Novel species of Cladosporium from environmental sources in Spain

Isabel Iturrieta-González; Dania García; Josepa Gené

doi:10.3897/mycokeys.77.60862

Abstract

Cladosporium is a monophyletic genus in Cladosporiaceae (Cladosporiales, Dothideomycetes) whose species are mainly found as saprobes and endophytes, but it also includes fungi pathogenic for plants, animals and human. Species identification is currently based on three genetic markers, viz., the internal transcribed spacer regions (ITS) of the rDNA, and partial fragments of actin (act) and the translation elongation factor 1-α (tef1) genes. Using this phylogenetic approach and from morphological differences, we have recognized six new species originating from soil, herbivore dung and plant material collected at different Spanish locations. They are proposed as Cladosporium caprifimosum, C. coprophilum, C. fuscoviride and C. lentulum belonging in the C. cladosporioides species complex, and C. pseudotenellum and C. submersum belonging in the C. herbarum species complex. This study revealed that herbivore dung represented a reservoir of novel lineages in the genus Cladosporium.

Keywords

Cladosporiales, Cladosporiaceae, hyphomycetes, phylogeny, Spain, taxonomy

Introduction

Cladosporium is a ubiquitous genus in the family Cladosporiaceae of the recently proposed order Cladosporiales in the Dothideomycetes (Abdollahzadeh et al. 2020). Their species inhabit a wide range of substrates and have been reported to be among the most common fungi in both indoor and outdoor environments, including in extreme ecological niches (Flannigan et al. 2002; Bensch et al. 2010, 2012, 2018; Sandoval-Denis et al. 2015; Temperini et al. 2018; Chung et al. 2019). Most Cladosporium species are saprobic, but some have also been reported as endophytes, hyperparasites on other fungi and plant as well as animal pathogens, including humans (Heuchert et al. 2005; Sandoval-Denis et al. 2016; de Hoog et al. 2017; Marin-Felix et al. 2017). Certain species show the ability to produce compounds of medical interest or are relevant as potential biocontrol agents for plant disease (Köhl et al. 2015; Khan et al. 2016; Adorisio et al. 2019).

Cladosporium is morphologically characterized mainly by its asexual morph, which shows differentiated conidiophores producing acropetal chains of conidia from mono- or polyblastic conidiogenous cells. Both conidiogenous cells and conidia exhibit conidiogenous loci (scars) with a unique coronate structure, which is composed of a central convex dome surrounded by a raised periclinal rim, usually thickened, refractive and darkly pigmented (David 1997). Based on these features and DNA phylogeny derived from the LSU nrRNA gene, the genus has been well-delineated and distinguished from other cladosporium-like genera such as Hyalodendriella, Ochrocladosporium, Rachicladosporium, Rhizocladosporium, Toxicocladosporium, Verrucocladosporium and the recently described genus Neocladosporium (Crous et al. 2007; Bezerra et al. 2017). Phylogenetic relationships among species of Cladosporium s. str. have been studied extensively over the last decade by a multi-locus sequence analysis approach with sequences of the internal transcribed spacers (ITS) of the rDNA and of the two protein encoding genes, translation elongation factor 1-α (tef1) and actin (act). The molecular approach combined with morphological features have allowed recognition of more than 230 species within the genus, which are split into three species complexes, i.e., the Cladosporium cladosporioides, Cladosporium herbarum and Cladosporium sphaerospermum complex (Schubert et al. 2007; Bensch et al. 2010, 2012, 2015, 2018; Sandoval-Denis et al. 2016; Marin-Felix et al. 2017).

While aiming to explore the diversity of microfungi from Spain, several interesting Cladosporium isolates have been recovered from different environmental samples. Using the above mentioned polyphasic approach and following the Genealogical Phylogenetic Species Recognition (GCPSR) criterion (Taylor et al. 2000), the taxonomy of those isolates has been resolved in six novel species for science; four pertaining to the C. cladosporioides species complex and two to the C. herbarum complex.

Material and methods

Samples and isolates

Samples of soil, plant debris and herbivore dung were collected between 2016 and 2018 at various Spanish locations. Dilution plating methods were used for isolating fungi from soil and dung samples following the procedure described by Crous et al. (2009) and a modified protocol described by Waksman (1922), respectively. In addition, soil samples were also processed by a baiting technique using small pieces of wood and filter paper as baits on the soil surface (Calduch et al. 2004). Samples of plant debris and also part of the herbivore dung were incubated in moist chambers following the procedures described by Castañeda-Ruiz et al. (2016) and Richardson (2001), respectively.

Among the cladosporium-like fungi found, we recovered eight isolates in pure culture which did not match any of the currently accepted species within the genus Cladosporium (Table 1). The isolates were deposited in the culture collection of the Universitat Rovira i Virgili (FMR, Reus, Spain) and, once phylogenetically and morphologically characterized, living cultures of the novel species and dry cultures for holotypes were also deposited in the Westerdijk Fungal Biodiversity Institute (CBS; Utrecht, the Netherlands). Nomenclatural novelties and descriptions were deposited in MycoBank (Crous et al. 2004).

Table 1.

Download as

CSV

XLSX

Cladosporium species, strain information and GenBank accession numbers for sequences obtained in this study.

Species	Strain number¹	Substrate	GenBank nucleotide accession no. for²:
Species	Strain number¹	Substrate	ITS	*act*	*tef1*
C. caprifimosum	FMR 16532^T	Goat dung	LR813198	LR813205	LR813210
C. coprophilum	FMR 16101	Unidentified herbivore dung	LR813199	LR813204	LR813211
C. coprophilum	FMR 16164^T	Unidentified herbivore dung	LR813201	LR813207	LR813213
C. fuscoviride	FMR 16385^T	Garden soil	LR813200	LR813206	LR813212
C. lentulum	FMR 16288^T	Unidentified leaf litter	LR813203	LR813209	LR813215
C. lentulum	FMR 16389	Unidentified herbivore dung	LR813202	LR813208	LR813214
C. pseudotenellum	FMR 16231^T	Garden soil	LR813145	LR813146	LR813196
C. submersum	FMR 17264^T	Submerged plant material	LR813144	LR813195	LR813197

DNA extraction, amplification and sequencing

Genomic DNA was extracted from cultures growing on potato dextrose agar (PDA; Pronadisa, Spain) after 7 days of incubation at 25 °C, following the modified protocol of Müller et al. (1998). Protocols listed previously in Sandoval-Denis et al. (2016) were used for amplification and sequencing. The primer pairs used were ITS5/ITS4 (White et al. 1990) to amplify the ITS region including the 5.8S gene of the rDNA, EF-728F/EF-986R to amplify a partial fragment of the tef1 gene, and ACT-512F/ACT-783R to amplify a partial fragment of act gene (Carbone and Kohn 1999). PCR products were purified and stored at -20 °C until sequencing. The sequences were obtained using the same primers at Macrogen Europe (Macrogen Inc. Amsterdam, The Netherlands). Finally, the software SeqMan v. 7.0.0 (DNAStarLasergene, Madison, WI, USA) was used to assemble, edit and obtain the consensus sequences, which were then deposited in GenBank of the National Center for Biotechnology Information (NCBI) (Table 1).

Sequence alignment and phylogenetic analysis

The sequences obtained were compared with other fungal sequences deposited in the NCBI database through the BLASTn tool. Alignment of those sequences and the phylogenetic analysis for each locus were performed with the MEGA (Molecular Evolutionary Genetics Analysis) program v. 6.0. (Tamura et al. 2013), using ClustalW algorithm (Thompson et al. 1994) and refined with MUSCLE (Edgar 2004) or manually if necessary, on the same platform. Since the isolates under study were related to the C. cladosporioides and C. herbarum species complexes, we also carried out alignments including sequence data of ex-type and reference strains of all the species from both complexes retrieved from the GenBank and mainly published by Schubert et al. (2007, 2009), Bensch et al. (2010, 2012, 2015, 2018), Sandoval-Denis et al. (2016) and Marin-Felix et al. (2017) (Suppl. material 1: Table S1).

Phylogenetic reconstructions were made with the three phylogenetic markers (ITS, act and tef1) recommended for an accurate identification at the species level (Bensch et al. 2010, 2018; Marin-Felix et al. 2017) using Maximum Likelihood (ML), Maximum Parsimony (MP), and Bayesian Inference (BI) analyses, with the Mega software v. 6.0. for the former two (Tamura et al. 2013) and with MrBayes v.3.2.6 for the latter one (Ronquist et al. 2012). Phylogenetic concordance of the three-locus datasets was evaluated through Incongruence Length Difference (ILD) implemented in the Winclada program (Farris et al. 1994) and also by visual comparison of the individual phylogenies in order to assess any incongruent results between nodes with high statistical support.

Determined by Mega software v. 6.0., the best nucleotide substitution model for ML analysis of the C. cladosporioides complex was General Time Reversible with Gamma distribution and invariant sites (GTR+G+I), and for the C. herbarum complex the best was the Kimura 2-parameter with Gamma distribution and invariant sites (K2+G+I). Bootstrap support value (MLBS) ≥ 70% was considered significant (Hillis and Bull 1993).

The MP analysis was performed using the heuristic search option with TBR (tree bisection and reconnection) branch swapping and 1,000 random sequence additions. Tree length (TL), consistency index (CI), retention index (RI), rescaled consistency index (RCI) were calculated. Bootstrap analysis was based on 1,000 replications. Maximum parsimony bootstrap support value (PBS) ≥ 70% was considered significant (Hillis and Bull 1993).

Determined by jModelTest (Posada 2008), the best nucleotide substitution models for the BI of the C. cladosporioides complex were Jukes Cantor with invariant sites (JC+I) for ITS, General Time Reversible with Gamma distribution (GTR+G) for tef1 and Hasegawa-Kishino-Yano with Gamma distribution (HKY+G) for act; and for the C. herbarum complex the best were the Kimura 2-parameter with Gamma distribution (K80+G) for ITS, Hasegawa-Kishino-Yano with Gamma distribution (HKY+G) for tef1 and act. The parameter settings used in these analyses were two simultaneous runs of 10,000,000 generations, and four Markov chains, sampled every 1,000 generations. The 50% majority rule consensus tree and posterior probability values (PP) were calculated after discarding the first 25% of the samples. A PP value of ≥ 0.95 was considered significant (Hespanhol et al. 2019).

Final sequence alignments and trees generated in this study were registered in TreeBASE under the submission number S27350 (http://treebase.org).

Phenotypic studies

Microscopic features of the Cladosporium isolates were obtained from cultures growing on synthetic nutrient-poor agar (SNA; 1 g of KH₂PO₄, 1 g of KNO₃, 0.5 g of MgSO₄ × 7H₂O, 0.5 g of KCl, 0.2 g of glucose, 0.2 g of sucrose, 14 g of bacteriological agar, 1 L of distilled water) after 7 to 14 days at 25 °C in the dark, mounted onto semi-permanent slides with Shear's solution (Bensch et al. 2018). At least 30 measurements were taken to calculate length and width ranges of the conidia and ramoconidia, given as the mean ± standard deviation in the descriptions. Macroscopic characterization of the colonies was made on PDA (Pronadisa, Spain), oatmeal agar (OA; 30 g of oatmeal, 13 g of bacteriological agar, 1 L distilled water) and SNA after 14 days of incubation at 25 °C in darkness. Colour notation of the colonies in descriptions were from Kornerup and Wanscher (1978). In addition, cardinal temperatures for the fungal growth were determined on PDA cultures after 14 days at temperatures ranging from 5 to 40 °C at intervals of 5 °C.

Results

Phylogeny

Three individual phylogenies (ITS, tef1 and act), carried out for the C. cladosporioides and C. herbarum species complexes, were visually very similar and the ILD test showed that the three loci datasets were congruent in both complexes (P = 0.16) and could be combined. Phylogenies obtained by ML, MP and BI also showed a visual topological congruence and were similar to that obtained by other authors (Marin-Felix et al. 2017; Bensch et al. 2018). The combined alignment of the three mentioned loci datasets encompassed 101 sequences in the C. cladosporioides complex and 58 sequences in C. herbarum complex. The alignment for the former group comprised 1,060 bp (ITS 484 bp, tef1 313 bp and act 263 bp), which included 424 bp variable sites (ITS 47 bp, tef1 239 bp and act 138 bp) and 319 bp phylogenetically informative sites (ITS 25 bp, tef1 193 bp and act 101 bp). Two species of the C. sphaerospermum complex, C. sphaerospermum CBS 193.54 and C. longissimum CBS 300.96, were included as outgroup in this first multi-locus phylogeny (Fig. 1). For the maximum parsimony analysis the maximum of 1,000 equally most parsimonious trees were saved (Tree length = 1614; CI = 0.294; RI = 0.666; RCI = 0.214).

Figure 1.

Maximum likelihood (ML) tree obtained from the combined analysis of ITS, tef1 and act sequences of 101 strains from the C. cladosporioides complex. The tree is rooted with C. sphaerospermum CBS 193.54 and C. longissimum CBS 300.96. Numbers on the branches represent ML bootstrap support values (MLBS) ≥70%, followed by Maximum Parsimony bootstrap support values (PBS) ≥70% and Bayesian posterior probabilities (PP) ≥0.95, lower values are indicate as “-“. Bold branches indicate MLBS/PBS/PP of 100/100/1. Names of species newly described are indicated in bold. Branch lengths are proportional to distance. ^T Ex-type strain. ^ET Ex-epitype strain.

Figure 1.

Continued.

For the C. herbarum species complex, the alignment comprised 1,057 bp (ITS 503 bp, tef1 309 bp and act 245 bp) with 407 bp variable sites (ITS 101 bp, tef1 186 bp and act 120 bp) and 240 bp phylogenetically informative sites (ITS 27 bp, tef1 123 bp and act 90 bp), using Cercospora beticola (CBS 116456) as outgroup (Fig. 2). For the maximum parsimony analysis the maximum of 1,000 equally most parsimonious trees were saved (Tree length = 898; CI = 0.537; RI = 0.661; RCI = 0.355).

Figure 2.

Maximum likelihood (ML) tree obtained from the combined analysis of ITS, tef1 and act sequences of 58 strains from C. herbarum complex. The tree is rooted with Cercospora beticola CBS 116456. Numbers on the branches represent ML bootstrap support values (MLBS) ≥70%, followed by Maximum Parsimony bootstrap support values (PBS) ≥70% and Bayesian posterior probabilities (PP) ≥ 0.95, lower values are indicate as “-“. Bold branches indicate MLBS/PBS/PP of 100/100/1. Names of species newly described are indicated in bold. Branch lengths are proportional to distance. ^T Ex-type strain. ^ET ex-epitype strain. ^NT ex-neotype strain.

The eight unidentified isolates did not match any known lineage of Cladosporium species, six were related to the C. cladosporioides species complex and two to the C. herbarum complex, and together they represented six new phylogenetic species in the genus.

In the combined phylogeny of the C. cladosporioides complex, 71 species were delineated (Fig. 1). The isolates FMR 16101 and FMR 16164 formed a strongly supported terminal clade representative for a unique taxon, but with an uncertain phylogenetic position due to the low statistical support (- MLBS / 77 PBS / - PP) for the nearest lineages of C. chasmanthicola and C. sinuatum. Both unidentified isolates were genetically identical and showed a percentage of identity with the ex-type strains of these latter species of 97.22% and 97.65% for act, and 96.79% and 97.50% for tef1, respectively. A second undescribed monophyletic terminal clade included FMR 16288 and FMR 16389, which grouped with the lineages of C. exasperatum, C. parapenidielloides and C. longicatenatum in a clade with highly supported values (94 MLBS / 84 PBS / 0.98 PP). However, both isolates showed a sufficient genetic distance to be considered a distinct species from the closest, C. longicatenatum and C. parapenidielloides, with a sequence similarity of 95.75% and 95.28% for act and 90.87% and 90.48% for tef1 respect to the ex-type strains of these two known species. FMR 16532 and FMR 16385 formed two distinct monophyletic branches. The former showed an uncertain phylogenetic position with the species in the complex; the comparison of its sequences with those of the GenBank dataset through the BLASTn tool showed that the ITS was 100% similar with several species of the C. cladosporioides complex, while sequences of act and tef1 were 99.12% and 89.02% similar with sequences belonging to C. asperulatum (UTHSC DI-13-216/GenBank LN834541 and CBS 113744/GenBank HM148237, respectively). FMR 16385 was closely related to the ex-type strain of C. alboflavescens (100 MLBS / 100 PBS / 1 PP). The percentages of identity between these latter two fungi (97.79% for act and 96.75% for tef1) together with morphological differences observed allow us to consider them distinct taxa.

In the C. herbarum complex, 40 species were phylogenetically well-delimited, including two novel lineages each represented by FMR 16231 and FMR 17264 (Fig. 2). Both were genetically and morphologically differentiated from their closest relatives, C. tenellum and C. subcinereum, respectively. The percentages of identity observed between the isolate FMR 16231 and the ex-type strain of C. tenellum (CBS 121634) were 97.78%, 83.76% and 100% for act, tef1 and ITS, respectively, and between FMR 17264 and the ex-type strain of C. subcinereum (CBS 140465) were 98.57%, 95.98% and 100% for act, tef1 and ITS, respectively.

The percentages of identity between the six putative new Cladosporium species and their relatives are summarized in Table 2. The novel taxa are described and illustrated in the taxonomy section below.

Table 2.

Download as

CSV

XLSX

Percentage of identity between the novel Cladosporium and their closest species.

Species	Closest taxa	Loci
Species	Closest taxa	ITS	act	tef1
C. caprifimosum (FMR 16532)	C. asperulatum ¹	100	99.12	89.02
C. coprophilum (FMR 16101 and 16164)	C. chasmanthicola	100	97.22	96.79
C. coprophilum (FMR 16101 and 16164)	C. sinuatum ¹	100	97.65	97.50
C. fuscoviride (FMR 16385)	C. alboflavescens	100	97.79	96.75
C. lentulum (FMR 16288 and 16389)	C. exasperatum	100	92.2	81.4–82.7
	C. longicatenatum	100	95.75	90.87
	C. parapenidielloides	100	95.28	90.48
C. pseudotenellum (FMR 16231)	C. tenellum ¹	100	97.5–100	83.2–84.1
C. submersum (FMR 17264)	C. subcinereum	100	98.57	95.98

Taxonomy

Cladosporium caprifimosum Iturrieta-González, Dania García, Gené, sp. nov.

MycoBank No: 836074

Fig. 3

Etymology

The name refers to goat dung, the substrate where the species was isolated (capra = goat and fimus = dung, with the adjectival Latin suffix -osus, indicating abundance or full or marked development).

Type

Spain, Catalonia, Tarragona province, La Fatarella, from goat dung, Mar. 2017, I. Iturrieta-González, M. Guevara-Suarez & J. Guarro (holotype CBS H-24469; cultures ex-type FMR 16532, CBS 146918).

Description

Mycelium in vitro superficial and immersed, composed of septate, branched, subhyaline, smooth to verruculose hyphae, 1–2 μm wide. Conidiophores dimorphic, micronematous or macronematous, arising from lateral or terminal hyphae, erect to slightly flexuous, non-nodulose, septate, branched or unbranched, 8–137 μm long, 2–4 μm wide, pale brown, slightly verrucose. Conidiogenous cells integrated, terminal, cylindrical, sometimes geniculate at the apex, 22–44 × 3–4 μm, bearing up to four conidiogenous loci, darkened and refractive. Ramoconidia aseptate, almost cylindrical, 10–24 × 2–4 μm [av. (± SD) 15.8 (± 3.4) × 3.1 (± 0.45)], olive to pale brown, smooth. Conidia forming branched chains, with up to five conidia in the terminal unbranched part, aseptate, olive to pale brown, smooth; small terminal conidia ellipsoidal to obovoid, 3–7 × 2–3.5 μm [av. (± SD) 5.7 (± 0.83) × 2.4 (± 0.43)]; intercalary conidia ellipsoidal to somewhat fusiform, 6–11.5 × 2–3 μm [av. (± SD) 7.8 (± 1.06) × 2.6 (± 0.39)]; secondary ramoconidia ellipsoidal to almost cylindrical, 9–14 × 2.5–3.5 μm [av. (± SD) 11.3 (± 1.6) × 2.9 (± 0.26)].

Culture characteristics

(14 d at 25 °C). Colonies on OA reaching 24–25 mm diam., dark green (30F8), flat, slightly dusty, aerial mycelium scarce, margin regular; reverse dark green (30F8) to black. On PDA attaining 34–35 mm diam., olive (3E6/3F4), slightly umbonate, radially folded, velvety, aerial mycelium scarce, margin slightly lobate; reverse dark green (30F8) to olive (3E4). On SNA reaching 25–26 mm diam., olive (3E8), flat, dusty, aerial mycelium scarce, margin regular; reverse dark green (30F8) to black.

Cardinal temperature for growth

Optimum 20 °C, maximum 30 °C, minimum 5 °C.

Distribution

Spain.

Notes

Although C. caprifimosum clearly belongs to the C. cladosporioides species complex, our multi-locus analysis did not reveal any phylogenetic relationship with other species in the complex. It is represented by a single branch placed distance from other Cladosporium species (Fig. 1). Cladosporium caprifimosum differs from the other novel species proposed here mainly by its aseptate and smooth conidia.

Figure 3.

Cladosporium caprifimosum (ex-type FMR 16532) a–c colonies on PDA, OA and SNA after 14 days at 25 °C d–e conidiophores f ramoconidia and conidia. Scale bars: 10 mm (a–c); 10 μm (d–f).

Cladosporium coprophilum Iturrieta-González, Dania García, Gené, sp. nov.

MycoBank No: 836075

Fig. 4

Etymology

Name refers to the substrate where the species was isolated, unidentified herbivore dung (ancient Greek, kópros = dung + phílos = loving).

Type

Spain, Extremadura, Badajoz province, Granja de Torrehermosa, unidentified herbivore dung, Jan. 2017, J. Cano (holotype CBS H-24470; cultures ex-type FMR 16164, CBS 144919).

Description

Mycelium in vitro superficial and immersed, composed of septate, branched, pale brown, smooth hyphae, 3–5 μm wide. Conidiophores macronematous, arising laterally or terminally from hyphae, erect to slightly flexuous, non-nodulose, septate, unbranched, up to 124 μm long, 3–4 μm wide, pale brown, smooth. Conidiogenous cells integrated, terminal, rarely intercalary, cylindrical, (7–)14–33 × (2–)3–4 μm, bearing up to 3 conidiogenous loci, slightly darkened and refractive. Ramoconidia 0(–1)-septate, subcylindrical to cylindrical, 9–19 × 3–5 μm [av. (± SD) 12.3 (± 2.8) × 3.9 (± 0.54)], pale brown, smooth. Conidia forming branched chains, with up to five conidia in the terminal unbranched part, aseptate, pale brown, smooth to verruculose; small terminal conidia ellipsoidal to slightly obovoid, 4.5–7 × 2.5–4 μm [av. (± SD) 6 (± 0.64) × 3.1 (± 0.31)]; intercalary conidia ellipsoidal, 6–10.5 × 2.5–4 μm [av. (± SD) 7.7 (± 1.32) × 3.3 (± 0.37)]; secondary ramoconidia subcylindrical to cylindrical, 7–12.5 μm long × 3–5 μm [av. (± SD) 9.6 (± 1.7) × 4.2 (± 0.51)].

Culture characteristics

(14 d at 25 °C). Colonies on OA reaching 21–22 mm diam., olive (2F6) to black, dark green margin (30F4), flat, slightly dusty at the center, aerial mycelium scarce, margin regular; reverse dark green (30F8) to black. On PDA attaining 36–37 mm diam., olive (2F6/2E3), greenish gray margin, slightly depressed and irregularly folded at the center, velvety, aerial mycelium scarce, margin regular; reverse dark green (30F8/27F3). On SNA reaching 27–28 mm diam., olive (3F6/2F8), flat, slightly dusty, aerial mycelium scarce, margin regular; reverse dark green (30F8) to black.

Cardinal temperature for growth

Optimum 20 °C, maximum 25 °C, minimum 5 °C.

Distribution

Spain.

Additional specimen examined

Spain, Extremadura, Badajoz province, Granja de Torrehermosa, unidentified herbivore dung, Mar. 2017, J. Cano (FMR 16101).

Notes

Based on the multi-locus analysis (Fig. 1), C. coprophilum is allocated to a terminal low-supported clade together with C. chasmanthicola and C. sinuatum, species recently described from leaf spots of Chasmanthe aethiopica in South Africa (Marin-Felix et al. 2017) and Alpine soil in China (Ma et al. 2017), respectively. The new species is distinguished from C. chasmanthicola by the production of smooth hyphae (smooth to distinctly verrucose or irregularly rough-walled in C. chasmanthicola), longer conidiogenous cells (up to 33 vs up to 24 μm), shorter ramoconidia (9–19 vs 15–33 μm) with fewer septa [(0(–1) vs 0–1(–3)-septate], and longer terminal conidia (4.5–7 vs 2.5–4.5 μm) (Marin-Felix et al. 2017). Cladosporium coprophilum differs from C. sinuatum by the production of aseptate intercalary conidia (0–1-septate in C. sinuatum). In addition, C. sinuatum is characterized by distinctive geniculate-sinuous conidiophores and a rather fast growth on OA (40–45 mm vs 21–22 mm in C. coprophilum after 14 d at 25 °C) (Ma et al. 2017).

Figure 4.

Cladosporium coprophilum (ex-type FMR 16164) a–c colonies on PDA, OA and SNA after 14 days at 25 °C d–e conidiophores f conidia. Scale bars: 10 mm (a–c); 10 μm (d–f).

Cladosporium fuscoviride Iturrieta-González, Dania García, Gené, sp. nov.

MycoBank No: 836076

Fig. 5

Etymology

Name refers to the dark green reverse of the colonies of the species growing in all agar media tested (fuscus = dark brown, blackish or figuratively dull and viridis = green).

Type

Spain, Catalonia, Tarragona province, Cambrils, Samà Park, garden soil, Feb. 2017, I. Iturrieta-González & J. Gené (holotype CBS H-24471; cultures ex-type FMR 16385, CBS 146920).

Description

Mycelium in vitro superficial and immersed, composed of septate, branched, subhyaline to pale brown, smooth to verruculose hyphae, 1–3 μm wide. Conidiophores semi-macronematous to macronematous, arising laterally and terminally from hyphae, sometimes reduced to conidiogenous cells, septate, erect to slightly flexuous, branched or unbranched, sometimes geniculate at the apex, up to 56 μm long, 3–4 μm wide, pale brown, smooth to verruculose. Conidiogenous cells terminal and subterminal, cylindrical to slightly clavate, 8–27 × 3–4 μm, bearing up to 4 conidiogenous loci, darkened and refractive. Ramoconidia 0–1(–3)-septate, subcylindrical to ellipsoidal, 7.5–22 × 2.5–4 μm [av. (± SD) 12.8 (± 3.9) × 3 (± 0.43)], pale brown, smooth to verruculose. Conidia in branched chains with up to 4 conidia in the terminal unbranched part, pale brown, smooth to verruculose, with protuberant, slightly darkened and refractive hila; small terminal conidia aseptate, globose, subglobose to obovoid, 3–6 × 2–3.5 μm [av. (± SD) 4.5 (± 0.66) × 3 (± 0.39)]; intercalary conidia aseptate, ellipsoidal to somewhat limoniform, 4.5–7 × 2.5–4 μm [av. (± SD) 5.7 (± 0.70) × 3.2 (± 0.36)]; secondary ramoconidia 0(–1)-septate, subcylindrical to ellipsoidal 6–11.5 × 2.5–4 μm [av. (± SD) 8.8 (± 1.64) × 3.1 (± 0.40)].

Culture characteristics

(14 d at 25 °C). Colonies on OA reaching 31–32 mm diam., olive (3F8) to dark green (30F5), olive final edge (2F8), flat, velvety, aerial mycelium scarce, margin regular; reverse dark green (30F5) to black. On PDA attaining 44–46 mm diam., gray to olive to olive yellow (3D1/2E5/2C6), white at the final edge, flat, velvety, aerial mycelium scarce, margin regular; reverse dark green (30F8) to black, with a whitish final edge. On SNA reaching 34–35 mm diam., olive (3F8), flat, velvety, aerial mycelium scarce, margin regular; reverse dark green (30F8), olive final edge (3F3).

Cardinal temperature for growth

Optimum 25 °C, maximum 30 °C, minimum 5 °C.

Distribution

Spain.

Notes

Cladosporium fuscoviride is closely related to C. alboflavescens (Fig. 1), a monotypic species described from an animal respiratory specimen collected in California (Sandoval-Denis et al. 2016). The species can be distinguished by their colony and microscopic features; i.e., C. fuscoviride has darker colonies and faster growth rates at 25 °C after 2 wk on the three media tested (OA, 31–32 vs 20–23 mm; PDA, 44–46 vs 34–36 mm; SNA, 34–35 vs 20–25 mm), shorter conidiophores (up to 56 μm vs up to 130 μm long in C. alboflavescens), and 0–3-septate (aseptate in C. alboflavescens) shorter (7.5–22 vs 11–36 μm) ramoconidia. Cladosporium iranicum is related with C. fuscoviride and C. alboflavescens, but can be easily distinguished from them by its larger conidiophores (40–180(–135) μm), with chains of up to 10 conidia in the terminal unbranched part, and a faster growth rate on PDA (56–60 mm after 14 d at 25 °C) (Bensch et al. 2010).

Figure 5.

Cladosporium fuscoviride (ex-type FMR 16385) a–c colonies on PDA, OA and SNA after 14 days at 25 °C d–g conidiophores h ramoconidia and conidia. Scale bars: 10 mm (a–c); 10 μm (d–h).

Cladosporium lentulum Iturrieta-González, Dania García, Gené, sp. nov.

MycoBank No: 836077

Fig. 6

Etymology

Name refers to its slower growth with respect to the phylogenetically related species (lentus = figuratively slow, with Latin adjectival suffix -ulus = diminutive).

Type

Spain, Catalonia, Tarragona province, Tarragona, unidentified leaf litter, Feb. 2017, I. Iturrieta-González (holotype CBS H-24472; cultures ex-type FMR 16288, CBS 146921).

Description

Mycelium in vitro superficial and immersed, composed of septate, branched, subhyaline to yellowish brown, smooth to verruculose hyphae, 1–4 μm wide. Conidiophores macronematous, arising laterally and terminally from hyphae, septate, erect to slightly flexuous, unbranched, sometimes geniculate at the apex, occasionally branched, up to 406 μm long, 3–4 μm wide, pale brown to brown, smooth to verrucose. Conidiogenous cells integrated, terminal and subterminal, cylindrical to subcylindrical, 11–27 × 2–4(–5) μm, bearing up to 5 conidiogenous loci, darkened and refractive. Ramoconidia 0(–2)-septate, subcylindrical to cylindrical, 10.5–23 × 2.5–4.5 μm [av. (± SD) 14.2 (± 2.61) × 3.2 (± 0.52)]; pale brown, smooth to verruculose. Conidia forming branched chains with up to 5 conidia in the unbranched part of the chain, pale brown, smooth to slightly verruculose, with protuberant, slightly darkened and refractive hila; small terminal conidia aseptate obovoidal to ellipsoidal, 4.5–7.5 × 1.5–2.5 μm [av. (± SD) 5.8 (± 0.81)) × 2.7 (± 0.29)]; intercalary conidia 0(–1)-septate, ellipsoidal to subcylindrical, 6–10.5 × 2–3 μm [av. (± SD) 8.4 (± 1.31) × 2.3 (± 0.34)]; secondary ramoconidia 0(–1)-septate, ellipsoidal to subcylindrical, slightly constricted at septum when present, 7.5–14.5 × 2–3 μm [av. (± SD) 10.5 (± 2.05) × 2.5 (± 0.30)].

Culture characteristics

(14 d at 25 °C). Colonies on OA reaching 19–20 mm diam., olive (3F8), flat, velvety, aerial mycelium scarce, margin regular; reverse dark green (30F8) to black. On PDA attaining 28–36 mm diam., dark green (27F8), with a whitish final edge, slightly umbonate, radially folded, velvety, aerial mycelium scarce, margin slightly lobulate; reverse olive brown (4E4), whitish at the edge. On SNA reaching 22–23 mm diam., olive (3F5), flat, slightly dusty, aerial mycelium scarce, margin fimbriate; reverse dark green (30F8) to black.

Cardinal temperature for growth

optimum 20 °C, maximum 30 °C, minimum 5 °C.

Distribution

Spain.

Additional specimen examined

Spain, Catalonia, Tarragona province, Poblet, unidentified herbivore dung, Mar. 2017, I. Iturrieta-González, M. Guevara-Suarez & J. Guarro (FMR 16389).

Notes

Our phylogeny shows C. lentulum included in a well-supported terminal clade together with the ex-type strains of C. exasperatum, C. parapenidielloides and C. longicatenatum, three species all described from plant material collected in Australia (Bensch et al. 2010, 2015). However, the genetic distance allows it to be considered a distinct species within the clade (Fig. 1). Phenotypically, C. lentulum can be distinguished from its counterparts mainly by its slower growth, especially on OA at 25 °C after 14 d (19–20 mm vs 39–54 mm for C. exasperatum, 42–55 mm for C. parapenidielloides and 43–54 mm for C. longicatenatum). In addition, our new species shows shorter ramoconidia (10.5–23 μm) than C. exasperatum and C. longicatenatum (19–40 μm and 22–42 μm, respectively); ramoconidia in C. parapenidielloides are absent; the conidia in C. lentulum are smooth or nearly so, while those of C. exasperatum and C. longicatenatum possess a unique verruculose-rugose conidial surface ornamentation, especially prominent in the former; and conidiophores in C. parapenidielloides are much shorter (up to 67 μm) than those observed in C. lentulum (up to 406 μm) (Bensch et al. 2010, 2015).

Figure 6.

Cladosporium lentulum (ex-type FMR 16288) a–c colonies on PDA, OA and SNA after 14 days at 25 °C d–e conidiophores f–g conidia. Scale bars: 10 mm (a–c); 10 μm (d–g).

Cladosporium pseudotenellum Iturrieta-González, Dania García, Gené, sp. nov.

MycoBank No: 836078

Fig. 7

Etymology

The name refers to “C. tenellum”, the closest phylogenetic species.

Type

Spain, Catalonia, Tarragona province, Reus, garden soil, Feb. 2017, I. Iturrieta-González (holotype CBS H-24473; cultures ex-type FMR 16231, CBS 146922).

Description

Mycelium in vitro superficial and immersed, composed of septate, branched, subhyaline to pale brown, smooth-walled, occasionally tuberculate and with abundant swellings, hyphae, 2–3(–4.5) μm wide. Conidiophores macronematous, arising laterally or terminally from hyphae, erect to slightly flexuous, non-nodulose, occasionally geniculate at the apex, septate, unbranched, occasionally branched, up to 146 μm long, 2.5–3 μm wide, pale brown, smooth to slightly verruculose. Conidiogenous cells integrated, terminal or intercalary, cylindrical, sometimes geniculate, 15–32 × 2.5–3 μm, with up to five conidiogenous loci, thickened, darkened and refractive, often crowded at the apex. Ramoconidia rarely formed, 0(–1)-septate, ellipsoidal to subcylindrical, 9–14.5 × 4–5.5 μm [av. (± SD) 11.6 (± 1.60) × 4.6 (± 0.44)], pale brown, verruculose. Conidia forming branched chains, with up to four conidia in the terminal unbranched part, aseptate, pale brown, verruculose to verrucose; small terminal conidia subglobose to obovoid, 4–7 × 3–5 μm [av. (± SD) 5.8 (± 0.77) × 3.9 (± 0.60)]; intercalary conidia ellipsoidal to limoniform, 6–8.5 × 3–5 μm [av. (± SD) 7.4 (± 0.73) × 3.8 (± 0.50)]; secondary ramoconidia 0(–2)-septate, ellipsoidal to subcylindrical, 7–12.5 × 4–5 μm [av. (± SD) 9.6 (± 1.76) × 4.4 (± 0.33)] with 1–3 distal hila.

Culture characteristics

(14 d at 25 °C). Colonies on OA reaching 21–22 mm diam., olive (2F8/2F4), flat, velvety, aerial mycelium scarce, margin fimbriate; reverse dark green (30F8) to black. On PDA attaining 29–30 mm diam., olive gray (3E2/3F2), paler at the periphery, radially folded, velvety, aerial mycelium scarce, margin slightly lobate; reverse dark green (30F8) to black. On SNA reaching 21–22 mm diam., olive (2F8), flat, slightly powdery, aerial mycelium scarce, margin fimbriate; reverse dark green (30F8) to black.

Cardinal temperature for growth

Optimum 20 °C, maximum 30 °C, minimum 5 °C.

Distribution

Spain.

Notes

Based on the phylogeny of the C. herbarum complex (Fig. 2), C. pseudotenellum is closely related with C. tenellum, a species originally described from hypersaline water in Israel, later found on Phyllactinia sp. (Erysiphaceae), and in indoor air samples collected in the USA (Schubert et al. 2007; Bensch et al. 2012, 2018). Our species differs from C. tenellum in the absence of micronematous conidiophores and in having shorter macronematous conidiophores (up to 146 μm vs up to 200 μm), shorter conidiogenous cells (15–32 μm vs 6–40 μm), with few conidiogenous loci (up to five vs up to 10 or more in C. tenellum), and shorter ramoconidia (9–14.5 vs up to 32 μm). In addition, terminal and intercalary conidia in C. pseudotenellum are aseptate, while those of C. tenellum are 0–1(–3)-septate (Schubert et al. 2007; Bensch et al. 2012).

Figure 7.

Cladosporium pseudotenellum (ex-type FMR 16231) a–c colonies on PDA, OA and SNA after 14 days at 25 °C d–e conidiophores f conidia. Scale bars: 10 mm (a–c); 10 μm (d–f).

Cladosporium submersum Iturrieta-González, Dania García, Gené, sp. nov.

MycoBank No: 836079

Fig. 8

Etymology

Name refers to the aquatic habitat where the substrate (submerged plant material) of the fungus was collected (submersus = submerged, verb in participle, from submergere).

Type

Spain, Catalonia, Tarragona province, Cornudella del Montsant, Siurana’s swamp, submerged plant material, Feb. 2018, I. Iturrieta-González, E. Carvalho & J. Gené (holotype CBS H-24474; cultures ex-type FMR 17264, CBS 146923).

Description

Mycelium in vitro superficial and immersed, composed of septate, branched, subhyaline to pale brown, smooth-walled to verruculose hyphae, 1–3 μm wide. Conidiophores dimorphic, micronematous or macronematous, arising laterally and terminally from hyphae, erect to slightly flexuous, nodulose, geniculate at the apex, septate, unbranched, occasionally branched with small prolongations just below the septum, up to 77 μm long, 3–5 μm wide, pale brown to brown, smooth to verruculose. Conidiogenous cells integrated, terminal and intercalary, geniculate, nodulose, 11–28 × 3–6 μm, bearing up to five conidiogenous loci, darkened and refractive. Ramoconidia rarely formed, 0(–1)-septate, sometimes constricted at the septum when present, cylindrical to subcylindrical, 10.5–24 × 4.5–7 μm [av. (± SD) 16 (± 3.6) × 6.1 (± 1.03)], pale brown, verruculose to verrucose. Conidia forming short branched chains, pale brown, verrucose, occasionally verruculose, with protuberant and slightly darkened hila; small terminal conidia aseptate, ovoid to ellipsoidal, 6–12.5 × 3.5–7 μm [av. (± SD) 7.8 (± 1.63) × 4.8 (± 0.79)]; intercalary conidia and secondary ramoconidia 0–1-septate, ellipsoidal or subcylindrical, 7.5–16 × 4.5–8 μm [av. (± SD) 11 (± 2.18) × 5.7 (± 0.99)].

Culture characteristics

(14 d at 25 °C). Colonies on OA reaching 22–23 mm diam., brownish gray to olive brown (4E2/4E4), umbonate, velvety, aerial mycelium scarce, margin slightly irregular and fimbriate; reverse dark green to olive brown (6F8/4E3). On PDA attaining 26–28 mm diam., olive (3F3/1F5), slightly umbonate, radially folded, velvety, aerial mycelium scarce, margin irregularly undulate; reverse dark green (30F9) to black with brownish red (9C6) areas observed between 15 and 20 °C and a white edge. On SNA reaching 21–22 mm diam., olive (3E3), slightly umbonate, loosely cottony, margin fimbriate; reverse dark olive brown to golden gray (3E3/4C2).

Cardinal temperature for growth

Optimum 20 °C, maximum 35 °C, minimum 5 °C.

Distribution

Spain.

Notes

Cladosporium submersum is related to C. subcinereum, and morphologically differentiated by having shorter conidiophores (up to 77 μm vs up to 140 μm), shorter conidiogenous cells (11–28 vs 16–38 μm), shorter ramoconidia (10.5–24 vs 19–59 μm), and longer terminal conidia (6–12.5 vs 5–7 μm), which are ovoid to ellipsoidal in our species and globose to subglobose in C. subcinereum (Sandoval-Denis et al. 2016). In addition, C. submersum exhibited a colony reverse on PDA with brownish red areas, a feature that is absent in C. subcinereum.

Figure 8.

Cladosporium submersum (ex-type FMR 16264) a–c colonies on PDA (front at 25 °C and reverse at 20 °C), OA and SNA at 25 °C after 14 days d, e conidiophores and conidia f conidia. Scale bars: 10 mm (a–c); 10 μm (d–f).

Discussion

Cladosporium is a well-delineated genus, the taxonomic structure and phylogenetic relationships of its species have been investigated in several studies over the last decade, so far giving rise to a genus of more than two hundred well-established species (Zalar et al. 2007; Bensch et al. 2010, 2012, 2018; Sandoval-Denis et al. 2016; Marin-Felix et al. 2017; Crous et al. 2009, 2019; Jayasiri et al. 2019). However, this species number will continue to expand through the study of soil, which is a proven pool of fungal species that remains undescribed, and other substrates poorly investigated by molecular tools for fungal diversity (Tedersoo et al. 2017; Hyde et al. 2018). In this context, a set of Cladosporium isolates were obtained in pure culture from samples of soil, dung from different herbivorous animals, and plant debris collected during a survey of microfungi in various Spanish locations. Using the molecular approach for species delineation in Cladosporium (Bensch et al. 2012; Marin-Felix et al. 2017), eight of those isolates represented six novel lineages for the genus which are proposed as C. caprifimosum, C. coprophilum, C. fuscoviride, C. lentulum, C. pseudotenellum and C. submersum. Of note is that almost all the specimens in the present study (7/8) were isolated directly from the natural substratum incubated in moist chambers or from baiting technique plates. Although Cladosporium isolates are commonly detected by plating methods, the slow growth rate or the low spore concentration of some cladosporium-like fungi compared to other fungi present in a given substrate is probably a handicap to detection and/or isolation of uncommon Cladosporium species. Therefore, as recommended by Crous (1998) for similar fungi, techniques based on fungal isolation directly from the natural substratum should be considered a choice for future studies of Cladosporium species diversity.

To our knowledge, Cladosporium species as dung inhabiting fungi have been reported in a very few studies, C. cladosporioides and C. herbarum being the most reported species (Bell 1975; Seifert et al. 1983; Jeamjitt et al. 2006; Masunga et al. 2006; Piontelli et al. 2006; Simões-Calaça et al. 2014; Thilagam et al. 2015). However, in all those studies, fungal identification was based exclusively on morphological features. Only C. herbarum has been reported recently from crown droppings and identified molecularly, but using only the ITS barcode (Torbati et al. 2016). In our case, the three new species isolated on herbivore dung (i.e., C. caprifimosum, C. coprophilum, and C. lentulum) showed the typical morphological features attributed to the C. cladosporioides species complex. However, their identifications would have been difficult with morphological features alone, even with the analysis of their ITS sequences (Table 2) since they are identical under the universal barcode for fungi as reported in previous studies for many other Cladosporium species (Bensch et al. 2010, 2012; Marin-Felix et al. 2017). Therefore, only sequence analysis with act and tef1 will allow us to know the real diversity of Cladosporium species from this understudied substrate by molecular tools.

Although no temperature studies have been systematically applied to characterize most Cladosporium species (Bensch et al. 2012, 2015, 2018), we agree with Ma et al. (2017) that cardinal temperatures for growth can help to differentiate certain species in their respective complexes. While species in the C. sphaerospermum complex show a maximum temperature for growth of no more than 30–32 °C, C. halotolerans was able to grow at 35 °C (Sandoval-Denis et al. 2015). Similarly, although most species of the C. cladosporioides complex do not tolerate high temperatures, C. angulosum, C. angustisporum, C. anthropophilum, C. flavovirens, C. funiculosum, C. pseudocladosporioides, C. subuliforme and C. tenuissimum were able to grow at 35 °C (Sandoval-Denis et al. 2015, 2016). To date, no member of the C. herbarum complex was found to be able to grow above 30 °C; however, one of the novel species of the complex described here, C. submersum, had a maximum growth at 35 °C. On the contrary, the recently described species C. neopsychrotolerans and C. tianshanense from the complex C. cladosporioides and C. psychrotolerans from the complex C. sphaerospermum showed a psychrophilic behavior (Zalar et al. 2007; Ma et al. 2017), demonstrating in part the ability of Cladosporium species to adapt to different environmental conditions.

Acknowledgements

The authors are grateful to Lei Cai of Institute of Microbiology of the Chinese Academy of Sciences (Beijing, China) for sending the sequences of C. tianshanense included in this study. This study was supported in part by the Spanish Ministerio de Economía y Competitividad, grant CGL2017-88094-P.

References

Abdollahzadeh J, Groenewald JZ, Coetzee MPA, Wingfield MJ, Crous PW (2020) Evolution of lifestyles in Capnodiales. Studies in Mycology 95: 381–414. https://doi.org/10.1016/j.simyco.2020.02.004

Adorisio S, Fierabracci A, Muscari I, Liberati AM, Cannarile L, Thuy TT, Sung TV, Sohrab H, Hasan CM, Ayroldi E, Riccardi C, Mazid A, Delfino DV (2019) Fusarubin and Anhydrofusarubin isolated from a Cladosporium species inhibit cell growth in human cancer cell lines. Toxins 11(9): e503. https://doi.org/10.3390/toxins11090503

Bell A (1975) Fungal succession on dung of the brush-tailed opossum in New Zealand. New Zealand Journal of Botany 13: 437–462. https://doi.org/10.1080/0028825X.1975.10430336

Bensch K, Groenewald JZ, Dijksterhuis J, Starink-Willemse M, Andersen B, Starink-Willemse M, Andersen B, Summerell BA, Shin H-D, Dugan FM, Schroers H-J, Braun U, Crous PW (2010) Species and ecological diversity within the Cladosporium cladosporioides complex (Davidiellaceae, Capnodiales). Studies in Mycology 67: 1–94. https://doi.org/10.3114/sim.2010.67.01

Bensch K, Braun U, Groenewald JZ, Crous PW (2012) The genus Cladosporium. Studies in Mycology 72: 1–401. https://doi.org/10.3114/sim0003

Bensch K, Groenewald JZ, Braun U, Dijksterhuis J, Yañez-Morales M, Dijksterhuis J, Yáñez-Morales MJ, Crous PW (2015) Common but different: The expanding realm of Cladosporium. Studies in Mycology 82: 23–74. https://doi.org/10.1016/j.simyco.2015.10.001

Bensch K, Groenewald JZ, Meijer M, Dijksterhuis J, Jurjevic Ž, Dijksterhuis J, Jurjević Z, Andersen B, Houbraken J, Crous PW, Samson RA (2018) Cladosporium species in indoor environments. Studies in Mycology 89: 177–301. https://doi.org/10.1016/j.simyco.2018.03.002

Bezerra JDP, Sandoval-Denis M, Paiva LM, Silva GA, Groenewald JZ, Silva GA, Groenewald JZ, Souza-Motta CM, Crous PW (2017) New endophytic Toxicocladosporium species from cacti in Brazil, and description of Neocladosporium gen. nov. IMA Fungus 8: 77–97. https://doi.org/10.5598/imafungus.2017.08.01.06

Calduch M, Gené J, Stchigel AM, Cano JF, Guarro J (2004) Ramophialophora, a new anamorphic genus of Sordariales. Studies in Mycology 50: 83–88.

Carbone I, Kohn LM (1999) A method for designing primer sets for speciation studies in filamentous ascomycetes. Mycologia 91: 553–556. https://doi.org/10.1080/00275514.1999.12061051

Castañeda-Ruiz RF, Heredia G, Gusmao LFP, Li DW (2016) Fungal diversity of central and south America. In: De-Wei L (Ed.) Biology of Microfungi. Springer International Publishing, Switzerland, 197–218. https://doi.org/10.1007/978-3-319-29137-6_9

Chung D, Kim H, Choi HS (2019) Fungi in salterns. Journal of Microbiology 57: 717–724. https://doi.org/10.1007/s12275-019-9195-3

Crous PW (1998) Mycosphaerella spp. and their anamorphs associated with leaf spot diseases of Eucalyptus. Mycologia Memoir 21: 1–170.

Crous PW, Gams W, Stalpers JÁ, Robert V, Stegehuis G (2004) MycoBank: an online initiative to launch mycology into the 21^st century. Studies in Mycology 50: 19–22.

Crous PW, Braun U, Schubert K, Groenewald JZ (2007) Delimiting Cladosporium from morphologically similar genera. Studies in Mycology 58: 33–56. https://doi.org/10.3114/sim.2007.58.02

Crous PW, Verkley GJM, Groenewald JZ, Samson RA (2009) Fungal Biodiversity. CBS Laboratory manual Series. Westerdijk Fungal Biodiversity Institute, Utrecht, The Netherlands, 269 pp.

Crous PW, Wingfield MJ, Lombard L, Roets F, Swart WJ, Roets F, Swart WJ, Alvarado P, Carnegie AJ, Moreno G, Luangsa-Ard J, Thangavel R, Alexandrova AV, Baseia IG, Bellanger J-M, Bessette AE, Bessette AR, Delapeña-Lastra S, García D, Gené J, Pham THG, Heykoop M, Malysheva E, Malysheva V, Martín MP, Morozova OV, Noisripoom W, Overton BE, Rea AE, Sewall BJ, Smith ME, Smyth CW, Tasanathai K, Visagie CM, Adamčík S, Alves A, Andrade JP, Aninat MJ, Araújo RVB, Bordallo JJ, Boufleur T, Baroncelli R, Barreto RW, Bolin J, Cabero J, Caboň M, Cafà G, Caffot MLH, Cai L, Carlavilla JR, Chávez R, Decastro RRL, Delgat L, Deschuyteneer D, Dios MM, Domínguez LS, Evans HC, Eyssartier G, Ferreira BW, Figueiredo CN, Liu F, Fournier J, Galli-Terasawa LV, Gil-Durán C, Glienke C, Gonçalves MFM, Gryta H, Guarro J, Himaman W, Hywel-Jones N, Iturrieta-González I, Ivanushkina NE, Jargeat P, Khalid AN, Khan J, Kiran M, Kiss L, Kochkina GA, Kolařík M, Kubátová A, Lodge DJ, Loizides M, Luque D, Manjón JL, Marbach PAS, Massolajr NS, Mata M, Miller AN, Mongkolsamrit S, Moreau P-A, Morte A, Mujic A, Navarro-Ródenas A, Németh MZ, Nóbrega TF, Nováková A, Olariaga I, Ozerskaya SM, Palma MA, Petters-Vandresen DAL, Piontelli E, Popov ES, Rodríguez A, Requejo Ó, Rodrigues ACM, Rong IH, Roux J, Seifert KA, Silva BDB, Sklenář F, Smith JA, Sousa JO, Souza HG, Desouza JT, Švec K, Tanchaud P, Tanney JB, Terasawa F, Thanakitpipattana D, Torres-Garcia D, Vaca I, Vaghefi N, Vaniperen AL, Vasilenko OV, Verbeken A, Yilmaz N, Zamora JC, Zapata M, Jurjevi Ž, Groenewald JZ (2019) Fungal Planet description sheets: 951–1041. Persoonia 43: 223–425. https://doi.org/10.3767/persoonia.2019.43.06

David JC (1997) A contribution to the systematics of Cladosporium. Revision of the fungi previously referred to Heterosporium. Mycological Papers 172: 1–157.

de Hoog GS, Guarro J, Gené J, Figueras MJ (2017) Atlas of clinical fungi. Electronic version 4.1.4. Westerdijk Fungal Biodiversity Institute / Universitat Rovira i Virgili, Utrecht / Reus. http://www.clinicalfungi.org/

Edgar RC (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research 32: 1792–1797. https://doi.org/10.1093/nar/gkh340

Hespanhol L, Vallio CS, Costa LM, Saragiotto BT (2019) Understanding and interpreting confidence and credible intervals around effect estimates. Brazilian Journal of Physical Therapy 23(4): 290–301. https://doi.org/10.1016/j.bjpt.2018.12.006

Heuchert B, Braun U, Schubert K (2005) Morphotaxonomic revision of fungicolous Cladosporium species (Hyphomycetes). Schlechtendalia 13: 1–78.

Hillis DM, Bull JJ (1993) An empirical test of bootstrapping as a method for assessing confidence in phylogenetic analysis. Systematic Biology 42: 182–192. https://doi.org/10.1093/sysbio/42.2.182

Farris JS, Källersjö M, Kluge AG, Bult C (1994) Testing significance of incongruence. Cladistics 10: 315–319. https://doi.org/10.1111/j.1096-0031.1994.tb00181.x

Flannigan B, Samson RA, Miller JD (2002) Microorganisms in home and indoor work environments: diversity, health impacts, Investigation and control. London, United Kingdom, 504 pp. https://doi.org/10.1201/9780203302934

Hyde KD, Norphanphoun C, Chen J, Dissanayake AJ, Doilom M, Dissanayake AJ, Doilom M, Hongsanan S, Jayawardena RS, Jeewon R, Perera RH, Thongbai B, Wanasinghe DN, Wisitrassameewong K, Tibpromma S, Stadler M (2018) Thailand’s amazing diversity: up to 96% of fungi in northern Thailand may be novel. Fungal Diversity 93: 215–239. https://doi.org/10.1007/s13225-018-0415-7

Jayasiri SC, Hyde KD, Jones EBG, McKenzie EHC, Jeewon R, McKenzie EHC, Jeewon R, Phillips AJL, Bhat DJ, Wanasinghe DN2, Liu JK, Lu YZ, Kang JC, Xu J, Karunarathna SC (2019) Diversity, morphology and molecular phylogeny of Dothideomycetes on decaying wild seed pods and fruits. Mycosphere 10: 1–186. https://doi.org/10.5943/mycosphere/10/1/1

Jeamjitt O, Manoch L, Visarathanonth N, Chamswarng C (2006) Diversity and distribution of hyphomycetes from dung in Thailand. Witthayasan Kasetsat 40: 890–901.

Khan MIH, Sohrab MH, Rony SR, Tareq FS, Hasan CM, Mazid MA (2016) Cytotoxic and antibacterial naphthoquinones from an endophytic fungus, Cladosporium sp. Toxicology Reports 3: 861–865. https://doi.org/10.1016/j.toxrep.2016.10.005

Köhl J, Scheer C, Holb IJ, Masny S, Molhoek W (2015) Toward an integrated use of biological control by Cladosporium cladosporioides H39 in apple scab (Venturia inaequalis) management. Plant Disease 99: 535–543. https://doi.org/10.1094/PDIS-08-14-0836-RE

Kornerup A, Wanscher JH (1978) Methuen Handbook of Colour (3^rd ed.). Methuen Publishing Ltd., London, 256 pp.

Ma R, Chen Q, Fan Y, Wang Q, Chen S, Wang Q, Chen S, Liu X, Cai L, Yao B (2017) Six new soil-inhabiting Cladosporium species from plateaus in China. Mycologia 109: 244–260. https://doi.org/10.1080/00275514.2017.1302254

Marin-Felix Y, Groenewald JZ, Cai L, Chen Q, Marincowitz S, Chen Q, Marincowitz S, Barnes I, Bensch K, Braun U, Camporesi E, Damm U, de Beer ZW, Dissanayake A, Edwards J, Giraldo A, Hernández-Restrepo M, Hyde KD, Jayawardena RS, Lombard L, Crous PW (2017) Genera of phytopathogenic fungi: GOPHY 1. Studies in Mycology 86: 99–216. https://doi.org/10.1016/j.simyco.2017.04.002

Masunga GS, Andersen Ø, Taylor JE, Dhillion SS (2006) Elephant dung decomposition and coprophilous fungi in two habitats of semi-arid Botswana. Mycological Research 110: 1214–1226. https://doi.org/10.1016/j.mycres.2006.07.004

Müller FM, Werner KE, Kasai M, Francesconi A, Chanock SJ, Francesconi A, Chanock SJ, Walsh TJ (1998) Rapid extraction of genomic DNA from medically important yeasts and filamentous fungi by high-speed cell disruption. Journal of Clinical Microbiology 36: 1625–1629. https://doi.org/10.1128/JCM.36.6.1625-1629.1998

Piontelli LE, Cruz CR, Toro SMMA (2006) Coprophilous fungal community of wild rabbit in a park of a hospital (Chile): a taxonomic approach. Boletín Micológico 21: 1–17. https://doi.org/10.22370/bolmicol.2006.21.0.239

Posada D (2008) jModelTest: phylogenetic Model Averaging. Molecular Biology and Evolution 25: 1253–1256. https://doi.org/10.1093/molbev/msn083

Richardson MJ (2001) Diversity and occurrence of coprophilous fungi. Mycological Research 105: 387–402. https://doi.org/10.1017/S0953756201003884

Ronquist F, Teslenko M, van der Mark P, Ayres DL, Darling A, Ayres DL, Darling S, Höhna S, Larget B, Liu L, Suchard MA, Huelsenbeck JP (2012) MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Systematic Biology 61: 539–542. https://doi.org/10.1093/sysbio/sys029

Sandoval-Denis M, Gené J, Sutton DA, Wiederhold NP, Cano-Lira JF, Wiederhold NP, Cano-Lira JF, Guarro J (2016) New species of Cladosporium associated with human and animal infections. Persoonia 36: 281–298. https://doi.org/10.3767/003158516X691951

Sandoval-Denis M, Sutton DA, Martin-Vicente A, Cano-Lira JF, Wiederhold N, Cano-Lira JF, Wiederhold N, Guarro J, Gené J (2015) Cladosporium species recovered from clinical samples in the United States. Journal of Clinical Microbiology 53: 2990–3000. https://doi.org/10.1128/JCM.01482-15

Schubert K, Groenewald JZ, Braun U, Dijksterhuis J, Starink M, Dijksterhuis J, Starink M, Hill CF, Zalar P, de Hoog GS, Crous PW (2007) Biodiversity in the Cladosporium herbarum complex (Davidiellaceae, Capnodiales), with standardisation of methods for Cladosporium taxonomy and diagnostics. Studies in Mycology 58: 105–156. https://doi.org/10.3114/sim.2007.58.05

Schubert K, Greslebin A, Groenewald JZ, Crous PW (2009) New foliicolous species of Cladosporium from South America. Persoonia 22: 111–122. https://doi.org/10.3767/003158509X449381

Seifert KA, Kendrick B, Murase G (1983) A Key to Hyphomycetes on Dung. Department of Biology, University of Waterloo, 62 pp.

Simões-Calaça FJ, Carvalho da Silva N, Xavier-Santos S (2014) A checklist of coprophilous fungi and other fungi recorded on dung from Brazil. Mycotaxon 128: 205–205. https://doi.org/10.5248/128.205

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. https://doi.org/10.1093/molbev/mst197

Taylor JW, Jacobson DJ, Kroken S, Kasugab T, Geiserc DM, Hibbettd DS, Fishera MS (2000) Phylogenetic species recognition and species concepts in fungi. Fungal Genetics and Biology 31: 21–32. https://doi.org/10.1006/fgbi.2000.1228

Tedersoo L, Bahram M, Puusepp R, Nilsson RH, James TY (2017) Novel soil-inhabiting clades fill gaps in the fungal tree of life. Microbiome 5: e42. https://doi.org/10.1186/s40168-017-0259-5

Temperini CV, Pardo AG, Pose GN (2018) Diversity of airborne Cladosporium species isolated from agricultural environments of northern Argentinean Patagonia: molecular characterization and plant pathogenicity. Aerobiologia 34: 227–239. https://doi.org/10.1007/s10453-018-9509-7

Thilagam L, Nayak BK, Nanda A (2015) Isolation and enumeration of saprophytic and coprophilous fungi from country cow dung. Journal of Chemical and Pharmaceutical Research 7: 474–477.

Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Research 22: 4673–4680. https://doi.org/10.1093/nar/22.22.4673

Torbati M, Arzanlou M, Bakhshi M (2016) Morphological and molecular identification of ascomycetous coprophilous fungi occurring on feces of some bird species. Current Research in Environmental & Applied Mycology 6: 210–217. https://doi.org/10.5943/cream/6/3/9

Waksman SA (1922) A method for counting the number of fungi in the soil. Journal of Bacteriology 7: 339–341. https://doi.org/10.1128/JB.7.3.339-341.1922

White TJ, Bruns T, Lee S, Taylor JW (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, New York, 315–322. https://doi.org/10.1016/B978-0-12-372180-8.50042-1

Zalar P, de Hoog GS, Schroers H-J, Crous PW, Groenewald JZ, Crous PW, Groenewald JZ, Gunde-Cimerman N (2007) Phylogeny and ecology of the ubiquitous saprobe Cladosporium sphaerospermum, with descriptions of seven new species from hypersaline environments. Studies in Mycology 58: 157–183. https://doi.org/10.3114/sim.2007.58.06

Supplementary material

Supplementary material 1

Table S1

Isabel Iturrieta-González, Dania García, Josepa Gené

Data type: Species name and strains data

Explanation note: Species, strain information and GenBank accession numbers of the sequences included in the phylogenetic analyses.

This dataset is made available under the Open Database License (http://opendatacommons.org/licenses/odbl/1.0/). The Open Database License (ODbL) is a license agreement intended to allow users to freely share, modify, and use this Dataset while maintaining this same freedom for others, provided that the original source and author(s) are credited.

Download file (53.85 kb)