What are Coelomycetes?
Spores are often produced inside specialized structures called fruitbodies, which are visible under the stereo microscope. Since spores can be produced either sexually or a sexually, fruitbodies are accordingly divided after to the type of spores they carry, either sexual (meiotic) or asexual (mitotic). In Ascomycetes and Basidiomycetes, fruitbodies with sexual spores are known as ascomata and basidiomata, respectively. Ascomata contain asci, which are sac-like structures in which the sexual ascospores are produced. The ascomata have different phenotypes, i.e. cleistothecia (closed), perithecia (with a preformed opening), or gymnothecia (surrounded by a hyphal network). Fruitbodies carrying asexual propagules (conidia) are named conidiomata (plural; conidioma = singular). In analogy to ascomata, conidiomata are also divided into morphological types: pycnidia (closed) or acervuli (open, saucer-shaped). Fungi producing conidiomata are referred to as coelomycetes (Sutton 1980; Stchigel & Sutton 2013). “Coelomycetes” thus is a phenotypic category, irrespective of the phylogenetic or taxonomic position of the species producing the conidioma. However, today we classify fungi according to phylogenetic relationships and no longer according to morphology. Therefore the “coelomycetes” have been abandoned as a taxonomic group; you can’t find it anymore as a section in the Atlas.
Not all ascomycetes are able to produce sexual forms readily; in fact, for many of them the sexual form is even unknown. Of the several hundreds of clinical species listed in the Atlas, only about 50 are able to produce ascomata under laboratory conditions (see pages 519 and 520). Cultures mostly display asexual forms of propagation. Traditionally, fungi which produce their conidia freely on the hyphae, without fruitbodies, were known as hyphomycetes, and these were classified as a separate group of the fungi, just as the coelomycetes. Main morphological criterion to distinguish species in both asxual groups concerned the conidial ontogeny (conidiogenesis) (Sutton 1960). With the introduction of genetic and molecular methods in fungal classification, the categories “hyphomycetes” and “coelomycetes” became obsolete. However, the terms are still in use for accurate description of fungal taxa, but not to determine the taxonomic position. Thus, a conidioma-producing fungus from a clinical specimen can still be called a “coelomycete”, as a descriptive term, but for precise species identification experimental diagnostics are needed (Stchigel & Sutton 2013).
Molecular and phylogenetic studies in fungi have revolutionized fungal taxonomy over the past decades. Relations between taxa are assessed on the basis of phylogenetic trees, built with one or multiple genetic markers. In the past, classification was done speculatively in coelomycetous fungi using morphology of conidiomata, conidia, and conidiogenesis (Wijayawardene et al. 2012). These phenotypic features were however found to be unstable and also non-specific: closely related species can produce phenotypically different fruitbodies, and in contrast, distantly related species might appear similar. That is why the DNA-based taxonomy resulted in dramatic changes in coelomycetous taxa (Valenzuela-Lopez et al. 2017). Large numbers of new species have been introduced, while elsewhere some large genera were reduced to just few species. An interesting example is the genus Phoma, a classical genus of coelomycetes previously containing about 200 species. Given its high phenotypic diversity, Boerema (1997) had grouped Phoma species into nine sections. But later phylogenetic studies revealed that the genus was highly polyphyletic, species belonging to different orders of the fungal Kingdom not matching the morphological sections (de Gruyter et al. 2009). Accordingly, many species were transferred to other, existing or novel genera. In the monophyletic lineage containing the type species of Phoma, P. herbarum, only five or six species remain. This little group is now referred to as Phoma sensu stricto (s. str.).
Despite the significant progress in phylogenetic taxonomy of coelomycetes, still numerous issues influencing stable nomenclature and routine clinical diagnostics remain. As discussed in the previous blog, methodical improvements have led to changing fungal names. The new systems reflect relationships between fungi better, but there are examples of repeated name changes within just a few years, simply due to the addition of new material or genetic markers. Phylogenetic trees are very sensitive to which taxa are analyzed, known as “taxon sampling effects”. Thus, good coverage of the fungal group, with sufficient taxa at both high and low taxonomic ranks is necessary (Chen et al. 2015). Given the enormous diversity of fungi producing conidiomata, this is not so easy (www.coelomycetes.org).
Many coelomycetes have an important role as plant pathogens or opportunists on humans. Because of their hidden diversity, responses to therapy may differ between species, and therefore identification is essential. We now know that specific identification of coelomycetes can only be achieved by DNA sequencing. Clinical isolates are often sterile, sporulate only after lengthy incubation, or show insufficient diagnostic morphology, but are much more diverse than we thought in the past. A study by Valenzuela-Lopez et al. (2017) of clinical isolates from a single laboratory revealed these strains belonged to 11 orders of Ascomycota. The number of agents is also increasing because molecular methods are independent from sporulation and allow identification of sterile cultures. The newly described species sometimes appear to be quite common. As an example, the recent description of a strain from a keratitis in the new genus Tintelnotia soon afterwards led to repeated discovery of this fungus as a cause of superficial infection in several European countries and in Australia (Ahmed et al. 2017). Below a Table is provided where you can find all clinically relevant coelomycetous species, where they belong phylogenetically, and under which name you can find them in the Atlas.
References:
- Sutton BC (1980) The coelomycetes. Fungi with pycnidia, acervuli and stromata. Kew, Surrey, England, Commonwealth Mycological Institute.
- Stchigel AM, Sutton DA (2013) Coelomycete fungi in the clinical lab. Curr Fungal Infect Rep 7: 171–191.Sutton BC. 1961. Coelomycetes I. Mycol. Pap. 80: 1–16.
- Wijayawardene NN, Mckenzie EHC, Chukeatirote E, et al. (2012) Coelomycetes. Crypt. Mycol. 33: 215–244.
- Valenzuela-Lopez N, Cano-Lira JF, Stchigel AM, Guarro J (2018) DNA sequencing to clarify the taxonomical conundrum of the clinical coelomycetes. Mycoses. 61: 708-717.
- Boerema GH (1997) Contributions towards a monograph of Phoma (Coelomycetes) – V. Subdivision of the genus in sections. Mycotaxon 64: 321-333.
- de Gruyter J, Aveskamp MM, Woudenberg JH, et al. (2009) Molecular phylogeny of Phoma and allied anamorph genera: towards a reclassification of the Phoma complex. Mycol. Res. 113: 508-519.
- Chen Q, Jiang JR, Zhang GZ, et al. (2015) Resolving the Phoma enigma. Stud. Mycol. 82: 137-217.
- Valenzuela-Lopez N, Sutton DA, Cano-Lira JF, et al. (2017) Coelomycetous fungi in the clinical setting: morphological convergence and cryptic diversity. J. Clin. Microbiol. 55: 552-567.
- Ahmed SA, Hofmüller W, Seibold M, et al. (2017) Tintelnotia, a new genus in Phaeosphaeriaceae harbouring agents of cornea and nail infections in humans. Mycoses 60: 244-253.
| Prevalent species name | Obsolete name | Order |
| Boeremia exigua | Phoma exigua | Pleosporales |
| Colletotrichum acutatum | Glomerellales | |
| Colletotrichum chlorophyti | Glomerellales | |
| Colletotrichum coccodes | Chaetomium coccodes, Vermicularia tramentarium, Colletotrichum atramentarium | Glomerellales |
| Colletotrichum dematium | Sphaeria dematium, Vermicularia dematia | Glomerellales |
| Colletotrichum gigasporum | Glomerellales | |
| Colletotrichum gloeosporioides | Vermicularia gloeosporioides | Glomerellales |
| Colletotrichum graminicola | Dicladium graminicola | Glomerellales |
| Colletotrichum truncatum | Vermicularia truncata | Glomerellales |
| Diaporthe bougainvilleicola | Phomopsis bougainvilleicola | Diaporthales |
| Diaporthe longicolla | Phomopsis longicolla | Diaporthales |
| Diaporthe oculi | Diaporthales | |
| Diaporthe phaseolorum | Phoma phaseoli, Phomopsis phaseoli, Sphaeria phaseolorum | Diaporthales |
| Diaporthe phoenicicola | Phomopsis phoenicicola, Subramanella arecae, Diaporthe arecae | Diaporthales |
| Diaporthe pseudooculi | Diaporthales | |
| Didymella macrostoma | Phoma macrostoma | Pleosporales |
| Didymella microchlamydospora | Phoma microchlamydospora | Pleosporales |
| Didymella glomerata | Coniothyrium glomeratum, Phoma glomerata, Peyronellaea glomerata,Phoma hominis, Alternaria hominis, Peyronellaea hominis | Pleosporales |
| Didymella gardeniae | Pyrenochaeta gardeniae, Phoma gardeniae, Peyronellaea gardeniae | Pleosporales |
| Didymella heteroderae | Phoma heteroderae, Peyronellaea heteroderae | Pleosporales |
| Emarellia grisea | Pleosporales | |
| Emarellia paragrisea | Pleosporales | |
| Epicoccum sorghinum | Phyllosticta sorghina, Phoma sorghina, Chaetophoma dermo-unguis | Pleosporales |
| Gloniopsis percutanea | Hysteriales | |
| Hongkongmyces pedis | Pleosporales | |
| Hongkongmyces snookiorum | Pleosporales | |
| Juxtiphoma eupyrena | Phoma eupyrena (used in the Atlas) | Pleosporales |
| Lasiodiplodia lignicola | Auerswaldia lignicola | Dothideales |
| Lasiodiplodia pseudotheobromae | Dothideales | |
| Lasiodiplodia theobromae | Diplodia gossypina, Physalospora rhodina, Botryosphaeria rhodina, Botryodiplodia theobromae | Dothideales |
| Macrophomina phaseolina | Macrophoma phaseolina, Botryodiplodia phaseoli | Dothideales |
| Medicopsis romeroi | Pyrenochaeta romeroi | Pleosporales |
| Microsphaeropsis arundinis | Alveophoma arundinis | Pleosporales |
| Microsphaeropsis olivacea | Coniothyrium olivaceum | Pleosporales |
| Neocucurbitaria cava | Phoma cava, Pleurophoma cava, Pyrenochaeta cava | Pleosporales |
| Neocucurbitaria keratinophila | Pyrenochaeta keratinophila | Pleosporales |
| Neocucurbitaria unguis-hominis | Pyrenochaeta unguis-hominis | Pleosporales |
| Neodeightonia subglobosa | Neodeightonia subglobosa, Botryosphaeria subglobosa | Dothideales |
| Neoscytalidium dimidiatum | Torula dimidiata, Scytalidium dimidiatum, Fusicoccum dimidiatum, Hendersonula toruloidea,Scytalidium hyalinum, Neoscytalidium hyalinum, Neoscytalidium hyalinum | Dothideales |
| Neoscytalidium novaehollandiae | Dothideales | |
| Nigrograna mackinnonii | Pyrenochaeta mackinnonii, Biatriospora mackinnonii | Pleosporales |
| Pallidocercospora crystallina | Pseudocercospora crystallina, Mycosphaerella crystallina | Dothideales |
| Paraconiothyrium cyclothyrioides | Pleosporales | |
| Paraconiothyrium fuckelii | Sphaeria coniothyrium, Leptosphaeria coniothyrium, Coniothyrium fuckelii | Pleosporales |
| Paraconiothyrium maculicutis | Pleosporales | |
| Phoma herbarum | Phoma cruris-hominis, Phoma hibernica, Atradidymella muscivora , Atradidymella muscivora | Pleosporales |
| Pleurophomopsis lignicola | Pleosporales | |
| Pseudochaetosphaeronema larense | Chaetosphaeronema larense | Pleosporales |
| Pseudochaetosphaeronemamartinelli | Pleosporales | |
| Parathyridaria percutanea | Roussoella percutanea (used in the Atlas) | Pleosporales |
| Roussoella solani | Neoroussoella solani | Pleosporales |
| Stagonosporopsis oculi-hominis | Phoma oculo-hominis, Phoma dennisii | Pleosporales |
| Tintelnotia destructans | Pleosporales | |
| Trematosphaeria grisea | Madurella grisea | Pleosporales |
| Westerdykella dispersa | Pycnidiophora dispersa, Preussia dispersa | Pleosporales |
| Westerdykella minutispora | homa oryzae, Phoma minutispora | Pleosporales |
