- Email: [email protected]
Reply: Two-step algorithm for the detection of Clostridium difficile from stool samples
96
Letters to the Editor / Journal of Hospital Infection 79 (2011) 93–98
than others of similar design to ours.2,4,5 This study supports the use of GDH as part of a two-step algorithm due to its high sensitivity.6 GDH is also cheap and confirmatory tests were only required in 14% of cases, which is similar to other studies.2 This study did not assess PCR alone, which some studies found superior to other methods, but PCR remains prohibitively expensive in the UK.7 Although we found that GDH improved the performance of EIA, we believe the low sensitivity of EIA prevents its use as both a screening test and as a confirmatory test. PCR following GDH appears to be the most sensitive test with the highest PPV and NPV overall but is the most expensive. The increased sensitivity (15 positive samples by GDH/PCR compared to 12 by EIA) may also adversely affect targets present in the UK. Further studies are required to determine the best method of diagnosis for patient care in terms of test performance and clinical outcome. Conflict of interest statement None declared. Funding sources Alere Ltd (Stockport, UK) provided the Dynex DS2 analyser and C. diff CheckÔ 60 kits for GDH assay. Cepheid provided GeneXpert C. difficile PCR assays. Internal funding was provided from the Clinical Microbiology & Public Health Laboratory, Peterborough & Stamford Hospitals NHSFT.
References 1. Bignardi GE, Settle C. Glutamate dehydrogenase as confirmatory test for Clostridium difficile toxin A/B-positive stools. J Hosp Infect 2010;75:327–328. 2. Goldenberg SD, Cliff PR, Smith S, Milner M, French GL. Two-step glutamate dehydrogenase antigen real-time polymerase chain reaction assay for detection of toxigenic Clostridium difficile. J Hosp Infect 2010;74:48–54. 3. Department of Health & Health Protection Agency. Clostridium difficile infection: how to deal with the problem. London: Department of Health; 2009. 4. Planche T, Aghaizu A, Holliman R, et al. Diagnosis of Clostridium difficile infection by toxin detection kits: a systematic review. Lancet Infect Dis 2008;8:777–784. 5. 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. 6. Fenner L, Widmer AF, Goy G, Rudin S, Frei R. Rapid and reliable diagnostic algorithm for detection of Clostridium difficile. J Clin Microbiol 2008;46:328–330. 7. Novak-Weekley SM, Marlowe EM, Miller JM, et al. Clostridium difficile testing in the clinical laboratory using multiple testing algorithms. J Clin Microbiol 2010; 48:889–893.
N.S. Clarka M.J. Doughtona J.A. Karasb D.A. Enocha,* a Clinical Microbiology & Public Health Laboratory, Peterborough & Stamford Hospitals NHS Foundation Trust, Peterborough City Hospital, Peterborough, UK b
Clinical Microbiology & Public Health Laboratory, Health Protection Agency, Papworth Hospital, Papworth Everard, Cambridge, UK *
Corresponding author. Address: Clinical Microbiology & Public Health Laboratory, Peterborough & Stamford Hospitals NHS Foundation Trust, Peterborough City Hospital, Bretton Gate, Peterborough PE3 9GZ, UK. Tel.: þ44 (0) 1733 678443; fax: þ44 (0) 1733 349721. E-mail address: [email protected] (D.A. Enoch). Accepted by S.J. Dancer Available online 2 June 2011
Ó 2011 The Healthcare Infection Society. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.jhin.2011.04.002
Reply: Two-step algorithm for the detection of Clostridium difficile from stool samples
Madam, We read with interest the letter by Clark et al. which comments on our previous letter and discusses the merits of various testing modalities for Clostridium difficile infection.1,2 When assessing the performance of C. difficile enzyme immunoassay (EIA) testing it is important to consider which EIA kit has been used, as the performance varies.3 We had used the Meridian Premier toxin AþB kit, because this is thought to be the toxin EIA with the highest sensitivity.3 When analysing the data from Clark et al. we notice that for the EIA test they used (VIDAS C. difficile toxin A & B) results are presented for 316 stools, whereas for the glutamate dehydrogenase (GDH) test (C. diff CheckÔ 60) results are presented for 322 stools.1 We presume that the difference is due to the fact that the VIDAS EIA kit can give equivocal results and that these were excluded from the analysis. Clark et al. commented on the fact that GDH screening followed by EIA testing improved the performance of EIA as a single test: this improvement was limited to the specificity and the positive predictive value (the number of false positives is reduced) but there is no gain in sensitivity.1 Toxin EIA testing cannot increase the overall sensitivity, in a two-step algorithm with GDH followed by toxin EIA, unless a third testing modality is added for the discrepant results. We have achieved a similar objective (reducing the number of false positives) with what we think is a more userfriendly protocol of toxin EIA followed by GDH immunochromatography, as opposed to the sequence used by Clark et al.1 of a GDH EIA followed by a toxin EIA. In order to estimate the frequency of false-negative results with our toxin EIA followed by GDH algorithm, we have recently retested 397 EIA toxinnegative stools and found that 26 were GDH positive but only one of these was positive with a cell culture cytotoxicity test with hep2 cells: thus if cell culture cytotoxicity was the ‘gold standard’ the number of Meridian Premier toxin AþB EIA false negatives would be relatively small. We have not, so far, encountered patients who were negative by EIA and subsequently represented with severe confirmed C. difficile disease: currently all our negative EIA results are issued with a warning about the potential for false negatives and a recommendation to discuss any concerns with a medical microbiologist. We definitely agree with the observation by both by Clark et al. and by Goldenberg et al. that GDH screening followed by PCR can lead to what seems to be a much increased sensitivity.1,4 We would like to find out more about the clinical significance of the positives with this testing algorithm. If isolation of a toxigenic C. difficile strain is used as a gold standard the GDH/PCR approach is much more sensitive: but are we detecting patients colonised by C. difficile (whose diarrhoea may sometimes be caused by other pathologies such as viral gastroenteritis or diverticulitis or laxatives) or patients whose diarrhoea is caused by C. difficile? And would all these patients really benefit from treatment with either metronidazole or vancomycin? The clinical trials of C. difficile treatment with
Letters to the Editor / Journal of Hospital Infection 79 (2011) 93–98
antibiotics (such as vancomycin or metronidazole) were typically done without a placebo arm and the patients included had a positive C. difficile stool toxin test with either EIA or cell culture cytotoxicity.5–8 The isolation in side-rooms of all patients who are positive with GDH/PCR algorithm is likely to be beneficial, but GDH/PCRpositive patients, particularly those who are stool toxin negative, may not benefit from specific therapy. Conflict of interest statement None declared. Funding sources None. References 1. Clark NS, Doughton MJ, Karas JA, Enoch DA. Two-step algorithm for the detection of Clostridium difficile from stool samples. J Hosp Infect 2011; 79:95–96. 2. Bignardi GE, Settle CS. Glutamate dehydrogenase as confirmatory test for Clostridium difficile toxin A/B positive stools. J Hosp Infect 2010;75:327–328. 3. 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. 4. Goldenberg SD, Cliff PR, Smith S, Milner R, French GL. Two-step glutamate dehydrogenase antigen real-time polymerase chain reaction assay for detection of toxigenic Clostridium difficile. J Hosp Infect 2010;74:48–54. 5. Musher DM, Logan N, Bressler AM, Johnson DP, Rossignol JF. Nitazoxanide versus vancomycin in Clostridium difficile infection: a randomized, double-blind study. Clin Infect Dis 2009;48:41–46. 6. Zar FA, Bakkanagari SR, Moorthi KM, Davis MB. A comparison of vancomycin and metronidazole for the treatment of Clostridium difficile-associated diarrhea, stratified by disease severity. Clin Infect Dis 2007;45:302–307. 7. Lagrotteria D, Holmes S, Smieja M, Smaill F, Lee C. Prospective, randomized inpatient study of oral metronidazole versus oral metronidazole and rifampin for treatment of primary episode of Clostridium difficile-associated diarrhea. Clin Infect Dis 2006;43:547–552. 8. Musher DM, Logan N, Hamill RJ, et al. Nitazoxanide for the treatment of Clostridium difficile colitis. Clin Infect Dis 2006;43:421–427.
G.E. Bignardi* C. Settle Microbiology Department, Sunderland Royal Hospital, Sunderland, UK * Corresponding author. Address: Microbiology Department, Sunderland Royal Hospital, Kayll Road, Sunderland SR4 7TP, UK. Tel.: þ44 (0)191 5656256; fax: þ44 (0)191 5410531. E-mail address: [email protected] (G.E. Bignardi).
Accepted by S.J. Dancer Available online 11 June 2011 Ó 2011 The Healthcare Infection Society. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.jhin.2011.04.001
Spread and persistence of Clostridium difficile spores during and after cleaning with sporicidal disinfectants Madam, The presence of Clostridium difficile spores on surfaces within the near-patient environment is a risk factor for the acquisition of C. difficile infection (CDI).1 Nosocomial transmission of C. difficile
97
can occur via the hands of healthcare workers.2 An increase in the level of C. difficile spores on environmental surfaces is mirrored by an increase in the number of spores present on the hands of healthcare workers.3 Therefore, effective routine surface disinfection is imperative in reducing the risk of patients acquiring CDI. Department of Health guidelines recommend the use of chlorinecontaining cleaning agents to prevent CDI.4 Commercially available sporicides include chlorine-releasing agents such as hypochlorites and sodium dichloro-isocyanurate (NaDCC), although other products exist. However, efforts to disinfect surfaces can be compromised. In one study, Acinetobacter lwoffii was inadvertently spread in a ward during cleaning when the manufacturer’s instructions for use of a disinfectant were not strictly followed.5 The choice of disinfectant must suit the cleaning regimen adopted. In this study, we assessed the spread and persistence of C. difficile spores during and after cleaning with two commercially available sporicidal disinfectants. Spore suspensions of C. difficile (polymerase chain reaction ribotype 027; clinical isolate) were prepared as described previously.6 A presanitized laminated work surface was marked with 50 25-cm2 test areas. One millilitre of the suspension (w106 spores) was inoculated on to a single test square and spread over the 25-cm2 surface area. The inoculated surface was wiped with a microfibre cloth and the same cloth was used to wipe four clean (uninoculated) test surfaces (i.e. four consecutive transfers). Prior to use, cloths were treated with tap water (control) or one of two commercially available sporicidal disinfectants [a chlorine-dioxide-generating or a chlorine-releasing (NaDCC) product]. Cleaning was performed by the same individual to avoid the introduction of technique bias, and test areas were sampled either immediately after cleaning or after leaving the wiped surfaces for 60 min. Test areas were sampled with a cotton-tipped swab premoistened with phosphate-buffered saline. Prior to plating, swabs were transferred to 10 mL of neutralizing solution [sodium thiosulphate (0.1% w/v), Tween 80 (3% w/v), lecithin (0.3% w/v) prepared in sterile phosphate-buffered saline solution]. Preliminary validation of the neutralizing solution showed that this was effective against any residual sporicidal activity of the disinfectants, and there was no inhibition of subsequent colony formation. All experiments were replicated five times. In the absence of a sporicide, wiping a surface with a damp microfibre cloth reduced the number of spores on the surface from 2.5 106 to 3.8 103 spores. Subsequent use of the cloth transferred between 1.0 103 and 5.4 103 spores to each of four clean surfaces (i.e. four consecutive wipes; Table I). Addition of a sporicide had no significant effect on the number of spores removed from a contaminated surface, nor the number of spores transferred during cleaning (P > 0.05; Table I). Sampling the surfaces immediately after cleaning permitted little or no contact time for effective sporicidal activity. However, when contact times were increased to 60 min, no significant reduction in spore counts was observed (P > 0.05). The persistence of spores may be attributed to the presence of suboptimal levels of sporicide on the surface after wiping. The volume of sporicide deposited by a cloth during cleaning may be too low to have effective activity against surface-associated spores, even after extended contact times (60 min). The potency of sporicides containing chlorine dioxide can fall to subsporicidal concentrations during use. Chatuev and Peterson showed that spreading a disinfectant containing chlorine dioxide on to surfaces during cleaning was ineffective against spores because the chlorine dioxide gas rapidly evaporated from the solution.7 This study highlights the limitations of two commercially available sporicidal disinfectants against C. difficile spores. Preliminary suspension tests confirmed that both sporicides exhibited sporicidal activity sufficient to pass EN 13704 standards (>3 log reduction in 60-min contact time) under clean conditions (data not shown).
Letters to the Editor / Journal of Hospital Infection 79 (2011) 93–98
than others of similar design to ours.2,4,5 This study supports the use of GDH as part of a two-step algorithm due to its high sensitivity.6 GDH is also cheap and confirmatory tests were only required in 14% of cases, which is similar to other studies.2 This study did not assess PCR alone, which some studies found superior to other methods, but PCR remains prohibitively expensive in the UK.7 Although we found that GDH improved the performance of EIA, we believe the low sensitivity of EIA prevents its use as both a screening test and as a confirmatory test. PCR following GDH appears to be the most sensitive test with the highest PPV and NPV overall but is the most expensive. The increased sensitivity (15 positive samples by GDH/PCR compared to 12 by EIA) may also adversely affect targets present in the UK. Further studies are required to determine the best method of diagnosis for patient care in terms of test performance and clinical outcome. Conflict of interest statement None declared. Funding sources Alere Ltd (Stockport, UK) provided the Dynex DS2 analyser and C. diff CheckÔ 60 kits for GDH assay. Cepheid provided GeneXpert C. difficile PCR assays. Internal funding was provided from the Clinical Microbiology & Public Health Laboratory, Peterborough & Stamford Hospitals NHSFT.
References 1. Bignardi GE, Settle C. Glutamate dehydrogenase as confirmatory test for Clostridium difficile toxin A/B-positive stools. J Hosp Infect 2010;75:327–328. 2. Goldenberg SD, Cliff PR, Smith S, Milner M, French GL. Two-step glutamate dehydrogenase antigen real-time polymerase chain reaction assay for detection of toxigenic Clostridium difficile. J Hosp Infect 2010;74:48–54. 3. Department of Health & Health Protection Agency. Clostridium difficile infection: how to deal with the problem. London: Department of Health; 2009. 4. Planche T, Aghaizu A, Holliman R, et al. Diagnosis of Clostridium difficile infection by toxin detection kits: a systematic review. Lancet Infect Dis 2008;8:777–784. 5. 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. 6. Fenner L, Widmer AF, Goy G, Rudin S, Frei R. Rapid and reliable diagnostic algorithm for detection of Clostridium difficile. J Clin Microbiol 2008;46:328–330. 7. Novak-Weekley SM, Marlowe EM, Miller JM, et al. Clostridium difficile testing in the clinical laboratory using multiple testing algorithms. J Clin Microbiol 2010; 48:889–893.
N.S. Clarka M.J. Doughtona J.A. Karasb D.A. Enocha,* a Clinical Microbiology & Public Health Laboratory, Peterborough & Stamford Hospitals NHS Foundation Trust, Peterborough City Hospital, Peterborough, UK b
Clinical Microbiology & Public Health Laboratory, Health Protection Agency, Papworth Hospital, Papworth Everard, Cambridge, UK *
Corresponding author. Address: Clinical Microbiology & Public Health Laboratory, Peterborough & Stamford Hospitals NHS Foundation Trust, Peterborough City Hospital, Bretton Gate, Peterborough PE3 9GZ, UK. Tel.: þ44 (0) 1733 678443; fax: þ44 (0) 1733 349721. E-mail address: [email protected] (D.A. Enoch). Accepted by S.J. Dancer Available online 2 June 2011
Ó 2011 The Healthcare Infection Society. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.jhin.2011.04.002
Reply: Two-step algorithm for the detection of Clostridium difficile from stool samples
Madam, We read with interest the letter by Clark et al. which comments on our previous letter and discusses the merits of various testing modalities for Clostridium difficile infection.1,2 When assessing the performance of C. difficile enzyme immunoassay (EIA) testing it is important to consider which EIA kit has been used, as the performance varies.3 We had used the Meridian Premier toxin AþB kit, because this is thought to be the toxin EIA with the highest sensitivity.3 When analysing the data from Clark et al. we notice that for the EIA test they used (VIDAS C. difficile toxin A & B) results are presented for 316 stools, whereas for the glutamate dehydrogenase (GDH) test (C. diff CheckÔ 60) results are presented for 322 stools.1 We presume that the difference is due to the fact that the VIDAS EIA kit can give equivocal results and that these were excluded from the analysis. Clark et al. commented on the fact that GDH screening followed by EIA testing improved the performance of EIA as a single test: this improvement was limited to the specificity and the positive predictive value (the number of false positives is reduced) but there is no gain in sensitivity.1 Toxin EIA testing cannot increase the overall sensitivity, in a two-step algorithm with GDH followed by toxin EIA, unless a third testing modality is added for the discrepant results. We have achieved a similar objective (reducing the number of false positives) with what we think is a more userfriendly protocol of toxin EIA followed by GDH immunochromatography, as opposed to the sequence used by Clark et al.1 of a GDH EIA followed by a toxin EIA. In order to estimate the frequency of false-negative results with our toxin EIA followed by GDH algorithm, we have recently retested 397 EIA toxinnegative stools and found that 26 were GDH positive but only one of these was positive with a cell culture cytotoxicity test with hep2 cells: thus if cell culture cytotoxicity was the ‘gold standard’ the number of Meridian Premier toxin AþB EIA false negatives would be relatively small. We have not, so far, encountered patients who were negative by EIA and subsequently represented with severe confirmed C. difficile disease: currently all our negative EIA results are issued with a warning about the potential for false negatives and a recommendation to discuss any concerns with a medical microbiologist. We definitely agree with the observation by both by Clark et al. and by Goldenberg et al. that GDH screening followed by PCR can lead to what seems to be a much increased sensitivity.1,4 We would like to find out more about the clinical significance of the positives with this testing algorithm. If isolation of a toxigenic C. difficile strain is used as a gold standard the GDH/PCR approach is much more sensitive: but are we detecting patients colonised by C. difficile (whose diarrhoea may sometimes be caused by other pathologies such as viral gastroenteritis or diverticulitis or laxatives) or patients whose diarrhoea is caused by C. difficile? And would all these patients really benefit from treatment with either metronidazole or vancomycin? The clinical trials of C. difficile treatment with
Letters to the Editor / Journal of Hospital Infection 79 (2011) 93–98
antibiotics (such as vancomycin or metronidazole) were typically done without a placebo arm and the patients included had a positive C. difficile stool toxin test with either EIA or cell culture cytotoxicity.5–8 The isolation in side-rooms of all patients who are positive with GDH/PCR algorithm is likely to be beneficial, but GDH/PCRpositive patients, particularly those who are stool toxin negative, may not benefit from specific therapy. Conflict of interest statement None declared. Funding sources None. References 1. Clark NS, Doughton MJ, Karas JA, Enoch DA. Two-step algorithm for the detection of Clostridium difficile from stool samples. J Hosp Infect 2011; 79:95–96. 2. Bignardi GE, Settle CS. Glutamate dehydrogenase as confirmatory test for Clostridium difficile toxin A/B positive stools. J Hosp Infect 2010;75:327–328. 3. 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. 4. Goldenberg SD, Cliff PR, Smith S, Milner R, French GL. Two-step glutamate dehydrogenase antigen real-time polymerase chain reaction assay for detection of toxigenic Clostridium difficile. J Hosp Infect 2010;74:48–54. 5. Musher DM, Logan N, Bressler AM, Johnson DP, Rossignol JF. Nitazoxanide versus vancomycin in Clostridium difficile infection: a randomized, double-blind study. Clin Infect Dis 2009;48:41–46. 6. Zar FA, Bakkanagari SR, Moorthi KM, Davis MB. A comparison of vancomycin and metronidazole for the treatment of Clostridium difficile-associated diarrhea, stratified by disease severity. Clin Infect Dis 2007;45:302–307. 7. Lagrotteria D, Holmes S, Smieja M, Smaill F, Lee C. Prospective, randomized inpatient study of oral metronidazole versus oral metronidazole and rifampin for treatment of primary episode of Clostridium difficile-associated diarrhea. Clin Infect Dis 2006;43:547–552. 8. Musher DM, Logan N, Hamill RJ, et al. Nitazoxanide for the treatment of Clostridium difficile colitis. Clin Infect Dis 2006;43:421–427.
G.E. Bignardi* C. Settle Microbiology Department, Sunderland Royal Hospital, Sunderland, UK * Corresponding author. Address: Microbiology Department, Sunderland Royal Hospital, Kayll Road, Sunderland SR4 7TP, UK. Tel.: þ44 (0)191 5656256; fax: þ44 (0)191 5410531. E-mail address: [email protected] (G.E. Bignardi).
Accepted by S.J. Dancer Available online 11 June 2011 Ó 2011 The Healthcare Infection Society. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.jhin.2011.04.001
Spread and persistence of Clostridium difficile spores during and after cleaning with sporicidal disinfectants Madam, The presence of Clostridium difficile spores on surfaces within the near-patient environment is a risk factor for the acquisition of C. difficile infection (CDI).1 Nosocomial transmission of C. difficile
97
can occur via the hands of healthcare workers.2 An increase in the level of C. difficile spores on environmental surfaces is mirrored by an increase in the number of spores present on the hands of healthcare workers.3 Therefore, effective routine surface disinfection is imperative in reducing the risk of patients acquiring CDI. Department of Health guidelines recommend the use of chlorinecontaining cleaning agents to prevent CDI.4 Commercially available sporicides include chlorine-releasing agents such as hypochlorites and sodium dichloro-isocyanurate (NaDCC), although other products exist. However, efforts to disinfect surfaces can be compromised. In one study, Acinetobacter lwoffii was inadvertently spread in a ward during cleaning when the manufacturer’s instructions for use of a disinfectant were not strictly followed.5 The choice of disinfectant must suit the cleaning regimen adopted. In this study, we assessed the spread and persistence of C. difficile spores during and after cleaning with two commercially available sporicidal disinfectants. Spore suspensions of C. difficile (polymerase chain reaction ribotype 027; clinical isolate) were prepared as described previously.6 A presanitized laminated work surface was marked with 50 25-cm2 test areas. One millilitre of the suspension (w106 spores) was inoculated on to a single test square and spread over the 25-cm2 surface area. The inoculated surface was wiped with a microfibre cloth and the same cloth was used to wipe four clean (uninoculated) test surfaces (i.e. four consecutive transfers). Prior to use, cloths were treated with tap water (control) or one of two commercially available sporicidal disinfectants [a chlorine-dioxide-generating or a chlorine-releasing (NaDCC) product]. Cleaning was performed by the same individual to avoid the introduction of technique bias, and test areas were sampled either immediately after cleaning or after leaving the wiped surfaces for 60 min. Test areas were sampled with a cotton-tipped swab premoistened with phosphate-buffered saline. Prior to plating, swabs were transferred to 10 mL of neutralizing solution [sodium thiosulphate (0.1% w/v), Tween 80 (3% w/v), lecithin (0.3% w/v) prepared in sterile phosphate-buffered saline solution]. Preliminary validation of the neutralizing solution showed that this was effective against any residual sporicidal activity of the disinfectants, and there was no inhibition of subsequent colony formation. All experiments were replicated five times. In the absence of a sporicide, wiping a surface with a damp microfibre cloth reduced the number of spores on the surface from 2.5 106 to 3.8 103 spores. Subsequent use of the cloth transferred between 1.0 103 and 5.4 103 spores to each of four clean surfaces (i.e. four consecutive wipes; Table I). Addition of a sporicide had no significant effect on the number of spores removed from a contaminated surface, nor the number of spores transferred during cleaning (P > 0.05; Table I). Sampling the surfaces immediately after cleaning permitted little or no contact time for effective sporicidal activity. However, when contact times were increased to 60 min, no significant reduction in spore counts was observed (P > 0.05). The persistence of spores may be attributed to the presence of suboptimal levels of sporicide on the surface after wiping. The volume of sporicide deposited by a cloth during cleaning may be too low to have effective activity against surface-associated spores, even after extended contact times (60 min). The potency of sporicides containing chlorine dioxide can fall to subsporicidal concentrations during use. Chatuev and Peterson showed that spreading a disinfectant containing chlorine dioxide on to surfaces during cleaning was ineffective against spores because the chlorine dioxide gas rapidly evaporated from the solution.7 This study highlights the limitations of two commercially available sporicidal disinfectants against C. difficile spores. Preliminary suspension tests confirmed that both sporicides exhibited sporicidal activity sufficient to pass EN 13704 standards (>3 log reduction in 60-min contact time) under clean conditions (data not shown).