Polyphasic identification of Sepedonium microspermum isolated from two genera of Boletales in Iran

Document Type: Original Article


Department of Plant Protection, Faculty of Agriculture, University of Tabriz, Tabriz, Iran


Moldy Boletus sp. and Xerocomus sp. were collected from several locations at the campus of the University of Tabriz, Iran. Fungicolous fungal isolates were recovered and characterized by the combination of morphological traits and phylogenetic analyses of combined ITS and LSU sequence data. Fungal isolates were identified as Sepedonium microspermum. This is the first report of S. microspermum on Xerocomus sp. from Iran which is comprehensively described and illustrated.


Main Subjects


The fungi that grow on macromycetes, rusts, powdery mildew, slime molds and etc., are called fungicolous fungi, even when the nature of the fungus-fungus association and its trophic relationships are obscure (Jeffries 1995). In nature, most of the fungicolous fungi are known as parasites (necrotrophs or biotrophs), commensals or saprobes (Hawksworth et al. 1995). Most of the mycologists such as Gilman and Tiffany (1952) and Barnett (1963, 1964), used the term for the fungi related to other fungi (Sun et al. 2019).

The taxonomy of fungicolous fungi has considerably progressed from the nineteenth century, especially with the findings of anamorph–teleomorph relations in the Hypocreales, as well as the reports of new mycoparasitic heterobasidiomycetous fungi (Sun et al. 2019). Regional surveys have mostly been limited to the taxa of sporocarp inhibiting fungi (e.g., Helfer 1991). Most of the fungicolous fungi that are related to mushrooms and plant soil-borne pathogens are scattered in temperate and sub-tropical regions (Sun et al. 2019). Rudakov (1978) study on fungicolous fungi resulted in the identification of about 1,700 (nonlichenicolous) species of this fungus. Moreover, he indexed fungicolous fungi occurring in the former Soviet Union, but many of his identifications need to be revised (Rudakov 1981). In a revision of the conidial fungicolous fungi, Hawksworth (1979, 1981) reported the number of 1,100 species grown on approximately 2,500 species of host fungi (including lichenized taxa).

The Hypocreaceae (Hypocreales) are the most important fungicolous fungi grown on fruit body of other fungi while Bionecteriaceae and Nectriaceae belonging to the Hypocreales, include mycoparasitic or mycosaprobic species (Rossman et al. 1999). Six genera of the Hypocreaceae include fungicolous fungi such as Trichoderma (=Hypocrea) and Hypomyces whose species are identified by morphological characteristics of teleomorph and anamorph (Põldmaa 2000).

Different genera of fungicolous fungi such as Cladobotryum, Mycogone, Stephanoma and Sepedonium are asexual forms of Hypomyces (Gams & Hoozemans 1970, de Hoog 1978, Rogerson & Samuels 1993, 1994, Rossman et al. 1999). Cladobotryum is one of the most important anamorphic fungicolous fungi of Hypomyces (Rogerson & Samuels 1993, Põldmaa 2000). Rogerson & Samuels (1985, 1989, 1993 and 1994) and Põldmaa (2000), classified the species of Hypomyces in the four fungal host groups including species that grow on Discomycetes, Boletus species (boleticolous species), Agaricales (agaricicolous species) and Aphyllophorales (aphyllophoricolous species). Other fungal groups that were previously reported before such as Discomycetes (Leotiales, Pezizales), Agarics (Russulales), Boletes and Aphylophorales are also the hosts of Hypomyces species (Zare & Asef 2008). One of the asexual forms of Hypomyces is the genus Sepedonium Link 1809based on S. mycophilum (Pers.) Link 1809 as the type. Until now, the number of 58 species has been reported and listed for Sepedonium in index fungorum (http://www.indexfungorum.org; accessed on 14th, January 2019). Sepedonium is characterized by the production of aleurioconida and phialoconidia and most of the species from this genus are parasites on Boletales. In addition to boletes, Sepedonium spp. have been reported on Scleroderma, Rhizopogon, agarics, air, soil, dung and etc. (Rogerson 1989).

In Iran, there are limited studies on the species diversity of Hypomyces and related asexual forms in which species identification has relied solely on the morphological characteristics. Asef & Mohammadi Goltapeh (2002) listed four species of Cladobotryum including C. dendroides, C. verticillatum, C. polypori, and C. varium, but there was no sexual form in the investigated samples. Recently, Asef & Zare (2006) recorded three species of Hypomyces as a sexual form of fungicolous fungi and a species of Cladobotryum from Iran, as a sexual form of fungicolous fungi and a species of Cladobotryum from Iran. Anamorphic forms of H. aurantius and H. rosellus which are called C. varium and C. dendroides respectively were reported as fungicolous fungi from Iran (Asef & Mohammadi Goltapeh 2002).  The occurrence of S. microspermum and Sepedonium sp. on Boletus sp. and Leccinum sp. were the only reported cases from Sepedonium species in Iran (Zare & Asef 2008). The hosts were identified according to the book written by Keizer (2004). In this study, we provide the first occurrence of S. microspermum on Xerocomus sp. in Iran. The identification of species was confirmed by a combination of morphological characteristics and sequence data of ITS-rDNA region and LSU gene.


Sample collection and fungal isolation

During a field excursion at the campus of the University of Tabriz in East Azerbaijan, Iran in 2016, samples were collected from fresh, mature and moldy specimens of some infected Boletales (five specimens for each species) and were stored separately in the paper bags to keep them clean for culture work. Host specimens were identified at genus level following Keizer (2004) protocol. Morphological observations and fungal isolations were done according to the protocols of Gams et al. (2004). Fungicolous fungal isolates from specimens were all recovered from the caps and hymenium of the collected hosts. Single-spore isolations were conducted on 2 % malt extract agar, according to the protocol of Torbati et al. (2018); in brief, using a sterile inoculation needle, a mass of conidia was picked up from the grown fungus on the host with the aid of a dissection microscope and suspended on 2 % malt extract agar (MEA; Merck, Darmstadt, Germany) plates supplemented by streptomycin sulphate (100 mg/l) containing 10 ml sterile water. The suspension was evenly spread on the surface of the medium and plates were kept overnight in an oblique position. The plates were then checked under the dissection microscope and germinating conidia were transferred to the potato carrot agar plates (PCA; freshly prepared according to Crous et al. 2009). Single-spore cultures were preserved on PCA in 2 ml agar slants at 4 °C.


Morphological identification

Morphological characteristics were examined both on the natural substrate and in vitro, following the protocol of Sahr et al. (1999). For all of the isolates, the colony color (surface and reverse) and growth rates were recorded on MEA after plate incubation at 25 °C in the darkness. Colour notations were conducted according to the Rayner (1970).

Microscopic characters were examined based on the shape and size of conidia and aleurioconidia on conidiophores and MEA medium, respectively (Sahr et al. 1999). Sample slides were prepared from agar cultures using a sterilized needle or with the oblique coverslip method (Nugent et al. 2006). All the microscopic characters were examined and measured using sterile water as a mounting medium. Whenever possible, a minimum of 25 measurements was made per structure with extreme values given in parentheses. Olympus digital camera system (DP 25) mounted on an Olympus BX41 light microscope was applied to take photographs of microscopic fungal structures. Adobe Photoshop CS6 (Adobe Systems Inc., USA) was used to edit the photos and prepare photo plates. Representative cultures were deposited in the culture collection of Tabriz University (CCTU), Tabriz, Iran and the culture collection of the Westerdijk Fungal Biodiversity Institute (CBS), Utrecht, Netherlands.


Molecular identification

Genomic DNA was isolated from fungal mycelium grown on MEA using the Moller et al. (1992) protocols. The internal transcribed spacers 1 and 2. the intervening 5.8S gene of the rDNA [ITS] and the large subunit gene of the rDNA [LSU] were amplified and sequenced using the following primer combinations: ITS1 plus ITS4 for ITS (White et al. 1990), and LR0R plus LR5 for LSU (Vilgalys & Hester 1990, Vilgalys & Sun 1994). Polymerase chain reaction amplifications were performed on the GeneAmp PCR System 9700 (Applied Biosystems, Foster City, CA) or a 2720 thermal cycler (Applied Biosystems, Foster City, CA). PCR amplification reactions were prepared with total volume of 12.5 µl and contained 0.1 µl Taq DNA polymerase (5 U/µl BIOTAQ™ DNA Polymerase, BioLine, Germany), 1.25 µl PCR buffer (10X NH4 reaction buffer, BioLine, Germany), 0.5 µl MgCl2 (50 mM, BioLine, Germany), 0.5 µl dNTP mix (10 mM, BioLine, Germany), 0.7 µl dimethyl sulfoxide (DMSO, Sigma-Aldrich, Germany), 0.25 µl of each primer (10 µM) and 1 µl of template DNA. The PCR reaction was carried out a 94 ºC for 180 s, 35 cycles of denaturation/extension at 94 ºC for 30 s, annealing for 60 s at 57 ºC for ITS and LSU, the 80 s at 72 ºC, and a final extension for 60 s at 72 ºC. The amplified products were purified using Sephadex® G-50 Fine (GE Healthcare, Sigma-Aldrich, Germany) and were sequenced by the BigDye Terminator v. 3.1 (Applied Biosystems, Foster City, CA, USA) Cycle Sequencing Kits and subsequently analysed on an ABI Prism 3700 or an AB 3730XL DNA Analyzer (Applied Biosystems, Foster City, CA, USA) according to the manufacturer's recommended instructions. Raw sequence files were edited manually and consensus sequences were generated from each forward and reverse sequence using SeqMan™ II (DNASTAR, Madison, WI, USA). The sequences were subjected to BLAST search at GenBank and sequences with high similarity were downloaded and included in the alignment file. Sequence alignments were performed independently for each gene using MAFFT (Katoh & Standley 2013, Li et al. 2015) under the European Bioinformatics Institute (EMBL-EBI) webserver. Alignments were visually inspected and manually edited using MEGA v. 6.06 (Tamura et al. 2013). Additional sequences were obtained from GenBank.


Phylogenetic analysis

The phylogenetic analyses included Bayesian (B) were conducted using XSEDE platform on the CIPRES Science Gateway Portal (Miller et al. 2012). Evolutionary models were calculated using MrModelTest v. 2.3 (Nylander 2004) and the gaps were coded as missing data. Bayesian analyses included two Markov chains of four incrementally heated runs each and lasted for 5 M generations with the stoprule option on, a stopval value set to 0.01 and a sampling frequency of every 1000 generations. After runs conversion, 50 % majority rule consensus tree and posterior probabilities were calculated after discarding 25 % of initial trees as a burn-in fraction. Statistical support for the branches was evaluated using bootstrap analysis (BS) of 1000 replicates.


A total number of ten specimens corresponding to two different fungal hosts i.e., Boletus sp. and Xerocomus sp. were collected. Ten Sepedonium isolates with similar cultural and morphological features were obtained from collected specimens. The isolates were identified as S. microspermum based on cultural and morphological characteristics.

Sepedonium microspermum Besl, Zeitschrift für Mykologie 64 (1): 46 (1998)

Colonies on MEA medium reached 9-10 mm in diameter after seven days; colony surface was initially white, then became yellow after seven days with flat to velvet and zonate with abundant aleurioconidia, margin entire (Fig. 1). Conidiophores arising from aerial mycelium, macronematous, hyaline, septate, with single or 2-3 verticillate and slender phialides, (45-)59-66(-75) × (2-)3-4(-5) µm, apex 1-2 µm width; phialoconidia ovoid to fusiform, unicellular, smooth, (9-)11-13(-17) × (3-)4-6(-7) µm. Aleurioconidia on short side branches were yellow, globose, 8-14 µm diam., with angular tubercles; mycelium developing irregular spots and pustules; optimum growth at 25 ˚C.

Our BLAST search of ITS sequence data against the nucleotide sequences at GenBank showed high similarity with S. microspermum. Because of low LSU sequence data, we did not include our LSU sequence (MH878229) in the phylogenetic analysis. A phylogeny inferred based on ITS sequence data obtained in this study together with 90 sequences from GenBank grouped our sequence along with S. microspermum (AF054847) as a reference sequence chosen by Kadri Põldmaa in 2014 (Nilsson et al. 2018) in a clade composed of S. microspermum collections from different hosts that are available in GenBank. Based on ITS phylogeny, four species clades could be recognized within S. microspermum isolates which might represent additional cryptic species in S. microspermum. However, analysis of additional isolates and more genomic loci are required to address this question (Valdez & Douhan 2012). Sepedonium microspermum has been described only recently and is well characterized by smaller aleurioconidia and more distinctive tubercles than S. chrysospermum (Besl et al. 1998). Sepedonium microspermum has been linked to Hypomyces microspermum; however, in the present study, the sexual state was not observed. This species has a worldwide distribution and is currently known from a diverse range of substrates including soil, plant materials and other fungi (Arellano-Galindo et al. 2017).

Additional material examined: Iran, East Azerbaijan province, the campus of the University of Tabriz, on Xerocomus sp., April 2016, M. Torbati (CCTUMO14=TuXe1)

Fig. 1. Sepedonium microspermuma. 7-d-old colony on MEA; b-d. Conidiophores and conidia; e-h. aleurioconidia. Scale bar — b-h = 10 μm.

Fig. 2. Bayesian inference phylogenetic tree of Sepedonium microspermum generated using sequences of the internal transcribed spacer (ITS-rDNA). The representative strain CBS 141557 in this study is bolded in blue. The values above branches show Bayesian posterior probability. The scale bar indicates the number of expected substitutions per site. Hypomyces aurantius (MH858568) was used as the out-group



This project was supported by the Research Deputy of the University of Tabriz, Iran.

Arellano-Galindo J, Eugenio VM, Elva JH, Jesus RS, Maria de los Angeles MR, Rodolfo Norberto JJ, Juan XC, Sara AO, Ariadna CC. 2017. A saprophytic fungus (Sepedonium) associated with fatal pneumonia in a patient undergoing stem cell transplantation. International Medical Research 45: 1430–1434.

Asef MR, Mohammadi Goltapeh E. 2002. Identification of fungicolous fungi of Iran I. Cladobotryum species. Rostaniha 11–22.

Asef MR, Zare R. 2006. Identification of fungicolous fungi of Iran. II. Teleomorphs belonging to the genus Hypomyces. Rostaniha 7: 35–42.

Barnett HL. 1963. The nature of mycoparasitism by fungi. Annual Review of Microbiology 17: 1–14.

Barnett HL. 1964. Mycoparasitism. Mycologia 56: 1–19.

Besl H, Hagn A, Jobst A, Lange U. 1998. Der Kleinsporige Goldschimmel, Sepedonium microspermum-ein Parasit an Röhrlingen der Xerocomus chrysenteron Gruppe. Zeitschrift für Mykologie 64: 45–52.

Crous PW, Verkley GJM, Groenewald JZ, Samson RA. 2009. Fungal Biodiversity. CBS Laboratory Manual Series 1: 1–269. Centraalbureau voor Schimmelcultures, Utrecht, the Netherlands.

De Hoog GS. 1978. Notes on some fungicolous Hypomyces and their relatives. Persoonia 10: 33–81.

Gams W, Diederich P, Põldmaa K. 2004. Fungicolous fungi. In: Biodiversity of fungi inventory and monitoring methods. (Mueller GM, Bills GF, Foster MS, eds). 343–392. Elsevier Academic Press, USA.

Gams W, and Hoozemans ACM. 1970. Cladobotryum Konidienformen von Hypomyces Arten. Persoonia 6: 95– 110.

Gilman JC, Tiffany LH. 1952. Fungicolous fungi from Iowa. Proceedings of the Iowa Academy of Sciences 59: 99–110.

Hawksworth DL. 1979. The lichenicolous hyphomycetes. Bulletin of the British Museum (Natural History) 6: 183–300.

Hawksworth DL. 1981. A survey of the fungicolous conidial fungi. Pp. 171 244. In: Biology of conidial fungi. (Cole GT, Kendrick B, eds). Vol. 1. Academic Press, USA.

Hawksworth DL, Kirk PM, Sutton BC, Pegler DN. 1995. Ainsworth and Bisby's Dictionary of the fungi, 8th ed. CAB International, Wallingford, UK.

Helfer W. 1991. Pilze auf Pilzfruchtkörpern. Untersuchungen zur Ökologie, Systematik und Chemie. IHW Verlag, Libri Botanici 1:33–35.

Jeffries P. 1995. Biology and ecology of mycoparasitism. Canadian Journal of Botany 73: S1284–S1290. (Suppl. 1)

Jiao P. 2006. Chemical investigations of freshwater and fungicolous fungi. PhD thesis, Chemistry Department, The University of Iowa, United States.

Katoh K, Standley DM. 2013. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Molecular Biology and Evolution 30: 772–780.

Keizer GJ. 2004. The complete encyclopedia of mushrooms: more than 700 pictures and descriptions of mushrooms. 2nd edn. Rebo, Netherlands.‏

Li W, Cowley A, Uludag M, Gur T, McWilliam H, Squizzato S, Park YM, Buso N, Lopez R. 2015. The EMBL-EBI bioinformatics web and programmatic tools framework. Nucleic Acids Research 43: W580–W584.

Miller MA, Pfeiffer W, Schwartz T. 2012. The CIPRES science gateway: enabling high-impact science for phylogenetics researchers with limited resources. In: Proceedings of the 1st Conference of the Extreme Science and Engineering Discovery Environment: Bridging from the extreme to the campus and beyond: 1–8. Association for Computing Machinery, USA.

Möller EM, Bahnweg G, Geiger HH. 1992. A simple and efficient protocol for isolation of high molecular weight DNA from filamentous fungi, fruit bodies, and infected plant tissues. Nucleic Acids Research20: 6115–6116.

Nilsson RH, Larsson KH, Taylor AFS, Bengtsson-Palme J, Jeppesen TS, Schigel D, Kennedy P, Picard K, Glöckner FO, Tedersoo L, Saar I, Kõljalg U, Abarenkov K. 2018. The UNITE database for molecular identification of fungi: handling dark taxa and parallel taxonomic classification. Nucleic Acids Research 47: D249–D264.

Nilsson RH, Larsson K-H, Taylor AFS, Bengtsson-Palme J, Jeppesen TS, Schigel D, Kennedy P, Picard K, Glöckner FO, Tedersoo L, Saar I, Kõljalg U, Abarenkov K. 2018. The UNITE database for molecular identification of fungi: handling dark taxa and parallel taxonomic classifications. Nucleic Acids Research, DOI: 10.1093/nar/gky1022

Nugent LK, Sangvichen EK, Sihanonth P, Ruchikachorn N, Whalley AJS. 2006. A revised method for the observation of conidiogenous structures in fungi. Mycologist 20: 111–114.

Nylander JAA. 2004. MrModeltest v2.0. Program distributed by the author. Evolutionary Biology Centre, Uppsala University, Uppsala, Sweden

Põldmaa K. 2000. Generic delimitation of fungicolous Hypocreaceae. Studies in Mycology 45: 83–94.

Rayner RW. 1970. A mycological colour chart. CMI and British Mycological Society, Kew, Surrey, England.

Rogerson CT, Samuels GJ. 1985. Species of Hypomyces and Nectria occuring on Discomycetes. Mycologia 77: 763–783.

Rogerson CT, Samuels GJ. 1989. Boleticolous species of Hypomyces. Mycologia 81: 413–432.

Rogerson CT, Samuels GJ. 1993. Polyporicolous species of Hypomyces. Mycologia 85: 231–272.

Rogerson CT, Samuels GJ. 1994. Agaricicolous species of Hypomyces. Mycologia 86: 839–866.

Rossman AY, Samuels GJ, Rogerson CT, Lowen R. 1999. Genera of Bionecteriaceae, Hypocreaceae and Necteriaceae (Hypocreales, Ascomycetes. Studies in Mycology 42: 1–248.

Rudakov OL. 1978. Physiological groups in mycophilic fungi. Mycologia 70:150­–159.

Rudakov OL. 1981. Mikofil'nye griby, ikh biologiya i prakticheskoe znachenie. Iz'vo Nauka, Moskva.

Sahr T, Ammer H, Besl H, Fischer M. 1999. Infrageneric classification of the boleticolous genus Sepedonium: species delimitation and phylogenetic relationships. Mycologia 935–943pp.

Sun JZ, Liu XZ, McKenzie EH, Jeewon R, Liu JKJ, Zhang XL, Zhao Q, Hyde KD. 2019. Fungicolous fungi: terminology, diversity, distribution, evolution, and species checklist. Fungal Diversity 1–94.‏

Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. 2013. MEGA6: Molecular Evolutionary Genetics Analysis Version 6.0. Molecular Biology and Evolution 30:2725–2729.

Torbati M, Arzanlou M, Sandoval-Denis M, Crous PW. 2018. Multigene phylogeny reveals new fungicolous species in the Fusarium tricinctum species complex and novel hosts in the genus Fusarium from Iran. Mycological Progress 18: 119–133.

Vilgalys R, Hester M. 1990. Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. Journal of Bacteriology 172: 4238–4246.

Valdez GU, Douhan GW. 2012. Geographic structure of a bolete-infecting cryptic species within the Hypomyces microspermus species complex in California. Mycologia 104: 14–21.

Vilgalys R, Sun BL. 1994. Ancient and recent patterns of geographic speciation in the oyster mushroom Pleurotus revealed by phylogenetic analysis of ribosomal DNA sequences. Proceedings of the National Academy of Sciences of the United States of America 91: 4599–4603.

White TJ, Bruns T, Lee S., Taylor J. 1990. Amplification and Direct Sequencing of Fungal Ribosomal RNA Genes for Phylogenetics. In: Innis MA, Gelfand DH, Sninsky JJ, White TJ, Eds., PCR Protocols. A Guide to Methods and Applications, Academic Press, San Diego, 315–322. Zare R, Asef M. 2008. Some phialidic fungicolous fungi from the Southern Caspian coasts. Rostaniha 9: 1–22.