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Candida auris is a multidrug-resistant nosocomial bloodstream pathogen that has been reported from Asian countries and South Africa. Herein, we studied the population structure and genetic relatedness among 104 global C. auris isolates from India, South Africa and Brazil using multilocus sequence typing (MLST), amplified fragment length polymorphism (AFLP) fingerprinting and matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS). RPB1, RPB2 and internal transcribed spacer (ITS) and D1/D2 regions of the ribosomal DNA were sequenced for MLST. Further, genetic variation and proteomic assessment was carried out using AFLP and MALDI-TOF MS, respectively. Both MLST and AFLP typing clearly demarcated two major clusters comprising Indian and Brazilian isolates. However, the South African isolates were randomly distributed, suggesting different genotypes. MALDI-TOF MS spectral profiling also revealed evidence of geographical clustering but did not correlate fully with the genotyping methods. Notably, overall the population structure of C. auris showed evidence of geographical clustering by all the three techniques analysed. Antifungal susceptibility testing by the CLSI microbroth dilution method revealed that fluconazole had limited activity against 87% of isolates (MIC90, 64 mg/L). Also, MIC90 of AMB was 4 mg/L. Candida auris is emerging as an important yeast pathogen globally and requires reproducible laboratory methods for identification and typing. Evaluation of MALDI-TOF MS as a typing method for this yeast is warranted.[1]
CDC is aware of one isolate of C. auris that was detected in the United States in 2013 as part of ongoing surveillance. Experience outside the United States suggests that C. auris has high potential to cause outbreaks in healthcare facilities. Given the occurrence of C. auris in nine countries on four continents since 2009.[2]
Candida auris is an emerging multidrug-resistant (MDR) yeast that can cause invasive infections and is associated with high mortality. It was first described in 2009 after being isolated from external ear discharge of a patient in Japan 1. Since the 2009 report, C. auris infections, specifically fungemia, have been reported from South Korea 2, India 3, South Africa 4, and Kuwait 5. Although published reports are not available, C. auris has also been identified in Colombia, Venezuela, Pakistan, and the United Kingdom.[2]
C. auris infections have most commonly been hospital-acquired and occurred several weeks into a patient’s hospital stay. C. auris has been reported to cause bloodstream infections, wound infections, and otitis 2. It has also been cultured from urine and the respiratory tract; however, whether isolation from these sites represented infection verses colonization in each instance is unknown. C. auris has been documented to cause infections in patients of all ages. Patients were found to have similar risk factors for infections with other Candida spp. 6, 7, including: diabetes mellitus, recent surgery, recent antibiotics, and presence of central venous catheters 3. Co-infection with other Candida spp. and detection of C. auris while the patient was being treated with antifungals have also been reported.[2]
Although no established minimum inhibitory concentration (MIC) breakpoints exist for C. auris, resistance testing of an international collection of isolates conducted by CDC demonstrated that nearly all isolates are highly resistant to fluconazole based on breakpoints established for other Candida spp. More than half of C. auris isolates were resistant to voriconazole, one-third were resistant to amphotericin B (MIC ≥2), and a few were resistant to echinocandins. Some isolates have demonstrated elevated MICs to all three major antifungal classes, including azoles, echinocandins, and polyenes, indicating that treatment options would be limited.[2]
C. auris phenotypically resembles Candida haemulonii 1. Commercially available biochemical-based tests, including API strips and VITEK-2, used in many U.S. laboratories to identify fungi, cannot differentiate C. auris from related species. Because of these challenges, clinical laboratories have misidentified the organism as C. haemulonii and Saccharomyces cerevisiae. Some clinical laboratories do not fully identify all Candida to the species level, and C. auris isolates have been reported as “other Candida spp.” Clinical, state, and public health laboratories should be aware of this organism and of the limitations in its identification.[2]
At least two countries have described healthcare outbreaks of C. auris infection and colonization involving more than 30 patients each. Analysis of isolates from these clusters demonstrate a high degree of clonality within the same hospital, supporting the idea that the organisms are being transmitted within those healthcare facilities. The precise mode of transmission within the healthcare facility is not known. However, experience during these outbreaks suggests that C. auris might contaminate the environment of rooms of colonized or infected patients. Good infection control practices and environmental cleaning may help prevent transmission.[2]
Infection Control — Until further information is available, healthcare facilities should place patients with C. auris colonization or infection in single rooms and healthcare personnel should use Standard and Contact Precautions.[2]
Environmental Cleaning – Anecdotal reports have suggested that C. auris may persist in the environment. Healthcare facilities who have patients with C. auris infection or colonization should ensure thorough daily and terminal cleaning and disinfection of these patient’s rooms using an EPA-registered hospital grade disinfectant with a fungal claim.[2]
Therefore it is important that any Candida spp isolates associated with invasive infections and isolates from superficial sites in patients from high intensity settings and those transferred from an affected hospital (UK or abroad) should be analysed to species level. If Candida haemulonii, Candida famata, Candida sake or Saccharomyces cerevisiae are identified, further work should be undertaken to ensure that they are not C. auris. This would involve either molecular sequencing of the D1/D2 domain or MALDI-TOF Biotyper analysis with C. auris either already present or added to the database.[3]
According to published data, commercially available biochemical-based tests, including API AUX 20C and VITEK-2 YST, used in many front line diagnostic laboratories can misidentify C. auris as Candida haemulonii, Saccharomyces cerevisiae or Rhodotorula glutini.[3]
Therefore, it is important that any Candida spp. isolates associated with invasive infections and isolates from superficial sites in patients from high intensity settings and those transferred from an affected hospital (UK or abroad) should be analysed to species level. As knowledge on the epidemiology and prevalence in the UK is as yet limited, PHE is currently not in a position to make specific recommendations with regards to screening policy. However, C. auris screening could be considered for patients at risk for Candida disease (ESCMID guidance developing group define such patients as “[…] mainly ICU patients, paediatric, HIV/AIDS and patients with malignancies including haematopoietic stem cell transplantation.”)[4]
Since April 2015, an adult critical care unit in England has been managing an outbreak of C. auris, with more than 40 patients either colonised or infected; approximately 20% with candidaemia. The hospital outbreak has been difficult to control, despite enhanced infection control interventions, including regular patient screening, environmental decontamination and ward closure[4]
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 pseudohyphae on cornmeal agar. 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. Growth on this and other chromogenic agars (which may display a different colour) cannot be used as a primary identification method. Chromogenic agars are useful to identify mixed cultures including the presence of C. albicans. If there is evidence of non - albicans on chromogenic agar these should be sub-cultured on Sabouraud’s agar and identified according to local laboratory protocols. It is unlikely that any of the currently available biochemical-based tests will include C. auris in their database as it is a newly recognised species so laboratories are advised to check the databases provided for their current methods. According to published data, commercially available biochemical-based tests, including API AUX 20C and VITEK-2 YST, used in many front line diagnostic laboratories can misidentify C. auris as Candida haemulonii, Saccharomyces cerevisiae or Rhodotorula glutinis (the latter species is pink on Sabouraud’s agar and is easily distinguished). Therefore it is important that any Candida spp isolates associated with invasive infections and isolates from superficial sites in patients from high intensity settings and those transferred from an affected hospital (UK or abroad) should be analysed to species level. If Candida haemulonii, Candida famata, Candida sake or Saccharomyces cerevisiae are identified, further work should be undertaken to ensure that they are not C. auris. This would involve either molecular sequencing of the D1/D2 domain or MALDI-TOF Biotyper analysis with C. auris either already present or added to the database.[3]
Antifungal susceptibility testing: There are no established minimum inhibitory concentration (MIC) breakpoints at present for C. auris. Using breakpoints for other Candida spp the Centers for Disease Control and Prevention (CDC) demonstrated that of the global outbreaks that they have been investigating, nearly all isolates are highly resistant to fluconazole. In their analysis, more than half of C. auris isolates were resistant to voriconazole, one- third were resistant to amphotericin B (MIC ≥2 mg/L), and a few were resistant to echinocandins. Some isolates have demonstrated elevated MICs to all three major antifungal classes, including azoles, echinocandins, and polyenes indicating that treatment options would be limited. Whole genome sequencing of the organism has found resistant determinants to a variety of antifungal agents. [3]
Treatment
Experience to date from the PHE Mycology Reference Laboratory indicates that so far no multi-drug resistant strains have been found in the UK but all isolates are resistant to fluconazole and often cross-resistant to other azoles. First-line therapy remains an echinocandin pending specific susceptibility testing which should be undertaken as soon as possible. However, there is evidence that resistance can evolve quite rapidly in this species, ongoing vigilance for evolving resistance is advised in patients who are found to be infected or colonised with C. auris. There is currently no evidence or experience to support combination therapy in invasive infections with this organism and clinicians are advised to make decisions on a case by case basis.[3]
Decolonisation:
Colonisation of patients has been reported from affected hospitals around the world. There is no evidence currently that can establish whether C. auris is susceptible or resistant to chlorhexidine. More work is being done in this area. Clinical experience to date has shown that colonisation is difficult to eradicate and colonisation tends to persist making infection prevention and control strategies particularly important. However it is still 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, skin decontamination and mouth gargles with chlorhexidine washes use of topical nystatin and terbinafine would be options for targeted topical management of key sites such as venous cannula entry sites.[3]
Screening
All Trusts are encouraged to develop a screening policy after local risk assessments are undertaken. Screening is recommended in units that have ongoing cases or colonisations. Screening is also advised for patients coming from other affected hospitals / units in the UK and abroad. Currently hospital outbreaks have been reported from India, Pakistan, Venezuela and Colombia; although UK and worldwide prevalence is still to be established due to problems with laboratory diagnosis. Suggested screening sites, based on the predilection of Candida species to colonise the skin and mucosal surfaces ie genitourinary tract, mouth and respiratory tract, are: nose, throat, groin urine / urethral swab perineal or low vaginal swab if appropriate. sputum / endotracheal secretions, drain fluid (abdominal/pelvic/mediastinal), cannula entry sites if clinically indicated, wounds.[3]
Routine wound swabs may be used to collect the screening sample. All screen positive patients should be isolated or cohorted as described below. As for other healthcare associated infections, a series of three negative screens taken 24 hours apart are needed to de-isolate the patient. As there is clinical experience of recurrence of colonisation, the need for ongoing vigilance in the form of weekly screens in certain clinical environments should be considered by performing local risk assessments.[3]
Infection, prevention and control (IPC)
Reports from India, Pakistan and Venezuela (CDC, personal communication) have described healthcare outbreaks of C. auris infection and colonisation involving more than 30 patients. The precise mode of transmission within the healthcare environment is not known. However, experience during these outbreaks suggests that C. auris might substantially contaminate the environment of rooms of colonised or infected patients. Transmission directly from fomites (such as blood pressure cuffs, stethoscopes and other equipment in contact with the patient) is a particular risk, however this does not
preclude transmission via hands of healthcare workers and hand hygiene needs to be strictly adhered to. Where possible equipment used for the infected/colonised patient should not be shared with other patients on the ward unless between-patient cleaning can be assured. It is essential that all healthcare providers work in a multi-disciplinary team with their Clinical Microbiologists and under the direction of their specialist IPC(infection prevention care) team.[3]
The patient
Key infection prevention and control measures should include: isolation of all patients colonised or infected with the organism in a single room with ensuite facilities wherever possible isolation of all patients who have been transferred from an affected UK hospital or a hospital abroad until screening results are available strict adherence to standard precautions including hand hygiene using soap and water followed by alcohol hand rub personal protective equipment in the form of gloves and aprons (or gowns if there is a high risk of soiling with blood or body fluids) these should be donned after hand washing and before entering the room and removed and discarded in the room followed by a thorough hand wash and application of alcohol hand rub visors and masks are not routinely required and should be worn only if there is a procedural risk of spillage or splashes visitors of infected or colonised patients need to be briefed about the infection and infection prevention and control precautions reinforced; including the need for robust hand hygiene and use of protective aprons.[3]
A chlorine releasing agent is currently recommended for cleaning of the environment at 1000 ppm of available chlorine.[3]
Terminal clean: once the patient has left the environment a terminal clean should be undertaken preferably using hydrogen peroxide vapour, all equipment should be cleaned in accordance with manufacturer’s instructions and where relevant returned to the company for cleaning. Particular attention should be paid to cleaning of multiple-use equipment (eg BP cuffs, thermometers, computers on wheels, ultra-sound machines) from the bed space of an infected/colonised patient.[3]
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Overview of Candida auris
Candida auris is a fungus, recently described as a rare cause of fungal infection with significant resistance to antifungal medications.[5] Candida auris isolates from north and south Indian hospitals, Japan and Korea were all found to be resistant to the antifungal medication fluconazole.[5] Some isolates were also noted to be resistant to flucytosine and voriconazole.[5] The high rate of therapeutic failure noted in cases of Candida auris fungemia poses significant concerns.[5] It's high potential for nosocomial horizontal transmission has been demonstrated.[6][7]An outbreak of fifty cases over a sixteen month period (April 2015-July2016) in a cardiothoracic center in London is the first reported case, and the largest outbreak in Europe.[7] It is recognized as a globally emerging fungal pathogen[7].
Historical Perspective
References
- ↑ Prakash A, Sharma C, Singh A, Kumar Singh P, Kumar A, Hagen F; et al. (2016). "Evidence of genotypic diversity among Candida auris isolates by multilocus sequence typing, matrix-assisted laser desorption ionization time-of-flight mass spectrometry and amplified fragment length polymorphism". Clin Microbiol Infect. 22 (3): 277.e1–9. doi:10.1016/j.cmi.2015.10.022. PMID 26548511.
- ↑ 2.0 2.1 2.2 2.3 2.4 2.5 2.6 2.7 Centers for Disease Control and Prevention. https://www.cdc.gov/fungal/diseases/candidiasis/candida-auris-alert.html Accessed on November 11th, 2016.
- ↑ 3.00 3.01 3.02 3.03 3.04 3.05 3.06 3.07 3.08 3.09 3.10 3.11 Public Health England.https://www.gov.uk/government/uploads/system/uploads/attachment_data/file/534174/Guidance_Candida__auris.pdf. Accessed on November 11th, 2016.
- ↑ 4.0 4.1 Schmoldt A, Benthe HF, Haberland G (1975). "Digitoxin metabolism by rat liver microsomes". Biochem Pharmacol. 24 (17): 1639–41. PMID HPR 10(21) Ref: HPR 10(21) Check
|pmid=value (help). - ↑ 5.0 5.1 5.2 5.3 Chowdhary A, Anil Kumar V, Sharma C, Prakash A, Agarwal K, Babu R; et al. (2014). "Multidrug-resistant endemic clonal strain of Candida auris in India". Eur J Clin Microbiol Infect Dis. 33 (6): 919–26. doi:10.1007/s10096-013-2027-1. PMID 24357342.
- ↑ Calvo B, Melo AS, Perozo-Mena A, Hernandez M, Francisco EC, Hagen F; et al. (2016). "First report of Candida auris in America: Clinical and microbiological aspects of 18 episodes of candidemia". J Infect. 73 (4): 369–74. doi:10.1016/j.jinf.2016.07.008. PMID 27452195.
- ↑ 7.0 7.1 7.2 Schelenz S, Hagen F, Rhodes JL, Abdolrasouli A, Chowdhary A, Hall A; et al. (2016). "First hospital outbreak of the globally emerging Candida auris in a European hospital". Antimicrob Resist Infect Control. 5: 35. doi:10.1186/s13756-016-0132-5. PMC 5069812. PMID 27777756.