ISSN 0097�8078, Water Resources, 2010, Vol. 37, No. 3, pp. 332–346. © Pleiades Publishing, Ltd., 2010.

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INTRODUCTION

The degradation of persistent organic matter(OM), in particular, lignin, is known to be imple�mented by the enzymatic systems of fungi in the pres�ence of oxygen. Enzyme molecules initiate the split�ting of bonds in organic macromolecules, thus causingthe formation of low�molecular products, which enterthe metabolism of microorganisms and undergo min�eralization. Lignin degradation, taking place duringthe degradation of wood remains, is well understood.Groups of terrestrial lignin�degrading fungi and theirenzymatic systems are described, and biochemicalmechanisms of enzyme action on substrate are pro�posed. The degradation processes in the aquatic envi�ronment are much less known. There is evidence thatfungi contribute to the degradation of anthropogeniclignins entering water bodies on land with effluents ofpaper and pulp mills industry by degrading persistentlignin fractions [25, 54]. Intense studies of persistentOM degradation in seawater and bottom sedimentshave started recently [51, 62, 101, 102, 112]. Thesestudies have found a number of interesting factsregarding the presence of oxygen in seawater at largedepth, the penetration of oxygen into deep�sea sedi�ments, the presence and diversity of fungi in thesemedia, and the spectrum of enzymes released by them.The obtained data is of extreme importance, first, inthe context of the problem of the World Ocean pollu�tion by persistent compounds; second, for the assess�ment of components of the global carbon cycle, pri�marily, the rate of OM mineralization and burial inbottom sediments (BS). It is clear that the OM occur�ring in the deep�sea environment is mostly persistent,since its labile fractions degrade during long�time sed�imentation of particles in the water body. In view of thenew data, there is reason to believe that the degrada�

tion of persistent OM in the aquatic environmentunder aerobic conditions is implemented, as well as onthe land, by fungi. The quantitative assessment of deg�radation requires the development of a kinetic model,reflecting all major biochemical regularities in theaction of enzymes on the substrate. These regularitiesshould be analyzed from kinetic viewpoint withemphasis placed on the molecular decompositionmechanisms.

Lignin, whose structure and properties weredescribed in detail in part 2 of this series of papers, willbe taken as a typical representative. The questions tobe answered are as follows: what are the fungi thatimplement the degradation of persistent OM in theaquatic environment, what are the enzymes involvedand the conditions required for the process to proceed;what is the chemical mechanism of lignin macromol�ecule degradation and how it is organized in space;what is the effect of the structure of macromolecules intheir volume and surface on the interaction withenzymes; what is the size ratio of the macromoleculeand enzyme; what fragments are produced in the ele�mentary acts of degradation and what is their fate. Letus consider the problems listed above, using materialsfrom the vast published data.

FUNGI IN THE AQUATIC ENVIRONMENT

The aquatic environment and sediments werefound to contain a wide diversity of fungi, mostlybelonging to terrestrial varieties [51]. Marine fungispecies were first isolated from the oceanic water of thenorthwestern subtropical region of the Atlantic Oceanfrom the depth of 4450 m [102]. Obligate marine fungiwere found on wooden panels at depths of 500–3000 m [79].

WATER QUALITY AND PROTECTION: ENVIRONMENTAL ASPECTS

Modeling the Biodegradation of Multicomponent Organic Matter in an Aquatic Environment: 3. Analysis of Lignin Degradation Mechanisms

T. N. Gubernatorova and B. M. DolgonosovWater Problems Institute, Russian Academy of Sciences, ul. Gubkina 3, Moscow, 119333 Russia

Received September 10, 2009

Abstract—Mechanisms of enzymatic degradation of a persistent organic substance are discussed in the caseof lignin. The major groups of lignin�decomposing microorganisms and their enzymatic systems aredescribed. The biochemical mechanisms of the action of main lignin�decomposing enzymes are analyzed.Typical sizes of enzyme molecules are estimated. The results of analysis are used to formulate the major reg�ularities of lignin destruction, which are required for the construction of a kinetic model of this process.

Keywords: persistent organic matter, lignin, aquatic environment, degradation mechanisms.

DOI: 10.1134/S0097807810030085

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MODELING THE BIODEGRADATION OF MULTICOMPONENT ORGANIC MATTER 333

One of the earliest reports about deep�sea fungusAspergillus ustus, isolated from calcareous BS of theBay of Bengal from the depth of 965 m was given in[100]. Experiments with cultivating yeast and myceliafungi under the conditions similar to those in deep�seasediments were described in [83, 128]. Fungi andyeasts were found to occur in sediment samples takenfrom Mariana Trench in the Pacific from the depth of10500 m [111]. Later [112] they were identified asPenicillium lagena and Rhodotorula mucilaginosa.The authors of [101] also report about the presence ofdeep�sea fungi, which were directly determined andisolated from BS core 4.7 m in length from ChagosTrench in the Indian Ocean taken from the depth of5000 m.

Studies of the fungi in deep�sea oceanic BS werecontinued in [51], resulting in the establishment of thepresence of fungi cultures, which were isolated andidentified. These cultures were studied in experimentsinvolving their cultivation under similar conditions.The comparison of some identified samples of deep�sea fungi with common terrestrial species has shownthe sporulating microorganisms to be able to developat large depths, where they exist not only as spores andhyphae, but also as mycelial agglomerations. Themycelial state of fungi is the consequence of active veg�etation growth under given conditions within BS or asdisseminations in OM particles. The amounts of fungimass in BS samples taken from different depths wereabout the same, suggesting the homogeneity of sedi�ments in the sampling region. Laboratory experimentsshowed that the terrestrial fungi species placed intodeep�sea BS, after a stress stage, gradually adapt to theambient conditions, in particular, to the high pressure,varying temperature, and unstable OM content. Somefungi cultures have adapted quite adequately by them�selves to the conditions of high pressure and low tem�perature (the authors studied four fungi species: threeterrestrial and one deep�sea).

The occurrence of fungi in deep�sea BS of the cen�tral Indian Ocean was studied in [51]. The sampleswere taken from the depth of 5000 m. During thisstudy, the authors isolated 181 fungi cultures, amongwhich terrestrial sporulating fungi were dominating.The microorganisms from deep�sea BS refer to thegenera of Aspergillus, Penicillium, Cladosporium,Curvularia, and Fusarium. Some non�sporulating andyeast fungi were also recorded. Aspergillus terreus spe�cies was found to be most typical of all isolated sam�ples. The authors established the presence of fungihyphae in BS. The total of twenty�five varieties offungi of Aspergillus genus produced considerable bio�mass under model conditions reproducing those ofdeep�sea environment (the pressure of 200 bar and thetemperature of 5 and 30°C). The low temperature wasfound to have a greater adverse effect on fungi growththan the high pressure. Earlier studies of Aspergillusgenus [55] showed this culture to be physiologicallyuniversal, most significant in sediments, and able to

colonize a wide range of substrates. These data wereconfirmed in [51], where it was also established thatthis genus morphologically refers to terrestrial forms offungi. However, the ability to adapt to deep�sea condi�tions can have resulted from a genetic modification.

As to the question of how terrestrial fungi formscould appear in deep�sea BS, it was supposed that theycould have been transported onto the oceanic bed ashyphae germinating on particles of BS of terrigenousorigin (partially decayed leaves and wood fragments)[51]. Moreover, fungal spores could be blown intowater from the land surface. The occurrence of woodfragments at large depths, resulting from their washoutduring inundation of trees in monsoon seasons in thetropics or during spring floods, was reported in [115].It is also possible that even substrates of purely oceanicorigin on water surface can be colonized by fungal cul�tures.

The discoloration of dyed water, polluted by efflu�ents of paper and pulp, textile, and paint industries, bymarine fungi isolated from BS and cultivated onremains of partially decayed wood from mangrove for�ests (no identification of fungi was made) was studiedin [54]. Laccase was found to be the dominating lig�nin�degrading enzyme among those released by fungi.In addition to laccase, a weak activity of manganeseperoxidase and lignin peroxidase was recorded. Therelease of laccase increased in the presence of phenoland non�phenol inductors. An increase in laccase pro�duction was also recorded after the processing of dyedeffluents. Water containing industrial pollutants andsynthetic dyes rapidly discolored after the introduc�tion of samples of growing fungi culture. The part ofthe culture floating on water surface, though having noconsiderable biomass, also effectively discolored pol�luted water during 6�hour incubation.

Studies of other enzymes produced by deep�seafungi species are also in progress now. An alkaline� andcold�tolerant protease produced by deep�sea fungi wasstudied in [52]. This protease refers to enzymes fromthe class of hydrolases and breaks the peptide bondbetween aminoacids in proteins. Two hundreds andtwenty�one deep�sea fungi species, isolated from BStaken from the depth of 5000 m in the central IndianOcean, were examined to determine their proteasecontent. This enzyme was produced in large amountsby Aspergillus ustus species. The fungal culture grew attemperatures of 5 and 30°C and the pressure increas�ing from 1 to 300 bar. The conditions were found atwhich the enzymatic activity attains its maximum.Interestingly, the peak of enzymatic activity does notcoincide with the peak of fungal culture biomass.

Fungi can exist only under aerobic conditions;therefore, the presence of fungi in aquatic environ�ment indirectly suggests the presence of oxygen in it.However, direct measurements of oxygen content ofdeep�sea BS have not been carried out until recently.The rate of oxygen consumption by benthic microor�ganisms is studied in [62]. BS samples, taken for this

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purpose in different regions of the central PacificOcean, were analyzed for oxygen content. Theobtained vertical oxygen profiles show the BS to be oxy�genated as deep as 8 m below the sediment surface. Oxy�gen concentration on the surface was ~200 μmol O2/l,while at depth of 8 m it varied within 150 to 180 μmol O2/l.The oxygen flux in BS was ~0.1 μmol O2/(m2 day).This flux remains about the same level deeper into thesediment, suggesting the extremely small rates of oxy�gen exchange, low microbial activity, and slow OMdegradation. The extrapolation of oxygen profilesdeeper into the sediment body, carried out by theauthors, allows the conclusion to be made that BS areoxygenated down to the basalt basement. The thicknessof sediments above the basement varied from 6 to 60 m.According to these estimates, the typical oxygen concen�tration near the basalt basement is ~100 μmol O2/l, i.e.,only half that on the sediment surface.

The presence of oxygen in BS enables the vitalactivity of not only fungi, but also aerobic bacteria,which utilize low�molecular products of destruction ofpersistent OM—a process, which is carried out byfungi. New bacterial species of Brevibacteriumgenus, inhabiting deep�sea BS, have been isolatedand identified recently [36]. BS samples were takenfrom the depth of 5904 m with deepening into thesediment by 4.6 m. The sediment at this depth isdated at 50000 years.

Thus, we can state that oxygen not only occurs atlarge depth in water but also penetrates into deep�seaBS, creating the necessary conditions for fungi exist�ence. The similar nature of aquatic and terrestrialfungi allows OM degradation in the aquatic environ�ment to be described with the use of data collected forterrestrial fungi, which are considered below.

LIGNIN�DEGRADING FUNGI AND THEIR ENZYMATIC SYSTEMS

The biodegradation of natural high�molecular sub�strates, including lignin, is a multistage polyenzymaticprocess implemented by fungi. Part of fungi enzymes(constitutive enzymes) forms in the process of ontoge�nesis, while the other part (adaptive enzymes) appearslater depending on the environmental conditions.Inductors of enzymes can be appropriate substrates ormonomer products of reactions they are involved in.The constitutive enzymes are produced not at once butin certain order; their list can widely vary in terms ofthe quantitative ratios and the qualitative composi�tion, depending on the age of the culture [3, 8, 9]. Theenzymatic systems of fungi are very flexible and canadapt to the environmental conditions and BS com�position, ensuring, on the one hand, the high adaptiv�ity of fungi to the changing ambient conditions, andon the other hand, the perpetual transformation ofpersistent OM under widely varying conditions.

Enzymes exhibit catalytic activity in the externalenvironment over a long time [10, 11, 27]. There is

some evidence that they can persist for a long time inseawater and BS [94–96]. In the aquatic environment,enzymes can diffuse into suspension mesopores,where they are protected against bacterial attack [49].Another protection mechanism is due to the incapsu�lation of biomolecules by the formation of an envelopefrom associated organic molecules [45, 76]. Severalother protection mechanisms are described in [95].

White�rot fungi, which have a developed enzy�matic system, are the main suppliers of lignin�degrad�ing enzymes among terrestrial fungi. This systemincludes a subsystem of lignin�degrading enzymes,such as laccases, peroxidases (lignin peroxidase, man�ganese peroxidase, versatile peroxidase, methane oxi�dase, etc.), tyrosinases, cellobiose dehydrogenases, aswell as a subsystem of hydrolases, which ensure thedegradation of polysaccharides [7, 14, 21, 22]. Thedegradation rates of lignin and polysaccharides aredifferent for different fungi, depending on the activityof enzymatic systems, such that different activities canbe recorded in the same fungi species [108]. Lignindegradation follows oxidation mechanism with oblig�atory participation of active oxygen. Oxidation reac�tions ensure multistage transformation of biopolymermacromolecules into low�molecular compounds uti�lized by the fungi and bacterial community [9, 22, 92,108, 114].

Brown�rot, soft�rot, blue stain, and some otherfungi also contribute to lignin transformation [2, 15,18–20, 28]. Brown�rot fungi change lignin onlychemically, reducing the number of methoxyl groupsand increasing the concentration of carbonyl and car�boxyl groups at the expense of oxidation degradationand demethylation. Oxidation results in the formationof a considerable amount of CO2 accompanied by car�bon losses from propane chains and methoxyl groups,however the loss of mass can be compensated for byoxygen introduction [75]. Soft�rot fungi produce aweak enzymatic system with respect to lignin [9].However, some fungi in this group were found to beable to utilize dehydropolymerizates. Methoxylgroups, propane chains, and aromatic rings also con�tribute to CO2 formation [21, 22, 26]. Some blue stainfungi have effects similar to soft�rot fungi. The pres�ence of lignin�degrading enzymes (phenoloxidases)was recorded in 19 blue stain fungi. The majority ofthem also produce laccase, mostly as an intracellularenzyme [19, 26].

The ability of bacteria to degrade lignin is very lim�ited, though it is greater in the case of mixed cultures.This was confirmed by experiments with dehydropoly�merizate. Bacterial metabolism incorporates succes�sive reactions of oxidation of α�carbon atom, the oxi�dation splitting of simple arylether bonds and thedetachment of two C atoms from propane chains.However, the degradation of the main lignin frame isweak [19, 26, 63]. Overall, under aerobic conditions,bacteria play a secondary role in lignin degradationrelative to fungi; however, under anoxia, when fungi

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cannot function, anaerobic bacteria, participating indenitrification, sulfidogenesis, and methanogenesis,become practically exclusive, though less effective,biodegrader of lignin, mostly of its low�molecularforms.

Note that lignin�degrading enzymes are commonlyproduced by fungi irrespective of whether lignin isavailable or not (unlike cellulase complex enzymes).However, this is not always the case. Thus, as noted in[1, 40, 42, 120], with the development of fungi and achange of metabolic mechanism (the change from tro�phophase to idiophase because of the depletion ofreadily degradable matter, the accumulation of metab�olites, and the depletion of medium), the lignolyticactivity of cultures manifests itself as the response toN, C, or S deficiency in the medium. The effectiveimpact of N limit on the manifestation of ligninperox�idase activity of lignin�degrading fungi [41, 113, 118]can be explained from the tropho–ecological view�point, since wood, which is a natural substrate forwhite�rot fungi, contains a minimal amount of N [19].

Among other features of the lignolytic system,noteworthy also are its nonspecificity with respect tosubstrate (unlike cellulases), and the fact that lignindegradation develops through oxidation processes,while in the case of other biopolymers (proteins, cel�lulose), degradation takes place through hydrolysis[78]. Thus, we can say that fungi are originally aimedto process persistent substrates and for this produceappropriate nonspecific enzymatic system with a widerange of action.

The degradation of lignin macromolecules underthe effect of enzymes incorporates the following reac�tion groups: the oxidation of lignin side chains by α�and β�carbon atoms with the formation of ketogroup�containing structures and phenol structures; thehydrolytic splitting of β–O–4 ether bonds with the for�mation of alcohol and phenol structures; the splitting ofalcylaryl C–C bonds, the formation of p�quinoid struc�tures and aldehyde or acid fragments; demethylationand hydroxylation of aromatic ring; splitting of aro�matic ring with the formation of aliphatic products,most often, carbonic acids.

ACTION MECHANISMS OF LIGNIN�DEGRADING ENZYMES

Ligninperoxidases

Ligninperoxidase (LiP) holds a central positionamong enzymes as the most effective enzyme, which isable to catalyze several lignin cleavage reactions. LiP isthe key enzyme in the oxidation of nonphenol ligninstructures to cation�radicals, which are next involvedin a series of nonenzymatic reactions, including therupture of C–C and C–O bonds and the fragmenta�tion of lignin three�dimensional lattice. The structureof LiP molecule is given in Fig. 1. A unique distinctivefeature of this enzyme group is its ability to perform

one�electron oxidation of a wide range of dimethox�yphenyl substrates with the formation of cation�radi�cal intermediates. By their structure, all LiP isoen�zymes are monomer hemoglycoproteins [64]. Theactive center of LiP is similar to the closed structure ofthe active center of horse�radish peroxidase (HRP)(the so�called heme pocket) and differs from otherheme�containing proteins: globins, catalase, chloroperox�idase, and cytochrome P450, in which the heme is exposedoutward [53]. In LiP isoenzymes, ion Fe3+, in addition tothe tetrapyrrol ring of protoporphyrin IX (1,3,5,8�tetram�ethyl�2,4�divinyl�6,7�carboxyethylporphyrin) (Fig. 2)coordinates also the remainder of imidazol of proteinhistidine. According to data of different authors, ironhas five [80] or six [32] ligands. The integrity of thestructure of enzyme active center is supported by cal�cium ions.

The catalytic cycle of LiP (Fig. 3) includes two�electron oxidation of ferry�(Fe3+) form of the enzymeby hydrogen peroxide to oxoferryl�(Fe4+O) cation�radical. This intermediate in peroxidase catalysis isreferred to as compound I (LiP I) [46]. The one�elec�tron reduction of compound I by an appropriate sub�strate (e.g., veratric alcohol—VA) leads to a second

Nt

Ca2–B' C

D

A

Ct

BJ

E

F

I HG

Ca2+

LiPH8

Fig. 1. Molecular structure of lignolytic peroxidase LiP(isoenzyme H8). Shown in the figure are the positions ofthe heme group (tetrapyrrol ring in the center); enzymedomains A–G; C and N terminals (Ct, Nt) and two struc�tural ions Ca2+ [86].

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intermediate, compound II (LiP II), in which there isno porphyrin cation�radical, but still persists a ferryl�derivative of the heme (oxoferryl�heme). CompoundII can further implement one�electron oxidation ofappropriate substrates, returning to initial ferri�formof the enzyme and thus completing the catalytic cycle[124]. In the absence of appropriate one�electronreducers, compound II can be involved in a series ofreactions with the excess of peroxide, forming oxy�complex (oxyferroheme), referred to as compound III(LiP III) (inactive form of enzyme) [105, 123].

Thus, the interaction of LiP with peroxide canresult in the formation of not less than three well�stud�ied intermediates, which feature different transforma�tion rates into the initial enzyme. The most activeintermediate is compound I, compound II ranks sec�ond, and compound III, which forms as the result ofthe reduction of the latter by the excessive peroxide, isthe least reactive.

The completion of the catalytic cycle of enzymeand the possibility for the catalytic reaction to proceedin a stationary regime require the conditions at whichcompound II and, all the more, compound III will notaccumulate and not become an irredundant form.Compounds I and II of the enzyme were shown to beable to some extent directly oxidize high�molecularprotein substrate [106, 107].

This disagrees with the crystallographic studies ofLiP structure [22], which show that heme is com�pletely submerged into protein globule and is unable todirectly interact with high�molecular substrates.

Nonetheless, ferrocytochrome c (Cc2+) is oxidizedby LiP/H2O2 system in the absence of VA. A mecha�nism involving direct impact of the enzyme on thepolymer substrate was proposed to explain this phe�nomenon based on electron transfer from the hemepocket onto globule surface through a long chain(long�range electron transfer (LRET)) (Fig. 4). Thetransfer of electron from cytochrome heme c onto LiPheme is assumed to take place via surface protein–

N

Fe

N

NN

OH

OOH

O

3+

1 2

3

4

56

78

1

δ�mezo

Fig. 2. Tetrapyrrol ring [5].

+O2–•

Fe3+ H2O2

Ferri�LiP

VA, Trp

O

VA, PhOH

VA+•, PhO•H2 O

2

+VA+•

+•

O

O2–•

Fe4+

Fe2+Fe2+

Fe4+

O

O2

OxoferrilhemeProtoporphyrin IX oxoferryl�cateonradical(LiP I)

Dimethoxybenzoletc.

ED

TA

dea

zafla

vin,

hν

Oxoferrilheme(LiP II)

Oxyferroheme(LiP III)

Ferri�LiP

1

2

3

Fig. 3. Catalytic cycle of LiP [46].

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protein interaction [126]. Such mechanism with theparticipation of LRET was proposed for cytochrome�c�peroxidase (CCP) [34, 93].

Another mechanism supposes that the depolymer�ization of lignin matrix and other high�molecular sub�strates is mediated by low�molecular dimethoxyphenylmediator of VA type or 1,4�dimethoxybenzol (14�DMB),which forms a stable radical cation particle under theeffect of enzyme (Fig. 5).

The oxidation depolymerization LiP of modeldimers of lignin with different bond types does notrequire the participation of a mediator. It has beenshown recently that not only dimers, but also higheroligomers, which are hardly able to penetrate into theaccess channel to the heme, as well as the dehydroco�niferyl polymer (DHP)—an analogue of spruce lig�nin—can be directly degraded by enzyme, supposedly,through the exposed outward redox�center [90]. LiP issupposed to have several substrate sites, and it is stillunclear how they are located relative to heme δ�mesosite (Fig. 2), which is connected with globule sur�face through a narrow channel. This is true for thebonding of not only polymer, but also low�molecularsubstrates [5].

Noteworthy, the action mechanism of lignin perox�idase is adequately understood only for model lignin

compounds [56, 68, 70, 125]. Lignin peroxidase isknown to catalyze the following reactions: cleavage ofaromatic ring [116], splitting of Cα–Cβ bonds in arylg�lycerin�β�aryl ethers [77], splitting of β–O–4 bonds[127], oxidation to quinines and alcohols [41], deme�thylation and oxidation to phenols [61]. The degrada�tion paths of dimer lignin compounds are described in[16], where exhaustive information is given about therole of lignin peroxidase in biooxidation reactions ofmodel compounds with α� and β–O–4 bonds domi�nating in lignin.

It should be emphasized that oxygen plays a keyrole in lignin peroxidase functioning. In the oxidationwith LiP of aromatic substrates, oxygen is responsiblefor the formation of peroxyradicals, which can laterinvolve new substrate molecules into chain nonenzy�matic reactions, thus considerably accelerating sub�strate transformation. Of particular importance, how�ever, is the availability of oxygen to outdo the recom�bination of carbon radicals that result from protondischarge by cation radicals—direct products of LiP.Otherwise, the degradation will be replaced by theinverse process—linking of radicals and the accumu�lation of dimers and oligomers, as was the case withlignin formation. Moreover, peroxides—the productsof peroxyradical reduction by ions Mn2+ or otherappropriate reducers—can be involved in secondary

Enzyme LiP LRET

Enzyme active center

Surface Red�Ox�active site to

whiche e– is transferred via LRET chain

Activeform of phenylpropane unit

Phenylpropanestructural

unit

Direct transfer of e– viasurface site onto

phenylpropane unit

e– transfer via the chain of conjugated bonds deeper into the matrix;C–

C and C–O bonds

Phenylpropaneuni Ph�OH

Phenylpropaneunit

detachment

e–

Ph ~ O•

Ph ~ OH

Ph ~ OH

Lignin

macromolecule

Fe3+

e–

Fig. 4. LRET mechanism of enzyme action.

⎯H⎯

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enzymatic processes, where they replace hydrogenperoxide during the formation of LiP compound I.

Thus, we can conclude that lignin peroxidase cata�lyzes the following reactions: splitting of C–C bondsbetween phenylpropane links, oxidation of benzylalcohols, splitting of ether bonds, oxidation of methylsubstituents in benzyl compounds, hydroxylation of ben�zyl methyl groups, cleavage of aromatic ring, decarboxy�lation of oxalic acid, polymerization of phenols.

Manganese Peroxidases

Manganese peroxidase (MnP) is the second mostimportant enzyme in the process of lignin degradationby white�rot fungi [122]. The ability to directly oxidizeMn2+ to Mn3+ is a unique property of MnP of P. chry�sosporium and many other fungi [71]. Some enzymes,such as LiP also can oxidize Mn2+, though not directlybut owing to superoxide anion radical, which forms inthe redox�cycle. Only some MnP are able to producemanganese�independent oxidation. The structure ofMnP is similar to that of LiP given in Fig. 1.

The specificity of MnP action. MnP I compoundoxidizes not only Mn2+, but also some organic sub�strates, whereas the presence of Mn2+ is absolutelynecessary for the reduction of compound II into thenative enzyme. Mn2+ oxidation is accompanied by theaccumulation of compound III and its reactivationwith Mn3+. When chelator is deficient or complex for�

mation is not efficient, the enzyme turnover is muchslower; however, the newly formed Mn3+ is more reac�tive and decomposes peroxidase following catalasetype, not allowing enzyme re�reduction by it and theformation of compound III. Thus, the main role ofMn2+ ions is the prevention of accumulation of MnPIII compound, with their protection effect followingdifferent mechanisms, depending on conditions[121, 123].

During the oxidation of oligomer phenol ligninmodels, compound I MnP is much more reactive thancompound II but much less reactive than LiP I. Thus,notwithstanding its ability to oxidize phenols, thisenzyme has only one physiologically significant sub�strate—Mn2+ ions—without which its compound IIis an irredundant form of enzyme, which hampers thebiocatalyst’s turnover. In other words, MnP can oxi�dize lignin oligomers in stationary regime only in thepresence of manganese ions as redox�mediators [33].At the same time, MnP can launch numerous nonen�zymatic processes initiated by Mn2+ complexes or sec�ondary mediators. Thus MnP can support free�radicalprocesses, following the Haber–Weiss or Fenton reac�tion type, and in some cases, it can provide itself withperoxide via dismutation of superoxide anion thatforms during oxygen reduction by carbonic acid radi�cals [31, 117].

The formation by manganese peroxidase of freelydiffusing low�molecular oxidizers, the main among

EnzymeLiP

Enzyme activecenter

Enzyme activation

ActiveformLiP+•

+ mediator(VA; PhOh;14DMB)

Activation of

Active form of Mediator (VA+•;PhO+•; 14DMB+•)

е– transfer; activation of phenylpropane unit

е– transfer via the chain of conjugated bonds deeper into the matrix; splitting of C–C andC–O bonds

Ph ~ OH

Ph ~ OH

Lignin macromolecule

Phenylpropanestructural units

Phenylpropane unit detachment

mediator

Phenylpropane unit Ph–OH

Active form ofphenylpropane unit

Ph ~ O•

–H•

е–

Fig. 5. Enzyme action mechanism through a mediator.

Fe3+ + H2O2

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which is chelated Mn3+ ion, allows it, along with LiP,to contribute to in vitro lignin depolymerization,though, unlike LiP, MnP participates in this processindirectly [31].

Thus, MnP is a powerful extracellular agent ofindirect impact on lignin. It has a valuable property—the ability to provide itself with peroxide, resultingfrom a cascade of conjugate nonenzymatic reactionsof oxygen reduction.

Versatile Peroxidases

Studying white�rot fungi of Pleurotus and Bjerkan�dera genera showed them to have specific forms ofmanganese peroxidases, which have a wide specificity[38, 43, 44, 48, 89, 103, 117]. These forms areincluded into an enzyme group referred to as versatileperoxidases (VP).

P. eryngii was found to have three types of extracel�lular peroxidases. One is allied to MnP of P.chrysospo�rium, another is a versatile enzyme VP, combining thecatalytic properties of LiP and MnP, including directMn2+ oxidation and direct oxidation of phenol andnonphenol dimers. The structure of VP is similar toLiP shown in Fig. 1. It differs from the typical MnPand LiP by the effective oxidation of phenols, forwhich the presence of Mn2+ is required for MnP ofP. chrysosporium and the presence of veratrole forLiP. The third type of enzyme (from a solid�phaseculture) is the intermediate phase which is active withrespect to Mn2+ and phenols, but features little activ�ity with respect to veratrole and is inactive to modeldimers [97].

Thus, one group of peroxidases can be consid�ered as analogue of MnP of P. chrysosporium, whilethe another group includes versatile enzymes VP[72, 73, 89].

Laccases

Laccases are most widespread enzymes capable oftransforming aromatic substrates and not limiting tolignin metabolism. Laccases were detected in manywhite�rot agents, soil saprotrophic and phytopatho�genic fungi (ascomycetes or their hyphomyceteanamorphs), in most polypores—white�rot agents, aswell as in many agaric fungi growing on soil or plantsubstrate.

Laccases participate in lignin degradation pro�cesses, humification of organic remains, and detoxi�cation of xenobiotics. They feature wide specificityand are active in a wide range of pH values.

This reasoning implies that the presence of differ�ent types of peroxidases in the enzymatic system of thesame fungus–destructor allows it to adapt to differentfunctioning conditions, including widely varying pHand the concentration of endogenic peroxide, thusensuring the continuous degradation of lignin under awide range of conditions. The stability of lignin degra�

dation process is maintained by the diversity of coex�isting species of fungi–destructors with varying enzy�matic systems.

Enzyme Sizes

Lignin macromolecule degradation involves aseries of elementary acts of bond splitting. This takesplace when enzyme molecule approaches the ligninmacromolecule surface. The sizes of macromolecules,which can reach several hundreds of nanometers, wereestimated in paper 2 of this series. The size of enzymemolecules given in the table (where, along withlignolytic enzymes, some other types of enzymes aregiven for comparison) is smaller than that. Accordingto these data, we can conclude that the size of lignot�lytic enzymes varies within 4.5 to 8.5 nm. Thus the sizeof lignin macromolecules is one–two orders of magni�tude greater than those of enzyme molecules. Thisimplies that enzyme macromolecules can penetrateinto folds in the relief of macromolecule surface andtransfer electrons into deeper phenylpropane units,thus causing the rupture of internal bonds, resulting inthat not only monomers, but oligomers as well canseparate from macromolecules, though the probabilityfor a monomer to tear off is appreciably less because ofits interaction with the nearest neighbors throughhydrogen bonds.

STRUCTURAL CHANGES DURING LIGNIN DEGRADATION

The main processes taking place during lignin deg�radation are given in Fig. 6.

At the initial stage of functioning of fungi lignolyticsystem, the concentration of carbonyl and carboxylgroups increases and that of aliphatic hydroxyl groupdecreases. The concentration of phenol hydroxyls caneither increase or decrease. In lignin molecules, themass fraction of oxygen increases, while the concen�tration of methoxyl groups decreases due to demethy�lation process. The increase in oxygen concentrationis due to the oxidation of α�carbon atoms and the oxi�dation destruction of bonds between β� and γ�carbonatoms in propane chains, as well as due to oxidationrupture of β–O–4 bonds with the terminal phenylpro�pane units [6, 15, 60, 87].

The reaction of rupture of α–O–4, β–5, β–1, andβ–β bonds results in the appearance of mono� and oli�golignols, most containing carboxyl groups. The deg�radation of lignin macromolecules under the effect ofenzymes involves hydrogen peroxide and free radicals.The peroxide mechanism of lignin degradation resultsin a successive chaotic splitting of numerous etherbonds and eventually ensures the deepest fragmenta�tion of macromolecules. The splitting is most rapid inacid environment for α�ether bonds, while in the caseof β�ether bonds, the splitting is equally fast in bothacid and alkaline environment [9, 23, 24, 29].

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Lignin degradation is not always complete: a part ofit can be transformed into a highly condensed product,because of the competition between destruction pro�cesses and reactions of polymerization and polycon�densation [50]. Condensed lignins, containing diphe�nyl bonds, are very tolerant to the impact of fungusenzymes. Syringyl elements in deciduous lignin andartificial guaiacyl–syringyl lignin are degraded fasterthan guaiacyl. This is due to the quaiacyl part of lignincontaining greater concentration of diphenyl struc�tures, in which phenol hydroxyl groups are not proneto form phenoxyl radicals [60, 87].

The distribution of oligolignol size depends on thestructure of macromolecules and bond energy. In thecase of dendritic macromolecule structure, the separa�tion of a fragment of any size from it would require therupture of a single C–C or C–O bond connecting phe�nylpropane links (the rupture energy is ~80 kcal/mol).Now the limiting factor for the size of the fragmentstorn off will be not the number of bonds broken (one,in this case), but the relief of macromolecule surface,i.e., the size of protrusions and hollows, which deter�mine the penetration depth of molecules of enzymesor radicals–mediators into the macromolecule vol�ume. This steric factor will regulate the size distribu�tion of oligolignols. However, there could be two addi�tional factors affecting oligolignol distribution. One isthe rare intramolecular cross�linking in lignin macro�molecules, resulting in the formation of cylces involv�ing different numbers of monomers. The separation ofa fragment from a cycle requires two bonds to be rup�tured with a total energy of ~160 kcal/mol, thus expo�nentially reducing the probability of this event. If thecross�linking is uniformly distributed over the macro�molecule volume, then, the greater the fragment, the

greater is the number of cycles its elements areinvolved in and the greater number of bonds is to beruptured. Another factor is associated with the hydro�gen bonds between macromolecule fragments (withthe energy of 5 ± 3 kcal/mol), which, though they aremuch weaker than C–C and C–O bonds, are morenumerous (on the average, three bonds per phenylpro�pane bond) and their number increases with the frag�ment’s size, thus reducing the probability that greaterfragment will separate. Thus, these causes contributeto a decrease in the probability of fragment separationwith increasing fragment’s size. The result is the for�mation of a decreasing oligolignol size distribution inthe aquatic environment.

The involvement of mono� and oligolignols in fun�gal metabolism can take place after the splitting of aro�matic rings. Aromatic rings are splitted by dioxigena�ses. This is accompanied by the inclusion of molecularoxygen into substrate. Bonds are splitted eitherbetween two neighboring hydroxyl groups (the so�called ortho�splitting), or between hydroxylated andadjacent nonhydroxylated carbon (meta�splitting).

Ortho�splitting results in the formation of dicarbonacid. It is likely that first O2 molecule combines withhydroxyl groups of neighboring C atoms with the for�mation of cyclic peroxide, and next intermolecularregrouping results in the splitting of the carbon–car�bon bond with the formation of cis,cis�muconic acid.Dioxygenase participates in this process.

Meta�splitting is also catalyzed by dioxygenases.The splitting products in this case are semialdehydes of2�hydroxymuconic acid, which later transform(depending on substitutes) into pyruvate, acetalde�hyde, hydroxyacetate, fumarate, acetoacetate, succi�

Sizes and molecular masses of some enzymes (dash means no data available)

EnzymeEnzyme parameters

LiteratureSize d, nm Molecular mass, Da

Lignin peroxidase (LiP) 4.5–8.2 37000–76800 [37, 47, 66, 99]

Manganese peroxidase (MnP) 4.6–5.3 39000 [109, 110]

Versatile peroxidase (VP) 7.55

3700034000

[81][67]

Laccase 4.5–8.5 54000–56000 [57, 69, 82, 84, 98]

Horse radish peroxidase (HRP) 44–6.7

4400034500–69800

[85][35, 74, 88]

Cytochrome�c�peroxidase (CCP) 5–7.5 34000–34400 [39, 91]

Microperoxidase 2 1900 [58]

Intradioldioxigenase 5 – [119]

Hypoxanthinephosphoribosyltransferaza (HPRT) 1.32 – [65]

Maltose phosphatase 3–4.5 – [59]

Mioglobine 1.5 17000 [17]

Hemoglobin 3 64000

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MODELING THE BIODEGRADATION OF MULTICOMPONENT ORGANIC MATTER 341

nate, or other intermediate products involved in inter�mediate metabolism [28, 104].

It is important to note that the time of cleavage andmineralization of low�molecular lignin degradationproducts is much less than the decay time of ligninmacromolecules.

The enzymatic splitting of aromatic rings is possi�ble not only in monomer degradation products but inlignin polymer as well. However, the probability of thisprocess is less than that of the splitting of other, lessstrong bonds. In this case, the aryl–ether bonds andpropane chains are splitted [9, 28, 29, 87].

Lignin degradation is accompanied by oxidationand degradation of side chains, demethylation,hydrolysis, opening of aromatic ring, and decarboxy�lation [4, 12, 13, 26, 30]. The result is the formation ofa group of substances with lesser molecular mass, suchas aromatic acids: ferulic, syringic, cinnamic, and van�illic, as well as appropriate aldehydes. Additionally,fragments can appear in the form of cyclic com�pounds, mostly phenols and cathechines. Next theacids and aldehydes will transform into appropriatealcohols, which, under favorable conditions, canagain couple with lignin macromolecules. In the caseof degradation of side chains, these aldehydes trans�form into vanillin, lilac aldehyde, and n�hydroxy�ben�zaldehyde. Experiments show that, in addition tothese substances, volatile monohydric phenols, poly�atomic phenols and their derivatives, carbonic acids,

and methyl alcohol also form [50]. Further degrada�tion is accompanied by changes in functional groups,opening of benzene ring and next up to the completemineralization of compounds.

CONCLUSIONS

Analysis of enzymatic degradation of lignin allowsus to identify the features of the process that are ofimportance for modeling the degradation kinetics;these are as follows:

—Biodegradation of lignin macromolecules is anextended process involving the enzymatic systems offungi in the presence of oxygen, required for the rup�ture of bonds between phenylpropane units of themacromolecule;

—Two mechanisms of enzyme action on ligninmacromolecules are known: the mechanism of directaction by transfer of electron from the heme pocketonto globule surface along a LRET chain; the mecha�nism of indirect action via stable cation�radical parti�cles that form as the result of enzyme action onto low�molecular dimethoxyphenyl mediators of VA or14DMB types;

—Oxygen plays a key role in the functioning of lig�nin peroxidase. During LiP oxidation of aromatic sub�strates, oxygen is responsible for the formation of per�oxy radicals, which can later involve new molecules inchain nonenzymatic reactions, thus appreciably

lignin macromoleculesSeparation of fragments with different

molecular mass with the formation of oligolignols andsome low�molecular substances

Rupture of bonds between with the formation of and their derivatives

Aromatic compounds: phenols, carbonic acids, aldehydes and their derivatives Separation of side chains

Processes of demethylation,hydroxylation,

substitution

Opening of aromatic ring

(ortho�, meta�splitting)

Processes of chain oxidation

and decarboxylation

Mineralization tolow�molecular compounds

Oxidation, formation of phenols of

coniferyl and lilac series

Oxidation to quinones and subsequent mineralization

Inverse processes of dimerization, polymerization, polycondensation, which take place parallel to

direct oxidation processes

Formation of molecules of low�molecular compounds

Humification

The processes take place withthe participation of enzyme

groups: peroxidases, laccases, oxidases,

oxitransferases,polyphenoloxidases,

decarboxylases

Fig. 6. Processes of lignin enzymatic degradation.

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accelerating its transformation. However, the avail�ability of oxygen is especially important for overdoingthe recombination of carbon radicals that form as theresult of proton discharge by cation�radicals—directproducts of LiP. This prevents the inverse reaction oflinking radicals, which counteracts macromoleculedegradation.

—The main lignin�degrading enzymes (LiP, MnP,VP, laccase) have molecular mass of 37–77 kDa andthe size of 4.5–8.5 nm.

—Llignin peroxidase catalyzes the following reac�tions: splitting of C–C bonds between phenylpropanelinks, oxidation of benzyl alcohols, splitting of etherbonds, oxidation of methyl substitutes in benzyl com�pounds, hydroxylation of benzyl methyl groups, cleav�age of aromatic ring, decarboxylation of oxalic acid;polymerization of phenols.

—Manganese peroxidase can oxidize lignin in sta�tionary regime only in the presence of manganese ionsas redox mediators. At the same time, MnP is able tolaunch numerous nonenzymatic processes initiated byMn3+ complexes or secondary mediators. MnP is apowerful extracellular agent of indirect impact on lig�nin. It can provide itself with peroxide resulting from aseries of conjugated nonenzymatic reactions of oxygenreduction.

—Laccases and versatile peroxidases feature widespecificity. Laccases are active within a wide range ofpH values.

—The presence of various types of peroxidases inthe enzymatic system of the same fungus–lignindegrader allows it to adapt to different functioningconditions, in particular the wide variations of pH andendogenous peroxide concentration, ensuring thecontinuous degradation of lignin under different con�ditions. The stability of lignin degradation process isdue to the diversity of the coexisting fungi–degraderswith different enzymatic systems, the ability of fungi toadapt to the environmental conditions and the avail�able substrate by appropriately changing (adjusting)the qualitative and quantitative composition of thesynthesized complex of enzymes.

—At the initial stage of the action of white�rotfungi enzymatic systems, the concentration of carbo�nyl and carboxyl groups in lignin increases, and that ofaliphatic hydroxyl groups decreases. The concentra�tion of phenol hydroxyl can either increase ordecrease. The mass fraction of oxygen in lignin mole�cules increases, and the concentration of methoxylgroups decreases due to demethylation.

—The peroxide mechanism of lignin degradationresults in chaotic splitting of numerous ether bondsand ensures deep fragmentation of macromolecules.The reactions of α–O–4, β–5, β–1, and β–β bondsplitting results in the formation of mono� and oligoli�gnols.

—The probability for a fragment to separatedecreases with an increase in its size. The result is the

formation of a decreasing oligolignol size distributionin the aquatic environment.

—The aromatic compounds that form during lig�nin biodegradation experience further cleavage. Thesplitting of aromatic ring is implemented by dioxygen�ase. This is accompanied by oxygen inclusion into thesubstrate. The splitting takes place either betweenneighboring hydroxyl groups (ortho�splitting), orbetween hydroxylated carbon and the neighboringnonhydroxylated one (meta�splitting).

—The enzymatic cleavage of aromatic rings in thepresence of oxygen is possible not only in monomerproducts of destruction, but in lignin polymer as well.However, this processes has lesser probability than thesplitting of other, less strong bonds.

—Under anaerobic conditions, lignin macromole�cules are almost not involved in metabolism, unlikethe oligomeric products of its degradation, which canbe metabolized by bacteria–anaerobes, which attackphenols in the denitrification, sulfidogenesis, andmethanogenesis processes.

—Lignin degradation is not always complete: apart of it can be transformed into a highly condensedproduct, because of the competition between destruc�tion processes and reactions of polymerization andpolycondensation. Condensed lignins, containingdiphenyl bonds, are very tolerant to the impact of fun�gus enzymes.

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