Fungal decolouration and degradation of azo dyes: A review

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journal homepage: www.elsevier.com/locate/fbr

Review

Fungal decolouration and degradation of azo dyes: A review Sudip Kumar SENa, Smita RAUTb, Partha BANDYOPADHYAYa, Sangeeta RAUTb,* a

Ingene Research Laboratory, Biostadt India Limited, Aurangabad, Maharashtra, India Department of Biotechnology, Gandhi Institute of Engineering and Technology, Gunupur, Odisha, India

b

article info

abstract

Article history:

The textile industry is a substantial consumer of water and produces enormous volumes of

Received 8 April 2016

contaminated water; the most important contaminants are azo dyes. Fungal processes for

Received in revised form

the treatment of textile wastewater have the advantage of being cost-effective and envi-

10 June 2016

ronmentally friendly and producing less sludge. Unlike bacteria, fungi possessed strong

Accepted 12 June 2016

ability of degrading complex organic compounds by producing extracellular ligninolytic enzymes including laccase, manganese peroxidase and lignin peroxidase, hence, researchers

Keywords:

paid more attention on fungi in recent years. The mechanism of fungal decolouration oc-

Azo dyes

curs from adsorption, enzymatic degradation or a combination of both. The goal of fungal

Bio-sorption

treatment is to decolorize and detoxify the dye contaminated effluents. In this review, we

Decolouration

summarize the methodologies used to evaluate the toxicity of azo dyes and their degrada-

Degradation

tion products. Recent studies on the decolouration or degradation of azo dyes with

Fungi

Advanced Oxidation Processes (AOPs) and Microbial Fuel Cells (MFCs) are discussed in

Toxicity

this review. ª 2016 British Mycological Society. Published by Elsevier Ltd. All rights reserved.

1.

Introduction

In recent years, greater attention has been paid to the discharge of effluents containing synthetic dyes (Miranda et al., 2013). Worldwide, 280,000 tons of textile dyes are discharged in industrial effluents every year (Jin et al., 2007). To dye 1 kg of cotton with reactive dyes, 0.6e0.8 kg NaCl, 30e60 g dyestuff and 70e150 L water are necessary; the wastewater produced has 20e30 % of the applied unfixed reactive dyes, with an average concentration of 2000 ppm, high salt content and dyeing auxiliaries (Babu et al., 2007).

The release of these effluents into the environment is undesirable due to the serious environmental problems linked with the dyes and their breakdown products (Ozdemir et al., 2013). Among commercial synthetic dyes, azo dyes are the largest class with a broad range of colours and structures and represents up to 70 % of the total textile dyestuffs used (Lang et al., 2013). Azo dyes are regularly used in various applications in food, pharmaceutical, paper, cosmetic, textile and leather industries (Saratale et al., 2013). These dyes belong to the class of aromatic and heterocyclic compounds having the azo bond (eN]Ne) which are recalcitrant and

* Corresponding author. Department of Biotechnology, Gunupur, Rayagada, Odisha, 765022, India. E-mail address: [email protected] (S. Raut). http://dx.doi.org/10.1016/j.fbr.2016.06.003 1749-4613/ª 2016 British Mycological Society. Published by Elsevier Ltd. All rights reserved.

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2

even possess carcinogenic properties (Saratale et al., 2011). As they are introduced into the environment due to the inefficiency during the dyeing process and subsequent discharge, they accumulate mainly in water bodies and have adverse effects in terms of dissolved oxygen (DO), Biological Oxygen Demand (BOD), Chemical Oxygen Demand (COD), colour, etc., (Saratale et al., 2013; Sol
ıs et al., 2012). Therefore, their removal from wastewater is of utmost importance before discharge of the wastewater into the environment (Ayed et al., 2011). Extensive research and development has focussed on biological methods as an eco-friendly alternative for remediation of dyes (Kaushik and Malik, 2009). Most studies on azo dye biodegradation have focused on bacteria and fungi, in which bacteria were widely used for azo dyes decolorization due to their high activity, extensive distribution and strong adaptability (Pearce et al., 2003; Dos Santos et al., 2007). However, decolorization of products such as aromatic amines can inhibit the activity of bacteria (Qu et al., 2010). By contrast, fungi can degrade complex organic compounds through catalysis with extracellular ligninolytic enzymes including laccase, manganese peroxidase and lignin peroxidase (Gomi et al., 2011). Many fungal species such as Pleurotus ostreatus, Pichia sp., Penicillium sp., and Candida tropicalis, have been confirmed to decolorize azo dyes through adsorption and/or degradation (Kalmis et al., 2008; Qu et al., 2010; Gou et al., 2009; Tan et al., 2013). Moreover, some fungi can partially or even completely mineralize azo dyes (Miranda et al., 2013; Qu et al., 2012; Tan et al., 2013). Compared with the dyes themselves, some decolorization intermediates such as aromatic amines and phenolics can be highly toxicity and lower biodegradability. Fungi have shown strong adaptability and efficiency in the removal of these aromatic compounds. For instance, the fungal strains belonging to Thamnidium elegans, Zygorhynchus moelleri and Yarrowia lipolytica were confirmed to efficiently degrade aromatic phenolics (Papanikolaou et al., 2008; Bellou et al., 2014). In this context, fungi offer an efficient system due to large surface area and easy solideliquid separation (Mishra and Malik, 2013). Fungi also possess multiple mechanisms for degradation of organic and inorganic contaminants (Awasthi et al., 2014). However, pollutant removal has been largely studied under single pollutant exposures and often using pure cultures (Singh and Singh, 2014; Mishra and Malik, 2012). These studies demonstrate the utility of a certain fungal strain for removing a particular pollutant. Moreover, there are wide variations in the dye uptake capacity among various strains. Industrial effluents are a cocktail of various metals and organic contaminants (Ruta et al., 2010; Yadav et al., 2010). For example, textile industry effluents contain both residual dyes (from dyeing operations) and metals (used as mordant). Likewise pulp and paper industries, tannery industries and dyeing industries are also generate effluents rich in metals as well as dyes. Therefore, in order to develop a biological system capable of remediating such wastewater, diverse types of microbial strains need to be used to form of a consortia (Mahapatra et al., 2014). The use of a microbial consortia has a clear advantage for bioremediation applications as a richer metabolic network can be preserved and exploited for the bioremediation of cocontaminated matrices. Therefore, in a mixed waste stream,

S. K. Sen et al.

each of the consortium partners can specialize in the uptake of a particular contaminant. As a result, simultaneous removal of several contaminants can be successfully accomplished. Therefore, exploitation and study of new fungal strains capable of degrading azo dyes efficiently are still necessary for field application. Though the use of specific contaminant-degrading fungi in the wastewater treatment system can provide an effective way to enhance the degradation of toxic organic pollutants, high degradation rate of pollutants would not persist for a long time in wastewater treatment systems due to the loss of degrading microorganisms through being washed out from the system (Li et al., 2013). Immobilization of fungi has been suggested as a strategy for maintaining efficient degradation biomass in the systems, and different methods of immobilization have been developed which lead to improved effectiveness in wastewater treatment (Moreno-Garrido, 2008). Entrapment is one of the most commonly used methods for immobilizing single-celled microorganisms such as yeast, which can effectively avoid the loss of microorganisms from treatment systems. In this way, the microbial cells are immobilized in alginate beads or polyvinyl alcohol (PVA) gel pellets (Samuel et al., 2013; Mart
ınez et al., 2013). Unlike the bacteria in conventional wastewater treatment systems, aerobic white-rot fungi (WRF) can degrade wide varieties of resistant compounds including textile dyes by non-specific extracellular enzymes (Yang et al., 2013). Several white rot fungi like Phanerochaete chrysosporium and Trametes versicolor, due to their efficient ligninolytic enzymatic systems, have been reported to degrade or sequester azo, heterocyclic, reactive or polymeric dyes (Sol
ıs et al., 2012). This review summarizes the recent achievements in the fungal technologies developed for the removal of azo dyes using simulated effluents and real textile industry effluents. The principal factors that affect dye removal are analysed, and the fungal decolouration systems based on the use of yeast, filamentous fungi, genetically modified strains, fungal consortia, and fungal processes in combination with AOPs and MFCs are discussed with particular reference to the analysis of the toxicity of the metabolic products of azo dye decolouration and the methodologies used to evaluate this toxicity.

2. Generalities in the fungal decolouration of azo dyes A wide variety of fungal organisms are capable of decolorizing a wide range of azo dyes (Fu and Viraraghavan, 2001). Many genera of fungi have been employed either in living or inactivated form. The use of white-rot fungi such as P. chrysosporium in decolorizing textile wastewater has been widely reported in literature (Bilgic et al., 1997; Cammarota and Sant Anna, 1992; Lankinen et al., 1991; Tatarko and Bumpus, 1998; Young and Yu, 1997; Gomaa et al., 2008; Sharma et al., 2009; Faraco et al., 2009). Apart from white-rot fungi, other fungi such as Aspergillus niger (Fu and Viraraghavan, 2000, 2001, 2002), Rhizopus arrhizus (Zhou and Banks, 1991), Rhizopus oryzae (Gallagher et al., 1997) can also decolorize and/or biosorb diverse dyes.

Please cite this article in press as: Sen, S.K., et al., Fungal decolouration and degradation of azo dyes: A review, Fungal Biology Reviews (2016), http://dx.doi.org/10.1016/j.fbr.2016.06.003

Fungal decolouration and degradation of azo dyes

3

Fungal bioreactors for dye decolorization Fungal bioreactors for dye decolorization and degradation have been developed recently. Both colour removal and degradation can take place individually or simultaneously in the reactor. White-rot fungi have been widely studied for its decolorization potential in bioreactor systems. At present no information on commercial applications of fungal bioreactor systems are available. Rotating drum, packed bed, fluidized bed, immobilized and membrane bioreactors have been used as decolorizing bioreactors (Table 1). A membrane bioreactor using T. versicolor combined with reverse osmosis was effective for decolorization of dye wastewater (Kim et al., 2004). A wood-rotting fungal strain F29 decolorized 95e99 % Orange II in a continuous packed bed and fluidized bed bioreactor systems (Zhang et al., 1999). Immobilized bioreactors have been found to exhibit good biological activities and abilities for long time operation (Singh, 2006). Irpex lacteus immobilized in a pine wood (PW) reactor decolorized Remazol Brilliant Blue R more rapidly than a polyurethane foam (PUF) reactor (Kasinath et al., 2003). However, contrary to the performance shown in aseptic batch tests, the application of white rot fungi in continuous bioreactors for dye wastewater treatment has been so far impeded by problems such as excessive growth of fungi causing reactor-clogging (Zhang et al., 1999), bacterial

contamination inhibiting fungal decolouration (Hai et al., 2009; Libra et al., 2003), and loss of the extracellular enzymes and mediators essential for dye degradation with treated effluent (Hai et al., 2012). Of major concern is the inhibition of enzymatic activity and decolouration performance due to bacterial contamination. White rot fungi grows slowly compared with bacteria. Once bacteria invade the system, bacteria can compete with white rot fungi for substrate and can thus inhibit fungal growth, rendering the white rot fungi system eventually ineffective for dye decolouration and degradation (Hai et al., 2009; Libra et al., 2003). The approaches that have been used to date to improve fungal decolouration under non-sterile environment include use of low pH and nitrogen-limited medium, omission of certain trace elements (e.g., iron) from the culture medium, periodic addition of fresh biomass, de-coupling of growth (sterile condition) and decolouration (non-sterile condition) stages, and use of bactericide (e.g. ozone) to control the bacterial contamination (Zhou and Wen, 2009; Blanquez et al., 2008; Gao et al., 2006; Libra et al., 2003). None of these approaches alone has been proven to be a long-term solution to the problem of bacterial contamination, and further research is deemed imperative. Different morphological forms of fungi can show different characteristics such as specific growth rate and enzyme secretion: one form may include several physiological states of fungi with the production of specific metabolites, which may not occur

Table 1 e Fungal bioreactors for dye decolorization. Fungus

Bioreactor configuration

Phanerochaete chrysosporium Trametes versicolor

Rotating drum

Poly R-478

19

MnP, LiP

15 m

Stirred-tank reactor

Poly R-478, 200 mg/L

80

41 d

Basic Blue 22, 200 mg/L Eerzol Turquoise Blue-G, 200 mg/L

98 82

Biotransformation and adsorption Lignolytic enzyme Lignolytic enzyme

24 h 20 d

Ge et al., 2004 Kapdan and Kargi, 2002

Poly R-478, 0.1 g/L

80

MnP

24 h

Mielgo et al., 2002

Remazol Brilliant Blue R, 150 mg/L Remazol Brilliant Blue R, 150 mg/L Indigo Carmine

100

MnP and laccase

6d

Kasinath et al., 2003

100

MnP and laccase

6d

Novotny et al., 2004

100

Laccase activity

3d

Rodr
ıguez Couto et al., 2004

Direct Blue 15, 20 mg/L

95e100

MnP

Methylene Blue 60 mg/L

100

MnP

Reactive Blue 4, 200 mg/L

Phanerochaete sordida Immobilized on RBC Coriolus versicolor Immobilized in form of biofilm, activated sludge, and wood ash P. chrysosporium Immobilized on Polyurethane foam, pulsed packed bed Irpex lacteus Immobilized on PUF or PW, packed bed reactor Irpex lacteus Immobilized on PW cubes, packed bed reactor Trametes hirsute Immobilized on stainless steel sponge, fixed bed bioreactor P. chrysosporium Immobilized on ZrOCl2activated pumice, packed bed reactor P. chrysosporium Immobilized on mineral kissiris T. versicolor Immobilized on birch wood, continuous RBC T. hirsute Immobilized on ground orange Orange peelings, fixed bed reactor

Dye

Percent Enzyme activity Duration removal

Reference Dominguez et al., 2001 Leidig et al., 1999

Pazarliogl et al., 2005

8d

Karimi et al., 2006

70

3d

Nilsson et al., 2006

Indigo Carmine

94

3d

Rodr
ıguez Couto et al., 2006

Methyl Orange Poly R-478

81.4 46.9

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4

S. K. Sen et al.

in another form. The effects of fungal morphology on fine chemical synthesis by pure fungal cultures has been well documented (Papagianni, 2004); however, very few articles have specifically considered the effects of morphology on the degree of removal of recalcitrant pollutants such as dye. Furthermore, studies available to date have focused mostly on a single type of morphologies (e.g., dispersed filaments, spherical pellets or attached biofilm) under aseptic conditions. For example, Bermek et al. (2004) reported that growth in dispersed filaments can increase the viscosity of the medium (low oxygen levels), limiting the mass transfer, causing a lower ligninolytic enzyme production. Borras et al. (2008) demonstrated high degree of dye decolouration using the ligninolytic fungus Trametes versicolor in pellet form; however, the performance was not compared with other morphological forms. Notably, compared with the morphologies encountered in suspended cultures (e.g., dispersed filaments and spherical pellets), immobilised cultures (entrapment into some matrix or attached growth onto some supports) tend to show a higher level of enzymatic activity and more resilience to environmental perturbations such as shear damage and pH/toxic shock (Rodriguez Couto, 2009). For example, Zhang et al. (1999) observed better decolouration of an azo dye by alginate-immobilised fungus than dispersed filaments in different reactor configurations; however, direct comparison with pellet morphology was not performed in their study. A range of techniques for pellet formation (Erdal and  Taskin, 2006; Yang et al., 2008a,b; Zmak et al., 2006) and carrier materials for fungal immobilisation (Rodriguez Couto, 2009) have been explored with encouraging dye removal performance; however, an important area of research requiring greater focus is the long term operation of bioreactors with pellets or attached growth fungi under non-sterile conditions. A comprehensive literature review could identify only a few studies where the removal performance of bioreactors containing pellets or attached growth fungi under non-sterile conditions has been investigated (Leidig et al., 1999; Nilsson et al., 2006; Tang et al., 2011). The advantages of membrane bioreactors (MBR) such as maintenance of high WRF concentration and prevention of enzyme washout have been demonstrated in previous studies (Hai et al., 2006, 2012).

3.

Fungal decolouration mechanisms

Biosorption Biosorption mechanisms may play an important role in the decolorization of dyes by living fungi. For dead cells, the mechanism is biosorption, which involves physico-chemical interactions such as adsorption, deposition, and ion-exchange. Decolorization of dye wastewater by fungal biomass has been extensively reviewed by Kaushik and Malik (2009) and Singh (2006). Limited information is available on interactions between dead fungal biomass and a variety of dyes with complex molecular structures. Fu and Viraraghavan (2002) studied the roles played by functional groups such as carboxyl, amino, phosphate and lipid fractions present in fungal biomass from Aspergillus niger in bio-sorption of four different dyes. In biosorption of Basic Blue 9 on A. niger, carboxyl and amino groups

were found to be the main binding sites while in bio-sorption of Acid Blue 29, only amino group was a major site and electrostatic attraction was believed to be the primary mechanism. In bio-sorption of Congo Red, the amino, carboxylic acid, phosphate groups and lipid fractions were all found to be important binding sites and in addition to electrostatic attraction, other mechanisms were also believed to be involved in biosorption. In biosorption of Disperse Red 1, physical and chemical adsorption along with electrostatic attraction was found to be the mechanism of biosorption while amino group and lipid fractions were the major binding sites. Research has shown that some pre-treatment processes can increase the adsorption capacity of biomass. The pretreatment methods include autoclaving or treating biomass with organic chemicals such as formaldehyde or inorganic chemicals such as NaOH, H2SO4, NaHCO3 and CaCl2. Fu and Viraraghavan (2000, 2001) used autoclaving and pretreatment with chemicals such as 0.1 M NaOH, 0.1 M HCl, 0.1 M H2SO4, of living A. niger biomass. It was found that autoclaving increased biosorption capacity of Basic Blue 9 from 1.17 mg/g to 18.54 mg/g, while 0.1 M H2SO4 pretreatment enhanced biosorption capacity from 6.63 mg/g to 13.83 mg/g for Acid Blue 29. It is suggested that autoclaving could rupture the fungal structure and expose potential binding sites for the dye while acid pre-treatment could change the negatively charged surface of fungal biomass to a positively charged surface, thus increasing the attraction between fungal biomass and Acid Blue 29, an anionic dye. Arica and Bayramoglu (2007) observed heating the biomass Lentinus sajor-caju at 100
C for 10 min enhanced the biosorption capacity while base-treatment with 0.1 M NaOH lowered the biosorption capacity of the fungi to remove Reactive Red 120.

Biodegradation (enzymatic degradation) of dyes For living cells, the major mechanism for decolorization of dyes is biodegradation due to the production of lignin modifying enzymes, laccase, manganese peroxidase (MnP) and lignin peroxidase (LiP) (Raghukumar et al., 1996). The relative contributions of LiP, MnP and laccase to the decolorization of dyes may be different for each fungus. Recently, there has been an increased interest in determining the exact mechanism of how these organic pollutants are broken down. Depending on the enzyme used, different mechanistic pathways have been reported in the literature and reviewed below:

Degradation of azo dyes by azo reductases Degradation of azo compounds by azo reductases has been shown to be almost exclusively anaerobic in nature. There are numerous reports of recalcitrant azo dyes being efficiently degraded by anaerobic microbes (due to their azo reductases) or purified azo reductase enzymes, but only under anaerobic conditions (Chen et al., 2005). These reactions require reducing co-factors like nicotinamide adenine dinucleotide (NADþ), nicotinamide adenine dinucleotide phosphate (NADPþ), etc. for catalysing the enzymatic reduction of azo dyes by azo reductases. Since these azo reductases are cytosolic in nature, it is assumed that azo dyes have to be transported into cell

Please cite this article in press as: Sen, S.K., et al., Fungal decolouration and degradation of azo dyes: A review, Fungal Biology Reviews (2016), http://dx.doi.org/10.1016/j.fbr.2016.06.003

Fungal decolouration and degradation of azo dyes

before they are degraded by dye-degrading anaerobic microbes. However, some studies have suggested an alternative mechanism for high molecular weight and highly charged dyes, which are unlikely to pass through the cell membranes. It has been hypothesized that some of the azo dye reducing activity of the dye may not be dependent on the intracellular uptake of the macromolecule (Robinson et al., 2001). This alternate proposed mechanism suggests the involvement of the electron reduction of these dyes in the extracellular environment of the microbes. For this to happen, the fungi should firstly link itself with the intracellular electron transport system and the dye molecules. This linkage requires that the electron transport component must be present in the outer membrane of the fungal cells, so that at the cell surface, the dye and the redox mediator comes in a direct contact. Moreover, it has been shown that redox mediators acting as electron carriers can dramatically increase the degradation of dye molecules by azo reductases (Robinson et al., 2001). The mediator compounds can be generated by the fungal metabolism or they may need to be added externally. As mentioned earlier these reactions work only in the absence of oxygen, as oxygen can inhibit the reduction mechanism and preferentially oxidize the redox mediators as compared to the dye molecule (Pricelius et al., 2007).

Azo dye degradation by laccases Laccases are copper-containing multimeric glycoproteins that have generic phenol oxidase activity (Majeau et al., 2010). The presence of copper gives a blue colour to these enzymes and is responsible for the actual oxidation of the substrate. Laccases use molecular oxygen to oxidize various aromatic and nonaromatic compounds by abstracting protons and creating radicals in the mechanistic process. These radicals can then further participate in other reactions such as polymerization, hydration or proton abstraction. Degradation of phenols and aromatics in effluents either in the free or immobilized form have been explored by using various enzymes.

Dye degradation by peroxidases Decolorization of Acid Orange 7 using peroxidase enzyme produced by the fungus Coprinus cinereus has also been reported in the literature. Under optimized conditions of pH, dye concentration and temperature, an overall removal of the dye was reported in a very short time (Yousefi et al., 2010). Although these enzymes are more genetically related to other fungal peroxidases including lignin peroxidase (LiP) and manganese peroxidase (MnP), they are very similar to classical plant peroxidases such as horseradish peroxidase (HRP) rather than the ligninolytic peroxidases in substrate specificity, pH optimum (nearly neutral) and specific activity.

4. Factors affecting fungal decolorization of dyes The degradation rate and efficiency of the enzymatic decolorization of the dye are highly dependent on a number of operational parameters that govern the degradation of the organic molecule. This section will briefly discuss the significance of each operational parameter.

5

Influence of nutrients on dye decolorization Biodegradation of dyes can be enhanced by improving the initial growth conditions. Glucose, starch, maltose, and cellobios were found to be good carbon sources for decolorization of cotton bleaching effluent by white-rot fungus (Zhang et al., 1999). P. chrysosporium showed superior performance at a glucose concentration of 5 g/L and an ammonium chloride concentration of 0.05 g/L in decolorizing Methyl Violet (Radha et al., 2005). A high dose of nutrient nitrogen was found to inhibit decolorization of Congo Red (Tatarko and Bumpus, 1998). Nitrogen was found to have no effect on the decolorization of dyes by Cyathus bulleri (Vasdev et al.,1995). As decolorization of dyes by P. chrysosporium occurs in secondary metabolic conditions, the enzyme LiP is released by fungal cells under either carbon or nitrogen limitation (Zhen and Yu, 1998).

Effect of oxygen, pH and temperature in the decolouration process Dye degradation can occur under anaerobic and aerobic conditions by different microbial organisms. Carbon sources such as glucose, starch, acetate, and ethanol affects the decolouration process of dyes under anaerobic conditions (der Zee and Villaverde, 2005). Under anaerobic conditions, reductive enzyme activities are generally higher; however, a small amount of oxygen is also required for the regeneration of reducing cofactors (e.g. NADH and NADPH) as well as oxidative enzymes which may also be involved in the degradation of azo dyes. The intermediates formed during azo dye reduction reaction, like the simple aromatic compounds, are broken down via hydroxylation and ring-opening in the presence of oxygen (Pandey et al., 2007). Hence, for the complete mineralization of the azo molecules, aerobic conditions are preferable. Thus, for the most effective effluent treatment an anaerobic process with subsequent aerobic treatment can be used to decolorize wastewaters containing dyes and improve their biodegradability (You and Teng, 2009). The main reasons limiting the degradation rates of dyes in wastewater streams are a lack of oxygen and the use of high energy cost electron acceptors needed for the mineralization processes (Gallizia et al., 2004). pH is an important factor for fungal growth. Fungi are generally found to grow at low pH, normally ranging from 4 to 5. The ionic forms of the dye in solution and the surface electrical charge of the biomass depend on solution pH. Therefore, solution pH influences both the fungal biomass surface dye binding sites and the dye chemistry in the medium. Fu and Viraraghavan (2000, 2001) reported that the effective initial pH of dye solution was 6 and 4, respectively, for Basic Blue 9 and Acid Blue 29. At pH of 2.0, no biosorption occurred for Basic Blue 9 due to the high concentration of protons, while at pH of 12, no biosorption occurred for Acid Blue 29. Arica and Bayramoglu (2007) reported that as the pH was decreased, the biosorption of Reactive Red 120 dye on the fungal biomass L. sajor-caju increased. Similarly, O’Mahony et al. (2002) observed maximum removal of reactive dye Remazol Black-B in the range of 1e2 with a sharp drop off at higher values. At lower pH values the fungal biomass will have a net positive charge. These charged sites become available for binding anionic groups such as reactive dyes.

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6

The optimum growth temperature for most of the fungi is found to be at about 25e35
C. Temperature is another parameter that appears to have an effect on the enzymatic degradation of dyes. For example, the decolorization ability of C. versicolor improved with an increase in incubation temperature to 30
C as the optimum temperature showing 92 % decolorization of Cibanon Blue GFJ-MD in 10 d (Asgher et al., 2008a,b). Similarly, the temperatures for optimum growth, ligninase activities and dye decolorization for most white rot fungi were found to be around 25e37
C (Asgher et al., 2007). We have also observed very similar results for in vitro experiments using SBP/H2O2 system, where increasing the temperature from 25
C to 45
C led to increased dye degradation. In fact, this was also found to hold for the bio-catalytic oxidation of environmental pollutant bisphenol A by HRP (Mei and Nicell, 2008). As expected for enzyme-based biological systems, increasing the incubation temperature too high would lead to microbial growth inhibition as well as to denaturation of enzymes, and hence to eventual decline in dye degradation efficiency. A simultaneous reduction in the efficiency of the decolorization of the dyes Bromophenol and Methyl Orange by about 50 % was shown in a recent study in the HRP/H2O2 system wherein the temperature was increased from 30 to 80
C (Liu et al., 2006). This behaviour is most likely due to the thermal denaturation and subsequent failure of enzymatic activity at higher temperatures. Most textile and other dye effluents are produced at relatively high temperatures and hence temperature will be an important factor in real application of biosorption in future. Arica and Bayramoglu (2007) found that biosorption of dye by L. sajor-caju increased with increasing temperature from 5 to 35
C. Aksu and Cagatay (2006) observed an increase in uptake of dye with increasing temperature up to 45
C for R. arrhizus showing endothermic character of biosorption.

Effects of enzyme and dye concentration on degradation The degradation of dyes is very much affected by the enzyme activity and the initial dye concentration. In some cases, e.g. Acid Red 27 (AR27), the absolute enzyme activity can be crucial and below a certain level, decolorization did not occur and an increased amount of purified laccase catalysed the complete decolorization of recalcitrant AR 27 within 24 h. Similarly in other studies, varying concentrations of different dyes were tested and lower concentrations (50e500 mg/L) were reported to be best decolorized (Wells et al., 2006; Levin et al., 2004). In another study, Lentinus crinitus cultured in Liquid Minimal Medium was successfully used to degrade 0.1 g/L concentration of Reactive Blue 220 (RB-220). However, the authors reported that increasing concentrations of RB-220 significantly delayed fungal growth, suggesting that higher concentration of dyes may result in decreased dye degradation because of their toxicity to the microbial organisms (Niebisch et al., 2010). Likewise, the effect of initial Methyl Red dye concentration (750, 800, 850, 900, 950 and 1000 ppm) on the percentage of decolorization by using an isolated Sphingomonas paucimobilis has also been published, which showed that the percentage of dye decolorization decreased with increase in the initial dye concentrations (Ayed et al.,2011). In summary, not only do higher dye concentrations adversely affect pure enzyme-based

S. K. Sen et al.

decolouration processes, they can negatively affect microbial growth and decrease the efficiency of microbe-based dye degradation processes as well. Dye concentration also affects the efficiency of colour removal. The decolorization efficiency by Coriolus versicolor decreased from 100 % to 80 % when the dye concentration was increased from 100 to 500 mg/L to 700e1200 mg/L (Kapdan et al., 2000).

Presence of redox mediators on dye degradation Although enzyme-mediated degradation of dyes [either in vivo (microbes) or in vitro (pure enzymes)] is very versatile and efficient. Redox mediators such as 1hydroxybenzotriazole (HOBT), veratryl alcohol, violuric acid, 2-methoxyphenothiazone, etc are frequently used. (Husain and Husain, 2011). More recently, natural compounds, such as syringaldehyde and acetosyringone, have also been explored as eco-friendly laccase mediators for various environmental applications (Cho et al., 2007). The oxidation of a substrate by a laccase/peroxidase enzyme occurs because the redox mediator forms cation radicals, which can be formed by two mechanisms. Firstly, the substrate can undergo a one-electron oxidation in the presence of a redox mediator and transform to a radical cation and secondly the mediator can abstract a H-atom from the substrate and convert it into a radical, which can then cause the substrate to co-oxidize (Fabbrini et al., 2002). There are numerous published examples on the use of mediators to efficiently degrade various classes of dyes. For example, SBP/H2O2 alone failed to degrade Rhodamine B, however as soon as the mediator HOBT was added to the reaction mixture, the dye degraded almost completely. Similarly, Immobilized turnip peroxidases in the presence of HOBT have been efficiently used to degrade Direct Red 23, Direct Red 239, Direct Blue 80 and Direct Yellow 4 dyes. Furthermore, researchers have reported examining the efficiency of six different mediators together with fenugreek seed peroxidase (with HOBT being the best), to decolorize textile effluents (Husain et al., 2010). In another study it was demonstrated that Malachite Green (MG) decolorization was enhanced in the presence of all mediators tested, however vanillin was found to be more effective in laccasemediated degradation of MG than HOBT (Bibi et al., 2011). Additionally, this study also demonstrated that laccase/mediators system could produce metabolites that were less toxic than the parent compound (Bibi et al., 2011). Interestingly, a different laccase-mediated study showed that HOBT was the best mediator in decolorizing textile wastewater effluent, but it was found that only laccaseeacetosyringone treated effluent was not toxic, whereas crude and laccaseeHOBT treated effluent remained toxic (Khlifi et al.,2010). Clearly, it appears that different mediators degrade different dyes via different mechanism, some of which may produce still toxic metabolites. Lastly, it is worth highlighting the amazing potential these microbes/enzymes offer for wastewater remediation.

Influence of azo dye structure in the decolouration process The molecular structure of dyes is found to have an effect on the extent of decolorization. Wong and Yu (1999) reported that

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Fungal decolouration and degradation of azo dyes

dye decolorization by T. versicolor was dependant on dye structures. Spadaro et al., (1992) observed that aromatic rings with substituents such as hydroxyl, amino, acetamido, or nitro functions were mineralized to a greater extent than unsubstituted rings in dye decolorization by P. chrysosporium. Since microbial and enzyme-based dye degradation involve binding of dyes to enzymes, it is not surprising that chemical structures of dyes strongly affect their decolorization efficiencies. However, it is difficult to suggest the actual molecular mechanism, because dye structure and efficiency of redox mediators contributes largely to peroxidase mediated catalysis. Azo dyes are electron-deficient molecules which can undergo degradation via azo reduction. Studies of model azo dyes have shown that dyes which have hydroxyl group either in the ortho- or para-position relative to azo-bond were the most reactive ones and are also more prone to oxidation when treated with peroxidase in the presence of redox mediators. Moreover, the efficiency of colour removal of dyes is strongly dependent on the steric effect of chemical substituents on dyes. The electron-withdrawing substituent may contribute towards recalcitrance of dyes which undergo redox mediated enzymatic decolorization (Almansa et al., 2004). The main function of the peroxidase/redox mediator thus consists of oxidatively rendering the azo-dye more susceptible to further nucleophilic attack and nitrogen is eliminated in molecular form. Strong electron withdrawing group such as sulfo groups at specific positions in an azo dye are more easily biodegradable as compared to the ones with a carboxyl group (Hsueh and Chen, 2008). Dyes with sulfonate group (such as Reactive Blue 15) were found to exhibit a strong electron-withdrawing effect and thus exhibited low overall reactivity. Literature findings have also indicated a higher affinity of laccase for anthraquinone dyes, for example, Jarosz-Wilko1azka et al. have shown that various fungi could degrade Basic Blue 22 much more efficiently than the azo dye, Acid Red 183 (Jarosz-Wilko1azka et al., 2002).

5. Toxicity of decolouration products and evaluation methods The metabolites produced from dye degradation are, in many cases, more toxic that the parent dye. For example, the products of the oxidation of indigo blue via electro incineration, coagulation with Al2(SO4)3 or the use of Lac are more toxic than the parent dye (Solis-Oba et al., 2009). Several azo dyes and the amines from their degradation have shown mutagenic responses in Salmonella and mammalian assay systems, and their toxicity depends on the nature and position of the substituents in the molecule. For example, the dyes Acid Red 18 and Acid Red 27 are non-mutagenic, whereas the structurally similar dye Acid Red 26 is carcinogenic because of the presence of a methyl group and the difference in the position of the sodium sulphonate. Similarly, 3methoxy-4-aminoazobenzene is a potent hepato carcinogen in rats and a strong mutagen in bacteria, whereas 2methoxy-4-aminoazobenzene is apparently noncarcinogenic and an extremely weak mutagen in bacteria. Methyl Red is mutagenic in nature, and most microbial degradation studies reveal the formation of N,N-dimethyl-

7

phenylenediamine (DMPD), a toxic and mutagenic aromatic amine (Wong and Yuen, 1998) that remains unchanged in the culture (Ayed et al.,2011). Therefore, in general, it becomes very important for any bioremediation technology to assess the toxicity of the pollutants and metabolites formed after dye degradation in order to study the feasibility of the method.

6. Fungal systems involved in the decolouration of azo dyes Decolouration of azo dyes using yeast Yeast decolorization and degradation of dyes has not been extensively studied. Yeast has mainly been studied with regard to biosorption. Yeast strains have been used in the decolouration of different azo dyes because they have many advantages for application in bioremediation, such as a high capacity to accumulate dyes and heavy metals, such as Pb(II) and Cd(II) (Ertugrul et al.,2008; Fairhead and Thony-Meyer, 2012), fast growth, faster decolouration than filamentous fungi and the ability to survive unfavourable environments (Martorell et al., 2012). Wastewater treatment sludge harbours abundant and diverse yeast strains compared to other environments, although yeasts are still a minor fraction of the microorganisms present in activated sludge (Yang et al., 2011a, b). Compared to bacteria and filamentous fungi, yeast has many advantages; they not only grow rapidly like bacteria, but like filamentous fungi, they also have ability to resist unfavourable environments (Yu and Wen, 2005). More recently, literature review have shown that yeast species acted as a promising dye adsorbent capable to uptake higher dye concentration, such as Galactomyces geotrichum, Saccharomyces cerevisiae, Trichosporon beigelii, etc. (Jadhav et al., 2008). Kluyveromyces marxianus IMB3 was reported to have role in decolorization of Remazol Black-B (Meehan et al., 2000). Trichosporon beigelii NCIM-3326 could decolorize various azo dyes such as Navy blue HER (100 %), Red HE7B (85 %), Golden yellow 4BD (60 %), Green HE4BD (70 %) and Orange HE2R (50 %), among which the decolorization rates of some dyes were not desirable (Saratale et al., 2008). Some studies also show that yeast species act as promising dye adsorbents which are able to uptake higher dye concentration (Safarikova et al., 2005). A number of simple azo dyes were degraded by yeast Candida zeylanoides after 22 h with colour removal of 46e67 % (Martins et al. 1999). Ramalho et al. (2002) reported that Candida zeylanoides could effectively decolorize four model azo dyes but with relatively long time (40e60 h). Similar to microalgae, the mechanisms of decolouration by yeast can involve adsorption (Yu et al., 2005; Aksu and Donmez, 2005; Safarikova et al., 2005; Safarik et al., 2007), enzymatic degradation, or a combination of both (Table 2). Adsorption on yeast biomass is more efficient at low pH. For example, maximal accumulation of Direct Violet 51 in Candida albicans occurs at pH 2.5 (Vitor and Corso, 2008) and that of Violet 3 in Candida tropicalis occurs at pH 4.0 (Charumathi and Nilanjana, 2010). Dissimilar azo dyes are adsorbed to different extents: C. tropicalis adsorbed 94 % of Remazol Blue and 44 % of Reactive Red (Donmez, 2002), whereas Trichosporon

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8

S. K. Sen et al.

Table 2 e Decolouration of azo dyes using yeast. Yeast and source

Decolouration process, conditions and enzyme

(Initial concentration) dye, % colour, COD, TOC removal and toxicity

Candida tropicalis from dye contaminated sludge Candida utilis

Adsorption; pH 3e7, 120 rpm, 28
C, 2d Adsorption; 150 rpm, 25
C, 10 d

Candida tropicalis and Debaryomyces polymorphus from wastewater treatment plant Pichia fermentans from a collection

Adsorption; 140 rpm, 28
C, 66 h

(10 ppm) Basic Violet 3, 85.3 % colour (50 ppm) Remazol Turquoise BlueG, 82 % colour (200 ppm) Reactive Black 5, 95 % colour

Rhodotorula mucilaginosa from textile effluent Candida albicans from industrial effluents Trichosporon akiyoshidainum from rainforest

Adsorption; pH 3e6, 100 rpm, 30
C, 6d Adsorption; pH 2.5, 150 rpm, 35
C, 72 h Adsorption; pH 2, 12 h. Aerobic degradation, pH 7, 250 rpm, 26
C, 16 h; MnP, Tyr

Candida rugopelliculosa from dye contaminated soil Galactomyces geotrichum from a collection

Anaerobic degradation; pH 2e8, 28
C, 48 h Aerobic degradation; pH 7, 120 rpm, 30
C, 24 h; Tyr, NADHDCIP reductase, Lac

Candida sp., Williopsis californica

Aerobic degradation; 250 rpm, 25
C, 24 h; MnP, Tyr

Paraconiothyrium variabile from soil

Aerobic degradation; 40
C, 3 h; Lac

Candida tropicalis from dye contaminated soil

Aerobic degradation; pH 3e9, 120 rpm, 28
C,

Adsorption; pH 3, 120 rpm, 28
C

akiyoshidainum adsorbed 63 % of Reactive Blue and 90 % of Reactive Red 141 (Pajot et al., 2007). Dye degradation is highly associated with the yeast growth process (Yang et al., 2008a,b; Lucas et al., 2006) and its primary metabolism (Martorell et al., 2012). Yeast cells do not grow without glucose or an easily metabolised carbon and energy source. Furthermore, the decolouration process requires a carbon source (Omar, 2008; Waghmode et al., 2011). The presence of azo dyes induces the production of oxidases and reductases, such as MnP, Tyr (Martorell et al., 2012; Pajot et al., 2011; Halaburgi et al., 2011; Aghaie-Khouzani et al., 2012) and NADH-DCIP reductase (Waghmode et al., 2011) by yeast. Increased Lac production reduces decolouration time. For example, the decolouration of the azo and non-azo dyes Sudan Black and Crystal Violet using a culture filtrate of Paraconiothyrium variabile in basal medium was observed to be 9.4 % and 16.8 % respectively, in 12 h. The Lac activity of P. variabile increases from 970 to 16,678 U/L upon the introduction of xylidine and copper to the culture medium and when using

(10 ppm each) Acid Blue 93, 100 % colour; Direct Red 28, 95 % colour; Basic Violet 3, 70 % colour (389 ppm) Remazol Blue, 96 % colour (100 ppm) Direct Violet 51, 73.2 % colour (200 ppm each) Reactive Blue 221, 65 % by adsorption at pH 2; Reactive Blue 221, Reactive Red 141, Reactive Black 5, 100 % (2000 ppm) Reactive Blue 13, 90 % colour (10 ppm each) Mixture of Remazol Red, Golden Yellow HER, Rubine GFL, Scarlet RR, Methyl Red, Brown 3 REL, Brilliant Blue, 88 % colour, 69 % COD, 43 % TOC. Phaseolus mungo and Sorghum vulgare 90 % germination in the presence of degradation products (200 ppm) Reactive Yellow 84, Reactive Black 5, Reactive Blue 221, Reactive Red 141, 96 % colour (200 ppm) Sudan Black, 84 % colour; (600 ppm), Remazol Brilliant Blue R, 93 % colour (50 ppm) Acid Blue 93, 100 % colour; Direct Red 28, 100 % colour; Basic Violet 3, 90 % colour, 30 h

Reference

(Das et al., 2011) (Gonen and Aksu, 2009) (Yang et al., 2008a,b)

(Das et al., 2010)

(Ertugrul et al., 2008) (Vitor and Corso, 2008) (Pajot et al., 2011)

(Liu et al., 2011a, b) (Waghmode et al., 2011)

(Grassi et al., 2011)

(Aghaie-Khouzani et al., 2012) (Charumathi and Nilanjana, 2010)

the filtrate of the optimised culture broth; the decolouration percentages of the mentioned dyes are 84 %, 94 %, 93 % and 87 %, respectively, after 3 h (Aghaie-Khouzani et al., 2012). The decolouration mechanism depends on the pH. T. akiyoshidainum adsorbs 63 % Reactive Blue 221 and 90 % Reactive Red 141 at pH 2.0, but at pH 7.0, an almost complete degradation of these dyes (Pajot et al., 2007) is achieved. Yeast degradation of dyes can be accomplished under aerobic or anaerobic conditions. One study (Waghmode et al., 2011) found that the decolouration and COD and TOC removal of a mixture of Remazol Red, Golden Yellow HER, Rubine GFL, Scarlet RR, Methyl Red, Brown 3 REL and Brilliant Blue using G. geotrichum is higher under aerobic than under anoxic or anaerobic conditions, while Candida rugopelliculosa has been shown to degrade Reactive Blue under static conditions (Liu et al., 2011a,b). Table 2 shows information on yeast decolorization of dyes available in literature. Biosorption of textile dyes has been found to occur by biomass derived from yeast Kluyveromyces marxianus IMB3 (Bustard et al., 1998). K. marxianus IMB3 was also found to

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Fungal decolouration and degradation of azo dyes

9

Table 3 e Decolouration of azo dyes using filamentous fungus. Filamentous fungus and source

Decolouration process, conditions and enzyme

Trametes versicolor from a collection Degradation; pH 4.5, 800 rpm, 25
C, 3h Trametes versicolor from a collection Adsorption; pH 2e7, 150 rpm, 45
C, 2h Trametes versicolor, Ganoderma Adsorption and degradation; pH lucidum & Irpex lacteus from a 4.5, 135 rpm, 25
C, 48 h; Lac collection Immobilized Trametes versicolor, Adsorption and degradation; pH Pleurotus ostreatus & Phanerochaete 3.5e5.2, 120 rpm, 30
C, 4 d; Lac, LiP, chrysosporium MnP Trametes versicolor from a Degradation; pH 3.5e6.5, 200 rpm, timberland 40
C, 7 d

Trametes versicolor

Degradation; pH 6.5, 240 rpm, 26
C, 384 h; Lac Degradation; 2000 ppm, 28
C, 10 d; Lac MnP and MiP, LiP

Immobilized Trametes pubescens Pleurotus ostreatus, from a collection Trametes hirsute, Phanerochaete Degradation; 150 rpm, 35
C, 72 h; chrysosporium, from a henequen by Lac products factory Trametes hirsute, Marasmius sp., Adsorption and degradation; 24
C, 7 d; Lac peroxidise from a collection

Trametes sp., from a compost

Degradation; 10 d; Lac, MnP

Trametes trogii, from a collection

Degradation; pH 4.5 and 7, 30
C, 30 min; Lac, MnP

Phanerochaete chrysosporium Immobilized Phanerochaete chrysosporium, from a collection

Degradation; pH 4.5, 150 rpm, 30
C, 72 h; LiP Degradation; pH 4.4, 25
C, 72 h; LiP, MnP

Immobilized Phanerochaete chrysosporium, from a collection

Degradation; pH 4.4, 150 rpm, 30
C, 120 h; MnP

Phanerochaete chrysosporium, from a Adsorption and degradation; 30
C, collection 24 h; MnP, LiP

(Initial con.) dye, % colour, COD, removal and toxicity

Reference

(100 ppm) Direct Brown 2, 100 % (Cano et al., 2012) colour (500 ppm) Sirius Blue K-CFN, (Erden et al., 2011) 62.62 mg dye/g fungi (150 ppm) Black Dycem, 90 % colour (Baccar et al., 2011) the degradation products not toxic (300 ppm) Reactive Black 5, 98 % colour

(Fernandez et al., 2009)

(50 ppm each) Blue 49, 94 % colour; Black 5, 88 % colour; Reactive Brilliant Blue R, 97 % colour; Orange 12, 83 % colour; Orange 13, 84 % colour (125 ppm) Reactive Blue 4, 90 % colour (2000 ppm) Remazol Brilliant Blue R, 95 % colour; Reactive Blue 49, 97 % colour. Toxicity reduction (0.01 %) Acid Blue, 90 % colour; Reactive Green 19, 95 % colour; Reactive Red 195, 83 % colour (2000 ppm each) Remazol Procion Blue H-EGN 125, 87 % colour; Levafixblue E-RA, 98 % colour; Levafixblue PN-3 R, 95 % colour; Remazol Golden Yellow 3R liquid 25 (4000 _L/L), 80 % colour. Degradation products showed no cellular toxicity (180 ppm each) Orange II, 100 % colour; Brilliant Blue R250, 100 % colour (133 mM) Remazol Brilliant Blue R, 82 % colour; (50 mM) Indigo Carmine, 84.5 % colour; (40 mM) Bromophenol Blue, 75 % colour (20 ppm) Direct Red 80, 100 % colour (100 ppm) Reactive Black 5, 90 % colour Degradation products not toxic (65 ppm) Direct Violet 51, 84 % colour; (120 ppm) Reactive Black 5, 90 % colour; (100 ppm) Ponceau Xylidine, 85 % colour; Bismark Brown 86.7 % colour (24 ppm) Acid Red 88, 99 % colour; Reactive 5, 100 %; Reactive Orange 16, 100 % colour; Acid Red 114, 90 % colour at 5 d; Direct Violet 55, 90 % colour at 5 d (50 ppm) Direct Red 80, 100 % colour (1600 ppm) Astrazon Red FBL, 87 % colour, 42 % COD (100 ppm) Orange II, 85 % colour

(Pilatin and Kunduhoglu, 2011)

Phanerochaete chrysosporium from a collection Phanerochaete chrysosporium

Degradation; 180 rpm, 39
C, 24 h; LiP Degradation; 37
C, 2 d

Phanerochaete chrysosporium, from a collection Aspergillus niger, from dye contaminated soil Aspergillus flavus, Alternaria sp. Penicillium sp., from dye contaminated sludge

Degradation; pH 4e7, 24e34
C, 7 d; MnP (10 ppm) Congo Red, 99 % colour Degradation; pH 3,11, 60
C, 36 h

(Yemendzhiev et al., 2009) (Casieri et al., 2008)

(Tapia-Tussell et al., 2011)

(Jadhav et al., 2010)

(Grinhut et al., 2011)

(Grassi et al., 2011)

(Sen et al., 2012) (Enayatizamir et al., 2011)

(Enayatizamir et al., 2010))

(Ghasemi et al.,2010)

(Sing et al., 2010) (Sedighi et al., 2009) (Sharma et al., 2009) (Karthikeyan et al., 2010)

Adsorption and degradation; 30
C, (20 ppm) Acid Red 151, 98 % colour; (Ali et al., 2010) 8d Orange II, 58 % colour (continued on next page)

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S. K. Sen et al.

Table 3 (continued) Filamentous fungus and source Aspergillus niger, Alternaria alternata, Chaetomium globosum, Curvularia lunata, Trichoderma virule, Drechslera rostrata, Fusarium oxysporum, Humicola fuscoatra, Leptostroma actaea, Mucor mucedo, Penicillium nutalum, Torula herbarum, Dactylosporium macropus, From dye contaminated soil Aspergillus niger, from a wastewater Treatment plant Coriolus versicolor, from a collection Trichoderma sp., from a collection Cunninghamella elegans, from mangrove sediments Bjerkandera adusta, from a collection

Ganoderma sp., from the forest

Fusarium oxysporum, from a collection Armillaria sp., wood from tropical rain forest Datronia sp., from a collection

Pleurotus sajor-caju, from a collection

Decolouration process, conditions and enzyme

(Initial con.) dye, % colour, COD, removal and toxicity

Reference

Degradation; room temperature, 21 (1 %) Scarlet Red 80 % colour; Fast d; K-amylase, protease and Greenish Blue90 % colour; Brilliant catalase Violet 89 % colour

(Laxminarayan et al., 2010)

(20 ppm) Acid Red 151, 98 % colour; Orange II, 84 % colour (33e100 ppm) Acid Orange II, 85 % colour Adsorption; 24 h (100 ppm) Acid Brilliant Red B, 100 % colour Adsorption; pH 5.6, 28
C, 120 h Reactive Orange II, Reactive Black 5, Reactive Red 198, 93 % colour Adsorption and degradation; pH (1250 ppm each) Mixture of 10,130 rpm, 28
C Reactive Yellow 145, Reactive Red 195, Reactive Blue 222, Reactive Black 5, 91 % colour. Degradation products not toxic to Pseudokirchneriella subcapitata and Cucumis. sativus Degradation; pH 5.5, 150 rpm, 28
C, (50 ppm) Methyl Orange, 96.7 % 72 h; Lac colour; Crystal Violet, 75 % colour; Bromophenol Blue, 90 % colour; (200 ppm) Malachite Green, 91 % colour Degradation; 160 rpm, 24
C, 144 h (100 ppm) Yellow GAD, 100 % colour Degradation; pH 4, 120 rpm, 40
C, (100 ppm) Reactive Black 5, 65 % 96 h colour; Remazol Brilliant Blue R, 70 % colour Adsorption and degradation; pH (1000 ppm) Reactive Blue 19, 95 % 3e9, 150 rpm, 30
C; Lac, MnP colour at 20 h; Reactive Black 5, 90 %, colour at 70 h Adsorption and degradation; 11 d; (50 ppm) Reactive Blue 220, 100 % Lac, MnP colour; Reactive Red 198,100 % colour; Reactive Yellow 15, 100 % colour

(Ali et al., 2009)

Adsorption and degradation; 100 rpm, 30
C, 24 h Degradation; 30
C

decolorize Remazol Black-B through physical adsorption (Meehan et al., 2000). The oxidative yeasts Rhodotorula sp. and Rhodotorula rubra were found to degrade crystal violet completely in four d (Kwasniewska, 1985). An yeast strain Candida zeylanoides was found to be able to degrade a number of azo dyes, whose reduction product include metanilic acid for azo dyes I and III, and sulfanilic acid for azo dyes II and IV. S. cerevisiae has been found to be effective in removing dye in molasses media (Aksu, 2003).

Decolouration of azo dyes using filamentous fungi Filamentous fungi are ubiquitous in the environment, and the fast adaptation of their metabolism to several carbon and nitrogen sources is important for their survival. The application of filamentous fungi in the decolouration process is an attractive alternative due to low cost and the possibility of total

(Hai et al., 2012) (Xin et al., 2012) (Ambrosio et al., 2012) (Anastasi et al., 2011)

(Porri et al., 2011) (Hadibarata et al., 2012)

(Vaithanomst et al., 2010)

(Munari et al., 2008)

mineralisation of the dye (Husain and Husain, 2007; Asgher et al., 2008a,b). Phanerochaete is the most widely studied genus, and others that are frequently applied are Trametes, Bjerkandera, Aspergillus, Pleurotus and Phlebia (Table 3). Decolouration can be accomplished by adsorption or enzymatic degradation. Inactivated mycelia of Cunninghamella elegans (Ambrosio et al., 2012), T. versicolor lyophilized biomass (Erden et al., 2011), T. versicolor (Baccar et al., 2011) or Fusarium solani non-viable cells (Abedin, 2008) are more efficient than materials such as activated carbon or amberlite. Adsorption is enhanced at pH 2e3 (Erden et al., 2011; Renganathan et al., 2006; Bakshi et al., 2006; Iqbal and Saeed, 2007; Pajot et al., 2011; Maurya et al., 2006), which is probably due to electrostatic attractions between charged dye molecules and the charged cell surface (Erden et al., 2011; Kaushik and Malik, 2009). The adsorption capacity of the fungal biomass increases with temperature as a result of the increase in surface activity

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Fungal decolouration and degradation of azo dyes

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Table 4 e Results of dye decolorization by live fungi. Culture Rhizopus oryzae

Dye Rhodamine B (xanthine dye)

Percent Experimental conditions removal 90

Schizophyllum commune Acid Orange 7 Acid Red 18 Reactive Black 5 Penicillium oxalicum Reactive Blue 19

44. 23a 127.53a 180.17a 91

Isolate Aspergillus 1

Direct Yellow Direct Brown Maxilon Red Erio Red Aspergillus ochrocous 5 Polar Red

72 85 49 21 96

Aspergillus niger 31

Polar Red

94

Umbelopsis isabellina

Reactive Black 5

>99

Penicillium geastrivorus Reactive Black 5

>99

Funalia trogii

92e98

Astrazon Red

Funalia trogii

>99

Reactive Blue 19 Reactive Blue 49 Acid Violet 43 Reactive Black 5 Reactive Orange 16 Acid Black 52 Aspergillus sp. Reactive Blue Reactive Black Trichoderma harzianum Erioglaucine

99 75 76e88

Pleurotus florida

Blue CA

93.54

Trametes hirusta Myrothecium sp.

Remazol Brilliant Blue

92.17 90 R

Aspergillus niger

Basic Blue 9

Aspergillus niger

Acid Blue 29

P. chrysosporium

Reactofix Gold Yellow

P. chrysosporium

Reactive Red 22

Pleurotus pulmonaris

Congo Red

Irpex lacteus

Methyl Red

C. versicolor

Indigo Carmine

Mechanism

Time of contact

Reference

Initial dye concentration 100 mg/L; pH 7.0; biomass dose 0.25 g/25 mL; temperature 40
C; point of zero charge pH 3.5 Initial dye concentration 100 mg/L; pH 2.0; temperature 30
C Initial concentration 100 mg/L; pH 2.0; biosorbent dose 0.25 g/100 mL Initial dye concentration range 1e100 mg/L

Chemical interaction, ionic 5 h interaction, physical forces

Das et al., 2006

Bioaccumulation

72 h

Renganathan et al., 2006

Biosorption

80 min

Zhang et al., 2003

Biosorption

24 h

Abd El-Rahim et al., 2003

Initial dye concentration 0.3 g/L Initial dye concentration 0.3 g/L Initial dye concentration 100 mg/L Initial dye concentration 100 mg/L Initial dye concentration range 0e1500 mg/L; initial pH 6e11; temperature 30
C Inoculum contain 80 mL of Kirk’s basal salts and 100 mg/L of dye; pH 4.5

Biosorption

8d

Biosorption

8d

MnP

48 h

Abd El-Rahim and Moawad, 2003 Abd El-Rahim and Moawad, 2003 Yang et al., 2003

Biosorption

48 h

Yang et al., 2003

Adsorption and microbial metabolism

24 h

Yesilada et al., 2002

Laccase, MnP

10 d

Park et al., 2007

Initial dye concentration Biodegradation 100 mg/L; pH 3.0 Initial dye concentration range 10e50 mg/L; biosorbent dose 1.5 g/50 mL; pH 4.0 Dye concentration 200 mg/L Laccase

Nutrient salt media; initial dye concentration 80 mg/L; pH 7.0; potato dextrose broth 10 Initial dye concentration 50 mg/L; initial pH 5.1; biosorbent dose 0.2 g/75 mL 80 Initial dye concentration 50 mg/L; initial pH 7.6; biosorbent dose 0.2 g/75 mL 73 Cell mass concentration 1.4 mg/mL; pH of growth medium 4.5; incubation at 39
C 92e100 Initial dye concentration 120e140 mg/L 93 Initial dye concentration 200 ppm; glucose ammonium tartratecorncob solid state cultures 56 Initial dye concentration 150 mg/g 98 Initial concentration 23 mg/L

24 h

Mohandass et al., 2007 105 min Sadhasivam et al., 2007

10 d

Sathiya Moorthi et al., 2007 Zhang et al., 2007

Biosorption and biodegradation

7d

Biosorption

2d

Fu and Viraraghavan, 2000

Biosorption

30 h

Fu and Viraraghavan, 2001

Lignin-degrading system and adsorption

3d

Capalash and Sharma, 1992

Lignin degrading system

30 h

Wu et al., 1996

Biodegradation, adsorption 6 d and laccase

Tychanowicz et al., 2004

Lignin degrading system

Novotny et al., 2001

14 d

Biodegradation and laccase 1 h

Levin et al., 2004 (continued on next page)

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S. K. Sen et al.

Table 4 (continued) Culture

Dye

Percent Experimental conditions removal

Trametes trogii

Anthraquinone

88

Geotrichum sp.

>99

P. chrysosporium

Reactive Black 5 Reactive Red 158 Reactive Yellow 27 Orange G Remazol Brilliant Blue R Direct dyes

Trametes versicolor

Direct Blue 1

63.2

Daedalea flavida

Coracryl Coracryl Coracryl Coracryl Reactive Reactive Reactive Rathidol Coracryl Coracryl Coracryl Coracryl Reactive Reactive Rathidol Coracryl Coracryl Coracryl Coracryl Reactive Reactive Reactive Rathidol Coracryl Coracryl Coracryl Reactive Reactive

13.9 52.9 26 35.2 11.7 4.8 39.8 39

Dichomitus squalens P. ostreatus

Dichomitus squalens

Irpex flavus

Polyporus sanguineus

Black Pink violet Red Yellow Orange Red Scarlet Black Pink violet Red Yellow Red Scarlet Black Pink violet Red Yellow Orange Red Scarlet Black Pink Red Orange Red

Glucose-asparagine and malt Blue extract/glucose medium Initial dye concentration 100 mg/L

Mechanism

4h

Levin et al., 2001

Biodegradation, MnP and laccase

10 d 20 d 20 d 14 d 9d

Maximo et al., 2003

Initial concentration 50 mMA Laccase and MnP Initial concentration 50 mM Laccase

100

Initial concentration 120 mg/L Initial concentration 800 mg/L; pH 6.0; biosorbent dose 250 mg/50 mL Initial concentration 30 mg/ L; fungal culture grown for 6 d

Biodegradation, adsorption and MnP Biosorption

15 d

Cell free enzyme based decolorization

3 3 3 3 3 3 3 3 1 2 1 1 2 1 3 2 3 5 2 1 2 1 2 5

Initial concentration 30 mg/ Cell free enzyme based L; fungal culture grown for 6 decolorization d

Initial concentration 30 mg/ Cell free enzyme based L; fungal culture grown for 6 decolorization d

Initial concentration 30 mg/ Cell free enzyme based L; fungal culture grown for 6 decolorization d

and kinetic energy of the dye (Bakshi et al., 2006; Kaushik et al., 2009). However, decolouration is decreased at very high temperatures, which is possibly due to the deactivation of the adsorbent surface or the destruction of some active sites (Erden et al., 2011; Iqbal and Saeed, 2007). Dye adsorption is dependent on dye concentration; at higher concentrations, adsorption is diminished. For example, the percentage of Acid Red 18 and Reactive Black 5 adsorbed by Schizophyllum commune diminished from 90 % to 27 % and from 92 % to 40 %, respectively, when the initial dye concentration was changed from 10 to 100 ppm (Renganathan et al.,2006). Filamentous fungi oxidise azo dyes via peroxidases and phenoloxidases (Majeau et al., 2010; Duran and Esposito, 2000; Baldrian, 2006; Pazarlioglu et al., 2005; Svobodova et al., 2007; Erkurt et al., 2007; Husain and Husain, 2011), avoiding the amine generation problem present during azo dye

Reference

Laccase

95 100

69 100 76 45 79.9 76 79 67.3 100 58 50.2 79 63.2 77 73.9 67 64.9 36.5 19.3 59.2

Time of contact

6h

h h h h h h h h h h h h h h h h h h h h h h h h

Eichlevora et al., 2005 Palmieri et al., 2005 Pazarlioglu et al., 2005 Bayramoglu and Arica, 2007 Chander and Arora 2007

Chander and Arora, 2007

Chander and Arora, 2007

Chander and Arora, 2007

reduction. The growth of filamentous fungi, enzyme production and subsequent dye degradation are affected by culture conditions, nutrient conditions, especially regarding N limitation, agitation, time, pH, temperature, carbon source, oxygen supply, additives and salts (Ayed et al., 2011; Gallizia et al., 2004; Asgher et al., 2008a,b; Khlifi et al., 2010; Parshetti et al., 2010; Karthikeyan et al., 2010; Mielgo et al., 2001; ZouariMechichi et al., 2006; Grinhut et al., 2011). Decolouration is more efficient under aerobic than anaerobic conditions. Higher shaking speed improves the process due to more efficient oxygen transfer, as in the decolouration of Blue 49, Orange 12, Orange 13, Black 5 and Remazol Brilliant Blue R by T. versicolor (Pilatin and Kunduhoglu, 2011); Drimaren Brilliant Blue by Trametes villosa and Pleurotus sanguineus (Machado et al., 2006); Congo Red by A. niger (Karthikeyan et al., 2010); and Acid Red 151 and Orange II by Aspergillus flavus, Alternaria

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Fungal decolouration and degradation of azo dyes

13

Table 5 e Results of dye decolorization by dead fungi. Culture

Dye

Percent removal

Aspergillus niger

Acid Blue

29 99

Aspergillus niger (immobilized)

Aspergillus niger

Acid Blue 29 Basic Blue 9 Congo Red Disperse Red 1 Congo Red

64.7a 8.3a 1.1a 0.1a 89.6

Aspergillus niger

Synazol

88

Aspergillus foetidus

Reactive Black 5

>99

P. chrysosporium

Astrazone Blue FGRL Cibracron Red Astrazone Blue FGRL Cibracron Red Direct Red 80 Reactive Blue 214 Reactive Blue 19 Direct Red 80 Reactive Blue 214 Reactive Blue 19 Direct Red 80 Reactive Blue 214 Reactive Blue 19 Reactive Orange 16 Reactive Red 4 Reactive Blue 19 Reactive Black 5

60 51

Funalia trogii

Cunninghamella elegans

Rhizomucor pusillius

Rhizopus stolonifer

Rhizopus arrhizus

Rhizopus arrhizus

48 38 100 99 57e63 100 >98 84e98 100 98 99 190a 150a 90a 62.5

Rhizopus arrhizus

Germazol Torquoise Blue-G

47.5

Rhizopus arrhizus

Gryfalan Black RL

59

Trametes versicolor Aspergillus niger Rhizopus nigricans

Reactive Green Reactive Blue

37 37.5 86 83

Penicillium chrysogenum

Acid Orange 8 Reactive Orange 16

70.4 67.6

Rhizopus stolonifer

Bromophenol Blue

88

Neurospora crassa

Acid Red 57

98.78

Trametes versicolor

Direct Blue 1

95.2

Agaricus bisporus þ Thuja orientalis (mixed)

Reactive Blue 49

72.86a

Experimental conditions

Mechanism

Initial dye concentration 50 mg/L; Biosorption initial pH 7.6; biosorbent dose 0.2 g/ 75 mL 4.5 g of beads; column dia 1.27 cm, height 40 cm; flow rate 6 mL/min

Initial dye concentration 50 mg/L; initial pH 6.5; biosorbent dose 0.2 g/ 75 mL Synazol Red 0.22%, Synazol Yellow 0.1% Initial dye concentration 100 mg/L; pH 2e3; biosorbent dose 0.2 g/ 100 mL Initial dye concentration 50 mg/L; biosorbent 0.2 g/50 mL; without adjusting pH Initial dye concentration 50 mg/L; biosorbent 0.2 g/50 mL; without adjusting pH Initial dye concentration 1000 ppm, 5000 ppm; biosorbent 3 g/30 mL

Biosorption

Time of contact 24 h

Reference Fu and Viraraghavan, 2001

5.0 (min) 5.2 (min)

42 h

Fu and Viraraghavan, 2002

18 h

Khalaf, 2008

Biosorption

2h

Adsorption

2h

Asma et al., 2006

Adsorption

2h

Asma et al., 2006

Biosorption

24 h

Prigione et al., 2008

Initial dye concentration 1000 ppm, Biosorption 5000 ppm; biosorbent 3 g/30 mL

24 h

Prigione et al., 2008

Initial dye concentration 1000 ppm, Biosorption 5000 ppm; biosorbent 3 g/30 mL

24 h

Prigione et al., 2008

Initial concentration 0e500 mg/L; pH 2.0; biosorbent dose 1 g/L

20 h

O’Mahony et al., 2002

24 h

Aksu and Tezer, 2000

Initial dye concentration 800 mg/L; biosorbent dose 1 g/L; pH 2.0 Initial dye concentration of 812.6 mg/L; pH 2.0; biomass dosage of 0.5 g/L; temperature 45
C Initial concentration range 1000 mg/L; biosorbent dose 1.0 g/L; pH 2.0; temperature 25
C (pH 1.0 and temperature 35
C for A. niger)

Initial dye concentration 50 mg/L; biomass loading:1 g% (w/v); pH 6; temperature: 29  1
C Polyethylenimine modified; initial concentration 500 mg/L; biosorbent dose 1 g/L; initial pH 6.8e7.2 Initial concentration 800 mg/L; pH 2.0; biosorbent dose 1 g/L Initial concentration range 100e400 mg/dm3; pH 1.0; temperature 20 _C; biosorbent dose 2 g/dm3 Initial concentration 800 mg/L; pH (Direct Blue 1) 6.0; biosorbent dose 250 mg/50 mL Initial concentration 150 mg/L; pH 1.0; point of zero charge 1.5; biosorbent dosage 3.0 g/L

Adsorption

Physical adsorption 24 h

Aksu and Cagatay, 2006

Adsorption and internal diffusion

400 min Aksu and Karabayur, 2008

Adsorption

1h

Kumari and Abraham, 2007

Sorption

14 h

Low et al., 2008

Biosorption

20 h

Zeroual et al., 2006

Biosorption

40 min

Akar et al., 2006

Biosorption

6h

Bayramoglu and Arica, 2007

Biosorption

90 min

Akar et al., 2009a

(continued on next page)

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14

S. K. Sen et al.

Table 5 (continued) Culture

Dye

Percent removal

Thuja orientalis

Acid Blue 40

48.5

Agaricus bisporus

Acid Red 44

59.80a

Aspergillus fumigatus Reactive Brilliant (carboxymethylcellulose Red K-2BP immobilized beads) Aspergillus niger Direct Blue 199

94.7

44.9

Cephalosporium aphidicola Acid Red 57

29.2

Experimental conditions

Mechanism

Initial concentration 200 mg/dm3; Biosorption pH 1.0; temperature 20
C; biosorbent dose 1 g/dm3 Initial concentration range Biosorption 50e300 mg/L; pH 2.0; point of zero charge w2.0; biosorbent dose 3 g/L; temperature 40
C Initial concentration 33.9 mg/L; pH Chemisorption 2.0; biosorbent dose 0.05 g/25 mL Initial concentration 400 mg/L; pH Biosorption 3.0; biosorbent dose 6 g/L, temperature 45
C Initial concentration 150 mg/dm3; Chemical pH 1.0; biosorbent dose 0.4 g/dm3; ion-exchange temperature 20
C

sp. and Penicillium sp. (Ali et al., 2009,2010). Decolouration can be increased if the pH is kept constant during the process at a value that is favourable for fungal growth (Arroyo-Figueroa et al., 2011). Carbon and nitrogen sources have important effects on decolouration, as seen with P. chrysosporium (Ghasemi et al., 2010; Urek and Pazarlioglu, 2005; Karimi et al., 2006) and Bjerkandera sp. (Axelsson et al., 2006), where optimal decolouration is reached in nitrogen-limited culture medium. The degradation of Orange II and Brilliant Blue R 250 by Trametes sp. is improved with a C/N ratio of 53 and is negligible with a C/N ratio of 10 (Grinhut et al., 2011). The addition of additives to the culture medium increases the efficiency of the decolouration process. MnP production from P. chrysosporium is enhanced in the presence of succinate (Ghasemi et al., 2010), veratryl alcohol raises LiP production and, therefore, the decolouration of Reactive Black 5 (Enayatizamir et al., 2011) and Tween 80 and Mn2þ elevate the MnP activity of P. chrysosporium (Urek and Pazarlioglu, 2005). The presence of veratryl alcohol induces enzymatic activity in immobilised P. chrysosporium depending on the support. Reactive Blue 220 can act as a redox mediator for the enzymatic reactions involved in the decolouration process in Pleurotus sajorcaju to enable azo dye degradation (Munari et al., 2008). The immobilization of fungal biomass is advantageous when the effluent has toxic substances and cellular growth is difficult, does not require a continuous supply of nutrients and can be regenerated and reused over many cycles. Fungi have been immobilised using different materials. P. chrysosporium has been immobilised on ZrOCl2-activated pumice (Pazarlioglu et al., 2005), polystyrene foam (Urek and Pazarlioglu, 2005), nylon sponges and sunflower seed shells (Enayatizamir et al., 2011), polyurethane foam and Luffa sponges (Iqbal and Saeed, 2007; Grinhut et al., 2011), Caalginate beads (Enayatizamir et al., 2010), lignitic xylite and lignite granules (Bohmer et al., 2010), and in a packed-bed bioreactor with Kissiris as the carrier (Karimi et al., 2006; Sedighi et al., 2009). T. versicolor has been immobilised on polyurethane foam (Erden et al., 2011) and Luffa sponges (Fernandez et al.,

Time of contact 90 min

Reference Akar et al., 2008

120 min Akar et al., 2009b

120 h

Wang et al., 2008

4h

Xiong et al., 2010

120 min Kiran et al., 2006

2009) and Trametes pubescens, Pleurotus ostreatus (Casieri et al., ar et al., 2006) have been immobi2008) and Irpex lacteus (Tavc lised on polyurethane foam. Another way to test the fungal biomass is via lyophilisation, as in the case of lyophilised mycelia of T. versicolor, which was shown to decolourate more than 85 % of 1000 ppm of indigo carmine (Cano et al., 2011). Information on the use of living and dead fungi to decolorize dyes is presented in Tables 4 and 5, respectively.

Decolouration of azo dyes using genetically modified microorganisms or enzymes Bioremediation is an environmentally friendly methodology for the treatment of textile wastewater, but the physicochemical characteristics of the effluents, including pH, the content of NaCl and other salts, temperature, and the presence of organic compounds, can result in the deactivation of enzymes and fungal cells. Therefore, it is necessary to have more active and versatile enzymes and fungi with high stability, high production and low cost that are suitable to meet the requirements of textile industry wastewater treatment. There are molecular biology methodologies, like cloning, heterologous expression, random mutagenesis, site directed mutagenesis, gene recombination techniques, directed evolution, rational design and metagenomics, to accelerate the evolution processes in such a way that the bioremediation process is enhanced. Additionally, advances in molecular genetics and genetic engineering have made it possible to clone and express virtually any gene in a suitable microbial host. The process can be iterative for the complete biodegradation of pollutants; the selected or screened enzymes can be subjected to further rounds of random mutagenesis or gene recombination to produce enzymes with different characteristics or biochemical pathway variants in a microorganism (Anga et al., 2005; Gopinath et al., 2009; Demarche et al., 2011; Zeyaullah et al., 2009; Desai et al., 2010). For example, the Lac gene lac48424-1 from white-rot fungi Trametes sp. 48424 was cloned in Pichia pastoris, and the purified recombinant Lac (rLAC48424-1) possesses a stronger capacity for de-

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Fungal decolouration and degradation of azo dyes

colourising different dyes, such as Methyl Orange and Bromophenol Blue, compared with some other known laccases (Fan et al., 2011). The expression of the Lac gene lcc1 from Trametes trogii in P. pastoris results in a higher production of Lac and improved decolouration of azo dyes in the presence of redox mediators (Colao et al., 2006). A Lac gene from Ganoderma lucidum was synthesised using optimised codons and a PCR-based two-step DNA synthesis method; the resulting recombinant Lac was over-expressed in P. pastoris, and showed high levels of Methyl Orange degradation (Sun et al., 2012). The purified recombinant Lac obtained from the heterologous production of Pleurotus sanguineus Lac in P. pastoris could efficiently decolourise synthetic dyes in the absence of mediators (Lu et al., 2009). Among 2300 randomly mutated variants of P. ostreatus POXA1b Lac, two mutants showed higher stability in a variety of environmental conditions and higher ability to decolourise azo dyes than the wild-type Lac; the mutant 2L4A also proved to be highly stable at both acidic and alkaline pHs (Miele et al., 2010). In contrast, fungal Lacs show optimal dye decolouration at acidic pHs and in the presence of redox mediators (Pereira et al., 2009; Martins et al.,2002).

Decolouration of azo dyes using consortia Use of multiple species consortia has proved advantageous for higher decolourization of azo dyes and more stability against environmental fluctuations; however, the investigations in this direction are very scanty. A fungal consortium-SR consisting of Trametes sp. SQ01 and Chaetomium sp. R01 was developed for decolorizing of triphenylmethane dyes, which were decolorized by individual fungi with low efficiencies. Consortium had a decolorization rate of 63e96 %, much higher than that of the monoculture of strain SQ01 (38e72 %) (Yang et al., 2011a, b). Pan et al. (2009) compared the growth and dye removal individually by two fungi (Penicillium sp. A1 and Fusarium sp. A19) with that obtained through their consortium and it was observed that higher efficiency in consortium than in pure culture.

Decolouration of azo dyes using advanced oxidation processes (AOPs) combined with fungal processes Advanced oxidation processes (AOPs) is a highly cost-effective process that has emerged as an important alternative for the elimination of several hazardous organic compounds from contaminated sludge and wastewater. In these processes, hydroxide radicals are produced to oxidise organic pollutants either completely into carbon dioxide, water and inorganic salts or incompletely into less hazardous intermediates (Rosales et al., 2012b). The traditional Fenton process, one of the AOPs, is widely used as a suitable treatment method for highly concentrated wastewaters. The reagents (Fe2þ and H2O2) are relatively inexpensive and environmentally benign; however, H2O2 could be readily decomposed to water and oxygen.

7.

Conclusions

Fungal assisted processes are important techniques for the transformation/degradation and decolorization of azo dyes. In this review article, we have reviewed the various

15

biochemical methods which are used for dye degradation. These include the fungal degradation methods using pure enzymes or biosorption. The various reports that shed light on the importance of these tools for handling the transformation of dyes to smaller, environmentally friendlier molecules have been reviewed and cited. Fungal enzymes and biosorbents capable of decolorizing azo dyes present in wastewater have been reviewed and proved to be an excellent candidate for removal of azo dyes within broad pH, temperature and aeration range. The findings indicate that degradation/decolouration rates can be influenced by operational parameters such as nutrients, pH, temperature, oxygen, dye structure and concentration of organic dyestuff besides the presence of mediators and other additives. Additionally, many of the cited studies focus on the optimization of the above mentioned conditions which vary from case to case. It is therefore clear that systematic and careful optimization studies as well as metabolite toxicity testing must be carried out for each system (dye and microbe (or enzyme) or mediator), as to-date it is not very clear how and why certain dyes are degraded while others are not. The presence of variety of functional groups in the biosorbents makes them selective and highly capable of biodegrading and biosorbing dyes from wastewater. The extensive research conducted on various biosorbents show that they are emerging as a promising alternative to conventional treatment systems. Fungal biomasses have shown excellent colour removal capabilities. This review focuses the biochemically assisted transformation methods which are presently being explored to tackle the problem of removing these organic pollutants from aqueous solutions. This review also demonstrates the potential application of an MFC system using a fungus as a cost-effective alternative to generate electricity and to treat dye wastewater. However, use of fungi to remove colour in a dye wastewater is still in the research stage. Efforts are needed to commercialize this research through (a) selection of suitable fungal biosorbents and enzymes based on economic and market analysis, (b) pilot-scale studies with actual wastewaters and (c) full-scale demonstration systems.

references

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Fungal decolouration and degradation of azo dyes

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