Nanofluidic Redox Cycling Amplification for the Selective Detection of

Anal. Chem. 2008, 80, 972-977

Nanofluidic Redox Cycling Amplification for the Selective Detection of Catechol Bernhard Wolfrum, Marcel Zevenbergen, and Serge Lemay*

Kavli Institute of Nanoscience, Section Molecular Biophysics, Technical University Delft, Delft, 2628 CJ, The Netherlands

We have developed a chip-based nanofluidic device to amplify the electrochemical signal of catechols by orders of magnitude. The amplification is based on rapid redox cycling between plane parallel electrodes inside a nanochannel. We show that it is possible to monitor the signal of only a few hundred molecules residing in the active area of the nanofluidic sensor. Furthermore, due to the nanochannel design, the sensor is immune to interference by molecules undergoing irreversible redox reactions. We demonstrate the selectivity of the device by detecting catechol in the presence of ascorbic acid, whose oxidized form is only stable for a short time. The interference of ascorbic acid is usually a challenge in the detection of catecholamines in biological samples. Catechols play a significant role in many biochemical processes. For example, catecholamines such as dopamine and serotonin are important neurotransmitters in the central nervous system. Abnormal levels of catecholamines in the brain are linked to diseases such as Parkinson’s,1 and changes in plasma concentrations of different catechols act as indicators for several diseases.2 The selective detection of catechol traces in biological samples remains an experimental challenge. Especially in neurochemical investigations, it is desirable to monitor catecholamine levels with high spatial and temporal resolution. Electrochemical approaches are particularly interesting in this respect, since they can fulfill both of these requirements.3-7 During electrochemical detection of catechols, molecules are oxidized to their corresponding quinone at an appropriately biased electrode. The resulting oxidation current is proportional to quinone generation. In most biological samples, this detection method is limited by interference of other molecules, in particular ascorbic acid. Since both catechols and ascorbic acid exhibit similar oxidation potentials, the selective detection of catechols in the presence of ascorbic acid at an untreated electrode is not straightforward. * To whom correspondence should be addressed. E-mail: [email protected]. (1) Dekker, M. C. J.; Bonifati, V.; van Duijn, C. M. Brain 2003, 126, 17221733. (2) Goldstein, D. S.; Eisenhofer, G.; Kopin, I. J. J. Pharmacol. Exp. Ther. 2003, 305, 800. (3) Wightman, R. M.; Strope, E.; Plotsky, P. M.; Adams, R. N. Nature 1976, 262, 145-146. (4) Stamford, J. A. Brain Res. Rev. 1985, 10, 119-135. (5) Oneill, R. D. Analyst 1994, 119, 767-779. (6) Michael, D. J.; Wightman, R. M. J. Pharm. Biomed. Anal. 1999, 19, 3346. (7) Wightman, R. M. Science 2006, 311, 1570-1574.

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Concentration ratios between ascorbic acid and the catechol of interest can be very large, effectively masking any signal from the target molecules. For example, the plasma level of 3,4dihydroyphenylalanine in healthy individuals is usually below 10 nM,2 while ascorbic acid is present at concentrations above 30 µM.8 Addressing this problem, several techniques have been developed to suppress the signal originating from the oxidation of ascorbic acid or to separate it from the oxidation of catechols. Most approaches reported in the literature involve some kind of electrode surface modification9-19 including polymer coatings,20-29 application of self-assembled monolayers,30-32 enzymatic pretreatment,33 or the use of special electrode materials such as diamond (8) Riemersma, R. A.; Wood, D. A.; Macintyre, C. C. A.; Elton, R. A.; Gey, K. F.; Oliver, M. F. Lancet 1991, 337, 1-5. (9) Gonon, F.; Buda, M.; Cespuglio, R.; Jouvet, M.; Pujol, J. F. Nature 1980, 286, 902-904. (10) Falat, L.; Cheng, H. Y. Anal. Chem. 1982, 54, 2108-2111. (11) Downard, A. J.; Roddick, A. D.; Bond, A. M. Anal. Chim. Acta 1995, 317, 303-310. (12) Wang, J.; Walcarius, A. J. Electroanal. Chem. 1996, 407, 183-187. (13) Zhou, D. M.; Ju, H. X.; Chen, H. Y. J. Electroanal. Chem. 1996, 408, 219223. (14) Zen, J. M.; Chen, P. J. Anal. Chem. 1997, 69, 5087-5093. (15) Koktysh, D. S.; Liang, X. R.; Yun, B. G.; Pastoriza-Santos, I.; Matts, R. L.; Giersig, M.; Serra-Rodriguez, C.; Liz-Marzan, L. M.; Kotov, N. A. Adv. Funct. Mater. 2002, 12, 255-265. (16) Domenech, A.; Garcia, H.; Domenech-Carbo, M. T.; Galletero, M. S. Anal. Chem. 2002, 74, 562-569. (17) Zen, J. M.; Chung, H. H.; Kumar, A. S. Anal. Chem. 2002, 74, 1202-+. (18) Mani, R. C.; Sunkara, M. K.; Baldwin, R. P.; Gullapalli, J.; Chaney, J. A.; Bhimarasetti, G.; Cowley, J. M.; Rao, A. M.; Rao, R. H. J. Electrochem. Soc. 2005, 152, E154-E159. (19) Gopalan, A. I.; Lee, K. P.; Manesh, K. M.; Santhosh, P.; Kim, J. H.; Kang, J. S. Talanta 2007, 71, 1774-1781. (20) Rice, M. E.; Oke, A. F.; Bradberry, C. W.; Adams, R. N. Brain Res. 1985, 340, 151-155. (21) Kristensen, E. W.; Kuhr, W. G.; Wightman, R. M. Anal. Chem. 1987, 59, 1752-1757. (22) Kawagoe, K. T.; Jankowski, J. A.; Wightman, R. M. Anal. Chem. 1991, 63, 1589-1594. (23) Hsueh, C. C.; Brajtertoth, A. Anal. Chem. 1994, 66, 2458-2464. (24) Ciszewski, A.; Milczarek, G. Anal. Chem. 1999, 71, 1055-1061. (25) Selvaraju, T.; Ramaraj, R. Electrochem. Commun. 2003, 5, 667-672. (26) Roy, P. R.; Okajima, T.; Ohsaka, T. Bioelectrochemistry 2003, 59, 11-19. (27) Brown, F. O.; Lowry, J. P. Analyst 2003, 128, 700-705. (28) Raoof, J. B.; Ojani, R.; Rashid-Nadimi, S. Electrochim. Acta 2005, 50, 46944698. (29) Chen, S. M.; Chen, J. Y.; Vasantha, V. S. Electrochim. Acta 2006, 52, 455465. (30) Malem, F.; Mandler, D. Anal. Chem. 1993, 65, 37-41. (31) Dalmia, A.; Liu, C. C.; Savinell, R. F. J. Electroanal. Chem. 1997, 430, 205214. (32) Raj, C. R.; Okajima, T.; Ohsaka, T. J. Electroanal. Chem. 2003, 543, 127133. 10.1021/ac7016647 CCC: $40.75

© 2008 American Chemical Society Published on Web 01/15/2008

films34-36 and carbon nanotubes or carbon composites.37-44 For example, commonly used anionic polymer coatings such as Nafion increase the selectivity of dopamine detection by repelling the negatively charged ascorbic acid molecules from the oxidizing electrode. However, inside the polymer films, the diffusion constant of dopamine is decreased, which effectively reduces the response time of the detection.21 Another method to discriminate between oxidation of ascorbic acid and catechols is fast-scan voltammetry.45-50 In this technique, the signal of ascorbic acid is suppressed due to its slow electrontransfer kinetics. Background subtraction is used in combination with this method to obtain improved sensitivity and selectivity. An alternative approach to reduce the influence of ascorbic acid without electrode coating is based on redox cycling.51-53 In redox cycling, at least two electrodes are biased independently, one below the reduction potential and the other above the oxidation potential of the redox-active compound. Catechols, like dopamine, undergo reversible oxidation, in the sense that the generated quinone can be reduced back to the original molecule.54 In this case, the same molecule contributes several electrons to the recorded current. The oxidation product of ascorbic acid on the other hand undergoes a fast irreversible hydration reaction, rendering the molecule inactive for reduction.55 This transition occurs on the order of milliseconds and can be exploited to remove the influence of ascorbic acid on redox cycling. Interdigitated arrays have been successfully employed to amplify electro(33) Lisdat, F.; Wollenberger, U.; Makower, A.; Hortnagl, H.; Pfeiffer, D.; Scheller, F. W. Biosens. Bioelectron. 1997, 12, 1199. (34) Popa, E.; Notsu, H.; Miwa, T.; Tryk, D. A.; Fujishima, A. Electrochem. Solid State Lett. 1999, 2, 49-51. (35) Cvacka, J.; Quaiserova, V.; Park, J.; Show, Y.; Muck, A.; Swain, G. M. Anal. Chem. 2003, 75, 2678-2687. (36) Shin, D. C.; Sarada, B. V.; Tryk, D. A.; Fujishima, A. Anal. Chem. 2003, 75, 530-534. (37) Wang, Z. H.; Liang, Q. L.; Wang, Y. M.; Luo, G. A. J. Electroanal. Chem. 2003, 540, 129-134. (38) Wu, K. B.; Fei, J. J.; Hu, S. S. Anal. Biochem. 2003, 318, 100-106. (39) Chen, R. S.; Huang, W. H.; Tong, H.; Wang, Z. L.; Cheng, J. K. Anal. Chem. 2003, 75, 6341-6345. (40) Valentini, F.; Amine, A.; Orlanducci, S.; Terranova, M. L.; Palleschi, G. Anal. Chem. 2003, 75, 5413-5421. (41) Zhang, M. N.; Gong, K. P.; Zhang, H. W.; Mao, L. Q. Biosens. Bioelectron. 2005, 20, 1270-1276. (42) Maleki, N.; Safavi, A.; Tajabadi, F. Anal. Chem. 2006, 78, 3820-3826. (43) Maldonado, S.; Morin, S.; Stevenson, K. J. Analyst 2006, 131, 262-267. (44) Ali, S. R.; Ma, Y. F.; Parajuli, R. R.; Balogun, Y.; Lai, W. Y. C.; He, H. X. Anal. Chem. 2007, 79, 2583-2587. (45) Stamford, J. A.; Kruk, Z. L.; Millar, J.; Wightman, R. M. Neurosci. Lett. 1984, 51, 133-138. (46) Baur, J. E.; Kristensen, E. W.; May, L. J.; Wiedemann, D. J.; Wightman, R. M. Anal. Chem. 1988, 60, 1268-1272. (47) Cahill, P. S.; Walker, Q. D.; Finnegan, J. M.; Mickelson, G. E.; Travis, E. R.; Wightman, R. M. Anal. Chem. 1996, 68, 3180-3186. (48) Hsueh, C.; Bravo, R.; Jaramillo, A. J.; BrajterToth, A. Anal. Chim. Acta 1997, 349, 67-76. (49) Venton, B. J.; Troyer, K. P.; Wightman, R. M. Anal. Chem. 2002, 74, 539546. (50) Heien, M.; Khan, A. S.; Ariansen, J. L.; Cheer, J. F.; Phillips, P. E. M.; Wassum, K. M.; Wightman, R. M. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 10023-10028. (51) Vandaveer, W. R.; Woodward, D. J.; Fritsch, I. Electrochim. Acta 2003, 48, 3341-3348. (52) Hayashi, K.; Iwasaki, Y.; Horiuchi, T.; Sunagawa, K.; Tate, A. Anal. Chem. 2005, 77, 5236-5242. (53) Paixao, T.; Richter, E. M.; Brito-Neto, J. G. A.; Bertotti, M. Electrochem. Commun. 2006, 8, 9-14. (54) Aoki, A.; Matsue, T.; Uchida, I. Anal. Chem. 1990, 62, 2206-2210. (55) Wehmeyer, K. R.; Wightman, R. M. Anal. Chem. 1985, 57, 1989-1993.

Figure 1. Scheme for catechol redox cycling inside a fluidic nanochannel. Catechol is oxidized at the upper electrode to the corresponding quinone. After diffusing to the bottom electrode, quinone is reversely reduced back to catechol. Every redox cycle, two electrons are shuttled from one electrode to the other, giving rise to an electrochemical current.

chemical detection of dopamine and similar catechol-related compounds by redox cycling.54,56-62 The efficiency of diffusionlimited redox cycling thereby depends critically on the distance of the reducing and oxidizing electrode.63-66 Here we show that redox cycling between plane parallel electrodes inside a nanochannel as illustrated in Figure 1 can be used to selectively amplify the signal of catechol by orders of magnitude. EXPERIMENTAL SECTION Reagents. Catechol and phosphate buffer were obtained from Sigma-Aldrich (Steinheim, Germany). Chromium etch (Selectipur) and ascorbic acid were purchased from Merck (Darmstadt, Germany). All chemicals were used without further purification. The solutions were passed through an Anotop 20-nm filter (Whatman, Madistone, England) before use. Device Fabrication. The nanofluidic redox cycling amplifier was fabricated on a silicon substrate employing five consecutive electron beam exposures on PMMA resist. Figure 2 shows a sketch of the fabrication scheme. The silicon substrate was thermally oxidized under wet conditions to grow 500 nm of SiO2. A bottom electrode of 4-µm width and ∼94-µm length was patterned via liftoff by evaporating 4 nm of Ti, 50 nm of Au, and 1 nm of Cr. Titanium and chromium were used to facilitate adhesion to the substrate and the insulating layer. In a next step, a sacrificial chromium layer of 3.5-µm width, ∼94-µm length, and ∼53-nm height was patterned via liftoff and e-beam evaporation on top of the bottom electrode. The sacrificial layer defined the geometrical shape of the nanochannel and was aligned in the center of the bottom electrode. Subsequently, the bottom electrode and sacrificial layer were insulated by sputtering 200 nm of silicon oxide. (56) Niwa, O.; Morita, M.; Tabei, H. Electroanalysis 1991, 3, 163-168. (57) Martin, R. S.; Gawron, A. J.; Lunte, S. M.; Henry, C. S. Anal. Chem. 2000, 72, 3196-3202. (58) Liu, Z. M.; Niwa, O.; Kurita, R.; Horiuchi, T. Anal. Chem. 2000, 72, 13151321. (59) Hayashi, K.; Iwasaki, Y.; Kurita, R.; Sunagawa, K.; Niwa, O. Electrochem. Commun. 2003, 5, 1037-1042. (60) Male, K. B.; Luong, J. H. T. J. Chromatogr., A 2003, 1003, 167-178. (61) Nebling, E.; Grunwald, T.; Albers, J.; Schafer, P.; Hintsche, R. Anal. Chem. 2004, 76, 689-696. (62) Dam, V. A. T.; Olthuis, W.; van den Berg, A. Analyst 2007, 132, 365-370. (63) Bard, A. J.; Crayston, J. A.; Kittlesen, G. P.; Shea, T. V.; Wrighton, M. S. Anal. Chem. 1986, 58, 2321-2331. (64) Fan, F. R. F.; Bard, A. J. Science 1995, 267, 871. (65) Ueno, K.; Hayashida, M.; Ye, J. Y.; Misawa, H. Electrochem. Commun. 2005, 7, 161-165. (66) Zevenbergen, M. A. G.; Krapf, D.; Zuiddam, M. R.; Lemay, S. G. Nano Lett. 2007, 7, 384-388.

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Figure 2. Sketch of the fabrication process of the nanofluidic device. First the bottom electrode is patterned (a) followed by deposition of a sacrificial chromium layer (b). SiO2 is sputtered to passivate the bottom layer (c). Access patterns are then etched (d) allowing the deposition of the top electrodes (e). The whole structure is insulated by sputtered SiO2 before access holes are etched (g) and the sacrificial chromium layer is removed (h).

Figure 3. Microscopic top-view image of a typical device just before the removal of the sacrificial chromium layer. Feed lines connect to the bottom electrode and three top electrodes. Access holes are visible at the end of the channels.

Three wells were etched through the passivation layer by reactive ion etching (CHF3,O2) to allow for a connection of the top electrodes to the sacrificial layer. Afterward, the top electrodes were deposited by e-beam evaporation (1 nm of Cr, 60 nm of Au, 4 nm of Ti) and liftoff on top of the sacrificial layer. The device was insulated again by sputtering 350 nm of silicon oxide. Two access holes with radii between 250 nm and 1.5 µm were etched at the outer ends of the bottom electrode for fluidic access. In a final step, the sacrificial layer was chemically removed using a wet chromium etch at room temperature. This step generates the nanochannel. The etching procedure was monitored either optically or electronically by measuring the resistance between the top and bottom electrodes. A microscope image of a device just before removal of the chromium layer is shown in Figure 3. Electrochemical Measurements. Electrochemical measurements were performed in aqueous solutions using a bipotentiostat (CHI 832B, CH Instruments). The solutions of catechol and ascorbic acid were prepared in a background electrolyte of 100 mM phosphate buffer adjusted to a pH of 7.2. A commercial Ag/ AgCl electrode (BASi, Stareton, UK) was used as a combined reference and counter electrode. Due to the low currents running through the reference electrode in our measurements (,1 nA), 974

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a separate counter electrode was unnecessary. One of the top electrodes and the bottom electrode inside the nanofluidic channel served as working electrodes. In all experiments shown, the bottom electrode was either disconnected (one-electrode mode) or set to a constant potential (redox cycling mode), while one of the top electrodes was swept to obtain a cyclic voltammogram. Although in principle all three top electrodes can be addressed independently, we used only one of the top electrodes for the measurements presented in this study. The two unused top electrodes were left floating and did not influence the results. We conditioned our devices for 30 s at the initial scan potential before recording electrochemical currents to ensure identical starting conditions for each experiment. A flow cell was connected to the chip to provide a fluidic interface and to integrate the reference electrode in the setup. After experiments, voltage sweeps from +800 to -300 mV versus Ag/AgCl were performed in pure phosphate buffer. The chips were then cleaned in MilliQ water, dried with argon gas, and stored for further reuse. Devices could be used multiple times. All the experiments were conducted in a Faraday cage to reduce electromagnetic interference. RESULTS AND DISCUSSION In a first series of experiments, we tested the response of the nanofluidic device to catechol in both one-electrode and redox cycling modes. Figure 4 shows cyclic voltammograms after flushing the nanofluidic device (channel height, ∼55 nm) with 100 µM catechol. The voltammogram was obtained by sweeping the center top electrode. If only one of the top microelectrodes was connected and the bottom electrode was left floating, redox cycling did not occur (Figure 4a). In this case, we monitored a small response with a maximum current of a few picoamperes on top of the background current. Apart from current passing through pinholes in the passivation layer, the maximum current is limited by the linear diffusion of molecules from the channel entrances to the electrode. Therefore, the length of the channel and the size of the entrance holes determine the ideal current response in this configuration. However, shape and size of the electrochemical signal depend strongly on the quality of the insulating layer. Due to the small signal obtained in one-electrode mode, leakage currents through the passivation cause deterioration of both the shape and the amplitude of the voltammogram. As expected for a system governed by linear diffusion, the example

Figure 4. Cyclic voltammogram of 100 µM catechol in 100 mM phosphate buffer. The top electrode was swept at 40 mV s-1, and the bottom electrode was either disconnected (black curve) or kept at a potential of -300 mV vs Ag/AgCl (top electrode: oxidation current, red curve; bottom electrode: reduction current, blue curve).

shown in Figure 4a displays an oxidative current peak as well as a reductive peak. The reductive peak is due to remaining quinone after oxidation of catechol. Redox cycling was enabled by additionally connecting the bottom electrode and biasing it below the reduction potential of quinone. Under this condition, molecules in the nanofluidic sensor underwent repetitive oxidation and reduction at the electrodes as sketched in Figure 1. The red and the blue curves in Figure 4b show the oxidation and reduction current of catechol and quinone inside the nanofluidic channel, respectively. The peak oxidation current is ∼3 orders of magnitudes larger than the oxidation current observed in single-electrode mode (black curve). This large difference in maximum currents illustrates the effectiveness of redox cycling amplification inside a nanochannel. As expected, the cycling efficiency inside the nanochannel is very close to 100%, which can be seen by comparing oxidation and reduction current: iox ≈ -ired. The background leakage observed in one-electrode mode is therefore rendered irrelevant. In principle, one would assume that the current response reaches a steady state above the oxidation potential of catechol, giving rise to a sigmoidal shape of the voltammogram. However, at potentials above ∼300 mV, we observed a decrease in redox cycling current with increasing voltage, which was especially pronounced while scanning in the oxidation direction (Figure 4b).

It suggests a reversible decrease of charge transfer at high potentials in the nanofluidic device. This voltammogram shape, including the current decrease and the shift to higher oxidation potential, was encountered for all scanning speeds (4-1000 mV/ s) and concentrations (100 nM-100 µM) investigated under redox cycling conditions. However, it did not occur in measurements performed with a standard 5-µm-radius gold microelectrode (BASi), where we observed a diffusion-limited plateau (data not shown). There are several possible causes that can contribute to this behavior, including potential induced changes at the electrode surface caused by modifications of the electrodes during the nanochannel fabrication, kinetic limitations of electron transfer, local pH changes due to fast proton liberation at the oxidizing electrode, and reversible adhesion of molecules to the electrode surface. The latter will have two effects, namely, the depletion of redox-active molecules from the nanochannel and the reduction of electron transfer due to blocking of the electrodes. A third effect, the oxidation and reduction of adsorbed molecules at the electrodes, is negligible because we observe almost the exact inverse current on the oxidizing and reducing electrode. Because the shape is preserved over a wide range of concentrations and scanning parameters, the deviation from the sigmoidal shape does not affect the use of the device in analytical applications. For these purposes, we measured the peak current of the catechol signature to quantify the concentration. The diffusion-limited maximum current in redox cycling mode is governed by the average number of molecules between the electrode and the height of the nanochannel. Due to the high background electrolyte of 100 mM phosphate buffer (Debye layer ∼1 nm), the electric field caused by the potential of the electrodes is not expected to have a significant influence on the current. Assuming that the electron transport is governed by the diffusion of catechol and neglecting effects of proton diffusion and possible breakdown of electroneutrality,67 we can estimate the diffusionlimited current in redox cycling mode to be

icycl )

< n > Dze h2

where ze is the transferred charge upon oxidation and reduction, h is the height of the channel, and D ) 7.6 × 10 - 10 m2 s-1 is the diffusion constant of catechol.68 For comparison, the steady-state oxidation current of a disk microelectrode with the same area (∼80 µm2, r ) 5.05 µm) has a radial diffusion-limited current of

ime )

4 h i π r cycl

()

Considering a channel of 55-nm height, the theoretical ratio of electrochemical currents is therefore icycl/ime ) 72. At a catechol concentration of 100 µM, we expect an average number of 2.65 × 105 molecules in the active volume of the employed channel (∼4.4 × 10-15 L). The estimated diffusionlimited current for a two-electron transfer in our channel is icycl ) 21 nA, which is more than twice as large as the observed peak (67) Smith, C. P.; White, H. S. Anal. Chem. 1993, 65, 3343-3353. (68) Silva Luz, R. D. C.; Damos, F. S.; de Oliveira, A. B.; Beck, J.; Kubota, L. T. Sens. Actuators, B: Chem. 2006, 117, 274-281.

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Figure 5. Peak oxidation current versus catechol concentration taken from cyclic voltammograms (third scan) at 10 (blue diamonds) and 40 mV s-1 (red circles). The line shows a linear fit with a sensitivity of 1.1 × 106 A M-1 m-2.

current in Figure 4b. The discrepancy is probably caused by kinetic limitations as discussed above. Partial insulation of the electrode interfaces by PMMA remains or inhomogeneities in the channel height might also contribute to a decrease in current. Figure 5 shows the dependence of peak current on catechol concentration in the range of 100 nM-100 µM for two different scan rates. The cyclic voltammograms from which the peak currents were derived were taken at 40 (red circles) and 10 mV/s (blue diamonds). The line depicts a linear fit with a sensitivity of 1.1 × 106 A M-1 m-2, which is to our knowledge the highest current density reported for catechol so far. Assuming that the whole electrode surface contributes to the signal, this corresponds to a current of 33 fA for each single molecule residing inside the active area of the nanochannel. Under the same conservative assumption, the data taken at a 100 nM concentration reflects the current of an average number of 265 molecules inside the nanochannel. For this concentration, we measured a signal-tonoise (rms) ratio on the order of 100 when scanning at 10 mV/s and applying an internal 15-Hz low-pass filter of the potentiostat. The noise is mainly caused by diffusive fluctuations of the number of molecules inside the channel.66 An important issue in the detection of catechols in biological samples is the interference of ascorbic acid. Since the oxidation product of ascorbic acid is only stable on the order of milliseconds,55 its influence on the amplified redox cycling currents will decay after very short time scales inside the nanochannel. Therefore, the large amplification factors achieved for catechol in redox cycling mode permit its detection even in the presence of high ascorbic acid concentration. In our device, the bottom electrode can be effectively used to deplete the channel of ascorbic acid by oxidation at high potentials. Sweeping the top electrode to low potentials, the reduction current of the generated quinone undergoes redox cycling amplification without interference of ascorbic acid. Depending on which interfering molecules are present in the sample, either the reduction or the oxidation current can therefore be used to monitor catechol concentration. Figure 6 shows current voltage scans performed in 45 µM catechol solutions in the presence of ascorbic acid. The voltammograms were recorded consecutively in the same nanofluidic device using either the one-electrode (a) or the redox cycling 976 Analytical Chemistry, Vol. 80, No. 4, February 15, 2008

Figure 6. Cyclic voltammograms (third scan) of 45 µM catechol in the presence of ascorbic acid in one-electrode (a) and redox cycling mode (b). The scans were performed with an outer top electrode of the nanochannel (scan rate 50 mV/s). Individual colors reflect different levels of ascorbic acid: blue, 500 µM; green, 100 µM; red, 0 µM. The black curve (b) was recorded in background electrolyte (100 mM phosphate buffer, pH 7.2). In (a), the bottom electrode remains floating; in (b) it is set to +600 mV.

mode (b). Ascorbic acid concentrations are 500 (blue curve), 100 (green curve), or 0 µM (red curve). After these scans, pure phosphate buffer was flushed through the device to reveal the background current (black curve). In one-electrode mode, the oxidation currents depend on the concentration of ascorbic acid. This is to be expected since ascorbic acid from solution can diffuse through the entrances of the device to the top electrode, where it is oxidized. Figure 6b shows data of the same solutions in redox cycling mode. The bottom electrode was set to +600 mV to oxidize catechol and ascorbic acid. The top electrode was swept toward negative potentials and monitored the electrochemical reduction of quinone previously generated at the bottom electrode. We can see in Figure 4b that the presence of ascorbic acid does not interfere with the reduction current of catechol. Although ascorbic acid will be oxidized at the bottom electrode,, it cannot be reduced back at the top electrode. The short lifetime of oxidized ascorbic acid makes it possible to detect catechol independent of ascorbic acid by monitoring the reduction of quinone in redox cycling mode. Since oxygen reduction is also not amplified during redox cycling, the potential window in which the device can be operated

ascorbic acid concentrations hiding the actual catechol signal at high concentrations of ascorbic acid. In redox cycling mode, we also observe an increase in the oxidation current with increasing ascorbic acid concentration (red curve). However, only the signal originating from catechol is amplified by redox cycling. This selective amplification allows for relatively stable overall oxidation currents even at high ascorbic acid concentrations. As shown in Figure 6b, the reduction current in redox cycling mode is independent of ascorbic acid concentration (Figure 7b, blue curve). In this case, only the previously generated quinone contributes to the overall signal, making it possible to detect catechol without interference of ascorbic acid even at high concentrations. For analytical purposes, the stability of the devices and reproducibility of redox cycling current is an important issue. As expected, a decrease in current response was observed during prolonged oxidation especially at high catechol concentrations caused by electrode fouling. However, the amplification factors of the devices could be regenerated within a few percent of their initial value by setting all electrodes to -300 mV Ag/AgCl for 30 s. A sophisticated cleaning procedure as developed by Manica et al.69 or antifouling coating on the electrodes70 will contribute to further enhancing stability.

Figure 7. Catechol (45 µM) oxidation and reduction currents depending on ascorbic acid concentration. (a) Absolute currents; (b) current normalized to catechol solution without ascorbic acid. Oxidation currents (red and magenta curve) were recorded at 350 mV vs Ag/AgCl; reduction currents (blue and black curve) were recorded at -200 mV vs Ag/AgCl. During redox cycling, the bottom electrode was biased to -300 and +600 mV vs Ag/AgCl, while measuring oxidation and reduction current at the top electrode, respectively.

is significantly larger than for conventional electrochemistry detection in the presence of oxygen. The operating potential is basically only limited by the oxidation and reduction of water, which starts to contribute significantly (>20 pA) below a potential of -700 mV and above +800 mV versus Ag/AgCl. Figure 7 shows oxidation and reduction currents in 45 µM catechol dependent on ascorbic acid concentrations. In oneelectrode mode, there is almost no reduction current (black curve) since the solution does not contain any quinone and consequently no reducible components. However, at oxidizing potentials (magenta curve), both catechol and ascorbic acid contribute to the signal. We therefore obtain increasing currents with increasing (69) Manica, D. P.; Mitsumori, Y.; Ewing, A. G. Anal. Chem. 2003, 75, 45724577. (70) Spegel, C.; Heiskanen, A.; Acklid, J.; Wolff, A.; Taboryski, R.; Emneus, J.; Ruzgas, T. Electroanalysis 2007, 19, 263-271.

CONCLUSION A technique based on redox cycling inside a nanofluidic channel of ∼55-nm height has been developed for the detection of low catechol concentrations in small volume samples (4.4 × 10-15 L). The large amplification factors resulting from redox cycling between closely spaced electrodes integrated in the nanochannel permit measuring the electrochemical signal of only a few hundred molecules. Further reduction of the electrode distance promises to allow single-molecule measurements. We have shown that our technique is capable of monitoring catechol concentrations without interference by ascorbic acid. Redox cycling selectively amplifies the catechol signal without interference by ascorbic acid. Additionally, the chip-based approach facilitates the integration of sensor arrays in combination with other on-chip detection methods. Although the prototypes in the present study are fabricated with electron beam lithography, it should be possible to transfer the process to optical lithography allowing large-scale wafer production for “lab on a chip” applications. ACKNOWLEDGMENT We thank Dirk Heering and Cees Dekker for their helpful discussion.

Received for review August 5, 2007. Accepted November 21, 2007. AC7016647

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