The following article is from the October 2019 issue of Healthcare Hygiene magazine.

Candida auris: A Stealth Enemy With Environmental Persistence

By Kelly M. Pyrek

Public health entities such as the Centers for Disease Control and Prevention (CDC) and the World Health Organization (WHO) have compiled lists of the problem pathogens that continue to represent some of the greatest challenges to infection prevention and control (IPC) efforts. Rising on that list is Candida auris, an emerging worldwide public health scourge that is growing in intensity as a healthcare-acquired organism. It is particularly fearsome because of its innate resistance to multiple anti-fungal drugs and its resilience in the face of traditional hygiene measures.

In contrast to other Candida species, C. auris is transmitted easily in the healthcare setting, and it is demonstrating the ability to persist both in the human host and on inanimate surfaces.
“Candida auris is an emerging fungus and one of the headaches with this species is it’s multidrug-resistant, meaning that it is resistant to multiple antifungal drugs commonly used to treat Candida infections,” confirms Rodney E. Rohde, PhD, MS, professor and chair of the Clinical Laboratory Science Program at Texas State University.

C. auris is also transcending traditional classification.

“Infection prevention experts tend to want to group pathogens into groups such as healthcare-acquired or community-acquired but when it comes to resistant organisms you really can't use those terms anymore,” Rohde says. “For instance, in some of the studies I have conducted with different organisms, what we used to think was only in the community, we see those strains or genotypes showing up in healthcare patients, and vice versa. So, even though microbiology and science tells us certain species are strictly found in the healthcare setting or strictly found in the community setting, in reality, it’s more of a global presence now due to so many vehicles and vectors that move in and out of healthcare settings.”

Rohde continues, “When we talk healthcare settings, we tend to only think hospitals, but we must think long-term care, outpatient and dialysis centers, clinics, and school and university health centers. I think it’s a communication problem; as scientists, we love to put things into pots and think we’ve described them adequately, and they are going to stay that way, but one of my favorite sayings is, organisms do not read the book of rules we’ve written for them. However, we teach it that way sometimes to try to make sense of it, but when you get into the real-world trenches, the truth is, we’re not able to do that. Organisms are crossing all boundaries.”

C. auris is insidious and still greatly unknown to hospitals and healthcare systems, much like the early days of Clostridium difficile. Experts are tending to agree that C. auris has operated under the radar. Rhodes and Fisher (2019) say that “Since its discovery, C. auris has caused a ‘stealthy pandemic,’ emerging across the globe and is now recorded in all continents except Antarctica. However, C. auris is thought to have been misidentified as C. haemulonii on several occasions, suggesting that C. auris has likely been circulating as a human pathogen before 2009.”

Candida auris as a newly recognized cause of fungal infection is catching healthcare professionals unawares, despite cases being reported for nearly a decade now. Since the first official isolation of Candida auris in 2009, scientific community has witnessed an exponential emergence of infection episodes and outbreaks in different world regions. According to the CDC as of June 30, 2019 (the most recent data available), there are 725 confirmed cases, plus an additional 1,474 patients have been found to be colonized with C. auris by targeted screening in 10 states with clinical cases.

“Candida, as a fungus, is an unusual and a difficult problem to deal with,” Rohde says. “It’s not typically something you would be looking for in a healthcare setting, and certainly not looking for it in the common population. But it is a factor when looking at the immunocompromised population in context of environmental and healthcare exposure; for example, could a patient have been working in a certain environment where fungi and yeast are present? But to have a fungus emerging and breeding in a healthcare setting is a novel challenge in our lifetime. So, I think we are trying to grasp what that means.”

He continues, “Microbiologists are very concerned about C. auris because you almost have to rethink everything from the laboratory perspective. If you are a clinician in a smaller facility or a rural hospital, you often must wait on sending specimens out to an off-site, more central clinical laboratory, and that eats up valuable diagnostic time. You can typically know pretty quickly if you have Staph or Klebsiella or any of these other multi-resistant organisms if you have access to rapid assays; but yeast and fungi are not typically, at least right now, on the radar for panel testing. Fortunately, it’s starting to get there, obviously, but if you don’t have a trained medical laboratory professional on staff who can look under a microscope or conduct some type of test to rule out yeast -- you could have some slippage with respect to turnaround time on a diagnosis, so there’s a significant challenge.”

Kenters, et al. (2019) confirms that C. auris is a budding yeast that forms white, pink, or purple colonies on CHROMagar and can be difficult to distinguish from C. glabrata: “Some strains form aggregates of cells while others do not. In contrast to most other Candida species, it grows well at higher temperature (40-42° C) … First attempts to identify C. auris using PCR directly from swabs, seem to produce frequent ‘false positive’ results -- positive in PCR, negative in culture swabs. The first report of three cases of nosocomial fungemia due to C. auris showed that this yeast is commonly misidentified as C. haemulonii and Rhodotorula glutinis using traditional phenotypic methods. These widely used routine identification methods for yeasts are based on phenotypic assimilation/fermentation tests using sets of carbon and nitrogen compounds. An investigation of 102 clinical isolates, previously identified as C. haemulonii or C. famata, showed that 88.2 percent of the isolates were in fact C. auris, when confirmed by ITS sequencing. Several studies have since reported that, in routine microbiology laboratories, C. auris remains a problematic, difficult to identify pathogen, because commercial biochemical identification systems lacked this yeast in their databases.”

“C. auris transmission might be happening in a healthcare facility where professionals are completely unaware; they haven’t been able to detect it, or they are just missing it due to false negatives, and it begs the issue of the importance of having access to clinical microbiology services,” Rohde says. “An expert in clinical microbiology and medical laboratory needs to be part of this equation -- and not just for Candida auris, but for all pathogens in the healthcare setting."

He continues, “People may argue with me on this, but physicians and others in healthcare who are responsible for patient outcomes, often do not have the kind of expertise, education and background to immediately identify problematic organisms; not to detract from their other medical diagnostic skills, but you cannot know intuitively which microbe is infecting a patient without a medical laboratorian conducting a test -- it’s just not going to happen. Unless of course you have those cases where a physician might have the training and the background to do so, but that’s rare. Unfortunately, healthcare has become so lean that many laboratories are being consolidated, with their full microbiology toolbox being moved to a central location. This makes sense from an economic perspective, but sometimes for clinicians, that’s not the best scenario.”

That becomes even more of an issue when considering the long list of risk factors for patients, including immunosuppressed state, significant medical comorbidities, central venous catheters, urinary catheters, recent surgery, parenteral nutrition, exposure to broad spectrum antimicrobials, intensive care unit admission, and residence in a high-acuity skilled nursing facility.

Sears and Schwartz (2017) point out that C. auris has been recovered in samples from blood, catheter tips, cerebrospinal fluid, bone, ear discharge, pancreatic fluid, pericardial fluid, peritoneal fluid, pleural fluid, respiratory secretions (including sputum and bronchoalveolar lavage), skin and soft tissue samples (both tissue and swab cultures), urine, and vaginal secretions. Clinically, it has been implicated as a causative agent in fungemia, ventriculitis, osteomyelitis, malignant otitis, complicated intra-abdominal infections, pericarditis, complicated pleural effusions, and vulvovaginitis. They add that, “Much like other Candida species, there is uncertainty about the ability of C. auris to cause true respiratory, urinary, and skin and soft tissue infections despite being isolated from such samples.”

Cortegiani, et al. (2019) observe that “It is likely that many cases are missed, due to its misidentification with other non-albicans Candida spp. (e.g., C. haemulonii) by common microbiological diagnostic methods. Most of the reports occurred in critically ill adults, with risk factors for invasive fungal infections, such as immunosuppression, surgery, or indwelling catheters. The most common form of infection was candidemia, with a crude mortality of nearly 30 percent, but up to 70 percent in some reports.”

Cortegiani, et al. (2019) emphasize the criticality of fighting C. auris in the intensive care unit (ICU): “Despite implementation of countermeasures to limit colonization and infections in ICUs, cases continue to be reported, with a tendency to an endemic pattern. This reflects the ability of C. auris to persist in the clinical environment, facilitating its transmission within the critical-care setting. Multidrug-resistant (MDR) pattern and has been frequently observed (around 40 percent) with serious and complex consequences for antifungal therapy.”

The researchers note that due to the progressive spread of C. auris and treatment-related concerns, attention should be focused on the following major issues: worldwide transmission, anti-fungal treatment resistance, resilience and mechanisms of transmission, Implementation of infection prevention and control measures, and surveillance.

Let’s review each issue:

Antifungal treatment resistance
Cortegiani, et al. (2019) note that, “To date, there are not established minimum inhibitory concentrations (MICs) breakpoints for susceptibility testing of C. auris. Antifungal susceptibility data from three continents demonstrated that nearly 40 percent were MDR, with strains being resistant to fluconazole (90 percent), amphotericin B (30 percent to 40 percent) and echinocandins (5 percent to 10 percent). Moreover, a small percentage were also resistant to all antifungals available. C. auris demonstrates a high propensity to develop antifungal resistance under selective pressure. Recent studies demonstrated mutations in ERG11 (encoding lanosterol demethylase, the target of azoles) and FKS1 genes (encoding 1,3-beta-glucan synthase, the target of echinocandins). The recommended antifungals for C. auris treatment are mainly based on in-vitro testing and on the most frequently retrieved resistance profiles. Echinocandins are the recommended first-line treatment, pending specific susceptibility testing. Lipid formulation of amphotericin B should be an alternative in patients not responding to echinocandins. Close monitoring to early detect therapeutic failures and evolution of antifungal resistance is needed. New antifungals (e.g., SCY-078, APX001A/APX001, and rezafungin) have been tested with success but they are not available to date for clinical use.”

Resilience and mechanisms of transmission
Cortegiani, et al. (2019) explain that, “Unlike others Candida species, C. auris can colonize different anatomical sites (e.g., skin, skin, rectum, axilla, stool) and contaminate hospital equipment and surfaces, creating a vicious cycle of acquisition, spreading, and infection, particularly in ICUs. Indeed, bed, chairs, and monitoring tools (e.g., pulse oximeters, temperature probes) were contaminated during outbreaks. Recently, Eyre et al. published the results of a patients’ and hospital environmental screening program in Oxford, UK, after 70 patients (66 admitted to a neuro-ICU) were identified as being colonized or infected by C. auris. Seven patients developed an invasive infection during hospital stay. C. auris was detected mainly on skin-surface axillary temperature probes and other reusable tools. In patients monitored with skin-surface temperature probes, the risk of C. auris infection/colonization was seven times higher. Adoption of specific bundles of infection control had no significant effects until removal of the temperature probes. Recent studies have confirmed that C. auris can form biofilms, with a high variation of capacity of production depending on the C. auris strain considered. Biofilm may present reduced susceptibility to hydrogen peroxide and chlorhexidine. Quaternary ammonium compounds and cationic surface-active products seem to be ineffective against C. auris. Chlorine-based products appear to be the most effective for environmental surface disinfection. Chlorine-based disinfectants (at a concentration of 1,000 ppm), hydrogen- peroxide, or other disinfectants with documented fungicidal activity are recommended for environmental cleaning by the European CDC (ECDC).”

Implementation of infection prevention and control measures
Cortegiani, et al. (2019): say that, “Usually, outbreaks follow an exponential increase in the number of affected patients. It is mandatory to trace contacts with the aim to achieve early identification and screening of possible colonized patients that might be responsible for persistence of C. auris. Patients potentially or already colonized should be placed in single rooms with contact isolation precautions. Screening should be applied for contacts and patients previously hospitalized in healthcare settings where C. auris isolation was confirmed. Hand hygiene (with alcohol or chlorhexidine handrubs), wearing of protective clothing, and skin and environmental/equipment decontamination should be performed to prevent ongoing transmission.”

Global surveillance
Cortegiani, et al. (2019) emphasize that the emergence of C. auris and progressive spread of infections caused by other resistant pathogens has strengthened the need for a surveillance network for antimicrobial resistance globally for critically ill patients’ safety. The researchers observe, “It is hard to predict future C. auris diffusion. There will be outbreaks also in countries in which C. auris has been not reported yet? Will new MDR clones continue to emerge? Will we be able to apply effective antifungal stewardship programs and control measures?”

So much about C. auris is still unchartered territory, and as Rhodes and Fisher (2019) observe, “The global emergence of C. auris testifies to the unmapped nature of Kingdom Fungi and represents a new nosocomial threat that will require enhanced infection control across diverse healthcare and community settings.” The researchers add, “Currently, nothing is known about the origins and initial emergence of C. auris; its propensity to survive on inanimate objects within the hospital alongside resistance to disinfection protocols suggests the existence of an unknown non-human environmental reservoir. However, similar to other Candida species, the true nature of C. auris’ ancestral reservoirs currently remains elusive. The detection of clonal C. auris isolates on multiple continents simultaneously with distinct geographical antifungal resistance mechanisms suggests at least four independent emergence events followed by clonal expansion and the ongoing evolution of resistance in response to antifungal therapy … As sequencing technology develops, it is likely rapid sequencing of C. auris isolates can be achieved in 48 hours or less leading to the potential for bedside diagnostics twinned with molecular epidemiology of nosocomial patterns of transmission. Currently, it is not often known when patients become colonized – whether from the hospital environment or endogenous carriers – and the extent of carriage in the community remains largely unexamined.”

Lockhart (2019) indicates that rapid identification of colonized patients followed by isolation and contact precautions can help stem the spread of resistant clones: “Real-time detection methods can not only rapidly identify colonized patients but may also contribute to the rapid detection of resistance. Besides the existing laboratory-developed tests, there is at least one commercially available PCR test for the rapid detection of C. auris. There are currently two real-time assays for detection of anti-fungal resistance in C. auris, one for detecting azole resistance and the other for echinocandin resistance, as well as a report that echinocandin resistance can be detected using MALDI-TOF. These rapid platforms may become essential for the rapid determination of appropriate therapy.”

Environmental Persistence of C. auris
Short, et al. (2019) are sounding the alarm about the environmental persistence of C. auris; in their study, they found show that the ability of this multidrug-resistant yeast to form cellular aggregates increases survival after 14 days, which coincides with the upregulation of biofilm-associated genes. The researchers also caution, “Additionally, the aggregating strain demonstrated tolerance to clinical concentrations of sodium hypochlorite and remained viable 14 days post treatment. The ability of C. auris to adhere to and persist on environmental surfaces emphasizes our need to better understand the biology of this fungal pathogen.”

The researchers explain, “A key attribute of its pathogenic repertoire is its ability to survive and persist in the environment, yet the methods employed by this multidrug-resistant pathogen to disseminate throughout healthcare environments are still not fully understood. This has profound implications for decontamination and infection control protocols. Therefore, understanding the mechanisms of spread and survival in the hospital environment is critical, particularly as it persists on hospital fomites, extensively colonize individuals, and to survive as biofilms. Although traditionally biofilms are associated with formation on an indwelling medical device or on a mucosal substrate, recent investigations have suggested that these communities can facilitate residence and survival upon surfaces within a clinical setting. Despite the lack of nutrients, these communities adapt to survive and display increased tolerance to both heat and conventional disinfection treatments compared to a free-floating, equivalent cell. C. auris has been shown to readily transmit between hospital equipment, such as reusable temperature probes, and patients suggesting limitations of current infection control strategies. Commonly used disinfectants have been shown to be highly effective when tested in suspension, yet our previous data indicate that adherent C. auris cells can selectively tolerate biocides, including sodium hypochlorite and peracetic acid, in a substrate-dependent manner.”

To test the theory of biofilm formation being employed as an endurance strategy of C. auris, Short, et al. (2019) performed survival studies using two phenotypically distinct isolates based on their ability to form cellular aggregates. The researchers report, “Similar to previous findings, C. auris was found to remain viable for at least two weeks within a dry environment, regardless of the organic material in which it was suspended. It was shown that aggregating cells survived considerably better than their single-cell counterparts in PBS (>2.5 log2 cfu/mL) and 10% FCS (>4 log2 cfu/mL).”

The researchers add, “To confirm a role for biofilms in facilitating environmental persistence, a panel of biofilm associated genes, selected according to our group's previous transcriptional characterization of C. auris biofilms, was assessed. These genes were highly expressed across both phenotypes; however, comparative analysis revealed increased expression of approximately two-fold of several of these genes, which are involved in adhesion, extracellular matrix (ECM) production, and efflux pumps. ECM production is a well-documented resistance mechanism in pathogenic fungal biofilms of Candida spp. Increasing ECM production could provide the necessary protection for C. auris to survive extended periods of desiccation and retain viability following terminal disinfection.”

Using Infection Prevention and Control to Fight C. auris
Case investigation by public health entities such as the CDC and others has demonstrated that C. auris patients within similar geographic regions commonly had overlapping stays in the same acute-care hospital or long-term care facility, further supporting healthcare exposure as a key method of transmission.

Given the risk of nosocomial transmission of this multidrug-resistant pathogen, Sears and Schwartz (2017) emphasize that, “…infection control measures are vital to slowing the spread of C. auris. CDC recommends that all hospitalized patients with C. auris infection or colonization be treated using both Standard Precautions and Contact Precautions and housed in a private room with daily and terminal cleaning with a disinfectant agent active against Clostridium difficile spores (Cadnum et al., 2017). Receiving healthcare facilities should also be notified prior to transfer of an infected or colonized patient. Infection control precautions should be maintained until a patient is no longer infected or colonized with C. auris although there is uncertainty as to how best to monitor for ongoing colonization (CDC, 2017). There are no clear data on the efficacy of decolonization measures for patients colonized with C. auris, however this has been attempted with chlorhexidine in healthcare facilities during outbreaks.”

Kean, et al. (2018) articulate one of the greatest worries about C. auris: “The ability of this organism to survive on surfaces and withstand environmental stressors creates a challenge for eradicating it from hospitals.”

An experience with surface cleaning and disinfection to help combat C. auris in a U.S. healthcare facility was documented by Marrs, et al. (2017) who reported on two patients with C. auris infections that were admitted to the University of Chicago Medicine (UCM). In their study, the researchers collected environmental samples to assess environmental contamination before and after cleaning. They sampled the following surfaces: Bathroom sink drain, bedside table, bedrail, mattress, chair and window ledge. Routine terminal cleaning included using a 10 percent sodium hypochlorite solution that was applied to high-touch surfaces of the patient room and bathroom. The enhanced terminal cleaning process also included removing and replacing privacy curtains, using a single UV disinfection cycle in the room and bathroom, as well as supervision of the process by the environmental services manager.

The researchers note that due to a delay in identification of C auris for the first patient, pre-cleaning samples were taken more than two weeks after the patient had been discharged. During the intervening weeks, multiple patients had occupied the room and there had been more than three routine terminal cleanings. None of these samples was positive for C auris. Pre-cleaning, in-residence samples indicated C auris contamination of multiple surfaces for the second patient. Because of transfers within the institution, there are three sets of post-cleaning cultures for the second patient. All post-cleaning environmental cultures were negative for both patients. The researchers concluded that while routine terminal cleaning may have been effective in removing C auris from surfaces in one patient’s room, the enhanced terminal cleaning strategy used here was effective in their facility.

In their study, Kean, et al. (2018) evaluated a panel of C. auris clinical isolates on different surface environments against the standard disinfectant sodium hypochlorite and high-level disinfectant peracetic acid. The researchers note that, “C. auris was shown to selectively tolerate clinically relevant concentrations of sodium hypochlorite and peracetic acid in a surface-dependent manner, which may explain its ability to successfully persist within the hospital environment.”

The implications for infection control are significant, and Kean, et al. (2018) add that, “Understanding the mechanisms of spread and survival of this pathogen in the hospital environment is therefore crucial, particularly as it may persist on plastics and steel, and survive as biofilms. Several recent investigations have confirmed that C. auris is capable of prolonged survival on surfaces and have shown that surface disinfection protocols have variable and unsatisfactory outcomes. Since it has been shown recently that 1,000 ppm of an active chlorine solution is highly effective against these organisms when tested in suspension, the interaction between the pathogen and surfaces is likely to be important in determining survival of C. auris in the hospital environment. Our own work confirms this, with C. auris biofilms being generally insensitive to a range of key antimicrobial agents, thus prolonging their survival capacity.”

Kean, et al. (2018) investigated the general disinfectant sodium hypochlorite (NaOCl), widely used for terminal cleaning within the hospital environment, and the high-level disinfection agent peracetic acid, on different substrate surfaces. Four C. auris isolates obtained from various clinical sites were used, and several test surface substrates were used: cellulose matrix, 304 stainless steel, and polyester coverslips.
The researchers report that, “Initially, a standard disinfectant challenge was performed against C. auris on different substrates relevant to the hospital environment. A cellulose substrate was included to act as control for porosity. All four C. auris were significantly killed by NaOCl challenge at 1,000 and 10,000 ppm, irrespective of substrate and strain, though differences were observed between these substrates. Complete eradication was only achieved on the cellulose substrate. On the non-porous materials, significant quantities of viable yeast cells were killed on the steel surface following NaOCl at all treatment parameters, with ∼2.5 log10 reduction, with no significant differences observed at each time-point and concentration tested. Notably, those isolates treated with 1,000 ppm for 5 minutes showed significantly more regrowth compared to the other test conditions. When C. auris was tested on a polymer substrate, 5-minute exposure at 1,000 ppm was the least effective overall; although significant activity was observed, 4.95 log10 was retained on the surface. However, following an increased contact time of 10 minutes, or increased concentration to 10,000 ppm, significantly enhanced activity was observed, with an approximate overall 3.5 log10 reduction. When comparing both increased treatment parameters, no significant differences were observed between the regimens, and no notable regrowth was detected.”

The researchers observe, “There was a significant difference in activity between polymer and steel, which could be explained by the general ability of Candida species to adhere and form biofilms that are inherently more resistant. Whereas the isolates on steel responded by ∼3 log10 equally to the treatment regimens, on plastic we demonstrated differential activity depending on concentration and time of exposure to NaOCl. Another study reported greater efficacy of chlorine-based products on steel, but differences in experimental design may explain this, e.g. products and inoculum. Taken together, these data suggest that the standard disinfection procedures are surface-dependent, and that the diversity of fomites in the hospital setting could pose a problem for disinfection.”
Kean and McKloud, et al. (2018) used a three-dimensional complex biofilm model to investigate the efficacy of a panel of antiseptic therapeutics, including povidone iodine (PVP-I), chlorhexidine (CHX) and hydrogen peroxide (H2O2). They hypothesized that the ability of C. auris to form biofilms may be a potential mechanism that results in reduced susceptibility to antiseptic agents.

Initially, the antiseptic efficacy of three agents was tested against four C. auris isolates. As Kean and McKloud, et al. (2018) report, “When biofilms were treated with PVP-I for 5 minutes, concentrations of 1.25-2.5 percent were required to inhibit biofilms, which is a 16- to 128-fold change compared with planktonic cells. Increasing the exposure time (10 and 30 minutes) was shown to increase susceptibility to 0.625-1.25 percent, which is an eight- to 64-fold change compared with planktonic cells. CHX was highly active against planktonic cells, whereas biofilms were less susceptible with MICs increasing 2- to 16-fold. Elevated biofilm MICs were also observed following H2O2 exposure, with concentrations ranging between 0.25 and >1 percent required to kill biofilms, i.e. a 16-fold increase in the planktonically-active concentration. Regardless of the antiseptic active used, minimal differences in susceptibility were observed between 24- and 48-hour biofilms.”

The researchers observe, “Interestingly, PVP-I, a commonly used pre-surgical wash, was shown to be equally active against both early and mature biofilms when assessed by culture. This agreement with other studies in which PVP-I demonstrated excellent fungicidal activity against C. auris. The use of 10 percent PVP-I for surgical skin preparation has been used clinically for C. auris, with no postoperative infection reported. An interesting finding from this study was the ineffectiveness of H2O2 against both planktonic and sessile cells of C. auris. This finding is inconsistent with previous studies in which H2O2 showed significant fungicidal activity. Discrepancies between these findings are likely due to the test methodologies employed, with vaporized H2O2 assessed in one of these studies.”

The researchers add, “Future studies assessing the efficacy of CHX diluted in alcohol may provide a potential anti-biofilm strategy for successful skin disinfection. Collectively these findings illustrate the need for a greater understanding of the survival strategies of C. auris. Considering the documented high transmissibility of C. auris between patients and the environment, the implementation of stringent infection prevention and control procedures, coupled with the biological understanding of the organism, will ultimately aid the intervention strategies for this emerging pathogen.”

Recommendations
It is quickly becoming evident that current evidence for pragmatic infection prevention and control (IPC) recommendations is lacking, according to Kenters, et al. (2019), who reviewed the epidemiology of C. auris and identified best practices to provide guidance and recommendations for IPC measures, based on available scientific evidence, existing guidelines and expert opinion. The IPC working group of the International Society of Antimicrobial Chemotherapy (ISAC) organized a meeting with infection prevention and mycology experts to review recommendations on IPC measures on C. auris in inpatient healthcare facilities for healthcare workers on the most common interventions including: screening, standard precautions, cleaning and disinfection, inpatient transfer, outbreak management, decolonization and treatment.

The working group identified these best practices to help combat C. auris:

1. Optimal diagnostics
It is crucial to identify and report C. auris correctly in order to provide optimal patient care, treatment and initiate appropriate IPC measures. All isolates should be susceptibility tested, from whatever body site, because of varying levels of resistance.

2. Patient screening upon admission
Outbreak investigations have revealed that screening sites most frequently culturing positive for C. auris were axilla, groin, rectum and urine. Other sites for screening, even if less sensitive to colonization are the nose, mouth, external ear canals, catheter urine and wounds. If a patient has open wounds and/or intravascular catheters they should be included for screening in addition to the swabbing sites. It is important to only swab open wounds if they are not sealed by wound dressing, as opening a sealed wound can cause a risk for patients to become colonized with C. auris if they are carriers. Risk groups admitted to the hospital are patients previously admitted into intensive care units in endemic countries and, transfers from hospitals known to have C. auris. Hospitals that have C. auris present should liaise with receiving hospitals infection control teams and ensure that appropriate is information is shared. If healthcare institutions have an existing surveillance screening protocol in place for patients at high-risk for colonization with MDRO, it might be more valuable to add C. auris to the existing testing panel in the lab, rather than to re-implement additional swabbing sites.

3. Infection prevention and control interventions
For healthcare facilities to be prepared for a first case of C. auris, it is important to have a screening protocol as well as adequate IPC procedures in place. Any detection of C. auris should be immediately reported to the IPC department, leading to timely implementation of strict IPC-related measures. Patients colonized or infected with C. auris should be isolated until discharge and flagged for at least one year after the first negative screening culture. When patients are transferred within the institution or to other healthcare facilities handing over of the patient C. auris status needs to be ensured. Patient contact screening of direct contacts should be initiated on the detection of a ‘first’ case, including those being discharged and this may involve tracking back throughout the patient admission if internal transfers have taken place. Every patient should be screened in the axilla and groin, including any other relevant sites (e.g. nose, urine, rectum, throat, wounds and catheter exit sites).

The following IPC measures should be implemented for any C. auris case:
- Standard precautions
Hand hygiene is key to prevent transmission of any microorganism, including C. auris. Special attention should be given to adequate compliance with hand hygiene while caring for patients in isolation. Hand hygiene should be performed at the point of care using alcohol-based handrub (ABHR). While ABHR is the preferable choice, water and soap should be used when hands are visibly soiled and a dedicated sink needs to be in place to wash hands.

- Patient environment
Patients colonized or infected with C. auris need to be placed in contact precautions in a single room, ideally with negative pressure, and preferably with an anteroom and in-suite bathroom/toilet. If the latter is not available, patients should use a dedicated washroom or a waterless washing product, as well as a dedicated commode. The use of an isolation room with anteroom might be preferable, not because airborne spread is assumed, but because compliance with isolation measures might possibly be higher, as the double doors function as a reminder. A flagging system indicating the isolation needs to be visible at the entry of the patient room and instructions for HCWs and visitors need to be available. All biomedical products and equipment should be used as disposables, or, if re-usable should be left in the patient's room until discharge and thorough disinfection. Sharing biomedical products and equipment to other wards poses a risk of additional transmission. For mattresses and pillows, HCWs should ensure that they are 100 percent sealed before using them for a C. auris patient and its integrity should be assessed upon discharge if they are to be used for another patient.

- Personal protective equipment (PPE)
It has become evident that the use of a long-sleeved gown and gloves are sufficient to enter the room of patients found positive for C. auris. Taking into consideration that people often (unconsciously) touch their face, a surgical mask could be considered to prevent colonization of healthcare staff, since one HCW has been found transiently positive in the nose in a previous outbreak.

- Environmental cleaning
Enhanced daily and terminal disinfection has been shown to be crucial to control the spread of C. auris within healthcare facilities. In addition, the frequency of cleaning and disinfection should be at least twice daily, of at least all high-touch surfaces. Terminal cleaning and disinfection of the rooms after patient discharge needs to be performed with great diligence. When selecting a product, users should keep in mind the toxicity of a product and select one that is safer to use near a patient. Innovative automated decontamination technologies, such as UV-C disinfection can be used to ensure optimal terminal cleaning of surfaces but are an additional safeguard and not a replacement of the routine cleaning method. Both methods – UV-C and HPV – require vigorous cleaning before being effective against microorganisms. If UV-C is used, the duration of exposure for efficacy is longer than that for vegetative bacteria and a cycle time effective for spores such as C. difficile should be selected.

Cleaning and disinfection of reusable equipment is particularly important, especially as these items may be decontaminated at the departmental level by clinical staff. Where possible, dedicated equipment should be used. If dedicated equipment is not an option equipment and devices must be disinfected thoroughly after every use in line with the manufacturer's instructions and considering materials compatibility. The surfaces of re-usable items should be periodically examined to check for surface integrity and the continued ability to be able to effectively decontaminate. Materials that cannot be disinfected should not be used or discarded after use. Where possible, single-use equipment is preferred to limit possible spread via inadequately disinfected equipment. Equipment that is cleaned by clinical staff should be audited to ensure that it is effective, and facilities may wish to consider whether formal training for clinical staff in decontamination has been or should be provided.

- Patient clothing
The role of patient clothing in the transmission of C. auris is unclear. In the experience of the hospitals dealing with outbreaks, patients were asked to use hospital garments or clothes that have been washed at high temperatures. The expert group was not able to give a recommendation on this topic. Seeing that C. auris has been found to survive on linen, it may be prudent to change bedding and patient attire daily if decolonization or skin suppression is being attempted.

- Patient movement in the facility
Transfer of colonized or suspected patients for C. auris should be done with great care. The treatment of patients should always come first, but if transfer can be prevented by using mobile equipment, this should be considered. When patients need to go to radiology, for example, they should ideally be placed at the end of the schedule to allow time for terminal decontamination of the area.

- Readmission of previous C. auris positive patient
If known, previous positive C. auris patients should be placed in contact isolation and screened on three consecutive days. Contact precautions may be stopped if all three screens are negative. Weekly screening is however recommended as C. auris may resurface after antibiotic therapy or other interventions such as chemotherapy. This is the minimum measure, as local MDRO-related guidelines that are stricter must be regarded.

- Outpatient management
Family members and healthcare providers can become colonized; however, this is of minimal risk to the “healthy” individual. There are no guidelines yet for management of C. auris-colonized patients; however, it is prudent that sharing items should be kept to a minimum in line with the principles for other fungal infections: in particular, towels and clothing should not be shared, as well as cosmetic items, creams, ointments, etc., even in the absence of studies demonstrating any effectiveness.

- Healthcare personnel education and training
Compliance with, as well as adequacy of using IPC measures is essential to prevent transmission of C. auris within the healthcare facility. To increase staff's awareness around IPC measures, on-site training and auditing is critical to contain C. auris. Training should focus on standard precautions, PPE, environmental cleaning and other IPC measures applied to control C. auris. In addition, compliance and correct execution of IPC measures should be monitored, and direct feedback to HCWs should be provided.

- Outbreak management
The index or any unexpected case colonized or infected with C. auris should be isolated in a single/isolation room and direct contacts should be placed in cohort isolation, with contact precautions and no new patients should be admitted to the affected room. Maintaining cohorts of “proven colonized”, “possibly colonized” and “no risk” patients is important under all circumstances, even if that would lead to lowered bed capacities, reduction of admitted patients or cancelation of operating procedures. As C. auris has been cultured from the hands of healthcare personnel, where possible they should be assigned to one of the cohorts, instead of working throughout the whole unit. If the outbreak is large, creation of a separate unit for all proven colonized patients might be advisable. Single use equipment or dedicated equipment should be used. A root cause analysis by Schelenz, et al. found that patients who had contact with a positive case or contaminated environment were likely to have contracted C. auris within just four hours of contact.

To confirm negative patients, three consecutive C. auris screenings should be negative. In the absence of published data, it is advisable to space out the three screening times (such as day 3-5-7), instead of doing them on day 1-2-3 or even all in a day. When de-isolated, weekly screening of the negative contact patients, until discharge, is recommended. Healthcare personnel have been identified as carriers in the nose and groin, so during an ongoing outbreak, screening of healthcare staff could be considered, as well as unannounced cultures from hands as an educational measure.

In an outbreak scenario, cleaning and disinfection should be increased to three times daily of at least all high-touch surfaces with a product effective against C. auris. Terminal cleaning and disinfection should be monitored with quality indicators that go beyond visual inspection, such as ATP or fluorescent markers, to ensure the quality of terminal cleaning and disinfection. If available, UV-C or HPV could be used after terminal cleaning and disinfection, as an additional assurance that the room has been adequately decontaminated and is safe for the next bed occupant.

No recommendations can be provided about to the effect of decolonizing patients; in theory, this may lead to a lower burden of yeast on the patient's skin and thus, a lower risk of transmission. This approach has been adopted in outbreaks; however, non-conflicted data are missing to draw that conclusion for C. auris. Currently, limited evidence on the use of topical agents for the control of skin colonization exists. In one UK major outbreak, 2 percent chlorhexidine washcloths or 4 percent chlorhexidine solution were used to control skin shedding as part of several interventions. However, despite daily CHX bathing, patients described in the UK continued to be colonized with C. auris. Chlorhexidine solutions may dry the skin in such a way that it may lead to prolonged colonization with C. auris. Some patients however remained persistently colonized, possibly due to recolonization from bedding, as Candida spp. have been demonstrated to survive on polyester textiles for up to eight days.
Mandatory national reporting of outbreaks in institutions should be considered, as well as mandatory reporting of infections with C. auris. In countries that do not have laws accounting for this, mandatory sharing of outbreak status data with regional healthcare providers is advisable.

References and Recommended Reading:
Cadnum JL, Shaikh AA, et al. Effectiveness of disinfectants against Candida auris and other Candida species. Infect Control Hosp Epidemiol, 38 (2017), pp. 1240-1243.
Chow NA, et al. Multiple introductions and subsequent transmission of multidrug-resistant Candida auris in the USA: a molecular epidemiological survey. Lancet Infect Dis, 18 (2018), pp. 1377-1384.
Chowdhary A, et al. Candida auris: a rapidly emerging cause of hospital-acquired multidrug-resistant fungal infections globally. PLoS Pathog, 13 (2017), Article e1006290.
Cortegiani A, Misseri G, Fasciana T, Giammanco A, Giarratano A, Chowdhary A. Epidemiology, clinical characteristics, resistance, and treatment of infections by Candida auris. J Intensive Care. 2018;6:69.
Cortegiani A, Misseri G, Chowdhary A. What's new on emerging resistant Candida species. Intensive Care Med. 2018. https://doi.org/10.1007/s00134-018-5363-x.
Cortegiani A, et al. The global challenge of Candida auris in the intensive care unit. BMC Critical Care. Vol. 23, No. 150. 2019.
Eyre DW, Sheppard AE, Madder H, Moir I, Moroney R, Quan TP, et al. A Candida auris outbreak and its control in an intensive care setting. N Engl J Med. 2018;379:1322–31.
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https://www.elsevier.com/connect/preventing-diagnostic-errors-by-uniting-the-clinical-laboratory-with-direct-patient-care
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