Biosensors and Bioelectronics 68 (2015) 303–309
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A simple and label-free aptasensor based on nickel hexacyanoferrate nanoparticles as signal probe for highly sensitive detection of 17β-estradiol Lifang Fan a,b, Guohua Zhao a,n, Huijie Shi a, Meichun Liu a a b
Department of Chemistry, Tongji University, 1239 Siping Road, 200092 Shanghai, China College of Chemistry and Chemical Engineering, Shanxi Datong University, Datong 037009, China
art ic l e i nf o
a b s t r a c t
Article history: Received 21 October 2014 Received in revised form 17 December 2014 Accepted 7 January 2015 Available online 8 January 2015
A simple and label-free electrochemical aptasensor was developed for detecting 17β-estradiol (E2). To translate the binding events between aptamer and E2 into the measurable electrochemical signal, the nickel hexacyanoferrate nanoparticles (NiHCF NPs) as signal probe was in situ introduced on the electrode by a simple two-step deposition method, exhibiting well-defined peaks with good stability and reproducibility. Subsequently, Au nanoparticles (Au NPs) was covered on the NiHCF NPs, which not only provided a platform for immobilizing the aptamer by S–Au interaction, but further enhanced the conductivity and stability of the signal probe. With the addition of E2, the formation of E2–aptamer complexes on the sensing interface retarded the interfacial electron transfer reaction of the probe, resulting in the decrease of the electrochemical signal. E2 could be readily examined by measuring the signal change. A linear range of 1 10 12–6 10 10 M was obtained with a low detection limit of 0.8 10 12 M. The aptasensor also exhibited high specificity to E2 in control experiments employing seven endocrine disrupting compounds as the interferents that had similar structure or coexisted with E2 in the environment. Besides, the applicability of the aptasensor was successfully evaluated by determining E2 in the real samples. & 2015 Elsevier B.V. All rights reserved.
Keywords: Aptasensor 17β-estradiol Nickel hexacyanoferrate nanoparticles
1. Introduction Endocrine disrupting chemicals (EDCs) are a class of ubiquitous contaminants in the environment, which seriously interfere with the normal endocrine function, further causing adverse effects on growth, metabolism and reproduction of the organisms, and increasing the incidence of cancer and tumors. (Lust et al., 2012; Racz and Goel, 2010; Vajda et al., 2008). Among the EDCs, 17βestradiol (E2) excreted by humans and animals, is a natural and most active estrogen, which can enter surface waters via discharge of municipal wastewater and industrial facility effluents, and discharge from animal feeding operations and septic system, etc. (Writer et al., 2012). The chronic exposure of human to E2, even at very low concentration will cause the drastic health problems (Wang et al., 2011). Therefore, it is necessary to establish a simple, rapid and sensitive method for the detection of E2 in the environment. So far, these methods, such as HPLC (Mahmoud et al., 2011; n
Corresponding author. Fax: þ 86 21 65982287. E-mail address: [email protected] (G. Zhao).
http://dx.doi.org/10.1016/j.bios.2015.01.015 0956-5663/& 2015 Elsevier B.V. All rights reserved.
Mishra and Joy, 2006; Yoon et al., 2003) or GC/MS (Rocha et al., 2011; Tsakalof et al., 2012) have been reported for detecting E2. These conventional methods are accurate and sensitive, but the analytical procedures are complicated and time-consuming. Considering these limitations, some biosensors based on human estrogen receptor (hER) and antibodies have been developed (Le Blanc et al., 2009; Liu et al., 2012, 2010). Although convenient and rapid, but the hER-based biosensors are lacking in selectivity for E2 detection because of the affinity of hER to other xenoendocrinesthes, whereas the immunosensors are better in the selectivity, but they are susceptible to harsh conditions, and the extraction of antibodies is complicated and laborious. Aptamers offer a new alternative to antibodies as biorecognition molecules, meanwhile, they have more advantages over antibodies, such as smaller size, easier synthesis and modification, and better stability. The aptamer-based biosensors have been reported via employing different techniques, such as fluorescence (Yildirim et al., 2012), surface plasmon resonance (Zhang et al., 2013), colorimetry (Kim et al., 2011) and electrochemistry (Lin et al., 2012). Among these aptasensors, electrochemical aptasensors have been widely concerned due to its outstanding advantages, including simple equipment,
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fast response, low cost and real-time detection. However, how to translate the binding events between aptamer and target into a measurable electrochemical signal is the key point in the development of electrochemical aptasensors. Generally, additional electroactive tags, for example, ferrocene derivatives (Song et al., 2012; Sosniak et al., 2009), methylene blue (Ferapontova and Gothelf, 2009; Liu et al., 2011), and ruthenium complexes (Vrabel et al., 2009), etc. have been used to label aptamer to obtain the measurable signal. But labeling aptamer is a complex and time-consuming process, even which might affect the affinity of aptamer toward target. Besides, electroactive compounds can also be directly added into test solution to generate a measurable signal. For instance, the aptasensors based on electrochemical impedance spectroscopy (EIS) by employing [Fe(CN)6]3 /4 as the indicator are commonly used owing to its simple and rapid response with high sensitivity (Gonzalez-Fernandez et al., 2011; Kashefi-Kheyrabadi and Mehrgardi, 2012; Tran et al., 2011). However, according to the literature reported (Bogomolova et al., 2009) and our previous works (Fan et al., 2013; Ke et al., 2014), the disadvantages of EIS aptasensors cannot be neglected. They are usually prone to false-positive results produced by the limitations, including initial sensing interface contamination, excess measurements on the same sensor as well as additional incubations in the [Fe(CN)6]3 / 4 between measurements for a long time (Bogomolova et al., 2009). So it is difficult to determine accurately targets except for performing vast and timeconsuming control or parallel experiments. In this work, in order to overcome effectively these drawbacks of labeling aptamer or adding additional probe into test system, a signal probe, the nickel hexacyanoferrate nanoparticles (NiHCF NPs) was chosen and in situ designed on the electrode due to its outstanding properties, such as easy synthesis, wonderful stability and good peak shape. Then Au nanoparticles (Au NPs) was deposited uniformly on the NiHCF NPs to act as the matrix for anchoring the aptamer and improve the conductivity of the electrode. Thus, a simple and label-free electrochemical aptasensor was fabricated for E2 detection. When it was exposed to E2 solution, the formation of the E2–aptamer complexes on the sensing interface by binding E2 hindered the electron transfer, leading to the decrease of the current. E2 could be quantified via measuring the current change. The sensing performance of the aptasensor for detecting E2 was investigated in detailed. Meanwhile, its practical application was evaluated by detecting the real samples.
2. Experiment 2.1. Reagents Aptamer, was chosen according to the prior literature (Kim et al., 2007). The oligonucleotides were purchased from Sangon Biotechnology Co. Ltd. (Shanghai, China).
ethanol, pH 8.0) was prepared and stored at 4 °C. All other chemical materials were of analytical reagent grade. 2.2. Apparatus Electrochemical measurements were performed with CHI 660C electrochemical workstation (CHI, USA) based on a conventional three-electrode system with a bare or modified gold electrode (d ¼2 mm) acted as the working electrode, a saturated calomel electrode (SCE) as the reference electrode and a platinum wire as the counter electrode. The morphology and energy dispersive X-ray spectroscopy (EDS) spectra of different modified electrodes were measured by scanning electron microscopy (FE-SEM, S-4800, Hitachi). 2.3. Preparation of the signal probe-modified electrode The gold electrode was polished by using 1.0, 0.3 and 0.05 μm alumina powers, respectively. The polished electrode was placed in the freshly Piranha solution for 5 min (Caution!), followed by ultrasonically cleaned. It was pretreated in 0.5 M H2SO4 by potential scanning between 0.2 and 1.5 V until a reproducible cyclic voltammogram was obtained. The cleaned electrode was put into 50 mM NiCl2 6H2Oþ 10 mM NH4Cl of pH 4.55, where metallic Ni was electrodeposited on the electrode at the potential of 0.9 V. The Ni-deposited electrode was transferred into 0.1 M NaNO3 and 5 mM K3[Fe(CN)6] and scanned by I–t technique at 1.0 V. The scanning was continued until the current decreased to almost zero to ensure that all of metallic Ni had been converted to NiHCF NPs. Thus, the signal probe-modified electrode was prepared by the two-step deposition method. 2.4. Design and construction of the aptasensor The as-prepared NiHCF NPs-modified electrode was immersed in 2.5 mM HAuCl4 containing 0.1 M KCl and 50 mM H2SO4, where Au NPs was deposited on the NiHCF NPs by cyclic voltammetry (CV) at N2 atmosphere (Au NPs/NiHCF NPs-modified electrode). Subsequently, it was incubated in 2.0 mM aptamer solution for over 12 h (aptamer/Au NPs/NiHCF NPs-modified electrode). Before incubation, 0.2 mM TCEP was used to reduce the disulfide bond of aptamer. Finally, 1 mM MCH was employed to block the unbound active sites of Au NPs (MCH/aptamer/Au NPs/NiHCF NPs-modified electrode). Any nonspecific binding was inhibited except for the aptamer–E2 interaction. The aptasensor was prepared and stored at 4 °C. Scheme 1 illustrates the schematic representation of the aptasensor construction and mechanism for detecting E2. To reuse the aptasensor, it was incubated in a 0.5% sodium dodecyl sulfate (SDS) solution (pH 1.9) for 3 min and washed with PBS (Yildirim et al., 2012). 2.5. Electrochemical measurements
Aptamer sequence The EIS for monitoring the different stages of the aptasensor 5′-SH-(CH2)6-GCT-TCC-AGC-TTA-TTG-AAT-TAC-ACG-CAG-AGG-GTA- construction was performed in 0.1 M PBS (pH 7.41) containing GCG-GCT-CTG-CGC-ATT-CAA-TTG-CTG-CGC-GCT-GAA-GCG-CGG- 5.0 mM K3[Fe(CN)6]/K4[Fe(CN)6] and 0.1 M KCl. EIS measurements AAG-C-3, were recorded between 1000 Hz and 0.1 Hz with a sinusoidal Random DNA sequence voltage perturbation of 5 mV amplitude. In the detection of E2, 5′-(SH)-(CH2)6-CTG-ACA-CCA-TAT-TAT-GAA-GA-3′ 20 μL different concentrations of E2 were dropped onto the sensing interface for 40 min at room temperature, following by E2 (99.9%), Hydrogen tetrachloroaurate (III) hydrate (HAuCl4) thoroughly rinsed to remove unbound E2. Before and after the binding of E2, the current of the aptasensor was measured by and Tris (2-carboxyethy1) phosphine (TCEP) were purchased from differential pulse voltammetry (DPV) in 0.1 M PBS (pH 7.41) as the Sigma-Aldrich (St. Louis, MO). 6-Mercap-1-hexanol (MCH) was supporting electrolyte. obtained from Adamas Reagent Co. Ltd. DNA sequences were dissolved in Tris–EDTA buffer (TE, pH 8.0). Tris–HCl buffer The two real samples, municipal wastewater, were collected from two living areas in Yangpu District, Shanghai. Prior to (100 mM Tris–HCl/200 mM NaCl/25 mM KCl/10 mM MgCl2, 5%
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Scheme 1. Illustration of the electrochemical aptasensor construction and the mechanism for detection of E2.
analysis, the samples were filtered through an ordinary filter paper and the membrane with the size of 0.22 μm, respectively, to remove the suspended solid matter and small particles. The substances dissolved in these samples were not changed by the pretreatment. The treated samples were detected as soon as possible to minimize microbial degradation. Furthermore, the samples were diluted to decrease the interference of complex matrices.
3. Results and discussion 3.1. Construction and properties of the signal probe The argenteous metallic Ni uniformly deposited on the gold
was oxidized to Ni2 þ and subsequently reacted with the hexacyanoferrate (III), forming the signal probe, NiHCF NPs. NiHCNFe NPs, similar to other nickel hexacyanoferrate, is an inorganic polymer analog from Prussian blue family (Steen et al., 2002). NiHCNFe complexes have two different types (NiHCNFe-II and NiHCNFe-III), which can be formed on the electrode together or alone when changing the preparation conditions (Hao et al., 2012). Here, NiHCF NPs-III was directly formed on the electrode by chemical reaction between Ni2 þ and [Fe(CN)6]3 in the deposition potential of 1.0 V and its molecular compositions is Ni3[Fe(CN)6]2 (Chen et al., 2009). Fig. 1A shows the SEM morphology of the NiHCF NPs, it is uniformly covered on the electrode, with the diameter of about 10 nm. The thickness of the probe depended on the amount of
Fig. 1. (A) SEM of the signal probe and the inset is SEM of the Au NPs/NiHCF NPs-modified electrode; (B) the EDS spectrum of the Au NPs/NiHCF NPs-modified electrode and the inset is the elemental mapping of Au (gray), Ni (green), Fe (red), C (purple) and N (dark red) with color superposition in the electrode; (C) CV curves of the signal probe at different scan rate (from a to h: 20, 40, 60, 80, 100, 120, 160, 200 mV/s); The inset is the dependence of the redox peak current on the scan rate and (D) CV curves of the signal probe in 0.1 M PBS (pH 7.41), Scan cycles: 30. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article).
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deposited Ni which could be controlled by the factors, including the concentration of NiCl2 and the deposition time. In order to achieve the excellent properties of the probe, the two factors were optimized in detailed and given in Figs. S1 and S2 in Supplementary information. The results demonstrate that the optimized concentration of NiCl2 is 50 mM, and the optimized deposition time is 60 s. Fig. 1C shows the CV curves of the probe at the different scan rate. It can be observed that a pair of well-defined redox peaks are present in the absence of any electroactive substance. The Ni2 þ in the NiHCF NPs cannot be reduced in the potential range of 0.0– 0.7 V, while the redox peaks correspond to the hexacyanoferrate (III:II) redox couple (Lin and Bocarsly, 1991) due to the formation of NiHCNFe-(II) on the electrode by electrochemical reduction of [Fe(CN)6]3 to [Fe(CN)6]4 and in turn coordination to Ni2 þ in the potential cycling scans. The peak width at half-height is approximately 96 mV, which is consistent with the literatures (Cai et al., 1995; Pournaghi-Azar and Razmi-Nerbin, 1998), approaching the theoretical value of a one-electron reaction mechanism (90.6/ nm V) (Bard and Faulkner, 1980). The relationship of the peak current with the different scan rate is shown in the inset of Fig. 1C. The anodic and cathodic current increase linearly with the scan rate range from 20 to 200 mV/s, as predicted for a diffusionless system. The peak separation ΔEp (ΔEp ¼ Ea Ec) is about 16 mV at the scan rate of 20 mV/s, and ΔEp is further enhanced with increasing the scan rate, for instance, 39 mV of ΔEp occurs approximately at 100 mV/s. For a fully reversible reaction in which no reactants from the solution take place, the separation of peaks should be zero (Cai et al., 1995). The relatively small ΔEp indicates fairly fast electrochemical reaction rate on the NiHCF NPs-modified electrode (Laviron, 1979a). Meanwhile, the surface coverage (Γ) of the NiHCF NPs on the electrode could be evaluated for 5.761 10 8 mol cm 2 by the equation Γ = Q /nFA (Laviron, 1979b). The detailed calculation for Γ was given in the supporting information. Besides, the stability of the signal probe was investigated by directly exposing it to air for a week or putting in 0.1 M PBS for three days, then recording the CV response. The results indicated that its stability was not affected by air. Furthermore, there was not any loss of its electroactivity after storing in 0.1 M PBS. The reproducibility of the probe was also measured, shown in Fig. 1D. The height and the ΔEp of the redox peaks were almost no change after 30 cycles of repetitive CV scanning, indicating that the signal probe had good reproducibility. Afterwards, Au NPs was deposited on the probe to act as a platform for immobilizing aptamer and further improve the conductivity and stability of the electrode. The SEM of the Au NPs/ NiHCF NPs-modified electrode is given in the inset of Fig. 1A. Au
NPs is uniformly dispersed on the probe, with the diameter of 50– 100 nm. The EDS spectrum in Fig. 1B demonstrates that the elements including Au, Ni, Fe, C and N are present in the Au NPs/ NiHCF NPs-modified electrode. Furthermore, the elemental mapping confirms that the five elements are highly dispersed on this modified electrode. In addition, the EDS element analysis shows that the atom percent of Ni and Fe is 4.88% and 3.27%, respectively. The atom ratio of Ni and Fe is 1.49:1, confirming that Ni3[Fe(CN)6]2 complex is present in the NiHCF NPs when Ni (II) reacts with [Fe(CN)6]3 . This result is agree with that of the literatures reported. (Chen et al., 2009; Carpani et al., 2006). 3.2. Design and construction of the aptasensor In order to characterize the immobilization results, the different stages of the aptasensor construction were investigated by CV and DPV. The CV curves are shown in Fig. 2A, there is no peak on the bare gold electrode and the current is almost zero in the absence of any electroactive substance (curve a). After depositing the signal probe on the electrode, there is a couple of well-defined redox peaks (curve b), corresponding to the hexacyanoferrate (III:II) redox couple, indicating that the probe has been successfully introduced on the electrode. Furthermore, the ΔEp of the peaks is about 40.0 mV and the ratio of the anodic and cathodic peak current (ia:ic) equals to one, revealing good reversibility of the probe. After depositing Au NPs, the current of the redox peaks decreases somewhat (curve c), owing to the decrease of some electroactive sites of the probe encased by Au NPs. The ΔEp decreases to 38.0 mV, implying that the conductivity of the modified electrode is improved. When aptamer was immobilized on the Au NPs/NiHCF NPs-modified electrode, the current of the redox peak decreased and had poor reversibility (curve d). The reason can be attributed to that the formation of a selfassembled monolayer of aptamer retard the electron transfer reaction of the signal probe. Finally, the aptamer-modified electrode was treated with MCH, and the current further decreased (curve e). Meanwhile, the DPV curves for characterization of the aptasensor construction are shown in Fig. 2B. There is nearly a line on the bare gold electrode (curve a). While the NiHCF NPs was electrodeposited, a large current of 146.9 μA (curve b) was recorded, implying the probe had good electroactivity. After depositing Au NPs, the current decreases to 131.2 μA, and the peak potential moves negatively (curve c). This result demonstrates that the electroactive sites of NiHCF NPs decrease due to the coating of Au NPs, which is consistent with the result of the CV measurements. After immobilizing aptamer, the current further decreases to 85.1 μA. Finally, the aptamer-modified electrode was blocked using MCH and the current decreased to 78.8 μA. Besides, EIS
Fig. 2. CV (A) and DPV (B) curves of the bare gold electrode (a), the NiHCF NPs-modified electrode (b), the Au NPs/NiHCF NPs-modified electrode (c), the aptamer/Au NPs/ NiHCF NPs-modified electrode (d) and the MCH/aptamer/Au NPs/NiHCF NPs-modified electrode (e) in 0.1 M PBS (pH 7.41).
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measurements were also carried out in the presence of [Fe(CN)6]3 /4 (Fig. S3) to evaluate the interfacial electrochemistry properties in the construction of the aptasensor. The characterization results of EIS totally agreed with those of CV and DPV, confirming that the aptasensor based on the NiHCF NPs as the signal probe was successfully fabricated. In addition, the load of aptamer on the sensing interface has great impact on the sensitivity of the aptasensor. Herein, the coverage density (Γ) of aptamer anchored was estimated by chronocoulometry employing 0.8 mM [Ru(NH3)6]3 þ as a cationic redox marker which bound electrostatically to the anionic phosphate groups of aptamer (Steel et al., 1998). Γ was calculated to be 7.67 1012 molecules cm 2. The control experiments were also performed utilizing the bare gold electrode as the matrix for modifying aptamer. The load of aptamer on the gold electrode was about 1.24 1012 molecules cm 2 (The detailed calculation was given in Fig. S4). The coverage density of aptamer on the Au NPs/ NiHCF NPs-modified electrode is almost six-fold more than that on the gold electrode. So Au NPs with the large surface area is more advantageous to load aptamer, which greatly improve the sensitivity of the aptasensor. 3.3. Analytical performance of the electrochemical aptasensor The analytical performance of the aptasensor was evaluated in different concentrations of E2 solution by DPV, shown in Fig. 3A. The current of the aptasensor decreases with increasing concentrations of E2. This is mainly attributed to the fact that more and more aptamer–E2 complexes formed on the sensing interface by capturing vast E2 hinder the electron transfer, resulting in the decrease of the current. Fig. 3B shows the dependence of the current change ΔI and different E2 concentrations. It is apparent that ΔI reach a plateau at E2 concentration of more than 600 pM, indicating that the amount of E2–aptamer complexes on the sensing interface reaches their saturation. A relationship curve between ΔI/I0 on the vertical ordinate and the logarithm of E2 concentrations on the horizontal axis is plotted, as is shown in the inset of Fig. 3B. It can be observed that ΔI/I0 is linear against the logarithm value of E2 concentrations and a linear dynamic range from 5.0 to 600.0 pM is obtained. The regression equation is ΔI/I0 ¼0.2894 log Cþ 0.1213 (unit of C, pM) with a correction coefficient of 0.9935. The limit of detection is estimated to be 0.8 pM based on the response of three times the standard deviation of zero-pose response (n ¼5) and the limit of quantitation is 5.0 pM. Some detailed comparisons of the analytical behavior of the aptasensor with other published methods for detecting E2 were given in Table S1. It can be seen that the
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sensitivity of the immunosensor or impedimetric aptasensor for E2 detection (Liu et al., 2010; Lin et al., 2012) is lower than that of the present aptasensor. Although high sensitivity was also achieved by HPLC (Farre et al., 2007; Rodriguez-Mozaz et al., 2004), compared to these conventional methods, the analysis progress by the present aptasensor is simpler, rapider and more low-cost without complicate sample pretreatments and expensive instruments. It is thus evident that the aptasensor can be applied to detect low level of E2 in the environment. In addition, the experimental results demonstrated that high sensitivity of the aptasensor was related with the parameters, such as the amount of Au NPs on the probe, controlled by the numbers of CV cycles in HAuCl4 solution, and the incubation time of the aptasensor in E2 solution. The parameters were optimized in detailed and given in Figs. S5 and S6. When the optimized number of CV cycles for Au NPs is 15 and the incubation time is 40 min, the aptasensor can arrive the best response for E2 detection. To investigate the selectivity of the aptasensor, seven endocrine disrupting chemicals including ethinylestradiol, bisphenol A, estriol, polychlorinated biphenyl (PCB 101), diethyl phthalate, 4-nonyl phenol and atrazine were used as the interferents, which could have similar structure or coexist with E2 in the environment. The aptasensor was incubated in 5.0 pM of each one among seven interferents, respectively and its current response was measured with the same method for determining E2. The current difference, ΔI obtained before and after incubation in 5.0 pM E2 and the interferent with the same concentration is shown in Fig. 4A. The results illustrate that ΔI is approximately 17 μA in 5.0 pM E2, whereas ΔI in any one of other interferents is less than 2.0 μA. The current response of the aptasensor for these interferents is less than 8.0% for that for E2, indicating high selectivity and specificity of the aptasensor to E2. Control experiments were performed with a random DNA sequence as the recognition element instead of aptamer for determining different concentrations of E2, shown in Fig. 4B. With increasing the concentrations of E2, the peak current is almost unchanged. This is attributed to the fact that the random DNA-based sensor has no recognition ability for E2, so that E2 is not captured on the sensing interface, further confirming that high specificity of the aptasensor toward E2 is closely related to the specific sequence of the aptamer. In addition, the long-term storage stability, the reproducibility and the reusability of the aptasensor were also investigated. When the aptasensor was stored in 0.1 M PBS at 4 °C for three days, it was used to determine 5.0 pM E2. The peak current was lower only 3.2% than the original response. After the storage for a week, the current decreased to 5.3% of the original response, suggesting that the aptasensor had good stability. The reproducibility of the
Fig. 3. (A) DPV response of the aptasensor in different concentrations of E2 (from a to j: 0, 1, 5, 10, 50, 100, 200, 400, 600, 1000 pM) and (B) the dependence of the peak current on the concentrations of E2; the inset is the linear calibration curve of ΔI/I0 with logarithm of E2 concentrations.
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Fig. 4. The selectivity and specificity test of the aptasensor for E2 detection.
Table 1 Detection of the wastewater samples with E2 at different concentrations. Sample
Sample 1
Sample 2
a b
Determined (pM)a
Recovery (%)
RSD (%)b
0.0 5.0 10.0 50.0 0.0
Undetected 4.8 9.8 50.1 Undetected
96.6 98.1 100.2
3.6 2.0 1.1
5.0 10.0 50.0
4.7 9.9 49.8
93.6 98.5 99.6
2.4 2.9 1.1
Spiked (pM)
Mean of three measurements. Relative standard deviation.
aptasensor was evaluated by an intraassay and an interassay relative standard deviation (RSD). The intraassay RSD was 8.0% by detecting 5.0 pM E2 with five replicated measurements. Then, five identical aptasensors were used to determine 5.0 pM E2, and the interasssay RSD was 7.2%, showing that the aptasensor had an acceptable reproducibility. Besides, the reusability of the aptasensor was also important in the practical application. The E2– aptamer complexes on the sensing interface could be effectively separated in 0.5% SDS solution (pH 1.9) for 3 min without any destruction of the binding affinity of aptamer toward E2. After regeneration, the current response as percent of the initial signal is given in Fig. S7. Over 80% of the initial response can be observed after repeated use for three times. 3.4. Application of the electrochemical aptasensor In order to evaluate the feasibility of the aptasensor for practical analysis, it was used to determine E2 in the real samples using the standard addition method. Three different concentrations of the standard solutions, 5.0, 10.0, 50.0 pM were added into the two pretreated samples, respectively. The recoveries results of E2 in the two different samples are shown in Table 1. It can be observed that the average recoveries range from 93.6% to 100.2% with RSD lower than 5.2% based on triplicate experiments at each concentration. The results indicate that the aptasensor can resist the interference of the complex matrices and be applied to determine E2 in different environmental wastewater.
4. Conclusions In summary, the electrochemical aptasensor was successfully developed for the detection of E2 based on the NiHCF NPs as the
in situ signal probe which overcame effectively the drawbacks of labeling aptamer or adding additional probe into test system. The aptasensor showed good analytical performance for E2 detection with high sensitivity, specificity and reproducibility. A low detection limit of 0.8 pM was obtained, which was comparable or even lower than the previous reports. Moreover, the aptasensor was successfully applied to detect the real samples, and the satisfactory recoveries were obtained.
Acknowledgments This work was financially supported by the National Natural Science Foundation of China (NSFC, No. 21277099).
Appendix A. Supplementary material Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.bios.2015.01.015.
References Bard, A.J., Faulkner, L.R., 1980. Electrochemical Methods: Fundamentals and Applications. Wiley, New York. Bogomolova, A., Komarova, E., Reber, K., Gerasimov, T., Yavuz, O., Bhatt, S., Aldissi, M., 2009. Anal. Chem. 81, 3944–3949. Cai, C.X., Ju, H.X., Chen, H.Y., 1995. Anal. Chim. Acta 310, 145–151. Carpani, I., Giorgetti, M., Berrettoni, M., Buldini, P.L., Gazzano, M., Tonelli, D., 2006. J. Solid State Chem. 179, 3981–3988. Chen, W., Tang, J., Xia, X.H., 2009. J. Phys. Chem. C 113, 21577–21581. Fan, L.F., Zhao, G.H., Shi, H.J., Liu, M.C., Li, Z.X., 2013. Biosens. Bioelectron. 43, 12–18. Ferapontova, E.E., Gothelf, K.V., 2009. Electroanalysis 21, 1261–1266. Farre, M., Kuster, M., Brix, R., Rubio, F., Lopez de Alda, M.J., Barcelo, D., 2007. J. Chromatogr. A 1160, 166–175. Hao, X., Yan, T., Wang, Z., Liu, S., Liang, Z., Shen, Y., Pritzker, M., 2012. Thin Solid Films 520, 2438–2448. Gonzalez-Fernandez, E., de-los-Santos-Alvarez, N., Lobo-Castanon, M.J., MirandaOrdieres, A.J., Tunon-Blanco, P., 2011. Biosens. Bioelectron. 26, 2354–2360. Kashefi-Kheyrabadi, L., Mehrgardi, M.A., 2012. Biosens. Bioelectron. 33, 184–189. Ke, H., Liu, M., Zhuang, L., Li, Z., Fan, L., Zhao, G., 2014. Electrochim. Acta 137, 146–153. Kim, Y.S., Jung, H.S., Matsuura, T., Lee, H.Y., Kawai, T., Gu, M.B., 2007. Biosens. Bioelectron. 22, 2525–2531. Kim, Y.S., Kim, J.H., Kim, I.A., Lee, S.J., Gu, M.B., 2011. Biosens. Bioelectron. 26, 4058–4063. Laviron, E., 1979a. J. Electroanal. Chem. 101, 19–28. Laviron, E., 1979b. J. Electroanal. Chem. 100, 263–270. Le Blanc, A.F., Albrecht, C., Bonn, T., Fechner, P., Proll, G., Proll, F., Carlquist, M., Gauglitz, G., 2009. Anal. Bioanal. Chem. 395, 1769–1776. Lin, C., Bocarsly, A.B., 1991. J. Electroanal. Chem. 300, 325–345. Lin, Z., Chen, L., Zhang, G., Liu, Q., Qiu, B., Cai, Z., Chen, G., 2012. Analyst 137, 819–822. Liu, S., Lin, Q., Zhang, X., He, X., Xing, X., Lian, W., Li, J., Cui, M., Huang, J., 2012. Sens.
L. Fan et al. / Biosensors and Bioelectronics 68 (2015) 303–309
Actuators B-Chem. 166-167, 562–568. Liu, X., Duckworth, P.A., Wong, D.K., 2010. Biosens. Bioelectron. 25, 1467–1473. Liu, Y., Yan, J., Howland, M.C., Kwa, T., Revzin, A., 2011. Anal. Chem. 83, 8286–8292. Lust, M., Makinia, J., Stensel, H.D., 2012. Water Sci. Technol. 65, 1130–1136. Mahmoud, I.Y., AlKindi, A.Y., Khan, T., Al-Bahry, S.N., 2011. J. Exp. Zool. Part A 315A, 170–174. Mishra, A., Joy, K.P., 2006. Gen. Comp. Endocrinol. 145, 84–91. Pournaghi-Azar, M.H., Razmi-Nerbin, H., 1998. J. Electroanal. Chem. 456, 83–90. Racz, L., Goel, R.K., 2010. J. Environ. Monit. 12, 58–70. Rocha, M.J., Ribeiro, C., Ribeiro, M., 2011. Int. J. Environ. An. Ch., 1191–1205. Rodriguez-Mozaz, S., de Alda, M.J.L., Barcelo, D., 2004. Anal. Chem. 76, 6998–7006. Song, M.J., Lee, S.K., Lee, J.Y., Kim, J.H., Lim, D.S., 2012. J. Electroanal. Chem. 677, 139–144. Sosniak, A.M., Gasser, G., Metzler-Nolte, N., 2009. Org. Biomol. Chem. 7, 4992–5000. Steel, A.B., Herne, T.M., Tarlov, M.J., 1998. Anal. Chem. 70, 4670–4677. Steen, W.A., Han, S.W., Yu, Q., Gordon, R.A., Cross, J.O., Stern, E.A., et al., 2002. Langmuir 18, 7714–7721. Tran, D.T., Vermeeren, V., Grieten, L., Wenmackers, S., Wagner, P., Pollet, J., Janssen,
309
K.P., Michiels, L., Lammertyn, J., 2011. Biosens. Bioelectron. 26, 2987–2993. Tsakalof, A.K., Gkagtzis, D.C., Koukoulis, G.N., Hadjichristodoulou, C.S., 2012. Anal. Chim. Acta 709, 73–80. Vajda, A.M., Barber, L.B., Gray, J.L., Lopez, E.M., Woodling, J.D., Norris, D.O., 2008. Environ. Sci. Technol. 42, 3407–3414. Vrabel, M., Horakova, P., Pivonkova, H., Kalachova, L., Cernocka, H., Cahova, H., Pohl, R., Sebest, P., Havran, L., Hocek, M., Fojta, M., 2009. Chem. Eur. J. 15, 1144–1154. Wang, S., Li, Y., Ding, M., Wu, X., Xu, J., Wang, R., Wen, T., Huang, W., Zhou, P., Ma, K., Zhou, X., Du, S., 2011. J. Chromatogr. B 879, 2595–2600. Writer, J.H., Ryan, J.N., Keefe, S.H., Barber, L.B., 2012. Environ. Sci. Technol. 46, 860–868. Yildirim, N., Long, F., Gao, C., He, M., Shi, H.C., Gu, A.Z., 2012. Environ. Sci. Technol. 46, 3288–3294. Yoon, Y., Westerhoff, P., Snyder, S.A., Esparza, M., 2003. Water Res. 37, 3530–3537. Zhang, Q., Wang, Y., Mateescu, A., Sergelen, K., Kibrom, A., Jonas, U., Wei, T., Dostalek, J., 2013. Talanta 104, 149–154.
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Biosensors and Bioelectronics journal homepage: www.elsevier.com/locate/bios
A simple and label-free aptasensor based on nickel hexacyanoferrate nanoparticles as signal probe for highly sensitive detection of 17β-estradiol Lifang Fan a,b, Guohua Zhao a,n, Huijie Shi a, Meichun Liu a a b
Department of Chemistry, Tongji University, 1239 Siping Road, 200092 Shanghai, China College of Chemistry and Chemical Engineering, Shanxi Datong University, Datong 037009, China
art ic l e i nf o
a b s t r a c t
Article history: Received 21 October 2014 Received in revised form 17 December 2014 Accepted 7 January 2015 Available online 8 January 2015
A simple and label-free electrochemical aptasensor was developed for detecting 17β-estradiol (E2). To translate the binding events between aptamer and E2 into the measurable electrochemical signal, the nickel hexacyanoferrate nanoparticles (NiHCF NPs) as signal probe was in situ introduced on the electrode by a simple two-step deposition method, exhibiting well-defined peaks with good stability and reproducibility. Subsequently, Au nanoparticles (Au NPs) was covered on the NiHCF NPs, which not only provided a platform for immobilizing the aptamer by S–Au interaction, but further enhanced the conductivity and stability of the signal probe. With the addition of E2, the formation of E2–aptamer complexes on the sensing interface retarded the interfacial electron transfer reaction of the probe, resulting in the decrease of the electrochemical signal. E2 could be readily examined by measuring the signal change. A linear range of 1 10 12–6 10 10 M was obtained with a low detection limit of 0.8 10 12 M. The aptasensor also exhibited high specificity to E2 in control experiments employing seven endocrine disrupting compounds as the interferents that had similar structure or coexisted with E2 in the environment. Besides, the applicability of the aptasensor was successfully evaluated by determining E2 in the real samples. & 2015 Elsevier B.V. All rights reserved.
Keywords: Aptasensor 17β-estradiol Nickel hexacyanoferrate nanoparticles
1. Introduction Endocrine disrupting chemicals (EDCs) are a class of ubiquitous contaminants in the environment, which seriously interfere with the normal endocrine function, further causing adverse effects on growth, metabolism and reproduction of the organisms, and increasing the incidence of cancer and tumors. (Lust et al., 2012; Racz and Goel, 2010; Vajda et al., 2008). Among the EDCs, 17βestradiol (E2) excreted by humans and animals, is a natural and most active estrogen, which can enter surface waters via discharge of municipal wastewater and industrial facility effluents, and discharge from animal feeding operations and septic system, etc. (Writer et al., 2012). The chronic exposure of human to E2, even at very low concentration will cause the drastic health problems (Wang et al., 2011). Therefore, it is necessary to establish a simple, rapid and sensitive method for the detection of E2 in the environment. So far, these methods, such as HPLC (Mahmoud et al., 2011; n
Corresponding author. Fax: þ 86 21 65982287. E-mail address: [email protected] (G. Zhao).
http://dx.doi.org/10.1016/j.bios.2015.01.015 0956-5663/& 2015 Elsevier B.V. All rights reserved.
Mishra and Joy, 2006; Yoon et al., 2003) or GC/MS (Rocha et al., 2011; Tsakalof et al., 2012) have been reported for detecting E2. These conventional methods are accurate and sensitive, but the analytical procedures are complicated and time-consuming. Considering these limitations, some biosensors based on human estrogen receptor (hER) and antibodies have been developed (Le Blanc et al., 2009; Liu et al., 2012, 2010). Although convenient and rapid, but the hER-based biosensors are lacking in selectivity for E2 detection because of the affinity of hER to other xenoendocrinesthes, whereas the immunosensors are better in the selectivity, but they are susceptible to harsh conditions, and the extraction of antibodies is complicated and laborious. Aptamers offer a new alternative to antibodies as biorecognition molecules, meanwhile, they have more advantages over antibodies, such as smaller size, easier synthesis and modification, and better stability. The aptamer-based biosensors have been reported via employing different techniques, such as fluorescence (Yildirim et al., 2012), surface plasmon resonance (Zhang et al., 2013), colorimetry (Kim et al., 2011) and electrochemistry (Lin et al., 2012). Among these aptasensors, electrochemical aptasensors have been widely concerned due to its outstanding advantages, including simple equipment,
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fast response, low cost and real-time detection. However, how to translate the binding events between aptamer and target into a measurable electrochemical signal is the key point in the development of electrochemical aptasensors. Generally, additional electroactive tags, for example, ferrocene derivatives (Song et al., 2012; Sosniak et al., 2009), methylene blue (Ferapontova and Gothelf, 2009; Liu et al., 2011), and ruthenium complexes (Vrabel et al., 2009), etc. have been used to label aptamer to obtain the measurable signal. But labeling aptamer is a complex and time-consuming process, even which might affect the affinity of aptamer toward target. Besides, electroactive compounds can also be directly added into test solution to generate a measurable signal. For instance, the aptasensors based on electrochemical impedance spectroscopy (EIS) by employing [Fe(CN)6]3 /4 as the indicator are commonly used owing to its simple and rapid response with high sensitivity (Gonzalez-Fernandez et al., 2011; Kashefi-Kheyrabadi and Mehrgardi, 2012; Tran et al., 2011). However, according to the literature reported (Bogomolova et al., 2009) and our previous works (Fan et al., 2013; Ke et al., 2014), the disadvantages of EIS aptasensors cannot be neglected. They are usually prone to false-positive results produced by the limitations, including initial sensing interface contamination, excess measurements on the same sensor as well as additional incubations in the [Fe(CN)6]3 / 4 between measurements for a long time (Bogomolova et al., 2009). So it is difficult to determine accurately targets except for performing vast and timeconsuming control or parallel experiments. In this work, in order to overcome effectively these drawbacks of labeling aptamer or adding additional probe into test system, a signal probe, the nickel hexacyanoferrate nanoparticles (NiHCF NPs) was chosen and in situ designed on the electrode due to its outstanding properties, such as easy synthesis, wonderful stability and good peak shape. Then Au nanoparticles (Au NPs) was deposited uniformly on the NiHCF NPs to act as the matrix for anchoring the aptamer and improve the conductivity of the electrode. Thus, a simple and label-free electrochemical aptasensor was fabricated for E2 detection. When it was exposed to E2 solution, the formation of the E2–aptamer complexes on the sensing interface by binding E2 hindered the electron transfer, leading to the decrease of the current. E2 could be quantified via measuring the current change. The sensing performance of the aptasensor for detecting E2 was investigated in detailed. Meanwhile, its practical application was evaluated by detecting the real samples.
2. Experiment 2.1. Reagents Aptamer, was chosen according to the prior literature (Kim et al., 2007). The oligonucleotides were purchased from Sangon Biotechnology Co. Ltd. (Shanghai, China).
ethanol, pH 8.0) was prepared and stored at 4 °C. All other chemical materials were of analytical reagent grade. 2.2. Apparatus Electrochemical measurements were performed with CHI 660C electrochemical workstation (CHI, USA) based on a conventional three-electrode system with a bare or modified gold electrode (d ¼2 mm) acted as the working electrode, a saturated calomel electrode (SCE) as the reference electrode and a platinum wire as the counter electrode. The morphology and energy dispersive X-ray spectroscopy (EDS) spectra of different modified electrodes were measured by scanning electron microscopy (FE-SEM, S-4800, Hitachi). 2.3. Preparation of the signal probe-modified electrode The gold electrode was polished by using 1.0, 0.3 and 0.05 μm alumina powers, respectively. The polished electrode was placed in the freshly Piranha solution for 5 min (Caution!), followed by ultrasonically cleaned. It was pretreated in 0.5 M H2SO4 by potential scanning between 0.2 and 1.5 V until a reproducible cyclic voltammogram was obtained. The cleaned electrode was put into 50 mM NiCl2 6H2Oþ 10 mM NH4Cl of pH 4.55, where metallic Ni was electrodeposited on the electrode at the potential of 0.9 V. The Ni-deposited electrode was transferred into 0.1 M NaNO3 and 5 mM K3[Fe(CN)6] and scanned by I–t technique at 1.0 V. The scanning was continued until the current decreased to almost zero to ensure that all of metallic Ni had been converted to NiHCF NPs. Thus, the signal probe-modified electrode was prepared by the two-step deposition method. 2.4. Design and construction of the aptasensor The as-prepared NiHCF NPs-modified electrode was immersed in 2.5 mM HAuCl4 containing 0.1 M KCl and 50 mM H2SO4, where Au NPs was deposited on the NiHCF NPs by cyclic voltammetry (CV) at N2 atmosphere (Au NPs/NiHCF NPs-modified electrode). Subsequently, it was incubated in 2.0 mM aptamer solution for over 12 h (aptamer/Au NPs/NiHCF NPs-modified electrode). Before incubation, 0.2 mM TCEP was used to reduce the disulfide bond of aptamer. Finally, 1 mM MCH was employed to block the unbound active sites of Au NPs (MCH/aptamer/Au NPs/NiHCF NPs-modified electrode). Any nonspecific binding was inhibited except for the aptamer–E2 interaction. The aptasensor was prepared and stored at 4 °C. Scheme 1 illustrates the schematic representation of the aptasensor construction and mechanism for detecting E2. To reuse the aptasensor, it was incubated in a 0.5% sodium dodecyl sulfate (SDS) solution (pH 1.9) for 3 min and washed with PBS (Yildirim et al., 2012). 2.5. Electrochemical measurements
Aptamer sequence The EIS for monitoring the different stages of the aptasensor 5′-SH-(CH2)6-GCT-TCC-AGC-TTA-TTG-AAT-TAC-ACG-CAG-AGG-GTA- construction was performed in 0.1 M PBS (pH 7.41) containing GCG-GCT-CTG-CGC-ATT-CAA-TTG-CTG-CGC-GCT-GAA-GCG-CGG- 5.0 mM K3[Fe(CN)6]/K4[Fe(CN)6] and 0.1 M KCl. EIS measurements AAG-C-3, were recorded between 1000 Hz and 0.1 Hz with a sinusoidal Random DNA sequence voltage perturbation of 5 mV amplitude. In the detection of E2, 5′-(SH)-(CH2)6-CTG-ACA-CCA-TAT-TAT-GAA-GA-3′ 20 μL different concentrations of E2 were dropped onto the sensing interface for 40 min at room temperature, following by E2 (99.9%), Hydrogen tetrachloroaurate (III) hydrate (HAuCl4) thoroughly rinsed to remove unbound E2. Before and after the binding of E2, the current of the aptasensor was measured by and Tris (2-carboxyethy1) phosphine (TCEP) were purchased from differential pulse voltammetry (DPV) in 0.1 M PBS (pH 7.41) as the Sigma-Aldrich (St. Louis, MO). 6-Mercap-1-hexanol (MCH) was supporting electrolyte. obtained from Adamas Reagent Co. Ltd. DNA sequences were dissolved in Tris–EDTA buffer (TE, pH 8.0). Tris–HCl buffer The two real samples, municipal wastewater, were collected from two living areas in Yangpu District, Shanghai. Prior to (100 mM Tris–HCl/200 mM NaCl/25 mM KCl/10 mM MgCl2, 5%
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Scheme 1. Illustration of the electrochemical aptasensor construction and the mechanism for detection of E2.
analysis, the samples were filtered through an ordinary filter paper and the membrane with the size of 0.22 μm, respectively, to remove the suspended solid matter and small particles. The substances dissolved in these samples were not changed by the pretreatment. The treated samples were detected as soon as possible to minimize microbial degradation. Furthermore, the samples were diluted to decrease the interference of complex matrices.
3. Results and discussion 3.1. Construction and properties of the signal probe The argenteous metallic Ni uniformly deposited on the gold
was oxidized to Ni2 þ and subsequently reacted with the hexacyanoferrate (III), forming the signal probe, NiHCF NPs. NiHCNFe NPs, similar to other nickel hexacyanoferrate, is an inorganic polymer analog from Prussian blue family (Steen et al., 2002). NiHCNFe complexes have two different types (NiHCNFe-II and NiHCNFe-III), which can be formed on the electrode together or alone when changing the preparation conditions (Hao et al., 2012). Here, NiHCF NPs-III was directly formed on the electrode by chemical reaction between Ni2 þ and [Fe(CN)6]3 in the deposition potential of 1.0 V and its molecular compositions is Ni3[Fe(CN)6]2 (Chen et al., 2009). Fig. 1A shows the SEM morphology of the NiHCF NPs, it is uniformly covered on the electrode, with the diameter of about 10 nm. The thickness of the probe depended on the amount of
Fig. 1. (A) SEM of the signal probe and the inset is SEM of the Au NPs/NiHCF NPs-modified electrode; (B) the EDS spectrum of the Au NPs/NiHCF NPs-modified electrode and the inset is the elemental mapping of Au (gray), Ni (green), Fe (red), C (purple) and N (dark red) with color superposition in the electrode; (C) CV curves of the signal probe at different scan rate (from a to h: 20, 40, 60, 80, 100, 120, 160, 200 mV/s); The inset is the dependence of the redox peak current on the scan rate and (D) CV curves of the signal probe in 0.1 M PBS (pH 7.41), Scan cycles: 30. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article).
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deposited Ni which could be controlled by the factors, including the concentration of NiCl2 and the deposition time. In order to achieve the excellent properties of the probe, the two factors were optimized in detailed and given in Figs. S1 and S2 in Supplementary information. The results demonstrate that the optimized concentration of NiCl2 is 50 mM, and the optimized deposition time is 60 s. Fig. 1C shows the CV curves of the probe at the different scan rate. It can be observed that a pair of well-defined redox peaks are present in the absence of any electroactive substance. The Ni2 þ in the NiHCF NPs cannot be reduced in the potential range of 0.0– 0.7 V, while the redox peaks correspond to the hexacyanoferrate (III:II) redox couple (Lin and Bocarsly, 1991) due to the formation of NiHCNFe-(II) on the electrode by electrochemical reduction of [Fe(CN)6]3 to [Fe(CN)6]4 and in turn coordination to Ni2 þ in the potential cycling scans. The peak width at half-height is approximately 96 mV, which is consistent with the literatures (Cai et al., 1995; Pournaghi-Azar and Razmi-Nerbin, 1998), approaching the theoretical value of a one-electron reaction mechanism (90.6/ nm V) (Bard and Faulkner, 1980). The relationship of the peak current with the different scan rate is shown in the inset of Fig. 1C. The anodic and cathodic current increase linearly with the scan rate range from 20 to 200 mV/s, as predicted for a diffusionless system. The peak separation ΔEp (ΔEp ¼ Ea Ec) is about 16 mV at the scan rate of 20 mV/s, and ΔEp is further enhanced with increasing the scan rate, for instance, 39 mV of ΔEp occurs approximately at 100 mV/s. For a fully reversible reaction in which no reactants from the solution take place, the separation of peaks should be zero (Cai et al., 1995). The relatively small ΔEp indicates fairly fast electrochemical reaction rate on the NiHCF NPs-modified electrode (Laviron, 1979a). Meanwhile, the surface coverage (Γ) of the NiHCF NPs on the electrode could be evaluated for 5.761 10 8 mol cm 2 by the equation Γ = Q /nFA (Laviron, 1979b). The detailed calculation for Γ was given in the supporting information. Besides, the stability of the signal probe was investigated by directly exposing it to air for a week or putting in 0.1 M PBS for three days, then recording the CV response. The results indicated that its stability was not affected by air. Furthermore, there was not any loss of its electroactivity after storing in 0.1 M PBS. The reproducibility of the probe was also measured, shown in Fig. 1D. The height and the ΔEp of the redox peaks were almost no change after 30 cycles of repetitive CV scanning, indicating that the signal probe had good reproducibility. Afterwards, Au NPs was deposited on the probe to act as a platform for immobilizing aptamer and further improve the conductivity and stability of the electrode. The SEM of the Au NPs/ NiHCF NPs-modified electrode is given in the inset of Fig. 1A. Au
NPs is uniformly dispersed on the probe, with the diameter of 50– 100 nm. The EDS spectrum in Fig. 1B demonstrates that the elements including Au, Ni, Fe, C and N are present in the Au NPs/ NiHCF NPs-modified electrode. Furthermore, the elemental mapping confirms that the five elements are highly dispersed on this modified electrode. In addition, the EDS element analysis shows that the atom percent of Ni and Fe is 4.88% and 3.27%, respectively. The atom ratio of Ni and Fe is 1.49:1, confirming that Ni3[Fe(CN)6]2 complex is present in the NiHCF NPs when Ni (II) reacts with [Fe(CN)6]3 . This result is agree with that of the literatures reported. (Chen et al., 2009; Carpani et al., 2006). 3.2. Design and construction of the aptasensor In order to characterize the immobilization results, the different stages of the aptasensor construction were investigated by CV and DPV. The CV curves are shown in Fig. 2A, there is no peak on the bare gold electrode and the current is almost zero in the absence of any electroactive substance (curve a). After depositing the signal probe on the electrode, there is a couple of well-defined redox peaks (curve b), corresponding to the hexacyanoferrate (III:II) redox couple, indicating that the probe has been successfully introduced on the electrode. Furthermore, the ΔEp of the peaks is about 40.0 mV and the ratio of the anodic and cathodic peak current (ia:ic) equals to one, revealing good reversibility of the probe. After depositing Au NPs, the current of the redox peaks decreases somewhat (curve c), owing to the decrease of some electroactive sites of the probe encased by Au NPs. The ΔEp decreases to 38.0 mV, implying that the conductivity of the modified electrode is improved. When aptamer was immobilized on the Au NPs/NiHCF NPs-modified electrode, the current of the redox peak decreased and had poor reversibility (curve d). The reason can be attributed to that the formation of a selfassembled monolayer of aptamer retard the electron transfer reaction of the signal probe. Finally, the aptamer-modified electrode was treated with MCH, and the current further decreased (curve e). Meanwhile, the DPV curves for characterization of the aptasensor construction are shown in Fig. 2B. There is nearly a line on the bare gold electrode (curve a). While the NiHCF NPs was electrodeposited, a large current of 146.9 μA (curve b) was recorded, implying the probe had good electroactivity. After depositing Au NPs, the current decreases to 131.2 μA, and the peak potential moves negatively (curve c). This result demonstrates that the electroactive sites of NiHCF NPs decrease due to the coating of Au NPs, which is consistent with the result of the CV measurements. After immobilizing aptamer, the current further decreases to 85.1 μA. Finally, the aptamer-modified electrode was blocked using MCH and the current decreased to 78.8 μA. Besides, EIS
Fig. 2. CV (A) and DPV (B) curves of the bare gold electrode (a), the NiHCF NPs-modified electrode (b), the Au NPs/NiHCF NPs-modified electrode (c), the aptamer/Au NPs/ NiHCF NPs-modified electrode (d) and the MCH/aptamer/Au NPs/NiHCF NPs-modified electrode (e) in 0.1 M PBS (pH 7.41).
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measurements were also carried out in the presence of [Fe(CN)6]3 /4 (Fig. S3) to evaluate the interfacial electrochemistry properties in the construction of the aptasensor. The characterization results of EIS totally agreed with those of CV and DPV, confirming that the aptasensor based on the NiHCF NPs as the signal probe was successfully fabricated. In addition, the load of aptamer on the sensing interface has great impact on the sensitivity of the aptasensor. Herein, the coverage density (Γ) of aptamer anchored was estimated by chronocoulometry employing 0.8 mM [Ru(NH3)6]3 þ as a cationic redox marker which bound electrostatically to the anionic phosphate groups of aptamer (Steel et al., 1998). Γ was calculated to be 7.67 1012 molecules cm 2. The control experiments were also performed utilizing the bare gold electrode as the matrix for modifying aptamer. The load of aptamer on the gold electrode was about 1.24 1012 molecules cm 2 (The detailed calculation was given in Fig. S4). The coverage density of aptamer on the Au NPs/ NiHCF NPs-modified electrode is almost six-fold more than that on the gold electrode. So Au NPs with the large surface area is more advantageous to load aptamer, which greatly improve the sensitivity of the aptasensor. 3.3. Analytical performance of the electrochemical aptasensor The analytical performance of the aptasensor was evaluated in different concentrations of E2 solution by DPV, shown in Fig. 3A. The current of the aptasensor decreases with increasing concentrations of E2. This is mainly attributed to the fact that more and more aptamer–E2 complexes formed on the sensing interface by capturing vast E2 hinder the electron transfer, resulting in the decrease of the current. Fig. 3B shows the dependence of the current change ΔI and different E2 concentrations. It is apparent that ΔI reach a plateau at E2 concentration of more than 600 pM, indicating that the amount of E2–aptamer complexes on the sensing interface reaches their saturation. A relationship curve between ΔI/I0 on the vertical ordinate and the logarithm of E2 concentrations on the horizontal axis is plotted, as is shown in the inset of Fig. 3B. It can be observed that ΔI/I0 is linear against the logarithm value of E2 concentrations and a linear dynamic range from 5.0 to 600.0 pM is obtained. The regression equation is ΔI/I0 ¼0.2894 log Cþ 0.1213 (unit of C, pM) with a correction coefficient of 0.9935. The limit of detection is estimated to be 0.8 pM based on the response of three times the standard deviation of zero-pose response (n ¼5) and the limit of quantitation is 5.0 pM. Some detailed comparisons of the analytical behavior of the aptasensor with other published methods for detecting E2 were given in Table S1. It can be seen that the
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sensitivity of the immunosensor or impedimetric aptasensor for E2 detection (Liu et al., 2010; Lin et al., 2012) is lower than that of the present aptasensor. Although high sensitivity was also achieved by HPLC (Farre et al., 2007; Rodriguez-Mozaz et al., 2004), compared to these conventional methods, the analysis progress by the present aptasensor is simpler, rapider and more low-cost without complicate sample pretreatments and expensive instruments. It is thus evident that the aptasensor can be applied to detect low level of E2 in the environment. In addition, the experimental results demonstrated that high sensitivity of the aptasensor was related with the parameters, such as the amount of Au NPs on the probe, controlled by the numbers of CV cycles in HAuCl4 solution, and the incubation time of the aptasensor in E2 solution. The parameters were optimized in detailed and given in Figs. S5 and S6. When the optimized number of CV cycles for Au NPs is 15 and the incubation time is 40 min, the aptasensor can arrive the best response for E2 detection. To investigate the selectivity of the aptasensor, seven endocrine disrupting chemicals including ethinylestradiol, bisphenol A, estriol, polychlorinated biphenyl (PCB 101), diethyl phthalate, 4-nonyl phenol and atrazine were used as the interferents, which could have similar structure or coexist with E2 in the environment. The aptasensor was incubated in 5.0 pM of each one among seven interferents, respectively and its current response was measured with the same method for determining E2. The current difference, ΔI obtained before and after incubation in 5.0 pM E2 and the interferent with the same concentration is shown in Fig. 4A. The results illustrate that ΔI is approximately 17 μA in 5.0 pM E2, whereas ΔI in any one of other interferents is less than 2.0 μA. The current response of the aptasensor for these interferents is less than 8.0% for that for E2, indicating high selectivity and specificity of the aptasensor to E2. Control experiments were performed with a random DNA sequence as the recognition element instead of aptamer for determining different concentrations of E2, shown in Fig. 4B. With increasing the concentrations of E2, the peak current is almost unchanged. This is attributed to the fact that the random DNA-based sensor has no recognition ability for E2, so that E2 is not captured on the sensing interface, further confirming that high specificity of the aptasensor toward E2 is closely related to the specific sequence of the aptamer. In addition, the long-term storage stability, the reproducibility and the reusability of the aptasensor were also investigated. When the aptasensor was stored in 0.1 M PBS at 4 °C for three days, it was used to determine 5.0 pM E2. The peak current was lower only 3.2% than the original response. After the storage for a week, the current decreased to 5.3% of the original response, suggesting that the aptasensor had good stability. The reproducibility of the
Fig. 3. (A) DPV response of the aptasensor in different concentrations of E2 (from a to j: 0, 1, 5, 10, 50, 100, 200, 400, 600, 1000 pM) and (B) the dependence of the peak current on the concentrations of E2; the inset is the linear calibration curve of ΔI/I0 with logarithm of E2 concentrations.
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Fig. 4. The selectivity and specificity test of the aptasensor for E2 detection.
Table 1 Detection of the wastewater samples with E2 at different concentrations. Sample
Sample 1
Sample 2
a b
Determined (pM)a
Recovery (%)
RSD (%)b
0.0 5.0 10.0 50.0 0.0
Undetected 4.8 9.8 50.1 Undetected
96.6 98.1 100.2
3.6 2.0 1.1
5.0 10.0 50.0
4.7 9.9 49.8
93.6 98.5 99.6
2.4 2.9 1.1
Spiked (pM)
Mean of three measurements. Relative standard deviation.
aptasensor was evaluated by an intraassay and an interassay relative standard deviation (RSD). The intraassay RSD was 8.0% by detecting 5.0 pM E2 with five replicated measurements. Then, five identical aptasensors were used to determine 5.0 pM E2, and the interasssay RSD was 7.2%, showing that the aptasensor had an acceptable reproducibility. Besides, the reusability of the aptasensor was also important in the practical application. The E2– aptamer complexes on the sensing interface could be effectively separated in 0.5% SDS solution (pH 1.9) for 3 min without any destruction of the binding affinity of aptamer toward E2. After regeneration, the current response as percent of the initial signal is given in Fig. S7. Over 80% of the initial response can be observed after repeated use for three times. 3.4. Application of the electrochemical aptasensor In order to evaluate the feasibility of the aptasensor for practical analysis, it was used to determine E2 in the real samples using the standard addition method. Three different concentrations of the standard solutions, 5.0, 10.0, 50.0 pM were added into the two pretreated samples, respectively. The recoveries results of E2 in the two different samples are shown in Table 1. It can be observed that the average recoveries range from 93.6% to 100.2% with RSD lower than 5.2% based on triplicate experiments at each concentration. The results indicate that the aptasensor can resist the interference of the complex matrices and be applied to determine E2 in different environmental wastewater.
4. Conclusions In summary, the electrochemical aptasensor was successfully developed for the detection of E2 based on the NiHCF NPs as the
in situ signal probe which overcame effectively the drawbacks of labeling aptamer or adding additional probe into test system. The aptasensor showed good analytical performance for E2 detection with high sensitivity, specificity and reproducibility. A low detection limit of 0.8 pM was obtained, which was comparable or even lower than the previous reports. Moreover, the aptasensor was successfully applied to detect the real samples, and the satisfactory recoveries were obtained.
Acknowledgments This work was financially supported by the National Natural Science Foundation of China (NSFC, No. 21277099).
Appendix A. Supplementary material Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.bios.2015.01.015.
References Bard, A.J., Faulkner, L.R., 1980. Electrochemical Methods: Fundamentals and Applications. Wiley, New York. Bogomolova, A., Komarova, E., Reber, K., Gerasimov, T., Yavuz, O., Bhatt, S., Aldissi, M., 2009. Anal. Chem. 81, 3944–3949. Cai, C.X., Ju, H.X., Chen, H.Y., 1995. Anal. Chim. Acta 310, 145–151. Carpani, I., Giorgetti, M., Berrettoni, M., Buldini, P.L., Gazzano, M., Tonelli, D., 2006. J. Solid State Chem. 179, 3981–3988. Chen, W., Tang, J., Xia, X.H., 2009. J. Phys. Chem. C 113, 21577–21581. Fan, L.F., Zhao, G.H., Shi, H.J., Liu, M.C., Li, Z.X., 2013. Biosens. Bioelectron. 43, 12–18. Ferapontova, E.E., Gothelf, K.V., 2009. Electroanalysis 21, 1261–1266. Farre, M., Kuster, M., Brix, R., Rubio, F., Lopez de Alda, M.J., Barcelo, D., 2007. J. Chromatogr. A 1160, 166–175. Hao, X., Yan, T., Wang, Z., Liu, S., Liang, Z., Shen, Y., Pritzker, M., 2012. Thin Solid Films 520, 2438–2448. Gonzalez-Fernandez, E., de-los-Santos-Alvarez, N., Lobo-Castanon, M.J., MirandaOrdieres, A.J., Tunon-Blanco, P., 2011. Biosens. Bioelectron. 26, 2354–2360. Kashefi-Kheyrabadi, L., Mehrgardi, M.A., 2012. Biosens. Bioelectron. 33, 184–189. Ke, H., Liu, M., Zhuang, L., Li, Z., Fan, L., Zhao, G., 2014. Electrochim. Acta 137, 146–153. Kim, Y.S., Jung, H.S., Matsuura, T., Lee, H.Y., Kawai, T., Gu, M.B., 2007. Biosens. Bioelectron. 22, 2525–2531. Kim, Y.S., Kim, J.H., Kim, I.A., Lee, S.J., Gu, M.B., 2011. Biosens. Bioelectron. 26, 4058–4063. Laviron, E., 1979a. J. Electroanal. Chem. 101, 19–28. Laviron, E., 1979b. J. Electroanal. Chem. 100, 263–270. Le Blanc, A.F., Albrecht, C., Bonn, T., Fechner, P., Proll, G., Proll, F., Carlquist, M., Gauglitz, G., 2009. Anal. Bioanal. Chem. 395, 1769–1776. Lin, C., Bocarsly, A.B., 1991. J. Electroanal. Chem. 300, 325–345. Lin, Z., Chen, L., Zhang, G., Liu, Q., Qiu, B., Cai, Z., Chen, G., 2012. Analyst 137, 819–822. Liu, S., Lin, Q., Zhang, X., He, X., Xing, X., Lian, W., Li, J., Cui, M., Huang, J., 2012. Sens.
L. Fan et al. / Biosensors and Bioelectronics 68 (2015) 303–309
Actuators B-Chem. 166-167, 562–568. Liu, X., Duckworth, P.A., Wong, D.K., 2010. Biosens. Bioelectron. 25, 1467–1473. Liu, Y., Yan, J., Howland, M.C., Kwa, T., Revzin, A., 2011. Anal. Chem. 83, 8286–8292. Lust, M., Makinia, J., Stensel, H.D., 2012. Water Sci. Technol. 65, 1130–1136. Mahmoud, I.Y., AlKindi, A.Y., Khan, T., Al-Bahry, S.N., 2011. J. Exp. Zool. Part A 315A, 170–174. Mishra, A., Joy, K.P., 2006. Gen. Comp. Endocrinol. 145, 84–91. Pournaghi-Azar, M.H., Razmi-Nerbin, H., 1998. J. Electroanal. Chem. 456, 83–90. Racz, L., Goel, R.K., 2010. J. Environ. Monit. 12, 58–70. Rocha, M.J., Ribeiro, C., Ribeiro, M., 2011. Int. J. Environ. An. Ch., 1191–1205. Rodriguez-Mozaz, S., de Alda, M.J.L., Barcelo, D., 2004. Anal. Chem. 76, 6998–7006. Song, M.J., Lee, S.K., Lee, J.Y., Kim, J.H., Lim, D.S., 2012. J. Electroanal. Chem. 677, 139–144. Sosniak, A.M., Gasser, G., Metzler-Nolte, N., 2009. Org. Biomol. Chem. 7, 4992–5000. Steel, A.B., Herne, T.M., Tarlov, M.J., 1998. Anal. Chem. 70, 4670–4677. Steen, W.A., Han, S.W., Yu, Q., Gordon, R.A., Cross, J.O., Stern, E.A., et al., 2002. Langmuir 18, 7714–7721. Tran, D.T., Vermeeren, V., Grieten, L., Wenmackers, S., Wagner, P., Pollet, J., Janssen,
309
K.P., Michiels, L., Lammertyn, J., 2011. Biosens. Bioelectron. 26, 2987–2993. Tsakalof, A.K., Gkagtzis, D.C., Koukoulis, G.N., Hadjichristodoulou, C.S., 2012. Anal. Chim. Acta 709, 73–80. Vajda, A.M., Barber, L.B., Gray, J.L., Lopez, E.M., Woodling, J.D., Norris, D.O., 2008. Environ. Sci. Technol. 42, 3407–3414. Vrabel, M., Horakova, P., Pivonkova, H., Kalachova, L., Cernocka, H., Cahova, H., Pohl, R., Sebest, P., Havran, L., Hocek, M., Fojta, M., 2009. Chem. Eur. J. 15, 1144–1154. Wang, S., Li, Y., Ding, M., Wu, X., Xu, J., Wang, R., Wen, T., Huang, W., Zhou, P., Ma, K., Zhou, X., Du, S., 2011. J. Chromatogr. B 879, 2595–2600. Writer, J.H., Ryan, J.N., Keefe, S.H., Barber, L.B., 2012. Environ. Sci. Technol. 46, 860–868. Yildirim, N., Long, F., Gao, C., He, M., Shi, H.C., Gu, A.Z., 2012. Environ. Sci. Technol. 46, 3288–3294. Yoon, Y., Westerhoff, P., Snyder, S.A., Esparza, M., 2003. Water Res. 37, 3530–3537. Zhang, Q., Wang, Y., Mateescu, A., Sergelen, K., Kibrom, A., Jonas, U., Wei, T., Dostalek, J., 2013. Talanta 104, 149–154.
