High enzymatic activity preservation of malate dehydrogenase immobilized in a Langmuir–Blodgett film and its electrochemical biosensor application for malic acid detection

Abstract

In this study, malate dehydrogenase (MDH) enzyme was immobilized on a cation octadecylamine (ODA⁺) monolayer at the air–water interface. Pressure–area (π–A) isotherm studies confirmed that the ODA–MDH system formed a stable monolayer at the air–water interface. The as-prepared MDH–ODA monolayer was transferred onto an indium tin oxide coated glass substrate (ITO) by the Langmuir–Blodgett (LB) method as an MDH–ODA/ITO LB film and characterized using FT-IR, SEM and UV-Vis absorption spectroscopy. The catalytic activity of the enzyme immobilized electrode (MDH–ODA/ITO) was assayed by sensing malic acid (MA) in the range of 10.0–50.0 mM. The high and preserved enzymatic activity of MDH in in vitro media was explored by following the absorbance (A_{340 nm}) of nicotinamide adenine dinucleotide (NADH). Moreover, the highly sensitive electrochemical biosensor behavior of the MDA–ODA/ITO electrode for MA detection was displayed by cyclic voltammetry studies. The electrochemical studies revealed that a voltammetric current from the MDA–ODA/ITO electrode was obtained, while the ODA/ITO electrode did not show this current response. The MDA–ODA/ITO ​demonstrated sensitive electrochemical sensor ability for quantification of MA in both standard solutions and real samples.

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Introduction

Investigation of optimized architectures for biosensors is still a key challenge. Control over molecular architectures based on nanostructured films is to preserve the catalytic activity of biomolecules such as enzymes. Enzymes serve as centers of each biochemical reaction and their catalytic activities are generally higher than synthetic and inorganic catalysts. The deficiency and higher catalytic activity of enzymes lead to some genetic diseases.^1–4 Therefore, the determination of enzyme activity in vivo and in vitro plays a key role for diagnosis of some diseases.^5,6 However, it is known that the enzyme activity in vitro is lower than that of the enzyme in vivo. Increasing the activity and lifetime of enzymes along with higher stability in vitro acts a key role for the diagnosis and treatment of some diseases. For this purpose, enzymes are immobilized into solid and liquid systems by several techniques, and the as-prepared systems are also used in selective and sensitive biosensor applications. A biosensor is a device that combines a selective and specific biological sensing material such as an enzyme, nucleic acid or antibody with a suitable transducer. Based on the transduction process, biosensors are classified into various categories such as electrochemical, optical, piezoelectric, thermal, and colorimetric. Among them, the electrochemical biosensors are most widely used and successfully commercialized.^7,8

Oxidative decarboxylation of malic acid (MA) is executed by malate dehydrogenase (MDH) (also called malic enzyme) using NAD⁺ as a coenzyme (cofactor) to produce oxaloacetate and NADH.⁹ Sources of NADP–malic enzymes are living organisms like microorganisms (both prokaryotic and eukaryotic), plants, animals and humans.¹⁰ Three types of malic enzyme found in mammals are cytosolic NADP⁺ dependent malic enzyme (c-NADP-ME), mitochondrial NAD(P)⁺ dependent malic enzyme (m-NAD-ME) and mitochondrial NADP⁺ dependent malic enzyme (m-NADP-ME).¹¹ In the present study, the malic enzyme which has a key role in many metabolic processes is immobilized on an ODA monolayer by the Langmuir–Blodgett method. As a result, this first application of an MDH–ODA/ITO-based biosensor will allow a new tool for enzymatic studies and open the way to biomolecular diagnostic applications.

Enzyme immobilization

General requirements for commercial enzymes are effective use of reactants, maximizing catalytic velocity, and longer lifetime.¹² When soluble enzymes are used, several disadvantages such as instability and sensitivity to process conditions compared to the optimal one appear.¹³ To enhance the economic applicability of soluble enzymes in various technological areas, they are generally immobilized onto solid supports.¹⁴ To date, various immobilization materials and methods have been reported in the literature. In these studies, it is highlighted that suitable materials and facile immobilization routes have a crucial role.¹⁵ In an immobilization process, enzymes are physically enclosed while maintaining their catalytic activities, and thus they can be repeatedly used for various applications.¹⁶ There are three types of enzyme immobilization: enzymes bound to a support, entrapment and cross-linking. Herein, an appropriate support binding can be physical, covalent or ionic. Covalent binding, which prevents enzyme leakage absolutely, is stronger than ionic, while physical binding is the weakest to keep the enzyme fixed to the support. In entrapment, enzymes are located in a polymer network without leakage.^9,17 Cross-linking bifunctional reagents are used to prepare carrierless macroparticles. For carrier-free immobilization of enzymes, two approaches are used viz. cross-linked enzyme crystals (CLEC) and cross-linked enzyme aggregates (CLEA). These approaches offer high enzyme activity, high stability and low-cost production.¹⁸ The immobilized enzymes solve several problems like loss of enzyme activity, stability and shelf-life.^19,20

Mechanism and activity

The active site of MDH is a hydrophobic cavity part within the protein complex, which has specific binding sites for the substrate and its coenzyme, NAD⁺. In the active site, it undergoes a conformational change, in which it surrounds the substrate to minimize solvent exposure and to locate key residues in closer proximity to the substrate.²¹ The three residues included in a catalytic trio are histidine (His-195), aspartate (Asp-168) and arginines (Arg-102, Arg-109, and Arg-171). The histidine and aspartate work together as a proton transfer system, while the arginines protect the substrate (Fig. S1, ESI†).²² The related kinetic studies display that the enzymatic activity of MDH is ordered. NAD⁺/NADH is bound before the substrate.²³ The NAD-specific MDH enzyme catalyzes the oxidative decarboxylation of L-malate to oxaloacetate with concomitant reduction of the cofactor NAD⁺ as shown in Fig. S2, ESI.†

In the present study, we report the immobilization of MDH on an indium tin oxide coated glass substrate (ITO) via amphiphilic ODA by the LB method and its high enzymatic activity is preserved in in vitro media as an LB film electrode. The MDH immobilized system is assayed as a sensitive electrochemical biosensor for MA detection.

Experimental

Materials

Malic enzyme (EC 1.1.1.37 from porcine heart), malic acid, sodium phosphate buffer, NAD⁺, octadecylamine (ODA), sodium hydroxide (NaOH) and chloroform were purchased from Sigma-Aldrich. The stock solution of MDH was prepared in 50% glycerol containing 0.05 M potassium phosphate buffer, pH 7.4. ITO was purchased from Sigma-Aldrich.

Apparatus

Absorbance of samples was measured using a Perkin-Elmer (Model Lambda 35) spectrophotometer. Langmuir–Blodgett film applications were executed with a commercially available KSV/Minithrough system. Deionized (DI) water was provided from a KrosClinic (KRS-R-75) system. The subphase pH was adjusted to 7.4 and its temperature kept constant at 18 °C. A Zeiss EVO40 operating at an acceleration voltage of 15 kV was used to obtain SEM (Scanning Electron Microscope) images. Fourier transform infrared (FT-IR) spectra were collected with a Spectrum One model Perkin-Elmer spectrophotometer by using a specular reflectance attachment.

Determination of enzyme activity

The activity of MDH (EC 1.1.1.37) was determined by following the absorbance at 340 nm for NADH as oxaloacetate reduction or malate oxidation (ε = 6.22 mM⁻¹ cm⁻¹).²⁴ All assays were performed in 50 mM potassium phosphate buffer, pH 7.4, 15 mM L-malate, 2.0 mM NAD⁺ and the immobilized enzyme. The reaction was started by the immersion of MDH immobilized on LB film into the solution.

Preparation of MDH–ODA/ITO LB electrode

For LB film applications, the KSV minitrough was repeatedly cleaned with ethanol and DI–water. Initially, the minitrough was filled with DI–water (pH = 7.4) and the subphase pH was adjusted with NaOH. Stable and floating layers of ODA and MDH–ODA at the air–water interface were confirmed by surface pressure–area (π–A) isotherms. The π–A isotherms studies proved that the stable and floating layers of ODA and MDH–ODA at the air–water interface were formed.²⁵ To prepare MDH–ODA/ITO LB films, 50 μL of ODA (0.5 mg mL⁻¹) in chloroform was spread on the subphase and then the ODA layer was compressed until 35 mN m⁻² surface pressure after allowing 10 min to evaporate the solvent. Afterwards, 600 μL of the MDH solution (0.5 EU/600 mL) was added into the subphase via a micro-syringe to immobilize the enzyme on the ODA monolayer at the air–water interface, by holding the surface pressure at 35 mN m⁻¹. Herein, it was observed that the protonation of ODA (as ODA⁺) based on the subphase pH increased the surface pressure due to the interaction of ODA⁺ with the negatively charged MDH.^26–29 To complete this interaction, it was left for 20 min. The monolayer was transferred onto the ITO at a surface pressure of 35 mN m⁻¹ by a dipping speed of 5 mm min⁻¹. Thus, the formation of the Y-type LB film on ITO as the MDH–ODA/ITO electrode is schematized in Scheme 1. The quality of the as-prepared LB film was controlled by analyzing the transfer ratio. It was calculated to be an average of 1.05.

Scheme 1 Preparation of MDH–ODA/ITO LB film electrode.

Electrochemical studies

Electrochemical experiments of ODA/ITO and MDH–ODA/ITO electrodes were performed using a Gamry potentiostat system connected to a three electrode cell. In the electrochemical studies, MDH–ODA/ITO electrodes performed as the working electrode for the biosensor studies. Effective electrode surface area was calculated using a cyclic voltammetry study with the prepared electrodes performing as the working electrode for a reversible Fe(CN)₆³⁻/Fe(CN)₆⁴⁻ redox system.³⁰ In all cases, an Ag/AgCl (3 M NaCl) (Bioanalytical Systems) electrode served as the reference electrode and a Pt wire was used as the counter electrode. The solutions were deoxygenated by passing dry nitrogen through the electrochemical cell for at least 15 min prior to each electrochemical study. Real sample analysis was performed for determination of MA concentration in an apple juice sample by using a standard addition method. For this purpose, an apple juice sample was collected from a local shop and used without pretreatment.

Results and discussion

The MDH enzyme was immobilized on an ODA monolayer at the air–water interface via a simple adsorption process, and the dual system was deposited on ITO by the Langmuir–Blodgett method as MDH–ODA/ITO. The formation of the MDH–ODA/ITO electrode was characterized using FT-IR, SEM and UV-Vis absorption spectroscopy. Afterward, the enzymatic activity of MDH immobilized on the electrode was explored and its electrochemical biosensor feature was tested in MA detection.

FT-IR studies

The binding of MDH on the ODA⁺ monolayer at the air–water interface was examined using FT-IR studies. Fig. 1 shows FT-IR spectra of ODA/ITO and MDH–ODA/ITO electrodes. As seen in Fig. 1, the peak corresponding to the primary amine group (–NH₂) of ODA in the ODA/ITO electrode was observed at 3331 cm⁻¹. After the immobilization of MDH on ODA, it shifted to 3496 cm⁻¹ for MDH–ODA/ITO. This means that the free amine groups are not present for the MDH–ODA/ITO electrode. The peaks observed at 2917 cm⁻¹ and 2851 cm⁻¹ for ODA/ITO were associated with C–H stretching vibrations of CH₂ and CH₃ groups, respectively. They were shifted to 2968 cm⁻¹ and 2895 cm⁻¹ for MDH–ODA/ITO. Consequently, the FT-IR studies confirmed that the MDH enzyme was attached onto the ODA monolayer as the MDH–ODA system.

Fig. 1 FT-IR spectra of ODA/ITO and MDH–ODA/ITO electrodes.

SEM studies

A morphological analysis of a thin film is readily performed using an SEM instrument. Therefore, the surface morphologies of ODA/ITO and MDH–ODA/ITO electrodes were explored using SEM images. Fig. 2 shows SEM images of ODA/ITO and MDH–ODA/ITO LB films with various magnifications.

Fig. 2 SEM images of (a) ODA/ITO and (b) MDH–ODA/ITO electrodes at different magnifications.

At a glance, the film morphology obviously changed with the adsorption of MDH onto the ODA monolayer. As seen from Fig. 2, the fibril-like morphology of the MDH–ODA/ITO electrode was observed, while the surface of the ODA/ITO electrode was composed of hierarchical islands. The varied surface morphology of the LB thin film after the immobilization process proved the immobilization of the MDH enzyme onto the ODA monolayer.

Enzyme activity of immobilized MDH

In industrial applications, several problems may be encountered in reactions using free enzymes. The recovery of enzymes is often very difficult for their reuse. Immobilized enzymes present many advantages such as higher catalytic activity and stability, reusability and low-cost application processes.³¹ The most simple immobilization route is nonspecific adsorption processes based on physical adsorption and ionic interactions.^32–34 In physical adsorption, hydrogen bonding, van der Waals forces and hydrophobic interactions play a key role to bind a matrix.³⁵ This binding does not influence the active sites of the enzyme and thus its activity is maintained.³⁶ In non-covalent immobilization, the binding strength is changed depending on reaction conditions comprising pH, ionic strength, temperature and solvent polarity. Adsorption-based immobilization supplies many advantages such as easy application and maintaining enzyme activity. In contrast, this type of method has some disadvantages such as removing the enzyme from the matrix due to the weak interaction.³⁷ In this study, the enzyme activity of the MDH–ODA/ITO electrode was monitored using UV-Vis absorption spectroscopy. The MDH-catalyzed NAD⁺/NADH reaction was followed with the absorbance at 340 nm at various time periods (Fig. 3). After the fabrication of the MDH–ODA/ITO electrode, the activity measurement was executed at the 1^st, 3^rd, 5^th and 7^th minutes by 1 min immersion of the electrode into the solution containing the enzyme substrate.

Fig. 3 The time dependent-activity of the MDH–ODA/ITO electrode.

As seen from Fig. 3, it was clearly observed that the enzymatic activity of MDH increased depending on the immersion time from 1 min to 7 min. During the time dependent measurement, the penetration of the immobilized MDH enzyme into the solution was checked using the absorption measurement at 340 nm after removal of the electrode from the solution (Fig. 4). After removal of the electrode for 1, 3, 5 and 7 min, the enzymatic activity of MDH–ODA/ITO was measured within 1 min every 20 seconds (0, 20, 40 and 60 seconds) from the solution. As seen from Fig. 4, the measured enzymatic activity did not change for the time period studied, where the absorbance at 340 nm was nearly the same. This strongly confirmed that the immobilized MDH enzyme, catalyzing NAD⁺/NADH, was not released in the solution containing the enzyme substrate when the electrode was dipped. That is, the immobilization process performed by the Langmuir–Blodgett method was successful. To test the stability of the immobilized MDH, the electrodes (MDH–ODA/ITO) were held at +4 °C for one week and their activity measurements were executed. The obtained activity results revealed that the activity of the MDH immobilized on the LB matrix was maintained.

Fig. 4 The enzymatic activities of MDH–ODA/ITO electrode in the solution after its removal.

Electrochemical studies

Electrochemical responses of both ODA/ITO and MDH–ODA/ITO LB film electrodes through MA detection were studied using cyclic voltammetry. Two successive cyclic voltammograms of the MDH–ODA/ITO electrode in 0.1 M phosphate buffer solution (PBS, pH = 7.4) containing 50 mM MA are depicted in Fig. 5a.

Fig. 5 Cyclic voltammograms of MDH–ODA/ITO electrode in 0.1 M PBS (pH = 7.4) with (a) and without (b) 50 mM MA. Cyclic voltammogram of ODA/ITO electrode in 0.1 M PBS (pH = 7.4) with 50 mM MA (c). Scan rate: 50 mV s⁻¹.

Scanning the potential of the MDH–ODA/ITO working electrode from −100 mV to 1100 mV at a scan rate of 50 mV s⁻¹, an anodic peak was observed at 910 mV, which was attributed to irreversible electrochemical oxidation of MA to CO₂ by the following reaction (eqn (1)),^38,39

HO₂CCH₂CHOHCO₂H + 3H₂O → 4CO₂ + 12H⁺ + 12e⁻ (1)

This peak was not observed at the second scan and this case exhibited loss of catalytic activity of MDH on the electrode surface. When this voltammetry approach was applied on the MDH–ODA/ITO electrode in a solution containing 0.1 M PBS without MA, no electrochemical response appeared, which verified that the observed peak at 910 mV was due to the electrooxidation of MA (Fig. 5b). We also performed cyclic voltammetry studies on the ODA/ITO electrode in 0.1 M PBS containing 50 mM MA. This voltammogram is illustrated in Fig. 5c. It is clear that a meaningful electrochemical response is not seen in this voltammogram, which indicates that the electrooxidation of MA does not occur without the enzyme on the electrode surface.

The performance of the MDH–ODA/ITO electrode toward MA was tested in various MA concentrations. Fig. 6 shows the cyclic voltammograms of the MDH–ODA/ITO electrode in 0.1 M PBS (pH = 7.4) with 10, 20, 30, 40, and 50 mM MA at a scan rate of 50 mV s⁻¹ and the calibration curve which is derived from the cyclic voltammograms.

Fig. 6 (a) Cyclic voltammograms of MDH–ODA/ITO electrode in 0.1 M PBS (pH = 7.4) with 10, 20, 30, 40, and 50 mM MA. Scan rate: 50 mV s⁻¹. (b) Calibration curve which is derived from the cyclic voltammograms.

Fig. 6a exhibits cyclic voltammograms of the MDH–ODA/ITO electrode in 0.1 M PBS (pH = 7.4) with various concentrations of MA. It was obvious that the current density of the anodic peak was increased by increasing the concentration of MA. Fig. 6b reveals the calibration curve (current density versus MA concentration), that is generated from the data in Fig. 6a. The anodic current density was linearly proportional to MA concentration in the range of 2.2 to 50 mM with a detection limit of 0.66 mM. The equation for this calibration curve was obtained as J (mA cm⁻²) = 1.83 + 0.04 CMA (mM) with a correlation coefficient of 0.996. The sensitivity of this method was calculated as 0.04 μA mM⁻¹ cm⁻². The principle of the malate biosensor is the oxidative decarboxylation of MA which is carried out by NADP-specific MDH using NADP⁺ as a cofactor to produce pyruvate, CO₂ and NADPH. The immobilization of MDH on ODA by the Langmuir–Blodgett method preserved the catalytic activity of MDH in in vitro media. Therefore, the surface area of the enzyme increased to bind and also act as an electron transfer mediator helping in enhancing the sensor response of the MDH–ODA/ITO electrode, which increased the sensitivity of the designed system.

The quality of the electrochemical sensor for voltammetric determination of MA was compared with that of previously published studies (Table S1, ESI†).^40–43 The results indicate that the MDH–ODA/ITO electrode reveals hopeful results with a very large linear range, acceptable detection limit, and high sensitivity. Moreover, this sensor can perform in neutral pH values, which may be suitable for in vivo studies for possible future applications. An apple juice sample was used for voltammetric determination of MA in real samples. For this aim, a standard addition method was executed by using the prepared MDH–ODA/ITO sensor as the working electrode. The obtained results are demonstrated in Table 1.

Table 1 Determination of MA in apple juice sample by using standard addition method

Added/mM	Expected/mM	Found/mM	Recovery%
0.0	—	4.50	—
10.0	14.50	14.35	98.96
50.0	54.35	54.75	100.74

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The good recoveries for this sample reveal that this electrochemical sensor can be effectively used for the quantification of MA in commercial samples.

Conclusions

In summary, we reported the successful immobilization of MDH on an ODA monolayer by the Langmuir–Blodgett method to prepare the MDH–ODA/ITO electrode and its sensitive electrochemical biosensor performance for MA detection. The formation of the MDH–ODA/ITO electrode was characterized using FT-IR, SEM, UV-Vis absorption and cyclic voltammetry techniques. The spectroscopic data confirmed that MDH on the ODA monolayer was successfully immobilized and the MDH–ODA/ITO electrode was formed in high quality. The high selectivity of the MDH–ODA/ITO electrode was due to MA being electrochemically transformed into oxalic acid through catalysis by the MDH immobilized on the ODA monolayer. The sensing performance data of the MDH–ODA/ITO electrode revealed that there was a good correlation between current response and MA concentration in the range of 2.2 to 50 mM. Consequently, the first application of the MDH–ODA/ITO-based biosensor provides a new tool for enzymatic studies and opens the way to biomolecular diagnostic applications.

Acknowledgements

The financial support by “Atatürk University Scientific Research Project Council (Project No. 2015/23)” is gratefully acknowledged.

Notes and references

1. F. Vogel and A. G. Motulsky, Vogel and Motulsky’s Human Genetics: Problems and Approaches, Springer Science & Business Media, 2013.
2. E. S. Allan, S. Winter, A. M. Light and A. Allan, Gut, 1996, 38, 886–893.
3. U. Muthane, Y. Chickabasaviah, C. Kaneski, S. K. Shankar, G. Narayanappa, R. Christopher and S. S. Govindappa, Mov. Disord., 2004, 19, 1334–1341.
4. L. Goth, A. Shemirani and T. Kalmar, Blood Cells, Mol., Dis., 2000, 26, 151–154.
5. İ. Yılmaz, J. Turgut Ozal Med. Cent., 2010, 17, 143–153.
6. M. Isik, Y. Demir, M. Kirici, R. Demir, F. Simsek and S. Beydemir, Arch. Physiol. Biochem., 2015, 121, 97–102.
7. E. Bakker and Y. Qin, Anal. Chem., 2006, 78, 3965–3983.
8. J. S. Rana, V. Beniwal and V. Chhokar, J. Am. Sci., 2010, 6, 353–375.
9. K. E. Franke and D. O. Adams, Plant Physiol., 1995, 107, 1009–1010.
10. V. Doubnerová and H. Ryšlavá, Plant Sci., 2011, 180, 575–583.
11. G. G. Chang and L. Tong, Biochemistry, 2003, 42, 12721–12733.
12. T. A. Kadima and M. A. Pickard, Appl. Environ. Microbiol., 1990, 56, 3473–3477.
13. P. Dunnill, Immobilized enzymes-research and development, Biochem. Educ., 1979, 7, 50–76.
14. M. I. G. Siso, E. Lang, B. Carreno-Gomez, M. Becerra, F. O. Espinar and J. B. Mendez, Process Biochem., 1997, 32, 211–216.
15. F. Vaillant, A. Millan, P. Millan, M. Dornier, M. Decloux and M. Reynes, Process Biochem., 2000, 35, 989–996.
16. P. J. Worsfold, Pure Appl. Chem., 1995, 67, 597–600.
17. P. S. J. Cheetham, Immobilized enzymes: an introduction and applications in biotechnology, Biochem. Educ., 1981, 9, 119.
18. R. A. Sheldon, Adv. Synth. Catal., 2007, 349, 1289–1307.
19. A. Amine, H. Mohammadi, I. Bourais and G. Palleschi, Biosens. Bioelectron., 2006, 21, 1405–1423.
20. S. Sanchez, M. Pumera, E. Cabruja and E. Fabregas, Analyst, 2007, 132, 142–147.
21. C. R. Goward and D. J. Nicholls, Protein Sci., 1994, 3, 1883–1888.
22. V. S. Lamzin, Z. Dauter, V. O. Popov, E. H. Harutyunyan and K. S. Wilson, J. Mol. Biol., 1994, 236, 759–785.
23. T. B. Shows, V. M. Chapman and F. H. Ruddle, Biochem. Genet., 1970, 4, 707–718.
24. V. H. T. Sukalovic, M. Vuletic, K. Markovic and Z. Vucinic, Plant Sci., 2011, 181, 465–470.
25. B. Gür and K. Meral, Colloids Surf., A, 2012, 414, 281–288.
26. M. Takahashi, K. Kobayashi, K. Takaoka, T. Takada and K. Tajima, Langmuir, 2000, 16, 6613–6621.
27. S. Choudhury, R. Chitra and J. V. Yakhmi, Thin Solid Films, 2003, 440, 240–246.
28. C. Hansda, S. A. Hussain, D. Bhattacharjee and P. K. Paul, Surf. Sci., 2013, 617, 124–130.
29. A. Pal, B. K. Mishra, S. Panigrahi, R. K. Nath and S. Deb, Incorporation of Erythrosin B Within the Restricted Geometry of Langmuir Monolayer and Langmuir–Blodgett Films of Octadecylamine, Soft Mater., 2013, 11, 85–89.
30. K. Dagci and M. Alanyalioglu, ACS Appl. Mater. Interfaces, 2016, 8, 2713–2722.
31. D. Brady and J. Jordaan, Biotechnol. Lett., 2009, 31, 1639–1650.
32. R. A. Messing, Adsorption and inorganic bridge formations, in Methods In Enzymology, Academic Press, New York, NY, 1976.
33. J. Woodward, Immobilized enzymes: adsorption and covalent coupling, in Immobilized Cells and Enzymes: A Practical Approach, IRL, Oxford, UK, 1985.
34. B. M. Brena and F. Batista-Viera, Immobilization of Enzymes and Cells, A product of Humana Press, Springer, 2006.
35. M. Persson, E. Wehtje and P. Adlercreutz, Biotechnol. Lett., 2000, 22, 1571–1575.
36. A. Mulchandam and K. R. Rogers, Methods in Biotechnology, Enzyme and Microbial Biosensors Techniques and Protocols, Humana Press Inc., New Jersey, 1998.
37. C. Mateo, O. Abian, R. Fernandez-Lafuente and J. M. Guisan, Biotechnol. Bioeng., 2000, 68, 98–105.
38. F. Epron, M. J. Chollier-Brym, E. Lamy-Pitara and J. Barbier, Appl. Catal., A, 1999, 176, 221–228.
39. E. Weiss, K. Groenen-Serrano, A. Savall and C. Comninellis, J. Appl. Electrochem., 2007, 37, 41–47.
40. G. Palleschi, G. Volpe, D. Compagnone, E. L. Notte and M. Esti, Talanta, 1994, 41, 917–923.
41. A. Arvinte, L. Rotariu and C. Bala, Sensors, 2008, 8, 1497–1507.
42. V. A. Tarasova, V. N. Kiryushov and R. Y. Bek, J. Anal. Chem., 2003, 58, 565–568.
43. A. Niaz, J. Fischer, J. Barek, B. Yosypchuk, Sirajuddin and M. I. Bhanger, Electroanalysis, 2009, 21, 1719–1722.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra17465e

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