Tyrosinase-catalyzed polymerization of L-DOPA (versus L-tyrosine and dopamine) to generate melanin-like biomaterials for immobilization of enzymes and amperometric biosensing

Abstract

Learning from nature emerges as one of the most promising ways to develop advanced functional materials and biodevices. Here, inspired by melanin formation, we report the tyrosinase (Tyr)-catalyzed polymerization of L-DOPA (versus L-tyrosine and dopamine) to immobilize enzymes for amperometric biosensing. The enzymatic polymerization is examined by UV-vis spectrophotometry, scanning electron microscopy and electrochemical methods. A poly(L-DOPA) (PD)-Tyr/glassy carbon electrode (GCE) prepared by casting an aqueous mixture of L-DOPA and Tyr on a GCE exhibits a linear cathodic amperometric response to catechol concentration from 0.4 to 57 μM (R² = 0.997) with a sensitivity of 4.29 mA mM⁻¹ cm⁻² and a limit of detection (LOD) of 70 nM (S/N = 3). A PD-glucose oxidase (GOx)-Tyr/Pt electrode prepared by casting an aqueous mixture of L-DOPA, GOx and Tyr on a Pt electrode exhibits a linear anodic amperometric response to glucose concentration from 2 to 5700 μM (R² = 0.998) with a sensitivity of 78.6 μA mM⁻¹ cm⁻² and a LOD of 0.1 μM (S/N = 3). The PD-based enzyme electrode shows better biosensing performance and higher bioactivity of the immobilized Tyr than those based on similarly biosynthesized poly(L-tyrosine) and polydopamine as well as well-established chitosan and Nafion systems, implying that the biosynthesized PD as a melanin-biomimetic material has promising application potential for biomacromolecular immobilization and biosensing.

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1. Introduction

The crucial step to construct various enzymatic electrochemical biosensors is the immobilization of enzymes with high load/bioactivity, and the biosensing performance is largely affected by the enzyme-immobilization matrices and strategies.^1–3 To date, many artificial matrices of biocompatible and environmentally benign properties, including polyaniline,⁴ poly(anilineboronic acid),³ and some polyelectrolytes, ionomers or insoluble detergents such as poly(diallyldimethylammonium chloride) and Nafion,^5–9 have been developed for immobilization of enzymes and biosensing. Meanwhile, the four kinds of natural biopolymers of polysaccharides, proteins, nucleic acids, and lipids exhibit inherent biocompatibility, controllability, and degradability, and are thus receiving great attention in biosensing,^8,10,11 regenerative medicine,¹⁰ green materials,¹² and drug delivery.¹⁰ Especially, polysaccharides such as cellulose,¹³ chitosan (CS),¹⁴ and alginate,^8,10 as well as proteins such as bovine serum albumin and fibrin,^2,15 have been widely used for immobilization of enzymes and biosensing. The development of efficient artificial and biogenic polymers as well as chemical and physical strategies to immobilize enzymes for biosensing is continuously a hot topic of scientific and industrial significance.^16–19

In our opinion, the bio-pigment melanins existing in most organisms (the skin, hair, eyes, ears, and brain) can be defined as a family of biopolymers different from the abovementioned four kinds of common biopolymers. Significant progress has been made in understanding melanins and their impact on human health.^20–23 Melanins play many critical biomedical roles, e.g., determining the skin color, playing an important role in the pathology of freckle and vitiligo, absorbing light to reduce the risk of skin cancer in humans, and acting as antioxidative species and free-radical scavenging species. There are three kinds of naturally occurring melanins, eumelanin of dihydroxyindole repeat units, pheomelanin of benzothiazine repeat units, and neuromelanin. Eumelanin is the most commonly found melanin, and its biogenesis includes the oxidation of L-tyrosine by tyrosinase (Tyr) to form 3-(3,4-dihydroxyphenyl)-L-alanine (L-DOPA, a rare α-amino acid) and the deep oxidation of L-DOPA finally to form eumelanin.²³ Accordingly, poly(L-DOPA) (PD) is the material base of eumelanin, which is actually an α-amino acid polymer like a protein, though the chemical bonding for L-DOPA polymerization are not based on the peptide bond. The recent years have witnessed a growing scientific interest on melanins and the melanin-like biomaterials,^22–25 especially the melanin-like polydopamine (PDA) of many attractive properties.²⁶ Since the inception of PDA as a functional material about ten years ago,^26,27 the electrochemically, chemically, and enzymatically synthesized melanin-like PDA materials and their composites have been so widely studied for diversified applications including facile modification of surfaces, immobilization of biomacromolecules, biotechnology, and environment analysis,^1,11,26–33 and over 700 records can be found after subject (polydopamine) search on Web of Science (Thomson Reuters).²⁰ In contrast, there are much less reports on researches and development of melanin-like PD materials to date, though L-DOPA and dopamine (DA) are both catecholamine neurotransmitters of similar chemical structures.^20,34 The Tyr-catalyzed synthesis of PD and its composite materials to immobilize enzymes is still a novel topic to date. In addition, the uses of biopolymers such as polysaccharides and proteins for immobilization of enzymes and biosensing are extensively and intensively studied, but relevant studies based on the melanin biopolymers are rather limited. We have proven that PDA is an excellent matrix to immobilize enzymes with higher efficiency versus many other common matrices,^{1,11,16,27,29} prompting that the melanin-like biomaterials, which belong to a biopolymer family of special structures and properties against the abovementioned four kinds of common biopolymers, can show high future potential for immobilization of enzymes and development of biosensors and bioreactors.

The polymers synthesized by chemical, electrochemical, or enzymatic oxidation of monomers have been widely used for immobilization of biomacromolecules.^26,35,36 Chemical oxidative polymerization is sometimes not suitable for immobilization of biomacromolecules due to their denaturation under possibly harsh reaction conditions.^26,37 The electrooxidation polymerization on the electrode surfaces may limit production of a large amount of polymers.³⁶ The enzymatic oxidative polymerization protocol of high bionics significance and green-chemistry characteristics can intrinsically offer mild procedures and biocompatible microenvironment for enzyme immobilization,^35,38 thus the enzyme-catalyzed synthesis of polymers has become an interesting and important topic in the field.^39–41 For example, horseradish peroxidase (HRP) can catalyze the oxidative polymerization of phenolic compounds in the presence of H₂O₂,^42–44 and the HRP-based synthesis of organic polymers have been used to construct mono-/bi-enzyme amperometric biosensors of glucose and uric acid.^45,46 Tyrosinase can catalyze the oxidative polymerization of fluorophenols.⁴⁷ Laccase can also catalyze the oxidative polymerization of phenolic compounds, phenol derivatives, and aromatic amines by molecular oxygen.^38,48–51 Obviously, the exploitation of more enzymes and monomers for green bionic synthesis of polymers and their biocomposites in diversified academic activities and technical applications continues to be one of the focuses in the field.

Herein, inspired by the bio-formation of melanins, we report the Tyr-catalyzed polymerization of L-DOPA, L-tyrosine and DA to immobilize enzymes and fabricate amperometric biosensors. It is found that thus-prepared PD can be used to immobilize Tyr and glucose oxidase (GOx) with higher bioactivity and better biosensing performance, in comparison with poly(L-tyrosine), PDA, and/or common CS and Nafion.

2. Experimental

2.1. Instrumentation and chemicals

All electrochemical experiments were conducted on a CHI660C electrochemical workstation (CH Instrument Co.). A disk glassy carbon electrode (GCE) or Pt electrode of 3.0 mm diameter (area = 0.0707 cm²) served as the working electrode, a KCl-saturated calomel electrode (SCE) as the reference electrode, and a graphite rod as the counter electrode. All potentials here are cited versus SCE. UV-vis spectra were collected on a UV2450 UV-vis spectrophotometer (Shimadzu Co., Japan). Scanning electron microscopy (SEM) studies were performed on a Hitachi S4800 scanning electron microscope with a field emission electron gun.

Tyr (EC 1.14.18.1; 2870 U mg⁻¹, from mushroom), GOx (EC 1.1.3.4; type II from Aspergillus niger, 150 U mg⁻¹), L-tyrosine, and DA were purchased from Sigma-Aldrich. L-DOPA was purchased from Aladdin. Glucose, Nafion, ascorbic acid and uric acid were purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). Multiwalled carbon nanotubes (MWCNTs, diameter 20–40 nm) were purchased from Shenzhen Nanotech Port Co., which were purified prior to use by stirring them in 2 mol L⁻¹ concentrated HNO₃ for 20 h. Graphene oxide (GO) was purchased from XianFeng Port Co. and used as received. Glucose stock solution was allowed to mutarotate overnight at room temperature before use. Phosphate buffer solution (PBS) consisting of 0.10 M KH₂PO₄–Na₂HPO₄ + 0.1 M K₂SO₄ (pH 7.0) served as the supporting electrolyte. 0.50 wt% CS solution was prepared in 0.10 M acetate buffer solution (pH 5.4). 0.50 wt% Nafion solution was prepared in alcohol, and 0.025 wt% Nafion solution was then prepared in 0.10 M PBS. All other chemicals were of analytical grade or better quality and used as received. Milli-Q ultrapure water (Millipore, ≥18 MΩ cm) was used throughout. All experiments were performed at room temperature around 25 °C.

2.2. Procedures

As shown in Scheme 1, a PD-Tyr/GCE for catechol biosensing was prepared by casting 3 μL 0.1 M PBS containing 5 mM L-DOPA and 1 mg mL⁻¹ Tyr on a GCE till air-dryness, followed by sufficient water-rinse. A PD-Tyr-MWCNTs/GCE (or PD-Tyr-GO/GCE) was similarly prepared in the presence of 1 mg mL⁻¹ MWCNTs (or GO) in the casting liquid. A poly(L-tyrosine)-Tyr/GCE (or PDA-Tyr/GCE) was prepared similarly to the PD-Tyr/GCE case, except that 5 mM L-tyrosine (or DA) was used instead. A CS-Tyr/GCE (or Nafion-Tyr/GCE) was prepared by casting 3 μL 0.1 M PBS containing 0.50 wt% CS (or 0.025 wt% Nafion, so that the molar concentration of monomer unit of CS or Nafion is equal to that of L-DOPA) and 1 mg mL⁻¹ Tyr on a GCE till air-dryness, followed by sufficient water-rinse. A chemically synthesized PD (PD_C)-Tyr/GCE was prepared by chemical oxidative polymerization of L-DOPA by adding 3 mM K₃Fe(CN)₆ into a stirred 0.1 M PBS containing 5 mM L-DOPA and 1 mg mL⁻¹ Tyr. The reaction was allowed to occur for 30 min to make the precipitation maximally saturated, and the upper solution became rather clear. The PD_C-Tyr precipitate was centrifuged and then redispersed in 100 μL PBS, and 2 μL of the redispersed precipitate was cast-coated on the GCE. A PD-GOx-Tyr/Pt electrode for glucose sensing was prepared by casting 3 μL 0.1 M PBS containing 5 mM L-DOPA, 1 mg mL⁻¹ Tyr and 1 mg mL⁻¹ GOx on a Pt electrode till air-dryness, followed by sufficient water-rinse. A PD-GOx-Tyr-MWCNTs/Pt (or PD-GOx-Tyr-GO/Pt) electrode was prepared similarly in the presence of 1 mg mL⁻¹ MWCNTs (or GO) in the casting liquid. A poly(L-tyrosine)-GOx-Tyr/Pt (or PDA-GOx-Tyr/Pt) electrode was prepared similarly to the PD-GOx-Tyr/Pt case, except that 5 mM L-tyrosine (or DA) was used instead.

Scheme 1 Schematic representation of the preparation of PD-Tyr-MWCNTs/GCE (a) and PD-GOx-Tyr/Pt electrodes (b).

During biosensing experiments, the steady-state current responses of the Tyr-based mono-enzyme electrodes after successively adding catechol into 10 mL stirred PBS were recorded in air at −0.1 V to cathodically detect the enzymatically generated o-benzoquinone (BQ). The steady-state current response of the PD-GOx-Tyr/Pt electrode after successively adding glucose into 10 mL stirred PBS was recorded in air at 0.50 V to anodically detect the enzymatically generated H₂O₂. The response current was marked with the change value between the steady-state current and the background current.

UV-vis spectrophotometry was performed for inspection of the oxidation/polymerization reaction and evaluation of the enzymatic specific activity (ESA) of Tyr.⁵² For ESA quantification, a stirred colorimetric system of 3 mL 0.1 M PBS containing 1 mM L-DOPA and 2.5 × 10⁻² mg native Tyr or encapsulated Tyr dispersion (the same amount as the native Tyr) was taken. The encapsulated Tyr dispersion was prepared as follows. 40 μL 0.1 M PBS containing 5 mM L-DOPA (or L-tyrosine, or DA, or CS and Nafion each of 5 mM monomer units) and 1 mg mL⁻¹ Tyr was allowed to react for 30 min to make the precipitation maximally saturated, and the upper solution became rather clear. The precipitate was centrifuged and then redispersed in 40 μL PBS. 25 μL of such dispersion was added to 3 mL of 0.1 M PBS containing 1 mM L-DOPA in a 4 mL cuvette to trigger the colorimetric reaction, and the time-dependent absorbance at 475 nm was recorded. The calculation of Tyr activity is detailed later.

3. Results and discussion

3.1. Tyr-catalyzed polymerization of L-DOPA, L-tyrosine or DA to generate melanin-like biomaterials

As shown in the digital pictures of Fig. 1, after 1 mg mL⁻¹ (final concentration) Tyr was added into 0.1 M PBS containing 5 mM L-DOPA (or L-tyrosine, or DA), the solutions turned red instantly, indicating the immediate onset of Tyr-catalyzed oxidation of L-DOPA, L-tyrosine or DA. The solutions turned black after 30 min quiescence, and some precipitates were obtained after 5 min centrifugation, implying the Tyr-catalyzed polymerization of L-DOPA, L-tyrosine or DA to generate water-insoluble melanin-like biomaterials.^22–25 After the Tyr-free monomer solution was exposed to air for 3 h, the solution color changed little, indicating that the oxidation and polymerization of L-DOPA, L-tyrosine or DA occurred very slowly here without Tyr. The UV-vis absorption spectrophotometry was also conducted. New absorption bands peaking at ca. 475 nm, originating from the characteristic electronic transition of the colorful oxidized states of L-DOPA, L-tyrosine or DA after intramolecular cyclization, appeared soon after addition of 2 μg mL⁻¹ Tyr (to avoid too large absorbance here) into 0.1 M PBS containing 5 mM L-DOPA (or L-tyrosine, or DA). The absorbance increase in the first two minutes follows the order L-DOPA > DA > L-tyrosine, implying a corresponding “monomer”-oxidation-rate order. For improved clarity, the possible pathways for Try-catalyzed oxidation and polymerization of L-DOPA, L-tyrosine and DA are depicted in Schemes S1 and S2,† similar to those reported previously for nonenzymatic polymerization.²⁰ Schemes S1 and S2† suggest a final indole-polymerization mechanism, and thus the melanin-like PD, PDA and poly(tyrosine) materials are probably polyindole derivatives.

Fig. 1 UV-vis absorption spectra and digital pictures (insets) of 0.1 M PBS (pH 7.0) containing 5 mM L-DOPA (A) or 5 mM L-tyrosine (B) or 5 mM DA (C) (samples 1 and curves a), 5 mM L-DOPA (A) or 5 mM L-tyrosine (B) or 5 mM DA (C) each after adding 1 mg mL⁻¹ (samples 2) or 2 μg mL⁻¹ (curves b–f, recorded every 2 min after Tyr addition) Tyr, 5 mM L-DOPA (A) or 5 mM L-tyrosine (B) or 5 mM DA (C) each containing 1 mg mL⁻¹ Tyr after 30 min quiescence (samples 3), samples 3 after 5 min centrifugation (samples 4), 5 mM L-DOPA (A) or 5 mM L-tyrosine (B) or 5 mM DA (C) after exposed to air for 3 h (samples 5), and 1 mg mL⁻¹ Tyr (samples 6). SEM pictures of bare GCE (D), PD-Tyr/GCE (E), poly(L-tyrosine)-Tyr/GCE (F), and PDA-Tyr/GCE (G).

Tyr-catalyzed oxidation of L-DOPA, L-tyrosine or DA was also validated by electrochemical techniques, by taking the newly emerging redox peaks of the intermediates at low potentials just after intramolecular cyclization as distinguishable markers, as shown in Fig. 2. During the first cathodic sweep in 0.1 M PBS (pH 7.0) containing 5 mM L-DOPA (A), small cathodic currents were observed at potentials negative to ca. −0.25 V, which is probably related to oxygen reduction. In the second potential cycle, a pair of redox peaks at 0.25/0.13 V from the dopaquinone/L-DOPA couple shown in Scheme S1† and a pair of redox peaks at −0.19/−0.25 V from the dopachrome/leucodopachrome couple shown in Scheme S1† were observed. In contrast, the pair of redox peaks at low potentials (−0.19/−0.25 V) was observed even in the first cathodic sweep in 0.1 M PBS (pH 7.0) containing 5 mM L-DOPA + 1 mg mL⁻¹ Tyr (B), highlighting the occurrence of intramolecular cyclization of dopaquinone by Tyr catalysis. Tyr-catalyzed oxidation of DA is similar to Tyr-catalyzed oxidation of L-DOPA, as shown in panels (A′) and (B′). In the first cathodic sweep for 0.1 M PBS (pH 7.0) containing 5 mM DA (A′), small cathodic currents were observed at potentials negative to ca. −0.25 V, which is also related to oxygen reduction. In the second potential cycle, a pair of redox peaks at 0.22/0.090 V from the dopaminequinone/DA couple shown in Scheme S1† and a pair of redox peaks at −0.22/−0.29 V from the dopaminechrome/leucodopaminechrome couple shown in Scheme S1† were observed. In contrast, the pair of redox peaks at low potentials (−0.21/−0.30 V) was observed even in the first cathodic sweep in 0.1 M PBS (pH 7.0) containing 5 mM L-DOPA + 1 mg mL⁻¹ Tyr (B′), highlighting the occurrence of intramolecular cyclization of dopaminequinone by Tyr catalysis. However, 5 mM L-tyrosine in PBS did not exhibit redox activity in the cyclic voltammogram shown in panel (A′′). Soon after addition of 1 mg mL⁻¹ Tyr, the Tyr-catalyzed oxidation of L-tyrosine can produce L-DOPA, as shown in Scheme S2,† thus two pairs of redox peaks similar to those in panel (B′) were obtained, as shown in panel (B′′). Tyr-catalyzed oxidation and polymerization of L-DOPA, L-tyrosine, or DA were allowed to occur on the electrodes for biosensing applications, as shown in Scheme 1. The biosynthesized melanin-like polymer films were in black or brown color on the GCE or Pt substrate, as shown in Fig. S1.† The SEM images showed that the PD-Tyr (or poly(L-tyrosine)-Tyr, or PDA-Tyr) composite film on the GCE was composed of many aggregated nanoparticles, in vivid contrast to the bare GCE surface (Fig. 1D–G).

Fig. 2 Cyclic voltammograms in 0.1 M PBS (pH 7.0) containing 5 mM L-DOPA (A), or 5 mM L-DOPA + 1 mg mL⁻¹ Tyr (B, recorded instantly after adding L-DOPA); 5 mM DA (A′), or 5 mM DA + 1 mg mL⁻¹ Tyr (B′, recorded instantly after adding DA); 5 mM L-tyrosine (A′′), or 5 mM L-tyrosine + 1 mg mL⁻¹ Tyr (B′′, recorded instantly after adding L-tyrosine). Initial potential: open circuit potential; initial scan: negative; scan rate: 20 mV s⁻¹, working electrode: GCE.

3.2. Immobilization of Tyr in melanin-like biomaterials for amperometric biosensing of catechol

Electroactive catechol is selected here to examine the immobilization of Tyr in the PD film for phenolic biosensing at PD-Tyr/GCE. As shown in Fig. 3, no obvious redox peak was observed in air-saturated catechol-free PBS. O₂-saturation of the PBS containing 1 mM catechol led to higher reduction currents and lower oxidation currents near the redox peaks, in comparison with N₂-saturation of the solution, as a result of the continuous Tyr-catalyzed oxidation of HQ by O₂ to generate BQ (Scheme 1).

Fig. 3 Cyclic voltammograms at PD-Tyr/GCE in deaerated (b, N₂ bubbling for 30 min) and O₂-saturated (c, O₂ bubbling for 30 min) PBS (pH 7.0) containing 1 mM catechol at 50 mV s⁻¹. Curve (a) shows cyclic voltammogram at PD-Tyr/GCE in air-saturated catechol-free PBS (pH 7.0). Scan rate: 50 mV s⁻¹.

The Tyr concentration for enzyme film preparation and the solution pH for biosensing were optimized to be 1 mg mL⁻¹ Tyr and pH 7.0, through variation of the examined one while others fixed, as shown in Fig. S2.† The amperometric responses of several Tyr electrodes to catechol under optimum conditions are shown in Fig. 4 and S3.† A linear detection range (LDR) was obtained from 0.4 to 57 μM (R² = 0.997) with a sensitivity of 4.29 mA mM⁻¹ cm⁻² and a limit of detection (LOD) of 70 nM (S/N = 3) on the PD-Tyr/GCE. The sensitivity here is higher than those of the poly(L-tyrosine)-Tyr/GCE (1.09 mA mM⁻¹ cm⁻²) and PDA-Tyr/GCE (1.75 mA mM⁻¹ cm⁻²), proving that PD is a better enzyme-immobilization matrix than poly(L-tyrosine) and PDA, probably because PD as an α-amino acid polymer like a protein is of higher biocompatibility to the enzyme, and/or its faster growth rate allows immobilization of more enzyme molecules, as shown in Fig. S4.† The sensitivity here is also higher than those of CS-Tyr/GCE (2.25 mA mM⁻¹ cm⁻²), Nafion-Tyr/GCE (1.43 mA mM⁻¹ cm⁻²) and PD_C-Tyr/GCE (2.86 mA mM⁻¹ cm⁻²), due to the in situ Tyr encapsulation by green synthesis and the excellent biocompatibility of PD. As given in Fig. S3 and Table S1,† co-immobilization of MWCNTs (PD-Tyr-MWCNTs/GCE) yielded a slightly higher sensitivity (4.66 mA mM⁻¹ cm⁻²), but co-immobilization of GO (PD-Tyr-GO/GCE) yielded a lower sensitivity (3.00 mA mM⁻¹ cm⁻²), probably resulting from the excellent electron-conductivity of MWCNTs but limited electron-conductivity of GO. Our enzyme electrodes gave comparable (or higher) sensitivity to some reported analogues (Table S1†). A relative standard deviation (RSD) of 3.5% (n = 5) was obtained for sensing 10 μM catechol on PD-Tyr/GCE. The PD-Tyr/GCE maintained 85% of its initial current response after one month storage (Fig. S5†), indicating the good storage stability of the PD-Tyr/GCE.

Fig. 4 Time-dependent current response (A) to successive additions of catechol into stirred 0.1 M PBS (pH 7.0) under air atmosphere and the calibration curves (B) at PD-Tyr/GCE (a), poly(L-tyrosine)-Tyr/GCE (b), PDA-Tyr/GCE (c), PD_C-Tyr/GCE (d), CS-Tyr/GCE (e) and Nafion-Tyr/GCE (f). Applied potential: −0.1 V vs. SCE.

In addition, spectrophotometry was used to measure the ESA of native and polymer-encapsulated Tyr. Usually, one unit (U) of enzymatic activity of Tyr is defined as consumption of 1 μmol L-DOPA in 60 s, and the ESA is defined as the ratio of the molar quantity of enzymatically consumed L-DOPA in μmol in 60 s (n_L-DOPA) to the mass of enzyme (m_e) in mg. ESA is calculated by ESA = n_L-DOPA/m_e = VΔA/(bεm_e), here n_L-DOPA is quantified from Lambert–Beer's law; ΔA is the change of absorbance at 475 nm at 60 s; V is the volume of reaction solution/dispersion (here 3 mL); b is the optical pathlength (here 1 cm), ε is the molar absorption coefficient of the product (3.7 × 10³ M⁻¹ cm⁻¹ for oxidized L-DOPA at 475 nm);⁵² and m_e is the amount of Tyr employed in the reaction (here 25 μg obtained from solution dilution). The ΔA at 60 s was experimentally obtained as 0.0712 ± 0.0009, 0.0651 ± 0.0006, 0.0512 ± 0.0003, 0.0314 ± 0.0005, 0.0261 ± 0.0007 and 0.0223 ± 0.0002 for native Tyr, Tyr-PD, Tyr-CS, Tyr-PDA, Tyr-poly(tyrosine), and Tyr-Nafion, respectively, as shown in Fig. 5. Hence, the ESA values of native, PD-encapsulated, CS-encapsulated, PDA-encapsulated, poly(L-tyrosine)-encapsulated and Nafion-encapsulated Tyr are quantified as 2309, 2111, 1661, 1018, 846 and 723 U mg⁻¹, respectively, and the five encapsulated Tyr can preserve 91.4%, 71.9%, 44.1%, 36.6% and 31.3% bioactivity of native Tyr (100%). This ESA order agrees fairly well with that of the sensitivities of their enzyme electrodes.

Fig. 5 Real-time monitoring of enzymatic kinetics of catalyzed oxidation of L-DOPA in 0.1 M pH 7.0 phosphate buffer solution by native (black curve, a), PD-encapsulated (cyan curve, b), CS-encapsulated (green curve, c), PDA-encapsulated (red curve, d), poly(tyrosine)-encapsulated (blue curve, e) or Nafion-encapsulated (pink curve, f) Tyr. Absorbance was measured at 475 nm. V = 3 mL; c_Tyr = 8.3 μg mL⁻¹.

3.3. Construction of a bienzyme electrode for glucose detection

Tyr-catalyzed polymerization of L-DOPA was also used to co-immobilize GOx as a second enzyme for biosensing of glucose, as shown in Scheme 1. The Pt electrode is well known to show a better electrocatalytic performance toward electrooxidation of H₂O₂ than GCE,⁵³ so the Pt electrode was employed here. The detection potential and solution pH for glucose biosensing on the PD-GOx-Tyr/Pt electrode were optimized to be 0.50 V and pH 7.0 PBS (Fig. S6†). Fig. 6 shows the steady-state current response at the PD-GOx-Tyr/Pt electrode to successive additions of glucose into PBS at 0.50 V. A LDR from 0.002 to 5.7 mM (R² = 0.998) with a LOD of 0.1 μM (S/N = 3) was obtained at the PD-GOx-Tyr/Pt electrode. Its sensitivity of 78.6 μA mM⁻¹ cm⁻² is equivalent to that at the PD-GOx-Tyr-GO/Pt electrode (74.3 μA mM⁻¹ cm⁻²) but somewhat higher than those at PD-GOx-Tyr-MWCNTs/Pt (64.3 μA mM⁻¹ cm⁻²) (Fig. S7†), poly(L-tyrosine)-GOx-Tyr/Pt (59.1 μA mM⁻¹ cm⁻²) and PDA-GOx-Tyr/Pt (45.7 μA mM⁻¹ cm⁻²) electrodes, as well as those of many reported GOx electrodes (Table S2†). The interesting finding of a smaller sensitivity on PD-GOx-Tyr-MWCNTs/Pt than on PD-GOx-Tyr/Pt (64.3 versus 78.6 μA mM⁻¹ cm⁻²) here is different from the above catechol-biosensing case, since the bioactivity of GOx can be somewhat inhibited by its direct adsorption on the rather hydrophobic surfaces of MWCNTs, as reported previously by us and other researchers.^54–56 The finding further proves that the hydrophobic surfaces/materials like MWCNTs are not good ones for immobilization of some rather hydrophilic enzymes like GOx.

Fig. 6 Time-dependent current responses (A) to successive additions of glucose into stirred 0.1 M PBS (pH 7.0) and the calibration curves (B, solid curves) at PD-GOx-Tyr/Pt (a), poly(L-tyrosine)-GOx-Tyr/Pt (b) and PDA-GOx-Tyr/Pt (c). Applied potential: 0.50 V vs. SCE.

The PD-GOx-Tyr/Pt electrode showed excellent anti-interference ability against ascorbic acid and uric acid, as shown in Fig. S8A.† The reproducibility and storage stability of the biosensor were investigated by amperometric measurements. The reproducibility of this biosensor is evaluated with a RSD of 5.0% (n = 5) for 1 mM glucose. This electrode maintained 80% of its initial current response after one month storage (Fig. S8B†). The potential of practical usage of the biosensor was validated with several human serum samples and the values measured at the PD-GOx-Tyr/Pt electrode agreed well with the hospital results, and the standard recoveries were between 92 to 105% (Table S3†).

4. Conclusions

In summary, inspired by the melanin bio-formation, we have successfully synthesized PD bionanocomposites by Tyr catalysis for biosensing of catechol and glucose. The PD-based enzyme electrode exhibited better biosensing performance than those based on poly(L-tyrosine), PDA, CS and Nafion, and the thus-immobilized Tyr also exhibited the highest mass-specific bioactivity. Since the enzymatic polymerization here mimics formation of natural melanin biopolymers of high biomedical significance and provides mild encapsulation process and biocompatible microenvironment for biomacromolecular immobilization, the Tyr-catalyzed polymerization of L-DOPA and the development of functional melanins-biomimetic materials constitute the best enzyme-immobilization systems as compared with the controls and are thus recommended for wider biosensor and bioreactor applications.

Acknowledgements

This work was supported by the National Natural Science Foundation of China (21475041, 21175042, 21305041, 21405042), Hunan Lotus Scholars Program (2011), and Foundations of Hunan Provincial Science/Technology Department (2014SK3096).

References

1. C. Chen, Y. Fu, C. Xiang, Q. Xie, Q. Zhang, Y. Su, L. Wang and S. Yao, Biosens. Bioelectron., 2009, 24, 2726–2729.
2. F. F. Han, X. Qi, L. Y. Li, L. J. Bu, Y. C. Fu, Q. J. Xie, M. L. Guo, Y. B. Li, Y. B. Ying and S. Z. Yao, Adv. Funct. Mater., 2014, 24, 5011–5018.
3. Y. Huang, X. Qin, Z. Li, Y. Fu, C. Qin, F. Wu, Z. Su, M. Ma, Q. Xie, S. Yao and J. Hu, Biosens. Bioelectron., 2012, 31, 357–362.
4. D. Lakshmi, A. Bossi, M. J. Whitcombe, I. Chianella, S. A. Fowler, S. Subrahmanyam, E. V. Piletska and S. A. Piletsky, Anal. Chem., 2009, 81, 3576–3584.
5. H. N. Choi, M. A. Kim and W. Y. Lee, Anal. Chim. Acta, 2005, 537, 179–187.
6. A. A. Karyakin, E. A. Kotel'nikova, L. V. Lukachova, E. E. Karyakina and J. Wang, Anal. Chem., 2002, 74, 1597–1603.
7. I. Mahmood, I. Ahmad, G. Chen and H. Z. Liu, Biochem. Eng. J., 2013, 73, 72–79.
8. C. Qin, C. Chen, Q. J. Xie, L. H. Wang, X. H. He, Y. Huang, Y. P. Zhou, F. Y. Xie, D. W. Yang and S. Z. Yao, Anal. Chim. Acta, 2012, 720, 49–56.
9. C. Qin, W. Wang, C. Chen, L. J. Bu, T. Wang, X. L. Su, Q. J. Xie and S. Z. Yao, Sens. Actuators, B, 2013, 181, 375–381.
10. M. Darder, P. Aranda, M. L. Ferrer, M. C. Gutierrez, F. del Monte and E. Ruiz-Hitzky, Adv. Mater., 2011, 23, 5262–5267.
11. Y. C. Fu, P. H. Li, Q. J. Xie, X. H. Xu, L. H. Lei, C. Chen, C. Zou, W. F. Deng and S. Z. Yao, Adv. Funct. Mater., 2009, 19, 1784–1791.
12. D. L. Graham, H. A. Ferreira, P. P. Freitas and J. M. S. Cabral, Biosens. Bioelectron., 2003, 18, 483–488.
13. J. P. Fu, Z. Y. Pang, J. Yang, Z. P. Yang, J. H. Cao, Y. Xu, F. L. Huang and Q. F. Wei, Electroanalysis, 2015, 27, 1490–1497.
14. Y. M. Tan, W. F. Deng, C. Chen, Q. J. Xie, L. H. Lei, Y. Y. Li, Z. F. Fang, M. Ma, J. H. Chen and S. Z. Yao, Biosens. Bioelectron., 2010, 25, 2644–2650.
15. L. S. Jasti, S. R. Dola, N. W. Fadnavis, U. Addepally, S. Daniels and S. Ponrathnam, Enzyme Microb. Technol., 2014, 64–65, 67–73.
16. C. Chen, Q. J. Xie, D. W. Yang, H. L. Xiao, Y. C. Fu, Y. M. Tan and S. Z. Yao, RSC Adv., 2013, 3, 4473–4491.
17. S. Cosnier, Biosens. Bioelectron., 1999, 14, 443–456.
18. B. W. Park, D. Y. Yoon and D. S. Kim, Biosens. Bioelectron., 2010, 26, 1–10.
19. S. Prakash, T. Chakrabarty, A. K. Singh and V. K. Shahi, Biosens. Bioelectron., 2013, 41, 43–53.
20. M. Dai, L. Sun, L. Chao, Y. Tan, Y. Fu, C. Chen and Q. Xie, ACS Appl. Mater. Interfaces, 2015, 7, 10843–10852.
21. Y. J. Li, J. J. Liu, Y. J. Wang, H. W. Chan, L. R. Wang and W. Chan, Anal. Chem., 2015, 87, 7958–7963.
22. L. K. Povlich, J. Le, J. Kim and D. C. Martin, Macromolecules, 2010, 43, 3770–3774.
23. J. D. Simon, L. Hong and D. N. Peles, J. Phys. Chem. B, 2008, 112, 13201–13217.
24. H. Satooka and I. Kubo, J. Agric. Food Chem., 2011, 59, 8908–8914.
25. M. Mandal, T. Das, B. K. Grewal and D. Ghosh, J. Phys. Chem. B, 2015, 119, 13288–13293.
26. H. Lee, S. M. Dellatore, W. M. Miller and P. B. Messersmith, Science, 2007, 318, 426–430.
27. Y. L. Li, M. L. Liu, C. H. Xiang, Q. J. Xie and S. Z. Yao, Thin Solid Films, 2006, 497, 270–278.
28. M. R. Li, C. Y. Deng, Q. J. Xie, Y. Yang and S. Z. Yao, Electrochim. Acta, 2006, 51, 5478–5486.
29. C. Chen, L. Wang, Y. Tan, C. Qin, F. Xie, Y. Fu, Q. Xie, J. Chen and S. Yao, Biosens. Bioelectron., 2011, 26, 2311–2316.
30. S. Chen, Y. Cao and J. Feng, ACS Appl. Mater. Interfaces, 2014, 6, 349–356.
31. S. Hong, Y. S. Na, S. Choi, I. T. Song, W. Y. Kim and H. Lee, Adv. Funct. Mater., 2012, 22, 4711–4717.
32. C. H. Tao, S. R. Yang, J. Y. Zhang and J. Q. Wang, Appl. Surf. Sci., 2009, 256, 294–297.
33. S. H. Yang, S. M. Kang, K.-B. Lee, T. D. Chung, H. Lee and I. S. Choi, J. Am. Chem. Soc., 2011, 133, 2795–2797.
34. S. Subianto, G. Will and P. Meredith, Polymer, 2005, 46, 11505–11509.
35. S. Kobayashi and A. Makino, Chem. Rev., 2009, 109, 5288–5353.
36. E. Shoji and M. S. Freund, J. Am. Chem. Soc., 2001, 123, 3383–3384.
37. B. A. Deore, I. Yu, J. Woodmass and M. S. Freund, Macromol. Chem. Phys., 2008, 209, 1094–1105.
38. Y. M. Tan, W. F. Deng, Y. Y. Li, Z. Huang, Y. Meng, Q. J. Xie, M. Ma and S. Z. Yao, J. Phys. Chem. B, 2010, 114, 5016–5024.
39. J. Kadokawa and S. Kobayashi, Curr. Opin. Chem. Biol., 2010, 14, 145–153.
40. J. E. Puskas, K. S. Seo and M. Y. Sen, Eur. Polym. J., 2011, 47, 524–534.
41. P. Walde and Z. Guo, Soft Matter, 2011, 7, 316–331.
42. R. Cruz-Silva, E. Amaro, A. Escamilla, M. E. Nicho, S. Sepulveda-Guzman, L. Arizmendi, J. Romero-Garcia, F. F. Castillon-Barraza and M. H. Farias, J. Colloid Interface Sci., 2008, 328, 263–269.
43. W. Liu, J. Kumar, S. Tripathy, K. J. Senecal and L. Samuelson, J. Am. Chem. Soc., 1999, 121, 71–78.
44. M. Thiyagarajan, L. A. Samuelson, J. Kumar and A. L. Cholli, J. Am. Chem. Soc., 2003, 125, 11502–11503.
45. Y. Huang, L. Bu, W. Wang, X. Qin, Z. Li, Z. Huang, Y. Fu, X. Su, Q. Xie and S. Yao, Sens. Actuators, B, 2013, 177, 116–123.
46. M. Dai, T. Huang, L. Chao, Q. Xie, Y. Tan, C. Chen and W. Meng, Talanta, 2016, 149, 117–123.
47. G. Battaini, E. Monzani, L. Casella, E. Lonardi, A. W. J. W. Tepper, G. W. Canters and L. Bubacco, J. Biol. Chem., 2002, 277, 44606–44612.
48. F. Hollmann, Y. Gumulya, C. Tolle, A. Liese and O. Thum, Macromolecules, 2008, 41, 8520–8524.
49. M. Mazur, A. Krywko-Cendrowska, P. Krysinski and J. Rogalski, Synth. Met., 2009, 159, 1731–1738.
50. X. J. Sun, R. B. Bai, Y. Zhang, Q. Wang, X. R. Fan, J. G. Yuan, L. Cui and P. Wang, Appl. Biochem. Biotechnol., 2013, 171, 1673–1680.
51. P. Xu, H. Uyama, J. E. Whitten, S. Kobayashi and D. L. Kaplan, J. Am. Chem. Soc., 2005, 127, 11745–11753.
52. L.-P. Xie, Q.-X. Chen, H. Huang, X.-D. Liu, H.-T. Chen and R.-Q. Zhang, Int. J. Biochem. Cell Biol., 2003, 35, 1658–1666.
53. K. J. Chen, K. C. Pillai, J. Rick, C. J. Pan, S. H. Wang, C. C. Liu and B. J. Hwang, Biosens. Bioelectron., 2012, 33, 120–127.
54. Y. H. Su, Q. J. Xie, C. Chen, Q. F. Zhang, M. Ma and S. Z. Yao, Biotechnol. Prog., 2008, 24, 262–272.
55. Y. Wang and Y. J. Yao, Microchim. Acta, 2012, 176, 271–277.
56. M. Wooten, S. Karra, M. G. Zhang and W. Gorski, Anal. Chem., 2014, 86, 752–757.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c5ra27478h

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