Abstract
Candida species bloodstream isolates were collected from institutions participating in an active, population-based surveillance for candidemia. Species identifications were performed locally and then confirmed at the Centers for Disease Control and Prevention (CDC) by phenotype-based methods. Discrepancies in species identification between the referring institution and the CDC were noted for 43 of 935 isolates (4.6%). A DNA probe-based species identification system (PCR-enzyme immunoassay [EIA]) was then used to resolve these discrepancies. The PCR-EIA result was identical to the CDC phenotypic identification method for 98% of the isolates tested. The most frequently misidentified species was Candida glabrata (37% of all discrepant identifications). Such misidentifications could lead to the administration of inappropriate therapy given the propensity of C. glabrata to develop resistance to azole antifungal drugs.
Candida species are the fourth most common cause of health care-associated bloodstream infections and are increasingly important causes of morbidity in hospitalized patients (5, 10). The emergence of non-Candida albicans species, including those innately or adaptively less susceptible to azole antifungals (14, 15), makes the identification of bloodstream isolates to the species level important for the implementation of appropriate antifungal therapy. Species identification is also important for an understanding of the epidemiology of candidemia, including trends in species distribution and antifungal drug susceptibility patterns.
Candida species have traditionally been identified by a combination of phenotypic tests that assess morphological characteristics and carbohydrate assimilation and fermentation patterns (9, 11). Whereas presumptive identification of C. albicans may be obtained in a few hours, identification of non-C. albicans species may require up to 72 h (8, 9, 17, 18). More recently, fungus-specific PCR primers and Candida species-specific DNA probes, directed to the internal transcribed spacer 2 (ITS2) region of ribosomal DNA (rDNA), have been used to detect PCR amplicons in a colorimetric enzyme immunoassay format (PCR-EIA) (6). This test has been shown to be highly specific, rapid, and easy to perform. Therefore, we used the PCR-EIA to resolve discrepancies in Candida species identifications between referring institutions and the Centers for Disease Control and Prevention (CDC) laboratory as part of an active, population-based surveillance for candidemia.
Collection and identification of bloodstream isolates.
Bloodstream isolates were obtained from institutions in the state of Connecticut and the city and county of Baltimore, Md., from October 1998 to September 2000 (10a). All blood cultures positive for Candida species were identified at the referring institution according to their standard methods. In describing their blood culture identification practices, 23 of 51 responding laboratories (45%) used the germ tube formation test to identify C. albicans. Forty-one respondents (80%) used some type of carbon assimilation/biochemical panel to identify non-C. albicans species, with 20 (39%) using the API 20C system (bioMerieux Vitek, Inc., Hazelwood, Mo.) and 13 (26%) using the Vitek yeast biochemical card (bioMerieux Vitek).
A total of 935 isolates were sent to the Mycotic Diseases Branch, CDC, for confirmation of species identification. At the CDC, the isolates were first subcultured onto Sabouraud dextrose agar (BBL Difco Laboratories, Detroit, Mich.) as well as onto CHROMagar Candida medium (DRG International, Mountainside, N.J.). Isolates were then identified to the species level with the API 20C AUX (bioMerieux Vitek) or RapID Yeast Plus system (Innovative Diagnostics, Norcross, Ga.) and by microscopic morphology on cornmeal-Tween 80 (Dalmau) plates.
Discrepancies in the phenotypic identification between the CDC and the referring institution were resolved by using species-specific DNA probes in an EIA detection format (PCR-EIA) (6). Candida species isolates were grown for 18 h at 35°C in 50-ml Erlenmeyer flasks containing 10 ml of YPD broth (1% yeast extract, 2% Bacto Peptone, 2% dextrose; BBL Difco Laboratories). DNA was isolated from these cultures with the PUREGENE DNA Purification kit for yeast and gram-positive bacteria (Gentra Systems, Inc., Minneapolis, Minn.) according to the manufacturer's directions. Universal fungus-specific primers ITS3 (5′ GCA TCG ATG AAG AAC GCA GC 3′) and ITS4 (5′ TCC TCC GCT TAT TGA TAT GC 3′) (19) were then used to amplify by PCR a portion of the 5.8S rDNA region, the entire ITS2 rDNA region, and a portion of the 28S rDNA region using a Perkin-Elmer (Emeryville, Calif.) model 9700 thermal cycler and Taq DNA polymerase (Roche Molecular Biochemicals, Inc., Indianapolis, Ind.). All other PCR reagents and thermal cycling conditions used were as previously described (6). Amplicons were captured onto a streptavidin-coated microtitration plate (Roche) with a biotinylated, all-Candida species DNA probe (5′ CAT GCC TGT TTG AGC GTC [GA]TT 3′) and were detected with digoxigenin-labeled species-specific DNA probes (C. albicans: 5′ AT TGC TTG CGG CGG TAA CGT CC 3′; C. glabrata: 5′ TT TAC CAA CTC GGT GTT GAT CT 3′; C. krusei: 5′ GG CCC GAG CGA ACT AGA CTT TT 3′; C. lusitaniae: 5′ CT CCG AAA TAT CAA CCG CGC TG 3′; C. parapsilosis: 5′ AC AAA CTC CAA AAC TTC TTC CA 3′; C. tropicalis: 5′ AA CGC TTA TTT TGC TAG TGG CC 3′) and horseradish peroxidase-labeled antidigoxigenin antibodies in a colorimetric EIA format (6). Oligonucleotide primers and probes were synthesized and labeled as described previously (12).
Discrepancies in species identification between the CDC's phenotypic methods and the PCR-EIA were resolved by rDNA sequencing on a Perkin-Elmer ABI Prism 310 automated capillary DNA sequencer as previously described (7). Briefly, universal fungus-specific primers ITS1 (5′ TCC GTA GGT GAA CCT GCG G 3′) (19) and ITS4 were used to amplify by PCR a portion of the 18S rDNA region; the entire ITS1, 5.8S, and ITS2 rDNA regions; and a portion of the 28S rDNA region. Amplicons were purified with the QIAquick PCR purification kit (Qiagen, Valencia, Calif.) and sequenced on both strands with primers ITS1 or ITS4 and the Big Dye Terminator Cycle Sequencing Ready Reaction kit (Applied Biosystems, Foster City, Calif.) (7). GenBank searches and comparative sequence analyses were assisted by using BLAST search tools (1) and GeneTool, version 1.0, software (BioTools, Inc., Edmonton, Alberta, Canada), respectively.
Distribution and resolution of discrepancies in Candida species identification.
Of the 935 isolates received at the CDC, there were discrepancies in species identification between the referring institution and the CDC laboratory for 43 (4.6%). In all but one case, the CDC identification based on biochemical and morphological criteria was validated by the PCR-EIA (Table 1). In the remaining case, an isolate reported as C. albicans by the referring institution and as C. parapsilosis or C. lusitaniae by the CDC was ultimately identified as C. lusitaniae by probe testing and DNA sequence analysis (GenBank accession number AY383555). All isolates identified at the CDC as C. albicans by phenotypic methods were subsequently screened by molecular identification methods (2, 10a ) to differentiate isolates of C. dubliniensis from those of C. albicans. Nine cases of C. dubliniensis candidemia were identified. All C. dubliniensis isolates were reported to be C. albicans by the referring institutions, but these identifications were not considered to be discrepant as C. dubliniensis is not routinely differentiated from C. albicans by most clinical laboratories.
TABLE 1.
Phenotypic identifications of Candida species by the referring hospital and by CDC compared to those by the PCR-EIA
| Hospital IDb | CDC ID | PCR-EIA ID | CHROMagar result | API 20C or RapID profile resulta (% IDc) | Morphologyd |
|---|---|---|---|---|---|
| C. albicans | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. albicans | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. albicans | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. albicans | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. albicans | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. albicans | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. albicans | C. glabrata | C. glabrata | Pink | Implicit CG (99.0) | BSP w/o PSH |
| C. krusei | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. krusei | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. krusei | C. glabrata | C. glabrata | Purple | Very good CG (99.4) | BSP w/o PSH |
| C. parapsilosis | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. tropicalis | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. tropicalis | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. tropicalis | C. glabrata | C. glabrata | Pink | Very good CG (99.4) | BSP w/o PSH |
| C. tropicalis | C. glabrata | C. glabrata | Pink | Satisfactory CG (99.2) | BSP w/o PSH |
| C. tropicalis | C. glabrata | C. glabrata | Pink | Low discrim. PW (70.7), CG (29.3) | BSP w/o PSH |
| C. albicans | C. parapsilosis | C. parapsilosis | Pale pink | Excellent CP (99.9) | BSP and PSH |
| C. albicans | C. parapsilosis | C. parapsilosis | Pale pink | Good CP (99.1) | BSP and PSH |
| C. albicans | C. parapsilosis | C. parapsilosis | Pale pink | Acceptable CP (97.4) | BSP and PSH |
| C. albicans | C. parapsilosis | C. parapsilosis | Pale pink | Satisfactory CP (95.7) | BSP and PSH |
| C. albicans | C. parapsilosis | C. parapsilosis | Pale pink | Low discrim. CP (83.3), CN (13.3) | BSP and PSH |
| C. albicans | C. parapsilosis | C. parapsilosis | Pale pink | Acceptable genus CT (57.1), CA (27.6), CP (12.2) | BSP and PSH |
| C. glabrata | C. parapsilosis | C. parapsilosis | Pale pink | Good CP (97.8) | BSP and PSH |
| C. glabrata | C. parapsilosis | C. parapsilosis | Pale pink | Acceptable CP (90.5) | BSP and PSH |
| C. glabrata | C. parapsilosis | C. parapsilosis | Pale pink | Low discrim. CN (50.2), CP (47.5) | BSP and PSH |
| C. guilliermondii | C. parapsilosis | C. parapsilosis | Pale pink | Good CP (98.0) | BSP and PSH |
| C. lusitaniae | C. parapsilosis | C. parapsilosis | Pale pink | Very good CP (99.9) | BSP and PSH |
| C. lusitaniae | C. parapsilosis | C. parapsilosis | Pale pink | Low discrim. CP (71.9), CN (18.9), CT (4.9) | BSP and PSH |
| C. tropicalis | C. parapsilosis | C. parapsilosis | Pale pink | Very good CP (99.0) | BSP and PSH |
| C. tropicalis | C. parapsilosis | C. parapsilosis | Pale pink | Good CP (95.9) | BSP and PSH |
| C. tropicalis | C. parapsilosis | C. parapsilosis | Pale pink | Low discrim. CN (50.2), CP (47.5) | BSP and PSH |
| C. albicans | C. tropicalis | C. tropicalis | Blue | Low discrim. CT (71.1), CN (18.5), CL (7.6) | BSP and PSH |
| C. albicans | C. tropicalis | C. tropicalis | Blue | Low discrim. CT (71.1), CN (18.5), CL (7.6) | BSP and PSH |
| C. albicans | C. tropicalis | C. tropicalis | Blue | Low discrim. CT (71.1), CN (18.5), CL (7.6) | BSP and PSH |
| C. albicans | C. tropicalis | C. tropicalis | Blue | Low discrim. CT (71.1), CN (18.5), CL (7.6) | BSP and PSH |
| C. glabrata | C. tropicalis | C. tropicalis | Blue | Adequate CT (97.0) | BSP and PSH |
| C. glabrata | C. tropicalis | C. tropicalis | Blue | Good CT (95.9) | BSP and PSH |
| C. glabrata | C. tropicalis | C. tropicalis | Blue | Low discrim. CT (71.1), CN (18.5), CL (7.6) | BSP and PSH |
| C. glabrata | C. albicans | C. albicans | Green | Low discrim. CA (58.7), CT (26.0), CN (14.4) | CHL and PSH |
| C. tropicalis | C. albicans | C. albicans | Green | Good genus CT (48.4), CA (44.4) | CHL and PSH |
| C. tropicalis | C. albicans | C. albicans | Green | Unacceptable profile | CHL and PSH |
| C. albicans | C. lusitaniae | C. lusitaniae | Tan | Good genus CT (60.7), CL (29.6) | BSP and PSH |
| C. albicans | C. parapsilosis or C. lusitaniae | C. lusitaniae | Pale pink | Very good genus CP (53.5), CL (38.8), CT (7.1) | BSP and PSH |
CA, C. albicans; CG, C. glabrata; CL, C. lusitaniae; CP, C. parapsilosis; CT, C. tropicalis; CN, Cryptococcus neoformans; PW, Prototheca wickerhamii. “Implicit CG” and “satisfactory CG” are RapID Yeast Plus results; all other results are from the API 20C AUX system. Low discrim., presumptive identification.
ID, identification.
%ID, percent identification likelihood.
BSP, blastospores; PSH, pseudohyphae; CHL, chlamydospores; w/o, without.
The most frequent misidentifications by the referring institutions were of C. glabrata (16 of 43 isolates or 37% of all discrepant identifications), followed by C. parapsilosis (15 of 43, 35%), C. tropicalis (7 of 43, 16%), C. albicans (3 of 43, 7%), and C. lusitaniae (2 of 43, 5%) (Table 1). These misidentifications represent 12 (15 of 123), 7 (16 of 226), 6 (7 of 118), and 0.7% (3 of 423) of all C. parapsilosis, C. glabrata, C. tropicalis, and C. albicans isolates, respectively. The most common misidentifications by the referring institutions were C. albicans for C. glabrata (7 of 43 isolates or 16% of all misidentifications), C. albicans for C. parapsilosis (6 of 43, 14%), C. tropicalis for C. glabrata (5 of 43, 12%), and C. albicans for C. tropicalis (4 of 43, 9%).
Of all isolates reidentified at the CDC that could be associated with a given type of institution, 618 (66%) were from nonacademic institutions and 316 (34%) were from academic institutions. Comparison of the misidentification rate between academic (university- or medical school-associated) and nonacademic institutions showed that, for the 41 of 43 isolates that could be associated with a particular category of institution, 34 (83%) of the misidentifications were from nonacademic institutions whereas 7 (17%) were from academic institutions. As a percentage of the total number of isolates received from academic versus nonacademic institutions, the overall misidentification rate for academic institutions was roughly one-half that for the nonacademic institutions (i.e., 2.2 versus 5.5%, respectively). Misidentifications were not associated with any one particular institution.
The vast majority (15 of 16, 94%) of misidentified isolates that were reidentified as C. glabrata in the CDC laboratory had typical biochemical profiles and morphologies on Dalmau plates and gave the expected colony color and appearance on CHROMagar Candida medium (Table 1). In contrast, 53% of C. parapsilosis isolates and 71% of C. tropicalis isolates showed profiles interpreted as “acceptable” to “low discrimination” by the API 20C AUX system; these isolates were differentiated from the alternative species choices listed in the API 20C AUX profile index by microscopic morphology and colony color on CHROMagar Candida medium (Table 1). C. albicans isolates could also be identified by their distinctive colony color on CHROMagar Candida medium and by their capacity to form chlamydospores (Table 1).
Specificity of DNA probes to identify Candida species.
The PCR-EIA generated results that were highly specific, the DNA did not cross-react with DNA from other Candida species tested, and the results were easy to interpret (Table 2). Mean positive EIA values ± standard errors (SE) ranged from 0.95 ± 0.10 for DNA from C. tropicalis to 0.38 ± 0.02 for DNA from C. lusitaniae. Inherent differences in absolute EIA values obtained for each of the probes may reflect differences in their G+C compositions and, as a result, their rate of denaturation and annealing during thermal cycling or their rate of hybridization during probe attachment to the PCR product. Nonetheless, the EIA values reported here were very similar to those reported previously for the identification of these same species (6) and were approximately 200 times above background values after subtraction of the water blank (Table 2). Testing of heterologous-species DNA gave no significant background reactivity (mean A650 ± SE = 0.002 ± 0.0001), making discrimination of a positive from a negative result unequivocal (Table 2). All Candida species DNAs were also tested with a probe specific for C. krusei DNA, and no reactivity with heterologous DNA was observed (mean A650 ± SE for the C. krusei-specific probe versus C. krusei DNA and versus all other Candida species DNAs, 0.49 ± 0.05 and 0.0014 ± 0.0004, respectively; n = 72).
TABLE 2.
Specificity of DNA probes for species identification by PCR-EIA
| DNA target species (no. of isolates tested) | Mean A650 ± SEa after reaction with probe for:
|
||||
|---|---|---|---|---|---|
| C. glabrata | C. parapsilosis | C. tropicalis | C. albicans | C. lusitaniae | |
| C. glabrata (16) | 0.91 ± 0.07 | 0 | 0 | 0 | 0 |
| C. parapsilosis (15) | 0 | 0.79 ± 0.05 | 0 | 0 | 0 |
| C. tropicalis (7) | 0 | 0 | 0.95 ± 0.10 | 0 | 0 |
| C. albicans (3) | 0 | 0 | 0 | 0.40 ± 0.05 | 0 |
| C. lusitaniae (2) | 0 | 0 | 0 | 0 | 0.38 ± 0.02b |
Mean A650 ± SE was calculated from spectrophotometric readings after target DNA was reacted with the DNA probes listed above. All samples were run in duplicate, and reagent blanks were run on each plate for each probe. Reagent blank values have been subtracted from test sample values above (mean reagent blank A650 = 0.038 ± 0.001; n = 72). Mean A650 ± SE for all control samples after subtraction of the reagent blanks for all probes was 0.002 ± 0.0001 (n = 226) and is represented in this table as 0 for ease of presentation.
Includes one C. lusitaniae isolate identified as C. albicans by the referring institution and as C. parapsilosis or C. lusitaniae by CDC phenotypic methods.
Conclusions.
Several population-based and sentinel surveillance studies have noted an increase in the proportion of Candida bloodstream infections caused by species other than C. albicans, and, in particular, an increase in the frequency of candidemia due to C. glabrata (reviewed in reference 15). Given the known propensity of C. glabrata to develop resistance to azole antifungals, the fact that this species was most frequently misidentified in this study is disturbing. Because each referring institution used its own method(s) for species identification, the reasons for the high rate of C. glabrata misidentification are not clear. However, for the 28% of non-C. glabrata isolates that were misidentified as C. albicans by the referring institution, two factors may provide some insight. First, in many referring institutions (45% of those surveyed), the germ tube test was performed as a primary screen for the identification of C. albicans. Second, a disproportionately greater number of misidentifications were received from nonuniversity laboratories than from university or medical school laboratories. These data suggest that the nonuniversity institutions in our study may have employed fewer specialists in mycology, who in turn had less experience in interpreting the germ tube test. This hypothesis is supported by the work of others (4) who found that, when the germ tube test was performed on a series of isolates tested in a blinded fashion by technicians who were not specialists in mycology, germ tube test specificity declined. Misinterpretation of results, particularly of pseudohyphal production, accounted for this drop in specificity (4). This might account for those isolates originally identified by the referring institution as C. albicans but ultimately identified as C. lusitaniae, C. parapsilosis, or C. tropicalis by the CDC. In addition, the CDC laboratory routinely employs CHROMagar Candida medium (16) as an adjunct to biochemical and morphological tests whereas none of the institutions polled in the surveillance area reported the use of this medium. The distinctive color of each species on this medium may be helpful in cases where the biochemical results are equivocal and the expertise for distinguishing various species based on morphology on cornmeal-Tween agar is lacking.
Most clinical laboratories do not differentiate isolates of C. albicans from those of C. dubliniensis. Therefore, the nine cases of C. dubliniensis candidemia identified at the CDC by molecular identification methods and reported as C. albicans by the referring institutions were not considered to be discrepant identifications. Nonetheless, it has been demonstrated previously that the PCR-EIA can unequivocally differentiate isolates of C. dubliniensis from those of C. albicans (6, 7).
Unlike current phenotypic identification methods, which may require a series of tests to confirm the identity of a given Candida species (8, 9, 17), the PCR-EIA is a single test that can be used to identify the majority of medically important Candida species (6). In contrast to phenotype-based identification methods, the PCR-EIA can be performed in a single day and the results are very easy to interpret. Use of a commercial kit for the isolation of Candida species DNA (6) makes this test a fast and reliable method for Candida species identification. Although other PCR-based methods for the identification of Candida species have been described (reviewed in references 3 and 13), these methods did not use a combination of (i) an EIA detection format; (ii) universal fungal, multicopy rRNA gene targets to increase test sensitivity; (iii) a commercial kit for rapid and simple sample preparation; and (iv) probes that could detect more than one or a few Candida species. This study demonstrates the usefulness of the PCR-EIA for the resolution of discrepant phenotype-based species identifications. Conversion of the DNA probes described in this study into either a real-time PCR format or an automated microarray format would reduce postamplification manipulation steps and further reduce the time required for accurate species identification.
Acknowledgments
We thank Hans Peter Hinrikson for assistance with the C. lusitaniae DNA sequence analysis.
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