Molecular phylogeny and spore evolution of Entolomataceae.

The phylogeny of the Entolomataceae was reconstructed using three loci (RPB2, LSU and mtSSU) and, in conjunction with spore morphology (using SEM and TEM), was used to address four main systematic issues: 1) the monophyly of the Entolomataceae; 2) inter-generic relationships within the Entolomataceae; 3) genus delimitation of Entolomataceae; and 4) spore evolution in the Entolomataceae. Results confirm that the Entolomataceae (Entoloma, Rhodocybe, Clitopilus, Richoniella and Rhodogaster) is monophyletic and that the combination of pinkish spore prints and spores having bumps and/or ridges formed by an epicorium is a synapomorphy for the family. The Entolomataceae is made up of two sister clades: one with Clitopilus nested within Rhodocybe and another with Richoniella and Rhodogaster nested within Entoloma. Entoloma is best retained as one genus. The smaller genera within Entoloma s.l. are either polyphyletic or make other genera paraphyletic. Spores of the clitopiloid type are derived from rhodocyboid spores. The ancestral spore type of the Entolomataceae was either rhodocyboid or entolomatoid. Taxonomic and nomenclatural changes are made including merging Rhodocybe into Clitopilus and transferring relevant species into Clitopilus and Entoloma.

INTRODUCTION

The euagaric family Entolomataceae Kotl. & Pouzar is very species-rich. It is composed of more than 1 500 species and occurs worldwide, from arctic to tropical habitats (Horak 1980, 2008, Baroni 1981, Largent 1994, Noordeloos 2004, Gates & Noordeloos 2007, Noordeloos & Hausknecht 2007). The family is highly variable in terms of sporocarp morphology (tiny to large; pleurotoid, omphalioid, collybioid, mycenoid, and tricholomatoid, as well as sequestrate), and micromorphology (spore shape, pileipellis structures, pigmentation types, cystidia presence and shape, etc.; Noordeloos 2004; Fig. 1). Lifestyles are equally varied: Most species are saprotrophic on soil, wood or moss, but some are parasitic on other mushrooms (Noordeloos 2004), parasitic on plants or ectomycorrhizal (Antibus et al. 1981, Agerer & Waller 1993, Agerer 1997, Kobayashi & Hatano 2001, Montecchio et al. 2006). The family traditionally contains three main agaricoid genera: Rhodocybe Maire, Clitopilus (Fr. ex Rabenh.) P. Kumm. and Entoloma (Fr.) P. Kumm. s.l. The latter genus is sometimes split into more genera (e.g. 13 genera; Largent 1994). Additionally, three smaller non-agaricoid genera have been distinguished on the basis of habit, namely, the monotypic Rhodocybella T.J. Baroni & R.H. Petersen (with a cyphelloid habit), Rhodogaster E. Horak (secotioid) and Richoniella Costantin & L.M. Dufour (gasteroid).

Fig. 1

Entolomataceae variation in basidiocarp morphology. a. Entoloma prunuloides; b. E. sinuatum; c. E. catalaense; d. E. conferendum; e. E. camarophyllus; f. E. roseum; g. Clitopilus prunulus; h. Rhodocybe gemina; i. Entoloma rodwayi; j. Richoniella pumila; k. E. uranochroum. — Photos by: a, b. Y. Deneyer; c. G. Consiglio; d. J. Vesterholt; e, g–i. M.E. Noordeloos; f. H. Huijser; j. M. Pilkington; k. M. Meusers.

It is no surprise that Entolomataceae, being such a large and highly variable family, raises questions that analysis of morphological characters alone cannot answer, either due to scarcity of characters and/or difficulty in interpreting the significance of the characters. Molecular phylogenetic methods are therefore used in our study to address four main systematic issues:

the monophyly of the Entolomataceae;

inter-generic relationships within the Entolomataceae;

genus delimitation of Entolomataceae and, with the addition of spore morphology;

spore evolution in the Entolomataceae.

Monophyly of the Entolomataceae and intergeneric relationships

The members of Entolomataceae have been classified together because they all share the property of spore prints that are pink to brownish or greyish pink in combination with spores that are bumpy, ridged, or angular in polar or in all views. The spore wall ornamentations are unique, being formed by local thickenings in the spore wall, the epicorium (Clémençon et al. 2004). The presence of pink, angular spores has been considered so unique that Entolomataceae, in contrast to many other Agaricales families, has been widely regarded a natural group (Kühner 1980, Singer 1986).

Species from other genera had, in the past, been placed within Entolomataceae. However, recent studies have excluded them. Macrocystidia Joss. and Rhodotus Maire had been classified in the family on account of their pink spores, but molecular phylogenetic studies have placed them outside the family (Moncalvo et al. 2002). Comparison of the spore wall of Rhodotus palmatus (Bull.) Maire and members of Entolomataceae showed that their bumps are not homologous (Clémençon 1997). Also, the phylogenetic study by Moncalvo et al. (2002) suggested that Catathelasma Lovejoy and a strongly supported clade containing Callistosporium Singer, Macrocybe Pegler & Lodge and Pleurocollybia Singer were best included in Entolomataceae. The more recent phylogenetic study by Matheny et al. (2006) has excluded Catathelasma and Callistosporium from Entolomataceae with strong support.

Both phylogenetic studies (Moncalvo et al. 2002, Matheny et al. 2006) were based on relatively small samples of Entoloma, Rhodocybe and Clitopilus and none of Rhodogaster, Richoniella and Rhodocybella. Thus, phylogenetic relationships among these six genera had remained unresolved.

Spore evolution

Spore characters have been important both to characterize the family (having pink, angular spores) but also to separate its three main agaricoid genera, Rhodocybe, Clitopilus and Entoloma, from each other. Rhodocybe has spores with ornamentations in the form of bumps and undulate ridges having various arrangements resulting in spores that are undulate to weakly angular in profile and face views, and angular in polar view (Baroni 1981). Clitopilus is characterized by spores with an ornamentation of longitudinal ridges. Entoloma has spores that are angular in all views due to its network of interconnected ridges that form facets and are highly varied in shape (Romagnesi 1974, Pegler & Young 1978, 1979).

There are two main theories on how spore shapes within Entolomataceae evolved. According to the first theory (Kühner 1980), rhodocyboid spores represent the plesiomorphic condition since they are the more similar to what he considered the closest relative of Entolomataceae, Lepista (Fr.) W.G. Sm. (Tricholomataceae). Species of that genus have pinkish, roughened spores. The spores of Clitopilus are the evolutionary intermediate between Rhodocybe and Entoloma. Entoloma spores are the most complex and represent the most evolved spore form. The second theory (Baroni 1981) is similar in that rhodocyboid spores are ancestral. However, Baroni based his argument that the rhodocyboid spore is the most primitive on the assumption that since pink angular spores do not exist elsewhere in the Agaricales, the first Entolomataceae evolved from an unknown member of Tricholomataceae with slightly rounded-angular, pinkish spores. More pronounced angularity then derived from this. Furthermore, in contrast to Kühner’s theory, clitopiloid and entolomatoid spores evolved independently from rhodocyboid spores. Note, however, that modern phylogenies support neither the sister relationship of Lepista nor any member of Tricholomataceae with a pink spore print. Rather, it is suggested that the Lyophyllaceae are the sister clade of Entolomataceae and that it is nested within other members of the Tricolomatoid clade (Hofstetter et al. 2002, Matheny et al. 2006).

These hypotheses can be evaluated through a phylogenetic reconstruction of Entolomataceae and its closest relatives and by subsequently mapping spore structure on the tree. In this framework, it is possible to reject hypotheses that are inconsistent with phylogeny. Hypotheses that are consistent with phylogeny can be further scrutinized and used as a basis for improved hypothesis formulation.

Genus delimitation of Entolomataceae

There are several problems with genus delimitation in Entolomataceae due to different interpretations of morphological evidence. Firstly, the genus Rhodocybella has rhodocyboid spores indicating a close relationship with Rhodocybe (Baroni & Petersen 1987). Secondly, Rhodogaster and Richoniella have entolomatoid spores and therefore these taxa are hypothesized to have been derived from or even to be members of Entoloma (Pegler & Young 1978, 1979, Kendrick 1994). Kuyper included Rhodogaster and Richoniella in Entoloma in the Dictionary of the Fungi (Kirk et al. 2008). Here we test this hypothesis and make the required nomenclatural changes.

Thirdly and most importantly, despite the generally agreed-upon distinction between the agaricoid genera due to spore shape, there are several taxa that are at the centre of dispute regarding the distinction between Rhodocybe and Entoloma. Baroni & Largent (1989) transferred E. trachyosporum Largent to Rhodocybe. They also proposed, although not formally, that E. nitidum Quél. be transferred to Rhodocybe (Baroni & Largent 1993, Castellano et al. 1999). Both proposals are based on their evaluation that the spores are rhodocyboid rather than entolomatoid. This evaluation implies that Entoloma species that are closely related to sect. Trachyospora and E. nitidum also belong to Rhodocybe, further blurring the distinction between the two genera. In contrast, Noordeloos (2004) retained E. nitidum in Entoloma, on the basis of his evaluation that the spores are entolomatoid rather than rhodocyboid.

Lastly, Entoloma is highly variable in terms of morphology. It is the second largest euagaric genus (after Cortinarius), with more than 1 500 species. Several mycologists therefore preferred to variously split this group into smaller genera (Orton 1960, 1991, Largent & Benedict 1971, Horak 1973, 1980, Largent 1977, 1994). These subdivisions do not show much consistency. Other mycologists have preferred to recognize one large genus (Romagnesi 1978, Noordeloos 1981, 1992, 2004). These authors noted problems with the delimitation of groups within Entoloma – particularly due to the existence of taxa with an intermediate position between subgenera. This study evaluates the existing proposals in a phylogenetic context. Proposals that are inconsistent with the phylogenetic relationships have to be rejected.

The present study uses molecular phylogenetic analyses of a 3-loci dataset together with spore characters scored from images captured using scanning electron microscopy (SEM) and transmission electron microscopy (TEM) to answer the following questions:

Is Entolomataceae monophyletic?;

Are the main genera Rhodocybe, Clitopilus and Entoloma monophyletic?;

What is the relationship of these three genera to each other?;

What theories on spore evolution in Entolomataceae are inconsistent with the phylogeny and should therefore be rejected, and what theories are consistent with it?;

How does the phylogeny inform the debate on the various taxonomic proposals for a possible generic delimitation within Entoloma s.l.?

MATERIALS AND METHODS

Taxon sampling

For outgroup, the analysis used 7 taxa from close relatives of Entolomataceae in the Tricholomatoid clade (sensu Matheny et al. 2006). Tricholoma vaccinum, Lepista ovispora and Collybia tuberosa were sequenced for this study. Sequences of Calocybe, Lyophyllum, Clitocybe and Collybia were downloaded from Genbank. For the ingroup, a total of 12 Rhodocybe, three Clitopilus, one Rhodogaster, one Richoniella, and 53 Entoloma accessions were sequenced. In addition, sequences of R. aureicystidiata and an AFTOL taxon identified as E. prunuloides were downloaded from Genbank. The AFTOL E. prunuloides turned out to be significantly different from the one sequenced for this study. Such discrepancies demonstrate the need for a system where sequences can be annotated (Bidartondo et al. 2008). It is here referred to as ‘Entoloma sp. 1’. The Entoloma species sampled represent 12 subgenera according to Noordeloos (2004). Subgenus Entoloma was more extensively sampled because preliminary analyses suggested that part of it is basal to the rest of the genus Entoloma and because it included E. trachyosporum and E. nitidum, taxa that have been at the centre of dispute regarding the distinction between Rhodocybe and Entoloma. We were not able to sequence Rhodocybella due to its insufficient collection. Table 1 summarizes taxonomic and collection information of the samples, as well as the DNA markers that were sequenced. The representatives of the Entolomataceae sequenced for this study were identified by the senior author, or are type-specimens.

Table 1

Species used in the phylogenetic analyses. All accessions were sequenced for this study except the last seven, which were downloaded from Genbank. The symbol * is placed before the names of species now transferred to Clitopilus and + for those transferred to Entoloma. Holotypes and isotypes are indicated with the collection number, as is the herbarium where the voucher specimen is deposited unless they are in L.

Species name	Genbank Accession numbers			Collection number	Country of collection
	mtssu	rpb2	lsu
*Rhodocybe caelata	GQ289348		GQ289208	“Exkursionsteilnehmer” 2005-08-28	Germany
*Rhodocybe fallax [1]	GQ289350	GQ289276	GQ289210	ME Noordeloos 200367	Slovakia
*Rhodocybe fallax [2]	GQ289349	GQ289275	GQ289209	ME Noordeloos 1997173	Italy
*Rhodocybe gemina	GQ289351	GQ289277		G. van Zanen 2003-09-14	Belgium
*Rhodocybe hirneola	GQ289352	GQ289278	GQ289211	ME Noordeloos 199956	Italy
*Rhodocybe mundula		GQ289280	GQ289213	ME Noordeloos 9867	Austria
*Rhodocybe nitellina [1]	GQ289355	GQ289282	GQ289215	ME Noordeloos 200435	Austria
*Rhodocybe nitellina [2]	GQ289354	GQ289281	GQ289214	ME Noordeloos 2002021	Austria
*Rhodocybe pallidogrisea	GQ289356	GQ289283	GQ289216	ME Noordeloos 2004032	Tasmania, Australia
*Rhodocybe pseudopiperita	GQ289357	GQ289284	GQ289217	ME Noordeloos 2004068	Tasmania, Australia
*Rhodocybe sp.	GQ289353	GQ289279	GQ289212	A. Gminder 2004-04-27	Germany
*Rhodocybe stangliana		GQ289285	GQ289218	N. Dam 05094	Switzerland
+Rhodocybe zuccherellii	GQ289346		GQ289206	A. Zuccherelli 1996-01-25 [holotype]	Italy
+Rhodogaster calongei	GQ289298		GQ289158	PM Pasabán [holotype, MA]	Spain
+Richoniella pumila	GQ289304	GQ289235	GQ289164	G.Gates E2031	Tasmania, Australia
Clitopilus cystidiatus	GQ289287	GQ289220	GQ289147	ME Noordeloos 200350	Slovakia
Clitopilus pinsitus	GQ289288		GQ289148	G. Immerzeel 1990-11	Netherlands
Clitopilus prunulus	GQ289289	GQ289221	GQ289149	ME Noordeloos 2003-09-14	Belgium
Entoloma abortivum	GQ289290	GQ289222	GQ289150	H den Bakker 92	Canada
Entoloma albidoquadratum	GQ289291	GQ289223	GQ289151	P. Manimohan 667 [holotype]	Kerala, India
Entoloma alcedicolor	GQ289292	GQ289224	GQ289152	E. Arnolds 0276 [holotype]	Netherlands
Entoloma araneosum	GQ289293	GQ289225	GQ289153	ME Noordeloos 200314	Belgium
Entoloma bloxamii	GQ289294	GQ289226	GQ289154	ME Noordeloos 200442	Austria
Entoloma caccabus	GQ289295	GQ289227	GQ289155	ME Noordeloos 200324	Belgium
Entoloma cephalotrichum	GQ289297	GQ289229	GQ289157	C. Ulje 1997-08-01	Netherlands
Entoloma cocles	GQ289299	GQ289230	GQ289159	J. Vauras 9770F	Finland
Entoloma coeruleogracilis [1]	GQ289309	GQ289240	GQ289169	G. Gates E1220	Tasmania, Australia
Entoloma coeruleogracilis [2]	GQ289308	GQ289239	GQ289168	G. Gates E1777	Tasmania, Australia
Entoloma haastii	GQ289307	GQ289238	GQ289167	ME Noordeloos 2004055	Tasmania, Australia
Entoloma conferendum	GQ289300	GQ289231	GQ289160	ME Noordeloos 200313	Belgium
Entoloma costatum	GQ289301	GQ289232	GQ289161	G. Immerzeel 2000-10-10	Netherlands
Entoloma cretaceum	GQ289302	GQ289233	GQ289162	G. Gates E1181 [holotype]	Tasmania, Australia
Entoloma excentricum	GQ289303	GQ289234	GQ289163	M. Meusers E 1705	Germany
Entoloma gelatinosum	GQ289305	GQ289236	GQ289165	G. Gates E792	Tasmania, Australia
Entoloma griseolazulinum	GQ289306	GQ289237	GQ289166	P. Manimohan 738 [holotype]	Kerala, India
Entoloma hebes	GQ289310	GQ289241	GQ289170	C. Hartman 1992-10-28	Netherlands
Entoloma indigoticoumbrinum	GQ289311	GQ289242	GQ289171	ME Noordeloos 200406 3 [holotype]	Tasmania, Australia
Entoloma indoviolaceum	GQ289312	GQ289243	GQ289172	P. Manimohan 700 [holotype]	Kerala, India
Entoloma kermandii	GQ289313	GQ289244	GQ289173	G. Gates E227 [holotype]	Tasmania, Australia
Entoloma myrmecophilum	GQ289314	GQ289245	GQ289174	G. Tjallingii-Beukers 1981-10-30	Netherlands
Entoloma nitidum	GQ289315	GQ289246	GQ289175	ME Noordeloos 200426	Slovakia
Entoloma pallideradicatum	GQ289316	GQ289247	GQ289176	A. Hausknecht [isotype ex WU 189010]	Austria
Entoloma parasiticum	GQ289317	GQ289248	GQ289177	ME Noordeloos 200330	Belgium
Entoloma perbloxamii	GQ289318	GQ289249	GQ289178	ME Noordeloos 2004071 [holotype]	Tasmania, Australia
Entoloma phaeomarginatum	GQ289319	GQ289250	GQ289179	ME Noordeloos 2004127	Tasmania, Australia
Entoloma pluteisimilis	GQ289320	GQ289251	GQ289180	C. Hermosilla 2001-12-08 [holotype]	Spain
Entoloma politum	GQ289321	GQ289252	GQ289181	ME Noordeloos 200325	Belgium
Entoloma porphyrescens	GQ289322	GQ289253	GQ289182	ME Noordeloos 2004113	Tasmania, Australia
Entoloma procerum	GQ289323	GQ289254	GQ289183	ME Noordeloos 2004070	Tasmania, Australia
Entoloma prunuloides	GQ289324	GQ289255	GQ289184	ME Noordeloos 200340	Slovakia
Entoloma pygmaeopapillatum	GQ289325	GQ289256	GQ289185	ME Noordeloos 200364	Slovakia
Entoloma readiae	GQ289326	GQ289257	GQ289186	ME Noordeloos 2004050	Tasmania, Australia
Entoloma rhodopolium var. nidorosum	GQ289327	GQ289258	GQ289187	ME Noordeloos 2003-09-16	Belgium
Entoloma sarcitum	GQ289328	GQ289259	GQ289188	A. Hausknecht 1994-04-20	Austria
Entoloma sericatum	GQ289329	GQ289260	GQ289189	ME Noordeloos 200328	Slovakia
Entoloma sericellum	GQ289330	GQ289261	GQ289190	ME Noordeloos 200315	Belgium
Entoloma sericeum	GQ289331	GQ289262	GQ289191	ME Noordeloos 200329	Slovakia
Entoloma serrulatum	GQ289332	GQ289263	GQ289192	ME Noordeloos 2004062	Tasmania, Australia
Entoloma sinuatum	GQ289333	GQ289264	GQ289193	J. Wisman 2003-09-19	Netherlands
Entoloma sordidulum	GQ289334	GQ289265	GQ289194	Co-David 2003	Belgium
Entoloma sp. [2]	GQ289296	GQ289228	GQ289156	TJ Baroni 9895 [CORT]	Belize
Entoloma sphagnetii	GQ289335		GQ289195	C. Bas 6.86	Netherlands
Entoloma tectonicola	GQ289336	GQ289266	GQ289196	P. Manimohan 741 [holotype]	Kerala, India
Entoloma tjallingiorum	GQ289337	GQ289267	GQ289197	J. Vauras 14318F	Finland
Entoloma trachyosporum [1]	GQ289338		GQ289198	H. den Bakker 1153	Canada
Entoloma trachyosporum [2]	GQ289339		GQ289199	H. den Bakker 1901	Canada
Entoloma transmutans	GQ289340	GQ289268	GQ289200	ME Noordeloos 2004155	Tasmania, Australia
Entoloma turbidum	GQ289341	GQ289269	GQ289201	ME Noordeloos 200351	Slovakia
Entoloma undatum	GQ289342	GQ289270	GQ289202	ME Noordeloos 200327	Belgium
Entoloma valdeumbonatum	GQ289343	GQ289271	GQ289203	M. Meusers E4565 [holotype]	Germany
Entoloma vezzenaense	GQ289344	GQ289272	GQ289204	A. Hausknecht [isotype, ex WU 14588]	Italy
Entoloma violaceovillosum	GQ289345	GQ289273	GQ289205	P. Manomohan 645 [holotype]	Kerala, India
Lepista ovispora	GQ289347	GQ289274	GQ289207	E. Arnolds 05-183	Netherlands
Tricholoma vaccinum	GQ289358	GQ289286	GQ289219	H. v.d. Burg 2004-11-03	Netherlands
Calocybe carnea [CBS552.50]	AF357097	DQ825423	AF223178
Clitocybe dealbata	AF357138	DQ825407	AF223175
Collybia tuberosa [AFTOL-ID 557]		AY787219	AY639884
Entoloma sp. [1; AFTOL-ID 523; identified as E. prunuloides]		DQ385883	AY700180
Lyophyllum leucophaeatum	AF357101	DQ367434	AF223202
*Rhodocybe aureicystidiata		AY337412	AY380407
Tephrocybe boudieri	AF357122	DQ825411	DQ825430

DNA markers

Three loci from three different parts of the genome were sequenced: RPB2 (nuclear RNA polymerase second largest subunit gene), LSU (nuclear ribosomal large subunit gene) and mtSSU (mitochondrial ribosomal small subunit gene). For RPB2, the primers bRPB2-6F and bRPB2-7R were used (Matheny 2005). For some samples these primers failed to amplify any fragment either due to its high degeneracy or degradation of the template DNA. Internal primers were thus designed particularly for Entolomataceae: rpb2-i6f (5’ GAA GGY CAA GCY TGY GGT CT 3’) and rpb2-i7r (5’ ATC ATR CTN GGA TGR ATY TC 3’). This new primer pair partially addressed the difficulty of amplification of RPB2 by being slightly less degenerate and by amplifying a shorter fragment. The large subunit of the nuclear ribosomal apparatus (LSU) was amplified and sequenced using LROR, LR16, LR3R and LR5 (more information from http://www.biology.duke.edu/fungi/mycolab/primers.htm). The primers used for mtSSU were MS1 and MS2 (White et al. 1990).

DNA extraction, amplification and sequencing

DNA was isolated from fresh lamellae preserved in a cetyl trimethylammonium bromide (CTAB) buffer and from dried herbarium material using a modified CTAB extraction method (Doyle & Doyle 1990). Varying amounts of fruitbody tissue were ground either by using a bleach-cleaned plastic pestle if the sample was CTAB-preserved, or by agitating the dried tissue with a 7 mm diam glass ball in a 2 mL microcentrifuge tube in a mixer mill (MM 200, Retch GmbH & Co, Germany) and adding a total of 500 μL 2XCTAB buffer afterwards. Proteinase K (2 μL of 20 mg/ml, 20 U/mg) and RNAase (1 μL of 10 mg/ml) were added and the tubes were incubated at 60–70 °C for 40 min. The material was twice mixed and centrifuged at 18 000 rpm for 15 min with an equal volume of chloroform-isoamyl alcohol (24 : 1), keeping the aqueous phase and placing it in a new tube each time. DNA was precipitated using an equal volume of isopropanol to the aqueous extract, an incubation time varying from 0 min to overnight in −20 °C, and centrifugation at 10 000 rpm for 15 min. The resulting pellet was washed with 70 % ethanol and air-dried, then re-suspended in 0.1X TE buffer. In the occasion that a thick brown liquid precipitated upon the addition of isopropanol, or the resulting DNA extract failed to work, the extract was either diluted up to 100 times or was further cleaned. The first two buffers (AP1 and AP2) of DNeasy Plant Mini Kit (QIAGEN, Germany) were added to remove further impurities. The resulting precipitate was separated from the solution by centrifugation, and the DNA once more precipitated, washed, dried and re-suspended.

Polymerase chain reaction amplifications (PCR) were generally performed in 25–50 μL reaction volumes. The recipe for a 25 μL volume is: 10 pmol for each primer for LSU or mtSSU or 30 pmol for RPB2, 1× PCR Buffer (QIAGEN, Germany), 1 μL DNTPs, 2 μL MgCl2, 0.5–1 μL BSA, 0.02 Taq polymerase. Products from multiple reactions were pooled if the products were low in concentration. The touch-down protocol used was: 5 min initial incubation at 94 °C, followed by cycles of 94 °C for 1 min, 67 °C annealing temperature for 1 min and 72 °C extension period of 1.5 min, with the annealing temperature decreasing by 1 °C every cycle until it reached 55–57 °C. A second round of 36 cycles was then used: 94 °C for 30–45 s, 55–57 °C for 1 min, and 73 °C for 1.5 min. The PCR protocol concluded with a 7-min final extension period. PCR products were visualized using agarose gel electrophoresis and ethidium bromide staining, and subsequently cleaned with the kit Nucleospin (Macherey-Nagel, Germany). When multiple bands were present, the bands of interest were cut out and cleaned according to kit instructions. When the resulting chromatograms were unreadable due to multiple signals and no better alternative specimen was available, cloning was performed to separate the strands of interest (using pGEM-T Easy Vector System and sequenced using the M13 primers with 35 cycles of 30 s at 95 °C, 30 s at 50 °C and 1 min at 72 °C).

Sequencing was performed either by cleaning with Sephadex G50 AutoSeq columns (GE Healthcare, Belgium) and run on an ABI 377 automated sequencer using the ABI BigDye Terminator chemistry for cycle sequencing (Applied Biosystems, USA), or by external services (using ABI 3730xl; Applied Biosystems, USA). The sequence chromatograms were processed using Sequencher v4.1.4 (Gene Codes Co., USA). The sequences generated for this study have been submitted to Genbank.

Multiple sequence alignments

Sequences were manually aligned in MacClade 4.06 (DR Maddison & WP Maddison, Sinauer Associates Inc., USA). In a few, small (< 12 bp), parts of the LSU and mtSSU alignment, it was difficult to unambiguously align the sequences across all taxa. If, in a section of the alignment, there was only a small fraction of taxa that could not be aligned with the remaining part, the unalignable portions (no more than 15 bp lengths) of these taxa were excluded from the analyses and were treated as missing. This procedure retained as much data as possible by preserving the information for the majority of the taxa where the alignment was unambiguous.

Conflict testing

Conflict between the RPB2, LSU and mtSSU datasets were evaluated in two ways: 1) using the ILD test (Farris et al. 1994) as implemented in PAUP* 4.b10 (Swofford 2002); and 2) by comparing phylogenetic analyses of single-locus datasets. In the second test, the results were compared to find conflicting branches in the tree that had > 70 % bootstrap support (both using maximum parsimony and maximum likelihood criteria) or that had 95 % posterior probability (p.p.) in Bayesian analyses. Some conflicts were found among the topologies of individual gene trees regarding the position of 6 species (see Results, Conflict testing). A second set of analyses that excluded these taxa was carried out for comparison to test if their inclusion had any effect on the phylogenetic reconstruction.

Phylogeny reconstruction

Maximum parsimony (MP) heuristic searches were performed using Parsimony Ratchet Analyses with PAUP* (PRAP) v1.21 using 200 ratchet replicates, 80 random addition cycles with 25 % of the characters weighted double (Müller 2004). Maximum parsimony bootstrap analyses were made with PAUP* 4.b10 (Swofford 2002) using 1 000 bootstrap replicates, each with 10 addition-sequence replicates using TBR branch swapping with a maximum of 10 trees saved per addition-sequence replicate. A bootstrap value of 70 % was considered significant.

Best-fit evolutionary models for the maximum likelihood and Bayesian analyses were selected for each single-locus dataset using MrModeltest 2.2 (Nylander 2004). The model GTR+I+G was indicated to be the best model to implement for all three loci.

Maximum likelihood (ML) analyses were performed using PHYML v3.0 (Guindon & Gascuel 2003). The following were implemented: GTR+G+I model of evolution, and four categories of the gamma distribution of the heterogeneity of the rates of evolution. SPR tree topology search was used and 1 000 bootstrap samples were used to calculate the maximum likelihood bootstrap support. A bootstrap value of 70 % was considered significant.

Bayesian analyses were performed using MrBayes v3.1.2p (Huelsenbeck & Ronquist 2001, Ronquist & Huelsenbeck 2003). In the 2- or 3-loci analyses, 2 or 3 partitions were set, respectively, each with a GTR+I+G model implemented. The prior on the gamma shape parameter was set to uniform ranging from 0.1 to 50. The following were implemented under the unlink command: revmat (substitution rates), pinvar (proportion of invariable sites), statefreq (character state frequencies) and shape (gamma shape parameter). Two runs, each with 11 chains were run with a temperature of 0.002 or 0.005, and three attempts at swapping every one or five generations. The topological convergence diagnostic (standard deviation of partition frequencies) was calculated every 10 000 generations but the stoprule was not implemented. The analyses were allowed to run up to 10 000 000 generations, sampling every 200 generations. More generations were added as necessary to reach convergence (as estimated by the topological convergence diagnostic equal to 0.01) between the two runs. In the case of the analysis of the LSU dataset, it reached an average standard deviation of split frequencies of only 0.014523 after 40 000 000 generations using 20 chains per run. Each LSU run was analysed separately and because the results were nearly identical in topology and support, one run was randomly chosen for the discussion and figures.

Scanning Electron Microscopy (SEM)

SEM pictures were taken to compare the taxa disputed to be either Rhodocybe or Entoloma (E. nitidum, E. trachyosporum) and compared with their supposed close relatives from Rhodocybe and Entoloma (see Table 1 where species samples for DNA and spores are noted). The spores were examined and scored for characters that define the difference between the classical entolomatoid and rhodocyboid spores:

the presence of isolated bumps and ridges (characteristic of rhodocyboid spores);

the presence of facets (characteristic of entolomatoid spores). If facets were present, then it was noted;

whether the facets were defined by a network of either incompletely or completely interconnected ridges. Ridges with an end that did not interconnect with another ridge were disregarded and not counted as irregular if the ridges ended towards the apiculus (and thus had no ridge to connect with) or if they symmetrically bisected a facet (with the argument that such bisections are regular).

Preparation of spores was from Baroni (1981), with the following modifications: preparations were washed twice in distilled water before dehydration for 20 min in 50 % acetone, followed by 20 min in 100 % acetone, then critical-point dried in a Balzers CPD 030 Critical Point Dryer (BAL-TEC, Liechtenstein) and sputter-coated in a SCD 005 Sputter Coater (BAL-TEC, Liechtenstein). Finally, SEM pictures were taken using JSM-530 Scanning Microscope (Jeol Ltd., Japan).

Transmission Electron Microscopy (TEM)

TEM photos were taken of some Entolomataceae as well as some of its close relatives in the Tricholomatoid clade sensu Matheny et al. (2006) within which Entolomataceae is nested (Hofstetter et al. 2002, Moncalvo et al. 2002, Matheny et al. 2006). These relatives sampled have bumpy or roughened spores: Tephrocybe tylicolor (Fr.) M.M. Moser, Tephrocybe ambusta (Fr.) Donk, Lepista irina (Fr.) H.E. Bigelow, Lepista nuda (Bull.) Cooke, and Omphaliaster asterosporus (J.E. Lange) Lamoure. It was noted what part of the spore wall formed the ornamentations to assess probably homologous structures.

Procedure follows Van der Ham (1990) with the following modifications: rehydration with glutaraldehyde was 3 w, fixing in OsO4 was for 1.5 h, pre-staining was with 1 % uranylacetate and soaking in lead citrate (Reynolds 1963) was for 10 min. The thickness of the cuts was 800 microns. The TEM machine was JEOL JEM-1010 Electron Microscope, JEOL Ltd. (Korea).

RESULTS

After sequencing and aligning, mtSSU yieled an alignment length of 360 bp and 95 parsimony-informative sites. RPB2 yielded an alignment length of 576 bp and 266 parsimony-informative sites. LSU yielded an alignment length of 707 bp and 169 parsimony-informative sites.

Of the three DNA loci sequenced, RPB2 yielded the best-resolved and best-supported phylogenetic reconstruction and LSU the least resolved and supported. The results of the 3-loci analyses are summarized in Fig. 2, while Fig. 3, 4, 5 show the Bayesian analyses of the single-gene datasets. The alignments of taxa with particularly long branch lengths (R. hirneola, C. prunulus) were examined. Both species were not misaligned and each had a large number of autapomorphies.

Fig. 2

Cladogram of Bayesian phylogenetic reconstruction using RPB2, LSU and mtSSU; phylogram of the same inset. Support is indicated with Bayesian p.p. / MP bootstrap percentage above the branches and ML bootstrap percentage below. Shapes at the ends of branches indicate spore type as determined using SEM except for Clitopilus spores which were determined through light microscopy: ★ regular entolomatoid spore, ▪ irregular entolomatoid spore, no bumps; / very irregular entolomatoid spore with bumps; rhodocyboid spore, clitopiloid spore. Subgenera of Entoloma species (according to Noordeloos 2004, Manimohan et al. 2006, Gates & Noordeloos 2007) is indicated as follows: ALB = Alboleptonia, ALL = Allocybe, CLA = Claudopus, CLI = Clitopiloides, ENT = Entoloma, INO = Inocephalus, LEP = Leptonia, NOL = Nolanea, PAR = Paraleptonia, POU = Pouzarella, TRI = Trichopilus.

Fig. 3

Phylogram of Bayesian analysis of RPB2. Bayesian p.p. indicated above the branch. Subgenera of Entoloma species (according to Noordeloos 2004, Manimohan et al. 2006, Gates & Noordeloos 2007) is indicated as follows: ALB Alboleptonia, ALL Allocybe, CLA Claudopus, CLI Clitopiloides, ENT Entoloma, INO Inocephalus, LEP Leptonia, NOL Nolanea, PAR Paraleptonia, POU Pouzarella, TRI Trichopilus.

Fig. 4

Phylogram of Bayesian analysis of mtSSU. Bayesian p.p. indicated above the branch. Subgenera of Entoloma species (according to Noordeloos 2004, Manimohan et al. 2006, Gates & Noordeloos 2007) is indicated as follows: ALB Alboleptonia, ALL Allocybe, CLA Claudopus, CLI Clitopiloides, ENT Entoloma, INO Inocephalus, LEP Leptonia, NOL Nolanea, PAR Paraleptonia, POU Pouzarella, TRI Trichopilus.

Fig. 5

Phylogram of Bayesian analysis of LSU. Bayesian p.p. indicated above the branch. Subgenera of Entoloma species (according to Noordeloos 2004, Manimohan et al. 2006, Gates & Noordeloos 2007) is indicated as follows: ALB Alboleptonia, ALL Allocybe, CLA Claudopus, CLI Clitopiloides, ENT Entoloma, INO Inocephalus, LEP Leptonia, NOL Nolanea, PAR Paraleptonia, POU Pouzarella, TRI Trichopilus.

Evaluation of conflicts by comparing the different trees from the single-gene analyses led to the conclusion that gene trees were mostly compatible and had no significantly supported conflicts (> 70 % MP or ML bootstrap or 0.95 Bayesian posterior probability) among the resulting reconstructions with the exception of two taxa: C. cystidiatus and E. sarcitum.

In the phylogenetic reconstructions with RPB2 and LSU, C. cys-tidiatus was strongly supported as sister group to C. prunulus in the Rhodocybe-Clitopilus clade. On the other hand, in the mtSSU reconstruction, C. cystidiatus was sister to E. prunuloides with 75 % MP bootstrap and 91 % ML bootstrap. These two taxa were placed within the insignificantly supported Entoloma clade.

The placement of E. sarcitum was different in the phylogenetic reconstructions of all three loci. It was placed in the Inocephalus-Cyanula clade in the ML and Bayesian analyses of mtSSU (57 % ML bootstrap and 0.99 Bayesian posterior probability; MP analyses were unresolved). Using LSU, E. sarcitum was strongly supported in a clade with E. pluteisimilis and R. zuccherellii (100 % MP bootstrap, 100 % ML bootstrap, and 1.00 Bayesian posterior probability), but with its relationship to others in the Entoloma clade was unresolved. In contrast, ML and Bayesian analyses of RPB2 placed it as sister to most of the genus Entoloma excluding the Prunuloides clade (that is, the rest of the crown Entoloma clade; 0.98 Bayesian posterior probability and 64 % ML bootstrap), while it was unresolved in the MP analysis.

Four other taxa had different positions in the single-gene analyses, but these placements did not receive significant support. Their strongly supported placements in the 3-loci analyses are more consistent with morphological studies. The first, E. kermandii, is in the Prunuloides clade in the Bayesian and ML analyses of RPB2 and mtSSU, while the same analyses of LSU place it among members of the Inocephalus-Cyanula clade. Second, Calocybe carnea (selected as outgroup) was in the Bayesian and ML analyses in one clade with Rhodocybe. Third and fourth, Bayesian and ML analyses of mtSSU had Lyophyllum boudieri in the Rhodocybe-Clitopilus clade while R. hirneola (Fr.) P.D. Orton was with the rest of the outgroup. The consensus tree of the most parsimonious trees of these analyses did not show the same unexpected placement, but it had poor resolution.

Analyses of single-gene and 3-loci datasets after the exclusion of these 6 taxa produced trees that were very similar with respect to topology and the level of support of the recognized clades, with some exceptions in the 3-locus dataset. The Bayesian posterior probability values for the monophyly of E. pluteisimilis, R. zuccherellii, E. sphagneti, and the rest of Entoloma excluding the Prunuloides clade changed from 0.96 to 0.84. Similarly, in the ML analyses, bootstrap support for the Nolanea-Claudopus clade decreased from 70 % to 60 %. On the other hand, support for the monophyly of a clade composed of Nolanea-Claudopus, Inocephalus-Cyanula, Pouzarella and the Rhodopolioid clades increased from 88 % to 100 %. Nonetheless, the phylogenetic reconstructions and conclusions drawn were not affected by these changes.

The ILD test indicated that while RPB2 and mtSSU were not significantly incongruent (P = 0.133), LSU was incongruent with both RPB2 and mtSSU (both P = 0.001). In order to test whether the taxa mentioned above caused the incongruence, the ILD test was once again performed without these taxa. The new analysis yielded the same conclusions.

— Fig. 6

Three main spore types could be recognized:

clitopiloid spores with longitudinal grooves and ridges;

rhodocyboid spores with irregular bumps and ridges;

entolomatoid spores with facets.

Entolomatoid spores are subdivided into three subtypes:

regular entolomatoid spores with ridges that completely interconnect to form facets and that have no isolated bumps;

irregular entolomatoid spores with at least one ridge end not connecting with another ridge to delineate a facet, giving a slightly to very irregular look, but without bumps and;

very irregular entolomatoid spores with bumps.

Irregular entolomatoid spores have not been reported before and bumps have previously not been known to occur in Entoloma (Baroni & Largent 1989). The degree of irregularity is highly variable among species. The regular and irregular entolomatoid spores grade into each other. Some very irregular spores with bumps are similar at first glance to some rhodocyboid spores with many ridges. However, regular and irregular entolomatoid spores are united by the presence of facets, and these are never present in rhodocyboid spores.

The spore wall of species with irregular entolomatoid spores is usually thinner than that of regular entolomatoid spores when viewed under a light microscope. We cannot exclude the possibility that the spores are truly regular, and that the apparent irregularity of the spore ridges is an artifact of spore preparation for SEM due to the thin spore walls. If this is the case, it is a phylogenetically informative artifact.

The spore types are mapped on the 3-loci phylogenetic reconstruction in Fig. 2. The Prunuloides clade is characterized by mostly irregular entolomatoid spores, with one subclade predominantly having very irregular spores with bumps and the other subclade having irregular spores without bumps. The Crown Entoloma clade, having regular entolomatoid spores (based on observations under the light microscope and SEM pictures by Pegler & Young 1978, 1979), contrasts with the rest of Entoloma (Prunuloides clade, the clade of E. plutei-similis and R. zuccherellii, and E. sarcitum), as the latter set of species have mostly irregular entolomatoid spores (with or without bumps).

— Fig. 7

Comparison of the spore wall structures in Entolomataceae and five representatives of the Tricholomatoid clade sensu Matheny et al. (2006) with ornamented spores showed that the structures forming the angularity or bumps in the Entolomataceae are different from those of the ornamentation of its closest relatives; none of the latter are formed with an epicorium and tunica.

DISCUSSION

The two tests for conflict yielded two kinds of apparent conflict. In the first instance, taxa had different positions according to different single-locus analyses. In the second instance, the ILD test indicated that analyses based on both mtSSU and RPB2 were incongruent with LSU. In the first case, phylogenetic analyses performed after the removal of the relevant taxa resulted in no significant changes to the phylogeny. In the case of the ILD test, the 2-loci phylogenetic analyses with just the supposedly congruent RPB2 and mtSSU yield phylogenetic reconstructions similar to the results of 3-loci analyses, supporting the conclusions of this study. We conclude that RPB2 and LSU informed the analyses more than LSU. Furthermore, there are cases where the ILD test fails (Dolphin et al. 2000, Darlu & Lecointre 2002) and this may have been the case here.

Monophyly of the Entolomataceae

The results of this molecular phylogenetic study are consistent with recent studies, based on both morphology and molecular phylogenetics data, that the family Entolomataceae is a natural group (Singer 1986, Noordeloos 1981, 1992, 2005) and is monophyletic (Matheny et al. 2006). Entolomataceae comprises the genera Entoloma, Rhodocybe, Clitopilus, Richoniella and Rhodogaster, and probably Rhodocybella as well. This study confirms that the presence of pink, angular or bumpy spores with a well-developed epicorium in the spore wall, which forms the facets, ridges or bumps and which was traditionally used to define the family Entolomataceae, has evolved only once among the euagarics. The ornamentations found among relatives, the members of the Tricholomatoid clade, are different and not homologous with those of Entolomataceae, as noted already before by Kühner (1980) and Clemençon et al. (2004).

Generic delimitation of Entolomataceae

Entolomataceae is divided into two monophyletic clades: one containing Rhodocybe and Clitopilus, and the other containing Entoloma, Richoniella and Rhodogaster.

In the Rhodocybe-Clitopilus clade, Rhodocybe is paraphyletic and Clitopilus is, with significant support, well-nested within Rhodocybe. Previous studies had already suggested this relationship (Moncalvo et al. 2002, Matheny et al. 2006). Some Rhodocybe spores have bumps and ridges arranged linearly along the spore length (Baroni 1981), presenting a spore shape that approaches clitopiloid spores. Kühner (1980) had already united Clitopilus with Rhodocybe in one genus on account of the characters these groups share: predominantly clitocyboid habit, decurrent lamellae, and resemblance in basal structure of the ornamentation of the spores. We formally merge Rhodocybe into Clitopilus (the oldest valid name) and make the necessary new combinations (see Appendix).

We consider it highly likely that future molecular phylogenetic studies will show that the cyphelloid Rhodocybella is nested within the Rhodocybe-Clitopilus clade. If so, its generic status will be untenable. This likelihood is indicated by the genus’ rhodocyboid spores and the existence of somewhat similar, reduced forms (pleurotoid rather than cyphelloid) in the clade.

Delimitation of the genus Entoloma

Phylogenetic analyses demonstrate that the sequestrate genera Richoniella and Rhodogaster arose from within Entoloma. Therefore, Entoloma is monophyletic only if these genera are included. Placement of these sequestrate genera within Entoloma had been suggested before (Kuyper in Kirk et al. 2008).

Richoniella and Rhodogaster do not form a monophyletic group. The sequestrate habit arose at least twice from agaricoid ancestors within Entoloma. Only one species of each genus was included in these studies and so the monophyly of each genus could not be tested. Future studies can address this possibility of parallel evolution. It is generally accepted that sequestrate taxa were derived many times from agaricoid forms and that this character was much overemphasized in the past (Peintner et al. 2001). We emend the description of Entoloma to include these sequestrate taxa (see Appendix).

The independent evolution of Rhodogaster and Richoniella shows that Rhodogaster (secotioid, and intermediate in form between agaricoid and gasteroid sporocarps) is not an evolutionary intermediate or the missing link between Entoloma and Richoniella.

Should Entoloma be maintained as one genus or should it be split, based on the results of our phylogenetic analyses? While a phylogeny per se allows different answers to that question, we propose it is best to retain Entoloma as a very large and morphologically variable genus in order to have a stable classification that is consistent with phylogeny. Our proposal to retain one large genus Entoloma conforms to the taxonomic tradition (Romagnesi 1974, 1978, Noordeloos 2004). Relevant nomenclatural changes are made (see Appendix).

The inclusion of Rhodogaster and Richoniella in a monophyletic Entoloma strengthens our argument. If Entoloma had not been monophyletic, alternative proposals of smaller genera would have been strengthened.

The currently proposed smaller subgenera are polyphyletic or paraphyletic, leaving one large genus indeed as the only viable option. Horak’s (1973, 1980) division of Entoloma s.l. into three genera is inconsistent with phylogeny. Horak separated the strikingly morphologically distinct Pouzarella and Claudopus (pleurotoid species only) from the rest of Entoloma and treated them on a generic level. However, this classification would result in a paraphyletic Entoloma s.s. and a polyphyletic Claudopus. Among the 13 proposed genera of Largent (1994), this study shows that at least five genera, viz. Entoloma, Leptonia, Nolanea, Alboleptonia, and Calliderma are not monophyletic. Of these, the first four are the largest genera containing most of the known species and the only major genus not demonstrably polyphyletic is Inocephalus. The monophyly of the other seven genera can also neither be disproven nor supported with the current phylogenetic reconstruction due to lack of sampling. Most species of Entoloma s.s. are in two different strongly supported clades, one of which, the Rhodopolioid clade, is more closely related to members of Nolanea or Leptonia than to the other, the Prunuloides clade. Leptonia species are in three different clades and Alboleptonia in two. Species, which show morphological characters of Calliderma, viz., E. indigoticoumbrinum and E. griseolazulinum, are nested and not monophyletic within the Prunuloides clade. Furthermore, Nolanea is paraphyletic since the well-supported clade containing Nolanea also contains members of Entoloma s.s. and Alboleptonia (Fig. 2).

A phylogenetic study of the genus Entoloma using a larger dataset is currently in progress (Co-David & Noordeloos, unpubl. data). This study will use both molecular and morphological data to distinguish well-supported monophyletic clades, with implications for infrageneric taxonomy and character evolution.

The distinction between Rhodocybe and Entoloma

We reject the proposed transfer of E. nitidum and E. trachyosporum to Rhodocybe (Baroni & Largent 1993, Castellano et al. 1999). These taxa belong, with significant support, to the Prunuloides clade of Entoloma. Rhodocybe zuccherellii also belongs to Entoloma and we make the transfer.

SEM pictures revealed that their spores have facets, which rhodocyboid spores do not (E. nitidum and E. trachyosporum on Fig. 6). The confusion regarding their generic placement likely arose because the pattern of their spores, and those of many of their close relatives, is irregular. The ridges that define the facets do not always interconnect, and sometimes there are bumps. This irregularity as well as the difficulty in seeing facets under the light microscope may be because the ridges themselves are thinner and not as distinct as those in spores with regular facets.

Fig. 6

SEM pictures of Entolomataceae spores. a, b. Regular entolomatoid spores. a. Entoloma indigoticoumbrinum; b. E. sp. — c, d. Slightly irregular entolomatoid spores. c. E. kermandii; d. E. bloxamii. — e, f. Very irregular entolomatoid spores with bumps. e. E. nitidum; f. E. trachyosporum. — g, h. Rhodocyboid spores. g. Rhodocybe nitellina; h. R. caelata.

The placement of these taxa in Entoloma also makes sense in the light of morphology. The presence of clamps distinguishes them from most of the taxa of Rhodocybe except for section Rhodophana, which does have abundant clamp-connections. A morphological character that distinguishes the relevant Entoloma species from Rhodophana is pileipellis structure: Species in Rhodophana have a compact, uni-layered cutis gradually passing into the trama without a well-developed subpellis, while these Entoloma species have a bi-layered pileipellis with a thin suprapellis of narrow hyphae, and a well-developed subpellis of inflated elements (Baroni 1981, Noordeloos 2004).

The monophyly of Entolomataceae shows that pink, angular spores arose once. Given that the family is nested in the Tricholomatoid clade (sensu Matheny et al. 2006), its ancestral spore likely arose from white, smooth spores that are common in the clade.

The ancestral spore of Entolomataceae is yet unknown. However, it was either rhodocyboid or entolomatoid. Both reconstructions are consistent with the phylogeny. Clitopiloid spores are definitely not the ancestral spores of Entolomataceae. Rather, clitopiloid spores arose from rhodocyboid spores. This conclusion is further supported by observations that some rhodocyboid spores are more elongate and have their ridges and bumps aligned in rows along the length of the spore, similar to clitopiloid spores (Baroni 1981). Consequently, we reject Kühner’s (1980) theory that the spores of Clitopilus are intermediate between Rhodocybe and Entoloma because it is inconsistent with the phylogenetic reconstruction, where Clitopilus, is well-nested within Rhodocybe. Clitopilus is also too distant from Entoloma to have shared an ancestor with clitopiloid spores. Baroni’s (1981) theory that clitopiloid and entolomatoid spores were both independently derived from rhodocyboid spores is consistent with the phylogenetic reconstruction.

Another hypothesis must be taken into account, because it is equally consistent with the phylogeny, namely that the entolomatoid rather than rhodocyboid spore type is ancestral. In this theory, entolomatoid spores (regular or irregular) gave rise to rhodocyboid spores that, in turn, gave rise to clitopiloid spores. We suggest two major reasons why this hypothesis (the entomolatoid spore as ancestral character state) has never been proposed before.

First is the assumption that Entolomataceae evolved from Tricholomataceae combined with the assumed homology between bumps on Rhodocybe spores and the bumps on the spores of the putative Tricholomataceae ancestral taxa (Kühner 1980, Baroni 1981). These bumpy-spored relatives of the Tricholomatoid clade are nested among their smooth-spored relatives, as is Entolomataceae (Moncalvo et al. 2002, Matheny et al. 2006), making it likely that these taxa all evolved spore wall ornamentations independently. Furthermore, the putative homology between species with bumpy spores itself is incorrect. Our study confirms that the spore walls of species in the Tricholomatoid clade with bumpy spores are not homologous to those found in Entolomataceae (as earlier implied by Clémençon 1997, Clémençon et al. 2004). This lack of homology refutes the possible argument for rhodocyboid ancestral spores that members of the Tricholomatoid clade have a spore feature that allows it to frequently, independently evolve rhodocyboid spores.

The second reason for the predominance of the hypothesis that rhodocyboid spores are ancestral is the implicit assumption on the direction of spore character evolution. The assumption is that, in evolution, complex structures are more derived (or advanced) than simple structures. This assumption is not always the case, the derivation of more simple sequestrate forms from more complex agaricoid ones being an example (Peintner et al. 2001, this study). Also, the assumption that if there is a morphological gradation (smooth spores to bumpy spores to faceted spores), then evolution must have occurred along the direction of the gradation is not necessarily true either. Therefore, we must consider both the rhodocyboid and entolomatoid spore type as possible ancestral states for Entolomataceae spores until further studies disprove either possibility.