Candida auris: An Emerging Yeast Pathogen Posing Distinct Challenges for Laboratory Diagnostics, Treatment, and Infection Prevention Open Access

In 2009, Satoh and colleagues⁷  isolated a novel ascomycetous yeast in discharge from the ear canal of a hospitalized patient in Tokyo, Japan. Sequencing results of the internal transcribed spacer (ITS) and D1/D2 regions of the species ribosomal DNA showed only an 87.5% and 85.7% similarity to Candida haemulonii. In addition, sequencing of the D1/D2 regions revealed only an 81.4% and 83.0% similarity to Candida pseudohaemulonii. Subsequent analysis of isolate sequencing data, 42°C growth, and unique patterns of carbon source assimilation as compared with other closely related yeast species supported establishment of a new species type designated as C auris, the Latin term auris referring to the ear. Candida auris is considered to have recently emerged as a pathogen based on retrospective epidemiologic analyses of 15 271 isolates collected from 2004 through 2015. Interrogation of an international surveillance culture collection (SENTRY) indicated a total of 4 isolates confirmed as C auris collected in the years 2009, 2013, 2014, and 2015. A single isolate in this group had been misidentified as C haemulonii during initial analysis; the other 3 isolates were identified only as Candida species. The absence of C auris isolates in the SENTRY collection before 2009 further supported evidence of relatively recent emergence of this new Candida species.⁸  Less than a decade after the initial identification, C auris was identified on 5 continents.^8–12 

In a study of global epidemiology of C auris, whole-genome sequencing allowed for geographic grouping of C auris into 4 distinct clades: East Asia, South Asia, Africa, and South America. Although isolates from different clades varied by tens of thousands of single-nucleotide polymorphisms, variation within a clade was minimal (0–70 single-nucleotide polymorphisms), suggesting independent clade evolution.⁸  Since the identification of these 4 clades, patients with C auris have been reported from more than 20 countries. The results of whole-genome sequencing of more than 1150 isolates from 15 countries performed at the Centers for Disease Control and Prevention (CDC) in the United States show that all of the isolates discovered worldwide originate from the previously described 4 clades (CDC, unpublished data, 2018). This indicates that after the simultaneous global emergence, C auris has been spreading from a few major epicenters, followed by regional clonal expansion of highly related isolates.^8,13,14 

Macroscopic characteristics of C auris may aid in identification but will not provide sufficient information for definitive differentiation of C auris from other species of Candida. On standard fungal media, such as Sabouraud dextrose agar, C auris appears as white to cream-colored, smooth, butyrous colonies (Figure 1). Growth of C auris can occur at 30°C; however, optimal development is noted at 37°C to 40°C. Growth of C auris at 42°C is slow and weak, and temperature cannot be used to differentiate this species from other closely related yeasts such as C haemulonii. Growth of C auris is inhibited in the presence of cycloheximide. Candida auris will ferment glucose, sucrose, and trehalose and will assimilate carbon sources such as glucose, sucrose, maltose, d-trehalose, d-raffinose, d-melezitose, d-mannitol, sorbitol, citrate, inulin, starch, ribitol, and galactitol.¹⁵ 

Microscopically, C auris has 2 cellular morphologic features; some strains form aggregates of cells, whereas others have single cells.¹⁵  Budding yeast cells are approximately 2.0 to 3.0 × 2.5 to 5.0 μm, with an ovoid to elongate appearance. Similar to Candida glabrata, pseudohyphae are typically not formed on cornmeal agar or rice–Tween 80 agar, although rudimentary pseudohyphae and true hyphae have been reported under certain growth conditions (Figure 2).^7,16  Although cellular morphology is traditionally used for the identification of many yeast species, C auris cannot be identified by morphology alone, and differentiation based only on phenotypic features should be discouraged.

Although useful for other species of Candida, chromogenic agar as a stand-alone method may not provide useful information for definitive identification of C auris. On chromogenic agar (eg, CHROMagar), C auris colonies appear beige or pink to red and may look similar to C glabrata (Figure 3).^17,18  Difficulties in isolation and identification of C auris may be addressed in part by supplementation of CHROMagar media with Pal (sunflower seed extract) agar. This method has been reported to be useful and rapid in combination with temperature tolerance for differentiation of C auris from the C haemulonii complex (C haemulonii, Candida duobushaemulonii, and C haemulonii var vulnera).¹⁹ 

Accurately identifying C auris using commercially available phenotypic methods is problematic, which has serious ramifications for treatment and infection control. Definitive identification of C auris and phenotypic differentiation from other closely related yeasts such as C haemulonii, C pseudohaemulonii, and C duobushaemulonii is challenging because of overlapping biochemical profiles and limitations in identification databases. Both nonautomated and automated biochemical methods commonly used in clinical microbiology laboratories have well-described performance issues that lead to inaccurate identification of C auris, primarily from lack of organism-specific biochemical profiles in system databases. This may result in a low confidence result, identification to Candida species only, or outright misidentification. The API 20C AUX clinical yeast system (bioMérieux Inc, Durham, North Carolina) misidentifies C auris as Candida sake, Candida famata, and Saccharomyces cerevisiae. In addition, this system may also misidentify C auris as Rhodotorula glutinis in isolates with a negative urease reaction that lack the characteristic pink-pigmented colonies of R glutinis. Automated identification systems also misidentify C auris as Candida catenulata and C haemulonii by BD Phoenix (Becton Dickinson and Company, Franklin Lakes, New Jersey); as C duobushaemulonii, C haemulonii, and C famata by Vitek 2 (bioMérieux); and as C catenulata, C famata, Candida guilliermondii, Candida lusitaniae, and Candida parapsilosis by MicroScan WalkAway Plus (Beckman Coulter, Inc, Brea, California).^15,20  Informational tables summarizing limitations of US Food and Drug Administration (FDA)–approved contemporary commercial identification systems may be found at the CDC Web site²¹  and in Mizusawa et al.²⁰  Laboratories with these organism identification/instrument combinations or that have a genus-only result of Candida should continue testing to determine if the isolate in question is C auris. Useful identification algorithms based on common identifications generated by commercial systems are available at the CDC Web site.²¹  Users of biochemical methods to identify yeasts should maintain awareness for updates in system databases that may allow for more accurate identification of C auris. Even if an updated database is used, identification of species commonly misidentified as C auris warrants additional investigation in order to verify results.

Proteomic methods such as MALDI-TOF MS can identify C auris rapidly and accurately with substantially better performance than phenotypic systems. The 2 MALDI-TOF platforms used most widely in clinical microbiology laboratories are the Bruker BioTyper (Bruker Daltonics, Billerica, Massachusetts) and the Vitek MS (bioMérieux). For MALDI-TOF MS, protein is first extracted from an isolate of interest in pure culture and applied to a target plate or slide. Then, MALDI-TOF MS generates a spectral profile of the protein and compares the profile with a reference database, identifying the isolate to the genus and species level. The method used to extract protein from the organism, coupled with the identification database used, affects the quality of identification. With respect to C auris, direct-on-target extraction methods have been shown to be successful with Vitek MS but may lead to a “no identification” result with the Bruker BioTyper. A tube-extraction method can be used with both systems but is slightly more time-consuming and labor-intensive.²⁰  A modified direct-on-target method for protein extraction has been described that generated acceptable spectral scores for identification of C auris without the need for full-tube extraction.²² 

As with biochemical methods, the quality of the reference database directly affects the identification accuracy of MALDI-TOF MS. Generally, both the Bruker and Vitek MS systems have the capability to accurately identify C auris using the research-use–only (RUO) spectral databases supplied with each instrument. Kathuria et al²³  reported excellent identification of C auris and differentiation from C famata, C haemulonii, C haemulonii var vulnera, and C duobushaemulonii using MALDI-TOF MS. If an RUO library is used, users should confirm that the corresponding database version specifically contains reference spectra for C auris. In some cases, updated databases or software patches may be required to obtain high-confidence spectral scores.^15,17  At this time, only the Bruker system contains C auris in the FDA-approved spectral database. Therefore, if a laboratory wants to use Vitek MS to identify C auris isolates, an RUO database must be used that includes reference spectra for C auris.¹⁷  It is also possible for a laboratory, using a manufacturer's protocols, to create its own custom database library.²⁴ 

In addition to these database options, the CDC, in collaboration with Bruker, offers an online tool to provide accurate classification of C auris to the species level (https://www.cdc.gov/microbenet/index.html).²⁵  The tool contains expert curated information and access to CDC spectral libraries. Users may upload laboratory-generated MALDI-TOF MS information to a CDC database containing proteomic data for organism identification by in silico comparison. If a laboratory elects to identify C auris using MALDI-TOF MS, a review of the contents of instrument databases (RUO or FDA-approved) for C auris is strongly encouraged in order to achieve optimal performance and accuracy with these systems. In addition, well-characterized isolates of C auris are available to qualified laboratories as part of the CDC antibiotic resistance organism bank to assist with verification or validation of FDA-approved or RUO organism identification databases (https://www.cdc.gov/drugresistance/resistance-bank/index.html).

Clearly, identification of C auris based solely on phenotypic, biochemical, or proteomic characteristics is a challenge because of the inherent characteristics of the organism and limitations of commercial identification systems. Candida auris can also be identified through molecular methods, such as conventional and real-time polymerase chain reaction (PCR) assays, which are rapid to perform and highly accurate. Kordalewska et al²⁶  developed an initial assay that specifically and accurately detected C auris. This group subsequently used a different set of PCR primers in an additional assay that enabled differentiation of C auris from other closely related species, such as C lusitaniae, C haemulonii, and C duobushaemulonii, by using SYBR Green (ThermoFisher Scientific, Waltham, Massachusetts) detection of amplicon and melting-point analysis. In an analysis of a 140-member proficiency panel, both PCR assays yielded an accuracy of 100% and complete concordance with sequencing results for ITS regions.²⁶ 

Definitive identification of C auris to species level may also be achieved through molecular sequencing of the 18S ITS regions or the D1/D2 regions of fungal ribosomal DNA.^17,23  However, the financial investment and technical requirements needed for sequencing may preclude the routine use of these methods for identifying organisms in most clinical laboratories.²⁵  A diagnostic modality using magnetic resonance for detection of specific Candida species directly from blood specimens is available, and an assay specific for identification of C auris using this technology is currently being developed.²⁷  In addition, isolates may be submitted to regional Antibiotic Resistance Laboratory Network facilities (https://www.cdc.gov/drugresistance/laboratories/AR-lab-network-testing-details.html) or to many commercial reference laboratories for identification of C auris.

Traditional methods of strain typing, such as amplified fragment-length polymorphism or multilocus sequence typing, have been used in an attempt to type C auris.¹⁰  However, the limited variability within C auris clades may impede the utility of strain typing for epidemiologic studies of local transmission or infection control within health care facilities.²⁸  Whole-genome sequencing remains state of the art for determination of clonality of C auris isolates.

The clinical significance of C auris is considerable because of its rapid worldwide emergence, ability to cause invasive infections associated with high mortality, propensity to show resistance to multiple classes of antifungal agents, and ability to spread in health care settings and cause outbreaks. Patients most at risk for C auris infections have had extensive exposure to health care (eg, multiple admissions to long-term and acute care facilities) in the months preceding C auris diagnosis; indwelling devices such as tracheostomy tubes, percutaneous gastrostomy tubes, or central vascular access ports/catheters; or recent exposure to broad-spectrum antibiotics and antifungal agents. Nosocomial transmission has been reported to occur in both acute and long-term care settings, although long-term care facilities, especially skilled nursing facilities with patients who require ventilators, have generally had the highest prevalence of patients with C auris colonization.^12,14  The ability of C auris to colonize in patients long-term and persist on both moist and dry surfaces for weeks is an additional factor considered highly contributory to transmission and recurrence within health care settings.^15,29,30  Fungal virulence factors such as proteinases, phospholipases, and hemolysins have been shown to express diverse levels of activity in vitro, suggesting inherent variability of virulence among different strains of C auris.¹⁵  However, mouse and invertebrate experimental models of infection suggest the overall virulence of C auris may be comparable to that of C albicans.^16,31–33 

Candida auris has been implicated in a variety of superficial and invasive infections from multiple body sites. After the first reported case (2009) of C auris in a Japanese patient with otitis media,⁷  another 15 cases were reported from 5 hospitals in South Korea; all the patients also had chronic otitis media.³⁴  Using the D1/D2 domain and ITS sequence analysis, all 15 isolates were determined to be a novel species (ultimately recognized as C auris), and 10 showed notable resistance to fluconazole (minimum inhibitory concentration [MIC] ≥32 μg/mL). Interestingly, all of these patients had a history of previous antibiotic treatment and manipulation of the ear canal.

The bloodstream is by far the most common site of isolation of C auris from clinical cultures. In a meta-analysis of reports from 2012 to 2017 describing 742 confirmed C auris isolates from 16 countries, 67% were isolated from blood.¹⁵  The bloodstream was the culture source for 5 of the first 7 patients infected with C auris in the United States (from 2013 to 2017); all of these patients had serious underlying medical conditions.³⁵  Reports from various locations worldwide have noted an association between C auris fungemia with indwelling catheters (venous or urinary), use of broad-spectrum antibiotics, parenteral nutrition, surgery, and extended stay in an intensive care unit, in addition to underlying comorbid conditions, such as diabetes mellitus and malignancy.^9,36–38  Determining overall mortality attributable to C auris fungemia is difficult because of multiple confounding comorbid conditions in many patients.³⁹  However, estimates of overall mortality from C auris fungemia range from 28% to 60%.^8,9 

Although bloodstream infections are most common, C auris has also been isolated from diverse body sites, such as wounds; bone; urine; vagina; and cerebrospinal, peritoneal, pleural, and pericardial fluids.¹⁵  Isolates of C auris have also been found in tissues excised from gangrenous feet and postoperative wounds.⁴⁰  Among US patients, approximately 46% of C auris infections have been identified from sites other than blood.⁶⁰ 

In a survey of cases with confirmed C auris isolated from bone, urine, peritoneal fluid, and cerebrospinal fluid, misidentification of the organism along with increased resistance of C auris to antifungal agents contributed to a 30-day mortality rate of 35.2%.³⁷  A case report of fatal C auris pericarditis in a patient with chronic liver disease who was receiving empiric fluconazole therapy is notable for the concurrent recovery of C auris from urine, bronchoalveolar lavage, and blood.⁴¹  A recent report of donor-derived, transplant-transmitted C auris associated with a bilateral lung transplant suggests reconsideration of the clinical significance of yeast in donor lung surveillance cultures.⁴² 

Regardless of the source of the infection, specimen, or underlying cause of disease, delayed diagnosis or misidentification of C auris will affect selection of appropriate antifungal therapy and impact infection control efforts, and may ultimately affect patient outcome. In the assessment of yeasts isolated from nonsterile sites (eg, skin, vaginal, upper respiratory), many laboratories will report “yeast, not Cryptococcus” with no additional speciation. At this time, there are no guidelines supporting routine assessment of nonsterile site specimens for the presence of C auris. However, in areas where C auris has been reported, both clinicians and laboratorians should be cognizant of this possibility and request additional identification studies, as appropriate.

Emerging evidence suggests that immune system suppression attributable to malignancy or immunosuppressive agents (eg, corticosteroids) is a risk factor for C auris infection and may contribute to dissemination of infection in the compromised host. In a 2013 report of C auris isolated from the bloodstream in 12 patients in India, 11 patients were considered immunosuppressed because of a wide variety of conditions, such as diabetes mellitus, chronic kidney disease, HIV, cancer chemotherapy, or a stay in the intensive care unit.¹⁰  A survey of the first C auris cases in the United States also showed that infection developed in association with serious underlying conditions, such as malignancy and corticosteroid use, that resulted in immunosuppression.³⁵ 

One of the most concerning features of C auris is its reduced susceptibility to azoles, polyenes, and, in some instances, echinocandins. This feature severely limits the antifungal treatment options for infected patients, especially given the diminished health status of many of these patients. Triazoles, including fluconazole, are an important therapy for Candida infections. Early susceptibility reports for C auris suggested the organism might be intrinsically resistant to fluconazole, as nearly all early isolates evaluated displayed elevated fluconazole MICs.⁸  For instance, a study of 350 Indian isolates showed 315 (90%) with fluconazole MICs greater than 16 μg/mL.³⁸  Further study of C auris showed that isolates from other geographic locations had fluconazole MICs of 2 to 8 μg/mL, similar to those for C glabrata, suggesting that azole resistance in C auris is acquired and not intrinsic.²⁵  The azoles prevent Candida cell growth by inhibiting the cell's sterol pathway, specifically the 14-α demethylation of ergosterol precursors. This enzyme is encoded by the Erg11 gene. Ergosterol is an essential component of the fungal cell membrane, and inhibition of ergosterol biosynthesis leads to accumulation of toxic methylated sterols, arresting cell growth. Substitutions to the Erg11p azole target have been described for C auris, leading to an altered protein structure, reduced azole binding affinity, and increased azole MICs. The specific mutations to Erg11p observed to date in C auris are associated with geographic clades, such that isolates from South Africa and Venezuela share F126L alterations, whereas Indian isolates harbor Y132F or K143R substitutions.⁸  Candida auris can exhibit reduced susceptibility to other triazole antifungals, including voriconazole, posaconazole, itraconazole, and isavuconazole, but most isolates have relatively low MIC values.⁴³ 

Variable susceptibility is exhibited by C auris isolates to the polyene amphotericin B. Again, susceptibility to this antifungal agent is defined geographically and may be affected when cases are imported from multiple geographic locations.⁴⁴  Most isolates from India have MICs less than 2 μg/mL,⁴³  but up to 64% of isolates in a Colombian study⁴⁵  had amphotericin B MICs greater than 1 μg/mL. The mechanisms that define amphotericin B resistance in C auris are not yet defined and are poorly defined for other Candida species; therefore, readers of case reports must be mindful of selection bias, which tends to present a picture of increased amphotericin B resistance.

Echinocandins are used as first-line therapy for C auris, pending antifungal susceptibility test results, unless the patient is an infant less than 2 months of age, in which case amphotericin B deoxycholate is recommended. Development of echinocandin resistance (ie, MICs >1 μg/mL for caspofungin) has been documented in C auris isolates recovered from patients in multiple geographic areas who were initially treated with an echinocandin. Echinocandins act by targeting the fungal-specific β-glucan synthase, which synthesizes a major cell wall polymer. Mutation to the Fks subunits of the glucan synthase in hot-spot regions decreases susceptibility of fungi to echinocandins. Isolates of C auris with elevated echinocandin MICs harbor the mutation S639F in the position equivalent to the well-characterized S645 FKS1 hot-spot 1 region.⁴³  General guidance on treatment recommendations may be found at https://www.cdc.gov/fungal/candida-auris/c-auris-treatment.html. It should be noted that for cases of C auris, treatment is indicated only if clinical disease is present or if the organism is isolated from a sterile site. Treatment is not recommended when C auris is identified in a noninvasive infection or if the patient exhibits no clinical signs of infection.

Interpretation of reduced susceptibility to azoles, polyenes, and echinocandins is challenging because no antifungal breakpoints for C auris have been defined by the Clinical and Laboratory Standards Institute (CLSI) or the European Committee on Antimicrobial Susceptibility Testing. However, tentative epidemiologic cutoff values (ECVs) have been established for both the CLSI and European Committee on Antimicrobial Susceptibility Testing methods, which differ slightly.⁴⁰  It should be cautioned that these ECVs are tentative, as the study used to derive them was not performed according to European Committee on Antimicrobial Susceptibility Testing or CLSI specifications for establishing ECVs. Furthermore, ECVs are not a clinical breakpoint, and isolates with MICs below the ECV should not be reported as susceptible, nor should isolates with MICs above the ECV be considered resistant. Insufficient pharmacodynamic and clinical data are available to establish clinical breakpoints for this species. The CLSI guidance on interpreting ECVs can be found in the CLSI M59 standard.⁴⁶  On the basis of ECVs and identified molecular mechanisms of resistance, the CDC has established tentative breakpoints. These breakpoints are informational only and have not been approved by any regulatory agency or standardization committee (Table).²¹ 

Analytical challenges can make evaluating antifungal susceptibility test results for C auris a daunting task. Kordalewska et al²⁶  documented a caspofungin Eagle effect for isolates of C auris that lacked the FKS1 mutation associated with echinocandin resistance. The Eagle effect, also known as a paradoxical effect, occurs when growth is observed at higher but not lower concentrations of drug, possibly because of dose-dependent echinocandin tolerance in response to cell-wall damage.⁴⁷  The isolates showed in vivo susceptibility to caspofungin at human therapeutic doses in a murine model of invasive candidiasis. Mixed results have been obtained when evaluating C auris susceptibility by commercial antifungal susceptibility test systems. Kathuria et al²³  noted that the Vitek 2 generated higher amphotericin B MICs than did the CLSI broth microdilution reference method for a collection of 90 Indian C auris isolates, with MIC₅₀ of 8 μg/mL and 1 μg/mL, respectively. In contrast, Etest (bioMérieux) was comparable to broth microdilution for this drug, with an MIC₅₀ of 0.5 μg/mL. Caspofungin and voriconazole MICs were similar among the 3 methods. Magobo et al¹¹  reported falsely high caspofungin MICs by using the Vitek 2, which is particularly concerning because caspofungin is recommended as first-line therapy for C auris infections.

Because resistance of C auris to antifungal agents has been shown to emerge while a patient is receiving therapy, repeated susceptibility testing should be performed if a laboratory isolates C auris on subsequent cultures. In the United States, isolates should also be referred to the local public health laboratory for confirmatory identification and antifungal susceptibility testing. As there are no CLSI- or FDA-approved specific interpretative breakpoints for C auris susceptibility testing, MIC-only results should not be reported.

Patients with C auris infections often become colonized with C auris at multiple body sites, which can include the axilla, groin, nares, ears, mouth, rectum, and vagina, but most often C auris is detected in the axilla and the groin.^12,14,48  A contaminated patient room or intensive care unit can lead to transmission of C auris and subsequent colonization or infection, or both, of other patients. In one hospital, more than 50% of C auris infections were preceded by colonization of the patient, which emphasizes the need to prevent colonization and to identify those who are colonized.⁴⁸ 

Both culture-based and molecular-based techniques for the detection of C auris colonization have been developed.^29,49  The culture technique is based on the ability of C auris to grow at 10% NaCl and 40°C and to use dulcitol as a carbon source to enrich for C auris over other Candida species that colonize the skin.²⁹  This technique has been successfully used during multiple surveys of health care facilities to detect patients colonized with C auris.^14,35  The molecular assay is a TaqMan-based real-time PCR assay that targets the ITS region of the ribosomal cistron.⁴⁹  Although it only has 89% sensitivity for detection of C auris from swabs compared with culture, it reduces the overall time to positivity from days to hours, allowing the rapid implementation of infection control procedures. There is an additional SYBR Green detection real-time PCR assay that has been developed, but this assay has not yet been optimized for use with specimens collected on swabs.²⁶ 

It is unlikely that persistently colonized health care personnel serve as a major source of transmission. During outbreaks in both England and Spain, health care personnel were screened for C auris colonization. Multiple sites of 258 health care workers in England were screened; sites included the hands, nares, axilla, and groin. Only a single positive nares swab was found, and colonization in that person was transient.¹²  In Spain, the hands and ears of 101 personnel who worked in a surgical intensive care unit were screened, and none were positive.⁴⁸  However, it is possible that health care personnel who are transiently colonized moving from a colonized or infected patient to a subsequent patient or from a contaminated environmental surface to a patient could transmit C auris in health care settings.

Decolonizing skin has proven to be an additional challenge. Chlorhexidine wipes (2%) with isopropyl alcohol are effective in eliminating C auris from surfaces, as is povidone iodine. However, their efficacy for skin has not been proven.^50,51  In vitro, 2% and 4% chlorhexidine have been shown to be effective against C auris, but the effect on decolonization of patients is not known. Even if the amount of C auris on the skin can be reduced through the application of these antiseptics, it is unclear what effect this will have on transmission because patients may continue to harbor reservoirs of C auris in the ears, nares, gastrointestinal, or genitourinary tracts.

In health care settings, C auris has been recovered from mattresses, beds, chairs, tables, floors, walls, carts, windowsills, equipment monitors and keypads, and countertops.^12,35,48  Several reports have shown that C auris can persist for 7 to 14 days on surfaces such as steel, ceramic, and plastic.^29,52  There is some evidence that C auris may enter a viable, nonculturable state for up to 4 weeks on some surfaces.²⁹  Quaternary ammonium compounds alone, which are the most common disinfectants used in health care settings, might not achieve the desired log reduction against C auris and are therefore not recommended for use. Many chlorine-based and peroxygen-based disinfectants appear to be effective against C auris and have shown substantial log reductions on surfaces and in solution (≥4 log₁₀).^51,53,54  Other disinfectant products that have been shown to achieve substantial reductions in C auris include some quaternary ammonium compounds with alcohol.^29,50,53,54  Mobile, short-wavelength ultraviolet light devices are also effective in reducing the bioburden of C auris substantially, and exposure times are less than those required for Clostridioides (Clostridium) difficile.⁵⁵ 

Candida species have not previously been considered organisms for which contact precautions were necessary. However, there is ample evidence that C auris can be transmitted from person to person in health care facilities.^12,14,48  Therefore, it is imperative that C auris is identified accurately by the laboratory so that infection control procedures can be put in place. The CDC has published guidance on infection control measures for C auris (https://www.cdc.gov/fungal/candida-auris/c-auris-infection-control.html).⁵⁶  Patients infected or colonized with C auris should be placed in a single-patient room and contact precautions instigated. Meticulous hand hygiene with an alcohol-based hand rub or soap and water is required for health care personnel caring for patients with C auris. Cleaning and disinfection of the patient room should take place daily, and then a terminal cleaning should be done before other patients use the room. Currently, the CDC recommends the use of an Environmental Protection Agency–registered disinfectant that is effective against C difficile spores (List K [https://www.epa.gov/pesticide-registration/list-k-epas-registered-antimicrobial-products-effective-against-clostridium])⁵⁷  until Environmental Protection Agency–registered products specifically designated for C auris are available. Lastly, close contacts of newly identified patients with C auris should be screened for colonization by swabbing of the axilla and groin.⁵⁸ 

Because of the clinical and epidemiologic implications of C auris infection, diagnostic laboratories are advised to identify to species level all Candida organisms isolated from sterile body sites, Candida isolates with unusual antifungal resistance profiles, isolates from patients with possible colonization based on known exposure or risk factors, or isolates from patients with prior C auris exposure.^59,60  If C auris is suspected, it is critical for clinical laboratories to partner with institutional infection preventionists to complete the clinical and epidemiologic details pertinent to the case in order to reduce the risk of nosocomial transmission. Isolates may be referred to reference laboratories, state public health laboratories, or the CDC for assistance with definitive identification. Recommendations for C auris identification may be found at https://www.cdc.gov/fungal/diseases/candidiasis/recommendations.html#identify. If a clinical laboratory has the capacity to confirm identification of suspect isolates, every report of C auris should be immediately conveyed to local and state health officials in addition to the CDC via [email protected].

In the clinical microbiology laboratory, identification of C auris in an accurate and timely fashion is challenging but essential for patient care and infection prevention. Clinical laboratories must assess their ability to identify C auris based on the methodology they use. Antifungal susceptibility profiles and interpretations in addition to treatment recommendations for patients with C auris must be understood for appropriate clinical management. Because of the ability of C auris to spread in health care facilities and cause outbreaks, rapid communication of suspected cases with institutional infection prevention providers and local, state, and national surveillance networks is essential to enhance awareness of this newly emerging yeast species.

We gratefully acknowledge the contributions of the College of American Pathologists Microbiology Committee for concept development, the Mycotic Diseases Branch at the Centers for Disease Control and Prevention for technical expertise, and Megan Bentz, BS, MS, for laboratory photography. We also gratefully acknowledge Diana M. Meza Villegas, MS, SM(ASCP)^CM, and Miranda E. Diaz, BS, SM(ASCP)^CM, for data collection and photography, and the Mayo Clinic Clinical Microbiology Laboratory in Jacksonville, Florida, for laboratory support.