The use of soybean peroxidase in the decolourization of Remazol Brilliant Blue R and toxicological evaluation of its degradation products

Journal of Molecular Catalysis B: Enzymatic 89 (2013) 122–129

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The use of soybean peroxidase in the decolourization of Remazol Brilliant Blue R and toxicological evaluation of its degradation products Maria Cristina Silva a,b,∗ , Juliana Arriel Torres a , Lívian Ribeiro Vasconcelos de Sá b,c , Pricila Maria Batista Chagas a , Viridiana Santana Ferreira-Leitão b,c , Angelita Duarte Corrêa a a

Biochemistry Laboratory, Department of Chemistry, Federal University of Lavras, CEP 37200-000, Lavras, MG, Brazil Biocatalysis Laboratory, Catalysis Division, National Institute of Technology, Ministry of Science and Technology, CEP 20081-312, Rio de Janeiro, RJ, Brazil c Federal University of Rio de Janeiro, Department of Biochemistry, CEP 21941-909, Rio de Janeiro, RJ, Brazil b

a r t i c l e

i n f o

Article history: Received 6 November 2012 Received in revised form 8 January 2013 Accepted 9 January 2013 Available online 17 January 2013 Keywords: Environmental biocatalysis Soybean peroxidase Decolourization Textile dyes Toxicity

a b s t r a c t This study evaluated the potential use of soybean peroxidase in the decolourization of reactive textile dye Remazol Brilliant Blue R (RBBR) and its synthetic effluent. The following parameters were studied: reaction time, dye concentration (10–60 mg L−1 ), enzyme load (4.96–140 U mL−1 ) and H2 O2 concentration (20–1100 ␮mol L−1 ). The maximum removal of RBBR (86%) was obtained after 13 min of reaction, using H2 O2 100 ␮mol L−1 , enzyme 70.4 U mL−1 and RBBR 40 mg L−1 . The toxicity of the products formed after enzymatic treatment was assessed by using Artemia salina and lettuce seeds (Lactuca sativa). Although soybean peroxidase was very efficient in colour removal, the products obtained after enzymatic decolourization presented higher toxicity. The inhibition concentration (IC50 ) obtained for lettuce seeds was 27.9%, and the lethal concentration (LC50 ) for A. salina was 59.3%. The aforementioned results emphasize the importance of toxicological evaluation after enzymatic treatment. The potential application of peroxidases for colour removal and the increase in the products’ toxicity reinforce the need of combined treatments. © 2013 Elsevier B.V. All rights reserved.

1. Introduction The removal of dyes from textile wastewater prior to its discharge or reuse is a challenging task. Chemical and photo stability are required characteristics for industrial dyes. As a consequence, these kinds of structure are recalcitrant in natural [1]. Unsuitable treatments of wastewater containing industrial dyes can damage natural systems, reducing water transparency and the incidence of solar radiation, which can modify photosynthetic activity and the dynamics of gas solubility [2]. Moreover, the degradation of some synthetic dyes may result in toxic and carcinogenic metabolites [3]. No single conventional technology can remove all classes of dyes [4,5]. In conventional biological treatment plants, the dyes are adsorbed into activated sludge and are poorly degraded. Additionally, this process also has the disadvantage of producing large amounts of solid waste [6]. Currently, the methods of textile wastewater treatment involve physico-chemical processes (coagulation/flocculation, adsorption, precipitation) and/or chemical

∗ Corresponding author at: National Institute of Technology, Ministry of Science, Technology and Innovation, Biocatalysis Laboratory, Av. Venezuela, 82, 302, 20081312, Rio de Janeiro, RJ, Brazil. Tel.: +55 21 2123 1108/1109; fax: +55 21 2123 1166. E-mail addresses: [email protected], cristina [email protected] (M.C. Silva). 1381-1177/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.molcatb.2013.01.004

processes (electrolysis, chemical reduction and advanced chemical oxidation). However, most of these processes are expensive, can generate large volumes of sludge and usually require the addition of environmentally hazardous chemical additives [5,7]. Remazol Brilliant Blue R (RBBR) dye has been used as a model substance in several studies on dye degradation and different kinds of physical, chemical and biological processes have been tested for its removal. Table 1 lists chemical and physical methods used for RBBR removal. Although these methodologies are effective, disadvantages of each process should be considered, such as: sludge generation (adsorption and fenton oxidation), short half life (ozonization) and high cost of electricity (electrochemical). Enzymes from different plants and microorganisms like peroxidases, lacases, mono and dioxegenases have shown a great capacity to degrade a wide range of persistent organic pollutants, which includes textile dyes [16,17]. Enzyme-based dye decolourization methods are very attractive, due to the minimal impact on the ecosystem [18]. However, the use of biocatalysts demands high production cost [5]. Moreover, enzymes are biodegradable and are easily removed from contaminated streams and are also able to efficiently convert complex chemical structures under mild conditions. The limitation for the use of plant peroxidase is the low yield and high production cost compared to microbial enzymes. Plant peroxidases, such as: horseradish [19–22], turnip [23,24], white radish

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Table 1 Chemical and physical methods studied for RBBR dye removal. Methodology Adsorption on agro-industrial waste materials (wheat bran) Adsorption on MgO nanoparticles Fenton oxidation Ozonization Hydrolysis Electrochemical Ultrasound assisted electrochemical process Biosorption on Candida sp.

Concentration range (mg L−1 )

Temperature (◦ C)

pH

50

20

1.5

50–300

20

8

100 800 100 400 50

20 25 120 25 –

203

25

[25] and soybean [26] have shown an excellent potential for dye decolourization. However, the high costs associated with biocatalysts production and application still hinder their large scale use under environmental purposes. The extraction of enzymes from agro industrial residues constitutes one alternative for reducing costs in biocatalysts production [27]. Soybean (Glycine max L.) is an agricultural product of great importance due to its versatile application in food and feed. It is also important to emphasize the economic relevance of soybean in the national and international markets. In addition, Brazil is among the largest producers of soybean in the world [28]. Soybean seed hulls have been identified as a rich source of peroxidases, and, as a soybean-processing industry by-product, they constitute one lowcost alternative [29]. Many treatments can be efficient in the decolourization, but it is essential to know if toxic products are formed during the process. The use of bioindicators is a valuable option to evaluate the toxicity of degradation products [30]. This work evaluated the use of a crude extract of soybean peroxidase from soybean seed hulls in order to catalyze colour removal from aqueous solution containing the reactive dye Remazol Brilliant Blue R (RBBR) and from simulated dyebath wastewater containing the same dye. This dye is widely used in Brazilian textile industries. The following parameters were studied: reaction time, enzyme load and dye and H2 O2 concentrations. The toxicity of the dye RBBR and its degradation products after enzymatic treatment were also evaluated through the use of A. salina and lettuce seeds (L. sativa) as bioindicators. Furthermore, a comparative study was performed among current methodologies and the present study. The most interesting point in this work lies on the promising use of a low cost enzyme obtained from an agro residue widely available in the Brazilian food industry. 2. Materials and methods 2.1. Dye The textile dye Remazol Brilliant Blue R (RBBR), which presents an anthraquinone as chromophore, was kindly provided by DyStar (Porto – Portugal) and used as received without further purification. Dye solution used for degradation experiments was prepared with distilled water. 2.2. Soybean seed hulls extracts The soybean hulls (25 g) were homogenized in a blender with 100 mL of 0.05 mol L−1 pH 6.5 phosphate buffer, containing NaCl 0.2 mol L−1 for 30 s. The homogenate was filtered in organza cloth and centrifuged at 10,000 × g for 15 min, at 4 ◦ C [31]. The enzymatic

3 3–10 4

Time reaction (min)

Decolorization (%)

300 5

8

60 45 120 80 120

2

5760

References

97.5

[8]

98

[9]

>99 100 74.39 100 90 69

[10] [11] [12] [13] [14] [15]

extract obtained was subjected to a precipitation by adding cold acetone until reaching 65% (v/v). After 12–14 h at −18 ◦ C, the sedimentation was concluded and the precipitate was separated by centrifugation at 11,000 × g for 15 min, at 4 ◦ C. The precipitate containing the peroxidases was left at 4 ◦ C for removal of waste acetone for approximately 72 h, and then, re-suspended in 25 mL pH 6.0 of citrate phosphate buffer 0.1 mol L−1 . The obtained suspension was stored at 4 ◦ C and used in the decolourization assays. The supernatant was collected, and the acetone was recovered by simple distillation in rotary evaporator with control temperature at 56◦ C. The recovered acetone can be reutilized in the enzyme obtention process, thus reducing the cost of the process, which yields the enzyme through an economically simple and viable process.

2.3. Enzymatic activity The enzymatic activity was determined according to Khan and Robinson [32] by using the reaction medium of: 1.5 mL of guaiacol 1% (v/v) (Vetec, 97%, v/v), 0.4 mL of H2 O2 0.3% (v/v) (Vetec, PA), 0.1 mL of enzyme (kept in ice bath) and 1.2 mL of 0.05 mol L−1 phosphate buffer pH 6.5. The reaction was carried out for 5 min at 30 ◦ C in a Spectrovision spectrophotometer coupled to a thermostatic bath. One unit of peroxidase activity represents the oxidation of 1 ␮mol of guaiacol per minute in the assay conditions and it was calculated by using data relative to the linear portion of the curve [32].

2.4. Decolorization assay The experiments were carried out at 30 ◦ C [33] by varying the following parameters: reaction time, dye concentration (10–60 mg L−1 ), enzyme activity (4.96–140 U mL−1 ) and H2 O2 concentration (20–1100 ␮mol L−1 ). All reactions were performed at pH 6.0, according to Silva et al. [33] (sodium citrate buffer 0.1 mol L−1 , 1.2 mL) containing dye RBBR (1.5 mL), enzyme (0.1 mL) and H2 O2 (0.4 mL), using multiple or single additions of H2 O2 . Controls were carried out in the absence of H2 O2 . The reaction mixture was analyzed by using a spectrophotometer coupled to a thermostatic bath. The consumption of RBBR was monitored at 596 nm, which corresponds to the maximum absorption wavelength of this dye. The amount of oxidized dye was estimated according to the Eq. (1): oxidized dye (mg L−1 ) =

 dye concentration  initial × removal percentage of dye 100 (1)

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2.5. Influence of chemical auxiliaries utilized in the dyeing process in the colour removal

90

A synthetic effluent containing the dye RBBR was prepared to simulate the real effluent produced by typical textile industries using reactive dyes. The preparation of synthetic effluent was carried out according to Santos [34]: (1) 6.25 g of NaCl were dissolved in 125 mL of warm tap water; (2) 125 ␮L of each of the auxiliaries Periwet WLV (anionic wetting agent) and Periquest T BSD (anionic sequestrant) were added; (3) addition of the RBBR dye (320 mg L−1 ) keeping the solution temperature at 55–60 ◦ C for 10 min to complete dissolution; (4) 0.25 g of NaOH and 1.25 g of Na2 CO3 were added and the solution was maintained at constant temperature for 1 h more; (5) the dye bath was diluted in 1 L of water, then the final concentrations of each component of the dye bath were: dye 0.040 g L−1 , NaCl 6.250 g L−1 , sequestrant agent 0.125 g L−1 , wetting agent 0.125 g L−1 and the salts NaOH 0.250 g L−1 and Na2 CO3 1.250 g L−1 . This simulated effluent was also used in biodegradation experiments.

80

2.6. Acute toxicity test with A. salina The tests with A. salina were carried out according to the methodology developed by Meyer and co-workers [35] and modified by Neto [36]. Larvae of brine shrimp (A. salina) were obtained after eclosion from dry eggs in synthetic sea water with aeration during 48 h. Subsequently, the larvae were transferred to each set of tubes (10 larvae/tube) with different concentrations of reaction media containing the dye (control) or their degradation products after enzymatic treatment. It was used the reaction media in different concentrations varying from 0 to 100% (30, 40, 50 and 100%), as showed in Fig. 5. After 24 h of incubation, the number of surviving organisms was counted and the lethal concentration, 50% (LC50 ) was calculated. The mortality percentage of A. salina was related to the concentrations of reaction media before and after enzymatic treatment. The following concentrations of reaction media were used: 100, 70, 50, 30 and 10% (v/v). Negative control (synthetic sea water) was performed in parallel. Tests were carried out in triplicate. 2.7. Acute toxicity test with lettuce seeds (L. sativa) The tests with lettuce seeds (L. sativa) were carried out according to Dutka [37]. Twenty lettuce seeds were placed in a polystyrene plate, containing a filter paper embedded in 2 mL of each reaction media. The plates were wrapped with tin foil and incubated in the dark. Radicle lengths were measured after 72 h of exposition and the inhibitory concentration 50 (IC50 ) was calculated. The inhibition percentage of growth of radicals was related to the concentrations of reaction media before and after enzymatic treatment. The following concentrations of reaction media were used: 75, 50, 25 and 5% (v/v). Negative control (distilled water) was performed in parallel. Tests were carried out in triplicate. 3. Results 3.1. Reaction time An important factor on enzymatic decolourization is the catalyst/substrate contact time which gives a maximum removal. According to Fig. 1, the decolourization of RBBR reached 60% in only 2 min of reaction and the highest percentage of decolourization was achieved after 13 min (86%). To evaluate the dye adsorption by the enzymatic extract, controls were carried out in the absence of H2 O2 . It was observed

Colour removal (%)

85

75 70 65 60 55 50 45 40 0

2

4

6

8

10

12

14

16

Time (min) Fig. 1. Reaction progress on the decolorization of the RBBR dye. Reaction conditions: enzyme load = 70.4 U mL−1 ; dye concentration = 50 mg L−1 and H2 O2 concentration = 100 ␮mol L−1 .

that there was no adsorption of the dye by the enzymatic extract because in the absence of hydrogen peroxide, there was no dye reduction 3.2. Dye concentration Table 2 shows the percentage of colour removal and the amount of oxidized RBBR in different dye concentrations (10–60 mg L−1 ). In the range from 10–40 mg L−1 , the decoulorization percentage varied from 75–85.7%. The soybean peroxidase ability to remove colour from reaction media was only affected in the concentration of 60 mg L−1 , which resulted in decolourization of 62.5%. The maximum colour removal (85.7%) was obtained at concentration of 40 mg L−1 , where 34.28 mg L−1 of RBBR were converted into its degradation products. However, it is important to note that in the lower percentage of colour removal (62.5%) in the concentration of 60 mg L−1 , the amount of dye oxidized (37.49 mg L−1 ) is very similar when compared to our best result, 85.7% of colour removal at 40 mg L−1 . 3.3. Enzyme activity The evaluation of enzyme concentration on RBBR decolorization is shown in Fig. 2. The increase in the enzyme load is accompanied by an increase in the decolourization percentage up to 70 U mL−1 , where the maximum removal is obtained (86%). Enzyme concentrations above 70 U mL−1 shows a reduction in the decolourization efficiency. Table 2 Effect of dye concentration on the decolorization process. Reaction conditions: enzyme load = 70.4 U mL−1 ; reaction time = 13 min and H2 O2 concentration = 100 ␮mol L−1 . Dye concentration (mg L−1 )

Colour removala (%)

10 20 30 40 50 60

75.00 80.33 83.94 85.70 84.15 62.48

a

Mean ± standard deviation.

± ± ± ± ± ±

0.2 0.31 0.36 0.44 2.97 1.01

Concentration of oxidized dye (mg L−1 ) 7.50 16.07 25.18 34.28 42.07 37.49

M.C. Silva et al. / Journal of Molecular Catalysis B: Enzymatic 89 (2013) 122–129

100

60 50

Colour removal (%)

80

Colour removal (%)

125

60

40

20

40 30 20 10 0

0 0

20

40

60

80

100

120

140

160

0

-1

enzyme load (U mL )

1

2

3

4

5

6

7

8

Time (min)

Fig. 2. Effect of enzyme concentration on the decolorization of RBBR dye. Reaction conditions: dye concentration = 40 mg L−1 ; reaction time = 13 min and H2 O2 concentration = 100 ␮mol L−1 .

Fig. 4. Reaction progress on the decolourization of synthetic effluent containing RBBR dye (40 mg L−1 ) by soybean peroxidase (31.87 U mL−1 ).

3.4. H2 O2 concentration

3.5. Influence of chemical auxiliaries utilized in the dyeing process in the colour removal

In this study, the concentration range of H2 O2 used showed no significant influence on decolourization efficiency. It was observed that in the absence of this coadjutant there was no decoulorization and in the low concentrations (20–60 ␮mol L−1 ) the percentage of decoulorization obtained was 9.34 and 56.4%, respectively (Fig. 3). In this case, a decrease of enzyme performance was observed at low H2 O2 concentration. The maximum colour removal (86%) was observed in the H2 O2 concentrations of 100 and 200 ␮mol L−1 . In the range from 100 to 500 ␮mol L−1 , the decoulorization percentage remained approximately constant. While, in higher H2 O2 concentrations (900–1100 ␮mol L−1 ), an inhibitory effect was observed. The H2 O2 stepwise addition to the reaction medium can avoid the inactivation of the enzyme by H2 O2 excess [38]. Thus, after 7 min of reaction (approximately half of the reaction total time), a new concentration of H2 O2 was added, which resulted in a final concentration of 100 ␮mol L−1 . In this case, the colour removal obtained was 86%, similar to the use of fixed concentration of H2 O2 .

The influence of salts and dyeing auxiliary chemicals on the decolourization potential of dye RBBR by soybean peroxidase was evaluated utilizing the synthetic textile effluent. The decolourization obtained was 53.4% after 6 min of reaction (Fig. 4). Approximately 5 min were sufficient to obtain a maximum decolourization for synthetic effluent; while, the reaction time required for achieving maximum decolourization of RBBR dye was 13 min (Fig. 1). 3.6. Toxicity tests Acute toxicity tests with A. salina showed an increase of toxicity of the reaction media after the enzymatic treatment with soybean peroxidase enzyme (Fig. 5). It was used the reaction media in different concentrations varying from 0 to 100% (30, 40, 50 and 100%), as showed in Fig. 5. The increased toxicity is evident from the LC50 values obtained before and after enzymatic treatment, 81 and 59%, respectively.

Mortality before enzymatic treatment Mortality after enzymatic treatment

100

100

90

80

70 60

Mortality (%)

Colour removal (%)

80

50 40 30

60 40 20

20 10

0

0 0

200

400

600

800

1000

1200

-1

H2O2 concentration (µmol L ) Fig. 3. Influence of H2 O2 concentration on the decolorization of RBBR dye. Reaction conditions: dye concentration = 40 mg L−1 ; reaction time = 13 min and load enzyme = 70 U mL−1 .

0

20

40

60

80

100

Reaction media (%) Fig. 5. Percentage of A. salina mortality according to the concentration of reaction media containing RBBR dye or RBBR degradation products after enzymatic treatment soybean peroxidase.

126

M.C. Silva et al. / Journal of Molecular Catalysis B: Enzymatic 89 (2013) 122–129

Fig. 6. Proposal mechanism for biocatalytic transformation of RBBR dye by A. flavus (Perlatti et al. [47]).

Table 3 Acute toxicity test with lettuce seeds (L. sativa), after 72 h of incubation in different reaction media concentrations. Reaction media (%)

Inhibition before enzymatic treatmenta (%)

5 25 50 75

0 0 0 0

a

± ± ± ±

0 0 0 0

Inhibition after enzymatic treatmenta (%) 35.6 47.6 68.5 72.5

± 2.3 ± 0.2 ± 3.0 ± 0.5

Mean ± standard deviation.

Based on the results for lettuce seeds (L. sativa) shown in Table 3, it can be observed that there is no inhibition of growth of radicals before enzymatic treatment. A significant increase in inhibition was observed with increasing concentration of the reaction media after enzymatic treatment, resulting in an IC50 of 28%. 4. Discussion Table 4 presents different studies on RBBR decolourization by using different enzymes. The decolourization of RBBR mediated by laccases was previously studied by Teixeira et al. [39]. Authors compared the action of a commercial laccase from Aspergillus oryzae and a rich extract from Pleurotus ostreatus cultivation residues in decolourization of dye RBBR. The decolourization of RBBR obtained was 80–90% with both laccases after 24 h in the presence of the mediator ABTS (2,2 -azino-bis(3-ethyl benzothiazoline-6-sulfonic acid)). As mentioned before, soybean peroxidase obtained from wasted seed hulls was able to remove the dye RBBR in only 13 min, which is a very significant result. Recent studies (Table 4) reported that 42, 10 and 4 h were the time required to catalyze the enzymatic decolourization of the dye RBBR by immobilized laccase from Trametes pubescens [40], enzymatic complex from Funalia trogii [41] and bilirubin oxidase [42], respectively. According to previous studies reported in Table 4, the enzymatic decolourization of the dye RBBR is promoted by different

microorganisms and plants. This work presents one of the most promising results regarding reaction time (13 min). Only the commercial enzyme Horseradish peroxidase, showed a faster reaction, 5 min [22]. Knuston et al. [26] also evaluated the potential of soybean peroxidase to decolorize the azo dye Direct Yellow 11 by treating with commercial enzyme (purchased from Sigma) for 2 h at 45 ◦ C. The colour removal was 48% after 90 min of reaction. Comparatively, the crude extract employed in the present study showed a promising performance. The knowledge of the metabolites formed during the decolorization of textile dyes by plant peroxidases is a way for the understanding of the break up mechanism of complex structures chemically stable, by the enzymes [46]. Perlatti et al. [47] identified the RBBR degradation products by Aspergillus flavus. It was possible to identify four metabolic degradation products, with molecular mass of 318, 336, 165 and 188 Da, respectively. Their identification was carried out by using their mass-to-charge ratio and temporal availability. The authors (Perlatti et al. [47]) identified the product of 336 Da observed at the chromatographic peak at 8.5 min as a product of the oxidation of the anthraquinone moiety, 2-amino-4-(2-carboxybenzoyl)-5hydroxybenzenesulfonate. According to Perlati et al. [47], the oxidation of the second carbonyl moiety in the anthraquinone ring would generate two products: o-phthalic acid and 2-amino1-phenol-4-sulfonic acid, which were attributed to degradation products of 165 Da and 188 Da, respectively (Fig. 6). The intracellular systems that are generally present in most fungi, such as cytochrome P-450 monooxygenase, may also be involved in organopolluant degradation [48]. It means without ligninolytic enzymes, RBBR decolourization can be attributed to the other enzymes such as P-450 monooxygenase or dioxygenase. The existence of several other kinds of monooxygenases in Aspergillus strains is also known, some of which belonging to the aflatoxin biosynthesis [49,50]. Thereby the degradation pathway of RBBR suggested by Perlatti et al. [47] probably involves P-450 monooxygenases, and the

Table 4 Different enzymatic treatments studied for RBBR decolourization. Enzyme

Source

Commercial laccase (Aspergillus oryzae) and crude enzymatic extract from shimeji residues

Aspergillus oryzae and Pleurotus ostreatus

Immobilized laccase Bilirubin oxidase Enzymatic complex Purified laccases (POXC and POXA3) Horseradish peroxidase (commercial enzyme) Free laccase Immobilized laccase Laccase (purified)

Trametes pubescens Myrothecium verrucaria Funalia trogii Pleurotos ostreatus

Soybean peroxidase (crude extract)

Armoracia rusticana Trametes versicolor Trametes trogii Soybean seed hulls

RBBR concentration (mg L−1 ) 60–120

133 100 100 31.3 31.3 60 22.5 100 40

Reaction time

Temperature (◦ C)

Decolorization (%)

References

24 h

Room

80–90

[39]

42 h 4h 10 h 1.7 h 1.7 h 5 min

Room 40 30 30 20 35

44 40 90 86 94 96

[40] [42] [41] [43]

0.5 h

23

[44]

2.5 h

50

90 77 97

[45]

13 min

30

86

Present work

[22]

Lettuce seeds (Lactuca sativa)

Bacillus cereus

a EC20 = sample concentration that causes an acute effect to 20% of organisms in the exposure time and conditions of the test (EC20 < 25% = very toxic; 25% < EC20 < 50% = moderately toxic; 50% < EC20 < 75% = toxic; 75% < EC20 < 100% = slightly non-toxic; EC20 > 100% = non-toxic); GI = germination index (GI < 50% = high toxicity; 50% < GI < 80% = moderate toxicity; GI > 80% = no toxicity); TR = percentage of toxicity reduction; LC50 = sample concentration that causes mortality to 50% of organisms in the exposure time and conditions of the test; IC50 = sample concentration that causes 50% inhibition of growth of radicals in the exposure time and conditions of the test.

–

IC50 = 28%

Present work LC50 = 59%

Percentage of mortality of microorganisms Growth of radicals inhibition Artemia salina Soybean peroxidase (soybean seed hulls)

86

LC50 = 81%

[57] EC20 > 100% Bacterial luminescence inhibition Vibrio fischeri 50 30

EC20 > 50–75%

[43] TR = 95% 90 31

Mixture of laccase POXC and laccase POXA3 (Pleurotos ostreatus) Trametes versicolor ATCC 20869

–

[40] 44

Ryegrass seeds (Lolium perenne)

Seed germination percentage and root length of seeds Bacterial growth inhibition 133

Artemia salina 97 120

Vibrio fischeri 77 112

Immobilized laccase (Trametes pubescens)

Approximately 45% of mortality of A. salina (90% of the reaction media) GI = 69% Approximately 75% of mortality of A. salina (90% of the reaction media) GI = 26%

[22]

EC20 = 11.7%

[44]

127

EC20 = 33.5%

[44] EC20 = 13.4% EC20 = 33.5%

Bacterial luminescence inhibition Bacterial luminescence inhibition Percentage of mortality of microorganisms 90 112

Free laccase (Trametes versicolor) Immobilized laccase (Trametes versicolor) Horseradish peroxidase (HRP)

Vibrio fischeri

Toxicological response (after treatment)a Toxicological response (before treatment)a Effects of bioindicators evaluated Bioindicators used in the toxicological tests RBBR decolorization (%) RBBR concentration (mg L−1 ) Treatment

Table 5 Different studies on RBBR dye decolorization mediated by enzymes and toxicological evaluation of degradation products.

degradation products or intermediates generated by the action of oxidase/peroxidase can be comparable. The capability of laccase from Polyporus sp. S133 to decolorize RBBR and the metabolic products resulting from the enzymatic treatment were also investigated by Hadibarata and coworkers [51]. In this study, it was proposed that laccase of Polyporus sp.S133 break the structure of RBBR became two subproducts (sodium 1-amino-9,10-dioxo-9,10-dihydroanthracene2-sulfonate and sodium 2-((3-aminophenyl)sulfonyl)ethyl sulfate). Others authors have been focusing their attention to identify the enzymatic degradation products of anthraquinone dyes [40,52]. These studies could contribute to a better comprehension of the mechanisms used by oxidative enzymes, as soybean peroxidase, to transform organic compounds. Substrate concentration is a key factor, which affects the oxidation velocity mediated by enzymes. Liu and co-workers [42] evaluated the dye-decolorizing potential of bilirubin oxidase (BOX) for dye RBBR. In this study, the decolourization efficiency decreased with increasing RBBR concentration, and a marked inhibition effect was exhibited when the dye concentrations were above 100 mg L−1 . The same result was observed in the present work when the dye concentrations were above 50 mg L−1 . The study of enzyme load used in dye decolourization is a critical point, since this parameter is directly involved in the overall cost of the bioprocess. The results obtained indicate that the use of a higher enzyme concentration (above 70 U mL−1 ) promoted a slight decrease on the decolourization of the RBBR dye, which may be attributed to a simultaneous decrease of all reactants involved (dye and co-substrate). Similar results were observed by Souza et al. [21] that studied the decolourization of the dye Remazol Turquoise Blue G 133% by H. peroxidase. It was found that in a concentration of 14.985 U mL−1 , the decolourization obtained was 58% and, when the concentration was doubled, the decolorization was 62%. Similar results were also reported in other studies that emphasize the effect of enzyme dose in the dye-decolorizing potential [19,53]. As hydrogen peroxide (H2 O2 ) acts as co-substrate in peroxidases reactions, its presence in low concentrations limits the reaction rate and its excess inhibits the enzyme activity [19,54]. However, the concentration range of H2 O2 used showed no significant influence on decolourization efficiency of RBBR dye. H2 O2 stepwise addition showed no significant influence on dye removal, which was probably due to the thermodynamic characteristics of the reaction. Similar results were also reported in previous studies that emphasize the decrease of decolourization potential of several oxidases (H. peroxidase, lignin peroxidase, manganese peroxidase and laccase) at high H2 O2 concentrations [19,20,38,53]. In the optimization of H2 O2 quantity, evaluations were carried out in the absence of H2 O2 and it was observed that in the absence of this coadjutant there was no decolorization. This result indicates that the decolorization occurs exclusively as a function of the catalytic activity of the enzyme. The efficiency of the soybean peroxidase obtained in this work on the decolourization of RBBR was compared to other previous studies reported in the literature, which are summarized in Table 4. According to the data presented, soybean peroxidase obtained in this study presents decolourization ability (86%) comparable to the commercial enzyme H. peroxidase (96%). The data obtained encourage further studies on crude extraction obtaining and application. The decrease of enzyme performance in the presence of salts and dyeing auxiliary chemicals, as observed in this study, can occur due to the following reasons: the formation of strong chemical bonds with the species involved in the degradation process, the creation of products during the process or even because of enzyme deactivation [55].

References

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When comparing the reaction times required achieving maximum decolourization for synthetic effluent and dyeing. This was probably due to changes in the dye structure, as secondary reactions may happen during dyeing process affecting directly the enzyme-substrate specificity. Probably happens the formation of strong chemical bonds with the species involved in the reaction medium (dye, enzyme, salts and chemicals auxiliary) as, for example, hydrolysis of dye [55]. In order to have the fixing between the reactive dye and cellulose, is added to the dyebath an alkali, usually sodium hydroxide, to promote the ionization of cellulose. However, hydrolysis of the dye can occur as a secondary reaction [56]. Cristóvão [55] studied the decolorization of dye reactive red 239 by commercial laccase from Aspergillus, and they observed a decrease of 68.23% to 42.04% on the decolourization of dye in the presence of dyeing auxiliary chemicals. Although soybean peroxidase showed effectiveness in removing colour from RBBR dye media, its performance decreased in the presence of dyeing auxiliary chemicals. However, the results presented in Fig. 4 encourage further studies. The impact of salts and auxiliary chemicals on decolourization efficiency can also be overcome by the use of enzymes in immobilized form, which can be used as catalysts with long lifetime [49–53]. Toxicity study was conducted before and after enzymatic treatment in order to evaluate the toxicity of the dye and its degradation products. Toxicity tests were performed using two types of bioindicators: A. salina and lettuce seeds (L. sativa). A. salina was used as bioindicator since the effluent from the textile industries has high salinity and, therefore, high conductivity, which makes this a critical parameter for a freshwater species [22,54]. Tests with lettuce seeds (L. sativa) are simple and low-cost. Although RBBR dye is widely used as a model in studies of textile effluents degradation, there is very little information regarding the toxicity of this dye and its degradation products. The toxicity bioassays are extremely important to evaluate not only the effectiveness of a treatment but also the degree of its environmental safety. Table 5 presents different studies on toxicological evaluation in studies of RBBR dye decolorization. Champagne and Ramsay [44] observed the increase of toxicity upon Vibrio fishseri after enzymatic decolourization of RBBR by free laccase from Trametes versicolor. The same was observed by authors utilizing immobilized laccase as biocatalyst [44]. All other enzymatic studies reported in Table 5 resulted in a reduction of toxicity after RBBR treatment. In this work, the obtention of metabolites more toxic than the parental molecule reinforces the concept of combined treatments effectiveness and emphasize the importance of toxicological evaluation after enzymatic treatment.

5. Conclusions In this study, the soybean peroxidase showed a promising performance as biocatalyst in the decolourization of a reactive anthraquinone dye. Nevertheless, the efficiency of this catalyst is associated to enzyme activity, substrate and H2 O2 concentrations used. The presence of chemical auxiliaries, which are commonly utilized in the dyeing baths in industrial processes, was detrimental to enzymatic decolourization. The acute toxicity tests for RBBR products with A. salina and lettuce seeds (L. sativa) showed an increase of toxicity after treatment with soybean peroxidase, which indicated the formation of toxic substances. The results obtained in this work encourage additional studies of soybean peroxidase in decolourization process. However, the enzymatic treatment should be associated with other types of treatment for further degradation and toxicity reduction.

Finally, it is important to highlight that soybean peroxidase was extracted from soybean seed hulls, an abundant residue from soybean-processing industries, which represents a cost reduction when compared to commercial enzymes normally used, such as H. peroxidase. Acknowledgements This study was supported by Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq, Brazil), Coordenadoria de Aperfeic¸oamento de Pessoal de Nível Superior (CAPES, Brazil), and National Institute of Technology – Ministry of Science, Technology and Innovation (INT-MCTI, Brazil) through financial funding and scholarships. References [1] P. Nigam, G. Armour, I.M. Banat, D. Singh, R. Marchant, Bioresour. Technol. 72 (2000) 219–226. [2] M.V. Zanoni, P.A. Carneiro, Ciên Hoje. 29 (2001) 61–64. [3] J.S. Knapp, P.S. Newby, L.P. Reece, Enzyme Microb. Technol. 17 (1995) 664–668. [4] Q. Husain, Crit. Rev. Biotechnol. 26 (2006) 201–221. [5] T. Robinson, G. McMullan, R. Marchant, P. Nigam, Bioresour. Technol. 77 (2001) 247–255. [6] N. Durán, E. Esposito (Eds.), Lignin Biodegradation and Effluent Treatment by Ligninolytic Fungi in Microbiologia Ambiental CNPMA/EMBRAPA, São Paulo, Brazil, 1997. [7] P.R. Gogate, A.B. Pandit, Adv. Environ. Res. 8 (2004) 501–551. [8] C. Fatma, O. Dursun, O. Ahmet, O. Ayla, J. Hazard. Mater. 146 (2007) 408–416. [9] G. Moussavi, M. Mahmoudi, J. Hazard. Mater. 168 (2009) 806–812. [10] S. Chang, S. Chuang, H. Li, H. Liang, C. Huang, J. Hazard. Mater. 166 (2009) 1279–1288. [11] A.R. Tehrani-Bagha, N.M. Mahmoodi, F.M. Menger, Desalination 260 (2010) 34–38. [12] M. Siddique, R. Farooq, Z.M. Khan, S.F. Shauka, Ultrason. Sonochem. 18 (2011) 190–196. [13] D. Raijkumar, B.J. Song, J.G. Kim, Dyes Pigm. 72 (2007) 1–7. [14] M. Siddique, R. Farooq, A. Khalida, A. Farooq, Q. Mahmooda, U. Farooq, I.A. Rajaa, S.F. Shaukatc, J. Hazard. Mater. 172 (2009) 1007–1012. [15] Z. Aksu, G. Dönmez, Chemosphere 50 (2003) 1075–1083. [16] D. Wesenberg, I. Kyriakides, S.N. Agathos, Biotechnol. Adv. 22 (2003) 161–187. [17] Q. Hussain, Rev. Environ. Sci. Biotechnol. 9 (2009) 117–140. [18] A. Michniewicz, S. Ledakowicz, R. Ullrich, M. Hofrichter, Dyes Pigm. 77 (2008) 295–302. [19] S.V. Mohan, K.K. Prasad, N.C. Rao, P.N. Sarma, Chemosphere 58 (2005) 1097–1105. [20] V.S. Ferreira-Leitão, J.G. Silva, E.P.S. Bon, Appl. Catal. B 42 (2003) 213–221. [21] S.M.A.G.U. Souza, E. Forgiarini, A.A.U. Souza, J. Hazard. Mater. 147 (2007) 1073–1078. [22] M.R. Da Silva, L.R.V. De Sá, C. Russo, E. Scio, V.S. Ferreira-Leitão, Enzyme Res. 2010 (2010) 1–7. [23] M. Matto, Q. Husain, Chemosphere 69 (2007) 338–345. [24] M.C. Silva, J.A. Torres, A.D. Corrêa, A.M.B. Junqueira, M.T.P. Amorim, C.D. dos Santos, Water Sci. Technol. 65 (2012) 669–675. [25] R. Satar, Q. Husain, Biochem. Eng. 46 (2009) 96–104. [26] K. Knutson, S. Kirzan, A. Ragauskas, Biotechnol. Lett. 27 (2005) 753–758. [27] J. Dec, J.M. Bollag, Biotechnol. Bioeng. 44 (1994) 1132–1139. [28] O.L. Mello Filho, C.S. Sediyama, M.A. Moreira, M.S. Reis, G.A. Massoni, N.D. Piovesan, Pesqui. Agropecu. Bras. 39 (2004) 445–450. [29] J.W. Gillikin, J.S. Graham, Plant Physiol. 96 (1991) 214–220. [30] T. Kusui, C. Blaise, in: S. Salem, Rao (Eds.), Impact Assessment of Hazardous Aquatic Contaminants, Ann Arbor Press, Michigan, 1999. [31] O. Fatibello Filho, I.C. Vieira, Quim. Nova 25 (2002) 455–464. [32] A.A. Khan, D.S. Robinson, Food Chem. 49 (1994) 407–410. [33] M.C. Silva, A.D. Corrêa, M.T.P. Amorim, P. Parpot, J.A. Torres, P.M.B. Chagas, J. Mol. Catal. B: Enzym. 77 (2012) 9–14. [34] S.C.R. Santos, Dissertation, Univ. Porto, 2009. [35] B.N. Meyer, N.R. Ferrigni, J.E. Putnam, L.B. Jacobsen, D.E. Nichols, J.L. MacLaughlin, J. Med. Plant Res. 45 (1982) 31–34. [36] A.I.S. Neto, Dissertation, Federal University of Juiz de Fora, 2003. [37] B. Dutka, Methods. For Toxicological Analysis of Waters, Wastewaters and Sediments, National water research institute (NWRI), Environment Canada, Burlington, Ottawa, 1989. [38] V.S. Ferreira-Leitão, M.E.A. Carvalho, E.P.S. Bon, Dyes Pigm. 74 (2007) 230–236. [39] R.S.S. Teixeira, P.M. Pereira, V.S. Ferreira-Leitão, Enzyme Res. (2010) 1–8. [40] J.F. Osma, J.L. Toca-Herrera, S. Rodríguez-Couto, Bioresour. Technol. 101 (2010) 8509–8514. [41] H.D. Özsoy, A. Ünyayar, M.A. Mazmanci, Biodegradation 16 (2005) 195–204. [42] Y. Liu, J. Huang, X. Zhang, J. Biosci. Bioeng. 108 (2009) 496–500. [43] G. Palmieri, G. Cennamo, G. Sannia, Enzyme Microb. Technol. 36 (2005) 17–24. [44] P.P. Champagne, J.A. Ramsay, Bioresour. Technol. 101 (2010) 2230–2235.

M.C. Silva et al. / Journal of Molecular Catalysis B: Enzymatic 89 (2013) 122–129 [45] T. Mechichi, N. Mhiri, S. Sayadi, Chemosphere 64 (2006) 998–1005. [46] A. Heinfling, M.J. Martínez, A.T. Martínez, M. Bergbauer, U. Szewzyk, Appl. Environ. Microbiol. 64 (1998) 2788–2793. [47] B. Perlatti, M.F.G.F. Silva, J.B. Fernandes, M.R. Forim, Bioresour. Technol. 124 (2012) 37–44. [48] C.-J. Cha, D.R. Doerge, C.E. Cerniglia, Appl. Environ. Microbiol. 67 (2001) 4358–4360. [49] K. Yabe, Y. Nakamura, H. Nakajima, Y. Ando, T. Hamasaki, Appl. Environ. Microbiol. 57 (1991) 1340–1345. [50] J. Yu, D. Bhatnagar, K.C. Eerlich, Rev. Iberoam. Micol. 19 (2002) 191–200. [51] T. Hadibarata, A.R.M. Yussoff, R.A. Kristanti, Water Air Soil Pollut. 223 (2012) 933–941.

129

[52] S. Andleeb, N. Atiq, G.D. Robson, S. Ahmed, Environ. Sci. Pollut. Res., http://dx.doi.org/10.1007/2011s11356-011-0687-x [53] F. Hoshino, T. Kajino, H. Sugivama, O. Asami, H. Takahashi, FEBS Lett. 530 (2002) 249–252. [54] J. Wu, J.K. Bewtra, N. Biswas, K. Taylor, Can. J. Chem. Eng. 72 (1994) 881–886. [55] R.O. Cristóvão, A.P.M. Tavares, J.M. Loureiro, R.A.R. Boaventura, E.A. Macedo, Bioresour. Technol. 100 (2009) 6236–6242. [56] L T.C. Beltrame, Thesis, Federal University of Rio Grande do Sul, Brazil, 2006. [57] J.A. Ramsay, T. Nguyen, Biotechnol. Lett. 24 (2002) 1757–1761.