Next Article in Journal
Engineered Production of Isobutanol from Sugarcane Trash Hydrolysates in Pichia pastoris
Previous Article in Journal
Heterologous Expressed NbSWP12 from Microsporidia Nosema bombycis Can Bind with Phosphatidylinositol 3-Phosphate and Affect Vesicle Genesis
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
JoF
Volume 8
Issue 8
10.3390/jof8080766
jof-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Species Diversity, Distribution, and Phylogeny of Exophiala with the Addition of Four New Species from Thailand

by
Tanapol Thitla
1,2,
Jaturong Kumla
2,3,
Surapong Khuna
2,3,
Saisamorn Lumyong
2,3,4,* and
Nakarin Suwannarach
2,3,*
1
Master of Science Program in Applied Microbiology (International Program), Faculty of Science, Chiang Mai University, Chiang Mai 50200, Thailand
2
Department of Biology, Faculty of Science, Chiang Mai University, Chiang Mai 50200, Thailand
3
Research Center of Microbial Diversity and Sustainable Utilization, Chiang Mai University, Chiang Mai 50200, Thailand
4
Academy of Science, The Royal Society of Thailand, Bangkok 10300, Thailand
*
Authors to whom correspondence should be addressed.
J. Fungi 2022, 8(8), 766; https://doi.org/10.3390/jof8080766
Submission received: 25 May 2022 / Revised: 19 July 2022 / Accepted: 20 July 2022 / Published: 24 July 2022
(This article belongs to the Topic Fungal Diversity)
Browse Figures
Versions Notes

Abstract

:
The genus Exophiala is an anamorphic ascomycete fungus in the family Herpotrichiellaceae of the order Chaetothyriales. Exophiala species have been classified as polymorphic black yeast-like fungi. Prior to this study, 63 species had been validated, published, and accepted into this genus. Exophiala species are known to be distributed worldwide and have been isolated in various habitats around the world. Several Exophiala species have been identified as potential agents of human and animal mycoses. However, in some studies, Exophiala species have been used in agriculture and biotechnological applications. Here, we provide a brief review of the diversity, distribution, and taxonomy of Exophiala through an overview of the recently published literature. Moreover, four new Exophiala species were isolated from rocks that were collected from natural forests located in northern Thailand. Herein, we introduce these species as E. lamphunensis, E. lapidea, E. saxicola, and E. siamensis. The identification of these species was based on a combination of morphological characteristics and molecular analyses. Multi-gene phylogenetic analyses of a combination of the internal transcribed spacer (ITS) and small subunit (nrSSU) of ribosomal DNA, along with the translation elongation factor (tef), partial β-tubulin (tub), and actin (act) genes support that these four new species are distinct from previously known species of Exophiala. A full description, illustrations, and a phylogenetic tree showing the position of four new species are provided.

1. Introduction

The genus Exophiala was initially described by Carmichael [1] in 1966 with Exophiala salmonis as the species type. Exophiala species are anamorphic ascomycete fungi belonging to the family Herpotrichiellaceae of the order Chaetothyriales [2]. The teleomorphic state of Exophiala has been classified in the genus Capronia [3,4]. Exophiala species are commonly known as black yeast-like fungi that are mainly characterized by annellidic conidiogenesis and yeast-like states [3,5,6]. However, several studies have indicated that Exophiala species are polymorphic fungi according to certain morphological variations that include budding cells, phialidic, catenate, or sympodial synanamorphs [3,7,8,9]. Due to the wide range of morphological variations, it is difficult to identify Exophiala by their morphological characteristics alone [6,10]. Frequently, when only morphological characteristics have been used to identify specimens within the Exophiala species, they can often be misidentified. Some species of Exophiala have been identified in the following genera: Graphium, Haplographium, Hormiscium, Phaeococcomyces, Phaeococcus, Phialophora, Pullularia, Sarcinomyces, Sporocybe, Trichosporum, and Torula [3,11,12,13,14,15,16,17]. Therefore, it is essential to identify Exophiala species by applying a molecular approach. Ribosomal DNA (ITS and nrSSU regions) and protein-coding (tef, tub, and act) genes have provided researchers with a powerful tool in the identification of the Exophiala species [3,4,5,7,11,17,18]. Currently, a combination of morphological characterization and multi-gene molecular phylogeny are being used for the accurate identification of the Exophiala species. From 1966 to the present, a total of 63 Exophiala species have been accepted and recorded in the Index Fungorum [19] and previous reports [4,20]. It has been revealed that the highest number of type species of the Exophiala were discovered during the period from 2012 to 2021 (26 species), followed by the periods from 2002 to 2011 (20 species) and 1972 to 1981 (7 species) (Figure 1). It can be expected that the trend of the discovery of new Exophiala species increasing will continue in the future.
Exophiala species have been successfully isolated in various habitats worldwide. This would indicate their capacity to adapt to different ecosystems as summarized in Table 1. Several species have been found in various natural environments [3,18,21,22,23,24,25,26]. Some species have been isolated from anthropogenic places and objects (e.g., bathrooms, gasoline tanks of cars, washing machines, and kitchen sponges) [7,27,28,29]. Moreover, some of these species have been isolated from diseased humans and animals [1,3,8,20,30,31]. Consequently, Exophiala species are known to be widely distributed in tropical, subtropical, temperate, and polar areas throughout the world (Figure 2). According to the outcomes of previous studies, it has been reported that the highest number of Exophiala species were found in Europe, accounting for 35 species. This is followed by North America (25 species), Asia (24 species), South America (18 species), the Oceania region (11 species), Africa (7 species), and Antarctica (2 species). Of these, only E. spinifera has been found to be distributed across the world [6,32,33,34,35,36,37,38,39,40,41,42,43,44]. Moreover, E. dermatitidis has been discovered in regions throughout the world, with the exception of Antarctica and the Oceania region [7,15,16,31,32,45,46,47,48,49,50,51,52,53]. Accordingly, E. jeanselmei has been found in Asia, Europe, North America, the Oceania region, and South America [6,17,27,31,32,33,34,54,55,56,57,58,59,60,61,62,63,64,65,66,67]. However, nine species (E. arunalokei, E. asiatica, E. calicioides, E. cinerea, E. clavispora, E. ellipsoidea, E. hongkongensis, E. nagquensis, and E. pseudooligosperma) have been recorded only in Asia [4,18,20,30,31,32,68]. Thirteen species, namely E. abietophila, E. bonariae, E. campbellii, E. italica, E. lacus, E. lavatrina, E. lignicola, E. mansonii, E. nidicola, E. psychrophila, E. quercina, E. radicis, and E. sideris, have been discovered in Europe [3,5,7,9,15,21,24,69,70,71,72,73]. However, E. xenobiotica has not been reported in Africa [6,7,27,31,34,57,74].
A search involving the keyword “Exophiala” retrieved 481 titles of research articles that had been published over the last 30 years (1992 to 2021) in the Scopus database [112]. The current upward trend associated with the research of Exophiala is expected to continue in the future (Figure 3A). It has been determined that the majority of applications for Exophiala have been reported in the medical field, accounting for 43.8%, followed by the fields of immunology and microbiology (18.7%), biochemistry and molecular biology (11.4%), agricultural and biological science (10.3%), veterinary medicine (5.7%), and pharmacology and toxicology (2.4%) (Figure 3B).
There are 26 Exophiala species (41.3%) that have been reported as causal agents of human diseases [1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43,44,45,46,47,48,49,50,51,52,53,54,55,56,57,58,59,60,61,62,63,64,65,66,67,68,69,70,71,72,73,74,75,76,77,78,79,80,81,82,83,84,85,86,87,88,89,90,91,92,93,94,95,96,97,98,99,100,101,102,103,104,105,106,107,108,109,110,111]. In addition, seven species of Exophiala (11.1%), namely E. angulospora, E. aquamarina, E. cancerae, E. equina, E. pisciphila, E. psychrophila, and E. salmonis, were identified as pathogens of sea creatures. However, the remaining 34 Exophiala species (54.0%) have not been associated with pathogenicity in humans or animals [1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43,44,45,46,47,48,49,50,51,52,53,54,55,56,57,58,59,60,61,62,63,64,65,66,67,68,69,70,71,72,73,74,75,76,77,78,79,80,81,82,83,84,85,86,87,88,89,90,91,92,93,94,95,96,97,98,99,100,101,102,103,104,105,106,107,108,109,110,111] (Table 1). However, in some previous studies, some Exophiala species have been effectively used in agricultural and biotechnological applications. Examples of these include E. pisciphila, which was able to promote the plant growth of maize by increasing phosphorus absorption, photosynthesis, and tolerance of cadmium [113,114]. Furthermore, by effectively suppressing Fusarium-wilt disease in strawberries, E. pisciphila could be considered a biocontrol agent [109]. In terms of drug discovery, exophillic acid and its derivative compounds derived from Exophiala species have exhibited activity against HIV-1 integrase [115,116]. Importantly, the antimicrobial property of chlorohydroaspyrones and exophilin A produced from Exophiala species has been reported [117,118]. Interestingly, Exophiala has demonstrated the ability to degrade hydrocarbons (e.g., benzene, toluene, and xylene) that can be employed in bioremediation applications [25,119]. Although Exophiala species have been researched in a variety of applications, certain risks still remain. Therefore, further research should be conducted in the future, particularly with regard to the aspects of management and safety.
Currently, only three Exophiala species have been identified in Thailand, namely E. dermatitidis, E. jeanselmei, and E. spinifera [43,46,67]. Accordingly, many studies have proposed that Thailand has proven to be a hot spot for novel microfungal species discovery [120,121,122]. During investigations of rock-inhabiting fungi in northern Thailand during the period of 2020 to 2021, we obtained fifteen Exophiala strains that are potentially representative of new species. In the present study, we describe four new species, namely E. lamphunensis, E. lapidea, E. saxicola, and E. siamensis. These four new species were identified based on morphological and molecular data. To confirm their taxonomic status, phylogenetic relationships were determined by analysis of the combined sequence dataset of ITS, nrSSU, tef, tub, and act genes.

2. Materials and Methods

2.1. Sample Collection and Fungal Isolation

Rock samples were collected from four natural forests located in Lamphun (three sites; 18°32′11″ N 99°07′29″ E, 18°32′10″ N 99°07′30″ E, and 18°32′11″ N 99°07′30″ E) and Sukhothai (17°32′58″ N 99°29′49″ E) Provinces, northern Thailand. The samples comprising flourishing black colonies were collected with a sterile chisel, kept in plastic bags, and carried to the laboratory in an ice box. All collected rock samples were processed for the isolation of fungi immediately after reaching the laboratory. Fungi were isolated using the method described by Selbmann et al. [123] with some modifications. Rock samples were washed in 1% sodium hypochlorite for 10 min and rinsed 5 times in sterile water. Fungal isolation was performed by pulverizing the rock samples and sprinkling rock powder onto 2% malt extract agar (MEA; Difco, Le Pont de Claix, France) and dichloran-rose bengal agar (DRBC; Difco, Le Pont de Claix, France) supplemented with chloramphenicol 100 ppm. Plates were incubated at 25 °C for 4 weeks. Plates were then inspected every day. Fungal colonies with dark pigments were transferred to fresh MEA. Pure fungal strains were kept in 20% glycerol and deposited in the Culture Collection of Sustainable Development of Biological Resources Laboratory (SDBR), Faculty of Science, Chiang Mai University, Chiang Mai, Thailand.

2.2. Morphological and Growth Observations

Agar plugs (5 mm in diameter) from the edges of each fungal strain were transferred onto plates containing potato dextrose agar (PDA; Condalab, Madrid, Spain), MEA, and oatmeal agar (OA; Difco, Le Pont de Claix, France) and then kept at 25 °C in the dark. After four weeks of incubation, relevant colony features, including aerial mycelium and pigment production, were recorded and the colony diameter was measured. Cardinal growth temperatures were studied on MEA for 4 weeks in the dark at 4, 10, 15, 20, 25, 28, 30, 35, 37, and 40 °C using the method described by de Hoog et al. [3] with some modifications. A light microscope (Nikon Eclipse Ni-U, Tokyo, Japan) was used to study the micromorphological features of each fungal strain. The anatomical structure related to size data (e.g., hyphae, budding cells, conidia, and chlamydospore) was based on at least 50 measurements of each structure using the Tarosoft (R) Image.

2.3. DNA Extraction, Amplification, and Sequencing

A Fungal DNA Extraction Kit (FAVORGEN, Ping-Tung, Taiwan) was used to extract genomic DNA from the 3-week-old fungal culture of each strain that grew on MEA at 25 °C. Ribosomal DNA (ITS and nrSSU regions) and protein-coding (tef, tub, and act) genes were amplified by polymerase chain reaction (PCR) using suitable primers (Table 2). PCR amplifications were performed using 20-µL reaction mixtures containing 1 µL of genomic DNA, 1 µL of 10 µM forward and reverse primers, 10 µL of Quick TaqTM HS DyeMix (TOYOBO, Osaka, Japan), and 7 µL of deionized water. PCR amplification conditions consisted of an initial denaturation step conducted at 95 °C for 5 min, followed by 35 cycles of denaturation at 95 °C for 30 s, an annealing step for 30 s, at appropriate temperatures (Table 2), and an elongation step at 72 °C for 1 min on a peqSTAR thermal cycler (PEQLAB Ltd., Fareham, UK). PCR products were checked on 1% agarose gel electrophoresis and were purified using a PCR clean up Gel Extraction NucleoSpin® Gel and a PCR Clean-up Kit (Macherey-Nagel, Düren, Germany). Purified PCR products were then sequenced by 1st Base Company (Kembangan, Malaysia).

2.4. Sequence Alignment

The resulting ITS, nrSSU, tef, tub, and act sequences were assessed for similarity analysis in the GenBank database via BLAST searching. The sequences from this study, and those of closely related fungi, were obtained from the nucleotide GenBank database and previous studies as listed in Table 3. Multiple sequence alignment was carried out using MUSCLE in MEGA v. 6 [128] and the results were enhanced, when necessary, using BioEdit v.6.0.7 [129].

2.5. Phylogenetic Analyses

Phylogenetic analyses were performed using combination datasets of ITS, nrSSU, tef, tub, and act genes. Cyphellophora eucalypti CBS 124764 and C. fusarioides MUCL 44033 were used as the outgroup. Maximum likelihood (ML) and Bayesian inference (BI) methods were used to generate a phylogenetic tree. For ML analysis, 25 categories and 1000 bootstrap (BS) replications under the GTRCAT model [145] were performed on RAxML-HPC2 version 8.2.12 [146] on the CIPRES web portal [147]. The evolutionary model of nucleotide substitution for BI analysis was selected independently for each gene using MrModeltest v. 2.1 [148]. The GTR + I + G substitution model was the best fit for the ITS and nrSSU genes while the HKY + I + G substitution model was the best fit for the tef and tub genes, and the HKY + G substitution model was the best fit for the act gene. MrBayes v.3.2.6 was used for BI analysis [149]. In total, 6 simultaneous Markov chains were run for 5 million generations with random initial trees, wherein every 1000 generations were sampled. A burn-in phase was used to eliminate the first 2000 trees while the remaining trees were utilized to create a phylogram with a 50% majority-rule consensus. The Bayesian posterior probability (PP) was then calculated. Branches with BS and PP values of more than or equal to 70% and 0.95, respectively, were deemed to have been substantially supported. The tree topologies were visualized in FigTree v1.4.0 [150].

3. Results

3.1. Fungal Isolation and Morphological Observations

A total of fifteen fungal strains were obtained in this study. Thirteen strains were isolated from rock samples collected from Lamphun Province and two strains were isolated from rock samples collected from Sukhothai Province. All fungal strains were cultivated on MEA at various temperatures (4–40 °C) and the diameter of the colonies was measured after 4 weeks of incubation. The results indicated that temperature had a significant effect on fungal growth. The average colony diameter of each fungal strain is shown in Table 4. It was found that that all fungal strains could not grow at 4 and 40 °C. However, all fungal strains grew well in temperatures ranging from 25–30 °C, with the exception of the strains SDBR-CMU417 and SDBR-CMU418. Five fungal strains (SDBR-CMU404, SDBR-CMU405, SDBR-CMU406, SDBR-CMU407, and SDBR-CMU408) showed the highest average value of the colony diameter at 28 °C while eight fungal strains (SDBR-CMU409, SDBR-CMU410, SDBR-CMU411, SDBR-CMU412, SDBR-CMU413, SDBR-CMU414, SDBR-CMU415, and SDBR-CMU416) showed the highest average value of the colony diameter at 30 °C. The results indicate that the highest average value of the colony diameter of two fungal strains, namely SDBR-CMU417 and SDBR-CMU418, was found at 20 °C; however, they did not grow at 35 and 37 °C. Based on the morphological characteristics, all fungal isolates were initially identified as belonging to the genus Exophiala. The identification was then further confirmed by the multi-gene phylogenetic analysis of the ITS, nrSSU, tub, tef, and act sequences.

3.2. Phylogenetic Results

A phylogenetic tree was constructed using a combination of the ITS, nrSSU, tub, tef, and act genes containing 3616 characters, including gaps (ITS: 1–739, nrSSU: 740–1829, tef: 1830–2454, tub: 2455–3045, and act: 3046–3616). The phylogram was constructed, consisting of 105 specimens of Exophiala and 2 specimens of the outgroup (Cyphellophora fusarioides MUCL 44033 and C. eucalypti CBS 124764). RAxML analysis of the combined dataset yielded the best scoring tree, with a final log likelihood value of −38,143.750648. The matrix was comprised of 1880 distinct alignment patterns with 53.45% undetermined characters or gaps. Estimated base frequencies were recorded as follows: A = 0.2297, C = 0.2831, G = 0.2311, T = 0.2561; substitution rates AC = 1.2386, AG = 4.4108, AT = 0.9986, CG = 0.8412, CT = 7.1418, and GT = 1.0000. The gamma distribution shape parameter alpha was equal to 0.3965 and the Tree-Length was equal to 12.0800. Using BI analysis, the final average standard deviation of the split frequencies at the end of the total MCMC generations was estimated to be 0.00513. In terms of topology, the phylograms of the ML and BI analyses were similar (data not shown). The phylogram generated from the ML analysis is shown in Figure 4. Our phylogenetic tree was constructed concordantly and is supported by previous studies [4,18]. The phylogram separated all fungal strains in this study into four monophyletic clades with high BS and PP support values. These clearly formed distinct lineages from previous known Exophiala species with high BS and PP support values. The results of our study revealed that two fungal strains, namely SDBR-CMU417 and SDBR-CMU418 (introduced as E. siamensis), were clearly separated from the previously known species of Exophiala. Moreover, five fungal strains, SDBR-CMU404, SDBR-CMU405, SDBR-CMU406, SDBR-CMU407, and SDBR-CMU408 (introduced as E. lamphunensis), formed a sister taxon to the two strains SDBR-CMU415 and SDBR-CMU416 (described here as E. saxicola), with 80% and 1.00 BS and PP support values, respectively. Notably, E. lamphunensis and E. saxicola formed a sister clade to E. xenobiotica, with high BS (98%) and PP (1.0) support values. Moreover, our six strains, SDBR-CMU409, SDBR-CMU410, SDBR-CMU411, SDBR-CMU412, SDBR-CMU413, and SDBR-CMU414 (introduced as E. lapidea), formed a sister taxon to E. moniliae (BS = 99% and PP = 1.0).

3.3. Taxonomic Descriptions

Exophiala lamphunensis Thitla, J. Kumla and N. Suwannarach sp. nov. (Figure 5).
MycoBank No.: 844209.
Etymology: “lamphunensis”, referring to Lamphun Province, the original place of fungus isolation.
Holotype: THAILAND, Lamphun Province, Mueang Lamphun District, Sribuaban Subdistrict, 18°32′11″ N 99°07′29″ E elevation 414 m, isolated from the rock of natural forest, July 2021, T. Thitla, dried culture: SDBR-LPN6_65; ex-type culture: SDBR-CMU404.
GenBank: ON555798 (ITS), ON555813 (nrSSU), ON544242 (tef), ON544227 (tub), and ON544257 (act).
Culture characteristics: Colonies on PDA, MEA, and OA were described at 25 °C after 28 days of incubation (Figure 5A). Colonies on PDA reached 20–24 mm in diameter, restricted, circular, flat, velvety, and greyish green to dark green with greyish-green edges. Reverse dark green at the center and dull-green to greyish-green entire margin. Colonies on MEA attained a diameter of 23–26 mm, restricted, circular, raised, and velvety with dull-green aerial mycelium and entire margins. Reverse dark green to greyish green. Colonies on OA reached a diameter of 22–24 mm, restricted, circular, and velvety with greenish-grey and dark-green margins. Reverse dull green. A soluble dark-green pigment was observed around the fungal colonies on PDA. Budding cells initially abundant, hyaline, subspherical to ellipsoidal, 2.8–7.2 × 2.0–4.4 µm (mean = 4.6 × 3.3 µm, n = 50) (Figure 5B). Germinating cells abundant, hyaline, subspherical to ellipsoidal, 3.1–7.3 × 2.4–5.8 µm (mean = 5.0 × 3.5 µm, n = 50) (Figure 5C). Hyphae smooth-walled, pale olive-brown, 1.2–3.2 µm wide. Hyphal coils abundant while anastomoses absent (Figure 5D). Conidiophores short, subcylindrical, and intercalary of hyphae (Figure 5E). Conidiogenous cells erect, cylindrical with short annellated zones emerging from both the conidiophore and the terminal or the intercalary of the hyphae (Figure 5E,F). Conidia attached in tiny clusters, subhyaline, obovoidal, and 2.7–5.3 × 1.5–3.2 µm (mean = 3.8 × 2.1 µm, n = 50) (Figure 5E–G). Chlamydospores absent. Torulose hyphae up to 6 µm wide in appearance. Teleomorph were not found in any culture media.
Growth temperature: growth occurred within a range of 10–37 °C, optimum at 28 °C, while no growth at 4 and 40 °C.
Additional specimens examined: THAILAND, Lamphun Province, Mueang Lamphun District, Sribuaban Subdistrict, 18°32′11″ N 99°07′29″ E elevation 414 m, isolated from rock in dipterocarp forest, July 2021, isolated by T. Thitla: SDBR-CMU405, SDBR-CMU406, SDBR-CMU407, and SDBR-CMU408.
Known distribution: Lamphun Province, Thailand.
Note: Colonies on MEA of E. lamphunensis were similar to E. nagquensis, E. oligosperma, E. saxicola, and E. xenobiotica. However, E. oligosperma, E. saxicola, and E. xenobiotica differed from E. lamphunensis in the way they did not produce any soluble pigment on PDA [17,18,27]. The conidial sizes (4.8–10.4 × 2.6–5.0 µm) of E. nagquensis were larger than E. lamphunensis while E. nagquensis could grow at 4 °C [18]. The size of the budding cells and conidia of E. lamphunensis were within the range of E. oligosperma, E. saxicola, and E. xenobiotica. However, E. oligosperma and E. saxicola produced chlamydospores that differed from E. lamphunensis. Moreover, the optimum growth of E. saxicola was observed at 30 °C, which was higher than for E. lamphunensis. Notably, E. xenobiotica differed from E. lamphunensis in it has a slightly shorter conidial size (3.3–4.0 × 1.6–2.0 µm) and chlamydospore formation [17,27].
The phylogenetic analyses of the combined ITS, nrSSU, tub, tef, and act sequences confirmed that E. lamphunensis formed a monophyletic clade that clearly distinguished it from E. nagquensis, E. oligosperma, E. saxicola, and E. xenobiotica. Exophiala lamphunensis formed a sister clade to E. saxicola. However, sequence similarity and pairwise nucleotide comparison of tef data also showed that E. lamphunensis differs from E. saxicola in 97% and 3.1% (5/162 bp), respectively. Differences in the morphological characteristics and the optimum growing temperature were found between E. lamphunensis and E. saxicola. Exophiala lamphunensis produces soluble pigment on PDA and chlamydospore production is absent while this was not the case for E. saxicola. The slightly wider size of the germinating cells in E. lamphunensis (3.1–7.3 × 2.4.–5.8 µm) distinguished it from E. saxicola (3.6–6.0 × 1.9–3.7 µm). Additionally, E. lamphunensis had a lower optimum temperature (28 °C) than E. saxicola (30 °C). Therefore, E. lamphunensis and E. saxicola were considered as different species based on their morphological, optimal growth temperature, and tef sequence data.
Exophiala lapidea Thitla, J. Kumla and N. Suwannarach sp. nov. (Figure 6).
MycoBank No.: 844211.
Etymology: “lapidea” referring to the fungi being isolated from rock.
Holotype: THAILAND, Lamphun Province, Mueang Lamphun District, Sribuaban Subdistrict, 18°32′10″ N 99°07′30″ E elevation 407 m, isolated from the rock of natural forest, July 2021, T. Thitla, dried culture: SDBR-LPN8_9; ex-type culture: SDBR-CMU409.
GenBank: ON555803 (ITS), ON555818 (nrSSU), ON544247 (tef), ON544232 (tub), and ON544262 (act).
Culture characteristics: Colonies on PDA, MEA, and OA were described at 25 °C after 28 days of incubation (Figure 6A). Colonies on PDA grew to 35–48 mm in diameter, restricted, flat, velvety, greyish brown to dark brown, with black slime at the center. Reverse black with brown margin. Colonies on MEA reached 32–42 mm in diameter, restricted, flat, dull green to greyish green with aerial mycelium at the middle. Reverse dark green with deep-green margins. Colonies on OA grew to a diameter of 30–33 mm, restricted, circular, flat, velvety, greenish grey with aerial mycelium at the middle and dark-green edge. Reverse dark green. Budding cells abundant, hyaline, spherical or ellipsoidal, 2.8–5.1 × 2.2–4.7 µm (mean = 4.2 × 3.7 µm, n = 50) (Figure 6B,C). Germinating cells ellipsoidal, 4.6–8.4 × 2.5–5.3 µm (mean = 6.6 × 3.8 µm, n = 50) (Figure 6D,E). Hyphae smooth, thin-walled, pale olive-brown, usually spiral, 1.2–2.1 µm wide. Anastomoses and hyphal coil abundant (Figure 6F–H). Conidiophores pale olivaceous brown, erect, cylindrical, inserted laterally on hyphae (Figure 6I). Conidiogenous cells erect, cylindrical, with short annellated zones emerging from hyphae, terminal or intercalary (Figure 6J–L). Conidia hyaline, thin-walled, obovoidal, 2.9–7.0 × 0.9–2.4 µm (mean = 4.3 × 1.5 µm, n = 50) with inconspicuous basal scars (Figure 6I–N). Chlamydospores absent. Torulose hyphae appeared, up to 7 μm wide (Figure 6O). Teleomorph not found in any culture media.
Growth temperature: growth occurred within a range of 10–37 °C, optimum at 28 °C, while no growth at 4 and 40 °C.
Additional specimens examined: THAILAND, Lamphun Province, Mueang Lamphun District, Sribuaban Subdistrict, 18°32′10″ N 99°07′30″ E elevation 407 m, isolated from rock in dipterocarp forest, July 2021, isolated by T. Thitla: SDBR-CMU410, SDBR-CMU411, SDBR-CMU412, SDBR-CMU413, and SDBR-CMU414.
Known distribution: Lamphun Province, Thailand.
Note: The colony characteristics of E. lapidea were similar to those of E. aquamarine, E. cancerae, and E. eucatypticola. However, the conidial size of E. lapidea (2.9–7.0 × 0.9–2.4 µm) was clearly smaller than E. aquamarine (6.7–19.2 × 4.0–4.8 µm) [3]. The wider size of the conidia in E. cancerae (4.9–8.0 × 2.7–4.8) and E. eucalypticola (4.0–7.0 × 2.0–3.0 µm) clearly distinguished them from E. lapidea [3,22]. Moreover, E. cancerae and E. eucalypticola could effectively grow at 4 °C.
The multi-gene phylogenetic analyses (ITS, nrSSU, tub, tef, and act genes) confirmed that E. lapidea formed a monophyletic clade that clearly separated it from the other previous known Exophiala species and closely related species. A phylogram showed that E. lapidea formed a sister taxon to E. moniliae (Figure 4). However, the shorter size of conidia in E. moniliae (2.3–3.9 × 1.6–2.2 µm) clearly distinguished it from E. lapidea [15].
Exophiala saxicola Thitla, N. Suwannarach and S. Lumyong sp. nov. (Figure 7).
MycoBank No.: 844212.
Etymology: “saxicola” referring to a stone inhabitant.
Holotype: THAILAND, Lamphun Province, Mueang Lamphun District, Sribuaban Subdistrict, 18°32′11″ N 99°07′30″ E elevation 413 m, isolated from the rock of natural forest, July 2021, T. Thitla, dried culture: SDBR-LPN6_71; ex-type culture: SDBR-CMU415.
GenBank: ON555809 (ITS), ON555824 (nrSSU), ON544253 (tef), ON544238 (tub), and ON544268 (act)
Culture characteristics: Colonies on PDA, MEA, and OA were described at 25 °C after 28 days of incubation (Figure 7A). All culture media restricted, circular, flat, and velvety. On PDA, PDA grew to 16–18 mm in diameter, dull green and dark green in reverse. Colonies on MEA reached 20–22 mm in diameter, dull-green and greyish-green margins. Reverse greyish green to dark green. Colonies on OA attained a diameter of 22–24 mm, greenish grey to dark green. Reverse dark green. Budding cells initially abundant, hyaline, subspherical to ellipsoidal, 4.0–7.0 × 2.7–5.2 µm (mean = 5.6 × 3.8 µm, n = 50) (Figure 7B,C). Germinating cells abundant, hyaline, ellipsoidal, 3.6–6.0 × 1.9–3.7 µm (mean = 4.8 × 2.6 µm, n = 50) (Figure 7D,E). Hyphae smooth-walled, pale olive-brown, 1.1–3.3 µm wide. Anastomoses abundant (Figure 7F). Conidiophores pale olivaceous brown, erect, cylindrical (Figure 7G). Conidiogenous cells obovoidal to clavate with short annellated zones, intercalary or terminal of hyphae (Figure 7H,I). Conidia adhering in small groups, hyaline, obovoidal, 2.8–6.2 × 1.3–3.5 µm (mean = 4.4 × 2.3 µm, n = 50) (Figure 7G–K). Chlamydospores are presented, subspherical, brown, 4.1–8.1 × 3.3–7.2 µm (Figure 7L). Torulose hyphae appeared up to 5 μm in width (Figure 7M). Teleomorph not found in any culture media.
Growth temperatures: growth occurred within a range of 10–37 °C, optimum at 30 °C, while no growth at 4 and 40 °C.
Additional specimens examined: THAILAND, Lamphun Province, Mueang Lamphun District, Sribuaban Subdistrict, 18°32′11″ N 99°07′30″ E elevation 413 m, isolated from rock in dipterocarp forest, July 2021, isolated by T. Thitla: SDBR-CMU416.
Known distribution: Lamphun Province, Thailand.
Note: The colony characteristics of E. saxicola on MEA were similar to those observed for E. xenobiotica, E. nagquensis, E. oligosperma, and E. lamphunensis. The production of soluble pigment on PDA was observed as was an absence of chlamydospore formation in E. lamphunensis, which clearly distinguished it from E. saxicola. The budding cells of E. saxicola (4.0–7.0 × 2.7–5.2 µm) were larger than the budding cells of E. oligosperma (3.0 × 2.5 µm) [17]. Notably, the small size of the germinating cells in E. saxicola (3.6–6.0 × 1.9–3.7 µm) clearly distinguished it from E. xenobiotica (7.0–10.0 × 3.0–5.0 µm) [27] and E. oligosperma (6.0 × 5.0 µm) [17]. Moreover, the conidia size of E. nagquensis (4.8–10.4 × 2.6–5.0 µm) was larger than E. saxicola [18]. The optimal growth temperature of E. saxicola (30 °C) distinguished it from E. lamphunensis (28 °C). Moreover, the maximum growth temperature of E. saxicola (37 °C) was higher than for E. xenobiotica (33–36 °C) [27] and E. nagquensis (28 °C) [18].
The phylogenetic analyses of the combined ITS, nrSSU, tub, tef, and act sequences confirmed that E. saxicola formed a monophyletic clade that clearly distinguished it from the other closely related species, namely E. nagquensis, E. oligosperma, and E. xenobiotica. Furthermore, E. saxicola formed a sister clade to E. lamphunensis. However, differences in the morphological characteristics, optimal growth temperature, and tef sequence data of E. saxicola and E. lamphunensis were observed and described above.
Exophiala siamensis Thitla, J. Kumla and N. Suwannarach sp. nov. (Figure 8).
MycoBank: 844213.
Etymology: “siamensis” referring to Siam (old name of Thailand), where this fungus was found.
Holotype: THAILAND, Sukhothai Province, Si Satchanalai District, 17°32′58″ N 99°29′49″ E elevation 153 m, isolated from the rock of natural forest, June 2021, T. Thitla, dried culture: SDBR-SKT3_3; ex-type culture: SDBR-CMU417.
GenBank: ON555811 (ITS), ON555826 (nrSSU), ON544255 (tef), ON544240 (tub), and ON544270 (act).
Culture characteristics: Colonies on PDA, MEA, and OA were described at 25 °C after 28 days of incubation (Figure 8A). Colonies on PDA were 14–21 mm in diameter, restricted, irregular, convex in elevation, and velvety with brownish-grey and dark-brown margins. Reverse black. Colonies on MEA and OA restricted, circular, flat, velvety. Colonies on MEA grew to 9–11 mm in diameter with dark-green to greyish-green and white margins. Reverse dark green. Colonies on OA reached a diameter of 15–16 mm with dark-green and greyish-green margins. Reverse black and olive margin. Budding cells rarely, hyaline, subspherical, 5.8–7.6 × 4.3–5.9 µm (mean = 6.7 × 5.3 µm, n = 50) (Figure 8B). Germinating cells ovoidal or obovoidal, 4.7–6.2 × 3.2–4.8 µm (mean = 5.5 × 3.9 µm, n = 50) (Figure 8C). Hyphae smooth, thin-walled, pale olive-brown, 1.2–3.0 µm in width, producing conidia apically and laterally. Anastomoses presence (Figure 8D). Conidiophores short, erect, cylindrical (Figure 8E). Conidiogenous cells cylindrical to ellipsoidal, terminal or intercalary (Figure 8F–H). Conidia hyaline, thin-walled, subspherical, 1.9–3.5 × 1.5–3.2 µm (mean = 2.7 × 2.2 µm, n = 50) (Figure 8E–I). Chlamydospores subspherical, pale brown, 7.4–16.5 × 3.1–6.7 µm (Figure 8J). Torulose hyphae appeared up to 6 μm in width (Figure 8K). Teleomorph not found in any culture media.
Growth temperatures: growth occurred within a range of 10–30 °C, optimum at 20 °C, while no growth at 4, 35, 37, and 40 °C.
Additional specimens examined: THAILAND, Sukhothai Province, Si Satchanalai District, 17°32′58″ N 99°29′49″ E elevation 153 m, isolated from rock in dipterocarp forest, June 2021, isolated by T. Thitla: SDBR-CMU418.
Known distribution: Sukhothai Province, Thailand.
Note: Morphologically, the colony characteristics of E. siamensis were similar to E. ellipsoidea, E. brunnea, E. polymorpha, and E. radicis. However, the wider size of the budding cells in E. siamensis (5.8–7.6 × 4.3–5.9 µm) clearly separated it from E. polymorpha (4.0–6.0 × 2.5–4.0 µm) [8]. The conidial size of E. siamensis (1.9–3.5 × 1.5–3.2 µm) was smaller than E. radicis (4.0–11.0 × 2.0–5.0 µm) [5]. In addition, the conidia size of E. siamensis was clearly shorter than E. brunnea (4.5–10.0 × 2.0–3.0 µm) [3], E. ellipsoidea (2.1–6.4 × 1.1–1.0 µm) [18], and E. polymorpha (3.5–4.0 × 1.5–2.5 µm) [8]. Exophiala siamensis produced chlamydospores that were different from E. polymorpha and E. radices [5,8]. The maximum growth temperature of E. ellipsoidea (33 °C) and E. polymorpha (30 °C) was higher than for E. siamensis (30 °C) [8,18]. The minimum growth temperature of E. brunnea (4–9 °C) was lower than E. siamensis (10 °C) [3].
Moreover, a multi-gene phylogenetic analysis confirmed that E. siamensis formed a well-supported monophyletic clade that was distinctly separated from other Exophiala species.

4. Discussion

Species of the genus Exophiala are known to be widely distributed around the world [3,7,33,151]. The traditional identification of Exophiala species has primarily been based on morphological characteristics [1,15,152]. However, identification can be difficult because some of the polymorphic characteristics are shared and some species have a similar appearance [3,5,8,15,89]. As a result, some previously identified Exophiala species were then transferred from the genera Graphium, Haplographium, Hormiscium, Phaeococcomyces, Phaeococcus, Phialophora, Pullularia, Sarcinomyces, Sporocybe, Trichosporum, and Torula [3,11,12,13,14,15,16,17]. Therefore, a combination of morphological and multi-gene data was used to concretely identify the Exophiala species [3,4,5,9,18]. Prior to conducting our study, a total of 63 species had been validated, published, and accepted into the genus Exophiala.
In this study, four new species of Exophiala, consisting of E. lamphunensis, E. lapidea, E. saxicola, and E. siamensis, were introduced. The different morphological characteristics identified between the four new species indicate that only E. lamphunensis produced soluble pigments around the colonies on PDA. Chlamydospore formations were observed in E. saxicola and E. siamensis, but this was not the case for E. lamphunensis and E. lapidea. Additionally, the budding cells of E. siamensis were larger and wider than those of E. lapidea and E. lamphunensis. However, the germinating cells and conidia of our four species were not observed to be different. The optimum growth temperature of E. lapidea and E. saxicola was 30 °C, which was higher than for E. lamphunensis (28 °C) and E. siamensis (28 °C). Additionally, the maximum growth temperature of E. siamensis (30 °C) was lower than for the other three new species (37 °C). Subsequently, our phylogenetic analyses of the combined five genes (ITS, nrSSU, tub, tef, and act) revealed that the four new species formed distinct lineages within the genus Exophiala. Therefore, a combination of the morphological characteristics and the molecular analyses conducted in our study strongly support the recognition of four new Exophiala species.
Exophiala species have been isolated in various habitats throughout the world as shown in Table 1. Several Exophiala species have been identified as potential agents of human and animal diseases. However, in some studies, certain Exophiala species have been employed in agricultural and biotechnological applications. In this study, four new Exophiala species were isolated from rock samples collected from natural forests located in northern Thailand. Our findings are similar to those of previous studies, which reported that some Exophiala species (e.g., E. bonaiae, E. cinerea, E. clavispora, E. ellipsoidea, and E. nagquensis) have been successfully isolated from rock samples. However, there have been no prior reports involving investigations of rock-inhabiting fungi in Thailand. Therefore, our study is the first of its kind to report on the discovery of Exophiala on rocks in Thailand. Prior to our study, a total of three Exophiala species (E. dermatitidis, E. jeanselmei, and E. spinifera) were known from Thailand [43,46,67]. Therefore, the successful identification of the Exophiala species in this study has increased the number of species found in Thailand to 7 species and has led to 67 global species. The outcomes of this present study will provide scientists and researchers with valuable information that can stimulate deeper investigations of rock-inhabiting fungi in Thailand. Ultimately, these findings will help researchers gain a better understanding of the distribution and ecology of Exophiala.

Author Contributions

Conceptualization, N.S., J.K. and S.L.; methodology, T.T., N.S., J.K. and S.K; software, T.T. and J.K.; validation, T.T., N.S., J.K. and S.L.; formal analysis, T.T., J.K. and N.S.; investigation, T.T., N.S. and J.K.; resources, T.T., J.K., N.S. and S.K.; data curation, T.T., N.S., J.K. and S.L.; writing—original draft, T.T., N.S., J.K. and S.K.; writing—review and editing, T.T., N.S., J.K., S.K. and S.L.; supervision, S.L. and N.S. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by TA&RA Scholarship, Graduate School, Chiang Mai University, and partially supported by Chiang Mai University, Thailand. This project was funded by Research Council of Thailand (N42A650198).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The DNA sequence data obtained from this study have been deposited in GenBank under the accession numbers; ITS (ON555798, ON555799, ON555800, ON555801, ON555802, ON555803, ON555804, ON555805, ON555806, ON555807, ON555808, ON555809, ON555810, ON555811, ON555812); nrSSU (ON555813, ON555814, ON555815, ON555816, ON555817, ON555818, ON555819, ON555820, ON555821, ON555822, ON555823, ON555824, ON555825, ON555826, ON555827); tub (ON544227, ON544228, ON544229, ON544230, ON544231, ON544232, ON544233, ON544234, ON544235, ON544236, ON544237, ON544238, ON544239, ON544240, ON544241); tef (ON544242, ON544243, ON544244, ON544245, ON544246, ON544247, ON544248, ON544249, ON544250, ON544251, ON544252, ON544253, ON544254, ON544255, ON544256) and act (ON544257, ON544258, ON544259, ON544260, ON544261, ON544262, ON544263, ON544264, ON544265, ON544266, ON544267, ON544268, ON544269, ON544270, ON544271).

Acknowledgments

The authors are grateful to Russell Kirk Hollis for the English correction of this manuscript.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Carmichael, J.W. Cerebral mycetoma of trout due to a phialophora-like fungus. Sabouraudia 1966, 5, 120–123. [Google Scholar] [CrossRef] [PubMed]
  2. Zhang, C.; Sirijovski, N.; Adler, L.; Ferrari, B.C. Exophiala macquariensis sp. nov., a cold adapted black yeast species recovered from a hydrocarbon contaminated sub-Antarctic soil. Fungal Biol. 2019, 123, 151–158. [Google Scholar] [CrossRef] [PubMed]
  3. de Hoog, G.S.; Vicente, V.A.; Najafzadeh, M.J.; Harrak, M.J.; Badali, H.; Seyedmousavi, S. Waterborne Exophiala species causing disease in cold-blooded animals. Persoonia 2011, 27, 46–72. [Google Scholar] [CrossRef] [Green Version]
  4. Yang, X.Q.; Feng, M.Y.; Yu, Z.F. Exophiala pseudooligosperma sp. nov., a novel black yeast from soil in southern China. Int. J. Syst. Evol. Microbiol. 2021, 71, e005116. [Google Scholar] [CrossRef]
  5. Maciá-Vicente, J.G.; Glynou, K.; Piepenbring, M. A new species of Exophiala associated with roots. Mycol. Prog. 2016, 15, e18. [Google Scholar] [CrossRef]
  6. Zeng, J.S.; de Hoog, G.S. Exophiala spinifera and its allies: Diagnostics from morphology to DNA barcoding. Med. Mycol. 2008, 46, 193–208. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  7. Borman, A.M.; Fraser, M.; Szekely, A.; Larcombe, D.E.; Johnson, E.M. Rapid identification of clinically relevant members of the genus Exophiala by matrix-assisted laser desorption ionization–time of flight mass spectrometry and description of two novel species, Exophiala campbellii and Exophiala lavatrina. J. Clin. Microbiol. 2017, 55, 1162–1176. [Google Scholar] [CrossRef] [Green Version]
  8. Yong, L.K.; Wiederhold, N.P.; Sutton, D.A.; Sandoval-Denis, M.; Lindner, J.R.; Fan, H.; Sanders, C.; Guarro, J. Morphological and molecular characterization of Exophiala polymorpha sp. nov. isolated from sporotrichoid lymphocutaneous lesions in a patient with Myasthenia Gravis. J. Clin. Microbiol. 2015, 53, 2816–2822. [Google Scholar] [CrossRef] [Green Version]
  9. Seyedmousavi, S.; Badali, H.; Chlebicki, A.; Zhao, J.; Prenafeta-Boldú, F.X.; De Hoog, G.S. Exophiala sideris, a novel black yeast isolated from environments polluted with toxic alkyl benzenes and arsenic. Fungal Biol. 2011, 115, 1030–1037. [Google Scholar] [CrossRef]
  10. Julou, T.; Burghardt, B.; Gebauer, G.; Berveiller, D.; Damesin, C.; Selosse, A. Mixotrophy in orchids: Insights from a comparative study of green individuals and nonphotosynthetic individuals of Cephalanthera damasonium. New Phytol. 2005, 166, 639–653. [Google Scholar] [CrossRef]
  11. Haase, G.; Sonntag, L.; Melzer-Krick, B.; de Hoog, G.S. Phylogenetic inference by SSU gene analysis of members of the Herpotrichiellaceae, with special reference to human pathogenic species. Stud. Mycol. 1999, 43, 80–97. [Google Scholar]
  12. McGinnis, M.R.; Padhye, A.A. Exophiala jeanselmei, a new combination for Phialophora jeanselmei. Mycotaxon 1977, 5, 341–352. [Google Scholar]
  13. Okada, G.; Jacobs, K.; Kirisits, T.; Louis-Seize, G.W.; Seifert, K.A.; Sugita, T.; Takematsu, A.; Wingfield, M.J. Epitypification of Graphium penicillioides Corda, with comments on the phylogeny and taxonomy of graphium-like synnematous fungi. Stud. Mycol. 2000, 45, 169–188. [Google Scholar]
  14. McGinnis, M.R. Exophiala spinifera, a new combination for Phialophora spinifera. Mycotaxon 1977, 5, 337–340. [Google Scholar]
  15. de Hoog, G.S.; Hermanides-Nijhof, E.J. The black yeasts and allied Hyphomycetes. Stud. Mycol. 1977, 15, e222. [Google Scholar]
  16. Matos, T.; Haase, G.; Gerrits van den Ende, A.H.G.; de Hoog, G.S. Molecular diversity of oligotrophic and neurotropic members of the black yeast genus Exophiala, with accent on E. dermatitidis. Antonie Van Leeuwenhoek 2003, 83, 293–303. [Google Scholar] [CrossRef]
  17. de Hoog, G.S.; Vicente, V.; Caligiorne, R.B.; Kantarcioglu, S.; Tintelnot, K.; Gerrits van den Ende, A.H.; Haase, G. Species diversity and polymorphism in the Exophiala spinifera clade containing opportunistic black yeast-like fungi. J. Clin. Microbiol. 2003, 41, 4767–4778. [Google Scholar] [CrossRef] [Green Version]
  18. Sun, W.; Su, L.; Yang, S.; Sun, J.; Liu, B.; Fu, R.; Wu, B.; Lui, X.; Cai, L.; Guo, L.; et al. Unveiling the hidden diversity of rock-inhabiting fungi: Chaetothyriales from China. J. Fungi 2020, 6, e187. [Google Scholar] [CrossRef]
  19. Index Fungorum. Available online: http://www.indexfungorum.org (accessed on 22 March 2022).
  20. Singh, S.; Rudramurthy, S.M.; Padhye, A.A.; Hemashetter, B.M.; Iyer, R.; Hallur, V.; Sharma, A.; Agnihotri, S.; Gupta, S.; Ghosh, A.; et al. Clinical spectrum, molecular characterization, antifungal susceptibility testing of Exophiala spp. from India and description of a novel Exophiala species, E. arunalokei sp. nov. Front. Cell Infect. Microbiol. 2021, 11, e686120. [Google Scholar] [CrossRef]
  21. Crous, P.W.; Schumacher, R.K.; Akulov, A.; Thangavel, R.; Hernández-Restrepo, M.; Carnegie, A.J.; Cheewangkoon, R.; Wingfield, M.J.; Summerell, B.A.; Quaedvlieg, W. New and interesting fungi. 2. Fungal Syst. Evol. 2019, 3, 57–134. [Google Scholar] [CrossRef]
  22. Crous, P.W.; Schumacher, R.K.; Wingfield, M.J.; Akulov, A.; Denman, S.; Roux, J.; Braun, U.; Burgess, T.I.; Carnegie, A.J.; Váczy, K.Z. New and interesting fungi. 1. Fungal. Syst. Evol. 2018, 1, 169–216. [Google Scholar] [CrossRef] [PubMed]
  23. Crous, P.W.; Schubert, K.; Braun, U.; Hoog, G.S.; de Hocking, A.D.; Shin, H.D.; Groenewald, J.Z. Opportunistic, human-pathogenic species in the Herpotrichiellaceae are phenotypically similar to saprobic or phytopathogenic species in the Venturiaceae. Stud. Mycol. 2007, 58, 185–217. [Google Scholar] [CrossRef] [PubMed]
  24. Tibpromma, S.; Hyde, K.D.; Jeewon, R.; Maharachchikumbura, S.S.N.; Liu, J.K.; Bhat, D.J.; Jones, E.B.G.; McKenzie, E.H.C.; Camporesi, E.; Bulgakov, T.S.; et al. Fungal diversity notes 491–602: Taxonomic and phylogenetic contributions to fungal taxa. Fungal Divers. 2017, 83, 1–261. [Google Scholar] [CrossRef]
  25. Estévez, E.; Veiga, M.C.; Kennes, C. Biodegradation of toluene by the new fungal isolates Paecilomyces variotii and Exophiala oligosperma. J. Ind. Microbiol. Biotechnol. 2005, 32, 33–37. [Google Scholar] [CrossRef] [Green Version]
  26. Libert, X.; Chasseur, C.; Packeu, A.; Bureau, F.; Roosens, N.H.; De Keersmaecker, S.C.J. Exploiting the advantages of molecular tools for the monitoring of fungal indoor air contamination: First detection of Exophiala jeanselmei in indoor air of air-conditioned offices. Microorganisms 2019, 7, e674. [Google Scholar] [CrossRef] [Green Version]
  27. de Hoog, G.S.; Zeng, J.S.; Harrak, M.J.; Sutton, D.A. Exophiala xenobiotica sp. nov., an opportunistic black yeast inhabiting environments rich in hydrocarbons. Antonie van Leeuwenhoek 2006, 90, 257–268. [Google Scholar] [CrossRef]
  28. Isola, D.; Selbmann, L.; de Hoog, G.S.; Fenice, M.; Onofri, S.; Prenafeta-Boldú, F.X.; Zucconi, L. Isolation and screening of black fungi as degraders of volatile aromatic hydrocarbons. Mycopathologia 2013, 175, 369–379. [Google Scholar] [CrossRef]
  29. Listemann, H.; Freiesleben, H. Exophiala mesophila spec. nov. Mycoses 1996, 39, 1–3. [Google Scholar] [CrossRef]
  30. Li, D.M.; Li, R.Y.; de Hoog, G.S.; Wang, Y.X.; Wang, D.L. Exophiala asiatica, a new species from a fatal case in China. Med. Mycol. 2009, 47, 101–109. [Google Scholar] [CrossRef] [Green Version]
  31. Woo, P.C.; Ngan, A.H.; Tsang, C.C.; Ling, I.W.; Chan, J.F.; Leung, S.Y.; Yuen, K.Y.; Lau, S.K. Clinical spectrum of Exophiala infections and a novel Exophiala species, Exophiala hongkongensis. J. Clin. Microbiol. 2013, 51, 260–267. [Google Scholar] [CrossRef] [Green Version]
  32. Li, D.M.; Li, R.Y.; de Hoog, G.S.; Sudhadham, M.; Wang, D.L. Fatal Exophiala infections in China, with a report of seven cases. Mycoses 2011, 54, 136–142. [Google Scholar] [CrossRef] [PubMed]
  33. Vitale, R.G.; de Hoog, G.S. Molecular diversity, new species and antifungal susceptibilities in the Exophiala spinifera clade. Med. Mycol. 2002, 40, 545–556. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Zeng, J.; Feng, P.; Gerrits van den Ende, A.H.G.; Xi, L.; Harrak, M.J.; de Hoog, G.S. Multilocus analysis of the Exophiala jeanselmei clade containing black yeasts involved in opportunistic disease in humans. Fungal Divers. 2014, 65, 3–16. [Google Scholar] [CrossRef]
  35. Barba-Gómez, J.F.; Mayorga, J.; McGinnis, M.R.; González-Mendoza, A. Chromoblastomycosis caused by Exophiala spinifera. J. Am. Acad. Derm. 1992, 26, 367–370. [Google Scholar] [CrossRef]
  36. Padhye, A.A.; Kaplan, W.; Neuman, M.A.; Case, P.; Radcliffe, G.N. Subcutaneous phaeohyphomycosis caused by Exophiala spinifera. J. Med. Vet. Mycol. 1984, 22, 493–500. [Google Scholar] [CrossRef]
  37. Padhye, A.A.; Hampton, A.A.; Hampton, M.T.; Hutton, N.W.; Prevost-Smith, E.; Davis, M.S. Chromoblastomycosis caused by Exophiala spinifera. Clin. Infect. Dis. 1996, 22, 331–335. [Google Scholar] [CrossRef] [Green Version]
  38. Lacaz, C.S.; Porto, E.; Andrade, J.G.; Telles Filho, F.Q. Feohifomicose disseminada por Exophiala spinifera. An. Bras. Derm. 1984, 59, 238–243. [Google Scholar]
  39. Rajam, R.V.; Kandhari, K.C.; Thirumalachar, M. Chromoblastomycosis caused by a rare yeast-like dematiaceous fungus. Mycopathol. Mycol. Appl. 1958, 9, 5–19. [Google Scholar] [CrossRef]
  40. Nielsen, H.S.; Conant, N.F. A new pathogenic Phialophora. Sabouraudia 1968, 6, 228–231. [Google Scholar] [CrossRef]
  41. Kettlewell, P.; McGinnis, M.R.; Wilkinson, G.T. Phaeohyphomycosis caused by Exophiala spinifera in two cats. J. Med. Vet. Mycol. 1989, 27, 257–264. [Google Scholar] [CrossRef]
  42. Conti-Díaz, I.A.; MacKinnon, J.E.; Civila, E. Isolation and identification of black yeasts from the external environment in Uruguay. Pan Am. Health Org. Sci. Publ. 1977, 356, 109–114. [Google Scholar]
  43. Song, Y.; Laureijssen-van de Sande, W.W.J.; Moreno, L.F.; Gerrits van den Ende, B.; Li, R.; de Hoog, S. Comparative ecology of capsular Exophiala species causing disseminated infection in humans. Front. Microbiol. 2017, 8, e2514. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Arenz, B.E.; Held, B.W.; Jurgens, J.A.; Farrell, R.L.; Blanchette, R.A. Fungal diversity in soils and historic wood from the Ross Sea Region of Antarctica. Soil Biol. Biochem. 2006, 38, 3057–3064. [Google Scholar] [CrossRef]
  45. Alimu, Y.; Ban, S.; Yaguchi, T. Molecular phylogenetic study of strains morphologically identified as Exophiala dermatitidis from clinical and environmental specimens in Japan. Med. Mycol. J. 2022, 63, 1–9. [Google Scholar] [CrossRef] [PubMed]
  46. Sudhadham, M.; Prakitsin, S.; Sivichai, S.; Chaiyarat, R.; Dorrestein, G.M.; Menken, S.B.; de Hoog, G.S. The neurotropic black yeast Exophiala dermatitidis has a possible origin in the tropical rain forest. Stud. Mycol. 2008, 61, 145–155. [Google Scholar] [CrossRef]
  47. Döğen, A.; Kaplan, E.; Ilkit, M.; de Hoog, G.S. Massive contamination of Exophiala dermatitidis and E. phaeomuriformis in railway stations in subtropical Turkey. Mycopathologia 2013, 175, 381–386. [Google Scholar] [CrossRef]
  48. Gümral, R.; Tümgör, A.; Saraçlı, M.A.; Yıldıran, Ş.T.; Ilkit, M.; de Hoog, G.S. Black yeast diversity on creosoted railway sleepers changes with ambient climatic conditions. Microb. Ecol. 2014, 68, 699–707. [Google Scholar] [CrossRef]
  49. Zupančič, J.; Novak Babič, M.; Zalar, P.; Gunde-Cimerman, N. The black yeast Exophiala dermatitidis and other selected opportunistic human fungal pathogens spread from dishwashers to kitchens. PLoS ONE 2016, 11, e0148166. [Google Scholar] [CrossRef]
  50. Yazdanparast, S.A.; Mohseni, S.; de Hoog, G.S.; Aslani, N.; Sadeh, A.; Badali, H. Consistent high prevalence of Exophiala dermatitidis, a neurotropic opportunist, on railway sleepers. J. Mycol. Med. 2017, 27, 180–187. [Google Scholar] [CrossRef]
  51. Sav, H.; Ozakkas, F.; Altınbas, R.; Kiraz, N.; Tümgör, A.; Gümral, R.; Döğen, A.; Ilkit, M.; de Hoog, G.S. Virulence markers of opportunistic black yeast in Exophiala. Mycoses 2016, 59, 343–350. [Google Scholar] [CrossRef]
  52. Jayaram, M.; Nagao, H. First report of environmental isolation of Exophiala spp. in Malaysia. Curr. Microbiol. 2020, 77, 2915–2924. [Google Scholar] [CrossRef] [PubMed]
  53. Klasinc, R.; Riesenhuber, M.; Bacher, A.; Willinger, B. Invasive fungal infection caused by Exophiala dermatitidis in a patient after lung transplantation: Case report and literature review. Mycopathologia 2019, 184, 107–113. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Naka, W.; Harada, T.; Nishikawa, T.; Fukushiro, R. A case of chromoblastomycosis: With special reference to the mycology of the isolated Exophiala jeanselmei. Mykosen 1986, 29, 445–452. [Google Scholar] [CrossRef] [PubMed]
  55. Langeron, M. Mycétome à Torula jeanselmei Langeron, 1928. Nouveau type de mycétome à grains noirs. Ann. Parasitol. Hum. Comp. 1928, 6, 385–403. [Google Scholar] [CrossRef] [Green Version]
  56. Murray, I.G.; Dunkerley, G.E.; Hughes, K.E.A. A case of madura foot caused by Phialophora jeanselmei. Sabouraudia 1964, 3, 175–177. [Google Scholar] [CrossRef] [PubMed]
  57. Noguchi, H.; Matsumoto, T.; Kimura, U.; Hiruma, M.; Kano, R.; Yaguchi, T.; Kubo, M.; Kashiwada-Nakamura, K.; Fukushima, S. Empiric antifungal therapy in patients with cutaneous and subcutaneous phaeohyphomycosis. J. Dermatol. 2022, 49, 564–571. [Google Scholar] [CrossRef]
  58. Pattanaprichakul, P.; Bunyaratavej, S.; Leeyaphan, C.; Sitthinamsuwan, P.; Sudhadham, M.; Muanprasart, C.; Feng, P.; Badali, H.; de Hoog, G.S. An unusual case of eumycetoma caused by Exophiala jeanselmei after a sea urchin injury. Mycoses 2013, 56, 491–494. [Google Scholar] [CrossRef]
  59. Valiente, C.; Quesada, E. Morphologic and physiologic characteristics of Costa Rica pathogenic fungi (Dermatiaceae). Rev. Biol. Trop. 1991, 39, 103–106. [Google Scholar]
  60. Ramírez Soto, M.C.; Malaga, G. Subcutaneous mycoses in Peru: A systematic review and meta-analysis for the burden of disease. Int. J. Dermatol. 2017, 56, 1037–1045. [Google Scholar] [CrossRef]
  61. Al-Tawfiq, J.A.; Amr, S.S. Madura leg due to Exophiala jeanselmei successfully treated with surgery and itraconazole therapy. Med. Mycol. 2009, 47, 648–652. [Google Scholar] [CrossRef] [Green Version]
  62. Thammayya, A.; Sanyal, M. Exophiala jeanselmei causing mycetoma pedis in India. Sabouraudia 1980, 18, 91–95. [Google Scholar] [CrossRef] [PubMed]
  63. Brownell, I.; Pomeranz, M.; Ma, L. Eumycetoma. Dermatol. Online J. 2005, 11, e10. [Google Scholar] [CrossRef]
  64. Nielsen Jr, H.S.; Conant, N.F.; Weinberg, T.; Reback, J.F. Report of a mycetoma due to Phialophora jeanselmei and undescribed characteristics of the fungus. Saboraudia 1968, 6, 330–333. [Google Scholar] [CrossRef]
  65. Negroni, R. Estudio micologico del primer caso de micetoma por Phialophora jeanselmei observado en la Argentina. Med. Cut. Iberolat. 1970, 5, 625–630. [Google Scholar]
  66. Simpson, A.; Singh, S.R. A case of Madura foot. J. Roy. College. Surg. Edinburgh 1984, 29, 326–328. [Google Scholar]
  67. Badali, H.; Najafzadeh, M.J.; van Esbroeck, M.; van den Enden, E.; Tarazooie, B.; Meis, J.F.; de Hoog, G.S. The clinical spectrum of Exophiala jeanselmei, with a case report and in vitro antifungal susceptibility of the species. Med. Mycol. 2010, 48, 318–327. [Google Scholar] [CrossRef] [Green Version]
  68. Seifert, K.A.; Okada, G. Graphium anamorphs of Ophiostoma species and similar anamorphs of other Ascomycetes. In Ceratocystis and Ophiostoma: Taxonomy, Ecology, and Pathogenicity; Wingfield, M.J., Seifert, K.A., Webber, J.F., Eds.; The American Phytopathological Society Press: St. Paul, MN, USA, 1993; pp. 27–41. [Google Scholar]
  69. Isola, D.; Zucconi, L.; Onofri, S.; Caneva, G.; de Hoog, G.S.; Selbmann, L. Extremotolerant rock inhabiting black fungi from Italian monumental sites. Fungal Divers. 2016, 76, 75–96. [Google Scholar] [CrossRef]
  70. Borman, A.M.; Fraser, M.; Schilling, W.; Jones, G.; Pearl, R.; Linton, C.J.; Johnson, E.M. Exophiala campbellii causing a subcutaneous palmar cyst in an otherwise healthy UK resident. Med. Mycol. Case Rep. 2020, 29, 43–45. [Google Scholar] [CrossRef]
  71. Madrid, H.; Hernández-Restrepo, M.; Gené, J.; Cano, J.; Guarro, J.; Silva, V. New and interesting chaetothyrialean fungi from Spain. Mycol. Prog. 2016, 15, 1179–1201. [Google Scholar] [CrossRef] [Green Version]
  72. Crous, P.W.; Wingfield, M.J.; Burgess, T.I.; Hardy, G.; Gené, J.; Guarro, J.; Baseia, I.G.; García, D.; Gusmão, L.; Souza-Motta, C.M.; et al. Fungal planet description sheets: 716–784. Persoonia 2018, 40, 240–393. [Google Scholar] [CrossRef]
  73. Crous, P.W.; Wingfield, M.J.; Schumacher, R.K.; Akulov, A.; Bulgakov, T.S.; Carnegie, A.J.; Jurjević, Ž.; Decock, C.; Denman, S.; Lombard, L.; et al. New and interesting fungi. 3. Fungal Syst. Evol. 2020, 6, 157–231. [Google Scholar] [CrossRef] [PubMed]
  74. Kochkina, G.A.; Ivanushkina, N.E.; Lupachev, A.V.; Starodumova, I.P.; Vasilenko, O.V.; Ozerskaya, S.M. Diversity of mycelial fungi in natural and human-affected Antarctic soils. Polar Biol. 2019, 42, 47–64. [Google Scholar] [CrossRef]
  75. Kondratiuk, T.O.; Kondratyuk, S.Y.; Khimich, M.V.; Beregova, T.V.; Ostapchenko, L.I. Confirmation of taxonomic status of black yeast-like fungus by three gene phylogeny. Acta Bot. Hung. 2016, 58, 287–302. [Google Scholar] [CrossRef] [Green Version]
  76. Nishimura, K.; Miyaji, M.; Taguchi, H.; Tanaka, R. Fungi in bathwater and sludge of bathroom drainpipes. 1. Frequent isolation of Exophiala species. Mycopathologia 1987, 97, 17–23. [Google Scholar] [CrossRef]
  77. Nyaoke, A.; Weber, E.S.; Innis, C.; Stremme, D.; Dowd, C.; Hinckley, L.; Gorton, T.; Wickes, B.; Sutton, D.; de Hoog, S. Disseminated phaeohyphomycosis in weedy seadragons (Phyllopteryx taeniolatus) and leafy seadragons (Phycodurus eques) caused by species of Exophiala, including a novel species. J. Vet. Diagn. Investig. 2009, 21, 69–79. [Google Scholar] [CrossRef] [Green Version]
  78. Saraiva, M.; Beckmann, M.J.; Pflaum, S.; Pearson, M.; Carcajona, D.; Treasurer, J.W.; West, P.V. Exophiala angulospora infection in hatchery-reared lumpfish (Cyclopterus lumpus) broodstock. J. Fish Dis. 2019, 42, 335–343. [Google Scholar] [CrossRef] [PubMed]
  79. Scholz, F.; Ruane, N.M.; Marcos-López, M.; Mitchell, S.; Bolton-Warberg, M.; O’Connor, I.; Mirimin, L.; MacCarthy, E.; Rodger, H.D. Systemic mycoses in lumpfish (Cyclopterus lumpus L.) in Ireland: Aetiology and clinical presentation. Bull. Eur. Ass. Fish Pathol. 2018, 38, 202–212. [Google Scholar]
  80. Kanchan, C.; MuraosaKishio, Y.; Hatai, H. Exophiala angulospora infection found in cultured Japanese flounder Paralichthys olivaceus in Japan. Bull. Eur. Ass. Fish Pathol. 2014, 34, 187–194. [Google Scholar]
  81. Gjessing, M.C.; Davey, M.; Kvellestad, A.; Vrålstad, T. Exophiala angulospora causes systemic inflammation in Atlantic cod Gadus morhua. Dis. Aquat. Organ. 2011, 96, 209–219. [Google Scholar] [CrossRef] [Green Version]
  82. Chermette, R.; Ferreiro, L.; De Bievre, C.; Camadro, J.P.; Mialot, M.; Vauzelle, P. Exophiala spinifera nasal infection in a cat and a literature review of feline phaeohyphomycosis. J. Mycol. Med. 1997, 7, 149–158. [Google Scholar]
  83. Bernhardt, A.; Bomhard, W.V.; Antweiler, E.; Tintelnot, K. Molecular identification of fungal pathogens in nodular skin lesions of cats. Med. Mycol. 2015, 53, 132–144. [Google Scholar] [CrossRef] [PubMed]
  84. Overy, D.P.; Martin, C.; Muckle, A.; Lund, L.; Wood, J.; Hanna, P. Cutaneous phaeohyphomycosis caused by Exophiala attenuata in a domestic cat. Mycopathologia 2015, 180, 281–287. [Google Scholar] [CrossRef] [PubMed]
  85. Silva, W.C.; Gonçalves, S.S.; Santos, D.W.; Padovan, A.C.; Bizerra, F.C.; Melo, A.S. Species diversity, antifungal susceptibility and phenotypic and genotypic characterisation of Exophiala spp. infecting patients in different medical centres in Brazil. Mycoses 2017, 60, 328–337. [Google Scholar] [CrossRef] [PubMed]
  86. Rakeman, J.L.; Bui, U.; LaFe, K.; Chen, Y.C.; Honeycutt, R.J.; Cookson, B.T. Multilocus DNA sequence comparisons rapidly identify pathogenic molds. J. Clin. Microbiol. 2005, 43, 3324–3333. [Google Scholar] [CrossRef] [Green Version]
  87. Figel, I.C.; Marangoni, P.R.; Tralamazza, S.M.; Vicente, V.A.; Dalzoto, P.; do Nascimento, M.M.; de Hoog, G.S.; Pimentel, I.C. Black yeasts-like fungi isolated from dialysis water in hemodialysis units. Mycopathologia 2013, 175, 413–420. [Google Scholar] [CrossRef]
  88. Vicente, V.A.; Orélis-Ribeiro, R.; Najafzadeh, M.J.; Sun, J.; Guerra, R.S.; Miesch, S.; Ostrensky, A.; Meis, J.F.; Klaassen, C.H.; de Hoog, G.S.; et al. Black yeast-like fungi associated with Lethargic Crab Disease (LCD) in the mangrove-land crab, Ucides cordatus (Ocypodidae). Vet. Microbiol. 2012, 158, 109–122. [Google Scholar] [CrossRef]
  89. Crous, P.W.; Groenewald, J.Z. Why everlastings don’t last. Persoonia 2011, 26, 70–84. [Google Scholar] [CrossRef]
  90. Bates, S.T.; Reddy, G.; Garcia-Pichel, F. Exophiala crusticola anam. nov. (affinity Herpotrichiellaceae), a novel black yeast from biological soil crusts in the Western United States. Int. J. Syst. Evol. Microbiol. 2006, 56, 2697–2702. [Google Scholar] [CrossRef] [Green Version]
  91. Katz, B.; McGinnis, R. A new species of Exophiala recovered from loblolly pine litter. Mycotaxon 1980, 11, 182–184. [Google Scholar]
  92. Crous, P.W.; Cowan, D.A.; Maggs-Kölling, G.; Yilmaz, N.; Larsson, E.; Angelini, C.; Brandrud, T.E.; Dearnaley, J.; Dima, B.; Dovana, F.; et al. Fungal Planet description sheets: 1112–1181. Persoonia 2020, 45, 251–409. [Google Scholar] [CrossRef]
  93. Persoonial Reflections. Pers. Mol. Phylogeny Evol. Fungi 2010, 25, 117–159.
  94. Ávila, A.; Groenewald, J.Z.; Trapero, A.; Crous, P.W. Characterization and epitypification of Pseudocercospora cladosporioides, the causal organism of Cercospora-leaf spot of olives. Mycol. Res. 2005, 109, 881–888. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Pollacci, G. Miceti del corpo umano e degli animali. Atti dell’Istituto Botanico Università di Pavia 1923, 18, 1–9. [Google Scholar]
  96. Manharth, A.; Lemberger, K.; Mylniczenko, N.; Pinkerton, M.; Pessier, A.P.; Kammeyer, P.; de Hoog, S. Disseminated Phaeohyphomycosis due to Exophiala species in a Galapagos tortoise, Geochelone nigra. J. Herpetol. Med. Surg. 2005, 15, 20–26. [Google Scholar] [CrossRef] [Green Version]
  97. Neubert, K.; Mendgen, K.; Brinkmann, H.; Wirsel, S.G.R. Only a few fungal species dominate highly diverse mycofloras associated with the common reed. Appl. Environ. Microbiol. 2006, 72, 1118–1128. [Google Scholar] [CrossRef] [Green Version]
  98. Najafzadeh, M.J.; Suh, M.K.; Lee, M.H.; Ha, G.Y.; Kim, J.R.; Kim, T.H.; Lee, H.J.; Choi, J.S.; Meis, J.F.; de Hoog, G.S. Subcutaneous phaeohyphomycosis caused by Exophiala equina, with susceptibility to eight antifungal drugs. J. Med. Microbiol. 2013, 62, 797–800. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  99. Crous, P.W.; Wingfield, M.J.; Burgess, T.I.; Hardy, G.E.; Crane, C.; Barrett, S.; Cano-Lira, J.F.; Le Roux, J.J.; Thangavel, R.; Guarro, J.; et al. Fungal planet description sheets: 469–557. Persoonia 2016, 37, 218–403. [Google Scholar] [CrossRef]
  100. Crous, P.W.; Wingfield, M.J.; Chooi, Y.H.; Gilchrist, C.; Lacey, E.; Pitt, J.I.; Roets, F.; Swart, W.J.; Cano-Lira, J.F.; Valenzuela-Lopez, N.; et al. Fungal planet description sheets: 1042–1111. Persoonia 2020, 44, 301–459. [Google Scholar] [CrossRef]
  101. Benedek, T.; Specht, G. Mykologisch-bakteriologische Untersuchungen über Pilze und Bakterien als Symbionten bei Kerbtieren. Zentbl. f. Bakt. Parasitenk. Infekt. Abt. I Orig. 1933, 130, 74–90. [Google Scholar]
  102. Lian, X.; de Hoog, G.S. Indoor wet cells harbour melanized agents of cutaneous infection. Med. Mycol. 2010, 48, 622–628. [Google Scholar] [CrossRef] [Green Version]
  103. McGinnis, M.R.; Sorrell, D.F.; Miller, R.L.; Kaminski, G.W. Subcutaneous phaeohyphomycosis caused by Exophiala moniliae. Mycopathologia 1981, 73, 69–72. [Google Scholar] [CrossRef] [PubMed]
  104. Tintelnot, K.; de Hoog, G.S.; Thomas, E.; Steudel, W.I.; Huebner, K.; Seeliger, H.P.R. Cerebral phaeohyphomycosis caused by an Exophiala species. Mycoses 1991, 34, 239–244. [Google Scholar] [CrossRef]
  105. Nucci, M.; Akiti, T.; Barreiros, G.; Silveira, F.; Revankar, S.G.; Sutton, D.A.; Patterson, T.F. Nosocomial fungemia due to Exophiala jeanselmei var. jeanselmei and a Rhinocladiella species: Newly described causes of bloodstream infection. J. Clin. Microbiol. 2001, 39, 514–518. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  106. Bossler, A.D.; Richter, S.S.; Chavez, A.J.; Vogelgesang, S.A.; Sutton, D.A.; Grooters, A.M.; Rinaldi, M.G.; de Hoog, G.S.; Pfaller, M.A. Exophiala oligosperma causing olecranon bursitis. J. Clin. Microbiol. 2003, 40, 4779–4782. [Google Scholar] [CrossRef] [Green Version]
  107. Nascimento, M.M.F.; Vicente, V.A.; Bittencourt, J.V.M.; Gelinski, J.M.L.; Prenafeta-Boldú, F.X.; Romero-Güiza, M.; Fornari, G.; Gomes, R.R.; Santos, G.D.; Gerrits Van Den Ende, A.H.G.; et al. Diversity of opportunistic black fungi on babassu coconut shells, a rich source of esters and hydrocarbons. Fungal Biol. 2017, 121, 488–500. [Google Scholar] [CrossRef] [PubMed]
  108. Gold, W.L.; Vellend, H.; Salit, I.E.; Campbell, I.; Summerbell, R.; Rinaldi, M.; Simor, A.E. Successful treatment of systemic and local infections due to Exophiala species. Clin. Infect. Dis. 1994, 19, 339–341. [Google Scholar] [CrossRef] [PubMed]
  109. Harsonowati, W.; Marian, M.; Surono, S.; Narisawa, K. The effectiveness of a dark septate endophytic fungus, Cladophialophora chaetospira SK51, to mitigate strawberry Fusarium-wilt disease and with growth promotion activities. Front. Microbiol. 2020, 11, e585. [Google Scholar] [CrossRef] [Green Version]
  110. Crous, P.W.; Cowan, D.A.; Maggs-Kölling, G.; Yilmaz, N.; Thangavel, R.; Wingfield, M.J.; Noordeloos, M.E.; Dima, B.; Brandrud, T.E.; Jansen, G.M.; et al. Fungal planet description sheets: 1182–1283. Persoonia 2021, 46, 313–528. [Google Scholar] [CrossRef]
  111. Crous, P.W.; Groenewald, J.Z.; Shivas, R.G.; Edwards, J.; Seifert, K.A.; Alfenas, A.C.; Alfenas, R.F.; Burgess, T.I.; Carnegie, A.J.; Hardy, G.E.; et al. Fungal planet description sheets: 69–91. Persoonia 2011, 26, 108–156. [Google Scholar] [CrossRef]
  112. Scopus Database. Available online: https://www.scopus.com (accessed on 9 May 2022).
  113. Xu, R.; Li, T.; Shen, M.; Yang, Z.L.; Zhao, Z.W. Evidence for a dark septate endophyte (Exophiala pisciphila, H93) enhancing phosphorus absorption by maize seedlings. Plant Soil 2020, 452, 249–266. [Google Scholar] [CrossRef]
  114. Xiao, Y.; Dai, M.X.; Zhang, G.Q.; Yang, Z.X.; He, Y.M.; Zhan, F.D. Effects of the dark septate endophyte (DSE) Exophiala pisciphila on the growth of root cell wall polysaccharides and the cadmium content of Zea mays L. under cadmium stress. J. Fungi 2021, 7, e1035. [Google Scholar] [CrossRef] [PubMed]
  115. Ondeyka, J.G.; Zink, D.L.; Dombrowski, A.W.; Polishook, J.D.; Felock, P.J.; Hazuda, D.J.; Singh, S.B. Isolation, structure and HIV-1 integrase inhibitory activity of exophillic acid, a novel fungal metabolite from Exophiala pisciphila. J. Antibiot. 2003, 56, 1018–1023. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  116. Cheikh-Ali, Z.; Glynou, K.; Ali, T.; Ploch, S.; Kaiser, M.; Thines, M.; Bode, H.B.; Maciá-Vicente, J.G. Diversity of exophillic acid derivatives in strains of an endophytic Exophiala sp. Phytochemistry 2015, 118, 83–93. [Google Scholar] [CrossRef]
  117. Zhang, D.; Yang, X.; Kang, J.S.; Choi, H.D.; Son, B.W. Chlorohydroaspyrones A and B, antibacterial aspyrone derivatives from the marine-derived fungus Exophiala sp. J. Nat. Prod. 2008, 71, 1458–1460. [Google Scholar] [CrossRef] [PubMed]
  118. Doshida, J.; Hasegawa, H.; Onuki, H.; Shimidzu, N. Exophilin A, a new antibiotic from a marine microorganism Exophiala pisciphila. J. Antibiot. 1996, 49, 1105–1109. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  119. Ide-Pérez, M.R.; Fernández-López, M.G.; Sánchez-Reyes, A.; Leija, A.; Batista-García, R.A.; Folch-Mallol, J.L.; Sánchez-Carbente, M.d.R. Aromatic hydrocarbon removal by novel extremotolerant Exophiala and Rhodotorula spp. from an oil polluted site in Mexico. J. Fungi. 2020, 6, e135. [Google Scholar] [CrossRef]
  120. Hyde, K.D.; Norphanphoun, C.; Chen, J.; Dissanayake, A.J.; Doilom, M.; Hongsanan, S.; Jayawardena, R.S.; Jeewon, R.; Perera, R.H.; Thongbai, B.; et al. Thailand’s amazing diversity: Up to 96% of fungi in northern Thailand may be novel. Fungal Divers. 2018, 93, 215–239. [Google Scholar] [CrossRef]
  121. Khuna, S.; Suwannarach, N.; Kumla, J.; Frisvad, J.C.; Matsui, K.; Nuangmek, W.; Lumyong, S. Growth enhancement of Arabidopsis (Arabidopsis thaliana) and onion (Allium cepa) with inoculation of three newly identified mineral-solubilizing fungi in the genus Aspergillus section Nigri. Front. Microbiol. 2021, 12, e705896. [Google Scholar] [CrossRef]
  122. Boonmee, S.; Wanasinghe, D.N.; Calabon, M.S.; Huanraluek, N.; Chandrasiri, A.K.U.; Jones, G.E.B.; Rossi, W.; Leonardi, M.; Singh, S.K.; Rana, S.; et al. Fungal diversity notes 1387–1511: Taxonomic and phylogenetic contributions on genera and species of fungal taxa. Fungal Divers. 2021, 111, 1–335. [Google Scholar] [CrossRef]
  123. Selbmann, L.; Isola, D.; Egidi, E.; Zucconi, L.; Gueidan, C.; de Hoog, G.S.; Onofri, S. Mountain tips as reservoirs for new rock-fungal entities: Saxomyces gen. nov. and four new species from the Alps. Fungal Divers. 2014, 65, 167–182. [Google Scholar] [CrossRef] [Green Version]
  124. Voigt, K.; Wostemeyer, J. Reliable amplification of actin genes facilitates deep-level phylogeny. Microbiol. Res. 2000, 155, 179–195. [Google Scholar] [CrossRef]
  125. White, T.J.; Bruns, T.; Lee, S.; Taylor, J.W. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In PCR Protocols: A Guide to Methods and Applications; Innes, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J., Eds.; Academic Press, Inc.: San Diego, CA, USA, 1990; pp. 315–322. [Google Scholar]
  126. Carbone, I.; Kohn, L.M. A method for designing primer sets for speciation studies in filamentous ascomycetes. Mycologia 1999, 91, 553–556. [Google Scholar] [CrossRef]
  127. Glass, N.L.; Donaldson, G.C. Development of primer sets designed for use with the PCR to amplify conserved genes from filamentous ascomycetes. Appl. Environ. Microbiol. 1995, 61, 1323–1330. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  128. Edgar, R.C. MUSCLE: A multiple sequence alignment method with reduced time and space complexity. BMC Bioinform. 2004, 5, e113. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  129. Hall, T. Bioedit Version 6.0.7. 2004. Available online: http://www.mbio.ncsu.edu/bioedit/bioedit.html (accessed on 20 November 2021).
  130. Zeng, J.S.; Sutton, D.A.; Fothergill, A.W.; Rinaldi, M.G.; Harrak, M.J.; de Hoog, G.S. Spectrum of clinically relevant Exophiala species in the United States. J. Clin. Microbiol. 2007, 45, 3713–3720. [Google Scholar] [CrossRef] [Green Version]
  131. Gueidan, C.; Villaseñor, C.R.; de Hoog, G.S.; Gorbushina, A.A.; Untereiner, W.A.; Lutzoni, F. A rock-inhabiting ancestor for mutualistic and pathogen-rich fungal lineages. Stud. Mycol. 2008, 61, 111–119. [Google Scholar] [CrossRef]
  132. Okada, G.; Seifert, K.A.; Takematsu, A.; Yamaoka, Y.; Miyazaki, S.; Tubaki, K. A molecular phylogenetic reappraisal of the Graphium complex based on 18 S rDNA sequences. Can. J. Bot. 1998, 76, 1495–1506. [Google Scholar] [CrossRef]
  133. Attili-Angelis, D.; Duarte AP, M.; Pagnocca, F.C.; Nagamoto, N.S.; De Vries, M.; Stielow, J.B.; de Hoog, G.S. Novel Phialophora species from leaf-cutting ants (tribe Attini). Fungal Divers. 2014, 65, 65–75. [Google Scholar] [CrossRef]
  134. Vicente, V.A.; Najafzadeh, M.J.; Sun, J.; Gomes, R.R.; Robl, D.; Marques, S.G.; Azevedo, C.M.P.S.; de Hoog, G.S. Environmental siblings of black agents of human chromoblastomycosis. Fungal Divers. 2014, 65, 47–63. [Google Scholar] [CrossRef]
  135. Untereiner, W.A.; Naveau, F.A. Molecular systematics of the Herpotrichiellaceae with an assessment of the phylogenetic positions of Exophiala dermatitidis and Phialophora americana. Mycologia 1999, 91, 67–83. [Google Scholar] [CrossRef]
  136. Vu, D.; Groenewald, M.; de Vries, M.; Gehrmann, T.; Stielow, B.; Eberhardt, U.; Al-Hatmi, A.; Groenewald, J.Z.; Cardinali, G.; Houbraken, J.; et al. Large-scale generation and analysis of filamentous fungal DNA barcodes boosts coverage for kingdom fungi and reveals thresholds for fungal species and higher taxon delimitation. Stud. Mycol. 2019, 92, 135–154. [Google Scholar] [CrossRef] [PubMed]
  137. Réblová, M.; Untereiner, W.A.; Réblová, K. Novel evolutionary lineages revealed in the Chaetothyriales (fungi) based on multigene phylogenetic analyses and comparison of its secondary structure. PLoS ONE 2013, 8, e63547. [Google Scholar] [CrossRef] [PubMed]
  138. Vasse, M.; Voglmayr, H.; Mayer, V.; Gueidan, C.; Nepel, M.; Moreno, L.; de Hoog, S.; Selosse, M.A.; McKey, D.; Blatrix, R. A phylogenetic perspective on the association between ants (Hymenoptera: Formicidae) and black yeasts (Ascomycota: Chaetothyriales). Proc. R. Soc. B 2017, 284, e20162519. [Google Scholar] [CrossRef] [PubMed]
  139. Prenafeta-Boldú, F.X.; Summerbell, R.; de Hoog, G.S. Fungi growing on aromatic hydrocarbons: Biotechnology’s unexpected encounter with biohazard? FEMS Microbiol. Rev. 2006, 30, 109–130. [Google Scholar] [CrossRef] [Green Version]
  140. Tischner, Z.; Sebők, R.; Kredics, L.; Allaga, H.; Vargha, M.; Sebestyén, Á.; Dobolyi, C.; Kriszt, B.; Magyar, D. Mycological investigation of bottled water dispensers in healthcare facilities. Pathogens 2021, 10, e871. [Google Scholar] [CrossRef]
  141. Haase, G.; Sonntag, L.; van de Peer, Y.; Uijthof, J.M.; Podbielski, A.; Melzer-Krick, B. Phylogenetic analysis of ten black yeast species using nuclear small subunit rRNA gene sequences. Antonie van Leeuwenhoek 1995, 68, 19–33. [Google Scholar] [CrossRef]
  142. Schoch, C.L.; Robbertse, B.; Robert, V.; Vu, D.; Cardinali, G.; Irinyi, L.; Meyer, W.; Nilsson, R.H.; Hughes, K.; Miller, A.N.; et al. Finding needles in haystacks: Linking scientific names, reference specimens and molecular data for fungi. Database 2014, 2014, bau061. [Google Scholar] [CrossRef]
  143. Cheewangkoon, R.; Groenewald, J.Z.; Summerell, B.A.; Hyde, K.D.; To-Anun, C.; Crous, P.W. Myrtaceae, a cache of fungal biodiversity. Persoonia 2009, 23, 55–85. [Google Scholar] [CrossRef] [Green Version]
  144. Crous, P.W.; Braun, U.; Hunter, G.C.; Wingfield, M.J.; Verkley, G.J.; Shin, H.D.; Nakashima, C.; Groenewald, J.Z. Phylogenetic lineages in Pseudocercospora. Stud. Mycol. 2013, 75, 37–114. [Google Scholar] [CrossRef] [Green Version]
  145. Felsenstein, J. Confidence intervals on phylogenetics: An approach using bootstrap. Evolution 1985, 39, 783–791. [Google Scholar] [CrossRef]
  146. Stamatakis, A. Raxml-vi-hpc: Maximum likelihood-based phylogenetic analyses with thousands of taxa and mixed models. Bioinformatics 2006, 22, 2688–2690. [Google Scholar] [CrossRef] [PubMed]
  147. Miller, M.A.; Holder, M.T.; Vos, R.; Midford, P.E.; Liebowitz, T.; Chan, L.; Hoover, P.; Warnow, T. The CIPRES Portals. CIPRES, 2009. Available online: http://www.phylo.org/sub_sections/portal (accessed on 4 April 2022).
  148. Nylander, J.A.A. MrModeltest 2.0; Program Distributed by the Author; Evolutionary Biology Centre, Uppsala University: Uppsala, Sweden, 2004. [Google Scholar]
  149. Ronquist, F.; Huelsenbeck, J.P. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 2003, 19, 1572–1574. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  150. Rambaut, A. FigTree Tree Figure Drawing Tool Version 131, Institute of Evolutionary Biology, University of Edinburgh. Available online: http://treebioedacuk/software/figtree/ (accessed on 4 April 2022).
  151. Taj-Aldeen, S.J.; El Shafie, S.; Alsoub, H.; Eldeeb, Y.; de Hoog, G.S. Isolation of Exophiala dermatitidis from endotracheal aspirate of a cancer patient. Mycoses 2006, 49, 504–509. [Google Scholar] [CrossRef] [PubMed]
  152. McGinnis, M.R. Taxonomy of Exophiala jeanselmei (Langeron) McGinnis and Padhye. Mycopathologia 1978, 65, 79–87. [Google Scholar] [CrossRef] [PubMed]
Jof 08 00766 g001 550
Figure 1. The discovery of Exophiala-type species since 1966 to the present time.
Figure 1. The discovery of Exophiala-type species since 1966 to the present time.
Jof 08 00766 g001
Jof 08 00766 g002 550
Figure 2. Global distribution of Exophiala species. Area and countries where Exophiala species have been discovered are indicated in dark blue color.
Figure 2. Global distribution of Exophiala species. Area and countries where Exophiala species have been discovered are indicated in dark blue color.
Jof 08 00766 g002
Jof 08 00766 g003 550
Figure 3. Number of research articles (A) and related field areas (B) between 1992 and 2021 with “Exophiala” as a keyword. The search was performed using the Scopus database (accessed on the 9 May 2022).
Figure 3. Number of research articles (A) and related field areas (B) between 1992 and 2021 with “Exophiala” as a keyword. The search was performed using the Scopus database (accessed on the 9 May 2022).
Jof 08 00766 g003
Jof 08 00766 g004a 550Jof 08 00766 g004b 550
Figure 4. Phylogram generated from maximum likelihood analysis of 105 specimens of the combined ITS, nrSSU, tub, tef, and act genes. Cyphellophora fusarioides MUCL 44033 and C. eucalypti CBS 124764 were used as the outgroup. The numbers above branches show bootstrap percentages (left) and Bayesian posterior probabilities (right). Bootstrap values ≥ 70% and Bayesian posterior probabilities ≥ 0.95 are shown. The scale bar reflects the estimated number of nucleotide substitutions per site. Color bands represent the sequences of fungal species obtained in this study. Type species are in bold.
Figure 4. Phylogram generated from maximum likelihood analysis of 105 specimens of the combined ITS, nrSSU, tub, tef, and act genes. Cyphellophora fusarioides MUCL 44033 and C. eucalypti CBS 124764 were used as the outgroup. The numbers above branches show bootstrap percentages (left) and Bayesian posterior probabilities (right). Bootstrap values ≥ 70% and Bayesian posterior probabilities ≥ 0.95 are shown. The scale bar reflects the estimated number of nucleotide substitutions per site. Color bands represent the sequences of fungal species obtained in this study. Type species are in bold.
Jof 08 00766 g004aJof 08 00766 g004b
Jof 08 00766 g005 550
Figure 5. Exophiala lamphunensis (SDBR-CMU404, holotype): (A) Colony at 25 °C for 28 days on PDA, MEA, and OA, respectively; (B) budding cells; (C) germinating cells; (D) hyphal coil; (E,F) subcylindrical conidiophore and conidiogenous cells; (E–G) conidia. Scale bars: (A) = 2 cm; (B–G) = 5 μm.
Figure 5. Exophiala lamphunensis (SDBR-CMU404, holotype): (A) Colony at 25 °C for 28 days on PDA, MEA, and OA, respectively; (B) budding cells; (C) germinating cells; (D) hyphal coil; (E,F) subcylindrical conidiophore and conidiogenous cells; (E–G) conidia. Scale bars: (A) = 2 cm; (B–G) = 5 μm.
Jof 08 00766 g005
Jof 08 00766 g006 550
Figure 6. Exophiala lapidea (SDBR-CMU409, holotype): (A) Colony at 25 °C for 28 days on PDA, MEA, and OA, respectively; (B,C) budding cells; (D,E) germinating cells; (F) hyphal coil; (G) spirally twisted hyphae; (H) anastomoses; (I) erect, cylindrical conidiophore; (J–M) conidial apparatus with conidia; (I–N) conidia; (O) torulose hyphae. Scale bars: (A) = 2 cm; (B–O) = 5 μm.
Figure 6. Exophiala lapidea (SDBR-CMU409, holotype): (A) Colony at 25 °C for 28 days on PDA, MEA, and OA, respectively; (B,C) budding cells; (D,E) germinating cells; (F) hyphal coil; (G) spirally twisted hyphae; (H) anastomoses; (I) erect, cylindrical conidiophore; (J–M) conidial apparatus with conidia; (I–N) conidia; (O) torulose hyphae. Scale bars: (A) = 2 cm; (B–O) = 5 μm.
Jof 08 00766 g006
Jof 08 00766 g007 550
Figure 7. Exophiala saxicola (SDBR-CMU415, holotype): (A) colony at 25 °C for 28 days on PDA, MEA, and OA, respectively; (B,C) budding cells; (D,E) germinating cells; (F) anastomoses; (G) erect, cylindrical conidiophore; (H,I) obovoidal conidiogenous cells with obovoidal conidia; (J) conidial apparatus with conidia; (G–K) conidia; (L) chlamydospore; (M) torulose hyphae. Scale bars: (A) = 2 cm; (B–M) = 5 μm.
Figure 7. Exophiala saxicola (SDBR-CMU415, holotype): (A) colony at 25 °C for 28 days on PDA, MEA, and OA, respectively; (B,C) budding cells; (D,E) germinating cells; (F) anastomoses; (G) erect, cylindrical conidiophore; (H,I) obovoidal conidiogenous cells with obovoidal conidia; (J) conidial apparatus with conidia; (G–K) conidia; (L) chlamydospore; (M) torulose hyphae. Scale bars: (A) = 2 cm; (B–M) = 5 μm.
Jof 08 00766 g007
Jof 08 00766 g008 550
Figure 8. Exophiala siamensis (SDBR-CMU417, holotype): (A) colony at 25 °C for 4 weeks on PDA, MEA, and OA respectively; (B) budding cells; (C) germinating cells; (D) anastomoses; (E–I) conidial apparatus with subspherical conidia; (J) chlamydospore; (K) torulose hyphae. Scale bars: (A) = 2 cm; (B–G) = 5 μm.
Figure 8. Exophiala siamensis (SDBR-CMU417, holotype): (A) colony at 25 °C for 4 weeks on PDA, MEA, and OA respectively; (B) budding cells; (C) germinating cells; (D) anastomoses; (E–I) conidial apparatus with subspherical conidia; (J) chlamydospore; (K) torulose hyphae. Scale bars: (A) = 2 cm; (B–G) = 5 μm.
Jof 08 00766 g008
Table
Table 1. Global distribution and isolation resources of Exophiala species.
Table 1. Global distribution and isolation resources of Exophiala species.
SpeciesIsolation ResourcesLocationReference
Exophiala abietophilaSilver fir (Abies alba)Norway[21]
Exophiala alcalophilaSoil, soap container, washing machine, bathwater from households, and human skinBrazil, Denmark, Germany, Japan, and Ukraine[3,75,76]
Exophiala angulosporaPolluted soil, drinking water, Tilia wood, fish nursery, weedy seadragon, lumpfish skin and spleen, olive flounder (Paralichthys olivaceus), Atlantic cod (Gadus morhua), and human skinBrazil, Denmark, Germany, Ireland, Japan, Netherlands, Norway, Russia, Scotland, and the USA[3,77,78,79,80,81]
Exophiala aquamarinaClown fish, leafy sea dragon, little tunnyfish, lumpfish, sand lance, weedy seadragon, and winter flounderCanada, the UK, and the USA[3,7]
Exophiala arunalokeiSubcutaneous lesion on humanIndia[20]
Exophiala asiaticaTonsil tissue of humanChina[30,32]
Exophiala attenuataSoil, nasal granuloma of cat, cutaneous phaeohyphomycosis of cat, and human diseaseColombia, France, Germany, and the USA[33,82,83,84]
Exophiala bergeriEye and skin of humanBrazil, Canada, Japan, Hong Kong, the UK, and the USA[6,7,31,85]
Exophiala bonariaeMarbleItaly[69]
Exophiala brunneaLeaf of Acacia karrooSouth Africa[3]
Exophiala calicioidesRotten woodJapan[68]
Exophiala campbelliiSubcutaneous lesion (foot ganglion) of human and human chest noduleGermany and the UK[7,70]
Exophiala canceraeWater, water from tank, fruit drink, dialysis water Mangrove crab (Ucides cordatus), liver of green toad, sputum of human, and human fingerAustralia, Brazil, Canada, Germany, Hong Kong, Israel, Netherlands, the UK, and the USA[3,7,31,86,87,88]
Exophiala capensisLeaf of Phaenocoma proliferaCanada and South Africa[7,30,31,32,33,68,69,70,82,83,84,85,86,87,88,89]
Exophiala castellaniiDecaying timber joinery, spoilt apple juice, drinking water, ice water, nematode, and human skinDenmark, Germany, Netherlands, Sri Lanka, Switzerland, and the UK[3,27]
Exophiala cinereaRockChina[18]
Exophiala clavisporaRockChina[18]
Exophiala crusticolaBiological soil crustthe USA[90]
Exophiala dermatitidisSoil, dishwasher’s rubber, wood, internal organs of bat, chromoblastomycosis, knee fluid, lung, finger, and central nervous system fluid of humanAngola, Brazil, China, Finland, Germany, Hong Kong, Iran, Iraq, Japan, Korea, Malaysia, Mauritius, Qatar, Slovenia, South Africa, Taiwan, Thailand, Turkey, the UK, the USA, and Venezuela[7,15,16,31,32,45,46,47,48,49,50,51,52,53]
Exophiala dopicolaLoblolly pine (Pinus taeda)the USA[91]
Exophiala ellipsoideaRockChina[18]
Exophiala embothriiRhizosphere of Embothrium coccineumChile[92]
Exophiala encephalartiOn leaves of Encephalartos transvenosusSouth Africa[93]
Exophiala equinaSoil, drinking water, bottled water, water from water machine, water system of packaging machine, wastewater, dialysis water bathroom-flask, bathroom-plate, silica gel, root mycorrhiza, Tilia root, Populus tremuloides, Cephalanthera damasonium, Phragmitis australis, Olea twig, nematode cyst (Heterodera sp.), subcutaneous infection of horse, Galapagos turtle, human stool, human sputum, human eye, and skin of humanAustralia, Brazil, Canada, Denmark, Germany, Italy, Japan, Netherlands, Korea, the UK, and the USA[3,7,10,87,94,95,96,97,98]
Exophiala eucalyptiLeaves of Eucalyptus sp.South Africa[99]
Exophiala eucalypticolaLeaf of Eucalyptus obliquaAustralia[22]
Exophiala eucalyptorumLeaf of Eucalyptus sp.New Zealand[23]
Exophiala exophialaeSoil, straw in armadillo’s burrow (Dasypus septemcinctus)Colombia and Uruguay[6,15,34]
Exophiala frigidotoleransSoilEcuador[100]
Exophiala halophilaSalty water, human skin, and human nailGermany and the USA[3]
Exophiala heteromorphaWood and humanSweden and the USA[7,15]
Exophiala hongkongensisBig toenail infection of humanChina and Hong Kong[31]
Exophiala italicaCytisus scoparius on dead branchItaly[24]
Exophiala jeanselmeiSubcutaneous abscesses, skin lesion, eumycetoma of human, peritoneal dialysis fluid, human blood, human sputum, and human eyeAustralia, Bangladesh, Brazil, Canada, China, Costa Rica, France, Hong Kong, Jamaica, Japan, Martinique, Pakistan, Paraguay, Peru, Philippines, Saudi Arabia, Thailand, Trinidad, the UK, Uruguay, and the USA[6,17,27,31,32,33,34,54,55,56,57,58,59,60,61,62,63,64,65,66,67]
Exophiala lacusLake water and river sedimentsNetherlands and Spain[3,71]
Exophiala lavatrinaDomestic bathroomthe UK[7]
Exophiala lecanii-corniLecanium corni, domestic bathroom, dialysis fluid, subcutaneous abscess, knee cyst, skin lesion, eye sclera, finger fluid, skin scales, and human nailAustria, Germany, Hong Kong, Japan, Netherlands, the UK, and the USA[27,31,57,101,102]
Exophiala lignicolaQuercus sp.Ukraine[21]
Exophiala macquariensisIsland soilAustralia[2]
Exophiala maliInner fruit tissue of Malus sp.South Africa[92]
Exophiala mansoniiPopulus tremulaSweden[15]
Exophiala mesophilaShower joint, swimming pool, dental waterline, bathroom, contact lens, phaeohyphomycotic cyst, subcutaneous nodule biopsy, immunosuppressed, bronchial endoscopy, finger, sinus, hip joint, hair, and nasal tissue of humanBrazil, France, Germany, Netherlands, the UK, and the USA[3,7,85]
Exophiala moniliaeBranch of Quercus sp., sludge in bathroom drainpipes, and medicated bathwaterAustralia, Japan, and Russia[15,76,103]
Exophiala nagquensisRockChina and Tibet[18]
Exophiala nidicolaNest of birdSpain[72]
Exophiala nishimuraeBark and human skinthe USA and Venezuela[17,33]
Exophiala oligospermaSoil, wood, swimming pool, water, polluted water, river sediments, sauna, silicone solution, ear swab, plastic foil, prosthetic contact lenses, cerebral mycosis, subcutaneous abscess, thigh abscess, skin lesion, sphenoid tumor, lung, sinus, and human sputumAustria, Brazil, Canada, Finland, France, Germany, Hong Kong, Italy, Japan, Netherlands, Spain, Switzerland, the UK, Ukraine, the USA, and Venezuela[6,7,17,31,34,57,71,104,105,106]
Exophiala opportunisticaDrinking water, rhizosphere (Triticum aestivum), polyvinyl alcohol, human nail, and human footAustralia, Denmark, Germany, and Netherlands[3]
Exophiala palmaeDecaying shell of babassu coconut (Orbignya phalerata)Brazil[107]
Exophiala phaeomuriformisNatural hot spring, sauna, tile floor of swimming pool, bathroom tap, bathroom sink, cutaneous mycosis, blood culture, external ear channel, oral mucosa, nail, and human sputum,Austria, Canada, Czech Republic, Germany, Japan, Netherlands, Slovenia, the UK, and the USA[7,16,108]
Exophiala pisciphilaSwimming pool, water pipe, dialysis water, catfish (Ictalurus punctatus), Potbelly seahorse, crocodile, and humanBrazil, Germany, Japan, Israel, and the USA[3,7,87,109]
Exophiala polymorphaSubcutaneous lesion of humanthe USA[8]
Exophiala prostantheraeLeaves of Prostanthera sp.Australia[92]
Exophiala pseudooligospermaKarst rocky desertification mountain soilChina[4]
Exophiala psychrophilaAtlantic salmon smolt (Salmo salar)Ireland and Norway[3]
Exophiala quercinaDead wood of Quercus sp.Germany[73]
Exophiala radicisSoil, root endophyte of Microthlaspi perfoliatum, plant roots, Olea sp. twig, nematode cyst (Heterodera sp.), toenail, tinea on leg, and foot of humanBulgaria, Denmark, France, Germany, Italy, the Netherlands, and Spain[5,71]
Exophiala salmonisDrinking water, drinking water tap and cerebral mycetoma of fingerling trout (Salmo clarkii)Canada and the Netherlands[1]
Exophiala siderisOak railway tie, creosoted tie, gold mine, and surface of wild berries of Sorbus aucupariathe Netherlands and Poland[9]
Exophiala spartinaeSpartina alterniflora root tissue in saltwater marshthe USA[110]
Exophiala spiniferaSoil, palm tree, wood, nest of Anumbius annumbi, armadillo burrow, maize kernel, apple juice, rotten cactus, skin lesion, foot abscess, neck lymph node, human sputum, and bark nasal granuloma of humanAntarctic, Argentina, Australia, Brazil, China, Colombia, Egypt, Germany, India, Mexico, Papua New Guinea, Senegal, Thailand, the UK, Uruguay, the USA, and Venezuela[6,32,33,34,35,36,37,38,39,40,41,42,43,44]
Exophiala tremulaePopulus tremuloides rootsCanada[111]
Exophiala xenobioticaSoil, wood, oil sludge, chromoblastomycosis on back, phaeomycotic cyst, subcutaneous cyst, elbow pus, and skin lesionsAntarctic, Australia, Brazil, Canada, Germany, Hong Kong, Japan, the Netherlands, New Zealand, Switzerland, Sweden, the UK, the USA, and Venezuela[6,7,27,31,34,57,74]
Table
Table 2. List of the primers, primer sequences, and annealing temperatures used for PCR amplification in each target gene.
Table 2. List of the primers, primer sequences, and annealing temperatures used for PCR amplification in each target gene.
Target GenePrimerPrimer Sequence (5′–3′)Annealing Temperature (°C)Reference
actAct1TGGGACGATATGGAIAAIATCTGGCA52[124]
Act5raTTAGAAGCACTTNCGGTG52[124]
ITSITS4TCCTCCGCTTATTGATATGC55[125]
ITS5GGAAGTAAAAGTCGTAACAAGG55[125]
nrSSUNS1GTAGTCATATGCTTGTCTC55[125]
NS4CTTCCGTCAATTCCTTTAAG55[125]
tefEF1-728FCATCGAGAAGTTCGAGAAGG57[126]
EF1-986RTACTTGAAGGAACCCTTACC57[126]
tubBt2aGGTAACCAAATCGGTGCTGCTTTC52[127]
Bt2bACCCTCAGTGTAGTGACCCTTGGC52[127]
Table
Table 3. DNA sequences used in the molecular phylogenetic analysis.
Table 3. DNA sequences used in the molecular phylogenetic analysis.
SpeciesStrainsGenBank Accession No.References
ITSnrSSUtubtefact
Exophiala abietophilaCBS 145038 TNR163357––––[21]
CBS 520.82 TJF747041JN856010JN112423JN128771JN112379[3]
CBS 122256JF747044–JN112425JN128773JN112381[3]
Exophiala angulosporaCBS 482.92 TJF747046JN856011JN112426JN128780JN112383[3]
CBS 120272JF747045–JN112427JN128781JN112382[3]
Exophiala aquamarinaCBS 119918 TJF747054JN856012JN112434–JN112388[3]
CBS 119916JF747055–JN112435–JN112389[3]
Exophiala arunalokeiNCCPF106033MW724320––––[20]
Exophiala asiaticaCBS 122847 TNR111332––––[30]
CBS 122848MW222182––––[30]
Exophiala attenuataCBS 101540 TAF549446––––[33]
UTHSC87-80EF025392––––[130]
Exophiala bergeriCBS 353.52 TEF551462FJ358308EF551497EF551524EF551464[131]
Exophiala bonariaeCBS 139957 TJX681046––––[69]
Exophiala brunneaCBS 587.66 TJF747062JN856013JN112442JN128783JN112393[3]
Exophiala calicioidesJCM6030–AB007655–––[132]
Exophiala campbelliiNCPF 2274LT594703––LT594739–[7]
Exophiala canceraeCBS 120420 TJF747064 –JN112444JN128800JN112394[3]
CBS 117491KF928439–KF928567JN128799JN112396[3]
Exophiala capensisCBS 128771 TJF499841––––[89]
Exophiala castellaniiCBS 158.58 TJF747070JN856014KF928586JN128766–[3,133]
CBS 120913JF747144–JN112506JN128750–[3]
Exophiala cinereaCGMCC 3.18778 TMG012695MG012724MG012745MG012704MG012714[18]
CGMCC 3.18779MG012696MG012725MG012746MG012705MG012715[18]
Exophiala clavisporaCGMCC 3.17512KP347940MG012733KP347931KP347909MG012712[18]
CGMCC 3.17517 TKP347942KP347967KP347932KP347911KP347893[18]
Exophiala crusticolaCBS 119970 TAM048755KF155199–––[90,134]
HM136MK281393––––Unpublished
Exophiala dermatitidisCBS 207.35 TAF050269–KF928572––[133,134,135]
CBS 120473MF320159–MF320217MF320196–[43]
Exophiala dopicolaCBS 537.94 TMH862483––––[136]
Exophiala ellipsoideaCGMCC 3.17348 TKP347955KP347965KP347921KP347901MG012713[18]
CGMCC 3.17522KP347954MG012735KP347919–KP347884[18]
Exophiala embothriiCBS 146558 TNR171982–MW055976MW055980–[92]
Exophiala encephalartiCBS 128210 THQ599588––––[93]
Exophiala equinaCBS 119.23 TJF747094JN856017JN112462JN128814JN112401[3]
CBS 120906JF747093–JN112461JN128813JN112400[3]
Exophiala eucalyptiCBS 142069KY173411––––[99]
Exophiala eucalypticolaCBS 143412 TNR158438–MH108039MH108016–[22]
Exophiala eucalyptorumCBS 121638 TNR132882KC455302KC455228––[137]
CPC 11261EU035417––––[23]
Exophiala exophialaeCBS 668.76 TAY156973KX822287EF551499EF551526EF551466[33,138]
CBS 671.76AY156975–EF551500EF551525EF551467[33]
Exophiala frigidotoleransCBS 146539 TLR699566––––[100]
Exophiala halophilaCBS 121512 TNR111628NG062077JN112473JN128774–[3]
Exophiala heteromorphaCBS 232.33 TAY857524––––[139]
U THSC87-67EF025400––––[130]
Exophiala hongkongensisCBS 131511JN625231–JN625236JN625246JN625241[31]
Exophiala italicaMFLUCC16-0245 TKY496744KY501114–KY514393-[24]
Exophiala jeanselmeiCBS 507.90 TAY156963FJ358310EF551501EF551530-[33,131]
CBS 528.76AY857530–EF551502EF551531EF551469[139]
Exophiala lacusFMR 3995KU705830––––[71]
CBS 117497 TJF747110––JN128776JN112407[3]
Exophiala lamphunensisSDBR-CMU404 TON555798ON555813ON544227ON544242ON544257This study
SDBR-CMU405ON555799ON555814ON544228ON544243ON544258This study
SDBR-CMU406ON555800ON555815ON544229ON544244ON544259This study
SDBR-CMU407ON555801ON555816ON544230ON544245ON544260This study
SDBR-CMU408ON555802ON555817ON544231ON544246ON544261This study
Exophiala lapideaSDBR-CMU409 TON555803ON555818ON544232ON544247ON544262This study
SDBR-CMU410ON555804ON555819ON544233ON544248ON544263This study
SDBR-CMU411ON555805ON555820ON544234ON544249ON544264This study
SDBR-CMU412ON555806ON555821ON544235ON544250ON544265This study
SDBR-CMU413ON555807ON555822ON544236ON544251ON544266This study
SDBR-CMU414ON555808ON555823ON544237ON544252ON544267This study
Exophiala lavatrinaNCPF 7893LT594696––LT594729–[7]
NCPF 7898LT594697––LT594731–[7]
Exophiala lecanii-corniCBS 123.33 TAY857528FJ358311–––[131,139]
B2242CMT320770––MZ190330–[140]
Exophiala lignicolaCBS:144622 TNR163358––MK442694–[21]
Exophiala macquariensisCBS 144232 TMF619956–MH297438MH297439–[2]
Exophiala maliCBS 146791 TMW175341––––[92]
Exophiala mansoniiCBS 101.67 TAF050247X79318–––[135,141]
Exophiala mesophilaCBS 402.95 TJF747111JN856016JN112476JN128761–[3]
CBS 119910JF747113–JN112478JN128753–[3]
Exophiala moniliaeCBS 520.76 TKF881967––––Unpublished
BMU00283MW222184––––Unpublished
Exophiala nagquensisCGMCC 3.17333 TKP347948KP347970KP347924KP347914KP347895[18]
CGMCC 3.17334KP347949MG012741KP347923KP347915KP347896[18]
Exophiala nidicolaCBS 138589 TNR161045––––[72]
Exophiala nishimuraeCBS 101538 TAY163560KX822288JX482552EF551523JX482553[33]
Exophiala oligospermaCBS 725.88 TAY163551FJ358313EF551508EF551534EF551474[17,131]
CBS 265.49MH856519–EF551507EF551536EF551473[136]
Exophiala opportunisticaCBS 109811 TJF747123–JN112486JN128792JN112408[3]
Exophiala palmaeCMRP1196 TKY680434–KY689829––[107]
CMRP1207KY680433–KY689828––[107]
Exophiala phaeomuriformisCBS 131.88 TAJ244259––––Unpublished
Exophiala pisciphilaCBS 537.73 TNR121269JN856018JN112493JN128788JN112412[3,142]
CBS 121500JF747134–JN112496JN128789JN112414[3]
Exophiala polymorphaCBS 138920 TKP070763––––[8]
Exophiala prostantheraeCBS 146794 TNR171990––––[92]
Exophiala pseudooligospermaYMFT 1.6741MW616557MW616558MZ127830––[4]
Exophiala psychrophilaCBS 191.87 TJF747135JN856019JN112497JN128798–[3]
CBS 256.92JF747136–JN112498––[3]
Exophiala quercinaCBS 146024 TNR170053––MT223713–[73]
Exophiala radicisP2854 TKT099204KT723453KT723463KT723458KT723443[5]
Exophiala salmonisCBS 157.67 TAF050274JN856020JN112499JN128747JN112415[3,135]
CBS 120274JF747138–KF928562JN128802JN112416[3]
Exophiala saxicolaSDBR-CMU415 TON555809ON555824ON544238ON544253ON544268This study
SDBR-CMU416ON555810ON555825ON544239ON544254ON544269This study
Exophiala siamensisSDBR-CMU417 TON555811ON555826ON544240ON544255ON544270This study
SDBR-CMU418ON555812ON555827ON544241ON544256ON544271This study
Exophiala siderisCBS 121818 THQ452311HQ441174HQ535833HQ452336–[9]
CBS 127096HQ452312HQ441175HQ535834HQ452337–[9]
Exophiala spartinaeCBS 147266 TNR174648––––[110]
Exophiala spiniferaCBS 899.68 TAY156976–EF551516EF551541EF551482[33]
Exophiala tremulaeCBS 129355 TFJ665274KT894147KT894148KT894149KT894146[5,89]
Exophiala xenobioticaCBS 118157 TDQ182587––––[27]
CBS 117646KP132146––––[27]
Cyphellophora eucalyptiCBS:124764 TGQ303274NG062860KF928601GU384510JQ325009[133,137,143,144]
Cyphellophora fusarioidesMUCL 44033NR132879NG065006KC455224––[137]
Note: species obtained in this study are in bold. Superscript “T” indicates type species and “–” represents the absence of sequence data in GenBank.
Table
Table 4. Colony diameter of 15 fungal strains on MEA at different temperatures for 28 days of incubation in the darkness.
Table 4. Colony diameter of 15 fungal strains on MEA at different temperatures for 28 days of incubation in the darkness.
Fungal StrainsColony Diameter (mm) *
10 °C15 °C20 °C25 °C28 °C30 °C35 °C37 °C
SDBR-CMU40410.25 ± 0.2717.92 ± 0.9218.17 ± 0.5223.83 ± 0.4124.42 ± 0.5822.83 ± 0.6813.08 ± 0.588.00 ± 0.55
SDBR-CMU40510.54 ± 0.3317.88 ± 0.5619.08 ± 0.1224.27 ± 0.4224.85 ± 0.5723.09 ± 0.4112.47 ± 0.528.12 ± 0.14
SDBR-CMU40611.42 ± 0.5216.12 ± 0.1619.78 ± 0.7224.05 ± 0.9725.41 ± 0.4422.79 ± 0.8512.55 ± 0.557.36 ± 0.22
SDBR-CMU40711.25 ± 0.4216.33 ± 0.4119.25 ± 0.5225.58 ± 0.58 25.92 ± 0.3823.17 ± 0.5212.33 ± 0.416.92 ± 0.20
SDBR-CMU40810.12 ± 0.2216.45 ± 0.8718.44 ± 0.6125.03 ± 0.4525.25 ± 0.6222.81 ± 0.4312.74 ± 0.407.45 ± 0.39
SDBR-CMU40917.75 ± 0.2720.08 ± 1.0728.33 ± 0.9836.24 ± 1.4437.00 ± 1.2640.83 ± 1.3310.08 ± 0.386.08 ± 0.20
SDBR-CMU41016.11 ± 0.1824.35 ± 0.8426.65 ± 0.8835.91 ± 1.3636.02 ± 1.3138.42 ± 1.448.27 ± 0.455.96 ± 0.22
SDBR-CMU41114.98 ± 0.1220.03 ± 0.4127.78 ± 1.2334.78 ± 0.9735.43 ± 1.2836.92 ± 1.969.04 ± 0.365.23 ± 0.27
SDBR-CMU41215.97 ± 0.5226.27 ± 0.9227.56 ± 0.7136.77 ± 1.2237.11 ± 1.4538.82 ± 0.799.19 ± 0.245.71 ± 0.13
SDBR-CMU41314.42 ± 0.3819.25 ± 0.2725.08 ± 1.0732.25 ± 1.4434.33 ± 2.04 35.42 ± 0.868.42 ± 0.495.58 ± 0.49
SDBR-CMU41414.23 ± 0.4725.78 ± 0.7425.71 ± 0.8835.04 ± 1.4735.47 ± 1.42 36.96 ± 0.6510.12 ± 0.565.44 ± 0.39
SDBR-CMU4159.92 ± 0.2014.42 ± 0.4915.50 ± 0.5521.33 ± 0.2623.75 ± 1.3724.17 ± 1.6612.75 ± 0.278.92 ± 0.20
SDBR-CMU4169.83 ± 0.4114.08 ± 0.2016.58 ± 0.3821.75 ± 1.1324.67 ± 0.41 26.88 ± 1.2811.42 ± 0.208.17 ± 0.26
SDBR-CMU4178.08 ± 0.3810.08 ± 0.4911.75 ± 1.17 10.42 ± 0.809.42 ± 0.587.33 ± 0.26––
SDBR-CMU4188.08 ± 0.3810.50 ± 0.7711.83 ± 0.6810.35 ± 1.179.92 ± 0.207.58 ± 0.20––
* The results are mean ± standard deviation and “–” represents no growth.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Thitla, T.; Kumla, J.; Khuna, S.; Lumyong, S.; Suwannarach, N. Species Diversity, Distribution, and Phylogeny of Exophiala with the Addition of Four New Species from Thailand. J. Fungi 2022, 8, 766. https://doi.org/10.3390/jof8080766

AMA Style

Thitla T, Kumla J, Khuna S, Lumyong S, Suwannarach N. Species Diversity, Distribution, and Phylogeny of Exophiala with the Addition of Four New Species from Thailand. Journal of Fungi. 2022; 8(8):766. https://doi.org/10.3390/jof8080766

Chicago/Turabian Style

Thitla, Tanapol, Jaturong Kumla, Surapong Khuna, Saisamorn Lumyong, and Nakarin Suwannarach. 2022. "Species Diversity, Distribution, and Phylogeny of Exophiala with the Addition of Four New Species from Thailand" Journal of Fungi 8, no. 8: 766. https://doi.org/10.3390/jof8080766

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop