Candida auris: a review of recent literature

Question 1

Introduction

Accepted Answer

Candida auris is an emerging fungal pathogen, first identified in 2009 from the external auditory canal of a patient in Japan (1). It has since been identified worldwide, across six continents, and in more than 40 countries (2). The recent inaugural World Health Organisation fungal priority pathogen list considers C. auris a critical priority pathogen, highlighting its threat to global public health (3). C. auris infections have been frequently reported from the bloodstream and also seen in association with bone, CSF, and intra-abdominal infections (4). Additionally, it has been isolated from wounds, ear and respiratory specimens, urine, bile, and jejunal biopsies (5). Detection in surveillance swabs from axilla and groin may indicate carriage rather than infection, with carriage posing a risk of transmission to others and of subsequent invasive infection.

Specialised laboratory methods are required to identify C. auris. On microscopy it can be indistinguishable from other Candida species. This can cause challenges with identification, outbreak detection, and control. As with other nosocomial infections, but unlike other Candida species, C. auris appears to be highly transmissible between patients and in healthcare settings from contaminated environments or equipment and is also associated with prolonged persistence in the environment (6, 7).

There have been five genetically distant clades of C. auris discovered to date. This includes the South Asian clade, which was first detected in India and Pakistan (clade I), the East Asian clade first detected in Japan (clade II), the South African clade first detected in South Africa (clade III), the South American clade first detected in Venezuela (clade IV), and a further clade recently detected in Iran (clade V) (8). Antifungal drug resistance is a common feature of C. auris isolates, and patterns vary by clade and region. Invasive infections from multidrug resistant C. auris have been associated with high mortality, and concerns have been raised regarding transmission, persistent colonisation, and need for effective disinfection (9, 10). Of concern, active community spread of independent clusters of multidrug-resistant C. auris has now been identified in the United States (U.S.) (11, 12).

There are many unanswered questions regarding C. auris (13). A review of the most recent literature has been conducted to consider any key updates in the following areas of epidemiology and genomic analysis, experience in the UK; the COVID-19 pandemic; identification and typing, resistance profiles and treatment, colonisation and infection, and infection prevention and control.

Question 2

Methods

Accepted Answer

In order to identify any recent updates in relation to C. auris since a previous review of the literature, a search of articles was conducted, between 2019 and 2023, with snowballing for any other significant studies. This included Medline, Embase, Scopus, NICE Evidence Search, Global Health, and CINAHL, and was limited to publications in the English language. The search terms Candida auris and C. auris were used. Articles were deduplicated and excluded if there was no reference to C. auris or if they did not contain information relevant to the key areas of the review, as described above.

Question 3

Background

Accepted Answer

C auris was first detected in 2009 in Japan, however, the first retrospectively identified isolate actually dates back to 1996 (1, 14). In South Korea, 15 patients with samples taken between 2006 and 2011 were found to have C. auris, all with a background of chronic otitis media and systemic antifungal use (15). Following additional epidemiological investigation, it was proposed that intra- and interhospital clonal transmission had occurred (16). In Europe the first imported case dates back to 2007, and this was of the South Indian clade (clade I) (17).

Despite these discoveries, retrospective analyses of international samples support that C. auris was a novel pathogen prior to 2009. From 15,271 retrospectively analysed Candida isolates taken between 2004 and 2015, only four were found to be C. auris, all of which had been collected since 2009 (18). A further retrospective analysis of unusual Candida species performed at the Mycology Reference Laboratory, UK Health Security Agency (UKHSA), formerly Public Health England (PHE), failed to find any evidence of C. auris in the United Kingdom (UK) prior to 2013 (19).

The first reports of invasive fungaemia associated with C. auris were from a multicentre surveillance study of candidemia conducted in South Korea in 2009, with one case dating back to 1996 from an incidental finding of unidentified yeasts in bloodstream isolates (14) . All cases had been hospitalised for at least 12 days before isolation of the organism. Invasive infections were also discovered in India dating back to 2009, initially in 12 inpatients from two hospitals in New Delhi, and clonal isolates suggesting interhospital transmission (20).

As of February 2021, according to data published by the CDC, C. auris had been identified in 47 countries across six continents (21). In the U.S. C. auris became notifiable in 2018, and reports of case detection subsequently increased more than two-fold. Between June 2020 and May 2021 more than 1,000 clinical cases had been identified across the U.S. and targeted screening had identified a further 2,386 colonised individuals (21). In response to a substantial rise in detection in European countries, a rapid risk assessment of C. auris in healthcare settings was updated by the European Centre for Disease Prevention and Control (ECDC) in 2018 (22), with a further risk assessment in 2022 following an outbreak in Northern Italy. Additionally, a survey by the ECDC in 2020 reported ongoing variability in laboratory capacity and overall preparedness for C. auris detection (23).

C. auris represents a significant burden of disease in certain countries and has become endemic in several (24, 25). In South Africa, the organism has been detected in almost 100 hospitals, causing large outbreaks, and is reportedly responsible for approximately 1 in 10 cases of candidemia (24). In India, C. auris has been implicated in 5% of candidemia cases across 27 intensive care units (26).

Healthcare outbreaks have been reported from hospitals in the UK, Spain, US, Algeria, Columbia, Greece, Italy, Kenya, Kuwait, Lebanon, Oman, Pakistan, Venezuela, Brazil, Qatar, Saudi Arabia, Turkey, as well as India and South Africa (24, 27-47). Sporadic cases have been reported from Norway, Denmark, Germany, France, Australia and recently, Bangladesh, Canada, China, Austria, Switzerland, Iran, Nigeria, Taiwan, and the Netherlands (48-60). In Australia, the first detection of C. auris was of the South African clade and occurred in Perth in 2015; since then, cases have been detected in Victoria and New South Wales (61).

Community prevalence of C. auris remains unknown, and screening on admission to hospital is not routine practice. A surveillance study in England was performed in 2017-18, involving screening of 998 admissions to eight intensive care units across three major cities. All screens were found to be negative, reflective of low prevalence on admission (62). Another study in the U.S. found that carriage was only identified in people previously exposed to the hospital environment (63). In areas where C. auris is endemic, however, community cases may be more prevalent. A recent study looking at C. auris amongst patients with chronic respiratory disease in India found that 9.3% (3/32) of patients who were colonised with C. auris were colonised at the time of admission (7). Many patients would have had previous contact with hospital services, prior to the admission due to the nature of their chronic illness.

Question 4

Genomic analysis and epidemiology

Accepted Answer

Whole genome sequencing (WGS) analyses indicate that C. auris emerged simultaneously in several different continents, with four distinct clades across three continents; South Asian (clade I), East Asian (clade II), African (clade III) and South American (clade IV) (64). A further genetically distinct clade has more recently been isolated in Iran from a 14-year-old girl who had never travelled abroad and was diagnosed with otomycosis, followed by a further fluconazole resistant case distinct from the first (65, 66).

Each of the clades are separated by thousands of single-nucleotide polymorphisms (SNPs), but strains are highly clonal within each clade, with on average fewer than 70 SNPs separating any two isolates within a given clade (13). This, along with various geographic resistance mechanisms support the hypothesis of independent clonal expansion and evolution. Potential drivers for the emergence of the pathogen include increasing antifungal selection pressures in humans, animals and the environment (64).

A retrospective analysis of 912 worldwide cases across 44 countries, between 2009 and 2020 found the South Asian strain (clade I) to be the most prevalent, having been identified in 17 countries (67). This was followed by the South African strain (clade III) found in eight countries (67). Only five countries reported presence of the East Asian strain (clade II) and three of the South American strain (clade IV).

Question 5

Experience in the UK

Accepted Answer

A retrospective analysis of historical isolates showed the first identified C. auris isolates in the UK were in 2013, from blood cultures in unrelated patients (19). In 2014, a single isolate from pleural fluid was recorded, and in 2015 there were 15 isolates identified, nine of which were from sterile sites. Between 2013 and 2021, 304 isolates of C. auris have been identified in England, with a total of 40 (13.2%) being from blood specimens. In 2020 there were just four colonisations identified, probably reflecting the lack of new introductions from endemic areas due to extreme travel restrictions associated with COVID-19 regulations. All clades except the South American clade have been identified to date. Sporadic introductions continue, many involving patients repatriated directly from international hospitals in endemic regions, including India, Qatar, Kuwait, Oman, Pakistan, and Kenya (68).

Outbreaks have been identified in three independent hospitals in England, each of which was prolonged and challenging to control (28, 69-71). The first was in a cardio-thoracic centre between 2015 and 2016, involving 50 patients over a 16-month period. The majority of cases were confined to colonisation of skin sites or mucosa, and 44% (n = 22/50) required anti-fungal therapy (69). The second outbreak started in 2016, and involved 34 patients including eight bloodstream infections. It was controlled within 13 months by an intensive screening and infection control approach maintaining a low mortality rate (178). The third outbreak in 2017, involved an intensive care setting with 70 patients having been colonised or infected, linked to reusable axillary temperature probes indicating that the pathogen can persist in the environment and be transmitted in health care settings (28).

A national incident management team (IMT) was established in 2016 in relation to C. auris in England, and this was led by the UKHSA. Recent transmission chains have been limited in light of early recognition, appropriate case isolation, enhanced infection prevention, and control measures, and wider screening (68).

Question 6

C. auris and COVID-19

Accepted Answer

Early in the pandemic patients with severe COVID-19 infection, including those in intensive care, were particularly vulnerable to bacterial and fungal co-infection (72-74). With studies having independently identified that underlying respiratory illness and mechanical ventilation in intensive care are significant predictors of C. auris colonisation and infection, it was highlighted that the SARS-CoV-2 pandemic could provide unfortunate conditions for widespread novel outbreaks of C. auris (19, 75).

Although the spread of resistant pathogens, including C. auris, has been observed amongst critically ill COVID-19 patients during peaks of the pandemic in certain countries, including Italy and the U.S., this has not been the experience of all areas (76, 77). In the UK, despite there having been large numbers of patients in intensive care settings, C. auris outbreaks were not identified during the pandemic.

The Americas

The Pan American Health Organisation (PAHO) released an epidemiological alert in 2021 in response to C. auris outbreaks in the context of the COVID-19 pandemic (78). In 2020, seven countries had documented cases of C. auris mostly in patients with COVID-19, including Brazil, Guatemala, Mexico, Peru, the U.S., Panama, and Columbia. C. auris had not been identified in four of these countries prior to the pandemic. Several outbreaks of C. auris have been identified in the U.S. in COVID-19 units of acute care hospitals, and new cases without links to known cases or healthcare abroad have also been identified across multiple states (77, 79). In Brazil, two colonised patients were identified in a COVID-19 intensive care unit in December 2020 (80). Screening was undertaken and an outbreak was subsequently identified with 8/47 (17%) patients testing positive from the axilla, as well as positive findings from environmental screening. Digital thermometers demonstrated the highest rate of positive cultures from environmental screening, followed by bed rails, vital sign monitors/intravenous infusion pumps, and tray tables (80). With travel restrictions during the pandemic and the absence of travel history among the colonised patients, it was hypothesised that the species was introduced several months before recognition of the first case (81).

In Guatemala, two isolates were detected on the same surgical unit, the first isolated in soft tissue and bone biopsies in a patient with osteomyelitis and the second from a surgical site infection, neither case being COVID-19 related (78). A hospital outbreak in Mexico also started with the identification of infection in a patient without COVID-19, however, it later spread to 12 patients within a COVID-19 intensive care unit, with three of four intensive care areas affected within three months (82). C. auris was isolated from blood in six patients (6/12; 50%), urine in eight patients (8/12; 66.6%), and from both sites in two patients (2/12; 16.6%). Mortality was high (5/6; 83.3%) amongst patients with candidaemia.

In Peru, two patients with respiratory illness, one with latent TB and one COVID-19 infection, were identified as having C. auris (78). Both had a central venous catheter in-situ, indwelling urinary catheter, and were on mechanical ventilation. In Panama, a significant association with COVID-19 was identified, with 124 isolates of C. auris having been identified prior to the PAHO epidemiological alert, of which 108 were in patients with COVID-19 infection (78). In Columbia, 340 cases of C. auris were identified in 2020, several of which were in patients hospitalised with SARS-CoV-2 (78).

Europe

In Spain, C. auris became the most isolated Candida species from blood cultures after the start of the pandemic, associated with a large outbreak in a tertiary hospital that commenced in 2017 but worsened during the pandemic (83). This was linked with overoccupancy in the intensive care unit, higher workload of healthcare workers, and poor compliance with IPC measures. In Italy, the index case of C. auris was identified in 2019, and a nosocomial outbreak subsequently declared in patients hospitalised in intensive care with COVID-19 infection (76). A single genetic lineage was identified suggesting a point source. There was also a high rate of multi-drug resistant organisms identified in patients with COVID-19 admitted to intensive care, in addition to C. auris. Again, concerns regarding IPC practices were raised and several patients required a high level of care, as well as longer hospital stays, and frequent use of broad-spectrum antibiotics (76). In response to 277 cases associated with this outbreak, spread across at least eight healthcare facilities in the Italian region of Liguria, and 11 cases in facilities in the neighbouring region of Emilia-Romagna, ECDC conducted a rapid risk assessment for the European Union (84).

Asia

In Lebanon, the first cases and subsequent outbreak of C. auris occurred during the midst of the pandemic in October 2020, with a total of 14 cases identified in critical care units over 13 weeks (85). Two patients were identified with C. auris during their stay in COVID intensive care after being admitted through the emergency department, with environmental contamination of the department remaining a possible source.

In India, an outbreak of C. auris was identified in an intensive care unit with COVID-19 patients (74). The outbreak was associated with high mortality, and a had a case fatality rate of 60%. Affected patients had been hospitalised in intensive care for prolonged periods of time (range 20 to 60 days), and all had indwelling invasive devises such as central venous and urinary catheters, and other risk factors such as the need for mechanical ventilation or steroid treatment and underlying chronic diseases including diabetes mellitus and hypertension.

Middle East

COVID-19 and C. auris co-infection was identified in the United Arab Emirates, where it was hypothesised that the increment in fungal infections was the result of acquisition in the hospital setting, and higher susceptibility of the patients, given treatment with broad-spectrum antibiotics and immunosuppressive therapies (86).

An epidemiological analysis of C. auris cases during the pandemic concluded that changes in practice were likely to have been relevant to the spread, including changes in prescribing practices, perceptions of appropriate use of personal protective equipment (PPE) including extended and excessive use, an increase in agency staffing with varied levels of training and experience in PPE use and care of COVID-19 patients (87).

It has been concluded in several regions that the pandemic created an opportunistic environment for spread of C. auris, posing pressures such as overwhelmed intensive care units, suboptimal infection control practices, PPE use, temporary staffing, and inadequate training (85, 87, 88). A review of 36 cases of co-infection with C. auris and COVID-19, found that most cases were males, with an age range of 25–86 years, and candidemia was the predominant presentation (89). Date of onset of infection post admission ranged from 4 to 45 days, and most patients had significant underlying comorbidities. Almost all had an indwelling urinary catheter, intra-venous catheters, were on broad-spectrum antibiotics, and most had received steroids. The mortality rate was 53% of 30 cases with documented outcomes (89).

Question 7

Risk factors

Accepted Answer

Acquisition of C. auris is commonly associated with high-risk healthcare settings and particularly high dependency contexts such as intensive care units, with findings of prospective screening on admission suggesting a high propensity for nosocomial acquisition (69, 90). Patients may be colonised rapidly during their inpatient stay, some within four days (90). A retrospective analysis of 912 cases worldwide found a higher proportion of men (61.4%) were affected (57). A population based study in Spain also showed incidence was higher in autumn and amongst the age group of 65–84 years (91).

Identified risk factors predisposing to C. auris infection are similar to that of other Candida species, being opportunistic pathogens. These factors include severe underlying disease with immunosuppression, such as HIV and bone marrow transplantation, corticosteroid therapy, neutropenia, malignancy, those with chronic kidney disease or diabetes mellitus, a prolonged stay in ICU, mechanical ventilation, presence of a central-venous catheter or urinary catheter, prolonged exposure to broad-spectrum antibiotic or antifungal use, underlying respiratory illness, vascular surgery, or surgery within the previous 30 days (19, 75, 76, 92, 93).

Review of clinical characteristics associated with C. auris found underlying diseases were high, however, kidney disease was the only significant risk factor for mortality in C. auris infected patients (67). Overall mortality with C. auris infection is reported as high in the literature, up to 40-60% worldwide, possibly due in part to severe underlying conditions in the at-risk populations, the multi-drug resistant nature of the pathogen, and limited availability of certain antifungal drugs in some countries (18, 29, 94).

Mechanical ventilation is an important risk factor. The rate of C. auris colonisation in residents of skilled nursing facilities caring for ventilated patients was found to be up to 10 times higher than the occurrence in nursing facilities without ventilator support (95). Additional risk factors for colonisation in this cohort of residents in a ventilator-capable nursing facility include one or more acute care hospital visits in the prior 6 months (adjusted OR 4.2; 95% CI, 1.9–9.6), antibiotic treatment with carbapenems in the prior 90 days (aOR 3.5; 95% CI, 1.6–7.6), and systemic fluconazole in the prior 90 days (aOR 6.0; 95% CI, 1.6–22.6) (95).

C. auris can affect both adult and paediatric populations. Nosocomial infection in children has been reported largely under the age of one year and in those with certain medical conditions including prematurity and malignancy, and in children with risk factors such as intravascular lines and those receiving parenteral nutrition (96). Transmission is also possible from a colonised mother to baby during delivery (95). Clusters of infection have been identified in neonates, with risk of mortality associated with invasive disease (97-99).

Question 8

Identification and typing

Accepted Answer

Identification of C. auris has been challenging using conventional laboratory diagnostic techniques (100). Genetic analyses have established that C. auris is closely related to C. haemulonii and C. lusitaniae (13). Other common misidentifications are described in Table 1. Isolates should therefore be identified to the species level. Options for species identification are available and include matrix-assisted laser desorption/ionisation time-of-flight (MALDI-TOF) mass spectrometry (101-103). MALDI-TOF can reliably differentiate C. auris from other Candida species (104, 105). Accuracy of this system is reliant on the spectra for the sample organism being present in the reference database and care being given to an appropriate extraction method. A novel chromogenic agar, CHROMagarTM Candida Plus, appears promising in the identification and differentiation of C. auris from other Candida species, with C. auris colonies appearing as a light blue colour with a blue halo, and obtaining a sensitivity and specificity of 100% at 36 hours incubation (2, 103, 106, 107). This is suggested as being an excellent alternative to conventional mycological media for the screening of patients who may be colonised or infected with C. auris and can be reliably used to identify this species.

The detection of C. auris has been enhanced by development of molecular tests (108). Turnaround times and diagnostic sensitivity are improved using molecular technologies, particularly for specimens with a high pre-test probability of C. auris colonisation or infection (2). Molecular identification can be performed by sequencing various DNA loci in specific domains of ribosomal genes (18S rDNA, 28S rDNA or internal transcribed spacers ITS1, ITS2), by conventional or real-time polymerase chain reaction (PCR) and loop-mediated isothermal amplification (LAMP) (13, 20).

The identification of C. auris using traditional phenotypic and biochemical methods is challenging, and these have largely been superseded by proteomics and molecular methods (2). Commercial biochemical identification systems commonly used in clinical microbiology laboratories may be unreliable for C. auris identification if their system has not been updated to include C. auris, due to phenotypic similarities with other species (105, 109). Checking the system for C. auris detection ability is therefore required.

Molecular typing of C. auris can be performed using a variety of methods, including sequencing of rDNA loci (D1/D2 or ITS regions) to differentiate between the major phylogeographic clades. Further delineation for outbreak analysis requires higher resolution methods, which can be performed by whole genome sequencing analysis and typing by amplified fragment length polymorphism (AFLP) (2, 13). International networks are being established to improve genomic capacity and develop analysis pipelines that will support ability to detect outbreaks, identify introductions, and characterize transmission of fungal infections (110, 111).

Table 1: Examples of misidentification of C. auris and the corresponding identification method used (table adapted from Elbaradei and others, 2020 (119) and Hardeep and others, 2021 (92)).

Candida auris	Identification method used	Reference
Candida albicans	MicroScan	(112)
	Vitek-MS	(113)
Candida catenulata	BD Phoenix	(105)
	Microscan	(105)
Candida famata	Vitek-2 [Note 1]	(20, 114)
	MicroScan	(105)
Candida guilliermondii	MicroScan	(105)
Candida haemulonii	BD Phoenix	(112) (105)
	Vitek-2 [Note 1]	(14, 20, 104, 105, 114, 115)
	Vitek-MS	(116)
Candida lusitaniae	MicroScan	(105)
	Vitek-MS	(116)
Candida parapsilosis	MicroScan	(105)
	RapID Yeast Plus	(117)
Candida sake	Api 20C AUX	(20, 116)
Candida tropicalis	MicroScan	(112)
Rhodotorula glutinis	Api 20C AUX	(14, 105)
C. duobushaemulonii	Vitek-2 [Note 1]	(114)
Saccharomyces kluyveri	API ID32C	(118)

Note 1. Software upgrade to version 8.01 includes C. auris. It is, however, recommended to confirm isolates identified as C. haemulonii and C. duobushaemulonii, C. famata and C. auris by MALDI-TOF or DNA sequencing.

Question 9

Antifungal resistance

Accepted Answer

C. auris shows high minimum inhibitory concentrations (MICs) for various antifungal agents. Fluconazole resistance is widespread but not universal, however, this is generally not a therapeutic option for most clinical cases (120). Additionally, many C. auris isolates also show high MICs for newer azoles such as voriconazole, likely indicating ineffectiveness of these agents for clinical cases (120). Clinical cut-off values have, however, not been described as yet.

In a collection of C. auris isolates (n = 54) from five continents, 41% were found to be resistant to two or more antifungal classes (64). Resistance to echinocandins remains uncommon; however, with widespread use of the drug as first-line therapy, it has been increasingly identified (121). Pan-resistant strains have also been described (12) and worringly evidence of their transmission between healthcare facilities is emerging (122).

Antifungal drug resistance is thought to be an acquired as well as a shared trait, with the potential to develop over time when fungi are exposed to antifungals (123, 124). The emergence of echinocandin resistance, detected by WGS and phenotypically, within an ongoing prolonged outbreak is concerning (125). C. auris has an intrinsic ability to exhibit or develop resistance very rapidly even while the patient is still undergoing treatment, which is why it is essential to use the antifungals at the right time and at the correct dosage (13). Antifungal susceptibility testing is therefore recommended on all Candida isolates, including of C. auris (126).

Question 10

Colonisation, infection and outcomes

Accepted Answer

The clinical spectrum associated with C. auris ranges from asymptomatic colonisation to invasive candidiasis, most commonly as healthcare–associated candidemia (100). Other healthcare associated infections reported include intravascular catheter infection, urinary tract and respiratory infection, meningitis, osteomyelitis, surgical site infection and otomastoiditis (30, 69, 112, 121, 127). C. auris colonises the axillae, groin, nares, respiratory and urinary tract of hospitalised patients (5, 128). Intestinal colonisation occurs less frequently, but has been associated with C. auris urinary infections (129). Colonisation results in invasive infections in nearly 10% of individuals, with mechanical ventilation and placement of invasive devices identified as two major risk factors (5).

The ability for rapid colonisation of skin and high transmissibility within the health care setting can result in prolonged and serious outbreaks (28, 69). Transmission can occur via contact with contaminated items or from affected individuals. Screening of the patient environment has yielded C. auris isolates with identical fingerprinting patterns, suggesting shedding of C. auris by colonised patients (5). Contaminated equipment identified during outbreak investigations have included thermometers, blood-pressure cuffs, a cloth lanyard and a call bell (130-132). The duration of patient colonisation remains uncertain; however, a recent study demonstrated the majority of patients with C. auris colonisation did not have detectable C. auris 12 months after discharge to the community setting (17).

Invasive disease is associated with high mortality in patients within an intensive care setting (92). Infection is also complicated by multidrug resistance, with bloodstream infections being more difficult to treat and risks of complications including spondylodiscitis, endocarditis and ventriculitis, and clinical syndromes including otomycosis and otomastoiditis (16, 127, 133). Involvement of skin, respiratory, urogenital, and abdominal sites has also been reported (48), including vulvovaginal candidiasis in the context of immunosuppression (134).

Different clades are associated with different clinical presentations and also appear to display differences in virulence in animal models (135, 136). A comparison study of in vivo pathogenicity of the four initial C. auris clades in a neutropenic bloodstream infection murine model found the highest overall mortality was observed for the South American clade (96%), followed by the South Asian (80%), South African (45%) and East Asian (44%) clades (135). The most virulent isolates appear to exhibit pathogenicity comparable to that of C. albicans. In an immunocompetent mouse model, C. auris was demonstrated to be highly virulent, but less virulent than C. albicans (137).

In relation to patterns of infection, South Asian and South American clades are commonly associated with bloodstream infections (47%-76%), whereas the South African clade is associated with a higher percentage of urinary tract infections or colonisation (38%) (138). Isolates from the East Asian clade are associated with otitis externa, which also shows higher genetic diversity compared to other clades and may indicate an older natural population (138). All except for the East Asian clade (and new Iranian clade) have been linked to outbreaks causing invasive infections (92).

Bloodstream infections can be followed by complications such as infective endocarditis and spondylodiscitis. Recurrence has also been reported (27). Case fatality associated with invasive infection is variably reported. In the U.S., among patients with blood isolates of C.auris, 30-day mortality was reported as 39% and 90-day mortality as 58% (139). In a UK outbreak, however, no mortality was directly attributed to C. auris infection (69).

Question 11

Treatment considerations

Accepted Answer

Antifungal agents commonly used for treatment of Candida infection include fluconazole and the echinocandins. C. auris isolates are commonly resistant to fluconazole;, however, resistance to the other antifungal agents is more variable.

Therapeutic recommendations for C. auris include echinocandin monotherapy as empirical treatment prior to results of susceptibility testing, in light of common resistance patterns identified to date (13, 140, 141). Although reports of echinocandin and pan-resistant isolates are increasing, in regions where most strains continue to remain susceptible it is reasonable for echinocandins to remain as first line treatment (141). Patients should, however, be monitored for clinical improvement, with follow-up cultures and susceptibility testing, as the organism can develop resistance quickly including during treatment (142).

Evidence is lacking about the most appropriate therapy for pan-resistant strains, where resistance to all three major classes of antifungals has been identified (echinocandins, amphotericin B and azoles) (9). There is in vivo evidence of inhibition of pan-resistant strains through combinations of two antifungal drugs using fixed concentrations, with an effective response noted from flucytosine combinations with amphotericin B, azoles, or echinocandins (143). There is additional in vitro evidence supporting combination therapy against C. auris with caspofungin in combination with posaconazole (144) or anidulafungin in combination with manogepix or flucytosine (145). A novel echinocandin, rezafungin, also appears promising from in vitro studies, including in subsets of echinocandin-resistant C. auris, and this is currently undergoing phase 3 trials (146, 147). Fosmanogepix, a first-in-class antifungal with a novel mechanism of action available in intravenous and oral formulations, has shown potential both in vitro and in phase 2 studies (148, 149). Both persistent and recurrent C. auris bloodstream infections have also been documented and animal studies and in vitro investigations suggest micafungin-based combination therapies appear promising in this context (140, 150, 151).

Question 12

Infection prevention and control

Accepted Answer

C. auris is able to grow at higher temperatures than other fungi and is able to tolerate high salt concentrations (5). These are important characteristics in its ability to persist in the environment and to survive for long periods of time. This provides opportunities for colonisation and transmission. There are additional challenges associated with rapid acquisition and prolonged colonisation, particularly in nosocomial settings, leading to an increased risk of contamination and transmission for weeks or months following exposure (96, 121).

Whole genome sequencing analyses of isolates from patients and their environment have shown that C. auris can contaminate surfaces at varying distances from the patient’s bed, including items not frequently touched and those further away from the patient (152, 153). Outbreak investigations have isolated the organism from environmental samples, including those taken from a mattress, bedside table, bed rail, chair, window sill, and from the air (69), from bed surfaces and equipment such as ventilators, a temperature probe, ECG leads (90), and pulse oximeters (28), as well as the floor, patient bedside trollies, air conditioners, a bed sheet, pillow, mobile phone, oxygen mask, and an intravenous pole (7). Biomedical products and equipment should therefore be used as disposables, or if re-usable equipment is being used, this should be single patient use ideally and should be left in the patient’s room, to ensure thorough terminal decontamination at the time of discharge and before reuse by another patient. (154).

Infection prevention and control (IPC) measures are crucial to containing transmission. Many of the major world health organisations have published guidance and recommendations regarding the isolation of patients, contact precautions, and cleaning of equipment and environments in contact with C. auris, including the UK, United States, Europe, South Africa, Australia, and Canada. These are listed in the Internationally published guidance and recommendations section.

Identification of C. auris and screening

C. auris can be transmissible whether an individual is infected or colonised, and thus infection control precautions are the same (114). Screening is widely recommended where transmission has occurred, or where there are close contacts of confirmed C. auris cases (2). Active screening of exposed and potentially exposed patients followed by strict infection control measures has been successful in outbreak management (155).

Evidence suggests frequently encountered sites for colonisation and therefore for screening, include the axilla and groin (90, 139). Screening sites also shown to culture positive for C. auris, and may be relevant, include the rectum, pharynx, urine, nose, mouth, ear, catheter-exit site, and wounds (27, 139, 154).

The CDC also currently recommend patient screening for colonisation should be of the groin (bilaterally) and axilla. The Pan American Health Organization/World Health Organization (PAHO/WHO) recommend when C. auris is identified in a healthcare facility, screening patients on the same hospital ward and direct close contacts with samples taken from the axilla, oropharynx, nostrils, groin, urine, and rectum, and if it is not feasible to obtain from all sites, to at least sample from the groin or axilla (78). A recent study has also demonstrated that adding nasal swabs to composite axilla and groin swabs may yield additional positive isolates, with 25% of positive findings being on nasal culture alone (139).

The benefits of routine screening of healthcare workers remain unclear, however screening in outbreak investigation has identified some positive findings (69, 152). C. auris has been found on skin, hands, and nares of care providers and healthcare workers and screening may therefore be initiated if risk factors are identified (2, 156, 157). One outbreak investigation also revealed genetically identical strains to a patient and environment (152).

The duration of colonisation with C. auris remains unknown, although this does appear to be protracted whilst patients remain in healthcare settings (114). Screening methodologies may not identify all sites of colonisation and de-isolation of contacts therefore remains challenging in this setting. An outbreak investigation reported having de-isolated close contacts after three consecutive negative screens, with one contact having subsequently tested positive again on screening continued until discharge (69). In response to high rates of detection in New York, a pilot case management program for people colonised with C. auris who were discharged to a community setting, found that although long-term colonisation was observed amongst some, serial C. auris assessments found that approximately two-thirds of patients colonised as inpatients and discharged to a community setting did not remain colonised indefinitely (158). The time taken for patients to become serially negative was 8.6 months (IQR 5.7–10.8) and for patients who became serially negative, the median time to the first negative was 4.7 months (IQR 3.5–7.5).

Transmission-based precautions

Given the evidence around transmission risk associated with C. auris, patients colonised or infected should be cared for in a single room with contact precautions, preferably with their own bathroom. A flagging system indicating the isolation should also be visible at the entry of the room and patients should have an alert on their medical records to ensure appropriate isolation on transfer and readmission (61).

If single room occupancy is not possible, patients with C. auris should at the very least be cohorted, ideally with single patient use commodes and single use equipment. Healthcare workers and staff should be advised of the importance of, and vigilance with hand hygiene. Hand hygiene washing with soap and water, alcohol-based, or chlorhexidine-based hand rub, have all been shown to be effective in eliminating C. auris from hands (90, 120). Transmission based precautions are also recommended, including personal protective equipment (PPE) and single use items, for the duration of the stay in a healthcare facility (114). PPE including gloves and a long-sleeved gown, is recommended for use in contacts with patients with C. auris or their environment (22, 154, 159). Special precautions are advised when cleaning or exposed to body fluids in C. auris affected areas. The CDC recommend the transmission-based precautions and enhanced barrier precautions are similar to their use for other multidrug-resistant organisms (159). Patients should ideally only be moved for necessary medical procedures and should ideally be last on the list to allow for a thorough clean following, with minimum staff involved (114).

Effective contact precautions are especially important in the context of invasive lines in high risk cohorts, due to the ability of fungal cells to adhere to the device, and their growth occurring in the form of a biofilm (92).

Disinfection of environmental surfaces

C. auris can resist certain disinfectants and is well adapted to nosocomial environments, having the potential to remain viable on vertical surfaces in the immediate environment of colonised patients for up to 28-days (90, 160). Additionally, the organism has been cultured from bedding for up to 7-days (90). C. auris can colonise and persist on surfaces for longer than C. albicans, and has shown a prolonged metabolic activity (161). Furthermore, the minimum time taken to acquire C. auris from a patient or their immediate environment is 4-hours or less, further reinforcing the importance of rigorous IPC measures (69).

Evidence supporting effective products and methods for disinfection of environmental surfaces contaminated by C. auris remains limited due to challenges with comparison of studies, with many applying different experimental techniques, the results of which also cannot readily be compared or directly translated to efficacy in real scenarios.

Chlorine-based disinfectants, in the form of sodium hypochlorite (NaOCl) and sodium dichloroisocyanurate, are commonly used in the healthcare setting for disinfection, especially for multi-drug resistant organisms, and therefore these have been the most investigated in relation to C. auris (162). A chlorine-based disinfectant was noted to be effective during an outbreak in the UK in 2015 (1,000 ppm Chlor-Clean, Guest Medical), having been used in the daily routine disinfection of the patient area and equipment, with a 10,000 ppm chlorine-based product (Haz-Tab, Guest Medical) having been used for terminal cleaning followed by further disinfection with hydrogen peroxide vapour (69). In vitro studies have also investigated efficacy of chlorine-based products. A chlorine-based disinfectant (Haz-Tab 1,000 ppm chlorine) was tested against different clinical isolates of C. auris with an exposure time of 5-minutes and all isolates had at least a 4.5 log10 reduction in growth (163). A further study evaluated chlorine-based products at 1,000 ppm (Chlor-Clean) and 10,000 ppm (Haz-Tab) against clinical isolates of C. auris and other Candida species, finding that C. auris isolates were effectively killed at all concentrations with a minimum of 3-minutes contact time (164).

Studies also conclude that sodium hypochlorite (NaOCl) with concentrations of 1,000 ppm or higher are effective in eradicating C. auris (154, 159, 165). In relation to effectiveness on surfaces, following application of 1 and 2% NaOCl to four different surfaces (stainless steel, ceramic, plastic, and glass) for a 10-min contact time, complete eradication of C. auris was reported on all surfaces (90). A further study then investigated efficacy of NaOCl at 1,000 and 10,000 ppm on cellulose matrix, stainless steel and polyester surfaces inoculated with clinical isolates of C. auris (165). At all concentrations, NaOCl demonstrated significant killing on all substrates at contact times of 5 and 10-minutes, however, of all materials tested complete eradication was achieved only on cellulose substrates. Several commercially available products containing NaOCl have also been found to be ineffective against dry biofilms containing C. auris (166).

The efficacy of peracetic acid has also been tested on stainless steel, polymer, and cellulose surfaces. Peracetic acid at 2,000 ppm was found to have significant killing activity against C. auris, however, like NaOCl, complete eradication was achieved on cellulose matrix but not with steel or plastic (165). As C. auris can survive on plastic surfaces, for efficient removal it has been recommended peracetic acid is added to NaOCl, peracetic acid (3,500 ppm) and sodium dichloroisocyanurate (1,000 ppm) having been effective against dry biofilms containing C. auris (166).

There is some evidence regarding effectiveness of hydrogen peroxide in disinfectant and vaporised form, but with lower levels of supportive evidence (164, 167). Application of hydrogen peroxide vapor appears promising for environmental decontamination in an in vitro study, based on a hospital outbreak (164, 168). This is currently recommended as a potential additional safety measure to manual cleaning and disinfection regimes, rather than replacing other regimes (169). A study of 0.5 and 1.4% hydrogen peroxide solutions were also found to be effective (167).

Quaternary ammonium compounds are commonly used disinfectants in healthcare settings, however, the overall evidence regarding their efficacy for C. auris is conflicting, and their use is therefore currently discouraged (170, 171).

Further studies highlighted that application in accordance with recommended concentrations and contact time were essential in terms of securing efficacy with glutaraldehyde, phenols, hydrogen peroxide, and ethanol (90, 170). Two percent glutaraldehyde and 5% phenol were found to be effective on multiple surfaces when the recommended contact times of 20 and 60 min, respectively, were used (90). Alcohol (29.4% Purell Healthcare Surface Disinfectant) did show some killing activity but not as effective as chlorine-based disinfectants or hydrogen peroxide (167).

The evidence regarding effectiveness of ultraviolet-C (UV-C) light is conflicting (170, 172). A number of UV surface distinction devices have been tested for efficacy against C. auris. These UV systems vary significantly, and each requires individual verification. Reports that found UV to be effective typically quote log reductions between 2.48 and 5.5 for cycle times between 10 and 20 minutes (173). This will vary depending on the UV emitter being tested, the distance between the emitter and the contaminated surface, the presence or absence of additional soil, exposure time, the angle of incident radiation and the degree of shadowing. Other studies have concluded that C. auris is significantly less susceptible to killing by UV-C in comparison than other Candida species (218), with one study stating the C. auris is not effectively killed by the standard UV-C disinfection (219) and two studies highlighting strain variability (206, 217). The CDC currently concludes that research is limited and the parameters for effective disinfection with UV-C are not yet well understood.

The CDC currently recommends use of EPA-registered hospital grade disinfectants that are effective against Clostridium difficile spores; primarily chlorine-based products (159). This has been supported by outbreak management success with stopping transmission (37) and by further investigation of chlorine-based disinfection, concluding effectiveness with appropriate application (163). If EPA-registered hospital grade disinfectants are unavailable, alternatives suggested by the CDC are hydrogen peroxide 0.5 – 1.4%, or quaternary ammonium compounds supplemented with isopropyl alcohol and/ or ethyl alcohol. The ECDC recommends terminal cleaning using disinfectants and methods with certified antifungal activity, including chlorine-based disinfectants (at a concentration of 1 000 ppm), hydrogen-peroxide or others. The Public Health Agency of Canada (PHAC) and South African Centre for Opportunistic, Tropical and Hospital Infections (COTHI) interim recommendations include “regular” and terminal cleanings with a chlorine-releasing agent at 1000 ppm, and COTHI suggests the addition of hydrogen peroxide vapor, when feasible. The Pan American Health Organization/ World Health Organization (PAHO/WHO) recommends cleaning with soap and water followed by disinfection with 0.1% bleach. PAHO/WHO are recommend the following ‘high activity’ compounds; sodium hypochlorite, hydrogen peroxide (and vaporised), peracetic acid and hydrogen peroxide. The Australian Society of Infectious diseases recommends use of products that claim to have sporicidal activity for disinfection (e.g., ≥1000 ppm bleach, peracetic acid or accelerated hydrogen peroxide).

Terminal cleaning and disinfection of the environment remains an essential element of the IPC precautions required to prevent transmission of C. auris from infected or colonised patients. Whilst antimicrobial surfaces and coating will never replace environmental decontamination, this is an area of active research. One example of this is a fast-acting, permanent antimicrobial surface made of compressed sodium chloride (CSC). Pilot data indicates at least 99% reduction of C. auris in one minute, which may be promising but also requires further investigation. There are in vitro data suggesting that C. auris strains can be killed on contact when exposed to caspofungin that is reformulated as a covalently-bound surface layer on glass and plastic (174), however the risk of selecting resistant strains is uncertain and an important concern.

Patient decolonisation

Several studies have investigated products for decolonisation in relation to C. auris, with effectiveness having been demonstrated for chlorhexidine gluconate, povidone-iodine, and alcohol (69, 90, 163). It remains unclear, however, of the benefits in relation to decolonisation or recolonisation risk in relation to C. auris. There are therefore no current protocols for decolonisation of patients with C. auris and recommendations regarding decolonisation were unable to have been made by an international expert working group due to a lack of evidence (154).

Chlorhexidine gluconate is widely used in healthcare for prevention. This includes in handwashing, pre-procedure skin preparation, indwelling catheter exit-site care, oral care for prevention of ventilator-associated pneumonia, and whole-body bathing (175). The evidence would suggest chlorhexidine gluconate is of benefit with decolonisation of Candida species, although there is little evidence specific to C. auris (162). One study demonstrated that chlorhexidine gluconate (<0.02% with a contact time of 24-hours), was effective in inhibiting growth of planktonic cells and biofilms of clinical isolates of C. auris (176). With concentrations of between 0.125% and 1.5%, and a 3-minute contact time, growth of C. auris was inhibited and increased efficacy seen at 3 and 30-hours (164). C. auris had consistently higher minimal inhibition concentrations (MIC) however when compared to other Candida species tested. A study of 2% chlorhexidine gluconate used on its own (i.e., 1:1 dilution of the antimicrobial hand and body wash) showed failure to eliminate C. auris at a contact time of two minutes. The addition of 70% isopropyl alcohol (IPA), however, did reduce all six strains of C. auris to undetectable levels within 2 minutes, suggesting that a chlorhexidine/IPA disinfectant could reduce colonisation if applied appropriately (163). This may however cause skin damage, and further evidence would be helpful.

The experience of several outbreaks has been that patients can remain colonised despite daily chlorhexidine washes (69, 90). Decolonisation may, however, have the potential to reduce propensity of C. auris on the skin surface of affected individuals, thus potentially reducing the risk of transmission (69, 121). The evidence suggests challenges result from persistent colonisation. It remains uncertain if this relates to reinfection from the environment, or reduced propensity to chlorhexidine. In one study, the two patients with persistent colonisation in the groin, suffered from diarrhoea and it was suggested that this possibly resulted in the persistence (90).

In a facility where C. auris was endemic, integrating microbial-genomic and epidemiologic data revealed occult C. auris colonisation of multiple body sites not commonly targeted for screening (177). Here, high concentrations of chlorhexidine were associated with suppression of C. auris growth but not with deleterious perturbation of commensal microbes. A murine model study observed that C. auris can enter the dermis without causing overt histopathologic signs of inflammation, expanding into deeper tissues, which could make decolonisation challenging (178). This could also explain the re-occurrence of C. auris in patients who had repeated interval negative swabs, and interestingly chlorhexidine antiseptic protected against colonisation of C. auris on mouse skin.

A study investigating the in vitro yeasticidal activity of povidone-iodine against C. auris, compared to C. albicans and C. glabrata, and according to the EN standard, were encouraging. The growth of all clinical C. auris isolates was inhibited at concentrations between 0.07 and 1.25%, which is below many of the commercially available concentration of 10%, with a minimum contact time of 3 minutes (164).

A murine skin colonization model was used to test fungal burden reduction following treatment with 1% terbinafine or 1% clotrimazole in a proprietary Advanced Penetration Technology formulation. This found both treatments significantly reduced fungal burden compared to that in control groups (179). Compounds with antimicrobial activity have been assessed for efficacy in antifungal decolonisation, including triclosan, boric acid and zinc oxide, which can be used for long periods of time without an abrasive skin effect. Antifungal activity of boric acid and triclosan was demonstrated against multiple Candida species, including a clade of C. auris (180).

Question 13

Discussion

Accepted Answer

C. auris is increasingly detected at widespread geographical locations, and prevalence is likely to be greater than what is currently known. Detection of the organism has improved in many countries known to be affected by C. auris. The organism should be considered when unidentified or unusual Candida species are isolated from patients who fail to respond to empiric antifungal therapy.

Colonisation and infection have been primarily detected in high-dependency settings to date, and important risk factors include invasive lines and mechanical ventilation. The COVID-19 pandemic therefore had an extremely detrimental impact in several countries worldwide, with widespread transmission and C. auris co-infection frequently associated with overcrowding, inadequate PPE use, and suboptimal IPC practice. This was also associated with high mortality, however, that directly attributed to C. auris cannot be established. Establishing prevalence outside of high-dependency settings has been a challenge, and an area that will likely require development being of importance to screening and control strategies.

It is clearly challenging to prevent transmission following identification of someone colonised with C. auris, even whilst employing strict IPC and isolation techniques, but control has been demonstrated (155, 181, 182).

Although treatment of asymptomatic colonisation is not currently widely recommended in the literature, this requires further review. Potential benefit to individuals and implications for wider transmission has been suggested; however, efficacy appears to be uncertain. The duration patients can remain colonised, and thus how long patients should be isolated after first detection, is also uncertain. Continuing isolation for the duration of an inpatient stay has been recommended given the potential implications of transmission. Despite the CDC currently recommending deisolation following two consecutive screening swabs, in reality, this criterion is reported as being uncommonly met and recolonisation is possible, potentially from sources of environmental contamination.

Antifungal susceptibility testing should be performed on all isolates, and empiric treatment with an echinocandin is still recommended whilst awaiting results. The development of increasingly resistant strains is, however, a concern. The future may be of combination therapies, novel, and alternative treatments, such as natural compounds (e.g., rocaglates), photodynamic therapy, and novel triazoles (e.g., PC945). Antimicrobial stewardship also plays a key role in limiting unnecessary antibiotic and antifungal use and ensuring appropriate management of cases. The risk-benefits of antifungal prophylaxis should be weighed up with every case and decisions based on local drug susceptibility patterns.

Multidisciplinary response is needed to control spread on C. auris in a healthcare setting including outbreak preparedness and response, rapid contact tracing and isolation, cohorting of patients and staff, hand-hygiene, and strict IPC, dedicated or single use equipment, appropriate disinfection, transfers, and discharge.

Question 14

Internationally published guidance and recommendations

Accepted Answer

This is an overview of the guidance and recommendations published by major world health organisations regarding the isolation of patients, contact precautions, and cleaning of equipment and environments in contact with C. auris.

UKHSA

(183)

Patient screening

Recommended in units with ongoing cases or colonisations; those arriving from affected units (UK and abroad). Screening sites such as groin, axilla, nose, throat, urine, perineal area, rectal area, and stool. Also consider screening, if indicated, LVS, sputum, endotracheal secretions, drain fluid, wounds, and cannula. Rescreening of patients known to have been previously colonized. Deisolation of screen-positive patients is not recommended apart from units with experience in managing C. auris

Contact precautions

Side room with en suite facilities where possible; isolation of all patients from affected UK or international hospital until screening is available; strict adherence to hand hygiene using soap and water, followed by alcohol rub to dry hands; PPE with gloves and aprons or gowns if there is a high risk of body or body fluid contact; briefing of visitors regarding contact precautions; single-patient-use items such as blood pressure cuffs should be considered; for cleaning C. auris-exposed areas, glove and apron use with subsequent appropriate hand decontamination

Contact screening

If there is novel detection in a unit, close contacts should be screened and isolated or cohorted; if the index patient is isolated, identify all Candida species isolates from the same unit to the species level using a method able to detect C. auris; review Candida spp. detected in the same ward areas in the 4 weeks prior to diagnosis of the index patient in case of unrecognized transmission; deisolation with 3 negative screens >24 h apart

Decolonisation

Strict adherence to central and peripheral catheter care bundles, urinary catheter care bundle, care of the tracheostomy site; skin decontamination with chlorhexidine washes in critically ill patients; consider use of mouth gargles with chlorhexidine and use of topical nystatin and terbinafine for topical management of key sites

Environmental management

Contact precautions

Single room with en suite or cohorting of patients; hand hygiene using soap and water or alcohol rub; gloves and aprons for patient contact; adherence to venous and urinary catheter and tracheostomy care bundles; advise visitors regarding hand hygiene and contact precautions.

Environmental management

Schedule affected patients last for theatre/ procedures/ imaging; regular cleaning and disinfection with chlorine-releasing agent at 1,000 ppm; terminal cleaning and disinfection of bed space; consider terminal cleaning with hydrogen peroxide vapor; clean multiuse equipment thoroughly; cleaning of all contact areas.

Community management

No specific recommendations.

Australasian Society for Infectious Diseases

(61)

Patient screening

Close contacts, patients transferred from facilities with endemic C. auris or admitted following stay in overseas healthcare institutions should be pre-emptively isolated and screened for colonisation. Composite swabs of the axilla and groin should be collected.

Contact precautions

All patients colonised or infected with Candida auris should be placed in a single room (with dedicated bathroom or commode or pans) and managed using Standard and Contact Precautions for the entire hospital stay and all subsequent hospital admissions.

Detection of C. auris in a non-isolated patient should trigger screening of close contacts, including those sharing the room or equipment (if uncertain screening all close contacts since admission of the case should be considered). Include all current ward mates if detection in a non-isolated patient during an intensive care stay or if secondary case. Healthcare worker screening not recommended unless substantial evidence as potential source or ongoing transmission.

Decolonisation

No recommendations.

Environmental management

For small surfaces, use 70% ethanol or alcohol based disinfectants as per manufacturer directions. For alcohol sensitive or large surfaces, use disinfectant compound with fungicidal activity. Twice daily.

Community management

No specific recommendations.

Question 15

References

Accepted Answer

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Prepared by: Claire Neill and Christopher Jones; with Colin Brown, Rebecca Guy, Andrew Borman, Elizabeth Johnston, Bharat Patel, Katie Jeffery, Surabhi Taori, Ginny Moore, Lesley Price, Martina Cummins, Mariyam Mirfenderesky, Karren Staniforth, Carol Fry.

Acknowledgements: The authors gratefully acknowledge the expert review and advice received from colleagues in the UK Health Security Agency, NHS, and health protection teams.

This document was approved and signed off by the UK Health Security Agency HCAI and AMR Division.

For queries relating to this document, please contact candidaauris@ukhsa.gov.uk

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Introduction

Methods

Background

Genomic analysis and epidemiology

Experience in the UK

C. auris and COVID-19

The Americas

Europe

Asia

Middle East

Risk factors

Identification and typing

Table 1: Examples of misidentification of C. auris and the corresponding identification method used (table adapted from Elbaradei and others, 2020 (119) and Hardeep and others, 2021 (92)).

Antifungal resistance

Colonisation, infection and outcomes

Treatment considerations

Infection prevention and control

Identification of C. auris and screening

Transmission-based precautions

Disinfection of environmental surfaces

Patient decolonisation

Discussion

Internationally published guidance and recommendations

UKHSA

Patient screening

Contact precautions

Contact screening

Decolonisation

Environmental management

Community management

CDC

Patient screening

Contact precautions

Contact screening

ECDC

Patient screening

Contact precautions

Contact screening

Decolonisation

Environmental management

Community management

South Africa (COTHI)

Patient screening

Contact precautions

Environmental management

Community management

Australasian Society for Infectious Diseases

Patient screening

Contact precautions

Contact screening

Decolonisation

Environmental management

Community management

WHO / PAHO

Patient screening

Contact precautions

Contact screening

Decolonisation

Environmental management

Community management

South Africa (FIDSSA guideline)

Patient screening

Contact precautions

Contact screening

Decolonisation

Environmental management

Community management

Public Health Ontario

Patient screening

Contact precautions

Contact screening

Decolonisation

Environmental management

Community management

Recommended actions in Switzerland

Patient screening

Contact precautions

Contact screening

Decolonisation