An overview of Candida auris: The emerging healthcare threat

Literature search

For the preparation of this review article, relevant scientific publications in the English language were searched on “Search Engines” such as “Google Scholar,” “PubMed,” and “Scopus.” Relevant MeSH/search terms such as “Candida auris,” “Candida auris infection,” “Transmission of Candida auris,” “Epidemiology of Candida auris,” “Molecular epidemiology of Candida auris,” “Laboratory diagnosis of Candida auris,” “Antifungal resistance in Candida auris,” “Prevention and control of Candida auris infection,” and other relevant search terms pertaining to C. auris. To extract more studies, the references cited in the publications referred to in preparing this manuscript were intensively studied.

Inclusion criteria

After assessing the title and the abstract of the selected studies, full texts were downloaded and meticulously reviewed for the preparation of the manuscript. Original research articles based on cross-sectional studies, reviews, case reports, and retrospective studies were included in this study, whereas letters to the editor, dissertations, theses, and abstracts of conferences were excluded from the study.

Microbiological characteristics

Morphologically, C. auris resembles other Candida spp and is oval and round to ovoid in shape and may exist as aggregate or non-aggregate cells. Unlike C. albicans and Candida dubliniensis and similar to many other non-albicans Candida (NAC) spp., C. auris lacks the ability to form a germ tube. Few strains of C. auris are capable of forming a germ tube when grown on cornmeal agar or incubated in fetal bovine serum.^[5] C. auris do not produce pseudohyphae. However, it is reported that when C. auris is grown at high concentrations of salt (NaCl), it may produce pseudohyphae.

C. auris grows at temperatures ranging from 37°C to 42°C. It is also reported that C. auris may demonstrate different morphologies on different media when grown at different temperatures. Cycloheximide in the media inhibits the growth of C. auris.^[6]

When grown on Lee’s glucose and Lee’s N-acetylglucosamine (GlcNAc) media, cells of C. auris demonstrate an oval shape at both 25°C and 37°C; on Lee’s glucose and Lee’s GlcNAc media, yeast cells of C. auris are oval-shaped, whereas they are round (relatively) at 40°C. C. auris cells appear round and comparatively smaller when grown on agar plus serum media. On routine Yeast Extract Peptone Dextrose (YPD) media, cells of C. auris are round, whilst when the same medium is supplemented with 10% NaCl, the cells are elongated and opaque, resembling those of C. albicans. On this medium (YPD with 10% NaCl), highly elongated cells, mimicking pseudohyphae, may be evident in small percentages. Colonies are smaller, relatively. C. auris cells are round on regular YPD Medium.^[7]

Upon staining with 4’6-diamidino-2-phenylindole (DAPI) multiple nuclei are evident with DAPI staining, multiple nuclei are observed. With Calcofluor stain, no septin/chitin rings are observed between the conjoined cells.^[8] On Sabouraud dextrose agar (SDA), the most common media used in mycology, C. auris produces colonies identical to other members of the genus. Colonies on SDA are smooth, pasty, and beige colored.

C. auris is capable of fermenting sugars such as glucose, sucrose (weak), and trehalose (weak). However, it does not ferment galactose, maltose, lactose, or raffinose. It can assimilate glucose, sucrose, maltose, d-trehalose, d-raffinose, d-melezitose, inulin (weak), soluble starch, ribitol (weak), galactitol, d-mannitol, sorbitol, and citrate.^[9]

It is capable of utilizing ammonium sulfate, cadaverine, and L-lysine as sole sources of nitrogen. However, it does not utilize sodium nitrite, potassium nitrate, and ethylamine. C. auris grows in vitamin-free media and glucose (50%) and NaCl (10%)/glucose media (5%). Urease test and diazonium B reaction are negative. It is inhibited in the presence of cycloheximide in concentrations of 0.1% and 0.01%.^[9]

Although C. auris produces pink/beige/pale rose/red color colonies on chromogenic media, chromogenic media often fail to accurately identify it [Figure 1]. However, when chromogenic (Chrom agar Candida) media are supplemented with Pal’s agar, it can be distinguished from Candida haemulonii. On this medium, when grown at 37°C, C. auris produces pink- or beige-colored smooth colonies, whereas C. haemulonii complex shows sparse growth and colonies are light-pink colored. This simple and cost-effective technique for identifying C. auris was suggested by Kumar et al.^[10]

Close

Figure 1:

Light-colored colonies of Candida auris on Hichrom Candida medium.

Export to PPT

CHROMagar Candida Plus (CHROMagar, France) appears to be a potential alternative both for presumptively identifying and differentiating C. auris from clinical specimens, with a sensitivity and specificity of 100%. On this medium, after 36 hours of incubation, C. auris produces characteristic colonies that are light blue in color and are surrounded by a blue halo.^[11]

As illustrated in Table 1, both conventional biochemical tests and automated systems like “API20C,” “Vitek 2 YST,” “BD Phoenix,” “MicroScan” and “RapID Yeast Plus” often misidentify C. auris owing to its close phylogenetic similarity to C. haemulonii complex and Candida ruelliae.

“The Vitek 2 XL” (BioMerieux version 8.01) is updated for the identification of C. auris. However, Ambaraghassi et al.^[12] reported that this software-based system can accurately identify only African and East Asian clades of C. auris, and therefore it is essential to confirm by molecular-based biotyping methods, such as matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) or Deoxyribonucleic acid (DNA) sequencing, all isolates that are identified as C. auris, Candida famata, or C. haemulonii complex by Vitek 2 XL.^[12]

Many researchers have highlighted the importance of MALDI-TOF for identifying C. auris isolates and recommended it as both an accurate and reliable technique for the identification of C. auris. However, this method of biotyping may be both lengthy and expensive, particularly when all colonies are to be tested in case of isolation from colonized patients.^[11]

Several novel methods are evaluated for both the identification and differentiation of C. auris from its closely related species. These include methods for detection of proteins and DNA by polymerase chain reaction (PCR)-based methods such as conventional PCR, real-time PCR, multiplex probe amplification, PCR-restriction fragment length polymorphism, T2 magnetic resonance (T2MR) system, loop-mediated isothermal amplification, and Germline Pathogen System Monodose Candida auris Detection Quantitative Polymerase Chain Reaction [GPS_TM MONODOSE dtec-qPCR kit (dried single-dose PCR tubes)]. CanAur dtecqPCR (dried single-dose PCR tubes). Molecular methods are more precise and accurate for identifying C. auris isolates, as their identification is based on sequencing the D1-D2 region of the 28S ribosomal DNA.^[13]

Similarly, commercial techniques such as AurisID (a species-specific molecular assay for C. auris), “Fungiplex (Fungiplex Candida auris PCR assay),” “the T2MR assay,” “the BioFire^® Blood Culture Identification 2 (BCID2) panel,” and “the GenMark Dx ePlex Blood Culture Identification Fungal Pathogen” panel are also recently available for the identification of C. auris from clinical specimens.^[14] The laboratory approach for the identification of C. auris has been shown in Figure 2.

Close

Figure 2:

Laboratory approach for identification of Candida auris from a clinical specimen. MALDI-TOF: Matrix-assisted laser desorption ionization–time of flight.

Export to PPT

Clinical manifestations

Although many studies on C. auris are focused on invasive infections, including candidemia, it is also reported to be involved in non-invasive infections such as vulvovaginitis and dermatological manifestations.

Researchers have reported isolation of this emerging Candida spp. from variety of clinical specimens including blood culture, catheter tips, cerebrospinal fluid, bone, ear discharge, various types of body fluids (pancreatic, pericardial, peritoneal, pleural) respiratory secretions such as sputum and broncho-alveolar lavage (BAL), skin, and soft-tissue samples (tissue and swab), urine and vaginal secretions.^[15]

In recent years, candidemia due to C. auris has garnered attention, particularly due to the high treatment failure rate and nosocomial dissemination. Bloodstream infection is the most common clinical form of invasive manifestations caused by C. auris.^[15]

Bloodstream infection (BSI) due to C. auris is associated with increased mortality rates. Studies have reported the crude mortality rates for C. auris BSI to range between 22% and 66% across a broad range of healthcare setups and patient populations.^[16,17]

The risk factors for C. auris invasive infections are the same as those for other species of Candida. These include immunocompromised status (acquired/induced), comorbidities, exposure to wide-spectrum antimicrobials, intensive care unit (ICU) admission, invasive medical procedures, and surgical interventions.^[18]

In a case-control study conducted in Indian ICUs, Rudramurthy et al. (2017)^[17] compared risk factors for BSI due to C. auris and other Candida spp. These authors noted that long hospital stay, presence of respiratory diseases, poor acute physiology and Chronic Health Evaluation II scores, vascular surgeries, and exposure (therapeutic/prophylactic) to antifungal drugs are risk factors for C. auris fungemia.^[17] The risk factors for C. auris BSI have been summarized in Table 2.

There are several reports available on colonization or infection or both of the urinary tract by C. auris. Diarrhea and gastrointestinal decompression are important risk factors for colonization/infection of the urinary tract by C. auris. It is postulated that migration of C. auris (colonized) from the intestinal tract to the urinary system leads to its colonization/infection/both. ICU admission, renal calculi, urinary catheterization, perirenal inflammatory conditions, and moderate hydronephrosis are risk factors for urinary tract colonization/infection due to C. auris. Biagi et al.^[19] reported that misidentification/incomplete identification of C. auris from urine cultures often complicates the treatment.^[19]

Although rare, pulmonary infections due to C. auris have been documented. The first case of severe pulmonary infection due to C. auris from India was reported by Choudhuri et al. in 2024.^[20] These authors reported severe pulmonary manifestation of C. auris complicated by hydropneumothorax and bronchopulmonary fistula in a 71-year-old male with diabetes mellitus and hypertension. Initially, it was considered a case of pulmonary tuberculosis, and in this patient anti anti-tuberculosis treatment was started despite the test for TB being negative. However, the condition of the patient deteriorated. Upon culture of endotracheal secretion, C. auris was isolated, and the patient was shifted from the antifungal regimen of caspofungin to posaconazole. With posaconazole therapy, the patient showed significant clinical improvement and was discharged from the hospital. Choudhuri et al. highlighted the importance of high clinical suspicion of C. auris infections in critically ill patients.^[20]

Wang et al. (2018)^[21] reported the first case of C. auris from a BAL specimen in China.^[21] This isolate was found to be susceptible to caspofungin, amphotericin B, and fluconazole.^[21] Rare involvement of C. auris in clinical conditions such as pericarditis, ophthalmitis, and otitis has also been documented.^[22]

Epidemiology

Several unique characteristics of persistence in a healthcare setup, marked resistance to antifungal and disinfectants, survival on a number of biotic and abiotic surfaces, including the hospital environment and medical devices, make C. auris a “super fungus.” The characteristic ability of forming “dry biofilms” and “aggregative phenotypes” makes the eradication of C. auris more difficult.^[23] These characteristics aid in nosocomial transmission (direct/indirect) of C. auris.^[24]

Research studies have revealed that once C. auris is introduced in a healthcare setting, it spreads rapidly, particularly among susceptible patients. Outbreaks are reported from various countries such as the United States of America (USA), the United Kingdom (UK), Canada, Mexico, Spain, India, Saudi Arabia, Oman, Kuwait, Kenya, Colombia, and South Africa [Figure 3].

Close

Figure 3:

Outbreak of Candida auris. Source: Author-created image with mapchart.net

Export to PPT

Lockhart et al. (2016)^[14] conducted a whole-genome sequencing (WGS) for analyzing the epidemiology of the emergence of multi-drug resistant (MDR) C. auris in three continents. Interesting findings were noted in the study. Results of WGS showed that C. auris isolated from cases of candidemia outbreaks in Pakistan had clonal genetic relatedness to Indian strains, whereas isolates from the USA had similarity to South America and South Asia. Some strains were phylogenetically different from India, Pakistan, and Venezuela but were similar to those isolated from the UK.^[14]

Epidemiological studies reveal C. auris has emerged simultaneously in East Asia and other parts of Asia, Africa, Europe, and South and North America.^[25] In the USA, the case of C. auris was reported in 2013, whereas in New York, it was identified in 2016. C. auris was incorporated in the list of national notifiable pathogens in 2018. It was documented in 24 states of the USA and approximately 55% of the cases (clinical and surveillance) from New York and the Metropolitan area of New Jersey.^[18,26]

Globally, the crude mortality rate in C. auris infection is reported between 33.33% and 100%.^[27] Many research studies have revealed that the incidence of C. auris is high in males in most countries except in South Africa, where female patients outnumber male patients. The reported global prevalence of C. auris BSI ranges between 5% and 30%.^[16,27,28]

In recent years, the incidence of healthcare-associated BSI due to C. auris has drastically increased; in some instances, it has even surpassed C. tropicalis and C. glabrata in some healthcare setups, even in a few Indian hospitals.^[29-31]

However, the exact prevalence of C. auris is yet to be correctly determined, primarily because of the misidentification of this species in laboratories that rely on conventional methods.^[27] The study conducted by us in a rural tertiary referral hospital revealed the prevalence of C. auris to be 9.4% among various NAC spp.^[32] Previously, isolates of C. auris were classified into four distinct clades that were considered to be geographically restricted. These include Clade I (South Asia), Clade II (East Asia), Clade III (South Africa), and Clade IV (South America). However, there is a possibility of the V^th Clade from Iran.^[8,33-35]

Various factors, such as detection methods along with a lack of screening protocols both for patients and healthcare workers, hinder the overall surveillance of C. auris across the world, especially in developing countries. A widespread variation is noted in the distribution of geographical clades, in addition to variation in several thousand nucleotide polymorphisms. Till recent days, there were five distinct geographic clades of C. auris.^[35] These include clades such as the South Asian, the South African, the South American, the East Asian, and the Iranian.^[36] The details of clades of C. auris reported from outbreaks in various countries are shown in Table 3.

Virulence traits

As compared to other common NAC spp., the virulence traits of C. auris are not completely elicited. The highly potent ability to disseminate and high rates of mortality directly provide evidence that C. auris has aggressive virulence properties.

Virulence traits of C. auris include polymorphism, production of exozymes (hemolysis, phospholipases, proteinases, lipases), biofilm formation, morphological plasticity, filamentation formation, osmo-tolerance, thermotolerance, and phenotypic switching.^[22]

Borman et al.^[37] reported that few strains of C. auris are capable of producing pseudohyphae, and these isolates are often more virulent than isolates incapable of producing pseudohyphae.^[37] The virulence trait of C. auris and its contribution to pathogenicity have been shown in Table 4.

Antifungal resistance

C. auris is considered a MDR fungal pathogen. Research studies focused on the evaluation of drug resistance mechanisms in C. auris have put forward several mechanisms responsible for resistance.^[38]

These include environmental factors such as climatic variation and agricultural practices, genetic variation that includes polyploidy, aneuploidy, and chromosome rearrangements, biofilm formation, mutation, and overexpression of drug target and restricted drug intake or efflux.^[38]

For treating and monitoring mycotic infections, till recent date, only three classes of antifungal drugs (azoles, echinocandins, and polyenes).^[39] In C. auris, resistance to two classes of antifungal drugs is observed in approximately 40% of the isolates, whereas 4% of isolates are reported to be resistant to all three classes of antifungal agents.^[39]

In the study by Chow et al.,^[40] C. auris demonstrated high antifungal resistance. A total of 80% isolates were noted to be fluconazole resistant, amphotericin resistance was also high (23%), whereas micafungin resistance was observed in 7% of the isolates. Resistance to two classes of antifungal drugs was observed in 24% of isolates, whereas 1% of isolates were resistant to all three classes of antifungals.^[40]

Fluconazole, the most widely used antifungal for the treatment of prophylaxis of various clinical manifestations, acts by restricting cell growth by inhibiting ergosterol synthesis, particularly the lanosterol 14-α-demethylase. The lanosterol 14-α-demethylase is encoded by the ERG11 gene and therefore the mutations of ERG11, TAC1b, Y132F, K143R, and F126L genes, as well as the ATB-binding cassette (ABC) and major facilitator superfamily transporters, are capable of conferring resistance to azole.^[41] In addition, in some C. auris strains, the substitution mutations, restricted to geographical-specific clades, are responsible for azole resistance. It is also observed that substitution mutations of certain strains of C. auris that are restricted to geographical clades are proven resistant to azoles.^[38]

It was noted that in a study based on 350 strains of C. auris from India and South Africa, in nearly 90% of isolates, the minimum inhibitory concentration (MIC) for fluconazole was greater than 16 µ/ML.^[39] As compared to Nakaseomyces glabrata (previously known as Candida glabrata) and C. haemulonii, the concentration of substrate, Rhodamine 6G, essential for the ABC-type efflux pumps, was higher in C. auris.^[42] Mutations in the TAC1b gene are also reported as a mechanism responsible for fluconazole resistance in C. albicans by researchers like Li et al.^[43]

As per Rhodes et al.,^[44] flucytosine resistance may occur because of a substitution in an F211I amino acid that is located in the FUR1 gene.^[44] Similarly, mutations in MEC3 elevate the MIC for polyene and caspofungin in C. auris.^[45] Amphotericin B resistance can also be attributed to alterations to the cell membrane sterol and/or a given point mutation.^[34]

As the resistance to azoles and polyenes is increasing, echinocandins are now considered the first line of drugs for treating C. auris infections. However, the mutations in the FKS1 gene encoding for a subunit of the β-D-glucan synthase are targeted by echinocandins.^[46] Emerging antifungal agents such as ibrexafungerp and rezafungin have shown promising activity against C. auris in vitro, highlighting their potential as future therapeutic options, particularly where resistance to current echinocandins is observed.^[47]

Infection prevention and control practices

C. auris bears every potential to spread in nosocomial environments, and hence it is imperative to practice adherence and strict compliance with infection prevention and control protocols that include identifying the cases, screening of suspected/at high-risk patients, strict adherence to standard and transmission-based precautions, environmental cleaning, and patient decolonization.^[48]

Disinfectants such as sodium hypochlorite and products containing hydrogen peroxide are most effective and recommended for disinfecting the rooms, equipment, and instruments of infected individuals. Hand hygiene is one of the most important protocols for preventing and limiting the nosocomial spread of C. auris. Hand wash (soap and water) and hand rubs (alcohol- and chlorhexidine-based) are important components of hand hygiene. The dissemination of C. auris can also be restricted/prevented by quarantining the patients in separate rooms.^[38]