Open Access

Studies on decolorization of reactive blue 19 textile dye by Coprinus plicatilis

Journal of Environmental Health Science and Engineering201412:49

DOI: 10.1186/2052-336X-12-49

Received: 23 July 2013

Accepted: 10 February 2014

Published: 24 February 2014

Abstract

Background

Studies were carried on the decolorization of the textile dye reactive blue 19 (RB 19) by a novel isolate of Coprinus plicatilis (C. plicatilis) fungi. We describe an in vitro optimization process for decolorization and its behavior under different conditions of carbon and nitrogen sources, pH, temperature and substrate concentration.

Results

The optimal conditions for decolorization were obtained in media containing intermediate concentrations of ammonium oxalate and glucose (10 g/L) as nitrogen and carbon sources, respectively, at 26°C and pH = 5.5. Maximum decolorization efficiency against RB 19 achieved in this study was around 99%. Ultra-violet and visible (UV-vis) spectrophotometric analyses, before and after decolorization, suggest that decolorization was due to biodegradation.

Conclusions

This effect was associated with laccase enzyme displaying good tolerance to a wide range of pH values, salt concentrations and temperatures, suggesting a potential role for this organism in the remediation of real dye containing effluents. In conclusion, laccase activity in C. plicatilis was firstly described in this study.

Keywords

Reactive blue 19 White rot fungi Decolorization Metabolite UV-vis

Introduction

Treatment of synthetic dyes in wastewater is a matter of great concern. Several physical and chemical methods have been employed for the removal of dyes [1]. However, these procedures have not been widely used due to high cost, formation of hazardous by products and intensive energy requirement [2]. Worldwide over 10,000 different dyes and pigments are used in dyeing and printing industries. The total world colorant production is estimated to be 8.00.000 tons per year and at least 10% of the used dyestuff enters the environment through wastes [3, 4]. Wastewater from textile industries constitutes a threat to the environment in many parts of the world. Although some of the dyes are not themselves toxic, after release into the aquatic environment their degradation products are often carcinogenic [5, 6]. The existing technologies for decolorization of textile dyeing effluents like adsorption, precipitation, membrane filtration, chemical degradation and photochemical degradation are relatively expensive and commercially unattractive [7, 8]. Coagulation-flocculation has major operational problems as it generates large amounts of sludge. Adsorption technique is also more expensive as it involves the use of powdered activated carbon as adsorbent and disposal of spent adsorbent [9] is still a problem. Microbial decolorization is a potential and an effective alternative for the decolorization of wastewater. Microbial decolorization methods of textile dye containing effluents have been reviewed and reported. White-rot fungi such as Phanerochaete chrysosporium[10, 11], Trametes versicolor[12, 13], Bjerkandera adusta[14], Pycnoporus cinnabarinus[15] and Phanerochaete sordida[16] have been shown to decolorize textile dyes or coloured effluents [17].

In recent years, the utilization of biodegradative abilities of some white rot fungi seems to be promising. They do not require preconditioning to particular pollutants and owing to their extracellular non-specific free radical-based enzymatic system they can degrade to nondetectable levels or even completely eliminate a variety of xenobiotics including synthetic dyes. Many white rot fungi (Phanerochaete chrysosporium, Pleurotus ostreatus, Bjerkandera adusta, Trametes versicolor, etc.) have been intensively studied in connection with their ligninolytic enzyme production and their decolorization ability [1825]. This capability is due to extracellular non-specific and non-stereoselective enzyme systems composed of laccases (EC 1.10.3.2), lignin peroxidases (EC 1.11.10.14) and manganese peroxidases (EC 1.11.1.13) [26]. Laccase based decolorization treatments are potentially advantageous to bioremediation Technologies since the enzyme is produced in larger amounts mainly by numerous fungi [27, 28]. Laccase belongs to a family of multi-copper oxidases that are widespread in nature. Laccases are related to oxidation of a range of aromatic, toxic and environmentally problematic substrates [29] and particular interest with industrial applications. The potential applications are in the textile industry [30], detoxification of pollutants and industrial effluents [31], pulp and paper industry [32], food and pharmaceutical industries [33] biosensor and biofuel applications [34].

The aim of the present work was to characterize the biodegradation of the textile dye Remazol reactive blue 19, by the action of soluble extracts from the white rot fungus C. plicatilis. We describe an in vitro optimization process for decolorization and its behavior under different conditions of carbon and nitrogen sources, pH, temperature and substrate concentration.

Materials and methods

Dyes and chemicals

Textile Remazol dye: reactive blue 19 (RB 19) was supplied by Dystar (Kocaeli, Turkey). 2,2-Azino-bis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) was obtained from Sigma Chemical Company (St. Louis, MO, USA). All chemicals used were of the highest purity available and of analytical grade.

Culture conditions

Mycelial suspension of Coprinus plicatilis isolated in our university fungus research Laboratory. The white rot fungi Coprinus plicatilis (C. plicatilis) were maintained on 2% (w/v) malt agar slants at 4°C and were then activated at 26°C for 3 days. The mycelium were harvested with a sterile 0.9% NaCl solution and were then inoculated into 100 mL of 2% malt extract broth (pH = 4.5) in 250 mL Erlenmeyer flasks at 26°C and 175 rpm for 4 days. Pellets were inoculated into the medium consisting of 10 g/L glucose, 1.0 g/L of NH4H2PO4, 0.05 g/L of MgSO4.7H2O, 0.01 g/L of CaCl2, 0.025 g/L of yeast extract. Cultivation was carried out in an orbital shaker incubator, at 26°C, 175 rpm [35]. At the beginning of the fourth day of incubation, dye solution was added to the flasks, aseptically, at desired concentrations. Aliquots were assayed for laccase activity. Experiments were performed in 250 mL Erlenmeyer flasks containing 50 mL of liquid medium. For biomass calculation, mycelia were filtered through previously dried and tared Whatman No. 1 filter papers, washed with distilled water and dried at 50°C to constant weight.

Spectrophotometric analysis

Aliquots of 1–2 mL volume of clear dye solution were taken from each reaction flask at regular time intervals and measured immediately using a UV-vis recording double beam spectrophotometer (Shimadzu UV-1601). Decolorization was determined spectrophotometrically by monitoring the absorbance at the wavelength maximum for this dye, and by the reduction of the major peak area in the visible region for dye. The percentage of in culture decolorization was calculated as;

Decolorization percentage, R (%)
R = C o - C C o . 100
(1)

Biodegradation Rate = (C0-C)/Incubation day

C0: Initial concentration of dye, C: Last concentration of dye

Optimization of carbon and nitrogen content for enzyme production

Six different carbon substrates (glucose, sucrose, starch, maltose, fructose and glycerol) and nitrogen sources (sodium nitrate, urea, ammonium tartrate, ammonium carbonate, ammonium oxalate and peptone) were used to substitute the original carbon and nitrogen sources. The assessed concentrations were 5, 10 and 15 g/L. All inoculated media were incubated for 15 days at 26°C in the dark and the supernatant or eluted extracts collected for further analysis.

Effect of pH, temperature and copper supplementation on enzyme production

Cultures optimized for carbon and nitrogen content were supplemented with copper sulfate (0.1-5 mM) and incubated for 15 days at 26°C. Supernatants or soluble extracts were then submitted to decolorization. Additionally, destaining activity was measured in samples derived from cultures grown at 26, 30 and 35°C for 15 days and at pH values from 5.5 to 7.0 at 26°C for 15 days.

Enzyme assays

Laccase (Lac) (EC 1.10.3.2) production was assessed by a measurement of the enzymatic oxidation of 2, 22-azinobis-(3 ethylbenzothiazoline- 6-sulphonic acid) (ABTS) at 420 nm (ϵ = 3.6 × 104 cm-1 M-1) [36]. The reaction mixture contained 300 μL of extracellular fluid, 300 μL of 1 μM ABTS and 0.1 M Na Acetate buffer (pH = 4.5, 400 μL). One unit of enzyme activity is defined as the amount of enzyme that oxidizes 1 μmol ABTS in one minute.

Effect of pH, temperature and salt concentration on decolorization

Soluble extracts derived from 15-day-old solid or liquid cultures, carried out under optimized conditions, were tested for decolorization. Decolorization of RB 19 (50 mg/L) was carried out at pH values from 2 to 9 (intervals of 0.5 units), adjusted by using 50 mM citrate-phosphate buffer. The temperatures assessed were 20, 30, 40, 50, 60 and 70°C. For testing susceptibility to salt, different NaCl concentrations (0.05, 0.1, 0.2, 0.4 and 0.6 M) were used in the assay.

Statistical analysis

Standard deviations of the results of triplicate samples from the flask studies were calculated using the Microsoft Excel Spreadsheet Program.

Results

Screening using reactive dyes

C. plicatilis cultures were initially exposed to RB 19 textile dye. In culture decolorization occurred only with RB 19. The total flattening of the UV-vis spectra of the samples indicates that decolorization was accompanied by biodegradation since the smoothening of absorbance peaks at 600 and at 280 nm is consistent with the reduction of azo linkages and the loss of aromatic rings (Figure 1).
Figure 1

UV-vis spectrum of a solution of RB19 before and after incubation with C. plicatilis cultures.

Effect of dye concentration and water content on fungal growth

When C. plicatilis was cultured in malt extract medium containing 50 mg/L RB 19, the percentage decolorization increased over time, reaching a peak of 99% decolorization after 15 days of incubation (Figure 2). The decolorization was not directly associated with fungal growth since over the period of 5–10 days the biomass level fell while the percentage decolorization was still increasing rapidly. In culture decolorization tests were then done with increasing concentrations of RB 19. Although these concentrations delayed fungal growth slightly, L. crinitus was able to develop in RB 19 concentrations as high as 200 or even 250 mg/L, giving percentage in culture decolorizations of 93.7% and 77.4%, respectively, after 15 days (Figure 3).
Figure 2

Decolorization of RB19 and growth of C. plicatilis in liquid minimal medium containing 50 mg/L dye. The absorbance of the culture supernatant was determined at 585 nm.

Figure 3

Effect of dye concentration on fungal growth and in culture decolorization. Absorbance at 585 nm was assessed after 15 days of culture.

In order to determine whether the fungal products responsible for dye decolorization were secreted or mycelium-associated, supernatants and mycelia from dead liquid cultures and eluted extracts from dead solid cultures were subjected to decolorization. Decolorization was highest in supernatants and eluted extracts, with relatively low decolorization being detected in mycelia (results not shown). Decolorization eluted from solid cultures was very similar to that obtained in the supernatants obtained from liquid cultures. Also, an RB 19 concentration of 50 mg/L was selected. At this concentration the dye was totally consumed and slightly higher decolorizations were obtained in the culture extracts.

Effect of carbon and nitrogen content on decolorization

In the case of white rot fungi, it has been reported that the production of lignin modifying enzymes (LMEs) and decolorization varies greatly, according to the type and concentration of carbon and nitrogen sources in culture media. These effects were therefore investigated. C. plicatilis was grown on solid minimal media that contained glucose, fructose, maltose, starch, sucrose or glycerol as the sole carbon source. For each substrate, three concentrations (5, 10 and 15 g/L) were tested. After 15 days of incubation, decolorization was performed using extracts from the solid medium. Drastic differences in the decolorization were observed. C. plicatilis produced the highest levels of decolorization when grown in 10 g/L glucose; 5 g/L maltose or 10 g/L fructose, with the decolorization ranging from 63.7% to 84.8%. Higher concentrations of these carbon sources did not lead to higher decolorization. With some carbon sources the yield of decolorization was very low. For example, with 15 g/L glycerol and 10 g/L starch decolorization of only 5-15% were produced (Figure 4A).
Figure 4

Effect of carbon and nitrogen on decolorization. A; Effect of carbon type and concentration on decolorization in solid medium. B; Effect of nitrogen source and concentraation on decolorization in solid media containing fructose (10 g/L) and glucose (10 g/L) as carbon source.

When nitrogen sources were assessed, using fructose (10 g/L) and glucose (10 g/L) as carbon sources (maltose was omitted due to its high cost), soluble extracts derived from cultures containing 5 g/L sodium nitrate, 15 g/L ammonium tartrate and 10 g/L ammonium oxalate performed the highest decolorization (Figure 4B). Again, some substrates promoted the decolorization and some of them, such as urea and ammonium chloride, were inhibitory. A significant observation is that the effect of a nitrogen source can depend on the accompanying carbon substrate. Thus, soluble extracts derived from cultures containing 5 g/L urea led to a 10% decolorization of this dye when combined with 5 g/L fructose in the culture medium, but when combined with 5 g/L glucose, this same concentration of urea gave a 5-fold higher decolorization. A similar effect was observed for 10 and 15 g/L ammonium oxalate with fructose and glucose.

Effect of pH, temperature and copper amount of cultures on decolorization of dye

Cultures were done in solid medium with optimized carbon and nitrogen contents (10 g/L glucose and 10 g/L ammonium oxalate) in five different initial pH values (at 26°C) and three different temperatures (with initial pH = 5.5). Other cultures were performed (at 26°C and initial pH = 5.5) in the presence of three different copper sulfate concentrations. Extracts from 4 day cultures were tested for decolorization. Decolorization of dye was better in extracts derived from cultures incubated at 26°C and with initial pH values ranging from 5.5 to 6.5 (Table 1). Extracts derived from copper supplemented cultures did not display an improvement in decolorization and at copper sulfate concentrations above 1 mM the decolorization was relatively low (Table 1).
Table 1

Effect of pH, temperature and copper concentration on decolorization (initial dye concentration; 50 mg/L)

T°C % Decolorization

26 95.1 ± 0.6

30 21.4 ± 2.3

35 0.04 ± 0.04

 

Cu 2+ (mM) % Decolorization

0 79.7 ± 2.2

0.1 84.3 ± 3.6

1.0 51.2 ± 1.3

2.0 49.6 ± 1.1

pH % Decolorization

5.5 99.2 ± 0.6

6 81.3 ± 1.2

6.5 68.7 ± 2.4

7 11.4 ± 1.7

Laccase activities

Figure 5 shows variations of laccase activities during decolorization within the ten days period for this organism. In an attempt to determine the possible role of ligninolytic enzymes on dye decolorization, laccase enzyme activity was monitored during the decolorization of this dye. In dye containing liquid cultures, laccase specific activity of this fungi was observed to have decreased by increasing dye concentration. These results emphasized the role of laccase in RB 19 decolorization. At higher concentrations levels than 50 mg/L RB 19, considered enzyme activities were diminished sharply, which is likely due to the toxicity of synthetic dyes as mentioned before (Figure 5).
Figure 5

Alteration of laccase enzyme activities.

Discussions

Several species and strains have been assessed for biodegradation of different pollutants such as crude oil [37], pentachlorophenol [38], DDT [39], trinitrotoluene [40] and some textile dyes [41, 42]. Here we report laccase activity in C. plicatilis, a relatively unexplored Coprinus species, and its participation in the decolorization of the textile dye reactive blue 19. Reactive blue 19 (RB 19) is a vinyl sulfone azo dye. Laccase activity in C. plicatilis was firstly described. Copper supplementation can stimulate laccase synthesis since the enzyme uses copper as cofactor, but the ion can also inhibit the growth of the organism [43]. Previous attempts to degrade it have focused on photo-catalytic and chemo-oxidative processes [4446]. Although these processes may degrade the dye partially or even totally, they have several drawbacks such as the generation of by-products, including chemical sludge, and high investment and operating costs [1].

It used to be generally accepted that carbon and nitrogen limitation favored the production of lignolytic enzymes in white rot fungi [47]. However, more recent results are somewhat contradictory. For example, in L. edodes, Buswell et al. [48] obtained 5-fold higher laccase levels under high nitrogen conditions than in low-nitrogen cultures, while Hatvani and Mécs [49], working on the biodegradation of dyes by Lentinus sp. grown in solid media, found that faster decolorization occurred at very low NH4Cl, peptone and malt extract concentrations. On the other hand, the lignolytic activity of L. edodes grown in liquid culture was stimulated by high N concentrations [50]. In the present work, the decolorization was affected by the type and concentration of the nitrogen source, but different trends occurred for different sources: higher concentrations of ammonium salts resulted in higher decolorization, while for sodium nitrate and urea higher production levels were obtained at the lower concentrations. Moreover, the final result varied drastically with the type of carbohydrate present in the culture medium. For example, the high decolorization obtained in cultures containing 5 g/L urea was almost lost when glucose was replaced with fructose as the main carbon source. A similar effect was observed for ammonium oxalate.

Conclusions

C. plicatilis cultures were able to decolorize and biodegrade the textile dye reactive blue 19. The decolorization was highly influenced by medium composition and culture conditions, being higher in media containing intermediate concentrations of ammonium oxalate and glucose. Decolorization of dye was associated with laccase displaying good tolerance to a wide range of pH values and temperatures, suggesting a potential role for this organism and enzyme in the remediation of real dye containing effluents. In conclusion, laccase activity in C. plicatilis was firstly described in this study.

Declarations

Acknowledgements

The author also would like to thank Shimadzu Co. for their advice.

Authors’ Affiliations

(1)
Department of Chemistry, Faculty of Science and Arts, Pamukkale University

References

  1. Robinson T, McMullan G, Marchant R, Nigam P: Remediation of dyes in textile effluents: a critical review on current treatment technologies with a proposed alternative. Bioresour Technol 2001, 77: 247–255. 10.1016/S0960-8524(00)00080-8View ArticleGoogle Scholar
  2. Hai FI, Yamamoto K, Fukushii K: Hybrid treatment systems for dye. Crit Rev Environ Sci Technol 2007, 37: 315–377. 10.1080/10643380601174723View ArticleGoogle Scholar
  3. Palmieri G, Cennamo G, Sannia G: Remazol brilliant R decolourisation by the fungus pleurotus ostreatus and its oxidative enzymatic system. Enzyme Microb Technol 2005, 36: 17–27. 10.1016/j.enzmictec.2004.03.026View ArticleGoogle Scholar
  4. Levin L, Papinutti L, Forchiassin F: Evaluation of Argentinean white rot fungi for their ability to produce lignin-modifying enzymes and decolorize industrial dyes. Bioresour Technol 2004, 94: 169–176. 10.1016/j.biortech.2003.12.002View ArticleGoogle Scholar
  5. Banat IM, Nigam P, Singh D, Marchant R: Microbial decolorization of textiledye- containing effluents: a review. Bioresour Technol 1996, 58: 217–227. 10.1016/S0960-8524(96)00113-7View ArticleGoogle Scholar
  6. Shyam SD, Satyshari D, Bhattacharyya BC: Dye decolorization in a column bioreactor using wood-degrading fungus Phanerochaete chrysosporium . Indian Chem Eng A 1995, 37: 176–180.Google Scholar
  7. Sarasa J, Roche MP, Ormad MP, Gimeno E, Puig A, Ovelleiro JL: Treatment of a wastewater resulting from dyes manufacturing with ozone and chemical coagulation. Water Res 1998, 32: 2721–2727. 10.1016/S0043-1354(98)00030-XView ArticleGoogle Scholar
  8. Konsowa AH: Decolorization ofwastewater containing direct dye by ozonation in a batch bubble column reactor. Desalination 2003, 158: 233–240. 10.1016/S0011-9164(03)00458-2View ArticleGoogle Scholar
  9. Blanquez P, Casas N, Font X, Gabarrell X, Sarra M, Caminal G, Vicent T: Mechanism of textile metal dye biotransformation by Trametes versicolor . Water Res 2004, 38: 2166–2172. 10.1016/j.watres.2004.01.019View ArticleGoogle Scholar
  10. Kirby N, McMullan G, Marchant R: Decolorization of an artificial textile effluent by Phanerochaete chrysosporium . Biotechnol Lett 1995, 17: 761–764. 10.1007/BF00130365View ArticleGoogle Scholar
  11. Swamy J, Ramsay J: The evaluation of white rot fungi in the decolorization of textile dyes. Enzyme Micro Technol 1999, 24: 130–137. 10.1016/S0141-0229(98)00105-7View ArticleGoogle Scholar
  12. Nilsson I, Moller A, Mattiasson B, Trubindamayugi MS, Welander U: Decolorization of synthetic and real textile wastewater by the use of white-rot fungi. Enzyme Microb Technol 2006, 38: 94–100. 10.1016/j.enzmictec.2005.04.020View ArticleGoogle Scholar
  13. Mehna A, Pratima B, Pramod KB: Studies on decolorization of effluent from a small pulp mill utilizing agri-residues with Trametes versicolor . Enzyme Microb Technol 1994, 22: 18–22.Google Scholar
  14. Heinfling A, Martinez MJ, Martinez AT, Bergbauer M, Szewzyk U: Transformation of industrial dyes by manganese peroxidases from Bjerkandera adusta and Pleurotus eryngii in a manganese-independent reaction. Appl Environ Microbiol 1998, 64: 2788–2793.Google Scholar
  15. Schliephake K, Lonergan GT: Laccase variation during dye decolorization in a 200-L packed-bed bioreactor. Biotechnol Lett 1996, 8: 881–886.View ArticleGoogle Scholar
  16. Koichi H, Yoshio W, Kazunori N: Decolorization of azo dye by the white rot Basidimycete Phanerochaete sordida by itsmanganese peroxidase. J Biosci Bioeng 2003, 95: 455–459.View ArticleGoogle Scholar
  17. Srinivasan SV, Murthy DVS: Statistical optimization for decolorization of textile dyes using Trametes versicolor . J Hazard Mater 2009, 165: 909–914. 10.1016/j.jhazmat.2008.10.072View ArticleGoogle Scholar
  18. Borchert M, Libra JA: Decolorization of reactive dyes by the white rot fungus Trametes versicolor in sequencing batch reactors. Biotech Bioeng 2001, 75: 313–321. 10.1002/bit.10026View ArticleGoogle Scholar
  19. Conneely A, Smyth WF, McMullan G: Study of the white-rot fungal degradation of selected phthalocyanine dyes by capillary electrophoresis and liquid chromatography. Anal Chim Acta 2002, 451: 259–270. 10.1016/S0003-2670(01)01415-5View ArticleGoogle Scholar
  20. Jarosz-Wilkołazka A, Rdest-Kochman’ska J, Malarczyk E, Wardas W, Leonowicz A: Fungi and their ability to decolorize azo and anthraquinonic dyes. Enzyme Microbiol Technol 2002, 30: 566–572. 10.1016/S0141-0229(02)00022-4View ArticleGoogle Scholar
  21. Martins MAM, Lima N, Silvestre AJD, Queiroz MJ: Comparative studies of fungal degradation of single or mixed bioaccessible reactive azo dyes. Chemosphere 2003, 52: 967–973. 10.1016/S0045-6535(03)00286-8View ArticleGoogle Scholar
  22. Moldes D, Couto SR, Cameselle C, Sanroma’n MA: Study of the degradation of dyes by MnP of Phanerochaete chrysosporium produced in a fixed-bed bioreactor. Chemosphere 2003, 51: 295–303. 10.1016/S0045-6535(02)00406-XView ArticleGoogle Scholar
  23. Moreira MT, Palma C, Mielgo I, Feijoo G, Lema JM: In vitro degradation of a polymeric dye (Poly R-478) by manganese peroxidase. Biotech Bioeng 2001, 75: 362–368. 10.1002/bit.10052View ArticleGoogle Scholar
  24. Rodrı’gues E, Pickard A, Vazquez-Duhalt R: Industrial dye decolorization by laccases from ligninolytic fungi. Curr Microbiol 1999, 38: 27–32. 10.1007/PL00006767View ArticleGoogle Scholar
  25. Andleeb S, Atiq N, Ali MI, Rehman F, Hameed A, Ahmad S: Biodegradation of anthraquinone dye by Aspergillus niger sa1 in self designed fluidized bed bioreactor. Iran J Environ Health Sci Eng 2010, 7: 371–376.Google Scholar
  26. Heinzkill M, Bech L, Halkier T, Schneider P, Anke T: Characterization of laccases and peroxidases from wood rotting fungi (Family Coprinacea e). Appl Environ Microbiol 1998, 64: 1601–1606.Google Scholar
  27. Abadulla E, Tzanov T, Costa S, Robra KH, Cavaco-Paulo A, Georg MG: Decolourization and detoxification of textile dyes with a laccase from Trametes hirsuta . Appl Environ Microbiol 2000, 66: 3357–3362. 10.1128/AEM.66.8.3357-3362.2000View ArticleGoogle Scholar
  28. Li FJ, Hong YZ, Xiao YZ, Xu YH, Fan W: High production of laccase B from Trametes sp. in Pichia pastoris . World J Microbiol Biotechnol 2007, 23: 741–745. 10.1007/s11274-006-9286-2View ArticleGoogle Scholar
  29. Canas AI, Camarero S: Laccases and their natural mediators: biotechnological tools for sustainable eco-friendly process. Biotechnol Adv 2010, 28(6):694–705. 10.1016/j.biotechadv.2010.05.002View ArticleGoogle Scholar
  30. Baldrian P: Fungal laccases – occurrence and properties. FEMS Microbiol Rev 2006, 30: 215–242. 10.1111/j.1574-4976.2005.00010.xView ArticleGoogle Scholar
  31. Trovaslet-Leroy M, Jolivalt C, Froment M, Brasme B, Lefebvre B, Daveloose D, Nachon F, Masson P: Application of laccase-mediator system (LMS) for the degradation of organophosphorus compounds. Chem Biol Interact 2010, 187: 393–396. 10.1016/j.cbi.2010.02.001View ArticleGoogle Scholar
  32. Widsten P, Kandelbauer A: Laccase applications in the forest products industry: A review. Enzym Microb Technol 2008, 42: 293–307. 10.1016/j.enzmictec.2007.12.003View ArticleGoogle Scholar
  33. Desai SS, Nitvanand C: Microbial laccases and their applications: a review. Asian J Biotechnol 2011, 3: 98–124. 10.3923/ajbkr.2011.98.124View ArticleGoogle Scholar
  34. Odaci D, Timur S, Pazarlioglu N, Montereali MR, Vastarella W, Pilloton R, Telefoncu A: Determination of phenolic acids using Trametes versicolor laccase. Talanta 2011, 71: 312–317.View ArticleGoogle Scholar
  35. Ardag Akdogan H, Kasikara Pazarlioglu N: Fluorene Biodegradation by P.ostreatus -I: Biodegradation by Free Cells. Process Biochem 2011, 46: 834–839. 10.1016/j.procbio.2010.12.011View ArticleGoogle Scholar
  36. Eggert C, Temp U, Dean JF, Eriksson KE: A fungal metabolite mediates degradation of non-phenolic lignin structures and synthetic lignin by laccase. FEBS Lett 1996, 391: 144–148. 10.1016/0014-5793(96)00719-3View ArticleGoogle Scholar
  37. Adenipekun CO, Fasidi IO: Bioremediation of oil-polluted soil by Lentinus subnudus, a Nigerian white-rot fungus. Afr J Biotechnol 2005, 4: 796–798.Google Scholar
  38. Machado KMG, Matheus DR, Monteiro RTR, Bononi VLR: Biodegradation of pentachlorophenol by tropical basidiomycetes in soils contaminated with industrial residues. World J Microbiol Biotechnol 2005, 21: 297–301. 10.1007/s11274-004-3693-zView ArticleGoogle Scholar
  39. Van Chung T, Khue DN, Minh DB, Cheng F: Use of fungal humus for 1,1,1- trichloro-2,2-bis(4-chlorophenyl)ethane (DDT) polluted soil treatment. Chem Asian J 2009, 21: 5967–5972.Google Scholar
  40. Samson J, Langlois E, Lei J, Piche Y, Chenevert R: Chenevert, Removal of 2,4,6-trinitrotoluene and 2,4-dinitrotoluene by fungi (Ceratocystis coerulescens, Lentinus lepideus and Trichoderma harzianum). Biotechnol Lett 1998, 20: 355–358. 10.1023/A:1005371112287View ArticleGoogle Scholar
  41. Minussi RC, de Moraes SG, Pastore GM, Duran N: Biodecolorization screening of synthetic dyes by four white-rot fungi in a solid medium: possible role of siderophores. Lett Appl Microbiol 2001, 33: 21–25. 10.1046/j.1472-765X.2001.00943.xView ArticleGoogle Scholar
  42. Gomes E, Aguiar AP, Carvalho CC, Bonfa MRB, da Silva R, Boscolo M: Ligninases production by basidiomycetes strains on lignocellulosis agricultural residues and their application in the decolorization of synthetic dyes. Braz J Microbiol 2009, 40: 31–39. 10.1590/S1517-83822009000100005View ArticleGoogle Scholar
  43. Shutova VV, Revin VV, Myakushina YA: The effect of copper ions on the production of laccase by the fungus Lentinus (Panus) tigrinus. Appl Biochem Microbiol 2008, 44: 619–623. 10.1134/S0003683808060100View ArticleGoogle Scholar
  44. Kurbus T, Le-Marechal AM, Voncina DB: Comparison of H 2 O 2 /UV, H 2 O 2 /O 3 and H 2 O 2 /Fe 2+ processes for the decolorisation of vinylsulphone reactive dyes. Dyes Pigm 2003, 58: 245–252. 10.1016/S0143-7208(03)00085-8View ArticleGoogle Scholar
  45. Voncina DB, Le-Marechal AM: Reactive dye decolorization using combined ultrasound/H 2 O 2 . Dyes Pigm 2003, 59: 173–179. 10.1016/S0143-7208(03)00101-3View ArticleGoogle Scholar
  46. Mahmoodi NM, Arami M, Limaee NY, Tabrizi NS: Kinetics of heterogeneous photocatalytic degradation of reactive dyes in an immobilized TiO 2 photocatalytic reactor. J Colloid Interface Sci 2006, 295: 159–164. 10.1016/j.jcis.2005.08.007View ArticleGoogle Scholar
  47. Kirk K, Farrell RL: Enzymatic “combustion”: the microbial degradation of lignin. Ann Rev Microbiol 1987, 41: 465–505. 10.1146/annurev.mi.41.100187.002341View ArticleGoogle Scholar
  48. Buswell JA, Cai Y, Chang ST: Effect of nutrient nitrogen and manganese on manganese peroxidase and lactase production by Lentinula ( Lentinus ) edodes . FEMS Microbiol Lett 1995, 128: 81–88. 10.1111/j.1574-6968.1995.tb07504.xView ArticleGoogle Scholar
  49. Hatvani N, Mécs I: Effect of the nutrient composition on dye decolorisation and extracellular enzyme production by Lentinus edodes on solid medium. Enzyme Microbiol Technol 2002, 30: 381–386. 10.1016/S0141-0229(01)00512-9View ArticleGoogle Scholar
  50. Leatham GF, Kirk TK: Regulation of ligninolytic activity by nutrient Nin white rot basidiomycetes. FEMS Microbiol Lett 1983, 7: 16–65.Google Scholar

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