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Mycoscience (2011) 52:363–375 DOI 10.1007/s10267-011-0117-4

FULL PAPER

Anatomical and ITS rDNA-based phylogenetic identification of two new West African resupinate thelephoroid species Nourou Soulemane Yorou • Atsu Kudzo Guelly Reinhard Agerer

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Received: 20 September 2010 / Accepted: 23 March 2011 / Published online: 10 April 2011 ! The Mycological Society of Japan and Springer 2011

Abstract An anatomical approach coupled with molecular phylogeny of 84 sequences of thelephoroid taxa have been used to describe two new West African resupinate Thelephorales, namely, Tomentella agereri and Tomentella maroana. T. agereri presents a maximal sequence similarity of 94% with its genetically closest species, Tomentalla pilosa, according to a Blastn search in public GenBanks. By molecular phylogenetics, it is nested within the T. pilosa complex, a well-supported (bootstrap support of 100%) monophyletic clade composed of cystidiate and differentiated rhizomorphic species, although it presents contrasting anatomical features including the lack of cystidia, the presence of undifferentiated rhizomorphs, and basidiospores with very short aculei, up to 0.5 lm. Tomentalla maroana is close, by molecular phylogenetic study, to T. ellisii, T. pisoniae, and T. hjortstamiana. The phylogenetic proximity between T. maroana and T. ellisii is supported by morphological characters between the two species, namely, a crustose adherent basidiocarp, a differentiated sterile margin, and a granular hymenium. The two species deviate from each other by 11.38–12.37% with regard to the ITS rDNA sequences, whereas the intraspecific genetic distances vary from 1.68% to 2.9% among the three specimens assigned to T. maroana. Discriminating characters as well

N. S. Yorou (&) ! R. Agerer Department Biology I, Organisms Biology: Mycology, University of Munich, Menzinger Str. 67, 80638 Mu¨nchen, Germany e-mail: [email protected] A. K. Guelly De´partement de Botanique et E´cologie Ve´ge´tale, Faculte´ des Sciences, Universite´ de Lome´, BP1515, 081 Lome´, Togo

as genetic distance between the new species and the closely related species are discussed in detail. Keywords Anatomy ! Genetic distance ! Phylogenetic position ! Taxonomy ! Tropical Africa Introduction Thelephorales encompass a total of about 180 accepted species (Kirk et al. 2008) that are accommodated in 14 genera and two families: the Bankeraceae and the Thelephoraceae (Donk 1964; Ju¨lich 1981; Stalpers 1993). The family Bankeraceae has a predominantly temperate distribution and is frequently reported from Europe and North America (Harrison 1964, 1968; Maas Geesteranus 1975; Baird 1986a, b; Harrison and Grund 1987; Arnolds 1989, 2003; Pegler et al. 1997; Parfitt et al. 2007). However, representatives of the genera Hydnellum P. Karst., Sarcodon Que´l. ex. P. Karst., and Phellodon P. Karst. have been reported from tropical and subtropical Asia (Maas Geesteranus 1971). In contrast, Thelephoraceae have a worldwide distribution (Patouillard 1897; Malenc¸on 1952, 1954; Wakefield 1966; Corner 1968; Hjortstam and Ryvarden 1988, 1995; Stalpers 1993; Ko˜ljalg 1996; Martini and Hentic 2005; Yorou 2008), although they have been frequently reported from Europe, North America, and temperate Asia with the highest species richness in coniferous forests (Larsen 1964, 1968, 1974; Wakefield 1966, 1969; Ko˜ljalg 1996). Taxonomically, members of the family Thelephoraceae in general and the resupinate species in particular display a limited number of anatomical characters, often overlapping, making their delimitation very difficult (Ko˜ljalg 1996). However, the size, shape, and type of ornamentation

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of the basidiospores coupled with the presence or absence of cystidia have been traditionally considered as the most discriminating features (Larsen 1968, 1974; Stalpers 1993; Ko˜ljalg 1996; Da¨mmrich 2006). The taxonomic relevance of rhizomorphs was for a long time underestimated in characterizing them simply as either mono- or dimitic (Ko˜ljalg 1996; Da¨mmrich 2006), although consistent anatomical features at the species level have been confirmed for rhizomorphs and detailed anatomical patterns have proven to play a paramount discriminating role, mostly within closely related species (Yorou and Agerer 2008, 2011a). The tropical African resupinate Thelephorales are particularly difficult to identify, not only because of the scarcity of available, scientifically reliable documentation but mostly because there is a limited number (still with no clear hiatus) of discriminating anatomical characters between specimens of different or contrasting biogeographic origins (Yorou et al. 2007; Yorou and Agerer 2007a). In such cases, the combination of traditional anatomically and morphologically based taxonomy and molecular techniques (barcoding threshold and phylogenetic inferences) were invaluable for an unambiguous discrimination and identification of tropical African thelephoroid taxa and in tracing their affinities with temperate or boreal taxa (Yorou et al. 2007; Tedersoo et al. 2007; Yorou and Agerer 2007a, 2008). The present paper is part of a series exclusively devoted to tropical African resupinate Thelephorales and their ectomycorrhizae. Nine new species were described in previous investigations (Yorou et al. 2007; Yorou and Agerer 2007a, 2008; Yorou et al. 2011a, b), using a combination of anatomo-morphological and molecular data. The present article aims at anatomical and morphological characterization of two additional new tropical African thelephoroid species and addressing their phylogenetic placement with regard to temperate and boreal species.

Materials and methods Specimen sampling Specimens were collected in various vegetation types in the Northern Guinean seasonal forests of West Africa (White 1983). Specimens were dried using a propane gas-heated field dryer (De Kesel 2001). Preliminary notes were recorded using fresh material. Color codes of the dried basidiocarps are given according to Kornerup and Wanscher (1978). All specimens used for descriptions and the holotypes are deposited in M (Holmgren et al. 1990) with the following herbarium labels: SYN878, SYN879, SYN892, and RA13792.

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Light microscopy and SEM investigations We refer to Yorou and Agerer (2007a,b, 2008) for the light microscopy protocol. For scanning electron microscopy (SEM) investigations, small hymenial parts taken from dried basidiocarps were fixed in 300–400 ll glutaraldehyde–cacodylate buffer and afterward treated as follows: 60 min in 2.5% glutaraldehyde, gradual washing (5, 15, 30, 60 min) in a neutral cacodylate buffer (75 mM cacodylate, 2 mM MgCl2, 100 ll H2O, pH 7), 1–2 h incubation in 1% OsO4 buffer (2.5 ml OsO4, 7.5 ml H2O), with subsequent washing in distilled water. The samples were then gradually dehydrated in a series of acetone solutions as follows: 10, 20, 40, 60, 80% (each 15 min), and 2 9 100% (15 and 30 min). Samples were then stored in 100% acetone overnight followed by critical point drying (Anderson 1951). The samples were sputter-coated with gold (using argon gas, under 0.05 mbar) for 3 h and 30 min, until a layer of approximately 15 nm was obtained. SEM was then carried out using a JEOL 5800 LV with a tension of 25 kV and working distance of 10–12 mm. Digital SEM images were captured using Orion V (vers. 5.22) Image Management System. DNA extraction and sequencing DNA was extracted from dried basidiocarps after Gardes and Bruns (1993) using a Quiagen DNeasy plant Mini Kit (Quiagen, Hilden, Germany), according to the manufacturer’s instructions. Polymerase chain reaction (PCR) amplification was performed for internal transcribed spacers ITS1, ITS2, and for the 5.8S region of the nuclear ribosomal DNA, using fungi-specific primer ITS1F and basidiomycete-specific primer ITS4B. PCR amplification was performed using Ready To Go beads (Amersham Pharmacia Biotech, Piscataway, NJ, USA), with 24 ll PCR solution (125.2 lg ddH2O, 20 ll buffer, 6.00 ll MgCl, 20 ll dNTP mix, 10 ll ITS1F, 10 ll ITS4B, 0.8 ll Taq polymerase at 5 U/ll) and 1 ll extracted DNA. The PCR was programmed as follows: 94"C for 3 min, 60"C for 1 min, 72"C for 1 min (1 cycle), and consecutively 94"C for 1 min, 60"C for 1 min, and 72"C for 1 min 30 s (28 cycles), 94"C for 1 min, 60"C for 1 min, and 72"C for 10 min (1 cycle). Amplified PCR products (2 ll) were run with bromophenol blue (2 ll) on 1% agarose gels for 30 min at 95"C, then stained in ethidium bromide for 10 min and afterward in ddH2O for 1 min. PCR products were then visualized under UV light. Successful DNA bands were purified using the QIAquick-PCR purification Kit (Qiagen) according to manufacturer’s instructions. DNA sequencing was performed by the sequencing service of Department of Biology I (Ludwig-Maximilians-Universita¨t, Mu¨nchen, Germany), using BigDye Terminator

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Ready Reaction Cycles Sequencing Kit v 3.1 (Applied Biosystems, Foster City, CA, USA). Sequencing was performed on 1 ll purified DNA probes plus 0.3 ll ITS1F (forward primer) and 0.3 ll ITS4B (reverse primer). Four sequences of both new species were generated and were deposited in GenBank NCBI with accession numbers EF507250, EF507251, EF507252, and EF538424. Sequence edition and estimation of similarities and identities among studied specimens The sequences were edited and the consensus sequence of each investigated specimen was executed in BioEdit v. 7.0.5 (Hall 2005) using the Cap. Contig. Assembl. Program option. The generated sequences were then submitted to a local similarity analysis to check to what extent they deviate from each other and to test if our anatomo-morphological discrimination is supported by molecular data. To do this, generated sequences were automatically aligned using ClustalW Multiple Alignment (BioEdit v. 7.0.5). We activated the ‘‘full multiple alignment,’’ ‘‘bootstrap NJ tree,’’ and ‘‘number of bootstrap = 1,000’’ options. As sequences have unequal lengths, the automatic alignment resulted in many insertion or deletion gaps. For anatomically close specimens, we included gaps in the similarity tests to highlight the maximal deviation rate. For the opposite condition, gaps are excluded in the calculation of similarity between evidently different anatomo-morphological specimens to estimate the minimal sequence deviation. Identity/ similarity was calculated using the ‘‘pairwise alignment, calculation of the similarity/identity’’ option of BioEdit v. 7.0.5. In the second step, we compared the newly generated sequences with ITS sequences deposited in public databanks. The most similar sequences were searched for in UNITE (Ko˜ljalg et al. 2005; Abarenkvov et al. 2010; http:// unite.ut.ee) using the ‘‘Blastn’’ search option. Sequences of the top ten similar taxa identified to species level were downloaded. Unknown or unidentified Tomentella or Thelephora species were disregarded. In addition, most similar sequences were searched for in GenBank NCBI (http://www.ncbi.nlm.gov) using the Megablast (Zhang et al. 2000) search option. We deactivated the ‘‘uncultured/ environmental sample sequences’’ search option to avoid acquiring sequences of taxonomically uninformative/unidentified taxa. Molecular phylogenetic analysis The most similar ITS sequences were downloaded from the public GenBanks (UNITE and NCBI). Additional sequences addressed during recent studies on tropical African resupinate Thelephorales (Yorou and Agerer 2007a, 2008; Yorou et al. 2007) and from the Seychelles (Tedersoo et al.

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2007; Suvi et al. 2010) were added to the dataset. All sequences were automatically aligned in BioEdit v. 7.0.5. The alignment was manually checked and optimized. Ambiguous columns that we could not align with absolute certainty were excluded. The final dataset include a total of 84 ITS rDNA sequences with an alignment length of 500 characters. The most likely tree was searched for using the program RAxML v.7.0.4 (Stamatakis 2006). The best tree was obtained by executing 100 rapid bootstrap inferences and thereafter a thorough search for the most likely tree using one distinct model/data partition with joint branch length optimization (Stamatakis et al. 2008). The generalized time-reversible (GTR) model of substitution was applied having maximum likelihood as the optimal criterion. In addition, the most parsimonious trees were searched for by executing batch files generated with PAUPRat (Sikes and Lewis 2001) in PAUP* v4.0 (Swofford 2002), with weighting mode set to multiplicative. The parsimony analysis is set as follows: heuristic search option, addition sequence random, tree-bisection-reconnection (TBR) swapping, all characters unordered, of equal weight, and gaps are treated as missing data. A consensus tree was calculated of all trees with equal minimal length, and posterior probabilities were recorded for each branch.

Results ITS rDNA-based differences between the new species Specimens SYN878 (accession number EF507250), SYN879 (accession number EF507251), and SYN892 (accession number EF507252) deviate from each other by 1.68–2.9% with regard to their ITS rDNA sequences. All three specimens are accommodated under the new species Tomentella maroana. The sequence (accession number EF538424) obtained from the specimen RA13792 deviates from those of T. maroana by 18.28–18.70%. It is described here as the new species T. agereri. The top five best matches of T. maroana are composed of Tomentella sublilacina (Ellis & Holw.) Wakef. according to Blastn search in UNITE. Sequences similarity between both species ranges from 88% to 91%. In contrast to T. maroana, for which the top five best matches are T. sublilacina, four different species with sequences identities varying from 90% to 94% are most similar to T. agereri according to ‘‘Megablast’’ search in NCBI. Tomentella agereri is 94% similar to T. atroarenicolor Nikol. and T. pilosa (Burt) Bourdot & Galzin, whereas lower identity rates (90% and 92%) are obtained with T. umbrinospora M.J. Larsen and T. ferruginea (Pers.) Pat., respectively. Differing from ‘‘Blastn’’ search in UNITE, the ‘‘Megablast’’ search in NCBI shows more diversity in

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the best matches of both new species. However, both searches generated T. sublilacina as the closest species to T. maroana whereas T. pilosa and T. atroarenicolor are most similar to T. agereri. Phylogenetic position of the two new species A total of 4,341 most parsimonious trees (shortest tree length = 972, CI = 0.568, RI = 0.876) were generated from 10,020 trees. Of 500 reliably aligned characters of the ITS rDNA, 199 were variable and 173 characters were parsimony informative. The final alignment has 234 distinct alignment patterns, and the proportion of gaps and completely underestimated characters in the alignment is estimated at 0.13%. The likelihood of the most likely tree found is -5295.424348 (tree length = 4.510765). The following substitution rates were estimated by RAxML: A $ C, 1.476325; A $ G, 8.882025; A $ T, 1.025856; C $ G, 0.498660; C $ T, 8.280101; G $ T, 1.000000; base frequencies were Freq. (A) = 0.224580, Freq. (C) = 0.247659, Freq. (G) = 0.242133, and Freq. (T) = 0.285627; gamma shape parameter = 0.898260; and proportion of invariable sites = 0.451289. In general, the topology is very similar in all trees generated by RAxMl and PAUP. Therefore, we present only the most likely tree obtained from RAxML (Fig. 1). The major clades of the most likely tree and their constitutive species are similar to those generated during previous molecular phylogenetic investigations on resupinate Thelephorales (Ko˜ljalg et al. 2000, 2001; Yorou and Agerer 2007a; Yorou et al. 2007). In all phylogenetic trees generated, Tomentella agereri forms a sister species to Tomentella pilosa, whose representatives all cluster together with 100% bootstrap support. The terminal clade comprising T. agereri and T. pilosa is well supported by a high bootstrap value of 98%. Both species form, together with T. capitata Yorou & Agerer, Tomentella brunneocystidia Yorou & Agerer, and Tomentella atroarenicolor, a strongly supported monophyletic clade with a bootstrap value of 100%. So far as T. maroana is concerned, all three investigated specimens cluster together with a bootstrap support of 100%. T. maroana forms a sister species of the group comprising Tomentella ellisii (Sacc.) Ju¨lich and Stalpers, Tomentella pisoniae Suvi & Ko˜ljalg, and Tomentella hjortstamiana Suvi & Ko˜ljalg, with, however, no bootstrap support. The sister group of all four species (T. ellisii, T. maroana, T. pisoniae, and T. hjortstamiana) is composed of Tomentella terrestris (Berk. & Broome) M.J. Larsen and Tomentella sublilacina, whose representatives cluster all together with 98% bootstrap support. There is, however, no bootstrap support between the T. sublilacina clade and that composed of T. maroana, T. ellisii, T. pisoniae, and T. hjortstamiana.

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Description of the new species Tomentella agereri Yorou sp. nov. Mycobank MB560122

Figs. 2–4

Basidiocarpi resupinati, tenues, ad 0.5 lm alti, a substrato pro partibus minoribus separabiles, arachnoidei, continui; rhizomorphis margine sterili absentibus. Hymenophorae laeves, continues; hymeniis brunneis usque ad rufo-brunneis; subiculis tenuissimis, arachnoideis, flavis, margine strerili determinato, aurantiaco-flavo, quam hymenio pallidiore. Rhizomorphae in subiculo presentes, sub stereomicroscopio flavae, in aqua et in 2.5% KOH flavae, monomiticae; omnibus hyphis similibus, uniformibus, laxis, in aspectu plano paene paralleliter dispositis, fibuligeris, septis simplicibus deficientibus, 4–7 lm diametro, tenuitunicatis, in aqua et in 2.5% KOH incoloratis usque ad pallide flavis, in aqua incrustatissimis, sed in 2.5% KOH incrustatione dissolventi, non cyanescentibus, subcongophilis, subcyanophilis, non amyloideis. Hyphae subiculi fibuligerae, septis simplicibus deficientius, 3.5–6(–7) lm latae, frequenter regulares, interdum inflatae ad 7–8 lm, in 2.5% KOH infrequenter sinuosae, cruciformibus ramificationibus raris, anastomosibus infrequentibus, in aqua incrustatissimae, in 2.5% KOH incrustatione dissolventi, in aqua et in 2.5% KOH incoloratae usque ad pallide flavae, non cyanescentes, subcongophilae, subcyanophilae, non amyloideae. Hyphae subhymenii fibuligerae, 2.5–4.5(–6) lm diametro, cellulis interdum brevissimis sed numquam inflatis, tenuitunicatae (0.2–0.5 lm), in aqua incrustatissimae, incrustatione in 2.5% KOH dissolventi, hyphae in reagente Melzeri laeves, in aqua et in 2.5% KOH pallide flavidae, non cyanescentes, subcongophilae, subcyanophilae, non amyloideae. Basidia fibuligera, (30–)35–55(–60) lm longa, apice (6–) 6.5–7.5(–8) lm lata, basi (5–)5.5–6.5(–7) lm lata, utriformia, non stipitata, semper sinuosa, infrequenter cum septis transversibus, in aqua et in 2.5% KOH incolorata, non cyanescentia, fere semper congophila sed in Congo red nonnulla basidia incolorata si basidia septata et in partibus distalibus distincte congophila, cyanophila, non amlyoidea, 4-sterigmatica, sterigmatibus 3–7(–7.5) lm longis, basi 1.5–2.5 lm latis. Basidiosporae (6–)6.5–7(–8) 9 (5–) 5.5–6.5(–7) lm in aspectu frontali, (6–)6.5–7.5(–8) 9 (5–) 5.5–6(–6.5) lm in aspectu laterali, in aspectu frontali triangulares, parte proxima dilatata usque ad lobata, ejus latitudine basidiosporae longitudine aequanti, ellipsoideae in aspectu laterali, semper guttulatae, verrucosae usque ad echinulatae, aculeis brevissimis, 0.2–0.5 lm altis, interdum binis ternisve aggregatis et primo aspectu 2- vel 3-furcatis, in aqua incoloratae usque ad pallide subflavae, in 2.5% KOH pallide flavae, non cyanescentes, non congophilae, non cyanophilae, non amyloideae. Chlamydosporae absentes.

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Fig. 1 Maximum likelihood tree showing placement of the two new species among Tomentella species. Bootstrap values higher than 50% are shown above the branches. GenBank (UNITE or NCBI) accession numbers and country of origin of selected species are indicated after species names. Both new species are highlighted in bold. Some clades have been collapsed to reduce the span of the tree

Basidiocarp resupinate, thin, up to 0.5 mm thick, separable from the substrate as small plaques, arachnoid, continuous. Rhizomorphs absent at the sterile margin. Hymenophore smooth, continuous, hymenium brown (7E6) to reddish brown (8E6), subiculum very thin, arachnoid, yellow, sterile margin determinate, paler than the hymenium, orange yellow. Rhizomorphs present in the subiculum, yellow under stereo microscope, yellow in water and

in 2.5% KOH, monomitic; rhizomorphs undifferentiated (Fig. 2), uniform-loose (Agerer 1999), or of type A (according to Agerer 1987–2008), nearly parallel arrangement in plane view, clamped, simple septa absent, 4–7 lm wide, thin walled, colorless to pale yellow in water and in 2.5% KOH, heavily encrusted in water, encrustation dissolving in 2.5% KOH, not cyanescent, slightly congophilous, slightly cyanophilous, not amyloid. Subicular hyphae

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R. Agerer, 22.08.2003, det. N.S. Yorou, herb. RA13792 (M), holotype. Genbank NCBI, accession number EF538424. Etymology The epithet is proposed in honor of Prof. Dr. Agerer, who collected the type material. Habitat

Fig. 2 Tomentella agereri. Optical section through the rhizomorph. Bar 10 lm

clamped, simple septa absent, 3.5–6(–7) lm wide, usually regular, sometimes inflated, then up to 7–8 lm wide, rarely sinuous in 2.5% KOH, cross-shaped branching and anastomoses rare, heavily encrusted in water, encrustation dissolving in 2.5% KOH, colorless to pale yellow in water and in 2.5% KOH, hyphae not cyanescent, slightly congophilous, slightly cyanophilous, not amyloid. Subhymenial hyphae clamped, 2.5–4.5(–6) lm wide, the cells sometimes very short but never inflated (Fig. 3), thin walled (0.2–0.5 lm), heavily encrusted in water, encrustation dissolving in 2.5% KOH, hyphae smooth in Melzer’s reagent, colorless to very pale yellow in water and in 2.5% KOH, not cyanescent, slightly congophilous, slightly cyanophilous, not amyloid. Cystidia absent. Basidia clamped at base, (30–)35–55(–60) lm long, (6–)6.5–7.5(–8) lm at apex, (5–)5.5–6.5(–7) lm at base, utriform, not stalked, always sinuous, rarely with transverse septa, colorless in water and in 2.5% KOH, not cyanescent, usually congophilous, some basidia remaining colorless in Congo red, if septate then the upper part distinctly congophilous, cyanophilous, not amyloid, 4-sterigmate, sterigmata 3–7(–7.5) lm long and 1.5–2.5 lm at base. Basidiospores (6–)6.5–7(–8) 9 (5–) 5.5–6.5(–7) lm in frontal face, (6–)6.5–7.5(–8) 9 (5–) 5.5–6(–6.5) lm in lateral face, in frontal view triangular to lobed, with a widened proximal part, proximal part as large as the length of the basidiospores, ellipsoid in lateral view (Fig. 4), always with oil drops, warted to echinulate, aculei very short, 0.2–0.5 lm, sometimes grouped in two or three, giving an impression of bi- or trifurcation, basidiospores colorless to very pale yellow in water, pale yellow in 2.5% KOH, not cyanescent, not congophilous, not cyanophilous, not amyloid. Chlamydospores absent. Material studied Benin, central part, Borgou Province, Sinende´ region, forest close to Foˆ-Bouko village, 10"80 46.600 N, 002"150 6.200 E, leg.

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On the underside of dead bark, in woodlands and forests dominated by Euphorbiaceae (Uapaca togoensis Pax), Caesalpiniaceae (Isoberlinia doka Craib & Stapf, Isoberlinia tomentosa Craib & Stapf, Burkea africana Hook. F., Afzelia africana Smith). Tomentella maroana Yorou sp. nov. Mycobank MB560123

Figs. 5–7

Basidiocarpi resupinati, tenues, ad 0.5 mm alti, adherentes, crustosi, continues, fusco-brunnei usque ad obscure rubrobrunnei. Hymenophorae laeves usque ad granulosae, continuae; hymeniis pallidiore, brunneis; subiculis arachnoideis obscuro-badio usque ad fere nigro, margine sterili determinato, cinereo-brunneo, pallidiore quam hymenio. Rhizomorphae in subiculo presentes, sub stereomicroscopio brunneae, tenues (usque ad 20 lm diametro) in aqua et in 2.5% KOH brunneae, crassiores (quam 20 lm) in aqua et in 2.5% KOH usque ad obscuro-brunneae, monomiticae, omnibus hyphis similibus, compactis, uniformibus, fibuligeris, plano aspectu compacte fere paralleliter dispositis, septis simplicibus presentibus, 4–6(–7) lm diametro, semper crassitunicatis (1.5–2.5 lm), in aqua et in 2.5% KOH non incrustatis, in aqua et in 2.5% KOH brunneis usque ad obscuro-brunneis, non cyanescentibus, non congophilis, non cyanophilis, non amyloideis. Hyphae subiculi fibuligerae, septis simplicibus infrequentibus, 4–6(–7) lm diametro, crassitunicatae (1.5–2.5 lm crassae), muribus flavis, regulares, in aqua et in 2.5% KOH frequenter tortuosae; hyphae marginis sterilis regulares, 4–5 lm diametro; hyphae subiculi infrequenter inflatae et infrequenter tortuosae, anastomosibus cruciformibus, non incrustatae, in aqua et in 2.5% KOH brunneae usque ad obscuro-brunneae; hyphae marginis sterilis in aqua et in 2.5% KOH brunneae, non cyanescentes, non congophilae, non cyanophilae, non amyloideae. Hyphae subhymenii fibuligerae, 4–7(–8) lm diametro, cellulis non brevibus, interdum inflatis (usque ad 10 lm), crassitunicatae (1–2 lm), muribus flavis, in aqua et in 2.5% KOH non incrustatae, interdum cyanescentes, non congophilae, non cyanophilae, non amyloideae. Cystidia absentia. Basidia fibuligera, (40–) 45–65(–70) lm longa, apice (8–)9.5–11(–12) lm lata, basi (5.5–)6–7.5(–8) lm lata, clavata, non stipitata, interdum sinuosa, infrequenter cum septis transversibus; basidiis et

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Fig. 3 Tomentella agereri. a Basidiospores in lateral view. b Basidiospores in frontal view. c Section through the basidiocarp. Bars 10 lm

basidiolis marginis basidiocarpi interdum distincte crassitunicatis (1–1.5 lm); basidiis in aqua et in 2.5% KOH incoloratis usque as pallide brunneis, interdum cyanescentiis, congophilis, subcyanophilis; basidiis juvenilibus distincte cyanophilis, non amyloideis, 4-sterigmatica, sterigmaticis 5–9(10) lm longis, basi 1.5–3 lm latis. Basidiosporae (8–) 8.5–11(–12) 9 (7–)7.5–8.5(–9) lm in aspectu frontali, (8–) 8.5–11(–11.5) 9 (7–)7.5–8(–8.5) lm in aspectu laterali, triangulares et parte proxima dilatata usque ad sublobata in

aspectu frontali, ellipsoideae in aspectu laterali, non guttulatae, verrucosae usque ad echinulatae, aculeis brevissimis, 0.2–0.5 lm altis, interdum binis ternisve aggregatis, in aqua et in 2.5% KOH pallide brunneae, interdum cyanescentes, non congophilae, non cyanophilae, non amyloideae. Chlamydosporae absentes. Basidiocarp resupinate, thin, up to 0.5 mm thick, adherent to the substrate, crustose, continuous, dark brown (7F7) to reddish dark brown (8F7). Hymenophore smooth

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Fig. 6 Tomentella maroana. Anastomoses of subicular hyphae. Bar 10 lm

Fig. 4 Tomentella agereri. Scanning electron microscopy (SEM) of basidiospores. a Basidiospore in nearly proximal view. b Basidiospore in lateral view. c General view of basidiospores. Bars a, b 1 lm; c 5 lm

Fig. 5 Tomentella maroana. Optical section through rhizomorph. Bar 10 lm

to granular, continuous, hymenium paler, brown (6F4), subiculum arachnoid, dark brown to almost black, sterile margin determinate, paler than the hymenium, greyish brown. Rhizomorphs present in the subiculum, brown under stereo microscope, brown (young rhizomorphs up to 20 lm) to dark brown (old rhizomorphs thicker than 20 lm) in water and in 2.5% KOH, monomitic, undifferentiated (Fig. 5), uniform compact (Agerer 1999), or of type B (according to Agerer 1987–2008), with a parallel compact arrangement in plane view, individual hyphae

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clamped, simple septa present, 4–6(–7) lm wide, always thick walled (1.5–2.5 lm), walls yellowish, without encrustation in water and in 2.5% KOH, brown to dark brown in water and in 2.5% KOH, not cyanescent, not congophilous, not cyanophilous, not amyloid. Subicular hyphae clamped, simple septa rare, 4–6(–7) lm wide, always thick walled (1.5–2.5 lm), walls yellowish, hyphae usually regular in outlines, commonly tortuous in 2.5% KOH, hyphae from the sterile margin regular, 4–5 lm wide, rarely inflated and rarely tortuous, cross-shaped branching and anastomoses present (Fig. 6), without encrustation, brown to dark brown in water and in 2.5% KOH, hyphae from the sterile margin brown in water and in 2.5% KOH, not cyanescent, not congophilous, not cyanophilous, not amyloid. Subhymenial hyphae clamped, 4–7(–8) lm wide, never short, sometimes inflated (up to 10 lm) (Fig. 7), thick walled (1–2 lm), walls yellowish, without encrustation in water and in 2.5% KOH, brown to dark brown in water and in 2.5% KOH, sometimes cyanescent, not congophilous, not cyanophilous, not amyloid. Cystidia absent. Basidia clamped at base, (40–)45–65(–70) lm long, (8–)9.5–11(–12) lm at apex, (5.5–)6–7.5(–8) lm at base, clavate, not stalked, sometimes sinuous, rarely with transverse septa, some basidia and basidioles from the sterile margin distinctly thick walled (1–1.5 lm), basidia colorless to pale brown in water and in 2.5% KOH, partly cyanescent, congophilous, slightly cyanophilous, young basidia distinctly cyanophilous, not amyloid, 4-sterigmate, sterigmata 5–9(–10) lm long and 1.5–3 lm at base. Basidiospores (8–)8.5–11(–12) 9 (7–)7.5–8.5(–9) lm in frontal face, (8–)8.5–11(–11.5) 9 (7–)7.5–8(–8.5) lm in lateral face, triangular with a widened proximal part to slightly lobed in frontal view, ellipsoid in lateral view, oil drops absent, warted to echinulate, aculei very short,

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Fig. 7 Tomentella maroana. a Basidiospores in lateral view. b Basidiospores in frontal view. c Section through basidiocarp. Bars 10 lm

0.2–0.5 lm, sometimes in groups of two or three, basidiospores pale brown in water and in 2.5% KOH, partly cyanescent, not congophilous, not cyanophilous, not amyloid. Chlamydospores absent. Materials studied Benin, central part, Borgou Province, Wari-Maro region, forest reserve of Wari-Maro, 08"200 21.700 N, 002"490 32.600 E, leg and det. N.S. Yorou, 06.08.2005, herb.

SYN878 (M), holotype, GenBank NCBI, accession number EF507250.—Benin, central part, Borgou Province, WariMaro region, forest reserve of Wari-Maro, 08"200 22.100 N, 002"490 32.900 E, leg and det. N.S. Yorou, 06.08.2005, herb. SYN879 (M), GenBank NCBI, accession number EF507 251; SYN892 (M), GenBank NCBI, accession number EF507252.—Togo, central western part, Sotouboua Prefecture, Fazao-Malfakassa National Parc, 08"420 58.600 N, 000"460 22.200 E, leg. and det. N.S. Yorou, 05.06.2008, herb. SYN1677 (M).

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Etymology The epithet is given in reference to Wari-Maro, the collecting site of the type material. Habitat The forest reserves of Wari-Maro and Fazao-Malfakassa are dominated by Caesalpiniaceae (Isoberlinia spp., Afzelia africana…) and Euphorbiaceae (Uapaca spp.). Specimens of T. maroana were collected on dead logs and barks under A. africana individuals.

Discussion Tomentella agereri is characterized by the complete lack of cystidia, the presence of uniform loose rhizomorphs, and small basidiospores not longer than 7(–8) lm with very short aculei (0.2–0.5 lm). By molecular phylogenetics, it falls in the clade formed by Tomentella pilosa and its allied species. The monophyly of the clade formed by T. pilosa, T. capitata, T. brunneocystidia, and T. atroarenicolor, in which T. agereri is placed, has been constantly confirmed in various molecular phylogenetic studies on resupinate Thelephorales and their ectomycorrhizae with strong bootstrap support of 84% (Ko˜ljalg et al. 2001), 83% (Yorou and Agerer 2008), 98% (Yorou et al. 2007), 90% (Yorou and Agerer 2007a), and 99% (Jakucs et al. 2005). This placement of T. agereri is in accordance with a Blastn search undertaken in UNITE and NCBI. Anatomically, this clade is composed, except for T. agereri, of cystidiate species (Stalpers 1993; Ko˜ljalg 1996; Da¨mmrich 2006; Melo et al. 2006; Yorou et al. 2007), which present either capitate cystidia (T. pilosa, T. capitata, T. brunneocystidia) or hyphoid cystidia (T. atroarenicolor). In this clade, cystidia are present on rhizomorphs of T. pilosa, T. capitata, and T. atroarenicolor (Melo et al. 1998, 2006; Jakucs and Agerer 1999; Yorou et al. 2007). Further investigation confirmed the presence of capitate cystidia on the ectomycorrhizal mantle of T. pilosa also (Jakucs and Agerer 1999). Anatomical description of the ectomycorrhizae of other species of this clade (namely, T. capitata and T. brunneocystidia) is not available for comparison, but the shape of the cystidia present in the hymenium is one of the anatomical features that support the clustering of T. pilosa, T. capitata, and T. brunneocystidia in a well-supported clade (bootstrap support of 93%). The separation of T. atroarenicolor as a sister species (with a strong bootstrap support of 100%) of these three capitate cystidiate species is justified by the hyphoid shape of the cystidia in the latter species. All members of this well-supported clade (except for T. agereri with uniform rhizomorphs) are

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characterized by slightly differentiated rhizomorphs, composed of central, wider hyphae that are covered by smaller, entwined and tortuous, multi-branched peripheral hyphae forming a kind of dense rind around the rhizomorphs (Jakucs and Agerer 1999; Agerer 2006; Yorou et al. 2007). The clustering of T. agereri in the anatomically and molecular phylogenetically well-defined clade (T. pilosa complex) is astonishing because of the lack of cystidia, the uniform loose rhizomorphs, and tiny aculei, but unambiguous evidence is provided that it is a new species within this clade. Although this clade is robust and supported by a bootstrap value of 100%, strong anatomical similarities that could explain the placement of T. agereri within cystidiate species are currently lacking, except the common triangular to slightly lobed shape and the yellowish color of the basidiospores. Tomentella agereri could have lost the capacity to form both cystidia and differentiated rhizomorphs. Tomentella maroana is characterized by a brown to dark brown crustose basidiocarp. Important anatomical features are the distinctly thick-walled subicular and subhymenial hyphae and the triangular to slightly lobed (in lateral view) pale brown basidiospores with short aculei of 0.2–0.5 lm. Although the most similar species is T. sublilacina according to Blastn search in both UNITE and NCBI, it is molecular phylogenetically close to T. ellisii, T. pisoniae, and T. hjortstamiana. The molecular phylogenetic proximity between T. ellisii and T. sublilacina has been repeatedly reported in many molecular phylogenetic studies (Ko˜ljalg et al. 2000, 2001; Yorou and Agerer 2007a; Yorou et al. 2007). From all aforementioned molecular studies, the delimitation between T. ellisii and T. sublilacina is consistent; T. sublilacina is more closely related to T. terrestris than to T. ellisii, which appears to be closely related to T. maroana, T. pisoniae, and T. hjortstamiana in this study. In this group, some species have been reported to present undifferentiated rhizomorphs (Agerer 1987– 2008, 1999), namely, T. sublilacina (Agerer 1996 under T. albomarginata (Bourdot & Galzin) M.P Christ. and T. radiosa (Agerer and Bougher 2001; Yorou and Agerer 2007b). Morphologically, T. pisoniae deviates from T. maroana, T. ellisii, and T. hjortstamiana by its arachnoid separable basidiocarp and the dark grey hymenophore (Suvi et al. 2010) with an indeterminate sterile margin. All three species—T. maroana, T. ellisii, and T. hjortstamiana—present crustose adherent basidiocarps with a differentiated sterile margin, but T. hjortstamiana deviates morphologically from both T. ellisii and T. maroana by the smooth grey yellowish hymenium (Suvi et al. 2010). Morphologically, T. ellisii is the closest species to T. maroana. Anatomical dissimilarities could be highlighted between both species. Tomentella ellisii differs by having thin-walled, colorless to only very pale brown, subicular hyphae (Ko˜ljalg 1996; Da¨mmrich 2006; Yorou and Agerer

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2011b), whereas T. maroana presents constantly brown to dark brown thick-walled (1.5–2.5 lm) hyphae. Short and inflated, thin-walled subhymenial hyphae up to 12 lm are common in T. ellisii (Da¨mmrich 2006; Yorou and Agerer 2011b), whereas those of T. maroana are never short or inflated but always thick walled (1.5–2.5 lm). Also, reniform (in lateral view) basidiospores are reported for T. ellisii (Da¨mmrich 2006; Yorou and Agerer 2011b), but we could not observe such basidiospores in all three specimens of T. maroana examined. In addition, basidiospores of T. ellisii present aculei of up to 1 lm, whereas those of T. maroana never exceed 0.5 lm. In contrast to T. ellisii, in which rhizomorphs are absent (Yorou and Agerer 2011a) or infrequent (Ko˜ljalg 1996; Da¨mmrich 2006), T. maroana is characterized by the presence of undifferentiated (uniform compact) rhizomorphs with thick-walled brown to dark brown individual hyphae. The molecular phylogenetic placement of T. maroana is thus supported by rhizomorph anatomy because some species in this clade present undifferentiated rhizomorphs (see above). The barcoding threshold has become a tool for a rapid screening of morphologically close specimens and the detection of cryptic species (Tedersoo et al. 2008, 2009). DNA barcoding has been widely used in ectomycorrhizal community studies to obtain the most similar species of a given environmental sample (Tedersoo et al. 2003; Izzo et al. 2005; O’Brien et al. 2005; Parrent et al. 2006). However, the ITS rDNA-based identification of thelephoroid species in public GenBanks (UNITE, NCBI, or EMBL) commonly results in either a large number of insufficiently identified environmental samples as best matches, or in a large number of structurally different species with, however, equal similarity percentage to the query (Nilsson et al. 2006, 2009; personal observation). On one hand, it may be a consequence of unequal sequence length and query coverage, which results in an overestimation of sequence similarity (see explanation in Tedersoo 2007; Nilsson et al. 2009) but also insufficient taxonomic annotation of specimens in the public GenBank (Nilsson et al. 2006). In such cases, the barcoding threshold loses its value, and only anatomical comparison can ensure a reliable identification to species level of the query. In resupinate Thelephorales, specimens (either fruit bodies or environmental samples) accommodated in one and the same species and clustering together in a well-supported terminal clade actually present sequence deviations that are generally less than 5% (Ko˜ljalg et al. 2000; Yorou et al. 2007; Yorou and Agerer 2008). Sequence deviations higher than 6–7% suggest the presence of at least two different species in the clade (Ko˜ljalg et al. 2000). Presently, however, a 3–4% threshold is discussed or accepted for discerning species (Izzo et al. 2005) to simplify interpretations of environmental samples, a procedure still waiting for justification (Nilsson et al. 2008). In the present

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study, all three specimens assigned to T. maroana deviate only by 1.68–2.9%; they cluster together in a terminal clade supported by 100% bootstrap. All three specimens of T. maroana deviate from those of T. ellisii, T. pisoniae, and T. hjortstamiana by at least 11.38–12.37%, 13.29–13.69%, and 14.03–15.52%, respectively. By molecular phylogenetics, T. agereri deviates from the closest species, T. pilosa, by at least 10.4–11.3%. Acknowledgments We are much indebted to the Deutsche Forschungsgemeinschaft (DFG) for financial support (project Ag7/19-1). Frank Van Caekenberghe (National Botanic Garden of Belgium, Meise) is thanked for his skilful assistance with the SEM micrographs.

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Mycoscience (2011) 52:376–382 DOI 10.1007/s10267-011-0120-9

FULL PAPER

Epitypification of Colletotrichum musae, the causative agent of banana anthracnose Yuan-Ying Su • Parinn Noireung • Fang Liu • Kevin D. Hyde • Mohamed A. Moslem • Ali H. Bahkali Kamel A. Abd-Elsalam • Lei Cai

•

Received: 19 December 2010 / Accepted: 7 April 2011 / Published online: 11 May 2011 ! The Mycological Society of Japan and Springer 2011

Abstract Colletotrichum musae is an important pathogen causing banana anthracnose. The type material (K) had no conidia or sclerotia, and DNA could not be extracted from a darkened area of the herbarium sample. This sample thus provides few characters to delimit this species from other closely related taxa in the ‘‘gloeosporioides’’ species complex. An epitype is therefore designated for C. musae to stabilize the application of the species name. A detailed morphological description is provided from the epitype. Multilocus phylogenetic analysis indicates that C. musae clusters in a distinct lineage in the ‘‘gloeosporioides’’ species complex and is most closely related to Colletotrichum fructicola. Keywords Disease ! Morphology ! Phylogeny ! Plant pathogen ! Taxonomy Introduction Anthracnose of banana, caused by Colletotrichum musae (Berk. & M.A. Curtis) Arx, is one of the most important and widely distributed diseases of ripe banana fruit Y.-Y. Su ! F. Liu ! L. Cai (&) Key Laboratory of Systematic Mycology and Lichenology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, People’s Republic of China e-mail: [email protected] P. Noireung ! K. D. Hyde School of Science, Mae Fah Luang University, Thasud, Chiang Rai 57100, Thailand M. A. Moslem ! A. H. Bahkali ! K. A. Abd-Elsalam Botany and Microbiology Department, College of Science, King Saud University, Riyadh 1145, Saudi Arabia

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(Meredith 1960a; Stover and Simmonds 1987). Colletotrichum musae may form lesions on fruits without skin bruising but produces larger lesions when fruits are damaged (Meredith 1960a). This species is also an important pathogen on wounded green banana fruits (Meredith 1960b; Stover and Simmonds 1987), infecting banana fruits at any time during the growing season in the field (Simmonds and Mitchell 1940). Colletotrichum musae is also responsible for crown rot, blossom end rot, and tip rot of banana (Nazriya et al. 2007). This taxon has been found on fruits, leaves, and roots of Musa spp. (Meredith 1960a; Simmonds 1965; Israeli and Temkin-Gorodeiski 1977; Pereira et al. 1999; Photita et al. 2001; Anthony et al. 2004; Chillet et al. 2006; Nazriya et al. 2007; Nuangmek et al. 2008). Colletotrichum musae appears to be host specific to Musa species; on the other hand, Mahadtanapuk et al. (2007) reported this taxon as an anthracnose pathogen on flowers of Curcuma alismatifolia Gagnep. in Thailand. Identification of this fungus on C. alismatifolia was based on morphology and should be confirmed by molecular analysis. Colletotrichum musae was described by Berkeley (1874) as Myxosporium musae Berk. & M.A. Curtis with a brief protologue and transferred to Colletotrichum by von Arx (1957). Sutton (1980, 1992) accepted it as a distinct species and provided a brief morphological description. Later, Hyde et al. (2009) pointed out that epitypification is needed to clarify its relationships with other closely related taxa. Traditionally, identification of C. musae was based on morphological characters, e.g., the abundant sporulation, straight and cylindrical conidia, and irregularly shaped appressoria (Sutton 1980). These morphological characters, however, are often overlapping and ambiguous among the ‘‘gloeosporioides’’ species complex. Thus, molecular

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analyses are needed for a precise diagnosis. Although molecular data can provide useful information for species delimitation, studies are generally flawed by the fact the type specimens have not been sequenced (Hyde et al. 2009). Then, we examined type material of C. musae borrowed from K for morphological characters. Unfortunately, no conidia or sclerotia could be found in this specimen, which consists of a 136-year-old dried banana pericarp. It was also not possible to extract fungal DNA from darkened areas on the banana pericarp of this type material. Therefore, the type of C. musae provided few characters to demarcate this species from other closely related taxa in the ‘‘gloeosporioides’’ species complex. An epitype of C. musae is therefore designated to stabilize the application of the species name. The objective of this study was to (1) designate a suitable specimen with living culture as epitype; (2) morphologically characterize the designated epitype; and (3) infer phylogenetic relationships of C. musae with closely related taxa.

Materials and methods Isolation, morphological examination, and reinoculation experiment A culture of C. musae was obtained from CBS (CBS 116870). This culture was isolated from banana fruit in North America, the original geographic locality of this taxon. Three additional strains of C. musae were isolated from banana fruits from Bandu, Chiang Rai Province in northern Thailand to provide a comparison with those from North America. Infected fruits were incubated in moist chambers at room temperature to induce sporulation. Strains were isolated by single-spore isolation (Choi et al. 1999). Pure cultures were stored on potato dextrose agar (PDA) slants and deposited in the culture collection of Mae Fah Luang University Culture Collection (MFLUCC), Chiang Rai, Thailand and BIOTEC Culture Collection (BCC), Pathumthani, Thailand. Mycelial discs (5 mm diameter) were taken from active sporulating areas near the growing edge of 5-day-old cultures, transferred to PDA, and incubated at 25"C, following the methods of Cai et al. (2009). Colony diameter of three replicate cultures growing on PDA was measured daily for 7 days. Growth rate was calculated as the 7-day average of mean daily growth (mm/day). Appressoria were produced using a slide culture technique in which 10 mm2 squares of PDA were placed in an empty Petri dish. The edge of the agar was inoculated with spores taken from a sporulating culture, and a sterile cover slip was placed over the inoculated agar (Cai et al. 2009). After 3–7 days, the shape and

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size of the appressoria formed across the underside of cover slips were studied. To confirm the pathogenicity of the ex-epitype strain, spore suspensions were inoculated back to banana, apple, pear, jujube, and tomato (three replicates). Sterilized distilled water was used as a control. Inoculated fruits were kept in plastic boxes to maintain humidity. Symptoms were examined after 7 days incubation at room temperature. Detailed protocols follow that of Cai et al. (2009). DNA extraction, polymerase chain reaction, and sequencing Isolates were grown on PDA and incubated at 27"C for 7 days. Genomic DNA was extracted by using a Biospin Fungus Genomic DNA Extraction Kit (BioFlux) according to the manufacturer’s protocol. Quality and quantity of DNA were estimated visually by staining with GelRed on 1% agarose gel electrophoresis. Partial actin (ACT), calmodulin (CAL), b-tubulin (TUB2), glutamine synthetase (GS), glyceraldehyde-3phosphate dehydrogenase (GPDH) genes, and the complete rDNA-internal transcribed spacer (ITS) region from the strains were amplified by polymerase chain reaction (PCR). Primer pairs and PCR amplification conditions were followed as previously described (Prihastuti et al. 2009; Crouch et al. 2009a). DNA sequencing was performed at the SinoGenoMax Company, Beijing. Sequence alignment and phylogenetic analyses Sequences from forward and backward primers were aligned to obtain a consensus sequence by using BioEdit (Hall 1999). Sequences of the ex-epitype isolate, along with reference sequences obtained from GenBank (Table 1), were aligned by Clustal X (Thompson et al. 1997). Alignments were optimized manually in BioEdit (Hall 1999). To compare C. musae with other Colletotrichum species, a combined ACT, CAL, GPDH, TUB2, GS, and ITS sequences dataset was used for phylogenetic reconstruction. Phylogenetic analyses were performed by using PAUP* 4.0b10 (Swofford 2002). Ambiguously aligned regions were excluded from all analyses. Unweighted parsimony (UP) analysis was performed. Trees were inferred using the heuristic search option with TBR branch swapping and 1,000 random sequence additions. Maxtrees were unlimited, branches of zero length were collapsed, and all multiple parsimonious trees were saved. Descriptive tree statistics such as tree length (TL), consistency index (CI), retention index (RI), rescaled consistency index (RC), homoplasy index (HI), and log likelihood [-ln L] (HKY model) were calculated for trees generated under different optimality criteria. Clade stability was assessed in a

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Table 1 Sources of isolates and GenBank accession numbers used in this study for genus Colletotrichum Culture collection no.a

GenBank accession numberb

C. asianum

MFLUCC 090232

FJ 903188

FJ 907434

FJ 917501

FJ 972586

FJ 972571

FJ 972605

Prihastuti et al. 2009

C. asianum

MFLUCC 090233b FJ 907424

FJ 907439

FJ 917506

FJ 972595

FJ 972576

FJ 972612

Prihastuti et al. 2009

C. asianum

MFLUCC 090234

FJ 907436

FJ 917503

FJ 972598

FJ 972573

FJ 972615

Prihastuti et al. 2009

Species name

ACT

b

FJ 907421

TUB2

Reference

CAL

GS

GPDH

ITS

C. cordylinicola

BCC 38872

HM470234 HM470249 HM470237 HM470243 HM470240 HM470246 Phoulivong et al. 2010

C. cordylinicola

BCC38864

HM470233 HM470248 HM470236 HM470242 HM470239 HM470245 Phoulivong et al. 2010

C. fructicola

MFLUCC 090227

FJ 907425

FJ 907440

FJ 917507

FJ 972594

FJ 972577

FJ 972611

Prihastuti et al. 2009

C. fructicola C. fructicola

MFLUCC 090228b FJ 907426 MFLUCC 090226 FJ 907427

FJ 907441 FJ 907442

FJ 917508 FJ 917509

FJ 972593 FJ 972592

FJ 972578 FJ 972579

FJ 972603 FJ 972602

Prihastuti et al. 2009 Prihastuti et al. 2009

C. gloeosporioides CORCG4

HM034800 HM034810 HM034802 –

HM034806 HM034808 Phoulivong et al. 2010

C. gloeosporioides CORCG5

HM034801 HM034811 HM034803 –

HM034807 HM034809 Phoulivong et al. 2010

C. gloeosporioides CBS 953.97b

FJ 907430

FJ 972582

C. horii

TSG001

GU133374 GU133375 GU133376 GU133377 GQ329682 AY787483 Wikee et al. 2011

C. horii

TSG002

FJ 907445

FJ 917512

FJ 972589

FJ 972609

Phoulivong et al. 2010

GU133379 GU133380 GU133381 GU133382 GQ329680 AY791890 Wikee et al. 2011

C. jasmini-sambac MFLUCC10-0277

b

HM131507 HM153768 HM131492 HM131502 HM131497 HM131511 Wikee et al. 2011

C. jasmini-sambac HLTX-01

–

C. jasmini-sambac CLTA-01

HM131510 HM153772 HM131496 HM131506 HM131501 HM131515 Wikee et al. 2011

HM153769 HM131493 HM131503 HM131498 HM131512 Wikee et al. 2011

C. kahawae

IMI 319418b

GU133374 GU133375 GU133376 GU133377 GQ329682 AY787483 Prihastuti et al. 2009

C. kahawae

IMI 363578b

GU133379 GU133380 GU133381 GU133382 GQ329680 AY791890 Prihastuti et al. 2009

C. musae

CBS116870b

HQ596284 HQ596280 –

C. musae

MFLUCC 10-0976 HQ596285 HQ596281 HQ596296 HQ596289 HQ596300 HQ596293 This study

C. musae

MFLUCC 10-0977 HQ596286 HQ596282 HQ596297 HQ596290 HQ596301 HQ596294 This study

C. musae C. siamense

MFLUCC 10-0978 HQ596287 HQ596283 HQ596298 HQ596291 HQ596302 HQ596295 This study MFLUCC 090231 FJ 907422 FJ 907437 FJ 917504 FJ 972597 FJ 972574 FJ 972614 Prihastuti et al. 2009

C. siamense

MFLUCC 090230b FJ 907423

C. simmondsii

BRIP 28519

C. simmondsii

CBS 294.67

b

HQ596288 HQ596299 HQ596292 This study

FJ 907438

FJ 917505

FJ 972596

FJ 972575

FJ 972613

Prihastuti et al. 2009

FJ 907428

FJ 907443

FJ 917510

FJ 972591

FJ 972580

FJ 972601

Wikee et al. 2011

FJ 907429

FJ 907444

FJ 917511

FJ 972590

FJ 972581

FJ 972610

Wikee et al. 2011

ACT actin, TUB-2 b-tubulin (tub2), CAL calmoudulin, GS glutamine synthetase, GDPH glyceraldehyde-3-phosphate dehydrogenase, ITS rDNAinternal transcribed spacer (ITS) region a

BRIP: Plant Pathology Herbarium, Department of Primary Industries, Queensland, Australia; CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; IMI: CABI Europe–UK, Bakeham Lane, Egham, Surrey TW209TY, UK; MFLU: Mae Fah Luang University, Thailand

b

The ex-type cultures: the newly generated sequences in this study are shown in bold

bootstrap analysis with 1,000 replicates, each with 10 replicates of random stepwise addition of taxa. Kishino– Hasegawa tests (Kishino and Hasegawa 1989) were performed to determine whether trees were significantly different. Trees were figured in Treeview (Page 1996). Model of evolution (HKY?G) was estimated by using MrModeltest 2.2 (Nylander 2004). Posterior probabilities (PP) (Rannala and Yang 1996; Zhaxybayeva and Gogarten 2002) were determined by Markov Chain Monte Carlo sampling (BMCMC) in MrBayes 3.0b4 (Huelsenbeck and Ronquist 2001), using the estimated model of evolution. Six simultaneous Markov chains were run for 1,000,000 generations, and trees were sampled every 100th generation (resulting in 10,001 total trees). The first 2,001 trees,

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which represented the burn-in phase of the analyses, were discarded, and the remaining 8,000 trees were used for calculating posterior probabilities (PP) in the majority rule consensus tree.

Results Sequences of the ex-epitype strain of C. musae (CBS 116870) and three strains from Thailand were obtained and deposited in GenBank (see Table 1), except CAL from the ex-epitype strain, which failed in several attempts of amplification. Phylogenetic relationships were inferred using combined ACT, CAL, TUB2, GS, GPDH, and ITS.

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379 C. kahawae IMI 363578*

100

C. kahawae IMI 319418*

100

C. cordylinicola BCC 38864

100

C. cordylinicola BCC 38872∗ C. gloeosporioides CORCG4

100

C. gloeosporioides CORCG5

62 91

C. gloeosporioides CBS 953.97*

98 C. siamense MFLU 090231

C. siamense MFLU 090230*

100 99

92 69

64

C. jasmini-sambac CLTA-01 b MFLUCC 10 0277* C i i C. jjasmini-sambac 10-0277* C. jasmini-sambac HLTX-01 C. asianum MFLU 090233*

99 100

C. asianum MFLU 090234 81

72

98 51

C. asianum MFLU 090232

C. fructicola MFLU 090227 C. fructicola MFLU 090226 C. fructicola MFLU 090228*

51

C. musae CBS116870 100

EPITYPE

C. musae MFLU10-0976 C. musae MFLU10-0978 C. musae MFLU10-0977

100

C. horii TSG001 C. C hhorii ii TSG002 C. simmondsii BRIP 28519 C. simmondsii CBS 294.67

10

Fig. 1 Phylogram of tree generated from maximum parsimony analysis based on combined actin (ACT), glyceraldehyde-3-phosphate dehydrogenase (GPDH), internal transcribed spacer (ITS), b-tubulin (tub2) (TUB2), glutamine synthetase (GS), and calmoudulin (CAL) sequences, showing the phylogenetic relationships of Colletotrichum

musae. Values above the branches are parsimony bootstrap ([50%); thickened branches represent significant Bayesian posterior probability (C95%). The tree is rooted with Colletotrichum simmondsii. Asterisks indicate the ex-type strains; arrow, epitype

Maximum parsimony analysis generated only one tree (TL = 1,776, CI = 0.882, RI = 0.900, RC = 0.794, HI = 0.118) (Fig. 1). Multilocus sequences analysis shows that C. musae appears as a distinct lineage among the ‘‘gloeosporioides’’ species complex of Colletotrichum. A close phylogenetic relationship between C. musae and C. fructicola is supported by this analysis. Cultural characteristics and conidial and appressorial morphology are illustrated in Fig. 2. Three banana fruits inoculated with spore suspensions from the ex-epitype all developed typical anthracnose disease (dark brown necrotic lesions). Apples, pears, jujubes, and tomatoes did not develop symptoms after 7 days of inoculation.

n = 6); circular, with sparse to abundant, white to grey floccose aerial mycelium, conidial masses well developed, salmon orange; reverse grey-yellowish. Sclerotia absent. Setae absent. Conidiophores cylindrical, tapered toward the apex, hyaline, subhyaline toward the base, up to 31 lm long, 3–5 lm wide. Conidia 11.5–19.5 9 4–5 lm (x = 14.7 ± 2.13 9 4.6 ± 0.41, n = 30), abundant, hyaline, aseptate, guttulate, oval, elliptical or cylindrical, often with a flattened base, apex obtuse. Appressoria in slide cultures 7.5–12.5 9 5–8.75 lm (x = 11.5 ± 2.5 9 7.4 ± 1.5, n = 30), abundant, medium to dark brown, irregular, crenate or lobed. Epitype designated here: USA, Florida, on Musa sp., isolated by M. Arzanlou, dried culture deposited in CBS-H20515, ex-epitype living culture CBS 116870. Other materials examined: USA, North Carolina, on Musa sp. fruit skin, M.A. Curtis, K(M) 166978 (holotype). Thailand, Chiang Rai, Bandu, on fruit of Musa sp., 7 Nov. 2009, MFLU10-0976, living strain isolated by P. Noireung, MFLUCC 10-0115. Thailand, Chiang Rai, Bandu, on fruit of Musa sp., 18 July 2009, MFLU10-0977, living strain

Taxonomy Colletotrichum musae (Berk. & M.A. Curtis) Arx, Verh. K. Akad. Wet., tweede sect. 51(3): 107 (1957). Fig. 2a–k MycoBank no.: MB295348. Colonies on PDA attaining 80 mm diameter in 5 days at 27"C, growth rate 16.9–18.4 mm/day (x = 17.6 ± 0.6,

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Fig. 2 Colletotrichum musae (from CBS 116870). a, b Upper and reverse view of culture on potato dextrose agar (PDA) 4 days after inoculation. c, d Conidia. e, f Conidiophores and conidia. g–k Irregularly shaped appressoria. Bars c, e, f 20 lm; d, g–k 10 lm Fig. 3 Colletotrichum musae holotype K(M) 166978

isolated by P. Noireung, MFLUCC 10-0120. Thailand, Chiang Rai, Bandu, on fruit of Musa sp., 9 May 2009, MFLU10-0978, living strain isolated by P. Noireung, MFLUCC 10-0121. Note: The holotype of Myxosporium musae K(M) 166978 (Fig. 3) consists of a dried banana pericarp. No conidia or

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sclerotia could be found on the specimen. Three attempts at DNA extraction from the darkened areas from this specimens were not successful. Berkeley (1874) provided only a brief description of the taxon with very few practical data, and it is impossible to be confident that this specimen is the same as our collections. We have the option of introducing a

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new species or using our collection to epitype this old name. The latter option both is sensible and allows us to stabilize the use of the taxon name sensu Sutton (1980, 1992), who treated C. musae with conidia with similar characteristics to those of our epitype and thus have been used by most authors since Sutton’s publications.

Discussion Colletotrichum musae has been shown to belong to the ‘‘gloeosporioides’’ species complex, and it is widely recognized as a separate species limited to banana (Musa spp.) (Du et al. 2005). Previous phylogenetic analyses of C. musae and Colletotrichum fragariae A.N. Brooks based on the ITS and the 28S-D2 rDNA gene sequences failed to separate them from C. gloeosporioides (Penz.) Penz. & Sacc. (Sreenivasaprasad et al. 1992, 1994; Johnston and Jones 1997). Multilocus phylogeny is now widely applied for the understanding of species relationships in Colletotrichum (Damm et al. 2009; Crouch et al. 2009b). For example, Prihastuti et al. (2009) introduced three new species—Colletotrichum asianum Prihastuti, L. Cai & K.D. Hyde, Colletotrichum fructicola Prihastuti, L. Cai & K.D. Hyde, and Colletotrichum siamense Prihastuti, L. Cai & K.D. Hyde—based on multilocus phylogeny and polyphasic phenotypic characters. In the phylogenetic tree, the epitype and three strains from Asian banana constitute a strongly supported monophyletic clade (see Fig. 1). Colletotrichum musae is most closely related to C. fructicola, which was originally isolated from coffee berries (Prihastuti et al. 2009) but has since been shown to occur on several other hosts (Yang et al. 2009; Phoulivong et al. 2010). The conidial size of C. musae overlaps with that of C. fructicola. However, the mean conidial length of C. musae is significantly greater (14.7 vs. 11.53 lm), and the shape and size of the appressoria are also different (ovoid and clavate, 7.0 9 4.5 lm, in C. fructicola vs. crenate or lobed, 11.5 9 7.4 lm, in C. musae). The Colletotrichum gloeosporioides species complex has been shown to contain several genetically and biologically separated species (e.g., Colletotrichum asianum, Colletotrichum fragariae, Colletotrichum fructicola, Colletotrichum gloeosporioides sensu stricto, Colletotrichum horii B. Weir & P.R. Johnst., Colletotrichum kahawae J.M. Waller & Bridge, and Colletotrichum siamense). However, these species have few distinguishable morphological characters. Therefore, it is essential to generate sequence data and compare these sequence data to those generated from type specimens. Before this study, there were no sequences generated from the type specimen of C. musae, and the current GenBank sequences under the name ‘C. musae’ differ one from another (details not shown). Comparison with strains from type specimens is essential

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for the systematics study and pathogen diagnosis in this group of fungi, and the potential ‘‘standard’’ sample should be based on a properly identified specimen. The ex-epitype strain infected the banana fruit in a short time and showed typical symptoms. Similar results also occurred in the study of Lim et al. (2002). By formally establishing an ex-epitype culture for C. musae that is consistent with the original type with respect to morphology, host and geographic derivation, and generating a multilocus sequence dataset for this strain, we have taken the first and vital step toward informative study of the taxa responsible for this important disease of anthracnose. Acknowledgments This work was financially supported by CAS (Nos. KSCX2-YW-Z-1026 & KSCX2-EW-J-6) and the National Natural Science Foundation of China (NSFC 31070020). This study was also partly funded by Mae Fah Luang University (51101010029 and 52101010002) and the National Plan of Science and Technology, Saudi Arabia (10-BIO 965-10). Dr. P.W. Crous is sincerely thanked for providing fungal strains used in this study.

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Mycoscience (2011) 52:376–382 is not a common pathogen on tropical fruits. Fungal Divers 44:33–43 Prihastuti H, Cai L, Chen H, McKenzie EHC, Hyde KD (2009) Characterization of Colletotrichum species associated with coffee berries in northern Thailand. Fungal Divers 39:89–109 Rannala B, Yang Z (1996) Probability distribution of molecular evolutionary trees: a new method of phylogenetic inference. J Mol Evol 43:304–311 Simmonds JH (1965) A study of the species of Colletotrichum causing ripe fruit rots in Queensland. Queensl J Agric Anim Sci 22:437–459 Simmonds TH, Mitchell RS (1940) Black end and anthracnose of the banana with special reference to Gloeosporium musarum Cke. & Mass. Bull Coun Sci Industr Res Aust 131:6–63 Sreenivasaprasad S, Brown AE, Mills PR (1992) DNA sequence variation and interrelationships among Colletotrichum species causing strawberry anthracnose. Physiol Mol Plant Pathol 41:265–281 Sreenivasaprasad S, Mills PR, Brown AE (1994) Nucleotide sequence of the rDNA spacer 1 enables identification of isolates of Colletotrichum as C. acutatum. Mycol Res 98:186–188 Stover RH, Simmonds NW (1987) Bananas, 3rd edn. Longman, London Sutton BC (1980) The coelomycetes. Commonwealth Mycological Institute, Kew Sutton BC (1992) The genus Glomerella and its anamorph Colletotrichum. In: Bailey JA, Jeger MJ (eds) Colletotrichum: biology, pathology and control. CAB International, Wallingford, pp 1–26 Swofford DL (2002) PAUP*. Phylogenetic analysis using parsimony (*and other methods), 4th edn. Sinauer Associates, Sunderland Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The Clustal X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 24:4876–4882 Wikee S, Cai L, Pairin N, McKenzie EHC, Su YY, Chukeatirote E, Thi HN, Bahkali AH, Moslem MA, Abdelsalam K (2011) Colletotrichum species from jasmine (Jasminum sambac). Fungal Divers 46:171–182 Yang YL, Liu ZY, Cai L, Hyde KD, Yu ZN, McKenzie EHC (2009) Colletotrichum anthracnose of Amaryllidaceae. Fungal Divers 39:123–146 Zhaxybayeva O, Gogarten JP (2002) Bootstrap, Bayesian probability and maximum likelihood mapping: exploring new tools for comparative genome analyses. Genomics 3:1–15

Mycoscience (2011) 52:383–391 DOI 10.1007/s10267-011-0123-6

FULL PAPER

Diversity and community structure of ectomycorrhizal fungi in Pinus thunbergii coastal forests in the eastern region of Korea Keisuke Obase • Jong Kyu Lee • Sang Yong Lee Kun Woo Chun

•

Received: 8 November 2010 / Accepted: 6 May 2011 / Published online: 27 May 2011 ! The Mycological Society of Japan and Springer 2011

Abstract We investigated the diversity and community structure of ectomycorrhizal (EcM) fungi in Pinus thunbergii stands on the eastern coast of Korea. We established two 10 9 10-m plots in six forest stands and sampled soil blocks containing rootlets of mature P. thunbergii trees. EcM roots were classified into morphological groups, and the fungal taxa associated with each morphotype were identified by sequencing the nuclear rDNA internal transcribed spacer region. Cenococcum geophilum and the Atheliales, Clavulinaceae, Russulaceae and Thelephoraceae species were the main members of the EcM fungal community, which included a total of 68 observed fungal taxa. As a whole, the community consisted of a few dominant fungal taxa, such as C. geophilum (28.6% relative abundance), and a large number of rare fungal taxa that showed low abundances and local distributions. Colonization patterns at the local site scale and at the scale of the study plots greatly differed among the EcM fungal taxa; C. geophilum was distributed extensively and was dominant in several study sites, whereas a certain Lactarius sp. was distributed locally but dominated in a given study site. We conclude with a discussion of the relationship between colonization patterns of EcM fungi and soil and environmental conditions. Keywords Atheliales ! Cenococcum ! Clavulinaceae ! Distribution ! Thelephoraceae

K. Obase ! J. K. Lee (&) ! S. Y. Lee ! K. W. Chun College of Forest and Environmental Sciences, Kangwon National University, Chuncheon 200-701, Korea e-mail: [email protected]

Introduction Coastal forests perform functions important for humans, such as alleviating damage from salt and sand dispersion to inland areas and providing places for recreation (Konta 2001). Pinus thunbergii Parl., which grows naturally in China, Korea and Japan, is one of the major constituents of the coastal forests established on sand dunes and oceanfacing mountain slopes in Korea. This tree’s prevalence is possibly due to its excellent tolerance of high salinity, drought stress and wind. In recent years, however, coastal pine forests have declined because of harmful pests such as the pine wood nematode Bursaphelenchus xylophilus (Steiner and Buhrer) Nickle (Yi et al. 1989) and also because of recent severe fire disasters in eastern Korean coastal areas (Son et al. 2006). Several studies and projects, including afforestation projects, have begun to recover and properly maintain coastal pine forests in Korea (Chun et al. 2008). Ectomycorrhizal (EcM) associations are common in pine forests and play a significant role in plant establishment by promoting nutrient and water uptake in host plants and enhancing host tolerance of stressful situations (Smith and Read 1997). Consequently, EcM fungi are considered to be one of the most effective biological resources and have been put to practical use for the reforestation of degraded areas (Quoreshi 2008). The effect of EcM associations on host plant growth differs among different species of EcM fungi (van der Heijden and Kuyper 2003), possibly because of differences in growth and physiological characteristics, such as enzymatic activity (Courty et al. 2005). Therefore, diversity and community structure of EcM fungi are hypothesized to be important for the establishment of coastal pine forests. A few studies have been conducted, mostly in Japan, describing the EcM fungal communities in P. thunbergii

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coastal pine forests. Using molecular tools, several studies have revealed belowground EcM fungal communities in naturally regenerating P. thunbergii seedlings (Taniguchi et al. 2007; Matsuda et al. 2009b) and mature trees (Obase et al. 2009) of coastal pine forests. Matsuda et al. (2009a) have described the abundance and distribution of the dominant EcM fungus Cenococcum geophilum Fr. in soils of four coastal stands of P. thunbergii in Japan. All of these studies were conducted on forest stands established on maritime sand dunes. However, several coastal pine forests on ocean-facing mountain slopes are rooted in forest soil. It has been shown that EcM fungal communities can differ between host plants in different habitats (Iwan´ski and Rudawska 2007). Therefore, it is possible that EcM fungal communities differ among forest stands established on different soil conditions, e.g., maritime sands and forest soils. In this study, we investigated the diversity and community structure of EcM fungi in six stands of P. thunbergii coastal forests established on maritime sand dunes or on ocean-facing slopes on the eastern coast of Korea. We established two 10 9 10-m study plots in each forest stand and sampled soil blocks containing rootlets of mature P. thunbergii trees. EcM roots were classified by morphology, and the fungal taxa associated with each morphotype were identified by sequencing the nuclear ribosomal DNA internal transcribed spacer region. As previously reported, we expected C. geophilum to be the most prevalent and dominant species in the coastal pine forests and that the species composition of EcM fungi would differ among stands, especially between stands established on different soil conditions.

Mycoscience (2011) 52:383–391

sand, which was classified as Regosols (IUSS Working Group WRB 2007), and organic horizons measuring 1–6 cm in thickness (Table 1). At the Samcheok 2 and Uljin sites, the soils consisted of forest soil, which was classified as Podzols (IUSS Working Group WRB 2007), and organic horizons measuring 5–10 cm in thickness. Gangneung, which was located roughly in the center of the study sites, had an annual precipitation of 1,401 mm and an annual average temperature of 12.9"C (the Korean Meteorological Administration). In each study site, we established two 10 9 10-m square plots in areas where other EcM hosts besides P. thunbergii were absent and general environmental factors, such as vegetation and soil conditions, appeared to be similar. The two plots in each study site were separated by a distance of approximately 20–1,000 m. The density of mature trees ranged from 10.5 to 22.0 (/100 m2) in the plots. Three soil cores (5 cm in diameter and depth) were collected at the soil surface in each plot, and soil pH (H2O) and water content (w/w) were determined. Soil pH ranged from 5.0 to 6.0 in each plot (Table 1). Soil water content was low in Sokcho, Gangneung and Samcheok 1 (3.4–4.8%), intermediate in Yangyang (8.4%) and relatively high in Samcheok 2 and Uljin (12.8–14.0%). Sampling methods and observation of EcM colonization

Study site

In September and October 2009, we sampled 16 soil blocks (5 9 5 9 20 cm length, width and depth, respectively) at regular intervals of 3 m in each study plot. We recognized that most of the EcM fungi in the soil blocks were associated with mature trees. All samples were stored in plastic bags at 4"C until further analysis. Roots were separated from the adhering soil by careful washing in tap water over a 0.5-mm sieve. All root segments were viewed under a dissecting microscope after

We established six study sites in the following P. thunbergii coastal forest sites on the eastern coast of Korea in the province of Kangwon-do: Sokcho, Yangyang, Gangneung, Samcheok (2 locations) and Uljin (Fig. 1). All of the coastal pine forests are located 60–250 m inland from the shoreline and measure from a few hundred to several thousand meters in width. The distances between adjacent study sites ranged from 15 to 40 km. The mature forests consisted primarily of P. thunbergii that were estimated to be approximately 40–50 years old. Several woody and herbaceous plants, such as Artemisia spp., Commelina communis L., Festuca ovina L., Quercus dentata Thunb., Rosa rugosa Thunb. and Robinia pseudoacacia L., were sparsely distributed in the understory. At the Sokcho, Yangyang, Gangneung and Samcheok 1 sites, the soils consisted entirely of maritime

Fig. 1 Locations of the six study sites on the eastern coast of Korea

Materials and methods

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385

Table 1 Soil properties such as soil type, texture, pH, water content and depth of humus layer in each study site Study sites

Sokcho

Yangyang

Gangneung

Samcheok 1

Samcheok 2

Uljin

Soil typea

Regosols

Regosols

Regosols

Regosols

Podzols

Podzols

Soil texture

Sandy

Sandy

Sandy

Sandy

Loamy

Sandy/loamy

5.4 ± 0.7

5.0 ± 0.5

5.0 ± 0.2

6.0 ± 0.4

5.0 ± 0.4

5.3 ± 0.4

4.8 ± 1.0

8.2 ± 2.2

4.2 ± 3.2

3.4 ± 2.4

14.0 ± 3.6

12.8 ± 4.1

1–2

3–5

2–4

2–6

5–10

5–6

pH (H2O)b Soil water content (%)

b

Depth of humus layer (cm) a

Soils were classified according to IUSS Working Group WRB (2007)

b

Averages and standard deviations were indicated

being cut into 5–10-cm sections. Vital EcM roots per soil block were classified into distinguishable groups by gross morphology. One to five root tips per group and soil block were then selected, and their microscopic characteristics, such as hyphal arrangement on the surface of mantle and emanating hypha, were observed (Ingleby et al. 1990) to further characterize the EcM root tips. EcM roots were classified into morphotypes that possessed distinguishable gross and/or microscopic morphologies. One EcM root tip of each morphotype was placed into 1.5-ml microtubes at -20"C for DNA extraction. The number of replicates ranged from 1 to 9 per morphotype, according to the abundance and/or frequency of each. Identification of EcM fungi EcM fungal DNA was extracted from one EcM root tip of each morphotype using the DNeasy Plant Mini kit (QIAGEN, USA) according to the manufacturer’s instructions. DNA amplification of ITS regions, including the 5.8S rDNA region, was performed using Takara Ex Taq (TAKARA, Japan) with a specific primer for higher fungi ITS1F (Gardes and Bruns 1993) and basidiomycete ITS4B (Gardes and Bruns 1993). In case the above primer pair failed to amplify the DNA, a primer set for ITS1F and the universal primer ITS4 (White et al. 1990) were used. We used the following polymerase chain reaction (PCR) amplification conditions: 94"C for 3 min, 30 cycles of 94"C for 30 s, 50"C for 30 s and 72"C for 2 min, with a final hold at 72"C for 10 min. Purification of PCR products and sequencing reactions were entrusted to Macrogen Inc. (Seoul, Korea). Sequencing reactions were conducted using the ABI 3730xl Analyzer (Applied Biosystems) with ITS1 and ITS4 primers. EcM root DNA sequences were submitted to the National Center for Biotechnology Information (NCBI: http://blast.ncbi.nlm.nih.gov) and compared with the GenBank database using the nucleotide-nucleotide basic local alignment search tool algorithm (BLAST program). Only library sequences that were derived from sporocarps and assigned names at the species or genus level were used for the comparison. The only exceptions to

this criterion were identifications of C. geophilum and Ceratobasidiaceae, which were based on sequences from sclerotia or isolates from roots. Identifications of EcM fungal taxa followed the criteria described below. The EcM root sequences that matched library sequences with the greatest similarity (C98%) to the full lengths of the ITS1 and ITS2 regions were designated by their species names. If the aligned sequences lacked a part or the full length of the ITS1 or ITS2 regions, genus or family names were assigned, respectively. If the highest sequence similarities were less than 98%, EcM root sequences and library sequences were submitted together to neighbor-joining analysis. EcM root sequences with affinities to only one genus were designated by the genus name. EcM root sequences with affinities to several genera within a family were designated by the family name. Each designation was followed by the numbers. Estimation of mycorrhizal colonization Soil blocks that contained \100 EcM root tips were excluded from all analyses. The EcM abundance, as indicated by the number of EcM root tips, of each morphotype was recorded separately for each soil block. The relative abundance was calculated in each study site as a ratio of abundance of a given morphotype to total EcM abundance and then averaged over all the study sites. The colonization frequencies of EcM fungal taxa, indicated by the ratio of the number of soil blocks that contained a given EcM morphotype to the total number of soil blocks, were also recorded in all the study sites. Statistical analysis To estimate whether sampling efforts were sufficient to describe EcM fungal flora, species accumulation curves were drawn by plotting the mean of the accumulated number of expected species in pooled samples after 1,000 randomizations without replacement using EstimateS program version 8.0 (Colwell 2005). The study site was selected as a sampling unit. In addition, the estimated

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measurements of EcM fungal species richness were calculated using Chao 2 and Jackknife 2 estimators. Principal component analysis (PCA) and redundancy analysis (RDA) were carried out to reveal trends within the EcM fungal community and the potential effects of measured environmental variables (soil type, pH, water content and depth of humus layer) on EcM fungal community structure, respectively. Relative abundances of EcM fungal taxa in each study site were standardized by arcsine square root transformation. Soil types were transformed to dummy variables (0 for Regosols and 1 for Podzols). Forty-two rare taxa that occurred at only one study site and showed\1.0% relative abundance and \10.0% colonization frequency were excluded from the analysis. Within the RDA, permutation tests (n = 999) were performed to test the significance of the relationship between community data and the environmental variables. The PCA and RDA were conducted within the Vegan package (Oksanen et al. 2008) in the statistical program R version 2.11.1 (R Development Core Team 2010). To calculate the correlations between community similarities and spatial distances among study sites, a Mantel test was conducted within the Ecodist package (Goslee and Urban 2007) in the statistical program R version 2.11.1 (R Development Core Team 2010) with 10,000 permutation tests. Horn-Morisita distances based on the abundance of each EcM morphotype in each study site and Euclidian distances based on geographic coordinates of each study site were used in the analysis.

Results Identification of EcM fungi Because all sequences obtained from a given EcM morphotype were categorized into the same taxa, we judged that EcM root tips with identical morphologies might be formed by identical EcM fungal taxa. The ITS regions of all EcM roots were successfully amplified; however, we could not obtain complete sequences from 36% of the samples. Consequently, we identified a total of 68 EcM fungal taxa from 121 sequenced root tips (Table 2). Of these, we identified 11 and 30 taxa at the species and genus levels, respectively. The remaining 26 taxa were categorized at the family, order or phylum level. We were unable to obtain a clear sequence from the one remaining EcM morphotype, which was referred to as unidentified EcM fungi. Diversity of EcM fungi All soil samples contained EcM roots of P. thunbergii. The total number of EcM root tips was 120,762. The number of

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EcM root tips per soil block ranged from 12 to 3,767 and averaged 629. Six soil cores that contained \100 EcM root tips were excluded from further analysis. EcM fungal species richness per soil block ranged from 1 to 9 species with an average of 4.3. Between 15 and 22 species were observed at each study site. The species accumulation curve was ascending when all study sites were randomly sampled (not shown). The estimated species richness values using Chao 2 and Jackknife 2 estimators were 164.8 and 128.7, respectively. Taxa belonging to Thelephorales (e.g., Tomentella) were the most species rich (21 taxa), followed by Agaricales (11 taxa), Russulales (7 taxa) and Boletales (6 taxa) (Table 2). Colonization pattern of each EcM taxon Only one fungal taxon, C. geophilum, showed a high relative abundance (28.6%), while the others were found at relatively low abundances (Table 2). Fourteen taxa ranged from 1.0 to 8.8% in prevalence, and the remaining 53 taxa comprised \1.0% of the community. In addition, C. geophilum showed the highest colonization frequency (79.0%), while the others showed intermediate to low colonization frequencies; 8 taxa ranged from 10.8 to 33.9%, and the remaining 59 taxa made up \10.0%. Seven taxa were observed at multiple study sites. Two taxa (C. geophilum and Atheliales 1) were found in six sites, three taxa (Clavulinaceae 1, Russula sp. and Thelephoraceae 2) at five sites and two taxa (Atheliales 2 and Lactarius hatsudake Tanaka) in four sites. Forty-four taxa were observed only in one study site. Among the 14 taxa aside from C. geophilum that had more than 1.0% relative abundance, Atheliales 1 showed the highest colonization frequency (33.9%), followed by Russula sp. (25.8%) and Clavulinaceae 1 (25.3%); these taxa were observed in 6, 5 and 5 study sites, respectively (Table 2). Clavulinaceae 2, Thelephoraceae 3 and Lactarius sp. were observed in two, two and one study sites, respectively, and they showed relatively low colonization frequencies of 8.6, 8.1 and 12.9%, respectively. However, these taxa showed high relative abundance and colonization frequencies in one study site (Fig. 2). Fifty-three rare taxa showing \1.0% relative abundance also had low colonization frequencies, ranging from 0.5 to 10.8%. Of the rare taxa, six [Sistotrema sp., Pseudotomentella tristis (P. Karst.) M. J. Larsen, Suillus granulatus (L.) Roussel, Tomentella sp. 1, Tricholoma flavovirens (Pers.) Lundell and Tuber sp.] were observed in three study sites, and the others were observed in two (5 taxa) or one study sites (42 taxa). The Mantel test found no significant correlation (Mantel r = -0.03; P = 0.51) between community similarities and spatial distances among study sites.

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Table 2 Possible identities of EcM fungal taxa based on comparisons of obtained sequences with references in Genebank database at NCBI, using BLAST program Possible identity

Obtained sequence

BLAST match with high similarity

EcM colonization

Accession no.

Length (bp)

Definition

Accession No.

Amanita spissa

AB587730

661

Amanita spissa

AJ889924

658/661 (99)

0.7

2.2

1

Atheliales 1

AB587731

560

Amphinema byssoides

AY219839

494/585 (85)

4.6

33.9

6

Atheliales 2

AB587732

569

Amphinema byssoides

AY219839

378/439 (87)

3.4

17.2

4

Similarity (%)

RA (%)

CF (%)

Stand (/6)

Atheliales 3

AB587733

576

Amphinema byssoides

AY838271

501/597 (84)

3.7

22.6

3

Basidiomycetes 1

AB587734

632

Tylospora asterophora

AF052558

239/256 (94)

0.2

5.9

2

Basidiomycetes 2

AB587735

599

Alloclavaria purpurea

AY228345

470/630 (75)

\0.1

0.5

1

Boletus sp.

AB587736

581

Boletus hiratsukae

EU231960

581/581 (100)

0.7

4.3

1

Cenococcum geophilum

AB587740

451

Cenococcum geophilum

AF495462

446/451 (99)

28.6

79.0

6 1

Ceratobasidiaceae 1

AB587741

643

Ceratobasidium sp.

GQ175300

590/650 (91)

0.7

5.4

Ceratobasidiaceae 2

AB587742

687

Vouchered mycorrhizae

AB303056

601/605 (99)

0.3

1.6

1

Clavulinaceae 1

AB587737

612

Clavulina sp.

FN669173

576/638 (91)

7.8

25.3

5

Clavulinaceae 2

AB587738

597

Clavulina sp.

FN669173

486/555 (88)

5.3

8.6

2

Cortinarius sp. 1

AB587743

588

Cortinarius californicus

FJ039588

585/588 (99)

0.4

3.2

1

Cortinarius sp. 2 Cortinarius sp. 3

AB587744 AB587745

623 651

Cortinarius aff. pauperculus Cortinarius aff. pauperculus

GQ159858 GQ159858

594/623 (96) 627/648 (97)

0.3 0.1

2.2 2.2

1 1

Entolomataceae

AB587746

923

Entoloma sinuatum

GQ397994

775/894 (87)

0.4

3.2

1

Hymenochaetaceae

AB587747

936

Coltricia cf. oblectans

AM412246

413/416 (99)

\0.1

1.6

1

Inocybe sp. 1

AB587748

660

Inocybe sp.

GQ892990

587/618 (95)

0.9

9.1

2

Inocybe sp. 2

AB587749

638

Inocybe flocculosa var. flocculosa

HQ604084

625/653 (96)

0.2

1.6

1

Inocybe sp. 3

AB587750

689

Inocybe arthrocystis

AM882856

632/664 (96)

\0.1

0.5

1

Laccaria sp.

AB587751

542

Laccaria amethystina

AB211270

534/543 (98)

\0.1

1.1

1

Lactarius akahatsu

AB587752

704

Lactarius akahatsu

AB301609

704/704 (100)

1.1

3.8

2

Lactarius hatsudake

AB587753

677

Lactarius hatsudake

AB301611

677/677 (100)

2.7

9.1

4

Lactarius sp.

AB587754

807

Lactarius alnicola

DQ099898

750/809 (93)

8.8

12.9

1

Otidea sp.

AB587756

580

Otidea bufonia

EU784387

563/583 (97)

0.5

1.6

1

Peziza sp.

AB587757

578

Peziza sp.

FN669234

565/579 (98)

0.1

2.7

1

Pezizales

AB587758

910

Helvella elastica

AF335455

435/505 (87)

0.1

1.6

1

Pseudotomentella tristis

AB587759

708

Pseudotomentella tristis

GQ267480

703/711 (99)

0.3

4.8

3

Psudotomentella sp. Pyronemataceae

AB587761 AB587763

706 560

Pseudotomentella tristis Trichophaea cf. hybrida

AJ889968 DQ200834

523/556 (95) 509/560 (91)

1.3 0.1

5.4 2.2

1 1

Rhizopogon roseolus

AB587764

683

Rhizopogon roseolus

HM036649

678/682 (99)

0.4

1.1

1

Rhizopogonaceae

AB587765

819

Rhizopogon succosus

AF062933

505/538 (94)

0.3

3.8

1

Russula nauseosa

AB587768

637

Russula cf. nauseosa

GU371293

629/637 (99)

0.1

0.5

1

Russula sp.

AB587766

640

Russula cf. fuscorubroides

HQ604842

613/645 (96)

6.2

25.8

5

Russulaceae 1

AB587755

640

Lactarius cf. piperatus

AB459515

569/667 (86)

0.3

1.6

1

Russulaceae 2

AB587767

646

Russula mariae

EU819426

593/653 (91)

\0.1

1.1

1

Sebacina sp. 1

AB587769

582

Sebacina incrustans

EU819442

550/584 (95)

2.7

9.7

2

Sebacina sp. 2

AB587770

579

Sebacina sp.

DQ974768

557/579 (97)

0.1

1.1

2

Sebacinaceae 1

AB587771

638

Sebacina sp.

DQ974768

516/620 (84)

0.1

1.1

2

Sebacinaceae 2

AB587772

511

Sebacina sp.

FN669252

453/511 (89)

\0.1

0.5

1

Sistotrema sp.

AB587739

581

Sistotrema sp.

FN669255

548/588 (94)

0.6

7.0

3

Suillus granulatus

AB587774

653

Suillus granulatus

AY898617

643/656 (99)

0.3

7.0

3

Suillus sp. 1

AB587773

510

Suillus bovinus

AB284446

509/519 (99)

0.9

7.0

1

Suillus sp. 2

AB587775

409

Suillus luteus

AB284448

409/409 (100)

1.1

3.2

2

Thelephora terrestris

AB587776 AB587762

625 631

Thelephora terrestris Pseudotomentella sp.

EU427330 GQ267479

619/625 (99) 566/636 (89)

\0.1 \0.1

1.6 1.6

1 1

Thelephoraceae 1

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Table 2 continued Possible identity

Obtained sequence

BLAST match with high similarity

EcM colonization

Accession no.

Length (bp)

Definition

Accession No.

Similarity (%)

Thelephoraceae 2

AB587777

632

Thelephora penicillata

U83484

592/632 (94)

3.8

14.0

5

Thelephoraceae 3

AB587778

620

Tomentella ferruginea

EU819497

564/634 (89)

3.2

8.1

2

RA (%)

CF (%)

Stand (/6)

Thelephoraceae 4

AB587781

628

Tomentella cf. sublilacina

AJ889982

574/630 (92)

0.3

1.6

1

Thelephoraceae 5 Thelephoraceae 6

AB587789 AB587790

625 627

Tomentella sp. Tomentella cinerascens

EF644116 U83483

568/635 (90) 547/633 (87)

0.3 \0.1

1.1 0.5

1 1 1

Thelephoraceae 7

AB587791

633

Tomentella sp.

DQ974780

571/634 (91)

\0.1

0.5

Thelephorales

AB587760

374

Pseudotomentella tristis

AJ889979

306/375 (81)

0.2

0.5

1

Tomentella badia

AB587779

636

Tomentella badia

AF272917

574/588 (98)

0.8

6.5

2

Tomentella sp. 1

AB587780

634

Tomentella sp.

DQ822830

599/636 (95)

0.6

9.7

3

Tomentella sp. 2

AB587782

419

Tomentella cf. coerulea

AY010274

404/416 (98)

0.2

3.2

1

Tomentella sp. 3

AB587783

628

Tomentella ellisii

DQ974775

583/627 (93)

0.1

0.5

1

Tomentella sp. 4

AB587784

628

Tomentella fuscocinerea

GU214810

581/627 (93)

\0.1

0.5

1

Tomentella sp. 5

AB587785

626

Tomentella sp.

FM955848

575/618 (94)

\0.1

0.5

1

Tomentella sp. 6

AB587786

629

Tomentella sp.

EF655702

581/625 (93)

0.8

5.4

1

Tomentella sp. 7

AB587787

628

Tomentella sp.

DQ822830

594/628 (95)

0.6

5.9

1

Tomentella sp. 8

AB587788

629

Tomentella ramosissima

U83480

607/629 (97)

0.5

4.3

1

Tomentellopsis sp.

AB587792

652

Tomentellopsis submollis

AJ410774

574/606 (95)

\0.1

1.1

1

Tremellodendron sp.

AB587793

591

Tremellodendron pallidum

GQ166897

570/590 (97)

0.4

0.5

1

Tricholoma flavovirens

AB587794

666

Tricholoma flavovirens

AF377181

652/667 (98)

0.3

4.3

3

Tricholoma sp. Tuber sp.

AB587795 AB587796

666 650

Tricholoma flavovirens Tuber separans

AF377181 HM485385

634/668 (95) 575/615 (94)

0.2 0.7

1.1 10.8

1 3

Unidentified EcM fungus

–

–

–

–

–

\0.1

1.1

1

Relative abundance (RA), colonization frequency (CF) of each fungal taxa and frequency of forests stands in which each fungal taxon was observed were also indicated

PCA revealed no clear impact of soil type on the EcM fungal community, probably because of soil type variations among the study sites (Fig. 3a); one of the study sites containing Podzol soil (Samcheok 2) was largely segregated from study sites with Regosol soil, whereas the other study sites (Uljin) appeared close together on a plot of the first two axis scores with Sokcho, where the soil type was Regosols. The RDA showed similar study site distributional patterns to those of the PCA on the ordination (Fig. 3b). No measured environmental variables significantly influenced the EcM fungal community composition.

Discussion Ectomycorrhizal fungal diversity We found species-rich and diverse EcM fungal communities in mature P. thunbergii coastal forests; 68 total taxa of ectomycorrhizal (EcM) fungi were observed in 6 stands of P. thunbergii coastal forests (Table 2). However, it appeared that our results only showed a small fraction of

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the total EcM fungal diversity in coastal pine forests. An ascending species accumulation curve that estimated a species richness approximately 2–3 times higher (128.7 or 164.8 taxa) than the actual richness (68 taxa) indicated that much of the new additional fungal taxa would be detected if sampling efforts were increased. Additionally, the low ratio of EcM roots subjected to DNA analysis (121 root tips) to the total number of EcM roots observed (120,762 root tips) may have increased errors in accurately discriminating EcM roots (i.e., two or more closely related species could be classified together into one group by morphotyping because of their similarity in morphology), perhaps resulting in an underestimation of the actual fungal diversity. Increasing the number of EcM root tips for DNA analysis is preferable in order to more accurately understand the fungal community and its diversity. Several dozen species of EcM fungi have been recorded in pine forests dominated by a single woody plant, such as natural stands of bishop pine (Pinus muricata D. Don) (20 taxa; Gardes and Bruns 1996), P. thunbergii coastal forests (27 taxa; Obase et al. 2009), red pine (Pinus resinosa Ait.) plantations (39 taxa; Koide et al. 2005), boreal Scots

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Cenococcum geophilum and species of Atheliales, Clavulinaceae, Russulaceae and Thelephoraceae were the main members of the EcM fungal communities (Table 2). Cenococcum geophilum is one of the most common EcM fungi in the world (Trappe 1964; LoBuglio 1999) and often abundantly colonizes in several forest ecosystems (e.g., Valentine et al. 2004; Blom et al. 2009). Cenococcum geophilum has also been reported as a dominant fungus in the roots of P. thunbergii seedlings (Taniguchi et al. 2007; Kataoka et al. 2008; Matsuda et al. 2009b) and mature trees (Matsuda et al. 2009a; Obase et al. 2009) in coastal pine forests. Some fungal taxa that were presumed to belong to Atheliaceae (e.g., Tylospora and Amphinema) (Erland 1995), Clavulinaceae (e.g., Clavulina) (Koide et al. 2005), Russulaceae (e.g., Russula and Lactarius) and Thelephoraceae (e.g., Thelephora and Tomentella) (Horton and Bruns 2001) have often been detected in inland forest ecosystems and have also been observed as the second or third most dominant species or a rare species in coastal pine forests of Japan (Taniguchi et al. 2007; Kataoka et al. 2008; Matsuda et al. 2009b). It appears that the EcM fungi that have colonized in P. thunbergii coastal forests are similar between Korea and Japan, and that the structures of the EcM fungal communities comprising those fungal taxa are likely similar among P. thunbergii coastal forests. Community structure of ectomycorrhizal fungi

Fig. 2 Relative abundance (bars) and colonization frequency (black circles) of ectomycorrhizal fungal taxa that showed an average relative abundance [1.0% over all of the study sites

pine (Pinus sylvestris L.) forests (43 taxa; Jonsson et al. 1999a, 135 taxa; Jonsson et al. 1999b), Pinus muricata D. Don coastal forests (48 taxa; Peay et al. 2007) and old stands of lodgepole pine (Pinus contorta Douglas ex Louden) (81 taxa; Douglas et al. 2005). It is difficult to compare the species richness of EcM fungi among various studies because of differences in sampling strategies; however, EcM fungal species richness in P. thunbergii coastal forests was as high, as has been reported in other pine forests.

Overall, C. geophilum was the most common and dominant taxon at the study sites (Table 2). A few fungal taxa, such as C. geophilum, Lactarius sp., Clavulinaceae 1 and Russula sp., showed high relative abundances, but the others showed intermediate to low relative abundances. A similar pattern was also observed for colonization frequency. The presence of a few dominant fungi and a large number of rare fungi is a common pattern in EcM fungal communities associated with P. thunbergii coastal forests (Matsuda et al. 2009b; Obase et al. 2009) or other tree species (e.g., Valentine et al. 2004). Most of the rare fungal taxa were distributed locally at both the stand scale and plot scale, and tended to show low relative abundances. These results indicate that the local distributions of rare EcM fungal taxa contributed to the species-rich EcM fungal communities in P. thunbergii coastal forests. Colonization patterns at the local scale of study sites and the small scale of study plots differed among EcM fungal taxa. Cenococcum geophilum was distributed extensively at the local scale and showed various levels of colonization, often being the dominant taxon, among study sites, while Lactarius sp. was distributed locally but dominated at only one study site (Fig. 2). Different distribution patterns among EcM fungal taxa have been well documented in several studies at small scales (e.g., within 20 9 20 m

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Fig. 3 a Principal component analysis displaying the positions of study sites. Axis 1 and axis 2 explained 38.7 and 24.3% of the variability, respectively. b Redundancy analysis displaying the positions of study sites (squares) and EcM fungal taxa (circles), and the effects of soil type, soil pH, soil water content and depth of humus layer

(arrows) on ectomycorrhizal fungal community structure. Axis 1 and axis 2 explained 33.3 and 24.1% of the variability, respectively. Cla1 Clavulinaceae 1, Cla2 Clavulinaceae 2, Ce Cenococcum geophilum, Lac Lactarius sp., Ru Russula sp., So Sokcho, Ya Yangyang, Ga Gangneung, S1 Samcheok 1, S2 Samcheok 2, Ul Uljin

plots) (Lilleskov et al. 2004; Pickles et al. 2010), but only a few studies have described the distribution of certain EcM fungi at a local scale. In P. thunbergii coastal forests, Matsuda et al. (2009a) investigated the distribution of C. geophilum at the local scale (several tens of kilometers in extent) in coastal pine forests of Japan and found that this fungus was distributed ubiquitously and dominant, but its colonization ratio varied among study sites (20.0 to 62.6% of EcM roots). The presence of colonization pattern differences among EcM taxa and in certain EcM fungal taxa among study sites could be related to several factors, such as spatial heterogeneity of soil conditions (e.g., Bruns 1995) and interspecific interactions among EcM fungi (i.e., competence and coexistence) (Pickles et al. 2010). Although we could not find distinct effects of the measured environmental variables on EcM fungal colonization, possibly because the most important soil parameters went unmeasured or because the number of replicated study sites was insufficient to understand the trends within the whole EcM fungal community (Fig. 3), field observations provided several insights into the distribution patterns of given EcM fungal taxa. In this study, low abundance and local colonization of C. geophilum were observed in one forest stand (Samcheok 2) where Lactarius sp. was dominant (Fig. 2). In study sites where C. geophilum was dominant, the soils consisted entirely of maritime sand with relatively shallow litter and low water content. Previous studies have suggested that C. geophilum is an excellent colonizer in areas where drought stress is high and where the soil is very weakly developed (Obase et al. 2009). In contrast, in a study site where Lactarius sp. was dominant, soils consisted of forest soil that appeared to be compacted, had poor drainage and contained a relatively thick and humid

humus layer. Most EcM roots extended into the humus layer, and Lactarius sp. exhibited distinct clumps in this layer. Dense root colonization by Lactarius species in organic soil layers has also been reported in a previous study (Genney et al. 2006).

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Conclusions We investigated the colonization status of EcM fungi in six stands of P. thunbergii coastal forests and found speciesrich and diverse communities in which each EcM fungal taxon showed various distribution patterns at local scales of the study sites and at small scales of the study plots. Estimation of species richness by an extrapolation method revealed that high sampling efforts are required to elucidate the entire EcM fungal community in coastal pine forests within an area that extends for several hundred kilometers. The dominance of C. geophilum is likely to be a common occurrence in P. thunbergii coastal forests; however, these results also show that EcM fungal communities could differ among study sites and that other taxa could be superior to C. geophilum in colonization at the study plot scale. It is important to understand the ecological characteristics of EcM fungi and their colonization patterns in various environmental conditions when attempts are undertaken to apply EcM fungi for revegetation techniques in disturbed coastal pine forests. Acknowledgments This study was carried out with the support of ‘Forest Science & Technology Projects (project no. C1002315)’ provided by the Korea Forest Service. We acknowledge special supports from the members of the Laboratory of ‘‘Forest Resources Development’’ and ‘‘Resources Protection’’ in Kangwon National University.

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Mycoscience (2011) 52:392–400 DOI 10.1007/s10267-011-0125-4

FULL PAPER

Two new species of Agaricales and a new Japanese record for Chaetocalathus fragilis from Ishigaki Island, a southwestern island of Japan Haruki Takahashi

Received: 9 October 2010 / Accepted: 17 May 2011 / Published online: 5 June 2011 ! The Mycological Society of Japan and Springer 2011

Abstract Two new species of Agaricales and a new Japanese record for Chaetocalathus fragilis from Ishigaki Island, a southwestern island of Japan, are fully described and illustrated: (1) Crinipellis (section Grisentinae) rhizomorphica sp. nov. produces brownish orange, fibrillosesquamulose basidiomata accompanied by white thread-like rhizomorphs on the dead twig, olivaceous hairs in KOH, and oblong-ellipsoid, relatively long basidiospores; (2) Chaetocalathus (section Holocystis) fragilis is a new record for Japan, growing on the dead twig; (3) Psilocybe (section Cubensae) capitulata sp. nov. forms a furfuraceous-squamulose pileus, cyanescent flesh, a persistent, membranous annulus, capitulate pilocystidia, and has a coprophilous habit on cow dung. Keywords Crinipellis rhizomorphica ! Japanese mycobiota ! Psilocybe capitulata ! Taxonomy Introduction Ishigaki Island is the second largest island of the Yaeyama Island group and is situated in the southwestern part of the Ryukyu Archipelago. The forest types of the Yaeyama Islands exist in subtropical climates and include predominantly laurel-leaved forest in the mountain region and a mangrove forest community developed on saline, muddy soil of estuaries or inlets under the high tide mark. The most intensively sampled area is the laurel-leaved forest type dominated by Quercus miyagii Koidz. and Castanopsis sieboldii (Makino) Hatus. ex T. Yamaz. et Mashiba H. Takahashi (&) 284-1 Ouhama, Ishigaki, Okinawa 907-0001, Japan e-mail: [email protected]

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from 100 to 250 m altitude in Mt. Banna and the surrounding region. As for the agaric flora of Yaeyama Islands, no intensive investigation has hitherto been carried out, with the exception of the studies by Miyagi (1958, 1964, 1971), who reported 53 species of agarics and boletes from Ishigaki Island and Iriomote Island. During studies on the diversity of macrofungi in the southwestern islands of Japan, a new distributional record and two species representing undescribed Agaricales were found, namely, Chaetocalathus fragilis, Crinipellis rhizomorphica sp. nov., and Psilocybe capitulata sp. nov., which are fully described and illustrated here.

Materials and methods Macroscopic features are all based on fresh materials. For microscopic observations, free-hand sections of the fresh basidiomata were examined in Melzer’s reagent, 5% KOH, or distilled water. Basidiospores were measured in the side view in Melzer’s reagent; for each collection, at least 20 basidiospores were measured. Color notations in parentheses are taken from Kornerup and Wanscher (1983). Specimens cited here are deposited in the Kanagawa Prefectural Museum of Natural History, Japan (KPM).

Taxonomic descriptions 1. Crinipellis rhizomorphica Har. Takah., sp. nov. Figs. 1–5 MycoBank no.: MB 519031 Pileo 5–8(–12) mm lato, primo hemisphaerico, dein planoconvexo, centro depresso umbonato zona leniter elevato,

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fibrilloso-squamuloso, brunneolo-aurantiaco; stipite 8–15 (–20) 9 0.7–1.3 mm, subaequali vel ad basim leniter incrassato, centrali, cavo, brunneolo-aurantiaco, strigoso-fibrilloso, mycelio basali non affixo; rhizomorphis filiformibus, albis; lamellis adnexis, albis; basidiosporis (9.5–)11–13 9 (4–)4.5–5 lm, oblongo-ellipsoideis, levibus, hyalinis, inamyloideis; basidiis 25–27 9 3–8 lm, tetrasporis; cheilocystidiis 17–33 9 6–10 lm, subclaviformibus, aliquot breviter lobatis; pleurocystidiis nullis; pilis pilei 400–600 9 4–6 lm, crassitunicatis, pseudo-

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amyloideis, in KOH olivaceis, septatis, pilis stipitis similibus; hyphis fibulatis. Holotypus: In ramulis dejectis in silva, Banna-dake, Ishigaki-shi, Okinawa Pref., Japonia, 25 July 2010, H. Takahashi and Y. Terashima (KPM-NC0017526). Etymology: from Latin, rhizomorphica, referring to the thread-like, white rhizomorphs on the substratum. Pileus (Fig. 1) 5–8(–12) mm in diameter, at first hemispherical to convex with incurved margin, then broadly

Figs. 1–5 Crinipellis rhizomorphica (Holotype): Fig. 1 Basidiomata. Fig. 2 Rhizomorphs. Fig. 3 Hairs of pileus. Fig. 4 Cheilocystidia. Fig. 5 Basidiospores. Bars 1 10 mm; 2 15 mm; 3 20 lm; 4 10 lm; 5 13 lm. Color photos of Crinipellis rhizomorphica can be seen at http://www7a.biglobe.ne.jp/ *har-takah/page128.html

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convex to nearly plane, sometimes with a depressed center, often with a brownish orange (6C7–8 or 7C7–8) or blackish, broad umbo at the center; surface dry, radially fibrillose-squamulose with strigose, brownish orange (6C7–8 or 7C7–8) to brown (7D7–8) hairs projecting beyond the margin, often concentrically zoned, dull, opaque. Flesh very thin (up to 0.5 mm), white, pliant but easily broken, odor and taste not distinctive. Stipe 8–15 (–20) 9 0.7–1.3 mm, subequal or slightly enlarged at the base, central, slender, terete, hollow, entirely strigosefibrillose with brownish orange (6C7–8 or 7C7–8) to brown (7D7–8) hairs, basal mycelium not seen. Lamellae adnexed, 19–23 reach the stipe, with 1–3 series of lamellulae, up to 1.5 mm broad, white; edges ciliate to fimbriate, concolorous. Rhizomorphs (Fig. 2) independent of the formation of basidiomata, growing into the air, scattered on the substratum, 5–15 (–30) 9 0.02–0.6 mm,

thread-like, gradually tapering toward the apex, tough, not branched, dense, parallel, white, inamyloid in the apical portion but otherwise dextrinoid, hyaline or turning pale olivaceous in KOH; constituent hyphae similar to the hairs of basidiomata, 2–5 lm diameter, cylindrical, with rounded apex, with smooth walls 0.5–2 lm thick, often with several secondary septa near the apical portion. Basidiospores (Fig. 5) (9.5–)11–13 9 (4–)4.5–5 lm (Q = length/breadth: 2.44–2.6), oblong-ellipsoid, smooth, colorless, inamyloid, rarely septate, thin-walled. Basidia 25–27 9 3–8 lm, clavate, 4-spored; basidioles fusiform to subclavate. Cheilocystidia (Fig. 4) 17–33 9 6–10 lm, gregarious, forming a compact sterile edge, projecting from the hymenium, subclavate, with two or three short cylindrical apical appendages 1–5 9 1–2.5 lm, smooth, colorless, inamyloid, hyaline in KOH, thin-walled. Pleurocystidia, none.

Figs. 6–10 Chaetocalathus fragilis (KPM-NC0017300): Fig. 6 Basidiomata. Fig. 7 Basidiospores. Figs. 8, 9 Cheilocystidia (metuloid) mounted in KOH. Fig. 10 Cheilocystidia (metuloid) mounted in

Melzer’s reagent. Bars 6 1 mm; 7 8 lm; 8–10 10 lm. Color photos of Chaetocalathus fragilis can be seen at http://www7a.biglobe.ne. jp/*har-takah/page134.html

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Hyphae of hymenophoral trama 3–8 lm wide, parallel, subcylindrical, not inflated, smooth, colorless, inamyloid, hyaline in KOH, thin-walled. Pileipellis a hypotrichial layer of subcylindrical cells 5–7 lm wide, with smooth, colorless walls 1–2 lm thick, inamyloid or weakly dextrinoid; hairs of pileus (Fig. 3) 400–600 9 4–6 lm, arising directly from the hypotrichium, repent or erect, cylindrical, with a rounded apex, sometimes flexuous, with smooth, brownish orange (6C7–8) walls 1–2 lm thick, strongly dextrinoid, turning olive yellow (3C6–8) in KOH, occasionally with several secondary septa. Hyphae of pileitrama 5–10 lm wide, similar to those of the hymenophoral trama. Hairs of stipe similar to those of the pileus. Stipe trama composed of longitudinally running, cylindrical hyphae 5–13 lm wide, smooth, colorless, inamyloid, hyaline in KOH, thin-walled. Clamps present in all tissues. Habitat: Gregarious or scattered, on dead twigs (unidentified substrata), June to July. Specimens examined: KPM-NC0017526 (holotype), Banna-dake, Ishigaki-shi, Okinawa Pref., on dead twigs, 25 July 2010, coll. Takahashi, H. and Terashima, Y.; KPMNC0017297, same place, 3 July 2006, coll. Takahashi, H.; KPM-NC0017298, same place, 13 June 2006, coll. Takahashi, H.; KPM-NC0017299, same place, 15 June 2007, coll. Takahashi, H.; KPM-NC0017527, same place, 11 August 2010, coll. Takahashi, H. Known distribution: Japan (Okinawa). Japanese name: Midori-nisehouraitake. Comments: Crinipellis rhizomorphica has the characteristics of the brownish orange, fibrillose-squamulose basidiomata accompanied by the white thread-like rhizomorphs on the dead twig, the dextrinoid hairs turning olivaceous in KOH, the oblong-ellipsoid, relatively long basidiospores averaging 12 9 4.75 lm, the subclavate cheilocystidia with two or three short cylindrical apical appendages, and the absence of pleurocystidia. Its olivaceous-colored hairs in KOH suggest that the present fungus is best accommodated in the section Grisentinae (Singer) Singer (Singer 1976, 1986). Within the section Grisentinae, C. rhizomorphica has macromorphological similarities to the following four species: C. sapindacearum Singer from Brazil (Singer 1976); C. trichialis (Le´v.) Pat. ex Antonı´n R. Ryoo & H.D. Shin (Le´veille´ 1846; Saccardo 1887; Antonı´n et al. 2009), redescribed by Singer from Venezuela (Singer 1976) and by Kerekes and Desjardin from Indonesia and Malaysia (Kerekes and Desjardin 2009); C. tucumanensis Singer from Argentina (Singer 1976); and C. rhizomaticola

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Antonı´n from the Republic of Korea (Antonı´n et al. 2009). These taxa mainly differ from C. rhizomorphica in having well-developed pleurocystidia and lacking rhizomorphs. Furthermore, C. sapindacearum has much smaller basidiospores: 7.5–8.2 9 3–3.5 lm (Singer 1976) and a habit on dead fallen coriaceous leaves of Sapindaceae. Crinipellis trichialis produces shorter and broader basidiospores: (8.5–) 9.6–11.5 9 (5.5–)6–7(–7.4) lm (Kerekes and Desjardin 2009, Holotype: FH!). Crinipellis tucumanensis forms much shorter basidiospores: 5.5–8.5 lm long (Singer 1976). Crinipellis rhizomaticola is distinct in having a chestnutbrown, larger pileus: 12–22 mm in diameter (Antonı´n et al. 2009), and significantly shorter basidiospores: 8.5–10 lm long (Antonı´n et al. 2009). Crinipellis rhizomorphica also shares characteristics such as olivaceous-colored hairs in KOH, copious rhizomorphs, and a radially fibrillose-strigose pileus with a minute blackish papilla in umbilicus in common with the following two species: Southeast Asian C. actinophora (Berk. and Broome) Singer (Berkeley and Broome 1874; Singer 1955; Pegler, 1986; Corner 1996; Kerekes and Desjardin 2009); and C. nigricaulis Har. Takah. from Japan (Takahashi 2000) and Republic of Korea (Antonı´n et al. 2009). These two taxa, however, can be discerned from C. rhizomorphica by forming a dark brown stipe occasionally associating with the much longer, dark brown, ‘hair-blight’ rhizomorphs, significantly shorter basidiospores: 6–10 lm long (Kerekes and Desjardin 2009), and cheilocystidia with numerous apical appendages. 2. Chaetocalathus fragilis (Pat.) Singer, Lilloa 8: 520, 1943 Figs. 6–10 Basionym: Crinipellis fragilis Pat., Philipp. J. Sci., C, Bot. 10:97, 1915 : Marasmius fragilis (Pat.) Sacc. and Trotter, Syll. fung. (Abellini) 23:153, 1925 : Lachnella fragilis (Pat.) Locq., Bull. trimest. Soc. mycol. Fr. 68:166, 1952 Pileus (Fig. 6) (0.5–)1–2(–2.5) mm in diameter 9 0.5–1 mm tall, very small, membranous, astipitate and dorsally attached to a substratum, discoid-cyphelloid, with pilose margin; surface not hygrophanous, smooth, appressed fibrillose to wooly-tomentose, pure white overall. Flesh very thin (up to 0.2 mm), white. Odor and taste not distinctive. Rudiment of stipe 0.1–0.2 mm diameter, white, reduced to a central small papilla (columella), situated in the underside of the pileus and not directly attached to a substratum. Lamellae adnexed to almost free, subdistant (13–18) with 1–2 series of lamellulae, medium broad (0.2–0.6 mm), thin, white; edges even, concolorous. Basidiospores (Fig. 7) 8–8.5 9 6–7 lm (Q = length/ breadth: 1.2–1.3), subglobose to shortly ovoid-ellipsoid, smooth, colorless, inamyloid, thin-walled. Basidia 16–22 9

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Figs. 11–14 Basidiomata of Psilocybe capitulata (Holotype): Fig. 11 Primordia and immature basidiomata. Fig. 12 Immature basidioma. Fig. 13 Mature basidiomata. Fig. 14 Surface of the mature pileus. Bar 11 2 mm; 12 5 mm; 13, 14 20 mm. Color photos of Psilocybe capitulata can be seen at http://www7a.biglobe.ne. jp/*har-takah/page112.html

4–6 lm, clavate, 4-spored; basidioles subclavate to subfusoid. Cheilocystidia (Figs. 8–10) 15–23 9 6–12 lm, gregarious, forming a compact sterile edge, projecting from the hymenium, metuloid-type, clavate to subclavate, usually with one or two cylindrical apical appendages

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5–11 lm long, with an obtuse apex, smooth, colorless, hyaline in KOH, with strongly dextrinoid, moderately thickened walls (up to 1 lm thick), usually in the upper portion encrusted with coarse hyaline crystals. Pleurocystidia scattered, similar to cheilocystidia. Hyphae of

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hymenophoral trama 2–9 lm wide, entangled, cylindrical, smooth, colorless, inamyloid, thin-walled, occasionally with clamped septa. Pileipellis a hypotrichial layer of subcylindrical cells 3–6 lm wide, with smooth, colorless walls up to 1 lm thick, weakly dextrinoid, occasionally with clamped septa; hairs of pileus 200–300 9 2–5 lm, arising directly from the hypotrichium, densely entangled, cylindrical, with a subacute apex, sometimes flexuous, hyaline in KOH, with smooth, hyaline walls 1–2 lm thick, strongly dextrinoid, without a secondary septum. Hyphae of pileitrama similar to those of the hymenophoral trama.

subaequali vel ad basim leniter incrassato, centrali, cavo, albo vel brunneolo, fibrilloso vel squamuloso; annuli tenuis, albis; mycelio basali albo affixo; lamellis adnexis, brunneis; basidiosporis (13–)13.5–15(–19) 9 8–9(–9.5) lm, ellipsoideis vel subhexagonoideus, levibus, rubro-brunneis, crassitunicatis; basidiis bisporis vel tetrasporis; cheilocystidiis 24–35 9 3–9 lm, abundantibus, fusoideo-ventricosis; pleurocystidiis nullis vel infrequens, 28–39 9 10–15 lm, basidioformibus vel late clavatis,; pilocystidiis 40–60 9 2–13 lm, clavato-cylindraceis, capituliformibus; hyphis fibulatis.

Habitat: Usually abundant, on dead twigs (unidentified substrata), June to August.

Holotypus: Ad fimum bovis, Ishigaki-shi, Okinawa Pref., Japonia, 9 Feb. 2002, H. Takahashi (KPM-NC0017300). Etymology: from Latin, capitulata = capitulate, referring to the capitulate pilocystidia.

Specimens examined: KPM-NC0017520, Banna-dake, Ishigaki-shi, Okinawa Pref., on dead twigs, 25 July 2010, coll. Takahashi, H. and Terashima, Y.; KPM-NC0017521, same place, 28 July 2010, coll. Takahashi, H.; KPMNC0017522, same place, 11 Aug. 2010, coll. Takahashi, H.; KPM-NC0017523, same place, 6 Jun. 2009, coll. Takahashi, H.; KPM-NC0017524, same place, 8 Jun. 2009, coll. Takahashi, H.; KPM-NC0017525, same place, 15 Jun. 2009, coll. Takahashi, H. Known distribution: Japan (Okinawa). Japanese name: Hida-fuurintake. Comments: The diagnostic features of this species are the minute (1–2 mm in diameter on average), white, membranous, discoid-cyphelloid basidiomata having a central, small, papillate columella, the subglobose to shortly ovoidellipsoid basidiospores, the dextrinoid hymenial metuloids with an obtuse apex, the strongly dextrinoid hairs covering the pileus surface, and the basidiome formation on the dead twig. The Japanese material fully conforms to C. fragilis originally described from the Philippines (Patouillard 1915; Saccardo and Trotter 1925; Singer 1943; Locquin 1952). At the collection locality in Japan (Ishigaki-shi, Okinawa Pref.), the species commonly occurs on dead twigs in the mixed forest of Q. miyagii and C. sieboldii in summer. The present species seems to be distributed widely in eastern Asia and the Pacific region. 3. Psilocybe capitulata Har. Takah., sp. nov. Figs. 11–18 MycoBank no.: MB 519033 Primordio 1–2 mm lato, ovoideis vel oblongo-ellipsoideis, cum flocculo albo obvolvato; pileo 42–93 mm lato, primo hemispherico vel conico-campanulato, dein late convexo vel applanato, saepe umbonato, furfuraceo-squamuloso, subviscido, hygrophano, brunneo; carne cyanescente; odore saporeque nullo; stipite 48–63 9 5–20 mm,

Primordium (Fig. 11) 1–2 mm in diameter, ovoid to oblong-ellipsoid, enveloped in white, floccose universal veil. Pileus (Figs. 12–14) 42–93 mm in diameter, at first hemispherical to conic-campanulate, expanding to broadly convex to almost plane, sometimes broadly umbonate, with straight margin, not appendiculate; surface at first spotted with fugacious, white remnants of the veil, soon glabrescent, eventually covered overall with brownish, furfuraceous squamules in age, subviscid when wet, hygrophanous, brownish orange (7C6–7) to brown (7D6–7) when moist, mottled with darker areas (near brownish red: 8C6–7) at the center, drying to paler (6C4–5) or whitish from the margin, gradually changing to blue where handled. Flesh up to 9 mm, whitish, gradually changing to blue when cut; odor and taste not distinctive. Stipe 48–63 9 5–20 mm, subequal or at times somewhat thickening toward the base, central, terete, hollow; surface fibrillose above, squamulose below, sometimes striate above the annulus, whitish to pale brownish, gradually changing to blue where handled; annulus 8–15 mm wide, thin, membranous, persistent, fragile, white, striate; base covered with white strigose mycelial hairs. Lamellae adnate to adnexed, close (42–51 reach the stipe) with 1–3 series of lamellulae, up to 12 mm broad, dark-brown (8F6–8) to reddish-brown (9F7–8); edges subfloccose, greyish. Basidiospores (Fig. 15) (13–)13.5–15(–19) 9 8–9 (–9.5) lm (Q = length/breadth: 1.66-1.68), subellipsoid in side view, subhexagonal in face view, smooth, greyish red (10D5) to brownish red (10D6) in H2O, orange–yellow (4A7–4B7) in KOH, thick-walled (0.5–1 lm), truncated at the apex, with a distinct germ pore. Basidia 25–35 9 10–12 lm, clavate, 2- to 4-spored. Cheilocystidia (Fig. 16) 24–35 9 3–9 lm, abundant, forming a compact sterile edge, fusoid-ventricose with a subcapitate apex, smooth, hyaline in KOH, thin-walled. Pleurocystidia (Fig. 17) none or infrequent, 28–39 9 10–15 lm, basidiomorphous, broadly clavate, with a rounded apex, sometimes with a

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Figs. 15–18 Micromorphological features of Psilocybe capitulata (Holotype). Fig. 15 Basidiospores mounted in KOH. Fig. 16 Cheilocystidia. Fig. 17 Pleurocystidia. Fig. 18 Pilocystidia. Bar 15 15 lm; 0.16 10 lm; 17 15 lm; 18 13 lm

slight median constriction, smooth, hyaline in KOH, thinwalled. Hyphae of hymenophoral trama 5–22 lm wide, subcylindrical to fusoid, often inflated, parallel, smooth,

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hyaline in KOH, thin-walled. Pileipellis a loose cutis of entangled, gelatinous, subcylindrical hyphae 2–10 lm wide, encrusted with orange (6A7–6B7) pigment in

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KOH; pilocystidia (Fig. 18) 40–60 9 2–13 lm, clavatecylindrical, often with a capitulate apex, smooth, hyaline in KOH, thin-walled. Squamules of the pileus similar to the pileipellis. Hyphae of pileitrama 5–10 lm wide, parallel, subcylindrical, not inflated, smooth, hyaline in KOH, thinwalled. Stipitipellis a loose cutis of entangled, non-gelatinous, cylindrical hyphae 2–5 lm wide, smooth, hyaline in KOH, thin-walled, lacking differentiated terminal cells. Stipe trama composed of longitudinally running, cylindrical cells 5–12 lm wide, smooth, hyaline in KOH, thinwalled. Elements of annulus 2–3 lm wide, loosely interwoven, subcylindrical, not inflated, smooth, hyaline in KOH, thin-walled. Clamp connections present in all tissues. Habitat: Solitary to scattered, coprophilous on cow dung, almost year-round, common. Specimens examined: KPM-NC0017300, Ishigaki-shi, Okinawa Pref., on cow dung, 9 Feb. 2002, coll. Takahashi, H., Uehara, S., and Sakamoto, H.; KPM-NC0017301, same place, 2 March 2002, coll. Takahashi, H.; KPMNC0017302, same place, 26 Feb. 2008, coll. Takahashi, H. & Sakamoto, H.; KPM-NC0017303, same place, 28 Feb. 2008, coll. Takahashi, H.; KPM-NC0017304, same place, 3 March 2008, coll. Takahashi, H.; KPM-NC0017305, same place, 2 June 2008, coll. Takahashi, H.; KPM-NC0017306, the same place, 14 June 2008, coll. Takahashi, H.; KPMNC0017307, same place, 2 Feb. 2009, coll. Takahashi, H.; KPM-NC0017308, same place, 9 March 2010, coll. Takahashi, H. Known distribution: Japan (Okinawa). Japanese name: Nanyou-sibiretake. Comments: Psilocybe capitulata is well characterized by the brownish orange to brown pileus covered overall with brownish, furfuraceous squamules in age, the cyanescent flesh, the persistent, white, membranous annulus, the capitulate pilocystidia, and the coprophilous habit on the cow dung. Its cyanescent basidiomata on cow dung, the subhexagonal, thick-walled basidiospores, and the welldeveloped, persistent annulus indicate alignment of the present fungus with the section Cubensae Guzma´n (Guzma´n 1983). Because of its typically furfuraceous-squamulose pileus in age, the capitulate pilocystidia, and the infrequent, less-developed pleurocystidia, P. capitulata can be distinguished from the other previously known taxa of the section Cubensae such as pantropical P. cubensis (Earle) Singer (Earle 1906; Guzma´n 1978, 1983; Pegler 1983; Stamets 1996; Thomas et al. 2002; Cortez and Coelho 2004) and P. subcubensis Guzma´n from Mexico (Guzma´n 1978, 1983). Apart from the section Cubensae, P. subaeruginascens Ho¨hn. from Indonesia (Ho¨hnel 1914; Guzma´n 1983; Horak

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and Desjardin 2006) and Japan (Nagasawa 1987) bears a superficial resemblance to P. capitulata, although it is distinct in possessing rhomboid basidiospores, producing well-developed, broadly fusoid-ventricose pleurocystidia, and lacking pilocystidia. Psilocybe subannulata E. Horak and Guzma´n from Puerto Rico (Guzma´n et al. 2009) is also similar to P. capitulata in appearance. The former is easily separated from P. capitulata in having rhombic basidiospores and lacking pilocystidia. The present species may possibly be related to P. magnispora E. Horak, Guzma´n and Desjardin from Thailand (Horak et al. 2009) in having a submembranous annulus, cyanescent flesh, and a coprophilous habit. Psilocybe magnispora, however, is differentiated from P. capitulata in possessing rhomboid basidiospores, forming well-developed pleurocystidia with refrigent incrustations, and lacking pilocystidia. Acknowledgments I am grateful to Mrs. Kanade Otsubo (KPM) for allowing the specimens cited to be kept in the Kanagawa Prefectural Museum of Natural History.

References Antonı´n V, Ryoo R, Shin HD (2009) Marasmioid and gymnopoid fungi of the Republic of Korea. 1. Three interesting species of Crinipellis (Basidiomycota, Marasmiaceae). Mycotaxon 108:429–440 Berkeley MJ, Broome CE (1874) Enumeration of the fungi of Ceylon. Part II. J Linn Soc Bot 14:29–141 Corner EJH (1996) The agaric genera Marasmius, Chaetocalathus, Crinipellis, Heimiomyces, Resupinatus, Xerula, and Xerulina in Malesia. Nova Hedwigia Beih 111:1–164 Cortez VG, Coelho G (2004) The Stropharioideae (Strophariaceae, Agaricales) from Santa Maria, Rio Grande do Sul, Brazil. Mycotaxon 89:355–378 Earle FS (1906) Algunos hongos cubanos. Inf An Estac Centr Agron Cuba 1:225–242 Guzma´n G (1978) The species of Psilocybe known from Central and South America. Mycotaxon 7:225–255 Guzma´n G (1983) The genus Psilocybe, a systematic revision of the known species including the history, distribution and chemistry of the hallucinogenic species. Nova Hedwigia Beih 74:1–439 Guzma´n G, Horak E, Halling R, Ramı´rez-Guille´n F (2009) Further studies on Psilocybe from the Caribbean, Central America and South America, with descriptions of new species and remarks to new records. Sydowia 61(2):215–242 Ho¨hnel FXR (1914) Fragmente zur Mykologie, XVI. Mitteilung, Nr. 813–875. Sitzungsber Kais Akad Wiss Wien Math Naturwiss Kl 123:49–155 Horak E, Desjardin DE (2006) Agaricales of Indonesia. 6. Psilocybe (Strophariaceae) from Indonesia (Java, Bali, Lombok). Sydowia 58(1):15–37 Horak E, Guzma´n G, Desjardin DE (2009) Four new species of Psilocybe from Malaysia and Thailand, with a key to the species of sect. Neocaledonicae and discussion on the distribution of the tropical and temperate species. Sydowia 61(1):25–37 Kerekes J, Desjardin DE (2009) A monograph of the genera Crinipellis and Moniliophthora from Southeast Asia including a molecular phylogeny of the nrITS region. Fungal Divers 37:101–152

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400 Kornerup A, Wanscher JH (1983) Methuen handbook of colour. Eyre Methuen, London Le´veille´ JH (1846) Descriptions des champignons de l’herbier du Muse´um de Paris. Ann Sci Nat Bot, se´r 3 5:111–167 Locquin M (1952) Sur la non-validite´ de quelques genres d’Agaricales. Bull Trimest Soc Mycol Fr 68:165–169 Miyagi G (1958) On the fungi of the Ryukyu Islands, vol 2 (in Japanese). Bulletin of Arts and Science Division, University of the Ryukyus, Mathematics and sciences, pp 35–40 Miyagi G (1964) On a luminous fungus, Pleurotus lunaillustris from the Yaeyama Islands, vol 7 (in Japanese). Bulletin of Arts and Science Division, University of the Ryukyus, Mathematics and sciences, pp 54–56 Miyagi G (1971) Notes on the Agaricales of Iriomote Island and Ishigaki Island (1) (in Japanese). Biol Mag Okinawa 7:33–37 Nagasawa E (1987) Strophariaceae. In: Imazeki R, Hongo T (eds) Colored illustrations of mushrooms of Japan I (in Japanese). Hoikusha, Osaka, pp 190–211 Patouillard NT (1915) Champignons des Philippines communique´s par C.F.Baker, II. Philipp J Sci C Bot 10:85–98 Pegler DN (1983) Agaric flora of the Lesser Antilles. Kew Bulletin Add Ser IX. HMSO, London

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Mycoscience (2011) 52:392–400 Pegler DN (1986) Agaric flora of Sri Lanka. Kew Bulletin Add Ser XII. HMSO, London Saccardo PA (1887) Sylloge Hymenomycetum, Vol. I. Agaricineae. Syll Fung 5:1–1146 Saccardo PA, Trotter A (1925) Supplementum Universale, Pars X. Basidiomycetae. Syll Fung (Abellini) 23:1–1026 Singer R (1943) A monographic studies of the genera of Crinipellis and Chaetocalathus. Lilloa 8:441–534 Singer R (1955) Type studies on Basidiomycetes VIII. Sydowia 9:367–431 Singer R (1976) Marasmieae (Basidiomycetes—Tricholomataceae). Fl Neotrop Monogr 17:1–347 Singer R (1986) The Agaricales in modern taxonomy, 4th edn. Koeltz, Koenigstein Stamets P (1996) Psilocybin mushrooms of the world. Ten Speed Press, Berkeley Takahashi H (2000) Three new species of Crinipellis found in Iriomote Island, southwestern Japan, and central Honshu, Japan. Mycoscience 41:171–182 Thomas KA, Manimohan P, Guzma´n G, Tapia F, Ramı´rez-Guille´n F (2002) The genus Psilocybe in Kerala State, India. Mycotaxon 83:195–207

Mycoscience (2011) 52:401–412 DOI 10.1007/s10267-011-0126-3

FULL PAPER

Taxonomic revision of Lophiostoma and Lophiotrema based on reevaluation of morphological characters and molecular analyses Kazuyuki Hirayama • Kazuaki Tanaka

Received: 29 March 2011 / Accepted: 23 May 2011 / Published online: 11 June 2011 ! The Mycological Society of Japan and Springer 2011

Abstract Lophiostoma and Lophiotrema share several morphological and ecological features. They have been regarded as closely related genera within the family Lophiostomataceae, but their morphological circumscriptions have been uncertain. To clarify the generic definitions of Lophiostoma and Lophiotrema, we conducted phylogenetic analyses of 29 isolates of these genera based on the SSU and LSU nrDNA sequences, and also reevaluated several key characters previously used for their generic characterization. Our results clearly confirmed that Lophiostoma and Lophiotrema are distinct genera belonging to different families; the ascus shape, including length of the ascus stipe, is a reliable taxonomic indicator to allow discrimination between the genera. In Lophiostoma species, asci are clavate with relatively long stipes [mostly (10–) 15–30 lm in length], whereas in Lophiotrema the asci are cylindrical with short stipes (up to 15 lm long). A new family, Lophiotremataceae, is proposed to accommodate species in the Lophiotrema clade that was distantly placed from the Lophiostomataceae within the Pleosporales. Lophiostoma quadrisporum, collected from twigs of Liriodendron tulipifera, is described as a new species with distinctive 4-spored asci. Lophiotrema vitigenum, which has clavate asci with long stipes, is transferred to Lophiostoma. Keywords Ascomycota ! Dothideomycetes ! Lophiotremataceae ! Pleosporales ! Systematics K. Hirayama ! K. Tanaka (&) Faculty of Agriculture and Life Science, Hirosaki University, 3 Bunkyo-cho, Hirosaki, Aomori 036-8561, Japan e-mail: [email protected] K. Hirayama The United Graduate School of Agricultural Sciences, Iwate University, 18-8 Ueda 3 chome, Morioka 020-8550, Japan

Introduction Lophiostoma Ces. & De Not. and Lophiotrema Sacc. are bitunicate ascomycetes in the Pleosporales, Dothideomycetes. Most species within these genera occur mainly on the twigs or bark of various woody plants (Holm and Holm 1988; Tanaka et al. 2010a). Some species, however, are frequently found on the culms of herbaceous plants, such as reeds (Tanaka and Harada 2003a), palms (Hyde et al. 2000), and bamboos (Cai et al. 2003). These species are considered as saprobes on the foregoing substrates in terrestrial (Holm and Holm 1988), freshwater (Hyde and Aptroot 1998), and marine environments (Hyde et al. 1992). The two genera share several morphological features (Figs. 1–27), such as carbonaceous ascomata with a laterally compressed apex (termed as a crest-like beak with a slit-like ostiole; Figs. 1–4, 19, 20), fissitunicate asci (Figs. 11–14, 24, 25), and hyaline to dark brown, one- to multiseptate ascospores (Figs. 15–18, 26, 27) (Holm and Holm 1988). These two genera have, therefore, been regarded as closely related genera within the Lophiostomataceae (Barr 1992). Lophiostoma and Lophiotrema have been distinguished based on Saccardo’s sporological principles: ‘‘phaeophragmiae’’ in Lophiostoma and ‘‘hyalophragmiae’’ in Lophiotrema (Saccardo 1878). Chesters and Bell (1970a), however, synonymized Lophiotrema under Lophiostoma because they considered that neither ascospore color nor number of transverse septa could be used for generic delimitation. Although this opinion was accepted by Leuchtmann (1985), these were reestablished as separate genera by Holm and Holm (1988), based primarily on peridial structure of ascomata and ascus shape. Namely, Lophiostoma has an ascomatal wall that is broader laterally at the base (*50 lm) and composed of parallel, long,

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b Figs. 1–27 Morphological features of representative species of Lophiostoma (1–18) and Lophiotrema (19–27). 1–4, 19, 20 Ascomata erumpent or immersed on host surface. 5, 6, 21 Longitudinal sections through ascomata. 7–10, 22, 23 Sections through ascomatal walls, composed of parallel rows of rectangular cells (7, 8), rectangular to prismatic cells (9, 10, 22) and prismatic cells (23). 11–14 Asci clavate with a long stipe. 15 Ascospore hyaline, 1-septate, with terminal short appendages (arrowheads). 16 Ascospore pigmented, 9-septate, with terminal long appendages (arrowheads). 17 Ascospore hyaline, 1-septate, with terminal long appendages (arrowheads). 18 Ascospore hyaline, 1-septate, without appendage or sheath. 24, 25 Asci cylindrical with a short stipe. 26 Ascospore hyaline, 3-septate, with an entire sheath (arrowheads). 27 Ascospore hyaline, 1-septate, with an entire sheath (arrowheads). 1–18 Lophiostoma species: 1, 7, 11, 15 from L. macrostomum (1, 15 = HHUF 27290; 7, 11 = HHUF 27293); 2, 8, 12 from L. arundinis (2 = HHUF 27305, 8 = HHUF 27413, 12 = HHUF 27304); 3 from L. fuckelii (HHUF 27325); 4, 6, 10, 14, 18 from L. quadrisporum (HHUF 27321); 5 from L. caulium ‘‘var. d’’ (HHUF 27310); 9 from L. sagittiforme (HHUF 29754); 13 from L. caudatum (HHUF 27319); 16 from L. caulium ‘‘var. f’’ (HHUF 27313); 17 from L. vitigenum (HHUF 26930). 19–27 Lophiotrema species: 19, 21, 23, 24, 26 from L. neohysterioides (19, 24, 26 = HHUF 27328; 21, 23 = HHUF 27331); 20, 22 from L. neoarundinaria (HHUF 27547); 25, 27 from L. vagabundum (HHUF 27323). Bars 1–4, 19, 20 200 lm; 5, 6, 21 100 lm; 7–10, 22, 23 20 lm; 11–18, 24–27 10 lm

prismatic cells, and it has clavate asci. In contrast, Lophiotrema has an ascomatal wall of entirely equal thickness (*25 lm) composed of textura angularis to globosa, and it has cylindrical asci. These generic circumscriptions have been followed by later authors (Barr 1992; Mathiassen 1993; Yuan and Zhao 1994; Tanaka and Harada 2003a,b; Tanaka and Hosoya 2008; Eriksson 2009). Taxonomic revision of Lophiostoma and Lophiotrema has been carried out on morphological grounds (e.g., Lehmann 1886; Berlese 1894; Chesters and Bell 1970a). Recent molecular analyses have revealed phylogenetic relationships and species validities of the lophiostomatoid fungi in the Pleosporales (Schoch et al. 2006, 2009; Tanaka and Hosoya 2008; Mugambi and Huhndorf 2009a). Several unrelated species without any compressed crest-like beak on their ascomata, previously described as Lophiostoma, have recently been excluded from the genus. For example, L. breviappendiculatum Kaz. Tanaka et al. (Tanaka et al. 2005) and L. ingoldianum (Shearer & K.D. Hyde) Aptroot & K.D. Hyde (Shearer and Hyde 1997; Hyde et al. 2002), occurring in freshwater habitats, were transferred to Lindgomyces K. Hiray. et al. based on analyses of the small and large subunit nuclear ribosomal DNA (SSU and LSU nrDNA) and morphological reevaluation (Shearer et al. 2009; Hirayama et al. 2010). Lophiostoma mangrovei Kohlm. & Vittal, found on marine mangroves (Kohlmeyer and Vittal 1986), was treated as a species in Rimora Kohlm. et al. based on analyses of four genes (Suetrong et al. 2009). In general, typical species in accordance with the generic concept of Lophiostoma (Holm and Holm 1988) appear to represent a natural group derived from a

403

single ancestor. On the other hand, taxonomic circumscription of the genus Lophiotrema would be problematic. Zhang et al. (2009b) clearly indicated that Lophiostoma and Lophiotrema are phylogenetically distinct genera based on molecular study. However, they considered that morphological criteria, particularly the peridial structure of ascomata formerly used to separate these lophiostomatoid genera, are unable to provide differentiation between Lophiostoma and Lophiotrema because the ascomata within these genera are almost identical (Zhang et al. 2009b). Then, they described two new species of Lophiotrema based on their close phylogenetic relationship to the type of Lophiotrema (L. nucula Rehm) (Zhang et al. 2009b), but morphological differentiation between Lophiostoma and Lophiotrema has consequently remained unclear. The phylogenetic study of lophiostomatoid genera conducted by Zhang et al. (2009b) further revealed that Lophiotrema is not a member of Lophiostomataceae; this observation contrasted with the traditional classification of the genus (Saccardo 1883; Clements and Shear 1931; Barr 1992). Subsequently, on the basis of further molecular analyses using five DNA regions [SSU and LSU nrDNA, the translation elongation factor-1 alpha (TEF1), and the largest and second largest subunits of RNA polymerase II (RPB1 and RPB2)], Zhang et al. (2009a) found that two species previously placed in Lophiostoma should be transferred to Lophiotrema, and that the monophyletic clade of Lophiotrema is related to the Testudinaceae rather than the Lophiostomataceae. Familial placement of Lophiotrema, however, remained uncertain, mostly because of the lack of a morphological circumscription of the genus. In this study, we carried out phylogenetic analyses of Lophiostoma and Lophiotrema based on SSU and LSU nrDNA sequences, using 29 isolates from these genera. The taxonomic significance of several key characters previously used for the morphological delimitation of these genera was reevaluated. Our purpose was to clarify the morphological circumscriptions of Lophiostoma and Lophiotrema and to reveal the familial placement of Lophiotrema.

Materials and methods Morphological studies and fungal isolates Specimens of Lophiostoma and Lophiotrema from the herbarium of Hirosaki University (HHUF) (Table 1) were used for microscopic observation following the method described by Hirayama et al. (2010). Special attention was given to key characters used in the delimitation of

123

123 – HHUF 30077 HHUF 30078 HHUF 27323 –

KH 172 KT 664 –

HHUF 27330

KT 756 – KH 164

HHUF 27328

KT 713

HHUF 30014 HHUF 27331

KT 2200 KT 686-2

HHUF 27547 HHUF 30015

HHUF 27316

KT 764 KT 856 KT 1034

HHUF 27317

KT 740

HHUF 26931

HH 26931

JPN Japan, SWE Sweden

The sequences determined in this study are in bold

Lophiotrema nucula Lophiotrema vagabundum

Lophiotrema neohysterioides

Lophiotrema neoarundinaria

Lophiostoma winteri

HHUF 26930

HHUF 27300

KT 828 HH 26930

HHUF 27298

KT 652

Lophiostoma vitigenum

HHUF 29754 HHUF 27299

HHUF 27321

KT 1934 KT 622

HHUF 27293

KT 709 KT 843

HHUF 27290

KT 635

Lophiostoma sagittiforme Lophiostoma semiliberum

HHUF 27288

HHUF 27325

KT 508

HHUF 30076

KT 634

HHUF 27315 HHUF 27311

KT 686-1 KT 794 KH 161

HHUF 27313

KT 573

HHUF 27310

KT 777

Lophiostoma quadrisporum

Lophiostoma macrostomum

Lophiostoma fuckelii

Lophiostoma caulium ‘‘var. f’’

HHUF 27309

KT 604

HHUF 27307

KT 633

Lophiostoma caulium ‘‘var. d’’

HHUF 27306

HHUF 27319

KT 530 KT 603

Lophiostoma caulium ‘‘var. a’’

HHUF 27305 HHUF 27413

KT 651 KT 668

Lophiostoma caudatum

HHUF 27304

KT 606

Lophiostoma arundinis

Specimen no.

Original no.

Species name

CBS 113975/JCM 14138

JCM 17675 MAFF 239456

CBS 113826/JCM 14132 JCM 17674

MAFF 239457

JCM 17673

–

NBRC 106238

MAFF 239461 NBRC 106239

MAFF 239454

JCM 17648

JCM 17676

JCM 13534/MAFF 239459

JCM 13549/MAFF 239448

JCM 13547

JCM 15100 JCM 13548

MAFF 239455

JCM 13546/MAFF 239447

JCM 13545

JCM 13544

MAFF 239458

JCM 17672

JCM 17670 JCM 17671

MAFF 239452

MAFF 239451

JCM 17668

JCM 17669

MAFF 239450

MAFF 239453

JCM 13551/MAFF 239449 –

JCM 13550

Culture collection no.

Uppland, SWE

Hokkaido, JPN Aomori, JPN

Uppland, SWE Hokkaido, JPN

Hokkaido, JPN

Aomori, JPN

Aomori, JPN

Kagoshima, JPN

Nagano, JPN Tochigi, JPN

Hokkaido, JPN

Hokkaido, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Kagoshima, JPN Aomori, JPN

Iwate, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Hokkaido, JPN

Aomori, JPN Aomori, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN

Aomori, JPN Aomori, JPN

Aomori, JPN

Collection site

Table 1 Specimens, isolates, and GenBank accession numbers of Lophiostoma and Lophiotrema used in this study

AB433273

AB619014

AB618696

AB619022

AB618704 AB618705 AB618706 AB618707

Vitis coignetiae Woody plant Epilobium angustifolium

AB619025

AB619023 AB619024

AB619020 AB619021

AB618702 AB618703 Fraxinus excelsior Vitis coignetiae

Woody plant

AB619019

AB618701

Robinia pseudoacacia

–

AB524597 –

AB524456

Phyllostachys bambusoides

AB524596 AB524598

AB619018

AB619017

AB619016

Woody plant

AB524455 AB524457

AB618700

Polygonum sp. Phyllostachys bambusoides Phyllostachys bambusoides

AB618699

AB618698

Vitis coignetiae Unknown plant

AB618697

Vitis coignetiae

AB619015

AB619013

AB618695

Phragmites australis Herbaceous plant

AB369267 AB619012

AB619011

AB433274 AB618693 AB618694

AB618692

AB521732

AB521731

AB619010

AB619009 AB618691

AB619008 AB618690

AB619006 AB619007 AB618689

AB619005 AB618687 AB618688

AB619004 AB618686

AB619003 AB618685

AB619002

AB618683 AB618684

AB619001

AB618682

AB619000

AB618999 –

AB618998

Machilus japonica Harbaceous plant

Liriodendron tulipifera

Unknown plant

Herbaceous plant

Morus bombycis

Unknown plant

Vitis coignetiae

Woody plant Dactylis glomerata

Herbaceous plant

Herbaceous plant

Herbaceous plant

Herbaceous plant

Herbaceous plant

AB618681

AB618680 –

Dactylis glomerata

AB618679

Phragmites australis Phragmites australis

GenBank no.

Phragmites australis

Substrate

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Lophiostoma and Lophiotrema, including size and peridial structure of ascomata, ascus shape, and ascospore morphology. Fungal cultures used were deposited in the Japan Collection of Microorganisms (JCM), the National Institute of Agrobiological Sciences (MAFF), and the NITE Biological Resource Center (NBRC) (Table 1). Phylogenetic analyses DNA was extracted from a total of 29 isolates, including two strains obtained from the Centraalbureau voor Schimmelcultures (CBS) (see Table 1). Approximately 1,300 nucleotides at the 50 -end of the partial SSU and LSU nrDNA were amplified by polymerase chain reaction (PCR) using the primer pairs NS1–NS4 for the SSU (White et al. 1990) and LROR–LR7 for the LSU (Rehner and Samuels 1994). Methods of DNA extraction and PCR amplification have been described by Hirayama et al. (2010). The SSU and LSU sequences of Lophiostoma and Lophiotrema species were aligned alongside those of related species from GenBank (Table 2). Sequences of Dothidea insculpta Wallr., an outgroup taxon, were used to root trees. Preliminary multiple alignment of sequences was conducted using MEGA 4 (Tamura et al. 2007). Gaps and ambiguous regions were excluded from analyses. The aligned dataset was subjected to three phylogenetic analyses: maximum parsimony (MP) using a close-neighbor-interchange heuristic search with an initial tree obtained by random addition sequence (100 replicates), neighbor-joining (NJ) analysis based on the Kimura two-parameter model, and Bayesian analyses using MrBayes version 3.1.2 (Ronquist and Huelsenbeck 2003). The final alignment was deposited in TreeBASE (http://www.treebase.org). Bootstrap values (BV) for MP and NJ analyses were computed from 1,000 replicates. MrModeltest version 2.3 (Nylander 2004), in conjunction with PAUP version 4.0b10 (Swofford 2003), were used to select substitution models for Bayesian analyses. On the basis of the Akaike information criterion, a general time-reversible, invariant, g-distributed (GTR ? I ? G) model was applied. Two runs with ten chains of Markov Chain Monte Carlo (MCMC) iterations were performed for 5 million generations, keeping 1 tree every 100 generations. Runs were deemed to have converged if the mean standard deviation of split frequencies became less than 0.01. The first 0.8 million generations of the dataset were discarded as burn-in, and the remaining 42,000 trees were used to calculate 50% majority rule trees and to determine Bayesian posterior probabilities (BPP) for individual branches.

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Results Analyses of combined SSU and LSU nrDNA sequences A combined alignment of the SSU (884 bp) and LSU (723 bp) regions consisting of 74 strains was generated. SSU region insertions found in Delitschia didyma Auersw. (512–808, 1247–1591) and Neottiosporina paspali (G.F. Atk.) B. Sutton & Alcorn (487–841) were excluded from the alignment. Of 1,607 characters, 425 (26.4%) were variable, and of these 306 (19.0%) were parsimony informative. A MP analysis of the dataset resulted in 51 equally parsimonious trees with a length of 1,212 steps (consistency index = 0.461, retention index = 0.797). The trees obtained from NJ and Bayesian analyses were topologically similar to the MP tree. One of the 51 MP trees is shown in Fig. 28. Lophiostoma and Lophiotrema formed distinct monophyletic clades (Fig. 28). All Lophiostoma species and Lophiotrema vitigenum Kaz. Tanaka & Y. Harada (HH 26930 and 26931) grouped with Lophiostoma macrostomum (Tode) Ces. & De Not. (the type species of the genus Lophiostoma) in a strongly supported clade (99% BV, 1.00 BPP). A new species, Lophiostoma quadrisporum, and L. fuckelii Sacc. were sister to all other taxa in the Lophiostoma clade. Species in Lophiotrema, including the type species of the genus (L. nucula), clustered in a well-supported lineage (91–97% BV, 1.00 BPP) in a basal position of a main pleosporalean clade composed of the Lophiostomataceae and several other families (Fig. 28).

Taxonomy Several characters, such as size and peridial structure of ascomata, shape and stipe length of asci, and ascospore morphology, were examined (Figs. 1–27) and are shown on our tree (Fig. 28). The shape and stipe length of asci appear to have diagnostic value for the separation of Lophiostoma and Lophiotrema. A new family, Lophiotremataceae, is proposed to accommodate the genus Lophiotrema, based on morphological and molecular evidence. One new species and one new combination within Lophiostoma are described below. Their detailed descriptions and illustrations are found in Tanaka and Harada (2003b). Lophiotremataceae K. Hiray. & Kaz. Tanaka, fam. nov. MycoBank no.:

MB 561063

Ascomata subglobosa vel globosa. Rostrum compressum, cum ostiolo rimiformi. Pseudoparaphyses copiosae, septatae, ramificantes et anastomosantes. Asci fissitunicati, cylindrici, brevistipitati vel sessiles. Ascosporae fusiformes

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406 Table 2 Additional sequences obtained from Genbank

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Species

Strain

GenBank accession no. SSU

LSU

Dothideomycetes Amniculicola immersa

CBS 123083

GU456295

FJ795498

Amniculicola parva

CBS 123092

GU296134

GU301797

Arthopyrenia salicis

CBS 368.94

AY538333

AY538339

Ascochyta fabae

CBS 524.77

EU754034

EU754133

Delitschia didyma

UME 31411

AF242264

DQ384090

Delitschia winteri

CBS 225.62

DQ678026

DQ678077

Didymella exigua

CBS 183.55

EU754056

EU754155

Helicascus nypae

BCC 36751

GU479754

GU479788

Lentithecium arundinaceum

CBS 619.86

GU296157

DQ813509

Lentithecium fluviatile Leptosphaeria doliolum

CBS 122367 CBS 505.75

GU296158 GU296159

FJ795451 GU301827

Leptosphaeria maculans

DAOM 2229267

DQ470993

DQ470946

Lindgomyces cinctosporae

Raja R56-1

AB522430

AB522431

Lindgomyces ingoldianus

ATCC 200398

AB521719

AB521736

Lophiostoma compressum

IFRD 2014

FJ795480

FJ795437

Lophiostoma crenatum

CBS 629.86

DQ678017

DQ678069

Lophiostoma heterosporum

CBS 644.86

AY016354

AY016369

Lophiostoma scabridisporum 1

BCC 22835

GQ925831

GQ925844

Lophiostoma scabridisporum 2

BCC 22836

GQ925832

GQ925845

Lophiotrema lignicola

CBS 122364

FJ795488

FJ795445

Massaria inquinans

M 19

HQ599444

HQ599402

Massaria platanoidea

M7

HQ599457

HQ599420

Massarina eburnea

JCM 14422

AB521718

AB521735

Montagnula opulenta

CBS 168.34

AF164370

DQ678086

Morosphaeria ramunculicola Neotestudina rosatii

JK 5304B CBS 690.82

GU479760 DQ384069

GU479794 DQ384107

Neottiosporina paspali

CBS 331.37

EU754073

EU754172

Phaeodothis winteri

CBS 182.58

GU296183

GU301857

Phaeosphaeria avenaria

CBS 602.86

AY544725

AY544684

Phaeosphaeria juncophila

CBS 575.86

GU456307

GU456328

Pleospora herbarum

CBS 714.68

DQ767648

DQ678049

Preussia terricola

DAOM 230091

AY544726

AY544686

Pseudotetraploa curviappendiculata

JCM 12852

AB524467

AB524608

Setosphaeria monoceras

CBS 154.26

AY016352

AY016368

Triplosphaeria maxima

JCM 13172

AB524496

AB524637

Ulospora bilgramii

CBS 110020

DQ384071

DQ384108

Westerdykella cylindrica

CBS 454.72

AY016355

AY004343

Botryosphaeria dothidea

CBS 115476

DQ677998

DQ678051

Dothidea insculpta

CBS 189.58

DQ247810

DQ247802

Spencermartinsia viticola

CBS 117009

DQ678036

DQ678087

Outgroup

SSU small subunit, LSU large subunit

vel cylindricae, uni-vel multiseptatae, hyalinae vel brunneae, cum vel sine tunicis gelatinosis. Ascomata subglobose to globose, scattered to crowded. Beak compressed, with a slit-like ostiole. Ascomatal wall

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composed of pale brown, small, thin-walled cells. Pseudoparaphyses filamentous, numerous, septate, branched, anastomosing. Asci fissitunicate, cylindrical, with a short stipe or sessile, rounded at the apex, with an apical

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chamber. Ascospores fusiform to cylindrical, 1- to multiseptate, hyaline to brown, with or without an entire gelatinous sheath. Lophiotrema Sacc.

Typus genus:

Lophiostoma quadrisporum K. Hiray. & Kaz. Tanaka, sp. nov. Figs. 4, 6, 10, 14, 18 MycoBank no.:

MB 561064

Ascomata 300–360 lm alta, 300–435 lm diametro, subglobosa vel globosa. Rostrum 130–200 lm latum, cristatum. Paries ascomatis 10–20 lm crassus, ex cellulis prismaticis compositus. Pseudoparaphyses copiosae, 1–2 lm latae. Asci (70–)80–110(–120) 9 (8–)9–11.5 lm, fissitunicati, clavati, cum longistipitibus, quadrispori. Ascosporae 19–24.5 9 6–9.5 lm, ellipsoidei-fusiformes, uniseptatae, hyalinae. Misapplied name: Lophiotrema nucula auct. non (Fr.) Sacc.,: Kaz. Tanaka & Y. Harada, Mycoscience 44: 116, 2003. Etymology:

In reference to the 4-spored asci.

Specimen examined: Japan, Iwate, Morioka, Ueda, campus of Iwate Univ., on twigs of Liriodendron tulipifera L., 11 Jan. 2002, coll. Y. Harada, KT 843 (HHUF 27321, holotype designated here; ex-holotype isolate MAFF 239455). Lophiostoma vitigenum (Kaz. Tanaka & Y. Harada) K. Hiray. & Kaz. Tanaka, comb. nov. Fig. 17 MycoBank no.:

MB 561065

:Lophiotrema vitigenum Kaz. Tanaka & Y. Harada, Mycoscience 44: 119, 2003 (basionym). Specimens examined: Japan, Aomori, Hirosaki, Kudoji, on twigs of Vitis coignetiae Pulliat ex Planch., 27 Oct. 2001, coll. S. Hatakeyama (HHUF 26930 holotype of basionym; ex-holotype isolate JCM 13534 = MAFF 239459); ibid (HHUF 26931 isotype of basionym; ex-isotype isolate JCM 17676).

Discussion Monophylies of Lophiostoma and Lophiotrema All phylogenetic trees obtained in our study confirmed clearly that Lophiostoma and Lophiotrema are separate genera belonging to different families (see Fig. 28); this has been indicated in several previous papers (Schoch et al. 2009; Zhang et al. 2009a, b). Zhang et al. (2009b) suggested that species in Lophiostoma could be divided

407

phylogenetically into two distinct lineages based on analyses of SSU ? LSU nrDNA and RPB2. They described these as ‘‘Lophiostoma clade I,’’ including several melanommataceous genera (e.g., Melanomma Nitschke ex Fuckel and Herpotrichia Fuckel), and ‘‘Lophiostoma clade II,’’ including most Lophiostoma species. They introduced two species in Lophiostoma clade I as new species of the genus (L. rugulosum Yin. Zhang et al. and L. glabrotunicatum Yin. Zhang et al.), because a sequence of L. macrostomum (the type species of Lophiostoma) retrieved from GenBank (DQ384094; voucher Lundqvist 20504 in S) also nested within this clade (Zhang et al. 2009b). However, species in clade I, as well as L. rugulosum and L. glabrotunicatum (Zhang et al. 2009b), appear to be more closely related to the Melanommataceae recently redefined (Mugambi and Huhndorf 2009a) rather than lophiostomatoid fungi, based on morphological feature of ascomata without a laterally compressed crest-like beak or slit-like ostiole. Mugambi and Huhndorf (2009a) suggested that the GenBank sequences of L. macrostomum (DQ384094) may be based on a misidentification. Taxa in ‘‘Lophiostoma clade II’’ sensu Zhang et al. (2009b) are currently accepted as Lophiostoma sensu stricto by several authors (Mugambi and Huhndorf 2009a; Schoch et al. 2009; Suetrong et al. 2009; Tanaka et al. 2010b). In the other genus, Lophiotrema, all species used in our study formed a highly supported monophyly with the exception of Lophiostoma vitigenum, and this clade was distantly placed from the Lophiostoma within the Pleosporales (Fig. 28). Morphological circumscriptions of Lophiostoma and Lophiotrema There has been some controversy about the circumscription of both Lophiostoma and Lophiotrema. The following morphological criteria have been used in the delimitation of both genera: (1) ascomatal size [large (200–700 lm) in Lophiostoma versus small (up to 200 lm) in Lophiotrema; Tang et al. 2003], (2) thickness of ascomatal peridium (*50 lm vs. 25 lm; Holm and Holm 1988), (3) peridial cell type (parallel, long, prismatic cells vs. textura angularis to globosa; Holm and Holm 1988), (4) ascus shape (clavate vs. cylindrical; Holm and Holm 1988), (5) ascospore color (pigmented vs. hyaline; Saccardo 1878), (6) ascospore septation (1- to several septate vs. 1-septate; Holm and Holm 1988), and (7) ascospore appendages (with or without appendages vs. with or without a gelatinous sheath; Holm and Holm 1988). In addition to these characters, we have noted length of ascus stipe for each species to help elucidate generic boundaries (see Fig. 28). A classical understanding following a generic concept based on Saccardoan spore morphology such as color and septation is obviously uninformative (Fig. 28). Saccardo

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b Fig. 28 One of the 51 most parsimonious (MP) trees based on a combined dataset of small subunit (SSU) (884 bp) and large subunit (LSU) (723 bp) rDNA. Most parsimonious (MP) and neighborjoining (NJ) bootstrap values greater than 50% and Bayesian posterior probabilities above 0.90 are indicated at the nodes as MPBV/NJBV/ BPP. Hyphen (‘‘-’’) indicates values lower than 50% (BV) or 0.90 (BPP). Tree length = 1,212, consistency index = 0.461, retention index = 0.797. The tree was rooted to Dothidea insculpta (Dothideales). Taxonomic criteria of Lophiostoma and Lophiotrema are noted after the species name as follows. Ascomatal size: filled triangles, up to 350 lm; open triangles, more than 350 lm. Thickness of ascomatal peridium: filled squares, up to 25 lm; open squares with a dot, 25–50 lm; open squares, more than 50 lm. Peridial cell type: open diamonds, parallel, long, prismatic cells; filled diamonds, small cells of textura angularis to globosa. Ascus shape: open circles, clavate; double circles, clavate to cylindrical; filled circles, cylindrical. Ascus stipe length: filled five-pointed stars, up to 15 lm; open five-pointed stars, more than 15 lm. Ascospore color: P pigmented, H hyaline. Ascospore septation: 1 1-septate, M multiseptate. Ascospore appendage: A appendage, S sheath, N none. Morphological data of the taxa with asterisks were obtained from Abdel-Wahab and Jones (2000) (*) and from Holm and Holm (1988) and/or Zhang et al. (2009a) (**)

(1878) erected Lophiotrema to include fungi that have ascomata with a crest-like beak similar to those of Lophiostoma, but with hyaline, multi-septate ascospores. Spore color was not considered of primary taxonomic importance in defining genera (Chesters and Bell 1970a), but spore septation was used as a criterion in the key to lophiostomatoid genera provided by Holm and Holm (1988). However, the presence of species with pigmented ascospores in the Lophiotrema clade, e.g., L. rugulosum and L. glabrotunicatum (Zhang et al. 2009b), and species with 3-septate ascospores, e.g., L. neohysterioides M.E. Barr (Fig. 28), rejects the diagnostic value of spore color and septation in the separation of these genera. Another major generic concept, emphasizing differences in ascomatal peridium (thickness and cell structure), was proposed by Holm and Holm (1988); this distinction has been widely accepted by many authors (e.g., Barr 1992; Mathiassen 1993; Yuan and Zhao 1994; Tanaka and Harada 2003a,b; Tanaka and Hosoya 2008; Eriksson 2009). Zhang et al. (2009b), however, concluded, from morphological comparison of both genera based on type specimens, that peridial thickness and structure do not have taxonomic significance. This conclusion is generally confirmed by our analyses (Fig. 28). The protruding appendages at the ends of ascospores were sometimes regarded as reliable features for the characterization of Lophiostoma species (Holm and Holm 1988; Tang et al. 2003). Appendages are certainly found only in species within the Lophiostoma clade, and not in the Lophiotrema clade (Fig. 28). It seems likely that the reason for many species with appendaged ascospores within the Lophiostoma clade relates to their habitats. Many are reported from aquatic environments, including,

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for example, L. armatisporum (K.D. Hyde et al.) E.C.Y. Liew et al., L. bipolare (K.D. Hyde) E.C.Y. Liew et al., and L. frondisubmersum (K.D. Hyde) E.C.Y. Liew et al., and all these have bipolar appendages on their ascospores (Hyde et al. 1992; Hyde 1994, 1995). Although these aquatic species are not included in our analyses, they have previously been phylogenetically verified as Lophiostoma (Liew et al. 2002; Tanaka and Hosoya 2008). Several typical Lophiostoma species (e.g., L. macrostomum and L. caulium (Fr.) Ces. & De Not.) have been found frequently on reeds or herbaceous debris in riparian habitats (Tanaka and Harada 2003a); these also have ascospores with protruded appendages (Fig. 28). These extracellular structures are generally considered adaptations to aquatic or humid habitats, better enabling the ascospores to attach to substrates (Shearer 1993; Jones 2006; Vijaykrishna et al. 2006). The phylogenetic significance of spore appendages for generic separation is therefore considered doubtful, because these may evolve convergently among unrelated taxa in aquatic habitats (Hirayama et al. 2010). Our results strongly confirm that ascus shape is a reliable taxonomic indicator to differentiate between Lophiostoma and Lophiotrema. This character has already been proposed for this purpose; Lophiostoma is usually characterized by clavate asci and Lophiotrema by cylindrical asci (Holm and Holm 1988). However, the border between ‘‘clavate’’ and ‘‘cylindrical’’ has sometimes been ambiguous or confusing. We have thus provided the additional character of ascal stipe length (from the base of the ascospore arranged at the lowest position to the base of the stipe) to help define ascus shape. In Lophiostoma species, the clavate asci were found to have a relatively long stipe [mostly (10–) 15–30 lm in length], whereas in Lophiotrema the cylindrical asci have a sessile to short stipe (up to 15 lm). The stipe length of two species, L. quadrisporum and L. vitigenum, previously reported as Lophiotrema (Tanaka and Harada 2003b) but actually belonging to Lophiostoma, is also relatively long (15–33 lm and 12–24 lm, respectively). Although this character has never previously been used for the differentiation of lophiostomatoid genera, we suggest that it should be used and tested in further taxonomic revisions of these genera. Familial placement of Lophiotrema Because there is no appropriate family in current Dothideomycetes classification (Lumbsch and Huhndorf 2010), we have established a new family, Lophiotremataceae, on morphological and molecular grounds, to accommodate Lophiotrema species. Traditionally, Lophiotrema has been placed in the Lophiostomataceae along with Lophiostoma (Saccardo 1883; Clements and Shear 1931; Barr 1992; Lumbsch and Huhndorf 2007; Kirk et al. 2008). However,

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its distant relationship with the Lophiostomataceae has been suggested (Schoch et al. 2009; Zhang et al. 2009a,b) and confirmed by our study, using sequences of more than ten strains of Lophiotrema (Fig. 28). Phylogenetically, the Lophiotrema clade is close to the families Testudinaceae (Tanaka et al. 2009; Zhang et al. 2009a,b) and Tetraplosphaeriaceae (Schoch et al. 2009) in the Pleosporales. Morphologically, however, species in the Lophiotrema clade are quite different from members of the Testudinaceae; the latter is characterized by cleistothecial ascomata and sculptured ascospores (von Arx 1971; Hawksworth 1979). Similarly, Lophiotrema species do not have the Tetraploa-like hyphomycetous anamorphs of taxa in the Tetraplosphaeriaceae (Tanaka et al. 2009). Lophiostomalike fungi having ascomata with a long slit-like ostiole are known from other families, such as the Platystomaceae [Ostropella (Sacc.) Ho¨hn. and Xenolophium Syd.; Mugambi and Huhndorf 2009a] and the Aigialaceae (Rimora; Suetrong et al. 2009). Molecular analyses of five genes (SSU, LSU nrDNA, TEF1, RPB1, and RPB2), however, do not support a close phylogenetic relationship between Lophiotrema and these families (Schoch et al. 2009). This finding indicates that fungi with lophiostomatoid ascomata may have evolved multiple times independently within the Dothideomycetes, as is the case for genera with hysterothecial ascomata with a slit-like ostiole, e.g., Glonium Muhl., Hysterium Pers., and Hysterographium Corda (Mugambi and Huhndorf 2009b). Familial circumscription placing much importance on the slit-like ostiolar opening (e.g., Lophiostomataceae sensu lato; Barr 1992) should not therefore be applied to the Lophiotremataceae, although this character may have taxonomic significance at generic level. Further molecular evidence and morphological evaluation of many species in related genera may be required to more clearly define the phenotypic circumscription of the Lophiotremataceae.

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Lophiostoma semiliberum (Desm.) Ces. & De Not. has been interpreted to be an immature stage of L. arundinis (Pers.) Ces. & De Not. (Munk 1957; Eriksson 1967; Eriksson and Yue 1986) because of the similarities in their ascospore sizes and habitats. We have revealed that these are different taxa, as was suggested from their anamorphic (possibly spermatial) morphology in vitro (Tanaka and Harada 2003a). Lophiostoma vitigenum and L. fuckelii have been treated as Lophiotrema because of their rather small ascomata and peridia of equal thickness (Tanaka and Harada 2003b). As already mentioned, however, these characters do not have phylogenetic significance for generic separation. Because these species both have clavate asci with long stipes, they should be treated as Lophiostoma species. Placement of these taxa in Lophiostoma was verified by the molecular work of Mugambi and Huhndorf (2009a) and our own work. Lophiostoma quadrisporum, described here as a new species, is based on a misidentified material of Lophiotrema nucula (HHUF 27321). Despite several discrepancies between the material and the description of L. nucula in terms of ascospore number and cultural characteristics, Tanaka and Harada (2003b) tentatively judged the specimen to be L. nucula on the basis of ascospore similarity. However, our reexamination of this specimen indicates that the fungus HHUF 27321 is neither congeneric nor conspecific with L. nucula. It is distinguished by ascospores that are somewhat wider than those of L. nucula [6–9.5 lm vs. (4–) 5–6.5 lm wide; Zhang et al. 2009b] and consistently 4-spored asci; L. nucula in contrast has 8-spored asci (Holm and Holm 1988). The clavate asci with long stipes (15–33 lm in length) of L. nucula sensu Tanaka and Harada (2003b) indicate its phylogenetic affinity with Lophiostoma rather than with Lophiotrema. This interpretation is evidently supported by our phylogenetic analyses (Fig. 28).

Notes on the species examined Further study Lophiostoma caulium has been considered to be a species complex of several related species (Chesters and Bell 1970a). Holm and Holm (1988) divided this ‘‘species’’ into five ‘‘varieties,’’ labeling them with the letters ‘‘var. a’’ to ‘‘var. e,’’ mainly on the basis of ascospore size and septation. Tanaka and Harada (2003a) followed these provisional decisions without formal taxonomic status and proposed a new taxon, ‘‘var. f,’’ for species with 9-septate ascospores. We analyzed three varieties (L. caulium ‘‘var. a, d, and f’’) phylogenetically, and these are clearly shown to be three distinct species in our MP tree (see Fig. 28). Further taxonomic revision based on type specimens of the L. caulium complex and phylogenetic analyses using their epitype strains will be necessary to establish species names.

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We have revealed that shape and stipe length of ascus are indicative in differentiating between Lophiostoma (Lophiostomataceae) and Lophiotrema (Lophiotremataceae). However, we would consider it equivocal to separate these at familial level based on this ascus character alone; more fundamental differences should define families. We considered that there were no differences in ascomatal wall anatomy between these genera; however, this opinion should be reexamined using a method of making precise ascomatal sections (Huhndorf 1991). Peridial anatomy is recognized as a good predictor of generic or familial relationship among some Ascomycota (Miller and Huhndorf 2005; Boehm et al. 2009). Leptosphaeria Ces. & De

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Not. (Leptosphaeriaceae) and Phaeosphaeria I. Miyake (Phaeosphaeriaceae), for example, can be distinguished by their wall structure (scleroplectenchymatous in Leptosphaeria and pseudoparenchymatous in Phaeosphaeria) (Caˆmara et al. 2002). These wall traits can be observed only in well-made, thin ascomatal sections (3–4 lm thick) (Huhndorf 1992). Several species in Lophiostoma (e.g., L. arundinis, L. fuckelii, L. macrostomum, and L. sagittiforme Kaz. Tanaka & Hosoya) and Lophiotrema [e.g., L. neoarundinaria (Ellis & Everh.) Yin. Zhang et al., L. neohysterioides, and L. vagabundum (Sacc.) Sacc.] are known to produce ascomata in culture (Leuchtmann 1985; Tanaka and Harada 2003a,b,c; Tanaka and Hosoya 2008). An in vitro developmental study of the ascomata of these homothallic species based on the semi-thin sectioning, as was partly carried out for L. fuckelii (Chesters and Bell 1970b), may help to further resolve important morphological differences between Lophiostoma and Lophiotrema. Acknowledgments This work was partially supported by grants from the Japan Society for the Promotion of Science (JSPS, 22770074), the Sasakawa Scientific Research Grant from the Japan Science Society, and the Hirosaki University Grant for Exploratory Research by Young Scientists (2008–2010). We thank Dr. Vadim A. Mel’nik (Komarov Botanical Institute, Russia) for reviewing the draft manuscript and our reviewers for their valuable comments.

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Mycoscience (2011) 52:413–418 DOI 10.1007/s10267-011-0118-3

SHORT COMMUNICATION

Bioactivity of secondary metabolites and thallus extracts from lichen fungi Mark Kowalski • Georg Hausner Michele D. Piercey-Normore

•

Received: 4 August 2010 / Accepted: 31 March 2011 / Published online: 17 April 2011 ! The Mycological Society of Japan and Springer 2011

Abstract Fungal secondary compounds and total extracts are known to affect growth of bacteria, fungi, and plants. This study tested the effects of purified compounds and total extracts from three lichens on the growth of two plant pathogens, Ophiostoma novo-ulmi ssp. americana and Sclerotinia sclerotiorum. Usnic acid showed no reduction in relative growth rates (RGR), whereas vulpinic acid reduced RGR for both fungi and atranorin reduced RGR of S. sclerotiorum only. However, purified vulpinic acid showed stronger effects than total extracts on fungal growth. The results suggest that these lichens show further promise as a source for bioactive compounds against fungi. Keywords Cladonia amaurocraea ! Hypogymnia physodes ! Plant pathogen ! Vulpicida pinastri More than 800 secondary compounds have been discovered from lichen-forming fungi, and most of these are unique to lichens (Huneck and Yoshimura 1996; Miao et al. 2001), being produced entirely within the fungal partner (Culberson et al. 1985; Culberson and Armaleo 1992). Lichen secondary compounds have historically been used in the taxonomy of lichens (Hawksworth 1976), and many have known ecological functions or medicinal properties (Lawrey 1986; Huneck 1999). In some cases the purified compound alone was bioactive, and in other cases the total M. Kowalski ! M. D. Piercey-Normore (&) Department of Biological Sciences, University of Manitoba, Winnipeg, MB R3T 2N2, Canada e-mail: [email protected] G. Hausner Department of Microbiology, University of Manitoba, Winnipeg, MB R3T 2N2, Canada

extract was reported to be bioactive, suggesting a synergistic effect with other compounds in the extract. Among common secondary metabolites, atranorin has been shown to inhibit spore germination of lichenized fungi (Whiton and Lawrey 1984). Atranorin is produced in the cortex of Hypogymnia physodes where it is thought to change the light wavelength so as to promote photosynthesis (Rao and LeBlanc 1965). Hypogymnia physodes, one of the most common tree lichens in conifer forests in North America, also produces five other secondary compounds (Culberson et al. 1977; Brodo et al. 2001). Usnic acid, another secondary compound, has been shown to inhibit biofilm formation of Staphylococcus aureus (Francolini et al. 2004) and to inhibit growth of eight fungal and Oomycota species in the genera Pythium, Phytophthora, Rhizoctonia, Botrytis, Colletotrichum, Fusarium, Stagonospora, and Ustilago (Halama and Van Haluwin 2004). Usnic acid is produced by many species of Cladonia including Cladonia amaurocraea, which is a ground-dwelling boreal forest species that produces barbatic acid in addition to usnic acid. Vulpinic acid, another secondary compound, is a deep yellow metabolite that is poisonous to mollusks and insects (Elix and Stocker-Wo¨rgo¨tter 2008) and has also been reported to inhibit spore germination of some lichenized fungi (Whiton and Lawrey 1984). Vulpinic acid is produced by Vulpicida pinastri along with two other important secondary compounds. V. pinastri is common in moist habitats on trees and sometimes rocks in the boreal forest of North America (Brodo et al. 2001). The bioactive nature of lichen secondary compounds suggests possible strategies to manage plant pathogens, which may include interference with different developmental stages, such as prevention of spore germination and inhibition of sexual reproduction, or interference directly with mycelial growth. These strategies are perhaps

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achievable with application of a purified bioactive compound or a suite of components with a synergistic effect (reviewed in Huneck 1999). The effects of lichen secondary metabolites on mycelial growth can be determined by measuring the growth of the mycelial mass over time (relative growth rate, RGR) or by measuring the prevention of growth near the source of an inhibitory compound (inhibition). Ophiostoma novo-ulmi ssp. americana Bras. et Kirk, a causative agent for a severe wilt disease (Dutch elm disease), has devastated American elm populations in North America (Brasier 2000). The fungus is also found in Europe and Eurasia and has recently been reported from Japan (Masuya et al. 2010). The application of plant breeding and fungicides to control O. novo-ulmi ssp. americana has been expensive and limited in use (Brasier 2000), and few efforts have been devoted to resistance breeding of native elms. Alternative control strategies would be economically and biologically beneficial for preserving urban forests composed of Ulmus sp. Sclerotinia sclerotiorum (Lib.) de Bary, commonly referred to as ‘‘white mold’’ of many agricultural crops (e.g., beans, canola, carrots, lettuce, sunflowers) and some woody ornamentals, devastates crops and causes spoilage of fruits and vegetables during storage (Agrios 2005). Although there are several fungicides available for S. sclerotiorum, the cost of these chemicals can again be high and timing of application is usually very important (Saharan and Mehta 2008). A combination of biological control agents or fungicides with secondary metabolites might be valuable in developing effective integrated control strategies against this disease (Budge and Whipps 2001; Saharan and Mehta 2008). In this study we examine whether compounds from three lichen-forming fungi (C. amaurocraea, H. physodes, and V. pinastri) are useful as potential antifungal compounds. The effects of three secondary metabolites (atranorin, usnic acid, vulpinic acid) and total lichen thallus extracts on the mycelial growth and inhibition of two unrelated plant pathogens (Sclerotinia sclerotiorum and Ophiostoma novoulmi ssp. americana) were studied. It was hypothesized that (1) all three compounds would reduce the RGR and inhibit growth of these plant pathogenic fungi; (2) that the higher concentrations of the compounds would show greater reduction in RGR and greater inhibition than the lower concentrations; and (3) that the purified compounds would show greater reduction in RGR and greater inhibition than the thallus extracts. The three compounds and total extracts were isolated from the lichen-forming fungal species Cladonia amaurocraea (Flo¨rke) Schaerer (Canada, Manitoba, Wapusk National Park; 58"310 16.600 N, 93"340 22.900 W; 2008, Normore 8652 (WIN(C)), Hypogymnia physodes (L.) Nyl.

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(Canada, Manitoba, Wapusk National Park; 58"300 37.700 N, 93"280 16.400 W, 2008, Normore 8565 (WIN(C)), and Vulpicida pinastri (Scop.) J.-E. Mattsson and M.J. Lai (Canada, Manitoba, Sherridon Rd., 548420 24.700 N, 1018330 53.100 W; 2008 Normore 8728 (WIN(C)). These lichens were selected because they are common in North America (Brodo et al. 2001) and the secondary metabolites are well known and readily isolated. The two plant pathogenic fungi are Sclerotinia sclerotiorum (WIN(M)) 1647, isolated from Daucus carota L. in Manitoba), and Ophiostoma novo-ulmi ssp. americana (WIN(M)) 903, isolated from Ulmus americana Planch. in Manitoba. These pathogens were selected for this study because their growth in culture was suitable for measuring growth rates over a period of 1–2 weeks, and the radial growth on the medium allowed for accurate growth measurements. A total of ten treatments were tested: three purified compounds at two concentrations each (crudely 20 mg/ml and 10 mg/ml from silica-coated plates); total extracts of the three lichens C. amaurocraea, H. physodes, and V. pinastri at a concentration of 20 mg/ml total extract; and a control consisting of acetone, which was the solvent for all treatments. Each treatment was tested on the two plant pathogenic fungi and was replicated five times for each pathogenic fungus with three inoculations per Petri dish. Usnic acid, atranorin, and vulpinic acid were isolated from C. amaurocraea, H. physodes, and V. pinastri, respectively. To obtain 50–100 mg dry weight of the target compounds, clean tissues of the lichens were ground with 25 ml acetone, and the crude extracts obtained were applied to preparative thin-layer chromatography (TLC). The developing solvent was toluene/dioxane/glacial acetic acid (180:45:5) (Orange et al. 2001). A ‘‘purified’’ extract is defined as the extract retrieved from the TLC isolation even though traces of other compounds may be present. A ‘‘crude’’ extract refers to the entire complement of compounds soluble in acetone from the entire lichen thallus. No TLC was performed to obtain the thallus crude extract. Test fungi were cultured on malt yeast agar (MYA) (20 g malt extract, 2 g yeast extract, 15 g agar per 1,000 ml distilled water) at 20"C for 24 h in the dark. A 50-ll portion of test extract or acetone as negative control was dropped on a sterilized triangular filter paper placed on the center of the culture plate; 2-mm fragment of mycelium was then inoculated onto the plate at a 1-cm distance from each of the three edges of the filter paper. Colony diameters were measured every 2 days (Soylu et al. 2007), and the inhibition zone was recorded as the distance between the filter paper and the leading front of mycelial growth. The colony area was calculated based on the diameter measurements of mycelial growth and area = pab (where p = 3.14; a = major radius; b = minor radius). Growth curves were produced by log10 transforming the average

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values for each treatment, and these values were plotted against time. The relative growth rates were then calculated from the interval between day 2 and day 4 because this interval represents the exponential growth phase, providing the most accurate representation of growth (Hunt 1978). Because colony growth was not consistent along the edge of the filter paper for the inhibition assays, three measurements were averaged for each side of the filter paper. The RGR between the treatments were assessed using a two-way analysis of variance (ANOVA) (Sokal and Rohlf 1995) with a 95% confidence interval implemented in Microsoft Excel (ver. 12.2.5) to examine the relationship between the compound (atranorin, usnic acid, vulpinic acid) and the concentration (10 or 20 mg/ml) for each of O. novo-ulmi ssp. americana and S. sclerotiorum. Another two-way ANOVA examined extract type (total extract and purified extract) and lichen species (V. vulpicida, H. physodes, C. amaurocraea) to test the relationship between lichen species and whether the total extract or a single compound affected growth. An interaction suggests these are not independent of one another. Each treatment was also compared to the control treatment to determine if a

significant difference occurred, using a t test with a 95% confidence interval. The same treatments were analyzed for the inhibition measurements. Diagnostic secondary metabolites for the three lichen species are well known (Culberson et al. 1977), with Rf values and spot characteristics described by Orange et al. (2001). Cladonia amaurocraea produces usnic acid (Rf = 0.70) and barbatic acid (Rf = 0.44). Hypogymnia physodes produces atranorin (Rf = 0.75) and chloroatranorin (Rf = 0.74). Five additional compounds are produced with Rf values less than 0.55 in H. physodes. Vulpicida pinastri (=Cetraria pinastri) produces vulpinic acid (Rf = 0.71), pinastric acid (Rf = 0.70), and usnic acid (Rf = 0.70). Therefore, it is likely that the usnic acid isolated from C. amaurocraea is pure, but the isolated atranorin from H. physodes may contain small amounts of chloroatranorin, and the isolated vulpinic acid from V. pinastri may also contain some usnic and pinastric acids. To identify these compounds, the Rf values and the spot characteristics were compared with those in Orange et al. (2001). The first analysis in Table 1 tested the hypothesis that the type of compound (atranornin, usnic acid, vulpinic

Table 1 Summary of analysis of variance (ANOVA) results for relative growth rates (RGR) of Ophistoma novo-ulmi ssp. americana and Sclerotinia sclerotiorum showing effects of purified compound (atranorin, usnic acid, vulpinic acid) and concentration (20 and

10 mg/ml) in one series of tests; and comparing lichen species (Cladonia amaurocraea, Vulpicida pinastri, Hypogymnia physodes) and extraction type (purified, crude extract) in the second series of tests

Source

O. novo-ulmi ssp. americana

S. sclerotiorum

df

F ratio

P value

df

F ratio

P value

Compound

2

65.258

\0.0001

2

174.950

\0.0001

Concentration

1

3.326

0.0807

1

1.096

0.3060

Interaction

2

1.003

0.3817

2

2.598

0.0950

Lichen

2

13.861

\0.0001

2

20.362

\0.0001

Extraction

1

0.389

0.5390

1

21.450

0.0001

Interaction

2

28.022

\0.0001

2

6.343

0.0061

A P value less than 0.05 indicates a statistically significant difference between treatments Table 2 Summary of t-test results for comparison of mean relative growth rates (RGR, mm2/mm2/days) among treatments for Ophistoma novo-ulmi ssp. americana and Sclerotinia sclerotiorum

For both species, treatments were compared with the acetone controlAsterisk (*) indicates a significant difference from the control at a = 0.05 based on a t test

Treatment

Mean relative growth rate (mm2/mm2/days) O. novo-ulmi ssp. americana

S. sclerotiorum

Acetone control

0.49

0.53

Usnic acid (20 mg/ml)

0.46

0.53

Usnic acid (10 mg/ml)

0.40

0.48

Atranorin (20 mg/ml)

0.45

0.37*

Atranorin (10 mg/ml)

0.40

0.37*

Vulpinic acid (20 mg/ml)

0.21*

0.18*

Vulpinic acid (10 mg/ml) Total extract (Cladonia amaurocraea)

0.21* 0.30*

0.20* 0.49

Total extract (Hypogymnia physodes)

0.37

0.46

Total extract (Vulpicida pinastri)

0.37

0.41*

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acid) and the concentration (20 or 10 mg/ml) had an effect on fungal growth. Results show that the type of compound had a significant effect on growth but the concentration of the compound did not affect growth. The second analysis tested the hypothesis that lichen species (C. amaurocraea, H. physodes, V. pinastri) and extraction type (purified or total extract) had an effect on mycelial growth. Although type of extract had no effect on mycelial growth, the lichen species did have an effect. That both compound and lichen species had a significant effect on mycelial growth is supported by the taxonomic concept of chemical species within lichens (Hawksworth 1976). The analysis in Table 2 tested the hypothesis that each compound and each extract

Growth rates

A

Growth rates

B

0.5 0.4 0.3 0.2 0.1 0

0.6 0.4 0.2 0

Growth rates

C

D Growth rates

Fig. 1 Comparison among treatments showing relative growth rates (mm2/mm2/day) of Ophistoma novo-ulmi ssp. americana at both concentrations for each of the three purified compounds (a); relative growth rates of S. sclerotiorum for both concentrations for each of the three purified compounds (b); relative growth rates of O. novo-ulmi ssp. americana for pure and total thallus extracts (c); relative growth rates of S. sclerotiorum for pure and total thallus extracts (d); growth inhibition (mm) for S. sclerotiorum for both concentrations for each of the three purified compounds (e); and growth inhibition (mm) for S. sclerotiorum for pure and total thallus extracts (f). C. am, Cladonia amaurocraea; H. ph, Hypogymnia physodes; V. pi, Vulpicida pinastri

Inhibition

E

0.6 0.4 0.2 0

0.6 0.4 0.2 0

3 2 1 0

F Inhibition

3 2 1 0

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could limit mycelial growth in the plant pathogens. Table 2 shows that vulpinic acid alone and the total extract from C. amaurocraea reduced the growth of O. novo-ulmi. However, two compounds alone (atranorin and vulpinic acid), as well as the total extract from V. pinastri, reduced the growth of S. sclerotiorum. Figure 1 shows that vulpinic acid reduced RGR for both species (Fig. 1a–d) whereas atranorin reduced RGR only of S. sclerotiorum. The data also show that vulpinic acid inhibited growth of both pathogens and the crude extract from C. amaurocraea inhibited growth of S. sclerotiorum (Fig. 1e–f). The decrease in RGR of O. novo-ulmi ssp. americana and the inhibition of S. sclerotiorum growth with the total Usnic 10 Usnic 20 Atranorin 10 Atranorin 20 Vulpinic 10 Vulpinic 20 Usnic 10 Usnic 20 Atranorin 10 Atranorin 20 Vulpinic 10 Vulpinic 20 C. am pure C. am total H. ph pure H. ph total V. pi pure V. pi total C. am pure C. am total H. ph pure H. ph total V. pi pure V. pi total Usnic 10 Usnic 20 Atranorin 10 Atranorin 20 Vulpinic 10 Vulpinic 20 C. am pure C. am total H. ph pure H. ph total V. pi pure V. pi total

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extract from C. amaurocraea suggest that other components within the thallus extract, in addition to usnic acid, affected mycelial growth. This finding is supported by Halama and Van Haluwin (2004), who reported greater inhibition of fungal growth in the total extract treatment and only slight inhibition by purified usnic acid. Previously, usnic acid was shown to inhibit fungal growth at concentrations greater than 32 lg/ml (Lauterwein et al. 1995) and with 50 lg of the chemical added to the plate (Ko¨nig and Wright 1999). The concentrations used in this study were greater than the reported effective concentrations, but the extraction method likely included impurities and fine silica particles that inflated dry weight measurements. Although no inhibition was observed with the purified atranorin treatment or the total H. physodes extract for S. sclerotiorum, the RGR of S. sclerotiorum decreased with the purified atranorin treatment in this study. Atranorin alone was previously shown to be a weak growth inhibitor (Lauterwein et al. 1995; Halama and Van Haluwin 2004), but the crude extract of H. physodes was effective against several plant pathogenic fungi (Halama and Van Haluwin 2004). The purified vulpinic acid compound showed greater activity than the total extract from V. pinastri, which is the opposite result of that shown by usnic acid and the extract from C. amaurocraea. Our findings are supported by previous reports that vulpinic acid, at 50 lg/plate, inhibited the growth of several fungi (Ko¨nig and Wright 1999). The greater growth inhibition observed in S. sclerotiorum compared to O. novo-ulmi spp. americana in many of the treatments in this study is consistent with findings by Land and Lundstro¨m (1998). They reported that yeasts, molds, blue stain fungi, and rot fungi behave differently when exposed to total extracts from Nephroma arcticum. Total extracts from species of Cladonia, Hypogymnia, and Evernia have also been reported to differentially reduce mycelial growth of various ascomycete and basidiomycete species, depending on the species tested (Halama and Van Haluwin 2004). The results showed that some lichen fungi produce bioactive secondary metabolites, which were either purified compounds or combinations with other components contained in the total extract. Although optimal concentrations of their antifungal activities were not determined in this study, the relative evaluation between total extracts and purified compounds may provide a direction for assessing antifungal activity. Usnic acid alone did not inhibit or reduce RGR in either of the plant pathogens tested, whereas the total extract of C. amaurocraea containing usnic acid showed synergistic activity. Vulpinic acid had the greatest effect among the three compounds studied, but we cannot exclude the possibility that our TLC preparation included traces of other

417

compounds in addition to vulpinic acid. The apparent low bioactivity for V. pinastri total extracts on O. novo-ulmi ssp. americana may result from the low concentrations of vulpinic acid. Our findings on the two plant pathogens justify further work that might include a larger number of isolates and species. The identification of bioactive compounds against S. sclerotiorum and O. novo-ulmi ssp. americana offers an opportunity to develop alternative strategies to control these fungi. Acknowledgments The authors thank S. Athukorala for help with fungal culturing, N. Kenkel for statistical advice, and T. Booth and K. Fontaine for checking the English. Lichens were collected with Parks Canada (permit number WAP-2008-67). This project was funded by National Science and Engineering Research Council of Canada (NSERC). Experiments comply with current Canadian laws.

References Agrios GN (2005) Plant Pathology, 5th edn. Elsevier Academic Press, Burlington, MA Brasier CM (2000) Intercontinental spread and continuing evolution of the Dutch elm disease pathogen. In: Dunn CP (ed) The elms: breeding, conservation, and disease management. Kluwer Academic, Boston Brodo IM, Sharnoff S, Sharnoff S (2001) Lichens of North America. Yale University Press, New Haven, 795 pp Budge SP, Whipps JM (2001) Potential for integrated control of Sclerotinia sclerotiorum in glasshouse lettuce using Coniothyrium minitans and reduced fungicide application. Phytopathology 91:221–227 Culberson CF, Armaleo D (1992) Induction of a complete secondary product pathway in a cultured lichen fungus. Exp Mycol 16:52–63 Culberson CF, Culberson WL, Johnson A (1977) Second supplement to ‘‘Chemical and botanical guide to lichen products’’. American Bryological and Lichenologial Society, Missouri Botanical Garden, St. Louis Culberson CF, Culberson WL, Johnson A (1985) Does the symbiont alga determine chemotype in lichens? Mycologia 77:657–660 Elix JA, Stocker-Wo¨rgo¨tter E (2008) Biochemistry and secondary metabolites. In: Nash TH III (ed) Lichen biology, 2nd edn. Cambridge University Press, Cambridge Francolini I, Norris P, Piozzi A, Donelli G, Stoodley P (2004) Usnic acid, a natural antimicrobial agent able to inhibit bacterial biofilm formation on polymer surfaces. Antimicrob Agents Chemother 48:4360–4365 Halama P, van Haluwin C (2004) Antifungal activity of lichen extracts and lichenic acids. BioControl 49:95–107 Hawksworth DL (1976) Lichen chemotaxonomy. In: Brown DH, Hawksworth DL, Bailey RH (eds) Lichenology: progress and problems. Academic Press, London Huneck J (1999) The significance of lichens and their metabolites. Naturwissenschaften 86:559–570 Huneck J, Yoshimura I (1996) Identification of lichen substances. Springer, Berlin Hunt R (1978) Plant growth analysis. In: The Institute of Biology’s Studies in Biology, no. 96. Arnold, London Ko¨nig G, Wright A (1999) 1H and 13C-NMR and biological activity investigations of four lichen-derived compounds. Phytochem Anal 10:279–284

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418 Land CJ, Lundstro¨m H (1998) Inhibition of fungal growth by water extracts from the lichen Nephroma arcticum. Lichenologist 30:259–262 Lauterwein M, Oethinger M, Belsner K, Peters T, Marre R (1995) In vitro activities of the lichen secondary metabolites vulpinic acid, (?)-usnic acid, and (-)-usnic acid against aerobic and anaerobic microorganisms. Antimicrob Agents Chemother 39:2541–2543 Lawrey J (1986) Biological role of lichen substances. Bryologist 89:111–122 Masuya H, Brasier C, Ichihara Y, Kubono T, Kanzaki N (2010) First report of the Dutch elm disease pathogens Ophiostoma ulmi and O. novo-ulmi in Japan. Plant Pathol 59:805 Miao V, Coeffet M, Brown D, Sinnemann S, Donaldson G, Davies J (2001) Genetic approaches to harvesting lichen products. Trends Biotechnol 19:349–355

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Mycoscience (2011) 52:413–418 Orange A, James PW, White FJ (2001) Microchemical methods for the identification of lichens. British Lichen Society, London Rao DN, LeBlanc F (1965) A possible role of atranorin in the lichen thallus. Bryologist 68:284–289 Saharan GS, Mehta N (2008) Sclerotinia diseases of crop plants: biology, ecology and disease management. Springer, Heidelberg Sokal RR, Rohlf FJ (1995) Biometry: the principles and practice of statistics in biological research, 3rd edn. Freeman, New York Soylu S, Yigitbas H, Soyle E, Kurt S (2007) Antifungal effects of essential oils from oregano and fennel on Sclerotinia sclerotiorum. J Appl Microbiol 103:1021–1030 Whiton JC, Lawrey JD (1984) Inhibition of crustose lichen spore germination by lichen acids. Bryologist 87:42–43

Mycoscience (2011) 52:419–424 DOI 10.1007/s10267-011-0119-2

SHORT COMMUNICATION

Boletus kermesinus, a new species of Boletus section Luridi from central Honshu, Japan Haruki Takahashi • Yuichi Taneyama Akito Koyama

•

Received: 5 December 2010 / Accepted: 7 April 2011 / Published online: 27 April 2011 ! The Mycological Society of Japan and Springer 2011

Abstract Boletus kermesinus, a new species of Boletus section Luridi, is fully described and illustrated based on the materials collected in subalpine coniferous forests of central Honshu, Japan. It has distinctive features of darkred basidiomata having distinct viscidity in the pileus surface, usually unchanging flesh, discolorous red pores, and an entirely reticulate stipe becoming coarsely laceraterimose with age. Keywords

Boletales ! Japanese mycobiota ! Taxonomy

Boletus section Luridi Fr. is well delimited in the genus macromorphologically in the boletoid habit, the small, often discolored, pores, and the reticulate or finely furfuraceous stipe surface (Singer 1986). The species of the section Luridi are mainly distributed in North America (Singer 1947; Snell and Dick 1970; Smith and Thiers 1971; Thiers 1975; Bessette et al. 1997, 2000), Europe (Singer 1967; Alessio 1985), and Southeast and East Asia (Corner 1972; Zang 2006). Excluding poorly known species and invalid names (http://www.indexfungorum.org), 14 taxa of the section Luridi have hitherto been recorded from Japan, namely, B. brunneissimus W.F. Chiu (=B. umbriniporus Hongo) (Hongo 1969; Wang and Liu 2002), B. fuscopunctatus Hongo and Nagas. (Hongo and Nagasawa 1976),

B. obscureumbrinus Hongo (Hongo 1968), B. pseudocalopus Hongo (Hongo 1972), B. sensibilis var. sensibilis Peck (Hongo and Nagasawa 1980), B. subcinnamomeus Hongo (Hongo 1977), B. venenatus Nagas. (Nagasawa 1996), B. laetissimus Hongo (1968), B. luridiformis var. luridiformis Rostk. (=B. erythropus Pers. sensu Imazeki 1952), B. subvelutipes f. subvelutipes Peck, (Nagasawa 1989), B. bannaensis Har. Takah. (Takahashi 2007), B. quercinus Hongo (Hongo 1967), B. generous Har. Takah. (Takahashi 1988), and B. rhodocarpus Uehara and Har. Takah. (Takahashi 2001). In this article, a peculiar new species of Boletus section Luridi is presented based on the materials collected in subalpine coniferous forests of central Honshu, Japan. Macroscopic features are all based on fresh materials. For microscopic observations, free-hand sections of the fresh or dried materials were examined in Melzer’s reagent, 30% NH4OH, 3% KOH, phloxine B, Congo red, or distilled water using a Nikon Eclipse 50i microscope. Basidiospores were measured in side view, excluding the hilar appendix; ± standard deviation, Q (length/width) range, and each average (median length, median width, median Q) were statistically derived from a random selection of all basidiospores measured. Color notations in parentheses are taken from Kornerup and Wanscher (1983). Specimens cited here are deposited in the National Museum of Nature and Science in Tsukuba (TNS).

H. Takahashi (&) 284-1 Ouhama, Ishigaki, Okinawa 907-0001, Japan e-mail: [email protected]

Boletus kermesinus Har. Takah., Taneyama & Koyama, sp. nov. Figs. 1–16 MycoBank no.: MB 519342.

Y. Taneyama 392-3 Yashikida, Nagano, Nagano 381-0055, Japan

Pileo (20–)52–68(–87) mm lato, convexo dein late convexo, viscido in humidis, glabro, rubro, margine leniter pallidiore; carne pallide flavida, immutabili, odore saporeque nullo; stipite 50–80 (–120) 9 8–20 mm, subaequali vel

A. Koyama 12,291-2 Otiai, Fujimi-cho, Nagano 399-0214, Japan

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Figs. 1–4 Basidiomata of Boletus kermesinus. 1 Basidioma in habitat. 2 Mature basidioma. 3 Vertical section of the mature basidioma (TNS-F37404). 4 Close-up of the somewhat raised reticulations on the young stipe. Bars 1–3 20 mm; 4 2 mm. Color photographs of Boletus kermesinus can be seen at http://www7a.biglobe.ne.jp/ *har-takah/page137.html

subventricosis, solido, rubro, manifeste rubro-reticulato, dein rimoso-areolato; tubulis depressis ad stipitem, flavis, fractis immutabilis; poris subrotundatis, parvis, rubris, immutabilis; basidiosporis (13.4–)15.5–18.6(–21.1) 9 (4.9–) 5.6–6.6(–7.2) lm, fusoideis vel subfusoideis, levibus, depressione suprahilari praeditis, sub microscopio in KOH hyalinis vel melleis; basidiis (32.1–)39.7–50.1(–53.9) 9 (9.2–)10.4–12.5(–14.8) lm, (1–2–)3–4 sporis; cheilocystidiis (34.6–)52.7–71.0(–76.5) 9 (3.8–)4.5–6.3(–7.1) lm, subcylindricis; pleurocystidiis (36.6–)44.9–58.4(–75.2) 9 (5.9–)6.9–8.6(–9.0) lm, fusoideo-ventricosis, apicem versus attenuatis; tramate hymenophori bilaterali subtypo

123

Boleto; pileipelle ex hyphis ixotrichodermialibus subcylindricis vel subclavatis composita; stipitipelle hymeniformi ex caulocystidiis fusoideo-ventricosis vel subcylindricis, rubris composita; hyphis defibulatis. Holotypus: Ad terram in silvis Abietis mariesii Mast., Sakuho-cho, Minamisaku-gun, Nagano Pref., Japonia, 29 Aug. 2009, M. Taneyama (TNS-F-37407). Etymology: from Latin, kermesinus = crimson-red. Pileus (Figs. 1, 2) (20–)52–68(–87) mm in diameter, at first hemispherical, expanding to broadly convex, with straight margin, often slightly appendiculate; surface glabrous,

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Figs. 5–14 Micromorphological features of Boletus kermesinus (holotype). 5 Vertical section of the pileipellis showing the ixotrichoderm consisting of loosely interwoven hyphae. 6 Terminal elements of the pileipellis. 7 Cheilocystidia. 8 Pleurocystidia.

421

9 Basidia. 10–13 Terminal elements of the stipitipellis (10, 11 from between the reticula; 13 from the reticulum). 12 Caulobasidia. 14 Basidiospores. Bars 5 50 lm; 6–13 20 lm; 14 10 lm

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smooth, viscid when wet, brownish red (10C7) to dark red (10C8) overall when young, often with a high red (10A8) to red (10B7–8) margin, becoming violet brown (10E7–8 to 10F7–8) in age, at times with faded portions in patches. Flesh (Fig. 3) up to 12 mm thick, yellowish white (3A2) to pale yellow (3A3), red (10A6 to 10B7) immediately beneath the pileus and the stipe surface, unchanging or rarely slightly changing to blue only around the tubes when cut; odor and taste indistinct or somewhat sour. Stipe (Figs. 1, 2, 4) 50–80(–120) 9 8–20 mm, subcylindrical or subventricose, often tapering toward the base, central, terete, solid; surface dry, concolorous with the pileus, initially entirely covered with a somewhat raised reticulum colored brownish red (10C7) to dark red (10C8) or blackish when older; as the stipe grows, the reticulated surface disrupting into large to small scaly patches particularly at the lower portion; base covered with pale yellowish mycelium. Tubes to 10 mm deep, depressed around the stipe, light yellow (3A5) to yellow (3A6), changing to blue when bruised; pores 2–3/mm when young, 1–2/mm in age, subcircular, brownish red (10C7) to dark red (10C8), changing to blue when bruised. Basidiospores (Fig. 14) (13.4–)15.5–18.6(–21.1) 9 (4.9–) 5.6–6.6(–7.2) lm (n = 100, median length = 17.03 ± 1.52, median width = 6.11 ± 0.51, Q = (2.4–)2.6–3.0 (–3.2), median Q = 2.8 ± 0.2), inequilateral with a distinct suprahilar depression in profile, fusoid to subfusoid in face view, with a rounded apex, smooth, with dull yellow (3B4) to greyish yellow (3B5) contents (in water) turning greyish yellow (4B6) to orange yellow (4B7) in KOH, inamyloid or weakly dextrinoid, with slightly thickened walls (up to 0.5 lm). Basidia (Fig. 9) (32.1–)39.7–50.1 (–53.9) 9 (9.2–)10.4–12.5(–14.8) lm, clavate, (1–2–)3–4spored; sterigmata (2.8–)3.7–6.9(–9.0) 9 (1.3–)1.8–2.5 (–2.9) lm. Basidioles clavate. Cheilocystidia (Fig. 7) abundant, forming a compact sterile edge, (34.6–)52.7– 71.0(–76.5) 9 (3.8–)4.5–6.3(–7.1) lm (n = 36), subcylindrical with an obtuse apex, smooth, with intracellular orange red (8A6–7) to brownish red (8C6–7) pigments, inamyloid, thin-walled. Pleurocystidia (Fig. 8) scattered, (36.6–)44.9–58.4(–75.2) 9 (5.9–)6.9–8.6(–9.0) lm (n = 38), narrowly fusoid-ventricose to ventricose-rostrate with an obtuse apex, smooth, hyaline in water and KOH, inamyloid, thin-walled. Hymenophoral trama bilateraldivergent of the Boletus subtype; elements 4.9–6.6 lm wide, cylindrical, smooth, gelatinized, colorless in lateral strata, pale melleous in a mediostratum, inamyloid, thinwalled. Pileipellis (Fig. 5) of an ixotrichoderm consisting of loosely interwoven hyphae (2.0–)2.2–3.4(–4.9) lm wide, cylindrical, smooth, gelatinized, with orange red (8A6–7) to brownish red (8C6–7) contents (in water), which instantly turn olivaceous (near to greyish yellow: 3B5–6) and then gradually fade to more yellowish (near to

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4B5–6) in KOH, inamyloid, thin-walled; terminal cells (Fig. 6) (25.4–)34.2–65.4(–75.3) 9 2.9–5.2(–6.4) lm, subcylindrical to subclavate, occasionally mucronate at the apex. Pileitrama of cylindrical, loosely interwoven hyphae 3.4–6.2 lm wide, smooth, colorless in water, inamyloid, thin-walled. Stipitipellis (Figs. 10–13, 15, 16) hymeniform, consisting of subclavate to fusiform cells that are distributed over the entire stipe surface, 32.1–41.2 9 5.9–7.2 lm, smooth, with orange red (8A6–7) contents (in water) turning greyish yellow (4B5–6) in KOH, inamyloid, thinwalled, and of scattered caulobasidia (Fig. 12) (29.9–) 36.4–44.6(–47.1) 9 (7.8–)8.1–10.0(–11.2) lm, (1–)2–3– 4-spored; edges of the reticulum mainly made up of subcylindrical to ventricose-rostrate caulocystidia (Figs. 13, 16) (40.1–)47.5–67.8(–85.2) 9 (4.5–)5.2–7.6(–9.7) lm (n = 32), smooth, with obtuse apices, with orange red (8A6–7) contents (in water) turning greyish yellow (4B5–6) in KOH, inamyloid, thin-walled. Stipe trama composed of longitudinally running, cylindrical cells (3.2–)4.6-7.7 (–11.2) lm wide, unbranched, smooth, colorless in water, greyish yellow (4B5–6) in KOH, inamyloid, thin-walled. Clamp connections absent. Habitat: Solitary to scattered, on ground in highland (subalpine) forests dominated by Tsuga diversifolia

Figs. 15, 16 Terminal elements of the stipitipellis. 15 Terminal elements between the reticulum (in annular illumination, holotype). 16 Terminal elements on the edge of the reticulum (in phloxine B, TNS-F-37404). Bars 20 lm

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(Maxim.) Mast., A. mariesii, and Abies veitchii Lindl., 1,800–2,100 m alt., July to October, not common. Known distribution: Japan (Nagano). Specimens examined: TNS-F-37407 (holotype), Sakuhocho, Minamisaku-gun, Nagano Pref., on ground in highland forests dominated by A. mariesii, 29 Aug. 2009, coll. Taneyama, M.; TNS-F-37408, same place, 6 Sept. 2009, coll. Arano, T.; TNS-F-37409, same place, 29 Aug. 2009, coll. Kitahara, K.; TNS-F-36806, same place, on ground in highland forests dominated by T. diversifolia and A. veitchii, Oct. 3, 2010, coll. Koyama, A.; TNS-F-36805, same place, Aug. 3, 2010, coll. Koyama, A.; TNS-F-36804, same place, July 25, 2010, coll. Koyama, A.; TNS-F-36802, same place, Sept. 5, 2008, coll. Koyama, A.; TNS-F-37404, Azumino-shi, Nagano Pref., on ground in highland forests dominated by A. mariesii, 6 Sept. 2008, coll. Itahana, K.; TNS-F-37405, Nagano-shi, Nagano Pref., on ground in highland forests dominated by A. mariesii, 16 July. 2009, coll. Fujisawa, T.; TNS-F-37406, Yamanouchi-cho, Simotakai-gun, Nagano Pref., on ground in highland forests dominated by A. mariesii, 8 Aug. 2009, coll. Taneyama, Y.; TNS-F-37410, Suzaka-shi, Nagano Pref., on ground in highland forests dominated by A. mariesii, 29 Aug. 2009, coll. Fujisawa, T.; TNS-F-36803, Ina-shi, Nagano Pref., on ground in highland forests dominated by T. diversifolia and A. veitchii, Sept. 7, 2008, coll. Koyama, A.; TNS-F-36801, same place, Sept. 18, 2006, coll. Koyama, A. Japanese name: Akane-amiasi-iguchi. Comments: The diagnostic features of this species are the dark red to violet brown, boletoid basidiomata having a viscid pileus and an entirely reticulate stipe, the usually unchanging flesh, the discolorous red pores, the olivaceous, fusiform basidiospores, and the habitat in subalpine, coniferous forests. In addition, the coarsely lacerate-rimose stipe surface in aged specimens is its unique characteristic. In the combination of the features mentioned previously, B. kermesinus can be referable to the section Luridi Fr. sensu Singer (1986). The young basidiomata of B. kermesinus, which possess an entirely reticulate stipe, bear a striking resemblance to the following two species: B. frostii J.L. Russell from eastern North America (Snell and Dick 1970; Smith and Thiers 1971; Bessette et al. 1997, 2000) and Costa Rica (Halling and Mueller 2005), and B. pseudofrostii B. Ortiz from Belize (Ortiz-Santana et al. 2007). Boletus frostii is distinct in its characteristics of distinctly cyanescent flesh and a consistently alveolate-reticulate stipe in which the reticulum is much coarser and deeper than that of B. kermesinus. Boletus pseudofrostii is easily separated from B. kermesinus because it produces much smaller basidiomata with pilei 17–20 mm in diameter (Ortiz-Santana et al.

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2007), with yellow ground color on the stipe surface, ellipsoid basidiospores without a suprahilar depression in profile, and pileipellis consisting of entangled, repent hyphae encrusted with red pigmentation. Leccinum rubrum M. Zang from Tibet (Zang 1986) also has dark red basidiomata with unchanging flesh and a distinctly viscid pileus in common with B. kermesinus. However, L. rubrum can be discerned from B. kermesinus by forming a non-reticulate, squamulose-punctate stipe becoming yellow or orange at the apex, having reddish hymenophoral trama, and lacking cheilocystidia. Among the Japanese members of the section Luridi, B. generous Har. Takah. (Takahashi 1988) is most similar macromorphologically, but apparently differs from B. kermesinus in the stipe being finely reticulated and densely dotted with brownish red on the yellowish ground, distinctly cyanescent flesh, and habitat in deciduous oak forests. Acknowledgments We are grateful to Professor Dr. Zhu Liang Yang, Kunming Institute of Botany, the Chinese Academy of Sciences, for his valuable information about Leccinum rubrum, and to Dr. Tsuyosi Hosoya for allowing the specimens cited to be kept in the National Museum of Nature and Science in Tsukuba (TNS).

References Alessio CL (1985) Boletus Dill. ex L. (sensu lato). Fungi Europaei 2. Biella Giovanna, Saronno Bessette AE, Bessette AR, Fischer DW (1997) Mushrooms of northeastern North America. Syracuse University Press, New York Bessette AE, Roody WC, Bessette AR (2000) North American boletes. A color guide to the fleshy pored mushrooms. Syracuse University Press, New York Corner EJH (1972) Boletus in Malaysia. Government Printing Office, Singapore Halling RE, Mueller GM (2005) Common mushrooms of the Talamanca Mountains, Costa Rica. Mem N Y Bot Gard 90:1–195 Hongo T (1967) Notulae Mycologicae (6). Mem Fac Educ Shiga Univ Nat Sci 17:89–95 Hongo T (1968) Notulae Mycologicae (7). Mem Fac Educ Shiga Univ Nat Sci 18:47–52 Hongo T (1969) Notes on Japanese larger fungi (20). J Jpn Bot 44:230–238 Hongo T (1972) Notulae Mycologicae (11). Mem Fac Educ Shiga Univ Nat Sci 22:63–68 Hongo T (1977) Notulae Mycologicae (15). Mem Fac Educ Shiga Univ Nat Sci 27:20–25 Hongo T, Nagasawa E (1976) Notes on some boleti from Tottori II. Rep Tottori Mycol Inst 14:85–89 Hongo T, Nagasawa E (1980) Notes on some boleti from Tottori V. Rep Tottori Mycol Inst 18:133–141 Imazeki R (1952) The Boletaceae of Japan. Nagaoa 2:30–46 Kornerup A, Wanscher JH (1983) Methuen handbook of colour. Methuen, London Nagasawa E (1989) Boletaceae. In: Imazeki R, Hongo T (eds) Colored illustrations of mushrooms of Japan II (in Japanese). Hoikusha, Osaka, pp 1–44

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424 Nagasawa E (1996) A new poisonous species of Boletus from Tottori. Rep Tottori Mycol Inst 33:1–6 Ortiz-Santana B, Lodge DJ, Baroni TJ, Both EE (2007) Boletes from Belize and the Dominican Republic. Fungal Divers 27(2):247–416 Singer R (1947) The Boletineae of Florida with notes on extralimital species III. Am Midl Nat 37:1–135 Singer R (1967) Die Ro¨hrlinge, vol II. Klinkhardt, Bad Heilbrum Singer R (1986) The Agaricales in modern taxonomy, 4th edn. Koeltz, Koenigstein Smith AH, Thiers HD (1971) The boletes of Michigan. University of Michigan Press, Ann Arbor Snell WH, Dick EA (1970) The boleti of northeastern North America. Cramer, Vaduz

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Mycoscience (2011) 52:419–424 Takahashi H (1988) A new species of Boletus sect. Luridi and a new combination in Mucilopilus. Trans Mycol Soc Jpn 29:115–123 Takahashi H (2001) Notes on new Agaricales of Japan 2. Mycoscience 42:347–353 Takahashi H (2007) Five new species of the Boletaceae from Japan. Mycoscience 48:90–99 Thiers HD (1975) California mushrooms: a field guide to the boletes. Hafner Press, New York Wang X-H, Liu P-G (2002) Notes on several boleti from Yunnan. Mycotaxon 84:125–134 Zang M (1986) Notes on the Boletales from Eastern Himalayas and adjacent areas of China. Acta Bot Yunn 8(1):1–22 Zang M (ed) (2006) Flora Fungorum Sinicorum, vol 22. Boletaceae. Science Press, Beijing

Mycoscience (2011) 52:425–430 DOI 10.1007/s10267-011-0121-8

SHORT COMMUNICATION

The mycorrhiza of Schizocodon soldanelloides var. magnus (Diapensiaceae) is regarded as ericoid mycorrhiza from its structure and fungal identities Ayako Okuda • Masahide Yamato Koji Iwase

•

Received: 12 January 2011 / Accepted: 11 April 2011 / Published online: 1 May 2011 ! The Mycological Society of Japan and Springer 2011

Abstract Structure and fungal identities were examined in the mycorrhizal roots of Schizocodon soldanelloides var. magnus (Diapensiaceae) to determine the mycorrhizal category. Previous studies had suggested the mycorrhizae of Diapensiaceae could be categorized as ericoid, but the mycorrhizal fungi have never been identified. The diameter of the fine lateral roots, in which coiled hyphae were found in epidermal cells, was mostly less than 100 lm. Molecular analyses identified the fungal isolates to be Helotiales and Oidiodendron. From the structure and fungal identities, we confirmed that the mycorrhiza of S. soldanelloides is an ericoid mycorrhiza. Keywords Angiosperm Phylogeny Group ! Ericales ! Helotiales ! ITS rDNA ! Oidiodendron Ericoid mycorrhizae are generally recognized in Ericoideae, Cassiopoideae, Vaccinioideae, and Styphellioideae in Ericaceae (Smith and Read 2008). The ericaceous plants have specialized narrow lateral roots, called ‘‘hair roots,’’ that consist of one or two layers of cortical cells and an epidermal layer of enlarged cells (Peterson et al. 2004). Ericoid mycorrhizae are characterized by hyphal coils in the epidermal cells. The mycorrhizal fungi enable host

A. Okuda Graduate School of Agriculture, Tottori University, 4-101 Koyama-Minami, Tottori 680-8553, Japan M. Yamato (&) ! K. Iwase Fungus/Mushroom Resource and Research Center, Faculty of Agriculture, Tottori University, 4-101 Koyama-Minami, Tottori 680-8553, Japan e-mail: [email protected]

plants to utilize complex organic nitrogen, which is unavailable to the plants themselves (Stribley and Read 1980). This mechanism is advantageous for the growth of plants in cool-temperate climates because decomposition of soil organic materials is usually slow (Straker 1996). Fungal isolation and molecular studies of ericoid mycorrhizae have identified a diverse assemblage of fungi from hair roots of ericaceous plants, and most of them are in Helotiales or Oidiodendron in Ascomycota (Smith and Read 2008). Diapensiaceae is a small plant family consisting of five genera, and most species are restricted to eastern Asia and the Appalachian Mountain system of eastern North America (Scott and Day 1983; Ro¨nblom and Anderberg 2002). This family belongs to the order Ericales and is closely related to Symplocaceae and Styracaceae (Angiosperm Phylogeny Group 2003). These two plant families are known to form arbuscular mycorrhiza (Janse 1897; Zangaro et al. 2003); however, the ericoid mycorrhiza was reported in Diapensia lapponica L. in Diapensiaceae by Hesselman (1900), as cited by Harley and Harley (1987). Kagawa et al. (2006) surveyed mycorrhizal formation in an alpine plant community in the Japanese South Alps, in which they classified the mycorrhiza of Shortia soldanelloides f. alpina, a synonym of Schizocodon soldanelloides Sieb. et Zucc. f. alpinus Maxim., in Diapensiaceae as ericoid mycorrhiza. They determined the mycorrhizal category according to the criteria of Harley (1989) by observation of hyphal coils in roots stained with 0.05% trypan blue. However, no structural description of the mycorrhiza was shown, and no mycorrhizal fungi were identified in their study. In the present study, we examined the structure and fungal identities in mycorrhizae of S. soldanelloides var. magnus (Makino) Hara in Diapensiaceae to confirm the mycorrhizal category of S. soldanelloides.

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The sampling site was in a deciduous broad-leaved forest in Yazu-cho, Tottori Prefecture, approximately 740 m above sea level. The dominant tree species of the forest was Fagus crenata Blume, with Quercus crispula Blume and Abies firma Sieb. et Zucc. sparsely intermixed. In vegetation of the understory, S. soldanelloides var. magnus was dominant. Soil core samples including stem and roots of S. soldanelloides var. magnus, 10 cm 9 10 cm 9 5 cm deep, were collected. In total, ten samples (SC1-10) were collected in October 2008, April 2009, and June 2009 (Table 1). The collected samples were kept in plastic bags in cool conditions until processed. Within 2 days, the fine roots attached to the stem were collected and washed with running water to remove soil particles. For each collected sample, a small portion of fine roots were stained with Clorazol black E according to Yamato and Iwasaki (2002) to observe hyphal coils under a Table 1 Plant samples of Schizocodon soldanelloides var. magnus with sampling date and identified fungal isolates with DDBJ accession numbers Plant no.

Sampling date

Fungal isolate

Fungal identity

Accession no.

SC1

5/10/2008

SC1-1

Helotiales

AB598082

SC1-2

Helotiales

AB598083

SC2-1

Helotiales

AB598084

SC2-2

Helotiales

AB598085

SC2-3

Helotiales

AB598086

SC2-4

Helotiales

AB598087

SC2-5

Helotiales

AB598088

SC2

SC3

SC4

5/10/2008

5/10/2008

5/10/2008

SC2-6

Helotiales

AB598089

SC3-1

Helotiales

AB598090

SC3-2

Helotiales

AB598091

SC3-3

Helotiales

AB598092

SC3-4

Helotiales

AB598093

SC4-1

Helotiales

AB598094

SC4-2

Helotiales

AB598095

SC4-3 SC4-4

Helotiales Helotiales

AB598096 AB598097

SC5

5/10/2008

SC5-1

Helotiales

AB598098

SC6

25/4/2009

SC6-1

Helotiales

AB598099

SC7

25/4/2009

SC7-1

Helotiales

AB598100

SC8

17/6/2009

SC8-1

Helotiales

AB598101

SC9

17/6/2009

SC9-1

Oidiodendron AB598102

SC10

17/6/2009

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SC9-2

Oidiodendron AB598103

SC9-3

Helotiales

AB598104

SC9-4

Oidiodendron AB598105

SC9-5

Oidiodendron AB598106

SC9-6

Oidiodendron AB598107

SC10-1

Helotiales

AB598108

SC10-2

Helotiales

AB598109

differential interference contrast microscope (Eclipse 80iRT-DIC-1; Nikon, Tokyo, Japan). All collected samples were used for fungal isolation. The mycorrhizal fungi were isolated from fine roots according to Leake and Read (1991) with slight modifications as follows. In each sample, the washed fine roots were cleaned thoroughly by ultrasonication; then, a small portion of the fine roots, about 20 fine roots around 2 cm in length randomly collected, were put into a Petri dish with 1.0 ml sterilized distilled water. The root surface was scraped with the back side of a scalpel to peel off the epidermal cells. From the suspension of the epidermal cells, those having hyphal coils were selected under a dissecting microscope (Leica MZ125; Leica, Tokyo, Japan) to transfer onto 1.5% water agar plates. The plates were incubated at 25"C in the dark, and hyphal growth from the hyphal coil was observed after 2 or 3 days. The growing hyphae were transferred onto plates of potato dextrose agar (PDA) medium (Difco, Detroit, MI, USA) and incubated at 25"C in the dark to obtain fungal cultures. DNA was extracted from each of the obtained fungal cultures using a PrepMan Ultra Reagent (Applied Biosystem, Foster City, CA, USA) according to the manufacturer’s instructions. The internal transcribed spacer (ITS) region of nuclear ribosomal RNA gene (rDNA) was amplified by polymerase chain reaction (PCR) for each of the extracted DNA using the primer ITS1F and ITS4 (Gardes and Bruns 1993) in TaKaRa Ex Taq Hot Start Version (Takara Bio, Otsu, Japan). The PCR reaction mixture contained 2 ll template DNA, 0.75 units of Taq polymerase, 0.25 lM of each primer, 200 lM of each deoxyribonucleotide triphosphate, and 3 ll supplied PCR buffer in 30 ll of the total amount. The PCR program performed on Program Temp Control System PC-818S (Astec, Fukuoka, Japan) was as follows: initial denaturation step at 94"C for 2 min, followed by a step of 30 cycles at 94"C for 25 s, 55"C for 30 s, 72"C for 1 min, then a final elongation step at 72"C for 10 min. The amplicons were directly sequenced using BigDye Terminator v. 3.1 Cycle Sequencing Kit (Applied Biosystem) using ITS1F and ITS4 as sequencing primers. If the sequence data were not available by direct sequencing, the PCR products were cloned using a pGEM-T Easy Vector System I (Promega, Madison, WI, USA) according to the manufacturer’s instruction, and plasmid DNAs were extracted from the cloned products using MagExtractor Plasmid (Toyobo, Osaka, Japan). The cloned products were sequenced with T7 and SP6 promoter primers. The obtained data were subjected to BLAST searches (Altschul et al. 1997), and homologous data were downloaded from the GenBank database. To compare with previous ericoid mycorrhizal studies, some sequence data of ‘‘Hymenoscyphus ericae aggregate,’’ a fungal group containing many ericoid

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Fig. 1 Roots of Schizocodon soldanelloides var. magnus. a Intact thin roots growing from a thick root. b Intact thin root. c A thin root stained by Clorazol black E showing coiled hyphae in the epidermal cells. Bars a 1 cm; b 1 mm; c 50 lm

mycorrhizal fungi, were also downloaded from GenBank. For all sequenced and downloaded data, multiple sequence alignment was carried out using CLUSTAL X version 2 (Larkin et al. 2007). The aligned sequences data were analyzed by the neighbor-joining method (Saitou and Nei 1987), and the topology was tested with 1,000 bootstrap trials. The phylogenetic tree was drawn using TreeView (Page 1996). Roots of S. soldanelloides var. magnus were mostly found in litter layers. No other plant roots were found in the collected soil cores except those of F. crenata. The roots of S. soldanelloides var. magnus were easily distinguished from the ectomycorrhizal roots of F. crenata. For the root morphology and fungal colonization, representative images from one sample (SC6) are shown in Fig. 1. The root system consists of main thick roots and highly branched fine lateral roots (Fig. 1a,b). Diameter of the fine roots was mostly less than 100 lm. Hyphal coils were present in epidermal cells (Fig. 1c). The same kind of root morphology and fungal structure were confirmed in all the other samples. Mycorrhizal fungi were successfully isolated from peeled root epidermal cells on the water agar plate. From 10 individuals of S. soldanelloides var. magnus, 28 fungal isolates were obtained (Table 1). The ITS rDNA sequences of the fungal isolates were deposited to the DNA Data Bank of Japan (DDBJ) database with accession numbers

AB598082–AB598109. Most of the isolated fungi were identified as Helotiales based on the DNA sequences. In the phylogenetic analysis (Fig. 2), the fungal isolates were shown to be multi-lineages, and most of them were closely related to some fungi isolated from ecto-, ericoid, or arbutoid mycorrhiza. No fungi were included in the Hymenoscyphus ericae aggregate. The five other fungi collected from SC9 (see Table 1) were identified as Oidiodendron (Fig. 3). All the sequences formed a clade with O. maius including some ericoid mycorrhizal fungi. The diameter of the ericaceous fine root is less than 100 lm, which is referred to as a hair root (Smith and Read 2008). The diameter of the fine lateral root of S. soldanelloides var. magnus was also mostly less than 100 lm. These lateral roots were highly branched, which was a feature different from the hair root of ericaceous plants. Fungal hyphae grew from coiled hyphae in some root epidermal cells of S. soldanelloides var. magnus on water agar medium. This feature was also similar to that of ericoid mycorrhizae. The fungi grown from coiled hyphae were the subjects of this study, which avoided including other fungal endophytes and surface-colonizing fungi. Most of the obtained fungi were identified as belonging to Helotiales based on the ITS rDNA sequences. The fungi were shown to be multi-lineages in the phylogenetic

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Fig. 2 A neighbor-joining phylogenetic tree based on partial internal transcribed spacer (ITS) sequence of rDNA of Helotiales. The DNA sequences of cultured fungi obtained from roots of Schizocodon soldanelloides var. magnus in this study are examined with those in the Genbank database to infer the phylogenetic relationships. The tree is rooted to Sclerotinia sclerotiorum (Sclerotiniaceae). The obtained fungal DNA sequences are shown with fungal number. The identifiers of the fungal number are shown in Table 1. Bootstrap values are shown where they exceed 70% (1,000 replicates). A scale is shown to infer the evolutionary distances. Accession numbers are given for all sequences. ECM, ectomycorrhiza; ERM, ericoid mycorrhiza; ARM, arbutoid mycorrhiza

analysis, suggesting relatively low specificity in this plant– fungal relationship. Some of them were closely related to the fungi isolated from ectomycorrhizae in the phylogenetic analysis, which suggested these fungi could connect between the roots of S. soldanelloides var. magnus and the ectomycorrhizal root tips of surrounding trees. In many ericaceous plants, H. ericae has been identified as the fungal symbiont (Straker 1996). Vra˚lstad et al. (2000) showed that the genus Hymenoscyphus (Rhizoscyphus) forms a clade with genetically related fungi in a

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phylogenetic study based on the ITS rDNA region, and this grouping is called the Hymenoscyphus aggregate. The fungi in this aggregate are ecologically diverse; some are known to form ectomycorrhizae (Vra˚lstad et al. 2002). No fungi belonging to the H. ericae aggregate were detected in S. soldanelloides var. magnus in this study, but this result does not deny the possibility that further studies may detect the fungal group from Diapensiaceae. A few mycorrhizal fungi were found to belong to the genus Oidiodendron, and all of them were closely related

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Fig. 3 A neighbor-joining phylogenetic tree based on partial ITS sequence of rDNA of Oidiodendron. The DNA sequences of cultured fungi obtained from roots of Schizocodon soldanelloides var. magnus in this study are examined with those in Genbank database to infer the phylogenetic relations. The tree is rooted to Byssoascus striatosporus (Myxotrichaceae). The obtained fungal DNA sequences are shown

with fungal number; the fungal numbers are identified in Table 1. Bootstrap values are shown where they exceed 70% (1,000 replicates). A scale is shown to infer the evolutionary distances. Accession numbers are given for all sequences. ERM, ericoid mycorrhiza

to O. maius (see Fig. 3). Some species in Oidiodendron, such as O. maius, have been frequently recorded as ericoid mycorrhizal fungi in Ericaceae (Usuki et al. 2003; Addy et al. 2005; Bougoure and Cairney 2005). In this study, Oidiodendron fungi were isolated from only one sample of S. soldanelloides var. magnus (SC-9) among the ten samples examined, but they were dominantly detected in this sample (see Table 1). The structure of the mycorrhizae having coiled hyphae in epidermal cells of thin lateral roots is very similar to that of ericoid mycorrhizae. The affiliation of the mycorrhizal fungi Helotiales and Oidiodendron was also the same as the ericoid mycorrhizae. From the structure and fungal identities, we confirmed that ericoid mycorrhizae occur in the roots of S. soldanelloides.

generation of protein database search programs. Nucleic Acids Res 25:3389–3402 Angiosperm Phylogeny Group (2003) An update of the Angiosperm Phylogeny Group classification for the orders and families of flowering plants. Bot J Linn Soc 141:399–436 Bougoure DS, Cairney JWG (2005) Fungi associated with hair roots of Rhododendron lochiae (Ericaceae) in an Australian tropical cloud forest revealed by culturing and culture-independent molecular methods. Environ Microbiol 7:1743–1754 Gardes M, Bruns TD (1993) ITS primers with enhanced specificity for basidiomycetes: application to the identification of mycorrhizae and rusts. Mol Ecol 2:113–118 Harley JL (1989) The significance of mycorrhiza. Mycol Res 92:129–139 Harley JL, Harley EL (1987) A check-list of mycorrhiza in the British flora. New Phytol 105:1–102 Hesselman H (1900) Mycorrhizal formation in arctic plants (in Swedish). Bihang Till K Svensk Vet-Akad Handlingar 26:1–46 Janse JM (1897) Les endophytes radicaux de quelques plantes Javanaises. Ann Jardin Bot Buitenzorg 14:53–201 Kagawa A, Fujiyoshi M, Tomita M, Masuzawa T (2006) Mycorrhizal status of alpine plant communities on Mt. Maedake Cirque in the Japan South Alps. Polar Biosci 20:92–102 Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ, Higgins DG (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23:2947–2948 Leake JR, Read DJ (1991) Experiments with ericoid mycorrhiza. Methods Microbiol 23:435–459 Page RDM (1996) An application to display phylogenetic trees on personal computers. Comput Appl Biosci 12:357–358 Peterson RL, Massicotte HB, Melville LH (2004) Mycorrhizas: anatomy and cell biology. NRC Research Press, Ottawa Ro¨nblom K, Anderberg AA (2002) Phylogeny of Diapensiaceae based on molecular data and morphology. Syst Bot 27:383–395

Acknowledgments We express our gratitude to Yazu-cho in Tottori Prefecture for the permission to collect the plant samples. This study was supported by Global COE Program ‘‘Advanced utilization of fungus/mushroom resources for sustainable society in harmony with nature’’ from the Ministry of Education, Culture, Sports, Science and Technology of Japan.

References Addy HD, Piercey MM, Currah RS (2005) Microfungal endophytes in roots. Can J Bot 83:1–13 Altschul SF, Madden TL, Schaffer AA, Zhang J, Zhang Z, Miller W, Lipman DJ (1997) Gapped BLAST and PSI-BLAST: a new

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430 Saitou N, Nei M (1987) The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4:406–425 Scott PJ, Day RT (1983) Diapensiaceae: a review of the taxonomy. Taxon 32:417–423 Smith SE, Read DJ (2008) Mycorrhizal symbiosis, 3rd edn. Academic Press, New York Straker CJ (1996) Ericoid mycorrhiza: ecological and host specificity. Mycorrhiza 6:215–225 Stribley DP, Read DJ (1980) The biology of mycorrhiza in the Ericaceae. VII. The relationship between mycorrhizal infection and the capacity to utilize simple and complex organic nitrogen sources. New Phytol 86:365–371 Usuki F, Abe JP, Kakishima M (2003) Diversity of ericoid mycorrhizal fungi isolated from hair roots of Rhododendron

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Mycoscience (2011) 52:425–430 obtusum var. kaempferi in a Japanese red pine forest. Mycoscience 44:97–102 Vra˚lstad T, Fossheim T, Schumacher T (2000) Piceirhiza bicolorata—the ectomycorrhizal expression of the Hymenoscyphus ericae aggregate? New Phytol 145:549–563 Vra˚lstad T, Schumacher T, Taylor AFS (2002) Mycorrhizal synthesis between fungal strains of the Hymenoscyphus ericae aggregate and potential ectomycorrhizal and ericoid hosts. New Phytol 153:143–152 Yamato M, Iwasaki M (2002) Morphological types of arbuscular mycorrhizal fungi in roots of understory plants in Japanese deciduous broadleaved forests. Mycorrhiza 12:291–296 Zangaro W, Nisizaki SMA, Domingos JCB, Nakano EM (2003) Mycorrhizal response and successional status in 80 woody species from south Brazil. J Trop Ecol 19:315–324

Mycoscience (2011) 52:431–435 DOI 10.1007/s10267-011-0122-7

NOTE

Molecular breeding of a novel Coprinopsis cinerea strain possessing a heterologous laccase gene, lccK, driven by a constitutive promoter Hajime Muraguchi • Manami Kondoh Yasuhiro Ito • Sonoe O. Yanagi

•

Received: 21 February 2011 / Accepted: 6 May 2011 / Published online: 21 May 2011 ! The Mycological Society of Japan and Springer 2011

Abstract Genomic DNA encoding the Pleurotus ostreatus LccK laccase was fused with the Coprinopsis cinerea b-tubulin promoter and terminator, and introduced into a C. cinerea strain. Linkage analysis, native PAGE separations, substrate specificity investigations and expression profiling indicated that C. cinerea transformants secrete P. ostreatus LccK, suggesting that the introns of the lccK gene are correctly spliced and the signal peptide for secretion is functional in C. cinerea. Transformants constitutively expressing laccase may be useful for the degradation of aromatic compounds. Keywords Heterologous expression ! Pleurotus ostreatus ! Laccase secretion

Laccases (EC 1.10.3.2) are copper-containing polyphenol oxidases that are widely found in plants, fungi and bacteria. Although laccases are narrowly defined as oxidoreductases of p-diphenol, they exhibit low substrate specificity and can oxidize a wide range of aromatic compounds, with the concomitant reduction of molecular oxygen to water. When laccases oxidize phenolitic hydroxyl substrates, the radicals produced can polymerize substrates to produce pigments

H. Muraguchi (&) ! M. Kondoh ! S. O. Yanagi Department of Biotechnology, Faculty of Bioresource Sciences, Akita Prefectural University, Shimoshinjo-nakano, Akita 010-0195, Japan e-mail: [email protected] Y. Ito National Food Research Institute, National Agriculture and Food Research Organization (NARO), 2-1-12 Kannondai, Tsukuba, Ibaraki 305-8642, Japan

(Bavendamm 1928), allowing us to easily detect laccase activity (Tanabe et al. 1989). The laccase secreted by P. ostreatus mycelia has been purified and characterized (Okamoto et al. 2000). Cloning and characterization of the P. ostreatus lccK gene suggested that this encodes the major extracellular laccase secreted by mycelia of P. ostreatus K16-2 (Okamoto et al. 2003). Pleurotus ostreatus laccase is constitutively expressed and secreted into culture media under normal cultural conditions (Okamoto et al. 2000). In contrast, laccase secretion by most wild-type strains of C. cinerea is not observed under normal cultural conditions, but appears to occur in response to environmental stresses. The different expression profiles of laccases in these basidiomycetes allowed us to detect heterologous expression of P. ostreatus laccase in C. cinerea. This has been suggested in a previous report, where P. ostreatus genomic DNA fragments digested with the restriction enzyme Eco RI were introduced into C. cinerea without cloning (Okamoto et al. 1995). To express the P. ostreatus laccase, LccK, in C. cinerea, the C. cinerea b-tubulin promoter and terminator were fused to the genomic fragment encoding LccK. Genomic DNA was extracted from strain 39 of P. ostreatus (Mori Sangyo Co., Ltd., Gunma, Japan) for amplification of the laccase gene. Primers were designed based on the sequence of the lccK gene (Genbank accession no.: AB089612). Primer sequences used are listed in Table 1. The promoter of the C. cinerea b-tubulin gene (Genbank accession no.: AB000116) was amplified using primers 1 and 2, and the terminator was amplified using primers 5 and 6, from pPHT1; this carries a bacterial hygromycin B resistance gene fused to the promoter and terminator regions of the C. cinerea b-tubulin gene (Cummings et al. 1999). Amplifications were performed using PfuUltra high-fidelity DNA

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432 Table 1 Primer sequences used

The HindIII site is indicated by underlining

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Primer name

Sequence

#1-HindIII-t(P)-For

GCGCAAGCTTCATTTAAACGGCTTC HindIII

#2-POt(P)-Rev

TGCGCCTGGAAACATGCTGGGAACGCGAGG

#3-t(P)PO-For

CCTCGCGTTCCCAGCATGTTTCCAGGCGCA

#4-t(T)PO-Rev

GAACTAACGAATCATTTAGGACGGAACGAT

#5-POt(T)-For

ATCGTTCCGTCCTAAATGATTCGTTAGTTC

#6-t(T)HindIII-Rev

GCGCAAGCTTCAATATTCATCTCTC HindIII

polymerase (Stratagene, La Jolla, CA) following the manufacturer’s instructions. A genomic region coding for P. ostreatus laccase was amplified using primers 3 and 4, based on the P. ostreatus K16-2 lccK sequence (Okamoto et al. 2003). DNA fragments were separated on an 0.8% agarose gel, and the desired fragments were purified using a GENECLEAN II kit (BIO 101, Vista, CA). Purified fragments were used to provide template DNA for subsequent amplifications. The promoter and the laccase coding region were mixed and fused via amplification with primers 1 and 4, and then gel-purified. This fused fragment and the terminator were then mixed and fused via amplification with primers 1 and 6, and gel-purified. The final fusion product was digested with HindIII, gel-purified and ligated into the HindIII site of the pBACTZ vector that was constructed by insertion of the C. cinerea trp1 gene into the pBeloBAC11 vector (Muraguchi et al. 2005). Ligation samples were used to electroporate 20 ll of electro-competent Escherichia coli DH10B cells; transformed cells were selected as described previously (Muraguchi et al. 2005), yielding pB-glccK. Protoplasts were prepared from oidia of C. cinerea trpstrain 292, as previously described (Zolan et al. 1992), with the modification of 50 mg/ml of lysing enzyme (L1412, Sigma-Aldrich, St. Louis, MO) instead of Novozyme 234. Transformation of protoplasts was performed as previously described (Binninger et al. 1987). One hundred microliters of protoplasts (109/ml) was transformed by 10 ll of pB-glccK isolated with alkaline lysis. Three trp? transformants were obtained using pB-glccK and examined for expression of laccase using the Bavendamm test (Bavendamm 1928), involving the inoculation of mycelia onto MYG medium (Rao and Niederpruem 1969) containing 1 mM guaiacol. Two of the transformants exhibited the red phenotype signifying laccase activity, as in the original P. ostreatus strain (Fig. 1). To examine the dominance of laccase expression in C. cinerea, transformants were crossed with a wild-type monokaryon to obtain dikaryons. The resulting dikaryons also showed the red phenotype when grown on guaiacol medium (Fig. 1). Since phenol oxidase activity has been linked to fruit-body

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Fig. 1 Laccase activity of transformants in guaiacol medium. Mycelia of the indicated strains were grown on MYG medium containing guaiacol in order to monitor laccase activity. KF32 is a wild-type strain. 292 was transformed with the LccK gene cassette inserted into the pBACTZ vector, yielding TF2 and TF3 transformants. Red coloration indicates laccase activity. The dikaryons formed by crossing transformants with KF32 exhibit the dominant phenotype with laccase activity

formation in C. cinerea (Vnenchak and Schwalb 1989), we observed fruiting of the dikaryons. These were able to form normal fruit bodies, suggesting that the laccase activity of LccK does not affect physiological processes in C. cinerea reproduction. To confirm that the laccase activity of transformants was due to expression of the P. ostreatus lccK gene integrated into the C. cinerea genome, we firstly examined whether the red phenotype in F1 progeny cosegregates with insertion of the vector sequence into the genome. In F1 progeny derived from a cross between transformant TF2 and a wildtype, the red phenotype was segregated (Fig. 2a). Ten white strains and ten red strains were selected from the F1 progeny; their genomic DNAs were extracted and subjected to Southern analysis. Five micrograms of genomic DNA was digested with HindIII and fractionated by electrophoresis in 0.8% agarose gels. Fractionated fragments were transferred onto nylon membranes (Hybond-N?, Amersham) by capillary blotting, using 0.4 N NaOH as the transfer solution, and subjected to Southern hybridization (Sambrook and Russell 2001). Probe labeling and hybridization were performed according to the Gene Images (GE

Mycoscience (2011) 52:431–435

433

Fig. 2 Cosegregation in F1 progeny. a Segregation of the red phenotype in F1 progeny derived from crossing TF2 and KF32. b Southern analysis of 10 white progeny and 10 red progeny using pBeloBAC11 as a probe. Lanes 1–10 white progeny, lanes 11–20 red progeny. M indicates a size marker, k DNA digested with HindIII. The hybridized bands in the marker lane indicate 23 and 4.4 kb

Healthcare) instructions. The DNA of the pBeloBAC11 vector, which was part of pBg-lccK and did not contain C. cinerea DNA, was labeled and used as a probe to detect the introduced vector DNA (Muraguchi et al. 2005). All white strains showed no signal, and all red strains showed signals on the Southern blots (Fig. 2b), suggesting that the red phenotype is due to integration of pBeloBAC11 carrying the P. ostreatus lccK gene. Secondly, we used native polyacrylamide gel electrophoresis (PAGE) (Davis 1965) to examine the properties of laccases secreted by mycelia into liquid media. Five agar cubes containing mycelia were inoculated into 20 ml of liquid MYG medium and incubated for 3 days at 28"C. Four milliliters of the liquid medium was dialyzed against 200 ml of Tris-HCl buffer (pH 6.8) at 4"C overnight. Dialyzed solutions were concentrated 40 times using a Mizubutori-kun kit (Atto, Tokyo, Japan). Samples were subjected to native PAGE (Fig. 3). A single band with laccase activity was detected in the P. ostreatus sample, as reported previously (Okamoto et al. 2000). The laccase secreted by TF2 had the same mobility as the P. ostreatus laccase, suggesting that TF2 secreted the P. ostreatus laccase, LccK. We isolated tsh (temperature-sensitive defect in hyphal growth) mutants (Muraguchi et al. 2008) and found that when the temperature was increased from 28 to 42"C, the tsh3-1 mutant (324) expressed the red phenotype in MYG medium containing guaiacol. We therefore used native PAGE to examine the laccases secreted by the tsh3-1

Fig. 3 Native PAGE analysis of secreted laccases. Laccases secreted into MYG liquid media were concentrated and subjected to native PAGE analysis. Lane 1 P. ostreatus, lane 2 C. cinerea 324 (tsh3-1), lane 3 C. cinerea TF2. Strain C. cinerea 324 (tsh3-1) is a temperature-sensitive mutant defective in hyphal growth at 42"C, which secretes intact laccase during incubation at 42"C. The laccase of 324 (tsh3-1) was collected after overnight culturing at 42"C

mutant using native PAGE. The tsh3-1 mutant secreted a single laccase with a mobility different from either TF2 or the P. ostreatus strain (Fig. 3). Although we could not rule out the possibility that TF2 secreted another endogenous laccase with the same mobility as the P. ostreatus laccase, TF2 secreted a laccase different from that secreted by strain 324 under heat-shock stress, and it seems likely that it secreted the LccK laccase. Thirdly, we examined the substrate specificity of the secreted laccases using 15 different substrates (Table 2). Mycelia of P. ostreatus, TF2, and 324 (tsh3-1) were

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inoculated onto plates of each substrate, and the pigments produced by secreted laccases were observed. To encourage secretion of the intact C. cinerea laccase from the tsh3-1 mutant, culture plates were transferred from 28 to 42"C. At 28"C, the substrate specificity of TF2 was the same as that of the P. ostreatus strain. After incubation at 42"C for 24 h, the pigments produced by TF2 at 28"C disappeared from some substrates (Table 2). This may have been due to degradation of the pigments by proteins secreted by TF2. The substrate specificity of the laccase at 28"C suggests that the laccase secreted by TF2 is the P. ostreatus laccase, LccK. Finally, the physiological modes of secretion of laccases from TF2, 292 and P. ostreatus strains were observed in liquid media for 40 days. Five agar cubes with mycelia were inoculated into 20 ml of MYG liquid medium and cultured. One hundred microliters of liquid medium was sampled and added to 1 ml of reaction mixture containing 1 mM guaiacol and 50 mM of potassium phosphate buffer (pH 6.5). The mixture was incubated at 30"C for 30 min in the dark, and absorbance at 470 nm was measured. The laccase activity of TF2 peaked at 4 days after inoculation and then decreased. The laccase activity of P. ostreatus increased for 10 days and peaked at double the level of TF2 (Fig. 4). The laccase activity of P. ostreatus then declined and had disappeared by 30 days, while the activity

of TF2 was maintained at low levels until 40 days. The expression property of TF2 suggested that the secreted laccase was constitutively expressed by the b-tubulin promoter. Transformants with constitutive expression of laccase may be useful for the degradation of aromatic compounds in the future. Thus, four lines of evidence, linkage analysis, native PAGE separations, substrate specificity investigations and expression profiling strongly suggested that the laccase activities of transformants were due to the introduced genomic lccK gene. Coprinopsis cinerea laccases and their genes have been extensively characterized, and the genome project has revealed that this fungus contains a large family of laccases (Kilaru et al. 2006). We were not able to rule out the possibility that TF2 secreted an endogenous laccase with the same mobility as P. ostreatus laccase. This could be caused by the integration of the introduced DNA fragment containing the b-tubulin promoter into the 50 region of the endogenous laccase. Further study using antibodies against LccK is required to confirm that the laccase secreted by TF2 is the P. ostreatus laccase, LccK. If the laccase secreted by TF2 is indeed the P. ostreatus laccase, LccK, this suggests that the introns of the P. ostreatus lccK gene are correctly spliced and the signal peptide for secretion is functional in C. cinerea. Our results are consistent with previous research in which P. ostreatus

Table 2 Substrate specificity of laccases secreted by mycelia Substrate

P. ostreatus TF2

324

28"C 42"C 28"C 42"C 28"C 42"C Guaiacol

?

?

?

-*

-

?

a-Naphtol

?

?

?

?

-

-

Catecol

?

?

?

?

-

-

Pyrogallol

?

?

?

?

-

?

Vanillin

-

-

-

-

-

?

Vanillic acid

?

?

?

-*

-

?

Gallic acid

?

?

?

?

-

?

3,4-Dihydroxy cinnamic acid

?

?

?

?

-

?

p-Hydroxy benzoic acid

-

-

-

-

-

?

Tannic acid

?

?

?

-*

-

?

p-Hydroxy benzaldehyde

-

-

-

-

-

?

4-Hydroxy-3methoxycinnamic acid

?

?

?

?

-

?

p-Coumaric acid

-

-

-

-

-

?

Protocatechuic acid

?

?

?

-*

-

?

Syringic acid

?

?

?

?

-

?

Mycelia were inoculated onto plates containing different substrates (1 mM) and incubated for 3 days. After incubation at 28"C, plates were transferred to 42"C and incubated for 1 or 2 days ‘‘?’’ indicates coloration based on laccase activity ‘‘*’’ at 42"C may be due to degradation of pigments produced at 28"C

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Fig. 4 Changes in laccase activity in liquid culture. Mycelia of P. ostreatus (filled triangle), C. cinerea TF2 (filled square) and 292 (filled diamond) were cultured in MYG liquid media at 28"C for 40 days. Laccase activities in the liquid media were measured daily

Mycoscience (2011) 52:431–435

genomic fragments digested with Eco RI were introduced into C. cinerea, and C. cinerea strains that appeared to express the P. ostreatus lccK emerged (Okamoto et al. 1995). It has also been reported that the heterologous promoter and terminator function in C. cinerea to express native laccase genes with introns (Kilaru et al. 2006) and heterologous manganese (II) peroxidase (MnP) cDNA (Ogawa et al. 1998). It would appear that C. cinerea could be used as a host to express genes in heterologous genomic DNAs. Since some mushrooms are extremely difficult to grow in pure culture, functional analyses using easily cultivated hosts such as C. cinerea seem significant for both basic and practical purposes.

References ¨ ber das Vorkommen den Nachweis von Bavendamm W (1928) U Oxydasen bei holzzersto¨renden. Pilzen Z Pflanzenkrank Pflanzenschutz 38:257–276 Binninger DM, Skrzynia C, Pukkila PJ, Casselton LA (1987) DNAmediated transformation of the basidiomycete Coprinus cinereus. EMBO J 6:835–840 Cummings WJ, Celerin M, Crodian J, Brunick LK, Zolan ME (1999) Insertional mutagenesis in Coprinus cinereus: use of a dominant selectable marker to generate tagged, sporulation-defective mutants. Curr Genet 36:371–382 Davis BJ (1965) Disc electrophoresis-II. Method and application to human serum proteins. Ann N Y Acad Sci 121:404–427 Kilaru S, Hoegger PJ, Majcherczyk A, Burns C, Shishido K, Bailey A, Foster GD, Ku¨es U (2006) Expression of laccase gene lcc1 in Coprinopsis cinerea under control of various basidiomycetous promoters. Appl Microbiol Biotechnol 71:200–210

435 Muraguchi H, Kamada T, Yanagi SO (2005) Construction of a BAC library of Coprinus cinereus. Mycoscience 46:49–53 Muraguchi H, Abe K, Nakagawa M, Makamura K, Yanagi SO (2008) Identification and characterisation of structural maintenance of chromosome 1 (smc1) mutants of Coprinopsis cinerea. Mol Genet Genomics 280:223–232 Ogawa K, Yamazaki T, Hasebe T, Kajiwara S, Watanabe A, Asada Y, Shishido K (1998) Molecular breeding of the basidiomycete Coprinus cinereus strains with high lignin-decolorization and -degradation activities using novel heterologous protein expression vectors. Appl Microbiol Biotechnol 49:285–289 Okamoto K, Sakamoto H, Sakai T, Yanagi SO (1995) Phenoloxidase activity in Coprinus cinereus due to protoplasting and insertion treatment with Pleurotus ostreatus DNA. Biotechnol Lett 17:1127–1130 Okamoto K, Yanagi SO, Sakai T (2000) Purification and characterization of extracellular laccase from Pleurotus ostreatus. Mycoscience 41:7–13 Okamoto K, Ito Y, Shigematsu I, Yanagi SO, Yanase H (2003) Cloning and characterization of a laccase gene from the whiterot basidiomycete Pleurotus ostreatus. Mycoscience 44:11–17 Rao PS, Niederpruem DJ (1969) Carbohydrate metabolism during morphogenesis of Coprinus lagopus (sensu Buller). J Bacteriol 100:1222–1228 Sambrook J, Russell DW (2001) Molecular cloning: a laboratory manual, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor Tanabe N, Sagawa I, Ohtsubo K, Iijima Y, Yanagi SO (1989) Comparison of phenoloxidase activities during the cultivation of several basidiomycetes. Agaric Biol Chem 53:3061–3063 Vnenchak P, Schwalb MN (1989) Phenol oxidase activity during development of Coprinus cinereus. Mycol Res 93:546–548 Zolan ME, Crittenden J, Heyler NK, Seitz LC (1992) Efficient isolation and mapping of rad genes of the fungus Coprinus cinereus using chromosome-specific libraries. Nucleic Acids Res 20:3993–3999

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Mycoscience (2011) 52:436–438 DOI 10.1007/s10267-011-0124-5

NOTE

Two noteworthy Phallus from southern Brazil Vagner G. Cortez • Iuri G. Baseia Rosa Mara B. da Silveira

•

Received: 3 November 2009 / Accepted: 10 May 2011 / Published online: 8 June 2011 ! The Mycological Society of Japan and Springer 2011

Abstract During a survey of gasteroid fungi from the state of Rio Grande do Sul, in southern Brazil, two noteworthy species of the genus Phallus were identified: P. duplicatus (new record from Brazil) and P. granulosodenticulatus. The latter is a poorly known species described by Braun in 1932 that was recently recollected, and its taxonomy is discussed based on the examination of fresh and type specimens. Keywords Gasteromycetes ! Neotropical fungi ! Phalloids ! Phallales ! Phallomycetidae Phallaceae Corda (Basidiomycota) comprises gasteroid fungi that have basidiomata with a phallic shape, and gelatinous gleba which usually spread an unpleasant smell of rotten meat that attracts insects for basidiospore dispersal. Phalloids (or ‘‘stinkhorns,’’ as they are called) are currently classified in Phallales E. Fisch., subclass Phallomycetidae K. Hosaka et al. (Hosaka et al. 2006). In Brazil, the following genera are known to occur: Phallus Junius ex L. (= Aporophallus Mo¨ller, Dictyophora

V. G. Cortez (&) Universidade Federal do Parana´, R. Pioneiro 2153, Palotina, PR 85950-000, Brazil e-mail: [email protected] I. G. Baseia Departamento de Botaˆnica, Ecologia e Zoologia, Universidade Federal do Rio Grande do Norte, Natal, RN 59072-970, Brazil R. M. B. da Silveira Departamento de Botaˆnica, Universidade Federal do Rio Grande do Sul, Av. Bento Gonc¸alves 9500, Porto Alegre, RS 91501-970, Brazil

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Desv., Itajahya Mo¨ller, Ityphallus Fr.), Mutinus Fr., and Staheliomyces E. Fisch. (Baseia et al. 2006). Southern Brazilian phalloids were studied by Mo¨ller (1895); Braun (1932) and Rick (1961). Some species were also reported in macrofungi surveys (Guerrero and Homrich 1999; Meijer 2006; Cortez et al. 2008) and broader revisions of phalloids (Lloyd 1909; Wright 1949, 1960; Dring 1980). Recently, Trierveiler-Pereira et al. (2009) emended the description of Phallus glutinolens (Mo¨ller) Kuntze, known only from southern Brazil and Argentina, and provided a key for seven Brazilian Phallus. In addition, new species (Baseia et al. 2003; Baseia and Calonge 2005) and records (Leite et al. 2007; Ottoni et al. 2010) from northeastern Brazil have increased the knowledge of this group of fungi. In the present paper, we discuss the record of two noteworthy members of the genus Phallus, collected during a survey of the gasteroid fungi from Rio Grande do Sul State, in southern Brazil (Cortez et al. 2008, 2009, 2011a, b). Specimens collected by the authors are kept at the ICN herbarium (Universidade Federal do Rio Grande do Sul, Instituto de Biocieˆncias) and specimens from the PACA herbarium (Fungi Rickiani) were checked. Colors are from Kornerup and Wanscher (1978). Phallus duplicatus Bosc, Mag. Ges. Naturf. Fr. Berl. 5: 86, 1811. Fig. 1a : Dictyophora duplicata (Bosc) E. Fisch. in Sacc., Syll. Fung. 7: 6, 1888. Eggs not seen. Basidioma up to 60 mm high when expanded. Receptacle 18 9 25 mm, campanulate, surface strongly rugose, color olive (3F8) due to the presence of gleba, apex covered with remnants of the peridium, perforated, margin appendiculate with a thin and yellowishwhite (3A2) indusium, about 16 mm in length. Pseudostipe

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Fig. 1 Basidiomata and basidiospores of Phallaceae. a Phallus duplicatus b P. granulosodenticulatus Fig. 2 Phallus granulosodenticulatus. Photo: J. M. Oliveira

up to 54 9 13 mm, cylindrical, white (3A1), surface spongy, hollow; base with a saccate volva, 22 9 26 mm, color reddish white (11A2) to pale red (11A3), with a few basal rhizomorphs attached to the soil. Gleba olive (3F8), slimy, fetid. Basidiospores 3.5–4.2 9 1–1.5 lm, cylindrical, hyaline, smooth and thin-walled. Examined materials Brazil, Rio Grande do Sul State, Porto Alegre, UFRGS, solitary on soil among grasses, 5 February 2009, leg. M.A. Sulzbacher & V.G. Cortez 003/09 (ICN); Sa˜o Leopoldo, 25 February 1932, leg. B. Braun (PACA 15049, 15052). Remarks The fungus is similar to P. indusiatus Vent., but differs in the shorter and less developed indusium, in contrast to the large indusium of P. indusiatus, which also presents broader basidiospores (Guzma´n et al. 1990; Calonge 2005). Phallus duplicatus is a new record from Brazil and probably South America. Phallus granulosodenticulatus B. Braun, Relat. Gin. Anch. 1932: 12, 1932. Figs. 1b, 2 Eggs subglobose to ovoid, 30–34 9 20–28 mm, grayish yellow (4B3) to grayish orange (5B4). Basidiomata 90 mm high when expanded. Receptacle 20 9 10 mm, campanulate, surface granular to slightly rugulose, color grayish green (28C4), deep green (29E8) to dark green (29F8), apex perforated, margin cogged to uneven. Pseudostipe 70 9 9 mm, subcylindrical, white (28A1) to greenish white (28A2), surface spongy, hollow; base with a saccate volva, 36 9 18 mm, grayish orange (5B4), bearing abundant rhizomorphs attached to the soil. Gleba deep green

(29E8), slimy, fetid. Basidiospores 3.8–5 9 2–3 lm, ellipsoid, hyaline, smooth and with slightly thickened walls, guttulate. Examined specimens Brazil, Rio Grande do Sul State, Santa Maria, UFSM, in meadow, 16 June 2008, leg. V.G. Cortez 117/08 (ICN). Sa˜o Leopoldo, 1929–1932, leg. B. Braun (PACA 15042, 15043, 15044, 15046, 15047-holotype, 15048, 15050, 15051). Remarks This species was described by Braun (1932) in a local journal, and it was later included in the posthumous checklist of Rick’s gasteromycetes (Rick 1961). Nevertheless, the species remained unknown to most mycologists who revised South American phalloids, and its name was forgotten. In his obscure paper, Braun (1932) described and discussed in detail the new species, and provided an excellent description and photos that allowed its easy recognition. Since the original description, no other collections of this fungus were reported. The main features of the species are the granular to rugulose surface of the fertile portion of the receptacle, white pseudostipe and campanulate receptacle with a conspicuously cogged margin (Braun 1932). This species is macroscopically similar to the North American P. ravenelii Berk. & M.A. Curtis, but the latter has larger basidiomata, a smooth and welldefined pileus margin, and cylindrical basidiospores (1–1.5 lm width, Smith 1951). Some collections (PACA 15042, 15043, 15044, 15046) were identified by Braun himself as P. ravenelii before the formal description of

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P. granulosodenticulatus. With the re-examination of Braun’s authentic materials and an additional collection, we consider it a good Phallus species. Acknowledgments The authors thank to the curator of the PACA herbarium for loaning specimens. Professor Francisco D. Calonge is thanked for providing useful literature. CNPq is acknowledged for financial support.

References Baseia IG, Calonge FD (2005) Aseroe¨ floriformis, a new species with a sunflower receptacle. Mycotaxon 91:169–172 Baseia IG, Gibertoni T, Maia LC (2003) Phallus pygmaeus, a new minute species from a Brazilian tropical rain forest. Mycotaxon 85:77–79 Baseia IG, Maia LC, Calonge FD (2006) Notes on phallales in the neotropics. Bol Soc Micol Madrid 30:87–93 Braun B (1932) Estudo sobre as Phalloideas Riograndenses. Relat Gin Anchieta (Porto Alegre) 1932:5–28 Calonge FD (2005) A tentative key to identify the species of Phallus. Bol Soc Micol Madrid 29:9–18 Cortez VG, Baseia IG, Silveira RMB (2008) Gasteromicetos (Basidiomycota) no Parque Estadual de Itapua˜, Viama˜o, Rio Grande do Sul, Brasil. Rev Bras Bioc 6:291–299 Cortez VG, Baseia IG, Silveira RMB (2009) Gasteroid mycobiota of Rio Grande do Sul, Brazil: Tulostomataceae. Mycotaxon 108:365–384 Cortez VG, Baseia IG, Silveira RMB (2011a) Gasteroid mycobiota of Rio Grande do Sul, Brazil: Lysuraceae (Basidiomycota). Acta Scient Biol Sci 33:87–92 Cortez VG, Baseia IG, Silveira RMB (2011b) Gasteroid mycobiota of Rio Grande do Sul, Brazil: Calvatia, Gastropila and Langermannia. Kew Bull (in press) Dring DM (1980) Contributions towards a rational arrangement of the Clathraceae. Kew Bull 35:1–96

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Mycoscience (2011) 52:436–438 Guerrero RT, Homrich MH (1999) Fungos macrosco´picos comuns no Rio Grande do Sul—Guia para identificac¸a˜o, 2nd edn. UFRGS, Porto Alegre Guzma´n G, Montoya L, Bandala VM (1990) Las especies y formas de Dictyophora (Fungi, Basidiomycetes, Phallales) en Mexico y observaciones sobre su distribucion en America Latina. Acta Bot Mex 9:1–11 Hosaka K, Bates ST, Beever RE, Castellano ME, Colgan W, Domı´nguez L, Nouhra ER, Geml J, Giachini AJ, Kenney SR, Simpson NB, Spatafora JW, Trappe JM (2006) Molecular phylogenetics of the gomphoid-phalloid fungi with an establishment of the new subclass Phallomycetidae and two new orders. Mycologia 98:949–959 Kornerup A, Wanscher JH (1978) Methuen handbook of colour, 3rd edn. Eyre Methuen, London Leite AG, Silva BDB, Araujo RS, Baseia IG (2007) Espe´cies raras de Phallales (Agaricomycetidae, Basidiomycetes) no Nordeste do Brasil. Acta Bot Bras 21:119–124 Lloyd CG (1909) Synopsis of the known Phalloids. Cincinnati Meijer AAR (2006) Preliminary list of the macromycetes from the Brazilian State of Parana´. Bol Mus Bot Mun (Curitiba) 68:1–59 Mo¨ller A (1895) Brasilische Pizlblumen. Gustav Fischer, Jena Ottoni T, Silva BDB, Fazolino EP, Baseia IG (2010) Phallus roseus, first record from the neotropics. Mycotaxon 112:5–8 Rick J (1961) Basidiomycetes Eubasidii in Rio Grande do Sul– Brasilia 6. Iheringia Se´r Bot 9:451–479 Smith AH (1951) Puffballs and their allies in Michigan. University of Michigan, Ann Arbor Trierveiler-Pereira L, Loguercio-Leite C, Calonge FD, Baseia IG (2009) An emendation of Phallus glutinolens. Mycol Prog 8:377–380 Wright JE (1949) Los Gasteromycetes del Museo Argentino de Ciencias Naturales Bernardino Rivadavia, I Phallales. Com Inst Nac Inv Cienc Nat (Cienc Bot) 1:3–15 Wright JE (1960) Notas sobre Faloideas Sud y Centroamericanas. Lilloa 30:339–359

Mycoscience (2011) 52:439 DOI 10.1007/s10267-011-0161-0

INDEX

Index to new names, combinations and emendation appearing in Mycoscience 52 (6)

! The Mycological Society of Japan and Springer 2011

Crinipellis rhizomorphica Har. Takah., sp. nov. …………………………………………………………………………. 392 Psilocybe capitulata Har. Takah., sp. nov. ………………………………………………………….…………………… 397 Tomentella agereri Yorou sp. nov. ………………………………………………………………………………………. 366 Tomentella maroana Yorou sp. nov. …………………………………………………………………………………….. 368

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