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Environmental source of Candida dubliniensis.

We isolated Candida dubliniensis from a nonhuman source, namely, tick samples from an Irish seabird colony. The species was unambiguously identified by phenotypic and genotypic means. Analysis of the 5.8S rRNA gene showed that the environmental isolates belong to C. dubliniensis genotype 1.


The ever-increasing number of immunosuppressed humans has led to a marked rise in opportunistic infections, particularly those caused by fungi (1). Candida albicans is the yeast species most commonly associated with oropharyngeal and systemic candidiasis in immunocompromised persons. However, the last 2 decades have seen an increase in infections by other Candida species, including C. dubliniensis, which was first recognized as distinct from C. albicans in 1995 in Ireland (2,3). C. dubliniensis has been recovered mainly from the oral cavities of HIV-infected persons (4) but also from lungs, vaginas, blood, and feces; occasionally this organism causes fatal systemic infections (5). Isolates are assigned to 4 genotypes, defined by the sequence of the internal transcribed spacer regions of the rRNA gene (6).

C. dubliniensis is globally distributed. In HIV-infected patients, the oral prevalence is 1.5%-32% (5). In healthy persons not infected with HIV, C. dubliniensis is absent or rare, but 14% of healthy Caucasians had oral C. dubliniensis in a South African study (7). Like C. albicans, C. dubliniensis may be a member of the normal oral microbial flora of humans, and oral candidosis may result from overgrowth of resident strains. In contrast to other Candida species, some of which are associated with birds (8,9), C. dubliniensis has not been found to date in nonhuman environmental sources. This has led to speculation that the species may be restricted to humans, possibly occupying sites deep within the oropharynx or upper respiratory tract (5).

The Study

Fungal strains were obtained from Ixodes uriae ticks (as part of a National Environment Research Council--funded study of a tickborne virus) at a seabird breeding colony on Great Saltee Island, Ireland (52[degrees] 07'N, 6[degrees]36'W). The ticks were taken from cracks in cliffs used by common guillemots (Uria aalge). Tissue cultures of tick homogenates undertaken for virus isolation were occasionally contaminated with yeastlike fungi.

To investigate this, individual adult ticks were homogenized in 1 mL minimum essential medium (MEM). After centrifugation (30 s, 10,000x g), 0.2 mL of supernatant was added to 4 mL MEM, 5% fetal bovine serum, and 100 [micro]g/mL penicillin-streptomycin. Cultures incubated at 37[degrees]C were examined microscopically daily for up to 6 days. Positive cultures were plated twice on Sabouraud dextrose agar (SAB) with chloramphenicol (bioMerieux, Marcy l'Etoile, France) before phenotypic testing. Isolates were identified by using API identification kits (bioMerieux) and by conventional methods (10). Antifungal drug susceptibility was tested according to the Clinical Laboratory Standards Institute guidelines (11). The control strains were C. albicans ATCC 90028, C. glabrata ATCC 90030, C. krusei ATCC 6258, and C. dubliniensis NCPF 3949.

Internal transcribed spacer 1 and 2 regions (ITS1/ ITS2) and the 5.8S rRNA gene were amplified with primers ITS1 and ITS4, described by White et al. (12). Template DNA was prepared by boiling single SAB-grown colonies in 50 [micro]L ultrapure water for 10 min. After centrifugation (5 min 10,000x g), 15 [micro]L supernatant was added to 50 [micro]L PCRs containing 1x reaction buffer, 1 [micro]mol/L ITS primers, 1.5 mmol/L Mg[Cl.sub.2], 400 [micro]mol/L deoxynucleoside triphosphates, and 2.5 U Immolase (BiolineLtd, London, England, UK). Cycling parameters were 7 min at 95[degrees]C, 30 cycles of 1 min at 95[degrees]C, 1 min at 58[degrees]C, 1 min at 72[degrees]C, and a final extension of 5 min at 72[degrees]C. Products were purified (QiaQuick kit, QIAGEN Ltd., West Sussex, England, UK) and sequenced (BigDye kit and ABI 377 sequencer; Applied Biosystems, Foster City, CA, USA) by using the ITS primers. Sequences were assembled by using Lasergene 6, Seqman version II, and aligned by using BioEdit software (13).

Fungal isolation was undertaken on 2 separate days with samples from 2 distinct sites on the island (Table 1). On both days, Happy Hole West (HHW) ticks were processed immediately after Labour in Vain (LIV) ticks in the same class II microbiologic cabinet. No fungi were detected in HHW ticks, whereas 16.7%-27.6% of ticks sampled from 2 locations within LIV gave positive cultures (Table 1). Twenty-two isolates were obtained (Table 1); SL370-429 were from LIV-1 and SL495-531 from LIV-2 (SL = Saltee).

On SAB the colonies from positive cultures were a creamy white color with a glabrous appearance similar to C. albicans. SL375 had a mixed phenotype (large and small colonies, designated SL375 1 and SL375-2). Like C. albicans, all SL isolates were germ-tube positive and produced chlamydospores at 37[degrees]C on Corn Meal Tween 80 agar (Oxoid Ltd, Basingstoke, England, UK) and Czapek Dox (1%) Tween 80 agar (Oxoid) (Figure 1A). None of the SL isolates grew at 43[degrees]C on SAB (Figure 1B), which suggested that they might be C. dubliniensis (14). This was confirmed by carbohydrate assimilation tests (Table 2) and by sequencing the 5.8S rRNA gene (Figure 2). With the API 20C AUX kit, all SL isolates yielded the same profile at 48 h, interpreted as 99.9% C. dubliniensis (Table 2). Eleven isolates from LIV-1 and the C. dubliniensis (NCPF 3949) reference strain were also tested with API 32C. All had an identical profile (7143100015), interpreted as 81.9% C. dubliniensis and 16.9% C. albicans.


ITS sequences of isolates SL375, SL397, SL407, SL410, SL411, SL417, and SL422 were identical to that of C. dubliniensis CD33 genotype 1 (Figure 2). Nevertheless, phenotypic variation among the SL isolate was evident. In addition to variation in trehalose assimilation rates, 3-4 distinct types were apparent in the germ tube test. Three independent inoculations ([10.sup.5] CFU/mL, mid log growth phase) of each isolate gave consistent morphologic differences. One group produced very long germ tubes (SL375-1, SL422, SL417 SL387, SL397, and SL413); another, medium size (SL411, SL407); a third, mainly long germ tubes with cells that aggregate like C. parapsilosis (SL375-2 and SL410); and the last, elongated cigarlike cells that clamped together with few germ tubes (SL370 and SL371).

To determine whether C. dubliniensis was present on their outer surface LIV-2 and HHW-2 ticks (n = 102) were individually washed in 1 mL MEM before homogenization. C. dubliniensis was cultured from the wash supernatants of 8 of 9 ticks that proved positive after homogenization but not from any of the negative ticks. These findings strongly suggest that the fungus is present in particles adhering to the ticks.

The SL isolates (n = 11) were extremely sensitive to antifungals. MIC90s ([micro]g/mL) were as follows: amphotericin B 0.031 (Sigma-Aldrich, Saint Louis, MO, USA), flucytosine 0.031 (Valeant Pharmaceuticals International, Aliso Viejo, CA, USA), fluconazole 0.125 (Pfizer Inc., Sandwhich, Kent, England, UK), itraconazole 0.007 (Ortho Biotech, Bridgewater, NJ, USA), voriconazole 0.007 (Pfizer), posaeonazole 0.007 (Schering-Plough, Kenilworth, N J, USA), ketoconazole <0.007 (Ortho Biotech), and caspofungin <0.007 (Merck & Co., Inc., Rahway, NJ, USA).


Our serendipitous isolation of C. dubliniensis from the environment ends speculation (5) that the species might be confined to humans. Because ticks were collected from cracks filled with guillemot guano and the fungal isolates were associated with the surface of ticks, the most likely source of the fungus is bird excrement. Thus, like C. albicans (8), C. dubliniensis may inhabit the digestive tract of marine birds. Only C. dubliniensis was isolated from the Great Saltee ticks; however, we have detected other Candida spp. from both soil and tick samples (unpub. data) at other seabird colonies, which suggests that marine as well as terrestrial birds (9) carry a variety of yeast species (15).

The environmental isolates are members of human C. dubliniensis genotype 1, commonly associated with HIV patients (6). Sequencing of the genotype 1 CD36 isolated from an HIV patient is nearly complete ( sequencing/Candida/dubliniensis). Phylogenetic comparison of variable loci from environmental and human genotype 1 isolates could be used to estimate when they last shared a common ancestor and thus how often C. dubliniensis is transmitted from the environment to humans.


We thank Smart Murray for collecting tick and soil samples from Great Saltee Island and Guido Paesen for reviewing the manuscript.

This work was supported by the National Environment Research Council Environmental Genomics and Biodiversity Programs.


(1.) Pfaller MA, Diekema DJ. Rare and emerging opportunistic fungal pathogens: concern for resistance beyond Candida albicans and Aspergillus fumigatus. J Clin Microbiol. 2004;42:4419-31.

(2.) Sullivan DJ, Westerneng TJ, Haynes KA, Bennett DA, Coleman DC. Candida dubliniensis sp. nov.: phenotypic and molecular characterization of a novel species associated with oral candidosis in HIV-infected individuals. Microbiology. 1995; 141:1507-21.

(3.) Sullivan D, Coleman D. Candida dubliniensis: characteristics and identification. J Clin Microbiol. 1998;36:329-34.

(4.) Sullivan D, Haynes K, Bille J, Boerlin P, Rodero L, Lloyd S, et al. Widespread geographic distribution of oral Candida dubliniensis strains in human immunodeficiency virus-infected individuals. J Clin Microbiol. 1997;35:960-4.

(5.) Sullivan DJ, Moran GP, Coleman DC. Candida dubliniensis: ten years on. FEMS Microbiol Lett. 2005;253:9-17.

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(7.) Blignaut E, Pujol C, Joly S, Soll DR. Racial distribution of Candida dubliniensis among South Africans. J Clin Microbiol. 2003;41: 1838-42.

(8.) Buck JD. Isolation of Candida albicans and halophilic Vibrio spp. from aquatic birds in Connecticut and Florida. Appl Environ Microbiol. 1990;56:826-8.

(9.) Cafarchia C, Camarda A, Romito D, Campolo M, Quaglia NC, Tullio D, et al. Occurrence of yeast in cloacae of migratory birds. Mycopathologia. 2006; 161:229-34.

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(11.) National Committee for Clinical Laboratory Standards. Reference method for broth dilution antifungal susceptibility testing of yeast, approved standard M27-A. Wayne (PA): The Committee, 1997.

(12.) White TJ, Bruns TD, Lee SB, Taylor JW. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis MA, Gelfand DH, Sininsky JJ, White TJ, editors. PCR protocols: a guide to methods and applications. San Diego (CA): Academic Press Inc.; 1990. p. 315-22.

(13.) Hall TA. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucl Acids Symp Ser. 1999;41:95-8.

(14.) Gales AC, Pfaller MA, Houston AK, Joly S, Sullivan DJ, Coleman DC, et al. Identification of Candida dubliniensis based on temperature and utilization of xylose and alpha-methyl-D-glucosidase as determined by API 20C AUX and Vitek YBC systems. J Clin Microbiol. 1999;37:3804-8.

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Dr Nunn heads the Tick and Tick-Borne Pathogen Research Group at the Centre for Ecology and Hydrology in Oxford. His research interests focus on the evolution and transmission of tick-borne pathogens and functional characterization of tick immuno-modulators.

Address for correspondence: Miles A. Nunn, National Environment Research Council Centre for Ecology and Hydrology, Mansfield Rd, Oxford OXI 3SR, UK; email:

All material published in Emerging Infectious Diseases is in the public domain and may be used and reprinted without special permission; proper citation, however, is required.

Miles A. Nunn, * Stefanie M. Schafer, * Michael A. Petrou, ([dagger]) and Jillian R.M. Brown *

* National Environment Research Council Centre for Ecology and Hydrology, Oxford, United Kingdom; and ([dagger]) Hammersmith Hospital National Health Service Trust, London, United Kingdom
Table 1. Male and female ticks positive for fungi in culture *

 No. positive/no. examined
Site Male Female Both sexes

Happy Hole West (HHW)-1 0/26 0/25 0/51
HHW-2 0/17 0/23 0/40
Labour in Vain (LIV)-1 5/30 8/29 13/59 ([dagger])
LIV-2 5/23 4/20 9/43 ([double dagger])

* Adult Ixodes uriae ticks were collected from within 2 guillemot-
breeding colonies on Great Saltee on August 25, 2004. The ticks were
stored frozen and processed by M.A.N on November 11, 2005. (HHW-1
and LIV-1) and 20. 07.06 (HHW-2 and LIV-2).

([dagger]) Isolates SL370, SL371, SL375, SL387, SL397, SL407, SL410,
SL411, SL413, SL414, SL417, SL422, and SL429.

([double dagger]) Isolates SL495, SL497, SL500, SL501, SL509, SL510,
SL522, SL529, and SL531.

Table 2. Substrate assimilation by Great Saltee fungi and
Candida albicans

 API 20C AUX assimilation
 profile code *

 C. albicans
Substrate SL407 (ATCC90028)

 L-arabinose - -
 D-xylose - +
 D-glucose + +
 D-galactose + +
 [alpha]-methyl-D-glucoside - +
 D-cellobiose - -
 D-lactose - -
 D-maltose + +
 D-saccharose + +
 D-trehalose ([dagger]) - +
 D-melezitose - -
 D-raffinose - -
 Glycerol + -
 Adonitol + +
 Xylitol + +
 Inositol - -
 D-sorbitol + +
Organic acids
 2-keto-gluconate + +
Amino acids
 N-acetylglucosamine + +
Identification C. dubliniensis C. albicans
API 20C AUX profile code 6172134 2566174
Predictive value 99.9%, excellent 99.2%,
 very good

* Results for 48 hours are shown. All 22 Great Saltee isolates
gave similar profiles.

([dagger]) By 72 hours all Saltee isolates showed some assimilation
of trehalose; the degree of assimilation varied between isolates.
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Author:Brown, Jillian R.M.
Publication:Emerging Infectious Diseases
Date:May 1, 2007
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