Letter pubs.acs.org/NanoLett
Low-Copy Number Protein Detection by Electrode Nanogap-Enabled Dielectrophoretic Trapping for Surface-Enhanced Raman Spectroscopy and Electronic Measurements Leonardo Lesser-Rojas,†,‡,§ Petra Ebbinghaus,∥ Ganesh Vasan,∥ Ming-Lee Chu,§ Andreas Erbe,*,∥ and Chia-Fu Chou*,§,⊥,¶ †
Nanoscience and Technology Program, Taiwan International Graduate Program, Academia Sinica, Taipei 11529, Taiwan Department of Engineering and System Science, National Tsing Hua University, Hsinchu 30013, Taiwan § Institute of Physics, Academia Sinica, Taipei 11529, Taiwan ∥ Interface Spectroscopy Group, Max-Planck-Institut für Eisenforschung GmbH, Düsseldorf 40237, Germany ⊥ Research Center for Applied Sciences, and ¶Genomics Research Center, Academia Sinica, Taipei 11529, Taiwan ‡
S Supporting Information *
ABSTRACT: We report a versatile analysis platform, based on a set of nanogap electrodes, for the manipulation and sensing of biomolecules, as demonstrated here for low-copy number protein detection. An array of Ti nanogap electrode with sub-10 nm gap size function as templates for alternating current dielectrophoresis-based molecular trapping, hot spots for surface-enhanced Raman spectroscopy as well as electronic measurements, and fluorescence imaging. During molecular trapping, recorded Raman spectra, conductance measurements across the nanogaps, and fluorescence imaging show unambiguously the presence and characteristics of the trapped proteins. Our platform opens up a simple way for multifunctional low-concentration heterogeneous sample analysis without the need for target preconcentration. KEYWORDS: Protein analysis, electrode nanogap, dielectrophoresis, conductance, surface-enhanced Raman spectroscopy (SERS)
D
ization may be performed on passively adsorbed small organic molecules on electrode nanogap, proteins as analytical targets with potential biomedical implications are much less, or not being attempted, neither active sample transport scheme has been deployed for real-time monitoring and detection in this scenario.15 As the overall sensitivity and/or detection efficiency for lowcopy number target is frequently limited by analyte mass transport and diffusion to the actual sensing element,16,17 strategies are required for directing the transport of the few sample molecules toward the sensing elements, to enable detection on practical time scales.18 In this study, to overcome the limitations of analyte transport by diffusion, we fabricated an array of sub-10 nm Ti nanogap electrode that would serve as templates for alternating current (ac) dielectrophoresis-based molecular trapping, SERS hot spots, and electronic detectors. We demonstrated that these metal electrode nanogaps offer the advantage that they can be simultaneously applied as a
etection of proteins at low-copy numbers in a solution is challenging, particularly when it is far below the sensitivity of conventional protein analysis methods, such as enzyme-linked immunosorbent assays.1 However, detection and analysis of low-copy-number biological specimens down to the single- or few-molecule level has become possible due to the advances of techniques and methods in device fabrication that enable micro- and nanofluidics,2,3 nanopore technologies,4 molecular electronics,5 single-molecule fluorescence,6 as well as surface- and tip-enhanced Raman spectroscopy (SERS and TERS, respectively).7−9 To this aim, nanodevices combining multiple functionalities and detection schemes are desirable for fast and reliable label-free analysis of heterogeneous targets at ultralow concentrations. For example, Raman scattering is strongly enhanced for molecules at metal nanogaps due to the confinement of electromagnetic waves near corners, edges, and in gaps,10 enabling detection of low-copy number of molecules in the “hotspot”.11 Further, measurements of the tunneling current across nanometer-sized gaps have identified signals characteristic of single molecules present in the gaps.12 Narrowing of the nanogap distance has been reported to lead to stronger electrode-molecule electronic coupling and an accompanying rise in tunneling current.13,14 Though it has been demonstrated that simultaneous SERS and electronic character© 2014 American Chemical Society
Received: September 1, 2013 Revised: February 21, 2014 Published: February 28, 2014 2242
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
Further, we fabricated an array of Ti nanogaps of widths less than 10 nm to capture freely diffusing protein molecules present in an aqueous solution using high-frequency DEP. Captured proteins are detected in real time via SERS and by the conductance measurement across the nanogap. As in lowfrequency DEP, that is, less than tens of kilohertz, ac electrohydrodynamic (EHD) flow may rise due to the longrange electro-osmotic flow of electrolytes in the electrical double layer across the electrode surface which may interfere with the DEP force,49,50 rendering the particle trapping generally not at the tips of the nanogap electrodes where the DEP trapping potential minimum, hence the strongest field gradient and the corresponding DEP force, lies. To avoid this complication of low-frequency field induced EHD and its associated electro-osmosis, we use high-frequency (1 MHz) ac field to obtain DEP trapping, thus molecular trapping relies exclusively on the polarizability contrast between the analyte and the solvent and particles can be positioned at the nanogap. Because of the nature of the short-ranged interaction volume of DEP, ∼1/r3, the trapping and detection occurs almost instantly (approximately seconds) if there are single molecules at the close proximity of the nanogap. This strategy ensures the trapping is reversible and the detection may be orders of magnitude faster than previously reported methods.35,36 In this context, the combination of electronic and Raman spectroscopic detection of the DEP-captured molecules makes the device attractive as a label-free bioanalytical tool. An array of Ti nanoelectrode pairs with gap dimensions less than 10 nm was fabricated with electron-beam lithography, egun metal evaporation, and a lift-off process on an open top 7 × 7 mm2 Si chip with a 1.2 μm thick thermal oxide layer. The array is shown in Figure 1A−C. (See Supporting Information for detailed fabrication process.) Examples of the resulting nanoelectrode structures are shown in Figure 1D,E. Three identical groups, each consisting of five different electrode pairs, were prepared. Within the group of five wires, each gap has a distinctive geometrical design, to investigate the effect of the tip geometry on the trapping of molecules and the enhancement of
molecular trap as well as an analytical device for signal transduction by optical and electronic means. Dielectrophoresis (DEP) is an electrokinetic effect that has been employed to attract and separate polarizable dielectric particles in aqueous media, depending on the dielectric response of the particle in the presence of a nonuniform electric field.19,20 Dielectrophoretic force acting on a polarizable spherical particle in a nonuniform field may be described as FDEP = 2πR3εmRe[K(ω)]∇Erms2, where R is the radius of the particle, εm is the absolute permittivity of the suspending medium, Erms is the root-mean-squared amplitude of the applied ac field, and Re[K(ω)] is the real part of the ClausiusMossotti (CM) factor, representing the frequency-dependent dielectric contrast between the particle and the suspending medium in an external driving field. Aided by advances in micro- and nanofabrication of structures for field localization, DEP has been successfully used to manipulate bioparticles such as DNA,21,22 RNA,23 viruses, and bacterial spores24 and provide means for rapid enrichment and mass transport of proteins with the implication of enhanced sensitivity and sensing kinetics.25−27 Though there is increasing interest in recent years by applying electrodeless or insulator-based DEP for biomolecule manipulation,18,28−32 the most commonly implemented scenario is the thin metal electrode-based DEP (mDEP), where DEP field is generated by applying a voltage across metal electrodes.19,21,22 MDEP is well-suited for the capture and concentration of low-copy numbers of small biomolecules, such as single short DNA fragments33 and proteins34 using metal electrode nanogaps, if mDEP is operated at the positive scenario where Re[K(ω)] > 0. However, the electrode nanogaps used in mDEP are not necessarily suitable to serve as a hotspot for SERS unless its gap size is in the order of less than 10 nm. On the other hand, various nanoscale metallic structures found in the literature may serve as good SERS substrates,10,11,35,36 but to our knowledge none has been applied simultaneously as a mDEP molecular trap for active sample transport, particular for protein analysis.15 To be noted, many of the SERS substrates previously introduced were fabricated using gold.11,34 Gold is, however, prone to corrosion especially in chloride-containing electrolytes, which include almost all biologically relevant buffers.37,38 Chloride reduces the standard reduction potential of the Au/AuIII redox pair from 1.4 to 1.0 V, making gold less noble.39 In the presence of oxygen, for example, nanogaps as used here will inevitably undergo crevice corrosion.40 Metal dissolution in general starts from exposed surface atoms,41 which must be present in large amounts near sharp structures. In analytical devices, flow will additionally lead to removal of corrosion products and hence a dissolution of nanostructures. Corrosion control can be achieved by the use of a passive material,42 that is, a metal with an oxide film, such as titanium (Ti). The use of such a material might be beneficial for use in practical bioanalytic applications, as it limits nanoelectrode degradation. Literature shows few reports on the observation of SERS near metallic Ti surfaces,43−45 which in all cases are covered with a native oxide under the conditions used in the experiments. On the other hand, there are several reports of (comparatively weak) SERS of molecules strongly interacting with TiO2.43,46−48 For this work, we start from the hypothesis that field enhancement, and consequently the observation of SERS, in nanogaps between metallic tips is an effect intrinsic to all metals. For the special case of Ti, this hypothesis will be tested by finite element simulations.
Figure 1. Fabricated electrode nanogap device. (A) A 7 × 7 mm2 silicon chip with a 1.2 μm thick top layer of thermally grown SiO2 on which (B) 15 parallel trapping microelectrode pairs with contact pads were defined using standard photolithography, thermal evaporation of Ti/Au, and lift-off process. (C) E-beam nanolithography, e-gun metal evaporation of a 40 nm Ti thin layer and lift-off process, define a 200 nm wide electrode pair with interelectrode gaps of different sizes. SEM of electrode nanogap structures: (D) 5 nm gap and (E) 9 nm gap. 2243
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
Figure 2. (A) Finite element method simulated distribution of the modulus of electric field (|E|) at 633 nm around a nanogap of 5 nm between Ti tips with an internal angle of 120°, resembling the gap used in experiments. (B) Plot of the SERS enhancement factor |E|4 at point of maximum field enhancement versus gap distance between tips for 120° internal angle tip. (C) Plot of the SERS enhancement factor |E|4 at point of maximum field enhancement versus angle of the tip for 5 nm gap.
thick) was used to encapsulate a 2 μL droplet of protein solution. The chamber was sealed against evaporation by a circular coverslip of 5 mm diameter (Knittel Gläser) and vacuum-grade grease applied on the edges to keep the solution from evaporation. Raman spectra were obtained using a Horiba Labram 2 confocal Raman microscope with a HeNe laser (633 nm) as light source and 50× (N.A. 0.5) and 100× (N.A. 0.9) objectives (Olympus) where appropriate. The recorded wavenumber axis of all spectra were subjected to a fine calibration by setting the frequency of the dominating Si phonon peak to 520 cm−1.53 All peak wavenumbers are reported with an error of ±3 cm−1. The polarization of the incident light was along the axis defined by the direction of the electrode nanogaps. Typical spectral mapping used confocal point illumination by a laser spot whose dimensions are diffraction-limited. Spatial sampling density was defined using the Nyquist criterion. Time for sampling of one voxel was 1 s. Single-voxel, time-lapsed Raman spectra acquisition at the nanogap hotspot was also performed with 1 s exposure time. Fluorescence microscopy was performed using an inverted fluorescence microscope (Leica DMI6000) with a high-stability mercury lamp as light source and a K3 filter cube (Ex/Em: BP 470−490 nm/LP 515, Leica). A 100× oil-immersion objective (Leica, N.A. 1.4), and an EMCCD camera (IXon-888, Andor Technologies) with exposure time of 25 ms/frame were used to make the observations. To study the distribution of electric fields at optical frequencies in the nanogaps, the time-harmonic Maxwell equations were solved numerically using the finite element method (FEM)-based commercial electromagnetic solver JCMsuite (www.jcmwave.com). The nanogap tips were modeled in a domain with cylindrical coordinates by creating a half-plane geometry of the tips and appropriate boundary condition applied along the rotational symmetry axis (see Supporting Information Section II and Figure S1.) The result of FEM solutions of the Maxwell equations for the modulus |E| of the electric field at 633 nm around the electrode nanogap predicts a region of high electric field at the rim of the nanogap between the Ti metal tips (Figure 2A). The nominal field modulus |E| and SERS enhancement factor |E|4 versus interelectrode distance and tip internal angle is shown in Figure 2B,C and Supporting Information Figure S2. In our experiments, real-time Raman, electronic, and fluorescence detection of the trapped protein molecules were performed. To achieve that, we positioned the laser focus at a
Raman scattering. The internal angle of the electrode pairs were typically 120° for the sub-10 nm gap (simulation results of the enhancement factor of these geometries will be discussed later). Every chip was inspected using scanning electron microscopy (SEM, FEI Nova) at various magnifications with acceleration voltage of 20 kV, UHR mode, and a working distance of 5 mm before an experiment to look for fabrication defects and confirm the quality of the gaps. Ohmic contact was inspected in vacuum using an in situ probe system that locates a pair of probe needles with the aid of a nanomanipulator (Zyvex S2004) inside an SEM (FEI-Inspect) coupled to a semiconductor characterization system (Keithley 4200) and a typical resistance measured in air was in the order of 1011−1012 Ω, indicating an excellent insulation of the underlying thermally grown SiO2 layer and the absence of leakage currents between tips. DEP trapping and electronic detection was conducted using a custom-made printed circuit board with signal amplification, filtration, and data acquisition capabilities with 100 pA current measurement resolution and 1 ms temporal resolution. Different ac field conditions were applied, ranging from hundreds of kilohertz up to 4 MHz and various amplitudes from 0.1 to 15 Vp‑p, coupled with a direct current (dc) bias typically between 10 and 100 mV, where ac field is responsible for the DEP trapping, and the dc bias leads to the net current (∼pA−nA) across the nanogaps. The ac field was applied by a function generator (33220A, Agilent) and monitored by an oscilloscope. Electronic detection of trapped molecules relies on the observation of current measurement across the nanogaps. Signals were retrieved through a transimpedance amplifier with a low-pass filter as key component. R-phycoerythrin (RPE), a 240 kDa disk-shaped protein with a diameter of 11 nm and a thickness of 6 nm derived from red algae,51 was used for the trapping experiments. The RPE is autofluorescent from the chromophore embedded in its molecular structure.52 RPE (4.0 mg/mL, ThermoScientific) was cassette dialyzed in PBS buffer (100 mM Na2HPO4, 150 mM NaCl, pH 7.2), then diluted to 0.8, 0.26, and 0.12 nM final protein concentrations. Protein concentrations were determined using a spectrophotometer (Nanodrop, Thermo Scientific) from Beer’s Law with extinction coefficient 2.0 × 106 LM−1 cm−1 at 567 nm. For further verification, a Bradford Microassay, calibrated against BSA, was also used to determine the protein concentration. For confocal Raman and fluorescence microscopy, a small fluidic chamber defined in a double-sided adhesive tape (70 μm 2244
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
point of maximum Raman intensity that corresponds to the location of the nanogap. In a spectral map without baseline subtraction, that is, including luminescence contributions, of the Si phonon at 520 cm−1, a region of high intensity was observed (see Supporting Information Section III and Figure S3). This high-intensity region was only observed when the resulting spectra included both Raman and luminescence contributions, that is, no baseline correction performed. The high-intensity region was observed in both dry and wet chip conditions. The phenomenon was only observed on wires with gap sizes below 10 nm and only on those gaps that were previously imaged through scanning electron microscopy (SEM). The observation of SERS signals come only from those electrodes being bombarded by electrons during SEM has been reported previously.11 Origins for the luminescence are possibly inelastic electronic effects in the chips generated by intraband transitions mediated by the strongly confined fields near the metal nanostructures, which cause a broad visible and infrared photoluminescence continuum background.11,54 The luminescence is enhanced in the vicinity of the metal nanogaps due to increased excitation intensity, consistent with an electromagnetic enhancement mechanism. No luminescence enhancement was detectable in larger nanogaps and in samples that were not investigated by SEM. An initial scan over the surface therefore provided an indication of the location of the Raman hotspot on the chip. The same hotspot also provided enhancement of the Raman signal of molecules at place, which was demonstrated by introducing a drop of paraffin on the gap and sealing it with a coverslip (Supporting Information Figure S3C). The existence of the hotspot also validates the simulation result in Figure 2A. The located hotspot was used for subsequent point illumination, time-dependent and DEP-assisted Raman measurements with RPE samples in solution. Figure 3A shows the Raman spectra of trapped RPE in a 5 and 9 nm gap compared against the RPE sample adsorbed in the hotspot of an Ag colloid hydrosol55,56 showing the match in the 9 nm gap of several distinctive vibrational modes for the protein under SERS conditions. Figure 3B shows the time-dependent Raman spectra during a trapping event. In the first few seconds, after switching the field to DEP trapping conditions just the typical silicon substrate spectrum is observed (see also Supporting Information Figure S4A). Between 10 and 20 s after the start of the experiment, and the increase of the fluorescence signal is observed. At the same time, a number of narrow Raman peaks appear on top of the fluorescence background, indicating the molecular Raman signature becomes more pronounced in the course of the experiment. It has been noted that peak positions in the spectra of molecules in the gap can differ substantially from the spectra of molecules in free solution,57 while conduction of electric current leads to changes in the intensity of the different bands.58 As shown in Figure 3A, there is a difference between observed spectra in gaps of width of 5 and 9 nm. In the 9 nm gaps (gap distance ∼ hydrodynamic diameter of RPE), several spectral features resemble the spectrum obtained after adsorption of RPE to Ag hydrosols (Figure 3A, inset) with several peaks in the range of the amide III modes.59,60 Differences between the spectra could either be attributed to stretching changes of RPE after adsorption to the Ag hydrosol,61 or the spatial distribution of the electric field in the gap may stress different spectral features in the protein spectra compared to the Ag hydrosol experiment. On the other hand, in 5 nm gaps the intensity is considerably lower, as only
Figure 3. (A) Raman spectra of RPE as trapped in a 5 nm (blue) and 9 nm gap (red) compared against the RPE sample adsorbed in the hotspot of an Ag colloid hydrosol (inset), showing the most distinctive vibrational modes for the protein (see text for corresponding peak assignment) under SERS conditions. (B) Time evolution of Raman spectra obtained from a 5 nm gap hotspot during a trapping event with RPE concentration 0.8 nM in 1× PBS buffer, ac 10 Vp‑p at 1 MHz and dc bias 100 mV. (C) Analysis of peak height versus time before and after (inset) the baseline correction for the 715 cm−1 peak (CH deformation mode in aromatic systems in (B)). (D) ac amplitude dependent on−off signature of the peak height (715 cm−1) versus time where distinctive Raman signatures are present after a DEP threshold value of 6 Vp‑p at 1 MHz and dc bias 100 mV (cf. the electronic data in Figure 4A). To be noted, the “off” condition here applies to both the ac/dc components.
protein chains can penetrate the gap. The different penetration into the gap also significantly affects the shape of the spectra, as 2245
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
spectroscopic evidence in accordance with the established DEP theory discussed in the previous session. In addition to the Raman measurements, electronic measurements were also performed. Supporting Information Figure S4B shows the current versus time signature in the absence of trapping (negative control). This is observed when just the dc detection bias is applied to the solution containing no molecules, or when the applied ac frequencies/amplitude were far from the threshold causing DEP trapping at the presence of proteins.22 The zoomed-in plot of Supporting Information Figure S4B shows the short time behavior similar, but not equal, to capacitor-like charging curve right after switching the field on. We attribute this to the presence of ionic species near the gap vicinity that redistribute locally in the beginning according to their charge until the ionic current flow reaches a stable value. Though dc bias can contribute to the capture of molecules by electrostatic trapping,67 the required field is orders of magnitude higher than the 10s mV/nm applied here. This explains why there are no observable changes in the current measurement when the dc bias alone is applied. Electrochemical effects on the electrodes and electro-osmosis (as discussed earlier) are minimized through superimposing a high-frequency modulation with a much larger amplitude on the applied dc bias.68 When DEP conditions are applied, a current fluctuation and a baseline increase is observed for both 5 and 9 nm gaps (Figure 4A), relating to proteins bridging the gap, and due to molecular rearrangements/reorientations allowing the ionic current to flow and to be blocked repeatedly through the junction. Though tunneling current may also contribute to the current increase with trapped proteins, its contribution cannot be readily separated at the moment. It is noted there are distinctive features of the electronic signals observed, that is, the current jumps in a stepwise manner when the amplitude of the ac signal is increased during the acquisition suggesting that the trapping of RPE may occur once the trapping threshold is reached, namely, 10 Vp‑p and 6 Vp‑p for 10 and 100 mV dc bias applied, respectively. This is due to the limited number of protein molecules in the vicinity of the trap where the DEP force exceeds the thermal drag force,19,20 or threshold force, which is defined from the diffusion path during the experiment.69 As different protein concentrations have been used in the experiments, namely 0.8, 0.26, and 0.12 nM, these correspond to an average of 1 protein molecule per every 2.0, 6.12, and 12.75 μm3, respectively. The increase in the amplitude of the ac field component reflects as an increase in the magnitude of the field gradient, hence the DEP force, ∼Vrms2, and the depth of the associated trapping potential well, and thus effectively extending the volume where DEP forces are effective, therefore bringing more molecules to the trap and improving the current conduction pathways. The scenario of EDL overlapping70 in the nanogap is excluded in the high ionic strength buffers used here, but it could be occurring in low ionic strength buffers (10 μS/cm and below) where it could be a reality in the beginning. But as ion separation effects take place upon the application of a dc detection bias (the current decrease observed at the beginning of the measurement in Figure 4A and Supporting Information Figure S4B), this might facilitate the ac trapping mechanism after the base ionic conduction is reached. To further estimate the number of proteins in the trap, we followed a similar strategy such as the one used by Hölzel et al.,34 on which for a successful spatial manipulation, electric forces must exceed diffusion and friction. Minimum force that is
vibrational modes from only a limited number of molecular groups contribute to the Raman spectrum significantly. Nevertheless, the spectrum contains a number of bands that can be tentatively assigned to typical protein structural elements: peaks at 1645 cm −1 (Amide I), 1580 cm −1 (antisymmetric stretching mode of side chain COO−), 1465 cm−1 (CH deformation mode of aliphatic CH), 1345/1285/ 1245 cm−1 (tentatively Amide III mode), and 815/715 cm−1 (CH deformation mode in aromatic systems) are observed.59,60 Aromatic and COO−-containing side chains are abundant in RPE.62 The strong peak at 1005 cm−1 may originate from a mode of phenylalanine.63,64 The two peaks observed at 1710 and 1765 cm−1 only in the 9 nm channels are characteristic CO stretching modes.65 The mode at 1715 cm−1 is observed in other proteins as well63 and may be related to COOH groups of side chains.66 As at pH 7.2, these groups are expected to be deprotonated, this assignment of the peak at 1715 cm−1 to a CO stretching mode from a COOH group is unrealistic. Instead, both modes at 1715 and 1765 cm−1 are tentatively assigned to modes from the chromophore, which contains unsaturated lactams.62 Indeed, amide groups in the vicinity of CC double bounds show vibrational frequencies up to 1770 cm−1.65 A full and thorough assignment of the Raman bands is beyond the scope of this work as it would involve computations, isotope exchange, solvent dependent measurements, and so forth. Figure 3C shows the time evolution of the peak height at 715 cm−1 before and after the baseline correction. The increase in Raman and luminescence intensity suggests that an increasing number of protein molecules within the vicinity of the gap were trapped at the nanogap and eventually reached a saturation level. Furthermore, SERS measurements during the on−off of DEP field show evidence that the molecules do not completely go away from the tip area when one switches off the ac field component, as shown in Supporting Information Figure S5A for the nonbackground- and background-subtracted signatures. In this case, the small dc bias at presence (needed for the electronic measurement) could be enough to keep some originally trapped molecules to stay around the hotspot by mere electrostatics where the negatively charged protein may be retained at the positively charged electrode side of the dc bias, which is not from the pure dc-DEP effect from the applied bias as it is far below the threshold of DEP (see also discussion later), but not bridging the electrode gap. Hence one can still observe some residual SERS signals (Supporting Information Figure S5A) but not the electronic and fluorescence signals when the ac field is off, where the majority of the fluorescence intensity goes away, and if a molecule is still present near the gap it is indistinguishable from the background (see more discussion later). However, when one switches off the experiment completely (both ac/dc components) and then applies the trapping condition again, it is easy to see that the Raman spectra (Figure 3D) of the first few seconds will be free from the molecular fingerprints, suggesting the proteins are not adsorbed on the hot spot, and that only after reaching the onset of DEP trapping, one will see the apparition of the protein vibrational modes, which indicates the trapping of proteins at the hotspot and supports the notion of reversibility of our operation. It is also interesting to observe in this data that the time to reach the onset of DEP decreases as one increases the ac amplitude, meaning that with higher field the readily available proteins get collected faster, and thus providing an 2246
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
CFD, North America, Huntsville, AL). Then the space in which RPE molecules are trapped and drawn toward the electrodes could be derived from the spatial distribution of ∇Erms2. From our computing results for an 8 nm gap, assuming the free space to remain a continuum, the value of ∇Erms2 along the central line perpendicular to the midpoint of nanogap axis has a maximum of 3 × 1023 V2/m−3 for an applied amplitude of 6 Vp‑p and 1.9 × 1024 V2/m−3 for 15 Vp‑p (the range of the field amplitude where trapping signatures were observed in our experiments with 100 mV dc bias), corresponding to a maximum holding force of 30 and 187 pN, respectively. Apparently, the space where the dielectrophoretic force exceeds molecular diffusion could extend to a larger volume above the size of the nanogap, however effects that could result in the reduction of dielectrophoretic forces have also been discussed.34 Here the extended range (rex) over the threshold force is 0.6 μm for 6 Vp‑p and 1.12 μm for 15 Vp‑p applied amplitude from the center of the nanogap. We could then estimate in average the lower bound of ∼0.5 molecule within the threshold volume (half dome with radius rex) at 6 Vp‑p and ∼3 molecules at 15 Vp‑p with 0.8 nM RPE protein loading concentration. It is also interesting to observe that the equilibrated current level (determined by projected histogram analysis on the current axis) and the corresponding conductance versus the applied ac amplitude from the threshold and above may be well fitted by a parabola, that is ∼(ac amplitude)2, coincident with the DEP force dependence on ac amplitude, for both the 10 and 100 mV applied dc bias (Figure 4B). However, the origin of this dependence is not clear. Hence, it is not straightforward to obtain the exact number of proteins bridging the nanogap from this observation, because there is no direct linking of the conductance to the number of molecules at the trap being established by theory or experiments for proteins, as the complex form of proteins, the orientation and the alignment of proteins (single or multiple), the chromophore embedded in the protein, all contribute to the conduction pathway. Though there are single molecule techniques that could potentially quantify the number of molecules, such as fluorescence72 or electrical73,74 correlation spectroscopy, the challenges lie in the fact both these methods are based on the assumption that the molecules do not interact with the probing optical or electric field, respectively, that is, freely diffusing. In our case, however, the molecules do interact with the nonuniform field (gradient), hence is not freely diffusing (when trapped) or diffusion is spatially dependent in the close proximity of the nanogap. These facts render the above methods inapplicable and an analytical/numerical model needs to be developed for correlation/fluctuation spectroscopy in a nonuniform interacting field. The corresponding current measurement of a repetitive on− off switching of the ac field for DEP during the acquisition indicates the reversibility of the trapping process with a typical current difference from 0 to 0.7 nA between the trapping (15 Vp−p ac amplitude) and nontrapping (ac off) conditions, if the applied dc bias is 10 mV across the nanogap (Figure 4C). It is noted that the 70 nS conductance indicated here (0.7 nA with 10 mV dc bias) is within the range of molecular conductance reported by others. For example, Tao’s group have reported conductance of single organic molecules, such as 4,4′ bipyridine, to be 800 nS (10 nA with 13 mV dc bias) and hexanedithiol (which has a large band gap) 100 nS,75 though the comparison may not be straightforward as we are using
Figure 4. (A) Electronic signatures of RPE trapped by DEP across a 9 nm gap with RPE concentration 0.8 nM in 1× PBS buffer, ac 1−15 Vp−p at 1 MHz with dc bias of 100 (red) and 10 mV (inset, green) (see also the negative controls in Figure S4B in the Supporting Information). The threshold ac amplitude for DEP trapping is greater than 6 and 10 Vp−p for the 100 and 10 mV dc bias, respectively, indicating by the baseline increases that suggest the trapping of proteins (data displayed with a 5-point running average). (B) The current signature and the corresponding conductance (y-axis on the right, constructed from the projected current histograms) versus the applied ac amplitude from the threshold and above in (A) may be fitted by a parabola (I ∼ Vac2) for both the 100 (red) and 10 mV (inset, green) dc bias with the goodness of fit R2 = 0.99 in both cases. (C) Reverse operation of ac dielectrophoresis (repetitive on−off switching of the ac field between 0 and 15 Vp−p, while keeping the dc bias at 10 mV) during the current acquisition for the 0.8 nM RPE solution indicating the reversibility of the trapping process (data displayed with 5-point running average).
needed to overcome these effects can be defined as an observable deterministic threshold force and is described by the relationship Fth = (2/DΔt)1/2kBT,71 where kBT is thermal energy and Fth is in the order of 3 × 10−16 N for a 10 s (Δt) experimental time interval, given diffusion constant (D) of RPE 40 μm2/s, and this corresponds to a threshold ∇Erms2 ∼ 3 × 1017 V2/m3. Dielectrophoretic forces could then be estimated using the FDEP equation on the basis of the gradient of the root-meansquare of the electric field strength, ∇Erms2, for which a multiphysics model was computed using finite element methods with the commercial software ESI-CFD ACE + (ESI 2247
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
filtered and pseudocolored for illustration purpose (see details of the imaging processing and the corresponding movie, nonfiltered, in Supporting Information). There are similar fluorescence phenomena conducted with 0.8 nM RPE concentrations in low conductivity buffers (≤15 μS/cm), as increasing the ac amplitude will make the inhomogeneous electric field gradient stronger and also expand the volume where it is effective, being therefore able to trap more molecules and thus increase the fluorescence intensity in the region near the gap. Figure 5B resembles a staircase-like increase (though not a perfect one) in fluorescence, which is in correspondence to the electronic data, being the first fluorescence increase visible only after reaching the threshold ac amplitude of 6 Vp−p and increasing thereafter the ac amplitude is increased, in this case up to 10 Vp−p. However, as the conductivity of the solution increases, it is more difficult to observe such phenomena due to poor signal/background ratio (even after heavy postimage processing operations). We attribute this observation to the charge screening effects that requires higher field, thus higher ac amplitude, to achieve the same degree of polarization of particle by DEP,19,20 that is, higher signal/background ratio, and the possible photobleaching and/or quenching effects of the low copy number of molecular species trapped at the gap interface. If the fluorescence measurements are performed at the same protein concentration (0.8 nM) in a higher conductivity buffer (38 μS/ cm), though there is a constant increase in the fluorescence intensity after the threshold amplitude value (data not shown), the stepwise increase is not directly mirrored in a likewise manner of the current (Figure 4A). In addition, there may have complication involving surface energy transfer mechanism76 and/or field-induced fluorescence quenching effect77 (while these effects may be investigated by fluorescence lifetime measurement, it is beyond the scope of the current study) causing the fluorescence signals being reduced when trapped molecules are sitting on top of the Ti/TiO2 electrode, which may render the lower number of trapped proteins unobservable in higher conductivity buffer, particularly at 1× PBS and the same field conditions where Raman and electronic measurements were performed. In our opinion, SERS/current measurements are much more sensitive probes to locating the molecule than fluorescence. Likewise, as performed in Raman and electronic experiments, we also acquired the fluorescence signatures of the proteins by switching the ac DEP field on and off. Supporting Information Figure S5B is obtained by such operation without background and photobleaching correction while Figure 5C is photobleaching but not background corrected. It also resembles the on−off signatures of the applied ac field in the Raman (Figure 3D) and electronic (Figure 4C) measurements and again indicates the reversibility of the trapping process. In summary, we have demonstrated a novel real-time analytical device involving well-patterned sub-10 nm electrodes nanogap array for active protein transport through DEP. During molecular trapping, three independent measurements, that is, recorded Raman spectra through SERS, current/ conductance across the nanogaps, and fluorescence imaging, all indicate the presence of the trapped proteins and its characteristics are mostly revealed by Raman spectroscopy. Other than common molecular electronics devices where a small organic molecule bridges a pair of nanoelectrode, we demonstrated low-copy number of protein analysis may be performed by DEP-assisted trapping and simultaneous SERS or
proteins here. On the other hand, if multiple proteins trapped in between the nanogap they could form multiple conduction pathways and the chromophores of RPE proteins could also play a role in the conductance. Additionally, as RPE is an intrinsically fluorescent protein, fluorescence microscopy during trapping may also be performed to further verify the presence of the molecules. Figure 5A shows a typical fluorescence image of RPE proteins being captured in the interelectrode region when applied field is above the onset of DEP, further supporting the presence of the trapped molecules. To be noted, the image of Figure 5A is
Figure 5. (A) Fluorescence imaging of RPE molecules (0.8 nM) in 24 μM PBS (15 μS/cm conductivity) trapped in a 9 nm electrode nanogap after 30 s of field application of ac 8 Vp−p at 1 MHz and dc bias 10 mV (image filtered and pseudocolored for illustration purpose, see details of the imaging processing and the corresponding movie in Supporting Information). (B) Fluorescence intensity (cropped from a 5 × 5 pixels area, ∼1 μm2, around the hot spot) versus time of RPE protein trapping (background and photobleaching corrected) with the same condition in (A) except the ac amplitude was ramped from 1 to 10 Vp−p (at 1 Vp−p increase every 10 s interval). Note the threshold of ac amplitude for DEP trapping is greater than 6 Vp−p. (C) Fluorescence intensity versus time (photobleaching, but not background, corrected) for 10 s interval of on−off cycles of the ac field (between 0 and 15 Vp‑p) at 1 MHz with dc bias 10 mV in a RPE containing solution as in (A). Data are presented with 10-point moving average. 2248
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
(10) Alvarez-Puebla, R.; Liz-Marzán, L. M.; de Abajo, F. J. G. J. Phys. Chem. Lett. 2010, 1, 2428−2434. (11) Ward, D. R.; Grady, N. K.; Levin, C. S.; Halas, N. J.; Wu, Y.; Nordlander, P.; Natelson, D. Nano Lett. 2007, 7, 1396−1400. (12) Albrecht, T. Nat. Commun. 2012, 3, 829. (13) Tsutsui, M.; Taniguchi, M.; Yokota, K.; Kawai, T. Nat. Nanotechnol. 2010, 5, 286−290. (14) Tsutsui, M.; Rahong, S.; Iizumi, Y.; Okazaki, T.; Taniguchi, M.; Kawai, T. Sci. Rep. 2011, 1, 46. (15) Natelson, D.; Li, Y. J.; Herzog, J. B. Phys. Chem. Chem. Phys. 2013, 15, 5262−5275. (16) Sheehan, P. E.; Whitman, L. J. Nano Lett. 2005, 5, 803−807. (17) Sparreboom, W.; van den Berg, A.; Eijkel, J. C. T. Nat. Nanotechnol. 2009, 4, 713−720. (18) Swami, N.; Chou, C. F.; Ramamurthy, V.; Chaurey, V. Lab Chip 2009, 9, 3212−3220. (19) Pohl, H. A. Dielectrophoresis: the Behavior of Neutral Matter in Nonuniform Electric Field; Cambridge University Press: Cambridge, 1978. (20) Pethig, R. Biomicrofluidics 2010, 4, 022811. (21) Washizu, M.; Kurosawa, O. IEEE Trans. Ind. Appl. 1990, 26, 1165−1172. (22) Washizu, M.; Suzuki, S.; Kurosawa, O.; Nishizaka, T.; Shinohara, T. IEEE Trans. Ind. Appl. 1994, 30, 835−843. (23) Giraud, G.; Pethig, R.; Schulze, H.; Henihan, G.; Terry, J. G.; Menachery, A.; Ciani, I.; Corrigan, D.; Campbell, C. J.; Mount, A. R.; Ghazal, P.; Walton, A. J.; Crain, J.; Bachmann, T. T. Biomicrofluidics 2011, 5, 024116. (24) Lapizco-Encinas, B. H.; Rito-Palomares, M. Electrophoresis 2007, 28, 4521−4538. (25) Lapizco-Encinas, B. H.; Ozuna-Chacon, S.; Rito-Palomares, M. J. Chromatogr., A 2008, 1206, 45−51. (26) Liao, K.-T.; Chou, C.-F. J. Am. Chem. Soc. 2012, 134, 8742− 8745. (27) Liao, K. T.; Tsegaye, M.; Chaurey, V.; Chou, C. F.; Swami, N. S. Electrophoresis 2012, 33, 1958−1966. (28) Chou, C. F.; Tegenfeldt, J. O.; Bakajin, O.; Chan, S. S.; Cox, E. C.; Darnton, N.; Duke, T.; Austin, R. H. Biophys. J. 2002, 83, 2170− 2179. (29) Chou, C.-F.; Zenhausern, F. IEEE Eng. Med. Biol. Mag. 2003, 22, 62−67. (30) Cummings, E. B.; Singh, A. K. Anal. Chem. 2003, 75, 4724− 4731. (31) Regtmeier, J.; Eichhorn, R.; Viefhues, M.; Bogunovic, L.; Anselmetti, D. Electrophoresis 2011, 32, 2253−2273. (32) Srivastava, S. K.; Gencoglu, A.; Minerick, A. R. Anal. Bioanal. Chem. 2011, 399, 301−321. (33) Tuukkanen, S.; Kuzyk, A.; Toppari, J. J.; Hytonen, V. P.; Ihalainen, T.; Torma, P. Appl. Phys. Lett. 2005, 87, 183102−3. (34) Hölzel, R.; Calander, N.; Chiragwandi, Z.; Willander, M.; Bier, F. F. Phys. Rev. Lett. 2005, 95, 128102. (35) Zheng, G.; Qin, L.; Mirkin, C. A. Angew. Chem. 2008, 120, 1964−1967. (36) Zheng, G.; Chen, X.; Mirkin, C. A. Small 2009, 5, 2537−2540. (37) Moslavac, K.; Lovrecček, B.; Radeka, R. Electrochim. Acta 1972, 17, 415−420. (38) Frankenthal, R. P.; Thompson, D. E. J. Electrochem. Soc. 1976, 123, 799−804. (39) Vanýsek, P. Thermochemistry, Electrochemistry, and Solution Chemistry. In CRC Handbook of Chemistry and Physics, 94th ed. (Internet Version 2014); Haynes, W. M., Ed.; CRC Press/Taylor and Francis: Boca Raton, FL, 2014; Section 5, pp 5−80. (40) Kaesche, H. A note on crevice corrosion. In Corrosion of Metals: Physicochemical Principles and Current Problems; Springer: Berlin, 2003; pp 381−384. (41) Landolt, D. Non-Uniform Corrosion Governed by Metallurgy. In Corrosion and Surface Chemistry of Metals; CRC Press: Boca Raton, FL, 2007; Chapter 7, pp 291−293.
electronic confirmation using electrode nanogap. As the chip and electronic detection technology could be easily mass produced with high yield and coupled to widely used spectroscopic instruments, our platform demonstrated a simple way for low-copy number protein detection that may promise applications and, if further coupled to fluidic channels, for lowconcentration heterogeneous sample and small molecule analysis (e.g., electrochemical or enzymatic) at the single or few molecules level.
■
ASSOCIATED CONTENT
S Supporting Information *
Detailed information related to device fabrication, FEM simulation, negative controls, and fluorescence imaging processing with supporting figures and movie. This material is available free of charge via the Internet at http://pubs.acs.org.
■
AUTHOR INFORMATION
Corresponding Authors
*E-mail: (C.-F.C.)
[email protected]. *E-mail: (A.E.)
[email protected]. Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS The authors are grateful to Professor Nathan Swami for critical reading of the manuscript, Professors Naomi Halas and Peter Nordlander and Drs. Chii-Dong Chen and Yi-Ren Chang for helpful discussions, Shuei-Jin Lai and Yi-Luan Li (AS Nano Core Facility) for technical assistance on e-beam lithography, Yii-Lih Lin on image processing, Katrin Vu on fluorescence measurements, and National Center for High-Performance Computing for computer time and facilities. This work was supported by AS Nano Program, AS Integrated Thematic Project (AS-103-TP-A01), National Science Council, Taiwan (102-2112-M-001-005-MY3 and 103-2923-M-001-007-MY3), and Asian Office for Aerospace Research and Development (FA2386-12-1-4002). L.L.-R. thanks the Max-Plack-Institut für Eisenforschung GmbH for a short-term visit scholarship granted by Professor Dr. Martin Stratmann and the complementary support by the Incentive Fund Program of Costa Rica’s National Council for Science and Technology Research-(CONICIT) and the Ministry of Science and Technology (MICIT).
■
REFERENCES
(1) Huang, B.; Wu, H.; Bhaya, D.; Grossman, A.; Granier, S.; Kobilka, B. K.; Zare, R. N. Science 2007, 315, 81−84. (2) Eijkel, J. C. T.; van den Berg, A. Microfluid. Nanofluid. 2005, 1, 249−267. (3) Tegenfeldt, J. O.; Prinz, C.; Cao, H.; Huang, R. L.; Austin, R. H.; Chou, S. Y.; Cox, E. C.; Sturm, J. C. Anal. Bioanal. Chem. 2004, 378, 1678−1692. (4) Venkatesan, B. M.; Bashir, R. Nat. Nanotechnol. 2011, 6, 615− 624. (5) Tao, N. J. Nat. Nanotechnol. 2006, 1, 173−181. (6) Joo, C.; Balci, H.; Ishitsuka, Y.; Buranachai, C.; Ha, T. Annu. Rev. Biochem. 2008, 77, 51−76. (7) Kneipp, J.; Kneipp, H.; Kneipp, K. Chem. Soc. Rev. 2008, 37, 1052−1060. (8) Le Ru, E. C.; Etchegoin, P. G. Annu. Rev. Phys. Chem. 2012, 63, 65−87. (9) Pettinger, B.; Schambach, P.; Villagómez, C. J.; Scott, N. Annu. Rev. Phys. Chem. 2012, 63, 379−399. 2249
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
Nano Letters
Letter
(42) Schultze, J. W.; Hassel, A. W. Passivity of Metals, Alloys, and Semiconductors. In Encyclopedia of Electrochemistry; Bard, A., Stratmann, M., Frankel, G., Eds.; Wiley-VCH: Weinheim, 2007; Vol. 4, pp 216−235. (43) Yamada, H.; Yamamoto, Y. Surf. Sci. 1983, 134, 71−90. (44) Chen, Y.; Fang, Y. Spectrochim. Acta, Part A 2008, 69, 733−737. (45) Qin, W.; Yao, J. L.; Gu, R. A. Guangpuxue Yu Guangpu Fenxi (Spectrosc. Spect. Anal., Beijing, China) 2009, 29, 3300−3303. (46) Yang, L.; Jiang, X.; Ruan, W.; Zhao, B.; Xu, W.; Lombardi, J. R. J. Phys. Chem. C 2008, 112, 20095−20098. (47) Musumeci, A.; Gosztola, D.; Schiller, T.; Dimitrijevic, N. M.; Mujica, V.; Martin, D.; Rajh, T. J. Am. Chem. Soc. 2009, 131, 6040− 6041. (48) Yang, L.; Zhang, Y.; Ruan, W.; Zhao, B.; Xu, W.; Lombardi, J. R. J. Raman Spectrosc. 2010, 41, 721−726. (49) Green, N. G.; Morgan, H. J. Phys. D: Appl. Phys. 1998, 31, L25− L30. (50) Green, N. G.; Ramos, A.; Morgan, H. J. Phys. D 2000, 33, 632. (51) Goulian, M.; Simon, S. M. Biophys. J. 2000, 79, 2188−2198. (52) Glazer, A. N.; Stryer, L. Trends Biochem. Sci. 1984, 9, 423−427. (53) Parker, J. H.; Feldman, D. W.; Ashkin, M. Phys. Rev. 1967, 155, 712−714. (54) Beversluis, M. R.; Bouhelier, A.; Novotny, L. Phys. Rev. B 2003, 68, 115433. (55) Creighton, J. A.; Blatchford, C. G.; Albrecht, M. G. J. Chem. Soc., Faraday Trans. 2 1979, 75, 790−798. (56) Xu, Y.; Liang, L. Appl. Spectrosc. 1994, 48, 1147−1149. (57) Ward, D. R.; Halas, N. J.; Ciszek, J. W.; Tour, J. M.; Wu, Y.; Nordlander, P.; Natelson, D. Nano Lett. 2008, 8, 919−924. (58) Galperin, M.; Ratner, M. A.; Nitzan, A. J. Chem. Phys. 2009, 130, 144109. (59) Barth, A.; Zscherp, C. Q. Rev. Biophys. 2002, 369−430. (60) Mikhonin, A. V.; Ahmed, Z.; Ianoul, A.; Asher, S. A. J. Phys. Chem. B 2004, 108, 19020−19028. (61) Maeda, Y.; Yamamoto, H.; Kitano, H. J. Phys. Chem. 1995, 99, 4837−4841. (62) Contreras-Martel, C.; Martinez-Oyanedel, J.; Bunster, M.; Legrand, P.; Piras, C.; Vernede, X.; Fontecilla-Camps, J. C. Acta Crystallogr., Sect. D 2001, 57, 52−60. (63) Frushour, B. G.; Koenig, J. L. Biopolymers 1974, 13, 1809−1819. (64) Tuma, R. J. Raman Spectrosc. 2005, 36, 307−319. (65) Nyquist, R. A. Interpreting Infrared, Raman, and Nuclear Magnetic Resonance Spectra; Academic Press: San Diego, 2001; Vol. 1. (66) Goormaghtigh, E.; Cabiaux, V.; Ruysschaert, J. M. Subcell Biochem. 1994, 23, 329−362. (67) Porath, D.; Bezryadin, A.; de Vries, S.; Dekker, C. Nature 2000, 403, 635−638. (68) Schwan, H. P. Ann. N.Y. Acad. Sci. 1968, 148, 191−209. (69) Castellanos, A.; Ramos, A.; González, A.; Green, N. G.; Morgan, H. J. Phys. D 2003, 36, 2584. (70) Kirby, B. J. Micro- and Nanoscale Fluid Mechanics: Transport in Microfluidic Devices; Cambridge University Press: New York, 2010. (71) Ramos, A.; Morgan, H.; Green, N. G.; Castellanos, A. J. Phys. D 1998, 31, 2338. (72) Hess, S. T.; Huang, S. H.; Heikal, A. A.; Webb, W. W. Biochemistry 2002, 41, 697−705. (73) Zevenbergen, M. A. G.; Singh, P. S.; Goluch, E. D.; Wolfrum, B. L.; Lemay, S. G. Anal. Chem. 2009, 81, 8203−8212. (74) Hwang, T. W.; Branagan, S. P.; Bohn, P. W. J. Am. Chem. Soc. 2013, 135, 4522−4528. (75) Xu, B.; Tao, N. J. Science 2003, 301, 1221−1223. (76) Saraswat, S.; Desireddy, A.; Zheng, D. S.; Guo, L. J.; Lu, H. P.; Bigioni, T. P.; Isailovic, D. J. Phys. Chem. C 2011, 115, 17587−17593. (77) Hsu, H.-Y.; Chiang, H.-C.; Hu, J.-Y.; Awasthi, K.; Mai, C.-L.; Yeh, C.-Y.; Ohta, N.; Diau, E. W.-G. J. Phys. Chem. C 2013, 117, 24761−24766.
2250
dx.doi.org/10.1021/nl4046685 | Nano Lett. 2014, 14, 2242−2250
