Improvement of catalytic efficiency of chloroperoxidase by its covalent ...

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Jun 20, 2012 - Abstract. Chloroperoxidase from the fungus Caldariomyces fumago was covalently immobilized on SBA-15 mesoporous material and assayed ...

J Porous Mater (2013) 20:387–396 DOI 10.1007/s10934-012-9608-8

Improvement of catalytic efficiency of chloroperoxidase by its covalent immobilization on SBA-15 for azo dye oxidation E. Guerrero • P. Aburto • E. Terre´s • O. Villegas • E. Gonza´lez • T. Zayas • F. Herna´ndez • E. Torres

Published online: 20 June 2012 Ó Springer Science+Business Media, LLC 2012

Abstract Chloroperoxidase from the fungus Caldariomyces fumago was covalently immobilized on SBA-15 mesoporous material and assayed for the enzymatic oxidation of four azo dyes. All dyes were oxidized by the free and the immobilized enzyme to different extent. Acid Blue 120 and Direct Blue 85 dyes were decolorized almost completely. The catalytic efficiency, kcat/KM, of the immobilized enzyme was 27, 2.9, 137 and 28 times higher than the free enzyme for Basic Blue 41, Disperse Blue 85, Acid Blue 120 and Direct Black 22 oxidation, respectively. The immobilized enzyme displayed improved thermostability and a similar pH profile compared to the free enzyme. The immobilized chloroperoxidase showed excellent storage stability and maintained 100 % catalytic activity after 90 days at both 4 and 25 °C. Keywords Chloroperoxidase  Azo dyes  SBA-15  Enzymatic decoloration

1 Introduction The intensive use of textile dyes has increased their concentration in water effluents due to leaks or spills, and also due to the low efficiency of the dyeing process; in addition,

E. Guerrero  P. Aburto  O. Villegas  E. Gonza´lez  T. Zayas  F. Herna´ndez  E. Torres (&) Centro de Quı´mica-ICUAP, Beneme´rita Universidad Auto´noma de Puebla, Edificio 103 G, Ciudad Universitaria, 72570 Puebla, Mexico e-mail: [email protected] E. Terre´s Instituto Mexicano del Petro´leo, Eje Central La´zaro Ca´rdenas 152, Col. San Bartolo Atepehuacan, 07730 Mexico D.F., Mexico

the low efficiency of some environmental treatment processes causes the dye to reach some bodies of water without being transformed. It is estimated that every year, 10 % of the amount of dyes used (about 80,000 tons) is discharged as effluents [1]. The textile industry is one of the main sources of polluting dyes; it discharges wastes containing a large variety of dyes: azo, disperse, basic and acid dyes. The azo dyes are the most commonly used dyes in textile processes and thus they constitute the largest group of all organic dyes on the market [2, 3]. Their chemical structure is characterized by the presence of the azo group N=N as a chromophore, associated with an auxochrome such as amine or hydroxyl groups. These compounds become contaminants with the production of mutagenic amines by natural degradation processes [4–7]. In addition, they affect the water quality by preventing the passage of light, by increasing the chemical oxygen demand (COD), and also they have a drastic visual effect at concentrations as low as 2 mg/L [8]. It is estimated that around 90 % of the azo dyes discharged to the environment is not degraded by conventional biological treatment processes [1, 9–11]. Other treatment processes, such as adsorption or chemical oxidation, generate contaminant-saturated adsorbents or sludges and oxidizing substances which are harmful to the environment [12, 13]. The enzymatic treatment of pollutants is an innovative process performed in mild conditions of reaction without generating toxic sludge or discharging aggressive chemicals into the environment, and that could result in less toxic effluents [14]. Enzymes with the ability to oxidize dyes include peroxidases and laccases from plants or fungi [10, 14, 15]. The toxicity of azo dyes may be drastically reduced after their enzymatic oxidation by peroxidases and



laccases where no mutagenic amines are produced, and more biodegradable products are formed [16, 17]. However, due to the susceptibility of enzymes to inactivation by the presence of other chemicals in water effluents, it is likely that enzymatic treatment would be most effective in those conditions where the highest concentration of target contaminants and the lowest level of other contaminants are present, which is unusual in industrial wastes. Additionally, the fluctuating conditions of pH or temperature can affect enzyme functionality. Therefore, it is necessary to produce or find more robust enzymes for a viable enzymatic treatment. An effective method for improving the functionality of enzymes is their immobilization in inorganic supports such as mesoporous materials [18–20]. Immobilization of enzymes within a pore or onto a surface with certain physical and chemical characteristics, has enhanced enzyme performance and produced more robust and adaptable biocatalysts [21, 22]. Enzyme immobilization on these materials has resulted in the stabilization of different peroxidases to temperature, pH, and some denaturing agents like organic solvents and urea and it has also allowed its reuse for several reaction cycles [21, 23], which it is important to the economy of the process [24]. In this work we report the oxidative capacity of the chloroperoxidase (CPO) against four different dyes from the textile industry and its immobilization on modified SBA-15 mesoporous material to improve its stability to temperature and pH.

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and metals [26–34]. Briefly, 4 g of Pluronic P123 were dissolved in 105 mL of water with HCl 37 % (20 mL) and the solution was stirred for a few hours. Then, 9 g of tetraethylorthosilicate were added and the resulting solution was first heated at 40 °C for 5 days and subsequently at 100 °C for another 6 days. The SBA-15 material was obtained by filtration, dried and calcined in air flow at 550 °C for 2 h. 2.3 SBA-15 modification Two hundred milligram SBA-15 were incubated with 2 mL of APTES for 24 h at room temperature in 10 mL anhydrous methanol. This modification at room temperature has proven to be successful in several reports [35–39]. After that, it was washed 10 times with methanol to remove the un-reacted APTES. The modified material was then dried and subjected to a second modification with glutaraldehyde. To this end, 100 mg SBA-15 were mixed with 3 mmol glutaraldehyde in 10 mL 60 mM acetate buffer pH 4 for 4 h with agitation. Finally, the material was washed 10 times with acetate buffer to remove the un-reacted glutaraldehyde. This last step was done to assure that not free glutaraldehyde was in the solution at the time the enzyme was added, avoiding enzyme crosslinking. SBA-15 functionalization was checked through FTIR spectroscopy on a 470 FTIR Nicolet spectrophotometer operating in diffuse reflectance mode using KBr powder. All spectra consisted on 32 consecutive scans with 2 cm-1 resolution.

2 Experimental 2.4 CPO immobilization 2.1 Reagents 3-Aminopropyl triethoxysilane (APTES), hydrogen peroxide, glutaraldehyde, and poly (alkyleneoxide) triblock Pluronic F127 (EO106PO70EO106, 99 %, MW = 12,600) were purchased from Sigma Chemical Company. Tetraethylorthosilicate (TEOS, 98 %) was obtained from Aldrich Company. Buffer salts were purchased from J.T. Baker. CPO from Caldariomyces fumago (42 kDa, Rz = 1.4, specific activity 22,000 min-1 based on monochlorodimedone assay), was kindly provided by Dr. Michael A. Pickard from the University of Alberta, Canada. All dyes—Direct Black 22, Acid Blue 120, Direct Blue 85 and Basic Blue 41—were purchased from BASF Chemicals Company. 2.2 Synthesis of SBA-15 mesoporous material The SBA-15 material was synthesized using Pluronic P123 as a structure-directing agent according to the method reported originally by Zhao et al. [25], which has been applied in several recent reports to immobilize enzymes


Enzyme immobilization was carried out by mixing 50 mg of the modified material (SBA-15-glutaraldehyde) with 100 nmol CPO (ratio 30:1 glutaraldehyde:CPO) at 4 °C, pH 4 for 4 h with stirring. The material was then recovered by centrifugation and washed with 60 mM acetate buffer pH 4. The amount of protein adsorbed was measured by difference, with protein concentrations determined at 398 nm (e = 85,000 M-1 cm-1) before and after CPO adsorption [40]. The final enzyme preparation was kept in 1 mL 60 mM acetate buffer pH 4. 2.5 Enzymatic dye oxidation The catalytic activity of the free and the immobilized CPO against the dyes was determined spectrophotometrically by measuring the change in absorbance at the wavelength of maximum absorption of each dye (Table 1). Reactions were carried out at room temperature in 1 mL reaction mixtures containing different dye concentrations to reach an absorbance of 0.5 in 60 mM acetate buffer pH 3.0

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Table 1 Structure and maximum absorption wavelength (kmax) of azo dyes tested Dye

kmax (nm)


Direct Black 22


Basic Blue 41


Acid Blue 120


Disperse Blue 85


containing 20 mM KCl, except for Direct Black 22, whose solubility in water allows only to reach an absorbance of 0.2. For reactions with the immobilized enzyme, 10 lL of material solution (0.5 mg enzyme preparation). Reactions were started by the addition H2O2 (1 mM final concentration). Two control experiments were done following the absorbance changes of all dyes during 10 min with the enzyme preparation in the absence of hydrogen peroxide (to check for dye adsorption) and with hydrogen peroxide in the absence of the enzyme preparation (to check for chemical oxidation). For all reactions, the amount of free and immobilized enzyme was 2.14 nmol.

2.6 Kinetic constants determination Reactions were performed in 1 mL of 60 mM acetate buffer pH 3.0, 20 mM KCl, 2.14 nmol CPO (free or immobilized) and different hydrogen peroxide concentrations, while the azo dye concentration was kept constant. The initial reaction rates were obtained by following the decrease in absorbance at the wavelength of maximum absorption of each dye. The values of all kinetics parameters were adjusted to Michaelis– Menten equation with an iteration procedure following the Marquardt–Levenberg nonlinear least-squares algorithm using Origin 7.0 software.



2.7 Effect of pH and temperature on enzyme activity The effect of pH on the catalytic activity of the free and the immobilized CPO against the dyes was determined spectrophotometrically by measuring the changes in absorbance at the wavelength of maximum absorption of each dye at different pH values, from 3 to 7, keeping the temperature constant at 25 °C. The catalytic activity data were normalized to the catalytic activity at 25 °C, which was considered as 100 % activity. The same methodology was used to study the effect of temperature on the catalytic activity, while keeping the pH constant at 3 and changing the temperature from 25 to 70 °C. The catalytic activity data were normalized to the catalytic activity at 25 °C, which was considered as 100 % activity. 2.8 Storage stability The immobilized enzyme was kept at 4 and 25 °C and the substrate conversion was measured throughout. The conversion was measured as the change of fluorescence of a model substrate, 4,6-dimethyl-dibenzothiophene (DMDBT), after incubation with immobilized enzyme for 5 min. Conversion at time zero corresponds to 100 % transformation of 20 nmol of DMDBT. Reactions were carried out at room temperature in 3 mL reaction mixtures containing 20 lM DMDBT, 1 mM H2O2, 20 mM KCl in 60 mM acetate buffer pH 3. 2.9 Biocatalyst reusability The reusability of the prepared biocatalysts were assayed in three sequential cycles for the oxidation of Acid Blue 120 as mentioned earlier. The immobilized CPO was recovered by centrifugation at 14,000 rpm during 1 min after each reaction cycle and placed in a fresh reaction media. The residual activity data were normalized to the first cycle assay, which was considered as 100 % activity.

3 Results and discussion Chloroperoxidase (EC has the capacity to oxidize a large variety of organic pollutants such as phenols, polycyclic aromatic hydrocarbons, and pesticides [41–43]. However, the application of enzymes as biocatalysts in environmental treatment processes is limited because the low activity and the fragile nature of these enzymes. In order to improve its biocatalytic functionality, we covalently immobilized the enzyme on a SBA-15 mesoporous material. To this end, we first modified a SBA-15 with an organosilane derivative (APTES) bearing a free amino


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group (Scheme 1). The resulting material was then reacted with glutaraldehyde, which subsequently reacts with the amino groups of the only three lysines on enzyme surface (Lys 112, Lys 145, Lys 211) (Scheme 1). Such lysines are exposed to the environment and located on the opposite side of the substrate access to the heme group [44]. Such immobilization procedure would allow a proper orientation of the enzyme active site towards the bulk medium [45]. As a first step we synthesized, modified and characterized the mesoporous material SBA-15. 3.1 Material characterization The X-ray diffraction (XRD) pattern of the parent material (Fig. 1a) showed three well-resolved peaks with distances ˚ , that correspond to (1 0 0), (1 1 0) of 93.9, 54.5 and 47.4 A and (2 0 0) reflections. The latter indexed to a hexagonal ˚ (a0 = 2d(1 0 0)/H3), structure with lattice constant of 107.8 A which is typical for a SBA-15 pore structure, as reported elsewhere [32]. The N2 isotherm adsorption/desorption curve of the SBA-15 sample displayed a type IV isotherm with a H1 hysteresis typical of mesoporous materials having cylindrical parallel channels (Fig. 1b). This last result was confirmed by the narrow pore size distribution displayed (Fig. 1c). The surface area of SBA-15 mesoporous silica was 935 m2/g; the total pore volume was 1.1 cm3/g and pore size 6.7 nm, as calculated by the desorption branch through the BJH method. Chemical modification of SBA-15 with APTES and glutaraldehyde was confirmed by FTIR (Fig. 2). Infrared spectroscopy has been one of the main spectroscopic tools used to probe functionalization of mesoporous materials [46–50]. The spectrum of the unmodified SBA-15 has a broad band with a maximum around 3,400 cm-1 attributed to the stretching surface silanol groups and the remaining adsorbed water molecules [51], one more around 990 cm-1 due to Si–OH stretching [46] and one at 800 cm-1 due to Si–OH bending [46] (Fig. 2a). After chemical modification with APTS, the O–H band (3,400 cm-1) was less intense in comparison with that of the SBA-15 sample, indicating differences in the content of O–H groups (Fig. 2b). The presence of APTES in the modified SBA-15 was further corroborated by a broad band at 3,000–3,300 cm-1, attributed to the NH stretching vibration [51]. Some other evidences of SBA-15 modification were detected, such as the disappearance of the band at 990 cm-1 due to Si–OH stretching and the appearance of a small band at 690 cm-1 due to N–H bending [52]. In addition, a characteristic band for aliphatic C–H stretching vibrations attributed to alkyl chains at around 2,930 cm-1 [53]. Finally, a small band at 1,520 due to N–H bonds vibration appeared [30, 53]. All this evidence confirms that silanol groups from SBA-15 reacted with APTS. Additional changes were observed after

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Scheme 1 Covalent immobilization of CPO on mesoporous material SBA-15 using a spacer arm (APTES) and glutaraldehyde

incubation with glutaraldehyde. A signal at 1,647 cm-1 (–C=N– bonds), due to the reaction between the –NH2 of APTS-modified SBA-15 and –CHO of glutaraldehyde, was detected (Fig. 2c) [53]. In addition, the band at 1,520 cm-1, due to N–H vibration, almost disappeared. One limitation of our FTIR analysis of SBA-15 material is that it was not possible to quantitatively assess the extent of functionalization with APTES and glutaraldehyde, so just a qualitative analysis was performed. The derivatized SBA-15 was incubated with CPO for 4 h, obtaining a load of 214 nmol CPO/g SBA-15. 3.2 CPO-mediated dye oxidation The oxidation of four azo dyes, whose structures are shown in Table 1, catalyzed by the free and the immobilized CPO, was assayed. All dyes—Direct Black 22, Acid Blue 120, Direct Blue 85 and Basic Blue 41—were oxidized by the enzyme to different extent. Acid blue 120 and Disperse blue 85 were the best substrates for both enzyme preparations (Fig. 3). The percentage of dye oxidation was

higher for the free enzyme; however, the specific activity (DAbs/min nmol CPO) of the immobilized enzyme was up to 20 times higher for the oxidation of azo dyes (Table 2), which means that the same amount of CPO transforms the dye faster when it is immobilized (Table 2). Control assays showed that neither dye oxidation nor adsorption took place in the absence of the enzyme. The mechanism for the peroxidase-catalyzed oxidation of phenolic-azo dyes involves two successive one-electron oxidations of the phenolic ring by the oxidized form of the enzyme, which produces a carbonium ion. Then, a water molecule reacts with the phenolic carbon bearing the azo linkage and an unstable hydroxyl intermediate is formed, which breaks down into a quinone and an amidophenyldiazine. The latter compound is then oxidized by oxygen into the corresponding phenyldiazene radical, which after elimination of nitrogen gives a phenyl radical finally oxidized by oxygen. This mechanism leads to the detoxification of azo dyes since no aromatic amines are formed [17]. To learn more about the kinetic capabilities of the prepared biocatalyst, the kinetic constants of the soluble and



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Fig. 2 FTIR spectra of neat (a), silanized (b) and glutaraldehydemodified (c) SBA-15

Fig. 1 a Small angle XRD pattern, b pore size distribution and c N2 adsorption–desorption isotherms of SBA-15 mesoporous material

the ing ing the

immobilized enzyme were determined under increasconcentration of hydrogen peroxide while maintainconstant the dye concentration. Table 3 summarizes results. It can be concluded that the covalent


immobilization of CPO on the SBA-15 material resulted in a much more active biocatalyst with kcat values up to 40 times higher than the free enzyme. The KM values were slightly higher for the immobilized CPO and the catalytic efficiency, as measured by the ratio kcat/KM, was up to 137 times higher for the immobilized enzyme (Table 3). To our knowledge, very few works have demonstrated this effect in catalytic activity when CPO is immobilized. Our own previous reports and others have shown significant decreases in CPO catalytic activity for oxidation of various substrates [45, 54–56]. However, very recently Aguila et al. [26] reported an increase in the catalytic activity of CPO through its immobilization on three silica nanostructured

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glutaraldehyde may provide a more suitable microenvironment for the catalysis of this enzyme. This positive outcome reinforces a potential enzymatic treatment of azo dyes mediated by a more active biocatalyst. 3.3 Effect of pH and temperature on catalytic activity of free and immobilized CPO

Fig. 3 Percentage of oxidation of azo dyes by free and immobilized CPO

Table 2 Biocatalytic activity of azo dye oxidation by the free and in the immobilized CPO Azo dye

Specific activity (DAbs/min nmol CPO) Free CPO

Immobilized CPO

Direct Black 22

0.84 ± 0.078

16.39 ± 1.11

Acid Blue 120

2.00 ± 0.095

35.23 ± 3.89

Disperse Blue 85 Basic Blue 41

1.35 ± 0.056 0.73 ± 0.073

24.02 ± 0.89 11.84 ± 1.86

supports with different pore sizes MCM-41 (3.3 nm), SBA15 (6.4 nm) and MCF (12.1 nm). The adsorbed CPO into SBA-15 and MCF showed total turnover numbers of 48,000 and 54,000 times higher than free CPO, respectively for styrene oxidation [26]. In another work, Hartmann and Streb [57] reported higher conversions of indole when the CPO was immobilized in SBA-15, although no values of catalytic activity were reported. Finally, Jung et al. [58] reported the crosslinking of chloroperoxidase molecules with glutaraldehyde (CLEAS of CPO) in the cages of mesocellular foam material, resulting in biocatalysts that were four times catalytically more active than the conventional catalyst prepared by physisorption of CPO. In our case, modification of amino groups of the enzyme, located on the opposite side of the entrance channel to the active site, and the presence of a larger spacer such as

As mentioned earlier, industrial wastewaters normally have physicochemical properties unsuitable for enzymatic catalysis, such as the presence of a wide variety of organic and inorganic compounds, metals, microorganisms and varying conditions of pH and temperature. As it is already known, the activity of an enzyme is affected by pH and temperature. Usually, enzymes display a bell-shaped activity versus these two variables, with a drastic reduction after its optimum value due to enzyme denaturation. Thus, it is necessary to develop robust biocatalysts for application in the environmental treatment of pollutants. Therefore, the catalytic profiles of our biocatalyst preparation at different pH and temperatures, two parameters of relevance towards the development of stable biocatalyst, were determined [59]. Figure 4 shows the pH profile of both immobilized and free enzyme in the oxidation of the different azo dyes. As shown, the optimum pH of the free enzyme is 3.5 except for the dye Disperse Blue 85, which showed higher oxidation rate at pH 4 with a significant increase in the specific activity of oxidation from 1.35 to 6.7 DAbs/min nmol CPO. The free enzyme was catalytically active just in acidic media, up to pH 5.5 for disperse blue 85, showing only 5 % of the catalytic activity with respect to pH 3. For the immobilized enzyme, the catalytic profile is similar, displaying an optimum pH of 4 for the oxidation of Disperse Blue 85, and 3.5 for the rest of the dyes. For Disperse Blue 85 the oxidation rate increased 5.85-fold, when the pH was changed from 3 to 4. For all dyes, the catalytic activity was higher for the immobilized enzyme in the whole range of pH tested. It seems that above pH 5, the change in the ionization state of some amino acid residues affect the catalytic activity, and no structural changes are expected, until pH 7.5 is reached [60]. Therefore, it seems that ionization state

Table 3 Kinetic constants for azo dye oxidation by the free and the immobilized CPO Dye

Basic Blue 41

kcat (DAbs/min nmol CPO)

KM (mM)

Free CPO

Free CPO

Immobilized CPO

kcat/KM (DAbs/min nmol CPO mM) Immobilized CPO

Free CPO

Immobilized CPO







Disperse Blue 85







Acid Blue 120







Direct Black 22









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140 Immobilized CPO

Catalytic activity (%)


Free CPO

100 80 60 40 20 0 25












Temperature ( oC)

Fig. 5 Effect of temperature on catalytic activity of free and immobilized CPO for Acid Blue 120 oxidation. The 100 % activity values are 0.84 and 16.39 DAbs/min nmol for free and immobilized CPO, respectively

Fig. 4 Effect of pH on catalytic activity of free and immobilized CPO for the oxidation of all azo dyes. The 100 % activity values for Acid Blue 20, Direct Black 22, Disperse Blue 85 and Basic Blue 41 are, respectively: (free CPO) 0.84, 2.00, 1.35, and 0.73 DAbs/ min nmol; (immobilized CPO) 16.39, 35.23, 24.02, and 11.84 DAbs/ min nmol

of the amino acid residues are not affected in the immobilized enzyme. Although an improvement on the catalytic behavior of CPO was not observed after its immobilization, some reports in the scientific literature have evidenced that mesoporous materials may alter the effect of pH on catalytic profile of some enzymes. For example, Hartmann and Streb [57] observed an improvement of CPO in the conversion of indole at different pH when the enzyme was physically immobilized on SBA-15. In another report, the optimum pH for free and immobilized laccase on magnetic mesoporous silica nanoparticles was observed at pH 4.0 and pH 5.0, respectively [61]. The shift of the optimum pH after enzyme immobilization was related to the ionic interaction between the enzyme and the charged surface of the support. In comparison with the free laccase, the immobilized laccase exhibited more than 95 % of the maximum activity with a wider pH range between 4.0 and 5.5 [61]. Figure 5 shows the temperature profile of the free and the immobilized CPO for the oxidation of Acid Blue 120, the best substrate tested in this work. The free enzyme was


catalytically active in a temperature range of 25–75 °C, exhibiting low sensitivity to temperature changes up to 40 °C. From 45 °C, the catalytic activity decreased linearly, down to 50 % at 60 °C. On the other hand, the immobilized enzyme was active up to 80 °C and showed higher catalytic activities throughout the temperature range tested. At 60 °C, the immobilized enzyme still maintained 77 % of the activity, while at 80 °C, 50 % of the catalytic activity was still observed. In addition, the catalytic activity at 80 °C was similar in magnitude to that shown by the free enzyme at 25 °C, about 30 DAbs/min nmol of enzyme. Therefore, it is evident that the immobilized enzyme is more stable with regards to temperature than the free enzyme. Scientific reports have confirmed the tendency of mesoporous materials to protect enzymes from inactivation by temperature [62, 63]. In our previous works, we reported a slight improvement of CPO when it was immobilized by ˚ pore size, close to physical adsorption on SBA-16 of 90 A the diameter of the enzyme [37]. As has been reported by Takahashi et al. [62, 63] pores for the immobilization of horseradish peroxidase should be large enough to accommodate the enzyme but should be matched to size in order to stabilize it by providing a restricted area of movement, reducing its flexibility and avoiding denaturation. Additionally, the functionalization of mesoporous silica support has been employed to improve the thermostability of peroxidases. Montiel et al. [32] reported that the residual thermal activity of CPO was improved by covalent ˚ pore size, which was immobilization on SBA-15 of 143 A chemically derivatized using 3-aminopropyl triethoxysilane and succinic anhydride. Wang et al. [61] researched the thermal stabilization of laccase by its immobilization via metal affinity adsorption on large-pore magnetic

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inactivate the enzyme. To check for enzyme leaching as reason for the lost of activity we assayed the oxidation of acid blue 120 with the supernatant after each centrifugation step, however no enzyme activity was detected, suggesting no enzyme leaching from SBA-15.

4 Conclusions

Fig. 6 Storage stability of immobilized CPO at 4 and 25 °C. 100 % conversion means complete transformation of 20 nmol of 4,6 dimethyl-dibenzothiophene

mesoporous silica nanoparticles. The material was derivatized with iminodiacerate through a silane-coupling agent and chelated with Cu. Therefore, not only the pore size of the mesoporous material is important for thermostabilization of enzymes, but also the organic spacer used to attach the protein. 3.4 Storage stability Figure 6 shows the catalytic activity of the immobilized CPO after several weeks of incubation at two different temperatures, 4 and 25 °C. As can be concluded, the immobilization on SBA-15 allowed the enzyme to retain the totality of the activity after three months of storage at both temperatures. Therefore, immobilization on SBA-15 provides not only a more active but also a stable biocatalyst with respect to temperature and storage. 3.5 Biocatalyst recyclability We finally tested the reuse of the enzyme preparation in the oxidation of acid blue 120. The immobilized enzyme oxidized 100 % of the dye in the first cycle; however, in the second and third cycles the conversion dropped to 75 and 40 %, respectively. In the fourth cycle no catalytic activity was detected. The enzyme is deactivated significantly after each reaction cycle probably due to inactivation by hydrogen peroxide, which is considered a suicide substrate of peroxidases [64, 65], or by a reaction between some reactive intermediate and the enzyme aminoacids [66, 67]. This last inactivation process occurs predominantly during phenol oxidation by peroxidases through binding with phenoxyl free radicals and the aminoacids residues of the enzyme [68, 69]. As mentioned, azo dye oxidation by peroxidases produces reactive intermediates like unstable hydroxyl intermediate and quinones, which might

The results show that it is possible to improve the functionality of CPO by its covalent immobilization on SBA15. Immobilization on the inorganic support SBA-15 increased the catalytic efficiency of CPO as well as its performance at different pH and temperatures for the oxidation of four textiles azo dyes. Due to industrial wastewaters having physicochemical properties unsuitable for enzymatic catalysis, such as changing conditions of pH and temperature, the immobilized enzyme might perform well in conditions close to those found in real effluents, where the pH and temperature normally fluctuate. Acknowledgments The authors are thankful for the financial support given by SEP-CONACyT 80986, PROMEP 103.5/11/5880.

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