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ABSTRACTS AND REFERENCES

Diagnosis of superficial dermatophyte and mould infections
Dr Elizabeth Johnson (The British Society of Medical Mycology)
Learning points
 Laboratory diagnosis of superficial fungal infection relies on direct microscopy of a skin, nail or hair sample and culture of the infecting organism.  Positive direct microscopy is proof of infection but culture is required to establish the causative organism which is desirable for epidemiological and treatment purposes.  The arrangement of spores in or around a hair shaft can give important information regarding  It is important to culture samples at 25-30C on a selective agar such as Sabourauds agar containing an antibacterial agent (e.g. chloramphenicol) and a selective antifungal agent such as cycloheximide (actidione) unless a non-dermatophyte mould is suspected.  There are few non-dermatophyte moulds that can cause infection of skin, the most common of these is Scytalidium dimidiatum but Phaeoannellomyces werneckii and Stenella agaruata can cause a skin infection known as tinea nigra.  Non-dermatophyte moulds can often be recovered as contaminants from nail tissue, they should only be considered significant causes of infection if hyphae are seen on direct microscopy and a mould is isolated in pure culture from a significant number of inoculum points in the absence of a dermatophyte. The dermatophytes comprise a group of fungi which cause tinea or ringworm infections of the keratinised tissue of skin, nail and hair. They belong to three different genera: Trichophyton, Microsporum and Epidermophyton and within these genera are organisms that are anthropophilic, zoophilic or geophilic. There are also some non-dermatophyte moulds such as Phaeoannellomyces werneckii, Stenella agaruata and Scytalidium dimidiatum which are capable of causing skin infections although infections with these organisms are rare. A large number of non-dermatophyte moulds can cause infection in traumatised nails, the most common are: Scopulariopsis brevicaulis, the Aspergillus versicolor group and Fusarium spp. However, nail tissue is often contaminated with fungal spores and isolation of a non-dermatophyte mould should only be considered significant if cultured from at least four fragments of a microscopically positive nail in the absence of a dermatophyte isolate. Laboratory diagnosis is by microscopy and culture of scrapings of skin or nail, sellotape strippings of skin or short lengths of plucked hairs. Samples should be softened in 20% potassium hydroxide which has been left for 10–20 min to digest the keratin. Use of an optical brightener such as Calcofluor white or Blankophor may enhance the detection of fungal hyphae particularly in nail tissue. In order to identify the causal fungus the specimen should be cultured on a plate of Sabouraud agar containing chloramphenicol to suppress the growth of contaminating bacteria and an agent such as actidione to suppress mould growth unless non-dermatophyte mould infection is suspected. Plates should be incubated aerobically at 25–30ºC for up to three weeks. After this time isolates can be identified by macroscopic and microscopic examination. Two types of conidia are formed by dermatophytes: small unicellular microconidia and larger septate macroconidia. The type and microscopic appearance of the spores together with an examination of gross colonial morphology enables species identification. Molecular methods for dermatophyte identification will also be discussed. 1. Padhye AA, Summerbell RC. The dermatophytes In Merz WP and Hay RJ (Eds) Topley and Wilson’s Microbiology and Microbial Infections: Medical Mycology (10th Edition). Hodder Arnold ASM Press 2005, pp. 220-224. 2. Campbell CK, Johnson EM. Dermatomycotic molds In: Merz WP and Hay RJ (Eds) Topley and Wilson’s Microbiology and Microbial Infections: Medical Mycology (10th Edition). Hodder Arnold ASM Press 2005, pp. 244-255. 3. Midgley G, ClaytonYM, Hay RJ. Diagnosis in Color: Medical Mycology Mosby-Wolfe 1988. 4. Ellis DH, Watson AB et al. Non-dermatophytes in onychomycosis of the toenails. Br J 5. Campbell CK, Johnson EM, Philpott C and Warnock DW Identification of Pathogenic Fungi 6. Denning DW, Evans EGV et al. Fungal nail disease: a guide to good practice: report of a working group of the British Society for Medical Mycology Br Med J 1995;311:1277–1281.


Epidemiology and treatment of dermatophyte infections
Dr Susan Howell (St Thomas’ Hospital, London)

Learning points
 Trichophyton rubrum is the commonest cause of fungal infection of human skin and nail.  The commonest causes of scalp infection of children in urban areas is Trichophyton tonsurans.  Identification of the causative organism has important implications for patient management.  Topical treatment with terbinafine, azole or imidazoles is appropriate for localised skin infections of the body, face and limbs, however, extensive lesions, dry type tinea pedis, or follicular involvement should be treated orally when possible.  Nail infections should be treated orally when possible with terbinafine or itraconazole for 6-8 weeks for finger nails and 3-4 months for toe nails. Topical treatment with amoralfine or azoles may be useful, especially if the causative agent is a non-dermatophyte mould.  Scalp infections should be treated orally if possible with griseofulvin for 6-8 weeks. Terbinafine is not yet licensed for treatment of scalp infection in children. Topical therapy is a useful adjunct to reduce possible transmission. The dermatophytes are the commonest cause of superficial skin infection of humans. Worldwide the anthropophilic species Trichophyton rubrum is responsible for the majority of infections of the body, limbs, face, hands, feet and nails, with fewer infections caused by T. interdigitale and Epidermophyon floccosum. Skin infections due to zoophilic species occur, and identification enables the source to be determined and management may include diagnosis and treatment of an animal. The commonest cause of tinea capitis in cities is now T. tonsurans, while in suburban and rural locations other species such as M. canis or M. audouinii may be more frequently isolated. This difference reflects changes in the populations and movement of people as T. soudanense and T. violaceum may also be isolated in urban areas, with the latter species commoner in children from the Indian subcontinent or East Africa. T. tonsurans is more frequently isolated from black children although children from all ethnic groups are susceptible. Dermatophytosis responds well to treatment with terbinafine, azoles or imidazoles, however, Microsporum species tend to respond poorly to terbinafine. For limited skin infections topical treatment with any of these drugs is appropriate. Nail and scalp infections should be treated systemically whenever possible as topical preparations may fail to penetrate to the focus of infection. However, topical treatment is a useful adjunct to oral therapy for tinea capitis and may help reduce transmission. For nail infections with non-dermatophyte moulds topical therapy with removal of the affected nail may be appropriate. 1. Report of the First International Meeting of the Taskforce on Onychomycosis Education. Eur Acad Derm Venereol 2005;19(supplement 1). 2. Fernandez-Torres B, Carrillo AJ, Martin E et al. In vitro activities of 10 antifungal drugs against 508 dermatophyte strains. Antimicrob Ag Chemother 2001;45:2524–2528. 3. Warshaw EM, Fett DD, Bloomfield HE et al. Pulse versus continuous for onychomycosis: a randomised, double-blind, controlled trial. J Am Acad Dermatol 2005;53:578–584. 4. Weitzman I, Summerbell RC. The dermatophytes. Clin Microbiol Rev 1995;8:240–259. 5. Fuller LC, Smith CH, Cerio R et al. A randomised comparison of 4 weeks of terbinafine vs. 8 weeks of griseofulvin for the treatment of tinea capitis. Brit J Dermatol 2001;144:321–327. 6. Fuller LC, Child FJ, Midgley G, Higgins EM. Diagnosis and management of scalp ringworm.

Diagnosis of Candida infections
Professor Frank Odds (University of Aberdeen)
No Abstracts and references submitted

An introduction to the UK Clinical Mycology Network and a Case study- a treatment dilemma
Dr Chris Kibbler (Royal Free Hospital, London)
Learning points
 There is now a national mycology qualification – the BSMM/UCL MSc/Diploma in Medical  The UK Clinical Mycology Network was established in 2005.  Mycology service provision is to be achieved through a number of regional mycology centres, supporting teaching hospital and district general hospital laboratories. The British Society for Medical Mycology set up a manpower working party in 2000 in response to the declining numbers of UK mycologists. This working party identified the need to establish a national mycology qualification and develop training networks. In parallel with this, the HPA Advisory Committee on Fungal Infection and Superficial Parasites concluded that there was a need to tackle the inconsistent and precarious mycology service within the UK by means of a mycology network. The consequence of these two initiatives has been the establishment of the BSMM/UCL MSc/Diploma in Medical Mycology, now in its second year, and the inauguration of the UK Clinical Mycology Network in 2005. This session will discuss the process by which this came about and examine the current and future roles for the Network. References 1. The Advisory Committee on Fungal Infections: 2. The UK Clinical Mycology Network Steering Group: 3. Fungal Diseases in the UK. The current provision of support for diagnosis and treatment: assessment and proposed network solution. Report of a working group of the HPA Advisory Committee for Fungal Infection and Superficial Parasites. London: Health Protection Agency April 2006.
Declaration on interest
Chair of the UKCMN Steering Group

Diagnosis of Invasive Aspergillosis
Dr Tom Harrison (St Georges Hospital Tooting)

No Abstracts and references submitted

In vitro laboratory susceptibility testing
Dr Michael Petrou (Hammersmith Hospital, London)
Learning points
 Standardised MICs Methods available for both yeasts and Filamentous Fungi.  All methods, user-friendly commercial or reference have advantages and disadvantages.  Choose the method you feel comfortable in performing and interpreting.  MICs must be performed on all isolates from sterile sites.  MICs for Haematology, Oncology, Neonates, Geriatric and ITU Patients ideal.  Correlation between MIC and clinical outcome depends on the drug and the disease. The choice of systemic antifungal drugs more than doubled in the last five years and so did the pressure to perform susceptibility on clinically important isolates. The standardised methods published by the Clinical and Laboratory Standard Institute (CLSI, formerly NCCLS) and the European Committee on Antibiotic Susceptibility Testing (EUCAST) make it easier to test any fungus deemed important.1,2 A method for dermatophytes is now going through its various phases under the auspices of the CLSI.3 The methods can be divided into agar and liquid based such as Etest and disk diffusion for the former and microdilution for the latter. Good correlation has been shown for the Etest when compared with the liquid based method and though disks exist for a number of antifungals, only fluconazole disc susceptibility is licensed (voriconazole is almost ready).4,5 Many other methods have been used and included Flow Cytometry, Viability Dyes and measurements of ergosterol, adenylates or substrate consumption. Generating MICs or sensitivity is less important than the ability to interpret its clinical meaning. Though there is clear evidence for fluconazole MICs and clinical outcome for diseases such as oesophageal Candidiasis, it is well accepted that in vitro susceptibility does not necessarily predict clinical success in vivo as in vitro resistance will nor always predict treatment failure.6 It is recommended that all isolates from sterile sites, all true pathogens, those suspected of being resistant and unusual isolates should be tested. It is advisable to test isolates from patients with treatment failure, recurrent infections and at least once the isolates from patients in intensive care, burns unit, neonates and immunocompromised patients. References 1. National Committee for Clinical and Laboratory Standards. Reference method for broth dilution antifungal susceptibility testing of yeasts. Approved standard, 2nd ed. M27-A2. National Committee for Clinical and Laboratory Standards, 2002, Wayne, Pa. 2. Cuenca-Estrella M et al. Multicenter evaluation of the reproducibility of proposed antifungal susceptibility method for fermentative yeasts of the Antifungal Susceptibility Subcommittee of the European Committee on Antimicrobial Susceptibility Testing (AFST-EUCAST). Clin Microbiol Infect 2003;9:467–474. 3. Ghannoum MA et al. Intra- and interlaboratory study of a method for testing the antifungal susceptibility of dermatophytes. J Clin. Microbiol 2004;42:2977–2979. 4. Petrou MA, Shanson DC. Susceptibility of Cryptococcus neoformans by the NCCLS microdilution and Etest methods using five defined media. J Antimicrob. Chemother 2000;46:815–818. 5. Pfaller MA et al. Results from the ARTEMIS DISK Global antifungal surveillance study: a 6.5- year analysis of the worldwide susceptibility of yeasts to fluconazole and voriconazole using standardized disk diffusion testing. J Clin Microbiol 2005;43:5848–5859. 6. Pfaller MA et al. Correlation of MIC with outcome for Candida species tested against Voriconazole: Analysis and proposal for interpretive breakpoints. J Clin Microbiol 2006;44:819–826.

Drugs: when should they be used and when shouldn’t they?
Dr Rosemary Barnes (Cardiff University)
Learning points
 Antifungal drug use can be prophylactic, pre-emptive or definitive.  Prophylaxis should be reserved for selected groups of patients following risk stratification.  There is no evidence of benefit for empirical therapy in patients on effective antifungal.  Better diagnostic techniques are allowing us to move towards a pre-emptive antifungal strategy.  The choice of effective antifungal drugs is increasing.  Minimizing drug toxicity is fundamental in determining choice of agent. With a growing number of antifungal drugs at our disposal and expenditure increasing year by year, rational usage depends on identifying patients who will benefit. Candid and Aspergillus infections predominate in the UK. Severely immunocompromised patients with haematological malignancy and those undergoing stem cell and organ transplantation are most at risk. However, candidal infections are most commonly seen in critically ill patients in the Intensive care and surgical wards1 highlighting the proven efficacy of azole prophylaxis in haematology and transplant patients2. Within the ICU it is possible to identify groups who would benefit from prophylaxis3 but widespread usage is likely to drive antifungal resistance and pathogen shifts. Improved diagnostic criteria4 and standardisation of sensitive molecular techniques5 can identify patients with markers of infection at an earlier stage allowing us to move pre-emptive therapy. For definitive therapy of proven invasive disease there is now a broad range of effective agents including polyenes, triazoles and echinicandins. Conventional amphotericin B is associated with significant toxicity which contributes to increased morbidity and mortality and this agent should be avoided. Lipid-based alternatives may reduce the incidence of (but not eliminate) these adverse events6, whilst the newer triazoles and echinocandins offer rational choices. References 1. Kibbler CC, Ainscough S, Barnes RA. Management and outcome of blood stream infections due to Candida species in England and Wales. J Hosp Infect 2003;54:18–24. 2. Marr KA, Crippa F, Leisenring W, et al. Itraconazole vs. Fluconazole for antifungal prophylaxis in allogeneic HSCT patients. Blood 2004;103:1527–1533. 3. Sinha J, Barnes RA. Fungal infections in critical care: the appropriate use of antifungal agents. Brit J Intens Care Med 2003;89–98. 4. Ascioglu S, Rex JH, de Pauw B, et al. Defining opportunistic invasive fungal infections in immunocompromised patients with cancer and haematopoietic stem cell transplants: an international consensus. Clin Infect Dis 2002;34:7–14. 5. White PL, Barton R, Guiver M, et al. A consensus on fungal PCR diagnosis – A UK-Ireland evaluation of PCR methods for the detection of systemic fungal infections. J Mol Diagnost 2006;8. 6. Ullman AJ, Sanz MA, Tramarin, et al. Prospective study of Amphotericin B formulations in immunocompromised patients in four Europeans countries. Clin Infect Dis 2006 In press.

Source: https://www.rcpath.org/Resources/RCPath/Migrated%20Resources/Documents/A/AMM_abs12Jul2006.pdf