Biosensors and Bioelectronics 58 (2014) 48–56

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Homogeneously ultrasensitive electrochemical detection of adenosine triphosphate based on multiple signal amplification strategy Xiaojun Chen a,n, Lingna Ge a, Buhua Guo a, Ming Yan b, Ning Hao b, Lin Xu b,n a b

College of Sciences, Nanjing Tech University, Nanjing 211816, PR China Biotechnology and Pharmaceutical Engineering, Nanjing Tech University, Nanjing 211816, PR China

art ic l e i nf o

a b s t r a c t

Article history: Received 23 December 2013 Accepted 6 February 2014 Available online 28 February 2014

An ultrasensitive electrochemical aptasensor was successfully fabricated for the detection of adenosine triphosphate (ATP). For the first time, one detection system combined several elements: magnetic aptamer sequences for target recognition and separation, a DNAzyme assisted cyclic signal amplification strategy, layer-by-layer (LBL) quantum dots (QDs) composites for promoting square wave anodic stripping voltammetric (SWASV) analysis and Bi, Nafion (Nf) and three-dimensional ordered macroporous polyaniline-ionic liquid (Bi/Nf/3DOM PANI-IL) film modified glassy carbon electrode (GCE) for monitoring enhanced SWASV signal. The modification of Nf/3DOM PANI-IL on GCE showed that the preconcentration efficiency was improved by the electrostatic absorption of Cd2 þ with negative Nf layer with the enhanced analytical sensitivity due to a large active surface area of 3DOM structure. The increased SWASV peak current values of the label (CdS)4@SiO2 composites were found to be proportional to the logarithmic value of ATP concentrations in the range of 1 pM–10 nM and 10 nM–1 mM, with the detection limit as low as 0.5 pM. The proposed aptasensor has shown an excellent performance such as high sensitivity, good selectivity and analytical application in real samples. The results demonstrated that the multiple signal amplified strategy we developed was feasible for clinical ATP assay and would provide a promising model for the detection of other small molecules. & 2014 Elsevier B.V. All rights reserved.

Keywords: Electrochemical aptasensor Layer-by-layer quantum dots Three-dimensional ordered macroporous Au Three-dimensional ordered macroporous polyaniline-ionic liquid composite DNAzyme cyclic amplification Square wave anodic stripping voltammetry

1. Introduction Small organic molecules in body fluids or tissues, with a molecular weight of less than 1000 Da, are important analytes of various molecular and cellular researches, as well as clinical diagnostics (D’Orazio, 2011). For example, adenosine-50 -triphosphate (ATP, molecular weight of 507.2 Da) is one of the most important metabolites in biological systems. It is not only the energy source for biological reactions, but also an extracellular signaling agent in biological processes such as photosynthesis, enzyme catalysis, biosynthesis, DNA replication, and cellular respiration (Knowles, 1980; Brandon et al., 2006). Furthermore, ATP also serves as a marker for evaluating micro-fungal contamination in food industry (Davidson et al., 1999). Therefore, the determination of ATP has become very important. Many techniques have been developed to detect ATP, including chromatography (Stratford, Dennis (1994)), bioluminescence and chemiluminescence (Ribeiro et al., 1998). Recently, new designs of ATP detection have emerged based on the advancement of analytical technologies employing aptamers.

n

Corresponding authors. Tel./fax: þ86 25 858139527. E-mail addresses: [email protected] (X. Chen), [email protected] (L. Xu).

http://dx.doi.org/10.1016/j.bios.2014.02.043 0956-5663 & 2014 Elsevier B.V. All rights reserved.

Aptamers are in vitro selected functional single-stranded DNAs, RNAs or even chemically modified nucleic acids, which could fold into special structures and possess high recognition ability to specific targets ranging from metal ions, organic and inorganic small molecule, proteins and even whole cells (Li et al., 2013). In particularly, aptamer provides excellent advantages of reversible thermal denaturation and unlimited shelf life (Xu et al., 2013). A variety of aptamer sensors (aptasensors) based on electrochemistry (Chen et al., 2013; Wang et al., 2012), fluorescence (Tedsana et al., 2013), luminescence (Lu et al., 2013), surface plasmon resonance (SPR) (Liu et al., 2012), colorimetry (Guo et al., 2011) and quartz crystal microbalance (QCM) (Ruslinda et al., 2012) has been developed for the detection of ATP. Among these aptasensors, electrochemical aptasensors are the most attractive due to their advantages of fast response, portability, high sensitivity, simple instrumentation, low cost (Xiao et al., 2005; Du et al., 2011; Yin et al., 2012). In order to improve the sensitivity of ATP electrochemical aptasensors, many signal amplification strategies generally rely on the transduction technology and the signal-reporting pattern (Lubin and Plaxco, 2010). Nanomaterial assisted amplification is an effective route and has attracted considerable attention for the construction of ultrasensitive aptasensors. Among various nanomaterials, three dimensional ordered macroporous (3DOM) Au is

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flavored in new generation of aptasensors, due to its good biocompatibility with immobilized biomolecules and large specific surface area for charge and mass transportation in electrochemistry (Walcarius, 2012). Zhu constructed an electrochemical aptasensor based on 3DOM Au for the detection of ATP, with the detection limit of 0.01 nM (Zhou et al., 2010). Quantum dots (QDs) are nanostructured semiconductor materials with a size typically between 1 and 12 nm that provides unique optical and electronic properties due to the quantum confinement effect (Weller, 1993). Since the first studies applying QDs as electrochemical label were published by Wang et al. (2002), QDs have gained interests for the sensitive detections in electrochemical bioassays (Wang et al., 2003a, 2003b). Typically, this procedure consists of dissolving the QDs by acid attack for releasing metal ions, which can be easily determined by square wave anodic stripping voltammetry (SWASV) with a mercury film glassy carbon electrode. Significantly, acid dissolution of QDs tags releases numerous metal ions, indicating the built-in amplification nature of the QDs labels (Hansen et al., 2006a, 2006b). However, the conventional SWASV method has a major limitation for wider application because of the requirement to use, manipulate and dispose metallic mercury or mercury salts. Since the year 2000, bismuth-film electrode (BiFE) has become an attractive subject in heavy metal analysis replacing mercury-film electrode (HgFE), due to its environmental friendly nature (Wang et al., 2000). Recently, electroanalysis of trace heavy metal ions at the conducting polymer modified electrodes has received considerable attention, as the polymer can enrich the target heavy metals and improve the analytical performance (Imisides et al., 1991). In this study, Bi/Nafion (Nf)/3DOM polyanline (PANI)-ionic liquid (IL) composite electrode with enlarged surface area, good stability and conductivity was achieved to enhance SWASV electrochemical signal of CdS QDs for the first time. Further, most electrochemical aptasensors are based on heterogeneous assays involving the immobilization of the aptamers on the electrode surface prior to collecting the target recognition induced electrochemical signal. In this way, the recognition event of aptamer to analyte occurs on the interface between the solution and electrode, and the configurational freedom of the immobilized aptamer is often restricted owing to the steric hindrance of the electrode surface, accompanied by the relatively low binding efficiency and rate of reaction of aptamer toward the substrate compared with homogeneous assays. Until very recently, Li's group first demonstrated an electrochemical aptamer-based strategy for ATP detection in a homogeneous solution phase, with a detection limit of 1 nM (Lu et al., 2013). Herein, we proposed another homogeneous electrochemical bioassay for the detection of ATP combining target-induced recycling DNAzyme catalysis and SWASV signal amplification. As shown in Scheme 1, the novel bioassay involves three steps (1) the duplex of S1 and 50 -thiol (SH) modified ATP aptamer was chemi-absorbed onto the surface of Au–SiO2@Fe3O4. In the presence of target ATP, the aptamer formed stable tertiary structure with ATP, which accordingly denatured the duplex and liberated the complementary S1; (2) molecule beacon (MB) molecules were immobilized onto the surface of Au–SiO2@Fe3O4 by S–Au interaction. The MB–Au–SiO2@Fe3O4 composites were added into the S1 solution of step 1, and then double stranded DNA modified Au–SiO2@Fe3O4 particles were obtained by hybridization of MB with its partially complementary S1. After Zn2 þ ions triggered the catalytic reaction with MB at the scissile rA, T1 fragments of MB dissociated from S1/MB–Au–SiO2@Fe3O4. The released S1 hybridized to another MB to trigger a series of DNAzyme catalyzed scission reactions under the cofactor Zn2 þ , and another batch of T1 fragments was released, which then recycled to the headstream; (3) The multi-layered (CdS)4@SiO2 nanocomposite was

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synthesized by firstly the covalent combination between aminoterminated SiO2 particles and carboxyl-stabilized CdS QDs, and then polyelectrolyte-attended electrostatic self-assembling between neighboring QDs. Single stranded S2 could be combined with (CdS)4@SiO2 via the acylamide binding in the presence of EDC as the activator, forming S2-(CdS)4@SiO2. Further, 50 –SH terminal S3 was immobilized onto the surface of 3DOM Au film modified electrode. In the presence of T1, S2-(CdS)4@SiO2 gained the chance of hybridization with S3 on the electrode surface. Subsequently, (CdS)4@SiO2 was dissolved with HNO3 for SWASV analysis. Therefore, the target ATP induced strand displacement was successfully monitored and ATP detection could be realized through the electrochemical signal using electrodeposited Bi/Nf/3DOM PANI-IL modified electrode.

2. Experimental 2.1. Chemicals and materials N-(3-dimethylamminopropyl)-N0 -ethylcarbodiimide hydrochloride (EDC), N-hydroxysulfosuccinimid sodium salt (NHS), sodium oleate (SO) was purchased from Aladdin chemistry Co. Ltd. (Shanghai, China). CdCl2  2.5H2O, Bi(NO3)3  5H2O, Zn(NO3)2  6H2O, FeSO4  7H2O, FeCl3  6H2O, sodium dodecyl benzene sulfonate (SDBS), tetraethoxysilane (TEOS, 98%) and Na2S  9H2O were all obtained from Shanghai Lingfeng chemical reagent Co. LTD (Shanghai, China). Thioglycolic acid (TGA) and poly(diallyldimethylammonium chloride) (PDDA, 20 wt%) were purchased from Aldrich. 1-Butyl-3-methylimidazolium tetrafluoroborate ([BMIm][BF4], purity 499%) was purchased from Lanzhou Institute of Chemical Physics, Chinese Academy of Sciences. Nafion (Nf) was obtained from DuPont as 5 wt% solution and diluted to 0.5% in water. Aniline (AN) was distilled twice under reduced pressure and stored in dark at low temperature before use. 0.1 M phosphate buffer saline (PBS) was prepared by mixing the stock solutions of KH2PO4 and K2HPO4, and adjusted to appropriate pH by addition of 0.1 M KOH or H3PO4 solution. Adenosine triphosphate (ATP), cytidine triphosphate (CTP), guanosine triphosphate (GTP), uridine triphosphate (UTP), diethyl pyrocarbonate (DEPC) and 6-Mercaptohexanol (MCH) were obtained from Shanghai Baoman Biotech. The SiO2 spheres with the diameter of 500 nm were obtained from Alfa Asear. The amino functionalized SiO2 spheres with the diameter of 20 nm were purchased from Haitai NANO Co. Ltd. (Nanjing, China). Au coated glass substrate was provided by Shanghai Institute of Microsystem and Information Technology (SIMIT), Chinese Academy of Sciences (CAS). The DNA sequences were ordered from Shanghai Sangon Engineering Technology and Services Co., Ltd., as follows: DNA

Sequence (from 50 to 30 )

ATP aptamer

SH-(CH2)6-AAAAAATGGAAGGAGGCGTTATGAGGGGGTCCACGCCAACTATTTCG CATCTCTTCTCCGAGCCGGTCGAAATAGTGGGTG NH2-(CH2)6-T10-CCACCACCGCCTC SH-(CH2)6-T10-CCTTCTCTACAATG SH-(CH2)6-CCACCACATTCAAATTCACCAACTATrAGGAAGAGATGTTACGAGGCGGTGGTG

S1 S2 S3 MB

S1, S2, S3 and MB stock solutions were all prepared in 25 mM pH 8.0 Tris–HAc solution containing 300 mM NaCl. ATP aptamer and ATP stock solution were prepared in 25 mM pH 8.0 Tris–HAc solution containing 300 mM NaCl and 5 mM MgCl2. The DNA strands hybridization and aging solution was 25 mM pH 8.0 Tris–HAc solution containing 300 mM NaCl. 5 mM MCH solution was prepared in 10 mM pH 7.4 Tris–HAc solution. In order to

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Scheme 1. Schematic illustration of electrochemical detection of ATP based on multiple signal amplification strategy. (1) Step 1: ATP induced S1 releasing; (2) Step 2: cyclic amplification procedure; and (3) Step 3: fabrication of the sensing interface and SWASV detection of Cd2 þ . Inset in the dashed box: preparation of S2-(CdS)4@SiO2.

create and maintain an RNase-free environment, all the samples and buffer solutions were dealed with 0.1% DEPC and stored at 4 1C before use. Notably, water was treated with DEPC before dissolving Tris to make the appropriate Tris buffer (Yin et al., 2012). All other chemicals, such as anhydrous ethanol (EtOH), acetone, H2SO4, HCl, HF, NaOH, HAc, NaAc, and NH3  H2O (25%) were of analytical grade. Double distilled water was used in all runs.

For CdS QDs solution, UV–vis spectra were measured on a TU1901 double beam UV–vis spectrophotometer (Beijing). Photoluminescence (PL) spectra were obtained on an RF-540 spectrophotometer (Shimadzu). Fluorescent microscopy images of different (CdS)n@SiO2 samples were detected on inverted fluorescence microscope (Nikon Ti) using 380–420 nm excitation light source. 2.3. Preparation of (CdS)n@SiO2 nanocomposite

2.2. Instruments All electrochemical experiments were carried out on a CHI 660D electrochemical workstation (Shanghai CH Instruments Co.) using a traditional three-electrode system. A platinum foil and a saturated calomel electrode (SCE) were used as counter electrode and reference electrode, respectively. The 3DOM PANI-IL composite film modified glassy carbon electrode (GCE, ϕ¼3 mm) was used as working electrode for square wave anodic stripping voltammetric (SWASV) measurements. The cyclic voltammetry (CV) and electrochemical impedance spectroscopy (EIS) measurements were implemented to characterize the modification process of 3DOM Au electrode surface. The morphologies of 3DOM PANI-IL film were characterized by field-emission SEM (FESEM, Hitachi S4800). The morphologies of the Fe3O4, Fe3O4@SiO2 and Au–Fe3O4@SiO2 were characterized by transmission electron microscopy (TEM, JEOL JEM-200CX). The morphologies and dimensions of the as-formed CdS QDs and (CdS)4@SiO2 samples were observed by a JEM-2100 high resolution TEM (HRTEM, JEOL).

TGA-stabilized CdS QDs were synthesized according to the literature with some minor modification (Yu et al., 2003). Briefly, 20 mL of 5 mM CdCl2 solution was adjusted with mercaptoacetic acid to pH 2, resulting in a turbid blue solution. Then 1.0 M NaOH solution was added to adjust the pH value to about 11. During the process, N2 was bubbled throughout the solution to remove O2. Then, 20 mL of 5 mM Na2S aqueous solution was injected into this solution to obtain TGAcapped water-soluble CdS QDs, and the reaction mixture was refluxed under N2 atmosphere for 4 h. The obtained TGA-capped CdS QDs was stored in a refrigerator at 4 1C for future use. The (CdS)n@SiO2 nanocomposite was obtained by using a slightly modified procedure reported by Wang et al. (2009). In a typical synthesis, 15 mL diluted CdS QDs solution was stirring with 100 mM EDC and 25 mM NHS for 2 h to activate carboxyl groups on the surface of CdS QDs. And then, 0.02 g amino-functionalized SiO2 nanospheres (ϕ¼20 nm) were dispersed in this solution and stirred for another 2 h. After that, it was centrifuged (9000 rpm) for 5 min to remove residual CdS QDs. The resulting (CdS)1@SiO2 nanocomposite was washed with 0.1 M PBS-T (pH 7.4 PBS containing 0.1% Tween 20) for three times to

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remove excess QDs, and resuspended in 2.0 mL PBS. To fabricate a larger nanocomposite containing more QDs, 10 mL 3% PDDA aqueous solution was added into (CdS)1-SiO2 nanocomposites under stirring to achieve positive charged modification. After 30 min, residual PDDA was removed by centrifugal separation and the nanocomposite was rinsed with water at least three times. Then, 15 mL diluted CdS QDs solution was added into the positive charged (CdS)1@SiO2 nanocomposites with stirring, resulting in (CdS)2@SiO2 nanocomposite. (CdS)n@SiO2 nanocomposite was obtained by repeating the operational procedure of PDDA modification and CdS QDs adsorption for several times, keeping the same dosage of PDDA and CdS QDs solution for every cycle. The each-stepped product of (CdS)n@SiO2 (n¼1–5) was collected and dispersed in PBS solution, respectively. 2.4. SWASV detection of ATP by DNA cycle amplification strategy 2.4.1. ATP recognition Firstly, ATP aptamer and S1 were hybridized in equal proportions (25 μL, 5.0 μM) and incubated at 37 1C for 1 h to form the duplex aptamer-S1. Then, the mixture of above aptamer-S1 solution and 50 μL 20 mg/mL of Au–SiO2@Fe3O4 dispersion was shaken for 12 h, forming S1/aptamer–Au–SiO2@Fe3O4. Afterwards, 20 μL of the ATP solution with different concentrations was added to S1/aptamer–Au–SiO2@Fe3O4 and incubated at 37 1C for 60 min, some S1 strands were released. The suspension was magnetic separated, and the supernatant containing S1 was sampled for the step 2.

2.4.2. Cyclic amplification procedure 50 μL of 4 μM MB was firstly heated at 95 1C for 2 min, then quickly cooled down from 95 1C to room temperature over 10 min. Afterwards, shake the mixture of MB and 50 μL of 20 mg/mL Au–SiO2@Fe3O4 dispersion overnight to obtain MB–Au–SiO2@ Fe3O4 composite. Then, 50 μL S1 solution obtained from step 1 and 1 mM Zn2 þ were added to MB–Au–SiO2@Fe3O4 dispersion and incubated at 37 1C for 30 min to make the cleavage of substrate MB strands. After the suspension was magnetic separated, and supernatant containing T1 was sampled for the step 3.

2.4.3. Fabrication of the sensing interface and SWASV detection of Cd2 þ After 30 μL of (CdS)4@SiO2 nanocomposite was activated by 0.2 M EDC and 0.1 M NHS for 30 min, 40 μL 5 μM of 50 –NH2 functionalized S2 was added into the (CdS)4@SiO2 dispersion and shaken overnight to covalently form S2-(CdS)4@SiO2. T1 solution obtained in step 2 was added to the S2-(CdS)4@SiO2 dispersion and incubated at 37 1C for 1 h to obtain the precipitate of T1/S2(CdS)4@SiO2, which was collected by centrifugation at 10,000 rpm and washed with PBS solution. Afterwards, it was dispersed in 25 mM pH 8.0 of Tris–HAc buffer containing 0.3 M NaCl for further use. Immersing S3/3DOM Au electrode into the dispersion of T1/S2(CdS)4@SiO2 for 1 h to immobilize (CdS)4@SiO2 onto the 3DOM Au surface through the partially complementation between T1 and S3. Put the (CdS)4@SiO2-S2/T1/S3-3DOM Au into a 400 μL of 0.1 M HNO3 for 30 min to dissolve CdS QDs. Subsequently, all the solution was added into a 4 mL glass vial containing 3.6 mL of acetate buffer (0.1 M, pH 4.6) spiked with 1.5 μM Bi3 þ . The SWASV detection involved pre-concentration at  1.2 V for 300 s under stirring. After an equilibrium period of 15 s, the SWASV curves were recorded from  1.1 to  0.5 V. Finally, the electrode was electrochemically cleaned at 0.3 V for 30 s to remove the residual metals and Bi film. All experiments were conducted at room temperature.

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3. Results and discussion 3.1. Characterization of (CdS)n@SiO2 nanocomposites and their stability evaluation Signal amplification is critical for fabrication of ultrasensitive electrochemical aptasensor. Layer-by-layer (LBL) assembled (CdS)n@SiO2 as a new class of signal amplification label, containing a larger number of CdS QDs, was involved in the ATP recognition event. PL and UV–vis techniques were applied to characterize the CdS QDs colloid solution, as shown in Fig. 1A. The UV–vis absorption maximum at 370 nm and the PL emission peak at 575 nm (λex ¼360 nm) indicated the consequence of quantum confinement. Additionally, the narrow and symmetric line width of PL spectrum also suggested that the CdS QDs were nearly monodisperse and homogeneous. Accordingly, the particle size of the CdS QDs was estimated using the empirical equation (Yu et al., 2003): D¼(  6.6521  10  8)λ3 þ(1.9557  10  4)λ2  (9.2352  2 10 )λþ 13.29. Here, D (nm) is the particle size of CdS QDs; λ is the UV–vis first absorption peak position (380 nm). The estimated result showed that the average size of the as-prepared CdS QDs was about 2.5 nm, which was in accordance with that from HRTEM characterization (Fig. 1E). Fluorescence microscopy images of (CdS)n@SiO2 (n ¼0–5) with different layers of CdS QDs, which assembled in conjunction with positively-charged polyelectrolyte polymer (PDDA), were used to evaluate the optimal number of layers. It can be seen from Fig. 1B that the (CdS)4@SiO2 exhibited the brightest image. Alternatively, after immobilizing different (CdS)n@SiO2 onto the bare GCE, SWASV was also implemented to characterize the electrochemical signals. As concluded from Fig. 1C, the highest SWASV current was also obtained from (CdS)4@SiO2. In this study, small sized SiO2 particles with the diameter of about 20 nm were used as the substrate, and it was postulated that the available space for incoming CdS QDs would be reduced as the number of LBL cycle increased due to the space hindrance and electrostatic repulsion. Thus, (CdS)4@SiO2 composite with four layers of CdS QDs was selected as the most efficient label for the subsequent assays. Additionally, from the TEM (Fig. 1D) and HRTEM images of (CdS)4@SiO2 (Fig. 1F), it can be observed that this core–shell structure contained a dark core of SiO2 and gray shell of uniformly distributed CdS QDs, with the size of about 36 nm. The structure stability of (CdS)4@SiO2 was evaluated by recording the fluorescence microscopy images and SWASV current of dissolved Cd2 þ . The (CdS)4@SiO2 composite was stored in PBS solution at 4 1C when it was not in use. After 6 months, the brightness and the SWASV current nearly kept constant, indicating the good long-term stability of the (CdS)4@SiO2 composite, due to the covalent coupling between the amino-functionalized SiO2 nanosphere and the first layered TGA-capped CdS QDs, and the electrostatic interaction between outer positively charged PDDA and negatively charged QDs. It also suggested that the LBL method may afford a more stable coating than that prepared by physical adsorption (Ma et al., 2011). 3.2. Characterization of Au–Fe3O4@SiO2 magnetic microspheres The morphologies of the Fe3O4, Fe3O4@SiO2 and Au–Fe3O4@SiO2 samples were characterized by TEM, as shown in Fig. S1. The TEM image of the Fe3O4 was shown in Fig. S1A, with the average diameter of about 20 nm. Besides, the pristine Fe3O4 nanoparticles tended to aggregate because of the magnetic properties. After coated with SiO2 shell, the dispersion of Fe3O4 beads was improved (Fig. S1B). Fig. S1C showed the TEM image of the Au–Fe3O4@SiO2 sample. Many Au NPs with a diameter of about 20 nm were adhered to the surfaces of the Fe3O4@SiO2

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Fig. 1. (A) PL (a) and UV–vis (b) spectra of the CdS QDs in aqueous solution. Inset was the fluorescence image obtained under UV light. (B) Fluorescence images of (CdS)n@SiO2: (a) pure CdS solution and (b–f) corresponding to the number of CdS layers (n) from 1 to 5, respectively. (C) SWASV curves (a–f) of (CdS)n@SiO2 corresponding to n¼ 0–5. Inset of (C) was the relationship between the SWASV current and n. HRTEM images of (D) SiO2, (E) CdS QDs and (F) (CdS)4@SiO2 particles, respectively.

nanocomposites. The EDS spectrum (Fig. S1D) showed that O, Fe, Si and Au elements were included in the Au–Fe3O4@SiO2 nanocomposite. The inset photographs of Fig. S1C showed that the prepared Au–Fe3O4@SiO2 was dispersed uniformly in water solution due to its small size, and the particles could be separated easily from solution with the help of an external magnetic force due to their strong magnetism. Furthermore, the outer Au nanoparticles could facilitate the combination with DNA strands through the well developed Au–S linkage.

3.3.1. Optimization of the recognition time of aptamer with ATP Since the concentration of ATP and the reaction time were closely related, the concentration of ATP was fixed at 50 nM when the effect of binding time of the aptamer with ATP on SWASV peak current was investigated. The interaction time includes both the binding time of ATP with the aptamer and the departure time of S1 from the Au–Fe3O4@SiO2 composite surface. From Fig. 2A, it was obvious that that peak current increased with the increased interaction time from 15 to 45 min and then reaches a plateau in 60 min, which suggested that 60 min was sufficient to allow ATP to bind its aptamer and induce the displacement of S1. Thus, 60 min was the interaction time chosen in the following experiments.

3.3. Optimization of experimental variables In order to maximize the detection sensitivity of proposed ATP aptasensor, various conditions were optimized, including the binding time of aptamer with ATP, reaction time between MB and Zn2 þ , the dosage of Zn2 þ , the electro-deposition charge of 3DOM PANI-IL and S3 immobilization quantity, via altering one condition while the others were fixed.

3.3.2. Optimization of the reaction time between MB and Zn2 þ Toward our goal for highly sensitive electrochemical detection of ATP, we optimized the reaction time between MB and Zn2 þ . Longer reaction time is expected to produce more T1 fragments and further lead to higher current output. As shown in Fig. 2B, the SWASV peak current increased with increasing the interaction

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Fig. 2. Effects of (A) binding time of the aptamer with ATP, (B) DNAzyme cleavage time, (C) the dosage of Zn2 þ and (D) the electro-polymerization charge of 3DOM PANI-IL film on the SWASV current responses of 1 μM Cd2 þ solution. (E) Effect of S3 immobilization concentration on the EIS response of S3/3DOM Au modified electrode.

time and then reaches a plateau in 30 min. Thus it was conceived that 30 min was sufficient for the cleavage reaction in the subsequent experiments. 3.3.3. Optimization of the dosage of Zn2 þ The S1 was chosen as the catalytic unit for amplifying the sensing signal by adopting Zn2 þ as cofactors (Lu et al., 2011). Upon the addition of Zn2 þ , the enzymatic recycling cleavage of MB strands was triggered successively, and then T1 segments were collected after removal of DNA–Au–Fe3O4@SiO2 composites. As shown in Fig. 2C, it indicated that the dosage of Zn2 þ would significantly affect the efficiency of cleavage. It was also found that the SWASV peak current increased with an increase of Zn2 þ dosage in the range from 0.1 to 0.5 mM and approximately reached a relative plateau over 1 mM. Therefore, 1 mM Zn2 þ was chosen as the optimal dosage (Fig. 2C). 3.3.4. Optimization of the electro-polymerization charge of 3DOM PANI-IL film It was reported that conducting polymer deposited on electrode surface could enhance SWASV response in detection of trace metal

ions (Rahman et al., 2003). And recently, ILs were found to be used as new types of dopant in conducting polymer synthesis due to their unique physicochemical properties, such as high thermal stability, negligible vapor pressure, good electrochemical stability and conductivity at room temperature (Saheb et al., 2008; Yang et al., 2008). Thus, herein we combine PANI-IL composite with 3DOM nanostructure to enhance the SWASV signal of CdS QDs. 3DOM PANI-IL modified electrode was fabricated by a potentiostatic technique (see supporting information). After the template of SiO2 spheres was removed by diluted HF solution, a well-ordered 3DOM PANI-IL was obtained. By adjusting the quantity of charge in electropolymerization, the formation of 3DOM PANI-IL could be controlled. As seen from Fig. 2D, when the concentration of Cd2 þ analyte was constant as 1 μM, the deposition charge increased from 0 to 5 mC, and then decreased at 10 mC. When the deposition charge exceeded 5 mC, some of the pores at the top layer might start to close and lead to the decrease of the surface area of 3DOM PANI-IL. Alternative explanation is that when the PANI-IL layer is beyond a critical thickness, the SWASV current of Cd2 þ may become weaker, because the thicker PANI-IL film may reduce the conductivity of the film (Wang et al., 2010). Thus, the deposition charge of 5 mC was chosen as the optimal electro-polymerization charge in this work.

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3.3.5. Optimization of S3 concentration The immobilized quantity of S3 on the 3DOM Au film electrode could influence the combined amount of the S2-(CdS)4@SiO2 via the linkage of T1, and then the sensitivity of the aptasensor. EIS technique was applied to detect the electron transfer resistance (Ret) of S3/3DOM Au modified electrode in pH 7.0 PBS solution containing 0.1 M KCl and 2.5 mM Fe(CN)36  /4-. From Fig. 2E, it was observed that the Ret value increased with the immobilized concentration of S3 solution from 0.1 to 5 μM, and then reached to a stable state until 10 μM. Since the backbone of DNA strands is negatively charged, the immobilized S3 could electrostatic repulse the same negatively charged probe of Fe(CN)36  /4  , causing the Ret value to increase accordingly. However, the immobilized amount of S3 strands was limited on a fixed surface of 3DOM Au film electrode, due to the steric hindrance and electrostatic repulsion among the immobilized S3 strands. Therefore, from the economical perspective, 5 μM of S3 solution was chosen for the modification of 3DOM Au film. 3.4. Amplification function of Bi/Nf/3DOM PANI-IL film From Fig. 3A, it was observed that the 3DOM PANI-IL structure was assembled in a hexagonal arrangement and connected to each other by symmetrical holes, with the center-to-center distance of about 460 nm. With respect to the diameter of the template SiO2 spheres (about 500 nm), the linear shrinkage was about 8%. Herein, the Bi/Nf/3DOM PANI-IL film was employed for the first time to enhance the SWASV peak current of Cd2 þ greatly. Nf film could facilitate the non-faradaic preconcentration of metal cations and improve the SWASV analysis sensitivity, due to the electrostatic absorption between the negatively charged sulfonate groups in Nf structure and the positively charged metal ions (AgraGutieÂrrez et al., 1999; Dam and Schroeder, 1996). In addition, Bi/Nf/PANI (curve a), Bi/Nf/PANI-IL (curve b) and Bi/Nf/3DOM PANI (curve c) modified electrodes with the same geometric surface area were compared as sensing interfaces in the detection of 1 μM Cd2 þ . As shown in Fig. 3B, the SWASV signal obtained from Bi/Nf/ 3DOM PANI-IL film (curve d) increased remarkably compared with the other three films. It suggested that the 3DOM PANI-IL structure possessed a larger active surface area. Further, it was shown that the SWASV signal became larger with the IL doping, indicating that the electron transfer was enhanced. 3.5. Sensitivity of ATP detection The target analyte ATP could trigger a series of scission reactions under the Zn2 þ assisted DNAzyme catalysis, which could be cycled for multiple rounds. Then, a large number of T1 segments could be associated from the system and numerous

(CdS)4@SiO2 could be immobilized onto the 3DOM Au electrode surface via the hybridization of S3/T1/S2, which led to greatly amplified increase of SWASV signal of Cd2 þ . As more concentration of target ATP released more T1 products, resulting in the immobilization of more (CdS)4@SiO2, thus more increase of SWASV signal was obtained. The change of SWASV signal was related to the concentration of target ATP molecules, which could be applied to the detection of ATP. Fig. 4A showed the SWASV signals for detecting different concentrations of target ATP. In the absence of ATP, only few S2-(CdS)4@SiO2 composite could be physically adsorbed onto the S3-3DOM PANI-IL surface, displaying very weak signal (curve a). When target ATP was present, the SWASV peak signal gradually increased with increasing ATP concentrations (curve b–k), indicating that the proposed strategy could be used to detect the ATP concentrations. The electrochemical ATP aptasensor displayed well-defined concentration dependence. As shown in Fig. 4B, the changes of Cd2 þ SWASV peak current (Δip) was linearly proportional to the logarithm value of ATP concentrations (log cATP) in the range between 1 pM and 10 nM, and between 10 nM and 1 μM, which could be quantified by the equations Δip ¼0.7506þ0.254 log cATP (R1 ¼ 0.98), and Δip ¼  2.7215þ3.114 log cATP (R2 ¼0.97), respectively. The detection limit was about 0.5 pM at 3s, suggesting that this method is highly sensitive and has great potential for accurate detection of ATP. As shown in Table S1, the sensitivity of our proposed aptasensor was much higher compared with the other ATP electrochemical aptasensors reported previously in literatures. The excellent performance of the aptasensor could be attributed to the following aspects: (1) the nanocomposites of the combined Au–SiO2@Fe3O4, (CdS)4@SiO2 and 3DOM Au were likely to allow more DNA strands on each surface; (2) the DNAzyme-assisted amplification technique increased the released amount of T1 strands; (3) large-sized (CdS)4@SiO2 containing much more CdS QDs than the usual single QD label may enhance the SWASV signal; (4) Bi/Nf/3DOM PANI-IL was efficient for detecting trace SWASV signal of Cd2 þ . In particular, it was noticeable that the SWASV peak potential of Cd2 þ shifted in the positive direction with increasing the ATP concentration. More ATP contained in the solution, more Cd2 þ was released. Due to the Cd2 þ absorbed in the porous structure of Bi/Nf/3DOM PANI-IL was poorly conductive, the resistance of the electrode may become larger after Cd2 þ absorption. Therefore, the SWASV peak shift of Cd2 þ might be due to the IR drop effect (Juárez et al., 2005; Armstrong et al., 2010). 3.6. Selectivity, regeneration and reproducibility of the aptasensor To evaluate the feasibility and reliability of the developed method, the aptasensor was applied to the determination of

Fig. 3. (A) SEM image of 3DOM PANI-IL film. Inset was the enlarged SEM image. (B) SWASV comparison on different sensing films in the detection of 1 μM Cd2 þ : (a) Bi/Nf/ PANI, (b) Bi/Nf/PANI-IL, (c) Bi/Nf/3DOM PANI and (d) Bi/Nf/3DOM PANI-IL films. Data has been processed by subtracting the blank signal.

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Fig. 4. (A) SWASV signal responses for detection of different concentrations of ATP. The concentrations of ATP from (a) to (k) were 0, 0.001, 0.01, 0.1, 1, 10, 50, 100, 500, 1000 and 2000 nM, respectively. (B) Detection linear relationship of Δip and log cATP in the range of 1 pM  10 nM and 10 nM  1 μM.

4. Conclusion

Table 1 Recovery of ATP from urine samples. Sample

Added (nM)

Found (nM)a

Recovery (%)b

1 2 3

0.05 5 50

0.0517 0.02 4.917 0.21 45.3 7 0.45

102 98.2 90.6

a The standard deviation of measurements were taken from five independent experiments. b Recovery means the ratio of Found (cATP) and Added (cATP).

several analogs with ATP: CTP, GTP and UTP (Huizenga, Szostak (1995)). Belonging to the nucleoside triphosphate family, CTP, GTP and UTP usually coexist with ATP in real biological samples. Differentiation of the other nucleoside triphosphate from ATP is of significant importance in bioassays. Target (ATP) and the control analytes (CTP, GTP and UTP) at the same concentration (100 nM) were spiked into the detection system and then measured using the SWASV method. Other conditions were the same as those described in experimental section. As seen from Fig. S2, only slight SWASV signal changes were observed in the addition of CTP, GTP and UTP, which indicated that the ATP aptamer possessed high affinity with ATP than the other ATP analogs. The high specificity of the aptasensor allowed the direct determination of ATP by SWASV method without the need for other separation methods. The regeneration of the 3DOM Au film was developed by treating the modified electrode with piranha solution (30% H2O2: 98% H2SO4 ¼3:7, V/V) for 1 min to violently peel all the adsorbed DNA strands from the 3DOM Au film substrate surface, rinsed with water, and electrochemically cleaned in 0.5 M H2SO4 prior to use. The reuse lifetime of regenerated 3DOM Au film was more than 10 assay runs. A series of three duplicate measurements of 10 pM and 100 nM ATP were used for estimating the precision, and the relative standard deviation (RSD) values were 5.6% and 3.7%, respectively, showing good reproducibility. 3.7. Analytical application of the aptasensor The electrochemical aptasensor was further evaluated by the standard addition method. A series of samples were prepared by adding ATP standard solution of different concentrations to human urine samples. From the analytical results shown in Table 1, it could be concluded that the recovery values of three samples for the added ATP with 0.05, 5 and 50 nM were 108%, 98.2% and 90.6%, respectively, which implies that the aptasensor has a promising feature for the analytical application in complex biological samples.

In summary, an ultrasensitive electrochemical aptasensor for the detection of ATP was developed by DNAzyme assisted cyclic amplification technique using QDs composite of large size as signal probe and 3DOM Bi/PANI-IL film as sensing platform. The SWASV peak current change (Δip) of QDs was found to be linear proportional with the concentration of ATP, with a detection limit of 0.5 pM. This low detection limit was primarily attributed to the “signal on” configuration of our method, which has shown an improved sensitivity over the traditional “signal off” schemes for small molecules based on target-induced strand displacement or target-induced conformational changes of aptamers. As a proof-ofconcept, we demonstrated that the aptasensor could sensitively and selectively detect ATP in real samples, such as in urine. This detection system has provided a universal approach for the detection of any types of target molecules by simply changing the aptamer sequences. Acknowledgments We greatly appreciate the support from the National Natural Science Foundation of China (20905035), State Key Laboratory of Materials-Oriented Chemical Engineering (KL10-15) and 973 Program (No. 2011CBA00807). This work is also supported by Qing Lan Project and Overseas Research & Training Program of Education Department of Jiangsu Province. Appendix A. Supporting information Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.bios.2014.02.043. References Agra-GutieÂrrez, C., SuaÂrez, M.F., Compton, R.G., 1999. Electroanalysis 11, 16–22. Armstrong, K.C., Tatum, C.E., Dansby-Sparks, R.N., Chambers, J.Q., Xue, Z.L., 2010. Talanta 82, 675–680. Brandon, M., Baldi, P., Wallace, D.C., 2006. Oncogene 25, 4647–4662. Chen, J.R., Jiao, X.X., Luo, H.Q., Li, N.B., 2013. J. Mater. Chem. 1, 861–864. Dam, M.E.R., Schroeder, K.H., 1996. Electroanalysis 8, 1040–1050. Davidson, C.A., Griffith, C.J., Peters, A.C., Fielding, L.M., 1999. Luminescence 14, 33–38. D’Orazio, P., 2011. Clin. Chim. Acta 412, 1749–1761. Du, Y., Chen, C.G., Zhou, M., Dong, S.J., Wang, E.K., 2011. Anal. Chem. 83, 1523–1529. Guo, S.J., Du, Y., Yang, X., Dong, S.J., Wang, E.K., 2011. Anal. Chem. 83, 8035–8040. Hansen, J.A., Mukhopadhyay, R., Hansen, J., Gothelf, K.V., 2006a. J. Am. Chem. Soc. 128, 3860–3861. Hansen, J.A., Wang, J., Kawde, A.N., Xiang, Y., Gothelf, K.V., Collins, G., 2006b. J. Am. Chem. Soc. 128, 2228–2229. Huizenga, D.E., Szostak, J.W., 1995. Biochemistry 34, 656–665.

56

X. Chen et al. / Biosensors and Bioelectronics 58 (2014) 48–56

Imisides, M.D., John, R., Riley, P.J., Wallace, G.G., 1991. Electroanalysis 3, 879–889. Juárez, A.V., Baruzzi, A.M., Yudi, L.M., 2005. J. Electroanal. Chem. 577, 281–286. Knowles, J.R., 1980. Annu. Rev. Biochem. 49, 877–919. Li, Z., Wang, Y.J., Liu, Y., Zeng, Y.Y., Huang, A.M., Peng, N.C., Liu, X.L., Liu, J.F., 2013. Analyst 138, 4732–4736. Liu, Q., Jiang, C., Zheng, X.X., Gu, Z., Li, D., Li, D.W., Huang, Q., Long, Y.T., Fan, C.H., 2012. Chem. Commun. 48, 9574–9576. Lu, J.J., Yan, M., Ge, L., Ge, S.G., Wang, S.W., Yan, J.X., Yu, J.H., 2013. Biosens. Bioelectron. 47, 271–277. Lu, L.M., Zhang, X.B., Kong, R.M., Yang, B., Tan, W.H., 2011. J. Am. Chem. Soc. 133, 11686–11691. Lu, S.F., Wang, Y., Zhang, C.X., Lin, Y., Li, F., 2013. Chem. Commun. 49, 2335–2337. Lubin, A.A., Plaxco, K.W., 2010. Acc. Chem. Res. 43, 496–505. Ma, Q., Ha, E.N., Yang, F.P., Su, X.G., 2011. Anal. Chim. Acta 701, 60–65. Rahman, Md. A., Won, M.S., Shim, Y.B., 2003. Anal. Chem. 75, 1123–1129. Ribeiro, A.R., Santos, R.M., Rosario, L.M., Gil, M.H., 1998. J. Biolumin. Chemilumin. 13, 371–378. Ruslinda, A.R., Ishiyama, Y., Wang, X., Kobayahi, T., Kawarada, H., 2012. J. Electrochem. Soc. 159, J182–J187. Saheb, A., Josowicz, M., Janata, J., 2008. Anal. Chem. 80, 4214–4219. Stratford, M.R., Dennis, M.F., 1994. J. Chromatogr. B Biomed. Sci. Appl. 662, 15–20. Tedsana, W., Tuntulani, T., Ngeontae, W., 2013. Anal. Chim. Acta 783, 65–73.

Walcarius, A., 2012. TrAc Trends Anal. Chem. 38, 79–97. Wang, J., Liu, G., Polsky, R., Merkoçi, A., 2002. Electrochem. Commun. 4, 722–726. Wang, J., Liu, G., Rivas, G., 2003a. Anal. Chem. 75, 4667–4671. Wang, J., Liu, G., Merkoçi, A., 2003b. J. Am. Chem. Soc. 125, 3214–3215. Wang, J, Lu, J., Hǒ cevar, S.B., Farias, P.A.M., Ogorevc, B., 2000. Anal. Chem. 72, 3218–3222. Wang, L., Xu, M., Han, L., Zhou, M., Zhu, C.Z., Dong, S.J., 2012. Anal. Chem. 84, 7301–7307. Wang, Z.M., Guo, H.W., Liu, E.G., Yang, C., Khun, N.W., 2010. Electroanalysis 22, 209–215. Weller, H., 1993. Angew. Chem. Int. Ed. Engl. 32, 41–53. Xu, S.X., Zhang, X.F., Liu, W.W., Sun, Y.H., Zhang, H.L., 2013. Biosens. Bioelectron. 43, 160–164. Xiao, Y., Piorek, B.D., Plaxco, K.W., Heeger, A.J., 2005. J. Am. Chem. Soc. 127, 17990–17991. Yang, W.R., Liu, J.Q., Zheng, R.K., Liu, Z.W., Dai, Y., Chen, G.N., Ringer, S., Braet, F., 2008. Nanoscale Res. Lett. 3, 468–472. Yin, B.C., Guan, Y.M., Ye, B.C., 2012. Chem. Commun. 48, 4208–4210. Yu, W.W., Qu, L.H., Guo, W.Z., Peng, X.G., 2003. Chem. Mater. 15, 2854–2860. Zhou, J.J., Huang, H.P., Xuan, J., Zhang, J.R., Zhu, J.J., 2010. Biosens. Bioelectron. 26, 834–840.