Enzymatic electrochemical biosensor for urea with a polyaniline grafted conducting hydrogel composite modified electrode

Abstract

Conducting polymer hydrogels (CPH) are a unique class of materials, which have both the advantageous features of hydrogels and organic conductors. Herein, we report a new urea biosensor in which the working electrode is modified by a CPH, specifically a composite made of conducting electro-polymerized aniline in the non-conducting hydrogel of polyacrylamide and polyvinyl alcohol for immobilization of the enzyme urease. Scanning electron microscopy (SEM) and atomic force microscopy (AFM) studies depict the porous structure of the hydrogel, which favours the high density immobilization of enzymes. Fourier transform infrared spectroscopy (FTIR) studies confirmed the grafting of polyaniline within the matrix. Electrochemical impedance spectroscopy (EIS), differential pulse voltammetry (DPV) and cyclic voltammetry (CV) predict high enzyme loading in the hydrogel. Bode plot analysis with the [FeCN₆]^3−/4− redox couple shows the partial capacitive behaviour of the bioelectrode in the frequency range of 10² to 10³ Hz. Factors, such as scan rate and pH of the electrolyte, were optimized to give the best results. DPV could detect 1.5–1000 μM urea with the lowest detection limit (LOD) of 60 nM and sensitivity of 878 μA mM⁻¹ cm⁻². The relatively low Michaelis–Menten constant (K_m) value of 113 μM indicates the high affinity of the enzyme for urea. EIS depicts an increase in charge transfer resistance (R_CT) with an increase in urea concentration (LOD = 14 μM). The proposed sensor exhibits a very good performance with respect to detection limit, sensitivity, K_m value, preparation method, cost, sample preparation, stability, selectivity and reproducibility. This sensor was successfully applied for the detection of urea in milk, puffed rice, soil and human blood. Statistical analysis of the results shows good agreement with commercial spectrophotometric analysis.

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1. Introduction

Urea is widely distributed in nature and its analysis is of utmost relevance for clinical diagnostics, environmental monitoring and in food chemistry.^1,2 In humans, urea is present in blood and urine. A urea concentration in blood serum that is higher or lower than the physiological range (1.33–3.33 mM) causes renal failure, hepatic failure, nephritic syndrome, cachexia, urinary tract obstruction, dehydration, shock, burns and gastrointestinal problems.³

Urea is used as an adulterant in milk for commercial benefit. It increases the level of fat in milk and at the same time lowers the nutritive value of the milk and may cause a great threat to human health. The cut off limit for the urea level in milk is 11.7 mM.⁴

When used as fertilizer, an excess dosage may lead to pest problems by increasing the birth rate, longevity and overall fitness of certain pests and thus poses major threats to annual food production. Urea may cause a reduction in soil pH and hence reduce its natural fertility which results in the lower productivity of food grains.⁵ A number of methods are available for the detection of urea including e.g. spectrophotometry, calorimetry and fluorimetric analysis.⁶ However, most of these laboratory based procedures need complicated sample pre-treatment steps which restrict on-site monitoring. On the contrary, electrochemical biosensors may overcome these problems since they are reliable, portable, highly sensitive, selective, cost effective and very responsive.

Various matrices, such as conducting polymers,^7,8 non-conducting polymers,⁹ metal oxides,^5,10–12 and metal nanoparticles,¹³ have been developed using several techniques, such as sol–gel,¹⁴ Langmuir–Blodgett,¹⁵ chemical deposition and sputtering,¹⁶ to construct urea biosensors. Recently Khan and co-workers¹⁷ used a transmission electron microscopy (TEM) grid filled with 4-cyano-4′-pentylbiphenyl (5CB) on an octadecyltrichlorosilane-coated glass substrate in aqueous media to construct a urea biosensor. Shalini et al.⁶ developed an amperometric urea biosensor based on the covalent immobilization of urease on an N₂ incorporated diamond nanowire electrode. Verma and Gupta¹⁸ detected urea using a surface plasmon resonance (SPR) based optical fibre multianalyte sensor. Ahmad et al.¹⁹ developed a urea biosensor using vertically aligned zinc oxide nanorods (ZnO NRs) grown on an Ag sputtered glass electrode. The use of sophisticated instruments or accessories, such as TEM, SPR, diamond nanowire electrodes and Ag glass electrodes, makes all these processes costly and also restricts their onsite performance. Tyagi et al.³ used NiO nanoparticles for the development of a urea biosensor, however the Michaelis–Menten constant (K_m) value of 0.34 mM for their system was high. A functionalized multilayered graphene platform based urea sensor was developed by Srivastava et al.²⁰ using dual enzymes, namely urease and glutamate dehydrogenase. Kaushik et al.²¹ used an iron-oxide chitosan nanobiocomposite for urea sensor using the same enzymes. However, the use of dual enzymes made these processes expensive. Some researchers tried to make urea sensors with a poly(N-glycidylpyrrole-co-pyrrole) conducting film,²² and biocompatible zirconia (ZrO₂) for urease immobilization.²³ Jha et al.²⁴ developed a urea biosensor by the entrapment of live microbial cells in an electropolymerized polyaniline (PANI) matrix. It should be noted that traditional PANI did not show a better enzyme loading capacity than its chemically modified form. Also, the potentiometric determination of urea in serum by a polyvinyl alcohol modified electrode has been reported.⁹ Jha et al.²⁵ used a polyvinyl alcohol–polyacrylamide composite membrane for the immobilization of urease in the potentiometric sensing of urea. Gabrovska et al.²⁶ used a nano-rhodium coated polymer membrane composed of chitosan and acrylonitrile for the immobilization of urease to develop a urea biosensor.

Conducting polymer hydrogels (CPH) are unique materials that combine the advantages of both soft hydrogel materials and organic conductors and hence may be used as excellent electrode modifiers. The potential advantages of CPH are the combination of solvation and diffusion, which makes the conducting hydrogel permeable to water-soluble molecules; good biocompatibility of hydrogels, which facilitates the immobilization of biomolecules (e.g. high enzyme loading capacity) and preserves their bioactivity; excellent electronic properties of conducting polymers due to their long π-conjugated backbone, which facilitate rapid electron transfer; and scalable and rapid synthesis and fabrication of CPH, which enable low cost electrochemical biosensor technologies.

In the present study, a urea biosensor using a unique conducting polymer hydrogel matrix is reported, in which the conducting monomer aniline is electropolymerized within a non-conducting hydrogel composed of polyacrylamide (PAM)–polyvinyl alcohol (PVA) to form a conducting PANI grafted PAM–PVA hydrogel membrane (CPHM). We used this CPHM for urease immobilization on the working electrode of a three-electrode electrochemical cell and performed electrochemical measurements (CV, DPV and EIS).

2. Experimental

2.1. Chemicals and electrochemical instruments

The details are given in the (ESI†).

2.2. Preparation of polymer hydrogel membrane

PVA (2.5%) and acrylamide (15%) were mixed in a beaker in 20 mL of 18.2 MΩ deionized (DI) water. The mixture was heated to 75–80 °C with gentle stirring until it turned into a clear solution. Then, 5 μL of 12.5% glutaraldehyde (crosslinker) and 10 μL of potassium peroxodisulfate (initiator, 10 mg in 200 μL DI water) were added to 400 μL of this mixture. An aliquot of 10 μL of this final preparation was dispensed onto a glassy carbon electrode (GCE) and kept at a temperature of 45–50 °C for 25–30 minutes to cure the hydrogel membrane.

2.3. Electrochemical synthesis of conducting polymer hydrogel membrane (CPHM)

The CPHM was synthesized by electropolymerization of aniline from a 1 M HCl solution containing 0.1 M aniline monomer on the hydrogel modified GCE by running 20 cycles from −0.2 V to +0.1 V at a scan rate of 100 mV s⁻¹ [(ESI) Fig. S1†].²⁷ The modified electrode was then immersed in DI water and −0.2 V applied for 35 minutes to remove the unbound aniline monomer and HCl residue remaining on the electrode surface.

2.4. Immobilization of urease

The enzyme urease was covalently immobilized over the glutaraldehyde activated CPHM matrix modified electrode, which was prepared by dipping the membrane modified electrode in 0.1% glutaraldehyde solution for 90 minutes. This was then carefully washed with DI water before the immobilization of urease. 10 μL (10 U) of urease solution in 100 mM phosphate buffer, pH = 7.1, was dispensed onto the glutaraldehyde activated CPHM surface and kept incubated overnight in a humid chamber at 4 °C. Glutaraldehyde acts as a cross-linker to covalently bind with the end amine groups of PANI and urease. The detailed procedure is depicted in Fig. 1.

Fig. 1 Detailed description of the process for the preparation of the CPHM.

2.5. Characterization of polymer composite by AFM, SEM and FTIR analysis

AFM analysis was carried out at room temperature using a Pico plus 5500 ILM AFM (Agilent Technologies, USA) with a piezo scanner maximum range of 9 micrometres (μm). Micro fabricated silicon cantilevers of 225 μm in length with a nominal spring force constant of 21–98 N m⁻¹ were used. Topographic images were observed using the tapping mode of the instrument. The samples were deposited onto freshly cleaved muscovite Ruby mica sheets (ASTM V1 Grade Ruby Mica from MICAFAB, Chennai) for 2–5 minutes and air dried before measurements.

SEM experiments were conducted on an EVO-18 Special Edition-CARL ZEISS (GERMANY) instrument. SEM images of the hydrogel membrane, CPHM and urease immobilized CPHM (UCPHM) were examined. Samples were prepared by freeze-drying at −20 °C and lyophilizing in a vacuum chamber at −110 °C for 12 h. After drying, they were placed in a QUORUM Q 150 TES-QUORUM (UK) sputter coater and covered by a thin platinum film (10 nm). The samples were then fixed in one of the stubs of a 9 stub holder by carbon tape and the 9 stub holder was placed within the microscope. Work distance, spot size, pressure and accelerating voltage were controlled for each sample in order to obtain the best resolution.

FTIR spectroscopy was used to characterize the grafting of PANI within the PAM–PVA composite hydrogel membrane and its interaction with urease. FTIR spectra were recorded on a Thermo Scientific Nicolet iS10 (UK) spectrophotometer.

All experiments were performed in compliance with the relevant laws and institutional guidelines. The institutional committee of the University of Calcutta also approved the experiments. No human or animal handling was done in these experiments. The samples of human serum and urine were collected from a pathological laboratory (Drs Trivedi and Roy diagnostic Laboratory, 93, Park Street, Kolkata-700016, India).

3. Results and discussion

3.1. Mechanism of formation of CPHM and immobilization of urease

The proposed mechanism relating to the preparation of the CPHM and immobilization of urease over the composite film onto the GCE is shown in Fig. 1. Formation of the composite hydrogel membrane was achieved by crosslinking PAM–PVA copolymers²⁸ with glutaraldehyde deposited onto a GCE and subsequently heated at 45 °C. Acetal bridges were formed between the pendant hydroxyl groups of the PVA chains and this gave rise to a PAM–PVA composite hydrogel polymer. The reaction between PVA and acrylamide in the presence of the potassium peroxodisulfate and glutaraldehyde producing PAM–PVA hydrogel is provided in Fig. 2.²⁹ The possible structure of the composite and the formation of the acetal crosslinked copolymer PAM–PVA structure are presented in Fig. 2.²⁹

Fig. 2 Chemistry of the formation of the PAM–PVA hydrogel.

When the aniline monomer was electropolymerized through the hydrogel membrane onto the GCE, the aniline initially became protonated with HCl and propagated to form an intermediate called PANI radical cation. This PANI radical cation formed bonds with either the –OH or –NH₂ groups of the composite hydrogel membrane and this led to the formation of the CPHM on the GCE (Fig. 1). Urease has multi-active sites which are composed of a 3 : 3 (α : β) stoichiometry with a 2-fold symmetric structure. Urease was covalently immobilized over the CPHM matrix using glutaraldehyde as a linker (Fig. 1). The enzyme immobilization mechanism is schematically illustrated in ESI Fig. S2.† The CPHM possessed an abundance of amino groups at the surface of the PANI fibers, and glutaraldehyde acted as a cross-linker. The aldehyde group in one end of glutaraldehyde bound covalently with the end amino group of the PANI and the other aldehyde group of glutaraldehyde bound with the end amino group of urease with the formation of covalent C=N bonds. Thus urease enzyme could be efficiently immobilized on the CPHM matrices.³⁰

3.2. Structural and morphological studies

The morphology of electrodeposited PANI was analyzed by SEM (Fig. 3a), which exhibited a nanofibrous porous rod like structure. Fig. 3b shows the highly porous structure of the PAM–PVA composite hydrogel. Pore sizes were observed to be between 10.50 and 20.42 micrometres (μm), as shown in Fig. 3c. The SEM image in Fig. 3d displays the rough surface of the composite upon grafting of interconnected PANI nanofibers; however, the pores remain the same. The porous and rough structure of the CPHM favors the high density immobilization of urease and penetration of water-soluble molecules. The grafting also imparts conductivity to the composite polymer. The morphology of the CPHM was again changed after immobilization of urease, as revealed in Fig. 3e. Fig. 4a and b show 2D and 3D AFM images of the PAM–PVA hydrogel membrane and CPHM, respectively. From the extracted profile of the 2D images and the corresponding graph in Fig. 4a and b, it could be observed that the CPHM structure is more rough than only the PAM–PVA hydrogel matrix. Thus, the former favors considerable urease loading and the penetration of water soluble molecules.

Fig. 3 SEM images of (a) PANI fibrous rod like structure, (b) and (c) porous PAM–PVA composite hydrogel membrane showing a smooth surface, (d) CPHM showing a rough surface due to the grafting of PANI in the porous PAM–PVA hydrogel, and (e) UCPHM.

Fig. 4 AFM images of (a) porous PAM–PVA polymer hydrogel membrane having a smooth surface, and (b) CPHM having a rough surface which confirms the proper grafting of PANI within the highly porous composite hydrogel membrane.

The FTIR spectra of the PAM–PVA composite hydrogel, CPHM and urease immobilized CPHM (UCPHM) are presented in Fig. 5. The FTIR spectrum of the PAM–PVA composite hydrogel [Fig. 5a] displays bands at 3197.76 cm⁻¹ and 3323.52 cm⁻¹ which might be due to the stretching vibrations of unreacted –OH groups of the PVA polymer. The peak at 2942.22 cm⁻¹ is due to the C–H stretching vibration of –CH₂ groups. The signature peak at 1651.69 cm⁻¹ is for the C=O bending (amide band I), 1605.42 cm⁻¹ is attributed to N–H in-plane bending in the –CONH₂ group (amide band II) and 1423.47 cm⁻¹ corresponds to C–N stretching (amide band III) for PAM and this confirms the proper synthesis of the PVA–PAM composite hydrogel membrane.³¹ Proper grafting of PANI within the hydrogel membrane and formation of the CPHM was confirmed by the presence of a characteristic band at 1505.83 cm⁻¹ [Fig. 5b] which is due to the C=N stretching vibration mode of the quinonoid structure of PANI. The peak at 1446.05 cm⁻¹ is due to the C=C stretching vibration mode of the benzenoid structure of PANI and the band at 1119.17 cm⁻¹ is assigned to the vibration mode of the –NH⁺= structure formed during protonation of PANI.²³ By comparison with the spectrum of the PAM–PVA hydrogel membrane in Fig. 5a, the absorption peak at 1651.69 cm⁻¹ (corresponding to the νC=O group, amide band 1) became broader and shifted to 1650.52 cm⁻¹ for the UCPHM GCE in Fig. 5c. The peak at 1605.42 cm⁻¹ (attributed to N–H in-plane bending in the –CONH₂ group, amide band II) disappeared, and also the peak at 1423.47 cm⁻¹ (corresponding to C–N stretching, amide band III) shifted to 1413.90 cm⁻¹ for the spectrum of UCPHM shown in Fig. 5c. All the above mentioned findings confirm the proper attachment of urease within the CPHM. The peak broadening at 3323.52 cm⁻¹ in Fig. 5a to 3279.29 cm⁻¹ in Fig. 5c was due to the addition of N–H stretching vibration due to the immobilization of urease onto the CPHM. The presence of peaks at 1547.89 cm⁻¹ (corresponding to the C=N stretching vibration mode of the quinonoid structure of PANI) and 1450.86 cm⁻¹ (corresponding to the C=C stretching vibration mode of the benzenoid structure of PANI) in Fig. 5c confirm the presence of PANI within the UCPHM.

Fig. 5 FTIR spectra of (a) PAM–PVA composite polymer hydrogel membrane having a smooth surface, (b) CPHM, and (c) UCPHM.

3.3. Effect of pH

To optimize pH for the UCPHM coated GCE, observations by CV (−0.4 V to 0.4 V at a scan rate of 30 mV s⁻¹) were considered at various pH (5–8) for 5 mM urea in 100 mM phosphate buffer [inset (b) in ESI Fig. S3†]. An optimum current was observed at pH 7.1 and hence this was chosen as the optimum pH for all electrocatalytic experiments.

3.4. Effect of scan rate

In order to analyze the bioelectrocatalytical process, cyclic voltammograms were recorded for the UCPHM-GCE at varying scan rates of 10 mV s⁻¹ to 100 mV s⁻¹ in 100 mM phosphate buffer, at pH = 7.1 containing 5 mM urea, which are shown in ESI Fig. S3.† The anodic (I_pa) as well as cathodic (I_pc) peak currents showed linear behaviour with the square root of the scan rate (ν^1/2) according to eqn (1) and (2) [inset (a) ESI Fig. S3†] which suggests that the deprotonation reaction of PANI by ammonium hydroxide, a product of the hydrolysis reaction of urea by the enzyme urease on the surface of the CPHM-GCE, followed a diffusion controlled electron transfer process.⁵ The anodic peak potential is +0.14 V and the cathodic peak potential is +0.081 V, and the change in the anodic peak current is less than that of the cathodic peak.³²

I_pa = 2.13 μA (s mV⁻¹)^1/2 × [scan rate (mV s⁻¹)]^1/2 − 3.01 μ(A) (1)

with the regression coefficient value of 0.961,

I_pc = −2.51 μA (s mV⁻¹)^1/2 × [scan rate (mV s⁻¹)]^1/2 + 2.61 μ(A) (2)

with the regression coefficient of 0.997.

The surface concentration (I*) of the ionic species per unit surface area of the UCPHM bioelectrode was estimated using the Brown–Anson model³³ equation:

I_pa = n²F²I*Aν/4RT (3)

where n is the number of electrons transferred (1), F the Faraday constant (96 485 C mol⁻¹), R the gas constant (8.314 J mol⁻¹ K⁻¹), A the surface area of the bioelectrode (0.07065 cm²), ν is the scan rate in V s⁻¹, and T is the temperature (298 K). From the slope of the plot of anodic peak current vs. scan rate, the estimated surface concentration of the ionic species (I*) on the UCPHM bioelectrode was 2.40 × 10⁻⁹ mol cm⁻², which is a higher value than that reported for urea biosensors using other matrices.³⁴ I* is the surface concentration of the active species involved in the redox reaction, as depicted in eqn (4) and (5). In this case it is the concentration of NH₄⁺ and OH⁻ (details are discussed in Section 3.6.1.). This high value clearly indicates large numbers of redox species are available in the bioelectrode for the redox reaction.

3.5. Electrochemical characterization studies using EIS, CV and DPV analysis

CPHM has a better urease loading capacity than the PANI matrix. This fact could be supported by the EIS, DPV and CV studies. In all the experiments with CPHM and the PANI matrix modified electrodes, 10 μL (10 U) of urease was immobilized onto the modified electrode.

The details of the electrochemical characterization using EIS are given in the (ESI†).

DPV experiments were performed with a three-electrode system in 1 mM [Fe(CN)₆]^3−/4− in 10 mM KCl in the potential range of 0.4 to −0.4 V at a scan rate of 30 mV s⁻¹. Fig. 6a shows the DPV of the UCPHM-GCE (curve a) and urease immobilized PANI (UPANI) GCE (curve c). The decrease in electron transfer in the UCPHM GCE (Fig. 6a, curve a) compared to the only urease immobilized PANI modified GCE (Fig. 6a, curve c) is due to the better binding of urease and hence greater electrostatic repulsive force between the redox probe of [Fe(CN)₆]^3−/4− ions and the negatively charged urease. The decrease in electron transfer from the CPHM GCE [Fig. 6a, curve b] to UCPHM GCE [Fig. 6a, curve a] confirms the proper immobilization of urease onto the CPHM.

Fig. 6 (a) DPV of (a) UCPHM GCE, (b) CPHM and (c) UPANI GCE in 1 mM [Fe(CN)₆]^3−/4− and 10 mM KCl in the potential range of 0.4 to −0.4 V at a scan rate of 30 mV s⁻¹. (b) CV of (a) UCPHM GCE (b) UPANI GCE in 1 mM [Fe(CN)₆]^3−/4− and 10 mM KCl in the potential range of −0.4 to 0.4 V at a scan rate of 30 mV s⁻¹.

CV studies were carried out using the enzyme modified CPHM GCE and PANI GCE with Fe[(CN)₆]^3−/4−, as above, in the potential range of −0.4 to 0.4 V at the scan rate of 30 mV s⁻¹ [Fig. 6b]. A decrease in electron transfer was observed in the CPHM GCE [Fig. 6b, curve a] compared to the PANI modified GCE [Fig. 6b, curve b]. This was obviously due to the greater electrostatic repulsive force between the redox probe of [Fe(CN)₆]^3−/4− ions and the negatively charged urease in the case of the CPHM GCE compared to the PANI modified GCE, which also proves the better urease binding capacity of the CPHM GCE than the PANI GCE. The obtained results are in agreement with the EIS and DPV studies discussed earlier.

3.6. Electrochemical sensing response

3.6.1. DPV analysis. Fig. 7 shows the DPV [(+0.4 V) to (−0.4 V) working potential range, 30 mV s⁻¹ scan rate] obtained for the urease immobilized CPHM bioelectrode as a function of urea concentration in 100 mM phosphate buffer solution, at pH = 7.1. It should be noted from Fig. 7 that the peak reduction current gradually increased with an increase in urea concentration from 0 μM to 1 mM.

Fig. 7 Electrochemical response of the UCPHM GCE as a function of urea concentration in the range of 0 to 1000 μM in 100 mM phosphate buffer solution, at pH = 7.1, and the corresponding calibration curve is shown in the inset as (a–c).

The biosensor response without the presence of the analyte (urea) was also investigated in a control experiment, in which a sharp peak was obtained at −0.05 V. This could be attributed to the de-doping nature of the PANI emeraldine salt (most conducting form) in the buffer system. We conducted control experiments with only CPHM and PANI electrodes in the presence of urea along with UCPHM in buffer, and the result is shown in ESI Fig. S5.† ESI Fig. S5† predicts that the CPHM and PANI electrodes alone have no effect on urea, which therefore proves that there were no discrepancies in the response of these control experiments.

The enzymatic reactions involved in the hydrolysis of urea are shown in the following equations:³⁵

PANI is a pH-sensitive conducting polymer. PANI exists in various forms, which differ in chemical and physical properties. The electroactivity and the electronic conductivity of PANI are strongly dependent on its electrochemical state and solution pH.³⁶ The conductivity of PANI is sufficiently high in acidic solution but becomes weaker when the solution pH increases. Thus in alkaline solution, PANI becomes electro-inactive and non-conducting. The hydrolysis of urea by urease produced ammonium hydroxide (NH₄⁺, OH⁻, eqn (4)) which increased the solution pH and hence protonated PANI (e.g., PANI hydrochloride or green PANI emeraldine salt) was converted to a non-conducting blue emeraldine base (neutral form) (eqn (5)). It is due to this deprotonation or dedoping reaction of PANI,³⁷ i.e. gain of electron of PANI, PANI⁺ → PANI, that the flow of the reduction current would be induced.³⁸ There is an increase in the reduction peak current in the DPV for the UCPHM bioelectrode with an increase in urea concentration from 0 to 1000 μM in 100 mM phosphate buffer solution, at pH = 7.1. The observed increase in the reduction peak current an increase in the concentration of urea (Fig. 7) could therefore be attributed to the increase in the number of accepted electrons during the reduction of PANI. Fig. 7 shows the variation in peak reduction current with urea concentration for the UCPHM.

Good linearity was obtained for urea concentrations in the range of 1.5–6 μM, which is represented by the linear regression equations: current (μA) = 0.062 × urea concentration (μM) + 0.004, R² = 0.992 [inset (a)]; 6–100 μM with current (μA) = 0.007 × urea concentration (μM) + 0.333, R² = 0.983 [inset (b)]; and 100–1000 μM with current (μA) = 0.001 × urea concentration (μM) + 0.636, R² = 0.987 [inset (c)]. It was also observed that with an increase in the concentration of urea the reduction peak potential of PANI shifted towards a higher potential.³⁹ The linear enhancement in the peak reduction current with an increase in urea concentration suggests that the CPHM bioelectrode provides a favourable microenvironment to the immobilized enzyme.

Although in real samples (e.g. cow milk, puffed rice, blood serum and urine) urea is present in higher concentrations (generally in mM levels), we performed the test in the low concentration range by diluting the samples in order to get high accuracy. Most importantly, only a drop of blood would be needed to do the test as compared to several mL of blood in conventional procedures.

The sensitivity of the UCPHM based biosensor was 878 μA mM⁻¹ cm⁻². This was calculated by evaluating the ratio of the slope of the calibration curve (i.e. in the concentration range of 1.5–6 μM) to the surface are of the electrode. Due to its high sensitivity, a very small amount of sample was needed and the actual concentration could be evaluated by multiplying by the dilution factor (as shown in Table 1).

Table 1 Real sample analysis

Sample no. and description	Urea concentrations detected by the proposed biosensor method (mean ± std dev) (n = 4)	Urea concentrations detected by a spectrophotometric method (mean ± std dev) (n = 4)
a Higher ranges detected via the dilution of samples.
01 (milk sample)	(10 ± 0.16)^a mM	(11 ± 0.82)^a mM
02 (puffed rice ∼85 mg, 5 grains soaked in buffer)	(3 ± 0.08)^a mM	(3.5 ± 0.29)^a mM
03 (soil seepage sample)	(625 ± 0.24)^a mM	(626 ± 0.41)^a mM
04 (human urine sample)	(120 ± 0.41)^a mM	(121 ± 0.41)^a mM
05 (human blood sample)	(4 ± 0.08)^a mM	(4.65 ± 0.29)^a mM

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The lowest detection limit (LOD) and limit of quantification (LOQ) for the bioelectrode were obtained using the following expressions:

LOD = (3 × SD)/S (6)

LOQ = (10 × SD)/S (7)

where SD is the standard deviation in the peak reduction current of the blank and S is the slope of the calibration plot for the urea concentration range of 1.5–6 μM [Fig. 7, inset (a)].

The LOD and LOQ of urea were found to be 60 nM and 199 nM, respectively, for the UCPHM bioelectrode. All experiments were carried out in triplicate on different electrodes fabricated under similar conditions and the obtained results revealed that the reproducibility of the system was under 3% error. The apparent Michaelis–Menten constant (K_m) gives an idea of the enzyme–substrate kinetics and it is obtained from the Lineweaver–Burk plot (1/I versus 1/[C]),²² which is given in the [ESI Fig. S6†]. The response time of the UCPHM bioelectrode was found to be about 10 s.

The value of K_m is highly dependent on the nature of the matrix, which often exhibits favourable conformational changes in the immobilized enzyme.⁴⁰ In the present work, the estimated value of K_m for the UCPHM-GCE (113 μM) was much lower than the reported values by other researchers (Table 2). This low value of K_m advocates high catalytic efficiency at low substrate concentrations and hence depicts fast biochemical reaction kinetics. Thus, the CPHM based matrix offers a better platform for enzyme immobilization for the biosensing of urea.

Table 2 Comparison of the performance of the proposed sensor with other urea sensors published in the literature

Transducer	Urease immobilization matrix	Linear detection range (mM)	Technique	Detection limit (μM)	Sensitivity (μA mM^−1 cm^−2)	Km (μM)	Shelf-life (days)	Ref.
(1) Indium-tin-oxide (ITO) electrode	Iron oxide–chitosan nanobiocomposite	0.83–6.65	DPV	831	12.5	560	56	21
(2) Platinum electrode	Rhodium nanoparticle incorporated chitosan grafted acrylonitrile copolymer membrane	1.60–8.20	Amperometry	500	3.19	52 600	10	26
(3) Platinum electrode	Electrodeposited rhodium incorporated acrylonitrile–methylmethacrylate–sodium vinylsulfonate copolymer membrane	0.1–1.75	Amperometry	50	1.85	6500	27	41
(4) Diamond nanowire electrode	N2 incorporated diamond nanowire film	1.66–16.63	Amperometry	643	37.2	—	<30	6
(5) Indium-tin-oxide (ITO) electrode	Zinc oxide–multiwalled carbon nanotubes hybrid nanocomposite	0.8–16.63	Cyclic voltammetry	230	43	850	>112	5
(6) Indium-tin-oxide (ITO) electrode	Functionalized multilayered graphene platform	1.66–16.63	Amperometry	647	32	—	40	20
(7) Pencil graphite disk electrode	Polyaniline–multiwalled carbon nanotube composite	0.01–10	Amperometry	40	120	2020	12	43
(8) ITO glass electrode	Pt–Rh multiwalled carbon nanotube composite	0.05–20	Amperometry	50	1.7	—	20	44
(9) Carbon electrode	Copper–polyaniline nano-composite	0.001–0.125	Amperometry	0.5	112	139	—	45
(10) Glassy carbon electrode	PANI grafted PAM–PVA conducting hydrogel membrane	0.0015–1	DPV EIS	0.060	878	113	60	Present work

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3.6.2. EIS analysis. The impedimetric response of the UPCHM bioelectrode was measured as a function of urea concentration. Fig. 8a shows the Nyquist plots for the UCPHM GCE in the presence of different concentrations of urea, which were obtained in pH 7.1 phosphate buffer in the frequency range of 0.08 Hz to 10⁵ Hz at a sample potential of −0.004 V (vs. Ag/AgCl) with an AC amplitude of 5 mV. The corresponding fitted Randles equivalent circuit for the respective Nyquist plot could be represented as [R_S(R_CTCPE)Z_W] (inset of Fig. 8a). To characterize the performance of the biosensor, R_CT values were plotted against urea concentrations (Fig. 8b). It could be seen that the charge transfer resistance value (R_CT) value increased with an increase in urea concentration (0–100 μM).

Fig. 8 (a) Nyquist plot of UCPHM as a function of urea concentration in the range of 0 to 100 μM in 100 mM phosphate buffer solution, at pH = 7.1 in the frequency range of 0.08 Hz to 10⁵ Hz at a sample potential of −0.004 V (vs. Ag/AgCl) with an AC amplitude of 5 mV, and the corresponding best fitted Randle's circuit is shown in the inset. (b) Variation of R_CT with increasing concentrations of urea (0 to 100 μM) in 100 mM phosphate buffer solution, at pH = 7.1, (c) Bode phase angle plot of UCPHM as a function of urea concentrations in the range of 0 to 100 μM and (d) Bode plot of UPCHM as a function of urea concentration in the range of 0 to 100 μM.

The calibration curve obtained for the UCPHM bioelectrode was fitted to the linear regression equation as follows:

R_CT (kΩ) = 0.021 × urea concentration (μM) + 15.1; R² = 0.942 (8)

The LOD for this process was calculated to be 14 μM of urea using eqn (8).

The variation of the capacitive behavior of the UCPHM GCE with the change in urea concentrations in phosphate buffer, at pH = 7.1, could be explained by the Bode phase angle diagram (Fig. 8c). The frequency range of 10 to 10³ Hz (log f = 1 to log f = 3) produced the highest phase angle value by the UCPHM GCE (50.4°) with 100 μM urea. Also, the phase angle of the UCPHM GCE gradually increased with an increase in urea concentration (0–100 μM), which signifies the increase in capacitive behaviour within this frequency range. Fig. 8d shows the Bode plot of the UCPHM GCE in varying urea concentrations. The different positions of the Bode plot corresponding to Z_W, R_CT, C_dl and R_S are shown in Fig. 8d. Thus with increasing concentrations of urea, the dominant C_dl value was observed in the frequency range of 10 to 10³ Hz in the case of the UCPHM GCE.

3.7. Interference studies

The effect of interfering substances on the prepared bioelectrode was tested via the DPV technique. In this experiment, the effects of a number of metabolites that are present in human blood and also animal fluids (e.g. cow milk), such as cholesterol (5 mM), glucose (5.6 mM), uric acid (0.1 mM), ascorbic acid (0.05 mM),²¹ lactose (3–7.2%), Na⁺ (45%), and K⁺ (6.79 mg in 5 mL milk), which might interfere with the response were studied. The selectivity of the bioelectrode was carried out by monitoring the DPV obtained after the addition of various interfering substances, separately, to 1 mM urea solution in 100 mM phosphate buffer (pH = 7.1). The amount of interference was calculated according to eqn (9):

Interference (%) = (|I_i − I_u|/I_u) × 100 (9)

where I_i and I_u are the peak reduction currents recorded for the mixed analyte (interfering substance + urea) and urea alone, respectively. The peak reduction currents obtained after the addition of various interfering agents are shown the bar graph in Fig. 9.

Fig. 9 Effect of interferences on the electrochemical response of the UCPHM GCE bioelectrode.

The obtained results clearly demonstrate that the presence of common interfering substances has a negligible effect on the biosensing response of the UCPHM-GCE bioelectrode for urea with only 1.65% and 2.48% of interference from Na⁺ and K⁺, respectively. It might be inferred that the prepared bioelectrode is highly selective for the measurement of urea.

3.8. Reproducibility and storage stability of the proposed sensor

The reproducibility of the UCPHM-GCE urea biosensor was investigated by measuring the DPV responses of three independent sets of modified electrodes for 500 μM urea by four repeat measurements each. The peak current was determined four times with the same electrode (within the batch) and three different electrodes (between batches). The proposed sensor showed acceptable reproducibility in terms of its relative standard deviation of 0.33%, 0.84% and 0.64% (within batches) and 0.32% (between batches) with 500 μM of urea (ESI Tables 1 and 2†), indicate that the senor demonstrates good reproducibility.

The stability of the prepared sensor was evaluated by storage in clean and dry conditions at 4 °C in a refrigerator. Periodical (once a week) measurement of the electrode current density for 1 mM urea was conducted (Fig. 10) to assess the stability of the electrode. The UCPHM-GCE showed a little decrease in response up to a period of one month. The UCPHM bioelectrode could retain 97% and 94% of its initial enzyme activity after 2 weeks and 2 months, respectively, when stored at 4 °C in a refrigerator. This long-term stability of the UCPHM bioelectrode is either comparable with or higher than the reported biosensors.^41,42

Fig. 10 Stability of the UCPHM GCE (fresh, after 2 weeks and after 2 months) in the presence of 1 mM urea in 100 mM phosphate buffer solution, at pH = 7.1.

3.9. Reusability

The reusability of a single UCPHM GCE was determined by comparing the DPV responses of 6 successive measurements of 1 mM urea using the same electrode. The % R.S.D was evaluated as 0.26%. Thus, the proposed urea sensor shows excellent reusability for the same electrode.

3.10. Real sample analysis

To demonstrate the applicability of the developed UCPHM-GCE for the analysis of urea in real samples, such as cow milk, puffed rice (collected from local market), human blood serum and urine (collected from pathological laboratory) samples were analysed. DPV analysis was performed to obtain the peak reduction current and the concentration of urea peak reduction was evaluated from the calibration curve. The cow milk sample was skimmed in the laboratory by centrifugation and then subjected to urea detection after 400 times dilution. In case of puffed rice, 5 grains of puffed rice (∼85 mg) were crushed and mixed with 3 mL of 100 mM phosphate buffer solution, pH = 7.1 and then filtered, diluted and subjected to biosensing for the detection of urea.

To check the applicability of the developed biosensor in soil urea compost seepage, some amount of soil sample was collected from fields where urea was applied as a compost to grow vegetables. About five grams of this soil was mixed with 15 mL of 100 mM phosphate buffer and filtered through Whatman filter paper twice. The filtrate was diluted and subjected to urea measurement by the proposed biosensor, and the results are shown in Table 1. The urea content in a human urine sample was also determined by the proposed biosensor. The urine sample was diluted 4000 times and applied to the proposed biosensor.

In the case of the blood samples, the serum was extracted and diluted 100 fold before analysis. In all these cases, the actual concentrations were evaluated by dilution factor correction.

To validate the accuracy of the fabricated urea biosensor, the results from the present biosensor were compared with that measured by the commercial spectrophotometric method.³ A statistical t-test [independent two-sample t-test, degrees of freedom − (2n − 2)] was conducted considering the P-value of 0.05 (i.e., 95% confidence limit) and the results are tabulated in Table 3. In all the cases we found that the calculated t-values did not exceed the t-value from the table with the number of samples 8 (4 for the present method and 4 for the conventional spectrophotometric method) and degrees of freedom 6 (2n −2) at the 95% confidence level. The t-test predicted that there was no significant difference between the spectrophotometric results and the proposed biosensing, which yielded a confidence limit of 95%.

Table 3 ‘T’ test results with t-values from the t-table considering a 95% confidence limit

Sample no.	Calculated t-value	Tabulated t-value (P = 0.05) for degrees of freedom = 6 (n = 4)
01	1.2	2.45
02	1.63	2.45
03	2.1	2.45
04	0.87	2.45
05	2.21	2.45

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3.11. Comparison of the performances of urea sensors

Table 2 presents a detailed comparison of the performances of the proposed urea biosensor and other methods reported in the literature. The UCPHM-GCE exhibits a better biosensing response in terms of higher sensitivity (878 μA mM⁻¹ cm⁻²), low K_m value (113 μM), long shelf-life (retained 94% of activity even after two months), high surface coverage (2.40 × 10⁻⁹ mol cm⁻²) and wide urea detection range (1.5–1000 μM) than the corresponding parameters of other urea biosensors based on urease immobilization.

4. Conclusions

This paper presents a unique urea sensor comprising a PANI grafted PAM–PVA polymer composite loaded urease modified electrode. The method for the preparation of the CPHM bioelectrode and modification of the GCE was simple. This imparts high urea sensitivity, stability and enzyme loading capacity to the bioelectrode. The immobilization procedure did not give the peak reduction potential of the UCPHM GCE in the presence of interfering agents, such as uric acid, glucose, and ascorbic acid. Electrochemical methods, e.g. EIS, CV and DPV, were successfully employed to evaluate its sensitivity, the physicochemical interaction on the modified electrode and interference due to the presence of other species. Several factors (e.g. electrode modification methods, scan rate and pH of the electrolyte) that affect the electrochemical response of the electrode were optimized. When compared with other reported urea sensors, the present sensor shows superiority with respect to sensitivity, detection limit, K_m value of the enzyme, sample preparation and stability. The prepared CPHM bioelectrode produced a high surface concentration of the ionic species (2.40 × 10⁻⁹ mol cm⁻²), and this also produced good linearity over a range of urea concentrations (1.5–1000 μM), high sensitivity of 878 μA mM⁻¹ cm⁻² and a low detection limit of about 60 nM. The low K_m value (113 μM) suggests the strong affinity of the enzyme towards the substrate. EIS studies of the UCPHM bioelectrode with increasing concentrations of urea resulted in an increase in the R_CT value. Real samples, such as milk, puffed rice, urine, soil and blood, were satisfactorily assessed for urea content by this newly developed sensor and the results were in good agreement with a commercial spectrophotometric method, as revealed by the t-test.

Acknowledgements

The authors are thankful to CRNN-University of Calcutta, Kolkata, India for providing instrumental facilities.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra12159d

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