Rapid Identification of the Genus Fonsecaea by PCR with Specific Oligonucleotide Primers

Abstract

An oligonucleotide primer set based on internal transcribed spacer regions of ribosomal DNA for PCR which gives the amplicon for only the DNA from Fonsecaea species was designed. This set yielded an amplicon with 333 bp for all strains of Fonsecaea pedrosoi and Fonsecaea compacta examined but no amplicons for related dematiaceous fungi and pathogenic yeasts. PCR using this primer set was considered to be a useful method for the rapid identification of the genus Fonsecaea.

The five etiologic agents of chromoblastomycosis that have been recognized worldwide are Fonsecaea pedrosoi, the principal agent, and Phialophora verrucosa, Cladophialophora carrionii, Fonsecaea compacta, and Rhinocladiella aquaspersa, in order of detection (15, 20). In recent years, increasing numbers of chromoblastomycosis cases and invasive infections caused by F. pedrosoi have been reported (1, 4, 10, 18, 19, 21). According to some investigators, infections by F. pedrosoi account for the overwhelming majority of cases (90 to 93%) (1, 4). F. compacta is a rare cause of chromoblastomycosis (8), with only a few cases reported (1, 3). Traditional methods of identification based on morphological characteristics and antigen detection have been used clinically (7, 13). However, these methods are time-consuming and possess low specificity.

In recent years, PCR with specific primers has been used for the detection and identification of the pathogenic fungi (11, 17, 24). However, a rapid detection system for the identification of Fonsecaea species has not yet been developed. In this paper, we report the development of a rapid and specific PCR method for the detection of Fonsecaea species.

Eighty-four strains of 51 species (Table 1), including medically relevant dematiaceous fungal species and medically important pathogenic yeasts, were examined. DNA was prepared according to the method of Makimura et al. (16). About 50 mg of fungal elements was suspended in 600 μl of extraction buffer (200 mM Tris-HCl [pH 7.5], 25 mM EDTA, 0.5% [wt/vol] sodium dodecyl sulfate, 250 mM NaCl). After being mixed by vortex for 15 s, the mixture was incubated at 100°C for 15 min, kept on ice for 60 min, and centrifuged at 14,000 × g for 15 min. The supernatant was transferred to new tubes, and the solution was extracted once with phenol-chloroform-isoamyl alcohol (25:24:1 [vol/vol]). DNA was precipitated with cold isopropanol (−20°C), dried, and resuspended in 100 μl of distilled water. The internal transcribed spacer (ITS) regions were amplified with universal primers ITS5 (5′-GGAAGTAAAAGTCGTAACAAGG-3′) and ITS4 (5′-TCCTCCGCTTATTGATATGC-3′) (25). Direct sequencing of the PCR amplicons corresponding to the ITS1-5.8S-ITS2 region of ribosomal DNA (rDNA) was performed with an ABI PRISM 3100 sequencer with an ABI PRISM BigDye terminator sequencing kit (Applied Biosystems, Foster City, Calif.). External primers ITS1 (5′-TCCGTAGGTGAACCTGCGG-3′) and ITS4 were used for sequencing. Six strains of F. pedrosoi, including type strain CBS 271.37, three strains of F. compacta, and strains of Cladophialophora carrionii, Cladophialophora bantiana, Cladophialophora devriesii, Cladophialophora minourae, Phialophora verrucosa, Exophiala dermatitidis, Hortaea werneckii, R. aquaspersa, and Rhinocladiella atrovirens were sequenced. The sequences of the ITS1-5.8S-ITS2 regions of other dematiaceous fungal species and medically important yeasts were obtained from GenBank and were aligned. On the basis of these sequences, a set of oligonucleotide primers, Fon-F (forward; 5′-TAATGCGGGTGTTGCCTCTG-3′) and Fon-R (reverse; 5′-AGGGGTGGAAAGTGTGAACT-3′) was designed and was obtained from Sigma Genosys Japan kk (Tokyo, Japan). The PCR mixture (25 μl) was composed of 2.5 μl of template DNA, 2.5 μl (2 pmol) of each primer, 2 μl of (2.5 mM) deoxynucleotide triphosphate mixture (Nippon Gene, Tokyo, Japan), 0.125 μl (5 U/μl) of Taq polymerase (Nippon Gene), and 2.5 μl of 10× reaction buffer (Nippon Gene). The PCR was performed with a PCR Thermal Cycler MP (TaKaRa, Tokyo, Japan) according to the following program sequence: 95°C for 4 min, followed by 30 cycles consisting of 94°C for 1 min, 64°C for 2.5 min, and 72°C for 2.5 min, with a final extension at 72°C for 10 min. After thermal cycling, 2 μl of the amplified product was run on a 1.5% agarose gel, stained with ethidium bromide, and visualized with UV light.

TABLE 1.

Strains examined and PCR results

Species	Strain	Sourcea	Accession no.	PCR productb
Fonsecaea pedrosoi	IFM 4856d	DCU 677	AB091203	+
	IFM 4887c,d	CBS 271.37	AB087202	+
	IFM 4889d	ATCC 44356	AB091204	+
	IFM 4890	Patient		+
	IFM 4892	Patient		+
	IFM 4894	Patient		+
	IFM 4896	Patient		+
	IFM 4898	Patient		+
	IFM 4914d	Venezuela	AB091205	+
	IFM 4915	Patient		+
	IFM 4916	Patient		+
	IFM 41522	Patient		+
	IFM 41523	Patient		+
	IFM 41524	Patient		+
	IFM 41525	Patient		+
	IFM 41705d	Bark, China	AB091206	+
	IFM 41706	BMU 3339		+
	IFM 46410d	Soil, Brazil	AB091207	+
Fonsecaea compacta	IFM 4886d	KUM 911	AB091208	+
	IFM 41704d	BMU 4845	AB091209	+
	IFM 41931d	MTU	AB091210	+
Phialophora verrucosa	IFM 4928d	ATCC 38561	AB087203	−
	IFM 41710	Corn, China		−
	IFM 41871	Soil, Colombia		−
	IFM 41873	Venezuela		−
	IFM 41879	Soil, Colombia		−
	IFM 41898	Soil, Brazil		−
Phialophora alba	IFM 51363	IFO 31973		−
Phialophora americana	IFM 51361	CBS 273.37		−
Phialophora atrovirens	IFM 51364	IFO 6793		−
Phialophora bubakii	IFM 51365	IFO 6794		−
Phialophora cinerescens	IFM 51366	IFO 6849		−
Phialophora fastigiata	IFM 41577	IFO 6856		−
Phialophora heteromorpha	IFM 41578	IFO 6878		−
Phialophora lagerbergii	IFM 51367	IFO 8576		−
Phialophora melinii	IFM 51362	CBS 268.33		−
Phialophora oxyspora	IFM 51368	URM 2904		−
Phialophora repens	IFM 4925	CBS 423.73		−
Phialophora richardsiae	IFM 4926	KUM 1681		−
Phaeoacremonium parasitica	IFM 4924	KUM 1827		−
Lecythophora hoffmannii	IFM 4922	CBS 245.38		−
Lecythophora mutabilis	IFM 4923	ATCC 26223		−
Cladophialophora carrionii	IFM 4805d	ATCC 44535	AB087204	−
	IFM 4810	DCU 300		−
	IFM 4812	DCU 302		−
	IFM 41444	UNEFM/SR3		−
	IFM 41446	DCU 606		−
	IFM 41641	BMU 237		−
Cladophialophora bantiana	IFM 46165d	CBS 173.52	AB091211	−
Cladophialophora devriesii	IFM 51369c,d	CBS 147.84	AB091212	−
Cladophialophora minourae	IFM 4818d	DCU 428	AB091213	−
Cladosporium cladosporioides	IFM 41447	IFO 6368		−
Cladosporium colocasiae	IFM 51371	CBS 386.64		−
Cladosporium coralloides	IFM 41451	IFO 6536		−
Cladosporium elatum	IFM 41452	IFO 6372		−
Cladosporium fulvum	IFM 40703	IAM 5006		−
Cladosporium herbarum	IFM 41454	TMI		−
Cladosporium minusculum	IFM 51370	URM 721		−
Cladosporium resinae	IFM 51372	IFO 8588		−
Cladosporium resinae f. sp. avellaneum	IFM 41456	IFO 6367		−
Cladosporium sphaerospermum	IFM 41453	IFO 4458		−
Cladosporium variabile	IFM 41458	IFO 6378		−
Exophiala alcalophila	IFM 41654	Bark, China		−
Exophiala dermatitidis	IFM 41479c,d	CBS 207.35	AB087205	−
	IFM 41818	Venezuela		−
	IFM 41828	Soil, Brazil		−
	IFM 45986	Tap water		−
Exophiala jeanselmei	IFM 41691	BMU 2756		−
	IFM 45989	Patient		−/PICK>
Exophiala moniliae	IFM 41500c	CBS 520.76		−
Exophiala spinifera	IFM 4883c	ATCC 18218		−
Hortaea werneckii	IFM 51373d	URM 704	AB087201	−
Alternaria alternata	IFM 41348	TIMM 1289		−
Aureobasidium pullulans	IFM 4802	ATCC 15233		−
Rhinocladiella aquaspersa	IFM 4930d	CBS 313.73	AB091214	−
Rhinocladiella atravirens	IFM 4931c,d	CBS 317.33	AB091215	−
Candida albicans	IFM 40214	ATCC 90029		−
Candida dubliniensis	IFM 48313	CBS 7987		−
Candida glabrata	IFM 46888	Vagina		−
Candida parapsilosis	IFM 46863	CUH 2565		−
Candida tropicalis	IFM 46816c	CBS 94		−
Cryptococcus neoformans var. neoformans	IFM 40216	ATCC 90113		−
Malassezia furfur	IFM 48585	CBS 7019		−
Trichosporon asahii var. asahii	IFM 48429c	CBS 2479		−

^aStrain sources are as follows: ATCC strains, American Type Culture Collection, Marassus, Va.j BMU strains, Department of Dermatology, Beijing Medical University, Beijing, China; CBS strains, Central Bureau voor Schimmelcultures, Baarn, The Netherlands; CUH strains, Department of Laboratory Medicine, School of Medicine, Chiba University, Chiba, Japan; DCU, strains, Department of Dermatology, School of Medicine, Chiba University; IAM, strains, Institute of Applied Microbiology, University of Tokyo, Tokyo, Japan; IFM strains, Research Center for Pathogenic Fungi and Microbial Toxicoses, Chiba University; IFO strains, Institute for Fermentation, Osaka, Japan; KUM strains, Department of Dermatology, School of Medicine, Kanazawa University, Kanazawa, Ishikawa, Japan; MTU strains, Department of Bacteriology, Faculty of Medicine, University of Tokyo; TIMM strains, Research Center for Medical Mycology, Teikyo University, Tokyo, Japan; TMI strains, Tottori Mycological Institute, Tottori, Japan; UNEFM strains, Universidade Nacional Experimental Francisco de Miranda, Coro, Falcon, Venezuela; URM strains, Department of Mycology, Federal University of Pernambuco, Recife, Brazil.

^b+, obtained; −, not obtained.

^cType strain.

^dSequenced in this study.

The necessity of prompt and accurate identification of pathogenic fungi is increasing, since such information can directly influence therapy and prognosis. To aid in the rapid identification of Fonsecaea species, we designed a set of genus-specific PCR primers from the sequence of the rDNA ITS region. This set amplified DNA from only the two Fonsecaea species and yielded a 333-bp fragment for all strains of the two species, as shown in Fig. 1. We performed PCR amplification on multiple strains of medically important dematiaceous species, including C. carrionii, P. verrucosa, E. dermatitidis, and E. jeanselmei, and verified that none of these strains produced visible amplicons, nor did other relevant dematiaceous fungal species and medically important yeasts examined (Fig. 2).

FIG. 1.

Agarose gel electrophoresis of PCR products of Fonsecaea species. Lane 1, 100-bp ladder; lanes 2 to 4, F. compacta; lanes 5 to 22, F. pedrosoi.

FIG.2.

Agarose gel electrophoresis of PCR products of Fonsecaea species, relevant dematiaceous fungi, and medically important yeasts. (A) Lane 1, 100-bp ladder; lane 2, F. compacta; lane 3, F. pedrosoi; lane 4, Phialophora verrucosa; lane 5, Phialophora alba; lane 6, Phialophora americana; lane 7, Phialophora atrovirens; lane 8, Phialophora bubakii; lane 9, Phialophora cinerescens; lane 10, Phialophora fastigiata; lane 11, Phialophora heteromorpha; lane 12, Phialophora lagerbergii; lane 13, Phialophora melinii; lane 14, Phialophora oxyspora; lane 15, Phialophora repens; lane 16, Phialophora richardsiae; lane17, Phaeoacremonium parasitica; lane 18, Lecythophora hoffmannii; lane 19, Lecythophora mutabilis. (B) Lane 1, 100-bp ladder; lane 2, F. compacta; lane 3, F. pedrosoi; lane 4, Cladophialophora carrionii; lane 5, Cladophialophora bantiana; lane 6, Cladophialophora devriesii; lane 7, Cladophialophora minourae; lane 8, Cladosporium cladosporioides; lane 9, Cladosporium colocasiae; lane 10, Cladosporium coralloides; lane 11, Cladosporium elatum; lane 12, Cladosporium fulvum; lane 13, Cladosporium herbarum; lane 14, Cladosporium minusculum; lane 15, Cladosporium resinae; lane 16, Cladosporium resinae f. sp. avellaneum; lane 17, Cladosporium sphaerospermum; lane 18, Cladosporium variabile. (C) Lane 1, 100-bp ladder; lane 2, F. compacta; lane 3, F. pedrosoi; lane 4, Exophiala alcalophila; lane 5, E. dermatitidis; lane 6, E. jeanselmei; lane 7, Exophiala moniliae; lane 8, Exophiala spinifera; lane 9, H. werneckii; lane 10, Alternaria alternata; lane 11, Aureobasidium pullulans; lane 12, R. aquaspersa; lane 13, R. atrovirens; lane 14, Candida albicans; lane 15, Candida dubliniensis; lane 16, Candida glabrata; lane 17, Candida parapsilosis; lane 18, Candida tropicalis; lane 19, Cryptococcus neoformans var. neoformans; lane 20, Malassezia furfur; lane 21, Trichosporon asahii var. asahii.

The ITS region, located between the 18S and 26S nuclear rDNA sequences, includes two spacers (ITS1 and ITS2) separated by a 5.8S conserved region. Interspecies sequence differences in the ITS1 and ITS2 regions have been used to detect and identify fungal species (9, 14, 23). The ITS region contains diagnostic sequences that distinguish interspecific-level divergence of organisms (12). From the alignment of the ITS sequence data in our study, two species of Fonsecaea were found to show minimal intraspecies variation, whereas there was high diversity among the dematiaceous species examined. F. compacta has been reported to be a dysplastic variety of F. pedrosoi (5). In addition, 18S and ITS restriction fragment length polymorphism patterns for this species identical to those for F. pedrosoi have been demonstrated by the use of several restriction enzymes (2, 6). However, the two species show distinct morphological characteristics; F. pedrosoi produces slender conidia generated by loosely branched conidiogenous cells, whereas F. compacta produces spherical conidia generated by densely clustered conidiogenous cells (8). This variability in morphology supports the concept that these two species are distinct from each other, and confirming or denying that concept may require looking at several other genetic markers.

Our primer set rapidly discriminated between F. pedrosoi and R. aquaspersa, which had previously been assigned to the genus Acrotheca (7, 22).

In conclusion, we designed Fonsecaea genus-specific primers based on the rDNA ITS region. These, combined with simple PCR, permitted the specific and rapid detection of Fonsecaea species, indicating their potential use in the identification of Fonsecaea species.

Nucleotide sequence accession numbers.

Accession numbers for the ITS regions sequenced in this study are provided in Table 1.