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
1. Goudsmit, J. et al. J. Med. Virol. 10, 91–99 (1982).
2. Shinohara T, Matsuda M, Cheng SH, et al. J Med Virol. 1993;41:301-305.
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.
H. pylori infection: a laboratory perspective
, /in Featured Articles /by 3wmediaInfection with Helicobacter pylori is indicated in disorders of the upper gastrointestinal tract, from dyspepsia and ulcer disease to gastric carcinoma. It can be detected during endoscopy or non-invasively using breath, stool or blood samples, each of which has advantages and limitations depending on patient and population circumstances.
by Sarah Knowles and Julia M. Forsyth
Helicobacter pylori
Helicobacter pylori is probably the most common cause of bacterial infection in humans, present in up to 50% of the world’s population [1]. The presence of such a microorganism in the stomach was first reported almost 100 years ago [2] but was not taken seriously until Marshall and Warren demonstrated a strong association between the presence of an unidentified curved bacillus and inflammation on a gastric biopsy [3]. The organism was initially placed in the Campylobacter genus but as further morphological, structural and genetic information was determined a new genus called Helicobacter was created. After identification, Marshall demonstrated the role of H. pylori in antral gastritis by self-administration of the bacteria and also showed that it could be cleared by the use of antibiotics and bismuth salts.
H. pylori and disease
Infection with H. pylori is known to be a contributing factor in producing gastritis [4]. Studies have shown that infection with H. pylori leads to increased release of gastrin by the antral mucosa (through a mechanism that is as yet undefined), which resolves on eradication [5, 6]. There is a high prevalence of H. pylori positive chronic gastritis in patients with duodenal and gastric ulcers (70% and 90% respectively) [6, 7]. These ulcer patients have been shown to have an exaggerated response to the increased gastrin, even compared to asymptomatic H. pylori-positive patients [6], leading to excess acid production and a deterioration from the initial inflammation of gastritis to mucus layer erosion by peptic acid. H. pylori is also linked with gastric carcinomas and mucosa-associated lymphoid tissue (MALT) lymphomas with one study showing that 62% of patients with low-grade gastric MALT lymphoma entering complete remission 12 months after H. pylori eradication therapy [8]. The vast majority of individuals with H. pylori infection, however, do not have ulcerative disease, but are symptomless carriers [4].
The survival capabilities of H. pylori in the stomach make it difficult to eradicate. Effective treatment requires a multidrug ‘triple therapy’ regime consisting of two antibiotics from clarithromycin, metronidazole, amoxicillin and tetracycline combined with an acid suppressant or bismuth compounds [9], and in areas of high clarithromycin resistance, quadruple therapy including two antibiotics, proton pump inhibitors (PPIs) and bismuth [10].
Testing for H. pylori
The definitive identification of H. pylori is by histological examination of biopsies stained with Giesma stain or Warthin–Starry silver impregnation [4]. Other biopsy immunohistochemical techniques using specific antibodies against H. pylori are also available along with rapid biopsy urease tests (otherwise known as the CLO-test), where a biopsy specimen is placed in a urea broth containing a pH indicator. Testing by any of these methods requires endoscopy which, apart from being very unpleasant for the patient, carries risk and is very costly, and is therefore unsuitable for routine screening for H. pylori in patients with a low risk of cancer. In these cases a ‘test and treat’ strategy is recommended using non-invasive tests (Fig. 1) [10].
There are three common non-invasive tests available for the routine detection of H. pylori:
1. The urea breath test (UBT)
2. Stool test for H. pylori antigen
3. Serum test for H. pylori antibody.
Urea breath test
The UBT is generally considered to be the ‘gold standard’ non-invasive test and is recommended as the first-choice screening test both in the diagnosis of infection and post-eradication testing [9–11].
Urea labelled with 13C is taken as a drink or a capsule and urease produced by H. pylori acts upon the urea to produce 13C-labelled carbon dioxide. This is absorbed into the blood stream and exhaled in the breath. The ratio of 13CO2 : 12CO2 is measured using an isotope ratio mass spectrometer (IRMS), and the ratios before and after the administration of urea are compared to give the result.
Apart from being the ‘gold standard’ test the other main advantage of the UBT is the stability of the breath samples once collected. This allows storage without refrigeration and transportation to the central referral laboratory for analysis. In the UK this permits sample collection at local health centres rather than hospital settings, which is convenient for patients. Worldwide this can extend to large, remote areas with limited facilities or courier transport links to laboratories, for example rural Africa and Australia, permitting testing in all communities. Local studies have also shown the test to be very popular with patients as sample collection is simple and pain free [12].
A disadvantage of the UBT is that PPIs and antibiotics need to be withheld for 2 and 4 weeks respectively before testing to avoid potential false-negative results as a consequence of decreased bacterial load [10]. PPIs particularly are commonly prescribed to alleviate the symptoms of dyspepsia and patients can be reluctant to withhold such medication pending tests. As with all testing cost is also an important issue. IRMS is a specialized laboratory technique requiring expensive equipment and highly trained laboratory personnel. These fixed costs can make the UBT an expensive option but running costs and reagents for such equipment are relatively low, and therefore centralized high workload laboratories can provide a very cost-effective service [12].
Stool antigen test
The stool antigen test uses enzyme-linked immunosorbent assay (ELISA) methodology to detect H. pylori antigen. ELISA technology is available in many laboratories, thus making the test easy to implement at local laboratories. It is also reported to be more cost effective than the UBT [13], although local method validation of commercially available ELISA kits (for a workload of 3500 tests per year) showed the costs to be almost identical when samples were analysed in duplicate [12]. Duplicate analysis was recommended due to poor replicate precision, probably as a result of the heterogeneity of stool in the samples. As workload increases, e.g. at centralized laboratories, the UBT actually becomes the cheaper alternative because of the low running/reagent costs [12]. A final advantage of the stool antigen test is that there is no need to attend a clinic or phlebotomy appointment for sample collection. The patient can collect the sample at home and deliver it to their GP/local courier collection or laboratory at their convenience.
Current UK guidelines for the investigations do not recommend the stool antigen test in the post-eradication setting [9]. There is, however, no discrimination between polyclonal and monoclonal assays in these guidelines. Initial commercial ELISA assays were based upon polyclonal antibodies which showed reasonable performance in untreated patients with reported sensitivities of 91–93% and likewise specificities of 91–93% [14–16]. However, in the post-eradication setting results are less convincing with reported sensitivities ranging from 67–89% [14–19]. The introduction of monoclonal antibody assays has lead to an improvement in the diagnostic accuracy with reported sensitivities of 94–98% in both untreated adults and children [16, 20, 21], and pooled specificity of 97% [22]. Several more-recent studies using monoclonal antibody kits with favourable results have been published since the guidelines were drawn up with pooled sensitivities and specificities of 93% and 96% respectively [22]. This is reflected in the recent European study group findings which have recently been updated to include the monoclonal ELISA test as accurate in the post eradication setting, suitable for use if the UBT is not available [10], and local studies have validated the stool test as a suitable alternative to the UBT both in diagnosis and post-eradication settings [12].
The stool antigen test offers no further advantage to the UBT with regard to patient preparation as antibiotics and PPI medication should also be withheld before testing and locally the test is unpopular with patients who do not wish to provide stool specimens. Stability is also a disadvantage as specimens need to be refrigerated, and if not tested immediately, frozen within 48 hours [23]. Samples therefore need to reach the laboratory quickly or be transported frozen, providing logistical challenges in more remote locations.
Rapid point-of-care testing or ‘office’ stool antigen assays are available but have limited accuracy and are not recommended by current guidelines [10, 11].
Serum antibody test
The serum antibody test is an ELISA assay for IgG H. pylori antibodies, which again has the advantage of being a standard laboratory technique. It is also the cheapest testing option as reagent costs are low compared to the stool antigen test. Unfortunately this test has been shown to have poor diagnostic accuracy [11] especially in the post-eradication setting. This is thought to be due to the fact that the antibody levels persist in the blood for a long time and therefore lead to false-positive results after treatment. It may, however, have a place in the diagnosis of infection setting on occasions where PPI or antibiotic medications cannot be withheld, or for testing young children for whom the UBT is inappropriate and the likelihood of previous infection is low. Point-of-care or office-based serology tests are available, although their performance has been shown to be inadequate and they are not recommended for use in testing for H. pylori [10, 11].
Conclusion
In conclusion, H. pylori detection and eradication can lead to significant health benefits to the world’s population, not only alleviating life-restricting symptoms but also preventing the development of more serious disease. Several reliable methods of laboratory testing are now available, the choice of which depends on the patient population, facilities available, workload and networking possibilities and pre-existing medical conditions.
References
1. Cover TL, Blaser MJ. Adv Int Med. 1996; 41; 85–117.
2. Maden E, et al. Am J Clin Pathol. 1988; 90: 450–453.
3. Marshall BJ, Warren JR. Lancet 1984; 1(8390): 1311–1315.
4. Mera SL. Br J Biomed Sci. 1995; 52; 271–281.
5. Smith JTL, et al. Gut 1990; 31: 522–525.
6. El-Omar E, et al. Gut 1993; 34: 1060–1065.
7. Parsonner J, et al. N Engl J Med. 1991; 325: 1127–1131.
8. Fischbach W, et al. Gut 2004; 53: 34–37.
9. NICE. Dyspepsia. Management of dyspepsia in adults in primary care. Clinical Guideline 17 2004; http://www.nice.org.uk/nicemedia/pdf/CG017NICEguideline.pdf.
10. Malfertheiner P, et al. Gut 2012; 61: 646–664.
11. Malfertheiner P, et al. Gut 2007; 56: 772–781.
12. Research Studies, Pathology Department, Royal Derby Hospital. Data awaiting publication.
13. Elwyn G, et al. Br J Gen Pract. 2007; 57: 401–403.
14. Roth DE, et al. Clin Diagn Lab Immunol. 2001; 8: 718–723.
15. Gisbert JP, et al. Am J Gastroenterol. 2001; 96: 2829–2838.
16. Gisbert JP, et al. Helicobacter 2004; 9: 347–368.
17. Perri F, et al. Am J Gastroenterol. 2002; 97: 2756–2762.
18. Bilardi C, et al. Aliment Pharmacol Ther. 2002; 16: 1733–1738.
19. Erzin Y, et al. J Med Microbiol. 2005; 54: 863–866.
20. Koletzko S, et al. Gut 2003; 52: 804–806.
21. Weingart V, et al. J Clin Microbiol. 2004; 42: 1319–1321.
22. Gisbert JP, et al. Am J Gastroenterol 2006; 101: 1921–1930.
23. Oxoid Amplified IDEIA HPStAR Kit insert Ref K6630.
The author
Sarah Knowles MSc, FRCPath and Julia Forsyth MSc, FRCPath
Pathology Department, Royal Derby Hospital, Derby, UK
*Corresponding author
E-mail: Sarah.knowles@nhs.net
A new approach for diagnosis of Clostridium difficile based on evolution of a unique volatile organic compound identifier
, /in Featured Articles /by 3wmediaClostridium 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
MALDI-TOF mass spectrometry for rapid identification and subtyping of H. cinaedi strains isolated from humans and animals
, /in Featured Articles /by 3wmediaAlthough Helicobacter cinaedi infection is now recognized as an increasingly important emerging disease in humans, it is difficult to identify particular isolates due to their unusual phenotypic profiles and similarity of 16S rRNA sequences among closely related helicobacters. MALDI-TOF MS resolved the present difficulties associated with the identification of H. cinaedi.
by Takako Taniguchi and Prof. Naoaki Misawa
Helicobacter cinaedi infection
Helicobacter cinaedi, was first recognized as a Campylobacter-like organism (CLO), is a Gram-negative, spiral-shaped, motile, microaerobic bacterium, and is now classified into enterohepatic Helicobacter species [1]. This organism was first isolated from homosexual men and was initially recognized as a rectal and intestinal pathogen among members of that population [1]. The first case of H. cinaedi bacteremia in Japan was in an HIV-negative patient, but was receiving immunosuppressive therapy after renal transplantation [2]. Moreover, a few cases of infection with H. cinaedi isolated from feces and blood from an apparently non-immunocompromised child and adult have been reported [3]. Since then, H. cinaedi has become thought of as an opportunistic pathogen that causes bacteremia, cellulitis, septicemia and enteritis in immunocompromised patients [4, 5], immunocompetent patients and even healthy individuals [6]. Kitamura et al. reported an outbreak of nosocomial H. cinaedi infections caused by direct person-to-person transmission [6]. Therefore, healthcare workers need to pay attention to H. cinaedi infection as an increasingly important emerging disease in humans.
Epidemiology
H. cinaedi-like organisms have also been isolated from non-human sources such as dogs, cats, monkeys, hamsters and other rodents [7–10], suggesting that the organism may be widespread in a broad range of animal species. As Gebhart et al. reported that H. cinaedi was found in 75% of the healthy hamsters used in their study [9], it was hypothesized that hamsters might be an important reservoir for human infection [7, 9]. However, no reliable epidemiological evidence of zoonosis has been demonstrated for human cases of H. cinaedi infection [3].
Diagnosis
To isolate H. cinaedi from blood, blood was usually collected in BACTEC culture bottles and incubated in a BACTEC 9050 blood culture system (Becton Dickinson, BD Biosciences) for at least 5 to 7 days. When the incubation time was less than 5 days or other culture systems were used, the organism was not often isolated. Earlier research suggested that certain patients with H. cinaedi infection may remain undiagnosed or incorrectly diagnosed because of difficulties in detecting the bacteria by conventional culture methods [2].
We previously isolated at least six different spiral-shaped organisms including H. cinaedi and H. bilis in a puppy with bloody diarrhoea [11]. These organisms were identified based on their morphology, biochemical traits, whole-cell protein profiles, and the results of molecular analysis of their 16S rDNA sequences. However, the biochemical identification of Helicobacter strains based on a limited number of tests is difficult because helicobacters frequently exhibit unusual phenotypic profiles, even in the same species [10, 12]. Furthermore, H. cinaedi cannot be clearly discriminated from H. bilis on the basis of 16S rRNA sequences because of the high level of sequence similarity (greater than 98%) [12].
Application of MALDI-TOF MS for rapid identification and subtyping of H. cinaedi strains
Recently, matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS) has made it possible to analyse the protein composition of a bacterial cell based on intact-cell mass spectrometry (ICMS) profiles as a new technique for species identification. The technique is simple, rapid and accurate for identifying microorganisms regardless of their characteristics, such as Gram-negative and Gram-positive bacteria, mycobacteria, anaerobes and yeast species [13, 14]. MALDI-TOF MS has an advantage in that it has a low-cost performance and is independent of the age of the culture, growth conditions or medium selected, making it applicable for routine bacterial identification in clinical laboratories. Although there is a commercially available library that includes more than three thousand kinds of microorganism such as bacteria, yeast and fungus for identification and phylogenic analysis (MALDI Biotyper Reference Library, Bruker Daltonics), we use a library created in-house.
Therefore, we considered that MALDI-TOF MS might resolve the present difficulties with identification of H. cinaedi. Furthermore, we examined whether H. cinaedi strains isolated from different animals could be differentiated or subtyped by their ICMS profiles [15].
As shown in Figure 1, although common peaks were detected in the H. cinaedi and H. bilis strains examined, the m/z 5200 and 10400 peaks were detected only in strains of H. cinaedi. These peaks showed good reproducibility regardless of the isolate origins, different media and passage numbers on the same medium. Therefore, the ICMS profile of H. cinaedi could be completely differentiated from those of H. bilis. Furthermore, the ICMS profile of H. cinaedi was also distinguishable from those of H. mustelae, H. pylori, H. fennelliae and H. canis, indicating that ICMS profiling using MALDI-TOF MS is applicable for the identification of H. cinaedi.
Cluster analysis of H. cinaedi strains based on the ICMS profiles
Several papers report that direct contact with pets may be a possible route of infection in humans [3–5]; however, details regarding the pathogenesis and epidemiological features, including routes of infection of animal isolates in the context of both humans and animals, are not fully understood. No reliable epidemiological evidence of zoonosis has been demonstrated for human infections caused by H. cinaedi. Therefore, ICMS profiles of H. cinaedi strains isolated from humans and animals were measured, and a phyloproteomic tree was constructed in order to analyse the relationships between the strains. As a result, these H. cinaedi strains were clearly divided into two groups. All of the strains isolated from humans belonged to Cluster 2. All the other animal-derived strains belonged to Cluster 1 (Fig. 2). Interestingly, the ICMS-based phyloproteomic tree agreed with the phylogenetic tree that had been based on the nucleotide sequences of the hsp60 gene. These H. cinaedi strains were also clearly divided into two groups by the hsp60-gene-based phyloproteomic tree. Thus, the data from phyloproteomic and phylogenetic analysis suggest that human strains of H. cinaedi may be distinct from animal strains. Kiehlbauch et al. also reported that there may be subgroups within H. cinaedi isolated from humans, dogs, cats and hamsters that correlate with the host source on the basis of DNA–DNA hybridization and ribotyping analyses [12]. The present study appears to support the hypothesis that H. cinaedi from different host sources may form subgroups, which may prompt a revision of the classification of H. cinaedi.
Conclusion
In conclusion, the construction of ICMS profiles using the MALDI-TOF MS approach may be a useful tool for H. cinaedi identification and subtyping. Further investigations will be required to analyse additional strains from a broader area to confirm whether human strains belong to a distinct subtype of H. cinaedi.
References
1. Quinn TC, Goodell SE, et al. Ann Intern Med. 1984; 101: 187–192.
2. Murakami H, Goto M, et al. J Infect Chemother. 2003; 9: 344–347.
3. Orlicek SL, Welch DF, Kuhls TL. J Clin Microbiol. 1993; 31: 569–571.
4. Kiehlbauch JA, Tauxe RV, et al. Ann Intern Med. 1994; 121: 90–93.
5. Matsumoto T, Goto M, et al. J Clin Microbiol. 2007; 45: 2853–2857.
6. Kitamura T, Kawamura Y, et al. J Clin Microbiol. 2007; 45: 31–38.
7. Comunian LB, Moura SB, et al. Curr Microbiol. 2006; 53: 370–373.
8. Fernandez KR, Hansen LM, et al. J Clin Microbiol. 2002; 40: 1908–1912.
9. Gebhart CJ, Fennell CL, et al. J Clin Microbiol. 1989; 27: 1692–1694.
10. Kiehlbauch JA, Brenner DJ, et al. J Clin Microbiol. 1995; 33: 2940–2947.
11. Misawa N, Kawashima K, et al. Vet Microbiol. 2002; 87: 353–364.
12. Vandamme P, Harrington CS, et al. J Clin Microbiol. 2000; 38:.2261–2266.
13. Saffert RT, Cunningham SA, et al. J Clin Microbiol. 2011; 49: 887–892.
14. Stevenson LG, Drake SK, et al. J Clin Microbiol. 2010; 48: 3482–3486.
15. Taniguchi T, Sekiya A, et al. J Clin Microbiol. 2014; 52: 95–102.
The authors
Takako Taniguchi1 MSc and Naoaki Misawa2* DVM, PhD
1Laboratory of Veterinary Public Health, Department of Veterinary Science, Faculty of Agriculture, University of Miyazaki, Miyazaki, Japan
2Center for Animal Disease Control, University of Miyazaki, Miyazaki, Japan
*Corresponding author
E-mail: a0d901u@cc.miyazaki-u.ac.jp
Monitoring BK virus in kidney transplant patients
, /in Featured Articles /by 3wmediaA 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
1. Goudsmit, J. et al. J. Med. Virol. 10, 91–99 (1982).
2. Shinohara T, Matsuda M, Cheng SH, et al. J Med Virol. 1993;41:301-305.
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.
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