Microbiological surface sampling cannot be described as new, with reports of its use going back to the 1920s and 1930s (Saelhof and Heinekamp, 1920, Krogg and Dougherty, 1936), although precise methodological details are lacking. However, most of this early work was based on swabbing, with direct agar contact methods only developed later, although the future is likely to see greater use of molecular methods. The main microbiological methods in use within the food industry include the use of swabs, sponges, or wipes to recover organisms from the surface followed by their cultivation on/in nutrient media (effectively indirect). The rationale for such testing can be either to semiquantitatively estimate the residual number of general or indicator organisms present, that is, to provide evidence of cleaning efficacy. Indicator organisms can reflect surface microbiological quality and whether conditions may permit the presence/growth of more specific pathogens. Often the latter may be like looking for a “needle in a haystack,” but is of particular benefit if:
Testing surfaces for the presence of pathogens, for example, L. monocytogenes, which could get into the food and cause problems, is a fundamentally different philosophical approach and is used to indicate risk. In this latter case it is usually a qualitative value that is needed rather than a semiquantitative one, that is, is a specific pathogen present. In testing for pathogens it is usual to test a larger surface area, for example, 1000 cm2 rather than the more conventional 100 cm2 (Willes et al., 2013). The medium inoculated by the swab used can be solid, semisolid, or liquid. In the former colonies counted on the surface are assumed to originate from one organism and this can contribute to variability. When wanting a count to reflect cleanliness for comparison purposes as part of a routine testing program a specific area (often 100 cm2) should be swabbed. If looking for the presence of a pathogen a large surface area should be tested. The requirement being—is a pathogen present or not? For such purposes large sponges (with or without a handle) are usually superior to swabs. Crucial in any microbiological surface testing is the recovery efficiency (RE) (Trafny et al., 2014) and this can vary by method, the number and types of microorganisms, and with the nature of the surface. Methods where the nutrient medium is in direct contact with the surface tested (contact plates and dipslides) are easier to use and could theoretically give superior recovery. How the comparison trials are set up can influence the results but in two large-scale comparisons (Salo et al., 2000, Salo et al., 2002) contact methods did give superior results, although the differences were not always significant. A problem with all cultivation methods is to remove the organisms from the surface in order to cultivate them. This has led to “rinsing” the surface to be tested (the rinse fluid is now used as the source of microorganisms), and is widely used where access to the surface can be difficult, for example, in CIP systems. More recently efforts to remove surface microorganisms, especially in biofilms, by sonication have been tried (Ismail et al., 2013). Apart from practical problems it raises questions about the importance and validity of the numbers recovered in relation to product contamination. The choice of microbial method will depend on the precise information required and the prevailing circumstances (Table 44.7 ). Comparison of Main Microbiological Methods for Hygiene Monitoring
A problem with cultivation methods can occur due to viable microorganisms going undetected due to stress, giving rise to viable but nonculturable (VBNC) bacteria. However, such organisms may or may not be able to cause spoilage or be infective/retain their pathogenicity. Nevertheless a positive result indicates that the surface has a history of previous contamination and could present a risk Swabbing in one form or another remains the oldest and probably the most widely used method for “surface monitoring” (Moore and Griffith, 2002a, Moore and Griffith, 2007). It should be noted that although the term monitoring is widely used this does often not conform to the HACCP definition. For the latter, results must be obtained in time for corrective action to be taken and swabbing, like impression plates, relies on cultivation, which, depending on the organism can take hours, days, or weeks (eg, for TB). Most swabbing protocols are based upon the swab-rinse technique originally developed by Manheimer and Yheunez in 1917 (Favero et al., 1968). A sterile swab, consisting of a more or less flexible shaft with a fibrous bud or tip, is premoistened in an appropriate wetting agent and inoculated by rubbing over the surface to be tested. The microorganisms transferred to the swab can then be cultivated and counted, either by inoculating the swab directly onto an appropriate solid culture medium or by releasing captured microorganisms into a known quantity of sterile recovery diluent, which is then used to prepare pour plates. This description of swabbing also indicates some of the variability in the technique (Moore and Griffith, 2007, Ismail et al., 2013, Downey et al., 2012), which can considerably affect the apparent number of organisms recovered (Moore and Griffith, 2002a, Moore and Griffith, 2007). If the number of microorganisms on a surface is known (as in laboratory conditions), and compared with the number obtained from swabbing, there is low recovery particularly at low surface population densities below 104 cells per cm2 (Holah et al., 1988). Additionally the swabbing technique lacks reliability, that is, repeatability and reproducibility are poor (Moore and Griffith, 2002a, Moore and Griffith, 2002b, Moore and Griffith, 2007, Moore et al., 2001). Various “standard” methods are available, including ISO 18593:2004, although currently there is no universally accepted method of swabbing. Some of the possible variables are indicated in Table 44.8 . Hygiene Swabbing Variables (Moore and Griffith, 2007)
Swabbing is widely used in industry to assess surface contamination, although not for larger surface areas (Ismail et al., 2013) and as a reference for comparison with other methods. However, basic information is still lacking as to the optimum protocol and the effect that variations may have on recovery rates (Moore and Griffith, 2007). Overall recovery can be seen as a function of the removal of microorganisms from the test surface, their release from the swab and their subsequent ability to grow. Recovery rates can vary from 0.1% to 25% (Moore and Griffith, 2007) and will depend on the technique used but an optimum recovery rate of 10% for Dacron swabs is not uncommon. The type and number of microorganisms sampled can have a major effect on recovery (Rose et al., 2011, Downey et al., 2012) and they can become increasingly difficult to remove once they have adhered to a surface (Cunliffe et al., 1999), particularly in biofilms. Additionally, organism retention within the bud fibers results in poor repeatability and sensitivity. Techniques/variables that improve one element of the swabbing process may adversely affect another. One study (Moore and Griffith, 2002a) showed protocols that improved removal, adversely affected release. Optimum overall recovery may therefore be a trade-off or compromise between different components of the whole process. The lack of repeatability can make it difficult to interpret the results from a single environmental swab, especially between staff, from different plants and when different protocols are used. An apparent low surface count from a single swab may reflect swabbing technique as much as low contamination levels. This could lead to a false impression of cleaning efficacy and whether guidelines or company specifications have been achieved. Swabbing, as with other surface assessment techniques, is best used to establish trends in the performance of the cleaning and disinfection program using multiple test results when over a period of time, the program can be seen to be failing or improving. The food manufacturers’ view is that the variability in swabbing per se, especially if standardized (Moore and Griffith, 2007), is not sufficient to prevent the detection of high surface counts on a given day, that is, the results of a badly implemented cleaning operation. Understanding the problems associated with overall recovery rates can help to improve and control the process (Moore and Griffith, 2007). Sampling/wetting solutions, designed to maintain isotonic conditions and reduce physiological stress, can be used to maintain the viability of microorganisms recovered from surfaces (Campden BRI, 2003). Care needs to be taken in their selection to ensure they do not artificially increase the count by providing a medium in which recovered microorganisms can grow during transit (Moore and Griffith, 2007). Some surfaces may still have residual disinfectant present and neutralizing agents, appropriate for the disinfectant being used, can be added to the wetting solution. These help to prevent organisms, removed from the surface (where they may be more resistant), being killed by residual sanitizer and thereby giving an “artificially” reduced count. Ideally swabs should be processed as soon as possible, although this is often impractical, especially when outside laboratories are used. Under such conditions, samples should be transported nonfrozen at a low temperature (<5°C), this can result in minimal differences compared with real-time analysis (Campden BRI, 2003). Times of sampling and processing need to be recorded, as well as delivery temperature, so that any unusual results or significant differences from the norm can be identified and considered when interpreting the results. Variables of time and wetting agent also need to be considered and optimized in sampling for specific pathogens. Appropriate pre-enrichment media should be used, although overgrowth by more rapidly growing nonpathogens needs to be considered. Some manufacturers may add a surfactant to their wetting solutions to improve “pick-up” from the test surface. These can, in some cases, artificially increase the number of colonies counted by breaking up clumps of organisms and thereby increasing the number of “colony-forming units.” Concerns over the inability of swab buds to release recovered organisms have prompted one manufacturer to develop a radically new type of swab (Moore and Griffith, 2007). This lacks the normal fibrous bud, which is replaced with short textured flocked nylon in spatula or swab format. This device releases more of the organisms removed from a surface and can yield an approximate 1 log improved overall recovery compared with traditional swabs. An alternative approach has led to the development of a wet or dry vacuum bacterial collection system. This may be of particular use in pathogen testing as it allows a much larger surface to be assessed without the need/use of a swab to lift/remove the organism from the surface being tested. Another variation involves self-contained “all-in-one media and hygiene swab” in a tube with the potential to offer more rapid results (Moore and Griffith, 2002b). A swab, after testing a surface, is returned to its accompanying culture tube containing a liquid or semisolid agar incorporating an indicator system. Microorganisms removed from the surface and retained by the swab grow and, as they multiply their growth can be detected, for example, by a color change. The results are semiquantitative in that the number of bacteria is not recorded but the time taken for the indicator to change color is a measure of the original microbial load. Unclean surfaces, depending upon the extent of microbial contamination, can test positive within 12 hours. When nonspecific media are used a general aerobic colony count is obtained, alternatively a selective or enrichment medium is used to test for the indicator organism or pathogens. Indicator systems can be based on chromogenic, fluorogenic, or bioluminogenic detection principles. In chromogenic assays the medium changes color as a consequence of microbial metabolism, and depending on the test indicates either the presence or absence of pathogen/group of organisms or the approximate degree of microbial surface contamination. Traditionally this could detect relevant organisms within as short a time as 18 hours (depending on the organism being tested for). More recently this time has been reduced by combining cultivation with a bioluminogenic test using a luminometer which can considerably reduce the detection time (Easter et al., 2012). This offers, depending on shift patterns, an opportunity for corrective action to be taken before further food production takes place. Using this approach, which correlates well with traditional counts, the time taken for detection extends from 1 hour, if the surface is heavily contaminated, to 8 hours for lightly contaminated surfaces. Such bioluminogenic tests are available for coliforms, Enterobacteriaceae, E. coli, and Listeria. Being able to perform a combined microbial cultivation and ATP assay extends the usefulness of luminometers beyond the conventional approach for estimating ATP in surface residues. Sponges work on a similar principle to swabbing, in that microorganisms are removed, released, and cultivated. Recovery is by wiping a compressed sterile sponge (eg, cellulose acetate) of varying sizes over the test surface. Some have no swab shaft, and in order to avoid contamination, the sponge needs to be held using a sterile glove, usually provided with the sponge. Sponges may be premoistened or require the addition of a wetting agent. After inoculation the sponge is returned to a sterile envelope/packet and transported to a laboratory. After the addition of a suitable diluent to the envelope, usually followed by agitation/stomaching, the released organisms can be counted. Similar errors to those encountered in swabbing may occur, and there is some evidence to suggest that the sponge matrix retains even more of the recovered organisms than swabbing, resulting in lower overall recovery (Moore and Griffith, 2002b). However sponges if returned to an enrichment medium for pathogen detection offer superior sensitivity and are not affected by the microorganisms being attached to the sponge matrix. Any organisms in the sponge go on to grow and multiply and contribute to a positive result. Some sponges also offer the advantage of greater surface area: being much bigger than conventional swabs they allow larger surface areas to be tested, and may therefore be more useful in testing surfaces for pathogens. Greater pressure can also be applied than with swabs. Other variations include sponges on sticks, and in France, the use of gauze to swab surfaces. Recently research has indicated that electrostatic wipes offered a better overall performance than swabs (Lutz et al., 2013), however validation data on the effectiveness of some of these alternatives under a wide range of conditions and organisms are not widely available. All direct agar contact methods, or replicate organism direct area contact (RODAC), involve pressing sterile agar onto a surface to be sampled. For this reason they are sometimes called “printing methods” (Ismail et al., 2013). A contact time of 10 seconds with a force of 25 g/cm2, without lateral movement, is suggested (ISO 14698: 2004). Microorganisms are directly transferred onto the agar surface and, after incubation for an appropriate length of time, multiply and form colonies, which are visible and can be counted. In general this approach is best suited to smooth, flat surfaces. The methods vary in how the agar is dispersed. Contact plates resemble small plastic Petri dishes with a lid. The agar is poured into them, leaving a convex contact surface. After removing the lid the agar is pressed onto the test surface. The contact plates are then incubated and examined 24–48 hours later. Agar immersion, plating, and contact (AIPC) slides, more commonly referred to as dipslides or paddles (in the United States), were developed from “dip spoons” used in counting the numbers of organisms in urine samples. They comprise a double-sided hinged paddle with a neutral or selective agar, attached to both sides. The paddle is contained within a transparent cylindrical tube or plastic container. The dipslide is removed, then pressed onto the surface to be tested, replaced back into the tube and resulting colony growth counted, or compared with pictorial estimates/diagrams of surface counts. They can also be used for counting the number of organisms in liquid samples of food, water, or rinse water. Recently a flexible hybrid contact plate/dipslide, to test more irregular-shaped surfaces, has become available. Other variations include the use of petrifilm to replace traditional agar plates for cultivation. These are small, thin films coated with nutrients and gelling agents. After wetting the film with approximately 1 mL of deionized water to rehydrate the growth medium, it can be used to provide a surface count. More recently a novel roller sampler was found to give a higher yield than conventional contact plates (Lutz et al., 2013). Direct agar contact methods have a number of advantages and disadvantages compared with traditional swabbing. Advantages include ease of use, generally lower costs, and better recovery and repeatability (Salo et al., 2000, Salo et al., 2002, Moore et al., 2001, Moore and Griffith, 2002b). Disadvantages include being more suited to flat surfaces and on very contaminated surfaces overgrowth can occur and this can make any statistical analysis of the results more problematic. However, this is not problematic if only an indication of cleaning adequacy, that is, pass or fail is required, rather than the precise number of organisms. It is easy to count the individual colonies, obtained from marginally unclean/clean surfaces, based on clean surface counts currently considered attainable (see Section 44.5.1). If a more precise number of colonies from a heavily contaminated surface is required then agar contact methods may be inappropriate. Sampling methods discussed so far have included assessment of chemicals such as ATP or protein (primarily for cleaning) although these chemicals are also found in cells/debris of nonmicrobial origin as well as more specific cultivation of microbial cells. A range of molecular methods is now available for the detection of groups, strains, or even specific subtypes of microorganisms including pathogens. Cultivation methods still require time, usually one or more days (although as can be seen this is being reduced) and molecular methods offer faster speed (although hours as opposed to seconds), greater sensitivity, and specificity. Often based on either DNA or RNA these methods target and amplify specific sections of a microorganism’s nucleic acid to a detectable level. Studies utilizing molecular methods have revealed the diversity found on some surfaces and have resulted in the characterization of the entire microbial community of an environmental sample (NIST, 2012). Techniques include polymerase chain reaction (PCR), reverse transcriptase (RT-PCR), and nucleic acid sequence-based amplification (NASBA). In real-time PCR the processes of amplification and detection are simultaneous. One potential disadvantage is that such techniques often do not distinguish between living microorganisms and noninfective nucleic acid and therefore only indicate that at some stage the organism was present on that surface—although they may be capable of detecting VBNC bacteria. At present, molecular methods require more technical expertise and high-cost equipment, are more expensive and are primarily used in outbreak investigations or to trace/track microorganisms within plants. However, with protocol advances it is likely in the future they will be used more routinely in assessing the effectiveness of primarily disinfection or in estimating risk. Ideally risk assessment also requires knowledge of the number of organisms and some molecular techniques can be made quantitative, for example, quantitative real-time PCR (qPCR). One laboratory study (Buttner et al., 2007) compared conventional cultivation with qPCR on a range of surfaces, although only one organism was tested. Cultivation techniques yielded few viable cells, whereas qPCR gave much higher results but represented nucleic acid from viable and nonviable cells. Depending on the analysis method used, sample pretreatment may be necessary, which can add to costs and lengthen the time taken. Page 2PMC full text: Copyright/LicenseRequest permission to reuse Copyright © 2016 Elsevier Ltd. All rights reserved. Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active. Case Studies on Cleaning, Cross-Contamination, and Listeria
Source: Case Study 1: Adapted from Blatter, S., Giezendanner, N., Stephan, R., Zweifel, C., 2010. Phenotypic and molecular typing of Listeria monocytogenes isolated from the processing environment and products of a sandwich-producing plant. Food Control 21, 1519–1523; Case Study 2: Adapted from CDC, 2008. Outbreak of Listeria monocytogenes infections associated with pasteurised milk from a local dairy—Massachusetts, 2007. Morbidity and Mortality Weekly Report 57 (40), 1097–1100. |
What product is used to reduce microbial count on surface but not necessarily to clean the surface?
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