Identification of genes for lignin peroxidases and manganese peroxidases in ectomycorrhizal fungi


  • David M. Chen,

    1. Mycorrhiza Research Group, School of Science Food & Horticulture, University of Western Sydney, Parramatta Campus, Locked Bag 1797, PENRITH SOUTH DC, NSW 1797, Australia;
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  • Andrew F. S. Taylor,

    1. Department of Forest Mycology & Pathology, Swedish University of Agricultural Sciences, Box 7026, S-750 07 Uppsala, Sweden;
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  • Ron M. Burke,

    1. Department of Biomolecular Sciences, UMIST, PO Box 88, Manchester M60 1QD, UK
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  • John W. G. Cairney

    Corresponding author
    1. Mycorrhiza Research Group, School of Science Food & Horticulture, University of Western Sydney, Parramatta Campus, Locked Bag 1797, PENRITH SOUTH DC, NSW 1797, Australia;
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Author for correspondence: John W. G. Cairney Tel: +61 29685 9903 Fax: +61 29685 9915


  • • Genes for ligninolytic enzymes, normally associated with white-rot fungi, are shown to be widespread in a broad taxonomic range of ectomycorrhizal (ECM) fungi.
  • • ECM fungi were screened for lignin peroxidase (LiP) and manganese peroxidase (MnP) genes by PCR using primers specific for known isozymes in the white-rot fungus Phanerochaete chrysosporium, with DNA sequencing used to confirm the identity of the amplified fragments.
  • • Genes for LiPs were detected in ECM fungi representing the orders Agaricales, Aphyllophorales, Boletales, Cantharellales, Hymenochaetales, Sclerodermatales, Stereales and Thelephorales. MnP genes were detected in only Cortinarius rotundisporus and three ECM Stereales taxa.
  • • The presence of genes for decomposer activities supports putative evolutionary relationships between ECM and saprotrophic fungi. Expression of the lignolytic genes may facilitate ECM fungal access to nutrients associated with dead plant material in soil and potentially a supplementary carbon supply. Strict functional boundaries between ECM and decomposer fungi may be less clear-cut than previously thought.


Ectomycorrhizal (ECM) associations are regarded as important in tree nutrition and as key components of mineral nutrient and carbon cycling processes in temperate, boreal and some tropical forest habitats (Smith & Read, 1997). The fungi that form ECM associations are largely basidiomycetes and belong to a broad range of taxonomic groupings, many of which include extant wood-decomposing species (Bruns et al., 1998; Kretzer & Bruns, 1999; Hibbett et al., 2000). Phylogenetic analyses of DNA sequences from extant taxa suggest that ECM fungi evolved convergently from saprotrophic ancestors several times (Bruns et al., 1998; Kretzer & Bruns, 1999). We currently know little regarding the physiological ecology of most extant ECM fungi and, although generally treated as separate ecological guilds, the extent to which ectomycorrhizal fungi have diverged physiologically from their saprotrophic ancestors remains unclear. The apparent evolutionary patterns of ECM fungi, however, suggest that the diverse communities of ECM fungi in extant forest habitats may contain considerable functional diversity, particularly with respect to their abilities to acquire nutrients from dead plant material in soil (Bruns, 1995; Cairney, 2000). Furthermore, the observation that some evolutionary reversals from the ECM to the saprotrophic condition appear to have occurred in some basidiomycete taxa, strongly implies that, at least some, ECM fungi have retained genes for decomposer activities (Hibbett et al., 2000).

While it has been accepted for some time that ECM fungi are capable of absorbing and translocating inorganic nutrients to their plant hosts (Smith & Read, 1997), the abilities of some ECM fungi to access organic forms of nitrogen and phosphorus have been identified only relatively recently (Leake & Read, 1997). There is also increasing cognizance that saprotrophic capabilities of some ECM fungi extend beyond utilization of simple organic forms of these elements. Some ECM fungi can acquire nitrogen and/or phosphorus from patches of partly decomposed litter in soil and effect their transfer to the host (Entry et al., 1991; Bending & Read, 1995a; Pérez-Moreno & Read, 2000). The extent to which ECM fungi produce the oxidative and/or hydrolytic enzyme activities required to release nitrogen and phosphorus complexed with, or sequestered within, dead plant tissue, however, remains unclear (Cairney & Burke, 1996b; Sen, 2000).

Production of cellulolytic activities by several ECM fungal taxa can be inferred from the observed degradation of 14C-labelled substrates in sterile systems (Durall et al., 1994), and cellulolytic activities have been experimentally measured in culture filtrates of some ECM taxa (Cairney & Burke, 1994, 1996b). Several phenol oxidase and peroxidase enzyme activities, including tyrosinase (EC, catechol oxidase (EC, laccase (EC, nonspecific peroxidase (EC and manganese peroxidase (MnP) (EC, have been shown to be associated with extramatrical mycelia of ECM fungi in nonsterile soil (Griffiths & Caldwell, 1992; Bending & Read, 1995b; Colpaert & van Laere, 1996; Gramms, 1997; Timonen & Sen, 1998). Although these activities may have arisen from the ECM fungi, it is not possible to separate these from enzymes produced by mycorrhizosphere microflora (Sun et al., 1999).

A number of ECM fungi have been reported to produce nonspecific phenol oxidase activities in axenic cultures (Giltrap, 1982; Bending & Read, 1997; Günther et al., 1998; Kanunfre & Zancan, 1998). While the caveat outlined below regarding culture media may apply in some of these cases, production of laccase by Thelephora terrestris Ehrh. Fr. has been clearly demonstrated (Kanunfre & Zancan, 1998). Growth of some ECM fungal taxa in axenic cultures containing lignin or dehydrogenative polymers of lignin monomers has been shown to result in partial mineralization of the compounds (Trojanowski et al., 1984; Haselwandter et al., 1990). While this has often been taken to indicate an ability of these fungi to produce lignin peroxidase (LiP) (EC and MnP (Griffiths & Caldwell, 1992; Cairney & Burke, 1994, 1996b), this is not necessarily the case. The concentrations of iron included in the growth media were at levels known to facilitate chemical generation of hydroxyl radicals from H2O2, probably resulting in partial lignin degradation via a mechanism akin to that employed by brown rot fungi (Burke & Cairney, 1998; Cairney & Burke, 1998). Indeed, when assays for LiP activities in ECM fungi have been conducted so as to avoid these potential artefacts, no activity has been measured (Bending & Read, 1996; Burke & Cairney, 1998; Chambers et al., 1999). In the case of MnP, however, extracellular activity has been measured and a DNA sequence for part of a gene identified in Tylospora fibrillosa (Burt.) Donk (Chambers et al., 1999).

Expression of LiP and other ligninolytic enzymes in axenic culture is strongly influenced by the composition of the external medium, and screening for genes that encode the enzymes is regarded as a more reliable means of identifying potential enzymatic activities in diverse arrays of fungi (Varela et al., 2000). Furthermore, many ECM taxa that are numerically important in below-ground communities are difficult to culture, rendering biochemical assays of enzyme activities impossible. In this study we have adopted an isozyme-specific PCR approach (Brooks et al., 1993) to screen for genes encoding several LiP and MnP isozymes in a broad taxonomic range of ECM fungi.

Materials and Methods

Fungal material and DNA extraction

Forty-four ECM, along with three putative ECM, basidiomycetes (representing 20 families from eight orders) were screened (Table 1). For most fungi, DNA was extracted from dried basidiome material, however, cultured mycelia (maintained on Modified Melin-Norkrans agar medium (Marx & Bryan, 1975)) were used for extraction in some cases (see Table 1). Mycelium of Phanerochaete chrysosporium Burdsall, grown as described by Burke & Cairney (1998), was included for comparative purposes.

Table 1.  Details of fungi used in PCR-based screening for lignin peroxidase (LiP) and manganese peroxidase (MnP) genes
SpeciesOrderFamilyOriginTissueAccession code
  • *

    Fungi known to be ectomycorrhizal;

  • †fungi thought to be ectomycorrhizal.

Amanita fulva (Schaeff) Pers.*AgaricalesAmanitaceaeSwedenbasidiomeSLU AT99030
Amanita spissa (Fr.) Kumm.*AgaricalesAmanitaceaeSwedenbasidiomeSLU AT99001
Cortinarius rotundisporusAgaricalesCortinariaceaeAustraliabasidiomeUWS CR001
Cortinarius semisanguineus (Fr.) Gill.*AgaricalesCortinariaceaeSwedenbasidiomeSLU AT20134
Cortinarius violaceus (L.Fr.) S.F. Gray*AgaricalesCortinariaceaeSwedenbasidiomeSLU AT20133
Inocybe maculata Boud.*AgaricalesCortinariaceaeSwedenbasidiomeSLU AT20135
Rozites caperatus (Pers.Fr.) Karst*AgaricalesCortinariaceaeSwedenbasidiomeSLU AT20137
Hygrophorus erubescens (Pers.Fr.) Fr.*AgaricalesHygrophoraceaeSwedenbasidiomeSLU AT20080
Hygrophorus latitabundus Britz.*AgaricalesHygrophoraceaeSwedenbasidiomeSLU AT20080
Laccaria laccata (Scop.Fr.) Berk. & Br.*AgaricalesTricholomataceaeSwedenbasidiomeSLU AT20140
Tricholoma fulvum (DC.Fr.) Sacc.*AgaricalesTricholomataceaeSwedenbasidiomeSLU AT20130
Tricholoma terreum (Schaeff.Fr.) Kumm.*AgaricalesTricholomataceaeSwedenbasidiomeSLU AT20129
Lactarius rufus (Scop.Fr.) Fr.*RussulalesRussulaceaeSwedenbasidiomeSLU AM00006
Lactarius scrobiculatus (Scop.Fr.) Fr.*RussulalesRussulaceaeSwedenbasidiomeSLU AT20132
Russula adusta Fr.*RussulalesRussulaceaeSwedenbasidiomeSLU AT20052
Russula puellaris Fr.*RussulalesRussulaceaeSwedenbasidiomeSLU AT20039
Phanerochaete chrysosporium BurdsallAphyllophoralesCorticeaceaeunknowncultureATCC 34541
Ramaria cf flavicingula R.H. PetersenAphyllophoralesGomphaceaeSwedenbasidiomeSLU AT20111
Boletus impolitus Fr.*BoletalesBoletaceaeSwedenbasidiomeSLU AT20072
Boletus pinophilus Pilát & Dermek*BoletalesBoletaceaeSwedenbasidiomeSLU AM00008
Leccinum versipelle (Fr.) Snell*BoletalesBoletaceaeSwedenbasidiomeSLU AT20064
Leccinum vulpinum Watl.*BoletalesBoletaceaeSwedenbasidiomeSLU AM00005
Suillus flavidus (Fr.Fr.) J.S.Presl*BoletalesBoletaceaeSwedenbasidiomeSLU AT20128
Suillus grevillei (Klotzsch:Fr.) Sing.*BoletalesBoletaceaeSwedenbasidiomeSLU AT20096
Xerocomus badius (Fr.) Kühn. ex Gilb.*BoletalesBoletaceaeSwedenbasidiomeSLU AT20077
Xerocomus subtomentosus (L.Fr.) Quél*BoletalesBoletaceaeSwedenbasidiomeSLU AT20032
Chroogomphus rutilus (Schaeff.Fr.) O.K. Miller*BoletalesGomphideaceaeSwedenbasidiomeSLU AM00011
Gyroporus castaneus (Bull.Fr.) Quél*BoletalesGyrodontaceaeEnglandbasidiomeSLU AT96011
Paxillus involutus (Batsch:Fr.) Fr.*BoletalesPaxillaceaeSwedenbasidiomeSLU AT99106
Paxillus filamentosus (Scop.) Fr.*BoletalesPaxillaceaeSwedenbasidiomeSLU AT20042
Rhizopogon luteolus (Fr.)*BoletalesRhizopogonaceaeEnglandbasidiomeSLU AT96014
Strobilomyces floccopus (Scop.Fr.) Berk.*BoletalesStrobilomycetaceaeGermanybasidiomeSLU AT96028
Tylopilus felleus (Bull.Fr.) Karst.*BoletalesStrobilomycetaceaeSwedenbasidiomeSLU AT20138
Cantharellus cibarius Fr.*CantharellalesCantherellaceaeSwedenbasidiomeSLU AT99014
Hydnum repandum L. ex Fr.*CantharellalesHydnaceaeSwedenbasidiomeSLU AT5153
Albatrellus confluens (Alb. & Schwein.Fr.) Kolt. & Pouzar*CantharellalesScutigeraceaeSwedenbasidiomeSLU AT20054
Coltrichia perennis (L.Fr.) Murr.HymenochaetalesColtrichiaceaeSwedenbasidiomeSLU AT5105
Pisolithus (albus?)*SclerodermatalesSclerodermataceaeAustraliacultureUWS W52
Pisolithus (marmoratus?)*SclerodermatalesSclerodermataceaeAustraliacultureUWS LJ31
Piloderma croceum J. Erikss. & Ryvarden*SterealesAtheliaceaeSwedencultureSLU UP141
Piloderma byssinum (P.Karst.) Jülich*SterealesAtheliaceaeSwedencultureSLU UP185
Tylospora fibrillosa (Burt) Donk*SterealesAtheliaceaeNorwaycultureUWS TF01
Bankera fuligineo-alba (Schmidt ex Fr.) Pouz.*ThelephoralesBankeraceaeSwedenbasidiomeSLU AT5192
Phellodon tomentosus (L.Fr.) Banker*ThelephoralesBankeraceaeSwedenbasidiomeSLU AT20106
Hydnellum caeruleum (Horn. ex Pers.) Karst.*ThelephoralesThelephoraceaeSwedenbasidiomeSLU AT5193
Thelephora palmata (Scop.) Fr.*ThelephoralesThelephoraceaeSwedenbasidiomeSLU AT20136
Thelephora terrestris (Ehrh.) Fr.*ThelephoralesThelephoraceaeSwedenbasidiomeSLU AT5182
Tomentella fibrosa (Berk. & M.A. Curtis) Kõljalg*ThelephoralesThelephoraceaeEstoniabasidiomeSLU UK159737

Genomic DNA was extracted from dried basidiome material using the method of Cubero et al. (1999) and from cultured mycelia using the modified CTAB method of Gardes & Bruns (1993).

PCR analysis

PCR was conducted using primers that are specific for the N-terminal portions of genes for H8-like (LiP1 and LiP2), H2-like (LiP5) and H10-like (LiP6) LiP isozymes, along with H3-like (MnP2) and undefined (MnP1) isozymes from P. chrysosporium. Primer sequences have been published previously as follows: LiP1 = LIG1u/d, LiP2 = LIG2u/d, LiP5 = LIG5u/d, LiP6 = LIG6u/d, MnP1 = mnp1u/d (Broda et al., 1995); MnP2 = MP1u/d (Chambers et al., 1999). Amplifications were performed in a 50-µl reaction volume containing c. 50 ng genomic DNA, 10 pmol of the relevant primer pair, 50 mM KCl, 10 mM Tris-HCl, 0.1% Triton X-100, 2.5 mM MgCl2, 200 µM each of dATP, dCTP, dGTP and dTTP, and 2 units of Taq DNA polymerase (Promega, Madison, WI, USA). All amplifications were performed in a PTC-100 DNA Thermal Cycler® (MJ Research, Watertown, MA, USA) with 35 cycles of 94°C for 1 min, 60°C–70°C (MnP1, MnP2 = 60°C; LiP1, LiP5, LiP6 = 68°C; LiP2 = 70°C) for 1 min and 72°C for 1 min, followed by 72°C for 10 min (Broda et al., 1995). A negative control, containing no fungal DNA was included in each PCR reaction run to test for the presence of contaminating DNA. Amplification products were electrophoresed in 3% (w/v) agarose gels, stained with ethidium bromide and viewed under UV light. Where amplification products were faint, they were eluted and re-amplified by PCR using the same conditions.

DNA sequencing and sequence analysis

In order to confirm the identity of the bands obtained in the PCR amplifications, DNA sequences were obtained for products obtained using each primer pair from 1 to 12 randomly selected taxa. Before sequencing, each PCR product was purified using the Wizard PCR Purification System® (Promega) according to the manufacturer's instructions. Purified PCR products were cloned with the pGEM-T easy vector system® (Promega) and three clones for each isolate sequenced. Each strand was sequenced using pGEMf and pGEMr primers on an ABI 377 automatic sequencer (Applied Biosystems, Inc., Foster City, CA, USA). The final sequence was compared to the GenBank and EMBL nucleotide databases using the FASTA 3.0 program (Pearson & Lipman, 1988) and aligned with sequences having close identity using the Pileup program (within the EGCG extensions to the Wisconsin Package, Version 8.1.0 (Rice, 1996)). All sequences obtained were submitted to the GenBank nucleotide database (accession codes are provided in Table 3).

Table 3.  Closest matches from FASTA searches between DNA sequences of isozyme-specific PCR fragments amplified from various ectomycorrhizal fungi and sequences from the GenBank nucleotide database
SpeciesPrimer pairFragment sizeGenBank accession codeClosest match in FASTA searchSequence identityNucleotide overlap (bp)
T. terreumLiP1175 bpAF357258P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIG1A] 95.2%125 bp
P. filamentosusLiP1176 bpAF357259P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIG1A] 96.0%125 bp
C. rutilusLiP1186 bpAF357260P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIG1A] 98.5%135 bp
T. felleusLiP1176 bpAF357261P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIG1A] 92.0%125 bp
T. fulvumLiP1176 bpAF357262P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIG1A] 92.0%125 bp
L. scrobiculatusLiP1176 bpAF357263P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIG1A] 98.4%125 bp
C. rotundisporusLiP1177 bpAF357264P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIG1A] 95.2%126 bp
P. involutusLiP2209 bpAF357254P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIGH8]100.0%155 bp
T. fibrillosaLiP2209 bpAF357255P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIGH8]100.0%155 bp
P. byssinumLiP2209 bpAF357256P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLIGH8]100.0%155 bp
P. croceumLiP2209 bpAF357257P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCLP001] 98.1%155 bp
P. involutusLiP5348 bpAF357242P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 98.9%299 bp
G. castaneusLiP5348 bpAF357243P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 96.6%299 bp
T. fulvumLiP5348 bpAF357244P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 97.0%299 bp
C. rotundisporusLiP5348 bpAF357245P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 98.7%299 bp
Pisolithus. sp. ILiP5348 bpAF357246P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 99.0%299 bp
Pisolithus. sp. IILiP5348 bpAF357247P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 98.3%299 bp
T. terreumLiP5348 bpAF357248P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 97.9%299 bp
R. caperataLiP5349 bpAF357249P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 97.6%299 bp
P. filamentosusLiP5348 bpAF357250P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 97.3%299 bp
L. scrobiculatusLiP5348 bpAF357251P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 96.9%299 bp
P. byssinumLiP5348 bpAF357253P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 95.9%299 bp
P. croceumLiP5348 bpAF357252P. chrysosporium LiP gene (H2-like isozyme) [GenBank PCLIP2] 95.6%296 bp
H. repandumLiP6317 bpAF357265P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCGLG5] 99.2%266 bp
P. byssinumLiP6317 bpAF357266P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCGLG5] 97.7%266 bp
P. croceumLiP6316 bpAF357267P. chrysosporium LiP gene (H8-like isozyme) [GenBank PCGLG5] 95.1%266 bp
C. rotundisporusMnP1680 bpAF357268P. chrysosporium MnP gene (undefined isozyme) [GenBank PCMNP1] 86.8%296 bp
C. rotundisporusMnP2287 bpAF357269P. chrysosporium MnP gene (H3-like isozyme) [GenBank PCU70998] 98.7%236 bp
P. croceumMnP2291 bpAF357270P. chrysosporium MnP gene (H3-like isozyme) [GenBank PCU70998] 93.4%241 bp
T. fibrillosaMnP2260 bpAF107097P. chrysosporium MnP gene (H3-like isozyme) [GenBank PCU70998] 93.7%201 bp


PCR products of the predicted sizes (Brooks et al., 1993; Chambers et al., 1999) were obtained by amplification of DNA from P. chrysosporium using the LiP and MnP gene-specific primers (Fig. 1a–d, Table 2). Using the LiP1 primers, products (175–186 bp) were obtained from 22 taxa representing the orders Agaricales, Aphyllophorales, Boletales, Cantharellales, Sclerodermatales, Stereales and Thelephorales (Fig. 1a, Table 2). Similarly, products (348–349 bp) were obtained from 28 taxa (from seven orders) using the LiP5 primer pair (Fig. 1b, Table 2). By contrast to the LiP1-specific primers, no product was obtained for the putative ECM Aphyllophporales taxon (Ramaria sp.), but an appropriately sized product was obtained for the putative ECM Hymenochaetales taxon (Coltrichia perennis) (Table 2). Comparison of randomly selected sequences for the LiP1 and LiP5 fragments with sequences in the GenBank database indicated closest matches with sequences for H8- and H2-like LiP isozyme genes from P. chrysosporium, respectively (Table 3). Sequence comparisons indicated > 92% identity (over 125–126 bp) for LiP1 and > 95% identity (over 299 bp) for LiP5 with the P. chrysosporium sequences (Table 3), confirming the presence of equivalent LiP genes in the ECM and putative ECM fungi. For many taxa, appropriately sized fragments were obtained using both LiP1 and LiP5 primers (Table 2), suggesting that some ECM taxa possess genes for multiple LiP isozymes.

Figure 1.

PCR amplification of fragments from DNA of ectomycorrhizal (ECM) fungi using primer pairs specific for genes for lignin peroxidase (LiP) and manganese peroxidase (MnP) isozymes in Phanerochaete chrysosporium at optimal annealing temperatures. (a) LiP1 primers. M, pGEM molecular size marker (Promega); C, negative control containing no fungal DNA; 1, Cortinarius rotundisporus; 2, Tricholoma terreum; 3, Paxillus filamentosus; 4, Lactarius scrobiculatus; 5, Russula adusta; 6, Tricholoma fulvum; 7, Chroogomphus rutilus; 8, Xerocomus subtomentosus; 9, Phanerochaete chrysosporium. (b) LiP5 primers. M, pGEM molecular size marker (Promega); C, negative control containing no fungal DNA; 1, Paxillus involutus; 2, Gyroporus castaneus; 3, Tricholoma fulvum; 4, Cortinarius rotundisporus; 5, Pisolithus albus; 6, Pisolithus marmoratus; 7, Chroogomphus rutilus; 8, Lactarius scrobiculatus; 9, Tylopilus felleus; 10, Phanerochaete chrysosporium. (c) LiP2 primers (lanes 1–3), LiP6 primers (lanes 4 and 5); M, pGEM molecular size marker (Promega); C, negative control containing no fungal DNA; 1, Tylospora fibrillosa; 2, Paxillus involutus; 3 and 5, Phanerochaete chrysosporium; 4, Hydnum repandum. (d) MnP1 primers (lanes 1 and 2), MnP2 (lanes 3–5); M, pGEM molecular size marker (Promega); C, negative control containing no fungal DNA; 1 and 3, Cortinarius rotundisporus; 2 and 5, Phanerochaete chrysosporium; 4, Tylospora fibrillosa.

Table 2.  Fungi for which appropriately sized (✓) PCR products were obtained, and those for which no products were obtained (–) using the various lignin peroxidase(LiP)- and manganese peroxidase(MnP)-specific primers
Amanita. fulva
Amanita spissa
Cortinarius rotundisporus
Cortinarius semisanguineus
Cortinarius violaceus
Inocybe maculata
Rozites caperata
Hygrophorus erubescens
Hygrophorus latitabundus
Laccaria laccata
Tricholoma fulvum
Tricholoma terreum
Lactarius rufus
Lactarius scrobiculatus
Russula adusta
Russula puellaris
Phanerochaete chrysosporium 
Ramaria cf flavicingula
Boletus impolitus
Boletus pinophilus
Leccinum versipelle
Leccinum vulpinum
Suillus flavidus
Suillus grevillei
Xerocomus badius
Xerocomus subtomentosus
Chroogomphus rutilus
Gyroporus castaneus
Paxillus involutus
Paxillus filamentosus
Rhizopogon luteolus
Strobilomyces floccopus
Tylopilus felleus
Cantharellus cibarius
Hydnum repandum
Albatrellus confluens
Coltrichia perennis
Pisolithus albus
Pisolithus marmoratus
Piloderma byssinum
Piloderma croceum
Tylospora fibrillosa
Bankera fuligo-alba
Phellodon tomentosus
Hydnellum caeruleum
Thelephora palmata
Thelephora terrestris
Tomentella fibrosa

The LiP2 primers produced appropriately sized fragments (209 bp) from only Paxillus involutus (Boletales), along with the two Piloderma species and Tylospora fibrillosa (Stereales), while the LiP6 pair gave a 317-bp product for only Hydnum repandum (Cantharellales) and both Piloderma species (Fig. 1c, Table 2). DNA sequence comparisons indicated 100% identity (over 155 bp) for the P. involutus and T. fibrillosa LiP2 sequences and 99% identity (over 266 bp) for the H. repandum LiP6 sequence with H8-like isozyme gene sequences from P. chrysosporium (Table 3).

Appropriately sized products were obtained using the MnP2 primers from the three ECM Stereales taxa only (P. byssinum, P. croceum, and T. fibrillosa), along with the putative ECM taxon C. rotundisporus (Fig. 1d, Table 2). These products had > 93% identity (over 201–236 bp) with a sequence for an H3-like MnP isozyme from P. chrysosporium (Table 3). Only C. rotundisporus DNA yielded a product of the appropriate size (680 bp) using the MnP1 primers (Fig. 1d, Table 2). This product showed 86% identity (over 296 bp) with a gene for an undefined MnP isozyme from P. chrysosporium (Table 3).


Due to their perfect complementarity to the 3′-end of each specific LiP or MnP gene fragment and mismatches relative to nontarget sequences, the primers used in this study are considered specific (at optimal annealing temperatures) for individual LiP or MnP isozyme genes in P. chrysosporium (Brooks et al., 1993; Broda et al., 1995). Amplification of products of identical size to the corresponding P. chrysosporium products using each of the primer pairs thus strongly suggests the presence of equivalent genes for ligninolytic activities in a range of ECM and putative ECM fungi. The high percentage sequence identity (86–100%) between these products and corresponding fragments from the 5′-ends of the P. chrysosporium genes, provides confirmation of the presence of LiP- and MnP-type genes in these fungi. In all cases matches with bacterial sequences were < 70% (data not shown), confirming that the sequences from dried basidiome material were of fungal origin and were not associated with putative bacterial contaminants.

Based on phylogenetic analyses of rDNA sequence data from extant ECM and saprotrophic fungi, Hibbett et al. (2000) proposed that some ECM basidiomycetes have undergone evolutionary reversals to the saprotrophic habit. Such reversals require that the fungi have become independent from their symbiotic plant hosts in terms of carbon supply, relying instead upon carbon derived from decomposition of dead plant cell wall material in soil. Given the juxtaposition of ECM and litter-decomposing basidiomycete mycelia in organic soil layers, this would entail only a shift from the symbiotic to the free-living habit, without requirement for a change of habitat (Hibbett et al., 2000). It would, however, require that the antecedent ECM taxa had retained (from their ancestral saprotrophic states) genes for plant cell wall-degrading enzymes (Hibbett et al., 2000). Our data, suggesting that genes for ligninolytic enzymes are present in a widespread taxonomic range of ECM fungi, provides strong support for this hypothesis.

Genes equivalent to LiPs appear to be widespread in ECM basidiomycetes, with 68% of the taxa screened being found to possess at least one LiP gene. Of these, the genes identified using the LiP1 and LiP5 primers (with closest sequence identity to H8- and H2-like isozymes, respectively) were most prevalent, with 32 taxa found to possess genes similar to P. chrysosporium genes for one or both of the isozymes. Indeed, at least one species from each of the investigated families, with the exception of Scutigeraceae (Cantharellales) and Hygrophoraceae (Agaricales), was found to possess the H8- and H2-type LiP genes, indicating that similar LiP genes exist in phylogenetically diverse ECM taxa (Bruns et al., 1998; Hibbett et al., 2000). Even the two Pisolithus species, widely assumed to have little ability to degrade plant cell wall polymers (Chambers & Cairney, 1999), appear to possess H8- and H2-like genes. No obvious taxonomic pattern was observed in the distribution of the LiP genes, with taxa within a single order, family or even genus differing in the presence or apparent absence of LiP genes. Varela et al. (2000) recently reported a similar situation in white-rot basidiomycetes. By screening white-rot taxa using Southern blotting with a probe for a LiP gene from P. chrysosporium, they found evidence for equivalent genes in some, but not all, members of certain basidiomycete families. Similar inconsistencies were also observed at the genus level for Lentinus and Phellinus species, indicating that different genes may encode for ligninolytic activities, even in closely related taxa. We are mindful, however, that genes for lignin-degrading enzymes may be more widespread in ECM fungal taxa than our current data imply. While ITS amplifications were performed for all fungi to ensure viability of the isolated DNA (data not shown), the isozyme-specific nature of the primers used is likely to have limited amplification to those genes that show a high degree of identity with the P. chrysosporium isozymes. Further screening with degenerate primers may identify related genes in many of the fungi for which no genes were detected using the specific primers. Indeed, a similar approach using primers for other enzymes, such as laccases and other oxidases, that are thought to contribute to lignin degradation, may yield further evidence of ligninolytic enzyme systems in ECM fungi.

A P. chrysosporium H3-type MnP gene has previously been detected in T. fibrillosa (Chambers et al., 1999). In addition to T. fibrillosa, similar MnP genes were detected in Cortinarius rotundisporus, Piloderma byssinum and Piloderma croceum in the present study using the MnP2 primers. Only C. rotundisporus appeared to have a gene equivalent to the H3-like MnP isozyme amplified by the MnP1 primers. With the exception of the putative ECM taxon C. rotundisporus, the three ECM Atheliaceae (Stereales) fungi were the only taxa found to contain MnP genes. Although regarded as ECM (Bougher & Syme, 1998), the mycorrhizal status of C. rotundisporus remains to be confirmed. Given that Cortinarius is generally regarded as an ECM genus (Liu et al., 1997) and that patterns of mycelial growth in forest soils are similar to known ECM taxa (Sawyer et al., 1999), it is, however, probable that this fungus forms ECM with Eucalyptus spp. hosts. In fact, the large mycelial genets formed by this taxon in forest soils grow as dense mats in the litter layers (J. W. G. Cairney, unpublished) and are reminiscent of the mycelial mats formed by certain ECM taxa, within which enhanced peroxidative enzyme activities have been measured (Griffiths & Caldwell, 1992). The identification of MnP genes, along with several LiP genes in each of the three Stereales taxa is also of interest, and suggests that ECM fungi in this order, now known to be an important component of below-ground ECM communities in some coniferous forest ecosystems (Erland & Taylor, 1999), may have considerable potential for producing ligninolytic enzyme activities.

Clearly, to be important in the putative role of facilitating access to mineral nutrients associated with dead plant tissue, the genes we have identified must be expressed. There is no doubt that ECM fungi obtain a considerable supply of carbon from their symbiotic tree hosts (Högberg et al., 1999), and this may be seen as suppressive to expression of saprotrophic genes (Harley & Smith, 1983; Durall et al., 1994). It does not, however, mean that host-derived carbon is uniformly distributed within extramatrical mycelial systems of ECM fungi (Cairney & Burke, 1994, 1996a). In particular, translocation of host-derived carbon into ‘patches’ of organic matter in soil is known to cease following colonization of the organic matter, and is accompanied by a decline in the nitrogen content of the patches (Bending & Read, 1995a). Since carbon and/or nitrogen limitation is known to be important in expression of ligninolytic enzymes in P. chrysosporium, such conditions might favour their expression in regions of soil-borne mycelia of ECM fungi (Cairney & Burke, 1994, 1996a). We hypothesise that, under circumstances such as these, expression of ligninolytic activities in ectomycorrhizal fungi may contribute significantly to the decomposition of lignin in temperate forests, and are currently investigating expression of the genes using RT-PCR. This would provide the ectomycorrhizal fungi with access, not only to mineral nutrients sequestered within plant cell wall structures, but also potentially to a supplementary carbon source. In this way, the strict functional boundaries between ECM and decomposer fungi may be less clear-cut than we have previously envisioned.


This work was supported by an Australian Research Council Large Grant (A00000685) to J. W. G. C.