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Involvement of Lignin-Modifying Enzymes in the Degradation of Herbicides

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Jaqueline da Silva Coelho-Moreira, Giselle Maria Maciel, Rafael Castoldi, Simone da Silva Mariano, Fabíola Dorneles Inácio, Adelar Bracht and Rosane Marina Peralta

Submitted: October 23rd, 2012 Published: May 29th, 2013

DOI: 10.5772/55848

From the Edited Volume


Edited by Andrew J. Price and Jessica A. Kelton

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1. Introduction

The high demand for food due to the increase in the world population has led to an increasing use of plant protection products, also known as pesticides, in order to improve productivity. However, along with the success in food production, the accumulation of these persistent chemicals in soil and water is harmful to the environmental and human health [1]. In recent years, agricultural pesticide application has increased all over the world. A total of 5,197 mil lbs of pesticides were used worldwide in 2007 [2]. There are many different types of pesticides; each is meant to be effective against specific pests. The term "-cide" comes from the Latin word "to kill." Among them, herbicides account for the largest market (around 40%) share, followed by insecticides (17%) and fungicides (10%). Herbicides are chemicals used to kill undesired plants, such as weeds, and they are used extensively in home gardens and in agriculture. Due to the relative predominance of herbicides, the present review will focus on this class of compounds.

Herbicides have well defined pros and cons associated with their use. Their use tends to increase yields, and thus makes a significant difference in food production, particularly in countries that struggle periodically against famines. On the other hand, they can cause water pollution when erosion and/or rainwater carry the chemicals off the farms together with the eroded soils after each rainfall. Herbicides vary in their potential to persist in the soil. They are chemically heterogeneous and their structure is one of the main features that determine persistence. For example, some substitutions on aromatic rings (-F, -Cl, -NO2, -NH2, -CF3, -SO3H) [1] in picloram, atrazine, flazasulfuron and other herbicides (Table 1), have a marked influence on persistence. Furthermore, water insoluble molecules are less susceptible to microbial attack because they may sorb tightly to particles and thus stay less available [3].

The herbicide persistence in soil is also affected by the properties of soil such as organic matter content, moisture, pH, microbial activity, among other particularities. Generally, soils with high contents in organic matter have low loss of chemicals through volatilization as a consequence of a stimulated adsorption to soil particles [3]. On the other hand, herbicides that sorb tightly to soil particles have low mobility which decreases the possibility of groundwater contamination. The organic matter also serves as an energy source for microorganisms that degrade pesticides. Furthermore, the soil pH may affect the adsorption processes, changing the mobility and bioavailability of the herbicide [1].

The presence of native microorganisms (fungi, bacteria, protozoans, etc) can greatly influence the toxicity and persistence of herbicides. The degradation process by native microorganisms produces less toxic products and it is probably the most important pathway for the breakdown of herbicides. However, microorganisms require special environmental conditions such as adequate temperature, oxygen and nutrient supply for growth and herbicide degradation [1].

Table 1.

Chemical properties of the most common herbicides

* Water solubility at 25oC. ** Soil average persistence at recommended rates [4]


2. General bioremediation processes

The impact of hazardous herbicide residues on the environment has led to the necessity of finding feasible technologies to remediate these sites. The conventional remediation methods use physical and chemical processes, such as incineration, adsorption on resins and UV irradiation. These methods generally generate excellent contaminant removal; however, from an ecological viewpoint, they are not friendly since they produce unwanted by-products and hazardous residues, besides the danger of human exposure to contaminants.

An innovative technology for complementing or substituting the conventional methods and which presents the same or even improved efficiency is bioremediation. By definition, bioremediation is the use of biological processes to clean up polluted sites. Such biological methods have the potential of being less expensive and more eco-friendly than the physical and chemical treatments [5]. Bioremediation is based on biological systems such as living organisms (bacteria, fungi and plant) and enzymes. They are effective systems to treat a polluted site because they are able to modify the chemical structure of the contaminant into less hazardous end-products [5]. Among these possibilities, microorganisms have become the most attractive option in bioremediation strategies [1].

The naturally occurring bacteria, fungi or plants in a contaminated site are commonly used in bioremediation but microorganisms may be isolated elsewhere and inoculated into the site with the goal of accelerating the remediation. This process is called bioaugmentation [6]. Other terms are commonly used in bioremediation studies. When a chemical compound is broken down into water and CO2, the biodegradation is completed and the process is called mineralization. On the other hand, the term biotransformation usually refers to the process whereby a molecule is converted into a different molecule which is not always less toxic than the precursor. Another term usually used is bioaccumulation or biosorption. This process does not involve any transformation of the molecule and refers to the concentrative transfer of substances into living or dead biomass. The microorganisms can still transform the molecule but are unable to grow on it, in other words, the compound is not used as an energy source [7].

Bioremediation can be developed by two main methods: in situ and ex situ. In situ methods treat the materials in place, while ex situ methods are based on excavation or removal of the polluted materials for processing in another place, such as in bioreactors and biopiles, where optimal environmental conditions for rapid biodegradation may be provided. The in situ bioremediation is more advantageous because it avoids the transport of hazardous materials and is more cost-effective. On the other hand, this process can be limited in some cases since the remediation is not effective in low-permeability soils [1].


3. Herbicide bioremediation studies

Bacteria and fungi are the main microorganisms capable of degrading herbicides. Most bioremediation studies have been focused on bacteria. Among those, several species of Pseudomonas have been found to degrade a wide range of herbicide groups [8-10]. Generally, the bacteria strains utilize the herbicide as the sole carbon and/or nitrogen source. In order to metabolize pollutants, bacteria must take the molecule inside the cell where the enzymes are located [11].

Although bacteria are the main agents used in most commercial bioremediation processes, filamentous fungi have received much attention in recent years, especially white rot fungi (WRF) or ligninolytic fungi [12]. Most WRF belong to the Basidiomycete group and comprise the organisms that are capable of degrading lignin, a polymer found in wood [13]. These fungi have been extensively investigated since the mid-1980s as bioremediation agents because of some particular characteristics. First, as filamentous fungi, WRF present extensive branching and mycelial growth, hence they are able to spread out in the environment reaching the pollutants more efficiently than bacteria. Second, they possess non-specific and radical-based enzymatic systems for degrading lignin. These oxidative enzymes allow them to degrade a broad range of pollutants, including low-solubility compounds, without the necessity of incorporating them into the cells. Third, WRF prefers to grow on substrates such as agricultural crop wastes (straw, corn cobs, sawdust, peanut shells) which are inexpensive and may be used as additional nutrient sources [5,14]. In this context, the primary focus of this review is on the main white-rot species which have been studied for herbicide transformation.


4. White-rot fungi and their lignin-modifying enzymes

White-rot fungi are known as the most efficient lignin degraders. Lignin is a polymer found in wood and vascular tissues [15]. In the cell wall, lignin is covalently associated with carbohydrate components (cellulose and hemicelluloses) forming a matrix that protects the cell wall against microbial attack [16]. Unlike most natural polymers, lignin is irregular and non-repeating. Its biosynthesis is the result of oxidative polymerization of several phenylpropanoid precursors, such as coniferyl alcohol, sinapyl alcohol and p-coumaryl alcohol. The polymerization occurs at random by various carbon-carbon and ether bonds resulting in an irregular structure (Fig. 1)

Figure 1.

General structure of lignin

Lignin resists attacks by most microorganisms; anaerobic processes tend not to attack the aromatic rings at all and the aerobic breakdown of lignin is slow. In nature, only basidiomyceteous white-rot fungi (WRF) are able to degrade lignin efficiently. Wood decayed by white-rot fungi is pale in color because of oxidative bleaching and loss of lignin and often retains a fibrous texture.

In order to become the most effective wood degraders in nature, WRF developed non-specific and radical-based mechanisms for degrading lignin. WRF variously secrete one or more of four extracellular enzymes that are involved not only in the lignin degradation but also in the degradation of several pollutants. The four major lignin-modifying enzymes (LMEs) are: lignin peroxidase (LiP, EC, manganese peroxidase (MnP, EC, versatile peroxidase (VP, EC and laccase (Lac, EC [17]. All these enzymes act via generation of free radicals which represent an efficient way to reach their substrates.

The LMEs are highly non-specific and, thus, capable of degrading a wide range of recalcitrant compounds structurally similar to lignin. White-rot fungi usually secrete one or more of the LMEs in different combinations. Distribution of white-rot fungi into groups according to their enzymatic systems have been undertaken [18-20]. General classification is based on the capacity of different fungi to produce one or a combination of three ligninolytic enzymes: LiP, laccase and MnP. However, the absence or non-detection of these enzymes in some white-rot fungi, the sequencing of white-rot fungal genomes and the discovery of new enzymes has led to the distinction of different groups [21]. Generally, white-rot fungi can be distributed into four groups, according to their ability to produce laccases and peroxidases (LiP, MnP and VP):

  1. Laccase, MnP and LiP (Trametes versicolor, Bjerkandera adusta)

  2. laccase and at least one of the peroxidases (Lentinus edodes, Pleurotus eryngii, Ceriporiopsis subvermispora)

  3. only laccase (Schizophyllum commune)

  4. only peroxidases (Phanerochaete chrysosporium)

The most frequently observed LMEs among the white-rot fungi species are laccases and MnP and the least are LiP and VP.

A considerable number of reviews detailing the numerous characteristics and applications of ligninolytic enzymes have been published in past years [22-24]. A general description of the mechanisms and functions of the main ligninolytic enzymes of the white-rot fungi is given in Panel 1.

Active site/prosthetic group: four copper atoms.
Catalytic mechanism: catalysis of the one electron oxidation of phenolics, aromatic amines and other aromatic hydrogen donors with simultaneous reduction of O2 to H2O.
General reaction: 4 benzenediol + O2 → 4 benzosemiquinone + 2 H2O
Mediators: 3-HAA, ABTS, HBT, RBB
Function: Removal of polyphenols, degradation of pesticides and textile dyes, bleaching, removal of lignin, synthesis of melanin, formation of basidiomes, generation of hydroxyl radicals, stress defense
Main laccase producer fungi: Ganoderma lucidum, Pleurotus eryngii, Pleurotus ostreatus, Rigidoporus lignosus, Trametes versicolor
Active site/prosthetic group: Heme (Iron protoporphyrin IX)
Catalytic mechanism: oxidation of lignin side chains by one electron abstraction in the presence of H2O2, generating reactive radicals. Cleavage of C-C and ether bonds in lignin. Cleavage of aromatic rings.
General reaction: 1,2-bis(3,4-dimethoxyphenyl)propane-1,3-diol + H2O2 → 3,4-dimethoxybenzaldehyde + 1-(3,4-dimethoxyphenyl)ethane-1,2-diol + H2O
Mediators: VA, 2Cl-14DMB
Function: Degrades lignin (non-phenolic lignin units), mineralizes recalcitrant aromatic compounds.
Main LiP producer fungi: Bjerkandera adusta, Phanerochaete chrysosporium, Trametes cervina, Trametes versicolor
Active site/prosthetic group: Heme (Iron protoporphyrin IX)
Catalytic mechanism: Oxidize Mn2+ into Mn3+. Mn3+ acts as a diffusible oxidizer on phenolic or non-phenolic lignin units and amino-aromatic compounds
General reaction: 2 Mn2+ + 2 H+ + H2O2 → 2 Mn3+ + 2 H2O
Mediators: Chelated Mn3+
Co-oxidants: unsaturated fatty acids and thiols
Function: Oxidation and depolymerization of lignin, degradation of dyes, phenols and other recalcitrant compounds
Main MnP producer fungi: Ceriporiopsis subvermispora, Irpex lacteus, Lentinus edodes, Pleurotus ostreatus, Trametes versicolor
Active site/prosthetic group: Heme (Iron protoporphyrin IX). Hybrid molecular architecture with multiple binding sites
Catalytic mechanism: Combination of the catalytic properties of LiP and MnP; Oxidation of Mn2+ to Mn3+ and also phenolic and nonphenolic aromatic compounds.
General reaction: donor + H2O2 → oxidized donor + 2 H2O
Mediator: Chelated Mn3+
Function: Oxidizes phenols, hydroquinones, dyes, amines, aromatic alcohols and xenobiotics.
Main VP producer fungi: Bjerkandera adusta, Pleurotus eryngii

Panel 1.

Lignin-modifying enzymes of the white-rot fungi [17]

3-HAA (3-Hydroxyanthranilic acid), ABTS (2,2′-azinobis-(3-ethylbenzothiazoline-6-sulphonate)), HBT (1-hydroxybenzotriazole), RBB (Remazol brilliant blue), VA (veratryl alcohol), 2Cl-14DMB (2-Chloro-1,4-dimethoxybenzene).

The LMEs may function cooperatively or separately from each other, however, auxiliary enzymes (unable to degrade lignin on their own) are necessary to complete the process of lignin and/or xenobiotics degradation: aryl alcohol oxidase (AAO, EC, aryl alcohol dehydrogenase (AAD, EC, glyoxal oxidase (GLOX, EC, quinone reductase (QR, EC, cellobiose dehydrogenase (CDH, EC, superoxide dismutase (SOD, EC, glucose 1-oxidase (GOX, EC, pyranose 2-oxidase (P2Ox, EC and methanol oxidase (EC These are mostly oxidases generating H2O2 and dehydrogenases. Cytochrome P450 monooxygenases are also significant components involved in the degradation of lignin and many xenobiotics.

Recent additions to the enzymatic system of white-rot fungi include dye-decolorizing peroxidases or DyP involved in the oxidation of synthetic high redox-potential dyes and nonphenolic lignin model compounds [30] and aromatic peroxygenases (APOs) that catalyze diverse oxygen transfer reactions which can result in the cleavage of ethers [31-32].

Other important components of the lignin degradative system of white-rot fungi are low molecular mass oxidants such as hydroxyl radicals (•OH) and chelated Mn3+ (Mn3+ mainly chelated by oxalic acid), produced through the action of the ligninolytic enzymes. These are particularly important during the early stages of wood decay and can also act on the degradation of xenobiotics [24,33-35].


5. Environmental applications of ligninolytic enzymes

Ligninolytic enzymes are highly non-specific and, thus, capable of degrading a wide range of recalcitrant compounds structurally similar to lignin. In recent years, the capability of white-rot fungi and their enzymes to biodegrade several xenobiotics and recalcitrant pollutants has generated considerable research interest in the area of industrial/environmental microbiology. As a consequence, a considerable number of reviews detailing the numerous characteristics and applications of ligninolytic enzymes have been published [22-27,36-40]. Among the industrial applications of the ligninolytic enzymes found in WRF are: a). paper whitening, b). degradation of industrial dyes from the textile industry and c). degradation of xenobiotic compounds, including polycyclic aromatic hydrocarbons, phenolics, herbicides and other pesticides. In this review, we focused on the degradation of herbicides by white-rot fungi with emphasis on work done in the last decade.


6. Isolation and laboratory maintenance of white-rot fungi

There are about 10,000 species of white-rot fungi in the world. However, not more than 3 dozen have been more properly studied. Among these, the majority of studies have focused on Phanerochaete chrysosporium, followed by Trametes versicolor, Bjerkandera adusta and Pleurotus sp [40]. White-rot fungi have great potential for biotechnological applications. For this reason, there has been a growing interest in screening for new white-rot fungi species and strains. A classical strategy is to collect basidioma or mycelia of white-rot fungi in forests, dead trees, and lignocellulosic crop residues showing signs of attack by fungi. Samples of basidioma or mycelia are aseptically transferred onto potato dextrose agar or malt extract agar and sub-cultured until the obtainment of pure mycelia. Identification is based on morphological, physiological, biochemical and genetic characteristics of the basidioma, hyphae and spores. More recently, molecular biology techniques are being used in the identification of new isolates. Advancements in molecular methods have permitted a more rational study of the phylogenetic relationships within the various organisms. Non-coding internal transcribed spacer regions (ITS1 and ITS2) of the ribosomal DNA seem to be one of the most frequently employed analytical tools [41].

There are several options for the maintenance of white-rot fungi in the laboratory. Continuous growth methods of preservation in which white-rot fungi are grown on agar (e.g., malt extract agar, potato dextrose agar and yeast extract agar) are typically used for short-term storage. For long-term storage, preservation in distilled water (method of Castellani [42]), mineral oil and anhydrous silica gel are some of the indicated methods. These are low cost methods but none is considered permanent. Lyophilization and liquid nitrogen refrigeration (cryo-preservation) are expensive methods but are considered permanent.


7. Herbicide biotransformation and detoxification by white-rot fungi

Phanerochaete chrysosporium is the main white-rot fungus used in herbicide biodegradation studies. The lignin-degrading system of P. chrysosporium comprises the peroxidases LiP and MnP. Studies have demonstrated that these enzymes are responsible, at least in part, for the transformation of several persistent pollutant molecules including toxic herbicides [43]. The biotransformation of triazine by P. chrysosporium was studied in liquid cultures [44]. The herbicide atrazine was partly transformed into polar dechlorinated and/or N-dealkylated metabolites. The removal of the ethyl side chain was the preferred reaction and produced metabolites which were not further transformed by the fungus. They found that the N-dealkylation was supported by the mycelium and not by purified peroxidases. Thus, the authors suggested that cytochrome-P450 could have been involved in the degradation.

Using non-sterile substrates, P. chrysosporium demonstrated a satisfactory growth and significant effect on the degradation of several herbicides such as simazine and trifluralin [45], bentazon and 2-methyl-4-chlorophenoxyacetic acid (MCPA) [46-47]. Thus, the systems based on non-sterile substrates offer good perspectives of practical application since the preparation of large amounts of sterile substrates is expensive.

The herbicide diuron has also been degraded by P. chrysosporium in several culture conditions. Higher degradation efficiency (95%) was found in ashwood culture as compared to the liquid culture (75%) [48]. The authors suggested that MnP could be responsible for the degradation since it was the dominant enzyme detected in the cultures. In liquid medium, the degradation performance of P. chrysosporium is not very good when compared with other white-rot species in the same experiments. Within 7 days the depletion of diuron, linuron and propanil was not higher than 34% [49]. In another work, no Phanerochaete strain was able to degrade more than 42% of the herbicides chlortoluron, isoproturon and diuron during 5 days [50]. However, P. chrysosporium was more efficient than Trametes versicolor in cleaving diketonitrile, which is derived from the herbicide isoxaflotole [51]. Enzyme preparations derived from these organisms, namely LiP and MnP, were unable to catalyze the transformation even in the presence of mediators.

Solid state cultures (SSC) seem to be great systems for cultivation of ligninolytic fungi as well as for herbicide degradation. Isoproturon [52] and bentazon [46-47] were extensively degraded by P. chrysosporium in wheat straw cultures. In vitro tests suggest the action of LiP and MnP in the degradation of both herbicides although the cytochrome P450 activity can not be ruled out. For isoproturon, non-identified hydroxylated metabolites were formed in SSC as well as in pure enzyme tests. Furthermore, a second degradation reaction seems to occur with isoproturon since N-demethylated metabolites were also formed in both systems.

Bjerkandera adusta is one of the most promising strains of white-rot fungi for bioremediation. This fungus possesses high ligninolytic activity and produces several extracellular lignin-degrading enzymes, including LiP, MnP and laccase [53]. Also, it is one of the few species to produce VP [54]. B. adusta has demonstrated a great ability to degrade pollutant molecules, especially polycyclic aromatic hydrocarbons [55] and synthetic dyes [56-57]. The removal of the herbicides chlortoluron, isoproturon and diuron by B. adusta and other fungi was reported in shaking liquid cultures [50], B. adusta was the most efficient WRF able to degrade the three herbicides. Removal of chlortoluron, diuron and isoproturon reached 98%, 92% and 88%, respectively, after two weeks. On the other hand, it was reported that B. adusta was able to transform efficiently the herbicide linuron but to lower extents than propanil and diuron [49]. The authors suggested that the low depletion of some molecules may be due to the absence of phenolic or aromatic amine groups. Unfortunately, no efforts were made to evaluate which enzymatic system from B. adusta could be involved in these herbicide transformations. Nevertheless, in vitro experiments using purified versatile peroxidase from B. adusta proved that this enzyme is able to transform halogenated phenolic pesticides including the herbicide bromoxynil [54]. This enzyme polymerized bromoxynil molecules producing a dimer and a trimer as the main products with the loss of bromide atoms and hydroxyl groups. Nevertheless, VP from B. adusta was able to oxidize only three of 13 halogenated pesticides tested. It was suggested that the peroxidase-mediated reactions are limited by the presence of phenoxyl groups and by the electronic properties of the substituents on the aromatic ring.

The genus Pleurotus includes edible and medicinal species which are found both in temperate and tropical climates. They are one of the most important cultivated mushrooms in the world and are commonly referred to as “oyster mushrooms”. Pleurotus ostreatus and P. pulmonarius are the main species applied in bioremediation studies. These species are able to degrade PAH, chlorinated biphenyls and industrial dyes [36, 58-59]. Pleurotus species are known to be good producers of ligninolytic enzymes, mainly laccases [59], versatile peroxidase (VP) and manganese peroxidase (MnP) [24]. P. pulmonarius, cultured in pasteurized straw solid medium in association with bacteria, was able to degrade the highly persistent herbicide atrazine [60]. This study demonstrated that P. pulmonarius acts on the atrazine molecule producing chloro-metabolites which were further dechlorinated by bacteria, producing non-toxic metabolites. In liquid cultures, P. ostreatus was able to reduce the concentration of several herbicides. During five days, the herbicides chlortolurun, isoproturon and diuron were depleted 35, 42 and 33%, respectively [50]. Another study demonstrated maximum depletions of 42, 81 and 94% for diuron, linuron and propanil, respectively, during seven days [49]. In the latter studies no investigation about the enzymatic system involved in herbicide transformations was carried out.

Trametes versicolor seems to be highly promising for herbicide bioremediation. It is a good lignin degrader and produces Lac, MnP and LiP under certain culture conditions [61], being one of the most-studied laccase-producing fungus. Direct involvement of its laccase in herbicide transformation has been demonstrated. Recently, in vitro degradation experiments of glyphosate using several ligninolytic purified enzymes in combination with different mediators was carried out [62]. Laccase from T. versicolor was able to rapidly degrade the herbicide, but only in the presence of mediators such as ABTS, MnSO4 and Tween 80. The best glyphosate degradation by laccase (90.1%) was obtained in the presence of all these mediators in the same reaction. The laccase-mediator reactions produced and accumulated the metabolite AMPA, a compound frequently found in soil exposed to glyphosate. Another work demonstrated that the compound diketonitrile, the active principle of the herbicide isoxaflutole, was transformed by laccase, purified from T. versicolor cultures, in the presence of the mediator ABTS [51]. The main metabolite produced in the reaction was benzoic acid, an inactive compound also produced by tolerant plants. Laccase from this white-rot species also has been reported to degrade metabolites from phenylurea herbicides commonly found in the fields. These are hydroxylated metabolites produced by fungi and plants as part of their phenylurea herbicide metabolization. The hydroxylation of their aromatic rings makes phenylureas substrates for laccases and thus ready for further degradation [63]. Purified laccases were able to transform the metabolites 2-hydroxyphenyl-urea (2-HF) and 4-hydroxyphenyl-urea (4-HF) into several products without mediator addition. The results suggested that these enzymes from T. versicolor allow the formation of polymers between the hydroxyphenylurea products, as deduced from the formation of insoluble compounds in the reaction medium [64].

In vivo studies have also been carried out. T. versicolor strains were able to deplete diuron, linuron and propanil from the culture medium after seven days, reaching 37%, 47% and 94% pesticide transformation, respectively [49]. It was demonstrated that the herbicide diuron was the more extensively removed phenylurea in cultures of the Trametes species, with 72% transformation by Trametes sp and 33% by T. versicolor [50]. T. versicolor was able to degrade diuron and atrazine [65], chlornitrofen (CNP) and nitrofen (NIP) [66]. In the last work, the authors suggest that extracellular ligninolytic enzymes are not involved in the initial step of CNP and NIP degradation. However, the possibility of an effective participation of cytochrome P450 in the degradation of CNP and NIP has been emphasized.

Trametes versicolor has also been monitored for soil bioremediation in microcosms of non-sterile soils under water stress. Under this condition, this species was able to decompose atrazine without correlation with the production of ligninolytic enzymes [67] and to act on a pesticide mixture containing herbicides (simazine and trifluralin) and an insecticide (dieldrin) [45]. This type of study is very important because the capability of a given fungus to act in a soil microcosm can be strongly affected by competition with the native soil microflora and by the adverse conditions such as water stress.

Coriolopsis fulvocenerea, Cerrena maxima and Coriolus hirsutus were investigated as atrazine decomposers in liquid cultures [68]. Among the several fungi strains, the authors found that the best degraders were those that produce laccases. The results showed that these fungi degrade at least 80-92% of atrazine after 40 days. Further, cultures with addition of laccase inducers such as guaiacol and syringaldazine, were more effective in decreasing the atrazine concentration, suggesting an important role of these enzymes in atrazine degradation [69]. Recently, it was studied the effect of laccase from C. hirsutus on atrazine binding to soil [70]. Laccase introduction into the soil caused a rapid increase in irreversibly bound atrazine. This mechanism probably decreases the herbicide toxicity and thus contributes to atrazine detoxification in soil.

Recently, the decrease of the concentration of the herbicides bentazon and diuron in Ganoderma lucidum cultures was reported [71-72]. The action of the ligninolytic enzymes on bentazon was confirmed by in vitro experiments with the crude enzyme filtrates. After 24 hours, bentazon was completely degraded by the liquid culture filtrate. Also, solid state cultures using corn cobs as substrates were a good medium for the production of laccase and MnP as well as for the transformation of bentazon.

The white-rot fungus Phlebia brevispora metabolizes the herbicide CNP (chlornitrofen) by nearly 80% within 28 days of incubation with the production of five metabolites [73]. P. vispora was also able to degrade a highly toxic compound, a dibenzo-ρ-dioxin, present as a contaminant of commercially produced CNP herbicide [73]. The transformation of phenylurea herbicides by Phlebia radiate after five days has also been reported [50].

Hypholoma fasciculare is another white-rot fungus capable of degrading different herbicide classes [65]. This fungus possesses a great ability to degrade the triazine herbicides terbuthylazine and atrazine and the phenylurea herbicide diuron in liquid cultures and also in biobed bioremediation system. No relationship between the ligninolytic activity and the fungi capacity to degrade pesticides was found, although similar mechanisms could be involved in degradation of all the compounds.

Lentinus subnudus was highly efficient in removing atrazine (78%) and metalachlor (94%) from contaminated soils after 25 days [74]. No attempts were done to identify the participation of the ligninoloytic enzymes in the process.


8. Are ligninolytic enzymes actually involved in herbicide degradation?

WRF have been demonstrated to be capable of transforming and/or degrading a wide range of herbicide classes. Two mechanisms or systems have been proposed. The first is transformation in the extracellular space and it involves lignin-degrading enzymes. These powerful capabilities of WRF reside in the fact that many pollutants have structural similarities to lignin and because ligninolytic enzymes are non-specific they can also act on the pollutant molecules. Furthermore, the transformation of some compounds can be enhanced with the use of small molecules, called mediators, which can extend the enzymatic reactivity of enzymes towards the substrates; this process is often referred to as the enzyme-mediator system. The participation of extracellular enzymes in the transformation of several herbicides by WRF was conclusively demonstrated by studies performed with purified enzymes [46,47,49,51,54,62,63].

The second system of WRF involved in herbicide transformation is an intracellular enzymatic mechanism, represented mainly by cytochrome P450. Purification of fungal cytochrome P450, in order to obtain conclusive data, has been accomplished in only a few studies, due to the difficulties in keeping the activation of the enzymes during microsome preparation. Hence, most conclusions were drawn from the results of indirect experiments consisting in the addition of specific cytochrome P450 inhibitors to the culture medium, such as piperonyl butoxide and 1-aminobenzotriazole [66,75-76] (Fig. 2). Direct evidence is also available. Some experiments were carried out with the microsomal fraction isolated from Pleurotus ostreatus [77]. The authors found that the microsomes transformed the pesticides in vitro in a NADPH-dependent reaction. In Table 2 and 3, representative studies were listed in which ligninolytic enzymes of different WRF were used for removing herbicides. These tables show only those studies in which direct or strong indirect evidence is presented about the participation of enzymes (laccases, peroxidases and cytochrome P450 monooxygenases) for the removal of herbicides by WRF.

Figure 2.

Specific cytochrome P450 inhibitors

Pesticide/metabolite(s) Treatment conditions Fungal source Ref.
Glyphosate/AMPA In vitro assay. The presence of ABTS, Mn2+ and Tween 80 resulted in the removal of 90.1% of glyphosate. Trametes versicolor [62]
2,4-dichlorophenol (DCP): metabolite of 2,4-D In vitro assay. Laccase/mediator system. Laccase was immobilized on chitosan. Coriolus versicolor (syn. Trametes versicolor) [78]
Dymron In vitro assay. Laccase/mediator system. Over 90% of the herbicide was degraded by the addition of ABTS as mediator. Trametes sp. [79]
Bromofenoxim; Bromoxynil; Dichlorophen; Niclosamide In vitro assay. Laccase/mediator system. Addition of syringaldehyde or acetosyringone resulted in higher rates of enzymatic degradation. Coriolopsis gallica [49]
Bentazon In vitro assay. Crude enzymes from liquid and solid state cultures were applied. A synergistic effect of ABTS, Mn2+, H2O2, and Tween 80 was observed. Ganoderma lucidum [72]
Isoxaflutole. In plants and soils, isoxaflutole is rapidly converted to a diketonitrile derivative (DKN) The WRF are able to convert the DKN to inactive benzoic acid analogue when cultured in liquid media. The DKN derivative was incubated with fungal extracellular oxidases, without or with their specific redox m ediator. Both LiPs and MnPs presenting high specific activities were unable to catalyze the enzymatic transformation of DKN, with or without mediators, during 12-h incubations. Laccase transformed the herbicide when incubated with 1 mM ABTS. Phanerochaete chrysosporium and Trametes versicolor [51]
Pentachlorophenol (PCP) Submerged cultures in the presence and absence of laccase inducers. Cultures actively producing laccase removed 70% of initial PCP. The removal of PCP was less than 20 percent in the cultures with low laccase activity. Pleurotus pulmonarius [80]

Table 2.

Studies where strong evidences of participation of laccase in the biodegradation of herbicides were found

ABTS = 2,2′-azinobis-(3-ethylbenzothiazoline-6-sulphonate), AMPA ((aminomethyl)phosphonic acid).

Enzymes Pesticide/
Treatment conditions Fungal source Ref.
Manganese peroxidase
Glyphosate/AMPA In vitro assay. Addition of Mn2+ was essential for the oxidative activity of the enzyme. A reaction mixture containing Mn2+ and Tween 80 with or without H2O2 resulted in total degradation of the herbicide. Nematoloma frowardii [62]
Pentachlorophenol (PCP) Fungal inoculation in soil microcosms contaminated with PCP. Addition of wheat straw increased enzyme production and PCP degradation. Anthracophyllum discolor [82]
Bentazon In vitro assay. Crude enzymes from liquid and solid state cultures were applied. A synergistic effect of ABTS, Mn2+, H2O2, and Tween 80 was observed. Ganoderma lucidum [72]
Versatile peroxidase
Bromoxynil; Dichlorophen; PCP In vitro assay. Purified versatile peroxidase (VP) able to transform several pesticides. Analysis of reaction products by GC–MS showed the presence of 2,3,5,6-tetrachloroquinone among the products from pentachlorophenol oxidation, while the main product from dichlorophen was 4-chlorophenol-2,2-methylenequinone. Several polymers were obtained from the peroxidase oxidation of bromoxynil. In all cases, VP mediated dehalogenation reactions. Bjerkandera adusta [54]
CYP450 enzymes /
Cytochrome P450 monooxygenase
Azinphos-methyl; Phosmet; Terbufos In vitro assay. Pesticide transformation by microsomal fraction. Addition of NADPH in the reaction mixture was required. Pleurotus ostreatus [77]
trans-Chlordane Static liquid cultures. Addition of cytochrome P450 inhibitors reduced degradation activity and demonstrated involvement of CYP450 enzymes in the pesticide metabolism. Phlebia lindtneri,
Phlebia brevispora
Phlebia aurea
Cloronitrofen (CNP) and nitrofen (NIP) Extracellular LiP, MnP and Laccase did not catalyze the oxidation of either CNP or NIP. Piperonyl butoxide, an inhibitor of cytochrome P450, suppressed fungal oxidation of CNP and NIP to their hydroxylated products. The inhibition resulted in increasing the amount of reductively dechlorinated and nitro-reduced products. Trametes (Coriolus) versicolor [66]

Table 3.

Studies where evidence of participation of peroxidases and CYP450 enzymes in the biodegradation of herbicides


9. Concluding remarks

The ability of WRF to transform a wide variety of herbicide classes makes them suitable candidates to detoxify contaminated water and soils. Nevertheless, numerous WRF species and herbicide classes are still not explored. Ligninolytic enzymes, such as laccases and peroxidases, have shown to be responsible for many herbicide transformations, although their action remains to be confirmed in most studies. In this context, also the cytochrome P450 system has demonstrated to have a very important role in bioremediation of numerous types of pollutants including herbicides, although further studies must be done in order to understand the action of this system in transformation of herbicides by WRF.

The employment of ligninolytic enzyme preparations could be advantageous over the direct use of white-rot fungi, considering that these preparations can be more easily standardized, facilitating accurate dosage. Enzyme application is simpler than the use of microorganisms and can be rapidly modified according to the characteristics of the herbicide to be removed. In addition to this, analysis of metabolite compounds produced by enzyme preparations is easier than analysis of the metabolites produced by the whole white-rot fungi.

There is a need for introducing new methodologies for the elucidation of the chemical structures of the metabolites produced by the action of ligninolytic enzymes. This should also facilitate the elucidation of the metabolic routes used by the white-rot fungi in herbicide degradation. Furthermore, the proof that the process of degradation corresponds to a real detoxification is a crucial step that must be overcome before its practical use.



This work was supported by grants from Fundação Araucária (Proc. ) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq, Edital 047/2010-SISBIOTA, proc. 563260/2010-6). Rosane Marina Peralta and Adelar Bracht are fellowship holders of CNPq. Jaqueline da Silva Coelho-Moreira and Giselle Maria Maciel are fellowship holders of the Coordenação de Aperfeiçoamento de Pessoal de Ensino Superior (CAPES).


  1. 1. Gavrilescu M. Fate of pesticides in the environment and its bioremediation. Engineering in Life Sicence 2005;5(4),497-526.
  2. 2. United States Environmental Protection Agency. EPA. http;// (acessed 5 october 2012)
  3. 3. Chowdhury A, Pradhan S, Saha M, Sanyal N. Impact of pesticides on soil microbiological parameters and possible bioremediation strategies. Industrial Journal of Microbiology 2008;48,114-127.
  4. 4. Rodrigues BN, Almeida FS. Herbicides Guide. Iapar, Brazil. 1998. p648.
  5. 5. Gianfreda L, Rao MA. Potential of extra cellular enzymes in remediation of polluted soils: a review. Enzyme and Microbial Technology 2004;35,339-354.
  6. 6. Vidali M. Bioremediation. An overview. Pure and Applied Chemistry 2001;73(7),1163-1172.
  7. 7. Bennet JW, Wunch KG, Faison BD. Use of fungi in Biodegradation. In: Hurst CJ (eds) Manual of Environmental Microbiology. ASM Press Washington, 2002; p 960-971.
  8. 8. Bellinaso MDL, Greer CW, Peralba MC, Henriques JAP, Gaylarde CC. Biodegradation of the herbicide trifluralin by bacteria isolated from soil. FEMS Microbiology and Ecolology 2003;43,91-194.
  9. 9. Dwivedi S, Singh BR, Al-Khedhairy AA, Musarrat J. Biodegradation of isoproturon using a novel Pseudomonas aeruginosa strain JS-11 as a multi-functional bioinoculant of environmental significance. Journal of. Hazardous Materials 2011;185 (2-3),3938-944.
  10. 10. Mandelbaum RT, Allan DL, Wackett LP. Isolation and characterization of a Pseudomonas sp that mineralizes the s-triazine herbicide atrazine. Applied and Environmental Microbiology 1995;61,1451-1457.
  11. 11. Barr DP, Aust SD. Mechanisms white rot fungi use to degrade pollutants. Environmental Science and Technology 1994;28 (2)79-87.
  12. 12. Asgher M, Bhatti HN, Ashraf M, Legge RL. Recent developments in biodegradation of industrial pollutants by white rot fungi and their enzyme system. Biodegradation 2008;19,771-783.
  13. 13. Pointing SB. Feasibility of bioremediation by white-rot fungi. Applied Microbiogy and Technology 2001;57,509-526.
  14. 14. Reddy CA. The potential for white-rot fungi in the treatment of pollutants. Current Opinion in Biotechnology 1995;6:,320-328.
  15. 15. Have RT, Teunissen PJM. Oxidative mechanisms involved in lignin degradation by white-rot fungi. Chemical Reviews 2001;101,3397-3413.
  16. 16. Leonowicz AL, Matuszewska A, Luterek J, Ziegenhagen D, Wajtas-Wasilewska M, Nam-Seok C, Hofrichter M, Rogalski J. Biodegradation of Lignin by white rot fungi. Fungal Genetics and Biology 1999;27,175-185.
  17. 17. Maciel GM, Bracht A, Souza CGM, Costa AM, Peralta RM. Fundamentals, diversity and application of white-rot fungi. In: Silva AP, Sol M (Eds) Fungi: types, environmental impact and role in disease, Nova Science Publishers, Inc. p409-458
  18. 18. Hatakka A. Lignin-modifying enzymes from selected white-rot fungi: production and role in lignin degradation. FEMS Microbiology Reviews 1994;13,125-135
  19. 19. Nerud F, Misurcova Z. Distribution of ligninolytic enzymes in selected white-rot fungi. Folia Microbiologica 1996;41,264-266
  20. 20. Tuor U, Winterhalter K, Fiechter A. Enzymes of white-rot fungi involved in lignin degradation and ecological determinants for wood decay. Journal of Biotechnology 1995;41,1-17
  21. 21. Kuhar S, Kapoor M, Kapoor R, Sharma KK, Singh A, Kuhad RC. Biodiversity of ligninolytic fungi. In Kuhad RC, Singh A (Eds) Lignocellulose biotechnology: future prospects. New Delhi: I K International p37-53
  22. 22. Ruiz-Duenas FJ, Martinez AT. Microbial degradation of lignin: how a bulky recalcitrant polymer is efficiently recycled in nature and how we can take advantage of this. Microbial Technology 2009;2,164-177
  23. 23. Kersten P, Cullen D. Extracellular oxidative systems of the lignin-degrading basidiomycete Phanerochaete chrysosporium. Fungal Genetics and Biology 2007;44,77-87.
  24. 24. Hofrichter M Review: lignin conversion by manganese peroxidase (MnP). Enzyme and Microbial Technology 2002;30,454–466.
  25. 25. Mayer AM, Staples RC. Laccase: new functions for an old enzyme. Phytochemistry 2002;60,551-565.
  26. 26. Wesenberg D, Kyriakides I, Agathos SN. White-rot fungi and their enzymes for the treatment of industrial dye effluents. Biotechnology Advances 2003;22,161-187
  27. 27. Baldrian P. Fungal laccases – occurrence and properties. FEMS Microbiolology Review 2006;30,215-242.
  28. 28. Lundell TK, Makela MR, Hilden K. Lignin-modifying enzymes in filamentous basidiomycetes: ecological, functional and phylogenetic review. Journal of Basic Microbiology 2010;50,5-20
  29. 29. Conesa A, Punt PJ. van den Hondel C. Fungal peroxidases: molecular aspects and applications. Journal of Biotechnology 2002;93,143-158
  30. 30. Liers C, Bobeth C, Pecyna M, Ullrich R, Hofrichter M. DyP-like peroxidases of the jelly fungus Auricularia auricula-judae oxidize nonphenolic lignin model compounds and high-redox potential dyes. Applied Microbiology and Biotechnology 2010;85,1869-1879.
  31. 31. Liers C, Arnstadt T, Ullrich R, Hofrichter M. Patterns of lignin degradation and oxidative enzyme secretion by different wood- and litter-colonizing basidiomycetes and ascomycetes grown on beech-wood. FEMS Microbiology Ecology 2011;78(1),91-102
  32. 32. Hofrichter M, Ullrich R, Pecyna MJ, Liers C, Lundell T. New and classic families of secreted fungal heme peroxidases. Applied Microbiology and Biotechnology 2010;87,871-897
  33. 33. Hammel KE, Kapich AN, Jensen KA, Ryan ZC. Reactive oxygen species as agents of wood decay by fungi. Enzyme and Microbial Technology 2002;30,445-453
  34. 34. Watanabe T, Teranishi H, Honda Y, Kuwahara M. A selective lignin-degrading fungus, Ceriporiopsis subvermispora, produces alkylitaconates that inhibit the production of a cellulolytic active oxygen species, hydroxyl radical in the presence of iron and H2O2. Biochemical and Biophysical Research Communications 2002;297,918-923
  35. 35. ten Have R, Teunissen PJM. Oxidative mechanisms involved in lignin degradation by white-rot fungi Chemical Reviews 2001;101,3397-3413.
  36. 36. Pozdnyakova NN, Rodakiewicz-Nowak J, Turkovskaya OV, Haber J. Oxidative degradation of polyaromatic hydrocarbons catalyzed by blue laccase from Pleurotus ostreatus D1 in the presence of synthetic mediators. Enzyme and Microbial Technology 2006;39,1242–1249.
  37. 37. Aust SD, Swaner PR, Stahl JD Detoxification and metabolism of chemicals by white-rot fungi. In: Gan JJ, Zhu PC. Aust SD, Lemley AT (eds) Pesticide decontamination and detoxification, American Chemical Society, Washington, DC. Oxford University Press, 2003, p3-14.
  38. 38. Couto SR, Herrera JLT. Industrial and biotechnological applications of laccases: A review. Biotechnology Advance 2006;24,500-513.
  39. 39. Riva S. Laccases: Blue enzymes for green chemistry. Trends in Biotechnology 2006;24(5),219-226.
  40. 40. Dashtban M, Schraft H, Syed T, Qin W. Fungal biodegradation and enzyme modification of lignin. International Journal of Biochemistry and Molecular Biology 2010;1,36-50
  41. 41. Prewitt ML, Diehl S, McElroy TC, Diehl WJ. Comparison of general fungal and basidiomycete-specific ITS primers for identification of wood decay fungi. Forest Products Journal 2008;58,68-71
  42. 42. Castellani A. Viability of some pathogenic fungi in distilled water. Journal of Tropical Medicine and Hygiene 1939;42,225-226
  43. 43. Marcos-Urrea E, Reddy CA. Degradation of chloro-organic pollutantsw by white rot fungi. In: Sing SN (Ed) Microbial degradation ox xenobiotics. Springer Berlin Heidelberg 2012. p31-66
  44. 44. Mougin C, Laugero C, Asther M, Dubroca J, Frasse P, Asther M. Biotransformation of the herbicide atrazine by white rot fungus Phanerochaete chrysosporium. Applied and Environmental Microbiology 1994;60(2),705-708.
  45. 45. Fragoeiro S, Magan N. Impact of Trametes versicolor and Phanerochaete chrysosporium on differential breakdown of pesticide mixtures in soil microcosms at two water potentials and associated respiration and enzyme activity. International Biodeterioration and Biodegradation 2008;62,376-383.
  46. 46. Castillo MDP, Ander P, Stenstrom J, Torstensson L. Degradation of the herbicide bentazon as related to enzyme production by Phanerochaete chrysosporium in two solid substrate fermentation systems. World Journal of Microbiology and Biotechnology 2000;16,289-295.
  47. 47. Castillo MDP, Andersson A, Ander P, Stenstrom J, Torstensson L. Establishment of the white rot fungus Phanerochaete chrysosporium on unsterile straw in solid substrate fermentation systems intended for degradation of pesticides. World Journal of Microbiology and Biotechnology 2001;17,627-633.
  48. 48. Fratila-apachitei LE, Hirst JA, Siebel MA, Gijzen HJ. Diuron degradation by Phanerochaete chrysosporium BKM-F-1767 in synthetic and natural media. Biotechnology Letters 1999;21,147-154.
  49. 49. Torres-Duarte C, Roman R, Tinoco R, Vazquez-Duhalt R. Halogenated pesticide transformation by a laccase-mediator system. Chemosphere 2009;77,687-692.
  50. 50. Khadrani A, Seigle-Murandi F, Steiman R, Vroumsia T. Degradation of three phenylurea herbicides (chlortoluron, isoproturon and diuron) by micromycetes isolated from soil. Chemosphere 1999;38(13),3041-3050.
  51. 51. Mougin C, Boyer F-D, Caminade E, Rama R. Cleavage of the diketonitrile derivative of the herbicide isoxaflutole by extracellular fungal oxidases. Journal of Agricultural and Food Chemistry 2000;48:4529-4534.
  52. 52. Castillo MDP, Wirén-Lehr SV, Scheunert I, Torstensson L. Degradation of isoproturon by the white rot fungus Phanerochaete chrysosporium. Biology and Fertility Soils 2001;33,521-528.
  53. 53. Nakamura Y, Sungusia MG, Sawada T, Kuwahara M. Lignin-Degrading Enzyme Production by Bjerkandera adusta Immobilized on Polyurethane Foam. Journal of Bioscience and Bioengineering 1999;88(1),41-47.
  54. 54. Davila-Vazquez G, Tinoco R, Pickard MA, Vazquez-Duhalt R. Transformation of halogenated pesticides by versatile peroxidase from Bjerkandera adusta. Enzyme and Microbial Technology 2005;36,223-231.
  55. 55. Field JA, Jong ED, Costa GF, De Bont JAM. Biodegradation of Polycyclic Aromatic Hydrocarbons by New Isolates of White Rot Fungi. Applied Environmental and Microbiology 1992;58,2219-2226.
  56. 56. Heinfling A, Bergbauer M, Szewzyk U. Biodegradation of azo and phthalocyanine dyes by Trametes versicolor and Bjerkandera adusta. Applied Microbiology and Biotechnology 1997;48,261-266.
  57. 57. Eichlerová I, Homolka L, Nerud F. Decolorization of high concentrations of synthetic dyes by the white rot fungus Bjerkandera adusta strain CCBAS 232. Dyes and Pigments 2007;75,38-44.
  58. 58. Zeddel A, Majcherczyk A, Hu°ttermann A. Degradation of polychlorinated biphenyls by white-rot fungi Pleurotus ostreatus and Trametes in a solid state system. Toxicological and Environmental Chemistry 1993;40,255–266.
  59. 59. Tychanowicz GK, Zilly A, Souza CGM, Peralta RM. Decolorization of industrial dyes by solid-state cultures of Pleurotus pulmonarius. Process Biochemistry 2004;39,855–859.
  60. 60. Masaphy S, Levanon D, Henis Y.Degradation of atrazine by the lignocellulolytic fungus Pleurotus pulmonarius during solid-state fermentation. Bioresource Technology 1996;56,207-214.
  61. 61. Rogalski J, Lundell T, Leonowicz A, Hatakka A. Production of laccase, lignin peroxidase and manganese-dependent peroxidase by various strains of Trametes versicolor depending on culture conditions. Acta Microbiologica Polonica 1991;40,221-234.
  62. 62. Pizzul L, Castillo MDP, Stenstrom J. Degradation of glyphosate and other pesticides by ligninolytic enzymes. Biodegradation 2009;20,751-759.
  63. 63. Jolivalt C, Neuville L, Boyer FD, Kerhoas L, Mougin C. Identification and formation pathway of laccase-mediated oxidation products formed from hydroxyphenylureas. Journal of Agricultural and Food Chemistry 2006;54,5046-5054.
  64. 64. Jolivalt C, Raynal A, Caminade E, Kokel B, Le Goffic F, Mougin C. Transformation of N’, N’-dimethyl-N-(hydroxyphenyl)ureas by laccase from the white rot fungus Trametes versicolor. Applied Microbiology and Biotechnology 1999;51,676-681.
  65. 65. Bending GD, Friloux M, Walker A. Degradation of contrasting pesticides by white rot fungi and its relationship with ligninolytic potential. FEMS Microbiology Letters 2002;212,59-63.
  66. 66. Hiratsuka N, Wariishi H, Tanaka H. Degradation of diphenyl ether herbicides by the lignin-degrading basidiomycete Coriolus versicolor. Applied Microbiology and Biotechnology 2001;57,563-571.
  67. 67. Bastos AC, Magan N. Trametes versicolor: potential for atrazine bioremediation in calcareous clay soil, under low water availability conditions. International Biodeterioration and Biodegradation 2009;63,389-394.
  68. 68. Koroleva OV, Stepanova EV, Landesman EO, Vasilchenko LG, Khromonygina VV, Zherdev AV, Rabinovich ML. Enzyme immunoassay of herbicide decomposition by soil and wood decay fungi. Appl. Biochem. Microbiol., 2002;38, 355-360.
  69. 69. Gorbatova ON, Koroleva OV, Landesman EO, Stepanova EV, Zherdev AV. Increase of the detoxification potential of Basidiomycetes by induction of laccase biosynthesis. Applied Biochemistry and Microbiology 2006;42(4),414-419.
  70. 70. Davidchik VN, Kulikova NA, Golubeva LI, Stepanova EV, Koroleva OV. Coriolus hirsutus laccase effect on atrazine adsorption and desorpton by different types of soil. Applied Biochemistry and Microbiology 2008;45(4),405-410.
  71. 71. Coelho JD, Oliveira AL, Souza CGM, Bracht A, Peralta RM. Effect of the herbicides bentazon and diuron on the production of ligninolytic enzymes by Ganoderma lucidum. International Biodeterioration and Biodegradation 2010;64(2),156-161.
  72. 72. Coelho JD, Souza CGM, Oliveira AL, Bracht A, Costa MAF, Peralta RM. Comparative removal of bentazon by Ganoderma lucidum in liquid and solid state cultures. Current Microbiology 2010;60(5),350-355.
  73. 73. Kamei I, Kondo R. Simultaneous degradation of commercially produced CNP herbicide and of contaminated dioxin by treatment using the white rot fungus Phlebia brevispora. Chemosphere 2006;65,1221-1227.
  74. 74. Nwachukwu EO, Osuji JO. Bioremedial degradation of some herbicides by indigenous white rot fungus, Lentinus subnudus. Journal of Plant Sciences 2007;2(6),619-624.
  75. 75. Mougin C, Laugero C, Asther M, Chaplain V. Biotransformation of s-triazine herbicides and related degradation products in liquid culture by the white rot fungus Phanerochaete chrysosporium. Pesticide Science 1997;49,169-177.
  76. 76. Xiao P, Mori T, Kondo R. Biotransformation of the organochlorine pesticide trans-chlordane by wood-rot fungi. New Biotechnology, 2011;29(1),107-115.
  77. 77. Jauregui J, Valderrama B, Albores A, Vazquez-Duhalt R. Microsomal transformation of organophosphorus pesticides by white rot fungi. Biodegradation, 2003;14(6),397-406.
  78. 78. Zhang J, Xu Z, Chen H, Zong Y. Removal of 2,4-dichlorophenol by chitosan-immobilized laccase from Coriolus versicolor. Biochemical Engineering Journal, 2009;45(1),54-59.
  79. 79. Maruyama T, Komatsu C, Michizoe J, Ichinose H, Goto M. Laccase-Mediated Oxidative Degradation of the Herbicide Dymron. Biotechnology Progress 2006;22(2),426-430.
  80. 80. Souza DF, Costa SC, Dacome AS, Souza CGM, Bracht A, Peralta RM. Pentachlorophenol removal by Pleurotus pulmonarius in submerged cultures. Brazilian Archives of Biology and Technology 2011;54(2),357-362
  81. 81. Cea M, Jorquera M, Rubilar O, Langer H, Tortella G, Diez MC. Bioremediation of soil contaminated with pentachlorophenol by Anthracophyllum discolor and its effect on soil microbial community. Journal of Hazardous Materials 2010;181(1–3),315-323.

Written By

Jaqueline da Silva Coelho-Moreira, Giselle Maria Maciel, Rafael Castoldi, Simone da Silva Mariano, Fabíola Dorneles Inácio, Adelar Bracht and Rosane Marina Peralta

Submitted: October 23rd, 2012 Published: May 29th, 2013