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Candida auris: laboratory investigation, management and infection prevention and control (draft)

Published 14 June 2023

1. Introduction

Candida auris is an emerging fungal pathogen, which was first identified in 2009 from the external auditory canal of a patient in Japan. It has since been identified across 6 continents, in more than 40 countries worldwide. C. auris infections have been frequently reported from the bloodstream and also seen in association with bone, cerebrospinal fluid (CSF), and intra-abdominal infections. Additionally, C. auris has been isolated from wounds, ear, and respiratory specimens, urine, bile, and jejunal biopsies. Detection in surveillance swabs from the axilla and groin may indicate carriage rather than infection. This, however, poses a risk for invasive infection and transmission to others.

Five genetically distant clades of C. auris have been discovered to date, including:

• 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)
• a further clade that has recently been detected in Iran (clade V)

Each clade is associated with certain clinical presentations, resistance patterns, and differences in virulence.

C. auris can affect both adult and paediatric populations. This is largely within high-risk healthcare settings, especially high-dependency and intensive care units. Findings of prospective screening on admission suggests a high propensity for nosocomial acquisition. Additionally, there have been reports of detection within long-term care facilities in the US, with colonisation rates found to be highest in long-term nursing facilities providing ventilator support. As with other organisms associated with nosocomial outbreaks, C. auris appears to be easily transmitted between patients, and the clinical environment, including via multi-use equipment, highlighting the importance of prompt, effective, and sustained infection prevention and control (IPC) practices.

Possible risk factors for developing C. auris colonisation/infection include severe underlying disease with immunosuppression, such as HIV and bone marrow transplantation, corticosteroid therapy, neutropenia, malignancy, chronic kidney disease or diabetes mellitus, a prolonged stay in intensive care, mechanical ventilation, presence of a central-venous catheter or urinary catheter, extra-ventricular CSF drainage device, prolonged exposure to broad-spectrum antibiotic or antifungal use, and underlying respiratory illness, recent vascular surgery, or surgery within 30 days. Review of clinical characteristics in C. auris infections found underlying diseases were common; however, kidney disease was found to be the only significant risk factor associated with mortality in C. auris infected patients.

Mortality associated with invasive C. auris is variably reported, but may be up to 40 to 60%. This is likely, in part, due to severe underlying conditions in at-risk populations, with attributable mortality being difficult to determine. Additionally, there is variability between different countries, which may be associated with differences in epidemiology and clade, multi-drug resistance, and the limited availability of certain antifungal drugs in some countries. In the UK, no fatalities within related outbreaks have been attributed to C. auris infection to date.

A retrospective analysis of historical clinical and culture collection isolates in the UK showed C. auris was first identified from blood cultures in unrelated patients in 2013. There have since been sporadic introductions into several hospitals in England. Many new detections have involved patients repatriated from international hospitals including from India, United Arab Emirates, Qatar, Kuwait, Oman, Pakistan, and Kenya.

Between 2013 and 2021, a total of 304 C. auris isolates were reported through laboratory surveillance in England, with 40 (13.2%) isolated from blood culture specimens. In 2020, however, there were just 4 cases detected, all colonisations. This is likely a reflection of the extreme travel restrictions associated with the COVID-19 pandemic, limiting international introductions. There was a small increase in cases in 2021; 9 were reported in total and 5 of these were isolated from blood culture specimens.

There have been several international patient transfers in the independent sector and recently C. auris colonisation and infection has been detected in some of these individuals. One UK secondary care provider that offers a screening programme to international patients (admitted directly from home or healthcare facilities) reports that, of 3,404 patients screened (groin, axilla, nose) for asymptomatic C. auris carriage between January 2020 and February 2023, 56 (1.6%) screened positive. The majority of these positive patients with confirmed last location (25 out of 56) were direct healthcare facility transfers from the Middle East, notably Kuwait, Qatar and the United Arab Emirates. However, this may be an under representation as 18 out of 56 patients could not have their last known location confirmed. Patients from Nigeria, South Africa and France have screened positive following admission. UK residents repatriated from hospitals abroad have also been identified on admission screening as C. auris positive.

Three large nosocomial outbreaks that occurred in intensive care units of major tertiary referral centres in England between 2015 and 2017 are responsible for the majority of UK isolates to date, with further spread to other hospitals occurring via repatriation routes. Repatriation and entry to UK hospitals for medical care continues to be a source for new introductions of C. auris in England with potential to spread more widely.

All isolates identified in the UK to date have demonstrated reduced susceptibility to the antifungal agent fluconazole, and variable susceptibility to other antifungal agents. No pan-resistant isolates have been detected yet.

This guidance is relevant to the laboratory investigation, management, and IPC practices associated with C. auris in a healthcare setting. This supersedes the previous guidance, released in 2017. The guidance has been updated in keeping with latest available evidence from published literature, international best practice, and expert consensus, which was considered alongside national surveillance.

2. Investigation in clinical laboratories

Identification of C. auris has traditionally been challenging using conventional laboratory diagnostic methods. C. auris, on microscopy, is indistinguishable from most other Candida species. It is a germ tube test negative budding yeast; however, some strains can form rudimentary pseudo hyphae on cornmeal agar.

For screening and clinical samples, mycological culture remains a principal mode of laboratory diagnosis, with polymerase chain reaction (PCR) on direct samples being used for screening in some centres in the UK. Diagnostic accuracy has improved following the development of chromogenic medium specifically for C. auris, and advances in spectral databases of matrix-associated laser desorption ionisation time-of-flight mass spectrometry (MALDI-TOF MS) systems. Turnaround times and diagnostic sensitivity may be improved using molecular technologies, particularly for specimens with a high pre-test probability of C. auris colonisation or infection.

Isolates of Candida spp from invasive sites should be identified to the species level, as well as any Candida isolates from superficial sites in patients from high intensity or augmented care settings, those transferred from an affected hospital (UK or abroad), or those with an overnight stay in a healthcare facility in another country within the previous year (particularly if the country has had documented cases). If suspected Candida spp are identified, further work should be undertaken to assess for C. auris. This could involve either molecular sequencing of the D1/D2 domain or MALDI-TOF-MS analysis with C. auris either already present or added to the database. This is available at the UK Health Security Agency (UKHSA) Mycology Reference Laboratory, if required. Please send pure isolates on Sabouraud’s slopes accompanied by the appropriate testing request form.

2.1 Culture-based identification

C. auris can grow at temperatures of up to 42 to 45 degrees Celsius (°C), with the optimal incubation temperature described as being around 40°C. This may be useful in distinguishing C. auris from many other Candida species, especially those for which it is most commonly mis-identified as, such as Candida haemulonii. Most C. auris isolates are a pale purple or pink colour on the chromogenic agar, CHROMagar™ Candida, in common with several other non-C. albicans species. A new chromogenic agar, CHROMagar™ Candida Plus has, however, been developed on which C. auris grows as a blue colony with a blue halo and can be readily distinguished from a wide variety of other related and non-related yeast species. This may be particularly useful as a screening tool to identify suspicious colonies from mixed cultures, including the presence of C. albicans, and for differentiating from C. haemulonii. If there is evidence of non–C. albicans species on non-discriminatory chromogenic agar these should be sub-cultured onto Sabouraud’s agar and identified according to local laboratory protocols.

2.2 Biochemical identification

Laboratories are advised to check their databases for currently available biochemical based tests that include C. auris. If C. auris is not present within the database, there is a risk of misclassification of C. auris as a range of other Candida species and genera using biochemical based tests (most commonly as Candida haemulonii, Candida famata, Candida lusitaniae, Rhodotorula glutinis or Saccharomyces cerevisiae).

2.3 Molecular identification

Molecular technologies to identify C. auris may be employed in laboratories as an adjunct to culture-based techniques. It is unlikely that diagnostic laboratories in the UK would routinely use these kits, however, they may be employed in circumstances such as an outbreak scenario. The advantage of direct molecular techniques for screening is that they do not depend on culture, thus are quicker, and are rapidly scalable in case transmission is detected.

Laboratories should also ensure correct mapping of the species code for C. auris to facilitate reporting to UKHSA through the Second Generation Surveillance System (SGSS).

2.4 Antifungal susceptibility testing

There are no established minimum inhibitory concentration (MIC) breakpoints at present for C. auris. The high proportion of isolates with likely acquired resistance has further complicated this. Established breakpoints for Candida albicans are currently being tentatively applied for the interpretation of antifungal susceptibility testing of C. auris. Most C. auris isolates described worldwide have been resistant to fluconazole, subject to the use of different tentative breakpoints. Multidrug-resistance has been demonstrated at variable rates, including to other azoles, amphotericin B, and echinocandins. MIC distribution does, however, currently vary with geography and by clade.

Using tentative breakpoints suggested by the Centers for Disease Prevention and Control (CDC) in the US, shows around 90% of isolates are resistant to fluconazole, 30% to amphotericin B, and less than 5% to echinocandins. The corresponding values for isolates from India are reported as being 90 to 95%, 7 to 37%, and less than 2% respectively. Some isolates have also been identified as resistant to all 3 classes of antifungal drugs. Although resistance to echinocandins remains uncommon, with the use of the drug as first-line therapy, it has been increasingly identified.

Experience to date from the UKHSA Mycology Reference Laboratory indicates that so far very few multi-drug resistant strains have been found in the UK. All isolates have been resistant to fluconazole and often cross-resistant to other azoles, with variable resistance to polyenes (approximately 20% for amphotericin B) and echinocandins (approximately 10%). Development of resistance to various antifungals has been observed in previously susceptible isolates.

Antifungal susceptibility testing is recommended on all isolates of C. auris. Unexpected or unusual susceptibility profiles should be confirmed by a reference laboratory. The National UKHSA Mycology Reference Laboratory is also available to support typing of isolates, should this be required.

3. Treatment

Considering antifungal resistance patterns identified to date, first-line therapy remains an echinocandin, pending specific susceptibility testing, which should be undertaken as soon as possible. Patients should also be monitored for clinical improvement, with follow-up cultures and susceptibility testing, as there is evidence that the organism can develop resistance quickly. Treatment with fluconazole is not recommended for C. auris infection.

Treatment of C. auris identified from sites such as the respiratory tract, urine, or skin, is not recommended in the absence of signs of clinical disease. Combination therapy in bloodstream infections with this organism is not currently supported by existing evidence, although if the urinary tract or central nervous system (CNS) is involved, dual therapy may be necessary, as some antifungal classes do not have bioavailability in either urine or CNS. The evidence is also currently lacking in relation to treatment of persistent and recurrent infection. Animal studies and in vitro investigations would suggest echinocandin-based combination therapies appear promising in this context. This is also the suggestion from in vivo investigations of treatment for pan-resistant strains. Clinicians are advised to make decisions on a case-by-case basis depending on the site of infection.

The UKHSA Mycology Reference Laboratory can undertake susceptibility testing for amphotericin B, fluconazole, voriconazole, itraconazole, posaconazole, isavuconazole, anidulafungin, caspofungin, and micafungin. If an isolate is found to be resistant to all these agents, the reference laboratory will also test for susceptibility to flucytosine, nystatin and terbinafine. Currently, UK strains remain susceptible to the topical agents nystatin and terbinafine and it is possible that for the treatment of any future multi-drug resistant strains a regimen incorporating oral terbinafine could be considered.

3.1 Colonisation

Sites that can be commonly colonised by C. auris in hospitalised patients include the axillae, groin, nares, rectum, respiratory, and urinary tract. Contact with contaminated items appears to be a common source of colonisation. In the UK, colonisation of C.auris positive patients has been observed to be sustained for long periods including post discharge from intensive therapy unit (ITU).

The CDC reports cases of colonisation for more than a year and suggests colonisation may remain indefinitely even after treatment for invasive disease. The literature reports that among patients who had a positive C. auris screening result followed by one or more negative screening results, more than 50% had a subsequent positive screening result. UK experience would support the perception that routine screening of previously positive inpatients produces unreliable, intermittent negative screens. It is recommended that once positive, inpatients are considered C.auris positive for the duration of their care. Repeat screening of these patients is therefore probably only indicated in specific cases, for example to assess effectiveness of skin decolonisation regimes.

There are currently no established routine protocols for decolonisation of patients with C. auris. Clinical experience to date has shown that colonisation may persist despite a range of approaches and is difficult to eradicate, making IPC strategies particularly important. Persistence may also be associated with recolonisation from the environment.

According to in vitro evidence, C.auris is susceptible to chlorhexidine. Although the evidence base does not support routine decolonisation, experience indicates skin decontamination with chlorhexidine wash, mouth gargles with chlorhexidine, or chlorhexidine-impregnated pads for catheter exit sites could be considered on a case-by-case basis, such as for certain high-risk procedures or patient groups, or in settings where there is ongoing transmission despite IPC measures.

It is recommended that strategies to prevent and/or treat colonisation include:

• strict adherence to central and peripheral catheter care bundles, urinary catheter care bundle and care of the tracheostomy site
• prompt removal of venous cannulas if there is any sign of infection
• high standards of aseptic technique when undertaking wound care or invasive procedures
• consideration of skin decontamination with chlorhexidine wipes or washes in critically ill or other high-risk patients, particularly where there is ongoing transmission despite other infection control measures and interventions

There is limited evidence for use of topical nystatin and terbinafine in decolonisation; however, this may be considered for targeted management of key sites such as venous cannula entry sites. Additionally, there is limited in vitro evidence that shorter contact times with chlorhexidine (without alcohol) may not be as effective as povo-iodine based topical applications in reducing C. auris colonisation – this may also be considered when performing invasive procedures such as line insertions or surgical procedures in colonised patients. Contact time as advised by the manufacturer must be followed to optimise efficacy.

3.2 Screening policies

All trusts are encouraged to develop a screening policy after local risk assessments are undertaken, considering patient population case-mix and prior prevalence of C. auris.

3.2.1. Patient screening

Screening is advised for patients coming from other affected hospitals or units in the UK and abroad. Patients who have had an overnight stay in a healthcare facility outside of the UK in the past year should also be screened, particularly if coming from a country with documented cases. Tracking of cases was recorded by the CDC until February 2021; however, the map is not currently being updated, given how widespread C. auris has become. The CDC and European Centre for Disease Prevention and Control (ECDC) provide alerts to C. auris outbreaks, which may be helpful to provide context and inform risk assessments.

3.2.2. Contact screening

Contact screening for C. auris is recommended in a setting where a new infected or colonised patient has been detected, and in units that have ongoing cases and/or colonisations.

Recommendations are as follows.

Any novel detection in a trust should be an indicator to screen close contacts (see following points for definition).

If detected in an intensive care setting (or other vulnerable patient setting), a one-off point prevalence screening survey of the entire unit or ward is recommended.

If detected on a general ward, close contacts should be screened (for example patients from the same bay, and where there have been concerns about patient movement). This should include any roommates, or those within the same bay, within the previous month, if they remain hospitalised or have been transferred to another care setting, such as a care home. Where there is evidence of ongoing transmission, the entire ward area should then be screened.

More extensive contact tracing and screening may be considered on a case-by-case basis.

Consider screening historical contacts (for example a month if unexpected high prevalence) based on findings of point prevalence screening or vulnerability of patient group (such as renal dialysis).

If the patient has been isolated during admission on a ward other than an intensive care setting, trusts are advised to identify all Candida isolates from the same unit to the species level using an appropriate method that will detect C. auris for the subsequent 4 weeks.

If the index patient was not isolated, contacts who have been in the same bay or room as an affected patient in the 48 hours prior to first identification should be isolated or cohorted with other contacts and cared for with enhanced IPC measures, as detailed below for cases. Close contacts can be deisolated after 3 consecutive negative screens at least 24 hours apart.

In all cases, for the 4 weeks prior to diagnosis in the index patient, hospitals should look back to see if there has been an increase in the detection of Candida spp in the same intensive care setting or ward, as this may represent unrecognised transmission.

Suggested screening sites, based on the predilection of Candida spp to colonise the skin and mucosal surfaces (such as genitourinary tract, gastrointestinal, mouth and respiratory tract), are:

• groin and axilla (these sites should be screened as a minimum, based on experience of positivity from the literature)
• nose (as experience of nasal swabbing in addition to axilla and groin swabbing has yielded additional findings, this should also be considered for routine screening)

Also consider screening the following sites (if clinically indicated or previously positive):

• urine (especially if there is a urinary catheter in-situ)
• throat swab
• perineal swab
• rectal swab or stool sample
• low vaginal swab
• sputum / endotracheal secretions
• drain fluid (abdominal/pelvic/mediastinal)
• cannula entry sites
• wounds

Rectal swabs have been shown to be intermittently positive. These may be more useful to detect incident colonisation rather than transmission in hospital environments, but the role of gastrointestinal carriage is as yet unclear. Routine wound swabs may be used to collect screening samples, in addition to the other sites.

3.2.3. Screen positive patients

All screen positive patients should be isolated or cohorted as described below. There is currently no evidence to support the deisolation of patients found to be colonised or infected with C. auris during their hospital stay, as carriage appears to be protracted in this context. As there is also clinical experience of recurrence of colonisation, the need for ongoing vigilance in the form of weekly (or more frequent if highly vulnerable population) screens in clinical environments where C. auris colonised patients have been managed, should be considered by performing local risk assessments.

For those colonised as an inpatient, clearance may occur following discharge from hospital. It has been demonstrated that many carriers do not remain positive for C. auris indefinitely after discharge to the community, although continuous carriage for more than a year after initial isolation of C. auris has also been documented. Isolation and rescreening of patients known to have been previously colonised with C. auris is therefore recommended on readmission. Due to the uncertainty about how long people may remain colonised for, a precautionary approach is to remain isolated during readmission. If bed pressures prevent isolation, regular screening for re-emergence may allow deisolation while managing risk of transmission.

3.2.4. Healthcare worker screening

There is insufficient evidence to recommend routine screening of healthcare workers. This may be considered on a case-by-case basis in outbreak settings by recommendation of the incident management team, for example, if epidemiological investigations suggest a healthcare worker as a source.

All newly positive screens or clinical samples from patients unknown to be colonised should be reported to the local UKHSA health protection team (HPT). The HPT will follow their standardised operating procedure (SOP) and support the public health action associated with any new positive detections.

4. Infection, prevention, and control

C. auris represents a significant burden of disease in certain countries. Healthcare outbreaks have been reported in many countries including the UK, South Africa, India, Kuwait, Spain, the US, Columbia, Kenya, Oman, Pakistan, Venezuela, Brazil, Qatar, and Turkey.

C. auris can grow at higher temperatures than many other fungi and is able to tolerate high salt concentrations. These are important characteristics in its ability to persist in the environment and survive for prolonged periods of time. Acquisition can be rapid (as little as 4 hours from initial exposure to colonisation in affected units), and colonisation can be prolonged (weeks or months from acquisition). Environmental contamination is extensive, and survival times of 7 days have been recorded on general surfaces, increasing to 14 days for some plastic devices.

Environmental screening of occupied patient environments yields C. auris isolates with identical fingerprinting patterns, suggesting shedding of C. auris by colonised patients. Experience during outbreaks supports the hypothesis that environmental contamination with C. auris is significant with C. auris having been detected on beds, and bedside equipment including mattresses, bed sheets, air conditioning units, radiators, windowsills, sink drains, bedside tables, and equipment such as ventilators, skin-surface temperature probes, blood pressure cuff, ECG leads, stethoscope, and pulse oximeters.

Infection control measures are crucial to containing transmission of C. auris. Healthcare staff should work in multi-disciplinary teams, which include Clinical Infection Specialists and IPC teams to risk assess and support the management of patients colonised with C. auris. Where possible, equipment used for the infected/colonised patient should not be shared with other patients on the ward, unless effective decontamination can be assured between each patient use. Single-use equipment where feasible is recommended.

Hospitals must ensure that the bed space requirements between patients comply with the Health Building Note regulations to minimise the likelihood of transmission, with sufficient ensuite single rooms to segregate patients identified as infected or colonised. While direct fomite transmission is a particular risk for nosocomial transmission of C. auris, this does not preclude concurrent transmission via contact with healthcare workers. Adherence to hand hygiene needs to be consistently high and sustained. Alcohol based hand rubs (ABHR) are recommended when hands are not visibly soiled. Hands should always be washed with water and soap on removal of personal protective equipment (PPE) including gloves.

4.1 Detection of C. auris in a patient

Main IPC measures are detailed below.

All patients colonised or infected with the organism should be isolated in a single room, with ensuite facilities, wherever possible, or with their own commode as a minimum (where required, for example, this may not be relevant for patients in intensive care).

If many patients are colonised, cohorting groups of patients may be considered, as appropriate.

All patients who have been transferred in, or from, an affected UK hospital or a hospital abroad should be isolated until screening results are available.

Standard precautions including hand hygiene should be strictly adhered to. Soap and water should always be used where hands are visibly soiled; however, alcohol-based hand rub is effective against C. auris and can be used on visibly clean hands.

Transmission-based precautions should be implemented, as applicable, to other multi-drug resistant organisms.

Effective contact precautions are especially important in the context of invasive devices, and introduction of chlorhexidine impregnated protective disks for long intravenous lines may be useful in preventing invasive infection.

Personal protective equipment in the form of gloves and aprons are to be worn during contact with the patient or environment, or long-sleeved gowns where there is a risk of extensive splashing of blood and/or other body fluids (for example excessive wound exudate, diarrhoea, faecal incontinence, likely physical contact with patient’s skin, or for any rigorous activity involving the patient or such as bed changing). These should be donned after hand hygiene and before entering the room or patient area, and removed and discarded in the room or patient environment followed by a thorough hand wash or application of alcohol hand rub on dry hands before exit, as appropriate.

Visors and/or masks are not routinely required but should be worn if there is a procedural risk of spillage or splashes.

Caps may also be considered where skin squames are expected to be dispersed through activities involving the patient or their environment.

Patients should be fully briefed on their condition and advised as to how to prevent onward transmission and how to prevent entry of the organism into wounds and entry sites. English may not be a first or familiar language, therefore patient information should be made accessible to the patient in the preferred means.

Visitors of infected or colonised patients should be informed about the required IPC precautions; including the need for robust hand hygiene and use of protective aprons or gowns.

Single-patient use items are advised, such as temperature probes and blood pressure cuffs, especially in outbreak situations. Stock of single-use and disposable equipment brought into the infected patient’s room should be limited as any unused disposable item will need to be disposed of on discharge.

Reusable patient care equipment brought into the room of the infected patient must be thoroughly cleaned in accordance with manufacturers’ instructions before using on another patient. Pillows and mattresses should have an entirely impervious plastic cover that is decontaminated between patients. If an item cannot be effectively decontaminated, the cover of any impermeable item of patient care equipment is damaged or there is obvious strikethrough, then the item should be appropriately sealed and disposed of. Consideration should be given to the possibility of capacity for thorough decontamination versus replacement, for example for pillows whether surfaces are smooth or ridged.

Patients should ideally only be moved for necessary medical procedures to reduce the risk of environmental contamination elsewhere in the facility. Where portable imaging is available for example, this should be facilitated in the patient’s isolation room and equipment carefully cleaned on leaving the room. Required visits to other departments must be planned and communicated to the department to allow for thorough cleaning and decontamination to take place once the patient has left.

For surgical procedures, strict adherence to care bundles including skin decolonisation processes using an alcohol-based product is critical to reduce the risk of invasive C. auris infection. Placing the C. auris positive patient last on the list is recommended where feasible to enable thorough cleaning to be applied after the episode of care. Further reference should be made to the NICE guidelines on the prevention and treatment of surgical site infections.

Consider cohorting of dedicated staff caring for patients affected by C. auris, where possible, and particularly in an outbreak situation.

4.2 Environmental cleaning and decontamination

Increased frequency (at least daily) cleaning and disinfection of all surfaces and touch points of patient rooms and other areas where the patient receives care (radiology, therapy rooms) is recommended.

Mopheads and cleaning cloths must be discarded or changed after use.

Surface disinfectants must be applied for the correct contact time and applied according to the manufacturers’ instructions.

Some products with C. albicans or fungicidal claims may not be effective against C. auris. Hospital grade disinfectants effective against C.difficile spores are likely to be effective against C.auris.

Hydrogen peroxide vapour (HPV) and other no-touch irradiation systems including ultraviolet light should be used only as a supplement to standard cleaning and disinfection methods.

Healthcare and domestic staff must change their PPE and decontaminate their hands after contact with the patient or the environment, before attending to any other task.

4.2.1. Terminal clean

Once the patient has left the environment, terminal cleaning and disinfection should include all surfaces. Any reusable item that may have come into contact with the patient or the environment should be thoroughly cleaned before it leaves the room. For example, seal waste items within the room, undertake a first clean of bulky reusable items before leaving the room, followed by disinfection.

1,000 ppm of available chlorine with detergent is effective. Equivalent disinfectant products may be selected, with agreement of the local IPC team, and used in accordance with local policies and manufacturers’ guidance. The manufacturer’s recommended product ‘contact time’ and instructions for use must be followed for all disinfection solutions. Quaternary ammonium compounds are currently not recommended as the evidence regarding their efficacy for C. auris is not clear.

As different staff groups may be responsible for different items, attention should focus on all relevant items undergoing decontamination, and the importance of cleaning prior to disinfection should be emphasised. Application of disinfectant should ensure good surface contact before the disinfectant dries. Privacy curtains should be changed (with contaminated curtains being removed prior to disinfection and hanging of new curtains taking place after cleaning and disinfection is complete). All equipment (including patient monitoring devices and mobility aids) should be cleaned in accordance with manufacturer’s instructions and where relevant returned to clinical engineering or to manufacturers for specialised decontamination (for example dynamic mattresses). Stocks of single-use items in the immediate patient environment should be discarded. Consideration should be given to discarding less expensive re-usable items that are difficult to decontaminate effectively.

If any non-contact disinfection is used (such as gaseous hydrogen peroxide or ultraviolet), this should be as an additional safety measure, and not instead of full cleaning and disinfection. Individual trusts should adopt a local cleaning and disinfection policy and regimen depending on the level of contamination and case load.

Domestic staff will require training and supervision to ensure competence. This will include training to clean in a specified sequence (clean to less clean), training in use of equipment and cleaning products correctly, and training on the safe use of PPE (changing gloves and aprons), with appropriate hand decontamination after cleaning each C. auris area.

4.2.2. Cleaning and decontamination of equipment

It is important to ensure regular cleaning and disinfection of the patient environment, surfaces, and equipment to minimise risk of transmission between patients. All equipment (including patient monitoring devices and mobility aids) should be cleaned in accordance with manufacturer’s instructions and recommended product ‘contact time’ must be followed for all disinfection solutions. Single-patient use devices (such as blood pressure cuffs) are recommended for patients who are infected or colonised with C. auris. Particular attention should be paid to cleaning and disinfection of reusable equipment (such as pulse oximeters, thermometer probes, computers on wheels, ultrasound machines, call bells) from the bed space of any infected/colonised patients.

4.2.3. Waste and linen disposal

Guidance from the National Infection Prevention and Control manual for England should be followed regarding waste and linen disposal, as for any other multi-resistant healthcare-associated organism.

Read further infection control guidance in community care settings

5. Communications

An information leaflet for affected patients and relatives is available on GOV.UK.

C. auris colonisation information should be included in any discharge summary or patient transfer documents, ideally with direct communication to IPC representatives at receiving hospitals or to receiving community care facilities, such as supported living, custodial settings, care homes, mental health facilities, learning disability and care settings of working age people. If positive results become available after discharge or transfer, information should be relayed to the receiving hospital/GP for further communication to the patient and for relevant public health action.

If a patient dies and the cause of death is attributable to C. auris, this must be included in the death certificate and should be relayed to the National Incident Team (candidaauris@ukhsa.gov.uk). To date there has been no attributable C. auris mortality within the UK.

Systems permitting, each hospital should label colonised patients with an infection control flag on the patient electronic case record, so healthcare professionals are immediately alerted to the C. auris status if or when that patient is readmitted in future.

6. Acknowledgements

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

The authors gratefully acknowledge the expert review and advice received from colleagues in UKHSA, NHS, and HPTs.

This guidance was approved and signed off by UKHSA Healthcare Associated Infections (HCAI) and Antimicrobial Resistance (AMR) Division.