Fabrication of conductive oxidase-entrapping nanocomposite of mesoporous ceria–carbon for efficient electrochemical biosensor

Abstract

A conductive nanocomposite containing an immobilized oxidative enzyme in the pores of mesostructured ceria (CeO₂)–carbon was developed as an efficient electrochemical biosensing platform. The construction of the nanocomposite began with the incorporation of CeO₂ in a carbon matrix by the co-assembly of cerium nitrate, resol, and triblock copolymer via a facile evaporation-induced self-assembly method, which resulted in the formation of mesoporous ceria–carbon (denoted as Meso-CeO₂/C). Glucose oxidase (GOx) was subsequently immobilized in the vacant pores of the Meso-CeO₂/C by using glutaraldehyde crosslinking to prevent enzyme leaching from the matrix. H₂O₂ generated by the catalytic action of GOx in proportion to the amount of target glucose was rapidly converted into hydroxyl radicals by the catalytic activity of CeO₂, which induced subsequent anodic oxidation of Ce³⁺ into Ce(OH)₂²⁺ or Ce(OH)₄ with the anodic current. The constructed Meso-CeO₂/C exhibited higher resolution in electrochemical detection of H₂O₂ than pure mesoporous carbon without ceria owing to the catalytic activity of ceria. The anodic current responses by the nanocomposite containing GOx in Meso-CeO₂/C resulted in a linear increase in the concentration of target glucose (0.25–5 mM), which is suitable to measure the serum glucose, with excellent storage stability of over two months at room temperature. The biosensor also exhibited a high degree of precision and reproducibility when employing real human blood samples. Based on these results, we anticipate that this novel biosensing format can be readily extended to other oxidative enzymes for the detection of various clinically important target molecules.

---

Introduction

Mesoporous materials have become very attractive as potent support matrices in various biotechnological applications such as enzyme immobilization, immunoassays, drug delivery, and biosensors.^1–4 Their distinctive merits include an extremely large surface area and pore volume, which enable accommodation of biomolecules with a much higher loading capacity than other nanostructured forms.^5–10 Of these, ordered mesoporous carbon (OMC) has recently aroused much interest for the development of electrochemical biosensors because of its advantageous characteristics such as inherent high conductivity, low background current, tailored porous structure, and activity for a variety of redox reactions. For example, H₂O₂ could be detected without any mediators through direct electron transfer of a redox protein (hemoglobin) adsorbed on CMK-3 mesoporous carbon.⁵ L-Cysteine oxidation on an OMC-modified electrode was also successfully detected.⁶ After successful immobilization of glucose oxidase (GOx) in several kinds of mesoporous carbon, glucose was electrochemically detected with excellent selectivity and sensitivity.^11–14 These examples demonstrate the potential of mesoporous carbon as an electrode material and efficient host for enzyme immobilization.

Metal oxide nanostructures have recently become important materials in the field of biosensors owing to their high ionic conductivity, capacitive action, catalytic properties, and high isoelectric point. A number of oxide forms of inorganic metals such as zinc, iron, cerium, tin, zirconium, titanium, and magnesium have been found to exhibit interesting nanomorphological, biocompatible, and catalytic properties.¹⁵ Among these, ceria (CeO₂) has been considered as a promising material for the fabrication of mediator-less electrochemical biosensors because of its ability to act as a redox couple.¹⁶ Furthermore, CeO₂ is suitable for tight adsorption of most of the enzymes having an isoelectric point lower than ∼9, which is the isoelectric point of CeO₂, by inducing electrostatic interaction without any further treatment. Moreover, the excellent electrical conductivity makes CeO₂ an attractive matrix for biosensor applications. Based on these features, CeO₂ has already been used to construct electrochemical biosensors to detect H₂O₂ or glucose by coupling it with GOx with excellent selectivity and sensitivity.^16–18

In this work, in order to construct an electrochemical biosensor with the catalytic activity of ceria and the merits of electrically conductive carbon, a mesoporous ceria–carbon (denoted herein as Meso-CeO₂/C) composite was synthesized through a simple “one-pot” assembly method. Typically, to fabricate a metal oxide/mesoporous carbon composite, a metal oxide precursor is backfilled into pre-synthesized mesoporous carbon.¹⁹ However, the “one-pot” approach eliminates many steps, thereby simplifying synthetic procedures.^20,21 The constructed Meso-CeO₂/C exhibited higher resolution for the electrochemical detection of H₂O₂ than pure mesoporous carbon without ceria owing to the catalytic activity of ceria, and glucose could be successfully quantified without any additional mediators by the successful incorporation of GOx in Meso-CeO₂/C, proving its potential in the fabrication of reagentless glucose biosensors.

Experimental section

Materials

Glutaraldehyde (GA), β-D-glucose, GOx, and a sodium silicate solution were purchased from Sigma-Aldrich. 35% H₂O₂ was purchased from Junsei Chemical Co. (Japan). All other chemicals used in this study were of analytical grade.

Synthesis of mesoporous ceria–carbon

Meso-CeO₂/C was synthesized by the assembly of resol and a cerium precursor with the triblock copolymer F127 as a soft template. In a typical process, 1.15 g of F127 was dissolved in 4 mL of ethanol and stirred for 1 h to form a clear solution. After that, the inorganic sources were prepared by dissolving the desired amount of cerium nitrate and resol in ethanol. Here, we assumed that half amount of used resol was converted to carbon after heat treatment; we therefore calculated the amount of cerium nitrate based on the amount of carbon. Subsequently, inorganic sources were added to the triblock copolymer solution and the mixture was additionally stirred for 2 h. The mixed solution was cast on a glass dish for 5–8 h at room temperature, followed by heat treatment at 100 °C overnight. After all of the solvent was evaporated, the resulting sample was heated to 850 °C for 2 h under Ar gas to generate Meso-CeO₂/C.

Characterization of synthetic materials

Transmission electron microscopy (TEM) was performed by using a Jeol EM-2010 microscope (JEOL Co.). N₂ adsorption/desorption isotherms were measured at 77 K using a Micromeritics ASAP 2010 sorptometer (Micromeritics Co.). Small-angle X-ray scattering (SAXS) data were obtained with an apparatus consisting of an 18 kW rotating anode X-ray generator (Rigaku Co. Cu Kα = 0.15418 nm) and a one-dimensional position-sensitive detector (M. Braun Co.). X-ray diffraction patterns were collected with an X'Pert diffractometer (Cu Kα radiation, PANalytical) using an X'Celerator detector (PANalytical). Zeta potential measurements were performed on a Zetasizer (Zetasizer 2000, Malvern, UK).

Immobilization of glucose oxidase in mesoporous ceria–carbon

Meso-CeO₂/C with 20 or 60 wt% loading of CeO₂ [Meso-CeO₂ (20%)/C or Meso-CeO₂ (60%)/C, respectively] (10 mg) was mixed with 1.5 mL of free GOx (10 mg mL⁻¹) in a sodium phosphate buffer (0.1 M, pH 7.0), vortexed for 30 s, sonicated for 10 s, and incubated at room temperature with shaking at 250 rpm. After a 1 h incubation, the samples were briefly washed with aqueous buffer (0.1 M sodium phosphate, pH 8.0) and incubated in the same buffer containing GA (0.1%, w/w) at 200 rpm for 30 min. After GA crosslinking, the samples were washed with phosphate buffer (0.1 M, pH 8.0). Capping of unreacted aldehyde groups was performed in a Tris–HCl buffer (0.1 M, pH 8.0) at 200 rpm for 30 min. After Tris capping, the samples were washed once with phosphate buffer (0.1 M, pH 8.0), twice with sodium phosphate buffer (0.1 M, pH 7.4), and stored at 4 °C in the buffer (0.1 M sodium phosphate, pH 7.4).

The amount of protein leaching from the nanostructure into the supernatant was measured using the bicinchoninic acid (BCA) assay (PIERCE, Rockford, IL) method. The final enzyme loading in the nanocomposite was calculated from the difference between the initial and leached enzyme amounts.

Preparation of glassy carbon electrodes coated with mesoporous ceria–carbon or nanocomposite containing glucose oxidase in mesoporous ceria–carbon

A glass carbon working electrode (GCE) was polished to a mirror-like finish using 0.05 μm alumina slurry and sonicated for at least 5 min in distilled water. The electrode was thoroughly rinsed with methanol and distilled water and subsequently dried at room temperature. 1 mg of Meso-CeO₂/C or nanocomposite containing GOx in Meso-CeO₂/C was subsequently added into 250 μL of chitosan (1% prepared in 0.1 M acetic acid). 7 μL of this mixture was deposited onto the electrode surface and allowed to dry for 30 min at room temperature. To increase the stability of the layer, the electrode was incubated for 30 min with 0.5% GA to crosslink the chitosan. The electrode was thoroughly rinsed with distilled water and stored at 4 °C in 0.1 M sodium phosphate buffer at pH 7.4 until use.

Electrocatalytic measurements

All electrochemical measurements were carried out in sodium phosphate buffer (0.1 M, pH 7.4) at room temperature. A CH Instrument 620B electrochemical analyzer (Austin, TX) coupled with a desktop computer was used for cyclic voltammetry (CV) and amperometry measurements. The measurements were performed with a three-electrode system employing a GCE with a 3.0 mm diameter as the working electrode, a platinum counter electrode, and a silver/silver chloride reference electrode. CV measurements were performed at 0–0.8 V (vs. Ag/AgCl) with a 10 mV s⁻¹ scan rate. For amperometric signal measurements, 0.7 V was applied as the operating potential. After initial current stabilization for 30 s, small aliquots of H₂O₂ and glucose stock solution (1 M) were added at appropriate time intervals to obtain the signal between additions. The glucose stock solutions were allowed to mutarotate overnight before use.

Quantification of glucose in clinical blood samples

Human blood samples, provided by clinical hospital, were diluted for 5-fold with sodium phosphate buffer (0.1 M, pH 7.4) and applied to the CV measurements as described above. The actual concentration of glucose in the clinical blood samples were determined by using enzymatic methods (ADVIA 2400, Siemens Healthcare Diagnostics, USA) according to the manufacturer's instructions and protocols. Reliability and reproducibility of this method were assessed by determining the recovery rate [recovery (%) = measured value/actual value × 100] and the coefficient of variation [CV (%) = SD/average × 100].

Results and discussion

A conductive nanocomposite consisting of GOx in the vacant pores of Meso-CeO₂/C was developed. Meso-CeO₂/C was first induced by a facile evaporation-induced self-assembly method in which a mixture solution containing resol, cerium precursor, and triblock copolymer was cast onto a glass dish and evaporated by heat treatment at 100 °C. During evaporation induced self-assembly, the relatively hydrophilic resol used as a conducting carbon precursor and the cerium oxide precursor interacted with poly(ethylene oxide) part of Pluronic F127 to form ordered mesostructures. Heat-treatment at 850 °C under Ar gas resulted in a Meso-CeO₂/C composite having a highly conductive wall consisting of ceria particles in a carbon matrix, large mesopores, which provided a large surface area, and an interconnected structure, which is required for developing high-performance biosensors. GOx was subsequently immobilized in the vacant pore spaces of the Meso-CeO₂/C through the GA crosslinking method to prevent enzyme leaching during immobilization and to promote the reaction.^7,9,22 This enabled us to achieve a high loading capacity of over 18 wt% as measured by BCA protein assay.²³ We envisioned that the use of nanostructures to entrap GOx in the conductive ceria–carbon matrix would result in an efficient electrochemical biosensor of glucose. Specifically, in the presence of target glucose, the immobilized GOx in the nanostructure would generate H₂O₂ by its catalytic action, which would be rapidly converted into hydroxyl radicals, as previously discussed.^24,25 The radicals from H₂O₂ would induce subsequent anodic oxidation of Ce³⁺ into Ce(OH)₂²⁺ or Ce(OH)₄ mediated by the catalytic activity of CeO₂ (Fig. 1).^17,18

Fig. 1 Synthetic scheme of Meso-CeO₂/C and electrochemical biosensing mechanism of the conductive nanocomposite entrapping GOx in pores of Meso-CeO₂/C.

Three different amounts of CeO₂, 20, 40, and 60 wt%, were loaded when Meso-CeO₂/C was synthesized. TEM investigation revealed that Meso-CeO₂/C showed clear distinctions with different ratios of carbon to ceria, as shown in Fig. 2. Highly ordered hexagonal arrays of mesopores are shown in Meso-CeO₂ (20%)/C (Fig. 2a). The pore size and structure of Meso-CeO₂ (20%)/C are similar to those of OMC synthesized without a ceria precursor because a small amount of hydrophilic ceria does not critically affect the co-assembly process.²⁶ As the amount of ceria increased up to 60 wt%, the highly ordered mesoporous structure was replaced with a randomly oriented mesoporous structure with larger-sized pores. A large amount of cerium precursor easily aggregated during thermal treatment and generated the large ceria particles, which is typical of metal oxides. Therefore, large angular ceria particles could destroy the ordering of mesoporous materials. The SAXS patterns of the ceria–carbon material were investigated in order to obtain more detailed structural information. The SAXS pattern of Meso-CeO₂ (20%)/C has two intense peaks at q = 0.44 and 0.76 nm⁻¹ in the as-prepared condition. After heat treatment, the intensities of the two peaks decreased but did not disappear, indicating that the structural regularity of Meso-CeO₂ (20%)/C was retained even after harsh thermal treatment (Fig. 2c). The SAXS profile of the 60 wt% sample (Fig. 2d) did not display prominent peaks, revealing deficiencies in the structural regulation in both the as-made and post-heat-treated samples, which is in accordance with the TEM images. The regularity of the ceria–carbon composite morphology gradually decreased as the amount of ceria increased. The TEM image and SAXS pattern of Meso-CeO₂ (40%)/C shows that the structure was less ordered than the 20 wt% sample, as seen in Fig. S1.†

Fig. 2 TEM images of (a) Meso-CeO₂ (20%)/C and (b) Meso-CeO₂ (60%)/C. SAXS traces of (c) Meso-CeO₂ (20%)/C and (d) Meso-CeO₂ (60%)/C.

The nitrogen sorption isotherms of Meso-CeO₂/C exhibited typical type IV isothermals with clear capillary condensation at P/P₀ = 0.7–0.8, indicating uniform mesoporosity. A narrow pore-size distribution was obtained, approximately 4 nm and 13 nm for 20 wt% and 60 wt%, respectively, based on the Barrett–Joyner–Halenda (BJH) model. It is essential that sufficient pore volume be retained in order to accommodate GOx in the mesoporous structures; thus the 60 wt% sample, with its larger pore size, is more advantageous. The 20 wt% and 60 wt% Meso-CeO₂/C materials have Brunauer–Emmett–Teller (BET) surface areas of 384.9 m² g⁻¹ and 224.6 m² g⁻¹, respectively, and total pore volumes of 0.25 cm³ g⁻¹ and 0.38 cm³ g⁻¹, respectively (Fig. 3a). Nitrogen physisorption was also carried out on the nanocomposite containing entrapped GOx in Meso-CeO₂ (60%)/C (Fig. S2†). The pore volume of the nanocomposite was determined to be 0.23 cm³ g⁻¹, which is less than 60% of the corresponding Meso-CeO₂ (60%)/C, confirming that GOx molecules are immobilized inside the pore spaces of the Meso-CeO₂ (60%)/C. The presence of GOx on Meso-CeO₂ (60%)/C was also confirmed by the negative shift of zeta potential after the immobilization of GOx molecules, whereas the zeta potential of Meso-CeO₂ (20%)/C did not show any significant change before and after the immobilization procedure (Fig. S3†). This observation clearly indicates that GOx molecules were efficiently immobilized within Meso-CeO₂ (60%)/C having sufficiently large pore size (13 nm) but not within Meso-CeO₂ (20%)/C presumably due to its pore size (4 nm) smaller than the size of GOx.

Fig. 3 (a) The pore size distribution of Meso-CeO₂ (20%)/C and Meso-CeO₂ (60%)/C and their nitrogen adsorption/desorption isotherms (inset). (b) X-ray diffraction patterns of Meso-CeO₂ (20%)/C and Meso-CeO₂ (60%)/C.

The XRD diffraction patterns display the presence of a crystalline phase of CeO₂ (JCPDS: 34-0394). The characteristic diffraction peaks at 2θ = 28.5, 33.1, 47.5, and 56.3° correspond to the (111), (200), (220), and (311) planes, respectively, as shown in Fig. 3b. A typical carbon peak is also observed at approximately 2θ = 26°. The characteristic peaks of ceria are covered by broad carbon peaks for Meso-CeO₂ (20%)/C owing to the highly loaded carbon. The 60 wt% samples exhibit obvious ceria crystalline phases with low-intensity carbon peaks. High resolution TEM images of Meso-CeO₂ (20%)/C and Meso-CeO₂ (60%)/C show that the wall of mesostructured composites is composed of cubic CeO₂ nanocrystals (Fig. S4†). The crystallite size in Meso-CeO₂ (60%)/C is evidently larger than that in Meso-CeO₂ (60%)/C, which is consistent with the XRD result.

The potential of Meso-CeO₂/C as an electrochemical biosensor toward the oxidation of H₂O₂ was investigated. The determination of H₂O₂ is of practical importance in chemical, biological, and many other fields.^27–31 It also holds significant incentives for further applications in various oxidase-coupled biosensors.^32,33 To demonstrate the electrocatalytic oxidation behavior of H₂O₂ by Meso-CeO₂/C, CV experiments were conducted in the presence and absence of 10 mM H₂O₂ (Fig. 4a). When only chitosan-coated GCE was employed in the CV experiments as a control, an increase in current according to the addition of 10 mM H₂O₂ was observed starting from approximately 0.6 V (vs. Ag/AgCl), which is similar to the results for bare GCE without any coating.^34,35 On the other hand, with a coating of Meso-CeO₂/C mixed with chitosan on GCE, faster H₂O₂ oxidation was observed with increasing current starting from approximately 0.3 V, indicating the affirmative catalytic effect of Meso-CeO₂/C on the oxidation of H₂O₂.^17,18 The current peak intensity using Meso-CeO₂/C in the presence of 10 mM H₂O₂ was approximately 10 times higher than that without H₂O₂, proving its potent capability to quantify H₂O₂ concentration with high resolution. MSU-F-C without CeO₂ was also employed in the same CV experiments to see the effect of ceria on the oxidation of H₂O₂ (Fig. S5†).⁷ As a result, H₂O₂ oxidation from MSU-F-C started a little late (∼0.4 V) as compared to Meso-CeO₂/C, and the current peak intensity with 10 mM H₂O₂ was only two times higher than that without H₂O₂, proving that the addition of ceria in mesoporous carbon enhanced the resolution for H₂O₂ determination because of the catalytic activity of ceria. Based on the CV results, the amperometric response of the electrode containing Meso-CeO₂ (60%)/C and Meso-CeO₂ (20%)/C obtained at 0.7 V (vs. Ag/AgCl) on sequential additions of 1 mM H₂O₂ resulted in a rapid increase (response time of 3 s) in anodic current (Fig. 4b). Although the initial background current for Meso-CeO₂ (60%)/C was observed to be higher than that of Meso-CeO₂ (20%)/C presumably due to higher conductivity originated from the highly loaded CeO₂,¹⁷ the sensitivities of H₂O₂ detection for Meso-CeO₂ (60%)/C and Meso-CeO₂ (20%)/C were measured to be 198 nA mM⁻¹ and 182 nA mM⁻¹, respectively, indicating that detecting performance for H₂O₂ was not significantly varied from the loading amount of CeO₂. Finally, we demonstrated our strategy by detecting glucose, a vital sugar in human blood at a normal concentration of 3–7 mM,³⁶ employing the Meso-CeO₂/C nanostructure containing immobilized GOx. As demonstrated above, Meso-CeO₂ (60%)/C had a pore size (∼13 nm) large enough to accommodate GOx molecules (∼7 nm), whereas Meso-CeO₂ (20%)/C had a relatively small pore size (∼4 nm) that could not accommodate the corresponding enzyme molecules. Thus, the catalytic action of GOx is expected to occur only with Meso-CeO₂ (60%)/C. As shown in Fig. 5 and S6a,† the anodic current responses obtained rapidly from the cyclic voltammogram at 0.7 V (vs. Ag/AgCl) by employing the nanocomposite entrapping GOx in Meso-CeO₂ (60%)/C resulted in a linear increase in the concentration of target glucose. The dynamic range of this system was from 0.25 to 5 mM, which is suitable to measure the serum glucose in both normal and patients with diabetes after appropriate dilution. The detection limit was approximately 0.1 mM based on a signal-to-noise ratio of 3. The sensitivity was measured to be 50.3 nA mM⁻¹ for the nanocomposite using Meso-CeO₂ (60%)/C. In the case of Meso-CeO₂ (20%)/C, no oxidation current was measured owing to the absence of GOx, as expected (Fig. S6b†). The measured sensitivity and dynamic range of the current biosensor are among the best results reported for glucose biosensing based on nanostructured carbons.^37,38 We further evaluated the selectivity of the biosensor toward the common interfering species for serum glucose detection including acetaminophen (AP), uric acid (UA), ascorbic acid (AA), and dopamine (DOP).³⁹ Their concentrations were set as the real levels present in human serum. As a result, these substances have negligible effects on the current generated from the cyclic voltammogram at 0.7 V, demonstrating high specificity of the current biosensor (Fig. S7†). Long-term stability of the biosensor was also investigated by measuring its sensitivity from 1 to 5 mM glucose using the nanocomposite in Meso-CeO₂ (60%)/C. The current response of the biosensor was revealed to maintain over 90% of the initial current response for 2 months at room temperature, indicating that the current biosensor is very robust for long-term use (Fig. 6).

Fig. 4 (a) Cyclic voltammetry in the presence or absence of 10 mM H₂O₂. (b) Amperometric current response on successive additions of 1 mM H₂O₂. The applied potential was 0.7 V (vs. Ag/AgCl), and the measurement was performed in a sodium phosphate buffer (0.1 M, pH 7.4) with stirring. The inset represents calibration plots from amperometric responses presented in (b).

Fig. 5 The current response curves of Meso-CeO₂/C nanocomposite-based electrodes in samples containing different concentrations of glucose. Insets show the linear calibration plots corresponding to the current responses of different concentrations of glucose. Error bars represent standard deviations of three independent measurements.

Fig. 6 Long-term stability of the biosensor stored at room temperature by measuring the sensitivity for 1–5 mM glucose using the nanocomposite in Meso-CeO₂ (60%)/C.

Encouraged by the above results, we evaluated the diagnostic capability of the nanocomposite system by using two independent human blood samples that contained representative levels of glucose corresponding to the normal and high stage of hyperglycemia (normal; ≤5.6 mM, borderline; 5.6–7 mM, and high; >7 mM).⁴⁰ Blood samples that show the representative levels were selected based on the actual concentrations determined by using enzymatic methods (ADVIA 2400, Siemens Healthcare Diagnostics, USA). As a result, the serum glucose levels were quantified with excellent precisions yielding CVs in a range of 3.1–6.8% and recovery rates of 96–104% (Table 1), verifying the excellent reproducibility and reliability of the method. These observations show that the nanocomposite-based assay system should serve as a promising analytical tool to diagnose high level of glucose in clinical settings.

Table 1 Detection precision of the nanocomposite-based electrochemical biosensor for the determination of glucose levels in blood samples

	Sample 1	Sample 2
a Mean of three independent measurements.b Standard deviation of three measurements.c Coefficient of variation.d Measured value/expected value × 100.
Expected glucose (mM)	3.8	12.6
Average^a (mM)	3.6	13.1
SD^b	0.25	0.4
CV^c (%)	6.8	3.1
Recovery^d (%)	95.8	104.0

---

The key analytical performances of the nanocomposite in mesoporous ceria–carbon to detect H₂O₂ and glucose were summarized and compared with the recently published results based on the catalytic effect of CeO₂ (Table S1†). It can be seen that the present biosensor is the sole electrochemical method that can detect clinical serum with high storage stability for glucose detection.

Conclusions

We developed an efficient electrochemical biosensor by incorporating ceria and carbon in a synthesized mesoporous material with further immobilization of GOx in the vacant pores of the mesoporous material. Ceria showed a catalytic effect on the oxidation of H₂O₂ and enabled rapid detection of H₂O₂. Finally, the glucose biosensor fabricated by the nanocomposite entrapping GOx in mesoporous ceria–carbon showed high sensitivity, storage stability, reproducibility, and reliability with a broad detection range. Based on these results, we anticipate that this novel biosensing format can be readily extended to other oxidative enzymes for the detection of various clinically important target molecules, and hence, can be used to develop convenient and robust biosensors useful for practical applications.

Acknowledgements

This research was supported by the MSIP (Ministry of Science, ICT and Future Planning), Korea, under the “IT Consilience Creative Program” (NIPA-2014-H0201-14-1001) supervised by the NIPA (National IT Industry Promotion Agency). This work was also supported by the Gachon University research fund of 2014 (GCU-2014-0110).

Notes and references

1. A. Dyal, K. Loos, M. Noto, S. W. Chang, C. Spagnoli, K. Shafi, A. Ulman, M. Cowman and R. A. Gross, J. Am. Chem. Soc., 2003, 125, 1684–1685.
2. J. M. Perez, L. Josephson, T. O'Loughlin, D. Hogemann and R. Weissleder, Nat. Biotechnol., 2002, 20, 816–820.
3. X. N. Fang, P. Zhang, L. Qiao, X. Y. Feng, X. M. Zhang, H. H. Girault and B. H. Liu, Anal. Chem., 2014, 86, 10870–10876.
4. I. Willner and E. Katz, Angew. Chem., Int. Ed., 2003, 42, 4576–4588.
5. J. J. Feng, J. J. Xu and H. Y. Chen, Biosens. Bioelectron., 2007, 22, 1618–1624.
6. M. Zhou, J. Ding, L. P. Guo and Q. K. Shang, Anal. Chem., 2007, 79, 5328–5335.
7. J. Lee, J. Kim, H. F. Jia, M. I. Kim, J. H. Kwak, S. M. Jin, A. Dohnalkova, H. G. Park, H. N. Chang, P. Wang, J. W. Grate and T. Hyeon, Small, 2005, 1, 744–753.
8. K. Ariga, A. Vinu, M. Miyahara, J. P. Hill and T. Mori, J. Am. Chem. Soc., 2007, 129, 11022–11023.
9. M. I. Kim, J. Shim, T. Li, J. Lee and H. G. Park, Chem.–Eur. J., 2011, 17, 10699–10706.
10. Y. J. Jiang, L. L. Shi, Y. Huang, J. Gao, X. Zhang and L. Y. Zhou, ACS Appl. Mater. Interfaces, 2014, 6, 2622–2628.
11. D. Lee, J. Lee, J. Kim, H. B. Na, B. Kim, C. H. Shin, J. H. Kwak, A. Dohnalkova, J. W. Grate, T. Hyeon and H. S. Kim, Adv. Mater., 2005, 17, 2828–2833.
12. J. Lee, D. Lee, E. Oh, J. Kim, Y. P. Kim, S. Jin, H. S. Kim, Y. Hwang, J. H. Kwak, J. G. Park, C. H. Shin and T. Hyeon, Angew. Chem., Int. Ed., 2005, 44, 7427–7432.
13. M. I. Kim, Y. Ye, B. Y. Won, S. Shin, J. Lee and H. G. Park, Adv. Funct. Mater., 2011, 21, 2868–2875.
14. J. Lu, J. Ju, X. J. Bo, H. Wang and L. P. Guo, Electroanalysis, 2013, 25, 2531–2538.
15. P. R. Solanki, A. Kaushik, V. V. Agrawal and B. D. Malhotra, NPG Asia Mater., 2011, 3, 17–24.
16. S. Saha, S. K. Arya, S. P. Singh, K. Sreenivas, B. D. Malhotra and V. Gupta, Biosens. Bioelectron., 2009, 24, 2040–2045.
17. C. Ispas, J. Njagi, M. Cates and S. Andreescu, J. Electrochem. Soc., 2008, 155, F169–F176.
18. P. Chaturvedi, D. C. Vanegas, M. Taguchi, S. L. Burrs, P. Sharma and E. S. McLamore, Biosens. Bioelectron., 2014, 58, 179–185.
19. I. Grigoriants, L. Sominski, H. Li, H. Ifargan, D. Aubarch and A. Gedanken, Chem. Commun., 2005, 921–923.
20. C. Jo, J. Hwang, H. Song, D. H. Anh, Y. T. Kim, S. H. Lee, S. W. Hong, S. Yoon and J. Lee, Adv. Funct. Mater., 2013, 23, 3747–3754.
21. J. Hwang, S. H. Woo, J. Shim, C. Jo, K. T. Lee and J. Lee, ACS Nano, 2013, 7, 1036–1044.
22. M. I. Kim, J. Kim, J. Lee, H. Jia, H. B. Na, J. K. Youn, J. H. Kwak, A. Dohnalkova, J. W. Grate, P. Wang, T. Hyeon, H. G. Park and H. N. Chang, Biotechnol. Bioeng., 2007, 96, 210–218.
23. P. K. Smith, R. I. Krohn, G. T. Hermanson, A. K. Mallia, F. H. Gartner, M. D. Provenzano, E. K. Fujimoto, N. M. Goeke, B. J. Olson and D. C. Klenk, Anal. Biochem., 1985, 150, 76–85.
24. M. Das, S. Patil, N. Bhargava, J. F. Kang, L. M. Riedel, S. Seal and J. J. Hickman, Biomaterials, 2007, 28, 1918–1925.
25. R. W. Tarnuzzer, J. Colon, S. Patil and S. Seal, Nano Lett., 2005, 5, 2573–2577.
26. Y. Meng, D. Gu, F. Zhang, Y. Shi, H. Yang, Z. Li, C. Yu, B. Tu and D. Zhao, Angew. Chem., Int. Ed., 2005, 117, 7215–7221.
27. A. Ramanavicius, A. Kausaite and A. Ramanaviciene, Biosens. Bioelectron., 2005, 20, 1962–1967.
28. Y. Usui, K. Sato and M. Tanaka, Angew. Chem., Int. Ed., 2003, 42, 5623–5625.
29. S. K. Ujjain, A. Das, G. Srivastava, P. Ahuja, M. Roy, A. Arya, K. Bhargava, N. Sethy, S. K. Singh and R. K. Sharma, Biointerphases, 2014, 9, 031011.
30. Z. Q. Song, H. C. Chang, W. Q. Zhu, C. L. Xu and X. J. Feng, Sci. Rep., 2015, 5, 7792.
31. I. S. Hosu, Q. Wang, A. Vasilescu, S. F. Peteu, V. Raditoiu, S. Railian, V. Zaitsev, K. Turcheniuk, Q. Wang and M. Li, RSC Adv., 2015, 5, 1474–1484.
32. R. Maidan and A. Heller, Anal. Chem., 1992, 64, 2889–2896.
33. L. Gorton, G. Bremle, E. Csoregi, G. Jonssonpettersson and B. Persson, Anal. Chim. Acta, 1991, 249, 43–54.
34. P. Westbroek and E. Temmerman, J. Electroanal. Chem., 2000, 482, 40–47.
35. H. H. Yang and R. L. McCreery, J. Electrochem. Soc., 2000, 147, 3420–3428.
36. W. Zhilei, L. Zaijun, S. Xiulan, F. Yinjun and L. Junkang, Biosens. Bioelectron., 2010, 25, 1434–1438.
37. L. Zhu, C. Tian, D. Zhu and R. Yang, Electroanalysis, 2008, 20, 1128–1134.
38. M. L. Lozano, M. C. Rodriguez, P. Herrasti, L. Galicia and G. A. Rivas, Electroanalysis, 2010, 22, 128–134.
39. K. Yamamoto, H. S. Zeng, Y. Shen, M. M. Ahmed and T. Kato, Talanta, 2005, 66, 1175–1180.
40. D. R. Nair, M. Sharifi and K. Al-Rasadi, Curr. Opin. Cardiol., 2014, 29, 381–388.

---

Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c5ra17808h

---