nickel hexacyanoferrate nanotube assembly for biosensor applications

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Biomaterials 28 (2007) 3408–3417 www.elsevier.com/locate/biomaterials

Functional histidine/nickel hexacyanoferrate nanotube assembly for biosensor applications Minghui Yang, Jianhui Jiang, Yashuang Lu, Yan He, Guoli Shen, Ruqin Yu State Key Laboratory of Chemo/Biosensing and Chemometrics, Biomedical Engineering Center, Chemistry and Chemical Engineering College, Hunan University, Changsha 410082, PR China Received 7 February 2007; accepted 3 April 2007 Available online 19 April 2007

Abstract By exploring the properties of histidine, functional histidine/nickel hexacyanoferrate nanotube assembly was prepared using nanopore alumina template via a sequential deposition strategy and demonstrated for improved biosensing. The vertically oriented nanotube assembly has a nanotube density 8  108 cm2 and can be stably attached to the glassy carbon electrode surface. The nanotube assembly formed provides an ordered well-defined three-dimensional (3D) structure with good electron transfer efficiency and a large specific surface area with abundant electroactive sites. Gold nanoparticles were then absorbed onto nanotube surfaces through amino group provided by histidine, and were used for further absorption of glucose oxidase into the 3D matrix. The histidine and gold nanoparticles on the nanotube surface provide a favorable microenvironment to keep the bioactivity of the enzymes with a low apparent Michaelis–Menten constant (KMapp) of 2.15 mM, and the ordered orientation of the nanotube assembly facilitated enzyme–substrate contact. The biosensor was sensitive and selective toward glucose with a linear range covered from 2 mM to 20 mM of glucose. The biosensor was used to determine glucose concentration in real blood samples with satisfactory results. r 2007 Published by Elsevier Ltd. Keywords: Biosensor; Electrochemistry; Enzyme; Gold nanoparticle; Metal hexacyanoferrate; Nanotube assembly

1. Introduction The rapid progress in nanoscience and nanotechnology introduced a fast growth in the field of electrochemical biosensors during the past years [1–6]. As promising building blocks for biosensing platforms with high complexity and controlled structure, nanotubes such as carbon nanotubes (CNTs) have gained special attention [7–11]. Aligned nanotube assembly provides a well-defined structure and large specific surface area for enzyme modification. The vertically oriented ordered nanotubes allow enzymes to be fully accessible to the substrate, so that the sensitivity and the scope of analytes to be detected improved greatly [12–15]. However, the attachment of biomolecules to CNTs is difficult and the molecules are mainly restricted to the open end of CNTs [16,17]. Corresponding authors. Tel./fax: +86 731 8821355.

E-mail addresses: [email protected] (Y. He), [email protected] (G. Shen). 0142-9612/$ - see front matter r 2007 Published by Elsevier Ltd. doi:10.1016/j.biomaterials.2007.04.020

Moreover, it has been reported that the high electrochemical reactivity of CNT-modified electrode toward hydrogen peroxide (H2O2) is due to metal impurities present in the nanotubes rather than intrinsic properties of the nanotube itself [18]. Because of interesting properties such as high conductivity, optical and catalytic properties, inorganic nanotubes have attracted considerable attention [19–21]. For example, platinum nanotubes were prepared using Se nanowires as the template, and the nanotubes can serve as catalysts for various organic reactions [22]. Among the numerous methods, porous template-based method is the most widely used method for inorganic nanotube and nanotube array preparation [23–25]. Au nanotubules were deposited within the pores of polycarbonate membrane and used for separation of proteins [26]. Transition metal hexacyanoferrates (MHCFs) represent an important class of mixed-valence compounds that have been studied extensively for many years. Wide application have been found of these compounds due to interesting

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properties such as ion-exchange properties, electrochromism, mixed-valence electrical conductivity, and high catalytic activity [27–29]. The molecular formula of this kind of compound is as follows: XMII ½FeIII ðCNÞ6 ; where X is alkali cation like Li+, Na+, or K+, and M is transition metal such as FeIII , CoII, or NiII. Different ways have been explored to modify the electrode surface with these compounds, including electrochemical, layer-by-layer method and using MHCF nanoparticles [30–33]. But the MHCF film is brittle and not stable, and it is relatively difficult to assemble the MHCF nanoparticles [34]. So far little attention has been paid to the preparation of MHCF nanotubes for biosensing application. It will be a significant advancement if perpendicularly aligned MHCF nanotube assemblies can be used as the sensing materials. In this paper, we describe a simple route to the production of an ordered functional histidine/nickel hexacyanoferrate nanotube assembly using template method. The preparation method is simple, first nickel hexacyanoferrate nanoparticles were formed along the template pore wall, and with the increase of the amount of nanoparticles, nanoparticles connect with each other forming the nanotubes. The analytical performance of the resulting nanotube assembly toward the detection of H2O2 was greatly improved. Gold nanoparticles were then easily adsorbed onto the nanotube surface through amino groups due to the presence of histidine, and were used for further absorption of glucose oxidase (GOx) uniformly onto the nanotubes for the fabrication of glucose biosensors. The biosensor fabrication method used here have some advantages: employ mild condition allowing for large quantities of enzymes to be immobilized uniformly; the gold nanoparticles and histidine on the nanotube surface provide a favorable microenvironment to maintain the enzyme activity.

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Elmer UV–vis–near-IR spectrometer, Lambda 900. A three-electrode cell (10 mL) with the nanotube assembly modified glassy carbon (GC) electrode as the working electrode, a saturated calomel electrode (SCE) as the reference electrode and a platinum foil electrode as the counter electrode was used. All potentials were measured and reported versus the SCE and all experiments were carried out at room temperature.

2.2. Preparation of the nanotube assembly The procedure to deposit histidine/nickel hexacyanoferrate nanotube onto the alumina membrane consists of the following steps: (1) place the anodic membrane template membrane in 0.005 M histidine (pH 9.0) solution for 30 min, (2) clean the membrane in 0.2 M KCl solution for 10 min, (3) immerse the membrane 0.01 M NiCl2 solution for 30 min, (4) clean the membrane in 0.2 M KCl solution for 10 min, (5) immerse the membrane in 0.2 M KCl-0.01 M K3[Fe(CN)6] solution for 30 min. This procedure is herein referred to as one deposition cycle. The procedure involved careful stirring with a magnetic stirrer, and KCl solutions were renewed between each step. After the desired number of deposition cycles, the nanotube assembly was obtained by dissolving the template membrane in a 10% phosphoric acid at 4 1C for 12 h.

2.3. Modification of the nanotubes For modification of the nanotube assembly with gold nanoparticles, the liberated nanotube film was immersed into 13 nm diameter gold nanoparticle solution prepared by the citrate reduction of HAuCl4 for 4 h [35]. For modification of the nanotube assembly with protein GOx, the nanotube film modified with gold nanoparticles was immersed into GOx solution (5 mg mL1) at 4 1C for 12 h.

2.4. Fabrication of nanotube assembly modified electrode The nanotube assembly modified electrode was prepared by casting the bare nanotube film or the GOx-modified nanotube film onto the surface of GC electrodes (4 mm diameter), after which 3 mL of chitosan solution (0.05%, dissolved in 1% acetic acid) was coated onto the electrode for protection.

3. Results and discussion 3.1. Preparation and characterization of the nanotube assembly

2. Experimental section 2.1. Apparatus and reagents Anodic alumina membrane was purchased from Whatman (Anodisc 25, pore size 0.2 mm). GOx (from Aspergillus niger; EC 1.1.3.4, type VII-S; 196,000 unit g1), chitosan (CHIT, MW1  106; 80% deacetylation) were purchased from Sigma. 0.1 M phosphate buffer (pH 7.0) solutions were prepared with Na2HPO4 and NaH2PO4. All other reagents were of analytical grade, and doubly distilled water was used throughout. Cyclic voltammetric and amperometric measurements were carried out on CHI 760B electrochemical workstation (Shanghai, China). Scanning electron microscopy (SEM) analysis was performed using a JSM-5600LV microscope (JEOL Co. Ltd., Japan). Transmission electron microscope (TEM) image was taken with a JEM-3010 transmission electron microscope (JEOL Co. Ltd., Japan). The TEM microscope was equipped with an EDAX energy dispersive spectroscopy (EDS) analyzer. X-ray diffraction (XRD) analysis was performed on a Siemens D5000 diffractometer, using Cu Ka radiation. Fourier transform infrared spectra (FT-IR) measured by KBr pellets was obtained on a WQF-410 Fourier transform infrared spectrophotometer (Beijing Secondary Optical Instruments, China). UV–vis adsorption spectra were recorded on a Perkin-

In an earlier report, Prussian blue (PB) nanotubes were prepared using a sequential deposition method [36]. However, the nanotubes prepared in that way had variable lengths, and appeared as loose precipitates after the alumina template was etched away. In this work, the problem was overcome by modifying the sequential deposition procedure based on the properties of histidine. It has been reported that histidine can adsorb strongly to the alumina surface at pH 9.0 and it is also known that histidine has good affinity toward nickel ions [37,38], therefore high-density histidine molecules adsorbed on the alumina surface can serve as ‘‘anchoring’’ sites for nickel ions, which form precipitates with hexacyanoferrates according to the following equation [39]: Ni2þ þ ½FeðCNÞ6 3  þ Kþ ! KNi½FeðCNÞ6 : By sequentially dipping porous alumina template into histidine solution, nickel ion solution and hexacyanoferrate

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solution for a number of cycles, a large amount of PB-like nanoparticles cover densely not only the pore walls but also the open-end surface of the template membrane, leading to nanotubes inter-linked to each other. The histidine also makes it easier for further enzyme immobilization, which will be discussed later. Fig. 1 shows the SEM images of a nanotube assembly obtained after 10 deposition cycles. The film was coated onto a glass slide. The assembly has highly ordered 3-D structure with nanotubes interconnected to each other and the hollows cores aligned perpendicular to the glass slide surface. The outer diameters of the nanotubes are close to the pore diameter of the template, and the surface density of the nanotubes is calculated to be 8  108/cm2. To have a detailed examination of its structure, the nanotube assembly was sonicated in water and forced to fall apart as precipitates in the solution. SEM image (Fig. 2A) shows that the length of these nanotubes was more than 10 mm. Many of the tubes were broken due to the sonication process to collect individual nanotubes [24]. The highresolution TEM image (Fig. 2B) indicates that there were no visible defects on the nanotube, and its wall thickness

Fig. 2. SEM (A) and TEM (B) images of the liberated nanotube. (C): EDS profile of the nanotube in (B).

Fig. 1. SEM images of the nanotube assembly.

was measured to be approximately 30 nm (between two arrows). The energy dispersive spectroscopy (EDS) profile of the nanotube (Fig. 2C) reveals the presence of nickel, iron and potassium, which were the component elements of nickel hexacyanoferrates. The powder X-ray diffraction (XRD) pattern of the deposited nanotubes showed a typical face-centered cubic (fcc) stucture (Fig. 3) with the

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3.2. Performance of the nanotube assembly modified electrode The densely packed and vertically aligned nickel hexacyanoferrate nanotube assembly makes it an ideal material for electrochemical biosensing applications. The large surface area provides enormous number of electrocatalytic sites. The semi-conducting metal hexacyanoferric backbone ensures efficient electron transfer between immobilized redox materials and the electrode [43,44]. To characterize its electrochemical properties, a nanotube assembly obtained after 10 deposition cycles was cast onto a 4 mm diameter GC electrode. Fig. 5A shows the cyclic voltammograms (CVs) of the bare GC electrode and the nanotube assembly modified GC electrode in 0.1 M sodium phosphate buffer solution at potential scan rate of

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lattic constant calculated as a ¼ 10.16 A˚, indicating the presence of PB analog [40]. UV–vis spectroscopy and FT-IR were used to further characterize the nanotubes. The nanotube assembly possesses broad absorption peaks around 290 and 400 nm (Fig. 4A). The absorption at 290 nm can be assigned to the ligand to metal charge transfer (LMCT) band of [FeIII(CN)6], and that at 400 nm may be due to the d–d transitions of NiII [41]. FT-IR spectrum of the nanotube assembly display a sharp peak at 2175 cm1(Fig. 4B), which can be assigned to the stretching vibration of the CN group in nickel hexacyanoferrate structure, confirming the unit of Fe–CN–Ni exists in the nanotube structure [42]. The small peaks at 2098, 3024 and 3117 cm1 are due to the presence of histidine on the nanotube wall, which would be proved from the spectrum of the pure histidine (Fig. 4C).

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Fig. 3. XRD pattern showing the fcc crystal structure of the nanotube.

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Fig. 5. (A) Cyclic voltammograms (CVs) of (a) bare GC and (b) nanotube assembly modified GC electrodes in phosphate buffer at 50 mV s1. (B) Stability of the nanotube assembly modified GC electrode in phosphate buffer upon repeated potential scans at 50 mV s1 for 10 cycles. (C) Nanotube assembly modified GC electrodes in phosphate buffer at different scan rates. From inside to outside: 5–500 mV s1. (D) The anodic and cathodic current plotted against the square root of the scan rate.

50 mV s1. For the bare GC, no peak current was observed (curve a), while for the one modified with the nanotube assembly, there is one pair of reversible redox peaks with formal potential (E00 ¼ [(Ep,a+Ep,c)/2]) of about 0.17 V, which is attributed to the transformations between Fe(II) and Fe(III) in the nanotubes. Sweeping the potential from 0.4 V to 0.8 V at 50 mV s1 shows nearly no peak current and peak potential change after 10 cycles (Fig. 5B). The relative standard deviation (RSD) of peak current and peak potential is 1.8% and 0.78%, respectively, indicating

that the nanotube assembly is stably immobilized onto the electrode surface via chitosan. Fig. 5C shows the CVs in 0.1 M sodium phosphate buffer at different scan rates. It can be seen that both the anodic and cathodic peak currents and the peak-to-peak separation clearly increase with increasing potential scan rates, suggesting quasireversible behavior [45]. One very interesting result is that the peak currents increase linearly with the square root of the scan rate in a range from 5 to 500 mV s1 (Fig. 5D), which is characteristic of a diffusion-controlled process rather

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than a surface-confined process [46]. It has been reported that nickel hexacyanoferrate film electrodeposited on GC electrode show surface-confined behavior in 0.1 M KCl solution [47]. It is known that in redox reactions of metal hexacyanoferrates, the transfer of cations between the solid compound and the adjacent solution is required to maintain charge balance [48,49]. For the nanotube assembly, the majority of electroactive sites are buried densely and deeply inside the porous structure though they are freely accessible by the supporting solution. Even with bulk concentration as high as 0.1 M, the local supply of cations may not be enough due to either reduced diffusion coefficient or fast charge transfer through the 3-D metal hexacyanoferrate network. As a result, local cation concentration gradient may develop within the space occupied by the nanotube assembly and diffusion of cations through the porous network becomes the ratelimiting step. Further studies to map the local diffusion coefficients and concentrations of the cation are needed to confirm the hypothesis. The electrocatalytic performance of the nanotube assembly was demonstrated using H2O2. Fig. 6A shows the CVs of the modified GC electrode in different H2O2 solutions. It can be seen that with increasing H2O2 concentration the reduction peak current of the nanotube increased and the oxidation peak current decreased indicating that the nanotube assembly catalyzed H2O2 reduction. Fig. 6B displays a typical Current–time curve for 1 mM H2O2 (10 mL of 1 M H2O2 added into 10 mL buffer) at the nanotube assembly modified electrode and the insert is the calibration curve. In phosphate buffer, direct determination of H2O2 could be achieved at 0.2 V. As can be seen, fast and sensitive response could be obtained with steady-state current produced within 7 s, indicating good electron transfer efficiency. This is probably due to the nickel hexacyanoferrate framework and the vertical orientation of the nanotube assembly, which makes the catalytic sites fully accessible to the substrate. The lower detection limit is 1.0  107 M, which is two times lower than electrodeposited PB-modified electrodes (2.5  107 M) and five times lower than PB nanoparticles-modified carbon paste electrodes (5.0  107 M) [50]. For peroxidase enzyme-based H2O2 sensors, the main disadvantage is the saturation of the enzyme with substrate that affects the upper detection limit of the sensor. For our nanotube assembly modified electrode, however, linear response towards H2O2 can be observed (Fig. 6B) even after 30 successive additions of 1 mM H2O2. The linear range of the modified electrode covers five orders of magnitude of H2O2 concentration (1.0  107–5.0  102 M), which is wider than enzyme-based (5.0  106– 1.0  102 M) and PB modified H2O2 sensors (1.0  107–5.0  104 M) [51,52]. The wide linear range can be ascribed to the fact that the vertically oriented nanotube assembly provides a large amount of surface electroactive sites, and the well-defined surface structure facilitated

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substrate–electrode contact. Moreover, repeated use of the electrode did not affect its long-term stability and good repeatability was obtained. For example, 1 mM H2O2 was measured consecutively for 10 times, and a RSD of 4.6% was obtained. 3.3. Modification of the nanotube with gold nanoparticles The good performance of the nanotube assembly modified electrode toward the detection of H2O2 makes it

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attractive for the fabrication of oxidase-based biosensors, as enzymatic reaction between oxidase and its specific substrate will produce H2O2, and through the detection of H2O2 the concentration of substrate could be detected. In this study, we incorporated GOx within the three-dimensional (3D) nanotube assembly for the fabrication of biosensors. Direct adsorption of enzymes onto nanotube surfaces makes it easy to capture intermediates created in the enzymatic reaction and also facilitates enzyme–substrate contact. For enzyme-based nano-biosensors, however, one key issue is to maximize the amount of enzymes

immobilized on the nanostructure while keeping their bioactivity. In the present study, this is achieved again through hisditine, which not only ensures the formation of an ordered nanostructure assembly but also introduces abundant amino groups on the nanotube surface, making it easier for further modification. By simply immersing the nanotube assembly in gold nanoparticle solution for 4 h, nanopartilces were immobilized on nanotube surfaces via the amino groups provided by histidine. TEM studies (Fig. 7) confirmed the successful attachment of gold nanoparticles onto the nanotubes. Large quantity of gold nanoparticles decorates the walls of the ordered nanotubes uniformly. After gold nanoparticle attachment, the modified nanotube assembly was further immersed in GOx

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Fig. 8. (A) Current–time recording for successive additions of 0.5 mM glucose at the nanotube assembly modified glucose biosensor. The inset is the calibration curve towards glucose. Applied potential, 0.2 V. (B) Amperometric responses of the glucose biosensor to glucose (Glu), ascorbic acid (AA), uric acid (UA), and acetaminophen (AP) at a potential of 0.2 V.

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solution for enzyme immobilization. Previous works have shown that enzymes maintain their catalytic and electrochemical activity when immobilized onto gold nanoparticles [53,54].

3.4. Performance of the glucose biosensor Fig. 8A shows the amperometric response at the GOxmodified biosensor to successive addition of 0.5 mM glucose at 0.2 V and the inset is the calibration curve. Immediately after the addition of glucose, the cathodic current of the biosensor increased, reaching a steady state within 20 s. The biosensor was sensitive toward glucose with a linear range covered from 2 mM up to 20 mM of glucose. The detection limit is 1 mM based on S/N ¼ 3. The wide dynamic range of the glucose biosensor may be ascribed to the high quantity of GOx incorporated into the 3D matrix, and the good performance of the nanotube assembly modified electrode in H2O2 detection. The apparent Michaelis–Menten constant (KMapp), which gives an indication of the enzyme–substrate kinetics, can be calculated from the linear part of the calibration curve [55]. A value of 2.15 mM for KMapp was obtained using the Lineweaver–Burk equation. The KMapp for GOx in this work is lower than the 22 mM for GOx entrapped in sol–gel organic–inorganic hybrid material [56], the 4 mM for GOx bound to plasma-polymerized film [57], and the 17 mM for GOx immobilized through layer-by-layer method using avidin and biotin [58], indicating that GOx absorbed onto the nanotube surface shows higher catalytic activity. The repeatability of the biosensor was also investigated. A set of 10 different amperometric measurements for 1 mM glucose with a single biosensor yield a relative standard deviation of 5.5%, indicating reliable performance of the modified electrode. Interferences from electroactive compounds commonly present in physiological samples usually prevent accurate determination of glucose concentration. The use of lower detection potential in this study greatly reduced the responses of common interference. Fig. 8B shows the amperometric responses of the glucose biosensor with the addition of 1 mM glucose and interfering species

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(0.1 mM ascorbic, 0.1 mM uric acid, and 0.1 mM acetaminophen). Compared to the response toward glucose, the current generated due to the interfering species are negligible, indicating high selectivity of the biosensor. The long-term stability of the nanotube assembly modified biosensor was studied when it was stored in phosphate buffer at 4 1C for an extended period of time. The biosensor was tested every 3 days. After 1 month of storage, the response of the biosensor only decreased 15% from the initial value, which indicates that the nanotube assembly biosensor did provide a favorable microenvironment to maintain the activity of GOx. 3.5. Real sample analysis The applicability of the biosensor was assessed by the determination of glucose concentration in real blood samples. Results were compared with those determined by hospital using colorimetric method with glucose kit (Shanghai Kehua-Dongling Diagnostic Products Co. Ltd.). A total of 10 serum samples were analyzed (0.5 mL sample added into 10 mL buffer) with each sample analyzed three times and the result is shown in Table 1. Glucose contents determined by the two methods agreed well, and a plot of the glucose concentration obtained by the two methods gave a straight line with a correlation coefficient of 0.992, indicating the biosensor can be used clinically for the detection of glucose. 4. Conclusions By exploring the properties of histidine, we prepared functional histidine/nickel hexacyanoferrate nanotube assembly using nanopore alumina template via a sequential deposition strategy. The nanotube assembly prepared can be stably immobilized onto GC electrode surface with nanotube density 8  108/cm2. It provides an ordered well-defined three-dimensional (3D) structure with good electron transfer efficiency and a large specific surface area with abundant electroactive sites, resulting in improved sensitivity and broadened dynamic range toward hydrogen peroxide (H2O2) reduction. The nanotube assembly

Table 1 Compare of the glucose contents in blood samples determined with the biosensor and the hospital method Sample

Glucose content determined by the biosensor (mM)

Glucose content determined by hospital method (mM)

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7.3570.22 7.5470.24 7.8770.21 8.6970.28 8.7570.24 11.2470.34 11.5870.31 11.8770.33 12.7570.35 12.8970.43

7.4670.18 7.7770.20 7.6870.20 8.9170.22 8.5570.21 11.5470.34 11.7870.31 12.0570.30 13.1570.28 13.2970.38

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demonstrates low detection limit (107 M) and wide linear range (1.0  107–5.0  102 M) toward H2O2, which made it an idea platform for the fabrication of biosensors. Gold nanoparticles were absorbed onto the nanotube surface uniformly through amino group due to histidine at the nanotube surface, and were used for further absorption of protein glucose oxidase into the 3D matrix. The histidine and gold nanoparticles on the nanotube surface provide a favorable microenvironment to keep the bioactivity of the enzymes, and the ordered orientation of the nanotube array facilitated enzyme–substrate contact. The biosensor enables a highly selective determination of glucose with wide linear range (2  106–2  102 M). Acknowledgments This work was supported by the NNSF of China (Nos. 20435010, 20375012 and 20205005). References [1] Hrapovic S, Majid E, Liu Y, Male K, Luong JHT. Metallic nanoparticle–carbon nanotube composites for electrochemical determination of explosive nitroaromatic compounds. Anal Chem 2006;78:5504–12. [2] Majid E, Hrapovic S, Liu YL, Male KB, Luong JHT. Electrochemical determination of arsenite using a gold nanoparticle modified glassy carbon electrode and flow Analysis. Anal Chem 2006;78:762–9. [3] Wang JB, Profitt JA, Pugia MJ, Suni II. Au nanoparticle conjugation for impedance and capacitance signal amplification in biosensors. Anal Chem 2006;78:1769–73. [4] Joshi PP, Merchant SA, Wang YD, Schmidtke DW. Amperometric biosensors based on redox polymer–carbon nanotube–enzyme composites. Anal Chem 2005;77:3183–8. [5] Wang J, Musameh M. Carbon nanotube/Teflon composite electrochemical sensors and biosensors. Anal Chem 2003;75:2075–9. [6] Wang J, Musameh M, Lin YH. Solubilization of carbon nanotubes by nafion toward the preparation of amperometric biosensors. J Am Chem Soc 2003;125:2408–9. [7] Azamian BR, Davis JJ, Coleman KS, Bagshaw CB, Green MLH. Bioelectrochemical single-walled carbon nanotubes. J Am Chem Soc 2002;124:12664–5. [8] Liu GD, Lin YH. Biosensor based on self-assembling acetylcholinesterase on carbon nanotubes for flow injection/amperometric detection of organophosphate pesticides and nerve agents. Anal Chem 2006;78:835–43. [9] Zhang MG, Gorski W. Electrochemical sensing platform based on the carbon nanotubes/redox mediators-biopolymer system. J Am Chem Soc 2005;127:2058–9. [10] Zhang MG, Smith A, Gorski W. Carbon nanotube–chitosan system for electrochemical sensing based on dehydrogenase enzymes. Anal Chem 2004;76:5045–50. [11] Hrapovic S, Liu YL, Male KB, Luong JHT. Electrochemical biosensing platforms using platinum nanoparticles and carbon nanotubes. Anal Chem 2004;76:1083–8. [12] Wang J, Scampicchio M, Laocharoensuk R, Valentini F, GonzalezGarcia O, Burdick J. Magnetic tuning of the electrochemical reactivity through controlled surface orientation of catalytic nanowires. J Am Chem Soc 2006;128:4562–3. [13] Wei C, Dai LM, Roy A, Tolle TB. Multifunctional chemical vapor sensors of aligned carbon nanotube and polymer composites. J Am Chem Soc 2006;128:1412–3. [14] Lin YH, Lu F, Tu Y, Ren ZF. Glucose biosensors based on carbon nanotube nanoelectrode ensembles. Nano Lett 2004;4:191–5.

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