Äîêóìåíò âçÿò èç êýøà ïîèñêîâîé ìàøèíû. Àäðåñ îðèãèíàëüíîãî äîêóìåíòà : http://www.mgumus.chem.msu.ru/publication/2011/2011-kulikova-use.pdf Äàòà èçìåíåíèÿ: Mon Jan 20 15:02:53 2014 Äàòà èíäåêñèðîâàíèÿ: Thu Feb 27 20:55:21 2014 Êîäèðîâêà: Ïîèñêîâûå ñëîâà: space station

ISSN 0003 6838, Applied Biochemistry and Microbiology, 2011, Vol. 47, No. 6, pp. 565­579. © Pleiades Publishing, Inc., 2011. Original Russian Text © N.A. Kulikova, O.I. Klein, E.V. Stepanova, O.V. Koroleva, 2011, published in Prikladnaya Biokhimiya i Mikrobiologiya, 2011, Vol. 47, No. 6, pp. 619­634.

Use of Basidiomycetes in Industrial Waste Processing and Utilization Technologies: Fundamental and Applied Aspects (Review)
N. A. Kulikovab, O. I. Kleina, E. V. Stepanovaa, and O. V. Korolevaa
a

Bach Institute of Biochemistry, Russian Academy of Sciences, Leninskii pr. 33, Moscow, 119071 Russia b Faculty of Soil Science, Moscow State University, Moscow, 119992 Russia e mail: evst@inbi.ras.ru, koroleva@inbi.ras.ru
Received December 16, 2010

Abstract--This review provides an analysis of recent data on the mechanisms of degradation of lignocellu losic materials and xenobiotics by basidiomycetes. Special attention is given to the analysis of the current state of research of ligninolytic enzymes and their involvement in the degradation of xenobiotics. Data on the prac tical use of basidiomycetes for bioconversion of industrial wastes are systematized. The most promising areas of bioconversion technologies are considered, such as contaminated water purification (including wastewa ter), cleanup of soils contaminated with heavy metals and xenobiotics, and degradation of difficult to degrade substrates (lignin and lignocellulose wastes, low energy coal, and synthetic polymers). DOI: 10.1134/S000368381106007X

INTRODUCTION Basidiomycetes are higher fungi with multicellular mycelium numbering approximately 30 thousand spe cies of both microscopic fungi and fungi with large fruiting bodies. Although basidiomycetes are found in various ecosystems including meadows, steppes, and deserts, they are most diverse and common in forest ecosystems. The main function of basidiomycetes in nature is to decompose lignin and cellulose. This abil ity attracts the attention of researchers both in terms of elucidation of mechanisms underlying this process and in order to develop procedures for biotechnologi cal utilization of wood and plant waste [1­5]. A unique feature of basidiomycetes is the ability to synthesize extracellular enzymes lignin peroxidases, Mn peroxidase, versatile peroxidases, and laccases with a broad substrate specificity [6, 7], which allows them to decompose organic matter not only of natural origin but also various xenobiotics. The most danger ous organic pollutants whose degradation can be accelerated by basidiomycetes are polycyclic aromatic hydrocarbons, chlorophenols, polychlorinated biphe nyls, pesticides, and municipal wastes. To date, the main mechanisms of decomposition of xenobiotics by basidiomycetes are studied sufficiently well, and the use of basidiomycetes as biological agents for process ing and utilization of industrial waste has been consid ered in several reviews [8, 9]. However, new data are constantly being accumulated that clarify the mecha nisms of degradation of xenobiotics by basidiomycetes and describe examples of using basidiomycetes and their ligninolytic enzymes for detoxification and deg radation of pollutants in various industries.

The purpose of this review was to analyze the cur rent state of technology of bioconversion of lignocel lulosic materials and xenobiotics by basidiomycetes. Main pathways of transformation of lignocellulosic materials and xenobiotics by basidiomycetes. Investi gation of decomposition of lignocellulosic materials and xenobiotics by white rot fungi demonstrated the possibility of their use in technologies of processing and utilization of poorly degradable anthropogenic waste. Recent experimental data obtained in this field are summarized in several reviews [2, 6­10]. It was established that degradation of xenobiotics and ligno cellulosic materials by white rot fungi includes the action of a multienzyme complex, whose synthesis depends on the substrate on which the fungus grows, its physiological and biochemical characteristics, and genomic organization. The efficiency of degradation is ensured by a combination of extracellular ligninolytic enzymes, organic acids, mediators, and related enzymes. According to the modern notion, there are three main pathways of decomposition of natural polymers and xenobiotics by basidiomycetes: direct enzymatic degradation, indirect enzymatic degrada tion, and nonenzymatic degradation (Fig. 1). Each of these pathways is characterized by specific mechanism of decomposition of poorly degradable compounds. The enzymatic pathway includes molec ular transformation of the substrate with alteration of its properties and ultimate complete degradation, which are accompanied by de novo synthesis of com pounds. Indirect enzymatic degradation is based on the formation of radicals in the main products and by products of enzymatic reactions, with subsequent trig ger of radical processes. Nonenzymatic degradation is

565


566

KULIKOVA et al.

Natural polymers and xenobiotics

Indirect enzymatic degradation

Direct enzymatic degradation

Nonenzymatic degradation

Generation of radicals, triggering of chain reactions

Molecular transformation with alteration of properties

Transition metal ions (Mn, Cu)

De novo synthesis

Degradation/ mineralization
Fig. 1. Main pathways of degradation of natural polymers and xenobiotics by basidiomycetes.

performed at the expense of reactive radicals and ions of transition metals. Under natural conditions, basid iomycete degradation processes are multi stage and, as a rule, include all the above mechanisms. However, both direct and indirect types of enzymatic degrada tion are performed with the participation of oxi doreductases and hydrolases, which determines the significance of these enzymes in the degradation of xenobiotics and biopolymers. In our opinion, the enzymatic decomposition is of greatest practical importance. Examples to support this assumption will be discussed below. Characteristics of ligninolytic enzymes of basidio mycetes. Basidiomycetes are able to synthesize many extracellular enzymes involved in modification and degradation of lignin. Currently, the common name of these enzymes is ligninases [2, 12], although some authors assign this term to lignin peroxidase [10, 11]. Ligninases can be divided into two groups: phenoloxi dases (laccases (LAC, EC 1.10.3.2)) and heme con taining peroxidases, namely lignin peroxidase (LP, EC 1.11.1.14), manganese peroxidase (MP, EC 1.11.1.13), and multifunctional (versatile) peroxi dase (VP, EC 1.11.1.16) [11, 12]. These two groups of enzymes have different electron acceptors: molecular oxygen for laccase and hydrogen peroxide for heme containing peroxidases (Table 1). Lignin peroxidase (LP). LP is a glycoprotein con taining 1 mol of ferriprotoporphyrin IX per 1 mol of the enzyme and 6 to 20% of carbohydrates (Table 1). The molecular weight (MW) of LP ranges from 39 to

43 kDa, and isoelectric points of isozymes range from 3.0 to 4.5 [13, 14]. Lignin peroxidase was first detected in Phanerochaete chrysosporium in 1983 [15, 16]. Later, the presence of LP was established in various strains of P. chrysosporium and Trametes versicolor [17]. Screening of basidiomycetes revealed the pres ence of LP genes in Panus sp., P. coccineus, P. san guineus, and Perenniporia medulla panis [18]. LP is nonspecific with respect to substrates: it oxidizes a wide range of aromatic substrates of phenolic nature and nonphenolic components of lignin with a redox potential less than 1.4 V (relative to the normal hydro gen electrode) in the presence of hydrogen peroxide. The catalytic cycle of LP is similar to the catalytic cycles of other heme peroxidases (Fig. 2). A unique feature of LP, which distinguishes it from other peroxidases, is the ability to oxidize methoxy lated lignin substructures with high redox potentials. For phenolic substrates, the oxidation rate is higher than for nonphenolic substrates; as a result of oxida tion, phenoxyl radicals are formed. In the presence of oxygen, phenoxyl radicals can interact with various compounds, leading to rupture of the aromatic ring and/or polymerization. Veratryl alcohol (VA), which is produced by ligni nolytic fungi as a secondary metabolite, is of particular importance for the functioning of ligninases. This compound protects ligninase from inactivation by hydrogen peroxide. It can also induce the synthesis of ligninase in culture medium and serves as a redox mediator during the oxidation of various substrates,
Vol. 47 No. 6 2011

APPLIED BIOCHEMISTRY AND MICROBIOLOGY


Table 1. General characteristics of ligninolytic enzymes

Enzyme Mediators

Active site structure

Localization

Catalyzed reaction

MW, pH Glycosylation kDa optimum

Refer ences []

APPLIED BIOCHEMISTRY AND MICROBIOLOGY

Lac

Ensemble of four copper ions: T1 copper center and the copper cluster consist ing of T2 copper ion and T3 antiferromagnetic pair

Intracellular 4 benzenediol+ O2 = and/or extra = 4 benzene semiquinone + 2H2O cellular en zyme

50­70 N glycosyla tion

2­10

ABTS, GBT, TM [2], PO, transition metal [21], complexes [28], [31], [45]

LP

Ferriporphyrin IX

39­43

Extracellular 1. LP[Fe(III)] + H2O2 LP* I[Fe(IV) = O*+] + H2O enzyme 2. LP I+AH LP II[Fe(IV) = O*+] + A*+ LP II+AH LP + A*+

N glycosyla tion

1­5

Veratryl alcohol

USE OF BASIDIOMYCETES IN INDUSTRIAL WASTE PROCESSING

Vol. 47

[8], [10], [11], [20], [21]

No. 6

2011

MP

Ferriporphyrin IX

Extracellular MP + H2O2 = MP I + H2O MP I + Mn2+ = MP II + Mn3+ enzyme MP II + Mn2+ = MP + Mn3+ + H2O

38­ 62.5

N glycosyla 2.5­6.5 tion

Organic acids as ch [2], elators, toles, unsat [8], urated fatty acids [10], [21]

VP

Heme

Extracellular Donor+ H2O2 = Donor (oxidized) + 2H2O 42­45 enzyme

Glycosyla tion type not determined

3­5

Same compounds [2], [8], that for LP and MP [11], [29] 567


568

KULIKOVA et al. LP completely reduced form 2H2O 2VAL Fe
3+

H2O2 H2O

2VA

·+

·+ ·+

VA

Fe4+

O

+·

LP compound I Fe
3+

VA

O2

VA

2H2O H2O2 excess

O

Fe4+

·+

VA

LP compound II

Fig. 2. Catalytic cycle of lignin peroxidase (LP).

including the lignin polymer [19]. During catalysis, cation radicals of veratryl alcohol are formed, which are highly reactive and enter nonenzymatic reactions. Currently, the ability of ligninases to catalyze the following reactions has been established [10­12]: (1) Cleavage of C­C bonds in dimeric lignin models (2) Oxidation of benzyl alcohol. (3) Oxid ation of methyl substituents in benzyl compounds. (4) Hydroxylation of benzyl methyl groups. (5) Hydroxylation of olefinic bonds. (6) Decarboxylation of phenylacetic acid. (7) Cleavage of ester bonds (8) Opening of aromatic rings. (9) Polymerization of phenols. The crystalline structure of LP shows that the heme group is located inside the structure and is bound to the surface through a channel whose diameter is too small for penetration of large polymeric structures of lignin but quite sufficient for penetration of small mol ecules and their subsequent binding [20]. Manganese peroxidase (MP). Manganese peroxi dase, similarly to LP, is a glycoprotein containing pro toheme IX (ferriprotoporphyrin IX), which is easily separated from the apoenzyme during electrophoresis even under nondenaturing conditions. The molecular weight of MP ranges from 38 to 62.5 kDa; the majority of purified enzymes have an MW of ~45 kDa [21]. Basidiomycetes produce a large number of MP iso

forms. For example, 11 isoforms were described for Ceriporiopsis subvermispora [22]. The isoelectric point values of MP vary from 2.5 to 6.8 [23]. Manganese peroxidases are produced by the majority of white rot fungi (families Polyporaceae, Meruliaceae, and Coriolaceae) and certain fungi that inhabit the soil litter (families Strophariaceae and Tri cholomataceae). Today, as many as 56 fungi producing MP are known [23]. MP catalyzes the oxidation of n2+ to n3+ in the presence of hydrogen peroxide. The catalytic cycle of MP in the presence of a chelating agent (oxalate, mal onate, malate, tartrate, and lactate) leads to the for mation of highly reactive n3+­chelator complex, which can oxidize many phenolic substrates by the one electron mechanism, including phenolic com pounds with the formation of lignin phenoxyl radicals (Fig. 3) . The reaction is initiated by the binding of 22 to the native enzyme and formation of an iron­peroxide complex. Subsequent break of the O­O bond leads to the transfer of two electrons and formation of com pound MP I, which is an Fe4+ oxo porphyrin­radical complex. Then, after the bond is broken, one water molecule forms. Further reaction involves the forma tion of compound MP II (Fe4+ oxo porphyrin com plex). The monochelated Mn2+ ion functions as a one electron donor for this porphyrin complex and is oxi dized to Mn3+. The reduction of compound II pro ceeds similarly, and another Mn3+ ion forms from the
Vol. 47 No. 6 2011

APPLIED BIOCHEMISTRY AND MICROBIOLOGY


USE OF BASIDIOMYCETES IN INDUSTRIAL WASTE PROCESSING [R­OOH] H2O2 [R­OH] H2O H2O [R­OH] Fe3+
·

569

Mn Mn3+

Fe4+

O

Mn compound I RH

R·

+

H+

Mn2+ O

Mn2+ Fe
4+

R· + H+ Mn3+ RH

Mn compound II

Fig. 3. Catalytic cycle of manganese peroxidase (MP).

Mn2+, leading to the formation of the native form of the enzyme and release of a second water molecule. Mn3+ ion is stabilized by organic acids, such as oxalate, and functions as a low molecular weight redox medi ator that nonspecifically attacks the substrate with cleavage of the hydrogen ion and one electron. Phe nolic and aminoaromatic compounds are oxidized with the formation of phenoxyl and amino radicals, respectively [23]. The oxidation potential of the Mn3+­chelator complex is insufficient for oxidation of lignin structures of phenol. Nonphenolic substrates can be oxidized by MP only in the presence of a sec ond redox mediator with the formation of reactive radicals. Organic acids, such as oxalate and malonate, function as such redox mediators. In the absence of enzymatic system that generates 22, the formed radicals can be used as by MP a source of hydrogen peroxide and increase the efficiency of lignin degrada tion by the fungus. MP can catalyze bond breaking in nonphenolic lig nin structures at ­ bonds and alkyl­aryl and par ticipate in oxidation of model lignin structures of the syringyl type ­1. In addition, it is believed that MP oxidizes nonphenolic lignin structures by produc ing highly active radicals from unsaturated fatty acids and thiols [24]. The authors of some studies assumed that nonphenolic lignin structures can be oxidized by MP after preliminary removal of methanol from aro matic rings of lignin molecules with the participation of cellobiose dehydrogenase [25].
APPLIED BIOCHEMISTRY AND MICROBIOLOGY

The crystal structure of MP and the structure of its active center (heme) are largely similar to LP. The main difference of the classical peroxidases from LP is the presence of a manganese binding site. The bound Mn2+ ion is coordinated by three amino acid residues, the propionate residue at position 6 of heme and oxy gen atoms of two water molecules. The binding site is located on the surface of the enzyme and is readily accessible [26]. Laccase. Lac is a glycoprotein containing 10 to 45% of carbohydrates per enzyme molecule [27]. Many researchers believe that the carbohydrate moi ety of the molecule provides the conformational sta bility of the protein globule. Fungal laccases have an MW of 50­70 kDa [28] and isoelectric points at pH 3­5 [23, 30, 31]. Laccases were found in fungi, bacte ria, and insects [31]. Today, the main source of the enzyme, including the enzyme used for industrial pur poses, are fungi. A large number of fungi that produce this enzyme are known. The most comprehensively laccase producers are Podospora anserina, Agaricus bisporus, Rhizoctonia practicola, Pholiota aegerita, Trametes versicolor, Pleurotus ostreatus [32], Coriolus hirsutus [33, 34], and Neurospora crassa [35, 36]. All fungal laccases are monomers or dimers, except for isoform 1 of Podospora anserine, which is apparently a tetramer. The majority of fungi produce both intracel lular and extracellular enzyme. The family of laccases that were discovered more than a century ago remains a subject of basic research primarily because the detailed mechanism of action of
No. 6 2011

Vol. 47


570

KULIKOVA et al. LP completely reduced form H2O T2 T3 Cu +4e slowly H2O T2 T3 Cu2+ OH Oxidized form T1 Cu
2+ 1+

Cu1+ O2 Cu1+ Cu1+ k = 2â105
­1 s­1

H2O T1 T2 Cu1+ O Cu2+ Cu2+ +4e rapidly H2O T2 k = 0.03 s
­1

O T3 Cu2+ Cu2+

T1 Cu1+ Peroxide intermediate k 1000 s Cu2+
­1

Cu2+ O

T3 Cu2+ 2H­ OH

H2O

T1 Cu2+ Native intermediate

Fig. 4. Catalytic cycle of laccase (Lac) [28].

the enzyme is still obscure. The catalytic cycle include the oxidation of the substrate (an electron donor) and electron transfer to the T1 center of the enzyme. The transfer of four electrons from the copper ion in the T1 cluster to the T2/T3 cluster is followed by sequential recovery of all three copper ions in the cluster; the copper ion 3, which has the highest electron affin ity, is reduced first. The reduction of 3 is accompa nied by protonation of the 3 oxo center and ligand, with protons dissociating from the reduced 3 center. The next stage is the reduction of the T2 center. The key step in this process is the formation of a "bridge" hydroxyl between T2 and 3, through which electrons can be rapidly transferred to the T2 center. Therefore, this model suggests the formation of a pair of copper ions of a mixed valence. Further reduction of the 3 copper ion is postulated as a rapid process of electron transfer along the cysteine­histidine dipeptide between T1 and T3, which is accompanied by protonation of the bridge hydroxyl followed by dissocia tion of two water molecules from the cluster [28]. Laccases have a broad substrate specificity and cat alyze oxidation of various compounds, including o,p diphenols, aminophenols, polyphenols, polyamines,

lignin, certain inorganic ions, and aryl diamines, with concomitant reduction of molecular oxygen to water [37, 38]. Laccases are capable of direct bioelectroca talysis, that is, direct electron transfer from the elec trode to the active site. It is assumed that laccase oxidizes phenol hydroxyl of substrates to form a phenoxyl radical, which enters the nonenzymatic reactions of demethoxylation of lig nin and methoxyphenolic acids, as well as the reaction of quinone formation and oxidative elimination of carboxyl groups. The structures of laccases isolated from different sources are very similar [39­41]. Molecules of lacca ses are usually monomers consisting of three sequen tially linked cupredoxin like domains folded into a dense globule. The T1 copper center is located in the third domain and is coordinated by two imidazoles of histidine and the sulfhydryl group of cysteine, which form a trigonal structure. It is a component of the sub strate binding "pocket" and is located at a distance of 6.5 å from the protein surface. Usually the T1 center is located at a distance of 12 å from the T2/T3 cluster and is linked with it through the His­Cys­His tripep tide, which is highly conserved among laccases. Trinu
Vol. 47 No. 6 2011

APPLIED BIOCHEMISTRY AND MICROBIOLOGY


USE OF BASIDIOMYCETES IN INDUSTRIAL WASTE PROCESSING

571

Table 2. Possibility of degradation of natural compounds and xenobiotics by LP, MP, and laccase during (1) direct and (2) indirect oxidation LP Compound 1 Lignin and its model components Phenolic lignin components Nonphenolic lignin components (aromatic alcohols) Alcohols Amino acids, proteins Aromatic amines Hydroxyphenylacetic acid and its derivative Unsaturated fatty acids Cinnamic acids Carbohydrates and their derivatives Humic substances Inorganic ions Xenobiotics PAHs PCBs Pesticides Dyes Halogenated phenols Azo compounds, aniline, acrylamide, hydrazine, benzotriazoles Amines (aryldiamines, hydroxylamine) Naphthols Benzene homologues + + + + + + 2 + + + 1 + + + 2 + + + 1 + + 2 + + + + + + + + + + + + + + + + + + MP Laccase

+ + + + + + + + + + + + + + + + + + + + + + + + + + + +

+ + + + + + +

+ + + +

clear T2/T3 cluster is located between the first and third domains and has amino acid ligands in each of them. The three copper ions of oxygen reducing T2/T3 cluster form an almost equilateral triangle with distances 3.7 to 5.1 å. The T2 copper ion has two N2 ligands from two histidine residues and one oxygen ligand O2, which form a trigonal planar configuration. Water molecule or the OH group may serve as oxygen ligands. Each copper ion of the T3 pair is coordinated by three histidine residues and an oxygen ligand, which is located between two T3 ions. The coordina tion of each of them can be described as a distorted tet rahedron. Versatile peroxidase (VP). Polyfunctional peroxi dase is a glycoprotein exhibiting hybrid properties of LP and MP. There is still confusion about the defini tion of these enzymes: sometimes they are called hybrid peroxidases and sometimes they are abbrevi ated. Currently, the group of VP includes enzymes that catalyze the oxidation of typical peroxidase substrates including Mn2+ and veratryl alcohol. Versatile peroxi dases were isolated from Bjerkandera adusta, Bjerkan dera sp. (BOS55), Bjerkandera sp. (B33/3), B. fumosa, Pleurotus eryngii, P. ostreatus, and P. pulmonarius [42,
APPLIED BIOCHEMISTRY AND MICROBIOLOGY

43]. This group of enzymes is very attractive in terms of practical use because of their ability to oxidize Mn2+ as well as phenolic and nonphenolic aromatic com pounds. It is assumed that VP can oxidize a wide range of substrates with different potentials--from low to high, comparable to those for LP. Versatile peroxidases are more effective than LP and MP, which are not able to effectively oxidize phenolic components in the absence of veratryl alcohol and to oxidize phenols in the absence of Mn2+, respectively. This substrate spec ificity is determined to their hybrid molecular struc ture. The catalytic cycles of MPs are similar to those of VP and LP [29]. As in other heme containing peroxidases, heme in VP is located within the globule and is connected with the surface by two channels. The function of the first channel, which is highly conserved in heme peroxi dases, is similar to that of LP, and the second channel is characteristic of VP, and MP and serves for the oxi dation of Mn2+ to Mn3+. Biodegradation of biopolymers and xenobiotics with participation of enzymes of the ligninolytic complex. Degradation of biopolymers and xenobiotics in nature under the influence of ligninolytic enzymes produced
No. 6 2011

Vol. 47


572

KULIKOVA et al.

by basidiomycetes is a process whose intensive study is primarily determined by the need to design environ mentally friendly biotechnology. As a result, ample factual material on biodegradation with LP, MP, and laccase by both direct and indirect oxidation pathways has been accumulated to date. The list of compounds degraded by direct and indirect oxidation by lignin peroxidase, MP, and laccase is presented in Table 2. Analysis of published data led us to conclude that a compound can be directly oxidized by a ligninolytic enzyme only if it can be a substrate of this enzyme (judging by its chemical structure) and if its redox potential is below the redox potential of the enzyme [11]. For example, the comparison of the effectiveness of oxidation of a homologous series of methoxyben zenes with different redox potentials (from 0.81 to 1.76 V at pH 3.0) by horseradish peroxidase, LP, MP, and lac case showed that, out of the 12 compounds tested, ten are oxidized by LP four (redox potentials, 0.81­1.12 V) , are oxidized by horseradish peroxidase and MP, and only one compound (1,2,4,5 tetramethoxybenzene; redox potential, 0.81 V) is oxidized by laccase [44]. Thus, LP oxidizes a wide range of aromatic com pounds with redox potentials less than 1.4. By the increase in the efficiency of substrate oxidation, ligni nolytic enzymes can be arranged in the following order: laccase, MP, and LP. However, the situation changes when the possibility of using the enzyme­ redox mediator system for degradation and/or detox ification of biopolymers and xenobiotics is considered. The effectiveness of such a system is determined by the stability of the enzyme during catalysis and by the redox properties of the mediator, including the stabil ity, lifetime, and reactivity of generated free radicals. The most effective enzyme­redox mediator system is the laccase­redox mediator system [45]. This is pri marily due to the fact that hydrogen peroxide inacti vates all heme peroxidases after several catalytic cycles are completed. Laccase that uses molecular oxygen as a cosubstrate is quite stable. In addition, the pH and thermal inactivation of ligninolytic peroxidases, which is associated with the release of two Ca2+ ions from the enzyme molecule, also reduces the effectiveness of such a system. Versatile peroxidase is of special interest because of catalytic multifunctionality--the ability to degrade a wide range of compounds in reactions of direct oxida tion that cannot be oxidized by LP and VP. Recently, it was demonstrated that VP effectively degrades polycy clic aromatic hydrocarbons [42]; pollutants of phe nolic and nonphenolic nature [46]; pesticides [47]; industrial dyes [48]; Active Blue 38 and other azo dyes; Active Black 5 and other phthalocyanine pigments; anthracene and its derivatives; benzopyrene; pyrene; 2,4 dichlorophenol; and pentachlorophenol. How ever, all limitations on the use of peroxidases in xeno biotic degradation technologies also apply to VP. Thus, currently, the most promising system to be used

in detoxification and degradation technologies is the laccase­redox mediator system. Nonenzymatic degradation of lignocellulosic mate rials and xenobiotics by basidiomycetes. The nonenzy matic pathways of degradation of polysaccharides, lig nin, and xenobiotics has been attracting the attention of researchers for decades. All mechanisms of nonen zymatic degradation are based on radical processes [1, 11, 12]. Similarly to the processes of degradation of lignocellulosic materials, the first stage of degradation of xenobiotics by basidiomycete may include the pro duction of highly reactive low molecular weight com pounds that function as oxidants. Such compounds are involved in the "treatment" of wood and ensure the availability of wood lignin for enzyme attack. The main radicals involved in these processes are hydroxyl radicals (OH*). The main pathways of generating OH* by basidio mycetes are the reactions catalyzed by cellobiose dehydrogenase (CDG), low molecular weight pep tides/quinone redox cycle, and redox reactions cata lyzed by glycopeptides (Fenton reaction catalyzed by glycopeptides). In addition, almost all basidiomycetes have systems generating hydrogen peroxide, which not only is used by enzymes as a cosubstrate but also enters the Fenton reaction to form OH* radicals. The latter attacks lignocellulosic material and/or polysaccha rides of the cell wall and leads to cleavage of biopoly mers and facilitates penetration of ligninases. It can be assumed that xenobiotics are degraded by a similar mechanism. Main directions of use of basidiomycetes in technol ogies of utilization and processing of anthropogenic structures and waste. Currently, basidiomycetes are of most demand in technologies that require decomposi tion of lignin and its modifications. Lignin and ligno cellulosic waste is produced mainly during agricultural activity (straw) and constitutes a considerable part of household as well as wood processing and pulp and paper industry waste [49]. Decomposition of lignin and lignocellulosic agricul tural waste. The most common agricultural waste con taining lignin and lignocellulose is straw. Straw is a valuable organic fertilizer [50]; however, if left in the field and plowed on the spot, it is degraded too long because of the low content of the carbon source and the high content of fiber and organosilicon com pounds. In the arable soil horizon, its residues remain for three to five years. In view of this, to improve utili zation of straw, it was suggested to perform its inocula tion with an association of nitrogen fixing and polysaccharide producing bacteria and basidio mycetes with a high cellulolytic activity. In addition, agricultural lignocellulosic waste is poorly assimilated when fed to cattle [49, 51]. The presence of lignin impedes the access of hydrolytic enzymes--cellulase and hemicellulose--to their sub strates. Preliminary biodelignification of plant feeds is the most promising way to improve their quality. Justi
Vol. 47 No. 6 2011

APPLIED BIOCHEMISTRY AND MICROBIOLOGY


USE OF BASIDIOMYCETES IN INDUSTRIAL WASTE PROCESSING Table 3. Patented methods of pulp and paper mill wastewater treatment using basidiomycetes Fungal species Alternaria alternata Phlebia tremellosa Scytinostroma galactinum Scytinostroma galactinum strain F361 Schizophyllum commune, Trichaptum biforme, Phanerochaete gigantea White and brown rot fungi Degraded compound Water soluble lignin sulfate Lignin and lignin containing resins in paper pulp Degradation of lignin containing wastes, cellulose, and chlorine aromatics Degradation of lignin, cellulose, and chlorine aro matics Lignin and lignin containing resins in paper pulp Degradation of lignin in paper pulp Source [66] [69] [70] [71] [72] [73]

573

fication for such an approach was shown in numerous studies with rice and wheat straw, cotton stalks, and card board [52]. The method of producing feed product based on straw delignified by Panus tigrinus [53] and Pleurotus ostreatus [54] is described in patents [53, 54]. Recycling wastes of wood and wood processing industry. Chips, sawdust, and other wood processing wastes, as a rule, are never used or used only slightly. Large amounts of these raw materials are accumulated over years near wood processing enterprises. A com mon practice for disposal of waste wood using basidi omycetes is to grow edible mushrooms on them. The most widely spread is the Oyster mushroom Pleurotus ostreatus, a delicious and nutritious mushroom, which has earned worldwide recognition due to the relatively simple farming techniques of its cultivation and resis tance to pests and diseases. Among wood wastes, wooden railroad ties pose a particular problem. Their life varies from 6 to 40 years depending on the type of wood of which ties are made, the climatic conditions in the area where the railway line is located, and the extent of the workload of paths. A characteristic feature of this type of wood waste is that they are saturated with the antiseptic creosote, which prevents rotting and, therefore, decreases the possibility of biological degradation. Several methods of decomposition of creosote containing wood wastes were patented abroad. It was suggested to use a range of basidiomycetes, such as Antrodia radiculosa, Merul iparia incrassata, Neolentinus lepideus, Melanoporia niger, Polyporus sp., Crustoderma dryinum, Gloeophyl lum subferrugineum, Phanerochaete sordida, Peniphora pseudopini, and Ceriporia spissa as biological agents [55­57]. Of other lignocellulosic wastes that can be degraded by basidiomycetes, we should mention the utilization of coconut fibers with P. ostreatus [58]; degradation of park waste using Coriolus versicolor, P. ostreatus, and Gano derma applanatum [59]; and degradation of coconut fibers and mulching materials using a consortium of basidiomycetes including P. ostreatus, P. pulmonarius, P. dryinus, P. tuberregium, Piptoporus betulinus, Fomi topsis pinicola, F. officinalis, Trametes versicolor, Hyp
APPLIED BIOCHEMISTRY AND MICROBIOLOGY

sizygus ulmarius, Ganoderma lucidum, G. applanatum, G.curtisii, G. oregonense, and G. tsugae [60]. In conclusion, it should be mentioned that progress in studying the degradation of lignocellulosic waste has made it possible to develop their future application in space flights [49]. It is possible that, in the near future, lignocellulosic waste transportation to the space station will lead to significant cost savings. Lignocellulose can become a raw material for obtain ing everything necessary: fuel, energy, chemical raw materials, food, and water. Experiments conducted within the framework of the program "Closed Ecolog ical Life Support System" (CELSS) showed that the treatment of plant wastes with the white rot fungus P. ostreatus for this purpose is a promising approach [61]. Treatment of pulp and paper mill (PPM) wastewa ter. Enterprises that use molecular chlorine for cellu lose bleaching produce polychlorinated dibenzo n dioxins (PCDDs) and polychlorinated dibenzofurans (PCDFs)--highly toxic carcinogenic compounds that are chlorinated cyclic aromatic esters [62]. The ability to detoxify pulp produced by PPM was shown for a number of basidiomycetes, including Phanerochaete chrysosporium, T. versicolor, Fomes lividus, and Thele phora sp. [63, 64]. In our country, it was also recom mended to use the method of sludge lignin utilization using a wood decomposing basidiomycete Trametes pubescens strain as a biological method of utilization of solid PPM wastes containing chlorinated aromatic compounds [65]. The calculations of the authors of [66] showed that the efficiency of degradation of phe nols and chlorinated compounds contained in the Baikal Pulp and Paper Mill waste by T. pubescens was 100%. In addition to T. pubescens, the method of bio logical purification of pulp and paper industry waste water from water soluble lignin sulfate by Alternaria alternata was patented in the Russian Federation [66]. In most cases, fungal laccase plays the key role in the detoxification of PPM wastewater [62, 67]; the key role of MP was shown only for certain strains (e.g., T. versicolor) [68]. Basidiomycetes belonging to gen era Alternaria, Phanerochaete, Phlebia, Scytinostroma,
No. 6 2011

Vol. 47


574

KULIKOVA et al.

and Trichaptum are used in technologies of biological treatment of PPM wastewater (Table 3). Purification of wastewater from textile dyes. The ability of basidiomycetes to decolorize various dyes that are present in textile industry effluent was studied sufficiently well [43, 48, 74­83]. Analysis of available data showed that discoloration of dyes was demon strated for 31 species of basidiomycetes and 77 dyes and mixtures thereof. Among the basidiomycetes studied, genera Phanerochaete and Trametes were described in the most detail as capable of degradation of a wide range of dyes. However, the preparation based on Flavodon flavus is the only preparation pat ented to date [84], which indicates the necessity of fur ther study of basidiomycetes of genera Phanerochaete and Trametes, which can be used for bioremediation of textile industry wastewater. Purification of wastewater containing heavy metals and radionuclides. Biological treatment of water from heavy metals and radionuclides by using basidio mycetes is based on their ability to rapidly absorb tox icants. The absorption of metals by fungi can occur not only due to adsorption processes, as in the case of bacteria, but also due to active transport of metals into cells [85, 86]. This unique feature makes basidio mycetes, in some cases, the best agents for biological purification of water from metals and radionuclides. In our country, the trend to use fungi in bioremedi ation rapidly developed at the end of XX century, as evidenced from patents that now have ceased to have effect [87, 88]. Ascomycetes (Aspergillus, Penicillium, and Phizopus) were recommended to be used for this purpose as fungi capable of intensive absorption of radionuclides and heavy metals. In recent years, how ever, increasing attention is being paid to studying basidiomycetes as potential biosorbents, apparently due to their lower pathogenicity. The authors of [89] demonstrated that Schizophyllum commune is promis ing for removal of uranium, and the possibility of using Phanerochaete chrysosporium for removal of cadmium was shown in [90]. Since high concentrations of heavy metals in the environment are toxic to basidiomycetes, when selecting a strain for bioremediation, its sensitiv ity to heavy metals should be studied. According to data obtained in [91], high resistance to metals is char acteristic of P. ostreatus, P. cystidosus, Stereum hirsu tum (resistant to Cd and Hg), and T. versicolor (resis tant to Cd, Zn, Ni, Co, Cr, Mo, Pb, Hg, and Sn). The list of basidiomycetes that are able to accumulate dif ferent metals is given in Table. 4. The ability to accu mulate the most broad range of heavy metals is char acteristic of fungi belonging to genera Pleurotus, Trametes, and Phanerochaete, which makes the repre sentatives of these genera the most promising tools in terms of using in technologies of biological treatment of wastewater from heavy metals. Cleanup of environment contaminated with oil hydrocarbons. In the classical scheme of purification from crude oil and petroleum, biological methods are

used only at the final stages of treatment; however, today there is a tendency to replace the multistep puri fication schemes with one step schemes. The devel oped approaches are based on the use of microbial consortia, which include representatives of filamen tous fungi, yeasts, and bacteria, which effectively transform oil components into nontoxic and low toxic substances. At the same time, purification of contam inated media in situ can be performed by maintaining and stimulating the natural oil oxidizing microorgan isms by creating optimal conditions for their develop ment (aeration and introduction of nitrogen and phos phorus fertilizers to the contaminated focus) and by introducing an active strain of the destructor to the contaminated site. To clean the water surface from oil spills, a complex mycosorbent containing strains of ascomycete fungi Fusarium solani, F. moniliforme, Trichoderma har zianum, and Cladosporium resinae was developed in Russia. These fungi are immobilized on hydrophobic carriers and are used as oil sorbents and destructors [113]. To clean soil and water surfaces from crude oil and oil products, a complex product containing asco mycetes (Aspergillus niger) and basidiomycetes (Phan erochaete chrysosporium) was developed and is cur rently used. This preparation is sprayed over the water surface in a mixture with detergents and sorbents [114]. Similar preparations containing Phanerochaete chrysosporium strain and designed to clean up environ ment contaminated with petroleum hydrocarbons was registered in the United States [115]. To clean up soil from oil pollution, biological prep arations containing mostly bacteria, such as Pseudomonas, Rhodococcus, Bacillus, Arthrobacter, Acinetobacter, Azotobacter, Alkaligenes, and Mycobac terium, as well as Candida yeast and filamentous acti nomycetes Streptomyces, are used. In preparations of fungal origin, primarily ascomycetes of the genera Aspergillus and Penicillium are used [116]. Among the higher basidiomycetes, a high oil degrading ability was shown only for genera Phanerochaete, Pleurotus, and Trametes. According to [117], the amount of petro leum hydrocarbons in the presence of P. chrysospo rium, P. ostreatus, and T. (Coriolus) versicolor decreased 12 months after inoculation by 68.7, 53.1, and 78.1%, respectively. A characteristic feature of biodegradation of petroleum hydrocarbons by basidi omycetes is the ability of the latter to metabolize the aromatic fraction of aromatic hydrocarbons, whereas bacteria degrade primarily paraffin naphthenic hydrocarbons [118]. Patents containing the descrip tion of preparations based on basidiomycetes, which are intended to clean up soils contaminated with oil, are absent in the Russian Federation. In the United States, the only patent containing a description of purification of oil contaminated environment by using P. chrysosporium was registered [115]. Cleanup of contaminated soils. To date, approaches to remediation of soils contaminated with various
Vol. 47 No. 6 2011

APPLIED BIOCHEMISTRY AND MICROBIOLOGY


USE OF BASIDIOMYCETES IN INDUSTRIAL WASTE PROCESSING Table 4. Basidiomycete species and metals accumulated by them Fungus Phanerochaete chrysosporium, Phellinus sanguineus, Pleurotus os treiformis, Pleurotus sajor caju, Pycnoporus sanguineus, Trametes versicolor, Volvariella volvacea BDT 14 (DSM 15396), Phanerochaete chrysosporium, Pleurotus ostreiformis, Pleurotus sajor caju, Trametes versicolor, Volvariella volvacea Coriolopsis strumosa, Daedalea tenuis, Ganoderma lucidum, Lenti nus strigosus, Lenzites malaccenis, Lepista nuda, Oudemansiella mucida, Phanerochaete chrysosporium, Phellinus sanguineus, Phellinus xeranticus, Pycnoporus cinnabarinus, Pycnoporus san guineus, Rigidoporus lineatus, Rigidoporus microporus, Trametes lactenia, Trametes versicolor Phanerochaete chrysosporium Phanerochaete chrysosporium, Pleurotus sajor caju, Pleurotus os treiformis, Trametes versicolor, Volvariella volvacea Phanerochaete chrysosporium, Phellinus badius, Phellinus san guineus, Pleurotus ostreiformis, Pleurotus sajor caju, Pycnoporus sanguineus, Trametes versicolor, Volvariella volvacea Innonotus mikadoi, Tricholoma conglobatum Metal Cd [90], [92­103] Source

575

Cr

[92], [93], [104]

Cu

[86], [93­ 95], [97], [103], [105], [106]

Hg Ni Pb

[108, 109] [92], [93], [110], [111] [92­97], [103], [107]

U, Th

[112]

Table 5. Patented methods of degradation of various xenobiotics using basidiomycetes Fungus species Antrodia radiculosa Meruliporia incrassata Marasmiellus troyanus Phanerochaeta chrysosporium Gloeophyllum striatum Phanerochaete chrysosporium Degraded compound Pentachlorophenols (in wood) Benzo(a)pyrene Antibiotics and quinolone naftiridon Halogenated hydrocarbons, DDT Dioxin, heptachlor, DDT, dieldrin, toksofen PAHs PCBs Halogenated hydrocarbons, pentachlorophenol Dioxins, polihlorfenily, biphenyls Source [119] [120] [121] [122] [123] [124] [125] [126] [127]

Phanerochaete gigantea Resiniciun bicolor Pleurotus ostreatus

xenobiotics, including PAHs, polychlorinated biphe nyls (PCBs), nitroaromatic compounds, and pesti cides by basidiomycetes have been developed. The most comprehensively studied basidiomycete genera capable of degradation of xenobiotics of different nature are Phanerochaete, Trametes, and Pleurotus. The list of basidiomycetes used in patented prepara tions for decomposition of various xenobiotics is shown in Table 5. Fungus P. chrysosporium is the most widely used for degradation of various xenobiotics. Degradation of low energy coals. Basidiomycete species that can degrade coal wastes and humates
APPLIED BIOCHEMISTRY AND MICROBIOLOGY

extracted from them were isolated primarily from wood (tree trunks, logs, branches, and stumps) and, therefore, cannot be competitive under conditions of soil [128]. In other words, the problem of assessing the possibility of depolymerization of coal wastes in situ remains to be solved [129]. The currently existing pat ented methods of coal biosolubilization are based on the use of P. chrysoporium [130, 131] and Polyporus versicolor [132] and imply coal processing ex situ. Another urgent problem is the search for basidio mycetes that can not only degrade coal wastes but also actively participate in degradation processes in soils.
No. 6 2011

Vol. 47


576

KULIKOVA et al.

Currently, only one fungal species (Collybia dryophila) meeting these requirements is known [133]. During the screening, the ability of the fungus to synthesize and release extracellular enzymes should be taken into account, because these enzymes are responsible for coal degradation. Decomposition of synthetic polymers. Along with degradation of natural polymers (lignin, cellulose, and humic substances), published data describe the ability of basidiomycetes to degrade synthetic polymers. Synthetic polymers (plastics) are widely used in today's world. Because of their extreme stability and steady accumulation in the environment, it is impor tant to find methods for their biodegradation. The possibility of using basidiomycetes for this purpose has been little studied, but there are several studies in this field. In particular, seven species of white rot fungi were shown to be able to degrade polyvinyl chloride (PVC), a widespread synthetic fabric [134]. Marked depolymerization recorded by reducing the number of CH bonds was demonstrated for P. chrysosporium, P. sajor caju, and Polyporus versicolor; the least depo lymerization potential was observed for species belonging to the genus Pleurotus. According to [135], fungus Pycnoporus cinnabarinus can degrade polyvinyl alcohol, another synthetic polymer that is used as an adhesive. The authors of [135] showed the relationship between the polymer degradation and laccase produc tion. Fungi P. chrysosporium and Trametes versicolor were demonstrated to be able to degrade the polymer nylon (nylon 66), which is widely used in textile industry [136]. Later, isolation and characterization of the enzyme responsible for the degradation of this polymer showed its similarity to the MP [137]. The authors of [138] demonstrated the ability of basidiomycetes to degrade remains of rubber tires. It was found that the most effective is, apparently, Resin icium bicolor. The treatment of aromatics, which are used as additives to rubber, with Resinicium bicolor showed an increase in the growth of bacteria Thioba cillus ferrooxidans on rubber as well as acceleration of devulcanization. Based on these results, the authors of [138] concluded that cocultivation basidiomycetes and bacteria for biodegradation of waste rubber is a promising approach for purposes of resin waste bio degradation. Despite the demonstrated principal possibility of using basidiomycetes for the degradation of synthetic polymers, this direction of using basidiomycetes in technologies of processing and utilization of industrial wastes has not yet found practical application. Thus, in recent years, the interest in using basidio mycetes for degrading lignocellulosic materials and xenobiotics has significantly increased. Numerous studies are still devoted to ligninolytic enzymes, with the majority of works describing the development of approaches to the degradation of xenobiotics and lignocellulosic materials, obtaining recombinant

strains producing these enzymes, and increasing the efficiency of catalysis, pH, and thermal stability. The analysis revealed that, at present, basidio mycetes can be used in technologies of processing and utilization of anthropogenic structures and wastes in the following directions: (1) Purification of contaminated water (including sewage water of textile industry pulp and paper mills, water contaminated with petroleum hydrocarbons, sewage water formed during production of olive oil and sugar from sugar beets or sugar cane, water sus pension remaining after coagulation of latex in rubber industry, and waste water containing heavy metals and radionuclides). (2) Cleanup of contaminated soil (including con tamination with xenobiotics and heavy metals). (3) Degradation of difficult to degrade substrates (including lignin and lignocellulosic waste, low energy coal, and synthetic polymers). ACKNOWLEDGMENTS This study was supported by the Russian Founda tion for Basic Research (project no. 08 04 01612), Ministry of Education and Science of the Russian Federation State Contract P211 and no. 16.512.11.2028. REFERENCES
1. Sanchez, C., Biotechnol. Adv., 2009, vol. 27, pp. 185­ 194. 2. Dashtban, M., Schraft, H., and Qin, W., Int. J. Biol. Sci., 2009, vol. 5, no. 6, pp. 578­595. 3. Martinez, A.T., Speranza, M., Ruiz Duenas, F.J., Fer reira, P., Camarero, S., Guillen, F., Martinez, M.J., Gutierrez, A., and del Rio, J., C, Int. Microbiol., 2005, vol. 8, pp. 195­204. 4. Lacina, C., Germin, G., and Spiros, A., Afr. J. Bio technol., 2003, vol. 2, pp. 620­635. 5. Mougin, C., Boukcim, H., and Jolivalt, C., Advances in Applied Bioremediation (Soil Biology), Berlin, Heidelberg: Springer, 2009. 6. Kumar, R., Singh, S., and Singh, O.V., J. Ind. Micro biol. Biotechnol., 2008, vol. 35, pp. 377­391. 7. Baldrian, P. and Valaskova, V., FEMS Microbiol. Rev., 2008, vol. 32, pp. 501­521. 8. Asgher, M., Bhatti, H.N., Ashraf, M., and Legge, R.L., Biodegradation, 2008, vol. 19, pp. 771­783. 9. Roblez Hernandez, L., Cecilia Gonzalez Franco, A., Crawford, D.L., and Chun, W.W.C., Tecnociencia Chi huahua, 2008, vol. 2, no. 1, pp. 32­40. 10. Rabinovich, M.L., Bolobova, A.V., and Kondrash chenko, V.I., Drevesina i razrushayushchie ee griby (Wood and Wood Degrading Fungi), Moscow: Nauka, 2001. 11. Wong, D.W.S., Enzymes Appl. Biochem. Biotechnol., 2009, vol. 157, pp. 174­209.
Vol. 47 No. 6 2011

APPLIED BIOCHEMISTRY AND MICROBIOLOGY


USE OF BASIDIOMYCETES IN INDUSTRIAL WASTE PROCESSING 12. Ruiz Duenas, F.J. and Martinez, A.T., Microb. Bio technol., 2009, vol. 2, no. 2, pp. 164­177. 13. Tien, M., Crit. Rev. Microbiol., 1987, vol. 161, pp. 141­168. 14. Tien, M. and Kirk, T., Methods Enzymol., 1988, vol. 161, pp. 238­249. 15. Kuwahara, M., Glenn, J.K., Morgan, M.A., and Gold, M.H., FEBS Lett., 1984, vol. 169, no. 2, pp. 247­250. 16. Tien, M. and Kirk, T., Science, 1983, vol. 221, pp. 661­663. 17. Johanson, T., Welinder, K.G., and Nyman, P.O., Arch. Biochem. Biophys., 1993, vol. 300, pp. 57­62. 18. Pointing, S.B., Pelling, A.L., Smith, G.J., Hyde, K.D., and Reddy, C.A., Mycol. Res., 2005, vol. 109, pp. 115­ 124. 19. Tonon, F. and Odier, E., Eur. J. Biochem., 1995, vol. 233, pp. 650­658. 20. Piontek, K., Smith, A.T., and Blodig, W., Biochem. Soc. Trans., 2001, vol. 29, pp. 111­116. 21. Hatakka, A., FEMS Microbiol. Rev., 1994, vol. 13, pp. 125­135. 22. Lobos, S., Larram, J., Salas, L., Cullen, D., and Vicuna, R., Arch. Microbiol., 1994, vol. 14, pp. 2691­ 2698. 23. Hofrichter, M., Enzyme Microb. Technol., 2002, vol. 30, pp. 454­466. 24. Kawai, S., Jensen, K.A., and Hammel, K.E., Appl. Environ. Microbiol., 1995, vol. 61, pp. 3407­3414. 25. Hilden, L., Johansson, G., Pettersson, L.J., Ljungquist, P., and Henriksson, G., FEBS Lett., 2000, vol. 477, pp. 79­83. 26. Sundaramoorthy, M., Youngs, H.L., Gold, M.H., and Poulos, T.L., Biochemistry, 2005, vol. 44, pp. 6463­ 6470. 27. Malmstrom, B.G., Multi Copper Oxidases, Messer schmidt, A., Ed., Singapore: World Sci. Publ., 1997. 28. Solomon, E.I., Sundaram, U.M., and Machonkin, T.E., Chem. Rev., 1996, vol. 96, no. 7, pp. 2563­2606. 29. Ruiz Duenas, F.J., Morales, M., Garcia, E., Miki, Y., Martinez, M.J.., and Martinez, A.T., J. Exp. Bot., 2009, vol. 60, pp. 441­452. 30. Thurnston, C.F., Arch. Microbiol., 1994, vol. 140, no. 1, pp. 19­26. 31. Baldrian, P., FEMS Microbiol. Rev., 2006, vol. 30, no. 2, pp. 215­242. 32. Youn, H. D., Kim, K. J., Maeng, J. S., Han, Y.H., Jeong, I. B., Jeong, G., Kang, S. O., and Hah, Y.C., Arch. Microbiol., 1995, vol. 141, no. 2, pp. 393­398. 33. Koroljova (Skorobogat'ko), O., Stepanova, E., Gavri lova, V., Morozova, O., Lubimova, N., Dzchafarova, A., Jaropolov, A., and Makower, A., J. Biotechnol. Appl. Biochem., 1998, vol. 28, no. 1, pp. 47­54. 34. Gindilis, A., Zhazhina, E., Baranov, Yu., Karyakin, A., Gavrilova, V., and Yaropolov, A., Biokhimiya, 1988, vol. 53, no. 5, pp. 735­739. 35. Lerch, K., Deinum, J., and Reinhammer, B., Bio chem. Biophys. Acta, 1978, vol. 534, no. 1, pp. 7­14.
APPLIED BIOCHEMISTRY AND MICROBIOLOGY

577

36. German, U.A., Muller, G., Hunziker, P.E., and Lurch, K., J. Biol. Chem., 1988, vol. 263, no. 2, pp. 885­896. 37. Xu, F. and Shin, W., Brown. S.H., Wahleithner J.A., Sundaram U.M., Solomon E.I, Biochim. Biophys. Acta, 1996, vol. 1292, no. 2, pp. 303­311. 38. Quintanar, L., Stoj, C., Taylor, A.B., Hart, P.J., Kos man, D.J., and Solomon, E.I., Acc. Chem. Res., 2007, vol. 40, no. 6, pp. 445­452. 39. Piontek, K., M. Antorini, T. Choinowski, J. Biol. Chem., 2002, vol. 277, no. 40, pp. 37663­37669. 40. Lyashenko, A.V., Zhukhlistova, N.E., Gabdoul khakov, A.G., Zhukova, Y.N., Voelter, W., Zaitsev, V.N., Bento, I., Stepanova, E.V., Kachalova, G.S., Koro leva, O.V., Cherkashyn, E.A., Tishkov, V.I., Lamzin, V.S., Schirwitz, K., Morgunova, E.Y., Betzel, C., Lindley, P.F., and Mikhailov, A.M., Acta Crystallogr. Sect. F, 2006, vol. 62, no. 10, pp. 954­957. 41. Polyakov, K.M., Fedorova, T.V., Stepanova, E.V., Cherkashin, E.A., Kurzeev, S.A., Strokopytov, B.V., Lamzin, V.S., and Koroleva, O.V., Acta Crystallogr. D, 2009, vol. 65, no. 6, pp. 611­617. 42. Wang, Y., Vazquez Duhalt, R., and Pickard, M.A., Can. J. Microbiol., 2003, vol. 49, pp. 675­682. 43. Moreira, P.., Duez, G., Dehareng, D., Antunes, A., Almeida Vara, E., and Frere, J.M., J. Biotechnol., 2005, vol. 118, pp. 339­352. 44. Kersten, P.J., Kalyanaraman, B., Hammel, K.E., and Reinhammar, B., Biochem. J., 1990, vol. 268, pp. 475­ 480. 45. Kunamneni, A., Plou, F.J., Ballesteros, A., and Alcalde, M., Recent Pat. Biotechnol., 2008, vol. 2, no. 1, pp. 10­24. 46. Rodriguez, E., Nuero, O., Guillen, F., Martinez, A.T., and Martinez, M.J., Soil Biol. Biochem., 2004, vol. 36, pp. 909­916. 47. Davila Vazquez, G., Tinoco, R., Pickard, M.A., and Vazquez Duhalt, R., Enzyme Microb. Technol., 2005, vol. 36, pp. 223­231. 48. Heinfling, A., Martinez, M.J., Martinez, A.T., Berg bauer, M., and Szewzyk, U., Appl. Environ. Microbiol., 1998, vol. 64, pp. 2788­2793. 49. Malherbe, S. and Cloete, T.E., Environ. Sci. Biotech nol., 2002, vol. 1, pp. 105­114. 50. Dovban. K.I. Zelenoe udobrenie (Green Fertilizer), Moscow: Agropromizdat, 1990. 51. Cohen, R., Persky, L., and Hadar, Y., Appl. Microbiol. Biotechnol., 2002, vol. 58, pp. 582­594. 52. Huttermann, A., Hamza, A.S., Chet, I., Majcherczyk, A., Fouad, T., Badr, A., Cohen, R., and Persky, L., Hadar Y., Agro Food Ind. Hi Tech, 2000, vol. 6, pp. 29­32. 53. RF Patent No. 2001117048/13, 2003. 54. US Patent No. 20070227063, 2007. 55. US Patent No. 2003064502, 2003. 56. US Patent No. 20000541893, 2002. 57. US Patent No. 20020097810, 2003. 58. US Patent No. 20050176583, 2005. 59. US Patent No. 20060104939, 2006. 60. US Patent No. 20080264858, 2008.
No. 6 2011

Vol. 47


578

KULIKOVA et al. 89. Merten, D., Kothe, E., and Buche, G., Mine Water Environ., 2004, vol. 23, pp. 34­43. 90. Iqbal, M., Saeed, A., Edyvean, R.G.J., O'Sullivan, B., and Styring, P., Biotechnol. Lett., 2005, vol. 27 P, pp. 1319­1323. 91. Baldrian, P., Enzyme Microb. Technol., 2003, vol. 32, pp. 78­91. 92. Dey, S., Rao, P.R.N., Bhattacharyya, B.C., and Ban dyopadhyay, M., Bioprocess Eng., 1995, vol. 12, pp. 273­277. 93. Yetis, U., Ozcengiz, G., Dilek, F.B., Ergen, N., and Dolek, A., Water Sci. Technol., 1998, vol. 38, pp. 323­ 330. 94. Day, R., Denizli, A., and Arica, M.Y., Biores. Technol., 2001, vol. 76, pp. 67­70. 95. Say, R., Denizli, A., and Arica, M.Y., Biores. Technol., 2001, vol. 76, pp. 67­70. 96. Cihangir, N. and Saglam, N., Acta Biotechnol., 1999, vol. 19, pp. 171­177. 97. Mashitah, M.D., Zulfadhly, Z., and Bhatia, S., Immo bil. Biotechnol., 1999, vol. 27, pp. 441­445. 98. Gabriel, J., Vosahlo, J., and Baldrian, P., Biotechnol. Tech., 1996, vol. 10, pp. 345­348. 99. Zhou, J.L. and Kiff, R.J., J. Chem. Technol. Biotech nol., 1991, vol. 52, pp. 317­330. 100. Bayramoglu, G., Denizli, A., Bektas, S., and Arica, M.Y., Microchem. J., 2002, vol. 72, pp. 63­76. 101. Arica, M.Y., Kacar, Y., and Genc, O., Biores. Technol., 2001, vol. 80, pp. 121­129. 102. Yalcinkaya, Y., Soysal, L., Denizli, A., Arica, M.Y., Bektas, S., and Genc, O., Hydrometallurgy, 2002, vol. 63, pp. 31­40. 103. Zulfadhly, Z., Mashitah, M.D., and Bhatia, S., Envi ron. Pollut., 2001, vol. 112, pp. 463­470. 104. Trivedi, B.D. and Patel, K.C., J. Microbiol. Biotech nol., 2007, vol. 23, pp. 683­689. 105. Muraleedharan, T.R. and Venkobachar, L.I., Environ. Technol., 1994, vol. 15, pp. 1015­1027. 106. Muraleedharan, T.R., Iyengar, L., and Venkobachar, C., Appl. Environ. Microbiol., 1995, vol. 3507­3508. 107. Wu, J. and Li, Q., J. Environ. Sci., 2002, vol. 14, pp. 108­114. 108. Saglam, N., Say, R., Denizli, A., Patir, S., and Arica, M.Y., Process Biochem., 1999, vol. 34, pp. 725­ 730. 109. Saglam, A., Yalcinkaya, Y., Denizli, A., Arica, M.Y., Genc, O., and Bektas, S., Microchem. J., 2002, vol. 71, pp. 73­81. 110. Ceribasi, I.H. and Yetis, U., Water S.A., 2001, vol. 27, pp. 15­20. 111. Dilek, F.B., Erbay, A., and Yetis, U., Process Biochem., 2002, vol. 37, pp. 723­726. 112. Nakajima, A. and Sakaguchi, T., Appl. Microbiol. Bio technol., 1993, vol. 38, pp. 574­578. 113. RF Patent No. 2005125503/13, 2007. 114. RF Patent No. 2004123329/13, 2006. 115. US Patent No. 19990417571, 2001. 116. Zlotnikov, A.K., Sadovnikova, L.K., Balandina, A.V., Zlotnikov, K.M., and Kazakov, A.V., Vestn. RASKhN, 2007, no.1, pp. 65­67.
Vol. 47 No. 6 2011

61. Sarikaya, A. and Ladisch, M.R., Appl. Biochem. Bio technol., 1997, vol. 62, pp. 131­149. 62. D'Souza, D.T., Tiwari, R., Sah, A.K., and Raghuku mar, C., Enzyme Microb. Technol., 2006, vol. 38, pp. 504­511. 63. Selvam, K., Swaminathan, K., Myung, Hoon., Song, M.H., and Chae, K., J. Microbiol. Biotechnol., 2002, vol. 18, pp. 523­526. 64. Selvam, K., Swaminathan, K., Rasappan, K., Rajen dran, R., and Pattabhi, S., Ecol. Environ. Conserv., 2006, vol. 12, pp. 223­226. 65. Chkhenkeli, V.A. and Nikolaeva, L.A., in Tezisy Mezh dunarodnoi Nauchnoi Konferentsii "Mikroorganizmy I biosfera" (Abstr. Int. Sci. Conf. "Microorganisms and Biosphere, Moscow: Winogradsky Institute of Micro biology, Russian Academy of Science, 2007, pp. 147­ 149. 66. RF Patent No. 4071008/26, 1994. 67. Font, X., Caminal, G., Gabarrell, X., and Vicent, T., Environ.Technol., 2006, vol. 27, pp. 845­854. 68. Driessel, B.V. and Christov, L., J. Biosci. Bioeng., 2001, vol. 92, pp. 271­276. 69. US Patent No. 19940247130, 1994. 70. US Patent No. 19950471126, 1996. 71. US Patent No. 19940330874, 1996. 72. US Patent No. 19950536536, 1998. 73. US Patent No. 6923912, 2003. 74. Novotny, C., Rawal, B., Bhatt, M., Patel, M., Sasek, V., and Molitoris, H.P., J. Biotechnol., 2001, vol. 89, pp. 113­122. 75. Michniewicz, A., Ledakowicz, S., Ullrich, R., and Hofrichter, M., Dyes Pigm., 2008, vol. 77, pp. 295­ 302. 76. Gill, P.K., Arora, D.S., and Chander, M., J. Ind. Microbiol. Biotech., 2002, vol. 28, pp. 2001­2003. 77. Baldrian, P., Appl. Microbiol. Biotechnol., 2004, vol. 63, pp. 560­563. 78. Mazmanci, M.A. and Unyayar, A., Pichia stipitis, Proc. Biochem., 2005, vol. 40, pp. 337­342. 79. Cameron, M.D., Timofeevski, S., and Aust, S.D., Appl. Microbiol. Biotechnol., 2000, vol. 54, pp. 751­ 758. 80. Balan, D.S.L. and Monteiro, R.T.R., J. Biotechnol., 2001, vol. 89, pp. 141­145. 81. Chagas, E.P. and Durrant, L.R., Enzyme Microb. Technol., 2001, vol. 29, pp. 473­477. 82. Jain, N., Kaur, A., Singh, D., and Dahiya, S., J. Envi ron. Biol., 2000, vol. 21, pp. 179­183. 83. Kapdan, I., Kargi, F., McMullan, G., and Marchant, R., Bioproc. Eng., 2000, vol. 22, pp. 347­351. 84. US Patent No. 20020124580, 2005. 85. Gutnick, D.L. and Bach, H., Appl. Microbiol. Biotech nol., 2000, vol. 54, pp. 451­460. 86. Gabriel, J., Baldrian, P., Hladikova, K., and Hakova, M., Lett. Appl. Microbiol., 2001, vol. 32, pp. 194­198. 87. RF Patent No. 5048003/25, 1994. 88. RF Patent No. 5048033/25, 1994.

APPLIED BIOCHEMISTRY AND MICROBIOLOGY


USE OF BASIDIOMYCETES IN INDUSTRIAL WASTE PROCESSING 117. Yateem, A., Balba, M.T., Al Awadhi, N., and El Nawawy, A.S., Environ. Int., 1998, vol. 24, pp. 181­ 187. 118. Pozdnyakova, N.N., Nikitina, V.E., and Turkovskaya, O.V., Appl. Biochem. Microbiol., 2008, vol. 44, no. 1, pp. 69­75. 119. US Patent No. 20000541944, 2002. 120. US Patent No. 19960599260, 2001. 121. US Patent No. 19951039445, 2000. 122. US Patent No. 19880183114, 1990. 123. US Patent No. 19910687368, 1994. 124. US Patent No. 19860899000, 1988. 125. US Patent No. 19970939464, 2000. 126. US Patent No. 19930074643, 1994. 127. US Patent No. 2004067730, 2004. 128. Dix, N.J. and Webster, J., Fungal Ecology, London, U.K:. Chapman Hall, 1995. 129. Kastner, M. and Hofrichter, M., Biopolymers. Lignin, Humic Substances and Coal, Hofrichter, M. and Stein

579

130. 131. 132. 133. 134. 135. 136. 137. 138.

buchel, A., Weinheim: Wiley VCH, 2001, vol. 1, pp. 349­378. US Patent No. 19930065563, 1995. US Patent No. 19950477410, 1997. US Patent No. 19870069709, 1989. Steffen, K.T., Hatakka, A., and Hofrichter, M., Appl. Environ. Microbiol., 2002, vol. 68, pp. 3442­3448. Kirbas, Z., Keskin, N., and Guner, A., Bull. Environ. Contam. Toxicol., 1999, vol. 63, pp. 335­342. Larking, D.M., Crawford, R.L., Christie, G.B.Y., and Lonergan, G.T., Appl. Environ. Microbiol., 1999, vol. 65, pp. 1798­1800. Deguchi, T., Kakezawa, M., and Nishida, T., Appl. Environ. Microbiol., 1997, vol. 63 P, pp. 329­ 331. Deguchi, T., Kitaoka, Y., Kakezawa, M., and Nishida, T., Appl. Environ. Microbiol., 1998, vol. 64, pp. 1366­1371. Bredberg, K., Andersson, B.E., Landfors, E., and Holst, O., Biores. Technol., 2002, vol. 83, pp. 221­224.

APPLIED BIOCHEMISTRY AND MICROBIOLOGY

Vol. 47

No. 6

2011