APIC New England 2014 Conference Report: ‘The how, when, & where of C. diff – can you “C” the difference?’

apic new england logo

I was privileged to be asked to speak at the APIC New England Conference today in Springfield, Massachusetts. It was a vibrant day, and congratulations to the organizing committee for putting on such an enjoyable event.

Marie-Louise Landry MD – Continuing conundrums and controversies in the laboratory diagnosis of Clostridium difficile.

Dr Landy, a virologist by trade, began by reflecting on the fact that CDI is relatively new phenomenon, discovered in the late 1970s and initially thought to be viral! Having briefly presented the clinical problem and pathogenesis of CDI, Dr Landry got to the nitty gritty of how to test for CDI. You can choose to target the toxin, bacteria or bacteria capable of producing toxin:

  • Toxin: cytotoxicity cell culture assay (complex and requires overnight incubation) or enzyme immuno assay (terrible sensitivity).
  • Bacteria: culture (slow and doesn’t tell you much) or rapid GDH antigen assay (sensitive, but requires a confirmatory test of toxigenicity).
  • Bacteria capable of producing toxin: toxigenic culture (requires incubation) or Nucleic Acid Amplification Test (NAAT) such as PCR or LAMP (rapid, expensive).

To be honest, before Dr Landry’s talk, I thought that testing for CDI was pretty much sorted: GDH as a sensitive screening test following by PCR to detect the toxin gene for GDH positives. However, Dr Landry presented a compelling case that whilst GDH makes sense as a screening test, detecting the toxin gene via PCR is only half the story: the real gold-standard test is a cytotoxicity cell culture assay to confirm that the disease-causing toxin is present. Cost & clinical association makes compelling case for cell culture cytotoxicity assays; convenience for PCR! Indeed is the initial enthusiasm for PCR CDI testing waning as reality sets in (like the famous ‘Going and Coming’ by Rockwell)?

A final point for discussion: you can have the best laboratory diagnostics in the world, but if you’re testing inappropriate specimens, you’ll end up with false positives. We need a firm “no diarrhea, no CDI diagnostics” rule!

Curtis Donskey MD – Controlling the spread of C. difficile: a multifaceted approach

Dr Donskey began by considering that no healthcare facility is an island, and that long-term care facilities are an integral part of CDI spread. Dr Donskey spent most of the talk considering the environmental considerations related to CDI. Why does cleaning fail? Due to poor implementation: a research team with a bucket of bleach can eliminate C. difficile from surfaces! Various tools are available to help us tackle C. difficile environmental contamination. However, fluorescent markers and UVC did not eliminate C. difficile contamination whereas carefully enhanced disinfection did; bleach goes round corners better than UV, apparently. A related (and under-reported) unintended consequence of introduction a “no-touch” room disinfection (NTD) system such as UVC is that cleaners stop cleaning, mistaking UVC for magical cleaning robots! Plus, you could find yourself spending more time screening than cleaning, to the extent that those tasked with monitoring the cleaning process would be better deployed by getting their hands dirty! Dr Donskey covered a number of other important environmental issues: who cleans what (“the nurses thought EVS were doing it; EVS thought the techs were doing it; nobody was doing it”), the need for daily disinfection, pre-emptive and extended isolation, the potential and under-recognized importance of proper daily bathing for CDI patients, and the potential contamination risk from asymptomatic carriers. The final word: “getting doctors to prescribe antibiotics appropriately is like getting EVS to clean properly: an ongoing challenge.” Oh, and he finished on a song.

Jon Otter PhD (who invited him?) – No-touch room disinfection (NTD) systems: when to use them and how to choose between them (Can you ‘C’ the difference?)

You can download my slides from the talk here.

The talk was loosely based around a review paper recently published in JHI. The increased risk from the prior room occupant argues for doing a better job of terminal disinfection. The goal of hospital disinfection is controversial: the ‘Pragmatist’ says a reduction in contamination is good enough, whereas the ‘Prior room occupantist’ says elimination of pathogens is required. I presented some data suggesting that transmission risk ∝ contamination level; ergo reduction in transmission ∝ decontamination level? The NTD scene is a four-horse race currently, with hydrogen peroxide vapour (HPV), aerosolized hydrogen peroxide (AHP), ultraviolet C (UVC), and the relatively new kid on the block, pulsed xenon UV (PX-UV). Each system has its pros and cons so which is best? My view is that will depend on the scenario: if you have a carbapenem-resistant Acinetobacter baumannii in your ICU, then the ‘belt and braces’ approach of HPV is warranted. However, if you have MRSA colonization on a medical ward, a ‘quick and easy’ UV treatment may the only feasible option.

To try to keep everybody awake after lunch, I polled the audience on a few questions (Figure). I was not surprised that most people had not used an NTD system. However, I was surprised that so few people selected UV in the scenarios!

APIC NE q1 APIC NE q2 APIC NE scenarios

Figure. Question 1: Should all acute hospitals be using a ‘no-touch’ automated room disinfection (NTD) system for terminal disinfection of some patient rooms? Question 2: Has your hospital has used the following NTD systems? Scenario 1: A patient with carbapenem-resistant A. baumannii is discharged from the ICU. Scenario 2: A patient with MRSA colonization is discharged a general medical unit. Scenario 3: A patient with recently resolved CDI is discharged from a general medical unit (‘Enhanced’ = enhanced conventional methods).

Mike McCarthy – Sustaining your gains in infection control initiatives

Mike McCarthy rounded off the day with an engaging overview of his experience from a number of industries of how to ‘sustain your gains’. There’s a temptation from administrators to dismantle the team once it has been shown to work; clearly, the results will disappear with the people! Mike gave useful advice on how to embed change in an individual and organization. Do not confuse respect for people with respect for their bad practices. We need to be good coaches of best practice – reinforce proper execution; correct improper execution. The typical number of audits is “once and done”, but this not enough to form good habits. Establishing a new habit takes 60-90 days of work to reach the happy state of ‘unconscious competence’. People like data-led feedback (we’re all nerds at heart), which results in tangible performance management and improvement.  So, implement a checklist, audit it, provide positive reinforcement and feedback and your gain will be sustained!

Points for discussion:

  • Laboratory diagnostics are only part of the story. We need to focus on making sure only appropriate specimens are tested. Dr Donskey mentioned that a shocking 12% of their stool specimens were not tested due to sample leaking or labeling errors. Unfortunately, the stools most likely to be from CDI are also most likely to be liquid! Conversely, testing formed stools doesn’t do anybody any favours.
  • Do we need to focus on asymptomatic toxigenic C. difficile carriers and, if so, how?
  • How far can conventional methods go in tacking environmental contamination with C. difficile and is it time to turn to NTD systems, at least some of the time?
  • How best to sustain our gains?

The pitfalls of PCR for detecting pathogens on surfaces

PCR has proven an invaluable tool for the rapid diagnosis of a range of pathogens, including MRSA and C. difficile. Several studies have evaluated the potential use of PCR for the detection of pathogens on surfaces and have identified some issues that, frankly, seem pretty terminal for this application using currently available commercial PCR kits.

A study from Cleveland evaluated the use of a commercial RT-PCR test for detecting C. difficile on hospital surfaces. Three composite sites were sampled in 22 patient rooms, 41% of which housed a patient with CDI with the remaining 59% sampled after terminal cleaning and disinfection. Two swabs and a gauze were collected from each site; one swab was cultured directly onto selective agar and the other was tested using PCR. The gauze was cultured using broth enrichment. C. difficile that grew on the selective agar were tested for toxin production and only toxigenic C. difficile were included.

Overall, 23 (35%) of the 66 sites grew toxigenic C. difficile and only 4 of these were detected using the standard RT-PCR assay (sensitivity 17%, specificity 100%). The sensitivity of RT-PCR in rooms that had been cleaned and disinfected was even worse (10%). Increasing the CT threshold of the assay (making it less stringent) improved the overall sensitivity to 52% and did not affect the specificity.

The study has several important limitations. The RT-PCR assay detected only the Toxin B gene, whereas the toxigenic culture methodology would detect both Toxin A and B producers. More importantly, there was a crucial difference in sampling methodology: the gauzes used for broth enrichment culture had a 50% higher positivity rate than the swabs (in line with other findings), but only swabs were tested by both PCR and culture. Thus, if the gauzes are a more effective sampling device, this would make the RT-PCR methodology seems worse than it is. I would have liked to have seen the sensitivity of the RT-PCR assay for detecting C. difficile cultured from the swabs only, but I could not derive this from the data in the paper.

An older study from New Haven, Connecticut provides a contrasting view of the use of PCR to detect pathogens from surfaces. Here, 10 standardized sites were sampled in the rooms of 10 patients infected or colonized with MRSA, and 5 rooms of patients not known to be infected or colonized with MRSA. Swabs were directly plated onto selective agar for MRSA, then DNA was extracted from the swabs before a broth enrichment procedure using the same swabs. In this study, 40 (27%) of the 150 surfaces were positive by culture, but 90 (60%) were positive by PCR (sensitivity 93%, specificity 51%).

Deshpande 2013

Figure 1. Contrasting sensitivity and specificity when using PCR to detect C. difficile and MRSA on hospital surfaces.

It seems then that the sensitivity of PCR is too low for the environmental detection of C. difficile but the specificity is too low MRSA (figure 1). How could this be? Assuming that this is not due to experimental differences between the studies, it could be that the standard extraction procedure used for the C. difficile assay was not robust enough to liberate DNA from the mature environmental spores, resulting in low sensitivity. Conversely, the PCR assay was detecting DNA from dead MRSA on surfaces, resulting in low specificity.

So, in summary, the MRSA assay was too sensitive and the C. difficile assay was not sensitive enough! While the use of these “off the shelf” commercial assays doesn’t seem to be useful for detecting pathogens on surfaces, there may be hope for a PCR assay tailored specifically for an environmental application.

Article citations:

Deshpande A, Kundrapu S, Sunkesula VC, Cadnum JL, Fertelli D, Donskey CJ. Evaluation of a commercial real-time polymerase chain reaction assay for detection of environmental contamination with Clostridium difficile. J Hosp Infect 2013;85:76-78.

Otter JA, Havill NL, Boyce JM. Evaluation of real-time polymerase chain reaction for the detection of methicillin-resistant Staphylococcus aureus on environmental surfaces. Infect Control Hosp Epidemiol 2007;28:1003-1005.