Mycol Progress (2010) 9:575–583 DOI 10.1007/s11557-010-0665-6

ORIGINAL ARTICLE

Xeromphalina setulipes (hygrophoroid clade, Agaricales), a new Mediterranean species Fernando Esteve-Raventós & Gabriel Moreno & Jose Luis Manjón & Pablo Alvarado

Received: 13 January 2010 / Revised: 11 February 2010 / Accepted: 15 February 2010 / Published online: 7 March 2010 # German Mycological Society and Springer 2010

Abstract A new Xeromphalina species from Mediterranean evergreen forests of the southern part of the Iberian Peninsula is described. It is characterized by its habitat on soil, arcuate-decurrent lamellae, trama hyphae which turn distinctly orange-brown to red-brown in KOH solution, thick-walled caulocystidia with an attenuate apex reminding setulae, and versiform to irregularly cylindrical cheilocystidia, which are conspicuously branched forming coralloid excrescences. LM microphotographs illustrate its diagnostic features. Additionally, two nuclear ribosomal DNA regions from this new taxon were sequenced and compared with homologue sequences from existing species in the genus Xeromphalina, supporting its recognition as a new species. Keywords Basidiomycetes . Tricholomatales . Mediterranean mycobiota . Taxonomy . New species

Introduction During field work carried during the autumn of 2005 in the southern part of the Iberian Peninsula, we discovered a xeromphalinoid agaric which was later determined as an unknown species of the genus Xeromphalina; it is presented here as X. setulipes, because of its peculiar protruding caulocystidia with a shape reminiscent of thick-walled setulae, similar to those present in many species of the

F. Esteve-Raventós (*) : G. Moreno : J. L. Manjón : P. Alvarado Dpto. de Biología Vegetal (Botánica), Fac. Biología, Universidad de Alcalá de Henares, 28871 Madrid, Spain e-mail: [email protected]

genera Inonotus and Phellinus; this particular feature is only known in a few species of this genus. In addition, X. setulipes shows trama hyphae which turn dark orangebrown to red-brown in KOH solutions, and this combination of characters makes it very distinct. About eight species (and some varieties) of Xeromphalina are known from European countries (Antonín and Noordeloos 2004), but from these, only two have been unambiguously and regularly recorded from Europe (X. campanella (Batsch) Maire and X. cauticinalis (With.) Kühner & Maire (= X. fellea Maire & Malençon). A similar situation has occurred in the Iberian Peninsula, where a third species, known as X. junipericola G. Moreno & Heykoop, was described only a few years ago by Moreno and Heykoop (1996), growing on Juniperus thurifera rotten wood and showing a peculiar pigment composition. Finally, a fourth species, X. minutissima, was invalidly published by Esteve-Raventós (1995), and because of the very scanty material (only one tiny specimen), the author considered that more collections were needed to proceed to its validation. In spite of the few known taxa, Xeromphalina is, however, a rather well-studied genus in Europe, where it has been the object of some monographic continental and local contributions (Klán 1984; Gulden 1992; Watling and Turnbull 1998; Bon 1999; Ludwig 2001; Antonín and Noordeloos 2004; Noordeloos 2008). Also, monographic studies have been made by Smith (1953), Miller (1968), and Redhead (1988) on North American species (the last also dealing with the northern Eurasian taxa), and some additions were also contributed from the South American (Singer 1965; Redhead and Halling 1987) and Asian continents (Horak 1979). At the present time, following the phylogenetic study of Matheny et al. (2007), Xeromphalina is considered a

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member of the “hygrophoroid clade”, close to the gilled genera Sarcomyxa P. Karst. and Phyllotopsis E.-J. Gilbert & Donk ex Singer, as well as the, morphologically distinct, members of the families Pterulaceae Corner and Typhulaceae Jülich; previously, Moncalvo et al. (2002), had recognized a distinct clade xeromphalinoid (no. 69) among the core of the white-spored euagarics. Outdated classifications based on morphological characters situated Xeromphalina in the Tricholomataceae (Kirk et al. 2001, 2008) and Mycenaceae (according to the Index Fungorum) and Xerulaceae Jülich (Redhead 1987). Prior to these, both Singer (1986) and Kühner (1980) included this genus in the Tricholomataceae R. Heim ex Pouzar, in the tribes Myceneae and Marasmieae, respectively. Singer’s point of view was followed by Antonín and Noordeloos (2004) in their monograph on European taxa.

Materials and methods The only collection consists of four specimens, all growing on soil among vegetal debris in a Mediterranean Quercus suber forest. The material, though scanty, is preserved in excellent condition, and is fully mature and well sporulated. The type collection has been deposited at AH (Departamento de Biología Vegetal, Universidad de Alcalá, Spain), with color photographs of the basidiomes taken in the field. Microscopical slides of dried material were prepared with 5–10% NH4OH, 5–10% KOH and Melzer. Digital photographs for the plates were taken with a Nikon camera, model DS-5 M, mounted to a binocular Nikon Eclipse 80i microscope. Spore measurements are quoted according to Heinemann and Rammeloo (1985), where Q=L/l means the quotient between the averages of length (L) and breadth (l), established with a minimum of 20 spores measured. Colors of fresh and dried basidiomes were compared with reference colors in Munsell (1994). Terminology of microscopical elements, especially cystidia (for example, circumcystidia is referred to those pileocystidia largely confined to a band around the pileus margin), has been mainly adopted from Redhead (1988) and also from Antonín and Noordeloos (2004). Total DNA from Xeromphalina setulipes holotypus (AH36724) and also from several European representatives of the genus Xeromphalina deposited at AH (Table 1), was extracted by means of the MasterPureTM Plant Leaf DNA Purification Kit (Epicentre Biotechnologies, Madison, US) following the manufacturer’s instructions: 1 μl of resuspended DNA was added to each 50 μl PCR mixture with the following composition: 1u EcoTaq DNA polymerase with 1× EcoTaq Buffer (Ecogene), MgCl2 2 mM, and 0.2 mM of each of the DNTPs. The primers LR1 (5′-GCA TAT CAA TAA

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GCG GAG GA-3′, Van Tuinen et al. 1998) and LR7 (5′-TAC TAC CAC CAA GAT CT-3′, Vilgalys and Hester 1990) targeting the 5′ end of the 28S nrLSU, and primers NSI1 (5′-GAT TGA ATG GCT TAG TGA GG-3′) and NLB4 (5′-GGA TTC TCA CCC TCT ATG AC-3′) from Martin and Rygiewicz (2005) targeting the nuclear ribosomal internal transcribed spacer (nrITS), were added at 0.5 μM each in independent PCR reactions. PCR consisted of a hot start at 95°C for 5 min, five cycles at 94°C, 50°C, and 72°C (45, 30, and 45 s, respectively), followed by 30 cycles at 94°C, 54°C, and 72°C (45, 30, and 45 s, respectively) and a final 72°C step for 10 min. PCR products were checked in 1% agarose gels prior to purification with UltraClean PCR Clean-up DNA purification kit (MoBio Laboratories, Carlsbad CA). The same PCR primers were used for sequencing. Chromatograms and sequences were visually compared in MEGA4 in order to detect compromised data. Phylogenetic analyses were performed using about 800 bp of nrLSU 5′ end and 650 bp of the ITS1-5.8S-ITS2 region. Homologue sequences of both genes from species in the genera Xeromphalina, Heimiomyces, and Sarcomyxa deposited at GenBank (Table 1) were aligned in MEGA4 using the ClustalW application. Most of the sequences gathered from GenBank were properly identified by specialists before sequencing (Johnson and Petersen 1997). The sequences were used in this work as they were the only available for molecular comparison of the new species. No position was considered ambiguous enough to be deleted in the final alignments. A neighbor-joining phylogenetic tree was constructed for each gene (5,000 bootstrap replicates, maximum composite likelihood model, pairwise deletion of gaps). Additionally, alignments were uploaded in PAUP* 4.0b10 (Swofford 2001) and a bootstrap search under the maximum parsimony criterion was performed. A thousand bootstrap replicates were executed using the tree-bisection-reconnection swapping algorithm, 50 random sequence additions in each replicate, and the MulTrees option not in effect. Finally, a maximum likelihood analysis of each alignment was performed in MrBayes 3.1 (Ronquist, Huelsenbeck and van der Mark) under the model specified by a previous MrModeltest (Nylander 2004) run in PAUP* 4.0b10. The best models for ITS and nLSU alignments were HKY+Γ and GTR+Γ, respectively. Maximum likelihood analyses employed 4 metropolis-coupled Monte Carlo Markov Chains (MCMC) with the temperature of the chains set to 0.2. Data sampling was done once for each 100 generations. The analysis was run until convergence parameters were met in two simultaneous runs, after 400,000 and 900,000 generations for ITS and nLSU alignments respectively. A full search for best-scoring ML trees was also performed in RAxML (Stamatakis 2006) using the standard search algorithm, and 1,000 bootstrap replications.

Mycol Progress (2010) 9:575–583 Table 1 DNA data from the genera Xeromphalina, Heimiomyces, and Sarcomyxa

577 Taxon Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina Xeromphalina

cauticinalis AH37873 cauticinalis AH37874 aff. parvibulbosa TENN61775 aff. parvibulbosa TENN61775 cornui TENN6397 campanelloides TENN6368 setulipes AH36724 campanella TENN7250 campanella AFTOL-ID 1524 campanella 3922 campanella fraxinophila TENN6398 austroandina TENN7392 helbergerii TENN6255

Xeromphalina junipericola AH19695 Xeromphalina junipericola AH19696 Xeromphalina kauffmanii TENN6906 Heimiomyces Heimiomyces tenuipes TENN6908 Heimiomyces fulvipes TENN5864 Sarcomyxa Sarcomyxa serotina I354 Sarcomyxa serotina UASWS0313 Sarcomyxa serotina AFTOL-ID 536 Sarcomyxa serotina ACCC 50309 Sarcomyxa serotina TM03_367 Sarcomyxa serotina

Results Phylogenetic analyses BLAST searches of the sequences from Xeromphalina setulipes in NCBI web service resulted in more than 90% identity with species in the genus Xeromphalina in more than 75% (ITS) or 60% (nLSU) of the sequence length. The final alignment of the nLSU and ITS regions included 45 and 187 parsimony-informative sites, respectively. Both nLSU and ITS inference supported the differentiation of, at least, three clades within the genus Xeromphalina. Xeromphalina setulipes grouped with X. fraxinophila, X. cornui, X. campanelloides and X. cauticinalis in one of them. ITS inference proved X. setulipes and X. cauticinalis to be different species. Samples of X. cauticinalis from the Iberian Peninsula seemed also somewhat different from those of the North American X. aff. parvibulbosa. Xeromphalina aff. parvibulbosa was related to X. cauticinalis on the basis of neighbor-joining and maximum parsimony analyses, while maximum likelihood supported a closer

Reference

nLSU

ITS

This work This work Hughes et al. (2009) Hughes et al. (2009) Moncalvo et al. 2002 Moncalvo et al. 2002 This work Moncalvo et al. 2002 Matheny et al. 2007 This work Walther et al. 2005 Moncalvo et al. 2002 Moncalvo et al. 2002 Moncalvo et al. 2002

GU320011 – – – AF261463 AF261462 GQ890700 AF261469 DQ470826 GU320009 AY207312 AF261464 AF261466 AF261465

GU320004 GU320005 FJ596890 FJ596889 – – GQ890701 – DQ494702 GU320006 – – – –

This work This work Moncalvo et al. 2002

GU320010 GU320012 AF261467

GU320007 GU320008 –

Moncalvo et al. 2002 Moncalvo et al. 2002

AF261471 AF261470

– –

Arhipova et al., unpub. Belbahri et al., unpub. Matheny et al. 2007

– – AY691887

GU062305 EF174452 DQ494695

Gao et al., unp. Porter et al. 2008 Garnica et al. 2007

EU365678 EU522798 DQ071731

– – –

relationship between X. setulipes and X.aff. parvibulbosa (data not shown). Xeromphalina junipericola was here sequenced for the first time, and seemed to be unrelated to the X. cauticinalis group on the basis of ITS data, while nLSU inference placed it in close relationship with X. helbergerii. This should be further confirmed by ITS analysis of the latter due to the relatively scarce number of informative sites in the nLSU region. Taxonomy Xeromphalina setulipes Esteve-Rav. & G. Moreno, sp. nov.—MycoBank MBS 16618; Figs. 1 and 2 Etymology. From latin setula = stiff hair, and pes = foot, owing to the stipe covered with setose hairs. Pileus 8–15 mm latus, convexus vel plano-convexus, haud striatus, rufo-brunneus vel hepaticus, glaber vel rugulosus, margine pallescente, leviter sericeo. Lamellae subconfertae, arcuato-decurrentes, brunneolae. Stipes 30–45×1–2 mm, cylindraceus, purpureo-brunneus vel subniger, minute pubescens, corneus, basi tomentosa, tomento cinnamomeo. Caro

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Fig. 1 Xeromphalina setulipes holotype AH 36274. Basidiomes. Scale bar 10 mm

tenuis, cartilaginea, brunneola, odore nullo, sapore mitis, non acerbo. Sporae Q= L×l: 5.0–6.1–7.3×2.9–3.3–3.7 µm, Qm: 1.6–1.87–2.1 (n = 21), ellipsoideae vel subcylindricae, leves, hyalinae, amyloideae. Basidia 21–26(−30)×4.5– 5.5 µm, clavata, tetraspora; sterigmata usque ad 3 µm longa. Cheilocystidia 30–80 × 3–7 µm, hyalina, coralloideo-ramosa. Caulocystidia 30–60×5–10 µm, fusiformia vel setuliformia, crassitunicata, luteo-brunnea. Pileipellis incrustata, rubrobrunnea in KOH. Hyphae fibulatae. Ad terram in silvis mediterraneis frondosis (Quercus, Cistus). In Hispania. Holotypus AH 36274. Basidiomes epigeous, gregarious. Pileus 8–15 mm in diam., convex to plano-convex, with depressed centre, remaining so at maturity; not or scarcely hygrophanous, not translucently striate at margin, dark red-brown, hepatic-brown to chocolate-brown (2.5 YR 3/4–3/6; 10 R 3/3–3/6), somewhat pallescent on drying to tobacco-brown (2.5 YR 4/3–4/6; 5 YR 4/4–4/6), especially towards the edge; surface glabrous when young, becoming rugose to rugulose with age, slightly pallescent and becoming somewhat sericeous towards margin when dry; margin inflexed, typically undulate to crenate. Lamellae subdistant (L=15–18), with lamellulae (l=1–2), distinctly arcuate-decurrent, up to 2 mm broad, tobacco-brown (5 YR 4/4–4/6) when young and remaining so at maturity, at times bifurcate, with paler, whitish and crenulate edge. Stipe 30–45×1–2 mm, cylindrical, somewhat enlarged at extreme base, stiff or cartilagineous, uniformly dark purplish-brown (2.5 YR 3/2–3/4 or 2.5/1–2.5/4) to nearly blackish-brown; apparently smooth, but finely pubescent-floccose under the lens (owing to the setulose brownish caulocystidia); basal tomentum well developed, amber to ozonium-like colored (5 YR 5/6–5/8). Context

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cartilagineous, concolorous with the surface; smell indistinct, taste not bitter. Rhizomorphs present, chestnut-brownish. Spores Q=L×l: 5–6.1–7.3×2.9–3.3–3.7 µm, Qm: 1.6– 1.87–2.1 (n=21), amyloid, ellipsoid to subcylindrical, thinwalled, smooth. Basidia 21–26(−30)×4.5–5.5 µm, 4-spored (rarely 2-spored), narrowly clavate to subcylindrical, with sterigmata up to 3 µm long. Lamellar edge heterogeneous, composed by numerous cheilocystidia mixed with some scattered basidia. Cheilocystidia abundant, 30–80×3–7 µm, sinuose to irregularly cylindrical, conspicuously branched forming dense coralloid excrescences (Mycena-like) 3–10 µm long, thin-walled, hyaline. Hymenophoral trama made up of cylindrical, parallel, thin-to slightly thickwalled, 4–8 µm wide hyphae, with walls smooth or slightly encrusted with brownish pigment which turns orangebrown in KOH. Pileipellis a not or hardly gelatinized cutis, constituting of interwoven to more or less radially arranged, slightly thick-walled, cylindrical, 5–10 µm wide hyphae, with yellowish-brown parietal pigment which turns reddishbrown in KOH. Subpileipellis made up of parallel, 4–8 µm wide hyphae, with finely encrusting yellowish-brown pigment, which turns orange-brown with KOH. Circumcystidia numerous, similar to cheilocystidia, showing numerous lateral branches, rod-like projections or diverticulae, forming coralloid masses, usually thick-walled. Stipitipellis a cutis of parallel, cylindrical, thick-walled hyphae, with dark red-brown parietal pigment which turns darker in KOH. Caulocystidia very numerous, of similar shape along the entire stipe, strongly projecting, 30–60× 5–10 µm, fusoid-ventricose, often sinuose or curved, exceptionally simply branched but never lobed at distal ends, with attenuated obtuse apex ( setula-like ), distinctly thick-walled, walls up to 2.5 µm wide, brown-yellowish pigmented. Clamp-connections present. Substratum. On acid soil, among vegetal debris in a mixed Quercus suber/Q. faginea forest with Cistus ladanifer, Erica sp. and Juniperus cf. oxycedrus undergrowth. Distribution Only known from the type locality, in Ciudad Real province, Spain. Specimens examined. SPAIN, Ciudad Real Province, Fuencaliente, camino del Robledo de las Hoyas, 30SUH8553, 770 m asl, on soil among vegetal debris near Cistus, Erica and Juniperus shrub, in a Mediterranean mixed Quercus suber/Q. faginea forest, in acid soil, 17 Nov. 2005, F.D. Calonge, F. Prieto, M.A. González & F. Esteve Raventós, holotype Herb. AH 36274.

Discussion Xeromphalina setulipes shows a distinct color change in the pileipellis in KOH solutions, turning dark reddish-brown; this color change of the tissues has been emphasized

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Fig. 2 Xeromphalina setulipes holotype AH 36274. a Detail of the stipe apex, pileus margin and lamellae insertion. b Floccosevelutinous stipe due to the presence of setuloid caulocystidia. c Mycelial tomentum at the stipe base. d Pileipellis constituted by cylindrical incrusted hyphae with yellowish-brown pigment observed

in phase-contrast. e The same in Congo red. f–g Basidia and basidioles. h–i Clamp-connections. j Amyloid basidiospores. k– l Setula-like caulocystidia showing thick walls. m–o Coralloid cheilocystidia. Scale bars a 3 mm, b,c 1 mm, d–o 10 µm

previously by Smith (1953), Redhead (1988), and Antonín and Noordeloos (2004) to separate some groups of species. Based on this character, Smith (1953) proposed section Mutabiles, which includes X. campanelloides Redhead, X. cauticinalis, X. cirris Redhead, X. cornui (Quél.) J. Favre, X. fraxinophila A.H. Sm., X. parvibulbosa (Kauffman & A.

H. Sm.) Redhead and X. setulipes. Also in this section, the presence of rhizomorphs and irregular and branched circumcystidia are characteristic. Morphologically, X. setulipes is similar in habit to the circumboreal X. cauticinalis; both species also show a characteristic amber tomentum at the stipe base, and also

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grow on soil, not directly on stumps or rotten wood; X. cauticinalis also shows some fusiform caulocystidia which remind of those of X. setulipes, but these are mixed with typical coralloid or lobed ones, which we have not observed in the new species in any part of the stipe; moreover, they also differ in many characters, such as the overall colors, and the yellow paler stipe apex in X. cauticinalis, very different to the uniformly dark brownchestnut to blackish stipe of X. setulipes. Lastly, both species also differ in taste, which is very bitter in X. cauticinalis, even in dry material. On the basis of the overall dark colors and smooth caulocystidia, it would seem that X. brunneola O.K. Mill. could be close to X. setulipes but clearly differs by the nonbranched or non-coralloid cheilo- and pileocystidia, the pileus trama not reacting in KOH solutions, the narrower spores (2–3 µm wide) and habitat in coniferous wood (Redhead 1988); the species seem not to be related phylogenetically, as X. brunneola is closer to X. campanella (see Figs. 3 and 4). Among those species of section Mutabiles (trama showing a positive color change in KOH), X. campanelloides Redhead is represented by collybioid, mostly fasciculate basidiomes (Redhead 1988: 488; Antonín and Noordeloos 2004), but this shows lobed, branched to coralloid thin-walled caulocystidia, and the presence of yellow pigment granules at the stipe medulla seems to be a characteristic feature, which we have not observed in X. setulipes. Another close species of this section is X. cornui (Quél.) J. Favre, originally described from central Europe; it develops in conifer forests (Larix decidua, Pinus montana Mill., etc.), but is typically found growing among Sphagnum. Apart from the habitat, this can be recognized by the discrete yellow granules on the pileus and stipe apex, circumcystidia turning reddish to reddishbrown in KOH, irregularly contorted to lobed caulocystidia restricted only to the stipe apex and a mild taste as in X. Fig. 3 ITS Neighbor-joining consensus tree showing the relationships between Xeromphalina setulipes and its closest relatives. Bootstrap proportions from neighbor-joining and maximum parsimony, and posterior probabilities from bayesian maximum likelihood, and RAxML likelihood analyses are shown in this order next to nodes. Supporting values lower than 70% are not shown. Nodes supported by none or just a single inference method have no annotation

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setulipes. A lectotype was designated by Redhead (1988: 496) for this small species, which has been illustrated by Courtecuisse and Duhem (1994) and Ludwig (2001: pl. 186, fig. 88.2). According to Redhead (1988), who studied numerous samples, it is not a rare species in North America, but was often formerly confused with X. cauticinalis (see Smith 1953). Xeromphalina fraxinophila is another member of section Mutabiles, which is close in some features to X. cornui (circumcystidia turning reddish in KOH); it is a more robust species (pileus 1–3 cm broad and stipe−3.5 mm wide), with yellowish lamellae; it shows caulocystidia similar in shape and length to X. setulipes, but clearly differs in habit, colors and the practical absence of cheilocystidia, which bear, if present, few simple lobes (like in X. cauticinalis). Xeromphalina parvibulbosa may have similar caulocystidia to X. setulipes, but its taste is bitter or astringent, and circumcystidia do not become reddish in KOH. Another representative from the northern hemisphere is Xeromphalina aspera Maas Geest. which was described growing on wood from Nepal (Maas Geesteranus (1992: 50); it shows a fasciculate growth, brown-reddish colors and is microscopically characterized by the pileus trama that turns reddish in KOH solutions, large lageniform cheilocystidia, a gelified pileipellis of smooth hyphae, and clavate to fusiform caulocystidia. Screening the literature for Xeromphalina representatives from pantropical areas and the southern hemisphere, we have found that only a few species were described by Singer (1965) in a contribution on the genera Xeromphalina and Heimiomyces in South America (especially those from the east slope of the Andes and Brazil); he studied three species, X. helbergeri (which is also present in Central America and the Caribbean), X. tenuipes (widely distributed in the American continent), and X. austroandina; all these three are lignicolous, lack coralloid cheilocystidia and were

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Fig. 4 nLSU Neighbor-joining consensus tree showing the relationships between Xeromphalina setulipes and its closest relatives. Bootstrap proportions from neighbor-joining and maximum parsimony, and posterior probabilities from bayesian maximum like-

lihood, and RAxML likelihood analyses are shown in this order next to nodes. Supporting values lower than 70% are not shown. Nodes supported by none or just a single inference method have no annotation

gathered by Singer in diverse Patagonian forests (Nothofagus, Podocarpus, etc.). Later on, Singer (1989) contributed with two new lignicolous species (X. tropicalis and X. yungensis), from Brazil and Bolivia respectively. None of the taxa described by Horak (1979) from Indomalaya and Australasia, either in Xeromphalina or in Heimiomyces, are morphologically similar to X. setulipes; all Xeromphalina species mentioned and described in this contribution show very different cheilocystidia, which are fusoid to lageniform and never coralloid; Horak (1968, 1979) also recognized Heimiomyces as an independent genus from Xeromphalina; it would seem that all Heimiomyces species macroscopically differ from Xeromphalina by the adnexed to broadly emarginate lamellae. Corner (1996) cited Heimiomyces tenuipes (Schwein.) Singer from Malesia; it is a common pantropical taxon that had previously also been recorded from South America, Africa, and the Caribbean (Singer 1965), as well as from North America (Redhead 1988). More recently, Maas Geesteranus and Horak (1995) described the new species Xeromphalina

nudicaulis Maas Geest. & E. Horak from Papua New Guinea, growing on rotten roots of Fagaceae (Castanopsis and Lithocarpus), characterized by its “naked” covering (devoid of pileo- and caulocystidia), and fusoid-ventricose cheilocystidia. We conclude that, from a morphological point of view, the combination of dark colors (brownish lamellae and uniformly blackish-brownish stipe), mild taste, smooth fusiform thick-walled caulocystidia, and branched to coralloid cheilo-and circumcystidia seems to be diagnostic for X. setulipes. The ecological patterns could also be a distinct characteristic of the new species, as X. setulipes has been found on soil in a Mediterranean ecosystem, specifically in a Quercus suber ( alcornoque ) forest with an undergrowth of typical Mediterranean shrub, mainly composed by Cistus and Juniperus. Xeromphalina setulipes seems to be closely related to X. fraxinophila, X. cornui, X. campanelloides, and X. cauticinalis. This group shows clear genetic differences with the group formed by the species X. campanella, X. kauffmanii,

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X. brunneola, and probably X. junipericola, and also with the southern hemisphere taxa X. helbergeri and X. austroandina (Fig. 3). The analysis of more informative DNA regions such as the ITS or RPB2 loci from the whole group could help to clarify the internal organization of these clades. As mentioned above, a comprehensive morphological–molecular review of the genus is also needed in order to validate GenBank sequences, define the molecular species concept in this clade, and review the current morphological taxonomy of the genus. Acknowledgements We acknowledge L. Monje and A. Pueblas (Gabinete de Dibujo y Fotografía Científica at Alcalá University) for their valuable help in the treatment of digital images; the Spanish Ministry of Education and Culture for FPU grant AP2006-00890, the Consejería de Agricultura y Medio Ambiente de la Junta de Comunidades de Castilla-La Mancha for the Research Projects 2004X802, PAI08-0240-5097, and the Empleaverde Truficulture project from the Fundación Diversidad (cofinanced by the European Social Fund and the FGUA Cátedra de Medio Ambiente), which enabled us to carry out this study. We also want to thank Dr. G. Consiglio for his help in the Latin diagnosis and an anonymous reviewer for kindly helping in the grammatical correction of the manuscript.

References Antonín V, Noordeloos ME (2004) A monograph of the genera Hemimycena, Delicatula, Fayodia, Gamundia, Myxomphalia, Resinomycena, Rickenella and Xeromphalina (Tribus Myceneae sensu Singer, Mycena excluded) in Europe. IHW, Germany Bon M (1999) Flore Mycologique d’Europe 5. Les CollybioMarasmïoïdes et ressemblants. Documents Mycologiques. Mémoire hors-série 5:1–171 Corner EJH (1996) The agaric genera Marasmius, Chaetocalathus, Crinipellis, Heimiomyces, Resupinatus, Xerula and Xerulina in Malesia. Nova Hedwigia Beiheft 111. Cramer, Berlin/Stuttgart Courtecuisse R, Duhem B (1994) Guide des champignons de France et d’Europe. Eclectis, France Esteve Raventós F (1995) Xeromphalina minutissima nom. prov., una nueva especie a ser validada. Bol Soc Micol Madr 20:313– 316 Garnica S, Weiss M, Walther G, Oberwinkler F (2007) Reconstructing the evolution of agarics from nuclear gene sequences and basidiospore ultrastructure. Mycol Res 111:1019–1029 Gulden G (1992) Xeromphalina. In: Hansen L, Knudsen H (eds) Nordic macromycetes, vol. 2. Nordsvamp, Denmark Heinemann P, Rammeloo J (1985) De la mésure des spores et de son expression. Agarica 6:366–380 Horak E (1968) Synopsis generum Agaricalium (Die Gattungstypen der Agaricales). Beitr Kryptogamenflora Schweiz 13:1–741 Horak E (1979) Xeromphalina and Heimiomyces in Indomalaya and Australasia. Sydowia 32:131–153 Hughes KW, Petersen RH, Lickey EB (2009) Using heterozygosity to estimate a percentage DNA sequence similarity for environmental species' delimitation across basidiomycete fungi. New Phytol (in press) Johnson JE, Petersen RH (1997) Mating systems in Xeromphalina species. Mycologia 89:393–399 Kirk PM, Cannon PF, David JC, Stalpers JA (eds) (2001) Dictionary of the fungi, 9th edn. CABI Publishing, United Kingdom

Mycol Progress (2010) 9:575–583 Kirk PM, Cannon PF, Minter DW, Stalpers JA (eds) (2008) Ainsworth & Bisby’s dictionary of the fungi, 10th edn. CABI Publishing, United Kingdom Klán J (1984) The genus Xeromphalina (Tricholomataceae) in Europe. Ces Mykol 38:205–217 Kühner R (1980) Les Hyménomycètes agaricoides. Bull Soc Linn Lyon, spec num 49:1–1027 Ludwig E (2001) Pilzkompendium Band 1. IHW, Germany Maas Geesteranus RA (1992) Some Myceneae of the Himalayan foothills. Persoonia 15:33–53 Maas Geesteranus RA, Horak E (1995) Mycena and related genera from Papua New Guinea and New Caledonia. Bibl Mycol 159:143–229 Martin KJ, Rygiewicz PT (2005) Fungal-specific PCR primers developed for analysis of the ITS region of environmental DNA extracts. BMC Microbiol 5:28 Matheny PB, Hofstetter V, Aime MC, Moncalvo J-M, Ge Z-W, Yang Z-L, Slot JC, Ammirati JF, Baroni TJ, Bougher NL, Hughes KW, Lodge DJ, Kerrigan RW, Seidl MT, Aanen DK, DeNitis M, Daniele GM, Desjardin DE, Kropp BR, Norvell LL, Parker A, Vellinga EC, Vilgalys R, Hibbett DS (2007) Major clades of Agaricales: a multilocus phylogenetic overview. Mycologia 98:982–995 Miller OK Jr (1968) A revision of the genus Xeromphalina. Mycologia 60:156–188 Moncalvo J-M, Vilgalys R, Redhead SA, Johnson JE, James TY, Aime MC, Hofstetter V, Verduin SJW, Larsson E, Baroni TJ, Thorn RG, Jacobsson S, Clémençon H, Miller OK Jr (2002) One hundred and seventeen clades of euagarics. Mol Phylogenet Evol 23:357–400 Moreno G, Heykoop M (1996) Xeromphalina junipericola sp. nov. (Tricholomataceae, Agaricales) from Spain. Z Mykol 62:37–41 Munsell (1994) Munsell soil color charts. Macbeth Division of Kollmorgen Instruments, New York Noordeloos ME (2008) Xeromphalina. In: Knudsen H, Vesterholt J (eds) Funga Nordica. Nordsvamp, Denmark, pp 240–242 Nylander JAA (2004) MrModeltest v2. Evolutionary Biology Centre, Uppsala University Porter TM, Skillman JE, Moncalvo JM (2008) Fruiting body and soil rDNA sampling detects complementary assemblage of Agaricomycotina (Basidiomycota, Fungi) in a hemlockdominated forest plot in southern Ontario. Mol Ecol 17:3037–3050 Redhead SA (1987) The Xerulaceae (Basidiomycetes), a family with sarcodimitic tissues. Can J Bot 65:1551–1562 Redhead SA (1988) Notes on the genus Xeromphalina (Agaricales, Xerulaceae) in Canada: biogeography, nomenclature, taxonomy. Can J Bot 66:479–507 Redhead SA, Halling RE (1987) Xeromphalina nubium sp. nov. (Basidiomycetes) from Cerro de la Neblina, Venezuela. Mycologia 79:383–386 Singer R (1965) Monographs of South American Basidiomycetes, especially those of the East slope of the Andes and Brazil. X. Xeromphalina. Bol Soc Argent Bot 10(4):302–310 Singer R (1986) The Agaricales in Modern Taxonomy, 4th edn. Koenigstein, Germany Singer R (1989) New taxa and new combinations of Agaricales (Diagnoses Fungorum Novorum Agaricalium IV). Fieldiana 21:1–133 Smith AH (1953) New and rare agarics from the Douglas Lake Region and Tahquamenon Falls State Park, Michigan, and an account of the North American species of Xeromphalina. Pap Mich Acad Sci Arts Lett 38:53–87 Stamatakis A (2006) RAxML-VI-HPC: maximum likelihood-based phylogenetic analyses with thousands of taxa and mixed models. Bioinformatics 22:2688–2690

Mycol Progress (2010) 9:575–583 Swofford DL (2001) PAUP*: phylogenetic analysis using parsimony (and other methods). Version 4.0b10. Sinauer, Sunderland Van Tuinen D, Zhao B, Gianinazzi-Pearson V (1998) PCR studies of AM fungi, from studies to application. Springer, Berlin Vilgalys R, Hester M (1990) Rapid genetic identification and mapping of enzymatically ampliWed ribosomal DNA from several Cryptococcus species. J Bacteriol 172:4238–4246

583 Walther G, Garnica S, Weiss M (2005) The systematic relevance of conidiogenesis modes in the gilled Agaricales. Mycol Res 109:525–544 Watling R, Turnbull E (1998) British Fungus Flora. Agarics and Boleti. 8/Cantharellaceae, Gomphaceae and Amyloid-Spored and Xeruloid Members of Tricholomataceae (excl. Mycena). Royal Botanic Garden, Edinburgh