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Langmuir 2006, 22, 10796-10800
Comparison of the Signaling and Stability of Electrochemical DNA Sensors Fabricated from 6- or 11-Carbon Self-Assembled Monolayers† Rebecca Y. Lai,‡,§,| Dwight S. Seferos,‡,§ Alan J. Heeger,‡,§,|,⊥ Guillermo C. Bazan,‡,§,⊥ and Kevin W. Plaxco*,‡,§,# Center for Polymers and Organic Solids, Department of Chemistry and Biochemistry, Department of Physics, Materials Department, Biomolecular Science and Engineering Program, UniVersity of California, Santa Barbara, Santa Barbara, California 93106 ReceiVed April 29, 2006. In Final Form: June 13, 2006 We have characterized the solution-phase and dry storage stability of electrochemical E-DNA sensors fabricated using mixed self-assembled monolayers (SAMs) composed of 6- or 11-carbon (C6 and C11, respectively) R,ω-thiol alcohols and the analogous C6- or C11-thiol-terminated stem-loop DNA probe. We find that the solution-phase and dry storage stability of C6-based E-DNA sensors are limited and poorly reproducible. The use of stabilizing agents bovine serum albumin plus either glucose or trehalose significantly improves the dry storage shelf life of such sensors: when using these preservatives, we observe only 7-9% sensor degradation after 1 month of storage in air at room temperature. In comparison, the stability of C11-based E-DNA sensors is significantly greater than that of the C6-based sensors; we observe only minor (5-8%) loss of signal upon storing these sensors for a week under ambient solution conditions or for more than a month in air in the presence of preservatives. Moreover, whereas the electron-transfer rate through C11 SAMs is slower than that observed for C6 SAMs, it is rapid enough to support good sensor performance. It thus appears that C11 SAMs provide a reasonable compromise between electron-transfer efficiency and sensor stability and are well suited for use in electronic DNA-sensing applications.
Part of the Electrochemistry special issue. * Corresponding author. E-mail:
[email protected]. Phone: (805) 893-5558. Fax: (805) 893-4120. ‡ Center for Polymers and Organic Solids. § Department of Chemistry and Biochemistry. | Department of Physics. ⊥ Materials Department. # Biomolecular Science and Engineering Program.
2 to 11 carbons have been described,11-13 very few studies of the stability of these sensors have been reported.14,15 The optimal SAM thickness for electrochemical biosensor applications reflects a compromise between two competing effects. First, because of improved van der Waals interactions longer-chain alkanes pack more tightly and form more stable monolayers, and thus SAM stability increases monotonically with alkane chain length.3,4 It has also been suggested that this tight packing reduces the oxidation of the gold-sulfur bond, further enhancing the stability of longer-chain SAMs.16-18 In contrast to SAM stability, however, through-SAM electrontransfer efficiency drops off exponentially with alkane chain length.5-7 Reduced electron-transfer efficiency in turn reduces sensor current, potentially hindering sensor performance. The ideal SAM thickness for electrochemical biosensor applications will thus represent a compromise between these competing effects. Here we describe the electron-transfer characteristics, operational stability, and shelf lives of electronic E-DNA sensors11,19,20 fabricated using mixed SAMS composed of either C6 or C11 R,ω-thiol alcohols interspersed with the analogous C6 or C11 thiol-modified DNA probes. (Figure 1) The E-DNA sensor, along with its aptamer-based equivalent for protein and small-molecule detection,21-23 is the electrochemical analogue of optical mo-
(1) Love, J. C.; Estroff, L. A.; Kriebel, J. K.; Nuzzo, R. G.; Whitesides, G. M. Chem. ReV. 2005, 105, 1103-1170. (2) Gooding, J. J. Electroanalysis 2002, 14, 1149-1156. (3) Bain C. D.; Troughton, E. B.; Tao, Y.-T.; Evall, J.; Whitesides, G. M.; Nuzzo, R. G. J. Am. Chem. Soc. 1989, 111, 321-335. (4) Poirier, G. E.; Tarlov, M. J.; Rushmeier, H. E. Langmuir 1994, 10, 33833386. (5) Smalley, J. F.; Feldberg, S. W.; Chidsey, C. E. D.; Linford, M. R.; Newton, M. D.; Liu, Y.-P. J. Phys. Chem. 1995, 99, 13141-13149. (6) Miller, C.; Cuendet, P.; Graetzel, M. J. Phys. Chem. 1991, 95, 877-886. (7) Lee, T.; Wang, W.; Reed, M. A. Ann. N.Y. Acad. Sci. 2003, 1006, 21-35. (8) Xia, N.; Shumaker-Parry, J. S.; Zareie, M. H.; Campbell, C. T.; Castner, D. G. Langmuir, 2004, 20, 3710-3716. (9) Stuart, D. A.; Yonzon, C. R.; Zhang, X.; Lyandres, O.; Chah, N. C.; Glucksberg, M. R.; Walsh, J. T.; Van Duyne, R. P. Anal. Chem. 2005, 77, 40134019. (10) Flynn, N. T.; Tran, T. N. T.; Cima, M. J.; Langer, R. Langmuir 2003, 19, 10909-10915.
(11) Fan, C., Plaxco, K. W.; Heeger, A. J. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 9134-9137. (12) Radi, A.-E.; Sanchez, J. L. A.; Baldrich, E.; O’Sullivan, C. K. Anal. Chem. 2005, 77, 6320-6323. (13) Jhaveri, S. D.; Mauro, M.; Goldston, H. M.; Schauer, C. L.; Tender, L. M. Chem. Commun. 2003, 338-339. (14) Baldrich, E.; Acero, J. L.; Reekmans, G.; Laureyn, W.; O’Sullivan, C. K. Anal. Chem. 2005, 77, 4774-4784. (15) Wong, E. L. S.; Chow, E.; Gooding, J. J. Langmuir 2005, 21, 6957-6965. (16) Finklea, H. O.; Avery, S.; Lynch, M.; Furtsch, T. Langmuir 1987, 3, 409-413. (17) Li, Y.; Huang, J.; McIver, R. T.; Hemminger, J. C. J. Am. Chem. Soc. 1992, 114, 2428-2432. (18) Tarlov, M. J.; Newman, J. C. Langmuir, 1992, 8, 1398-1405. (19) Immoos, C. E.; Lee, S. J.; Grinstaff, M. W. J. Am. Chem. Soc. 2004, 126, 10814-10815. (20) Mao, Y.; Luo, C.; Ouyang, Q. Nucleic Acids Res. 2003, 31, e108.
Introduction The formation of self-assembled alkanethiol monolayers (SAMs) on gold offers a convenient means by which biomolecules can be grafted onto conductive substrates for electronic biosensor applications.1,2 In addition to ease of fabrication, the utility of SAMs is also a function of their stability in solution, and under dry storage conditions, these factors are often critical elements in defining the operational lifetimes and shelf lives of SAMbased biosensors. To date, however, the extent to which SAM thickness, which is linked to SAM stability3,4 and through-SAM electron-transfer efficiency,5-7 affects sensing and sensor stability has seen only limited investigation in the open literature.8-10 For example, whereas electron transfer-based biosensors fabricated using mixed alkanethiol SAMs (composed of DNA-modified thiols interspersed within a SAM of thiol-alcohols) ranging from †
10.1021/la0611817 CCC: $33.50 © 2006 American Chemical Society Published on Web 08/03/2006
Signaling, Stability of Electrochemical DNA Sensors
Langmuir, Vol. 22, No. 25, 2006 10797
Figure 1. E-DNA sensor fabricated by the self-assembly of a redox-tagged, thiol-modified DNA probe on a gold electrode surface. (The analogous C6 or C11 alkane thiol monolayer in which the alkane modification of the DNA is embedded has been omitted for clarity.) In the absence of the target, the stem-loop structure is thought to hold the methylene blue (MB) redox tag in proximity to the electrode surface, thus enabling efficient electron transfer. Upon hybridization with the target DNA, a large change in the reduction peak current of MB is observed.
lecular beacons. Because the E-DNA platform is reagentless and readily reusable24 (the latter allowing, for example, the repeated monitoring of a single sensor during storage), it is well suited to studies of the impact of SAM thickness on electronic DNA sensor performance. Materials and Methods Materials. The reagents iodine (I2), triethylamine (Et3N), ethanol (EtOH), methanol (MeOH), 4,4′dimethoxytrityl chloride (DMTCl), 4-(dimethylamino)pyridine (DMAP), pyridine, 2-cyanoethyl-N,N′diisopropylchlorophosphoramidite, diisopropylethylamine, dichloromethane, ethyl acetate, deuterated chloroform (CDCl3), and sodium bicarbonate (NaHCO3) (all reagent grade) and bovine serum albumin (BSA), glucose, trehalose, hydrogen peroxide (30%), concentrated sulfuric acid, guanidine hydrochloride, 20× saline-sodium citrate buffer (SSC), 6-mercapto-1-hexanol (C6-OH), 11-mercapto-1undecanol (C11-OH), and iron-supplemented fetal calf serum were all used as received (from Sigma-Aldrich, St. Louis, MO). The probe DNA, thiol-, and amine-modified oligonucleotides containing both the complement to the Salmonella gyrB gene and stem-forming termini were obtained from Biosource (Foster City, CA).25 A methylene blue (MB) reporter group was conjugated to the 3′-end of the amino- and thiol-modified stem-loop oligonucleotide through succinimide ester coupling (MB-NHS, EMP Biotech, Germany) during synthesis. C6-thiol-modified probe DNA was fabricated using a commercially available C6-thiol phosphoramidite. The equivalent C11 probe DNA was synthesized using the equivalent 11-carbon phosphoramidites11-O-(4,4′-dimethoxytrityl)undecanol11′-O-((2-cyanoethyl)-N,N′-diisopropylphosphoramidite)undecanol1,1′-disulfide (C11 linker)swhich was synthesized as described below and provided to Biosource for use in synthesis. Oligomer 1 (C6-DNA): 5′-HS-(CH2)6-GCAGTAACAAGAATAAAACGCCACTGC(CH2)7-NH2-3-MB. Oligomer 2 (C11-DNA): 5′-HS-(CH2)11-GCAGTAACAAGAATAAAACGCCACTGC(CH2)7-NH2-3-MB (21) Xiao, Y.; Piorek, B. D.; Plaxco, K. W.; Heeger, A. J. J. Am. Chem. Soc. 2005, 127, 17990-17991. (22) Baker, B. R.; Lai, R. Y.; Wood, M. S.; Doctor, E. H.; Heeger, A. J.; Plaxco, K. W. J. Am. Chem. Soc. 2006, 128, 3138-3139. (23) Radi, A.-E.; Acero Sanchez, J. L.; Baldrich, E.; O’Sullivan, C. K. J. Am. Chem. Soc. 2006, 128, 117-124. (24) Lubin, A. A.; Lai, R. Y.; Baker, B. R.; Heeger, A. J.; Plaxco, K. W. Anal. Chem., submitted for publication, 2006. (25) Lai, R. Y.; Lagally, E. T.; Lee, S.-H.; Soh, H. T.; Plaxco, K. W.; Heeger, A. J. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 4017-4021.
The target DNA sequence was obtained from Genosys (SigmaAldrich, St. Louis, MO). Oligomer 3 (target DNA): 5′-GGCGTTTTATTCTTGTT-3′. Synthesis and Characterization of 11-O-(4,4′-Dimethoxytrityl)undecanol-11′-O-((2-cyanoethyl)-N,N′-diisopropylphosphoramidite)undecanol-1,1′-disulfide. A solution of 11-mercapto-1undecanol (500 mg, 2.45 mmol) and triethylamine (1.1 mL, 7.6 mmol) in ice-cold ethanol/methanol (5 mL, 1:1) was treated dropwise with a solution of iodine (311 mg, 1.22 mmol) in ethanol (10 mL) in a round-bottomed flask equipped with a Teflon-coated magnetic stir bar maintaining rapid stirring. The reaction was allowed to warm to room temperature and was monitored by thin-layer chromatography. After complete conversion, the solvent was evaporated, and the crude material was purified by column chromatography on silica gel eluting with ethyl acetate in hexanes (1:1) to afford 450 mg (90%) of the desired product. 1H NMR (CDCl3): δ 3.647 (t, 3J ) 6.70 Hz, 4H), 2.684 (t, 3J ) 7.54 Hz, 4H), 1.674 (quint, 3J ) 7.54 Hz, 4H), 1.572, (m, 4H), 1.2-1.4 (m, 28H). 13C NMR (CDCl3): δ 63.3, 39.4, 33.0, 29.8, 29.72, 29.69, 29.44, 29.41, 28.71. HRMS-EI: 406.2955; ∆ ) 3.9 ppm. 11-O-(4,4′-Dimethoxytrityl)undecanol-11′-undecanol-1,1′-disulfide. 11,11′-Diundecanol-1,1′-disulfide (468 mg, 1.15 mmol) was dissolved in anhydrous pyridine (5 mL) and evaporated to dryness under reduced pressure in a round-bottomed flask equipped with a Teflon-coated magnetic stir bar and a needle valve. An argon atmosphere was created in the flask containing the solids, pyridine (8.0 mL) and DMAP (10 mg, 0.8 mmol) were added, and DMTCl (430 mg, 1.27 mmol) was added portionwise with constant stirring. After 24 h, the volatiles were removed under reduced pressure, and the crude material was dissolved in dichloromethane (50 mL), washed with several portions of a saturated solution of NaHCO3, dried, and concentrated. Purification by column chromatography on silica gel eluting with ethyl acetate in dichloromethane (1:19) afforded 377 mg (46%) of the desired product. 1H NMR (CDCl3): δ 7.452 (d, 3J ) 7.45 Hz, 2H), 7.338 (d, 3J ) 8.93 Hz, 4H), 7.288 (m, 2H), 7.206 (m, 1H), 6.831 (d, 4H), 3.788 (s, 6H), 3.644 (t, 3J ) 6.70 Hz, 2H), 3.036 (t, 3J ) 6.70 Hz, 2H), 2.688 (t, 3J ) 7.26 Hz, 4H), 1.5-1.7 (m, 8H), 1.2-1.4 (m, 28). 13C NMR (CDCl3): δ 158.4, 145.7, 137.0, 130.2, 128.4, 127.9, 126.7, 113.1, 85.8, 63.7, 63.3, 55.4, 39.4, 33.0, 30.3, 29.8, 29.8, 29.7, 29.6, 29.5, 29.4, 28.74, 28.71, 26.5, 25.9. HRMS-EI: 708.4236; ∆ ) 1.4 ppm. 11-O-(4,4′-Dimethoxytrityl)undecanol-11′-O-((2-cyanoethyl)N,N′-diisopropylphosphoramidite)undecanol-1,1′-disulfide. Working in a inert atmosphere drybox, a round-bottomed flask equipped with a Teflon-coated magnetic stir bar was charged with 11-O-
10798 Langmuir, Vol. 22, No. 25, 2006 (4,4′-dimethoxytrityl)undecanol-11′-undecanol-1,1′-disulfide (90 mg, 0.127 mmol), diisopropylethylamine (66 mg, 0.51 mmol), and dichloromethane (1.5 mL). A solution of 2-cyanoethyl-N,N′diisopropylchlorophosphoramidite (16 mg, 0.32 mmol) in dichloromethane (0.75 mL) was added dropwise over 10 min with constant stirring. After 5 h, the apparatus was removed and placed on a Schlenk line, and the volatiles were evaporated under reduced pressure. Triethylamine in dichloromethane (1:9) was added (10 mL), and the solution was washed with several portions of a saturated solution of NaHCO3 and then dried and concentrated. Purification by column chromatography on basic alumina eluting with ethyl acetate in dichloromethane (1:9) with triethylamine (5% v/v) afforded 60 mg (52%) of the desired product. 1H NMR (CDCl3): δ 7.447 (d, 3J ) 7.45 Hz, 2H), 7.333 (d, 3J ) 8.93 Hz, 4H), 7.285 (m, 2H), 7.205 (m, 2H), 6.828 (d, 4H), 3.829 (m, 2H), 3.798 (s, 6H), 3.603 (m, 4H), 3.030 (t, 3J ) 6.70 Hz, 2H), 2.665 (m, 6H), 1.6-1.7 (m, 6H), 1.3-1.4 (bm, 28H), 1.189 (m, 12H). 13C NMR (CDCl3): δ 158.5, 145.7, 137.0, 130.2, 128.4, 127.9, 126.7, 113.1, 85.8, 64.0, 63.9, 63.7, 58.6, 58.4, 55.4, 43.2, 43.1, 39.4, 31.5, 31.4, 30.3, 29.8, 29.53, 29.46, 29.43, 28.7, 26.5, 26.1, 24.9, 24.83, 24.80, 24.7, 20.59, 20.52. 31P NMR (CDCl3): δ 164.9. FAB-MS: calcd, 909; found, 910. (SI 1) Sensor Preparation. The patterned gold electrodes were cleaned by immersion in piranha (3:1 H2SO4/H2O2) for 5 min and then thoroughly rinsed in deionized water. Caution: piranha solution reacts Violently with many organic materials and should be handled with care. C6-DNA was dissolved in 6X SSC buffer to a final oligonucleotide concentration of 0.5 µM. The piranha-cleaned electrode was immersed in this aqueous solution for ∼30 min to allow the oligonucleotides to chemisorb onto the surface. The electrode was subsequently immersed in 2 mM C6-OH in 6X SSC for ∼3 h to displace nonspecifically bound oligonucleotides. Normal DNA coverage obtained on the basis of the above immobilization scheme is ∼6 × 10-12 mol/cm2. Although many previous studies on SAMs utilize monolayers formed overnight or over a few days to ensure optimal packing density,20,10 we find that longer deposition times do not enhance the storage stability of C6-based sensors (data not shown). Whereas our C6-based E-DNA sensors are fabricated by first depositing the C6-DNA followed by passivation with C6-OH, we fabricated our C11-based E-DNA sensors by first depositing the C11-OH and then backfilling with the C11-DNA. Note that we have explored this issue extensively and have found that these two protocols lead to the most reproducible surface coverage and the most stable films for C6- and C11-based sensors, respectively (data not shown). For C11-based sensor fabrication, C11-DNA was dissolved in 6X SSC buffer to a final oligonucleotide concentration of 0.5 µM. The piranha-cleaned electrode was first immersed in a 1 mM C11-OH ethanolic solution for ∼5 min. The electrode was subsequently immersed in C11-DNA for ∼2 h to chemisorb onto the surface. With this deposition method, a DNA coverage of ∼2 × 10-13 mol/ cm2 is readily obtained. Prior to storage or interrogation with target DNA, the electrodes modified with either C6-DNA or C11-DNA were incubated in the SSC buffer or serum in which the voltammograms were collected for ∼30 min. After hybridization, regeneration of the DNA-coated electrodes was achieved by rinsing with a copious amount of deionized water. For sensors in which hybridization was conducted in serum, regeneration was achieved by 1 min of incubation in 8 M guanidine hydrochloride. Sensor regeneration was verified by an AC voltammogram recorded 5 min after reimmersion in 6X SSC buffer or serum. Electrochemical Biosensor Measurements. All electrochemical measurements were performed at room temperature (∼21 °C) by using a CHI 730B electrochemical workstation (CH Instruments, Austin, TX). The electrolyte that we employed contains undiluted fetal calf serum or 6X SSC buffer (90 mM sodium citrate, 0.9 M NaCl, pH 7) unless specified in the figure captions. Gold working electrodes with a geometric area of 0.88 mm2 were fabricated on a glass plate using standard microfabrication techniques.26 A platinum wire was used as the counter electrode. All electrochemical potentials are reported versus a Ag/AgCl (3 M KCl) reference electrode.
Lai et al. Electrodes modified with DNA were analyzed by alternating current voltammetry (ACV). Alternating current voltammograms were recorded in 1 mL of 6X SSC buffer or fetal calf serum from -0.1 to -0.45 V versus Ag/AgCl with a 10 Hz, 25 mV AC potential. Stability Measurements. The solution-phase stability of the sensor was determined as follows. After collecting the initial scan, AC voltammograms were collected every 24 h to monitor the onset of sensor degradation. The buffer or serum in which the sensors were stored was changed immediately prior to each electrochemical interrogation. To determine the long-term dry storage stability, the sensors were prepared as follows. After collecting the initial AC voltammogram, the modified electrodes were briefly rinsed with either 6X SSC buffer or the same buffer containing 2.5% (w/v) each of a sugar (trehalose or glucose) and BSA as stabilizing agents, dried under a stream of dry nitrogen, and sealed in containers filled with air. After being stored in the dark at room temperature (∼21 °C) for 5, 7, or 32 days, the electrodes were rinsed with deionized water and incubated in buffer for ∼1-3 h prior to electrochemical interrogation. A longer incubation time was needed for the previously dried sensor surface to rehydrate fully and attain the appropriate stemloop DNA structure. C6-based sensors, however, require a shorter rehyrdration time (∼1 h) when compared to the C11-based sensors (∼3 h). Hybridization to the correct target DNA (200 nM) was used in determining the performance of all previously stored sensors.
Results and Discussion Some early E-DNA and E-AB sensors employed short 2-, 3-, or 6-carbon SAMs at the biomolecule-electrode interface.11,12,21 Whereas such short-chain SAMs are useful in the initial validation of new sensor architectures (in no small part because oligonucleotides modified with 3- and 6-carbon alkanethiols are readily available via commercial synthesis), the shorter alkanethiol SAMs are unlikely to endure moderate or long-term solution-phase or dry storage conditions. 11 Thus motivated, here we have studied the suitability of mixed alkanethiol SAMs composed of C6 or C11 R,ω-thiol alcohols and the analogous thiol-modified DNA to application in the E-DNA electronic sensing platform (Figure 1). To do so, we have monitored the stability of the sensor’s methylene blue reduction current, the sensor’s ability to detect a fully complementary target, and our ability to regenerate the sensor after target recognition, all as functions of time and storage conditions. The stability of C6-based E-DNA sensors is limited. For example, even the most stable of the many C6-based sensors that we have characterized (spanning a wide range of DNA surface coverage) lost ∼40% of its initial probe DNA over the course of 7 days storage in SSC buffer under ambient conditions (Figure 2, top). Consistent with this, the mean degradation that we observed for a set of C6-based sensors was 56 ( 9% (Table 1), with most of the sensors that we have characterized exhibiting detectable degradation after as little as 12 h of solution storage (data not shown). This rather poor stability presumably arises because shorter-chain alkanethiols do not readily form pristine monolayers.27 C6-based sensors are similarly unstable when stored dry in the absence of preservatives: we observed a 42 ( 6% decrease in sensor current for C6-based sensors coated only with SSC and stored for 1 week dry in air (SI 2; Table 2). In contrast to the above performance, C6-based sensors exhibit reasonable dry storage stability in the presence of preservatives BSA and either trehelose or glucose (Table 2).8,14 Under these conditions, the C6-based sensor exhibits only 7 to 9% degradation after 32 days of storage under ambient conditions (Figure 3, (26) Lai, R. Y.; Lee, S.; Soh, H. T.; Plaxco, K. W.; Heeger, A. J. Langmuir 2006, 22, 1932-1936. (27) Bain, C. D.; Troughton, E. B.; Tao, Y.-T.; Evall, J.; Whitesides, G. M.; Nuzzo, R. G. J. Am. Chem. Soc. 1989, 111, 321-335.
Signaling, Stability of Electrochemical DNA Sensors
Langmuir, Vol. 22, No. 25, 2006 10799 Table 2. Dry Storage Stability of C6- and C11-Based E-DNA Sensors in the Presence or Absence of Stabilizing Agents
system
stabilizers
C6 C6
6X SSC BSA/trehalose/ 6X SSC BSA/glucose/ 6X SSC 6X SSC BSA/trehalose/ 6X SSC BSA/glucose/ 6X SSC
C6 C11 C11 C11
% signal suppression % signal upon days decrease % signal stored poststoragea hybridizationa,b regeneration 7 32
42 ( 6 9(2
n.d.c 42 ( 3
n.d.c 94 ( 3
32
7(2
43 ( 4
95 ( 2
5 32
15 ( 12 8(1
40 ( 7 43 ( 2
93 ( 7 97 ( 2
32
5(1
42 ( 2
98 ( 1
a The % signal decrease poststorage and % signal suppression upon hybridization were averages (and standard deviations) obtained from three different sensors. b The sensors were allowed to hybridize to 200 nM of the full complement target for ∼30 min. c Not determined. Because of advanced sensor degradation, hybridization and regeneration were not attempted.
Figure 2. Whereas the solution-phase stability of C6-based E-DNA sensors (top) is relatively poor (at room temperature under air), the stability of C11-based sensors (bottom) is markedly improved. Shown are baseline-subtracted AC voltammograms for such an E-DNA sensor before and after storage for 1 week in 6X SSC buffer. Because of the poor stability of the C6-based sensors, hybridization with the target was not attempted. Table 1. Solution Storage Stability of C6- and C11-Based E-DNA Sensors in Buffer and Undiluted Blood Serum % signal suppression % signal upon decrease % signal days system solution stored poststoragea hybridizationa,b regeneration C6 C6 C11 C11
6X SSC serum 6X SSC serum
7 7 7 7
56 ( 9 73 ( 5 7(2 10 ( 2
n.d.c n.d.c 45 ( 3 20 ( 2
n.d.c n.d.c 97 ( 2 95 ( 3
a The % signal decrease poststorage and % signal suppression upon hybridization were averages (and standard deviations) obtained from three different sensors. b The sensors were allowed to hybridize to 200 nM of the full complement target for ∼30 min. c Not determined. Because of advanced sensor degradation, hybridization and regeneration were not attempted.
top). Note that the level of degradation observed upon dry storage (with or without preservatives) appears to depend on the initial DNA surface coverage, with higher DNA coverage improving storage stability (data not shown). In comparison to C6 thiols, 11- and 12-carbon alkanethiols form relatively well-ordered films,3,4 suggesting that the use of these longer-chain alkanethiols would result in more stable SAMs and longer sensor shelf life. Electron transfer through thicker alkane SAMs, however, is reduced,5 and thus the use of C11 SAMs might reduce the sensor signal and harm sensor perfor-
Figure 3. Dry storage stability of (top) C6- and (bottom) C11based E-DNA sensors is excellent when preservatives are employed. Shown are baseline-subtracted AC voltammograms of the sensor before and after 32 days of storage in air at room temperature in the presence of preservatives BSA and trehalose. Also shown are AC voltammograms of the sensors before and after 25 min of incubation with 200 nM target DNA.
mance. To better characterize the suitability of C11 SAMs for electronic DNA sensing applications, we have measured the hybridization kinetics, signal gain, and electron-transfer kinetics of E-DNA sensors fabricated using either C6 or C11 SAMs.
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Lai et al.
current after 7 days of storage in solution under air at room temperature (Figure 2, bottom). In addition, the stored sensor remains reusable upon rinsing with deionized water, indicating that the sensor components were still strongly attached to the electrode surface (Table 1). The C11-based sensor is similarly stable even when deployed in complex, grossly contaminated media: after incubation in undiluted fetal calf serum for 7 days at room temperature, we observe only a 10 ( 2% decrease in sensor current (SI 6). The shelf life of C11-based sensors is also improved. For example, whereas C6-based sensors showed reasonable dry storage stability only in the presence of preservatives (Table 2), the C11-based sensor is relatively stable even in the absence of these agents, with some electrodes exhibiting as little as a ∼2% decrease in the sensor current after 5 days of storage in air at room temperature (SI 2). This behavior, however, is not readily reproducible, as shown by replicate measurements producing a mean degradation of 15 ( 12% after such storage (Table 2). In contrast, after more than 1 month of storage under ambient conditions in the presence of the stabilizing agents, BSA mixed with either trehalose or glucose, the sensor loses only 8 ( 1% or 5 ( 1%, respectively, of its initial signal (Figure 3, bottom). Critically, upon rehydration all of these stored sensors retain the sensitivity and reusability that are the hallmarks of the E-DNA sensor (Table 2). Finally, the level of degradation observed upon dry storage (with or without preservatives) is independent of the initial DNA surface coverage (data not shown).
Conclusions
Figure 4. Comparison of the signaling of (top) C6- and (bottom) C11-based E-DNA sensors showing that, despite the slower electrontransfer rates associated with the thicker SAM, C11-based sensors exhibit excellent signal-to-noise and signal suppression. The voltammograms representing regenerated sensors (regenerated via short washes with room-temperature distilled water) demonstrate the excellent reusability of both sensing platforms.
Using a well-established AC voltammetric method28 (the details of which are included as Supporting Information - SI3), we have reproduced earlier6,29 findings that electron transfer through the C11 SAMs is relatively slow (7 s-1; SI 4). Despite this decrease in the electron-transfer rate, however, the AC voltammetric currents obtained lead to more than adequate signal-to-noise ratios even for the small 0.88 mm2 E-DNA electrodes employed here (Figure 4). Consistent with this, the sensitivity (Figure 4), regeneration efficiency (Figure 4), and hybridization kinetics (SI 5) of the C11-based sensor are closely comparable to those of the C6-based sensor. Nonetheless, with the possibility of scaling down the electrode size for future array applications (when the reduced currents associated with C11-based sensors would begin to impede signaling), passivating the surface with molecular wires such as the well-characterized thiolated oligo(phenylenevinylene)30 instead of C11-OH might circumvent the problem of low tunneling current, and this study is currently in progress. The solution-phase stability of C11-based E-DNA sensors is significantly improved relative to that of C6-based sensors (Table 1). For example, we observe only a 7 ( 2% drop in E-DNA (28) Creager, S. E.; Wooster, T. T. Anal. Chem. 1998, 70, 4257-4263. (29) Becka A. M.; Miller, C. J. J. Phys. Chem. 1991, 96, 2657-2668. (30) Sikes, H. D.; Smalley, J. F.; Dudek, S. P.; Cook, A. R.; Newton, M. D.; Chidesy, C. E. D.; Feldberg, S. W. Science 2001, 291, 1519-1523.
For sensors to function in real-world applications, they must achieve acceptable operational and storage stability. Here we show that whereas electronic DNA sensors fabricated using mixed C6 SAMs are relatively unstable under ambient conditions, sensors based on mixed C11 SAMs exhibit promising solutionphase and dry storage stability. Critically, the signaling current, hybridization kinetics, and reusability of these more stable C11based E-DNA sensors are also promising. Mixed C11 SAMs thus appear to provide a reasonable compromise between the two main criteria for electrochemical biosensor applications: stability and electron-transfer efficiency. The formation of C11 SAMs is also relatively rapid (about 2 h) and reproducible, indicating that C11-based sensors will be compatible with the fabrication of multianalyte arrays via, for example, electrochemical lithographic techniques.26 When coupled with the ease of fabricating C11 thiol-modified oligonucleotides via the protocol described here, these attributes suggest that mixed C11 SAMs are well suited for electronic DNA sensing applications. Acknowledgment. This research was supported by the Center for Nanoscience Innovation for Defense (CNID) under DMEA9002-2-0215, by the National Institutes of Health (U54 AI06535901) and by the Institute for Collaborative Biotechnologies (ICB) (DAAD19-03-D-0004). Supporting Information Available: Synthetic route to 11-O(4,4′-dimethyloxytrityl)undecanol-1,1′-O-((2-cyanoethyl)-N,N′-diisopropylphosphoramidite)undecanol-1,1′-disulfide. Baseline-subtracted AC voltammograms of C6- and C11-based sensors before and after 5 or 7 days of storage in air at room temperature in the absence of preservatives. AC voltammograms of the sensors before and after 20 min of incubation with 200 nM target DNA. Ipeak/Ibackground versus log(frequency) plots for C6- and C11-based E-DNA sensors. Indistinguishable rates at which C6- and C11-based E-DNA sensors respond to their targets. Baselinesubtracted AC voltammograms for C6- and C11-based sensors before and after storage for 1 week in undiluted fetal calf serum. This material is available free of charge via the Internet at http://pubs.acs.org. LA0611817
