RESEARCH LETTER
Discrepancies among three laboratory methods for Clostridium difficile detection and a proposal for their optimal use Alexandre A. Monteiro1, Renata N. Pires1,2, Ludmila F. Baethgen1, Lilian C. Carneiro1, Rejane G. ~o1, Steven Park5, David S. Perlin5, Edison M. Rodrigues Filho2,6 & Tavares3,4, Juliana Caiera Alessandro C. Pasqualotto1,2 rdia de Porto Alegre, Porto Alegre, de de Porto Alegre, Porto Alegre, Brazil; 2Santa Casa de Miserico Universidade Federal de Cie^ncias da Sau 3 Brazil; Universidade Federal De Pelotas, Pelotas, Brazil; 4Universidade Feevale, Porto Alegre, Brazil; 5Public Health Research Institute, New Jersey Medical School, New Jersey, NJ, USA; and 6Grupo Hospitalar Conceicß~ ao, Porto Alegre, Brazil
1
Correspondence: Alessandro C. Pasqualotto, Molecular Biology Laboratory, Santa Casa de Misericordia de Porto Alegre, Avenida Independ^ encia 155, 8º andar, 90035-074, Porto Alegre, Brazil. Tel.: +55 51 9995 1614; fax: +55 51 3213 7491; e-mail: [email protected] Received 16 October 2013; accepted 7 November 2013. Final version published online 11 December 2013. DOI: 10.1111/1574-6968.12330
MICROBIOLOGY LETTERS
Editor: Marilyn Roberts Keywords faecal samples; qPCR; algorithms.
Abstract Clostridium difficile is the major cause of nosocomial diarrhoea. Several detection methods are available for the laboratory diagnosis of C. difficile, but these vary in terms of sensitivity and specificity. In this study, we compared the performance of three following laboratory tests to detect C. difficile: in-house real-time PCR aiming for toxin B gene (tcdB), EIA for detection of toxins A and B (Premier Toxins A & B) and C. difficile culture in selective medium (bioMerieux). Our results were grouped into three categories as follows: (1) C. difficile-associated diarrhoea (CDAD); (2) asymptomatic carriers; and (3) negative results. Among the 113 patients included in the study, 9 (8.0%) were classified as CDAD, 19 (16.8%) were asymptomatic carriers, 76 (67.2%) had negative results and 9 (8.0%) could not be categorized (positive test for C. difficile toxins only). PCR was found to be the most sensitive diagnostic test in our study, with the potential to be used as a screening method for C. difficile colonization/CDAD. Diagnosis of CDAD would be better performed by a combination of PCR and EIA tests.
Introduction Clostridium difficile is a major cause of nosocomial diarrhoea in adults and is associated with significant morbidity and mortality. The incidence of C. difficile infection (CDI) and C. difficile-associated diarrhoea (CDAD) has increased significantly in recent years, partly associated with the emergence of more virulent strains (Pepin et al., 2004; Song et al., 2008; Zilberberg et al., 2008). Despite being a laborious and time-consuming technique, culture remains the gold standard for the detection of C. difficile in clinical samples (Cohen et al., 2010; Boyanton et al., 2012). Therefore, the laboratorial diagnosis of CDAD is frequently based on the detection of C. difficile toxins A and B in faecal samples by enzyme-linked immunosorbent assay (EIA), despite its known limited sensitivity (32– 73%; Humphries et al., 2013; Crobach et al., 2009; Peterson & Robicsek, 2009). More recently, molecular tests have been developed for the detection of genes encoding C. difficile toxins, with improved sensitivities (c. 95– 100%; de Jong et al., 2012; Luna et al., 2011). In this FEMS Microbiol Lett 350 (2014) 133–137
regard, algorithms have been proposed to diagnose CDAD by combining different diagnostic tests (Vasoo et al., 2012). Here, we compared three different methods for C. difficile detection in the clinical practice, in an attempt to determine their best use in the routine laboratory.
Methodology Selection of patients and samples
Between October 2011 and July 2012, we prospectively tested all adult (≥ 18 years old) patients with diarrhoea (≥ 3 liquid stools over a 24-h period) who had been admitted to two hospitals in Southern Brazil. Hospital 1 is a referral transplant centre with 65 beds and one intensive care unit (ICU). Hospital 2 is a large public hospital, and only patients admitted to the ICU in this hospital were considered for study. Faecal samples were collected in closed sterile containers and sent immediately to the laboratory for processing. At arrival, samples were kept at 4 °C for a maximum of ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
134
48 h, aliquoted and identified. Different researchers were responsible for the performance of specific tests to detect C. difficile – investigators remained blinded to the tests being performed by the others. All samples were frozen at 80 °C and distributed to the individual investigator, therefore passing through just one defrost cycle. Diagnostic tests used in the study
The following tests were performed: (1) detection of toxins A and B by EIA; (2) bacterial culture; and (3) realtime polymerase chain reaction (qPCR) for toxin B gene (tcdB). All discordant results were repeated to minimize the sample processing errors.
A.A. Monteiro et al.
by the Public Health Research Institute (PHRI, NJ). The reactions were optimized to 25 lL, using 0.4 lM of primers, 3 mM of magnesium, 0.5 lM of molecular beacon, 12.5 lL of Platinum Quantitative PCR SupermixUDG (Invitrogen), 5 lL of DNA extract and MilliQ H2O. The real-time PCR assay was performed on an ABI 7500 qPCR system (Applied Biosystems). The conditions used were as follows: 50 °C for 2 min (incubation step for the UDG enzyme), 95 °C for 10 min, followed by 45 cycles of 95 °C for 30 s, 56 °C for 30 s and 72 °C for 30 s. Negative (Milli-Q water) and positive controls (a 1185-bp PCR product received from the PHRI) were used in every reaction. Sensitivity of qPCR test
EIA
For detection of toxins A and B, a commercial EIA kit was used (Premier Toxins A & B; Meridian Bioscience, Cincinnati, OH). Tests were performed according to the manufacturer’s instructions. Washes were carried out on an automated plate washer (ImmunoWashTM 1575 microplate washer; Bio-Rad). Readings were performed on a spectrophotometer plate reader (iMarkTM microplate absorbance reader; Bio-Rad). Absorbance was read at 450 nm. Culture
Samples were inoculated with 1-lL disposable loops in C. difficile agar (CLO) selective medium (bioMerieux, France). Plates were incubated anaerobically for 48 h at 37 °C, using a generator anaerobic jar (Anaerocult A; Merck KGaA, Germany), following the manufacturer’s specifications. Clostridium difficile colonies exhibit typical morphology and odour in this selective medium. After the growth in culture, C. difficile identification was confirmed by qPCR as described hereafter. Extraction of bacterial DNA
DNA was extracted using a commercial kit (QIAamp DNA Stool Mini Kit; Qiagen, Hilden, Germany). As C. difficile is a Gram-positive bacillus, bacterial lysis was performed at 95 °C instead of 70 °C. DNA was stored at 80 °C, until processing. DNA concentration and purity were determined with NanoDrop 2000, reading the optical density by the ratio 280/260. In-house real-time PCR detection of tcdB
qPCR tests were used based on specific primers and probe (molecular beacons) targeting tcdB, both granted ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
The analytical sensitivity of the qPCR was experimentally determined by performing 10-fold dilutions of the PCR product of C. difficile used in this study as a positive control. This fragment had known size and therefore DNA concentration, allowing for the generation of the standard curve. The limit of detection was five copies (Ct of 41.0), and the average Ct for positive samples was 37.0 ( SD 2.0). Classification of the laboratory results
For the purpose of this study, laboratory results were grouped into the following categories: (1) CDAD: we defined a CDAD case as a patient with positive culture and/or PCR, positive EIA for toxins A/B and diarrhoea. These individuals were classified as harbouring C. difficile organisms that are producing toxins; (2) Asymptomatic carriers: positive culture and/or PCR, in association with negative EIA. These patients are colonized by nontoxigenic isolates of C. difficile; and (3) Negative results: all tests were negative. The tests that were positive only for EIA were not included in any of the categories above – as cytotoxicity assays were not performed, we were not able to exclude these as truly positive cases.
Results During the period of study, 113 patients were evaluated. Nine cases (8.0%) were classified as CDAD, 19 (16.8%) were considered asymptomatic carriers and the remaining 76 patients (67.2%) were negative for C. difficile. Moreover, 9 (8.0%) had a positive test for C. difficile toxins by EIA only, with negative PCR and/or culture and could not be categorized. Figure 1 presents the performance of the diagnostic tests evaluated in this study. PCR was found to be the FEMS Microbiol Lett 350 (2014) 133–137
Discrepancies in Clostridium difficile laboratorial tests
Fig. 1. Performance of distinct diagnostic tests for patients with nosocomial diarrhoea and suspected CDAD: culture in selective medium, EIA for toxin detection and PCR. Numbers inside circles correspond to positive results.
most sensitive test, as it was positive for a total of 22 patients, 10 of whom were culture negative. Culture was positive in 18 patients, and 6 of these were found to be PCR negative. We observed that a positive culture was associated with a higher increase in the odds (OR 2.13; 95% CI 0.57–7.99) of a positive EIA result, in comparison with a positive PCR (P = 0.261).
Discussion This study revealed a high frequency of discordance among laboratory tests commonly used to diagnose C. difficile colonization and infection: PCR, culture and EIA for toxin detection. When compared with culture, PCR was found to be the most sensitive method, a finding
135
that has already been published by others (de Jong et al., 2012). However, as seen in this study, PCR may also detect nontoxigenic isolates of C. difficile, considering that PCR aims for the detection of genes A and B and not toxins themselves (Crobach et al., 2009). These characteristics make PCR an interesting test to screen patients for C. difficile colonization, which might be of particular interest for infection control purposes. Despite recent studies showing that isolates belonging to the same strain type were not necessarily linked by person-to-person transmission (Rodriguez-Palacios et al., 2013), it has also been demonstrated that asymptomatic carriers of C. difficile may be the source of infection to other patients, sharing in the hospital environment (Samore et al., 1994; Gerding et al., 2008). Conversely, the detection of C. difficile toxins by EIA had greater clinical relevance and is possibly cheaper, in comparison with PCR. In a previous study, the detection of stool toxins by EIA was not associated with disease severity (Humphries et al., 2013). The main issue with EIA testing is limited sensitivity (Tenover et al., 2010; Dubberke et al., 2011) – therefore, this is not an ideal test for screening. To confirm the EIA-negative cases, we would recommend the test for cytotoxicity in laboratories that have the necessary infrastructure. One should also consider the feasibility of such testing in clinical practice. An additional limitation of EIA testing in the current investigation was the high rate of inconclusive results (9 of the 18 positive samples). In a previous study in which paediatric inpatients were evaluated for CDI, false-positive EIA results occurred in approximately one-third of cases (Toltzis et al., 2012). Therefore, as stated before, the main value of EIA testing seems to rely on the detection of symptomatic patients, in combination to other more sensitive tests.
Fig. 2. Proposed algorithm for the use of PCR and EIA for toxins detection in the diagnosis of both Clostridium difficile detection (CDAD) and colonization.
FEMS Microbiol Lett 350 (2014) 133–137
ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
136
Recent studies have shown differences in terms of toxin constitution related to the C. difficile ribotype, directly influencing the performance of the EIA tests. In some cases, EIA sensitivity may be as low as 15.4% (Tenover et al., 2010). One of the main limitations of our study was the lack of cytotoxic assays, in the absence of such experiments, we cannot conclusively conclude on the sensitivity of the EIA tests. Also, no ribotyping assays were performed in the current investigation. In this study, bacterial culture was less sensitive than PCR, for the detection of C. difficile. However, it should be noted that enrichment cultures were not performed. Studies have shown that the use of enrichment cultures increases the sensitivity of the test, which could at least partially explain the false-positive attributed to EIA in our study (Arroyo et al., 2005). Patients with positive cultures were more likely infected by toxigenic isolates of C. difficile, in comparison with PCR. However, culture is labour-intensive, and the interpretation of results may be quite tricky. A few patients were culture positive but PCR negative (n = 6). In the PCR-negative samples, a problem during DNA extraction may have occurred. As our PCR test did not have an internal control, the carryover of PCR inhibitors during DNA extraction in faecal samples is a potential explanation for these results. Due to the absence of new material for further DNA extractions, all discordant tests were actually repeated using the same DNA samples, for the purpose of test confirmation. Considering that an ideal diagnostic test must be highly sensitive and specific, easy and quick to perform (Lyerly et al., 1988), none of the methods evaluated in this study has the appropriateness to be used alone in CDAD diagnosis. EIA presented low sensitivity, while PCR and culture did not detect the presence of the active toxin in biologically samples. To overcome the limitations of each test, an algorithm to detect CDI/colonization is presented in Fig. 2. Finally, the diagnostic success of CDAD should be based in fast and accurate laboratorial tests, essential characteristics for the correct management of the patient as well as to the prevent nosocomial transmission of C. difficile (Novak-Weekley et al., 2010). Thus, knowledge of the available diagnostic methods, their strengths and limitations may allow for the adoption of earlier and more accurate measures of hospital infection control, facilitating the choice of the most appropriate methodology for the laboratory routine.
Acknowledgements We are grateful to all of those who were involved in sample collection and patient recruitment, particularly Carla S. Lincho and Marion W. Trombetta. This study ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
A.A. Monteiro et al.
was supported by an independent research grant by Merck Sharp and Dhome and CNPq (National Counsel of Technological and Scientific Development). All authors report no conflict of interest relevant to this article.
References Arroyo LG, Rousseau J, Willey BM, Low DE, Staempfli H, McGeer A & Weese JS (2005) Use of selective enrichment broth to recover Clostridium difficile from stool swabs stored under different conditions. J Clin Microbiol 13: 5341–5343. Boyanton BL Jr, Sural P, Loomis CR, Pesta C, Gonzalez-Krellwitz L, Robinson-Dunn B & Riska P (2012) Loop-mediated isothermal amplification compared to real-time PCR and enzyme immunoassay for toxigenic Clostridium difficile detection. J Clin Microbiol 50: 640–645. Cohen SH, Gerding DN, Johnson S, Kelly CP, Loo VG, McDonald LC, Pepin J, Wilcox MH; Society for Healthcare Epidemiology of America & Infectious Diseases Society of America (2010) Clinical practice guidelines for Clostridium difficile infection in adults: 2010 update by the Society for Healthcare Epidemiology of America (SHEA) and the Infectious Diseases Society of America (IDSA). Infect Control Hosp Epidemiol 31: 431–455. Crobach MJT, Dekkers OM & Wilcox MH (2009) European Society of Clinical Microbiology and Infectious Diseases (ESCMID): data review and recommendations for diagnosing Clostridium difficile infection (CDI). Clin Microbiol Infect 15: 1053–1066. de Jong E, de Jong AS, Bartels CJ, Der Rijt-Van Van, Den Biggelaar C, Melchers WJ & Sturm PD (2012) Clinical and laboratory evaluation of a real-time PCR for Clostridium difficile toxin A and B genes. Eur J Clin Microbiol Infect Dis 31: 2219–2225. Dubberke ER, Han Z, Bobo L, Hink T, Lawrence B, Copper S, Hoppe-Bauer J, Burnham CA & Dunne WM Jr (2011) Impact of clinical symptoms on the interpretation of diagnostic assays for Clostridium difficile. J Clin Microbiol 49: 2887–2893. Gerding DN, Muto CA & Owens RC (2008) Measures to control and prevent Clostridium difficile infection. Clin Infect Dis 46: S43–S49. Humphries RM, Uslan DZ & Rubin Z (2013) Performance of Clostridium difficile toxin enzyme immunoassay and nucleic acid amplification tests stratified by patient disease severity. J Clin Microbiol 51: 869–873. Luna RA, Boyanton BL, Mehta S, Courtney EM, Webb CR, Revell PA & Versalovic J (2011) Rapid stool-based detection of Clostridium difficile infection by real-time PCR in a children’s hospital. J Clin Microbiol 49: 851–857. Lyerly DM, Krivan HC & Wilkins TD (1988) Clostridium difficile: its disease and toxins. Clin Microbiol Rev 1: 1–18.
FEMS Microbiol Lett 350 (2014) 133–137
Discrepancies in Clostridium difficile laboratorial tests
Novak-Weekley SM, Marlowe EM & Miller JM (2010) Clostridium difficile testing in the clinical laboratory by use of multiple testing algorithms. J Clin Microbiol 48: 889–893. Pepin J, Valiquette L, Alary M-E, Villemure P, Pelletier A, Forget K, Pepin K & Chouinard D (2004) Clostridium difficile-associated diarrhea in a region of Quebec from 1991 to 2003: a changing pattern of disease severity. Can Med Assoc J 171: 466–472. Peterson LR & Robicsek A (2009) Does my patient have Clostridium difficile infection? Ann Intern Med 151: 176–179. Rodriguez-Palacios A, Borgmann S, Kline TR & LeJeune JT (2013) Clostridium difficile in foods and animals: history and measures to reduce exposure. Anim Health Res Rev 14: 11–29. Samore MH, DeGirolami PC, Tlucko A, Lichtenberg DA, Melvin ZA & Karchmer AW (1994) Clostridium difficile colonization and diarrhea at a tertiary care hospital. Clin Infect Dis 18: 181–187. Song X, Bartlett JG, Speck K, Naegeli A, Carroll K & Perl TM (2008) Rising economic impact of Clostridium difficile-associated disease in adult hospitalized patient population. Infect Control Hosp Epidemiol 29: 823–828.
FEMS Microbiol Lett 350 (2014) 133–137
137
Tenover FC, Novak-Weekley S, Woods CW et al. (2010) Impact of strain type on detection of toxigenic Clostridium difficile: comparison of molecular diagnostic and enzyme immunoassay approaches. J Clin Microbiol 48: 3719–3724. Toltzis P, Nerandzic MM, Saade E, O’Riordan MA, Smathers S, Zaoutis T, Kim J & Donskey CJ (2012) High proportion of false-positive Clostridium difficile enzyme immunoassays for toxin A and B in pediatric patients. Infect Control Hosp Epidemiol 33: 175–179. Vasoo S, Stevens J, Portillo L, Barza R, Schejbal D, Wu MM, Chancey C & Singh K (2012) Cost-effectiveness of a modified two-step algorithm using a combined glutamate dehydrogenase/toxin enzyme dehydrogenase/toxin enzyme immunoassay and real-time PCR for the diagnosis of Clostridium difficile infection. J Microbiol Immunol Infect. In press. doi:10.1016/j.jmii.2012.07.008. Zilberberg MD, Shorr AF & Kollef MH (2008) Increase in adult Clostridium difficile-related hospitalizations and case-fatality rate, United States, 2000–2005. Emerg Infect Dis 14: 929–931.
ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
Discrepancies among three laboratory methods for Clostridium difficile detection and a proposal for their optimal use Alexandre A. Monteiro1, Renata N. Pires1,2, Ludmila F. Baethgen1, Lilian C. Carneiro1, Rejane G. ~o1, Steven Park5, David S. Perlin5, Edison M. Rodrigues Filho2,6 & Tavares3,4, Juliana Caiera Alessandro C. Pasqualotto1,2 rdia de Porto Alegre, Porto Alegre, de de Porto Alegre, Porto Alegre, Brazil; 2Santa Casa de Miserico Universidade Federal de Cie^ncias da Sau 3 Brazil; Universidade Federal De Pelotas, Pelotas, Brazil; 4Universidade Feevale, Porto Alegre, Brazil; 5Public Health Research Institute, New Jersey Medical School, New Jersey, NJ, USA; and 6Grupo Hospitalar Conceicß~ ao, Porto Alegre, Brazil
1
Correspondence: Alessandro C. Pasqualotto, Molecular Biology Laboratory, Santa Casa de Misericordia de Porto Alegre, Avenida Independ^ encia 155, 8º andar, 90035-074, Porto Alegre, Brazil. Tel.: +55 51 9995 1614; fax: +55 51 3213 7491; e-mail: [email protected] Received 16 October 2013; accepted 7 November 2013. Final version published online 11 December 2013. DOI: 10.1111/1574-6968.12330
MICROBIOLOGY LETTERS
Editor: Marilyn Roberts Keywords faecal samples; qPCR; algorithms.
Abstract Clostridium difficile is the major cause of nosocomial diarrhoea. Several detection methods are available for the laboratory diagnosis of C. difficile, but these vary in terms of sensitivity and specificity. In this study, we compared the performance of three following laboratory tests to detect C. difficile: in-house real-time PCR aiming for toxin B gene (tcdB), EIA for detection of toxins A and B (Premier Toxins A & B) and C. difficile culture in selective medium (bioMerieux). Our results were grouped into three categories as follows: (1) C. difficile-associated diarrhoea (CDAD); (2) asymptomatic carriers; and (3) negative results. Among the 113 patients included in the study, 9 (8.0%) were classified as CDAD, 19 (16.8%) were asymptomatic carriers, 76 (67.2%) had negative results and 9 (8.0%) could not be categorized (positive test for C. difficile toxins only). PCR was found to be the most sensitive diagnostic test in our study, with the potential to be used as a screening method for C. difficile colonization/CDAD. Diagnosis of CDAD would be better performed by a combination of PCR and EIA tests.
Introduction Clostridium difficile is a major cause of nosocomial diarrhoea in adults and is associated with significant morbidity and mortality. The incidence of C. difficile infection (CDI) and C. difficile-associated diarrhoea (CDAD) has increased significantly in recent years, partly associated with the emergence of more virulent strains (Pepin et al., 2004; Song et al., 2008; Zilberberg et al., 2008). Despite being a laborious and time-consuming technique, culture remains the gold standard for the detection of C. difficile in clinical samples (Cohen et al., 2010; Boyanton et al., 2012). Therefore, the laboratorial diagnosis of CDAD is frequently based on the detection of C. difficile toxins A and B in faecal samples by enzyme-linked immunosorbent assay (EIA), despite its known limited sensitivity (32– 73%; Humphries et al., 2013; Crobach et al., 2009; Peterson & Robicsek, 2009). More recently, molecular tests have been developed for the detection of genes encoding C. difficile toxins, with improved sensitivities (c. 95– 100%; de Jong et al., 2012; Luna et al., 2011). In this FEMS Microbiol Lett 350 (2014) 133–137
regard, algorithms have been proposed to diagnose CDAD by combining different diagnostic tests (Vasoo et al., 2012). Here, we compared three different methods for C. difficile detection in the clinical practice, in an attempt to determine their best use in the routine laboratory.
Methodology Selection of patients and samples
Between October 2011 and July 2012, we prospectively tested all adult (≥ 18 years old) patients with diarrhoea (≥ 3 liquid stools over a 24-h period) who had been admitted to two hospitals in Southern Brazil. Hospital 1 is a referral transplant centre with 65 beds and one intensive care unit (ICU). Hospital 2 is a large public hospital, and only patients admitted to the ICU in this hospital were considered for study. Faecal samples were collected in closed sterile containers and sent immediately to the laboratory for processing. At arrival, samples were kept at 4 °C for a maximum of ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
134
48 h, aliquoted and identified. Different researchers were responsible for the performance of specific tests to detect C. difficile – investigators remained blinded to the tests being performed by the others. All samples were frozen at 80 °C and distributed to the individual investigator, therefore passing through just one defrost cycle. Diagnostic tests used in the study
The following tests were performed: (1) detection of toxins A and B by EIA; (2) bacterial culture; and (3) realtime polymerase chain reaction (qPCR) for toxin B gene (tcdB). All discordant results were repeated to minimize the sample processing errors.
A.A. Monteiro et al.
by the Public Health Research Institute (PHRI, NJ). The reactions were optimized to 25 lL, using 0.4 lM of primers, 3 mM of magnesium, 0.5 lM of molecular beacon, 12.5 lL of Platinum Quantitative PCR SupermixUDG (Invitrogen), 5 lL of DNA extract and MilliQ H2O. The real-time PCR assay was performed on an ABI 7500 qPCR system (Applied Biosystems). The conditions used were as follows: 50 °C for 2 min (incubation step for the UDG enzyme), 95 °C for 10 min, followed by 45 cycles of 95 °C for 30 s, 56 °C for 30 s and 72 °C for 30 s. Negative (Milli-Q water) and positive controls (a 1185-bp PCR product received from the PHRI) were used in every reaction. Sensitivity of qPCR test
EIA
For detection of toxins A and B, a commercial EIA kit was used (Premier Toxins A & B; Meridian Bioscience, Cincinnati, OH). Tests were performed according to the manufacturer’s instructions. Washes were carried out on an automated plate washer (ImmunoWashTM 1575 microplate washer; Bio-Rad). Readings were performed on a spectrophotometer plate reader (iMarkTM microplate absorbance reader; Bio-Rad). Absorbance was read at 450 nm. Culture
Samples were inoculated with 1-lL disposable loops in C. difficile agar (CLO) selective medium (bioMerieux, France). Plates were incubated anaerobically for 48 h at 37 °C, using a generator anaerobic jar (Anaerocult A; Merck KGaA, Germany), following the manufacturer’s specifications. Clostridium difficile colonies exhibit typical morphology and odour in this selective medium. After the growth in culture, C. difficile identification was confirmed by qPCR as described hereafter. Extraction of bacterial DNA
DNA was extracted using a commercial kit (QIAamp DNA Stool Mini Kit; Qiagen, Hilden, Germany). As C. difficile is a Gram-positive bacillus, bacterial lysis was performed at 95 °C instead of 70 °C. DNA was stored at 80 °C, until processing. DNA concentration and purity were determined with NanoDrop 2000, reading the optical density by the ratio 280/260. In-house real-time PCR detection of tcdB
qPCR tests were used based on specific primers and probe (molecular beacons) targeting tcdB, both granted ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
The analytical sensitivity of the qPCR was experimentally determined by performing 10-fold dilutions of the PCR product of C. difficile used in this study as a positive control. This fragment had known size and therefore DNA concentration, allowing for the generation of the standard curve. The limit of detection was five copies (Ct of 41.0), and the average Ct for positive samples was 37.0 ( SD 2.0). Classification of the laboratory results
For the purpose of this study, laboratory results were grouped into the following categories: (1) CDAD: we defined a CDAD case as a patient with positive culture and/or PCR, positive EIA for toxins A/B and diarrhoea. These individuals were classified as harbouring C. difficile organisms that are producing toxins; (2) Asymptomatic carriers: positive culture and/or PCR, in association with negative EIA. These patients are colonized by nontoxigenic isolates of C. difficile; and (3) Negative results: all tests were negative. The tests that were positive only for EIA were not included in any of the categories above – as cytotoxicity assays were not performed, we were not able to exclude these as truly positive cases.
Results During the period of study, 113 patients were evaluated. Nine cases (8.0%) were classified as CDAD, 19 (16.8%) were considered asymptomatic carriers and the remaining 76 patients (67.2%) were negative for C. difficile. Moreover, 9 (8.0%) had a positive test for C. difficile toxins by EIA only, with negative PCR and/or culture and could not be categorized. Figure 1 presents the performance of the diagnostic tests evaluated in this study. PCR was found to be the FEMS Microbiol Lett 350 (2014) 133–137
Discrepancies in Clostridium difficile laboratorial tests
Fig. 1. Performance of distinct diagnostic tests for patients with nosocomial diarrhoea and suspected CDAD: culture in selective medium, EIA for toxin detection and PCR. Numbers inside circles correspond to positive results.
most sensitive test, as it was positive for a total of 22 patients, 10 of whom were culture negative. Culture was positive in 18 patients, and 6 of these were found to be PCR negative. We observed that a positive culture was associated with a higher increase in the odds (OR 2.13; 95% CI 0.57–7.99) of a positive EIA result, in comparison with a positive PCR (P = 0.261).
Discussion This study revealed a high frequency of discordance among laboratory tests commonly used to diagnose C. difficile colonization and infection: PCR, culture and EIA for toxin detection. When compared with culture, PCR was found to be the most sensitive method, a finding
135
that has already been published by others (de Jong et al., 2012). However, as seen in this study, PCR may also detect nontoxigenic isolates of C. difficile, considering that PCR aims for the detection of genes A and B and not toxins themselves (Crobach et al., 2009). These characteristics make PCR an interesting test to screen patients for C. difficile colonization, which might be of particular interest for infection control purposes. Despite recent studies showing that isolates belonging to the same strain type were not necessarily linked by person-to-person transmission (Rodriguez-Palacios et al., 2013), it has also been demonstrated that asymptomatic carriers of C. difficile may be the source of infection to other patients, sharing in the hospital environment (Samore et al., 1994; Gerding et al., 2008). Conversely, the detection of C. difficile toxins by EIA had greater clinical relevance and is possibly cheaper, in comparison with PCR. In a previous study, the detection of stool toxins by EIA was not associated with disease severity (Humphries et al., 2013). The main issue with EIA testing is limited sensitivity (Tenover et al., 2010; Dubberke et al., 2011) – therefore, this is not an ideal test for screening. To confirm the EIA-negative cases, we would recommend the test for cytotoxicity in laboratories that have the necessary infrastructure. One should also consider the feasibility of such testing in clinical practice. An additional limitation of EIA testing in the current investigation was the high rate of inconclusive results (9 of the 18 positive samples). In a previous study in which paediatric inpatients were evaluated for CDI, false-positive EIA results occurred in approximately one-third of cases (Toltzis et al., 2012). Therefore, as stated before, the main value of EIA testing seems to rely on the detection of symptomatic patients, in combination to other more sensitive tests.
Fig. 2. Proposed algorithm for the use of PCR and EIA for toxins detection in the diagnosis of both Clostridium difficile detection (CDAD) and colonization.
FEMS Microbiol Lett 350 (2014) 133–137
ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
136
Recent studies have shown differences in terms of toxin constitution related to the C. difficile ribotype, directly influencing the performance of the EIA tests. In some cases, EIA sensitivity may be as low as 15.4% (Tenover et al., 2010). One of the main limitations of our study was the lack of cytotoxic assays, in the absence of such experiments, we cannot conclusively conclude on the sensitivity of the EIA tests. Also, no ribotyping assays were performed in the current investigation. In this study, bacterial culture was less sensitive than PCR, for the detection of C. difficile. However, it should be noted that enrichment cultures were not performed. Studies have shown that the use of enrichment cultures increases the sensitivity of the test, which could at least partially explain the false-positive attributed to EIA in our study (Arroyo et al., 2005). Patients with positive cultures were more likely infected by toxigenic isolates of C. difficile, in comparison with PCR. However, culture is labour-intensive, and the interpretation of results may be quite tricky. A few patients were culture positive but PCR negative (n = 6). In the PCR-negative samples, a problem during DNA extraction may have occurred. As our PCR test did not have an internal control, the carryover of PCR inhibitors during DNA extraction in faecal samples is a potential explanation for these results. Due to the absence of new material for further DNA extractions, all discordant tests were actually repeated using the same DNA samples, for the purpose of test confirmation. Considering that an ideal diagnostic test must be highly sensitive and specific, easy and quick to perform (Lyerly et al., 1988), none of the methods evaluated in this study has the appropriateness to be used alone in CDAD diagnosis. EIA presented low sensitivity, while PCR and culture did not detect the presence of the active toxin in biologically samples. To overcome the limitations of each test, an algorithm to detect CDI/colonization is presented in Fig. 2. Finally, the diagnostic success of CDAD should be based in fast and accurate laboratorial tests, essential characteristics for the correct management of the patient as well as to the prevent nosocomial transmission of C. difficile (Novak-Weekley et al., 2010). Thus, knowledge of the available diagnostic methods, their strengths and limitations may allow for the adoption of earlier and more accurate measures of hospital infection control, facilitating the choice of the most appropriate methodology for the laboratory routine.
Acknowledgements We are grateful to all of those who were involved in sample collection and patient recruitment, particularly Carla S. Lincho and Marion W. Trombetta. This study ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
A.A. Monteiro et al.
was supported by an independent research grant by Merck Sharp and Dhome and CNPq (National Counsel of Technological and Scientific Development). All authors report no conflict of interest relevant to this article.
References Arroyo LG, Rousseau J, Willey BM, Low DE, Staempfli H, McGeer A & Weese JS (2005) Use of selective enrichment broth to recover Clostridium difficile from stool swabs stored under different conditions. J Clin Microbiol 13: 5341–5343. Boyanton BL Jr, Sural P, Loomis CR, Pesta C, Gonzalez-Krellwitz L, Robinson-Dunn B & Riska P (2012) Loop-mediated isothermal amplification compared to real-time PCR and enzyme immunoassay for toxigenic Clostridium difficile detection. J Clin Microbiol 50: 640–645. Cohen SH, Gerding DN, Johnson S, Kelly CP, Loo VG, McDonald LC, Pepin J, Wilcox MH; Society for Healthcare Epidemiology of America & Infectious Diseases Society of America (2010) Clinical practice guidelines for Clostridium difficile infection in adults: 2010 update by the Society for Healthcare Epidemiology of America (SHEA) and the Infectious Diseases Society of America (IDSA). Infect Control Hosp Epidemiol 31: 431–455. Crobach MJT, Dekkers OM & Wilcox MH (2009) European Society of Clinical Microbiology and Infectious Diseases (ESCMID): data review and recommendations for diagnosing Clostridium difficile infection (CDI). Clin Microbiol Infect 15: 1053–1066. de Jong E, de Jong AS, Bartels CJ, Der Rijt-Van Van, Den Biggelaar C, Melchers WJ & Sturm PD (2012) Clinical and laboratory evaluation of a real-time PCR for Clostridium difficile toxin A and B genes. Eur J Clin Microbiol Infect Dis 31: 2219–2225. Dubberke ER, Han Z, Bobo L, Hink T, Lawrence B, Copper S, Hoppe-Bauer J, Burnham CA & Dunne WM Jr (2011) Impact of clinical symptoms on the interpretation of diagnostic assays for Clostridium difficile. J Clin Microbiol 49: 2887–2893. Gerding DN, Muto CA & Owens RC (2008) Measures to control and prevent Clostridium difficile infection. Clin Infect Dis 46: S43–S49. Humphries RM, Uslan DZ & Rubin Z (2013) Performance of Clostridium difficile toxin enzyme immunoassay and nucleic acid amplification tests stratified by patient disease severity. J Clin Microbiol 51: 869–873. Luna RA, Boyanton BL, Mehta S, Courtney EM, Webb CR, Revell PA & Versalovic J (2011) Rapid stool-based detection of Clostridium difficile infection by real-time PCR in a children’s hospital. J Clin Microbiol 49: 851–857. Lyerly DM, Krivan HC & Wilkins TD (1988) Clostridium difficile: its disease and toxins. Clin Microbiol Rev 1: 1–18.
FEMS Microbiol Lett 350 (2014) 133–137
Discrepancies in Clostridium difficile laboratorial tests
Novak-Weekley SM, Marlowe EM & Miller JM (2010) Clostridium difficile testing in the clinical laboratory by use of multiple testing algorithms. J Clin Microbiol 48: 889–893. Pepin J, Valiquette L, Alary M-E, Villemure P, Pelletier A, Forget K, Pepin K & Chouinard D (2004) Clostridium difficile-associated diarrhea in a region of Quebec from 1991 to 2003: a changing pattern of disease severity. Can Med Assoc J 171: 466–472. Peterson LR & Robicsek A (2009) Does my patient have Clostridium difficile infection? Ann Intern Med 151: 176–179. Rodriguez-Palacios A, Borgmann S, Kline TR & LeJeune JT (2013) Clostridium difficile in foods and animals: history and measures to reduce exposure. Anim Health Res Rev 14: 11–29. Samore MH, DeGirolami PC, Tlucko A, Lichtenberg DA, Melvin ZA & Karchmer AW (1994) Clostridium difficile colonization and diarrhea at a tertiary care hospital. Clin Infect Dis 18: 181–187. Song X, Bartlett JG, Speck K, Naegeli A, Carroll K & Perl TM (2008) Rising economic impact of Clostridium difficile-associated disease in adult hospitalized patient population. Infect Control Hosp Epidemiol 29: 823–828.
FEMS Microbiol Lett 350 (2014) 133–137
137
Tenover FC, Novak-Weekley S, Woods CW et al. (2010) Impact of strain type on detection of toxigenic Clostridium difficile: comparison of molecular diagnostic and enzyme immunoassay approaches. J Clin Microbiol 48: 3719–3724. Toltzis P, Nerandzic MM, Saade E, O’Riordan MA, Smathers S, Zaoutis T, Kim J & Donskey CJ (2012) High proportion of false-positive Clostridium difficile enzyme immunoassays for toxin A and B in pediatric patients. Infect Control Hosp Epidemiol 33: 175–179. Vasoo S, Stevens J, Portillo L, Barza R, Schejbal D, Wu MM, Chancey C & Singh K (2012) Cost-effectiveness of a modified two-step algorithm using a combined glutamate dehydrogenase/toxin enzyme dehydrogenase/toxin enzyme immunoassay and real-time PCR for the diagnosis of Clostridium difficile infection. J Microbiol Immunol Infect. In press. doi:10.1016/j.jmii.2012.07.008. Zilberberg MD, Shorr AF & Kollef MH (2008) Increase in adult Clostridium difficile-related hospitalizations and case-fatality rate, United States, 2000–2005. Emerg Infect Dis 14: 929–931.
ª 2013 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
