Part I RisksRisks
3.2 Pathogens in biofilms
is common in parts of the world where milk is neither boiled nor pasteurised. It occurs, but much less frequently, in developed countries where the main products implicated are pasteurised milk, powdered milk and certain cheeses (Mead, 1993). Formation of a biofilm by Salmonella on various types of surfaces used in the food processing industry has been reported by several groups (Mafu et al., 1990; Helke et al., 1993; Ronner & Wong, 1993; Joseph et al., 2001). These studies have shown that Salmonella spp. can form biofilms on food contact surfaces and that the cells in biofilms are much more resistant to sanitisers compared to planktonic cells (Ronner & Wong, 1993; Joseph et al., 2001; Stepanovic et al., 2003). Mokgatla and co-workers (1998) studied the resistance of Salmonella sp. isolated from a poultry abattoir and found out that it will grow in the presence of in-use concentrations of hypochlorous acid. The presence of Pseudomonas fluorescens in the biofilm resulted in the increased resistance of S. Typhimurium to chlorine (Leriche & Carpentier, 1995).
3.2.2 Escherichia coli biofilms
Escherichia coli is a Gram-negative, rod-shaped bacterium. Because many microbes from faeces are pathogenic in animals and humans, the presence of the intestinal bacterium E. coli in water and foods indicates a potential hygiene hazard. Most strains of E. coli are harmless. However, a few strains with well-characterised traits are known to be associated with pathogenicity (Venkitanarayanan & Doyle, 2003). Those of greatest concern in water and foods are the intestinal pathogens, which are classified into five major groups:
the enterohaemorrhagic E. coli (EHEC), the enterotoxigenic E. coli (ETEC), the enteroinvasive E. coli (EIEC), the enteropathogenic E. coli (EPEC) and the enteroaggregative E. coli (EAEC). Growth can occur at 7±46 ëC with the maximal growth rate at 35±37 ëC. The minimum awfor growth ranges from 0.94 and 0.97. The optimum pH for growth is approximately 7.0, with a minimum and maximum pH for growth of 4.5 and 9.0. EHEC has been shown to grow poorly at temperatures of 44 ëC (Venkitanarayanan & Doyle, 2003).
Escherichia coli has been isolated from a large number of foods and drinks, e.g. fermented meat sausage, dairy products, vegetables, meat, poultry and fish products, water and apple cider. These agents can cause diarrhoeal outbreaks (Junkins & Doyle, 1993; Venkitanarayanan & Doyle, 2003). Unpasteurised milk is a common vehicle of E. coli O157:H7 transmission to humans (Dontorou et al., 2004). E. coli can also survive for extended periods of time in several acidic foods, e.g. cheese and yogurt. Acid-adapted E. coli O157:H7 has shown enhanced survival and prevalence in biofilms on stainless steel surfaces (Stopforth et al., 2003a,b; Venkitanarayanan & Doyle, 2003). In a hygiene survey performed in the food industry by Holah et al. (2002), microbial strains, e.g. E. coli and L. monocytogenes, were found either on surfaces or in products or in both, and some of these strains were persistent. Faille et al. (2002, 2003) found out that E. coli cells were poorly adhered to surfaces. The cells were embedded in the organic matrix of the biofilm, which shows that the structure of Biofilm risks 51
the biofilm formed affects the way in which the surfaces should be cleaned.
Oulahal-Lagsir et al. (2003) showed in their studies that proteolytic and glycolytic enzyme treatment together with ultrasonics enhance the removal of E.
coli biofilm from stainless steel soiled with milk. These findings correspond with results obtained in the food industry.
3.2.3 Campylobacter biofilms
Campylobacter spp. are microaerophilic, very small, curved and thin Gram-negative rods (Price & Tom, 2003a). Growth can occur in a microaerophilic atmosphere containing 3±15% oxygen and 3±5% carbon dioxide at 30±48 ëC with a maximal growth rate at 42±43 ëC. The minimum awfor growth is above 0.987. The optimum pH for growth is approximately 6.5±7.5, with a minimum at 4.9 and a maximum at 9.0 (Stern & Kazami, 1989; Roberts et al., 1996; van Vliet & Ketley, 2001). C. coli and C. laridis can grow at 30.5 ëC while C. jejuni cannot. C. laridis tolerates slightly more salt than C. jejuni or C. coli and ceases growing in the presence of 2.5% sodium chloride (Roberts et al., 1996). Illness can be caused by ingestion of as few as 500±800 cells in milk. Since the infective dose is rather low and the food in many cases may contain only a few cells, liquid enrichment methods are normally required before plating on selective media in order to detect contamination with C. jejuni or C. coli.
Successful detection of these organisms requires incubation at 42 ëC under microaerophilic conditions (Roberts et al., 1996).
In laboratory tests Campylobacter has been shown to form a biofilm in optimum circumstances on stainless steel and glass beads in 2 days (Somers et al., 1994; Dykes et al., 2003). In studies performed by de Beer et al. (1994) biofilms are shown to form zones with low oxygen content in aerobic surroundings and Campylobacter spp. can therefore more easily survive in biofilms. Trachoo et al. (2002) showed that viable C. jejuni cells grown on polyvinyl chloride surfaces decreased with time and the greatest reduction occurred on surfaces without a pre-existing biofilm. The number of viable C.
jejuni determined by using a direct viable count was greater than by using culturing techniques, which indicates that C. jejuni cells can form a viable but non-culturable state within the biofilm. Both determination methods showed that biofilms enhance the survival of C. jejuni during a 7-day period at 12 ëC and 23 ëC (Trachoo et al., 2002). Taking the resistance of the viable but non-culturable C. jejuni cells into account is important in the optimisation of cleaning and decontamination procedures, especially in those food industrial processes in which raw meat products are processed (Rowe et al., 1998; Trachoo
& Frank, 2002). Organic soil, e.g. food residues, or moisture improve the survival of campylobacter on surfaces (Humphrey et al., 1995; Kusamaningrum et al., 2003). Boucher and co-workers (1998) showed that C. jejuni survived very well on wooden surfaces because the pores in the wood protect the cells from oxygen.
52 Handbook of hygiene control in the food industry
3.2.4 Listeria monocytogenes biofilms
Listeria monocytogenes is a facultatively anaerobic Gram-positive, non-spore-forming short rod that is widely distributed in nature (El-Kest & Marth, 1988;
Griffiths, 2003). It is a non-host specific pathogen (El-Kest & Marth, 1988;
LundeÂn, 2004). Listeriosis may occur sporadically or epidemically. The organism has been isolated from raw milk, mastitic milk and pasteurised milk. Foodstuffs associated with listeriosis outbreaks also include cold-smoked and gravad rainbow trout products, sliced cold meat, soft cheese, butter, ice-cream and coleslaw. Examples of epidemic sources are: coleslaw in Canada 1981, unpasteurised milk in the USA 1983, Mexican-style soft cheese in USA 1985, pork product in France 1992, chocolate milk in the USA 1994, soft cheese in Swizerland 1995, rainbow trout in Sweden 1997, corn in Italy 1997, hot dogs in the USA 1998±99 and butter in Finland 1999 (LyytikaÈinen et al., 2000; Weinstein & Ortiz, 2001). Treated wastewater can also be a source of L. monocytogenes. Of the 13 different L. monocytogenes serotypes only three (1/2a, 1/2b and 4b) have been predominantly implicated in human diseases (Chae & Schraft, 2000). It has been reported that healthy people can be carriers of L. monocytogenes (El-Kest & Marth, 1988). L.
monocytogenes is able to grow in many environments, at a low oxygen tensions, in high salt concentrations and over a wide range of pH (5±9.5) and temperatures (3±45 ëC) with an optimum at 30 ëC (Griffiths, 2003; LundeÂn, 2004). The bacterium can survive for a limited time in up to 25% salt at 4 ëC (El-Kest & Marth, 1988).
Hygiene monitoring in the food processing industry is important because L.
monocytogenes, in particular, can colonise and form biofilms in food processing environments and on surfaces (Husu et al., 1990; Eklund et al. 1995; Autio et al., 1999; Miettinen et al., 1999, 2001; LyytikaÈinen et al., 2000; Aarnisalo et al., 2003; LundeÂn, 2004; Miettinen & Wirtanen, 2005; Wirtanen & Salo, 2004).
Listeria sources in processing plants are conveyor belts, cutters, slicers, brining and packaging machines, coolers and freezers as well as floors and drains (Wirtanen, 2002; LundeÂn, 2004; Wirtanen & Salo, 2004). L. monocytogenes has been found to form biofilms on common food contact surfaces such as plastic, polypropylene, rubber as well as stainless steel and also on glass (Mafu et al., 1990; Helke et al., 1993; Ronner & Wong, 1993; Wirtanen, 1995; Chae &
Shraft, 2000; Borucki et al., 2003; LundeÂn, 2004).
3.2.5 Staphylococcus aureus biofilms
Staphylococcus aureus is a Gram-positive, aerobic, non-spore-forming catalase postitive rod. It is ubiquitous in the mucous membrane and skin of most warm-blooded animals. Nasal and skin carriage are frequent vehicles in the transporta-tion of S. aureus. It is an opportunistic pathogen causing infectransporta-tions via open wounds, for example (Roberts et al., 1996). The growth temperature for this bacterium is 7±48 ëC, with an optimum around 37 ëC. Growth has been demon-strated over the pH range 4±10, with an optimum at 6±7. The lower limit of aw
Biofilm risks 53
permitting growth is 0.83. It readily produces enterotoxins, which are not destroyed in heat treatment (Roberts et al., 1996).
Staphylococcus aureus is a pathogen that can also affect dairy products. Its occurrence in sour milk products such as yoghurt is worthwhile investigating as it is present in relatively high numbers in raw milk (Benkerroum et al., 2002).
According to studies by Benkerroum et al. (2002), staphylococci grew rapidly during the initial fermentation. Similar behaviour by S. aureus has previously been reported both in yoghurt and cheese (Ahmed et al., 1983; Attaie et al., 1987). Elvers et al. (1999) isolated S. aureus in a total of 7% of food contact sites and 8% of environmental sites from 10 small and medium sized enterprises producing high risk foods in their study, which was performed for the Ministry of Agriculture, Fisheries and Food (now DEFRA) in the UK. The source of S. aureus almost always originated from food handlers or from utensils previously contaminated by humans (Elvers et al., 1999; Peters et al., 1999). A survey revealed that S. aureus was involved in 15% of the recorded foodborne illnesses caused by dairy products in eight developed countries whereas L. monocytogenes was involved in 22% (Benkerroum et al., 2002). It is resistant to drying and may also colonise complex food-processing equipment, which is left in wet conditions (Bolton et al., 1988). It can also be found in the dust in ventilation systems (Roberts et al., 1996). Resistance to oxidative disinfectants has mainly been associated with biofilm formation (Bolton et al., 1988). Luppens et al. (2002) showed that S. aureus biofilm formed on stainless steel, polystyrene and glass in a nutrient flow needed concentrations of benzalkonium chloride that were 50 times higher and concentrations of hypochlorite that were 600 times higher to achieve 4 log killing of S. aureus compared with cells in suspensions. Supporting results were obtained by Mùretrù et al. (2003a).
3.2.6 Bacillus cereus biofilms
Bacillus cereus is a Gram-positive, aerobic, spore-forming rod, normally present in soil, dust, and water (Jay, 1996). It can also grow well anaerobically. Cells of B. cereus are large and motile. The growth temperature for this bacterium is 4±
50 ëC, with an optimum around 28±35 ëC. Growth has been demonstrated over the pH range 4.9±9.3 (Jay, 1996; Granum, 2003; Shelef, 2003; Svensson et al., 2004). The organism elaborates a number of toxins with distinct diarrhoeal and emetic syndromes (Shelef, 2003). B. cereus occurs extensively in the environment but despite the fact that it is a common contaminant in raw milk, food poisoning outbreaks caused by dairy products contaminated with B. cereus have been rare (Wirtanen et al., 2002; Svensson et al., 2004).
In a dairy product survey, Wong (1998) showed that B. cereus was found in 52% of ice-creams, 35% of soft ice-creams, 29% of milk powders, 17% of fermented milks and 2% of pasteurised milks and fruit-flavoured milks.
Svensson et al. (1999) found indications of a prolonged contamination problem caused by mesophilic B. cereus strains early in the production chain of one 54 Handbook of hygiene control in the food industry
dairy plant. Additional contamination of milk by the B. cereus biofilm was shown to occur in the filling machine. Different Bacillus spp., and among them B. cereus, have been found on liquid packaging boards and blanks and these could thus be an additional source of biofilms containing Bacillus spp.
(Svensson et al., 2004). Furthermore, spores of B. cereus are reported to possess a pronounced ability to adhere to the surface of stainless steel, which is commonly used in food processing. Both B. cereus and B. subtilis biofilms were detected on stainless steel and Teflon surfaces, and removal from stainless steel was more difficult than from Teflon because of surface roughness (Wirtanen et al., 1996). Te Giffel and co-workers (1997) showed that spores of B. cereus adhered, germinated and multiplied on the stainless steel surfaces of a tube heat exchanger. The cells of B. cereus were isolated from the individual tubes after cleaning. The attachment of B. cereus in process lines may act as a continual source of post-pasteurisation contamination (Elvers et al., 1999). Lindsay (2001) found that the biofilms of food spoilage Bacillus and Pseudomonas species attach themselves to liquid food processing equipment surfaces and cells in biofilms, even if treated with an in-use concentration of sanitisers, manage to survive and grow. This phenomenon is even stronger when mixed biofilms are involved. The attached B. cereus cells may subsequently form a biofilm on a stainless steel surface and present a major problem for the food industry (Peng et al., 2002).
3.2.7 Clostridium perfringens biofilms
Clostridium perfringens is a spore-forming, Gram-positive, anaerobic, non-motile rod which forms large, regular, round and slightly opaque and shiny colonies on the surface of agar (Brynestad & Granum, 2002). There are five types of C. perfringens: A, B, C, D and E, which produce different types of toxins (LabbeÂ, 2003). C. perfringens can grow between 10 ëC and 52 ëC, with a maximum of 45 ëC for most strains (Brynestad & Granum, 2002). It is often a cause of human food poisoning due to its ability to grow over a wide temperature range. Its spores can also survive several food processing procedures. Spores of some strains are resistant to temperatures of 100 ëC for more than 1 h (LabbeÂ, 2003).
Clostridium perfringens food poisoning from new food sources, because the bacterium is so adaptable and prolific, has helped to show how our perceptions and understanding of safe food change with new knowledge (Foster, 1997). C.
perfringens can be found as part of the normal flora of the intestinal tracts of both animals and humans, as well as in soil, clothing and skin. It has been found in virtually all environments tested, including water, milk and dust (Brynestad &
Granum, 2002). In view of its widespread presence in moist soil, its presence in air and dust in kitchens, catering and food processing environments is not surprising (LabbeÂ, 2003). C. perfringens serotypes commonly associated with human illness have been found on recently slaughtered carcasses. Other foods contaminated with C. perfringens are poultry, fish, vegetables and dairy Biofilm risks 55
products (Roberts et al., 1996). As with other agents of human food poisoning, the number of outbreaks of food poisoning attributable to C. perfringens is under-reported (LabbeÂ, 1993).
3.2.8 Mycobacterium biofilms
The genus Mycobacterium contains approximately 50 species, which are divided into rapid growers, slow growers and the human leprosy bacillus (Collins & Grange, 1993). Mycobacteria are weakly Gram-positive, non-motile, slender, non-spore-forming, rod-shaped, aerobic and free-living in soil and water (Payeur, 2000). They do not produce appreciable amounts of toxin substances and do not cause food poisoning (Collins & Grange, 2003).
Mycobacteria are widely distributed in nature and have been isolated from natural and piped waters, wet soil, mud, compost, grasses, vegetables, unpasteurised milk and butter. They have also been isolated from domestic water pipes from which they readily enter drinking water (Collins & Grange, 2003). M. tuberculosis, M. africanum, M. bovis, M. bovis BCG and M. microti are collectively referred to as the M. tuberculosis complex because these organisms cause tuberculosis (Payeur, 2000). Infections in humans and animals may be caused by most of the slowly growing mycobacteria such as M. avium, M. intracellulare, M. scofulaceum, M. kansasii, M. marinum, M. simiae, M.
ulcerans and M. xenopi. The only rapidly growing pathogenic species are M.
chelonae and M. fortuitum. The principal source of these infections seems to be water (Payeur, 2000).
A pilot plant pasteuriser was used to examine the heat resistance of M. avium subsp. paratuberculosis (M. paratuberculosis) during high temperature short time (HTST) pasteurisation using raw milk samples under various time and temperature conditions. Results indicated that low numbers of M.
paratuberculosis may also survive extreme HTST treatments (Hammer et al., 2002). Torvinen et al. (2004) studied 16 drinking water distribution systems in Finland for growth of mycobacteria by sampling water from waterworks and in different parts of the systems. In the experimental part, mycobacterial colonisation as biofilms on polyvinyl chloride tubes was studied. The isolation frequency of mycobacteria increased from 35% at the waterworks to 80% in the systems, and the number of mycobacteria in the positive samples increased from 15 to 140 cfu/l, respectively. The densities of mycobacteria in the developing biofilms were highest at the distal sites of the system. Over 90% of the myco-bacteria isolated from water deposits belonged to M. lentiflavum, M. tusciae, M.
gordonae and a previously unclassified group of mycobacteria. Dailloux et al.
(2003) investigated the ability of M. xenopi to colonise an experimental drinking water distribution system. M. xenopi was found to be present in the biofilm within an hour of introduction. After 9 weeks, it was constantly present in all outlet water samples (1±10 cfu/100 ml) and in biofilm samples (102±103 cfu/
cm2). Biofilms may be considered to be the reservoirs for the survival of M.
xenopi. Gao et al. (2002) studied the survival of M. paratuberculosis in 7 regular 56 Handbook of hygiene control in the food industry
batch and 11 HTST pasteurisation experiments using raw milk or ultra-heat treatment (UHT) milk samples spiked with M. paratuberculosis. E. coli and M.
bovis BCG strains were used as controls. No survivors were detected from any of the slants or broths corresponding to the 7 regular batch, but survivors were detected in 2 of the 11 HTST experiments. No survivors were detected after heat treatment for 15 min at 63 ëC. These results indicate that M. paratuberculosis may survive HTST pasteurisation (Gao et al., 2002).