Electrochemiluminescent lead biosensor based on GR-5 lead-dependent DNAzyme for Ru(phen)₃²⁺ intercalation and lead recognition

Abstract

An electrochemiluminescent (ECL) lead biosensor was developed based on GR-5 lead-dependent DNAzyme for lead recognition and intercalated ruthenium tris(1,10-phenanthroline) (Ru(phen)₃²⁺) as the ECL probe. The thiol-modified substrate was first immobilized on the surface of the gold electrode via gold–sulfur self-assembly. Subsequently, the hybridization of DNAzyme and its substrate and the automatic intercalation of Ru(phen)₃²⁺ proceeded. Intercalated Ru(phen)₃²⁺ can transfer electrons through double-stranded DNA to the electrode and its electrochemiluminescence was excited with a potential step using tripropylamine as the coreactant. In the presence of lead, the substrate cleaves at the scissile ribo-adenine into two fragments. The dissociation of DNAzyme occurs, leading to the releasing of intercalated Ru(phen)₃²⁺ accompanied by a decrease in the intensity of electrochemiluminescence. A quantity of lead can be calculated from this decrease. The biosensor is highly sensitive and specific, along with an ultra-low limit of detection of 0.9 pM and a dynamic range from 2 to 1000 pM. It enables analysis of trace amounts of lead in serum samples. The combination of the intercalated-Ru(phen)₃²⁺ ECL probe and the cofactor-dependent DNAzyme may push the performance of cofactor-sensing tactics to the extreme.

---

Introduction

Lead is a common composite of ceramics, oil paintings, and electronics.¹ It is a toxic metal, causing environmental pollution and severe risks to humans. Lead can enter the bloodstream, and blood lead level is an important index to show lead exposure. People show symptoms like kidney damage, hypertension, anemia, and nervous system dysfunction when exposed to high levels of lead.¹ Available protocols for the determination of lead include atomic absorption spectrometry,² inductively coupled plasma mass spectrometry,³ and anodic stripping voltammetry (ASV).⁴ However, they are expensive and complex, only suitable for laboratory analysis, or interfered with by surface adsorption. The exploration of cheap, uncomplicated and selective lead-determining methods are urgently needed.

DNAzymes, catalytic DNA, are regulated by various cofactors. For example, the cofactor hemin binds specific G-quadruplexes to form a horseradish peroxidase-mimicking conjugate.^5–7 Lead-dependent DNAzymes cleave specific, partially complementary DNA substrate in the presence of lead ions.^8,9 Using lead-dependent DNAzyme, several lead-sensing approaches have been developed. The first fluorescent lead biosensor using lead-dependent DNAzyme was contributed by Lu's group in 2000.¹⁰ Later, they developed a colorimetric lead biosensor based on lead-dependent DNAzyme and color changes upon aggregation and disaggregation of gold nanoparticles.¹¹ Great efforts have been made to improve the performance of fluorescent and colorimetric biosensors since.^12–17 Yet, the inherent drawbacks derive from the cumbersome equipment utilized and the potential false signals produced by fluorophores or colorants and their high background signals.

Recently, different types of transduction signals, such as chemiluminescence,^18,19 cytometry,²⁰ dynamic light scattering,^21,22 electrochemistry,^23–29 ECL,^30,31 surface plasmon resonance,²⁴ and surface-enhanced Raman spectroscopy,³² coupled with lead-dependent DNAzymes have been applied to build lead biosensors. Among them, novel electrochemical platforms are more popular, due to their low cost, simplicity, convenient operation, eminent specificity, and less electro-active contaminants, compared to the fluorescent and colorimetric approaches. In 2006, Plaxco's team announced the first electrochemical lead-dependent DNAzyme biosensor with its core component of a 3′-end-methylene-blue-tagged and 5′-end-thiol-group-modified DNAzyme.²⁹ Several signal-amplified electrochemical methods have been reported implementing gold–DNA nano-clusters,^24,25,28 quantum dots,²⁶ and hemin–G-quadruplexes.²⁴

ECL combines the merits of electrochemistry and chemiluminescence. The system of ruthenium complex with tripropylamine (TPA) as coreactant is of particular interest because it enables ultra-low limits of detection. Chemically labeled lead-dependent DNAzymes or their substrates with derivatives of ruthenium tris(2,2′-bipyridine) (Ru(bpy)₃²⁺) have been used to design lead biosensors, which achieved limits of detection of 1.4 pM and 11 pM, respectively.^30,31 However, labeling and purification of Ru(bpy)₃²⁺ derivative-labeled DNA is time-consuming and suffers from contamination and low-yields. While ruthenium tris(1,10-phenanthroline) (Ru(phen)₃²⁺) is highly ECL-active, it can intercalate into double-stranded DNA, which provides an alternative strategy for the building of ECL biosensors.^33–44 ECL biosensors based on intercalation avoid the shortcomings of chemical labeling. They also benefit from amplified detection caused by more than one Ru(phen)₃²⁺ signal molecule intercalated into each double-stranded DNA.

Some lead biosensors are established on the basis that lead induces the formation of G-quadruplexes.^23,27,45,46 Similar to aptamers,⁴⁷ guanine-rich DNA is in an equilibrium conformational state between the unfolded and folded G-quadruplex, and the equilibrium is merely driven to form the G-quadruplex upon the addition of lead, so false signals are a possibility. The ECL strategy using intercalated Ru(phen)₃²⁺ may also overcome the false positive signal existing in G-quadruplex-based biosensors because lead-dependent DNAzymes are less likely to encounter the problem of equilibrium. In this paper, we introduce an ECL lead biosensor based on GR-5 DNAzyme for Ru(phen)₃²⁺ intercalation and lead recognition with an ultra-low limit of detection of 0.9 pM.

Experimental section

Chemicals and materials

GR-5 DNAzyme and its substrate were synthesized and purified by Takara Biotechnology (Dalian, China). The sequences of this DNA are given below.

GR-5 DNAzyme: 5′-ACAGA CATCA TCTCT GAAGT AGCGC CGCCG TATAG TGAG-3′

Substrate: 5′-HS–(CH₂)₆–CTCAC TATrAG GAAGA GATGA TGTCT GT-3′

Tris(2-carboxyethyl)phosphine hydrochloride (TCEP) was purchased from Acros Organics, and 6-mercapto-1-hexanol (MCH) from J&K Scientific. Dichlorotris(1,10-phenanthroline)ruthenium(II) hydrate [Ru(phen)₃Cl₂·H₂O], lead(II) acetate trihydrate [Pb(OAc)₂·3H₂O], and TPA were purchased from Sigma-Aldrich. All other chemicals were of analytical grade. Gold disk electrodes of 2 mm diameter were employed. Deionized water of about 18.25 MΩ cm was used throughout the experiment. The standard lead solution was prepared by dissolving Pb(OAc)₂·3H₂O into a buffer of 50 mM Tris–acetate (pH 8.2).

Preparation of the ECL lead biosensor^29,34,48

The gold electrode was mechanically polished and electrochemically cleaned. TCEP in water was added to the diluted substrate in 10 mM Tris–HCl (pH 7.4) containing 1 mM EDTA to a final concentration of 0.2 μM substrate and 10 μM TCEP, and stored at 4 °C for 1 h while reduction took place to break di-sulfur bond formed between two substrate strands. TCEP was newly prepared to avoid its oxidation by air. 4.5 μL of 0.2 μM substrate solution was cast on the electrode surface for 1 h to immobilize the substrate. Next, 4.5 μL of 1 mM fleshly prepared MCH in water was dropped for 1 h at 4 °C to block the active sites on the electrode surface. By casting a 4.5 μL mixture of 0.4 μM DNAzyme and 2.4 mM Ru(phen)₃²⁺ in 50 mM Tris–acetate (pH 8.2) containing 500 mM NaCl at 4 °C for 16 h, the full hybridization of DNAzyme and its substrate and the intercalation of Ru(phen)₃²⁺ automatically proceeded on the electrode surface. To decrease the interference, deionized water and 50 mM Tris–acetate (pH 8.2) containing 500 mM NaCl were used to remove non-chemical adsorbed species.

ECL measurement of lead

For lead determination, 4.5 μL lead solutions of various concentrations were incubated on the ECL lead biosensor at 37 °C for 1 h for maximum cleavage. The working electrode was transferred to 0.1 M PBS (pH 7.4) containing 2.5 mM TPA with saturated Ag/AgCl as the reference and platinum wire as the counter electrodes. Cyclic voltammetry and potential step voltammetry were performed with a LK98BII microcomputer-based electrochemical analyzer (Lanlike Chemical and Electron High Technology Co., Ltd., Tianjin, China) and ECL was recorded by an MPI-A ECL analyzer (Remax Analyze Instrument Co., Ltd., Xi'An, China) biased at −900 V. The standard protocol of ASV using a LK98BI microelement analyzer (Lanbiao Electron and Technology Development Co., Ltd, Tianjin, China) was used to validate the practicability of the proposed ECL method.

Results and discussion

Design of the ECL lead biosensor

The lead DNAzymes, such as 8–17 (ref. 9) and GR-5 DNAzyme,⁸ are capable of catalyzing a phosphodiester bond cleavage reaction in the presence of lead. GR-5 DNAzyme has high selectivity for lead,⁸ but 8–17 DNAzyme is still active for other metal ions, such as zinc⁴⁹ and magnesium,⁵⁰ as they were selected under different conditions. GR-5 was confirmed to have higher activity for lead than 8–17 DNAzyme¹⁶ and was therefore selected for the development of the present ECL biosensor.

For the building of the lead biosensor, the complementary strand of GR-5 DNAzyme was first immobilized on the gold electrode via the thiol group labeled at the 5′ end (Scheme 1). The hybridization between DNAzyme and its complementary strand made sure that the DNAzyme–substrate complex maintained a double helical structure for the intercalation of Ru(phen)₃²⁺. Lead cleaved the phosphodiester bond of the RNA base. The cleavage resulted in the destabilization of base pairing and the dissociation of double strands in the DNAzyme–substrate complex. Correspondingly, the intercalated Ru(phen)₃²⁺ was released. The ECL signal from the intercalated Ru(phen)₃²⁺ using TPA as a co-reactant was recorded. The difference in ECL intensity before and after the addition of lead was used to quantify lead ions.

Scheme 1 Schematic representation of the ECL lead biosensor based on Ru(phen)₃²⁺ intercalation and GR-5 DNAzyme. Left: the hybridization of DNAzyme and the substrate with intercalated Ru(phen)₃²⁺ for high ECL emission. Right: the cleaved substrate along with released Ru(phen)₃²⁺ after the introduction of lead with the decreased ECL. DNA is drawn in rigid form for better illustration.

Of note is the fact that two substrates may be combined together through the formation of a di-sulfur bond by the thiol group prior to immobilization. TCEP was added into the buffer to cleave di-sulfur bonds, which benefits the self-assembly of the substrate on the gold electrode surface. To decrease the unspecific adsorption at the active site on electrode surface and to improve hybridization subsequently, the electrode modified with the substrate was immersed in an MCH solution to block the active sites. The self-assembled MCH can also maintain the perpendicular orientation of the substrate.

DNAzyme and its substrate are partially complimentary, and there is a big bump formed after hybridization. Enough area is expected to be occupied for the duplex of DNAzyme and its substrate to stand on the surface of the electrode. A relatively low density of immobilized substrate is therefore pursued. A 4.5 μL of 0.2 μM substrate-containing drop was cast on the electrode surface for 1 h to form the substrate monolayer. Since one Ru(phen)₃²⁺ molecule can be embedded in every four base pairs,^33,51 five Ru(phen)₃²⁺ molecules in total are expected in each duplex in this case. While the chemical label is not needed, the intercalation strategy can increase the number of Ru(phen)₃²⁺ for each duplex over the single site labeling for improved sensitivity.^30,31

Characterization of the ECL lead biosensor

The self-assembly of the substrate and the hybridization of DNAzyme and its substrate played an important role in the construction in the present biosensor. They were characterized by the cyclic voltammograms (CVs) of modified electrodes step-by-step in K₃[Fe(CN)₆] solution (Fig. 1). At the bare gold electrode, a pair of well-defined peaks was obtained with anodic and cathodic peak potentials of 0.28 and 0.18 V, respectively (blue line). The peaks were reduced in size and separated from 0.3 to 0.17 V after the immobilization of the substrate (red line). Peak currents were much more suppressed when double-stranded DNA formed between DNAzyme and the substrate (green line). These account for the fact that the backbone of DNA is charged negatively. Immobilized DNA impedes the approach of Fe(CN)₆³⁻ from the electrode, and the duplex does more so. This result validated the successful immobilization of the substrate and the hybridization of DNAzyme and the substrate.

Fig. 1 CVs of (blue) a bare gold electrode, (red) a substrate-modified gold electrode, and (green) a substrate–DNAzyme-modified gold electrode in 5 mM K₃[Fe(CN)₆] solution containing 0.1 M KCl at scan rate of 0.05 V s⁻¹.

The possibility of sensing lead by the proposed biosensor was also used to validate the preparation of the biosensor. As shown in Fig. 2, with the increase of the concentration of lead, the ECL intensity decreased. An evident intensity as much as 3463 was observed in lead-free solution adopting the ECL biosensor (blue line). It dropped to 1941 after the reaction with 50 pM lead (red line). The emission decreased sharply to 1086 when 1000 pM lead was introduced (green line). The change between the intensity and lead concentration indicates that the biosensor seems lead-responsive and the decrease of the intensity is related directly to the increase of lead concentration. All the results further validated the immobilization of the substrate and the hybridization of DNAzyme and the substrate. The observed ECL emission also illustrated that Ru(phen)₃²⁺ was intercalated into the duplex.

Fig. 2 ECL responses of the proposed biosensor to (blue) blank solution, (red) 50 pM lead, and (green) 1000 pM lead in 0.1 M PBS (pH 7.4) containing 2.5 mM TPA at a constant potential of 1.25 V vs. saturated Ag/AgCl.

Oxidation of TPA by the ECL lead biosensor

The oxidation of TPA, as coreactant, is critical to the intensity of ECL in the ruthenium complex system. CVs of stepwise modified electrodes in 0.1 M PBS (pH 7.4) containing 2.5 mM TPA were investigated (Fig. 3). The anodic peak at ca. 0.85 V vs. saturated Ag/AgCl is attributed to the oxidation of TPA. Improvement of TPA oxidation was observed after the self-assembly of the substrate as a clearly anodic wave was generated (blue and red line). More obvious oxidation related to an increase of peak current occurred once MCH was adsorbed on the electrode surface (green line). The formation of duplex between DNAzyme and its substrate caused the peak current to elevate prominently, indicating a better oxidation of TPA (purple line). Previous research has demonstrated that DNA attached to an electrode pre-concentrates and accepts protons from protonated TPA in weak basic solution by electrostatic interactions between the negatively charged DNA backbone and the positively charged protonated TPA.³⁴ Anchored duplex shows good conductivity through the stacked bases in the delocalized pi-system.^52,53 Meanwhile, immobilized alkane-thiol also pre-concentrates TPA in a hydrophobic manner.⁵⁴ The above factors facilitate the oxidation of TPA. The significant improvement in TPA oxidation facilitates the enhancement of sensitivity.

Fig. 3 CVs of (blue) a bare gold electrode, (red) a substrate-modified gold electrode, (green) a substrate–MCH-modified gold electrode, and (purple) a substrate–MCH–DNAzyme–Ru(phen)₃²⁺-modified gold electrode in 0.1 M PBS (pH 7.4) containing 2.5 mM TPA at scan rate of 0.05 V s⁻¹.

The amount of intercalated Ru(phen)₃²⁺ was relatively low due to the density of the substrate on the electrode surface. Thus, shown in purple line in Fig. 3, the peak at ca. 1.1 V vs. saturated Ag/AgCl did not appear which corresponds to the oxidation of Ru(phen)₃²⁺. An insufficient Ru(phen)₃²⁺ level makes the calculation of substrate density unavailable, upon whose oxidation peak area it is based.⁴⁰

To investigate the mechanism of the oxidation of TPA, CVs of the proposed biosensor in 0.1 M PBS (pH 7.4) containing 2.5 mM TPA at different scan rates were examined (Fig. 4A). At scan rates tested, only anodic peaks corresponding to the oxidation of TPA appeared but no cathodic ones. Hence, an irreversible electrochemical mechanism of TPA oxidation is suggested. Moreover, the peak currents were directly proportional to the square root of scan rates from 0.05 to 0.3 V s⁻¹ (Fig. 4B), which implies that a diffusion-controlled electro-oxidation process happens during TPA oxidation.

Fig. 4 (A) CVs of the proposed biosensor in 0.1 M PBS (pH 7.4) containing 2.5 mM TPA at scan rates of, from inner to outer, 0.05, 0.1, 0.15, 0.2, 0.25, and 0.3 V s⁻¹. (B) Relationship between the peak currents and the square root of scan rates with a correlation coefficient of 0.993.

Performance of the ECL lead biosensor

Strong emission can be obtained by applying the technique of potential step voltammetry. Unlike the common potential scan, potential step voltammetry produces excited Ru(phen)₃²⁺ through direct and TPA-assisted oxidation in just a twinkle. Additionally, the measurement of ECL is accomplished in seconds, so the potential step technique was preferred in this work. By applying a potential step from 0 to 1.25 V, TPA and Ru(phen)₃²⁺ are oxidized simultaneously. ECL responses of the proposed biosensor to different concentrations of lead in 0.1 M PBS (pH 7.4) containing 2.5 mM TPA are depicted in Fig. 5A. The intensity decreased as the lead concentration increased with a linear response to the logarithm of concentrations from 2 to 1000 pM of lead (Fig. 5B). The limit of detection of 0.9 pM was calculated based on a signal-to-noise ratio of 3 and this is lower than most of the reported lead biosensors (Table 1).

Fig. 5 (A) ECL responses of the proposed biosensor to 0, 2, 10, 50, 200, and 1000 pM lead in 0.1 M PBS, at pH 7.4 containing 2.5 mM TPA at a constant potential of 1.25 V vs. saturated Ag/AgCl. (B) Relationship between the intensity decrease and the logarithm of concentrations with a correlation coefficient of 0.992. Error bars represent the standard deviation of three measurements.

Table 1 Comparison of method and limit of detection of lead biosensors based on lead-dependent DNAzyme

Method	Limit of detection	Ref.
ECL	0.9 pM	Current work
Fluorescence	600 pM	15
Fluorescence	3.7 nM	16
Fluorescence	300 pM	17
Colorimetry	32 nM	18
Chemiluminescence	1 nM	18
Dynamic light scattering	35 pM	21
Surface plasmon resonance	0.005 pM	24
Electrochemistry	300 nM	29
ECL	1.4 pM	30
ECL	11 pM	31
Surface enhanced Raman scattering	20 nM	32

---

Employing ECL as the transduction signal is considerably favorable to reach such a low limit of detection due to the improved TPA oxidation on the DNA–MCH modified electrode. Two ECL lead biosensors were developed using 8–17 lead-dependent DNAzyme chemically labeled with a ruthenium complex,^29,30 and one of them used potential step to excite ECL. The performance of our biosensor is superior to them, because we take advantage of a more specific GR-5 DNAzyme, the potential step technique and the increased number of Ru(phen)₃²⁺ molecules per DNAzyme strand. The precision of three replicate ECL measurements using the proposed biosensor was tested. The relative standard deviation of the intensity of the biosensor to blank solution was 2.6%, and 5.9% to 1000 pM lead. To other concentrations, they were below 6.4%. The proposed biosensor is reproducible enough for the real applications.

Specificity and selectivity of the ECL lead biosensor

The specificity of the proposed biosensor is assessed with the five metal ions selected as representative interferants, and the ECL responses are shown in Fig. 6. With 50 pM calcium, cobalt, mercury, magnesium, or zinc individually, the ECL intensity was almost identical to that of lead-free blank solution, while 50 pM lead greatly suppressed the intensity. It is speculated that the biosensor is specific enough to distinguish lead from all the tested metals due to the specificity of lead-dependent DNAzyme.

Fig. 6 ECL measurements using the proposed biosensor in blank solution, 50 pM divalent metal ions including lead, calcium, cobalt, mercury, magnesium, and zinc. Error bars represent the standard deviation of three measurements.

The concentration of lead in the blood reflects the level of exposure to lead, which is useful for diagnostic purposes. Since there is no evidence to confirm that lead below a threshold amount will not hazard human health, devices that achieve ultra-low limits of detection with high sensitivity and precision are required. The ECL measurements of lead using the proposed biosensor in human serum are summarized in Table 2. The results of the standard protocol ASV, using a microelement analyzer that is operated in hospitals, were used to validate the precision by comparison. After the sample was digested and diluted with 50 mM Tris–acetate (pH 8.2), the lead level was determined with the same procedure as was used for the determination of lead in a standard lead solution, except the reported value of ECL is converted to the lead content in blood samples before digestion and dilution. The measured results from the two methods agreed with each other, implying that the proposed biosensor is selective and could be an alternative way for determining lead levels in blood.

Table 2 Measurements of lead using the proposed biosensor and microelement analyzer in human serum

Method	Measurement/μg L^−1
	#1	#2	#3	#4	#5
ASV	119	10.8	5.41	2.70	1.03
ECL	112	10.5	5.29	2.54	0.97

---

Conclusion

In summary, we constructed an ECL lead biosensor based on the intercalation of Ru(phen)₃²⁺ in duplex and GR-5 lead-dependent DNAzyme. The biosensor is highly sensitive and specific, along with an ultra-low limit of detection of 0.9 pM and a dynamic range from 2 to 1000 pM, which enables analysis of trace amounts of lead in serum samples. It also allows one-step measurement of lead. The combination of Ru(phen)₃²⁺-intercalated mode ECL and the cofactor-dependent DNAzyme illustrated in this paper may push the performance of other cofactor-sensing tactics to the extreme.

Acknowledgements

This work was supported by the National Basic Research Program of China (973 Program, Grant no. 2011CB707703), NSF of China (Grant no. 21075068), Tianjin Natural Science Foundation (Grant no. 11JCZDJC22200) and the Fundamental Research Funds for the Central Universities.

References

1. Toxicological Profile for Lead, In Agency for Toxic Substances and Disease Registry, Atlanta, Georgia, 2007.
2. C. R. T. Tarley, F. N. Andrade, F. M. de Oliveira, M. Z. Corazza, L. F. M. de Azevedo and M. G. Segatelli, Anal. Chim. Acta, 2011, 703, 145–151.
3. H.-F. Hsieh, W.-S. Chang, Y.-K. Hsieh and C.-F. Wang, Talanta, 2009, 79, 183–188.
4. D. I. Bannon and J. J. Chisolm, Clin. Chem., 2001, 47, 1703–1704.
5. I. Willner, B. Shlyahovsky, M. Zayats and B. Willner, Chem. Soc. Rev., 2008, 37, 1153–1165.
6. J. Kosman and B. Juskowiak, Anal. Chim. Acta, 2011, 707, 7–17.
7. P. Travascio, Y. Li and D. Sen, Chem. Biol., 1998, 5, 505–517.
8. R. R. Breaker and G. F. Joyce, Chem. Biol., 1994, 1, 223–229.
9. A. K. Brown, J. Li, C. M.-B. Pavot and Y. Lu, Biochemistry, 2003, 42, 7152–7161.
10. J. Li and Y. Lu, J. Am. Chem. Soc., 2000, 122, 10466–10467.
11. J. Liu and Y. Lu, J. Am. Chem. Soc., 2003, 125, 6642–6643.
12. J. Liu and Y. Lu, J. Am. Chem. Soc., 2004, 126, 12298–12305.
13. J. Liu and Y. Lu, J. Am. Chem. Soc., 2005, 127, 12677–12683.
14. J. Liu and Y. Lu, Anal. Chem., 2003, 75, 6666–6672.
15. X.-B. Zhang, Z. Wang, H. Xing, Y. Xiang and Y. Lu, Anal. Chem., 2010, 82, 5005–5011.
16. T. Lan, K. Furuya and Y. Lu, Chem. Commun., 2010, 46, 3896–3898.
17. X.-H. Zhao, R.-M. Kong, X.-B. Zhang, H.-M. Meng, W.-N. Liu, W. Tan, G.-L. Shen and R.-Q. Yu, Anal. Chem., 2011, 83, 5062–5066.
18. T. Li, E. Wang and S. Dong, Anal. Chem., 2010, 82, 1515–1520.
19. J. Elbaz, B. Shlyahovsky and I. Willner, Chem. Commun., 2008, 1569–1571.
20. D. Nie, H. Wu, Q. Zheng, L. Guo, P. Ye, Y. Hao, Y. Li, F. Fu and Y. Guo, Chem. Commun., 2012, 48, 1150–1152.
21. X. Miao, L. Ling and X. Shuai, Chem. Commun., 2011, 47, 4192–4194.
22. X. M. Miao, L. S. Ling and X. T. Shuai, Anal. Biochem., 2012, 421, 582–586.
23. Z. Lin, Y. Chen, X. Li and W. Fang, Analyst, 2011, 136, 2367–2372.
24. G. Pelossof, R. Tel-Vered and I. Willner, Anal. Chem., 2012, 84, 3703–3709.
25. L. Shen, Z. Chen, Y. Li, S. He, S. Xie, X. Xu, Z. Liang, X. Meng, Q. Li, Z. Zhu, M. Li, X. C. Le and Y. Shao, Anal. Chem., 2008, 80, 6323–6328.
26. H. Zhang, B. Jiang, Y. Xiang, J. Su, Y. Chai and R. Yuan, Biosens. Bioelectron., 2011, 28, 135–138.
27. F. Li, Y. Feng, C. Zhao and B. Tang, Chem. Commun., 2011, 47, 11909–11911.
28. X. Yang, J. Xu, X. Tang, H. Liu and D. Tian, Chem. Commun., 2010, 46, 3107–3109.
29. Y. Xiao, A. A. Rowe and K. W. Plaxco, J. Am. Chem. Soc., 2006, 129, 262–263.
30. F. Ma, B. Sun, H. Qi, H. Zhang, Q. Gao and C. Zhang, Anal. Chim. Acta, 2011, 683, 234–241.
31. X. Zhu, Z. Lin, L. Chen, B. Qiu and G. Chen, Chem. Commun., 2009, 6050–6052.
32. Y. Wang and J. Irudayaraj, Chem. Commun., 2011, 47, 4394–4396.
33. X.-B. Yin, TrAC, Trends Anal. Chem., 2012, 33, 81–94.
34. X.-B. Yin, Y.-Y. Xin and Y. Zhao, Anal. Chem., 2009, 81, 9299–9305.
35. D.-Y. Liu, Y.-Y. Xin, X.-W. He and X.-B. Yin, Biosens. Bioelectron., 2011, 26, 2703–2706.
36. C.-X. Tang, N.-N. Bu, X.-W. He and X.-B. Yin, Chem. Commun., 2011, 47, 12304–12306.
37. N.-N. Bu, C.-X. Tang, X.-W. He and X.-B. Yin, Chem. Commun., 2011, 47, 7689–7691.
38. Y. Zhao, X.-W. He and X.-B. Yin, Chem. Commun., 2011, 47, 6419–6421.
39. C.-X. Tang, Y. Zhao, X.-W. He and X.-B. Yin, Chem. Commun., 2010, 46, 9022–9024.
40. D.-Y. Liu, Y.-Y. Xin, X.-W. He and X.-B. Yin, Analyst, 2011, 136, 479–485.
41. D.-Y. Liu, Y. Zhao, X.-W. He and X.-B. Yin, Biosens. Bioelectron., 2011, 26, 2905–2910.
42. Y. Chen, J. Xu, J. Su, Y. Xiang, R. Yuan and Y. Chai, Anal. Chem., 2012, 84, 7750–7755.
43. Z. Liu, W. Zhang, L. Hu, H. Li, S. Zhu and G. Xu, Chem.–Eur. J., 2010, 16, 13356–13359.
44. T. Yuan, Z. Liu, L. Hu, L. Zhang and G. Xu, Chem. Commun., 2011, 47, 11951–11953.
45. W. Liu, H. Zhu, B. Zheng, S. Cheng, Y. Fu, W. Li, T. C. Lau and H. Liang, Nucleic Acids Res., 2012, 40, 4229–4236.
46. B. B. Sha, X. B. Yin, X. H. Zhang, X. W. He and W. L. Yang, J. Chromatogr., A, 2007, 1167, 109–115.
47. Y. Xiao, T. Uzawa, R. J. White, D. DeMartini and K. W. Plaxco, Electroanalysis, 2009, 21, 1267–1271.
48. Y. Xiao, R. Y. Lai and K. W. Plaxco, Nat. Protoc., 2007, 2, 2875–2880.
49. J. Li, W. Zheng, A. H. Kwon and Y. Lu, Nucleic Acids Res., 2000, 28, 481–488.
50. S. W. Santoro and G. F. Joyce, Proc. Natl. Acad. Sci. U. S. A., 1997, 94, 4262–4266.
51. H. Y. Ding, Y. Zhou, S. J. Zhang, X. B. Yin, Y. J. Li and X. W. He, Chin. J. Anal. Chem., 2008, 36, 839–842.
52. H.-W. Fink and C. Schonenberger, Nature, 1999, 398, 407–410.
53. S. O. Kelley, N. M. Jackson, M. G. Hill and J. K. Barton, Angew. Chem., Int. Ed., 1999, 38, 941–945.
54. Y. Zu and A. J. Bard, Anal. Chem., 2001, 73, 3960–3964.

---