A label-free electrochemical DNA sensor based on exonuclease III-aided target recycling strategy for sequence-specific detection of femtomolar DNA

Biosensors and Bioelectronics 28 (2011) 232–238

Contents lists available at ScienceDirect

Biosensors and Bioelectronics journal homepage: www.elsevier.com/locate/bios

A label-free electrochemical DNA sensor based on exonuclease III-aided target recycling strategy for sequence-specific detection of femtomolar DNA Di Wu 1 , Bin-Cheng Yin 1 , Bang-Ce Ye ∗ Lab of Biosystems and Microanalysis, State Key Laboratory of Bioreactor Engineering, East China University of Science & Technology, Shanghai, 200237, China

a r t i c l e

i n f o

Article history: Received 3 May 2011 Received in revised form 11 July 2011 Accepted 11 July 2011 Available online 23 July 2011 Keywords: Electrochemical method Exonuclease III Signal amplification DNA detection Femtomolar

a b s t r a c t The present work demonstrates a rapid, single-step and ultrasensitive label-free and signal-off electrochemical sensor for specific DNA detection with excellent discrimination ability for single-nucleotide polymorphisms, taking advantage of Exonuclease III (Exo III)-aided target recycling strategy to achieve signal amplification. Exo III has a specifical exo-deoxyribonuclease activity for duplex DNAs in the direction from 3 to 5 terminus, however its activity on the duplex DNAs with 3 -overhang and single-strand DNA is limited. In response to the specific features of Exo III, the proposed E-DNA sensor is designed such that, in the presence of target DNA, the electrode self-assembled signaling probe hybridizes with the target DNA to form a duplex in the form of a 3 -blunt end at signaling probe and a 3 -overhang end at target DNA. In this way, Exo III specifically recognizes this structure and selectively digests the signaling probe. As a result, the target DNA dissociates from the duplex and recycles to hybridize with a new signaling probe, leading to the digestion of a large amount of signaling probes gradually. A redox mediator, Ru(NH3 )6 3+ (RuHex) is employed to electrostatically adsorbed onto signaling probes, which is directly related to the amount and the length of the signaling probes remaining in the electrode, and provides a quantitative measure of sequence-specific DNA with the experimentally measured (not extrapolated) detection limit as low as 20 fM. Moreover, this E-DNA sensor has an excellent differentiation ability for single mismatches with fairly good stability. © 2011 Elsevier B.V. All rights reserved.

1. Introduction In recent years, there have been ever-growing interests in the development of highly sensitive and selective DNA sensors in a wide range of areas including clinical diagnosis, mutation detection, and forensic analysis (Hacia et al., 1996; Jobling and Gill, 2004; Park et al., 2002). Consequently, a variety of optical (Cao et al., 2002; Dai et al., 2008; Zhang et al., 2011a,b), acoustic wave (Chiu and Gwo, 2008; Cooper et al., 2001; Höök et al., 2001; Su et al., 1994) and electrochemical (Fan et al., 2003; Hsieh et al., 2010; Hu et al., 2008; Patolsky et al., 2000; Xiao et al., 2006; Zhang et al., 2009) techniques have been employed for the development of sensitive DNA sensors. Among them, electrochemical DNA (E-DNA) sensor has attracted increasing attention owing to their remarkable features of simple, portable and inexpensive instrumentation, high sensitive, selective and fast response as well as low fabrication cost. A typical E-DNA sensors for specific DNA detection takes advantage of hybridization-induced conformational change in a redox-modified signaling probe self-assembled on a gold elec-

∗ Corresponding author. Tel.: +86 21 64252094; fax: +86 21 64252094. E-mail address: [email protected] (B.-C. Ye). 1 These authors contributed equally to this work. 0956-5663/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.bios.2011.07.029

trode by a gold–thiol bond, which is a surface-confined stem-loop (hairpin) DNA structure and analogous to fluorescent “molecular beacons” (Tyagi et al., 1998; Tyagi and Kramer, 1996). The conformational change of redox-modified signaling probe leads to the change of the distance between the redox moiety and the electrode surface, which will cause a decrease or an increase of the faradaic current, often broadly categorized as signal-off and signal-on EDNA sensor, respectively. Each type has its own advantages and limitations. Signal-off E-DNA sensor is stable, readily reusable, and sequence-specific to perform measurements in multi-component samples such as blood serum. However, it is easily susceptible to false-positives and lacks sensitivity. In contrast to signal-off E-DNA sensor, great efforts have been taken to design signal-on sensor, which often offer improved sensitivity, but exhibit greater complexity and poorer stability and reusability (Cash et al., 2009). Therefore, there is a great desire to improve the performance of signal-off E-DNA sensor in both sensitivity and stability for specific DNA analysis. Signal amplification strategy aiming at the improvement of sensitivity of E-DNA sensor still poses a great challenge for researchers. Many E-DNA sensors have been developed based on enzyme-based amplification (Kavanagh and Leech, 2006; Kim et al., 2003; Liu et al., 2008; Wan et al., 2009; Wang et al., 2002; Zhang et al., 2008) and nanomaterial-based amplification strategy (Hansen et al., 2006;

D. Wu et al. / Biosensors and Bioelectronics 28 (2011) 232–238

Zhang et al., 2006; Hu et al., 2009). Here we proposed a rapid, singlestep and ultrasensitive E-DNA sensor for specific DNA detection using the enzyme, Exonuclease III (Exo III), to achieve the aim of signal amplification. Exo III can catalyze the stepwise removal of mononucleotides from the 3 -hydroxyl ends of DNA duplexes and its activity on DNA duplexes with 3 -overhang and single-strand DNA is limited. Firstly, the fabricated E-DNA sensor is exposed in RuHex buffer to record the initial redox signal arising from electron transfer between RuHex molecules and the gold electrode surface via the phosphates backbone of DNA. In the presence of target DNA, the signaling probe hybridizes with target DNA to form a stable duplex with a 3 -blunt end at signaling probe and a 3 -overhang end at target DNA, in which the signaling probe is specifically recognized and correspondingly selectively digested by Exo III. In contrast, the DNA target is protected from Exo III and is released to hybridize with another signaling probe to initiate a new round of digest process. Thereby a large number of signaling probes are digested by Exo III and only a small quantity of RuHex is adsorbed compared to the initial signal. The experiment results demonstrate that our proposed E-DNA sensor is stable, selective, and sensitive with a very low detection limit of 20 fM. 2. Materials and methods 2.1. Materials and instruments Oligonucleotides were synthesized and purified by Sangon Inc. (Shanghai, China). The sequences of these oligomers are listed in Table S1. The signaling probe was labelled with a thiol group with a 6-carbon spacer at 5 end for immobilisation onto a gold electrode. Hexaammineruthenium(III) chloride (Ru(NH3 )6 3+ , RuHex) and tris(2-carboxyethyl)phosphine hydrochloride (TCEP) were purchased from Alfa Aesar (Ward Hill, MA, USA) and were used as received. 6-Mercaptohexanol (MCH) was purchased from Sigma–Aldrich (St. Louis, MO, USA). Escherichia coli Exonuclease III (Exo III) was purchased from Fermentas Inc. (Vilnius, Lithuania). Other chemicals used in this work, purchased from Sinopharm Chemical Reagent Co. Ltd. (Shanghai, China), were of analytical grade and were used without additional purification. All solutions were prepared with ultrapure water (Milli-Q water, 18.2 M cm) from a Millipore Milli-Q system (Bedford, MA, USA). The buffers employed in this work were as follows: DNA immobilization buffer (I-buffer) was 10 mM Tris–HCl, 0.1 M NaCl, 10 ␮M TCEP (pH 7.4). MCH incubation buffer (M-buffer) was 5 mM MCH and 10 mM Tris–HCl (pH 7.4). Exo III digestion buffer (D-buffer) was 66 mM Tris–HCl, 0.4 M NaCl, 0.66 mM MgCl2 (pH 8.0) with 50 units ExoIII. The electrochemical detection buffer (E-buffer) was 10 mM Tris–HCl, 5 ␮M RuHex (pH 7.0). The buffer (Q-buffer) for quantitation determination of probe surface density on gold electrode was 10 mM Tris–HCl (pH 7.0) with 50 ␮M RuHex. Electrochemical characterisations including cyclic voltammetry (CV), chronocoulometry (CC) and square wave voltammograms (SWV) measurements were carried out on a CHI 660D electrochemical workstation (Chenhua Instrument Company, Shanghai, China) with a conventional three-electrode cell. The three-electrode electrochemical cell consisted of a bare gold working electrode (2 mm in diameter), a platinum counter electrode, and a saturated calomel electrode (SCE) as reference electrode (saturated with 3.0 M KCl). All electrochemical measurements were carried out at laboratory ambient room temperature (RT, 22–25 ◦ C). 2.2. E-DNA sensor fabrication The E-DNA sensors were fabricated following the reported protocols (Xiao et al., 2007; Zhang et al., 2007). Briefly, bare gold

233

electrodes were polished with two micropolish deagglomerated alumina suspensions (0.3 and 0.05 ␮m diameter) in sequence for 5 min each, followed by ultrasonic cleaning with absolute ethanol and Milli-Q water for 5 min each to remove the residual alumina powder. Then the electrodes were electrochemically cleaned to remove the remaining impurities by performing consecutive CV measurements in a freshly prepared 0.5 M H2 SO4 solution with following parameters: potential range from −0.3 V to 1.5 V at a scan rate of 0.1 V/s versus SCE. A cyclic voltammogram characteristic of a gold electrode was clearly observed with typical single sharp reduction peak located at ∼0.88 V and multiple overlapping oxidation peaks in the range of 1.1–1.5 V (Fig. S1). After that, the electrodes were washed thoroughly with Milli-Q water and dried in a nitrogen stream to obtain a clean gold surface. Then the electrodes were ready for DNA immobilization. Prior to immobilization onto the gold electrode, a thiolated oligonucleotide (P1) was dissolved in I-buffer and incubated for 30 min at RT to reduce disulfide bonds. Then, an aliquot of P1 solution was dispensed on the freshly cleaned electrode for 3 h at RT. After that the gold electrode was washed thoroughly with MilliQ water and dried in a nitrogen stream. Then the gold electrode was incubated in M-buffer for 2 h at RT to prevent nonspecific DNA adsorption on the gold surface in the subsequent experiments. Finally, the electrode was rinsed thoroughly with Milli-Q water and dried with a stream of nitrogen gas again, which was ready for the electrochemical experiments. For the control experiments, the MCH-modified gold electrode was fabricated by following the procedures above, but without immobilising the thiolated oligonucleotides.

2.3. Surface density of E-DNA sensors The surface density (the number of immobilized electroactive ssDNA moles per unit area of the electrode surface) was chronocoulometrically quantified from the redox charges of RuHex (Steel et al., 1998). It is based on the assumption that redox active RuHex cation stoichiometrically associates with anionic phosphate backbone of DNA and the saturated amount of RuHex in the DNA monolayer is directly proportional to the number of phosphate residues. Therefore, we can estimate the quantity of single- or double-stranded DNA on an electrode by measuring the charge passed during the reduction of RuHex. The charge corresponding to RuHex electrostatically bound to surface-confined ssDNA (Qss ) can be calculated from the following equation: Qss = Qtotal − Qdl , where Qtotal represents the total charge flowing through the electrode, comprising both faradaic (redox) charges and non-faradaic (capacitive) charges (Qdl ). The amount of electroactive oligonucleotide on the electrode surface ( ) is calculated based on the equation

 ss =

Qss NA/nFA z/m



where n is the number of electrons transferred in the reaction (RuHex3+ + e− → RuHex2+ , n = 1), F represents the Faraday constant (coulombs per equivalent), A is the effective surface area of gold electrode (square centimeters), m is the number of nucleotides in the DNA, z is the charge of the redox molecules and NA is Avogadro’s number. Chronocoulometry measurement was carried out with a pulse width of 0.25 s.

234

D. Wu et al. / Biosensors and Bioelectronics 28 (2011) 232–238

Scheme 1. Schematic illustration of the label-free signal-off electrochemical DNA sensor via Exo III-aided target DNA recycling for sequence-specific detection of DNA. Inset: The mechanism of Exo III to different DNA structures (perfect-matched DNA duplex, DNA duplex with a 3 -overhang and single-strand DNA).

2.4. Electrochemical DNA detection Prior to DNA detection, the fabricated P1-modified electrodes were first immersed to equilibrate in a single-compartment cell filled with 8-mL E-buffer for 5 min and then tested to gain the initial RuHex redox current using CV and SWV measurement. CV measurement was performed at a scan rate of 0.1 V/s between −0.4 V and −0.1 V. SWV measurement was performed with following parameters: initial potential of −0.1 V, final potential of −0.6 V, frequency of 25 Hz. After the initial measurement, the electrodes were washed thoroughly with ethanol and Milli-Q water to remove binding RuHex molecule, and dried in a nitrogen stream. A series of 10 ␮L DNA target solution dissolved in D-buffer with known concentrations were dispensed on different electrodes with similar probe surface density for 2 h at RT. After incubation, the electrodes were washed thoroughly with ethanol and Milli-Q water, and dried in a nitrogen stream. Then the electrodes were immersed in the single-compartment cell with 8-mL E-buffer again to allow equilibrate for 5 min prior to electrochemical measurements to obtain the final redox currents. Unless otherwise noted, each electrochemical measurement was repeated using at least three electrodes with similar probe density under the same conditions.

the perfectly matched target DNA to this system (Scheme 1C), the target DNA and signaling probe form a double-stranded structure (Scheme 1D). This duplex has unique characteristic of 3 -blunt end at the signaling probe and 3 -overhang end at the target DNA. It is reported that commercial Exo III product can specifically catalyze the stepwise removal of mononucleotides of duplex starting from a 3 -OH at nicks, blunt or recessed ends to yield nucleoside 5 phosphates. More importantly, Exo III is not active on 3 -overhang ends of duplex with at least four bases long and single-strand DNA. Based on the its catalytic property, Exo III specifically recognizes the duplex formed by target DNA and signaling probe, and selectively digests the signaling probe (Scheme 1E and F). In contrast, the DNA target is not digested due to 3 -overhang ends and subsequently dissociated from the duplex. Then it hybridizes with a new signaling probe to induce the next probe-digestion cycle, resulting in the digestion of a large amount of signaling probes and a significant decrease in the amount of the RuHex electrostatically adsorbed onto signaling probe (Scheme 1G). Thus via the targetrecycling event, the proposed E-DNA sensor provides an amplified electrochemical signal, which is proportional to the concentration of target DNA. 3.2. Effect of probe surface density on electrochemical signaling

3. Results and discussion 3.1. Design strategy of Exo III-aided E-DNA sensor In this work, a label-free E-DNA sensor via Exo III-aided analyte recycling strategy for sequence-specific detection of DNA is proposed. As depicted in Scheme 1, a sensor consisting of a signaling probe with a thiol group at 5 end to self-assemble onto the gold electrode (Scheme 1A) is exposed in E-buffer to entrap RuHex molecules (Scheme 1B), in which RuHex/DNA/electrode system gives a significant redox signal arising from electron transfer between RuHex molecules and the gold electrode surface via the phosphates backbone of DNA. Upon addition of the Exo III and

It is well known that the probe surface density of electrode is a crucial parameter in the design of a sensitive and selective E-DNA sensor (Markham and Zuker, 2005; Ricci et al., 2007; Lubin et al., 2009). According to the previous study (Steel et al., 1998), the probe surface density can be quantitatively determined by CC measurement, based on the electrostatic binding of the signaling transducer RuHex cation to the anionic phosphate backbones of DNA. Prior to the specific target DNA detection, we first investigated the effect of probe surface density on E-DNA signaling. The probe surface density was modulated by varying the concentration of capture probes P1 and quantified in Q-buffer by CC measurement. A series of chronocoulometric curves for RuHex at MCH and

D. Wu et al. / Biosensors and Bioelectronics 28 (2011) 232–238

235

experimental data demonstrate that the reproducible maximum hybridization signaling is obtained at the probe density of ∼6.0 × 1012 molecules/cm2 with 1 ␮M P1 employed to prepare the electrode, which presumably represents a balance between steric and electrostatic blocking. In contrast, the high probe density (1.22 × 1013 –1.50 × 1013 molecules/cm2 ) leads to low signaling that can be explained by the presence of steric crowding. Notably, the electrodes employed in the following experiments were fabricated using 1 ␮M P1 to ensure improved hybridization efficiency. 3.3. The feasibility of the proposed E-DNA sensor

Fig. 1. Chronocoulometric response curves for gold electrodes assembled with MCH (dot line) and P1with increasing concentrations (from bottom to top: 0.01, 0.05, 0.10, 0.5, 1.0, 2.0, 3.0, 5.0, 10.0 ␮M) in the Q-buffer, respectively. Inset: The dashed lines are the fits to the data used to determine the intercept at t = 0, for the chronocoulometric curves recorded by MCH-modified and the P1-modified gold electrodes, respectively. (B) Probe surface density of electrode as a function of P1 concentration employed during sensor fabrication process. (C) SWV current change (I) as a function of the probe surface density. Values represent the average and standard deviation of measurements conducted with three independent sensors at each surface density.

P1/MCH electrodes at different P1 concentrations are presented in Fig. 1A. According to equations as described in Section 2, probe surface density as a function of P1 concentration employed during sensor fabrication was calculated. As shown in Fig. 1B, the probe density increases monotonically from low-density surfaces (1.95 × 1011 –2.06 × 1012 molecules/cm2 ) to mediumdensity surfaces (3.78 × 1012 –8.98 × 1012 molecules/cm2 ) with increasing probe concentrations until the signal gain reaches a saturated value with high-density surfaces (1.22 × 1013 –1.50 × 1013 molecules/cm2 ). Intuitively, it is thought that an ever-increasing probe surface density would give an increasing signaling during hybridization process. However, this is not the case. Surface hybridization of signaling probes interacting with solution-phase targets strongly depends on the target sequence and probe density (Peterson et al., 2002). The hybridization efficiency of T1 and P1 is studied by measuring the electrochemical signal at different surface densities. As shown in Fig. 1C, the signal increases monotonically from low-density surface to medium-density surface, reaches a maximum value and then decreases monotonically at high-density surface. The

To explore the feasibility of the proposed Exo III-aided E-DNA sensor in detecting sequence-specific DNA, we studied the CV response of RuHex for P1-modified electrodes and MCH-modified electrodes. As shown in Fig. 2A, two pairs of well-defined peaks measured by CV for P1-modified (red line) and MCH-modified (black line) electrode were recorded. P1-modified electrode exhibits a pair of pronounced defined redox peaks, suggesting the successful electron communication between RuHex cation and the electrode. Similar to that obtained at P1-modified electrode, the redox peaks from MCH-modified electrode has two pairs of peaks with very small peak currents. This is because little RuHex molecules were adsorbed onto the short alkanethiol. In the presence of 10 nM T1 in D-buffer in this system, we observed substantially diminished redox peaks (green line). This observation clearly demonstrates the greater decrease of electron transfer between the RuHex cation and gold electrode, due to the digestion of T1; thereby reduce the adsorption amount of RuHex. The significant difference in the redox potentials of the two pair of peaks in the absence and presence of T1 offers the possibility to detect the sequence-specific DNA, providing a stronger evidence of the feasibility of this design. It is well-known that SWV measurement offers excellent resolution of the response with much higher sensitivity compared to the conventional sweep techniques such as CV. Therefore, we employed SWV measurement to further characterize the electron transfer (Fig. 2B). It is observed that the proposed E-DNA sensor gives a remarkable peak at −0.328 V versus SWV in the absence of T1, and the peak current decreased from 0.829 ␮A to 0.149 ␮A in response to 10 nM T1. In addition, there is a very small peak potential shift in the MCH-modified electrode (Fig. 2B inset). It is ascribed to the minor differences in electron transfer media of phosphates backbone of DNA and small molecule of MCH. The above results give immediate evidence for the successful degradation of signaling probe P1 by Exo III in the presence of target DNA T1. We further investigated the catalytic activity of Exo III on the single-strand DNA by incubating P1-modified electrode in the Dbuffer without adding T1. As expected, no significant change in the redox signal was observed compared with the initial signal. This result confirms that Exo III has little catalytic activity on the singlestrand DNA, which is one of the crucial preconditions for the Exo III-aided signal amplification strategy in this work. 3.4. Effect of the length of 3 -overhang ends on Exo III activity Exo III is an exonuclease specific for double-stranded DNA from 3 -phosphate ends; however, it hardly catalyzes the removal of mononucleotides of 3 -overhangs with at least 4 bases. In order to investigate the effect of the length of 3 -overhang end in the duplex structure on Exo III activity, seven DNA targets including perfectly matched (T1), 1-base (T2), 2-base (T3), 3-base (T4), 4-base (T5), 5base (T6), 6-base (T7) recessed DNAs (see Table S1 for details) were prepared to react with P1-modified electrodes in D-buffer, respectively. All tested target DNAs contained the same 5-base overhang sequences at 3 ends and different length sequences matched P1

236

D. Wu et al. / Biosensors and Bioelectronics 28 (2011) 232–238

Fig. 2. (A) Cyclic voltammograms of P1-modified electrodes before (solid line) and after reaction with 10 nM T1 (dash line), and MCH-modified electrode (dot line) in the D-buffer. (B) SWV curves for P1-modified electrodes before (solid line) and after reaction with 10 nM T1 (dash line), and MCH-modified electrode (dot line) in the D-buffer.

at 5 ends. Therefore, P1 has different length overhang from 0-base to 6-base at 3 ends to the different target DNAs correspondingly. Fig. 3A illustrates that SWV peak current decreased monotonously with the presence of the target DNAs from T5 to T1. The SWV peaks of T6 and T7 are close to that of T5. They are not included so that a clearer display of the SWV results is shown. More direct contrast for all the test targets is shown in Fig. 3B. Based on the above observations, the optimal 3 -overhang length in the duplex structure appears to be of at least 5 bases, such that degradation of electrode-bound P1 probe from Exo III can be avoided. These results are consistent with the information of commercial product Exo III, in which 3 -overhangs of ≥4 bases are protected from Exo III digestion. Thus, for better performance of protection, we chose a 5-base overhang at the 3 -end of a DNA target to ascertain a successful target recycling process.

3.5. Selectivity of the proposed E-DNA sensor Different kinds of target DNAs including the perfect complementary target T1 and mismatch T8 (1-base mismatch), T9 (2-base mismatch), T10 (3-base mismatch), T11 (4-base mismatch), T12 (5-base mismatch), T13 (6-base mismatch) were chosen to investigate the selectivity of the sensor (see Table S1 for details). Different redox response at P1-modified electrodes and at P1-modified electrodes hybridized with various target DNAs, and corresponding comparison of these SWV peak currents are shown in Fig. 3C and D. As the SWV peaks for T12 and T13 are close to that for T11, they were not included in Fig. 3C for clarity. The peak current for T1 is 0.183 ␮A. It is the lowest signal among other mismatch target DNAs whose peak currents were 0.382 ␮A (T8), 0.557 ␮A (T9), and 0.701 ␮A (T10), and 0.786 ␮A (T11) respectively. This could

Fig. 3. (A) SWV curves recorded in absence of target DNA and in the presence of T1, T2, T3, T4 and T5, respectively. (B) Comparison for the signal reduction for sensors in absence of target DNA and in the presence of T1, T2, T3, T4, T5, T6 and T7, respectively. The error bars represent the standard deviation of three independent measurements at each target DNA. (C) SWV curves recorded in absence of target DNA and in the presence of T1, T8, T9, T10 and T11, respectively. (D) Comparison for the signal reduction in absence of target DNA and in the presence of T1, T8, T9, T10, T11, T12 and T13 respectively. Error bars represent the standard deviation of three independent measurements at each target DNA. All tested target DNAs’ concentration was 2 ␮M.

D. Wu et al. / Biosensors and Bioelectronics 28 (2011) 232–238

237

Fig. 4. SWV curves corresponding to the detection of different concentrations of T1: (a) 0, (b) 20 fM, (c) 50 fM, (d) 100 fM, (e) 500 fM, (f) 1 pM, (g) 10 pM, (h) 20 pM, (i) 50 pM, (g) 100 pM, (k) 1 nM, and (l) 10 nM. (B) Linear fit plot of peak current as function of the logarithm of T1 concentrations (20 fM, 50 fM, 100 fM, 500 M, 1 pM, 10 pM, 20 pM, and 50 pM). Error bars represent the standard deviations of three independent measurements with three E-DNA sensors.

be ascribed to that Exo III has the strongest exonuclease activity on 3 -blunt structure formed by T1 and P1. The significant current difference between perfect-matched and mismatched target DNAs demonstrates that the proposed sensor readily discriminates between single nucleotide polymorphisms. 3.6. Sensitive detection of target DNA The sensitivity of the sensor was investigated by challenging it with T1 in a series of concentrations. Fig. 4A reveals SWV peak current is strongly dependent on the concentration of T1. As the T1 concentration increases from 20 fM to 1 (M (curves b–m), SWV peak monotonously decreases. It is clearly demonstrated that the introduction of T1 at different concentrations to the sensing interface results in different degrees of digestion of P1 anchored on the electrode, leading to significantly decreased peak currents associated with the amounts of RuHex. The SWV peak current is found to be a linear logarithmic function related to T1 concentration (Fig. 4B). T1 was quantified over a concentration range of 20 fM to 50 pM. The linear function is I = 0.5677 − 0.1287 lg cT1 . The experimentally measured detection limit is as low as 20 fM (>3SD/m, SD and m represent standard deviation and slope rate, respectively), indicating a high sensitivity. Hsieh et al. (2010) reported electrochemical DNA detection via ExoIII and target-catalyzed strategy, which employed conformational change of redox-modified signaling probe and obtained a detection limit of ∼2 nM. In contrast, our proposed E-DNA sensor is 100-fold lower. To the best of our knowledge, our sensor is superior in sensitivity compared with other reported electrochemical assays as listed in Table S2. Of note, we find that the signal for the sensor is reproducible, with an electrodeto-electrode variation of <5% (data from three independent tests with different electrodes).

4. Conclusion We have described a simple signal-off E-DNA sensor that employs an Exo III-aided target recycling strategy for sequencespecific DNA detection with improved sensitivity, selectivity and stability. Compared to other E-DNA sensors based on enzyme/nanoparticle-based amplification, our design has several significant advantages toward sensitivity and selectivity improvement. First, our sensor requires no external modification on the signaling probe. A redox mediator, RuHex, is selected as an electrical signal readout, which is easy to prepare, convenient to operate with a good stability, and is also cost-effective. Second, this assay is more straightforward and sensitive as a signal-off type sensor. Usually the signal-off sensor is simple, selective and stable but at a cost of insufficient sensitivity and falsepositive signals. As to signal-on sensor, it has the potential of improved sensitivity but it often required more complex design with poorer stability and reusability. In contrast, our sensor is proven to be a sensitive signal-off electrochemical DNA detection platform with remarkable stability and specificity. The crucial design of our sensor is that it employs an Exo III-mediated target recycling mechanism to achieve amplified electrochemical signals. This strategy is single-step and single-reagent that can be performed at room temperature. By employing the amplification strategy, the sensitivity of the sensor has been greatly improved to the detection limit of 20 fM, which is significantly lower than those achieved using signal-off and signal-on sensors. Third, given that Exo III does not require a specific recognition site, it is fairly easy to apply this strategy to detect different DNA sequences. These features will give rise to a promising alternative to the cumbersome femtomolar electrochemical assays reported to date.

3.7. Sensor response in blood serum Acknowledgments Besides sensitivity and selectivity, real-life application is also an important feature for E-DNA sensor. In order to assess the response of our sensor in a clinical sample, E-DNA sensors were incubated with 1 ␮M T1 in D-buffer and a complex sample matrix of 50% serum with 50 units Exo III, respectively. It is found that a higher signal in the 50% serum, compared to the peak current observed in the D-buffer (Fig. S2). It is attributed to the influence of serum on the catalytic activity of Exo III. In spite of this, the presence of biological fluids of 50% serum do not sacrifice the performance of our sensor and this sensing platform still performs well to allow being well-suited for clinical applications.

This work was financially supported by NSF (21075040), the Shanghai Project (09JC1404100, 11XD1401900), the SKLBE (2060204), and the Fundamental Research Funds for the Central Universities.

Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.bios.2011.07.029.

238

D. Wu et al. / Biosensors and Bioelectronics 28 (2011) 232–238

References Cao, Y.C., Jin, R., Mirkin, C.A., 2002. Science 297, 1536–1540. Cash, K.J., Heeger, A.J., Plaxco, K.W., Xiao, Y., 2009. Anal. Chem. 81, 656–661. Chiu, C.S., Gwo, S., 2008. Anal. Chem. 80, 3318–3326. Cooper, M.A., Dultsev, F.N., Minson, T., Ostanin, V.P., Abell, C., Klenerman, D., 2001. Nat. Biotechnol. 19, 833–837. Dai, Q., Liu, X., Coutts, J., Austin, L., Huo, Q., 2008. J. Am. Chem. Soc. 130, 8138–8139. Fan, C., Plaxco, K.W., Heeger, A.J., 2003. Proc. Natl. Acad. Sci. U. S. A. 100, 9134–9137. Hacia, J.G., Brody, L.C., Chee, M.S., Fodor, S.P., Collins, F.S., 1996. Nat. Genet. 14, 441–447. Hansen, J.A., Mukhopadhyay, R., Hansen, J.Ø., Gothelf, K.V., 2006. J. Am. Chem. Soc. 128, 3860–3861. Höök, F., Ray, A., Nordén, B., Kasemo, B., 2001. Langmuir 17, 8305–8312. Hsieh, K., Xiao, Y., Tom Soh, H., 2010. Langmuir 26, 10392–10396. Hu, K., Lan, D., Li, X., Zhang, S., 2008. Anal. Chem. 80, 9124–9130. Hu, K., Liu, P., Ye, S., Zhang, S., 2009. Biosens. Bioelectron. 24, 3113–3119. Jobling, M.A., Gill, P., 2004. Nat. Rev. Gene 5, 739–751. Kavanagh, P., Leech, D., 2006. Anal. Chem. 78, 2710–2716. Kim, E., Kim, K., Yang, H., Kim, Y.T., Kwak, J., 2003. Anal. Chem. 75, 5665–5672. Liu, G., Wan, Y., Gau, V., Zhang, J., Wang, L., Song, S., Fan, C., 2008. J. Am. Chem. Soc. 130, 6820–6825. Lubin, A.A., Vander Stoep Hunt, B., White, R.J., Plaxco, K.W., 2009. Anal. Chem. 81, 2150–2158. Markham, N.R., Zuker, M., 2005. Nucleic Acids Res. 33, W577–W581.

Patolsky, F., Lichtenstein, A., Willner, I.I., 2000. Angew. Chem. Int. Ed. Engl. 39, 940–943. Park, S.J., Taton, T.A., Mirkin, C.A., 2002. Science 295, 1503–1506. Peterson, A.W., Wolf, L.K., Georgiadis, R.M., 2002. J. Am. Chem. Soc. 124, 14601–14607. Ricci, F., Lai, R.Y., Heeger, A.J., Plaxco, K.W., Sumner, J.J., 2007. Langmuir 23, 6827–6834. Steel, A.B., Herne, T.M., Tarlov, M.J., 1998. Anal. Chem. 70, 4670–4677. Su, H., Kallury, K.M.R., Thompson, M., Roach, A., 1994. Anal. Chem. 66, 769–777. Tyagi, S., Kramer, F.R., 1996. Nat. Biotechnol. 14, 303–308. Tyagi, S., Bratu, D.P., Kramer, F.R., 1998. Nat. Biotechnol. 16, 49–53. Wan, Y., Zhang, J., Liu, G., Pan, D., Wang, L., Song, S., Fan, C., 2009. Biosens. Bioelectron. 24, 1209–1212. Wang, J., Kawde, A.N., Musameh, M., Rivas, G., 2002. Analyst 127, 1279–1282. Xiao, Y., Lai, R.Y., Plaxco, K.W., 2007. Nat Protoc. 2, 2875–2880. Xiao, Y., Lubin, A.A., Baker, B.R., Plaxco, K.W., Heeger, A.J., 2006. Proc. Natl. Acad. Sci. U. S. A. 103, 16677–16680. Zhang, J., Lao, R., Song, S., Yan, Z., Fan, C., 2008. Anal. Chem. 80, 9029–9033. Zhang, J., Song, S., Wang, L., Pan, D., Fan, C., 2007. Nat. Protoc. 2, 2888–2895. Zhang, J., Song, S., Zhang, L., Wang, L., Wu, H., Pan, D., Fan, C., 2006. J. Am. Chem. Soc. 128, 8575–8580. Zhang, M., Yin, B.C., Tan, W.H., Ye, B.C., 2011a. Biosens. Bioelectron. 26, 3260–3265. Zhang, M., Guan, Y.M., Ye, B.C., 2011b. Chem. Commun. 47, 3478–3480. Zhang, Y., Wang, Y., Wang, H., Jiang, J.H., Shen, G.L., Yu, R.Q., Li, J., 2009. Anal. Chem. 81, 1982–1987.