Developing species recognition and diagnostics of rare opportunistic fungi
Zeng, J.Publication date 2007
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Zeng, J. (2007). Developing species recognition and diagnostics of rare opportunistic fungi. IBED.
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Developing Species
Recognition and Diagnostics of
Rare Opportunistic Fungi
Developing Species Recognition and
Diagnostics of Rare Opportunistic Fungi
Promotor
Prof. Dr. G.S. de Hoog
Centraalbureau voor Schimmelcultures Fungal Biodiversity Centre, Royal Netherlands Academy of Arts and Sciences
Institute for Biodiversity and Ecosystem Dynamics, University of Amsterdam Co-promotor
Dr. Y. Gräser
Humboldt University, Germany Promotiecommissie
W. Admiraal (IBED, Amsterdam) S. Menken (IBED, Amsterdam) M. Sabelis (IBED, Amsterdam)
Developing Species Recognition and
Diagnostics of Rare Opportunistic Fungi
Jingsi Zeng
ACADEMISCH PROEFSCHRIFT ter verkrijging van de graad van doctor
aan de Universiteit van Amsterdam op gezag van de Rector Magnificus
prof. dr. J. W. Zwemmer
ten overstaan van een door het college voor promoties ingestelde commissie, in het openbaar te verdedigen in de Aula der Universiteit
Printed by Ponsen & Looijen B.V., Wageningen, the Netherlands
The work was financially supported by the joint research project ‘Comparative genomics in search of origins of human pathogenicity in the fungal Tree of Life focusing on species with high morbidity and mortality in Chinese patients’, Scientific cooperation between China and the Netherlands, China Exchange Programme, Royal Netherlands Academy of Arts and
This thesis is dedicated to my dear parents, husband and daughter, my tutors for Master’s degree Prof. Zhaoru Zhu and Prof. Yuechen Zheng
谨献给我亲爱的父母、丈夫及女儿， 以及恩师祝兆如教授和郑岳臣教授
Contents
Page
Chapter 1 Introduction and outline of thesis 1
Chapter 2 Intraspecific diversity of species of Pseudallescheria boydii 25
complex
Chapter 3 Exophiala xenobiotica sp. nov., an opportunistic black yeast 45
inhabiting environments rich in hydrocarbons
Chapter 4 Spectrum of clinically relevant Exophiala species in the U.S.A. 65
Chapter 5 Phylogeny of the Exophiala spinifera clade using multilocus 85
sequence data and exploring phylogenetic species concept
Chapter 6 Exophiala spinifera and its allies: diagnostics from 109
morphology to DNA barcoding
Chapter 7 Susceptibility of Pseudallescheria boydii and Scedosporium 133
apiospermum to new antifungal agents
Chapter 8 General dicussion 139
Appendix Summary 151
List of publications and abstracts 155
Acknowledgements 158
Historical overview of species recognition in Pseudallescheria and
Exophiala
The fungal genera Pseudallescheria and Exophiala include agents of opportunistic infection in humans. They are potentially able to cause a wide diversity of mycoses, varying from cutaneous infections to disseminated syndromes. Most infections are noted in otherwise healthy individuals and are of traumatic nature or concern colonization of cavities or the intestinal tract. Most are typical opportunists in that they expand or disseminate when the innate immunity of the host is impaired. However, some Exophiala species are repeatedly observed to cause fatal, systemic or disseminated infections in patients without any immune disorder. The pathology of these fungi is poorly understood since development of the
knowledge was long time hampered by inadequate diagnostics. Although they are among the first fungi reported from deep human infections [1], their taxonomy flourished only since the application of molecular methods [2;3]. In the routine laboratory, species recognition is still problematic.
Pseudallescheria Negroni et Fischer
The lineage of the genus Pseudallescheria (anamorph Scedosporium) is Eukaryota, Fungi, Ascomycota, Euascomycetes, Microascales, Microascaceae. The species thus far established in the genus are listed in Table 1. Colonies on cornmeal agar are rapid growing, floccose or lanose, initially whitish or grey, becoming grey or brown; ascomata are usually submerged, globose, nonostiolate, 100-200 μm in diameter width; peridium composed of flattened, brown, pseudoparenchymatous cells, 2-3 cells thick; asci are globose to ellipsoidal, symmetrical or slightly flattened, with 2 germ pores, light brown to yellowish; homothallic. Scedosporium, Graphium, or both anamorphs may be formed [4].
In 1893, Costantin described a species Eurotiopsis gayoni Cost. [5]. As the name Eurotiopsis had already been used for an entirely different fungus, Saccardo substituted the generic name Allescheria in 1899 [6]. Based on Costantin’s illustration and descriptions of E. gayoni, Malloch [7] concluded that it should be classified in the genus Monascus van Tiegh., 1884. Thus the genera Allescheria and Eurotiopsis became later synonyms of the genus Monascus. In 1922, a strain, isolated by M. F. Boyd from a patient with mycetoma, was
new species, Allescheria boydii Shear in 1922 [8]. When the members of the ascomycete family Microascaceae were studied by Malloch, the ascomata and ascospores produced by A. boydii were found to be significantly different from those produced by either the other species of Monascus or any other genus of cleistothecial ascomycetes [7]. Malloch proposed the new genus Petriellidium in the family Microascaceae for A. boydii. Consequently, six other
Petriellidium (Pe. africanum Arx & G. Franz, 1973, Pe. angustum Malloch & Cain, 1972, Pe. desertorum Arx & Moustafa, 1973, Pe. ellipsoideum Arx & Fassatiová, 1973, Pe. fimeti Arx, Mukerji & Singh, 1978 and Pe. fusoideum Arx, 1973) (throughout this chapter Pe will be used for Petriellidium, Ps for Pseudallescheria) were described in this genus [9-11]. In 1943 and again in 1944, Negroni & Fischer described the genus and species Pseudallescheria shearii (as ‘sheari’) for a cleistothecial ascomycete [12;13]. Comparing the type specimens of Petriellidium Malloch and Pseudallescheria Negroni et Fischer, McGinnis et al. indicated that these 2 genera were congeneric [4]. Because of priority, the proper name of this genus is Pseudallescheria. At that moment, a total of 7 Petriellidium species were reclassified on the basis of morphology as distinct species of Pseudallescheria (Ps. boydii, Ps. africana, Ps. angusta, Ps. desertorum, Ps. ellipsoidea, Ps. fimeti and Ps. fusoidea). Members of the genus Pseudallescheria typically produce Scedosporium or Graphimu anamorphs, or both. Until 1991, 2 anamorph species of Pseudallescheria had been accepted, which are Scedosporium
apiospermum Sacc. ex Castell. & Chalmers [14] and Scedosporium prolificans (Hennebert &
B.G. Desai) Guého & de Hoog [15].
The morphological circumscriptions of Ps. boydii, Ps. angusta, Ps. ellipsoidea and Ps. fusoidea are narrow. A part of isolates maintained as Ps. boydii failed to produce cleistothecia upon inspection from collection strains regardless of growth conditions. Ps. angusta, Ps. fusoidea and Ps. ellipsoidea were later reduced to synonymy with Ps. boydii (Ps. boydii complex) on the basis of identical Internal Transcribed Spacer (ITS) sequences of ribosomal DNA (rDNA) [16] and a ~300 bp fragment of the D1/D2 Large Subunit (LSU) region of rDNA [17]. The recently described species Ps. minutispora Gilgado et al., S. aurantiacum Gilgado et al. [2], and two more species proposed by Gilgado and co-workers [18] have been segregated from Ps. boydii on the basis of genealogical concordance of calmodulin, β-tubulin and ITS region of rDNA genes and phenotypic characters. At this moment, 9 species are accepted in the ascomycete genus Pseudallescheria, but thus far only few outstanding
differences have been found in features of pathogenicity, ecology and antifungal susceptibility between the new species and Ps. boydii [2;19;20]. Excluding molecular data, species
recognition still is difficult in the routine lab, and as a result existing differences may be overlooked. Therefore it is essential that more tools become available to recognize species in the genus Pseudallescheria.
Exophiala J.W. Carmichael
Exophiala is the main genus of black yeasts, characterised by annellidic
conidiogenesis. Colonies are mostly restricted, slimy at the centre due to yeast-like growth. Some cultures appear entirely yeast-like (synanam. Phaeococcomyces), or form phialidic collarettes (synanam. Phialophora), sympodial conidiophores (synanam. Rhinocladiella without conidiophores, Ramichloridium with conidiophores) or dry conidial chains (synanam. Cladophialophora).
The genus Exophiala was established in 1966 by J. W. Carmichael on the basis of Exophiala salmonis J.W. Carmichael, which type strain was isolated from cerebral mycetoma of Salmo clarkia [21]. The primarily described phialide-like sporogenous cells of this genus were recognized as annellides later [22-24]. The annellated zones are the main diagnostic feature of this genus. With this morphological criterion erected, a number of Exophiala species were consequently defined or re-classified, e. g. E. jeanselmei (Langeron) McGinnis & A.A. Padhye (including three morphological varieties), E. dermatitidis (Kano) de Hoog, E. pisciphila McGinnis & Ajello, E. mansonii (Castell.) de Hoog, E. moniliae de Hoog, E. spinifera (H.S. Nielsen & Conant) McGinnis, etc. [22;23;25-28]. The genus Wangiella McGinnis [29] was proposed for the dematiaceous hyphomycete originally invalidly published as Hormiscium dernatitidis Kano [30] and subsequently placed in the genera Fonsecaea [31], Hormodendrum [32]and Phialophora [33]. McGinnis placed Exophiala dermatitidis in the genus Wangiella as W. dermatitidis according to that the species formed conidia predominantly from phialides without collarettes. However, Nishmura et al.
confirmed by scanning electron microscopy that the conidia of E. dermatitidis arised mostly from annellides, but annellated zones were limited [34]. Until 1996, a total of twelve accepted Exophiala species were identified morphologically[35], including 6 known opportunists in humans. Table 2 gives the name list of Exophiala species.
In the past, taxonomic and diagnostic schemes for Exophiala were morphological, while later some physiological parameters proved to be useful [36;37]. Morphology is rather unreliable for species identification due to variable appearance. Their taxonomic coherence has been proven by sequencing of the ribosomal gene [38-41]. During the last ten years, taxonomic schemes have become re-ordered supplemented by molecular data, particularly sequences of the rDNA ITS regions [42-44]. Most species were redefined, re-designated or initially described mainly depending on genetic characteristics, combined with morphological, physiological and ecological features [39;44-46]. Only phenetically recognizable taxa such as E. dermatitidis and E. spinifera remained largely unaltered. Thus far, 26 species of the genus Exophiala have been erected (including 15 opportunists in humans), followed by several new species to be described soon (including 3 opportunists in humans) [PhD thesis, M. J. Harrak, 2008]. In the mean time, phylogenetic knowledge of the genus developed.
Though the anamorphs of major Capronia teleomorphs can be predicted to belong to Exophiala [34;47-51], the number of described species of Exophiala with a proven Capronia teleomorph, conspecific to or even close in morphological and physiological parameters [38;40;52;53] is very small.
A new tool for species recognition: GCPSR
Historically there are four prevalent species concepts [54]. These are Evolutionary Species Concept (ESC), Morphological Species Concept (MSC), Biological Species Concept (BSC) and Phylogenetic Species Concept (PSC). ESC defines a species as, “ . . . a single lineage of ancestor-descendent populations which maintains its identity from other such lineages and which has its own evolutionary tendencies and historical fate” [55]. When ESC comes to identifying species, it is not helpful because it has no recognition criteria. In contrast, the other 3 species concept do specify criteria for recognizing species, therefore the term ‘species recognition’ was proposed to replace ‘species concept’, namely Morphological Species Recognition (MSR), Biological Species Recognition (BSR) and Phylogenetic Species Recognition (PSR) [56]. Among these types of species recognition, PSR performs best. Once the evolutionary species has formed from an ancestor, changes in gene sequences occur and can be recognized before changes have occurred in mating behavior or morphology [56]. With fungi, Harrington & Rizzo proposed a type of PSR that diagnoses species as “ . . . the
smallest aggregation of populations with a common lineage that share unique, diagnosable phenotypic characters” [57]. With PSR, fungal species are diagnosed based on the
concordance of gene genealogies and phenotypic features. Some fungal studies demonstrate that PSR is well suited to fungi [58-60]. But it is difficult to place the limit of the species when a gene is polymorphic within a species or fix on for alternate alleles in more than one species. Given additional information, e.g. ability to interbreed, this difficulty could be solved [61]. PSR can avoid the subjectivity of determining the limits of a species by replying on the concordance of more than one gene genealogy [62]. This type of PSC was named as
Genealogical Concordance Concept (GCC) by Mayden [54]. Later, Taylor et al. used the term ‘Genealogical Concordance Phylogenetic Species Recognition’ (GCPSR) to indicate this principle related to operational species concept [56]. With GCPSR, more than one gene lineages need to be investigated and compared. When conflict occurs among lineages, the transition from concordance to conflict determines the limits of species [56]. However this transition can not occur for clonal populations. In this case, the boundary of species is determined by diagnosable phenotypic characters in morphology, physiology, ecology, pathogenicity and antifungal susceptibility; or as suggested by Taylor et al., clonal species could be accommodated via the GCPSR by defining them in relation to their recombining relatives [56].
Techniques for measuring and testing the significance of phylogenetic incongruence are used widely in systematic biology, and are necessary when considering multiple genes that may or may not have different histories [63-71]. One of the most intuitive measures of incongruence is the Incongruence Length Difference test (ILD) [63-64]. The version of the test available in the popular phylogenetic analysis program PAUP* [72] is called the ‘Partition Homogeneity Test’ (PHT).
Incongruence found by ILD indicates evolutionary historical heterogeneity among lineages. Normally evolutionary historical heterogeneity includes differences in evolutionary processes, such as different rates of evolution, or alternative evolutionary history. The latter mostly concerns non-vertical inheritance, e.g. gene duplication, loss, horizontal gene transfer, hybridization or recombination. ILD is meant to be an overall measure of incongruence between datasets. Though tests for locating incongruence among loci are available [65;71], most of them are not thoroughly tested and insufficiently understood. Detailed localization is
time consuming. Of the above mentioned historical events, recombination is most closely related to species recognition with GCPSR.
One of methods to detect recombination with multi-locus data sets is to calculate the Index of Association (IA, a measure of multi-locus linkage disequilibrium) for a
reproductively isolated population. Detecting population differentiation (index: ө) and computing IA and can be performed with Multilocus v. 1.2.2
(http://www.bio.ic.ac.uk/evolve/software/multilocus) software. Population structure can be inferred from phylogenetic analysis and from clustering analysis of genotype data sets with STRUCTURE software v. 2. [http://pritch.bsd.uchicago.edu].
The scheme for recognizing fungal species with the criterion of GCPSR in my studies is summarised in Figure 1.
Diagnostics of etiologic fungi
Conventional methods to identify etiologic fungi often rely on identification of disease symptoms, isolation and culturing of organisms, and laboratory identification by morphology and biochemical tests. These methods rely on experienced, skilled laboratory staff, the ability of the organism to be cultured, are time consuming, non-quantitative, and prone to
contamination. Although they are the cornerstone of fungal diagnostics, they can lead to problems in identification, resulting in incorrect interpretation, diagnosis and ultimately treatment. New, rapid screening methods are being developed and increasingly used in all aspects of fungal diagnostics. These methods include immunological methods, DNA/RNA probe technology and Polymerase Chain Reaction (PCR) technology. Based on PCR and probe technology, micro-array technology and real-time PCR methods bring a bright future for the development of accurate, quantitative diagnostic tools. DNA barcoding as a
standardized approach to identify species by a short gene sequence from a uniform region in the genome [http://www.barcoding.si.edu] was advocated several years ago. Barcoding provides tools that allow rapid and unambiguous definition and recognition, and has phylogenetical implications as it is directly based upon the evolutionary history of life.
A ss ess r e prod uct iv e m ode fo r ea ch c lu st er (m ea su re IA wit h L in ka ge D ise qu ili br iu m A na lys is in M ultilo cu s so ft wa re ) M ultil o cu s se que nc e da ta set s C on stru ct M P tree of ea ch g ene D efi ne geno typ e s fo r ea ch lo cu s ( ge noty p e d a ta se t) G ro up in g b a se d on ph en ot yp ic fe at ur es ( e . g . h o st pr ef er en ce , ge og ra ph ic al di str ibu tio n, pa th og en ic ity , an tif ung al susc ep tib ili ty ) Acc ordi ng t o PSR cr ite rio n Spe ci e s m a y be as si gn ed to m ono ph yl et ic cl ad es D e te ct p o p u la ti on di ff er en ti at io n (g en et ic is ol at io n) a m on g cl u ste rs (u si n g P opu la tion D iffe ren tia tio n A na lys is in M ulti lo cu s, ө ) In con gr u en t C lo na lity Ac co rd ing to GCP S C Inf e r p opu la tion ( o r c lus te r) n um be r (u si ng S tr uctur e sof twar e) C on grue n t (c lo na lity) R e co n stru ct ph yl og e neti c tr ee o f co mb in ed mul til oc us data s ets R e co m bin a tio n S peci es m ay be a ssi gn e d to re co m bi ni n g cl u ste rs De te c t c ongruenc e am ong ge ne lin e ages (p er fo rm ed with P a rtiti on H om o gene ity T e st in PA U P pr og ra m) D et ect hom op la sy with in lo cu s (ca lc u la te C I, H I in PA UP) C om par e c lus te rin g o bt ai ne d w ith d iff er en t met ho ds , de ci de cl u st er n um be rs an d in di vi du al di st rib ut ion in each cl ust er D e te ct r epr od uc tiv e m ode s of cl ade s in the tre es Fi g. 1. T he s che m e f or r eco gn iz ing fu ng al s pe ci es wi th th e cr it er io n of P S R a nd GCP SR
Outline of this thesis
The diagnoses which can leads to ultimately efficient treatment are based on correct recognition of species. Therefore, it is important to investigate not only genetic characteristics but also phenotypic features obtained by traditional taxonomic methods to gain a full
understanding of the species at hand. This involves at least morphology, physiology, detailed ecology (including route of infection and predilection), pathogenicity and antifungal susceptibility of species.
Chapter 2 of this Thesis delineates variability within the Ps. boydii complex which was investigated at different levels of diversity by multilocus gene sequencing and Restriction Fragment Length Polymorphism (RFLP) analysis. In order to explore new species and try to find more evidence supporting recently described species with the criterion of GCPSR, concordance among multi-locus lineages was tested by PHT; population differentiation among clusters and recombination within partial clusters were detected in evolutional history of the complex.
Chapter 3 contains a novel species E. xenobiotica, separated from E. jeanselmei on the basis of the divergency of ITS region of rDNA, partial elongation factor 1-α and β-tubulin genes with supporting by distinct ecology. The species is found to have a predilection in human infection (Chapter 4), in addition to occurrence with xenobiotic aromates.
With re-identification of 188 clinical isolates of Exophiala from the USA by sequencing the ITS region of rDNA gene, the spectrum of clinically relevant Exophiala species was investigated for the first time (Chapter 4). The involved Exophiala species showed different pathogenic predilections ranging from superficial infection to systemic mycoses. The main species are found to be clonal (Chapter 5), and the data largely support the present taxonomy of Exophiala species in the clinics.
Because taxonomy of Exophiala recently has been mainly based on sequence diversity of the ITS region of rDNA gene, it is necessary to know if the diversity of the ITS region is concordant with those of other genes, and if sequencing the ITS region can be used as a reliable tool for species identification. As a representative of the genus Exophiala, species in the E. spinifera clade were chosen for this research (Chapter 5). Sequences of four
algorithms. To find specific borderlines with GCPSR, reproductive isolation within the clade and reproductive modes of the species were also determined.
Summarizing diagnostic features of morphology, physiology, immunology and
genetics of species belonging to the E. spinifera clade, it is concluded that sequencing the ITS region is reliable method for identification of the species in this clade, and the ITS region is therefore a good candidate for barcodes of Exophiala (Chapter 6).
Having reliably identified the opportunistic taxa, tools should be developed to aid the clinician to provide an optimal therapy to the patients. To this aim, methods for antifungal susceptibility testing were adjusted to Pseudallescheria (Chapter 7).
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Table 1. Checklist of names in Pseudallecheria and Scedosporium with indication of type strains
and cross reference numbers (The present names are printed in bold. Names are listed in alphabetic order of epithet.)
• Allescheria boydii Shear, 1922, Shear [Mycologia 14: 239] • ≡ Petriellidium boydii (Shear) Malloch, 1970 [Mycologia 62: 727]
• ≡ Pseudallescheria boydii (Shear) McGinnis, A.A. Padhye & Ajello 1982 [Mycotaxon 14: 94] o Type strain: CBS 101.22 = IHEM 15933 = IMI 015407 = IP 1975.91 = JCM 7441 =
NCPF 2216 = UAMH 3982man, isolated from mycetoma of a human patient in Galveston, Texas, USA by M.F. Boyd in 1921.
• Petriellidium africanum Arx & G. Franz, 1973 [Persoonia 7: 367]
• ≡ Pseudallescheria africana (Arx & G. Franz) McGinnis, A.A. Padhye & Ajello, 1982 [Mycotaxon 14: 94]
o Type strai: CBS 311.72 = UAMH 4000, isolated from brown sandy soil, 25 km W of Tsintsabis , Namibia by G. Franz; herbarium: CBS H-15815.
• Petriellidium angustum Malloch & Cain, 1972 [Can. J. Bot. 50: 66]
• ≡ Pseudallescheria angusta (Malloch & Cain) McGinnis, A.A. Padhye & Ajello, 1982 [Mycotaxon 14: 94]
o Type strain: CBS 254.72 = ATCC 22956 = IHEM 4429 = RV 57007 = TRTC 45321 = UAMH 3984, isolated from sewage half digestion tank in Dayton, Ohio, USA by W.B. Cooke.
• Petriellidium desertorum Arx & Moustafa, 1973 [Persoonia 7: 367]
• ≡ Pseudallescheria desertorum (Arx & Moustafa) McGinnis, A.A. Padhye & Ajello, 1982 [Mycotaxon 14: 94]
o Type strain: CBS 489.72 = UAMH 3993, isolated from salt-marsh soil in Kuwait by A.F. Moustafa.
• Petriellidium ellipsoideum Arx & Fassatiová, 1973 [Persoonia 7: 367]
• ≡ Pseudallescheria ellipsoidea (Arx & Fassatiová) McGinnis, A.A. Padhye & Ajello, 1982 [Mycotaxon 14: 94]
o Type strain: CBS 418.73 = UAMH 3987, isolated from soil in Tajikistan by O. Fassatiová.
• Petriellidium fimeti Arx, Mukerji & N. Singh, 1978 [Persoonia 10: 23]
• ≡ Pseudallescheria fimeti (Arx, Mukerji & N. Singh) McGinnis, A.A. Padhye & Ajello, 1982 [Mycotaxon 14: 94]
o Type strain: CBS 129.78, isolated from dung of goat in Aligarh,India by K.G. Mukerji, 1976, herbarium: CBS H-7549.
• Petriellidium fusoideum Arx, 1973 [Persoonia 7: 367-375]
• ≡ Pseudallescheria fusoidea (Arx) McGinnis, A.A. Padhye & Ajello, 1982 [Mycotaxon 14: 94]
o Type strain: CBS 106.53 = ATCC 11657; CDC A-722; UAMH 3997, isolated from soil in Panama, Guipo by L. Ajello.
• Pseudallescheria minutispora Gilgado, Gené, Cano & Guarro, 2005 [J. Clin. Microbiol. 43: 4930]
o Type strain: IMI 392887 = FMR 4072 = IHEM 21148 = CBS 116911, isolated from sediment of Tordera river, Barcelona, Spain.
• Scedosporium apiospermum Sacc. ex Castell. & Chalm., 1919 [Manual of Tropical Medicine.
3rdedn. p. 1122]
• = Scedosporium sclerotiale (Pepere) Neveu-Lem., 1921 [Précis de Parasitologie humaine, ed 5, p. 86]
• telemorph Pseudallescheria boydii (Shear) McGinnis, A.A. Padhye & Ajello 1982 • Scedosporium aurantiacum Gilgado, Gené, Cano & Guarro, 2005 [J. Clin. Microbiol. 43:
4930]
o Type strain: IMI 392886 = FMR 8630 = IHEM 21147 = CBS 116910, isolated from ankle ulcer of a human patient by J.L. Taboada in Hospital Clinico Universitario de Santiago, Santiago de Compostela, Spain.
• ≡ Monosporium magalhaesii (H.P. Fröes) C.W. Dodge, 1935 [Dodge, C.W. 1935, Medical mycology]
o Type strain: probably not preserved
• Lomentospora prolificans Hennebert & B.G. Desai, 1974 [Mycotaxon 1: 45] • = Scedosporium inflatum Malloch & Salkin, 1984 [Mycotaxon 21: 247]
• ≡ Scedosporium prolificans (Hennebert & B.G. Desai) E. Guého & de Hoog, 1991 [J. Mycol. Méd. 1: 8]
o Type strain: CBS 467.74 = IHEM 5739 = IMI 188615 = MUCL 18141, isolated from greenhouse soil, from mixed forest litter in Heverlee, Belgium by B.G. Desai in 1971; herbarium: CBS H-7308 (isotype), herbarium: G.L.H. 18141 (holotype)
• Pseudallescheria mesopotamicum ined.(mentioned in database of the CBS collection) • Scedosporium frequentans (Gilgado et al, in press)
• Scedosporium dehoogii (Gilgado et al, in press)
Table 2. Checklist of names in Exophiala (26 species) with indication of type strains (This list was
made as a supplement to that from of J.M. J. Uijthof (Uijthof 1995). The present species names are printed in bold. Names are listed in alphabetic order of epithet.)
• Exophiala alcalophila Goto & Sugiy [Trans. Mycol. Soc. Japan 22: 430]
o Type strain: CBS 520.82 = IAM 12519, isolated from soil in Hirose, Wako-shi, Saitama Pref., Japan.
• Exophiala angulospora Iwatsu, Udagawa & T. Takase [Mycotaxon 41: 321-328] o Type strain: CBS 482.92 = NHL 3101, isolated from drinking well water in
• Exophiala attenuata Vitale & de Hoog, 2003 [Med. Mycol. 40: 554]
o Type strain: CBS 101540 = IFM 46115, isolated from soil, Colombia.
• Torula bergeri Langeron in berger & langeron, 1949 [Annls Parasit. Hum. Comp. 24: 597] • ≡ Pullularia bergeri (Langeron) Seeliger, Silva Lacaz & Ulson, 1959 [Proc. Int. Congr. Trop.
Med. Malar. 6: 641]
• ≡ Exophiala bergeri Haase & de Hoog, 1999 [Stud. Mycol. 43: 91]
o Type strain: CBS 353.52 = NCMH 159 = Duke 978, isolated from human chromomycosis in Canada.
• Exophiala salmonis J.W. Carmichael, 1966 [Sabouraudia 5: 120] • = Exophiala brunnea Papendorf, 1969 [Trans. Br. Mycol. Soc. 52: 487] • = Aureobasidium salmonis (J.W. Carmichael) Borelli, 1969 [ ? ]
o Type strain: CBS 587.66 = ATCC 32288 = PRE 43729, isolated from Acacia karroo (Leguminosae-mimosoideae) leaf litter in Potchefstroom, South Africa; herbarium: CBS H-12618.
• Sporocybe calicioides Fr. 1832 [Syst. Mycol. (Lundae) 3: 342] • ≡ Periconia calicioides (Fr.) Berk.
• ≡ Hypsotheca calicioides (Fr.) Ellis & Everh., 1885
• ≡ Graphium calicioides (Fr.) Cooke & Massee, 1887 [Grevillea 16: 11] • ≡ Exophiala calicioides (Fr.) G. Okada & Seifert, 2000
o Type strain: unknown
• Microsporum mansonii Castellani, 1905 [Brit. Med. J. 2: 1271]
• ≡ Foxia mansonii (Castellani) Castellani, 1908 [J. Trop. Med. Hyg. 11: 261]
• ≡ Malassezia mansonii (Castellani) Verdun, 1912 [Précis Parasitol. Hum., éd 2,p. 698] • ≡ Cladosporium mansonii (Castellani) Castellani & Chalmers, 1919 [Man. Trop. Med. p.
1100]
• ≡ Torula mansonii (Castellani) Vuillemin, 1929 [C. r. hebd. Séanc. Acad. Sci., Paris 89: 406] • ≡ Sporotrichum mansonii (Castellani) Toro, 1932 [Scient. Surv. P. Rico 8:222]
• ≡ Dematium mansonii (Castellani) C.W. Dodge, 1935 [Med. Mycol. p. 678]
• ≡ Rhinocladiella mansonii (Castellani) Schol-Schwarz, 1968 [Antonie van Leeuwenhoek 34: 122]
• ≡ Exophiala mansonii (Castellani) de Hoog, 1977 [Stud. Mycol. 15: 114]
• ≡ Wangiella mansonii (Castellani) McGinnis ex Bièvre & Mariat, 1979 [Bull. Soc. Fr. Mycol. Méd. 8: 127]
• ≡ Exophiala castellanii Iwatsu, Nishimura & Miyaji, 1984 [Mycotaxon 20: 307]
• ≡ Exophiala jeanselmei var. castellanii (Iwatsu, Nishimura & Miyaji) Iwatsu & Udagawa, 1990 [Mycotaxon 37: 292]
o Type strain: CBS 158.58 = ATCC 18657 = IFM 4702 = MUCL 10097, isolated from skin scrapings of human patient in Sri Lanka; herbarium: CBS H-7132 (isotype), CBS H-7133 (isotype)
• Exophiala crusticola Bates, Gundlapally & Garcia-Pichel, 2006 [Int. J. Syst. Evol. Microbiol. 56: 2697]
o Type strain: CBS 119970 = ATCC MYA-3639 = DSM 16793, isolated from biological soil crust sample in Colorado Plateau by S.N.R. Gundlapally, USA; herbarium: UAMH 10686 (holotype); MycoBank number MB501062.
• Hormiscium dermatitidis Kano, 1934 [Aichi Igakukwai Zasshi 41: 1668] • ≡ Fonsecaea dermatitidis (Kano) Carrión, 1950 [Arch. Derm. Syphil. 61:1008]
• ≡ Hormodendrum dermatitidis (Kano) Conant in Conant, 1954 [Man. Clin. Mtcol. ed. 2 p. 276]
• ≡ Phialophora dermatitidis (Kano) Emmons, 1963 [Med. Mycol. p. 291] • ≡ Exophiala dermatitidis (Kano) de Hoog [Stud. Mycol. 15: 118] • ≡ Wangiella dermatitidis McGinnis, 1977 [Mycotaxon 5: 355]
o Type strain: CBS 207.35 = ATCC 28869 = DUKE 2400 = IFO 6421 = IMI 093967 = LSHTM 1135 = NCPF 2422 = UAMH 3967, isolated from human facial
chromomycosis in Japan; herbarium: CBS H-7131. • Exophiala dopicola Katz & McGinnis [Mycotaxon 11: 182]
o Type strain: CBS 537.94 = BAK 978, isolated from Pinus taeda litter (Pinaceae) in Duke forest, Orange County, North Carolina, USA by B. Katz in 1977.
• ≡ Phaeococcomyces exophialae (de Hoog) de Hoog, 1979 [Taxon 28: 348] • ≡ Exophiala exophialae (de Hoog) de Hoog, 2003 [J. Clin. Microbiol. 41: 4767]
o Type strain: CBS 668.76 = ATCC 26088, isolated from straw in armadillo burrow (Dasypus septemcinctus), Uruguay by J.E. MacKinnon; herbarium: CBS H-7551 • Exophiala equina (still to be reported)
o Exophiala pisciphila clade
• Trichosporum heteromorphum Nannfeldt, 1934 [Svenska Skogsv.-Fören. Tidskr. 32: 397] • = Margarinomyces heteromorphus (Nannfeldt) F. Mangenot, 1952 [Revue. Gén. Bot. 59: 391] • ≡ Phialophora heteromorpha (Nannfeldt) C.J.K. Wang, 1964 [Can. J. Bot. 42: 1011]
• ≡ Exophiala jeanselmei var. heteromorpha (Nannfeldt) de Hoog, 1977 • ≡ Wangiella heteromorpha (Nannfeldt) McKemy, 1999
• ≡ Exophiala heteromorpha (Nannfeldt) de Hoog & Haase [J. Clin. Microbiol. 41: 4767] o Type strain: CBS 232.33 = CDC B-2823 = MUCL 9894 = NCMH 17 = VKM F-704,
isolated from wood pulp in Sweden by E. Melin, herb CBS H-17795. • Torula jeanselmei Langeron, 1928 [Ann. Paras. hum. Comp. 6: 385]
• ≡ Torula sp., Jeanselma, 1928 [Bull. Soc. Fr. Dermatol. Syphiligr. 35: 369]
• ≡ Pullularia jeanselmei (Langeron) C.W. Dodge, 1935 [Medical Mycology p. 675] • ≡ Phialophora jeanselmei (Langeron) C.W. Emmons, 1945 [Arch. Pathol. 39: 368] • ≡ Exophiala jeanselmei (Langeron) McGinnis & A.A. Padhye, 1977 [Mycotaxon 5: 345] • ≡ Exophiala jeanselmei var. jeanselmei, 1977 [Stud. Mycol. 15: 108]
o Type strain: CBS 507.90 = ATCC 34123 = CBS 664.76 = DUKE 2405 = IAM 14677 = IHM 283 = NCMH 123 = NCPF 2439, isolated from human mycetoma in Uruguay. • Torula lecanii-corni Benedek & G. Specht, 1933 [Zentbl. Bakt. Parasitkde, Abt. 1, 130: 74] • = Pullularia fermentans var. benedekii E.S. Wynne & Gott, 1956 [J. Gen. Microbiol. 14: 512] • ≡ Exophiala jeanselmei var. lecanii-corni (Benedek & G. Specht) de Hoog, 1977[Stud. Mycol.
15: 112]
• ≡ Exophiala lecanii-corni (Benedek & G. Specht) Haase & de Hoog, 1999 [Stud. Mycol. 43: 80]
o Type strain: CBS 123.33 = ATCC 12734 = IMI 062462, isolated as symbiont of
Lecanium corni (scale bug).
• Exophiala mesophila Listemann & Freiesleben, 1996 [Mycoses 39: 1]
o Type strain: CBS 402.95, isolated from silicone seal in shower room of hospital in Hamburg, Germany.
o Exophiala lecanii-corni clade
• Exophiala moniliae de Hoog, 1997 [Stud. Mycol. 15: 120]
o Type strain: CBS 520.76, isolated from twig of Quercus sp.(Fagaceae) in St. Petersburg, Russia; herb: CBS H-7134.
• Nadsoniella nigra Issatschenko, 1914 [Annu. Exped. Sci. Mourmansk, 1906, p. 273] • ≡ Exophiala nigra (Issatschenko) Haase & de Hoog, 1999 [Stud. Mycol. 43: 91]
o Type strain: CBS 535.94, isolated from seawater (5-10 m), Kolskiy near Murmansk, Harbour Ekaterninskaya (naval base Poliarny), Russia.
• Exophiala nishimurae Vitale & de Hoog [Med. Mycol. 40: 545]
o Type strain: CBS 101538 = IFM 41855, contaminant in Exophiala spinifera IFM 41855 from bark in Venezuela.
• Melanchlenus oligospermus Calendron, 1953 [C. R. Acad. Sci. 236: 1598]; invalid.
o Type strain: CBS 265.49 = MUCL 9905, isolated from honey near St. Domineuc, Ille & Vilaine, France by A. Calandron.
• ≡ Exophiala oligosperma Calendron ex de Hoog & Tintelnot, 2003 [J. Clin. Microbiol. 41: 4767]
o Type strain: CBS 725.88, isolated from human patient of tumour of sphenoidal cavity, Germany, Würzburg by K. Tintelnot, 1988.
• Sarcinomyces phaeomuriformis Matsumoto, A.A. Padhye, Ajello & McGinnis, 1986 [J. Med. Vet. Mycol. 24: 395]
• ≡ Exophiala phaeomuriformis (Matsumoto, A.A. Padhye, Ajello & McGinnis) Matos, Haase & de Hoog, 2003 [Antonie van Leeuwenhoek 83: 293]
o Type strain: CBS 131.88 = CDC B-3558 = NCMH 1215 = UAMH 4278, isolated from human phaeohyphomycosis, Japan.
• Exophiala pisciphila McGinnis & Ajello, 1974 [Mycologia 66: 518]
o Type strain: CBS 537.73 = ATCC 24901 = CDC B-1229 = IHEM 3404 = IMI 176060 = IP 1670.86 = NCMH 9 = UAMH 2981, isolated from Channel catfish (Ictalurus
punctatus) in Central Alabama, Alabama, USA; herb: CBS H-7135.
• Pullularia prototropha Bulanov & Malama, 1965 [Vesci Akad. Naruk Belarus. SSR 4: 116] • ≡ Exophiala prototropha (Bulanov & Malama) Haase, Yurlova & de Hoog, 1999 [Stud.
Mycol. 43: 93]
o Type strain: CBS 534.94, source unknown.
• Exophiala psychrophila Pedersen & Langvad, 1989 [Mycol. Res. 92: 153]
o Type strain: CBS 191.87 = ATCC 62848, isolated from kidney granuloma of Salmo
salar.
• Exophiala salmonis J.W. Carmichael, 1966 [Sabouraudia 5: 120]
• ≡ Aureobasidium salmonis (J.W. Carmichael) Borelli, 1969 [Medicina Cutanea 3: 588] • = Exophiala brunnea Papendorf, 1969 [Trans. Br. Mycol. Soc. 52: 487]
o Type strain: isotype CBS 157.67 = ATCC 16986 = IHEM 3405 = IMI 124165 = MUCL 10078 = UAMH 34 = VKM F-3000, isolated from cerebral mycetoma of
Salmo clarkia in Alberta hatchery, Calgary, Alberta, Canada; herb: CBS H-12617,
CBS H-7136.
• Phialophora spinifera H.S. Nielsen & Conant, 1968 [Sabouraudia 6: 228]
• ≡ Rhinocladiella spinifera (H.S. Nielsen & Conant) de Hoog, 1977 [Stud. Mycol. 15: 93] • ≡ Exophiala spinifera (H.S. Nielsen & Conant) McGinnis, 1977 [Mycotaxon 5: 337]
o Type strain: CBS 899.68 = ATCC 18218 = DSM 1217 = DUKE 3342 = IHM 1767 = NCMH 152 = NCPF 2358, isolated from nasal granuloma of a human patient form USA.
• Exophiala xenobiotica de Hoog, 2006 [Antonie van Leeuwenhoek 90: 257]
Reference:
J. M. J. Uijthof (1995). Taxonomy and phylogeny of the human pathogenic black yeast genus
Exophiala Carmichael. PhD. Dissertation, Centraalbureau voor Schimmelcultures Fungal Biodiversity
Centre, Royal Netherlands Academy of Arts and Sciences and Institute of Molecular Cell Biology, University of Amsterdam.
Collection acronyms:
ATCC: American Type Culture Collection, Rockville, Maryland, USA CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands CDC: Centers for Disease Control and Prevention, Atlanta, Georgia, USA
DSM: Deutsche Sammlung von Mikroorgannismen und Zellkulturen, Baunschweig, Germany DUKE: Duke Medical Center, North Carolina, USA
IAM: Institute of Applied Microbiology, Tokyo, Japan
IFM: Research Institute for Pathogenic Fungi and Microbial Toxicoses, Chiba University, Chiba, Japan
IFO: Institute for Fermentation, Osaka, Japan
IHEM: Institute of Hygiene and Epidemiology, Brussels, Belgium IMI: CAB International Mycological Institute, Egham, UK IP: Institute Pasteur, Paris, France
LSHTM: London School of Hygiene and Tropical Medicine, London, UK
MUCL: Mycothèque de l’Université Catholique de Louvain, louvain-la-Neuve, Belgium NCMH: North Carolina Memorial Hospital, Chapel Hill, USA
NCPF: National Collection of Pathogenic Fungi, Public Health Laboratory Service, Mycological Reference Laboratory, London, UK
NHL: national Hygiene laboratory, National Institute of Hygiene Services, Tokyo, Japan PRE: National Herbarium, Botanical Research Institute, Pretoria, South Africa
UAMH: University of Alberta, Microfungus Herbarium and Collection, Edmonton, Canada VKM: All-Union Collection of Micro-organisms, Moscow, Russia
Intraspecific diversity of species of the
Pseudallescheria boydii complex
J.S. ZENG a,b,c,d, K. FUKUSHIMA a, , K. TAKIZAWA a, Y. C. ZHENG b, K. NISHIMURA a, Y. GRÄSER e & G. S. DE HOOG c,d
a Research Center for Pathogenic Fungi and Microbial Toxicoses, Chiba University, Chiba,
Japan; b Department of Dermatology and Venereology, Union Hospital, Tongji Medical
College, Huazhong Science and Technology University, Wuhan, Hubei, P. R. China; c
Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; d Institute for Biodiversity
and Ecosystem Dynamics, University of Amsterdam, Amsterdam, The Netherlands; eInstitute
for Microbiology and Hygiene, Department of Parasitology (Charité), Humboldt University, Berlin, Germany
Accepted by Medical Mycology in May, 2007
Keywords: Pseudallescheria boydii, Scedosporium apiospermum, LSU, ITS, IGS, RFLP, elongation factor, phylogeny, population structure, recombination
Summary
In order to establish intraspecific diversity of Pseudallescheria boydii and Scedosporium apiospermum, and to develop tools for identification, variability within P. boydii and related species was investigated at different levels of diversity. Sequences of the D1/D2 region of large subunit (LSU) and of the internal transcribed spacer (ITS) region of ribosomal DNA (rDNA) gene were analyzed for a set of 57 strains, as well as partial sequences of the elongation factor 1-α (EF 1-α ) gene. Incongruence among 3 locus lineages was detected by partition homogeneity test (PHT). The maximum parsimony (MP) tree of the combined sequence data set, with the exception of strain CBS 499.90, formed 3 clades with high bootstrap support, correspondeding to previously described nuclear DNA (nDNA) /DNA reassociation groups. These groups are known to differ slightly in predilection and
temperature relations. Using STRUCTURE software, population genetic analysis revealed 3 clusters within the complex on the basis of multi-locus genotype data set. Strain distribution in the clusters was concordant with that in the 3 clades of combined multi-locus MP tree. Recombination among individuals of a clade in evolutional history was found in 2 of the 3 clades. There was population differentiation among the 3 clades. Restriction fragment length polymorphism (RFLP) analysis of the intergenic spacer (IGS) region of rDNA gene was added to further characterize subspecific entities. When the IGS regions of 22 strains were digested with the restriction endonucleases Hae III and Mbo I, seven and five distinct patterns were detected, respectively. This subtyping did not reveal any correspondence with
geographic origin or clinical appearance. Though we need more evidence to locate the 3 clades of the P. boydii complex at species or population level, the sequence of the D1/D2 region is sufficiently variable for identification of taxa belonging to the P. boydii complex.
Introduction
Pseudallescheria boydii and the related anamorph Scedosporium apiospermum are among the emerging agents of opportunistic mycoses [1]. Longtime the complex had been listed as a single species as an agent of human mycetoma, but colonization of the airways in patients with cystic fibrosis is increasingly reported [2], as well as systemic disorders such ascerebral infections after near-drowning [3,4]. It may be difficult to recognize the species in the clinic
because of its histopathological similarity to aspergillosis [5]. In vitro, considerable genetic variability within P. boydii is known, not only in data from genetic fingerprinting [6], but also in less variable genes such as small subunit (SSU) and large subunit (LSU) of ribosomal DNA (rDNA) [6-8] and genomic homology [9]. With the latter technique, three approximate
intraspecific groups were recognized, but thus far these could not be confirmed
unambiguously using other techniques [6]. Gilgado and coworkers [10] are in the process of attributing formal taxonomic entities to a number of these groups.
The D1/D2 domain of LSU region of rDNA gene is considered to be relatively conserved, allowing identification of fungi at the species level. Issakainen and coworkers [11, 8]
presented the phylogeny of P. boydii and related taxa on the basis of this region. Nevertheless intraspecific polymorphism was noted in the LSU region. In the present paper we will
establish whether this variability is also expressed at lower levels of diversity by sequenceing the internal transcribed spacer (ITS) region of rDNA gene and the elongation factor 1-α (EF 1-α) gene, and by restriction fragment length polymorphism (RFLP) analysis of the
intergenetic spacer (IGS) region of rDNA gene, and whether independent polymorphisms at these different levels of diversity are concordant. These data add to our understanding of breeding systems in the teleomorph species complex P. boydii that is known to produce different types of anamorph, morphologically classified in Scedosporium and Graphium.
Materials and Methods
StrainsFifty seven strains analyzed are listed in Table 1, 38 of which were previously identified as P. boydii or S. apiospermum on the basis of morphology. Seven reference strains of P. boydii and 12 reference strains of related species were included for comparison. Some additional
sequences were downloaded from GenBank.
DNA extraction
DNA was prepared with 6% InstaGene matrix kit (Bio-Rad,Hercules, Calif.). A small amount of fungal pellet was suspended in 200 μl of InstaGene matrix and incubated at 56°C for 30 min. The mixture was heated at 100°C in a water bath for 8 min and centrifuged at 12,000 r.p.m. for 3 min at room temperature. The supernatant was transferred to a new tube.
Hae III Mbo I
P. boydii IFM 52858 patient China Clade 5A Clade 5 Clade 5A A A
P. boydii IFM 52864 unknown Clade 5A Clade 5 Clade 5A A A
P. boydii IFM 48461 pleural fluid Japan Clade 5A Clade 5 Clade 5A B B
P. boydii IFM 52861 patient China Clade 5A Clade 5 Clade 5A B B
P. boydii IFM 52865 unknown Clade 5A Clade 5 Clade 5A B B
P. boydii IFM 47161 patient Japan Clade 5A Clade 5 Clade 5A C B
P. boydii IFM 41911 soil Colombia Clade 5A C B
P. boydii IFM 50914 bronchoalveolar lavage Japan Clade 5A Clade 5 Clade 5A D B
P. boydii IFM 49724 lung Japan Clade 5A Clade 5 Clade 5A D B
P. boydii IFM 52862 necrotic tissue of eye China Clade 5A Clade 5 Clade 5A D B
P. boydii IFM 41901 soil Colombia Clade 5A D B
P. boydii IFM 48046 unknown Clade 5A D B
P. boydii IFM 50907 lung and brain tissues Japan Clade 5A Clade 5 Clade 5A
P. boydii IFM 41585 patient Japan Clade 5A Clade 5 Clade 5A
P. boydii IFM 52863 cerebrospinal fluid China Clade 5A Clade 5 Clade 5A
P. boydii IFM 52875 patient Spain Clade 5A
P. boydii CBS 116892 sputum France Clade 5A Clade 5 Clade 5A 1
P. boydii CBS 101.22 T mycetoma USA Clade 5A Clade 5* Clade 5A 1
P. ellipsoidea CBS 418.73 T soil Tajikistan Clade 5A Clade 5* Clade 5A
P. boydii CBS 108.54 soil Zaire Clade 5B Clade 5 Clade 5B 3
P. boydii CBS 116894 soil Thailand Clade 5B Clade 5 Clade 5B 3
P. boydii IFM 52874 litter Brazil Clade 5B
P. boydii IFM 52860 patient China Clade 5B Clade 5 Clade 5B
P. boydii IFM 47302 skin Japan Clade 5B Clade 5 Clade 5B
P, angusta CBS 254.72 T sewage half digestion tank USA Clade 5B Clade 5* Clade 5B
P. fusoidea CBS 106.53 T soil Panama Clade 5B Clade 5* Clade 5B
P. boydii IFM 52873 cerebrospinal fluid China Clade 4 Clade 5 Clade 4 E E
P. boydii IFM 52859 paranasal sinus biopsy China Clade 4 Clade 4 Clade 4 E E
P. boydii IFM 52928 necrosis tissue of eye China Clade 4 Clade 4 Clade 4 F D
P. boydii IFM 52930 patient Italy Clade 4 G D
P. boydii IFM 50000 synovia Japan Clade 4 Clade 4 Clade 4 G D
P. boydii IFM 52028 paranasal sinus biopsy Japan Clade 4 Clade 4 Clade 4 G C
P. boydii IFM 49770 sputum Japan Clade 4 Clade 4 Clade 4 G C
P. boydii IFM 52929 patient Italy Clade 4 G C
P. boydii IFM 49731 unknown Clade 4 Clade 4 Clade 4 G C
P. boydii IFM 41921 soil Colombia Clade 4 G C
P. boydii IFM 46992 bronchoalveolar lavage Japan Clade 4 Clade 4 Clade 4
P. boydii IFM 51940 pleural fluid Japan Clade 4 Clade 4 Clade 4
P. boydii IFM 46993 bronchial polyp brushing Japan Clade 4 Clade 4 Clade 4
P. boydii IFM 49188 bronchoalveolar lavage Japan Clade 4 Clade 4 Clade 4
P. boydii IFM 50256 corneal scraping Japan Clade 4 Clade 4 Clade 4
P. boydii IFM 50231 corneal scraping Japan Clade 4 Clade 4 Clade 4
P. boydii CBS 116779 sinus France Clade 4 Clade 4 Clade 4 2
P. boydii CBS 695.70 nasal cavity of pig Ukraine Clade 4 Clade 4* Clade 4 2
P. boydii IFM 40911 unknown Clade 4 Clade 4 Clade 4
P. boydii IFM 41595 unknown Clade 4 Clade 4 Clade 4
P. boydii IFM 41923 soil Colombia Clade 3
P. boydii CBS 499.90 mud of pond Netherlands Clade 3 Clade 3* Clade 3 2
G. tectonae CBS 127.84 T tectona grandis, seed Jamaica *
P. desertorum CBS 489.72 T salt-marsh soil Kuwait *
P. minutispora CBS 116911 T river sediment Spain *
S. aurantiacum CBS 116910 T ulcer of ankle Spain *
S. aurantiacum CBS 118934 leg biopsy Netherlands
S. prolificans CBS 467.74 T soil Belgium *
S. prolificans CBS 452.89 blood of patient France
P, africana CBS 311.72 T soil Namibia *
P. fimeti CBS 129.78 T dung India *
D1/D2
groupb,# ITS groupb EF groupb
Table 1. Strains examined and results of sequencing and RFLP analysis.
nDNA/D NA d
Species Strain a Source Geography
RFLP type of IGSc
a Abbreviations: IFM = Research Center for Pathogenic Fungi and Microbial Toxicoses, Chiba University, Chiba, Japan; CBS = CBS Fungal
Biodiversity centre, Utrecht, The Netherlands; b Groups obtained by sequencing rDNA D1/D2, ITS region and partial EF 1-α gene; group
names, Clades 3-5 followed those in Gilgado's research (2005); c Restriction pattern of IGS region obtained with HaeIII and MboI.; d Data
from Guého & de Hoog (1991); # sub-groups Clades 5A and 5B based on substitutions in positions 140 and 406 in the alignment of D1/D2
region of rDNA gene; * ITS sequences were downloaded from GenBank. Accession number of strains CBS 101.22: AJ888435; CBS 418.73: AJ888426; CBS 254.72: AJ888414; CBS 106.53: AJ888428; CBS 695.70: AY877353; CBS 499.90: AY878952; CBS 127.84: AY228113;
Sequencing and data analysis
Three primer sets NL1 (5’-GCATATCAATAAGCGGAGGAAAAG-3’) and V3-12 (modified NL4, 5’-GGTCCGTGTTTCAAGACG-3’), ITS1 (5’-TCC GTA GGT GAA CCT GCG G-3’) and ITS4 (5’-TCC TCC GCT TAT TGA TAT GC-3’), EF1-728F (5’-CAT CGA GAA GTT CGA GAA GG-3’) and EF1-986R (5’-TAC TTG AAG GAA CCC TTA CC-3’) were used to amplify and sequence the D1/D2 and ITS regions of rDNA gene, and part of the EF 1-α gene, respectively. If no amplicon of the ITS region was obtained, the primers ITS1 and ITS4 were changed to V9G (5’-TTA CGT CCC TGC CCT TTG TA-3’) and LS266 (5’-GCAT TCC CAA ACA ACT CGA CTC-3’). Amplification of the D1/D2 region was performed as follows: 95°C for 4 min, followed by 35 cycles consisting of 94°C for 45 sec, 58°C for 30 sec and 72°C for 2 min. The annealing temperature was changed to 52 and 55°C, respectively when amplifying the ITS region and the partial EF 1-α genes. The amplified products were purified with
SUPRE-CTM-02 kit (Takara Shuzo Co., Shiga, Japan) and were subjected to direct sequencing with an ABIPRISM 3100 sequencer after labeled with ABI PRISM BigDyeTM terminator cycle sequencing standard (Applied Biosystems, Tokyo, Japan).
Sequences were adjusted using SeqMan Π of Lasergene software (DNASTAR, Inc.). Then they were aligned iteratively and cluster analyses were performed using Ward’s averaging in the BioNumerics package v. 4.0 (Applied Maths, Kortrijk, Belgium). Nearest neighbours were found by local Blast searches. By using the DCSE program [12], the sequences of each locus and combined multi-locus data set were re-aligned. Maximum Parsimony (MP) trees of each gene and combined gene data were constructed using PAUP v. 4.0b10 [13]. A heuristic search was performed for each dataset with 100 random taxon additions and tree bisection and reconstruction (TBR) as the branch swapping algorithm. Branches of zero-length were collapsed and all multiple, equally parsimony trees were saved. The maximum of trees saved was set as 5000. The robustness of the resulting phylogenetic tree was evaluated by 100 bootstrap replications and every replication used a maximum of 500 trees. The phylogenetic tree was printed with TreeView v. 1.6.6 [14].
The congruence of genealogies was assessed using partition homogeneity test (PHT) in PAUP v. 4.0b10 [13]based on sequences of the D1/D2 and ITS regions of rDNA gene and on partial sequence of the EF 1-α gene.
IGS RFLP analysis
Amplification of the IGS region of 45 P. boydii strains was performed with KOD-Plus-DNA Polymerase (Toyobo Co., Osaka, Japan) with primers IGSL.fw (5'-TAG TAC GAG AGG AAC CGT-3') and IGSR.bw (5'-GCA TAT GAC TAC TGG CAG-3') [15]. The resulting amplicons were purified with ethanol and digested with the restriction endonucleases HaeIII and MboI (Takara Shuzo Co., Shiga, Japan) as recommended by the manufacturer. The corresponding products were electrophoresed in 3% agarose gel at 100 V for 70 min.
Population genetic analysis
In order to confirm the intraspecific diversity shown in the MP trees, the number of populations in the P. boydii complex was inferred with STRUCTURE software v. 2.
[http://pritch.bsd.uchicago.edu] using genotype data of the D1/D2 and ITS regions of rDNA gene and of the partial EF 1-α gene. Genotypes of these 3 loci of 38 isolates in the P. boydii complex were sorted on the basis of similarity of the sequences. The burning period length and number of MCMC repetitions after burning were set as 10000 and 100000, and admixture model and allele frequencies correlated model were chosen for analysis. The number of populations (K) was assumed from 2 to 5. To test for reproductive mode in each population, index of association (IA, a measure of multi-locus linkage disequilibrium) was calculated with MULTILOCUS 1. 2. 2 [http://www.bio.ic.ac.uk/evolve/software/multilocus]. The null
hypothesis for this analysis is complete panmixia. The values of IA were compared between observed and randomized datasets. The hypothesis would be rejected when p < 0.05. Population differentiation (index: ө) was also detected using the same software and a null hypothesis for this analysis is no population differentiation. When observed ө is statistically significantly different from those of random data sets (p < 0.05), population differentiation will be considered.
Results
Sequencing and grouping based on phylogenetic analysis
100 60 55 79 96 74 100 51 1 CBS 467.74 T S. prolificans CBS 452.89 S. prolificans CBS 101.22 T P. boydii (1) IFM 50914 IFM 41585 IFM 52861 IFM 41901 IFM 41911 IFM 52865 IFM 52864 IFM 52875 IFM 49724 IFM 48461 IFM 48046 IFM 47161 IFM 50907 IFM 52858 IFM 52862 IFM 52863 CBS 418.73 T P. ellipsoidea CBS 116892 (1) CBS 108.54 (3) CBS 106.53 T P. fusoidea IFM 52874 CBS 116894 (3) CBS 254.72 T P. angusta IFM 47302 IFM 52860 CBS 695.70 (2) CBS 116779 (2) IFM 46992 IFM 49770 IFM 50256 IFM 46993 IFM 52859 IFM 41921 IFM 49731 IFM 41595 IFM 52930 IFM 52929 IFM 52928 IFM 40911 IFM 49188 IFM 50000 IFM 50231 IFM 51940 IFM 52028 IFM 52873 CBS 116910 T S. aurantiacum CBS118934 S. aurantiacum CBS 116911 T P. minutispora CBS 489.72 T P. desertorum CBS 499.90 (2) IFM 41923 CBS 127.84 T G. tectonae CBS 311.72 T P. africana Clade 5 Clade 4 Clade 3
Figure 1. One of 26 most parsimonious trees obtained from a heuristic search with 100 random taxon additions of the D1/D2 rDNA of 56 strain sequence alignment. The scale bar shows 1 change; bootstrap support values (>50%) from 100 replicates are shown at the nodes. Thickened lines indicate the restrict consensus branches. The tree was rooted to S. prolificans CBS 467.74 T. Specise name of
P.boydii isolates is omitted. Numbers in brackets are the group numbers based on nDNA homology data used by Guého & de Hoog
