Mycoscience
Online ISSN : 1618-2545
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Taxonomic and ecological significance of synnema-like structures/acanthophyses produced by Physisporinus (Meripilaceae, Polyporales) species
Ryotaro Shino , Kozue Sotome, Naoki Endo, Nitaro Maekawa, Akira Nakagiri
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Keywords: freshwater, Meripilus, oxygen concentration, phylogenetic analysis, Rigidoporus
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2023 Volume 64 Issue 6 Pages 136-149

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Abstract

Physisporinus, a genus in Polyporales, Basidiomycota, is a versatile fungus that lives as a wood decomposer, a potential pathogen of standing trees, and an orchid mycobiont. We previously reported that some Physisporinus species inhabiting wet wood in aquatic environments such as streams and waterfalls form synnema-like structures (SSs) bearing acanthophyses at their apices, and that they produce acanthophyses on vegetative hyphae when cultured on agar media. In this study, we investigated the acanthophysis-forming ability in Physisporinus and allied genera, and experimentally demonstrated the function of SSs. Phylogenetic analyses and observations of Meripilus, Physisporinus and Rigidoporus cultures showed that all of the strains forming acanthophyses belonged to Physisporinus, whereas strains of Meripilus and Rigidoporus did not produce acanthophyses. These findings suggest that SS/acanthophysis formation is a useful taxonomic character for members of Physisporinus. When Physisporinus strains were cultured under oxygen (O2) concentrations of 5, 10, 20 and 40%, most of those cultured under 20% O2 formed the most acanthophyses. According to these experimental data, the SSs/acanthophyses in Physisporinus were considered to have a respiratory function. Physisporinus probably acquired the SS/acanthophysis-forming ability to adapt to moist and/or aquatic habitats and to decay wet wood in which the O2 concentration is often low.

1. Introduction

Physisporinus P. Karst. is a poroid fungus in Meripilaceae, Polyporales, Agaricomycetes, Basidiomycota (Justo et al., 2017). This cosmopolitan genus mainly decays dead broad-leaved and coniferous trees (Breitenbach & Kränzlin, 1986; Dai, 2012; Gilbertson & Ryvarden, 1987; Núñez & Ryvarden, 2001; Ryvarden et al., 2022; Ryvarden & Gilbertson, 1994; Ryvarden & Melo, 2017). Some species may cause butt rot in living Japanese cedars [Noguchi et al., 2007 (as Basidiomycete-B)], or establish mycorrhizal relationships with mycoheterotrophic orchids (Yamashita et al., 2020). Based on morphological and phylogenetic studies of aquatic fungi inhabiting wet wood in streams, we previously reported that five clades of fungi in the genus Physisporinus, i.e., two new species (P. microacanthophysis Shino, Sotome & Nakagiri and P. rhizomorphae Shino, Sotome & Nakagiri) and three unidentified groups (P. cf. 1 eminens, P. cf. 2 eminens, and P. cf. furcatus) form synnema-like structures (SSs) and produce numerous acanthophyses at their apices (Shino et al., 2022). Since cultures isolated from SSs or basidiocarps produced several types of acanthophyses on agar media, we considered that these characters might be useful as a taxonomic trait of Physisporinus. However, our previous study did not examine whether genera that are closely related to Physisporinus also produce SSs/acanthophyses. Phylogenetically, Physisporinus is closely related to both Meripilus P. Karst. (Chen & Dai, 2021; Shino et al., 2022; Tomšovský et al., 2010), which is the type genus of the Meripilaceae (Binder et al., 2013; Jülich, 1981; Justo et al., 2017), and Spongipellis Pat. (Kotiranta et al., 2017; Spirin et al., 2022; Wang & Dai, 2022). Spongipellis forms basidiocarps composed of a duplex context and generative hyphae with clamp connections (Ryvarden, 1991; Spirin et al., 2022) and can be distinguished from Physisporinus having a simplex context and generative hyphae without clamp connections in the basidiocarps (Gilbertson & Ryvarden, 1987). Meripilus species produce pileate basidiocarps with single to numerous brownish pilei arising from a short stipe or a base. This type of basidiocarp differs from the whitish resupinate basidiocarps found in Physisporinus, but the micromorphological features in basidiocarps of Meripilus (monomitic hyphal system, generative hyphae without clamp connections, and smooth and broadly ellipsoid to subglobose basidiospores with inamyloid reaction in Melzer's reagent) are similar to those found in Physisporinus (Gilbertson & Ryvarden, 1987). These characteristics are also observed in Leucophellinus Bondartsev & Singer, Oxyporus (Bourdot & Galzin) Donk, and Rigidoporus Murrill, which have been phylogenetically assigned to Hymenochaetales (Wu et al., 2017). Leucophellinus, which produces clavate and occasionally septate cystidia and has distinctly thick-walled basidiospores (Núñez & Ryvarden, 2001), is distinguishable from Physisporinus and related genera. Many species of Oxyporus and Rigidoporus resemble each other in that they form cystidia in their basidiocarps, but these taxa differ from Physisporinus, which has no cystidia (Ryvarden & Gilbertson, 1994). Regarding the former two genera, Pouzar (1966) treated Oxyporus as a subgenus of Rigidoporus because of their morphological similarities including the above features. However, several mycologists (Corner, 1987; Donk, 1967; Ryvarden & Johansen, 1980) proposed that these genera should remain separate because Rigidoporus typically produces basidiocarps with bright colors and forms cystidia in tramae while Oxyporus has pale-colored basidiocarps with cystidia in hymenia (Ryvarden, 1991). Recently, Wu et al. (2017) integrated Oxyporus into Rigidoporus since their phylogenetic analysis showed that the type species of the two genera grouped in the same clade in Hymenochaetales. Moreover, they transferred part of the remaining species of Rigidoporus, which were found belonging to Polyporales, to Physisporinus. As a result of this and other studies, Physisporinus currently accommodates several species that form apically encrusted cystidia [e.g., P. eminens (Y.C. Dai) F. Wu, Jia J. Chen & Y.C. Dai, formerly treated as R. eminens Y.C. Dai (Dai, 1998); P. furcatus (Núñez & Ryvarden) F. Wu, Jia J. Chen & Y.C. Dai, formerly R. furcatus Núñez & Ryvarden (Núñez et al., 2001); P. lineatus (Pers.) F. Wu, Jia J. Chen & Y.C. Dai, formerly R. lineatus (Pers.) Ryvarden (Ryvarden, 1972; Ryvarden & Johansen, 1980); P. pouzarii (Vampola & Vlasák) F. Wu, Jia J. Chen & Y.C. Dai, formerly R. pouzarii Vampola & Vlasák (Vampola & Vlasák, 2012)] and have pore surfaces with vivid colors when fresh [e.g., P. lavendulus F. Wu, Jia J. Chen & Y.C. Dai (Wu et al., 2017); P. roseus Jia J. Chen & Y.C. Dai (Chen & Dai, 2021); P. sulphureus Y.C. Dai (Dai & Dai, 2018)]. Thus, Physisporinus and Rigidoporus have become difficult to clearly distinguish by the morphology of their basidiocarps. Hence, the first objective of this study is to evaluate the taxonomic significance of SS/acanthophysis formation among Physisporinus, phylogenetically related genus Meripilus and morphologically similar genus Rigidoporus, and to identify other taxonomically important characters found in their cultures, such as formation of clamp connections at hyphal septa, as well as the presence of conidia and plectenchymata in mycelia.

We previously reported that acanthophyses on the apices of SSs are not conidia because they neither easily detach from SSs nor germinate hyphae (Shino et al., 2022). Since SSs have been often found at the water-boundary part of wet wood in aquatic environments such as streams and waterfalls, we hypothesized that the SSs of Physisporinus may be associated with the respiration for mycelia creeping in the water-saturated wood tissue where oxygen (O2) levels tend to be lower than in the atmosphere. Therefore, as the second objective of this study, we aim to verify this hypothesis by experiments using cultures of Physisporinus and to discuss the ecological significance of SSs/acanthophyses.

2. Materials and methods

2.1. Samples

24 specimens and 41 strains were tested in this study (Figs. 1, 6; Table 1). Procedures for the establishment of dried specimens and living isolates followed Shino et al. (2022). We also used strains preserved in the Fungus/Mushroom Resource and Research Center (FMRC), Faculty of Agriculture, Tottori University, and strains obtained from the Westerdijk Fungal Biodiversity Institute, Utrecht, the Netherlands (Table 1).

Fig. 1 - Basidiocarps (A-C), SSs (D) bearing acanthophyses (E), and rhizomorphs (F) of Physisporinus in nature. A: Whitish basidiocarps produced on wet wood nearby streams (P. cf. 2 eminens TUMH 65445). B: Basidiocarps (P. cf. 1 eminens TUMH 65440). C: Pore surface of basidiocarps (P. cf. 1 eminens TUMH 65442). D: SSs on the water-boundary part of wood in streams (the source for P. pouzarii TUFC 101965). Arrowheads show parts forming acanthophyses. E: Acanthophyses on the apex of SS (P. microacanthophysis TUMH 64311). F: Rhizomorphs on the submerged part of wood in streams (P. rhizomorphae TUMH 64298). Bars: C, D 1 mm; E 30 µm.
Table 1. Data of samples used in this study.

Species name Strain No. a Herbarium Specimen No. Locality Collection date Isolation date Source b Habitat c
Meripilus giganteus CBS 421.48 - Germany - - - -
M. giganteus TUFC 100564 (TUMH 60367)d Tottori Pref., Japan 17 Aug 2012 17 Aug 2012 T F
Physisporinus cf. 1 eminens TUFC 101880 TUMH 64307 Miyagi Pref., Japan 22 Sep 2014 22 Sep 2014 S A
P. cf. 1 eminens TUFC 101957 TUMH 65440 Tottori Pref., Japan 05 Nov 2019 05 Nov 2019 B A
P. cf. 1 eminens TUFC 101958 TUMH 65441 Tottori Pref., Japan 05 Nov 2019 05 Nov 2019 B A
P. cf. 1 eminens TUFC 101959 TUMH 65442 Tottori Pref., Japan 05 Nov 2019 05 Nov 2019 B A
P. cf. 2 eminens TUFC 101881 TUMH 64308 Tottori Pref., Japan 25 Jun 2014 25 Jun 2014 S A
P. cf. 2 eminens TUFC 101960 TUMH 65443 Hokkaido Pref., Japan 26 Sep 2017 26 Sep 2017 B F
P. cf. 2 eminens TUFC 101961 TUMH 65444 Tottori Pref., Japan 14 Oct 2017 14 Oct 2017 B A
P. cf. 2 eminens TUFC 101962 TUMH 65445 Tottori Pref., Japan 30 Oct 2019 30 Oct 2019 B A
P. cf. furcatus TUFC 101883 TUMH 64310 Gifu Pref., Japan 29 May 2012 29 May 2012 S A
P. cf. furcatus TUFC 101884 TUMH 64310 Gifu Pref., Japan 29 May 2012 29 May 2012 S A
P. crocatus CBS 107806 - Canada - 1982 - -
P. lineatus CBS 167.65 - USA - 24 Aug 1960 - -
P. lineatus CBS 700.94 - Germany - - - -
P. lineatus CBS 109425 - Taiwan 27 Aug 1996 - - -
P. lineatus TUFC 13809 TUMH 60931 Tokyo Metropolis, Japan 19 Nov 2010 - T F
P. lineatus TUFC 13812 TUMH 60932 Tokyo Metropolis, Japan 19 Nov 2010 - T F
P. microacanthophysis TUFC 101885 TUMH 64311 Tottori Pref., Japan 07 Jul 2011 07 Jul 2011 S A
P. microacanthophysis TUFC 101888 TUMH 64312 Miyazaki Pref., Japan 24 Nov 2013 24 Nov 2013 S A
P. microacanthophysis TUFC 101889 TUMH 64313 T e Tottori Pref., Japan 23 Oct 2019 23 Oct 2019 B A
P. microacanthophysis TUFC 101963 TUMH 65446 Tottori Pref., Japan 04 Apr 2020 16 Apr 2020 S A
P. microacanthophysis TUFC 101964 TUMH 65447 Tottori Pref., Japan 19 May 2020 19 May 2020 B A
P. pouzarii TUFC 101965 No specimen Osaka Pref., Japan 01 Apr 2013 01 Apr 2013 S A
P. pouzarii TUFC 101966 TUMH 65448 Tottori Pref., Japan 05 Nov 2019 05 Nov 2019 B A
P. rhizomorphae TUFC 101870 TUMH 64297 Tottori Pref., Japan 24 Sep 2013 24 Sep 2013 S A
P. rhizomorphae TUFC 101871 TUMH 64298 Tottori Pref., Japan 19 Oct 2014 19 Oct 2014 R A
P. rhizomorphae TUFC 101876 TUMH 64303 T Tottori Pref., Japan 14 Oct 2017 14 Oct 2017 B A
P. rhizomorphae TUFC 101967 TUMH 65449 Tottori Pref., Japan 04 Apr 2020 08 May 2020 S A
“P. sanguinolentus” CBS 139.76 - Belgium Sep 1975 - - -
“P. sanguinolentus” CBS 193.76 - Netherlands - - - -
“P. sanguinolentus” CBS 679.70 - USA - - - -
“P. sanguinolentus” CBS 107146 - Denmark - 1980 - -
Physisporinus sp. TUFC 101892 TUMH 64316 Kagoshima Pref., Japan 04 Sep 2018 04 Sep 2018 B A
Physisporinus sp. TUFC 101968 TUMH 65450 Tottori Pref., Japan 05 Sep 2020 05 Sep 2020 T F
Rigidoporus ulmarius CBS 186.60 - USA 05 Nov 1952 - T f -
“R. vinctus” CBS 153.84 - New Zealand - 16 Oct 1973 - -
“R. vinctus” CBS 174.71 - Costa Rica 20 Jun 1963 - - -
“R. vinctus” TUFC 11175 (TUMH 60851) Kagoshima Pref., Japan 19 Sep 2007 19 Sep 2007 B F
“R. vinctus” TUFC 13815 (TUMH 63562) Tokyo Metropolis, Japan 20 Nov 2010 - B F
“R. vinctus” TUFC 35082 (TUMH 60896) Okinawa Pref., Japan 13 Jul 2003 13 Jul 2003 B A

a Strains in bold were used for the experiments incubating cultures under different O2 concentrations.

b “B”, “R”, “S”, and “T” mean basidiospores, a rhizomorph, a SS, and tissue of the basidiocarp as the source of isolation, respectively.

c “A” means that the sample was collected in or nearby an aquatic area. “F” means the sample from a forest area, not aquatic.

d Parenthesis means the specimen that we did not examine in the present study.

e T means the type specimen.

f FP 103737, other strain number of CBS 186.60, was isolated from tissue of the basidiocarp (Lombard et al., 1960).

2.2. Molecular phylogeny

2.2.1. DNA extraction, amplification and sequencing

DNA extraction from mycelia cultured on agar media was performed using a modified cetyltrimethylammonium bromide (CTAB) method (Shino et al., 2022). From the obtained genomic DNA, the internal transcribed spacer (ITS) region and D1/D2 domains of the large subunit (LSU) of nuclear ribosomal DNA (nrDNA) were amplified by the polymerase chain reaction (PCR) using a thermal cycler (PC-812 or PC-818; ASTEC Co., Ltd., Fukuoka, Japan). As primers, we used ITS5 and ITS4 for the ITS region (White et al., 1990), and LR0R and LR5 for the LSU region (Rehner & Samuels, 1994; Vilgalys & Hester, 1990). PCRs were conducted using the protocol described in Shino et al. (2022). Amplicons were purified using NucleoSpin Gel and PCR Clean-up (Takara Bio Inc., Shiga, Japan), and Fasmac Co., Ltd. (Kanagawa, Japan) was commissioned to perform the DNA sequencing. All of the sequences except the ITS regions of CBS 186.60 and TUFC 101965 were readable by direct sequencing. Of the above two samples showing partial heterogeneity between the gene copies, we performed a cloning for TUFC 101965 using pGEM-T Easy Vector Systems (Promega K.K., Tokyo, Japan) and competent bacterial cells (Escherichia coli (Migula) Castellani & Chalmers JM109). Sequence data were deposited at the DNA Data Bank of Japan (DDBJ; https://www.ddbj.nig.ac.jp/index-e.html).

2.2.2. Sequence alignment and phylogenetic analyses

Alignment of the data sets and creation of phylogenetic trees were performed online using MAFFT v. 7 (Katoh & Standley, 2013; https://mafft.cbrc.jp/alignment/server/; Jun 2023) and MEGA7 (Kumar et al., 2016). DNA sequences of the ITS and/or LSU regions of nrDNA retrieved from the GenBank database (https://www.ncbi.nlm.nih.gov/genbank/) were included in phylogenetic analyses which were performed using the maximum likelihood (ML) method. Based on the results of the best-fitting model test in MEGA7, the GTR+G+I model was adopted as a model of molecular evolution in the ML analyses using a combined data set of nrDNA ITS and LSU sequences for the Meripilaceae group (Meripilus, Physisporinus, and Spongipellis) and only nrDNA ITS for the Cerrenaceae group [Cerrena Gray, Irpiciporus Murrill, Pseudolagarobasidium J.C. Jang & T. Chen, Pseudospongipellis Y.C. Dai & Chao G. Wang, Radulodon Ryvarden, “Rigidoporus hypobrunneus” (Petch) Corner, and “R. vinctus” (Berk.) Ryvarden] in Polyporales, whereas the TN93+G model was applied to only nrDNA LSU for Rigidoporus in Hymenochaetales. The confidence coefficient of each node in the phylogenetic trees was confirmed by bootstrap (BS) analysis with 1,000 replicates (Felsenstein, 1985). Outgroups for the phylogenetic analyses of each data set for the above three clusters were selected as follows; Abortiporus biennis (Bull.) Singer (FD-319), Hyphoderma setigerum (Fr.) Donk (FD-312), and Hypochnicium sp. (FP-110227-sp) (Floudas & Hibbett, 2015; Justo et al., 2017) for the data set consisting of the nrDNA ITS and LSU sequences of the Meripilaceae, Polyporales group, following Yamashita et al. (2020); Cymatoderma sp. (OMC-1427), Panus conchatus (Bull.) Fr. (Miettinen 13966), and P. fragilis O.K. Mill. (HHB-11042-Sp) (Floudas & Hibbett, 2015; Justo et al., 2017; Miettinen et al., 2012) for the data set consisting of nrDNA ITS sequences of the Cerrenaceae, Polyporales group, referring to Justo et al. (2017); Bridgeoporus sinensis (X.L. Zeng) F. Wu, Jia J. Chen & Y.C. Dai (Cui 10013), Leucophellinus hobsonii (Berk. ex Cooke) Ryvarden (Cui 6468), and L. irpicoides (Bondartsev ex Pilát) Bondartsev & Singer (Yuan 2690) (Wu et al., 2017) for the data set consisting of nrDNA LSU sequences of Rigidoporus, Hymenochaetales, referring to Wu et al., 2017. The DNA sequences analyzed in this study and retrieved from GenBank to infer phylogenetic relationships among the Meripilaceae, Polyporales group are listed in Table 2. The lists of DNA sequences for the Cerrenaceae, Polyporales group and Rigidoporus, Hymenochaetales are shown in Supplementary Table S1 and S2. Sequence alignment data are added as Supplementary alignment S1 for the Meripilaceae, Polyporales group, S2 for the Cerrenaceae, Polyporales group, and S3 for Rigidoporus, Hymenochaetales.

Table 2. DNA sequence data newly obtained in this study (bold-face type) and employed from GenBank for the phylogenetic analysis of Meripilaceae in Polyporales.

Species name Sample No. Locality GenBank accession No.
nrDNA ITS nrDNA LSU
Abortiporus biennis FD-319 USA KP135300 KP135195
Hyphoderma setigerum FD-312 USA KP135297 KP135222
Hypochnicium sp. FP-110227-sp USA KY948804 KY948862
Meripilus giganteus CBS 421.48 Germany MH856418 a LC770099
M. giganteus FP-135344-Sp UK KP135307 KP135228
M. giganteus TUFC 100564 Japan LC643683 LC643708
Physisporinus castanopsidis Dai 20396 T b China MT309485 MT309470
P. castanopsidis Dai 20397 China MT309486 MT309472
P. castanopsidis Dai 20398 China MT840113 MT840131
P. cf. 1 eminens TUFC 101880 Japan LC643670 LC643695
P. cf. 1 eminens TUFC 101957 Japan LC770057 LC770082
P. cf. 1 eminens TUFC 101958 Japan LC770058 LC770083
P. cf. 1 eminens TUFC 101959 Japan LC770059 LC770084
P. cf. 2 eminens TUFC 101881 Japan LC643671 LC643696
P. cf. 2 eminens TUFC 101960 Japan LC770060 LC770085
P. cf. 2 eminens TUFC 101961 Japan LC770061 LC770086
P. cf. 2 eminens TUFC 101962 Japan LC770062 LC770087
P. cf. furcatus TUFC 101883 Japan LC643673 LC643698
P. cf. furcatus TUFC 101884 Japan LC643674 LC643699
P. cinereus Cui 3266 China KY131844 KY131903
P. crataegi Dai 15497 T China KY131845 KY131904
P. crataegi Dai 15499 China KY131846 KY131905
P. crocatus CBS 107806 Canada LC770074 LC770100
P. crocatus Dirks 374051 USA ON364084 ON369536
P. crocatus MJ 19/09 Slovakia JQ409466 OM669978
P. eminens Cui 9520 China KY131847 KY131906
P. eminens Cui 10341 China KY131849 KY131907
P. eminens Cui 10344 China KY131850 KY131908
P. eminens Cui 10475 China MT840114 MT840132
P. eminens Dai 11400 China KY131852 KY131909
P. eminens Dai 12685 Czechia MT840115 MT840133
P. eminens Dai 17200 Unknown MT279690 MT279911
P. eminens Dai 19861 China MT840116 MT840134
P. eminens Dai 20832 China MT279689 MT279910
P. eminens Dai 20868 China MT840117 MT840135
P. eminens Dai 22472 China OM669900 OM669983
P. furcatus Dai 2105 China KY131854 KY131911
P. furcatus Dai 2544 China KY131855 KY131912
P. furcatus Dai 11313 China KY131856 KY131913
P. furcatus Dai 12938 China KY131857 KY131914
P. furcatus Dai 20976 Belarus MT840118 MT840136
P. furcatus Dai 20977 Belarus MT840119 MT840137
P. furcatus TAA 15097 T Russia KY131853 KY131910
P. lavendulus Dai 9925 China KY131858 KY131915
P. lavendulus Dai 13587A T China KY131859 KY131916
P. lineatus CBS 167.65 USA LC770075 LC770101
P. lineatus CBS 700.94 Germany LC770076 LC770102
P. lineatus CBS 109425 Taiwan LC770077 LC770103
P. lineatus Dai 17986 China MT840121 MT840139
P. lineatus Dai 18280 Vietnam MT840122 MT840140
P. lineatus JV 1407/37 Costa Rica OM669903 OM669986
P. lineatus TUFC 13809 Japan LC770063 LC770088
P. lineatus TUFC 13812 Japan LC770064 LC770089
P. longicystidius Cui 16630 Australia ON417177 ON417227
P. longicystidius Cui 16725 Australia ON417178 ON417228
P. microacanthophysis TUFC 101885 Japan LC643675 LC643700
P. microacanthophysis TUFC 101888 Japan LC643678 LC643703
P. microacanthophysis TUFC 101889 T Japan LC643679 LC643704
P. microacanthophysis TUFC 101963 Japan LC770065 LC770090
P. microacanthophysis TUFC 101964 Japan LC770066 LC770091
P. pouzarii Dai 15005 China KP420014 KP420017
P. pouzarii Dai 21043 Belarus MT840124 MT840142
P. pouzarii JV 0511/23 Czechia JQ409465 KY131921
P. pouzarii TUFC 101965 Japan LC770067 LC770092
P. pouzarii TUFC 101966 Japan LC770068 LC770093
P. rhizomorphae TUFC 101870 Japan LC643660 LC643685
P. rhizomorphae TUFC 101871 Japan LC643661 LC643686
P. rhizomorphae TUFC 101876 T Japan LC643666 LC643691
P. rhizomorphae TUFC 101967 Japan LC770069 LC770094
P. roseus Dai 19877 T China MT840126 MT840144
“P. sanguinolentus” CBS 139.76 Belgium MH860969 c MH872738 c
“P. sanguinolentus” CBS 193.76 Netherlands LC770078 LC770104
“P. sanguinolentus” CBS 679.70 USA MH859899 c MH871689 c
“P. sanguinolentus” CBS 107146 Denmark LC770079 LC770105
“P. sanguinolentus” Dai 20995 Belarus MT309483 MT309480
“P. sanguinolentus” DM1068 Denmark MT644902 MT644902
“P. sanguinolentus” JV 1610/2 Czechia OM669921 OM669999
“P. sanguinolentus” KHL 11913 Sweden JX109843 JX109843
Physisporinus sp. 1 Dai 11693 China KY131865 KY131922
Physisporinus sp. 2 Dai 6720 China KY131867 KY131923
Physisporinus sp. 4 Dai 15184 Unknown KY131868 KY131924
Physisporinus sp. Cui 16852 Puerto Rico ON417179 ON417229
Physisporinus sp. CWU 3874 Ukraine OM971903 OM971889
Physisporinus sp. JV 0308/58 USA OM669909 OM669991
Physisporinus sp. JV 0509/47 USA OM669906 OM669988
Physisporinus sp. JV 0709/188 USA OM971904 OM971890
Physisporinus sp. JV 0909/3 Czechia OM669939 OM670011
Physisporinus sp. JV 1407/36 Costa Rica OM669933 OM670008
Physisporinus sp. Miettinen 15239 Indonesia KY948732 KY948867
Physisporinus sp. Miettinen 16699 USA KY948733 KY948863
Physisporinus sp. TUFC 101892 Japan LC643682 LC643707
Physisporinus sp. TUFC 101968 Japan LC770070 LC770095
P. subcrocatus Dai 12800 USA KY131869 KY131925
P. subcrocatus Dai 15917 T China KY131870 KY131926
P. sulphureus Dai 17839 T Singapore MG132179 MG132181
P. sulphureus Dai 17841 Singapore MG132180 MG132182
P. tibeticus Cui 9381 T China KY131871 KY131927
P. tibeticus Cui 9588 China KY131873 KY131929
P. tibeticus Cui 10478 China MT840128 MT840146
P. undatus JV 0110/48 Czechia OM669931 OM670005
P. undatus Miettinen 13591 Finland KY948731 KY948870
P. undatus MJ 129/04 Czechia OM669932 OM670006
“P. vinctus” Cui 16903 China MT840129 MT840147
P. vitreus Dai 21060 Belarus MT840130 MT840148
P. vitreus KHL 11959 Norway JQ031129 JQ031129
P. yunnanensis CLZhao 21583 China OP852341 OP852343
P. yunnanensis CLZhao 21647 T China OP852340 OP852342
Polyporales sp. 422b Japan AB470242 AB470242
Polyporales sp. TK-10 Japan AB716748 AB762089
Uncultured mycorrhizal fungus Polyporales218 Taiwan KP238183 KP238182
Uncultured mycorrhizal fungus Polyporales859 Taiwan KP238186 KP238184
Uncultured mycorrhizal fungus Physisporinus222 Taiwan KP238185 KP238181
Spongipellis ambiens Niemelä 6407 China ON979313 ON979313
S. ambiens Spirin 5389 Russia ON979322 ON979322
S. profissilis Dai 3934 China ON979321 ON979321
S. profissilis Kotiranta 26990 Russia OP104014 OP104014
S. spumea JV 1511/6 Czechia ON979318 ON979318
S. spumea Kotiranta 26889 Finland ON979311 ON979311
S. spumea Spirin 6741 Russia ON979326 ON979326
S. variispora Niemelä 6423 China ON979320 ON979320
S. variispora Spirin 3737 T Russia ON979312 ON979312

a The nrDNA ITS sequence of M. giganteus CBS 421.48 had been already registered with this accession number in GenBank before our study. We used it for the analysis of the Meripilaceae group because the sequence of this strain obtained in the present study corresponded to the above sequence with high homology (99%) by the Standard Nucleotide BLAST (Basic Local Alignment Search Tool) of the GenBank database.

b T means the type specimen or ex-type culture.

c The nrDNA ITS and LSU sequences of “P. sanguinolentus” CBS 139.76 and CBS 679.70 had been already registered with these accession numbers in GenBank before our study. We used them for the analysis of the Meripilaceae group because the sequences of the two strains obtained in the present study corresponded to the above sequences with high homology (Both were 100% in the ITS and 99% in the LSU region) by the BLAST.

2.3. Observation of cultures and specimens

Strains including new isolates were precultured on an antibiotics-added corn meal agar medium (Shino et al., 2022) at room temperature (20-25 °C) for 1-2 mo. After agar discs containing mycelia were cut out or stamped out from the precultured plates using a flame sterilized scalpel or autoclaved sterilized plastic straws (6 mm diam), they were inoculated on the following four media; a corn meal agar medium [CMA; Corn Meal Agar “Nissui” (Nissui Pharmaceutical Co., Ltd., Tokyo, Japan; containing 2 g/L cornmeal extract and 15 g/L agar)], a malt extract agar medium [MA; 15 g/L malt extract (Oriental Yeast Co., Ltd., Tokyo, Japan) and 15 g/L agar (FUJIFILM Wako Pure Chemical Corporation, Osaka, Japan)], a potato dextrose agar medium [PDA; Potato Dextrose Agar “Nissui” (Nissui Pharmaceutical Co., Ltd.; containing 3.9 g/L potato extract, 21 g/L glucose and 14.1 g/L agar)], and a starch agar medium [SA; 10 g/L starch, soluble (FUJIFILM Wako Pure Chemical Corporation), 5 g/L malt extract, and 20 g/L agar]. These plates were incubated at 25 °C for 7-31 d and the following cultural properties were examined using a Nikon ECLIPSE 80i differential interference contrast microscope (DICM) (Nikon Corporation, Tokyo, Japan); presence/absence and morphology of acanthophyses, clamp connections, conidia, and plectenchymata. Samples were mounted in 3% potassium hydroxide (KOH) on a slide glass, and the size was measured by PhotoRuler ver. 1.1.3 software (http://inocybe.info). Specimens of SSs and basidiocarps from which strains were isolated were observed by the above method to confirm the consistency to the results of phylogenetic analyses. Acanthophyses were also observed using a scanning electron microscope (SEM) (SU1510; Hitachi High-Tech Corporation, Tokyo, Japan). Preparation and observation of SEM samples followed Shino et al. (2022).

2.4. Investigation of acanthophysis production on agar media during incubation under different O2 concentrations

A schematic illustration of the following experiment is shown in Fig. 2. Selected strains (Physisporinus cf. 1 eminens TUFC 101880, P. cf. 2 eminens TUFC 101881, P. cf. furcatus TUFC 101883, P. lineatus TUFC 13809, P. microacanthophysis TUFC 101885, and P. pouzarii TUFC 101965; see Table 1) were preincubated at room temperature (20-25 °C). After mycelia covered the entire surface of the medium, they were stamped out as discs with agars using autoclaved sterilized plastic straws (6 mm diam) and the agar discs were used to inoculate CMA plates: six discs were inoculated per a plate (three discs were placed facing up and the other three discs were placed facing down). After checking the number of acanthophyses on the surface of the discs under the DICM, as they were already formed during preincubation before experiment, the CMA plates with their inoculated agar discs were placed in four desiccators [Vacuum Polycarbonate Desiccator 240G (or 240GA) or 300G (or 300GA) (AS ONE Corporation, Osaka, Japan)]. Then, the air in each desiccator was exhausted using a diaphragm type dry vacuum pump (DA-20D; ULVAC KIKO, Inc., Miyazaki, Japan) and each desiccator was filled with one of the following standard gas mixtures or air to prepare four different O2 conditions: 5% O2 (O2 4.95%, CO2 402 ppm, and N2 as a base gas), 10% O2 (O2 10.0%, CO2 405 ppm, and N2), 20% O2 (the atmospheric air: O2 21%, CO2 400 ppm, and N2 78%; all the values of concentrations are approximate), and 40% O2 (O2 39.5%, CO2 395 ppm, and N2). To prepare the 5, 10 and 40% O2 conditions, we used calibration gas cylinders that were prepared by Taiyo Nippon Sanso JFP Corporation (Kanagawa, Japan) and a gas regulator (GHN-3; CHIYODA SEIKI Co., Ltd., Hyogo, Japan). The gas in each desiccator was exchanged daily: the process of exhausting and filling of each gas was repeated twice per exchange. Uncovered plates in desiccators were incubated for 2-5 d at room temperature. After finishing the incubation, the number of acanthophyses that formed on the upper surface and side of each disc was counted under the DICM. The number of acanthophyses produced during preincubation was excluded from the data.

Fig. 2 - The outline of experiment for acanthophysis production under different O2 concentrations (For details, see section 2.4. in materials and methods).

3. Results

3.1. Relationships between molecular phylogeny and cultural characteristics with emphasis on acanthophysis formation

The phylogenetic analysis of Meripilaceae in Polyporales that was based on the combined sequences of the ITS and LSU regions of nrDNA showed that Meripilus, Physisporinus, and Spongipellis form a clade, and Meripilus was included within the Physisporinus clade (BS = 99%; Fig. 3). The topology of this phylogenetic tree was almost correspondent with trees estimated in several recent studies (Chen & Dai, 2021; Shino et al., 2022; Spirin et al., 2022; Wang & Dai, 2022). Acanthophysis-forming strains on agar media in the present study belonged only to the Physisporinus clade (Figs. 3, 4, 5). In this clade, a monophyletic cluster including P. castanopsidis Jia J. Chen & Y.C. Dai, P. crocatus (Pat.) F. Wu, Jia J. Chen & Y.C. Dai, P. microacanthophysis, P. pouzarii, “P. sanguinolentus” (Alb. & Schwein.) Pilát, P. subcrocatus F. Wu, Jia J. Chen & Y.C. Dai, P. tibeticus F. Wu, Jia J. Chen & Y.C. Dai, and P. vitreus (Pers.) P. Karst. (BS = 100%) formed shorter acanthophyses (10-30 µm long: the ornamented part with warts or spines, but not including spines) than the rest of species in this genus which are known to produce SSs/acanthophyses (20-70 µm long), except for P. cf. 1 eminens (12-29 µm long). Moreover, the strains in this cluster had sparse clamp connections at the septa of vegetative hyphae (Figs. 3, 4, 6E), whereas strains in other clades of Physisporinus lacked clamp connections on the hyphae. Meripilus strains did not produce acanthophyses on the agar media employed in this study. Both Meripilus and Physisporinus formed plectenchymata in cultures (Fig. 6B, F). Formation of the plectenchymata in both genera have been described previously, e.g., Larsen and Lombard (1988) and Lombard and Chamuris (1990).

Fig. 3 - Phylogenetic tree of Meripilaceae in Polyporales inferred from connected sequences of the ITS and LSU regions of nrDNA by ML method. A total of 1,171 sites in the final data set were used for this analysis. The values at nodes indicate BS in ML method (≥ 70%), and bold branches mean BS ≥ 90% in the above method. The species names and numbers of strains used in this study are shown in bold, and the strain number followed by “S” or “R” indicates the isolate from a SS or rhizomorph. T on the sample number means the sequence obtained from the type specimen or ex-type culture. Filled circles show the strains forming acanthophyses in culture. Open squares show the strains having clamp connections at the septa of vegetative hyphae on agar media (white arrow indicates a monophyletic clade characterized by this feature). The strains without sufficient cultural investigations in the present study are unmarked.
Fig. 4 - The relationship between molecular phylogeny and acanthophysis formation in Physisporinus. A figure at the upper left is the reduced Fig. 3, and a box in it shows a magnified part for this figure. As with Fig. 3, filled circles show the strains forming acanthophyses in culture, and open squares show the strains having clamp connections at the septa of vegetative hyphae on agar media (white arrow indicates a monophyletic clade characterized by this feature). The strains without sufficient cultural investigations in the present study are unmarked. The sizes of acanthophyses of Physisporinus species investigated in this study are described under the species name. The appearance of a typical acanthophysis of each species is exhibited by photographs using SEM at the right of this figure. Bars: 10 µm.
Fig. 5 - The relationship between molecular phylogeny and acanthophysis formation in Physisporinus. A figure at the upper left is the reduced Fig. 3, and a box in it shows a magnified part for this figure. As with Fig. 3, filled circles show the strains forming acanthophyses in culture. The strains without sufficient cultural investigations in the present study are unmarked. The sizes of acanthophyses of Physisporinus species investigated in this study are described under the species name. The appearance of a typical acanthophysis of each species is exhibited by photographs using SEM at the right of this figure. Bars: 10 µm.
Fig. 6 - Cultural characteristics of the strains of Meripilus giganteus (A-C) and Physisporinus species (D-F) in Meripilaceae. A: Colony on 1.5% MA (TUFC 100564). B: Plectenchymata in mycelia (CBS 421.48). C: Vegetative hyphae (CBS 421.48). Arrow heads show clampless septa. D: Colony on 1.5% MA (P. pouzarii TUFC 101965). E: Clamp connection on a septum of vegetative hyphae (P. pouzarii TUFC 101965). F: Plectenchymata and vegetative hyphae (P. lineatus CBS 167.65). Arrow heads show clampless septa. Bars: B, C, E, F 10 µm.

The phylogenetic analysis based on the nrDNA ITS or LSU region showed that “Rigidoporus” species in the traditional usage separated in two different lineages, Polyporales and Hymenochaetales (Supplementary Figs. S1, S2) as reported by Wu et al. (2017). The phylogenetic tree of the Cerrenaceae group in Polyporales (Supplementary Fig. S1) showed that this family clusters with Cerrena, Irpiciporus, Pseudolagarobasidium, Pseudospongipellis, Radulodon, and “Rigidoporus” (BS = 100%), as reported in previous studies (Justo et al., 2017; Wang & Dai, 2022; Westphalen & Motato-Vásquez, 2022), and a highly supported clade (BS = 99%) of “R. hypobrunneus”/“R. vinctus” accommodates five strains of “R. vinctus” examined in this study. Arthroconidia production in “R. vinctus” strains was observed (Supplementary Fig. S1) as reported previously [Setliff, 1972 (as oidia); Stalpers, 1978], but they did not produce any acanthophyses. The topology of the tree estimated for Rigidoporus in Hymenochaetales was similar to that in Wu et al. (2017) and Yuan et al. (2020). Rigidoporus ulmarius (Sowerby) Imazeki CBS 186.60 formed a clade together with nine sequences, including four sequences of R. microporus (Sw.) Overeem (BS = 98%; Supplementary Fig. S2) in Hymenochaetales. This strain did not form acanthophyses, but it did produce vesicular cells on vegetative hyphae laterally and terminally in culture (Supplementary Fig. S2), as described previously in Lombard et al. (1960) and Stalpers (1978; as terminal vesicles).

3.2. Acanthophysis production on agar media under different O2 concentrations

The results of the experiments using six strains (Physisporinus cf. 1 eminens TUFC 101880, P. cf. 2 eminens TUFC 101881, P. cf. furcatus TUFC 101883, P. lineatus TUFC 13809, P. microacanthophysis TUFC 101885, and P. pouzarii TUFC 101965) cultured under four different O2 conditions (5, 10, 20, and 40% O2) are shown in Fig. 7. In the case of the agar discs placed facing up, the three strains (TUFC 101880, TUFC 101881, and TUFC 101885) formed acanthophyses most abundantly under the atmospheric condition, i.e., 20% O2, whereas the two strains (TUFC 101883 and TUFC 101965) formed most acanthophyses under 40% O2. The four strains other than TUFC 13809 and TUFC 101880 tended to form more acanthophyses on the upper surface than on the sides of the agar discs in the situation placed facing up, probably because aerial vegetative hyphae on the discs are easy to contact to the air.

Fig. 7 - Differences in the number of acanthophyses produced on agar discs under four different O2 concentrations (5, 10, 20, and 40%) by six Physisporinus strains. The vertical axis of the bar graph shows the number of acanthophyses. “T” and “B” under the horizontal axis mean the agar discs inoculated on a CMA plate as top face up and back face up, respectively. “T” is shown by a blue bar and “B” by a white bar. “U” and “S” indicate the upper surface and side face of the agar discs.

In the case of the agar discs placed facing down, the four strains (TUFC 101880, TUFC 101881, TUFC 101885, and TUFC 101965) produced more acanthophyses under the 20% O2 condition than under the 40% O2 condition. TUFC 101883 produced acanthophyses only under the 40% O2 condition. When the agar discs were set on the plates with facing down, the aerial vegetative hyphae at the upper surface were facing the CMA plates, which resulted in most of the hyphae existing inside the discs. Hence, this latter situation in which the hyphae spreading inside the agar disc is somewhat analogous to that of hyphae growing within water-saturated wood tissue in the natural wet habitats of SS-forming Physisporinus species.

The number of acanthophyses formed by the five strains (TUFC 101880, TUFC 101881, TUFC 101883, TUFC 101885, and TUFC 101965) on the discs placed both facing up and down under the 5% and 10% O2 conditions was lower than 20% or 40% O2 conditions, but TUFC 13809 produced acanthophyses most abundantly under 10% O2 in the case of discs being placed facing up and under 5% O2 in the case of discs being placed facing down. The above five strains tended to mainly produce acanthophyses from the upper surface and/or side face of the agar discs placed facing up or down on CMA plates, while TUFC 13809 formed acanthophyses abundantly on the aerial vegetative hyphae that spread on the plates as well as on the entire surface of the agar discs.

4. Discussion

Some basidiomycetes have been known to produce acanthophyses on vegetative hyphae in culture, and most of these taxa are now placed in Physisporinus; for example, P. crocatus, which was formerly treated as Poria nigrescens Bres. (Nobles, 1958); P. lineatus, formerly treated as Polyporus zonalis Berk. [Bakshi et al., 1963; Davidson et al., 1942 (acanthophyses were described as hyphal ends covered with short knobs or definite spines); Nobles, 1958], as Rigidoporus zonalis (Berk.) Imazeki (Kobayashi, 1972), and as R. lineatus [Hood et al., 1997 (as acanthohyphidia); Motato-Vásquez et al., 2016 (as the spiny and clavate cystidia); Stalpers, 1978 (as acanthohyphidia)]; P. undatus (Pers.) Pilát, formerly treated as R. undatus (Pers.) Donk (Motato-Vásquez et al., 2016); P. vitreus, formerly treated as R. vitreus (Pers.) Donk (Lombard & Chamuris, 1990; Schmidt et al., 1996, 1997). In these previous studies, species identification was based mainly on the morphological characteristics of basidiocarps. Our phylogenetic studies showed that the above acanthophysis-forming species are accommodated in Physisporinus. Except for Physisporinus, some species of Xylobolus P. Karst. have also been reported to form acanthophyses on agar media; for example, X. frustulatus (Pers.) Boidin [Lombard & Chamuris, 1990; Nakasone, 1990 (termed as acanthohyphidia); Stalpers, 1978 (as acanthohyphidia)]; X. subpileatus (Berk. & M.A. Curtis) Boidin (Stalpers, 1978), although the two species scarcely produce acanthophyses (Stalpers, 1978). In addition to Xylobolus, Aleurodiscus Rabenh. ex J. Schröt. sensu lato (Wu et al., 2001), Megalocystidium Jülich [only M. diffissum (Sacc.) K.H. Larss. & Spirin (Spirin et al., 2021)], and Stereum Hill ex Pers. are also known to produce acanthophyses (or termed as acanthocystidia or acanthohyphidia) in their basidiocarps (e.g., Bernicchia & Gorjón, 2010; Larsson & Ryvarden, 2021). These genera belong to Stereaceae, Russulales (Miller et al., 2006; Wu et al., 2022), and acanthophysis formation on vegetative hyphae in culture has been known only in the above Xylobolus species. On the other hand, Physisporinus seldom or never produce acanthophyses in their basidiocarps. Among Physisporinus, the closely related Meripilus and morphologically similar Rigidoporus, only Physisporinus species produce SSs in nature and/or acanthophyses in culture. Previous studies on the culture of Meripilus and Rigidoporus species, except for the species currently transferred to Physisporinus, did not observe SS/acanthophysis formation (Campbell, 1937; Davidson et al., 1942; Go et al., 2021; Kaewchai et al., 2010; Larsen & Lombard, 1988; Lombard et al., 1960; Nobles, 1948, 1965; Setliff, 1972; Stalpers, 1978). These results support our previous suggestion that SS/acanthophysis formation could be a taxonomic character for defining the genus Physisporinus, which is currently difficult to distinguish from Rigidoporus based on the morphology of the basidiocarp (Shino et al., 2022). All the Physisporinus samples used in our previous study were collected from aquatic environments (Shino et al., 2022). However, in addition to samples from streams, this study includes new samples from terrestrial environments (Table 1). Therefore, the Physisporinus species that inhabit forest areas may also form acanthophyses on their vegetative hyphae. Physisporinus species that have not been proven to produce SSs or acanthophyses, especially those that form perennial and/or brightly colored basidiocarps, should be investigated for their SS/acanthophysis-forming ability on wet wood or on media. Moreover, we found that the species group producing small acanthophyses (10-30 µm long) and rare clamp connections on septa of vegetative hyphae formed a highly supported clade that harbored at least eight Physisporinus species (P. castanopsidis, P. crocatus, P. microacanthophysis, P. pouzarii, “P. sanguinolentus”, P. subcrocatus, P. tibeticus, and P. vitreus), whereas other acanthophysis-forming clades in this genus produce larger acanthophyses (20-70 µm long, except for P. cf. 1 eminens which forms acanthophyses of 12-29 µm long) and no clamp connections (Figs. 3, 4, 5). These characteristics suggest that the size of acanthophyses is related to the phylogeny of Physisporinus. In this study, we were unable to find any clearly distinctive characteristics in basidiocarps of the species group having short acanthophyses and clamp connections in culture in comparison with other Physisporinus species contained in different clades. Further studies focusing on both the basidiocarps and isolates are therefore needed.

We currently face a raft of challenges related to the taxonomy of Physisporinus and the allied genera, Meripilus and Rigidoporus. Physisporinus still contains taxonomically confused species probably composed of plural species [e.g., P. sanguinolentus (Runnel et al., 2021, refer to Additional file 5; Wu et al., 2017); P. furcatus group and P. undatus group (Chen & Dai, 2021); P. vitreus (Runnel et al., 2021)]. The question remains about the validity of P. subcrocatus from the perspective of the very close similarity to P. crocatus in terms of morphology of basidiocarps and phylogeny. The phylogenetic position of Meripilus (i.e., whether this genus is truly nested in the Physisporinus clade or not) has not been confirmed by multi-gene phylogenetic analyses with sufficient sequences yet, although this study showed that, unlike Physisporinus, Meripilus species do not produce acanthophyses. In the Cerrenaceae, Polyporales group, a taxonomic problem regarding the “R. hypobrunneus”/“R. vinctus” clade, which was also pointed out by Nakasone and Ortiz-Santana (2022), was more clearly highlighted by our phylogenetic analysis (Supplementary Fig. S1). This clade should be treated as a new or another genus, but we do not treat it as such in this study because we could not investigate the type specimens of these species. In addition, it is important to reexamine specimens that are currently treated as P. vinctus (Berk.) Murrill, a synonym of R. vinctus, in the phylogenetic trees by Chen and Dai (2021), Shino et al. (2022), Wu et al. (2017), and this study (Fig. 3). To solve these taxonomic issues, investigations of currently overlooked or underestimated characters, such as the cultural properties of asexual states and vegetative hyphae, as well as the ecological characteristics of these species should be conducted in addition to the currently dominant studies focusing on the morphology of sexual states (basidiocarps) and molecular phylogeny.

When we find SSs of Physisporinus species in freshwater areas, they are often formed on the water-boundary part of dead and wet wood of broad-leaved or coniferous trees. The wood substrate is carried by water flow as drift and caught between rocks, then exposed to the flow and splash for extended periods, mostly resulting in barkless and sometimes partly getting mossy. The insides of wet wood at the water-boundary and submerged parts are saturated with water and the dissolved O2 concentrations within the wood tissue most likely decrease due to the low level of gas exchange. The results of the present experiments that exposed cultures of Physisporinus to gas mixtures with different O2 concentrations suggested that acanthophyses were produced in response to higher O2 concentrations and that they probably play a role in obtaining O2. Our experiments clearly showed that the number of acanthophyses produced on agar discs was markedly increased when incubated under O2 concentrations of 20-40% compared to when incubated under O2 concentrations of 5-10% O2 (Fig. 7). The numerous spines of the acanthophyses probably function to increase the surface area of acanthophysis cells for gas exchange. This thought is supported by the fact that acanthophyses are formed on aerial vegetative hyphae, not on submerged hyphae in agar medium. Thus, it is possible that SSs furnished with numerous acanthophyses are formed at the water-boundary part that is exposed to the air and that they play a role in respiration at the closest site to the submerged part. The synnematous morphology of the SSs possibly serves to maintain the distance from the water surface and the water-saturated part of the wood substrate, so that the acanthophyses are exposed to the atmosphere, and also to withstand the force of the water flow. When the agar discs containing mycelia were placed on media facing down, four strains (P. cf. 1 eminens TUFC 101880, P. cf. 2 eminens TUFC 101881, P. microacanthophysis TUFC 101885 and P. pouzarii TUFC 101965) formed acanthophyses more abundantly under 20% O2 condition than under 40% O2 condition (Fig. 7). This finding might be explained as follows; under the 40% O2 condition, high levels of O2 permeate the agar media, so only a fewer number of acanthophyses need to be produced in order to obtain sufficient O2 for extending hyphae into the media. However, P. cf. furcatus TUFC 101883 responded differently, as producing more acanthophyses under the 40% O2 condition than under the 20% O2 condition (Fig. 7). Therefore, the sensitivity to O2 may differ among species and/or strains. In the present experiments, most strains formed less acanthophyses under the low O2 conditions, 5% or 10%. This is assumed that there was insufficient difference in oxygen concentration between inside and outside the culture medium to induce acanthophysis formation. Further detailed physiological study is required to verify this hypothesis. Interestingly, P. lineatus TUFC 13809 exceptionally formed numerous acanthophyses, even under 5% O2 condition (Fig. 7). This species is known to show a high level of mycelial growth rate even in low O2 concentrations (Hood et al., 1997) and to cause the decay in heartwood, especially root and butt rot of living trees in Asia and North and South America [Dai et al., 2007 (as Rigidoporus lineatus); Kobayashi, 1972 (as R. zonalis); Overholts, 1953 (as Polyporus zonalis); Rajchenberg & Robledo, 2013 (as R. lineatus)]. The internal part of trees is not normally exposed to the atmosphere, so such heart rot fungi must be adapted to the low oxygen condition. Though SS formation by P. lineatus has not been reported yet, the ability to produce large numbers of acanthophyses even under low O2 concentrations is likely to contribute to the high rate of hyphal growth and the heart rot in trees. Additional investigations on the ecology of this species are needed. Hyde and Goh (1998) reported an unidentified fungus that formed tufts of acanthophyses on the apices of root-like hyphal strands on wet wood collected in several tropical streams. Based on the habits and observations of the characteristics of the fungus, they guessed that the function of acanthophyses was to take up O2 in water. However, their discussion was speculative at the time. Because the fungus was not identified or observed in its sexual state, the taxonomic assignment of their fungus should be clarified by further study.

Basidiomycetous fungi have been considered to prefer terrestrial environments to aquatic environments; this is suggested by the fewer number of aquatic species compared to terrestrial species (Jones et al., 2014; Shearer et al., 2007). However, our findings showed that Physisporinus species have adapted to humid environments such as streams and waterfalls by acquiring the ability to form SSs/acanthophyses, which appear to function as respiratory organs. This may be a strategy for terrestrial fungi in origin to adapt to aquatic habitats and decay water-saturated wood with low O2 concentrations. Further research of the basidiomycetous fungi inhabiting wet habitats should be undertaken to better clarify their biodiversity and ecology.

Disclosure

The authors declare no conflicts of interest. All the experiments undertaken in this study comply with the current laws of the country where they were performed.

Acknowledgments

This study was supported, in part, by a Grant-in-Aid from the Institute for Fermentation, Osaka (IFO). We appreciate staff of Westerdijk Fungal Biodiversity Institute for providing living cultures of Meripilus, Physisporinus and Rigidoporus species. We also great thank Ms. Sachiko Ueta and Kaori Shimizu, FMRC, for taking on the cryopreservation and maintenance of strains investigated in the present study. Deposition and utilization of Tottori University Fungal Culture Collection (TUFC) strains were supported by FMRC through the National BioResource Project (NBRP) of the Ministry of Education, Culture, Sports, Science and Technology (MEXT), Japan (http://nbrp.jp).

References
 
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