Influence of different herbicides on soil fungi
Abstract
Understanding herbicide transformation is necessary for pesticide development for their safe and efficient use, as well as for developing pesticide bioremediation strategies for contaminated soil and water. Recent studies persuasively demonstrated the key role of soil white-rot fungi in biotransformation of various anthropogenic environmental contaminants. However, often this common knowledge is not associated with specific metabolic processes of fungi and therefore cannot be transformed into specific recommendations for agricultural practice. The given review offers a systematic collection and analysis of the current knowledge about herbicide transformation by white-rot fungi at the cellular and molecular levels. Special attention is given to the role of oxidative enzymes such as laccases, lignin peroxidases, and manganese peroxidases in the biotransformation processes.
Keywords
- White-rot fungi
- biotransformation
- herbicides
- oxidases
- metabolic fate
1. Introduction
Fungi are unique organisms that colonize all areas of the environment – air, water, and soil. This group lists more than 1.5 million species and is remarkably flexible, occupying all biocenoses from the arctic tundra to the deserts. Biodiversity and specific genetic and molecular organization of fungi provided background for their key role in nature, i.e., maintaining of ecosystems’ equilibrium. One of the most important groups playing a key role in the carbon cycle in nature is Wood Degrading Fungi, due to its ability to degrade or even mineralize lignin – widely present and one of the most stable biopolymers. They belong to Basidiomycota and Ascomycota and possess the unique ability to degrade components of xylem cell walls (cellulose, hemicellulose, lignin, and compounds forming these biopolymers). According to Anastasi et al. [1], this group is divided into white-rot fungi (WRF) or white rotters, brown-rot fungi, and soft-rot fungi because of the appearance of rotten wood.
The ability to degrade lignin and its aromatic compounds is mostly attributed to white-rot fungi [2]. The white-rot decay of wood is performed by the combined action of oxidoreductive metalloenzymes, heme peroxidases, and laccases, encoded by multigene families as well as organic acids, secondary metabolites, and surfactants secreted by WRF [3]. This assembly is considered the Lignin Modifying System (LMS) [4]. It should be mentioned that extracellular ligninolytic enzymes (laccases, lignin peroxidases, manganese peroxidases, and versatile peroxidases) are nonspecific and can act both alone as well as using a redox mediator that enhances the range of potential substrates and provides the possibility of effective oxidation of xenobiotics. The WRF also produces reactive oxygen species (ROS) such as superoxide anion radical (O2-), hydrogen peroxide (H2O2), and hydroxyl radicals (OH-) [5]. WRF, their LMS and ROS are involved in the degradation of lignin and carbohydrate components of wood commonly accomplished by production of carbon dioxide and water. Both WRF and LMS are capable of in vitro oxidizing and degrading a broad range of xenobiotics: polycyclic aromatic hydrocarbons (anthracene, benz[a]pyrene, naphthalene, and phenanthrene); polychlorinated phenols (2,4-di-, 2,4,5-, and 2,4,6-tri-, and pentachlorophenols), chlorinated guaiacol and benzoate derivatives, 2,4,6-trichlorophenoxyacetate, and chlorinated biphenyls; stable polymers (polyacrylate, polyacrylamide, polycaprolactam, and polyethylene), 2,4-dichloroaniline, dioxins, explosives nitrates, dyes. The degradation of xenobiotics by WRF as well as enzymatic aspects of these processes is well documented and summarized in several recent reviews [1,6-10]. However, there are contradictory data reported on the role of ligninolytic enzymes in pesticide degradation. There was no relationship between WRF degradation of the dye Poly R-478, a presumptive test for ligninolytic potential, and degradation of the highly available pesticides, diuron, metalaxyl, atrazine or terbuthylazine in liquid culture [11]. Moreover, it was also shown that no degradation of the herbicide picloram by
Recent findings have highlighted the molecular aspects of ligninolytic enzymes’ functioning [13-15]. Genes-encoding ligninolytic enzymes of the white-rot fungi have been found to undergo differential regulation in response to different environmental signals and stimuli such as carbon and nitrogen concentration in cultural media, presence of xenobiotics and heavy metals, temperature regime, and various lengths of daylight. The analysis of MnP, LiP, and laccase gene promoter regions revealed the presence of xenobiotic response mechanism (XRE – xenobiotic responsive element), suggesting that these enzyme expressions can be similar in the presence of xenobiotics [15]. It was shown that compounds such as paracetic acid, ethanol, sodium arsenite, 2,4-dichlorophenol, and N,N-dimethylformamide enhanced the MnP production [16]. Moreover, a list of available aromatic compounds including xenobiotics (1-hydroxybenzotriazole, 2,5-xylidine, o-toluidine, 3,5-dihydroxytoluene, dimethylphenol, caffeic acid, caffein, guaiacol, hydroquinone, etc.) that demonstrated the similar effect on laccase production was generated by Piscitelli et al. [17]. The data available confirmed that regulation of the expression of genes-encoded ligninolytic enzymes is a highly complex process. However, the constant progress in molecular and genomic techniques gave new insights on the role of regulating elements in the differential expression of ligninolytic enzymes in WRF. Further studies will elucidate the mechanisms of ligninolytic enzyme transcriptional regulation and provide deeper understanding of this complicated process.
Thus, the potential of ligninolytic enzymes in degradation of herbicides has not been well characterized yet, especially at the molecular level. Most of the data available correspond to the studies of different herbicide degradation by WRF, their individual ligninolytic enzymes and oxidative enzymes – redox mediator systems that are successful or less successful [6,9,10,18-21]. Few attempts have been made to propose the mechanisms of pesticide degradation (based on pentachlorophenol degradation pathways and ligninolytic enzymes action). The aim of this review is to summarize the data about herbicide degradation by WRF and their ligninolytic enzymes.
2. Modern herbicides and common regularities of their transformation
Approximately 2 million tons of pesticides are used worldwide each year [22] and play a significant role in modern agricultural practices. Approximately half of this volume is herbicides that are routinely applied to crops at rates varying from g to kg ha−1. In 2010, about 907 million kg of active ingredients of herbicides was applied in the world (FAO data), and this figure continues to grow. 2019 estimates demonstrate that the herbicides market will experience both the highest growth rate as well as the highest volume traded in the next years as compared with other pesticides. The expected annual growth rate of herbicides for the given period is computed to be 6.1% [23].
Although attempts to reduce pesticide use through organic agricultural practices and the use of other technologies continue, direct and indirect exposure to pesticides is still an important health risk factor. About one-third of the agricultural products are produced by using pesticides [24]. Without pesticide application the loss of fruits, vegetables, and cereals from pest injury would reach 78%, 54%, and 32%, respectively [25].
The emergence of herbicide-resistant (HR) genetically engineered crops in 1996 made it possible for farmers to use a broad-spectrum herbicide, glyphosate, in ways that were previously impossible. From 1996 through 2011, 0.55 billion hectares of HR corn, soybeans, and cotton were grown in the USA, and in 2011, an estimated 94% of the soybean area planted, 72% of corn, and 96% of cotton were planted to HR varieties, respectively, which led to a 239 million kg increase in herbicide use [26].
Priority pesticides vary significantly for different regions and crops, and have evolved with time. The era of organic synthetic pesticides started approximately 70 years ago from DDT, 2,4-D, and such compounds as HCH, dieldrin were added to the most actively used compounds at the second wave. The assortment of modern pesticides is highly variable in trademarks and based on relatively wide (but much shorter) row of successfully commercialized active ingredients. However, the bulk of the world market is formed by a very small number of compounds, even taking into consideration their variability for different regions (really – for main crops of these regions). Atrazine, glyphosate, acetochlor, metolachlor, tefluthrin, cyfluthrin, and, maybe, mesotrione should be considered as priority pesticides for environmental/health risks due to their wide application as protection tools in cereal agriculture. It should be noted, however, that the integral impact of two parameters, i.e., manufacturing volume and toxicity of active compounds, determines danger of different agrochemicals and necessity in their efficient decontamination.
Only a lesser part of applied pesticides reaches the target organism, with the remainder being deposited on the soil and nontarget organisms, as well as moving into the environment [27]. The metabolic fate of pesticides is dependent on their physico-chemical characteristics, field abiotic conditions, and plant and microbial communities. Transformation of pesticides includes abiotic processes (such as photolysis, hydrolysis, oxidation, and rearrangements) and chemical/biological reactions. The variety of biotransformation processes for herbicides should be considered in connection with specific features of the microenvironments in and near target organisms involved into metabolic pathways. So the key stage for determining further biotransformation of herbicides is their adsorption (and adsorption of their intermediate metabolites) to soil and soil colloids. These processes are highly important to regulate the dynamics of action for modern herbicide preparations. The ratio between free and adsorbed forms of herbicides determines the rate of their abiotic transformation. Nevertheless, enzymatic transformation (i.e., biotransformation) is the major driver of detoxification.
The classic concepts of pesticide metabolism [28,29] divide their transformation into three phases. In the first phase, the parent compounds are transformed through oxidation, reduction, or hydrolysis to more water-soluble and usually less toxic products. As a whole, oxidation (hydroxylation, dealkylation, and deamination), hydrolysis (esters, amides, and nitriles) and reduction reactions are considered as main factors for this phase. The main process of the second phase is the conjugation of the obtained derivatives to a sugar (typically glucose), glutathione, or amino acid with further increased water solubility, reduced toxicity, and support of internal transport of the metabolite for final transformation. The third phase provides further conjugation and results in nontoxic final products of metabolic pathways of the pesticides.
An important factor in the transformation of pesticides in soil is a complementary action of plants and microorganisms on them. The roles of plants may be simply characterized as reduction of toxicity, whereas microorganims are responsible for deep destruction and mineralization. The line of enzymes and catalyzed reactions for microorganisms is much wider as compared with plants. Several processes, such as dehalogenation or C-P bond cleavage, are associated mainly with microbial metabolism of pesticides. (On the other hand, however, glutathione conjugation is a typical tool for plant transformation of pesticides.) The common opinion in modern remediation biotechnology is that the tasks of detoxification cannot be solved at the plant level alone and should be based on the detailed analysis of the most efficient microbial participants of this process. A significant additional factor of interest concerning microbial detoxification is the lower cost of such technologies as compared with the alternative ones [30].
Pathways of pesticide destruction have been described in many works, both at the levels of the species responsible and the enzymes involved. Currently, the existing information is systematized in several sources, and Biocatalysis/Biodegradation Database of the University of Minnesota (EAWAG-BBD; http://eawag-bbd.ethz.ch) seems the most informative available tool. This database contains information on microbial biocatalytic reactions and biodegradation pathways for primarily xenobiotics. This permanently maintained and updated system collects data about hundreds of pathways, enzymes, and microorganisms, thousands of reactions and compounds of environmental interests. In addition, this database contains two supporting tools. The Pathway Prediction System predicts microbial catabolic reactions using substructure searching, a rule-base, and atom-to-atom mapping. The biotransformation rules are based on reactions found in the EAWAG-BBD or in the scientific literature. The Biochemical Periodic Table provides an overview of microbial interactions with different chemical elements. Individual element pages contain a summary of published data about microbial interactions with the selected element.
It should be noted that efficient recommendations for microbial remediation require integral knowledge about potential of individual enzymatic reactions and specific features of their interactions for different microbial species. Current information about genetic regulation of coupled reactions may significantly improve bioremediation technologies as well as contribute to empiric data about multistep detoxification with the use of different microorganisms. That is why the further consideration of fungal destruction of herbicides will provide data on integrated potential of multienzyme systems from different detoxificators rather than data about elementary catalytic reactions, but first of all about.
3. Effects of herbicides on soil fungi
There are two main teqniques of herbicide application in the field. The first one is foiliar spray, and the second is soil application. In case with soil application, the herbicide is introduced directly into the soil and so can affect soil microorganisms. However, even in case with foliar application, significant amounts of these chemicals reach the soil. Therefore, although herbicides are very useful in farming, under certain circumstances they may turn into pollutants, affecting soil microflora and deteriorating the quality of soil if there are sensitive organisms and/or if the degradation products are toxic. Among various indicators used in monitoring soil biological activity, microbial community structure seems to be the most preferred due to its sensitivity to the environmental changes. To address these concerns, the impacts of herbicides on soil microbial communities are widely studied and discussed.
In general, the recommended field rate of herbicide had no major effects on soil microorganisms, but excessive doses retard the reproduction rate of some groups of microflora and may reduce enzyme activity and populations of various microorganisms in soil, including fungi (Table 1) [31-35]. No significant changes in soil microflora were detected using phospholipid fatty acid (PLFA) profiles’ analysis after atrazine, bentazon, or glyphosate application by Banks and coauthors [35]. Crouzet and coauthors [33] tested the herbicide mesotrione in chernozem soil at the rates from 0.45 to 45 mg/kg and recorded only small genetic structural shifts in the bacterial and fungal communities. Maximum dissimilarity of the bacterial and fungal genetic structures between control and herbicide-treated soil did not exceed 12% and 28%, respectively. Martinez et al. [36] did not demonstrate any significant changes in the multiplication of bacteria and fungi following an application of sulfentrazone. Allievi and Gigliotti demonstrated no significant differences in number of aerobic bacteria in soil attributable to cinosulfuron treatment at the field rate 0.42 μg/kg after 1 and 4 weeks of incubation under laboratory conditions [31]. Possible effects of the herbicide on the specific group of microorganisms of the microbial community resulting in eventual counterbalance by the development of another group were further tested. To execute this, the individual microbial strains were isolated and their sensitivity in relation to cinosulfuron was tested. Among eighteen studied strains of aerobic bacteria from uncultivated soil, a fourth of the tested strains underwent some growth inhibition in the presence of the herbicide, and for one strain total and permanent inhibition was observed. In the case of fungi, however, only two of seventeen fungi strains underwent temporary growth inhibition. In the case of isolates from agricultural soil, neither bacterial nor fungal isolates were sensitive to the studied herbicide. The herbicide cinosulfuron was concluded to negatively affect only a few aspects of the microbial community in soil ecosystems, even at concentrations higher than those currently in use. Baćmaga and coauthors [37] also reported on the absence of adverse effects of the herbicide metazachlor at the recommended dose (0.3 mg/kg) on soil microorganisms including oligotrophic bacteria,
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2,4-D | No effect | Stimulation | [38] |
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Glyphosate | No effect | nd | [35] |
Stimulation | nd | [39] | |
No effect | No effect | [40] | |
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Cinosulfuron | No effect | Temporary growth inhibition or no effect | [31] |
Imazethapyr | No effect | Inhibition | [38] |
Metsulfuron-methyl | Stimulation | nd | [41] |
Nicosulfuron | Inhibition | Inhibition at intermediate doses, no effect at high doses | [42] |
Sulfosulfuron | Stimulation | Inhibition | [43] |
No effect | nd | [44] | |
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Glufosinate | Inhibition | Inhibition | [45] |
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Atrazine | No effect | nd | [35] |
Isoproturon | Inhibition | nd | [46] |
Inhibition | nd | [44] | |
Linuron | No effect | No effect | [47] |
Metribuzin | No effect | nd | [44] |
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Bentazon | No effect | nd | [35] |
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Brominal | Inhibition | Inhibition | [32] |
Temporary inhibition | Temporary inhibition | [48] | |
Sulfentrazone | No effect | nd | [36] |
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Clodinafop | No effect | nd | [44] |
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Mesotrione | No effect | Slightly modified the fungal genetic structures | [33] |
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Alachlor | No effect | Inhibition | [49] |
Butachlor | Stimulation | Inhibition | [50] |
Metazachlor | No effect | Inhibition | [37] |
Napropamide | Inhibition followed by stimulation | nd | [51] |
The negative influence of the herbicides on fungi was also reported by Kucharski and Wyszkowska [43], who tested herbicide Apyros 75 WG (a.i. sulfosulfuron), and by Zhang and coauthors [38], who studied the effect of imazethapyr in two agricultural soils. The ratio of fungi/bacteria in the imazethapyr-treated soil tended to decrease in the initial 15 d incubation period when compared to the control, and then recovered after 30 d of incubation. Stimulation of bacterial and suppression of fungal population due to isoproturon application was reported by Nowak et al. [46]. Omar and Abdel-Sater studied the effect of soil treatment with brominal on population counts of bacteria, actinobacteria, and cellulolytic fungi in soil and found out that the herbicide significantly decreased the total number of cellulolytic fungi and most fungal species while bacterial populations in soil treated with the herbicide was promoted at field application rates and inhibited only at higher levels [32]. Pampulha et al. demonstrated a significant decrease of soil bacteria, fungi, and actinobacteria populations 40 days after glufosinate application [45].
The evidences of no effect or positive effect of the herbicides on fungi growth were also numerously demonstrated. Araújo et al. [39] proved that soil pollution with glyphosate increased populations of fungi and actinobacteria while depressing counts of the other bacteria. Kucharski and Wyszkowska [43] demonstrated a stimulating effect of sulfosulfuron on fungi in the objects treated with the recommended dose of the herbicide 8.9 μg/kg. Treatment of soil with 2,4-D butyl ester at the extremely high dose of 1000 mg/g caused a decline in culturable microbial counts, with the exception of fungal numbers, which increased over the incubation time [34]. At that, when herbicide concentration increased, the Gram-negative/Gram-positive bacteria ratio decreased dramatically in the studied soils. Soil treatment with linuron at the dosages of 4–400 mg/kg did not change the fungal numbers significantly in two agricultural soils as compared to the corresponding controls [47]. Sørensen et al. explained the observed phenomenon by the presence of linuron-degrading fungi, including different species of
He et al. studied the effects of metsulfuron-methyl on soil microorganisms by the method of microbial inoculation culture and found an inhibiting effect of the herbicide on the aerobic heterotrophic bacteria, whereas the number of tolerant fungi increased greatly in the rhizosphere after the application of metsulfuron-methyl [41]. Impact of another sulfonylurea herbicide, nicosulfuron, on the structure, abundance, and function of the soil microbial community using standardized methodologies (PLFAs, taxa-specific qPCR, and enzyme activities) was investigated by [42]. Soil concentrations of nicosulfuron exceeding 1 μg/g resulted in significant reduction of the total PLFAs, although significant reductions of the bacterial PLFAs were observed only at nicosulfuron concentration levels above 10 μg/g. A different picture was evident for fungal PLFAs with significant reductions observed only at intermediate herbicide concentration levels (1–10 μg/g) compared to the control. Besides, qPCR analysis demonstrated that fungi showed the highest sensitivity to nicosulfuron and their abundance was reduced even at the lowest concentration levels of the herbicide (0.25–1 μg/g). Finally, field experiments showed that nicosulfuron applied to the field at dose rates ×1, ×2, and ×5 of the recommended did not significantly affect either the soil microbial biomass or the abundance of fungi and bacteria or enzymatic activity. No significant changes in fungal numbers due to clodinafop introduction into the soil were observed by [43]. Wardle and Parkinson [40] reported that bacterial propagules were temporarily enhanced while actinobacteria and fungal propagule numbers were unaffected by glyphosate. Min et al. [50] reported the influences of the herbicide butachlor on microbial populations, respiration, nitrogen fixation, and nitrification and on the activities of dehydrogenase and hydrogen peroxidase in paddy soil. The results showed that the number of actinobacteria declined significantly after the application of butachlor at different concentrations ranging from 5.5 to 22 mg/kg, while that of the other bacteria and fungi increased. However, at higher butachlor concentrations the growth of fungi was retarded, and the growth of anaerobic hydrolytic fermentative bacteria, sulfate-reducing bacteria, and denitrifying bacteria was stimulated. Treatment of soil with another acetanilide herbicide, napropamide, resulted in decrease of populations of bacteria, while the populations of fungi displayed the decreasing, recovering, and increasing trend [51].
Detailed examination of the observed effects of the herbicides on soil fungi associated with the mode of action of herbicides (Table 1) does not reveal any interrelationships between herbicide identity and their toxicity to fungi. Though herbicides inhibiting amino acid synthesis (ALS and glutamine synthetase inhibitors), contact herbicides inhibiting PPO, and seedling growth inhibitors are seemingly the most toxic, a detailed systematic study needs to be conducted to prove or disprove this observation. Moreover, currently, no general pattern of soil microbiota responses has been inferred regarding herbicide doses applied, exposure time, soil type, or other environmental factors [40,54]. The latter results very likely from the fact that up to now most studies dealing with pesticide soil microbial toxicity were performed using methods that were not well standardized, which did not allow their comparative meta-analysis, and focused on the independent assessment of effects on population, diversity, or functional endpoints, which did not provide a comprehensive view of the toxicity of the pesticide [42]. Standardization of the advanced methodologies available in soil microbial ecology is a necessary step toward harmonization of datasets and is a prerequisite for their integration in the regulatory framework of pesticide soil microbial toxicity assessment [55]. Standards for a number of methods have been already developed and others are under development at the International Standard Organisation (ISO) by TC190/SC4/WG4 and can be found elsewhere [42]. These include:
Measurement of enzyme activity patterns in soil samples using fluorogenic substrates in micro-well plates (ISO/TS 22939)
Determination of soil microbial diversity. Part 1: method by phospholipid fatty acid analysis (PLFA) and phospholipid ether lipids (PLEL) analysis (ISO/TS 22843 part 1)
Determination of soil microbial diversity. Part 2: method by phospholipid fatty acid analysis (PLFA) using the simple PLFA extraction method (ISO/TS 22843 part 2)
Method to directly extract DNA from soil samples (ISO11063)
Estimation of abundance of selected microbial gene sequences by quantitative real-time PCR from DNA directly extracted from soil (ISO/DIS 17601)
Therefore, there is a global need for more complex investigations of the functional diversity responses and degrading activity of soil microbial communities in order to provide deeper insight for herbicide risk assessment. The combined utilization of the above standardized molecular and biochemical methods that provide data of different resolution levels guarantee an accurate estimation of pesticide-driven effects on soil microbes [42].
4. Transformation of the herbicides by white-rot-fungi
It is well documented that a wide range of pollutants including pesticides are transformed and degraded by WRF: pentachlorophenols, isoproturon, derivative of isoxaflutole, atrazine, simazine, propazine, lindane, atrazine, diuron, terbuthylazine, metalaxyl, DDT, dieldrin, aldrin, heptachlor, chlordane, etc. [11,56-65]. This list may be expanded given the strong evidence for WRF degradation potential toward different classes of pollutants. The data on herbicide degradation by WRF are summarized partly in Table 2. It should be mentioned that a large number of works were performed using stationary conditions on liquid media and solid system fermentation conditions. However, there are contradictory data about level of herbicide degradation, role of ligninolytic enzymes in this procedure, and mechanism of degradation as well.
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Type | Days | ||||
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Atrazine | Stat | 42 | 40 | [11] |
Diuron | 42 | 70 | |||
Terbuthylazine | 42 | 60 | |||
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Atrazine | Stat | 42 | 16 | [11] |
Diuron | 42 | 10 | |||
Terbuthylazine | 42 | 37 | |||
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Atrazine | Sub | 40 | 83 | [66] |
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Atrazine | Sub | 40 | 78 | [66] |
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Atrazine | Sub | 40 | 88 | [66] |
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Atrazine | Sub | 40 | 91 | [66] |
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Atrazine | Stat | 42 | 86 | [11] |
Chloronitrofen | 12 | 30 | [67] | ||
Diuron | 42 | 99 | [11] | ||
Nitrofen | 12 | 80 | [67] | ||
Terbuthylazine | 42 | 63 | [11] | ||
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Atrazine | Stat | 42 | 25 | [11] |
Diuron | 42 | 21 | |||
Terbuthylazine | 42 | 52 | |||
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Diuron | Stat | 42 | 6 | [11] |
Terbuthylazine | 42 | 30 | |||
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Bentazon (5 mM) | Stat | 10 | 88 | [68] |
Bentazon (20 mM) | 10 | 55 | [69] | ||
Bentazon (50 mM) | Sol | 10 | 90 | [68] | |
Diuron (30 μM) | Stat | 10 | 55 | [69] | |
Picloram | 10 | 0 | [12] | ||
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Atrazine | Stat | 42 | 57 | [11] |
Diuron | 42 | 71 | |||
Terbuthylazine | 42 | 97 | |||
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Atrazine | Stat | 14 | 0 | [57] |
Stat | 10 | 60 | [70] | ||
42 | 20 | [11] | |||
Bentazon | Sol | 33 | 55 | [71] | |
20 | 65 | [72] | |||
Diketonitrile (derivative of isoxaflutole) | Stat | 15 | 42 | [74] | |
Diuron | Stat | 10 | 94 | [75] | |
42 | 3 | [11] | |||
Isoproturon | Bio-beds | 28 | 78 | [76] | |
100 | >99 | [76] | |||
MCPA | Sol | 20 | 75 | [73] | |
Propazine | 8 | 45 | [70] | ||
Simazine | 8 | 5 | [70] | ||
Terbuthylazine | 42 | 53 | [11] | ||
8 | 95 | [70] | |||
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Atrazine | Stat | 42 | 15 | [11] |
Diuron | 42 | 12 | |||
Terbuthylazine | 42 | 30 | |||
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Atrazine | Stat | 42 | 57 | [11] |
Diuron | 42 | 80 | |||
Terbuthylazine | 42 | 88 | |||
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Picloram | Stat | 10 | 0 | [12] |
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Diketonitrile (derivative of isoxaflutole) | Stat | 15 | 34 | [74] |
Several fungi, such as
White-rot fungi
It was shown that
Hiratsuka et al. [67] reported that
Based on the result obtained, the authors assumed that cytochrome P450 played an important role in lowering the ionization potential of environmentally persistent aromatics and in providing suitable substrates for ligninolytic one-electron oxidizing enzymes for effective degradation. When diphenyl ether, 4-chlorodiphenyl ether, and 4-nitrodiphenyl ether were added to the fungal culture, 4-hydroxydiphenyl ether, 4-chloro-4′-hydroxydiphenyl ether, and 4-nitro-4′-hydroxydiphenyl ether were identified as the major products, respectively. 4-chlorophenol and 4-nitrophenol were detected in trace amounts from 4-chlorodiphenyl ether and 4-nitrodiphenyl ether, respectively, but the counterpart hydroquinone was not observed. These data suggest that the formation of phenolic products from either the A or B ring of CNP might be derived via a different pathway, and that the direct ether cleavage might not have occurred. These findings gave evidence that fungi degraded herbicides via different pathways using their multiple metabolic systems.
The comparative study of herbicide bentazon degradation by
The most studied WRF is
To analyze data presented in Table 2, rate of herbicide disappearance was calculated as the ratio of disappearance (%) to the duration of degradation (days), followed by an average value calculation for every herbicide (Fig. 1). Taking into consideration the effect of cultivation conditions on herbicide degradation by fungi, only data on stationary conditions on liquid media were treated this way.
Obtained results correspond well to the study [70], where it was established that the presence of alkyl groups is necessary for the degradation of s-triazine herbicides by
The contradictory data about participation of ligninolytic enzymes in the herbicide degradation and transformation did not allow establishing their precise role in these processes [18,75,81,84,85,86]. We summarized the data about efficiency of individual ligninolytic enzymes, their mixtures, and enzymes – redox-mediator systems in herbicide degradation in Table 3. As can be seen, no degradation of diketonitrile, diuron, atrazine, chloronitrofen, nitrofen, glyphosate was observed for MnP and LiP crude extracts and purified enzymes from
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Laccase | Atrazine | No | 25°C, pH 4.5 | 240 | 0 |
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Koroleva & Gorbatova (unpublished data) |
[Ru(bpy)2Cl2] | 0 | ||||||
[Ru(phpy)(phen)2]PF6, | 0 | ||||||
HBT | 70 | ||||||
Syringaldezine | 0 | ||||||
Bentazon | Catechol | 25°C, pH 4.0 | 0.5 | 100 |
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[88] | |
Chloronitrofen | No | 0 |
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[67] | |||
HBT | 0 | ||||||
Diketonitrile (derivative of isoxaflutole) | ABTS | pH 3.0 | 0.3–0.4 nmol /(h unit) |
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[74] | ||
Dymron | No | 37°C | 24 | 0 |
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[89] | |
ABTS | 60°C | 24 | >90 | ||||
HBA | 90 | ||||||
MeHBA | 90 | ||||||
NNDS | >90 | ||||||
Glyphosate | No | pH 6.0, Mn2+ + H2O2 + Tween 80 | 24 | 90 |
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[81] | |
No | pH 6.0, Mn2+ + Tween 80 | 90 | |||||
Nitrofen | No | 0 |
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[67] | |||
HBT | 0 | ||||||
Laccase, immobilized | Chloroxuron | No | 30°C, pH 4.5 | 0.5 | 80 |
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[86] |
3-HAA | 0.5 | 80 | |||||
HBT | 0.3 | 100 | |||||
Syrinaldehyde | 0.5 | 80 | |||||
LiP | Atrazine | No | 30°C, pH 5, veratryl alcohol + Mn2+ + H2O2 | 1 | 0 |
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[70] |
Bentazon | No | pH 3.5, veratryl alcohol + H2O2 | 4 | ∼100 |
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[71] | |
Chloronitrofen | No | 0 |
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[67] | |||
No | 0 |
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Glyphosate | No | pH 3.0, veratryl alcohol + Mn2+ + H2O2 + Tween 80 | 24 | 0 |
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[81] | |
Nitrofen | No | 0 |
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[67] | |||
No | 0 |
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MnP | Atrazine | No | 30°C, pH 5, veratryl alcohol + Mn2+ + H2O2 | 1 | 0 |
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[70] |
Bentazon | No | pH 4.5, Mn2+ + Tween 80 | 168 | ∼700 |
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[71] | |
Chloronitrofen | No | 0 |
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[67] | |||
Glyphosate | No | pH 4.5, Mn2+ + H2O2 + Tween 80 | 24 | 100 |
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[81] | |
No | pH 4.5, Mn2+ + Tween 80 | 100 | |||||
Irgarol 1051 | No | 30°C, Mn2+ + glucose + glucose oxidase | 24 | 37 |
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[87] | |
Nitrofen | No | 0 |
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[67] | |||
Pesticide Mix 34 | No | 35°C, pH 4.5, Mn2+ + H2O2 + Tween 80 | 144 | 20-100 |
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[81] | |
Lac+MnP | Bentazon | ABTS | Mn2+ + H2O2 + Tween 80 | 24 | 98 |
|
[69] |
LiP+MnP | Atrazine | No | 39°C, veratryl alcohol + Mn2+ + H2O2 | 24 | 0 |
|
[57] |
No | 30°C, pH 5, veratryl alcohol + Mn2+ + H2O2 | 1 | 0 | [70] | |||
Diketonitrile (derivative of isoxaflutole) | No | 30°C, pH 3 or 5, H2O2 | 12 | 0 |
|
[74] | |
1-HBT | 0 | ||||||
3-HAA | 0 | ||||||
ABTS | 0 | ||||||
Diuron | No | pH 3.0, veratryl alcohol + Mn2+ + H2O2 | 24 | 0 |
|
[75] |
In the study of atrazine degradation with purified laccase from
During the nonenzymatic stage, a product consisting of atrazine and HBT is formed. As the substrates and the products in the “Atr/HBT” system are in equilibrium, the addition of laccase to the reaction causes the oxidation of HBT and the formation of HBT radical. The HBT radical reacts with the Atr-HBT compound and triggers the dissociation of the (-NH-CH-) bonds, resulting in the formation of DEA-HBT and ethyl alcohol. In turn, DEA-HBT decomposes to form two products: DEA and HBT. The ability of HBT to form tautomeric forms and to directly react with atrazine suggested that HBT would degrade in the reaction mixture. However, under the proposed scheme, during the hydrolysis of DEA-HBT, DEA and HBT formed. This may be one of the reasons for the effectiveness of HBT as a redox-mediator in laccase – redox-mediator system.
The high potential of WRF as well as their ligninolytic enzymes in herbicide transformation is well documented. Nevertheless, the mechanisms of degradation and degradation pathways for many herbicides are still not explored. Further studies are needed to elucidate the mechanism of herbicide degradation by WRF and ligninolytic enzymes and identify the metabolites formed.
5. Bioremediation technologies based on application of white-rot-fungi or their extracellular enzymes
The increasing use of agricultural chemicals including herbicides results in the accumulation of these compounds and their derivatives in soil and water. Many herbicides have medium- to long-term stability in soil and so their persistence has a significant impact on the functioning of soil ecosystems. Biological decomposition of herbicides is the most important and effective way to remove these compounds. Therefore, bioremediation is now regarded as a promising strategy for the rehabilitation of polluted environments because of its cost efficiency and environmental friendliness. A detailed examination of the advantages and the disadvantages of bioremediation as well as comparison of bacteria and white-rot fungi in terms of their usage for bioremediation can be found in [9,90]. Filamentous fungi in general and white-rot fungi in particular are generally more tolerant to high concentrations of organic and inorganic toxicants as compared to bacteria [8,91]. On the other hand, white-rot fungi possess great powers of endurance under environmental stresses [91-93]. Finally, white-rot fungi are unique among eukaryotic or prokaryotic microorganisms, because they possess a very powerful extracellular oxidative lignin-modifying enzyme system, which has broad substrate specificity and is able to oxidize a fair amount of organic pollutants [91]. So, white-rot fungi are likely to be powerful prospective agents in soil bioremediation technologies [90,91]. Table 2 gives some examples of white-rot fungi that have been demonstrated to be able to degrade herbicides effectively.
Currently, more than ten species of white-rot fungi can be considered as the effective degraders of different herbicides (Table 2). Among them,
In spite of high degradation potential of white-rot fungi demonstrated in lab settings, fungi are rarely agents of choice for environmental biotechnology. The most important problem is that many research studies examine only destruction of single xenobiotic, whereas in reality mixtures of xenobiotics differing in their structure and mode are subjects for detoxification in the environment [90]. The latter can be toxic for the fungi, resulting in significant inhibition of their growth and, in turn, in the target herbicide degradation. For example, Maceil and coauthors studied effects caused by picloram on the white-rot fungi
Bending et al. [11] studied degradation of the herbicides diuron, atrazine, and terbuthylazine in the so-called biobeds inoculated with white-rot fungi. Biobeds are on-farm pesticide bioremediation constructions developed in Sweden to retain pesticide spills occurring during filling the spraying equipment and facilitate natural attenuation and are currently being evaluated in a number of other European countries [96]. Biobed matrix was prepared by mixing together barley straw, topsoil, and compost [97]. When
To confine white-rot fungi within the toxic environment, a new methodology, which uses growing on potato dextrose agar only or dextrose agar enriched with adsorbent materials, was explored for the removal of xenobiotics from wastewaters [98-100]. This methodology, assuming combined adsorption of organic toxicants followed by their removal, debarred mycelium entrance in the contaminated medium, and excreted fungal enzymes could degrade only the contaminants that entered the medium. The advantage of this methodology is that it avoids additional contamination of the environment with fungal hyphae and exudates, scarce aeration for fungal activity, the continuous contaminant supplying for fungal activity, and the fungus can be easily removed with the agar medium. The developed methodology was successfully employed for simultaneous removal of five coexisting xenobiotics including herbicide linuron from wastewaters, using isolates of
An approach assuming an introduction preliminary inoculated matrix rather than fungal inoculum itself seems to be very promising with respect to the contaminated soil as well. Recently, some companies have included the use of ligninolytic fungi for soil remediation into their programs, for example, “EarthFax Development Corp.” in the USA and “Gebruder Huber Bodenrecycling” in Germany [90]. EarthFax Engineering, Inc. and its affiliate EarthFax Development Corp. have demonstrated the degradation of polychlorinated dibenzo-p-dioxins (PCDDs) and polychlorinated dibenzo furans (PCDFs) in soil under pilot-scale conditions through the use of sawdust thoroughly colonized with the white-rot fungus
Although the mechanisms involved in herbicide degradation by white-rot fungi are not clearly understood, most scientists emphasize the role of the extracellular enzymes of LMS in the degradation of the herbicides by WRF [8,9,103,104]. An alternate pathway of detoxification is the use of a cytochrome P450 monooxygenase system, independent of the production of ligninolytic peroxidase enzymes [105]. To date, the latter was clearly proved only for the fungus
The three principal classes of these enzymes, namely lignin peroxidases, manganese peroxidases, and laccase, are likely able to degrade not only phenols, chlorophenols, and aromatic amines but also non-phenolic compounds such as phenylureas, phenylamides, and s-triazines [100], and the presence of redox active mediators can enlarge the range of compounds that could be oxidized by these enzymes [106]. Table 3 gives some examples of herbicide degradation by the above enzymes. Laccase from
Although application of LiP, MnP, and laccase for degradation many organic pollutants including aromatic compounds, pentachlorophenol, dyes, chlorophenol, urea derivatives, etc., is well known [21,107], only a few papers concerning herbicide degradation specifically are available. Bollag suggested that it is possible to enhance the natural process of xenobiotic binding and incorporation into the humic substances by adding laccase to the soil [108]. Chlorinated phenols and anilines were transformed in soil by oxidative coupling reactions mediated by laccase or peroxidase [109]. The herbicide bentazon was incubated with laccase or peroxidase in the presence of guaiacol, which was used as a model humic monomer. Although bentazon did not react significantly with guaiacol in the presence of the enzymes solely, the reaction of the herbicide with guaiacol was almost complete in 30 min in the presence guaiacol and ferulic acid, which are the electron donor co-substrates in most of the oxidative coupling reactions [88]. Laccase from
Pizzul et al. conducted degradation tests using purified MnP from
However, real contaminated environments contain usually a wide number of different chemical species, some of which can inhibit fungal growth and/or reduce enzymatic activity [106]. To preserve the enzyme’s activity and stability over time, immobilization of the enzyme can be used. Immobilized enzymes have usually a long-term and operational stability, being very stable toward physical, chemical, and biological denaturing agents. Furthermore, they may be reused and recovered at the end of the process [85,91]. Immobilization of laccase from
Both LiP, MnP and laccase may behave as powerful catalysts in the biodegradation of herbicides. However, their full-scale application for remediation of polluted environments is still limited. The latter may derive from several drawbacks and disadvantages of the enzymes application such as enzyme instability in the environment and loss of their activity. Immobilization of the enzymes is likely to be a promising way to develop a successful approach for the remediation of the herbicide polluted sites.
6. Conclusion
The high potential of WRF as well as their ligninolytic enzymes in herbicide transformation is analyzed in the present review. Analysis of literature data on degradation rate of herbicides by WRF demonstrated enhancing WRF degradation capacity along with increase content of branched alkyl groups in the herbicide molecule. However, detailed quantitative structure–degradation activity studies should be conducted to prove or disprove this preliminary observation. Therefore, the mechanisms of herbicides degradation by WRF for many herbicides are still not explored and degradation pathways are not established, including the identification of the metabolites formed.
The ligninolytic enzymes MnP and laccase were shown to behave as powerful catalysts in the biodegradation of herbicides. However, their full-scale application for remediation of polluted environments is still limited. The latter may derive from several drawbacks and disadvantages of the enzymes application such as enzyme instability in the environment and loss of their activity. Immobilization of the enzymes is likely to be a promising way to develop a successful approach for the remediation of the herbicide polluted sites.
The potential of ligninolytic enzymes in the degradation of herbicides is beginning to be characterized at the molecular level. The constant progress in molecular and genomic techniques has provided new insights on the role of regulating elements in the differential expression of ligninolytic enzymes in WRF. Further studies will elucidate the mechanisms of ligninolytic enzymes’ transcriptional regulation and provide deeper understanding of this complicated process.
It should be noted that efficient recommendations for microbial remediation need integral knowledge about potential of individual enzymatic reactions and specific features of their interactions for different microbial species. Current information about genetic regulation of coupled reactions may improve significantly bioremediation technologies, as well as empiric data regarding multistep detoxification with the use of different microorganisms.
Analysis presented in this review confirms the important role of white-rot fungi as participants in herbicide decontamination in the environment and the prospects of the development of new biotechnological preparations on the basis of fungal enzymes. The most important tasks in the development of bioremediation technologies and recent results of key stakeholders in this field are discussed.
7. Abbreviations
ABTS – 2,2'-azinobis(3-ethylbenzthiazoline-6-sulfonic acid)
ACC – acetyl-CoA carboxylase
ALS – acetolactate synthase
DEA – deethylatrazine
DCPMU – 1-(3,4-dichlorophenyl)-3-methylurea
DCPU – 1-(3,4-dichlorophenyl)urea
1H-NMR – proton nuclear magnetic resonance
HPPD – p-hydroxyphenylpyruvate dioxygenase
HR – herbicide-resistant crop
EPSP – 5-enolpyruvylshikimate-3-phosphate
LiP – lignin peroxidase
LMS – lignin modifying system
MnP – Mn peroxidase
PCDD – polychlorinated dibenzo-p-dioxin
PCDF – polychlorinated dibenzo furan
PCR – polymerase chain reaction
PLEL – phospholipid ether lipids
PLFA – phospholipid fatty acid
PPO – protoporphyrinogen oxidase
PSII – photosystem II
ROS – reactive oxygen species
WRF – white-rot fungi
XRE – xenobiotic responsive element
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