Immunocompromised hosts are at special risk for invasive fungal infections (IFI). Traditional methods of diagnosis (e.g. culture and molecular methods) largely suffer from poor sensitivity, thus limiting their clinical utility. Enzyme immunoassay-based detection of fungal antigens represents an attractive, supplementary method for IFI identification and is the focus of this review.

by Phillip R. Heaton and Elitza S. Theel

Background
Invasive fungal infections (IFI) are a significant cause of morbidity and mortality in patients with hematologic malignancies and in hematopoietic stem cell transplant (HSCT) recipients. Although Aspergillus species and Candida albicans are among the most common agents of IFI, an increasing incidence of IFI due to other filamentous fungi (e.g. Fusarium, Zygomycetes) and non-albicans Candida (e.g. C. tropicalis, C. krusei, C. glabrata) has been reported. Currently, IFI diagnosis is based on clinical evaluation, radiologic imaging (both of which may lack clinical specificity) and culture-based laboratory findings. Unfortunately, culture of the aforementioned fungi from bronchoalveolar lavage fluid (BAL) and blood, the most commonly collected specimens in suspected IFI cases, suffers from poor sensitivity: only 45 to 60% of BAL specimens and up to 50% of blood cultures yield fungal growth [1]. Additionally, as some fungi are common in the environment (e.g. Aspergillus), providers must determine whether growth from BAL cultures is indicative of invasive disease or colonization of the respiratory tract. Finally, procedures to collect alternative specimens, including lung tissue, are often contra-indicated due to the critical state of the patient. These limitations have led to a clinical need for alternative methods to identify IFI – techniques independent of culture, which are both sensitive and specific. This demand has driven the development of novel assays to detect fungal biomarkers including the Aspergillus galactomannan (GM) antigen, the (1→3) β-D-glucan (BDG) polysaccharide common to many fungi, and the Candida mannan antigen (Mn-A). This brief review will discuss the clinical utility, advantages and limitations of GM, BDG and Mn-A detection assays in patients at risk for IFI.

Detection of the Aspergillus galactomannan antigen
Galactomannan, composed of a mannan core and highly immunogenic galactofuranosyl side chains, is a dominant cell wall component present in the majority of clinically relevant Aspergillus species and is released during hyphal growth into surrounding tissue (Fig. 1). Currently, the Platelia Aspergillus antigen (Bio-Rad, Marnes-la-Coquette, France) enzyme immunoassay (EIA) is the only FDA approved assay for GM detection of in serum and BAL fluid, though other kits are also available (e.g. Pastorex kit, Sanofi Diagnostics, Pasteur, Marnes-La-Coquette, France). The Platelia EIA is a quantitative assay with GM levels ≥0.5 ng/mL considered as positive. The presence of GM in patient specimens can be used as an aid, alongside other clinical studies, to specifically detect invasive aspergillosis (IA), a potentially devastating condition encountered in 5–20% of HSCT patients [2]. The performance characteristics of the Platelia GM assay have been widely evaluated with overall favorable outcomes. Briefly, one study reported a clinical sensitivity and specificity of 94.4% and 98.8% respectively, from serum of HSCT patients with proven or probable IA [as defined by the European Organization for Research and Treatment of Cancer (EORTC) criteria], with similar positive and negative predictive values [3]. While a subsequent meta-analysis of GM studies found a significantly lower sensitivity in this patient population (58%), specificity remained comparable at 95% [4]. Notably, these results are in stark contrast to the sensitivity of this assay in other immunocompromised (ICH) patient populations, specifically in solid organ transplant (SOT) recipients, where sensitivity can be as low as 22–41% [4]. Additionally, while the kinetics of GM clearance are not yet well defined, serial testing and trending of GM levels following initiation of antifungal therapy has been shown to correlate well with patient outcome. Specifically, while persistently elevated GM levels were associated with treatment failure, a decrease of GM levels by ≥35% between baseline and week one of antifungal treatment was associated with clinical improvement [5].

Despite the advantage of rapid GM testing in serum, a readily available specimen source, and the potential to monitor response to therapy, a number of limitations affecting assay specificity have been described. First, false-positive GM reactions have been associated with prior (<12 hours) administration of piperacillin/tazobactam, a fungal-derived antibiotic [6]. Recently, however, Mikulska and colleagues demonstrated negligible GM levels in patients on piperacillin/tazobactam therapy, suggesting that modern day manufacturing practices may have improved antibiotic purity [7]. Nonspecific reactions have also been noted in patients with non-Aspergillus IFI, including Fusarium and Penicillium (an exceedingly rare agent of IFI which also expresses GM) species infections and in individuals with either graft versus host disease (GVHD) or a damaged intestinal wall through which GM from food products can translocate [8].

Detection of β-D-glucan, a pan-fungal biomarker
(1→3)-β-D-glucan (BDG) is an abundant cell wall polysaccharide found in most fungi with the exception of Cryptococcus species, the Zygomycetes and Blastomyces dermatitidis (Fig. 2). The most commonly used BDG detection method, the Fungitell assay (Associates of Cape Cod, East Falmouth, MA), is a quantitative EIA (values ≥60 pg/mL considered positive) which detects BDG in serum using a modified version of the Limulus (horseshoe crab) clotting cascade. As a pan-fungal biomarker, BDG detection in patients with hematologic malignancies and HSCT recipients has been associated with high clinical specificity (76–99%) and negative predictive values (87–96%) for the presence of proven or probable IFI [9]. Similar to the GM assay however, inclusion of other ICH groups (e.g. SOT patients) dramatically lowers the performance characteristics of this assay. Interestingly, regardless of the patient population, the associated clinical sensitivity and positive predictive value of the BDG assay are generally poor (range 38 – 80%), collectively indicating that a single, negative BDG result should not be used to exclude the diagnosis of IFI [9]. Serial BDG testing, however, can significantly improve the clinical sensitivity of this assay and trending BDG levels during antifungal therapy has some prognostic value with respect to treatment failure or response, particularly in patients with disseminated candidiasis [9, 10]. Furthermore, BDG was shown to be detectable in critically ill patients prior to development of clinical symptoms, radiological findings or culture positivity, suggesting that in patients at increased risk for IFI, the presence of BDG should warrant further evaluation to identify an infectious process [11]. Finally, in patients with Pneumocystis jirovecii pneumonia (PjP), for whom invasive BAL or biopsy procedures are often precluded due to safety concerns, the demonstration of elevated BDG levels has been associated with high clinical sensitivity (>95%) [12]. Though this data is encouraging, especially in light of the limited sensitivity of current diagnostic methods to detect P. jirovecii, due to the pan-fungal nature of BDG, a positive result cannot be used to diagnose PjP pneumonia; a negative BDG finding can, however, be used to potentially exclude P. jirovecii as the causative agent.

The greatest limitation of BDG assays is their poor specificity. Many studies have now documented the generation of false-positive results in patients who received or have been exposed to albumin, intravenous immunoglobulin, amoxicillin-clavulanic acid, gauze during surgery, or cellulose based filters during dialysis. Additionally, infection with certain bacterial agents, including Alcaligenes faecalis, can also lead to false-positive results. Therefore, providers using these assays must confidently exclude these confounding factors prior to interpreting BDG results.

Detection of the Candida mannan antigen   
The Candida Mn-A is an oligomannan cell wall component which can be detected by multiple quantitative EIAs (Fig. 3). Currently, the Platelia Candida Ag Plus quantitative EIA (Bio-Rad) is most commonly used. A recent meta-analysis of 14 studies evaluating the utility of Mn-A detection found significant heterogeneity in clinical sensitivity for detection of invasive candidiasis (IC), which, interestingly, appeared to be species dependent. For example, among patients with disseminated C. albicans, C. glabrata or C. tropicalis, the sensitivities ranged from 58–70%, whereas for patients with invasive C. parapsilossis and C. krusei, sensitivity of the assay ranged between 25–30% [13]. Importantly, however, despite the low sensitivity of the assay, the majority of Mn-A positive patients were subsequently confirmed as culture positive, suggesting the utility of this assay as an early diagnostic marker in at risk patients. Notably, the specificity of this assay is high (>90%) with cross-reactivity reported in patients with Geotrichum or Fusarium species infections, both fairly uncommon [14]. Due to the described performance variability and the short duration of Mn-A circulation, many authors have suggested combination testing with anti-Mn antibodies (Platelia Candida Ab Plus, Bio-Rad), which are detectable in at risk patients >10 days prior to proven candidemia [15]. One study evaluated neutropenic patients using combination testing and found that Mn-A/anti-Mn outperformed traditional diagnostic methods (cultures, radiology, and histopathology) for detection of IC with a sensitivity and specificity of 89% and 84%, respectively [15]. Based on these and other studies, current ECIL-3 recommendations support using a combination of Mn-A and antibody testing as an aid to detect IC [13].

Conclusions
The diagnosis of IFI in ICHs remains a challenge, and despite the limited sensitivity and specificity of the various fungal antigen detection assays, in 2008 the EORTC included detection of GM and BDG as supportive evidence for proven or probable IFI in specific patient populations [3]. When used appropriately (i.e. serial testing of high risk patients) and by providers knowledgeable of the associated limitations, antigen detection can be crucial marker for the identification of IFI. Future advancement of IFI diagnostics lies in the molecular arena and real-time polymerase chain reaction (RT-PCR) assays to detect fungal nucleic acid.
 
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2. Tamma P. The Galactomannan antigen assay. Pediatr Infect Dis J. 2007; 26: 641-642 610.1097/INF.1090b1013e318070c318525.
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4. Pfeiffer CD, Fine JP, Safdar N. Diagnosis of invasive aspergillosis using a galactomannan assay: a meta-analysis. Clin Infect Dis. 2006; 42: 1417–1727.
5. Chai LY, Kullberg BJ, Johnson EM, Teerenstra S, Khin LW, Vonk AG, Maertens J, Lortholary O, Donnelly PJ, Schlamm HT, Troke PF, Netea MG, Herbrecht R. Early serum galactomannan trend as a predictor of outcome of invasive aspergillosis. J Clin Microbiol. 2012; 50: 2330–2336.
6. Machetti M, Majabo MJ, Furfaro E, Solari N, Novelli A, Cafiero F, Viscoli C. Kinetics of galactomannan in surgical patients receiving perioperative piperacillin/tazobactam prophylaxis. J Antimicrob Chemother. 2006; 58: 806–810.
7. Mikulska M, Furfaro E, Del Bono V, Raiola AM, Ratto S, Bacigalupo A, Viscoli C. Piperacillin/tazobactam (TazocinTM) seems to be no longer responsible for false-positive results of the galactomannan assay.
J Antimicrob Chemother. 2012; 67: 1746–1748.
8. Mennink-Kersten MASH, Donnelly JP, Verweij PE. Detection of circulating galactomannan for the diagnosis and management of invasive aspergillosis. Lancet Infect Dis. 2004; 4: 349–357.
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10. Jaijakul S, Vazquez JA, Swanson RN, Ostrosky-Zeichner L. (1,3)-β-D-Glucan as a prognostic marker of treatment response in invasive candidiasis. Clin Infect Dis. 2012; 55: 521–526.
11. Odabasi Z, Mattiuzzi G, Estey E, Kantarjian H, Saeki F, Ridge RJ, Ketchum PA, Finkelman MA, Rex JH, Ostrosky-Zeichner L. Beta-D-glucan as a diagnostic adjunct for invasive fungal infections: validation, cutoff development, and performance in patients with acute myelogenous leukemia and myelodysplastic syndrome. Clin Infect Dis. 2004; 39: 199–205.
12. Karageorgopoulos DE, Qu JM, Korbila IP, Zhu YG, Vasileiou VA, Falagas ME. Accuracy of β-D-glucan for the diagnosis of Pneumocystis jirovecii pneumonia: a meta-analysis. Clin Microbiol Infect. 2013; 19: 39–49.
13. Marchetti O, Lamoth F, Mikulska M, Viscoli C, Verweij P, Bretagne S. ECIL recommendations for the use of biological markers for the diagnosis of invasive fungal diseases in leukemic patients and hematopoietic SCT recipients. Bone Marrow Transplant. 2012; 47: 846–854.
14. Rimek D, Singh J, Kappe R. Cross-reactivity of the PLATELIA CANDIDA antigen detection enzyme immunoassay with fungal antigen extracts. J Clin Microbiol. 2003; 41: 3395–3398.
15. Prella M, Bille J, Pugnale M, Duvoisin B, Cavassini M, Calandra T, Marchetti O. Early diagnosis of invasive candidiasis with mannan antigenemia and antimannan antibodies. Diagn Microbiol Infect Dis. 2005; 51: 95–101.
 
The authors
Phillip R. Heaton PhD and Elitza S. Theel PhD*
Division of Clinical Microbiology,
Department of Laboratory Medicine and Pathology, Mayo Clinic, Rochester,
Minnesota, USA

*Corresponding author
E-mail: theel.elitza@mayo.edu

Whipple´s disease, a systemic and ultimately fatal infection with Tropheryma whipplei, is usually easily treated if diagnosed early enough. A novel real-time PCR protocol and fluorescence in situ hybridization provide an improved diagnosis. All results, however, need careful interpretation. We recommend involving specialized centres in the initial diagnosis and patient follow-up.

by Alexandra Wießner, Dr Annette Moter and Dr Judith Kikhney

Whipple’s disease: a fatal infectious disease
Tropheryma whipplei causes a rare, but fatal, bacterial infection: Whipple’s disease. This systemic disease can usually be cured by antibiotic therapy if detected early enough. The key challenge for physicians and microbiologists is to recognize the bacterial origin of various clinical symptoms in time. Diagnosis is still not trivial owing to the rarity of the disease, diverse and unspecific clinical symptoms, the fastidious nature of T. whipplei and the absence of non-invasive serological tests [1]. Improved diagnostic assays for the detection of T. whipplei are very valuable in combination with expertise to interpret the results for fast initiation of treatment.

T. whipplei belongs to the Gram-positive class of Actinobacteria and can be detected intracellularly in vacuoles or as extracellular bacteria in the tissue [2]. It is a slender rod shape, readily visible with Periodic Acid–Schiff (PAS) staining. T. whipplei strains can be cultured in an axenic culture medium supplemented with amino acids, but the slow growth rate means that culture is not an option for routine diagnosis of T. whipplei infection.

Symptoms of Whipple’s disease
In classical Whipple’s disease patients suffer from chronic diarrhoea, weight loss and fever. Molecular methods have detected isolated or systemic T. whipplei infection in almost every organ [joints, central nervous system (CNS), heart valves, skin, eye, lymph node, bone and lung], even in the absence of intestinal involvement. Depending on the location of infection, the symptoms may vary substantially. Often, the diagnosis of Whipple’s disease is delayed as the result of misdiagnosis as sero-negative rheumatoid arthritis, culture-negative endocarditis or neurological disorders. The involvement of the CNS is especially dramatic, as damage caused by the bacteria is often irreversible and antibiotic treatment may no longer be effective enough to cure the infection [3].

Transmission and asymptomatic carriage
To complicate the picture even more T. whipplei has been found in healthy carriers at an estimated prevalence in the population of <1–4% [4, 5]. This means that the detection of T. whipplei in stool or saliva may not be indicative of Whipple’s disease and in this case does not necessarily require antibiotic treatment. A higher prevalence has been found in high risk populations for direct or indirect faecal–oral transmission, such as sewage workers [5], homeless people and family members of Whipple´s disease patients. As T. whipplei is common in the environment, it is assumed that Whipple’s disease patients must have an immunological predisposition for developing a chronic infection instead of being only transiently colonized [6].

The current transmission model assumes that T. whipplei is taken up orally, probably in early childhood, leading to temporary asymptomatic carriage, self-limiting gastroenteritis, fever, or cough [1, 7, 8]. In most cases a protective humoral and cellular immune response prevents T. whipplei infection. However, in predisposed persons T. whipplei may spread systemically over the years resulting in Whipple´s disease.

Diagnosis of Whipple’s disease
Currently, Whipple’s disease is most often detected through PAS staining of biopsies from the lower duodenum or jejunum showing PAS-positive macrophages in the lamina propria. However, PAS staining can give false-positive results because of other infections, for example with nontuberculous mycobacteria, and also false-negative results because of low bacterial load [9]. Therefore, every positive PAS result should be confirmed by an independent method. Here, molecular techniques such as PCR are, so far, irreplaceable for providing a direct, valid species diagnosis. Several in-house PCR protocols are now successfully used to detect T. whipplei DNA [10, 11]. In patients without gastrointestinal manifestation of classical Whipple’s disease, sample specimens from the clinically affected organs, e.g. heart valves, lymph nodes, synovial tissue, cerebrospinal fluid (CSF) or brain biopsies, may be PAS-positive, whereas duodenal biopsies remain negative [1]. PCR was suggested for screening stool and saliva samples as the prevalence and load of T. whipplei is far higher in Whipple´s disease patients than in healthy controls [4]. Here, however, positive PCR results are no proof of infection compared to the direct detection of T. whipplei DNA in affected organs. Analysis of peripheral blood is also possible, but a negative PCR result will not rule out infection [12]. As with all PCR assays, results need to be carefully interpreted as the assay is prone to laboratory contamination (especially nested PCR protocols) or false-positive results because of nonspecific reaction conditions or primer design. Importantly, some positive PCR results in the past have been shown to be due to cross-reactivity, e.g. with Actinomyces odontolyticus [13].

Improved diagnostics of T. whipplei
A break-through for the diagnosis of Whipple’s disease that is specific and less prone to contamination is modern real-time PCR [5, 14]. We evaluated a real-time PCR assay targeting T. whipplei-specific segments within the rpoB gene on test strains and over 1000 clinical specimens in a national reference laboratory [14]. This assay proved to be specific, sensitive and substantially faster than a conventional in-house assay. The protocol includes two specific hybridization probes and, to our knowledge for the first time in T. whipplei diagnostics, a melting curve analysis. Both are crucial for the robustness and reliability of the assay. This applies especially to polymicrobial samples, such as saliva or stool, which contain numerous uncultured bacterial species with unknown DNA sequences. Here, the problem of unexpected probe binding with false-positive results remains and, therefore, PCR results should always be interpreted in the context of clinical and histopathological findings. An initial diagnosis of Whipple´s disease should not rely on only one isolated PCR result, and a confirmatory PCR (using a different target sequence, sequence analysis of ribosomal RNA sequence or genotyping PCR) is mandatory. In inconclusive cases a second PCR with an independent sample specimen is recommended.

Emerging techniques for the detection of T. whipplei
Besides PCR and PAS staining, additional methods such as immunohistochemistry or fluorescence in situ hybridization (FISH) are offered by specialized laboratories. These techniques are, as yet, not part of the routine work-up but provide promising insights. FISH uses fluorescently labelled probes that hybridize specifically with their target sequence in the intact bacterial cells (usually the 16S rRNA). Thus, FISH not only provides direct identification of T. whipplei but also visualizes the pathogen directly in the tissue context. Surprisingly, we found T. whipplei to be by far the most abundant cause of culture-negative endocarditis among the rare pathogens [15]. FISH revealed impressive infected areas in heart valves densely scattered with T. whipplei. In gut biopsies FISH reveals the amount and localization of single microorganisms in the tissue (Fig. 1). As with all microscopic techniques, however, FISH is less sensitive than PCR and will only give information on post-operatively obtained tissue and exclusively on the section investigated. Thus, a low bacterial load in the tissue might be missed. However, FISH is so far the only method bridging the gap between specific molecular biology and histopathology and, thus, might find broader application in the future.

Sampling for T. whipplei
Tissue specimens, such as small bowel biopsies in classical Whipple´s disease or samples of the affected organ in isolated T. whipplei infection, should be examined by PAS staining and PCR (Fig. 2). In the event of positive results, CSF should be tested by PCR to check for CNS involvement. For isolated T. whipplei infections gastrointestinal involvement should be controlled as well. Fluid samples, such as CSF, etc., should be examined by PCR.

For histological examination, PAS staining and FISH the samples should be fixed in 10% formalin and transported at room temperature. For PCR the samples need to be native (no formalin pre-treatment!) and can be transported at room temperature within one day. Samples can be stored for a few days at 4°C and should be kept at –80°C for long-term storage.

Conclusions
The recent development of real-time PCR protocols with hybridization probes for the specific detection of T. whipplei provides accurate and fast results in the challenging clinical situation of Whipple´s disease. However, due to the variety of clinical symptoms, asymptomatic carriage, isolated and systemic infection, as well as false positive and negative results all examinations need careful interpretation in specialized centres. Clinical and histopathological facts always have to be taken into account. Emerging techniques such as FISH might in the future close the gap between molecular biology and histopathology. Together clinical and microbiological expertise are the key to the fast and successful treatment of Whipple´s disease. Similarly, after initial diagnosis and initiation of treatment, it is highly recommended to follow each patient in specialized centres during and after antibiosis to keep relapses at bay.

Acknowledgements
We thank the Robert Koch Institute for its continuous support.

Funding Sources
This study was supported by the Robert Koch Institute (RKI). The epifluorescence microscope was a gift from the Sonnenfeld-Stiftung.

References
1. Moos V, Schneider T. Changing paradigms in Whipple’s disease and infection with Tropheryma whipplei. Eur J Clin Microbiol Infect Dis. 2011; 30: 1151–1158.
2. Raoult D, Birg ML, La Scola B, Fournier PE, Enea M, Lepidi H, et al. Cultivation of the bacillus of Whipple’s disease. N Engl J Med. 2000; 342: 620–625.
3. Lagier JC, Lepidi H, Raoult D, Fenollar F. Systemic Tropheryma whipplei: clinical presentation of 142 patients with infections diagnosed or confirmed in a reference center. Medicine 2010; 89: 337–345.
4. Fenollar F, Laouira S, Lepidi H, Rolain JM, Raoult D. Value of Tropheryma whipplei quantitative polymerase chain reaction assay for the diagnosis of Whipple disease: usefulness of saliva and stool specimens for first-line screening. Clin Infect Dis. 2008; 47: 659–667.
5. Fenollar F, Trani M, Davoust B, Salle B, Birg ML, Rolain JM et al. Prevalence of asymptomatic Tropheryma whipplei carriage among humans and nonhuman primates. J Infect Dis. 2008; 197: 880–887.
6. Martinetti M, Biagi F, Badulli C, Feurle GE, Müller C, Moos V et al.  The HLA Alleles DRB1*13 and DQB1*06 Are Associated to Whipple’s Disease. Gastroenterology 2009; 136: 2289–2294.
7. Moos V, Schneider T. The role of T cells in the pathogenesis of classical Whipple’s disease. Expert Rev Anti Infect Ther. 2012; 10: 253–255.
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9. Müller SA, Vogt P, Altwegg M, Seebach JD. Deadly carousel or difficult interpretation of new diagnostic tools for Whipple’s disease: case report and review of the literature. Infection 2005; 33: 39–42.
10. Hinrikson HP, Dutly F, Nair S, Altwegg M. Detection of three different types of ‘Tropheryma whippelii’ directly from clinical specimens by sequencing, single-strand conformation polymorphism (SSCP) analysis and type-specific PCR of their 16S-23S ribosomal intergenic spacer region. Int J Syst Bacteriol. 1999; 49: 1701–1706.
11. Relman DA, Lepp PW, Sadler KN, Schmidt TM. Phylogenetic relationships among the agent of bacillary angiomatosis, Bartonella bacilliformis, and other alpha-proteobacteria. Mol Microbiol. 1992; 6: 1801–1807.
12. Marth T, Fredericks D, Strober W, Relman DA. Limited role for PCR-based diagnosis of Whipple’s disease from peripheral blood mononuclear cells. Lancet 1996; 348: 66–67.
13. Rolain JM, Fenollar F, Raoult D. False positive PCR detection of Tropheryma whipplei in the saliva of healthy people. BMC Microbiol. 2007; 7: 48.
14. Moter A, Schmiedel D, Petrich A, Wiessner A, Kikhney J, Schneider T et al. Validation of an rpoB gene PCR assay for detection of Tropheryma whipplei: 10 years’ experience in a National Reference Laboratory. J Clin Microbiol. 2013; 51: 3858–3861.
15. Geißdörfer W, Moos V, Moter A, Loddenkemper C, Jansen A, Tandler R et al. High frequency of Tropheryma whipplei in culture-negative endocarditis. J Clin Microbiol. 2012; 50: 216–222.
16. Mallmann C, Siemoneit S, Schmiedel D, Petrich A, Gescher DM, Halle E et al. Fluorescence in situ hybridization to improve the diagnosis of endocarditis: a pilot study. Clin Microbiol Infect. 2010; 16: 767–773.
 
The authors
Alexandra Wießner¹, Annette Moter¹* MD, Judith Kikhney^1,2 PhD
¹ Center for Biofilms and Infection, German Heart Institute Berlin, Berlin, Germany
² Institut für Mikrobiologie und Hygiene, Charité University medicine Berlin, Berlin, Germany

*Corresponding author
E-mail: moter@dhzb.de

Clostridium difficile is a major cause of nosocomial infections and rapid diagnosis of the disease is essential for infection control. Several methods for C. difficile detection are employed in clinical laboratories; each method has its advantages and disadvantages. A novel method has recently been developed that allows differentiation between C. difficile-positive and -negative stool samples based on volatile organic compound evolution and their detection by headspace solid-phase microextraction gas chromatography–mass spectrometry.

by Dr Emma Tait, Prof. Stephen P. Stanforth, Prof John D. Perry and Prof. John R. Dean

Introduction
Clostridium difficile is a Gram-positive anaerobe and the causative agent of C. difficile infection (CDI). CDI is a major healthcare problem with a total of 14,687 cases reported in patients aged 2 years and over in England between April 2012 and March 2013 [1]. C. difficile is a spore-forming bacterium; dormant spores are resistant to antibiotics, heating and chemicals such as disinfectants and, therefore, can persist on surfaces and survive for long periods in the environment [2]. Ingestion of spores and their subsequent germination in the gut allows the proliferation of C. difficile in patients whose normal gut flora has been severely reduced following antibiotic treatment. Following germination, vegetative C. difficile can produce toxins and is susceptible to antibiotic treatment. Pathogenic C. difficile releases two types of toxins, toxin A and toxin B, and it is these toxins that cause the symptoms associated with CDI [3]. Clinical symptoms of C. difficile infection (CDI) include mild to severe diarrhoea, which can lead to pseudomembranous colitis and death. Diagnosis of CDI includes both clinical manifestations of symptoms supported by laboratory findings.

Diagnosis of C. difficile infection
Rapid diagnosis of CDI is essential to allow the most appropriate treatment to be prescribed, to enable proper use of hospital isolation facilities and to reduce the spread of the infection. Routine diagnostic methods in clinical microbiology laboratories employ a variety of techniques for diagnosis of CDI. These include immunoassays for detection of glutamate dehydrogenase (GDH) antigen and toxins, polymerase chain reaction (PCR) for detection of toxin B or isolation of C. difficile by culture (followed by confirmation of toxigenicity, e.g. using PCR). Immunoassays and PCR methods are typically highly automated and deliver rapid results and these have largely replaced the traditional cell cytotoxicity assay, which requires propagation of cell lines and takes days rather than hours to provide results.

Toxin immunoassays, although relatively inexpensive with rapid turnaround time, have limitations in terms of their sensitivity and specificity leading to false-negative results and false-positive results [4]. Immunoassays for GDH are typically highly sensitive for detection of C. difficile but lack specificity. Culture media for the isolation of C. difficile typically incorporate the antibiotics D-cycloserine and cefoxitin which suppress commensal bacteria; such media are based on the formulation recommended by George et al. [5]. Isolation of C. difficile via culture is sensitive but can take several days to obtain results and there may be a heavy growth of other fecal bacteria, particularly when reduced antibiotic concentrations are used [6]. Recent developments in C. difficile detection include using a chromogenic substrate which is structurally similar to naturally occurring substrate used by C. difficile toxins [7]. This allows the detection of toxigenic C. difficile, i.e. only pathogenic strains are targeted.

Identification of C. difficile-positive stool samples using gas chromatography has previously been explored [8]; volatile organic compounds (VOCs) such as p-cresol and short chain fatty acids were identified as potential markers for C. difficile. However, these methods suffered from a lack of specificity, particularly due to the high number of false positives obtained following the detection of p-cresol and isocaproic acid in stool samples without C. difficile. As a consequence, gas chromatography methods were deemed unsuitable for C. difficile detection [8]. More recent attempts to use bacterial VOC analysis as a tool for C. difficile identification have used headspace solid-phase microextraction (HS-SPME) as a VOC collection method coupled with gas chromatography–mass spectrometry (GC-MS) for VOC separation and detection [9].

Development of a novel method for detection of C. difficile in stools
A novel method for rapid identification of C. difficile in stool samples has been developed using the analysis of VOCs. Use of synthetic enzyme substrates is an effective means of differentiating bacteria, for example in chromogenic culture media [10]. These types of culture media incorporate chromogenic enzyme substrates where the action of a specific enzyme on the substrate liberates a molecule that is detectable visually, allowing the detection of pathogenic bacteria [10]. The philosophy behind the use of substrates in culture media can be applied to the analysis of bacterial VOCs, where a substrate is incorporated into a clinical sample inoculated in liquid media; the cleaved product is volatile and detectable using an analytical method such as HS-SPME-GC-MS. The detection of VOCs liberated following enzyme activity increases the specificity of bacterial VOC profiles, as these liberated VOCs act as markers for a particular species, hence aiding identification of bacteria. This approach was applied to the detection of C. difficile in stool samples.

p-Cresol is formed in C. difficile by the decarboxylation of p-hydroxyphenylacetic acid. The enzyme responsible for this decarboxylation is p-hydroxyphenylacetate decarboxylase. It has been established that the hydroxyl group in the para position on the phenyl ring is an essential requirement for decarboxylation to occur [11]. C. difficile is almost unique in its ability to form p-cresol using this pathway, with the exception of a Lactobacillus strain [12]. This was exploited in the development of the novel method that allows successful differentiation between C. difficile culture-positive and -negative stool samples based on VOC generation from an enzyme substrate [13]. 3-Fluoro-4-hydroxyphenylacetic acid was used as a substrate for p-hydroxyphenylacetate decarboxylase; the evolution of the VOC 2-fluoro-4-methylphenol indicated the presence of C. difficile (Fig. 1). VOCs were detected using HS-SPME-GC-MS.
 
The lack of specificity of previous GC methods was often due to the detection of VOCs in C. difficile culture-negative stool samples as these VOCs were generated by commensal bacteria. Techniques employed to reduce background flora, and therefore improve the selectivity and sensitivity of methods, include alcohol shock [14] and the inclusion of antibiotics [5]. The antibiotics D-cycloserine, cefoxitin and amphotericin were added to the sample matrix and an alcohol-shock step was included to suppress background flora present in stool samples. Alcohol shock kills vegetative cells but does not affect the viability of spores and incorporation of sodium taurocholate in a culture medium can subsequently aid germination of spores [6, 15]. Inoculation of stool samples into such a culture medium allows bacterial growth and concomitant generation of VOCs.

The method was tested with 100 stool samples, of which 77 were C. difficile culture-positive and 23 culture-negative. The generation of 2-fluoro-4-methylphenol indicated the presence of C. difficile after overnight incubation. Method specificity and sensitivity were 100 % and 83.1 %, respectively, using 2-fluoro-4-methylphenol as a marker for C. difficile identification (Table 1). The VOCs isocaproic acid and p-cresol were useful indicators for C. difficile-positive stool samples, although were insufficient for identification purposes. Both VOCs, particularly p-cresol, were generated by C. difficile-negative samples; this is in agreement with previous studies [8].
 
Advantages and disadvantages of VOC method
The method allows the detection of C. difficile with a very high specificity (100%), i.e. 2-fluoro-4-methylphenol was not generated by C. difficile culture-negative stool samples tested. Rapid detection of VOCs was possible with confirmation of the presence of C. difficile within 18 hours. This indicates that the method could be used to screen for C. difficile in stools allowing the prompt diagnosis of culture-positive samples by the detection of 2-fluoro-4-methylphenol. However, a study on method sensitivity in terms of the number of bacterial cells required to generate a positive signal confirmed that identification of C. difficile was possible provided the stool sample contained at least 150 colony forming units (CFU). It is entirely possible that some stool samples will contain much fewer CFU and therefore 2-fluoro-4-methylphenol would not be detected and a false-negative result would be obtained. This limitation is reflected in the method sensitivity (83.1%) after evaluation with 100 stool samples. The method targets all strains of C. difficile and further testing would be required (e.g. using PCR or immunoassay) to distinguish whether positive stool samples contain toxigenic strains. As a result, it is recommended that VOC analysis should be used alongside conventional methods for C. difficile detection, including toxin detection methods, which would allow any false negative results to be eliminated.

Conclusion
C. difficile is a common cause of nosocomial infections and therefore rapid, accurate diagnosis of CDI is of extreme importance for infection control and patient care. There are currently a number of methods used in hospital laboratories for the diagnosis of CDI; however, each method has its drawbacks. A novel approach has been developed for the identification of C. difficile in stool samples that involves the incubation of stool samples in the presence of 3-fluoro-4-hydroxyphenylacetic acid which acts as a substrate for the enzyme p-hydroxyphenylacetate decarboxylase. The success of this new approach is evaluated by its application to 100 stool samples and its ability to differentiate between C. difficile culture-positive and -negative stool samples. It is envisaged that the identification of C. difficile culture-positive stool samples by the analysis of VOCs could allow rapid diagnosis of CDI. In addition, the novel approach of using enzyme substrates that release VOCs that are not normally generated by bacteria, for example fluorinated VOCs, may find application in the identification of other bacterial pathogens in clinical microbiology.

References
1. Public Health England. Summary Points on Clostridium difficile Infection (CDI). 2013; http://www.hpa.org.uk/webc/HPAwebFile/HPAweb_C/1278944283388.
2. Kuijper EJ, Coignard B, Tull P. Emergence of Clostridium difficile-associated disease in North America and Europe. Clin Microbiol Infect. 2006; 12(Suppl 6): S2−S18.
3. Voth DE, Ballard JD. Clostridium difficile toxins: mechanism of action and role in disease. Clin Microbiol Rev. 2005; 18: 247–263.
4. Eastwood K, Else P, Charlett A, Wilcox M. Comparison of nine commercially available Clostridium difficile toxin detection assays, a real-time PCR assay for C. difficile tcdB, and a glutamate dehydrogenase detection assay to cytotoxin testing and cytotoxigenic culture methods. J Clin Microbiol. 2009; 47: 3211–3217.
5. George WL, Sutter VL, Citron D, Finegold SM. Selective and differential medium for isolation of Clostridium difficile. J Clin Microbiol. 1979; 9: 214–219.
6. Nerandzic MM, Donskey CJ. Effective and reduced-cost modified selective medium for isolation of Clostridium difficile. J Clin Microbiol. 2009; 47: 397–400.
7. Darkoh C, Kaplan HB, DuPont HL. Harnessing the glucosyltransferase activities of Clostridium difficile for functional studies of toxins A and B. J Clin Microbiol. 2011; 49: 2933–2941.
8. Levett PN. Detection of Clostridium difficile in faeces by direct gas liquid chromatography. J Clin Pathol. 1987; 37: 117–119.
9. Garner CE, Smith S, Costello BL, White P, Spencer R, Probert CSJ, Ratcliffe NM. Volatile organic compounds from feces and their potential for diagnosis of gastrointestinal disease. Faseb J. 2007; 21: 1675–1688.
10. Orenga S, James AL, Manafi M, Perry JD, Pincus DH. Enzymatic substrates in microbiology. J Microbiol Meth. 2009; 79: 139–155.
11. Selmer T, Andrei PI. p-Hydroxyphenylacetate decarboxylase from Clostridium difficile. A novel glycyl radical enzyme catalysing the formation of p-cresol. Eur J Biochem. 2001; 268: 1363–1372.
12. Yokoyama MT, Carlson JR. Production of skatole and para-cresol by a rumen Lactobacillus sp. Appl Environ Microbiol. 1981; 41; 71–76.
13. Tait E, Hill KA, Perry JD, Stanforth SP, Dean JR. Development of a novel method for detection of Clostridium difficile using HS-SPME-GC-MS. J Appl Microbiol. DOI: 10.1111/jam.12418.
14. Clabots CR, Gerding SJ, Olson MM, Peterson LR, Gerding DN. Detection of asymptomatic Clostridium difficile carriage by an alcohol shock procedure. J Clin Microbiol. 1989; 27: 3286–3287.
15. Wilson KH, Kennedy MJ, Fekety FR. Use of sodium taurocholate to enhance spore recovery on a medium selective for Clostridium difficile. J Clin Microbiol. 1982; 15; 443–446.

The authors
Emma Tait¹ PhD, Stephen P. Stanforth¹ PhD, John D. Perry² PhD and John R. Dean¹* DSc, PhD
¹Faculty of Health & Life Sciences, Department of Applied Sciences, Northumbria University, Newcastle-upon-Tyne, UK
²Department of Microbiology, Freeman Hospital, Newcastle-upon-Tyne, UK

*Corresponding author
E-mail: John.dean@northumbria.ac.uk

A new, benchtop molecular analyser allows renal transplant centres to monitor urine and serum BK viral loads in house, permitting earlier diagnosis and management of BK virus associated nephropathy (BKVAN) in renal transplant recipients.

BK virus infection
Named after the renal transplant patient (with initials B.K.) that it was first isolated from in 1971, BK virus (BKV) is a Polyomavirus, characterized by its nonenveloped, icosahedral capsid and its circular, double-stranded DNA genome.  Although BKV is prevalent around the world, estimated to have infected more than 80% of the global population, infection with the virus is usually asymptomatic or associated with only mild respiratory tract symptoms in healthy individuals [1].  Following primary infection, which typically occurs in early childhood, the virus persists in a latent form in the kidneys and urinary tract of its host [2].

Reactivation of BKV can occur in immunocompromised and immunosuppressed individuals [3].  In most cases reactivation of the virus is benign but it can pose a particular challenge for renal transplant patients.  In such cases immunosuppression can cause a lytic BKV infection that results in viruria in 30-50% and viremia in 13-22% of renal transplant recipients [4].  BKV infection is one of the most common viral complications to affect renal allografts [5].  It can lead to BKV associated nephropathy (BKVAN) in up to 10% of renal transplant recipients, and is associated with graft failure in 15-80% of affected patients [3,6-9].

Not every latent infection leads to viral reactivation and BKVAN in renal transplant patients.  In addition to immunosuppression, other risk factors, such as intragraft inflammation and host-specific immunity have been suggested [3].  The progression of BKVAN may occur without obvious signs or symptoms, other than raised serum creatinine, and so it is often misdiagnosed [9].

Management of BKVAN
Treatment of BKV infection and BKVAN in renal transplant patients usually involves a reduction or modification of immunosuppressive therapy.  It is generally agreed that early diagnosis and treatment are extremely important to prevent damage to the allograft [3,9].  At a later stage of infection, when intragraft inflammation has developed, reduction of immunosuppression may not be effective and may even be detrimental to the allograft [3].  For this reason, frequent monitoring of BKV in renal transplant patients, to detect early onset BKV infection, is recommended to ensure timely intervention [3].

Confirmation of BKVAN is performed by histological examination of an allograft biopsy sample.  However, clinical intervention is often based on the presence of viral replication as a surrogate marker and early indicator of BKV infection.  For this reason, non-invasive urine and blood tests have value in screening for BKV reactivation, monitoring the clinical course of infection, or monitoring response to therapy [3].

Urine cytology has been used to screen for BKV reactivation in renal transplant recipients.  However, since the virus may be shed in the urine of healthy individuals, quantitative results are required for this method to have diagnostic value [3].  Furthermore, accurate interpretation of cytology results requires training and expertise as it is often difficult to distinguish BKV from other viral infections [9].

Recently, molecular techniques for the detection and quantification of BKV in blood and urine have become available.  Such methods offer greater specificity for BKV and provide a valuable tool for identifying patients at risk of BKVAN before renal function deteriorates.

Monitoring BK viral loads
Quantitative measurements of BK viral load in urine and blood by molecular techniques are useful for monitoring the course of BKV infection [9] and for predicting the development of BKVAN [4,7,10,11].  Viral reactivation can be detected in the urine several weeks before the virus is detected in the blood, and viremia can be detected months before histological evidence of BKVAN is present [3].  Monitoring BKV loads in the urine and serum or plasma of renal transplant recipients, therefore, may be valuable in identifying those at risk of developing BKVAN, allowing further investigation and early intervention if necessary [3,9].  Such measurements are also valuable in monitoring response to therapy [3,9].

Although suggested BKV load thresholds for quantitative molecular measurements vary, and laboratories are encouraged to establish their own cut off values for the purpose of clinical management [9], BK viral loads of greater than 10,000 copies/mL in blood [6,11-14] and greater than 10 million copies/mL in urine are considered predictive for BKVAN [6,11,12].

Current guidelines recommend screening for BKV in the serum or plasma of kidney transplant patients monthly for the first 3-6 months after transplantation, and then every 3 months up to one year post-transplantation [15].  These guidelines also recommend that patients are screened for BKV if there is an unexplained rise in serum creatinine or following treatment for acute rejection [15].

Faster quantification of BKV
Due to the specialist nature of BKV testing and the resources and expertise required to perform BKV measurements by urine cytology or nucleic acid testing, many centres are required to send samples to a reference laboratory for analysis.  Some laboratories have adopted in-house polymerase chain reaction (PCR) BKV assays.  These can be labour intensive, variable in terms of specificity for BKV, and may require further confirmatory testing on positive samples, which can cause significant delays and can potentially impact patient management.

A new molecular method is now available that can reduce the turnaround time for quantitative BKV results significantly and provide the high specificity required for making important clinical decisions about the management of renal transplant patients.  The Diasorin Liaison® Iam benchtop instrument, with its small footprint and ease of operation, offers a cost effective and scalable solution for laboratories servicing renal transplant centres.  Demonstrating no cross reactivity with other significant pathogens, including JCV, the Diasorin Iam BKV assay provides reliable, quantitative results on the same day as sample receipt [16].

The Iam BKV assay uses loop-mediated isothermal amplification (Diasorin Q-LAMP) to measure BKV DNA in urine, plasma or serum.  Unlike conventional LAMP technology, Diasorin Q-LAMP is a rapid, real-time, fluorescent technique that allows quantitative analysis of individual or multiple targets in a single reaction [16]. 

Q-LAMP is based on the recognition of multiple primer binding regions on the target nucleic acid and amplification of the target sequence, which is facilitated by polymerase with strand displacement activity.  Quantification is achieved through the use of fluorophore-labelled primers and an observed decrease in fluorescence during amplification of the target sequence, together with known calibrators.  The Diasorin Iam BKV assay is a duplex Q-LAMP assay, designed to recognize a consensus sequence common to all known BKV subtypes. Integral controls provide verification of the efficiency of the extraction process and demonstrate the absence of inhibitors [16].

The Iam BKV Q-LAMP assay fits easily into daily laboratory routines.  Once samples are prepared and loaded onto the Liaison® Iam instrument, no operator intervention is required during an assay run, allowing staff to walk away until the routine is completed and the result is displayed.  The Iam BKV assay is extremely sensitive, with a limit of detection (defined as that concentration of BK virus with a 95% probability of detection by probit analysis) of 450 cps/mL in plasma (95% Confidence Interval 350 – 770 cps/mL) and 540 cps/mL in urine (95% CI 440 – 780 cps/mL) [16].  The BKV primers used represent all known BKV subtypes (Ia, Ib-1, IB-2, Ic, II,III and IV) and show no significant homology with a range of pathogens, including SV-40 virus and Herpes viruses, or cross reactivity with the closely-related Polyomavirus, JCV [16].

Improved management of renal transplant patients
The DiaSorin Iam BKV assay for the detection and quantification of BKV has been in use at the 975-bed Westmead Hospital in Sydney since July, 2013.  Westmead Hospital is a major teaching hospital for Sydney University and one of Australia’s largest centres for post-graduate training to specialist level in all fields.  The Department of Renal Medicine and Transplantation and the Centre for Transplant and Renal Research work closely with the Centre for Infectious Diseases and Microbiology (CIDM), which is part of Pathology West, a leading public pathology service in New South Wales.  The focus of the Transplant and Renal Research Group is to improve the lives of people with end-organ failure through transplantation. It also aims to reduce the number of people requiring dialysis by preventing the progression of chronic renal disease.

Senior Hospital Scientist at the Westmead CIDM laboratory, Dr Neisha Jeoffreys, comments, “BKV is an important pathogen in renal transplant patients.  It can cause serious complications and so early detection of viral reactivation and accurate monitoring of viral loads is a vital aspect of patient management.”
 
The Westmead CIDM laboratory provides a BKV testing service to the hospital’s renal transplant outpatient clinics as well as other specialist clinics associated with the centre.  They also test samples from other pathology groups in their reference capacity.  Dr Jeoffreys explains, “Renal transplant patients are tested routinely using the Iam BKV assay at 1, 2, 3, 6, 9 and 12 months post transplantation.  Patients that test positive for BKV are tested more frequently, every 2-4 weeks.”

“”The role of the quantitative Iam BKV assay is to determine if the patient is likely to develop BKVAN, which may lead to premature graft loss.  Patients with high BKV levels will have their immunosuppression regime modified in order to reduce BKV levels while preventing graft rejection.  Ongoing monitoring of BK viral load then assists the renal physicians to ensure the right amount of immunosuppression is delivered to reduce the risk of BKVAN and maintain a healthy graft.  Quantitative results allow the physicians to determine the appropriate point at which to modify the treatment.”

Previously, the laboratory used a qualitative in-house conventional PCR method for the detection of BKV followed by monthly quantification of viral loads in BKV-positive patients using a commercially available real-time PCR assay.

“We feel that the Iam BKV assay enables us to provide a better service for our renal specialists,” Dr Jeoffreys continues.  “We like the scalability of the Liaison® Iam instrument.   We have 3 instruments, which provide the flexibility to perform 1 or up to 21 samples at the same time, optimizing reagent usage.  This has allowed us to provide faster turnaround of results as we can now perform quantitative assays immediately and several times a week.  The Liaison® Iam method has also helped to improve workflows as it is fast and easy to perform, with less hands-on time than our previous methods, which makes it more cost effective.”

“It is hoped that the rapid quantitative results provided by the Iam BKV assay will allow our renal physicians to respond more quickly to high or escalating BK viral loads,” Dr Jeoffreys concludes.  “This will ultimately reduce the rate of graft loss due to BKVAN and allow for better patient management with reduced immunosuppression.”

The Iam BKV assay was the first assay to become available on the Diasorin Liaison® Iam instrument.  The growing portfolio of tests available on this platform includes assays for Varicella zoster virus, Parvovirus B19 and Toxoplasma gondii. 

^{Dr Neisha Jeoffreys is Senior Hospital Scientist at the Centre for Infectious Diseases and Microbiology (CIDM) based at Westmead Hospital, part of the Pathology West Institute of Clinical Pathology and Medical Research (ICPMR).}

For further information, please contact Tiffany Page, Global Marketing Manager, Molecular Infectious Disease, DiaSorin, tiffany.page@ie.diasorin.com
www.diasorin.com.

References
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3. Babel, N,  Volk, H and Reinke, P (2011).  Nat. Rev. Nephrol. 7: 399–406
4. Hirsch, H. H. et al. J. Med. 347, 488–496 (2002).
5. Ramos, E., Drachenberg, C. B., Wali, R. & Hirsch, H. H. Transplantation 87, 621–630 (2009).
6. Hirsch, H. H. et al. Polyomavirus-associated nephropathy in renal transplantation: interdisciplinary analyses and recommendations. Transplantation 79, 1277–1286 (2005).
7. Brennan, D. C. et al. Am. J. Transplant. 5, 582–594 (2005).
8. Hirsch HH. Clin Infect Dis. 2005;41:354-360.
9. Bechert, CJ, Schnadig, VJ, Payne, DA and Dong, J. (2010) Monitoring of BK Viral Load in Renal Allograft Recipients by Real-Time PCR Assays. Am J Clin Pathol 133:242-250.
10. Babel, N. et al. Transplantation 88, 89–95 (2009).
11. Dadhania, D. et al. Transplantation 86, 521–528 (2008).
12. Costa, C. et al. Nephrol. Dial. Transplant. 23, 3333–3336 (2008).
13. Hirsch HH, Steiger J. Polyomavirus BK. Lancet Infect Dis. 2003;3:611-623.
14. Ding R, Medeiros M, Dadhania D, et al. Transplantation. 2002;74:987-994.
15. KDIGO, Am. J. Transplant. 9 (Suppl. 3), S44–S58 (2009).
16. Diasorin Iam BKV assay Instructions for Use, BKV-524-02,EN 12/12.