1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 DOI: 10.1002/elan.201700332
Novel Enzymatic Rhodium Modified Poly(styrene-g-oleic
amide) Film Electrode for Hydrogen Peroxide Detection
Muhammet Samet Kilic,*
[a]Seyda Korkut,
[b]and Baki Hazer
[a]Abstract: Newly synthesized poly(styrene-g-oleic amide) was coated onto a rhodium nanoparticle modified glassy carbon (GC) surface for the fabrication of horseradish peroxidase based biosensor used for hydrogen peroxide detection. The rhodium modifed electrode presented ten times higher signal than unmodified electrode even at low elecrtroactive enzyme quantity by enhancing the electron transfer rate at the applied potential of 0.65 V. The biosensor designed by under the optimized rhodium
electrodeposition time exhibited a fast response less than 5 s, an excellent operational stability with a relative standard deviation of 0.6 % (n = 6), an accuracy of 96 % and a large linear range between 50 mM and 120 mM for hydrogen peroxide. Detection limit and the sensitivity parameters were calculated to be 44 mM and 57 mA mM1cm2, respectively by preserving its entire
initial response up to the 15 days, while only 20 % of its initial response was lost at the end of one month.
Keywords: Biosensor · horseradish peroxidase · hydrogen peroxide · polystyrene · rhodium modification
1 Introduction
Reliable, fast and accurate detection of hydrogen peroxide has an importance on many research areas such as pharmaceutical, clinical, industrial, and environmental [1]. Several hydrogen peroxide sensors have been fabricated as enzymatically and non-enzymatically up to now. Be-sides many advantages of non-enzymatic hydrogen peroxide sensors, enzymes based hydrogen peroxide sensors have more advantages such as higher sensitivity and selectivity [2]. Horseradish peroxidase a heme con-taining glycoprotein extracted from horseradish root is the most commonly used peroxidase in many analytical applications [1]. There have been an interesting attention on the third generation hydrogen peroxide biosensor based on direct electron transfer between electrode and immobilized peroxidase [3]. Even though, these studies have been useful for the investigation of enzyme structure, redox transformation mechanisms of protein molecules and metabolic processes, it is usually difficult to achieve the direct electron transfer between the enzyme and bare electrode owing to the deeply embedded redox active center and unfavorable orientation of enzymes. Therefore, researchers are focused on modifying the bare electrode with appropriate materials such as metal nanoparticles [4], metalsulfide [5], metaloxides [6–7] and carbon materials [8] to accelerate the electron transfer between the active center of the enzyme and the electrode by immobilizing enzymes on these materials. However, there is still a problem in fabricating such kinds of biosensor electrodes designed with bare electrode modification alone. It is how to immobilize enzymes onto the modified electrode surfaces without denaturation. To overcome this drawback usage of a biocompatable and flexible polymer/copolymer layer for the applications in biosensor is essential because
this layer allows enzyme immobilization onto the modi-fied electrode surface by preventing enzyme denaturation. Rhodium is a preferable material to obtain enhanced response from sensor/biosensor devices. A highly selective non-enzymatic rhodium nanoparticle-mesoporous silicon nanowire nanohybrid hydrogen peroxide sensor has been reported [9]. Rhodium/graphene hybrids have been used for hydrogen peroxide oxidation with enhanced activity in presence of rhodium [10]. Rhodium stabilized Prussian Blue-modified graphite electrodes have been designed for hydrogen peroxide reduction by researchers and they reported that the rhodium modified electrode exhibited a higher relative stability in comparison to the unmodified electrode [11]. A rhodium modified amperometric bio-sensor based on the introduced ceramic-carbon composite electrode has been developed. The sensors have been comprised of rhodium metal and glucose oxidase modified graphite particles embedded in a porous, organically modified silicate network. The electrode has been used for enzymatically generated hydrogen peroxide reduction with high in-use and storage stability [12]. To the best of our knowledge rhodium modification of a horseradish peroxidase based hydrogen peroxide biosensor electrode has not been reported before. In this study, GC carbon
[a] M. Samet Kilic, B. Hazer
Department of Chemistry, Bulent Ecevit University, 67100, Zonguldak, Turkey
Tel.: + 90 372 2911660 fax: + 90 372 2574023 E-mail: sametkilic_4274@hotmail.com [b] S. Korkut
Department of Environmental Engineering, Bulent Ecevit University, 67100, Zonguldak, Turkey
Supporting information for this article is available on the WWW under https://doi.org/10.1002/elan.201700332
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58
surface has been modified rhodium nanoparticles by electrodeposition process. Newly synthesized Poly(styr-ene-g-oleic amide) has been coated onto the modified GC, and then horseradish peroxidase has been adsorbed on the modified surface for hydrogen peroxide detection. The effect of rhodium on electrode response has been investigated. The results revealed that rhodium modified electrode surface proved to have improved electrocatalyt-ical behavior in combination with enzyme and the polymeric film layer.
2 Experimental
2.1 Reagents
Ethylenediamine ( 99 %), oleic acid ( 99 %), styrene, toluene (99.9 % HPLC grade), methanol ( 99.9 % HPLC grade), hydrochloric acid (36.5–38 %), potassium di-hydrogen phosphate, rhodium (III) chloride hydrate (RhCl3.xH2O, Rh 38–40 %), D-glucose monohydrate,
L-ascorbic acid and potassium chloride were purchased from Sigma. Horseradish peroxidase (E.C.1.11.1.7) with an activity of 10000 U vial1, hydrogen peroxide (35 %) and
di-potassium hydrogen phosphate were obtained from Merck. Stock solution of hydrogen peroxide was freshly prepared in 100 mM pH 7 phosphate buffer before the experiments.
2.2 Electrochemical Measurements
Electrochemical measurements were performed by using a CH Instruments 1040B electrochemical analyzer. A conventional three-electrode electrochemical cell, a GC working electrode (3 mm diameter), Ag/AgCl (1 M KCl) reference electrode and a platinum wire counter electrode were supplied from CH Instruments. All measurements were carried out by immersing the electrodes into the three-electrode electrochemical cell containing 10 mL of 100 mM pH 7 phosphate buffer solution at an applied potential of0.65 V under continuous stirring at 100 rpm to produce current-time recordings.
2.3 Synthesis of Poly(styrene-g-oleic amide) Graft Copolymer
Oleic acid polymerization was carried out according to the autoxidation procedure reported in a recent study [13]. 18 g of oleic acid was spread out into a petri dish and was exposed to daylight in the air at room temperature. After one month of autoxidation, a yellowish viscous liquid polymer was formed. Free radical polymerization of styrene was initiated by adding of poly(oleic acid) according to the procedure presented in Hazer and Akyol [13] to obtain poly(styrene-g-oleic amide) graft copoly-mer. In brief, 0.05 g of poly(oleic acid) and 4.52 g of styrene monomers were dissolved in 5 ml of toluene in a capped bottle. Argon gas was passed with the aid of a needle for 3 minutes to remove air from the reaction
medium. Then, reaction bottle was put into a water bath at 958C for 6 hours. After water bath, graft copolymer was precipitated in methanol and dried overnight under vacuum at 308C. 2.3 g of poly(styrene-g-oleic acid) graft copolymer and 3.2 g of ethylene diamine were dissolved in dichloromethane and stirred at room temperature for 1 hour. Then, the solvent of graft copolymer was evapo-rated and dried under vacuum at 1108C. 10 mg mL1 of
poly(styrene-g-oleic amide) graft copolymer solution was prepared in toluene daily to coat the GC electrode surface.
2.4 Preparation of the Working Electrode
The electrode fabrication step was preceded by a cleaning phase of the GC electrode using gamma alumina powder (1, 0.3 and 0.05 micron size) followed by rinsing with ultra pure water. GC pretreatment was shown to be very useful for preparing the bioelectrodes [14]. It is known that, pretreatment increases the surface roughness and density of GC, and this fact would contribute to a better deposition of metallic nanoparticles [15–16]. A cyclic voltammogram was carried out at a potential scan ranging between0.3 and + 0.8 V with scan rate of mV s1for the
pretreatment of the GC surface. Rhodium (III) chloride hydrate (RhCl3.xH2O) was electrodeposited on the
sur-face of the cleaned GC electrode in 10 mL of 100 mM hydrochloric acid solution containing 50 ppm of rhodium at a potential scan ranging between 0.8 and 0.2 V (vs. Ag/AgCl) with a scan rate of 10 mV s1. Rhodium
electro-deposited GC electrode was washed with ultra pure water and dried in the room temperature. Then, 2 mL of poly(styrene-g-oleic amide) solution was directly spread onto the surface of electrode and allowed to dry for solvent evaporation. After washing the electrode with ultra pure water, the rhodium modified and polymer coated electrode was immersed in 10 mg mL1 of
horse-radish peroxidase solution for enzyme adsorption at room temperature for 2 hours under continuous stirring at 50 rpm. Finally, horseradish peroxidase immobilized rho-dium modified poly(styrene-g-oleic amide) coated elec-trode was washed with ultra pure water. Working electrode surface was presented in Scheme 1.
3 Results and Discussion
3.1 Characterization Studies
Autooxidation process used in the synthesis of the grafted polymer was carried out by the reaction between air oxygen and unsaturated molecules under daylight at room temperature. Ecofriendly autooxidation of oleic acid was successfully carried out to prepare macroperoxide initia-tor [17–18]. Molar mass of oleic acid macroperoxide was determined by using size exclusion chromatography (SEC and GPC) (not shown). The molar mass and polydispersity of macroperoxide initiator was found to be 175 g mol1
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58
conducted by using Perkin Elmer Spectrum 100 spectrom-eter. The spectrum of the poly(styrene-g-oleic amide) was presented in Figure 1. The characteristic signal of poly-styrene was observed at 1601 cm1. Strong signal of
carboxylic acid or amide groups were observed between 1680 cm1 and 1738 cm1. C-H stretches and a tiny free
amine groups were determined at 2922—3025 cm1 and 3400 cm1, respectively. The SEM images of the rhodium
modified and unmodified poly(styrene-g-oleic amide) film
coated glassy carbon surfaces were taken with Quanta FEG 450 model SEM device. The images were presented in Figure 2. Different surface morphologies were observed from the SEM images between the modified and unmodi-fied electrodes. Clusters of rhodium nanoparticles on the surface were observed, and the size of these clusters were observed in the range of 615 nm 2.5 mm (Figure 2A). The Energy dispersive X-ray chromatograms (EDX) were taken with the same device for the clustered and non-clustered surface of the rhodium modified poly(styrene-g-oleic amide) film coated electrode (Supporting info-Figure S1). Chromatogram results revealed that the mod-ification took place on the entire surface of the electrode because typical rhodium peak was observed on the non-clustered surface (Supporting info-Figure S1 B). It can be attributed to the sole rhodium particles were placed on the electrode surface other than clustered regions.
The electrodeposition of rhodium on the GC surface was succeeded by applying repetitive (number of cycle) potentials in the range of 0.8 and 0.1 V (Figure 3). The rhodium electrodeposition potential varies depending on electrode substrate and electrodeposition conditions. A research group [19] was observed an increase in the amount of electrodeposited rhodium at negative poten-tials ranging between0.5 and 0.8 V. Rhodium (+ 3) is converted to rhodium (solid) particle by three electron reduction on electrode surface electrochemically during the deposition. In our study, the typical reduction peak of rhodium ion was observed at around 0.5 V during the electrodeposition experiment, and the reduction current decreased by the increasing number of electrodeposition cycle since more rhodium nanoparticles were accumulated on the electrode surface by consuming rhodium ion in the solution during the deposition process.
The electrochemical characteristic of the rhodium modified and unmodified poly(styrene-g-oleic amide) film coated electrodes was evaluated through cyclic voltamme-try (CV) at a potential scan ranging between 0.8 and +0.8 V with scan rate of 100 mV s1 in 100 mM pH 7
phosphate buffer (Figure 4). The presence of rhodium leaded to wider cyclic voltammetric curve in comparison to unmodified electrode. This can be explained with the augmentation of the working electrode electroactive sur-face. An observable effect of rhodium was at0.65 V by providing more forceful electrode surface characteristic. In addition, the current values were substantially higher than the unmodified electrode.
3.2 Electrochemical Response of the Rhodium Deposited Poly(styrene-g-oleic amide) Film Electrode to Hydrogen Peroxide
Cyclic voltammograms of the rhodium modified and unmodified poly(styrene-g-oleic amide) film coated en-zyme immobilized electrodes were obtained in the absence and presence of 2 mM hydrogen peroxide in 100 mM pH 7 phosphate buffer at a potential scan ranging between 0.8 and + 0.8 V with scan rate of 100 mV s1
Scheme 1. Schematic diagram of the working electrode surface.
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58
(Figure 5). The biosensor electrode without rhodium generated a hydrogen peroxide reduction current of 3 mA (Figure 5A), while rhodium modified generated 10 times higher reduction current (30 mA) for the same concen-tration of hydrogen peroxide (Figure 5B). It is obvious that, rhodium nanoparticles improved the signal gener-ation ability for hydrogen peroxide reduction. Rhodium nanoparticles created a bridge for the electron flow from
the electrode to the enzyme. The reaction mechanism of the rhodium modified electrode was presented in Support-ing info- Figure S2. Horseradish peroxidase has a redox active center with Fe+3
ion which is responsible of hydrogen peroxide reduction to water molecule. The enzyme catalyzes hydrogen peroxide to form the inter-mediate enzyme contains (Fe4 +=
O). The intermediate Fig. 2. SEM images of the rhodium modified (A), and unmodified (B) poly(styrene-g-oleic amide) film coated glasy carbon surfaces.
Fig. 3. The observed current values from the rhodium electro-deposition process.
Fig. 4. Cyclic voltammograms of the rhodium modified and unmodified poly(styrene-g-oleic amide) film coated electrodes at a potential scan ranging between0.8 and + 0.8 V with scan rate of 100 mV1 s1in 100 mM pH 7 phosphate buffer.
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58
horseradish peroxidase has catalytic activity and it gains one electron from the electrode to form another inter-mediate enzyme, which is subsequently reduced back to the native enzyme by accepting another electron from the electrode under a favorable applied potential [20]. Some-times, electrical communication between the electrode surface and enzyme is hindered [21] depending on polymeric film morphology which results in signal loss of the biosensor. To overcome this problem, using of an artificial electron relay which creates a bridge for facile electron transport between redox centre of enzyme and electrode is proposed. Hydrogen peroxide signal of the rhodium deposited electrode was observed at 0.65 V while unmodified electrode was at 0.5 V (vs. Ag/AgCl). The cyclic voltammogram results showed that the poten-tial shifted to0.65 V in presence of rhodium. This can be attributed to the electron flow was carried out via rhodium nanaoparticles at 0.65 V where rhodium pre-sented electroactive characteristic (Figure 4). Chandra et al. [22] used N,N-bis-succinamide-based dendrimer capped rhodium nanoparticles for hydrogen peroxide sensor. Similary they reported that presence of hydrogen peroxide significantly increased the peak current at 0.6 V, and the current values were substantially higher than those obtained with uncoated surface of GC electrode. They clarified that the dendrimer-encapsulated rhodium nanoparticles acted as mediator for the electro-catalytic reduction of hydrogen peroxide. Hydrogen
peroxide was sensed at various potentials with horseradish peroxidase based biosensor studies, for instance, at 0.57 V for graphene oxide, cobalt(III) oxide modified nafion film [23], at0.25 V for poly(aniline-co-N-methyl-thionine) [24], at 0.3 V for SnO2 sol-gel [25], at 0.4 V
for silica–hydroxyapatite hybrid film electrode [26]. Dif-ferent operational potentials can be attributed to the different surface morphologies of working electrodes. Potential value for a spesific analyte can shift or change significantly in biosensors prepared with various surface morphologies which can lead to loose or enhance of the electron transfer ability on the working electrode. The experiments were also performed for enzyme-less rho-dium modified poly(styrene-g-oleic amide) film coated electrodes. No signal was recorded from the enzyme-less electrode for 2 mM of hydrogen peroxide.
Rhodium electrodeposition time was investigated with cyclic voltammetry by changing the number of cycle at the potential scan ranging between0.8 and 0.2 V with a scan rate of 10 mV s1. A serie of GC electrode was prepared
at different rhodium electrodeposition time at 5, 10, 12 and 15 cycle number, respectively. The electrodes were then coated with poly(styrene-g-oleic amide) film and horseradish peroxidase was immobilized onto the elec-trode surfaces according to the procedure presented in electrode preparation section. Each of the electrodes was tested by immersing 10 mL of 100 mM pH 7 phosphate buffer solution containing 2, 4, 8, 16 and 32 mM hydrogen peroxide at 0.65 V, respectively. Amperometric hydro-gen peroxide signals of the each electrode were shown in Figure 6A. The response of the all tested concentrations of hydrogen peroxide were increased up to the deposition cycle number of 10, as expected due to the presence of a higher amount of rhodium on the electrode, and then decreased slightly. This experiment was repeated one more time for each cycle number. The relative standard deviation of the currents obtained from these two experi-ments was 2.1 % for the electrodes prepared at 10 cycle number. The decreasing of the electrode response at higher cycle numbers indicated that the overall response was not only function of the electrodeposited amount of rhodium. Distribution of rhodium on the GC surface played a crucial role for the enzyme adsorption onto the rhodium modified polymeric film. In fact, the electro-deposited rhodium amount has to be high enough to facilitate the biocatalytical reduction of hydrogen peroxide, but not too high to prevent the enzyme adsorption onto the surface. These results revealed that the decrease of the signal beyond 10 cycle number can be due a more difficult enzyme adsorption with higher rhodium quantity. Similar result was obtained in the case of other metallic nanoparticle deposited GC electrode [14, 19]. To proof this, cyclic voltammogram of each electrode was measured in phosphate buffer (pH 7) at various scan rates ranging between 10 and 150 mV s1 to
investigate electroactive enzyme concentration on the surface (supporting info-Figure S3 A). The cathodic peak currents increased linearly with increasing scan rate for Fig. 5. Cyclic voltammogram of the biosensor electrode without
rhodium (A) and rhodium modified (B) obtained in the absence and presence of 2 mM hydrogen peroxide.
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58
the all working electrodes prepared with different rho-dium quantities, as Ipc=0.282u–5.919 (mA, mV s1, r
2=
0.97) for the rhodium deposited electrode prepared with 10 cycle number. From the integration of the cathodic peak currents and using Faraday’s law, the surface concentration of electroactive enzyme could be estimated according to the following equation [27]:
Ipc¼
n2
F2
A G n
4RT ð1Þ
where Ipc was the cathodic peak current, A was the
electrode surface area, v was the scan rate, n was the number of electron, R, T and F had their usual meanings. By considering v = 0.1 Vs1, n = 1, A = 0.07068 cm2
, F = 96,485 C mol1, R = 8.314 J K1mol1 and T = 298 K, I
pc=
14.106 A for the rhodium deposited electrode prepared with 10 cycle number, amount of the electroactive enzyme was calculated to be 6 mg which was the maximum quantity in comparison to the other electrodes prepared with different rhodium electrodeposition times. The peak currents varied linearly with the square root of the scan rate over the entire range of 10–150 mV s1 (supporting
info- Figure S3 B), suggesting diffusion controlled mass transfer reactions. The Randles-Sevcik model (equation 2) was used to calculate the diffusion coefficient of the electrons within the polymeric film, where n is electron stoichiometry, C is the surface concentration (2.123
105mol/m2
). The slope of the linear regression (y = 4.41x + 8.52 (r2
=0.99) was used to calculate the diffusion coefficient (De) as 1.193106m 2 s1. Ip¼ ð2:69 10 5 Þ n3=2 A D1=2 e C v 1=2 ð2Þ Cyclic voltammogram of the rhodium deposited elec-trode with 10 cycles in the buffer containing 2, 4, 8, 16 and 32 mM hydrogen peroxide was presented in Figure 6B. The voltammogram showed that reduction current of hydrogen peroxide increased slightly at0.65 V owing to the participation of rhodium nanoparticles to the electron flow occurring between the enzyme and the electrode surface.
3.3 Analytical Figures of Merit
The typical amperometric response of the rhodium deposited poly(styrene-g-oleic amide)/horseradish peroxidase electrode was investigated by successive addition of increasing hydrogen peroxide concentration ranging between 50 mM and 126 mM at the applied potential of 0.65 V. The current-time recording of the biosensor was presented in Figure 7A (all responses not shown). The reduction current increased rapidly when the increasing concentrations of hydrogen peroxide were added, and reached a maximum steady state value with less than 5 s. The biosensor detected hydrogen peroxide at mA level for all concentrations. A linear calibration plot (Figure 7B) was constructed by using steady-state reduc-tion currents obtained from Figure 7A. The response of the biosensor had a linear relationship with hydrogen peroxide concentrations ranging between 50 mM and 120 mM (r2
= 0.997), and then the current increased nonlinearly, as expected in biocatalytic reactions. The linear range was wider than previously publishedmetallic nanoparticle modified horseradish peroxidase based hy-drogen peroxide biosensors. For example; it was wider than Fe3O4-Au magnetic nanoparticles coated graphene
sheets-nafion film (20 mM–2.5 mM) [28], sol-gel support-ing matrix containsupport-ing gold nanoparticles (12.2 mM– 1.1 mM) [29], ultrathin gold nanowires and nanoparticles prepared by oleylamine synthesis (20 mM–500 mM) [30], poly(N-[3-(trimethoxy silyl)propyl]aniline on gold nano-rods (10 mM–1 mM) [31], palladium nanoparticles contain-ing graphene modified graphite (25 mM–3.5 mM) [32], polypyrrole synthesized on platinum (0.49 mM–0.63 mM) [33], carbonnanotubes/chitosan composite (16.7 mM– 0.74 mM) [34], poly(glycidylmethacrylate-co-vinylferro-cene) film (2–30 mM) [35] and nafion-sonogel-composite film (4–100 mM) [36], NiFe2O4 nanoparticle modified film
(10 mM–20 mM) [37], Ag nanoparticle modified film (1– 147 mM) [38] and Ag@TiO2modified film electrode (0.83–
Fig. 6. Effect of cycle number in rhodium electrodeposition process on hydrogen peroxide response (A). Cyclic voltammo-grams of the rhodium deposited (with a cycle number of 10) electrode obtained in presence of 2, 4, 8, 16 and 32 mM hydrogen peroxide in 100 mM pH 7 phosphate buffer (B).
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58
43.3 mM) [39] . The detection limit was 44 mM which was calculated with 3 sb/m criteria [40], where m was the slope of the linear range of the calibration plot, and sb was the standard deviation of the currents of different solutions of hydrogen peroxide at the concentration level correspond-ing to the lowest concentration of the calibration plot. The biosensor sensitivity was found to be 57 mA mM1 cm2
which was higher than the previously published hydrogen peroxide sensors based on horseradish peroxidase or non-enzymatic electrodes (Supporting Information-Table S1). The measurement accuracy of the biosensor was 96 % calculated according to the formula given as (measured concentration/actual concentration)3100 [47].
Repeat-ability of the detection in one operation was monitored by the measurement of the response for a serie of 6 succesive additions of 2 mM hydrogen peroxide into 100 mM phosphate buffer (pH 7) at0.65 V (vs. Ag/AgCl). Well-defined reduction responses were obtained with relative standard deviation (RSD %) of 0.6 % which was an ignorable value in comparison to RSD % values ranging between 3.2 and 5.3 % [33, 34, 36] of the other horseradish
peroxidase based hydrogen peroxide biosensors. No response loss was observed up to 15 days when the electrode was stored in phosphate buffer + 48C, while only 20 % of its initial response was lost at the end of one month. It is well known that excess hydrogen peroxide concentration inhibits peroxidase-catalyzed reactions [48, 49]. Horseradish peroxidase in the intermediate state can be oxidized by excess peroxide. This enzyme state is not catalytically active but its formation does not represent a terminal inactivation of the peroxidase since the enzyme in this state decomposes spontaneously to native peroxidase. However, the return to the native enzyme is sufficiently slow. Therefore, in the presence of excess peroxide, this new state of the enzyme formation would increase resulting in suppressed catalyst perform-ance [50]. In this study, relatively large linear range, high operation and the storage stability can be attributed to the mild microenvironment of the polymeric film layer which preserved the enzyme effectively even at high concen-trations of hydrogen peroxide with its thrummy structure created by oleic amide chains.
3.4 Interference and Selectivity
Interference experiments were performed by comparing the amperometric responses of 0.5 mM hydrogen peroxide before and after the addition of some interferants into 100 mM pH 7 phosphate buffer. The possible interferants selected for hydrogen peroxide detection were glucose, ascorbic acid and potassium chloride. Each of the interferants (0.5 mM) was added into the reaction me-dium. The amperometric currents were recorded at 0.65 V, and presented in Figure 8. Results showed that the working electrode presented high selectivity for only hydrogen peroxide, and the tested possible interferants Fig. 7. Current-time recording of the biosensor to increasing
hydrogen peroxide concentrations ranging between 50 mM and 126 mM (schematic presentation up to the concentration of 50 mM) at the applied potential of 0.65 V (vs. Ag/AgCl) (A), and hydrogen peroxide calibration curve obtained from current-time recording in linear range from 50 mM to 120 mM (B).
Fig. 8. Interference effect of various substances (0.5 mM) on biosensor response at0.65 V.
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58
did not lead any observable interference effect on hydro-gen peroxide detection. Even though, hydrohydro-gen peroxide was added after the each interferent additon, no signal loss was observed for hydrogen peroxide.
4 Conclusion
In this report, a rhodium modified GC electrode was used for the first time for enzymatic hydrogen peroxide biosensor. The modified electrode surface was coated with newly synthesized poly(styrene-g-oleic amide) film to immobilize enzyme and protect enzyme from denatura-tion. Results showed that rhodium nanoparticles acceler-ated the electron transfer rate between the GC and the redox active center of the enzyme at 0.65 V. Good analytical characterictics of the biosensor were observed with regard to linear range, measurement accuracy, opera-tional and storage stability. Microenvironment of the polymeric film layer preserved the enzyme effectively even at high hydrogen peroxide concentrations with its long oleic amide chains.
References
[1] C. Kac¸ar, B. Dalkiran, P. E. Erden, E. Kilic¸, Appl. Surf. Sci. 2014, 311, 139.
[2] Y. Lv, F. Wang, H. Zhu, X. Zou, C. Tao, J. Wang, Nanomat. Nanotechnol. 2016, 6, 30.
[3] S. Xu, X. Qin, X. Zhang, C. Zhang, Microchim. Acta 2015, 182, 1241.
[4] L. Y. Chen, T. Fujita, M. W. Chen, Electrochim. Acta 2012, 67, 1.
[5] G. C. Fan, X. L. Ren, C. Zhu, J. R. Zhang, J. J. Zhu, Biosens. Bioelectron. 2014, 59, 45.
[6] Z. Yang, Y. Tang, J. Li, Y. Zhang, X. Hu, Biosens. Bioelectron. 2014, 54, 528.
[7] A. K. Yagati, T. Lee, J. Min, J. W. Choi, Biosens. Bioelectron. 2013, 47, 385.
[8] Y. Fang, E. Wang, Chem. Commun. 2013, 49, 9526.
[9] Z. Song, H. Chang, W. Zhu, C. Xu, X. Feng, Scientific Reports 2015, 5, 7792.
[10] H. Chang, D. Y. Lei, D. He, X. Sheng, Z. Song, X. Feng, Chem. Electrochem. 2014, 1, 1480.
[11] L. Mures¸an, G. L. Turdean, I. C. Popescu, J. Appl. Electro-chem. 2008, 38, 349.
[12] S. Sampath, O. Lev, J. Electroanal. Chem. 1997, 426, 131. [13] B. Hazer, E. Akyol, J. Am. Oil Chem. Soc. 2016, 93, 201. [14] M. C. Rodriguez, G. A. Rivas, Electroanal. 1999, 11, 558. [15] L. J. Kepley, A. J. Bard, Anal. Chem. 1988, 60, 1459. [16] C. Barbero, R. Ko¨tz, J. Electrochem. Soc. 1993, 140, 1. [17] B. Hazer, J. Polym. Sci. Part A Polym. Chem. 1987, 25, 3349. [18] A. Alli, B. Hazer, J. Am. Oil Chem. Soc. 2011, 88, 255. [19] S. A. Miscoria, G. D. Barrera, G. A. Rivas, Sens. Actuators B
2006, 115, 205.
[20] S. Korkut Ozoner, J. Chem. Technol. Biotechnol. 2012, 87, 146.
[21] M. S. Kilic, S. Korkut, B. Hazer, E. Erhan, Biosens. Bioelectron. 2014, 61, 500.
[22] S. Chandra, K. S. Lokesh, A. Nicolai, H. Lang, Anal. Chim. Acta 2009, 632, 63.
[23] S. Asim Bin Asif, S. B. Khan, A. M. Asiri, Microchim. Acta 2016, 183, 3043.
[24] C. Chen, X. Hong, T. Xu, A. Chen, L. Lu, Y. Gao, Synth. Met. 2016, 212, 123.
[25] N. Q. Jia, J. Xu, M. H. Sun, Z. Y. Jiang, Anal. Lett. 2005, 38, 1237.
[26] B. Wang, J. J. Zhang, Z. Y. Pan, X. Q. Tao, H. S. Wang, Biosens. Bioelectron. 2009, 24, 1141.
[27] E. Laviron, Journal of Electroanalytical Chemistry and Interfacial Electrochemistry 1979, 100, 263.
[28] Y. Xin, X. Fu-bing, L. Hong-wei, W. Feng, C. Di-zhao, W. Zhao-yang, Electrochim. Acta 2013, 109, 750.
[29] C. X. Lei, S. Q. Hu, N. Gao, G. L. Shen, R. Q. Yu, Bioelectrochem. 2004, 65, 33.
[30] E. Koposova, X. Liu, A. Kisner, Y. Ermolenko, G. Shumilova, A. Offenhausser, Y. Mourzina, Biosens. Bioelec-tron. 2014, 57, 54.
[31] S. Komathi, A. I. Gopalan, S. K. Kim, G. S. Anand, K. P. Lee, Electrochim. Acta 2013, 92, 71.
[32] S. Nandini, S. Nalini, R. Manjunatha, S. Shanmugam, J. S. Melo, G. S. Suresh, J. Electroanal. Chem. 2013, 689, 233. [33] S. S. Razola, B. L. Ruiz, N. M. Diez, H. B. Mark Jr, J. M.
Kauffmann, Biosens. Bioelectron. 2002, 17, 921. [34] L. Qian, X. Yang, Talanta 2006, 68, 721.
[35] M. S¸enel, E. C¸ evik, M. F. Abasıyanık, Sens. Actuators B 2010, 145, 444.
[36] M. ElKaoutit, I. Naranjo-Rodriguez, M. Dominguez, M. P. Hernandez-Artiga, D. Bellido-Milla, J. L. Hidalgo-Hidalgo de Cisneros, Electrochim. Acta 2008, 53, 7131.
[37] L. Luo, L. Zhu, Y. Xu, L. Shen, X. Wang, Y. Ding, Q. Li, D. Deng, Microchim. Acta 2011, 174, 55.
[38] Y. Wang, M. Tang, X. Lin, F. Gao, M. Li, Microchim. Acta 2012, 176, 405.
[39] M. M. Khan, S. A. Ansari, J. Lee, M. H. Cho, Mater. Sci. Eng. C 2013, 33, 4692.
[40] S. Korkut, B. Keskinler, E. Erhan, Talanta 2008, 76, 1147. [41] T. Tatsuma, Y. Okawa, T. Watanabe, Anal. Chem. 1989, 61,
2352.
[42] L. L. Zhang, G. Q. Han, Y. Liu, J. Tang, W. H. Tang, Sens. Actuators B 2014, 197, 164.
[43] K. Manivannan, C. C. Cheng, J. K. Chen, Electroanalysis 2017, 29, 1443.
[44] D. Shan, S. Wang, D. Zhu, H. Xue, Bioelectrochemistry 2007, 71, 198.
[45] A. Ralph, K. Wilson, G. S. Wilson, Anal. Chem. 1980, 52, 1198.
[46] N. H. Al-Hardan, M. A. A. Hamid, R. Shamsudin, N. K. Othman, L. K. Keng, Sensors 2016, 16, 1004.
[47] S. Korkut, S. Uzuncar, M. S. Kilic, B. Hazer, Instrum. Sci.Technol. 2016, 44, 614.
[48] F. W. Scheller, F. Schubert, J. Fedrowitz, Frontiers in Biosensorics I Fundamental Aspects, Birkhauser-Verlag, Basel, Switzerland 1997.
[49] V. A. Bogdanovskaya, V. A. Fridman, M. R. Tarasevich, F. Scheller, Anal. Lett. 1994, 27, 2823.
[50] J. A. Nicell, J. K. Bewtra, K. E. Taylor, N. Biswas, C. St. Pierre, Water Sci. Technol. 1992, 25, 157.
Received: June 4, 2017 Accepted: July 12, 2017 Published online on July 19, 2017
