red-shifted from 751 to 760 nm as shown in Figure 3b. Also, as the film thickness increased, the dip shift
using an optical fiber with a smaller core or by changing to a single mode optical fiber. This improvement is now easy to achieve as a result of the development of commercial optical devices. Finally, the Os-gel-HRP coated gold nanohole array was incorporated into a microfluidic channel, and it was applied to an EC-SPR biosensor by injecting various concentrations of H2O2 for the substrate of HRP. In this experiment, a relatively thin Os-gel-HRP film was used to monitor the reflection spectra because of its sharper peak. Before the measurement, the Os complex was completely reduced either by applying 0 V or injecting 100 μM of ascorbic acid (AA) solution for 5 min at a flow rate of 5 μL/min. After the H2O2 had diffused in the Osgel-HRP film, the HRP reduced the H2O2 to water and changed to an oxidized state. Then the HRP was returned to its reduced form by the electron transfer from the reduced form of the Os complex. Figure 4a depicts the SPR kinetic curves at different H2O2 concentrations from 0 to 1000 μM. In the absence of H2O2 (0 μM, red line), λSPR remained unchanged. In the presence of H2O2, the λSPR decreased because the reduced Os complex was gradually oxidized by HRP, which was first oxidized by H2O2. The ΔλSPR slope (ΔλSPR/Δt) became steeper as the H2O2 concentration increased by reflecting the reaction rate of the Os complex oxidation. Figure 4b shows the relationship between H2O2 concentration and the λSPR slope shown in Figure 4a. The ΔλSPR value exhibited a linear relationship between 10 and 250 μM as shown in the inset of Figure 4b. The correlation coefficient (R2) value in the linear part was 0.996. This value is comparable to the R2 values of 0.998 obtained by Perez et al. and 0.999 by Zong et al. with the electrochemical method.19 The equation for the linear line is shown in the inset of Figure 4b, and the standard deviations of the slope and intercept were 0.26 and 0.006, respectively, as obtained with three different measurement devices. When the H2O2 concentration exceeded 250 μM, the slope became saturated. When 1 mM of H2O2 solution was injected, ΔλSPR was estimated to be about 1 nm s−1 and the oxidization of the reduced state of the Os complexes was complete in around 5 s. This reaction time is about 6 times faster than that reported by one of the authors in collaboration with Koide et al.12b However, the average amount of Os-gel-HRP film that we obtained for the same surface area was about 2 times larger than the previously reported value. Also in the previous case, the slope of the response curve is insufficiently steep to obtain quantitative data even at an H2O2 concentration of 75 μM.12b In contrast, we can measure 10 μM of H2O2 with our chip. This is because the diffusion distance of H2O2 is decreased by integrating the sensing surface into the microfluidic channel, suggesting that the linear range of the sensor chip can be tuned by changing the flow rate without changing the Os-gel-HRP thickness. On the other hand, when the H2O2 concentration is very high, the linear range can be controlled by increasing the thickness of the Os-gel-HRP layer. By immobilizing one or several oxidase enzymes such as glucose oxidase, lactate, or glutamate oxidase on or upstream of the Os-gel-HRP, a greater variety of substances can be measured with small volume samples by employing our gold nanohole based SPR device.
Figure 3. (a) Schematic illustration of an Os-gel-HRP film on a bottom-filled gold nanohole array. (b) Reflection spectra at different positions on the array. The Os-gel-HRP film is thin (blue solid line), middle (red dots), and thick (light green dots and dashes).
induced by the oxidation and reduction of the Os complex became larger as shown in Table 1 and Figure S4 in the Table 1. Dependence of SPR Dip Properties of Os-Gel-HRP Modified Nanohole Array on Layer Thickness layer thickness
thin
middle
thick
λSPR (nm) dip shift (nm) fwhm (nm) reflection (%)
750.6 3.8 30.6 18.5
754.6 5.7 33.1 30.7
760.2 12.0 38.2 40.9
Supporting Information indicating that a wider dynamic range can be obtained with a thicker film. In contrast, a sharp peak or narrower fwhm is important in regards to improving the resolution in the dip shift measurement. The exactness of the estimated fwhm value from the thick film area might be inferior to that from the thin and middle film areas owing to the very broad dip in the reflection spectra. A thinner film is more suitable for detecting the substrate concentration with better resolution despite the smaller SPR dip wavelength change. In addition, the above results indicate that we can employ this system for multiple site measurement by modifying different positions on the device with different capture proteins or enzymes. In our current system, the core diameter of the fiber bundle is 400 μm, which limits the detection spots on the nanohole array. However, we can obtain reflection spectra from a narrower area and thus improve the spatial resolution by
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CONCLUSIONS In this paper, we described our development of a gold nanohole array based EC-SPR measurement device using Os-gel-HRP film as a mediator to transduce an electrochemical reaction to an SPR wavelength shift. The gold nanohole array was 3190
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Figure 4. (a) Time course of relative dip shift when injecting 0−1000 μM H2O2 onto the Os-gel-HRP modified gold nanohole array substrate. The flow rate was 5 μL/min. (b) Calibration curve for H2O2 from 0 to 1000 μM. The slope value was estimated from each sensorgram in part a. The linear range is shown in the inset. Methot, M. P.; Live, L. S. Analyst 2010, 135, 1483−1489. (e) Nuzzo, R. G.; Stewart, M. E.; Mack, N. H.; Malyarchuk, V.; Soares, J. A. N. T.; Lee, T. W.; Gray, S. K.; Rogers, J. A. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 17143−17148. (f) Sannomiya, T.; Scholder, O.; Jefimovs, K.; Hafner, C.; Dahlin, A. B. Small 2011, 7, 1653−1663. (g) Jonsson, M. P.; Dahlin, A. B.; Jonsson, P.; Hook, F. Biointerphases 2008, 3, Fd30− Fd40. (h) Murray-Methot, M. P.; Ratel, M.; Masson, J. F. J. Phys. Chem. C 2010, 114, 8268−8275. (i) Masson, J. F.; Murray-Methot, M. P.; Menegazzo, N. Analyst 2008, 133, 1714−1721. (7) Ebbesen, T. W.; Lezec, H. J.; Ghaemi, H. F.; Thio, T.; Wolff, P. A. Nature 1998, 391, 667−669. (8) Van Duyne, R. P.; Haes, A. J. J. Am. Chem. Soc. 2002, 124, 10596−10604. (9) (a) Klein, W. L.; Haes, A. J.; Hall, W. P.; Chang, L.; Van Duyne, R. P. Nano Lett. 2004, 4, 1029−1034. (b) Van Duyne, R. P.; Bingham, J. M.; Anker, J. N.; Kreno, L. E. J. Am. Chem. Soc. 2010, 132, 17358− 17359. (10) (a) Leroux, Y.; Lacroix, J. C.; Fave, C.; Stockhausen, V.; Felidj, N.; Grand, J.; Hohenau, A.; Krenn, J. R. Nano Lett. 2009, 9, 2144− 2148. (b) Stockhausen, V.; Martin, P.; Ghilane, J.; Leroux, Y.; Randriamahazaka, H.; Grand, J.; Felidj, N.; Lacroix, J. C. J. Am. Chem. Soc. 2010, 132, 10224−10226. (11) Hiep, H. M.; Endo, T.; Saito, M.; Chikae, M.; Kim, D. K.; Yamamura, S.; Takamura, Y.; Tamiya, E. Anal. Chem. 2008, 80, 1859− 1864. (12) (a) Iwasaki, Y.; Horiuchi, T.; Niwa, O. Anal. Chem. 2001, 73, 1595−1598. (b) Koide, S.; Iwasaki, Y.; Horiuchi, T.; Niwa, O.; Tamiya, E.; Yokoyama, K. Chem. Commun. 2000, 741−742. (13) Nelson, B. P.; Grimsrud, T. E.; Liles, M. R.; Goodman, R. M.; Corn, R. M. Anal. Chem. 2001, 73, 1−7. (14) Iwasaki, Y.; Tobita, T.; Kurihara, K.; Horiuchi, T.; Suzuki, K.; Niwa, O. Biosens. Bioelectron. 2002, 17, 783−788. (15) (a) Nakamoto, K.; Kurita, R.; Niwa, O.; Fujii, T.; Nishida, M. Nanoscale 2011, 3, 5067−5075. (b) Nakamoto, K.; Kurita, R.; Niwa, O. Micro Total Analysis Systems 2009 2009, 1, 1599−1601. (c) Nakamoto, K.; Kurita, R.; Niwa, O. Micro Total Analysis Systems 2009 2009, 2, 1752−1754. (16) Kang, X. F.; Cheng, G. J.; Dong, S. J. Electrochem. Commun. 2001, 3, 489−493. (17) (a) Gregg, B. A.; Heller, A. J. Phys. Chem. 1991, 95, 5970−5975. (b) Vreeke, M.; Maidan, R.; Heller, A. Anal. Chem. 1992, 64, 3084− 3090. (18) Sannomiya, T.; Dermutz, H.; Hafner, C.; Voros, J.; Dahlin, A. B. Langmuir 2010, 26, 7619−7626. (19) (a) Perez, E. F.; Neto, G. D.; Tanaka, A. A.; Kubota, L. T. Electroanalysis 1998, 10, 111−115. (b) Zong, S. Z.; Cao, Y.; Zhou, Y. M.; Ju, H. X. Langmuir 2006, 22, 8915−8919.
fabricated by nanoimprinting and the vacuum deposition of a gold film. We succeeded in obtaining the synchronized responses of current and SPR optical signals in association with the redox change of the Os complex. We obtained a sufficient spectral shift in the EC-SPR measurement, which allowed us to estimate the slope value from the wavelength shift. We also succeeded in measuring the thickness dependent reflection spectra simply by moving the stage. The H2O2 concentration was measured in a microfluidic channel, which improved the detection limit and time by increasing the H2O2 reaction time.
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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.
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AUTHOR INFORMATION
Corresponding Author
*Phone: +81 29 861 6158. Fax: +81 861 6177. E-mail: niwa.o@ aist.go.jp. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was supported by the Research Fellowships of the Japan Society for the Promotion of Science for Young Scientists, Grant No. 21-3300. Part of this work was conducted at the Nano-Processing Facility, supported by the IBEC Innovation Platform, AIST.
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