Characterization and immobilization of Trametes versicolor laccase on magnetic chitosan–clay composite beads for phenol removal

Abstract

Laccase from Trametes versicolor was immobilized on magnetic chitosan–clay composite beads by glutaraldehyde crosslinking. The physical, chemical, and biochemical properties of the immobilized laccase and its application in phenol removal were comprehensively investigated. The structure and morphology of the composite beads were characterized by SEM, TGA, and FTIR analyses. The immobilized laccase showed better storage stability and higher tolerance to the changes in pH and temperature compared with free laccase. Moreover, the immobilized laccase retained more than 75% of its original activity after 10 cycles. The efficiency of phenol removal by immobilized laccase was about 80% under the optimum conditions after 4 h.

Keywords::

• chitosan
• enzyme immobilization
• magnetic
• phenol removal
• Trametes versicolor laccase

Introduction

Contamination of water, underground water, and soil by aromatic organic pollutants in many areas has caused great concern around the world. Among these organic pollutants, phenol is well known for its high toxicity in humans and animals (Rangelov and Nicell 2015, Hou et al. 2014a, Xu et al. 2014). Phenolic pollutants can originate from agricultural and industrial activities, including the partial degradation of phenoxy herbicides, the use of wood preservatives, the generation of wastes by pulp and paper, petrochemicals, dyeing, and other organic chemical and textile industries. Conventional purification methods like solvent extraction, chemical oxidation, and adsorption on activated carbon suffer from serious drawbacks such as high cost, inefficient purification, and the formation of hazardous by-products, which are not characteristic of the adsorption process; on the other hand, the main drawback is the recovery of adsorbents.

Enzyme-based treatments for the removal of phenolic compounds have offered some distinct advantages over physical and chemical removal methods (Bezerra et al. 2015, Xu et al. 2015, Hou et al. 2014b, Başak and Aydemir 2013, Fernández et al. 2013). Thus, using enzymes as decontaminating agents has received great attention because of their potential to remove pollutants from the environment without creating the harsh side effects associated with other methods. The significant advantages of this enzymatic method include the mild condition of enzymatic treatment, the requirement of only trace amounts of enzymes, the ability to decontaminate low concentrations of contaminants, and the ability to handle large volumes of effluent as well. Peroxidases, laccases, and tyrosinases catalyze the oxidation of phenolics using either hydrogen peroxide or molecular oxygen, generating phenoxy radicals that react with themselves or other phenolics to form dimers. Laccase (EC 1.10.3.2 benzenediol:oxygen oxidoreductase) is a multi-copper oxidase able to catalyze the one-electron oxidation of several aromatic substrates. The substrates of the enzyme include ortho-, para-, amino-, and poly-phenols; and the low substrate-specificity of laccase makes it an attractive catalyst for industrial oxidative processes. However, enzymes have some limitations for use in non-biological applications. Thus, for many industrial applications, enzymes have to be immobilized, via very simple and cost-effective protocols, in order to improve their properties such as activity, stability, and selectivity (Bezerra et al. 2015, Xu et al. 2015, Şanlıer et al. 2013). Many different carriers have been used for enzyme immobilization. Among them, chitosan, as a macromolecular material, exhibits many interesting properties, namely biocompatibility, availability of reactive functional groups for chemical modifications, hydrophilicity, mechanical stability, regenerability, and ease of preparation in different geometrical configurations suitable for a chosen biotransformation. Recently, there has been a growing interest in chemical or physical modification of chitosan to improve its solubility and widen its environmental and biomedical applications (Başak and Aydemir 2013, Xu et al. 2013).

In this study, laccase was immobilized on magnetic chitosan–clay beads, and the conditions for immobilization and characterization of the immobilized enzyme were studied systematically. Application of the immobilized system in phenol removal was investigated in a batch system.

Materials and methods

Materials

Laccase from Trametes versicolor (EC 1.10.3.2) was supplied by Sigma. ABTS (2,2-azino-bis-(3-ethylbenzthiazoline-6-sulfonic acid), chitosan (deacetylated chitin, CAS 9012-76-4; medium molecular weight, deacetylation degree 75–85%), glutaraldehyde (25%), sodium triphosphate pentabasic (TPP), Coomassie Brilliant Blue G-250, nanoclay (hydrophilic bentonite, average particle size: ≤ 25 μm, H₂Al₂O₆ Si, 180.1 g/mol) were obtained from Sigma Chemical Co. (St. Louis, MO, USA). Acetic acid (100%) and all other chemicals were obtained from Merck AG (Darmstadt, Germany).

Preparation of magnetic chitosan–clay beads and laccase immobilization

Fe₃O₄ nanoparticles were prepared by coprecipitation of Fe²⁺ (FeSO₄•7H₂O) and Fe³⁺ (FeCl₃• 6H₂O) ions in ammonia solution (Yuanbi et al. 2008). To prepare the magnetic chitosan–clay beads, 2 g of chitosan flakes was dissolved in acetic acid (100 ml). Next, 1 g of clay, and 1 g of magnetic Fe₃O₄ were added into the chitosan solution and then were agitated by magnetic stirring at room temperature. After 10 min, the viscous solution was placed in an ultrasonic bath to remove air bubbles. The magnetic chitosan–clay solution was dropped through a syringe into cold TPP solution (pH 8.2) and was stirred for 4 h. Then, the beads were washed with deionized water until they became neutral, and were stored in deionized water for use afterwards.

Laccase from Trametes versicolor was immobilized on magnetic chitosan–clay composite beads covalently with glutaraldehyde crosslinking (Chang 1971). Adequate composite (1 g) was placed in 3% (v/v) of glutaraldehyde (100 ml) and crosslinked at 150 rpm and 30° C for 4 h. Then the beads were washed several times with deionized water to remove excess of glutaraldehyde. An amount of modified bead (0.05 g) was added to 2.5 ml of the laccase solution (2 mg/ml), and the immobilization reaction was accomplished in a rotary shaker for 4 h at 25°C. The beads were washed with 0.05 M acetate buffer and were stored in acetate buffer at 4°C until use.

Characterization of magnetic chitosan–clay beads

Fourier transform infrared (FTIR) analysis was accomplished on a Perkin Elmer spectrum BX, scanning from 4000 cm^{− 1}–400 cm^{− 1} at room temperature. Modified chitosan beads were mixed with KBr and pressed to plates for measurements. Thermal gravimetric analysis of the modified chitosan beads was performed with a Perkin Elmer SII 7300 TGA analyzer. The surface morphology of the beads was examined using scanning electron microscopy (SEM) (XL30-SFEG, FEI/Philips). The magnetic properties of the samples were studied by a vibrating sample magnetometer (Lake Shore VSM) at room temperature.

Laccase activity assay

The activity of free and immobilized laccase was determined spectrophotometrically at ʎ_max =420 nm with 1 mM ABTS as substrate in 0.05 M sodium acetate buffer (pH 4.0) at 25°C (Bourbonnais and Paice 1992). The oxidation of substrate to ABTS⁺ was measured using a UV–Vis spectrophotometer (UV-2250, SHIMADZU Corporation, Japan), with the molar extinction coefficient of 36 × 10³ M^{− 1} cm^{− 1}. One unit (U) of laccase activity was defined as the amount of enzyme needed to oxidize 1 μmol of ABTS per minute. The activity recovery of the immobilized laccase is calculated from the formula:

where R is the activity recovery of the immobilized laccase (%), Ai is the activity of the immobilized laccase (U), and Af is the activity of the same amount of free laccase in solution as that immobilized on support (U). The protein content of solutions was determined with the Bio-Rad Protein Assay (Bio-Rad, USA), based on the Bradford assay (Bradford 1976), using bovine serum albumin (BSA) as the protein standard.

Properties of immobilized enzyme

To determine the pH–activity profiles of the free and immobilized laccase, activities were measured in sodium acetate buffer (0.05 M, pH 2.0–4.0) and phosphate buffer (0.05 M, pH 5.0–7.0) at 25°C. Effects of temperature on the activities of the free and immobilized enzymes were conducted over the temperature range of 20–70°C at pH 5.0. Here, the activity was expressed as relative forms (%), with the maximal value of enzyme activity at a certain pH or temperature set as 100%.

Kinetic parameters of the Michaelis–Menten equation (K_M and V_max) for the free and immobilized laccase were tested by measuring the initial rates of ABTS oxidation over the concentration range of 0.01–1 mM at 25°C in sodium acetate buffer at pH 5.0. The Lineweaver–Burk plot was used to estimate kinetic parameters, and V_max and K_M were determined from the intercepts at X- and Y-axis of the plot, respectively.

For determination of thermal stability, either free or immobilized laccase were incubated in acetate buffer (0.05 M, pH 5.0) for 120 min at 60 and 70°C. Activity of samples was determined at optimum conditions.

The storage stability of the free and immobilized laccase was investigated by measuring their activities after being stored at 4°C for 8 weeks, and the measurement of remaining activity was performed once a week. All experiments were performed in triplicate.

Reusability

The immobilized enzyme beads were used several times for phenol removal, to test the reusability of immobilized laccase. After each cycle, the beads were separated with a magnet and washed with 0.05 M sodium acetate buffer (pH 5.0) and the solution replaced with fresh phenol solution. The activity of freshly prepared beads in the first run was defined as 100%. The reusability study was performed in triplicate.

Removal of phenol

Studies on the removal of phenol were performed at 25°C with immobilized laccase in acetate buffer (0.05 M, pH 5.0) with shaking at 100 rpm. In each set of the experiment, 50 mg of the immobilized laccase was added into 10 ml of the reaction medium with 10 ppm phenol, with an incubation time lasting 24 h. The residual phenolic compounds were analyzed by measurement of absorbance at a wavelength of 510 nm using a UV spectrophotometer after color development by the 4-aminoantipyrine (4-AAP) method (Wolfenden and Willson 1982). The phenol degradation rate was calculated from the initial and final concentrations of phenolic substrate by the following equation:

Results and discussion

Characterization of magnetic chitosan–clay beads

Microscopy photographs and SEM images of chitosan and chitosan–clay–Fe₃O₄ composite beads are shown in Figure 1. SEM analysis showed that the magnetic beads were well dispersed on the chitosan surfaces, indicating a homogeneous combination of the constituents. The average diameters of the dry and wet beads were 0.75 mm and 1.5 mm respectively. Similar morphological images were viewed by previous researchers on chitosan composites (Xu et al. 2013, Dinçer et al. 2012).

Figure 1. Microscopy photographs (A) and SEM micrograph of chitosan and magnetic chitosan–clay-beads (B) natural chitosan (30,000×) and (C,D) magnetic chitosan–clay-beads (C: 30,000×, D: 200×).

The FTIR spectra corresponding to chitosan, chitosan–clay, and magnetic chitosan–clay beads are shown in Figure 2. As seen in Figure 2A (a), the peak around 3400 cm^{− 1} is attributed to the –OH group. The C–H stretching vibration of the polymer backbone is manifested through strong peaks at 2925 cm^{− 1} and 2855 cm^{− 1}. The stretch vibrations of C–O are found at 1084 cm^{− 1}. The characteristic adsorption bands appeared at 1589 cm^{− 1} which can be assigned to N–H bending vibration, and peaks at 1399 cm^{− 1} appeared due to C–O stretching of the primary alcoholic group in chitosan. In Figure 2A (b), the peaks between 600–400 cm^{− 1} can be associated respectively with Si–O stretching, Al–O stretching, and Si–O bending. Similar results were reported for chitosan and for chitosan–clay (Auta and Hameed 2014, Bayramoglu and Arica 2008). In Figure 2A (c), a new sharp peak at 580 cm^{− 1} relates to the Fe–O group. Wu et al. (2012) reported that the peak around 608 cm^{− 1} is owing to Fe–O vibration, which confirms the presence of magnetite nanoparticles. Their spectrum also shows that there are many oxygen-containing groups on the surface of the nanocomposites.

Figure 2. Characterization of chitosan and magnetic chitosan–clay beads (A) FTIR spectra (B) TGA curves. [(a) Chitosan beads, (b) chitosan–clay beads (c) magnetic chitosan–clay-beads].

The thermal decomposition of the chitosan and its composites was examined using thermogravimetric analysis, and the results are shown in Figure 2B. All thermal decomposition profiles exhibit two main stages, with one starting at around 100°C and another at 230°C. At 230°C, the mass losses for samples of chitosan, chitosan–clay, and magnetic chitosan–clay were 26%, 21%, and 16%, respectively. Finisie et al. (2001) and Flores–Hernández et al. (2014) reported that the weight loss observed from chitosan at around 200–300°C probably corresponded to the decomposition and elimination of the polymeric constituent. Chitosan's mass loss was higher than that of the magnetic chitosan and chitosan–clay at high temperatures. The addition of clay and magnetic Fe₃O₄ nanoparticles in chitosan usually make the chitosan more resistant to degradation when compared to the chitosan. From the results of the TGA curve, it is evident that it was found that the thermal stability of the composite was higher than that of the pure chitosan, which was obviously related to the existence of thermally stable clay. These results agreed with those of Zhao et al. (2006), who reported that the temperature of the fourth peak is as high as 712°C, and the percentage of mass loss is about 5.8%, which is due to the phase transition from Fe₃O₄ to FeO, because FeO is thermodynamically stable above 570°C in the phase diagram of the Fe–O system.

The separation ability of the magnetic chitosan–clay beads by an external magnet are demonstrated in Figure 3. Magnetization of magnetic Fe₃O₄ nanoparticles and magnetic chitosan–clay beads was achieved at 25°C. The saturation magnetization of magnetic chitosan beads was about 7.32 emu/g, while for the magnetic Fe₃O₄ nanoparticles the magnetization was 43.66 emu/g. Compared with non-magnetic carriers, the magnetic property of the magnetic chitosan beads is a significant advantage for enzyme immobilization. Magnetic chitosan beads are used as the support material and they can be easily separated from the reaction medium with conventional permanent magnets for the purpose of reuse, thus the use of magnetic particles can reduce the capital and operational costs (Xiao et al. 2006).

Figure 3. Demonstration of magnetic separation of magnetic chitosan–clay beads.

Properties of free and immobilized laccases

After optimization, the percentage of immobilization yield and enzyme activity per gram of carrier were recorded as 75% and 2.90 U/g for magnetic chitosan–clay beads. Several reports have described new support for laccase immobilization. Lu et al. (2006) reported that activity of laccase immobilized in alginate-chitosan microcapsules was 6.84 U per 1.5 g of preparation. Cabana et al. (2007) obtained crosslinked aggregates with specific activity of 0.148 U/mg protein, and Roy and Abraham (2006) reported the value of specific activity of laccase-crosslinked crystals to be about 2300 U/mg protein. D’Annibale et al. (1999) reported that activity of laccase on chitosan beads was 85.5 U/g of support. All these immobilization studies published in the literature have been performed under different conditions, and different organisms were used as the laccase source. Therefore, it is almost impossible to compare these immobilization results. The effect of pH on the activity of the free and immobilized laccase on magnetic chitosan–clay was investigated at different pH values varying from 3.0 to 7.0 (Figure 4A). The free and immobilized laccase exhibit maximal activity at pH 4.0 and pH 5.0, respectively. The shift of optimum pH was attributed to the electrostatic interaction influenced by the carrier microenvironment (Jiang et al. 2005). The covalently immobilized laccase showed broadening in the pH–activity profile as compared to the native enzyme, which means that the immobilization methods preserved the enzyme activity over a wider pH range (Sarı et al. 2006).

Figure 4. Effects of (A) pH (0.05 mol/l acetate/phosphate buffer, 30°C) (B) temperature (0.05 mol/l acetate buffer, pH 5.0), Data were shown as mean ± SD (n = 3).

Catapane et al. (2013) also showed that the optimal pH for free and immobilized laccase on PAN (Poly Acrylo Nitrile) was about 5; however, the activity of immobilized enzyme in the pH range of 3–7 was retained significantly, in compare with free enzyme. Spinelli et al. (2013) immobilized (Trametes versicolor) laccase on Amberlite IR-120 H beads. According to their results, the optimum pH was shown to be 4.5. Kumar et al. (2012) reported that optimum pH was 4.0 for laccase using ABTS as a substrate, and the lower relative activity was obtained at pH 7.0. Furthermore, the majority of fungal laccases have been found to function as laccase under mild acidic condition (pH 4–6).

The effect of temperature on activity of free and immobilized laccases is shown in Figure 4B. Both laccases showed an optimum reaction temperature at about 40°C. Laccases from different white rot fungi have been reported to show optimum activities in the temperature range of 40–65°C. The catalytic activity of fungal laccases is optimum in the temperature range between 30 and 55°C. Kumar et al. (2012) reported that optimum temperature was 45°C for laccase, and that the laccase was active over a broad range of temperatures of 28–5°C. According to Spinelli et al. (2013) optimum temperatures were 40 and 45°C for free and immobilized laccase, respectively. The thermal stability is one of the most important features for the application of the biocatalyst. The covalent immobilization of an enzyme to a carrier often limits its freedom to undergo drastic conformational changes, thus resulting in increased stability towards denaturation thermal stability (Osma et al. 2010, Sarı et al. 2006). In the present study, laccase immobilization led to a significant stabilizing effect towards heat denaturation at 60°C and 70°C (Figure 5A). These results agreed with those of Kunamneni et al. (2007) and Forde et al. (2010), who reported that the immobilization process induces 10–15% enhancement within the tested temperature range. Moreover, the activity of the immobilized laccase was greater at high temperatures (50–60°C) compared with that of the free counterpart. Several studies have shown the temperature resistance of immobilized enzyme in contrast with the free form (Başak and Aydemir 2013, Dinçer et al. 2012). Nicolucci et al. (2011) also showed that the relative activity of immobilized laccase was about 80% at 70°C, while the free laccase was more or less inactivated.

Figure 5. (A) Temperature stability (0.05 mol/l acetate buffer pH 5.0, incubation time 2 h) and (B) storage stability (0.05 mol/l acetate buffer, pH 5.0, 4°C) of the free and immobilized laccase onto magnetic chitosan–clay beads. Data are shown as mean ± SD (n = 3).

Storage stability is one of the most significant indices for evaluating enzyme properties. Figure 5B shows the storage stabilities of free and immobilized laccase at 4°C. With increased storage time, a remarkable difference was observed between the activity variations of free and immobilized laccase. Free laccase completely lost its initial activity, whereas immobilized laccase retained about 55% after 6 weeks. Thus far, the mechanisms proposed for the storage stability of immobilized laccase are complex and unclear. One possible explanation may be the covalent bonding between the laccase molecules and support, which improved the enzyme's stability against structural denaturation (Osma et al.2010, Forde et al. 2010, Sarı et al.2006).

Kinetic parameters were estimated from the Lineweaver–Burk plot using ABTS as substrate. Immobilization decreased at Vmax value from 26. 96 U/mg to 12.05 U/mg. The K_M value of immobilized laccase (0.410 mM) was 1.4 times higher than that of free laccase (0.297 mM), which means the immobilized laccase had lower affinity towards the substrate, in agreement with other investigators reporting significant decreases in affinity in immobilized catalysts (Rekuc et al. 2009, Cabana et al. 2007). Immobilization of laccase on polypropylene membrane by covalent bonding was explored by Georgieva et al. (2010), and the K_M value increased 1.8-fold by using phenol as the substrate for the kinetic study. Bayramoglu et al. (2013) reported the K_M values increased 2.3-fold after laccase was immobilized on p(HEMA-g-GMA) film. The ratio Vmax/K_M is a measurement of the catalytic efficiency of an enzyme–substrate pair. In this study, the catalytic efficiencies of the free and immobilized laccase were found to be 0.090 s^{− 1} and 0.029 s^{− 1}, respectively. The catalytic efficiency of laccase was decreased by about 3-fold upon immobilization. The decreasing trend of the ratio Vmax/Km may have occurred due to inactivation of the enzyme and the production of intermediate compounds during degradation of the substrate (Nicolucci et al. 2011). The increase in K_M might be caused by the steric hindrance of the active site by the support, the loss of enzyme flexibility necessary for substrate binding, or diffusion limitation of substrate and products because of porous nature of the support (Silva et al. 2012).

Phenol removal and reusability

Application of the immobilized system in phenol removal was investigated in a batch system, and phenol removal values of about 80% were obtained at the end of 4 h at pH 5.0 and 25°C (Figure 6A). The optimum pH of 5 was achieved for enzymatic degradation of the pollutant, which is in agreement with the results of other research studies (Bayramoglu and Arıca 2008, Georgieva et al. 2010). However, the optimum pH reported by Okazaki et al. (2002) was 3 for laccase from Coriolus versicolor converting α-phenylenediamine. This difference may be due to differences in the type and concentration of the buffer used and the purity of the enzyme (Okazaki et al. 2002). To determine how carrier adsorption affected removal efficiency of phenol, the heat-denatured immobilized enzyme was used in place of the intact immobilized laccase under the same experimental condition. The adsorption of phenol by the support surface was around 18%. Therefore, the removal of phenol was due to the combined effect of biodegradation and adsorption. Many studies have already reported similar results where dyes and phenol were adsorbed by carriers (Rekuc et al. 2009, Bayramoglu and Arica 2008). Liu et al. (2012) claimed that nearly 20% of the removal of phenolic compounds was attributed to the adsorption by the mesoporous support.

Figure 6. (A) UV–Vis spectrum of phenol removal (10 ppm) before and after treatment by immobilized laccase. (B) Reusability of immobilized laccase in phenol removal (Reaction conditions: 10 ppm phenol, pH 5.0, 4 h of contact time, 30°C). Data shown as mean ± SD (n = 3).

Reusability of immobilized enzymes is an important aspect for industrial applications, because immobilized enzymes decrease the cost of production due to their repeated, continuous, or batch uses. After the tenth use, the residual activity for immobilized enzyme was found to be 76%, (Figure 6B). Decrease in the enzyme activity upon repeated usage is expected. Upon repeated uses, either blocking of some pores of support by substrate or product may take place, or enzyme may lose its activity and denature. In literature, there are activity values reported as retained at 60% after 10 batch uses for covalently immobilized laccase on activated polyvinyl alcohol (Yinghui et al. 2002), and at 80% after 5 consecutive operations for laccase immobilized on amino-terminated magnetic nanocomposites by the crosslinking method (Xiao et al. 2006). Other studies in the literature show that immobilized laccase on Eupergit C support retained 65% of initial activity after 10 cycles (Lloret et al. 2012), while immobilization on amine-terminated magnetic nanocomposites retained 80% of activity after 5 cycles of reaction (Xiao et al. 2006).

Conclusion

One of the most important aims of enzyme technology is to enhance the conformational stability of the enzyme through immobilization procedures. The extent of stabilization depends on the enzyme structure, immobilization methods, and type of support. In this paper, laccase from Trametes versicolor was immobilized by covalent attachment on magnetic chitosan–clay composite beads by glutaraldehyde crosslinking. The immobilized enzyme exhibited the maximal activity at pH 5.0. The K_M value of immobilized laccase was 0.410 mM, higher than that of free laccase (0.297 mM), which means that the immobilized laccase had lower affinity towards the substrate. The immobilized biocatalyst displays an improved thermal and storage stability paired with a good performance for reusability. The catalytic performance of immobilized laccase was evaluated by the degradation of phenol. The removal efficiency of phenol by immobilized laccase was about 80% under the optimum conditions (pH 5.0, 25°C) after 4 h.

Declaration of interest

The authors report no declarations of interest. The authors alone are responsible for the content and writing of the paper.