Novel Haloperoxidase from the Agaric Basidiomycete Agrocybe aegerita Oxidizes Aryl Alcohols and Aldehydes

Agrocybe aegerita, a bark mulch- and wood-colonizing basidiomycete,was found to produce a peroxidase (AaP) that oxidizes aryl alcohols,such as veratryl and benzyl alcohols, into the correspondingaldehydes and then into benzoic acids. The enzyme also catalyzedthe oxidation of typical peroxidase substrates, such as 2,6-dimethoxyphenol(DMP) or 2,2'-azinobis-(3-ethylbenzothiazoline-6-sulfonate)(ABTS). A. aegerita peroxidase production depended on the concentrationof organic nitrogen in the medium, and highest enzyme levelswere detected in the presence of soybean meal. Two fractionsof the enzyme, AaP I and AaP II, which had identical molecularmasses (46 kDa) and isoelectric points of 4.6 to 5.4 and 4.9to 5.6, respectively (corresponding to six different isoforms),were identified after several steps of purification, includinganion- and cation-exchange chromatography. The optimum pH forthe oxidation of aryl alcohols was found to be around 7, andthe enzyme required relatively high concentrations of H₂O₂ (2mM) for optimum activity. The apparent K_m values for ABTS, DMP,benzyl alcohol, veratryl alcohol, and H₂O₂ were 37, 298, 1,001,2,367 and 1,313 µM, respectively. The N-terminal aminoacid sequences of the main AaP II spots blotted after two-dimensionalgel electrophoresis were almost identical and exhibited almostno homology to the sequences of other peroxidases from basidiomycetes,but they shared the first three amino acids, as well as twoadditional amino acids, with the heme chloroperoxidase (CPO)from the ascomycete Caldariomyces fumago. This finding is consistentwith the fact that AaP halogenates monochlorodimedone, the specificsubstrate of CPO. The existence of haloperoxidases in basidiomycetousfungi may be of general significance for the natural formationof chlorinated organic compounds in forest soils.

INTRODUCTION

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Introduction

Heme peroxidases are found in plants, fungi, bacteria, and animalsand have been grouped on the basis of sequence similarity intotwo superfamilies; animal peroxidases form one superfamily (18),and plant, fungal, and bacterial peroxidases form another superfamily(45). Peroxidases belonging to the latter superfamily have beenproposed to have various functions in the individual organisms,including roles in the polymerization and detoxification ofphenolic compounds in secondary metabolism, as well as in thedecomposition of wood and humus (7, 32).

Over the last two decades, particular attention has been paidto the peroxidases of white rot fungi, which are involved inthe biodegradation of lignin, humic materials, and organopollutants(13, 16, 33). These enzymes are capable of oxidizing recalcitrantaromatic molecules by one-electron abstractions, resulting inthe formation of unstable radicals, which tend to disintegratespontaneously (19). Numerous isozymes of lignin peroxidase (LiP)and manganese peroxidase (MnP), as well hybrid forms of bothenzymes (the so-called versatile peroxidases), have been purifiedand characterized from different basidiomycetes, including Phanerochaetechrysosporium, Phlebia radiata, Trametes versicolor, Pleurotuseryngii, Bjerkandera spp., and Stropharia coronilla (14, 31).In addition to these ligninolytic enzymes, which preferentiallyoxidize nonphenolic aromatic compounds and/or Mn²⁺ ions, therehave been reports of other peroxidases from basidiomycetousfungi that resemble plant peroxidases and oxidize phenolic andamino aromatic compounds (e.g., peroxidases from Coprinus cinereusand related species and versatile peroxidases) (15, 25, 26).Heme haloperoxidases, which introduce chlorine, bromine, oriodine into organic molecules and also oxidize various aromaticand aliphatic compounds, are other examples of versatile fungalheme proteins; however, they have not been found in basidiomycetesto date, although they have been found in several ascomycetes(6, 12, 27).

In this paper, we describe a novel type of peroxidase from thebasidiomycetous fungus Agrocybe aegerita, a common edible mushroomin Mediterranean countries, which oxidizes aryl alcohols andaldehydes at neutral pH and shows haloperoxidase activity.

MATERIALS AND METHODS

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Materials and Methods

Organisms.
A. aegerita (Brig.) Sing. (a synonym of Agrocybe cylindracea;black poplar mushroom) is an agaric fungus that colonizes deciduouswood and bark mulch, preferably stumps of poplar trees. Thisfungus is found in North America, Europe, and Asia, and it seemsto prefer warm or mild climates. A. aegerita is a popular ediblemushroom in southern Europe, especially in Italy (Pioppino mushroom),where it is also commercially cultured (37). We performed mostexperiments with A. aegerita TM A1, which was isolated by G.Gramss from fungal fruiting bodies that developed on a wheatstraw pile in Jena, Germany. For purposes of comparison, thefollowing strains obtained from the Deutsche Sammlung fürMikroorganismen, Braunschweig, Germany (DSM), and the Centraalbureauvoor Schimmelcultures, Baarn, The Netherlands (CBS), or depositedin the culture collection of the Institute of Microbiology,University of Jena, Jena, Germany (TM strains) were used: A.aegerita TM A1, CBS 358.51, CBS 127.88, TM ae, and TM AAN (=DSM 9613), Agrocybe praecox DSM 3355, Agrocybe vervactii TMN18, and Agrocybe dura A71-2 (= DSM 8353).

Culture conditions.
Fungal stock cultures were maintained on malt extract agar inculture slants and were stored at 4°C in the dark. The funguswas routinely precultured on 2% malt extract agar plates for2 weeks. The contents of agar plates were homogenized in 40ml of a sterile NaCl solution (0.9% NaCl), and each mycelialsuspension was used to inoculate liquid cultures (2.5 to 5%,vol/vol). The basal liquid medium contained 30 g of soybeanmeal (Hensel Voll-Soja; Schoeneberger GmbH, Magstadt, Germany)per liter. In addition to soy meal, various concentrations ofBacto Peptone (Difco Laboratories, Detroit, Mich.) were tested.Fungal cultures were agitated in 500-ml flasks containing 200ml of medium on a rotary shaker (100 rpm) at 24°C in thedark for 14 days. Samples were taken every 1 to 3 days, andthe activities of peroxidases and laccase were measured.

The medium used for production of larger amounts of A. aegeritaperoxidase (AaP) in a 5-liter stirred-tank bioreactor (BiostatB; Braun Biotech International GmbH, Melsungen, Germany) consistedof 20 g of soybean meal per liter and 5 g of Bacto Peptone perliter and was inoculated with 200 ml of a fungal suspensionprecultured as described above (fermentation parameters: agitationat 300 rpm and 100% pO₂ for dissolved oxygen concentration).The peroxidase and laccase activities, as well as the pH, weredetermined every 1 to 2 days during a total fermentation periodof 11 days.

Chemicals.
2,2'-Azinobis-(3-ethylbenzothiazoline-6-sulfonate) (ABTS), 2,6-dimethoxyphenol(DMP), veratryl alcohol, veratraldehyde, veratric acid, benzylalcohol, benzaldehyde, and anisyl alcohol, as well as glucoseoxidase (from Aspergillus niger), were purchased from Sigma-Aldrich(Steinheim, Germany). Chloroperoxidase (CPO) from Caldariomyces(Leptoxyphium) fumago and 2-chloro-5,5-dimethyl-1,3-cyclohexanedione(monochlorodimedone [MCD]) were obtained from Fluka (Buchs,Switzerland).

Enzyme assays and UV-visible spectra.
The activity of AaP was measured at 310 nm by monitoring theoxidation of veratryl alcohol into veratraldehyde ( ${varepsilon}$ ₃₁₀ = 9.3mM^–1 cm^–1). Unlike the LiP reaction (40), the reactionwas carried out at pH 7 in sodium phosphate/citrate buffer (McIlvainebuffer) and was started by addition of 2 mM H₂O₂. AaP activitieswith benzyl alcohol and benzaldehyde were measured under identicalconditions by monitoring either the formation ( ${varepsilon}$ ₂₈₀ = 1.4 cm^–1mM^–1) or the conversion ( ${varepsilon}$ ₃₀₀ = 0.6 cm^–1 mM^–1)of benzaldehyde. Oxidation of DMP was determined by using thesame assay mixture and an ${varepsilon}$ ₅₆₉ of 49.6 cm^–1 mM^–1(44); for ABTS oxidation ( ${varepsilon}$ ₄₂₀ = 36 cm^–1 mM^–1) thepH of the buffer was adjusted to 5. Laccase activity was alsodetermined with ABTS, but H₂O₂ was omitted from the reactionmixture (9).

Haloperoxidase activity was measured quantitatively by monitoringthe chlorination or bromination of MCD ( ${varepsilon}$ ₂₉₀ = 20.1 cm^–1mM^–1) in potassium phosphate buffer (pH 2.75) (12).

For comparison, commercial CPO from the ascomycete C. (L.) fumago(Fluka) was tested by using conditions identical to those usedin the AaP assays. All enzyme activities were expressed in units(micromoles of product formed per minute or micromoles of substrateconverted per minute).

UV-visible spectra of resting, oxidized, and reduced AaP wererecorded in 10 mM sodium phosphate buffer (pH 7) in the wavelengthrange from 200 to 700 nm by using a Lambda 2 spectrophotometer(Perkin-Elmer, Fremont, Calif.). Oxidation and reduction ofAaP (3.3 µM) were achieved by addition of H₂O₂ (15 or100 mM) and a small crystal of sodium dithionite, respectively.

Enzyme purification and characterization.
Culture fluid (3.9 liters) from the liquid fermentation wasfrozen (–20°C) and then thawed to precipitate thefree extracellular glucan that was produced by the fungus underthe culture conditions used. Precipitated material was removedby centrifugation and subsequent filtration through glass fiberfilters. Filtrates were concentrated 60-fold by ultrafiltrationby using a tangential-flow cassette (Omega open channel; molecularmass cutoff, 10 kDa; Pall-Filtron, Dreieich, Germany) and a250-ml filter chamber unit equipped with a 10-kDa-cutoff polysulfonemembrane (Amicon, Beverly, Mass.).

Crude enzyme was purified further by several steps of ion-exchangechromatography by using Q Sepharose, SP Sepharose, and MonoS columns, as well as an ÄKTA fast protein liquid chromatography(FPLC) system (Amersham Biosciences, Freiburg, Germany). Separationwas carried out by using sodium acetate (10 mM, pH 4.25 to 6.0)as the solvent and an increasing sodium chloride gradient (0to 0.3 or 0.6 M) for protein elution (see Fig. 2).

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FIG. 2. FPLC elution profile of peroxidases from A. aegerita TM A1 on a Mono S column. Absorption at 405 nm (thick line) and 280 nm (thin line), AaP activity (•), and the NaCl gradient (dotted line) were determined. AaP activity was measured with veratryl alcohol as the substrate.

Molecular weights of proteins were determined by sodium dodecylsulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) (HoeferSE 600 Ruby; Amersham Biosciences) as described by Laemmli (21).After electrophoresis, the gels were stained, and protein bandswere visualized with AgCl. A low-molecular-weight protein calibrationkit (MBI Fermentas, St. Leon Roth, Germany) was used as thestandard. Two-dimensional (2-D) electrophoresis (SDS-PAGE followedby isoelectric focusing) was performed by using a MultiphorII electrophoresis system (Amersham Biosciences) and DryStripgels (pH 3 to 11), which were cut to obtain a final pH rangefrom 3 to 7; staining of 2-D gels was carried out with guaiacolor Coomassie blue.

For N-terminal amino acid analysis, purified AaP was separatedby 2-D electrophoresis, and the spots of the AaP II fraction(spots a and b) stained with Coomassie blue (see Fig. 3B) weretransferred by electroblotting with a Trans Blot cell (Bio-Rad,Munich, Germany) onto a polyvinylidene difluoride membrane (ProBlott;Perkin-Elmer, Applied Biosystems, Foster City, Calif.). N-terminalamino acid sequences were determined by Sequence LaboratoriesGmbH, Göttingen, Germany.

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FIG. 3. Electrophoretic characterization of purified AaP. (A) SDS-PAGE after each purification step. From left to right the lanes contained crude extract, AaP fractions after Q Sepharose separation, AaP I after SP Sepharose separation, AaP I after Mono S separation, protein standards, AaP II after SP Sepharose separation, and AaP II after Mono S separation. (B) 2-D plot of purified AaP II after isoelectric focusing and SDS-PAGE. Spots b (basic form) and a (acidic form) were plotted and used for determination of N-terminal amino acid sequences.

The apparent Michaelis-Menten constants (K_m) and catalytic constants(k_cat) of purified AaP II were determined for benzyl alcohol,veratryl alcohol, ABTS, DMP, and H₂O₂ (the expression apparentK_m is used because peroxidases do not operate by classic Michaelis-Mentenkinetics). Lineweaver-Burk plots were prepared from the initialrates obtained with various substrate concentrations while theconcentration of the second substrate was kept constant.

Substrate oxidation experiments.
The following substrates (200 µM) were treated with purifiedAaP (1 U [= 12 µg = 0.26 µM]) in stirred 3-ml vialscontaining 1 ml of a reaction solution: veratryl alcohol, anisylalcohol, benzyl alcohol, and ethanol. Each reaction mixturecontained sodium phosphate/citrate buffer (pH 7), 15 mM glucose,and 0.2 U of glucose oxidase (Sigma-Aldrich). The reaction mixtureswere incubated at 25°C, and the alcohol and aldehyde concentrationswere determined after 20 and 30 min. High-performance liquidchromatography analyses were performed by using a model HP 1090liquid chromatograph (Agilent, Waldbronn, Germany) equippedwith a Merck LiChrospher 5-µm RP-18 reversed-phase column(4.6 by 125 mm). A mixture of acetonitrile and 0.05% aceticacid (40:60, vol/vol) was used as the solvent at a flow rateof 1 ml min^–1 under isocratic conditions. Eluted substanceswere detected in the wavelength range from 190 to 550 nm andwere identified by using authentic standards.

RESULTS

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Results

Peroxidase production.
A. aegerita produced appreciable amounts of a peroxidase (AaP)that oxidized veratryl alcohol at pH 7 only in N-rich, complexliquid media that contained soybean meal. All strains of thefungus tested secreted the enzyme, but there were considerabledifferences in the individual enzyme levels (range, 5 to 1,336U liter^–1) (Table 1). A. aegerita TM A1 proved to be themost active strain and was used for all further studies. Theother Agrocybe spp. tested did not show any peroxidase activityin the soybean medium.

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TABLE 1. Peroxidase production by several strains of A. aegerita, as well as three other species of the genus Agrocybe, in soybean meal medium

Increasing the amount of soybean meal in the medium (from 12.5to 60 g liter^–1) resulted in an almost linear increasein the AaP activity; in the absence of soybean meal, AaP activitywas not detectable (Fig. 1A). The maximum AaP activity (2,021U liter^–1) was found in the presence of 60 g of soybeanmeal per liter. Bacto Peptone (0 to 20 g liter^–1) furtherstimulated enzyme production; however, the effect was less pronounced,and Bacto Peptone was not required (Fig. 1A).

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FIG. 1. (A) Effect of soybean meal ( ${blacktriangleup}$ ) and Bacto Peptone ( ${blacksquare}$ ) on the production of peroxidase by A. aegerita TM A1. The concentration of one component was varied in the presence of a constant amount of the other component (5 g of Bacto Peptone per liter or 20 g of soybean meal per liter). (B) Time course of peroxidase and laccase titers in a 5-liter bioreactor with a soybean meal-Bacto Peptone medium. Peroxidase activity (•) was measured by determining the oxidation of veratryl alcohol into veratraldehyde at pH 7; laccase activity ( ${blacklozenge}$ ) was measured with ABTS. The dotted line indicates the time course of pH.

To obtain larger quantities of AaP for the subsequent purificationprocedure, A. aegerita TM A1 was cultured in a stirred-tankbioreactor. Figure 1B shows the fermentation profile for a 5-literdevice. Due to the high viscosity of the growth medium, smalleramounts of soybean meal and Bacto Peptone (20 and 5 g liter^–1,respectively) were used. In the course of fermentation, thepH of the medium increased from 5.7 to 8.6. At the beginningof the experiment the amount of dissolved O₂ was kept at 100%by constantly bubbling air into the medium (2 liter min^–1),but due to the consistency of the soybean slurry, it droppedto 30% after 5 days of fermentation. Production of AaP startedon day 6, and the maximum activity (1,550 U liter^–1) wasreached after 10 days of cultivation. In addition to AaP, thefungus produced laccase, but the level of laccase was noticeablylower (maximum activity, 290 U liter^–1). Activities ofMnP and LiP were not detectable.

Purification of AaP.
Culture filtrate from the bioreactor was harvested after 10days and frozen to remove dissolved polysaccharides. This procedureled, after centrifugation, to loss of about 45% of the activity,which can be explained by the partial adsorption of AaP to theprecipitated polysaccharides (Table 2). After ultrafiltration,the concentrated crude extract was separated by three stepsof ion-exchange chromatography by using FPLC equipment. Figure2 shows the elution profile for the third purification stepwith a Mono S column (strong cation exchanger). Two fractionsof the enzyme (designated AaP I and AaP II) absorbing at 405nm (heme) and exhibiting peroxidase activity with veratryl alcoholwere distinguished. The final specific activities of both AaPfractions were around 165 U mg^–1, but the total activityof AaP II was considerably higher than that of AaP I (244 versus20 U); the Reinheitszahl values of AaP I and AaP II were 1.65and 1.85, respectively.

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TABLE 2. Purification of peroxidase from A. aegerita TM A1^a

Characterization of AaP.
SDS-PAGE revealed the same molecular mass (46 kDa) for bothAaP fractions (Fig. 3A). 2-D electrophoresis revealed that therewas little difference in the isoelectric points. The two AaPfractions each split into three spots; the pIs for AaP I were4.9, 5.2, and 5.4, and the pIs for AaP II were 4.9, 5.3, and5.7, indicating the presence of six isoforms of the enzyme (Fig.3B). The most intense spots of AaP II (spots b and a) were blottedand used for determination of N-terminal amino acid sequences.

The UV-visible spectrum of the resting enzyme (AaP II afterpurification) had a characteristic absorption peak at 420 nm(Soret band), which gave the purified enzyme its characteristicreddish color, and two absorption maxima at 540 and 572 nm (Fig.4). Addition of catalytic amounts of H₂O₂ (0.2 to 15 mM) ledto only slight changes in the spectrum, and the maxima did notshift (Fig. 4); adding excess H₂O₂ (100 mM) resulted in theimmediate formation of gas bubbles and in a drastic decreasein the absorption intensity, but again no shift of the absorptionmaxima was observed. Addition of sodium dithionite to the restingenzyme caused a shift in the peak from 420 to 409 nm, and inthe longer-wavelength range, the peak at 572 nm became a shoulderand a new maximum appeared at 625 nm (Fig. 4).

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FIG. 4. Spectral characteristics of AaP (3.3 µM), including the resting enzyme (thick line), the enzyme oxidized by 15 mM H₂O₂ (dotted line) or 100 mM H₂O₂ (dashed line), and the reduced enzyme (after addition of a sodium dithionite crystal) (thin line).

The N-terminal peptide sequencing showed that the spots cutout after 2-D electrophoresis belonging to the AaP II fractionwere almost identical, apart from a few uncertain amino acidpositions that probably resulted from obstacles during the sequencingprocedure (Fig. 5). The levels of sequence identity to otherperoxidases from basidiomycetes were very low ( $≤$ 7%), but thefirst three amino acids, as well as two additional amino acids(positions 7 and 9), were identical to amino acids at the Nterminus of CPO from the ascomycete C. (L.) fumago (35% sequenceidentity).

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FIG. 5. N-terminal sequence alignment of two AaP II isoforms and several fungal peroxidases, including A. aegerita peroxidase (AaP II spot a and AaP II spot b after 2-D electrophoresis), CPO from C. fumago (CP), C. cinereus peroxidase (CiP), LiP H8 b from P. chrysosporium (PcLiP), MnP from A. bisporus (AbMnP), and versatile peroxidase from P. eryngii (PeVP).

Oxidation of different substrates.
Purified AaP II was used to examine the substrate spectrum ofthe enzyme. When a combined assay mixture in which glucose oxidasecontinuously generated H₂O₂ was used, AaP was able to oxidizeseveral aryl alcohols and aldehydes; ethanol was not subjectto AaP attack (Table 3). High-performance liquid chromatographyanalysis showed that benzyl and veratryl alcohols were firstconverted into benzaldehyde and veratraldehyde and then intobenzoic and veratric acids, respectively. The latter findingwas confirmed by the direct conversion of benzaldehyde and veratraldehydeinto the corresponding benzoic acids. Anisyl alcohol was alsooxidized into the corresponding aldehyde; however, the acidproduct could not be identified (due to the lack of authenticanisic acid). The following trends were observed for the oxidationof aryl alcohols and aldehydes by AaP: anisyl alcohol > benzylalcohol >> veratryl alcohol and benzaldehyde > anisaldehyde>> veratraldehyde.

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TABLE 3. Oxidation of several alcohols and aldehydes by purified AaP^a

Besides aryl alcohols and aldehydes, AaP was found to oxidizetypical peroxidase substrates, such as ABTS and DMP. Table 4shows the apparent K_m and k_cat values determined photometrically,as well as those of veratryl alcohol, benzyl alcohol, and H₂O₂.The pH optima of AaP for the oxidation of ABTS, benzyl alcohol,and DMP differed noticeably. Whereas the curves for benzyl alcohol,veratryl alcohol, and DMP oxidation showed broad, almost neutralpH optima (pH 6.8 to 7.2), ABTS oxidation occurred in a narrowerpH range (pH 2 to 6.5), with an acidic maximum around pH 4.7(Fig. 6). Interestingly, AaP seems to have two pH optima forthe oxidation of benzyl alcohol and veratryl alcohol (pH 6.2and 7.2 and pH 5.5 and 7, respectively). AaP was found to beactive between pH 2.5 and 9 and at the latter pH and in thepresence of some substrates lost only 30 to 70% of its activity.The enzyme required relatively large amounts of peroxide; thus,in the presence of 5 mM veratryl alcohol, optimal activity wasobserved with 2 mM H₂O₂, but the enzyme still exhibited 35%of the maximum activity with 10 mM H₂O₂ (data not shown).

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TABLE 4. Apparent Michaelis-Menten constants (K_m) and catalytic constants (k_cat) of AaP II for veratryl alcohol, benzyl alcohol, ABTS, and DMP^a

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FIG. 6. Effect of pH on the oxidation of ABTS (0.3 mM) (•) and veratryl alcohol (5 mM) ( ${blacktriangleup}$ ) (A) and on the oxidation of DMP (1 mM) ( ${blacksquare}$ ) and benzyl alcohol ( ${blacklozenge}$ ) (B) by AaP II. Reactions were performed in sodium phosphate/citrate buffer in the presence of 2 mM H₂O₂ at 25°C. The data are means for three parallel experiments, and the error bars indicate standard deviations.

Comparison of AaP and CPO.
The halogenating activity of AaP was proved by monitoring thechlorination or bromination of MCD. For comparison, authenticheme CPO from C. fumago was tested under identical conditions.We found that AaP is in fact capable of chlorinating and brominatingMCD, although the specific halogenating activities (at leastat pH 2.75) were considerably lower than those of CPO (21- and14-fold lower for chloride and bromide, respectively) (Table5). On the other hand, the activity of AaP with ABTS at pH 5was 250 times higher than that of CPO, and the activities ofAaP with DMP and benzyl alcohol at neutral pH were 10 and 50times higher than those of CPO. Benzaldehyde, which was moderatelyoxidized by AaP, was not a substrate for CPO (even highly concentratedCPO was not capable of oxidizing benzaldehyde [Table 5]).

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TABLE 5. Oxidation of different substrates and halogenation of MCD by purified AaP and CPO from C. fumago^a

DISCUSSION

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Discussion

A. aegerita produces, in the presence of soybean meal, a peroxidase(AaP) that oxidizes aryl alcohols into the corresponding aldehydesand then into the corresponding benzoic acids. This enzyme alsocatalyzes the conversion of typical peroxidase substrates, suchas ABTS or DMP, and halogenates MCD. The optimum pH for nonhalogenatingAaP activities is around 7 (except for ABTS), and the enzymerequires relatively large amounts of H₂O₂. Two AaP fractions(AaP I and AaP II), which split further into six isoforms, canbe distinguished on the basis of the results obtained for theFPLC elution profiles and 2-D electrophoresis gels. The N-terminalamino acid sequence of the major AaP fraction (AaP II) showsalmost no homology to the sequences of other peroxidases frombasidiomycetes, but it shares the first three amino acids andtwo additional amino acids with the sequence of heme CPO fromthe ascomycete C. fumago.

CPOs are known from bacteria and ascomycetous fungi and areamong the most versatile oxidoreductases (7). Among the fungalenzymes, two types, vanadium and heme-thiol peroxidases, aredistinguished (5, 43). Both types are capable of chlorinatingand brominating organic substrates, but they also catalyze variousother oxidation reactions (7, 8, 38). So far, both fungal vanadiumand heme-thiol CPOs have been found only in ascomycetes andrelated deuteromycetes (2, 12, 24), although their presencein basidiomycetes was proposed some years ago (6) and very recentlythere have been indications that there is a putative CPO genein Agaricus bisporus (white button mushroom) (41). Brominatingactivities have been described for LiP and MnP, but these enzymesdo not chlorinate the assay substrate MCD (10, 36). On the otherhand, there is extensive information concerning absorbable organichalogens formed by basidiomycetes as secondary metabolites (6,11, 42). Most of the more than 80 halogenated metabolites identifiedfrom basidiomycetes to date are chlorinated, and they are producedby fungi belonging to 68 genera and 20 different families (6).Taxonomic hot spots are found in the order Agaricales, particularlyin the families Strophariaceae and Tricholomataceae (6, 42),which are closely related to the Bolbitiaceae, which is thefamily that A. aegerita belongs to.

Despite the fact that AaP shares the first three amino acidsof the N terminus with Caldariomyces CPO and halogenates MCD,the former enzyme differs from the known heme CPOs in its pHbehavior, substrate spectrum, and specificity, as well as inthe UV-visible spectrum. Thus, the spectrum of the resting enzymeshows the Soret maximum at 420 nm and additional absorptionmaxima at 359, 540, and 572 nm. Interestingly, these maximacorrespond with those reported for CPO compound II (358, 438,542, and 571 nm) rather than with those of the native enzyme(403, 515, 542, and 650 nm) (7, 12). AaP reduced by sodium dithioniteshowed more similarity to the corresponding form of CPO. Ithad a Soret maximum at 409 nm, a shoulder in the 460-nm region,and two peaks around 550 and 640 nm; only the shoulder at 420nm was lacking in the reduced form of AaP.

The substrate spectrum of AaP comprises phenolic compounds,ABTS, aryl alcohols, and aldehydes, as well as halogenides (Cl^–,Br^–). CPO is also able to oxidize most of these substrates;however, the specific activities (except those for halogenides)are lower, and aldehydes are apparently not CPO substrates.CPO-catalyzed oxidation of aryl alcohols into the correspondingaldehydes has been described for the Caldariomyces enzyme (1).Similar to AaP, this CPO converted benzyl and anisyl alcoholsinto the corresponding aldehydes; however, the specific constantsreported for CPO are considerably lower than those of AaP. TheCPO reaction with aryl alcohols has been proposed to occur viaan attack on the benzylic carbon, resulting in the formationof the benzylic radical and cation (1); a similar mechanismis also conceivable for AaP. Furthermore, it was found thatCPO possesses prochiral selectivity in the oxidation of arylalkanols that makes the enzyme interesting for applicationsin chemical syntheses (1, 30). This is also true for AaP, especiallywhen its broader substrate spectrum and higher pH optima aretaken into consideration.

AaP seems to act not only as a peroxidase but also as a catalase,which was shown by the intensive formation of gas bubbles (veryprobably O₂) after addition of larger amounts of H₂O₂ in theabsence of a second available substrate. CPO from Caldariomycesalso has strong catalase activity, and there is a whole classof so-called catalase-peroxidases that are found in both prokaryotesand eukaryotes (8, 39).

CPO production by C. fumago was carried out at a larger scalein complex media comprising fructose and Phytone (papaic digestof soybean meal) in airlift fermentors (3), and the enzyme cannow be produced as a recombinant protein (e.g., in A. niger)(4). Culturing A. aegerita in a heavily stirred bioreactor waspossible, and the enzyme production was only slightly reduceddue to shear stress. For AaP production, soybean meal was foundto be the key nutrient, and the concentration of the meal correlatedwith the AaP activity. Complex N-rich media were also foundto promote fungal growth and peroxidase production in otherbasidiomycetes. Thus, mixtures of Bacto Peptone, yeast extract,and sugars enabled the production of large amounts of versatileperoxidase (a hybrid of MnP and LiP) by Bjerkandera sp. strainBOS55 and P. eryngii (23, 25). On the other hand, nitrogen wasshown to repress the production of ligninolytic peroxidasesin other fungi (for example, P. chrysosporium or P. radiata)(20, 22). A medium quite similar to that used in our study,consisting of soybean meal, Bacto Peptone, and maltose, wassuccessfully used to produce laccases and peroxidases in a stirredbioreactor with dung- and grassland-dwelling agaric basidiomycetes(Panaeolus spp., Coprinus friseii) (15).

Two isoenzymes of Caldariomyces CPO, isoenzymes A and B, werereported to have molecular masses of 40 and 46 kDa; later workshowed that there are multiple forms that differ only in thecarbohydrate content, which varies between 20 and 30% (17, 34).We also found two fractions of AaP based on the FPLC elutionprofiles, and there were indications of multiple forms of theenzyme (at least six isoforms) that became visible in the SDS-PAGEand 2-D gel electrophoresis plots.

At the moment, it is difficult to speculate about the physiologicalrole of AaP. According to previously published reports, involvementof the enzyme in the production of halogenated metabolites thatact as antibiotics or as part of the ligninolytic system canbe proposed (6). This possibility is particularly interestingbecause A. aegerita apparently lacks MnP and LiP (32) but colonizeswood and causes, according to some authors, a moderate whiterot (35, 46). Moreover, the ability of Caldariomyces CPO tochlorinate and cleave recalcitrant lignin structures has beenreported very recently (29), suggesting that this type of enzymecould be involved in the biodegradation of lignocellulosic materials.

Hence, future studies on AaP should focus on two questions.(i) What is the physiological function of AaP, and is the enzymesomehow involved in lignin biodegradation? (ii) Do other basidiomycetes,particularly those secreting large amounts of absorbable organichalogens, produce AaP-like peroxidases? If so, what ecologicaland environmental significance does this type of halogenationof organic compounds, for example, have for the recently reportedformation of stable chlorinated hydrocarbons in weathering plantmaterial (28)? In this context, the substrate spectrum of AaPshould be studied more in detail and compared with the substratespectra of CPO and other peroxidases. The first attempts inthis direction are currently being undertaken in our laboratories.

ACKNOWLEDGMENTS

This work was supported financially by the German EnvironmentalFoundation (DBU) (grant 13663 to R.U., J.N., and K.S.), theAcademy of Finland (grant 52063 to M.H.), and the administrationof the International Graduate School of Zittau (R. Konschak,Zittau, Germany).

We thank G. Gramss (Jena, Germany) for providing the fungalstrain A. aegerita TM A1, R. Upadhyay (Solan, India) for usefulpreinvestigations, and I. Schwabe (Jena, Germany) and U. Schneider(Zittau, Germany) for excellent technical assistance.

FOOTNOTES

* Corresponding author. Mailing address: Unit of Environmental Biotechnology, International Graduate School of Zittau, Markt 23, D-02763 Zittau, Germany. Phone: 49 3583 771521. Fax: 49 3583 771534. E-mail: ullrich{at}ihi-zittau.de.

REFERENCES

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