Effects of physicochemical surface characteristics of Listeria monocytogenes strains on attachment to glass

ARTICLE IN PRESS FOOD MICROBIOLOGY Food Microbiology 23 (2006) 250–259 www.elsevier.com/locate/fm

Effects of physicochemical surface characteristics of Listeria monocytogenes strains on attachment to glass Min Seok Chaea,1, Heidi Schraftb,, Lisbeth Truelstrup Hansenc, Robert Mackerethd a

Department of Food Science, University of Guelph, Guelph, ON, Canada N1G 2W1 Department of Biology, Lakehead University, 955 Oliver Road, Thunder Bay, ON, Canada P7B 5E1 c Department of Food Science and Technology, Dalhousie University, P.O. Box 1000, Halifax, NS, Canada B3J 2X4 d Centre for Northern Forest Ecosystem Research, Ontario Ministry of Natural Resources, c/o Lakehead University, 955 Oliver Road, Thunder Bay, ON, Canada P7B 5E1 b

Received 9 December 2004; received in revised form 8 April 2005; accepted 8 April 2005 Available online 24 June 2005

Abstract Seven strains of Listeria monocytogenes frequently involved in foodborne disease (epidemic strains) and 14 sporadic strains were examined to compare the attachment and subsequent biofilm growth on glass slides at 37 1C. Epidemic strains at 3 h incubation had significantly higher attachment values than sporadic strains (Po0:001), but subsequent biofilm growth over 24 h was not dependent on initial attachment. To better understand this phenomenon, the surface hydrophobicity and charge, as well as the extracellular carbohydrate content of the 21 L. monocytogenes strains were studied to determine if these surface characteristics had an effect on bacterial attachment to glass. Hydrophobicity was measured by the bacterial adherence to hydrocarbon (BATH) and polystyrene adherence methods. Hydrophobicity values obtained with the BATH method were linearly correlated with those from the polystyrene adherence method (r ¼ 0:64, Po0:001), but no correlation was found between hydrophobicity and bacterial attachment to glass. Hydrophobicity and surface charge measured as electrophoretic mobility (EM) were correlated (r ¼ 0:77, Po0:001); however, there was no correlation between the degree of attachment and surface charge. Colorimetric measurements of the total extracellular carbohydrates revealed that attached cells produced significantly (Po0:05) higher levels than planktonic cells after a 3 h time period. Analysis of co-variance (Nested ANCOVA) furthermore demonstrated that total carbohydrates produced by planktonic cells had a significant positive effect on 24 h biofilm growth (P ¼ 0:006). This is the first report to indicate that the ability of a L. monocytogenes strain to produce high levels of extracellular carbohydrates may increase its ability to form a biofilm. Genetic studies targeting carbohydrate synthesis pathways of L. monocytogenes will be required to fully understand the importance of this observation. r 2005 Elsevier Ltd. All rights reserved. Keywords: Listeria monocytogenes; Biofilm; Attachment; Hydrophobicity; Electrophoretic mobility; Extracellular polymeric substances

1. Introduction Listeria monocytogenes is a Gram-positive, facultative anaerobe intracellular bacterium that is widely distributed in nature and is also frequently isolated in food Corresponding author. Tel.: +1 807 343 8351; fax: +1 807 346 7796. E-mail address: [email protected] (H. Schraft). 1 Present address: Truspex Inc., 61-240 London Rd. W., Guelph, ON, Canada N1 H 8N8.

0740-0020/$ - see front matter r 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.fm.2005.04.004

processing environments (Ryser and Marth, 1991). The highest incidences are associated with wet areas of food plants, such as floors, floor drains, and conveyer belts, and areas that are difficult to clean, including gaskets, joints, and crevices (Wong, 1998). CDC officials (1999) consider foodborne listeriosis to be from raw food of animal origin, as well as postprocessing contamination of heat-treated ready-to-eat foods, especially deli meats, hot dogs, and sausages. There are 13 serovars of L. monocytogenes, but almost

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all human cases of listeriosis have been associated with types 4b, 1/2a, and 1/2b (Schuchat et al., 1991; Gellin and Broome, 1989). Type 4b seems to be most often associated with large outbreaks of foodborne listeriosis (McLauchlin, 1996). Based on DNA pattern analyses and multi-locus enzyme analysis, a specific clone of serotype 4b is phenotypically and genetically closely related to the isolates responsible for the large outbreaks in California, Switzerland, and France (Vines and Swaminathan, 1998; Wiedmann et al., 1997). The high frequency of large outbreaks caused by this clone could be due to an increased virulence, a better adaptation to survival in foods, a broader distribution in the environment, or an increased ability to survive in food processing environments via biofilm formation. Biofilm formation in food processing environments has become a concern for food manufacturers because sessile bacteria within biofilms are highly resistant to common cleansers and sanitizers, compared to bacteria in a planktonic state (Dhir and Dodd, 1995; Gilbert and Brown, 1995; Frank and Koffi, 1990). In the sessile state, bacteria may express different genes, alter their morphology, change their growth rates, or produce large amounts of extracellular polymeric substances (EPS) (Gilbert and Brown, 1995; McCarter et al., 1992). L. monocytogenes cells rapidly attach and form biofilms on food contact surfaces such as plastic, polypropylene, rubber, stainless steel, and glass (Chae and Schraft, 2001; Jeong and Frank, 1994a, b; Mafu et al., 1990). It has also been shown that L. monocytogenes can grow within mixed species biofilms at 10 1C, and that these biofilm bacteria are highly resistant to sanitizers (Blackman and Frank, 1996; Wirtanen and Mattila-Sandholm, 1992). Biofilm formation by L. monocytogenes may enhance bacterial persistence in food processing environments, and consequently increase the chances of contributing to post-processing contamination. The cell surface is generally considered a significant factor in bacterial attachment to surfaces. Many studies suggest that microbial cell surface charge and hydrophobicity play an important role in the initial steps of microbial adhesion. Numerous methods have been developed that attempt to determine physicochemical properties of microbial cell surfaces and the substratum. These include electrophoretic mobility (EM), bacterial adherence to hydrocarbon (BATH) and adherence to polystyrene (Parkar et al., 2001; Champlin et al., 1999). The hydrophobicities and surface charges of bacteria can differ between species, serotypes or strains, and can change with variation in growth conditions, physiological state of cells, and composition of suspension media (Briandet et al., 1999; Chavant et al., 2002; Giovannacci et al., 2000). Net cell surface charge can be determined based on the cells’ zeta potential and EM has been used to assess this surface net charge of bacteria in many investigations

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(Champlin et al., 1999; van der Mei et al., 1993, 1997). In microelectrophoresis, micro-organisms are suspended in a liquid phase and a voltage is applied across the cell. Negatively charged bacteria are then attracted to the positive electrode, while positively charged organisms are attracted to the negative electrode. The velocity at which the micro-organism moves is a direct measure of EM, which can be used to calculate the zeta potential (van der Mei et al., 1993; James, 1991). Bacterial cell surface hydrophobicity has been demonstrated by BATH and adherence to polystyrene (Parkar et al., 2001; Li and McLandsborough, 1999). Using the BATH method, an aqueous cell suspension is mixed with an organic solvent (usually hexadecane) and the turbidity of the aqueous portion is measured before and after mixing, to determine the percentage of cells lost to the organic phase. The larger the percentage of cells adhering to the hydrocarbon phase, the more hydrophobic the cell is assumed to be. Polystyrene is a highly hydrophobic material (Absolom et al., 1983), and the percentage of particles adhering to the polystyrene substrate under the mixing action of a micropipettor provides a reliable qualitative measure of the particle– surface adhesion energy (Visser, 1976). Production of EPS plays an important role in biofilm development of many bacteria and polysaccharides are considered significant components of EPS, although other materials such as nucleic acids and protein may also be present (Laspidou and Rittmann, 2002). Only little information has been published on EPS in L. monocytogenes biofilms, but Borucki et al. (2003) reported correlation of exopolysaccharide production and three-dimensional biofilm matrix formation for some L. monocytogenes strains. However, the effects of physicochemical surface properties and polysaccharides of L. monocytogenes on bacterial adherence are not well studied, which means that the mechanisms of attachment and colonization still remain largely unknown. The objectives of this study were to compare initial attachment and biofilm growth for various strains of L. monocytogenes frequently involved in foodborne disease (epidemic strains) with those of other isolates (sporadic strains). We also examined the importance of the cell surface characteristics and production of extracellular polysaccharides for L. monocytogenes’ ability to attach to glass.

2. Materials and methods 2.1. Bacterial strains, growth conditions, and preparation of bacterial suspensions A total of 21 strains of L. monocytogenes were used in this study (Table 1). Stock cultures were suspended in 30% (V/V) glycerol and stored at 80 1C. For

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Table 1 List of strains of L. monocytogenes, indicating their serotypes and food sources Strains of L. monocytogenes

Serotype

Epidemic (E) & sporadic (S)

Source

Country

Alternate designationa

Obtained fromb

TS 23 TS 65 Scott A TS 55 ATCC 43256 1016-242 TS 27 TS 7 1033-232 TS 14 TS 17 1037-229 1022-238 TS 24 TS 13 TS 46 Murray 7163 5105-3 TS 8 7148

1/2a 4bx 4b 4b 4b 4b 4b 3b 1/2b 1/2a 1/2b 1/2b 1/2a 1/2b 4b 1/2b 4b 1/2a 3a 1/2c 1/2a

E E E E E E E S S S S S S S S S S S S S S

Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Rabbit Meat isolate Human Human Meat isolate

England England Not known Switzerland Not known Not known Canada USA Not known USA USA Not known Not known USA USA UK Not known Not known Not known England Not known

L745 L3238

C C B C B C C C C C C C C C C C A A A C A

L4486a

L4738 G0039 F6900 F7271

F2598 F7440 L6116

L940

a

Additional information on these strains can be found in Bille and Rocourt (1996). Cultures of L. monocytogenes were donated by: A: Dr. M. Griffiths, Department of Food Science, University of Guelph; B: Dr. R. McKellar, Agriculture and Agrifood Canada, Guelph; C: Dr. J. Odumeru, Laboratory Services Division, University of Guelph. b

measurement of microbial attachment and surface properties, a loop of frozen bacterial cells was subcultured twice on trypticase soy agar (TSA; Difco Laboratories, Detroit, Michigan, USA) for 24 h at 37 1C. A single colony was selected and propagated in tryptic soy broth (TSB; Difco Laboratories) for 24 h at 37 1C. The cells were washed 3 times by centrifugation (3088g at 4 1C for 10 min) in phosphate buffered saline (PBS) and standardized cell suspensions prepared by adjusting to OD600 ¼ 0.32470.007 (108 cfu/ml) using a UV-visible spectrophotometer (Mandel Scientific Co. Ltd., Guelph, Ontario). 2.2. Microbial attachment 2.2.1. Preparation of glass slides Glass slides were prepared according to BellonFontaine and Cerf (1990) with modifications. The slides (25  50 mm) were washed by a 10 min immersion with agitation in 1000 ml of an aqueous 2% RBS 35 Detergent Concentrate solution (20 ml of RBS 35 Concentrate per litre of tap water at 50 1C; Pierce, Rockford, Illinois), and rinsed by immersion in 1000 ml of tap water (initial temperature 50 1C) with agitation for 25 min. Five more 1 min immersions with agitation in 1000 ml of distilled water at ambient temperature were performed. The glass slides were placed on aluminium foil, covered, and dried in an oven. An area (1.3 cm  1.3 cm) was marked with a hydrophobic

marker (Dako pen; Dako, Mississauga, Ontario) on the glass slides and allowed to dry for 3 h at room temperature before the slides were autoclaved at 121 1C for 20 min.

2.2.2. Initial attachment and formation of biofilms Standardized cell suspensions (108 cfu/ml) were prepared for each strain and a 100 ml inoculum was deposited on the 1.3  1.3 cm marked area of the glass slide which was then placed in a humidity cabinet (approx. 95% relative humidity) and incubated at 37 1C for 3 h to allow attachment to occur. After 3 h incubation, the loosely attached bacteria were removed by carefully washing the marked, inoculated area with 20 ml of PBS. Before incubation at 37 1C for 24 h, 100 ml TSB was deposited onto the marked area to provide nutrients for the attached bacteria. After this 1-day incubation, the slides were removed from the incubator and rinsed as described above to remove unattached cells. After the 3 h attachment step and after the 24 h interval, biofilm bacteria were scraped off the slides by moving a sterile plastic swab approximately 100 times over the inoculated area. The swab was then placed in a tube containing 5 ml of sterile PBS. The tube was vigorously vortexed to suspend the bacteria into the PBS. The number of biofilm bacteria was determined by Standard Plate Counting. Each experiment was

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replicated 3 times with separately grown cultures and duplicate slides were analysed for each strain. 2.2.3. Standard Plate Counts Standard Plate Counts (PC) were performed by the modified drop plating method (McFeters et al., 1982). Serial 10-fold dilutions of each sample were prepared in PBS and five 10 ml drops of each dilution were placed onto TSA. The plates were enumerated after incubation for 48 h at 37 1C. 2.3. Measurements of microbial cell surface properties 2.3.1. Bacterial adherence to hydrocarbon test The BATH test was performed using a modification of the procedure described by Rosenberg et al. (1980). Grown cultures of each L. monocytogenes strain in TSB were harvested by centrifugation at 3088g for 10 min at 4 1C and standardized cell suspensions prepared as described above (OD600 ¼ 0.32470.007). One millilitre of standardized cell suspension was added to 0.15 ml of n-hexadecane. The tubes were vortexed for 120 s to ensure mixing and then left to stand for 15 min to allow for separation of the two phases. The absorbance of the aqueous phase was read at 400 nm with a UV-visible spectrophotometer (Mandel Scientific Co. Ltd., Guelph, Ontario) and the percentage of cells bound to the organic phase was calculated using the following formula: Adherence (%) ¼ (1A/A0)  100%, where A0 is the OD400 of the aqueous cell suspension before mixing and A is the OD400 after mixing. 2.3.2. Adherence to polystyrene An alternative method for measuring hydrophobicity measures the degree of adherence of cells to the surface of polystyrene microtiter plates (Rosenberg, 1981). This method was chosen because much smaller volumes could be used for hydrophobicity measurements and the results of adherence to polystyrene could be compared with those of BATH. One hundred microlitre aliquots of each standardized cell suspension were added to 96-well polystyrene microtiter plates (Corning Inc, Corning, New York). The plate was incubated at 22 1C for 30 min and then washed 3 times with sterile PBS. Subsequently, the plate was stained with Gram’s crystal violet and allowed to stand for 1 h. The plate was washed 6 times with PBS until the wash buffer became clear, dried and then the absorbance read at 595 nm in a microtiter plate reader (BioRad Model 550, BioRad Laboratories Inc., Hercules, California). 2.3.3. Surface charge measured by EM Cells were grown to stationary phase in TSB, then washed 3 times and suspended in TRIS buffered saline (0.01 M TRIS, 0.8 g/l NaCl, 0.2 g/l KCl, pH 7.2) to a final

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concentration of 108 cfu/ml. The EM, expressed as the zeta potential, of all strains was measured at 25 1C on a Pen Kem System 3000 Automated Electrokinetic Analyzer (PenKem Inc., Bedford Hills, New York). The electrophoretic velocities of particles were measured in the instrument using a helium–neon laser to project the image of the moving particles onto a rotating grating. Zeta potential values (using the Smoluchowski Equation) were calculated using the system software supplied with the Electrokinetic Analyzer. The EM of each strain was determined with duplicate samples from three independently prepared cultures. 2.4. Quantitative analysis of extracellular free glucose and extracellular total carbohydrates Cell suspensions for each L. monocytogenes strain were prepared and standardized as described above. To determine free extracellular glucose and total extracellular carbohydrates of planktonic cells, the cell suspensions in PBS were incubated for 3 h at 37 1C. The planktonic cells were then centrifuged at 3088g for 10 min at 4 1C to release EPS into the supernatant which was analysed for free glucose and total carbohydrate content using an enzymic and a colorimetric method, respectively (described below). To analyse free extracellular glucose and total extracellular carbohydrates of bacteria attached to glass, 5 ml of standardized cell suspensions were deposited into glass Petri dishes and incubated for 3 h at 37 1C. The Petri dishes had prior to inoculation been washed and sterilized following the procedure described above for glass slides. After a 3 h attachment period, planktonic cells were removed by rinsing with PBS and attached bacteria were harvested with a plastic swab (sterile DACRONs polyester tipped applicators, Hardwood Products Co., Maine) from the substratum and then suspended by vortexing into a tube containing 5 ml of sterile PBS as previously described. The supernatant containing the free glucose and total carbohydrates from the biofilm bacteria was subsequently prepared and analysed as described above for planktonic cells. 2.4.1. Enzymic determination of the glucose concentration The glucose concentration in samples was determined using a commercial glucose-oxidase-based test kit (Sigma diagnostics, Test Kit Procedure No. 510). Samples (25 ml of the provided glucose standard solution, PBS [blank control], supernatants from the 3 h planktonic or biofilm cultures suspended in 0.5 ml PBS) were added to test tubes containing a mixture of glucose oxidase, peroxidase and o-dianisidine, according to the instructions of the manufacturer. The reaction was allowed to proceed for 30 min at 37 1C and the

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absorbance was read at 475 nm. The determination of glucose was calculated with the following formula: Glucose (mg/ml) ¼ A/A0  100 where A0 is the OD475 of the glucose standard solution sample and A is the sample OD475. 2.4.2. Colorimetric determination of the total carbohydrate content Total carbohydrate concentrations were determined according to the phenol–H2SO4 method of Dubois et al. (1956). Two millilitres of PBS (blank control), twofold dilution series of glucose standards and supernatant samples from the 3 h planktonic or biofilm cultures was pipetted into test tubes followed by addition of 0.05 ml of 80% phenol. Then 5 ml of concentrated sulphuric acid was added rapidly and the samples were mixed well, allowed to stand for 10 min, mixed again and incubated for 20 min in a water bath at 30 1C before the absorbance was read at l ¼ 460 nm using a UV-visible spectrophotometer (Mandel Scientific Co. Ltd., Guelph, Ontario). The total carbohydrate content in samples was calculated from the glucose standard curve. 2.5. Statistical analysis All the studies were replicated 3 times and duplicate samples were analysed. One way analysis of variance (ANOVA) and the Tukey’s multiple range tests were used to evaluate differences between means for strains and treatments. This analysis included a comparison between strains of serotype 1/2 and serotype 4. To identify variables (factors) that may influence biofilm formation of epidemic and sporadic strains, a nested analysis of co-variance (ANCOVA) was performed for selected factors (BATH, EM and total extracellular carbohydrates produced by planktonic cells). Also, cell surface hydrophobicity, EM, and total carbohydrates were correlated with the attachment of L. monocytogenes to glass using the logarithmic regression model. All calculations were performed using either SigmaStatTM statistical software version 2.0 or SPSS for Windows version 11.0.1 (SPSS Inc., Chicago, Illinois).

3. Results 3.1. Attachment and subsequent biofilm growth of epidemic and sporadic strains Cell numbers for 3 h attachment and 24 h biofilms for the seven epidemic and 14 sporadic L. monocytogenes strains are shown in Table 2. Epidemic strains at 3 h incubation had higher initial relative attachment values (41.8–51.7) compared to sporadic strains (29–47.08),

and cell numbers for 24 h biofilms were significantly different between epidemic and sporadic strains (Po0:001). Initial 3 h attachment values were also different between strains of serotype 1/2 and serotype 4 (Po0:05). No statistically significant effect of initial attachment on biofilm growth between the 3 h and the 24 h time points was found. The initial ability of some L. monocytogenes strains to attach to the substratum was not dependent on the initial inoculum. For example, even though L. monocytogenes strains TS 23, TS 65, Scott A, TS 55, TS 7, 1033-232, TS 17, TS 24, TS 13, TS 46, and Murray had the same initial inoculum, the relative attachment values for L. monocytogenes TS 23, TS 65, Scott A, and TS 55 (44.95–51.78) were significantly higher than for the other five strains (36.20–41.78) (Po0:001).

3.2. The effects of L. monocytogenes surface physicochemical properties on attachment to glass 3.2.1. Hydrophobicity Table 3 shows the surface hydrophobicity of L. monocytogenes measured by the BATH and polystyrene adherence methods. Based on the values of percentage adhesion of L. monocytogenes strains to hexadecane, eight of 21 strains were moderately hydrophilic (10–29%) while eight were moderately hydrophobic (30–54%). Two strains (Scott A and Murray) were strongly hydrophilic (o10%), and three strains (TS 27, 1022-238, and 7148) were strongly hydrophobic (455%). Results from the microtiter plate assay measuring adhesion to polystyrene were also distributed over a wide range. Logarithmic regression calculated between BATH and polystyrene adherence showed a significant correlation between the two methods (r ¼ 0:64, Po0:001). Nested ANCOVA indicated that surface hydrophobicity did not significantly influence 3 h adhesion or 24 h biofilm growth.

3.2.2. Cell surface charge The EM of 21 L. monocytogenes strains in TRIS saline at pH 7.2 is shown in Fig. 1. The zeta potential values of all strains were negative, ranging from 1.65 to 4.48 mV. The hydrophobicity was logarithmically correlated with the EM of the bacteria, in that low hydrophobicity was associated with high surface charge and vice versa. This correlation was significant for both the BATH (r ¼ 0:64, Po0:001) and polystyrene adherence (r ¼ 0:77, Po0:001) methods. There was a significant difference in EM between strains of serotype 1/2 and serotype 4 (Po0:05). However, nested ANCOVA indicated that cell surface charge did not significantly influence attachment after 3 h or biofilm formed after 24 h.

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Table 2 Data obtained from the initial culture inocula, 3 h attachment and 24 h biofilms of L. monocytogenes Strains

Cell numbersa

OD600 ¼ 0.32 (Log10 CFU/ml) TS 23 (E) TS 65 (E) Scott A (E) TS 55 (E) ATCC 43256 (E) 1016-242 (E) TS 27 (E) TS 7 (S) 1033-232 (S) TS 14 (S) TS 17 (S) 1037-229 (S) 1022-238 (S) TS 24 (S) TS 13 (S) TS 46 (S) Murray (S) 7163 (S) 5105-3 (S) TS 8 (S) 7148 (S)

8.8270.12 8.8970.10 8.8570.06 8.9170.04 8.7070.15 8.4270.16 8.7470.17 8.8470.09 8.9270.13 8.5570.14 8.8970.09 8.6570.17 8.6670.22 8.8570.07 8.8670.07 8.7870.12 8.8270.08 8.7370.08 8.5070.14 8.5870.10 8.4570.13

(1,2,3) (1) (1) (1) (3,4,5) (6) (2,3,4,5) (1,2) (1) (4,5,6) (1) (3,4,5,6) (3,4,5,6) (1) (1) (1,2,3) (1,2) (3,4,5) (5,6) (4,5,6) (6)

Relative attachment valueb at 3 h 3 h attachment (Log10 CFU/cm2)

24 h biofilms (Log10 CFU/cm2)

4.5470.15 4.4470.08 4.1170.13 4.0570.18 3.8470.11 3.7170.16 3.6470.13 4.3470.16 3.9470.10 3.6770.10 3.6970.13 3.8770.22 3.7570.15 3.5670.16 3.4670.15 3.3970.15 3.2370.11 3.1570.09 2.7170.18 2.5270.20 2.5370.11

5.0670.11 5.6970.17 4.9170.10 4.2770.13 4.8970.18 5.2470.12 4.7970.09 4.8770.14 4.8670.11 4.7870.06 5.0370.16 4.6970.12 4.8970.19 4.3570.08 4.8970.05 4.7970.09 5.3970.10 5.3070.04 4.7470.06 4.2870.20 4.4170.24

51.7871.11 49.8770.39 46.3671.36 44.9571.58 44.5871.06 44.1871.43 41.8770.84 47.0870.87 43.7271.78 43.070.46 41.7870.53 41.6071.27 41.4271.53 40.2071.41 39.1071.39 38.6771.25 36.2070.61 35.9070.75 32.071.71 29.472.00 29.0970.61

(1) (1) (2,3) (3) (3) (3) (4) (2) (3,4) (3,4) (4,5) (4,5) (5,6) (5,6) (6) (6,7) (7) (7) (8) (8) (8)

Biofilm growth from 3 to 24 hc (Log10 CFU/cm2)

4.9170.14 5.6670.16 4.8070.08 3.8770.15 4.9170.16 5.2670.11 4.6470.15 4.4970.14 4.8170.12 4.7470.05 5.0470.19 4.6470.12 4.9370.15 4.2670.06 4.8770.04 4.7870.09 5.3870.09 5.3070.04 4.7370.05 4.2770.20 4.4070.29

(4,5,6) (1) (4,5,6) (9) (4,5,6) (2,3) (6,7) (7,8) (4,5,6) (4,5,6) (3,4) (5,6,7) (4,5) (8) (4,5,6) (4,5,6) (1,2) (2,3) (4,5,6) (7,8) (7,8)

‘‘E’’ stands for ‘‘Epidemic strain’’ and ‘‘S’’ stands for ‘‘Sporadic strain’’. (1) – (9) Within each column, values after superscripts with the same numbers are not significantly different (Po0:05). a Cell numbers are given in log10 cells per millilitre or cm27standard deviation. b A relative attachment value at 3 h was determined by dividing the log10 number of cells obtained using plate counting after 3 h attachment by the log10 cell numbers in the initial culture inoculum and multiplying by 100. c Biofilm growth from 3 to 24 h was determined by subtracting cell numbers enumerated after 3 h attachment from the number of cells counted after 24 h biofilm growth. The result was then converted to log10.

3.3. The effects of extracellular carbohydrates of L. monocytogenes on attachment to glass For planktonic cells, the extracellular carbohydrate content varied among strains from 0.035 to 0.154 mg/ log10 cfu and from 1.71 to 9.45 mg/log10 cfu for free glucose and total carbohydrates, respectively (Table 4). Glucose content of attached or sessile cells after 3 h remained low with 0.048–0.14 mg/log10 cfu. However, the concentration of total sugars produced by L. monocytogenes strains varied and extracellular carbohydrate contents of attached cells at 3 h incubation (6.26–12.12 mg/log10 cfu) were much higher than those of planktonic cultures at 3 h incubation (1.71–9.45 mg/ log10 cfu, Po0:001). Carbohydrate levels measured after 3 h attachment were significantly different between strains of serotype 1/2 and serotype 4 (Po0:001). Nested ANCOVA furthermore demonstrated that total carbohydrates produced by planktonic cells had a significant positive effect on 24 h biofilm growth within each strain (P ¼ 0:006). This means that, for certain strains, planktonic cells with the highest total carbohydrate production may also exhibit the best ability to

form 24 h biofilms. Examples of such strains include TS 17 and isolate 1016–242 (Tables 2 and 4).

4. Discussion The growth and presence of the foodborne pathogen L. monocytogenes in processing plants is of increasing concern to the food industry. The strains that have caused large outbreaks of listeriosis are most commonly of the serotype 4b, although strains of the 1/2a and 1/2b serotypes have also been isolated from patients (McLauchlin, 1996). Based on phylogenetic analyses, L. monocytogenes has been divided into two distinct groups (Mereghetti et al., 2002). Division I includes strains of serotypes 4b and 1/2b, associated with large listeriosis outbreaks (epidemic strains), while division II encompasses serotypes 1/2a and 1/2c which are only occasionally associated with listeriosis (sporadic strains). Most food and environmental isolates not linked to disease also cluster in division II. In the present study, epidemic strains of L. monocytogenes at 3 h incubation had significantly higher attachment values than sporadic

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256 Table 3 Hydrophobicity of L. monocytogenes strains Hydrophobicity BATHa (%)

TS 23 TS 65 Scott A TS 55 ATCC 43256 1016-242 TS 27 TS 7 1033-232 TS 14 TS 17 1037-229 1022-238 TS 24 TS 13 TS 46 Murray 7163 5105-3 TS 8 7148

30.872.3 36.972.9 8.271.0 30.372.3 21.372.9

(5,6)

32.373.2 93.472.7 21.772.8 15.672.3 46.275.5 26.374.5 36.075.0 65.674.4 38.875.4 10.671.6 28.174.0 5.373.4 37.875.2 10.371.9 14.772.2 66.574.5

(4,5,6)

(4,5) (10) (6) (8)

(1) (7,8) (9) (3) (6,7,8) (4,5) (2) (3,4) (10) (6,7) (10) (3,4,5) (10) (9) (2)

Polystyrene (absorbance at 595 nm)b 0.0970.01 (2,3) 0.1070.02 (1,2) 0.0270.001 (7) 0.0670.004 (5,6) 0.0770.01 (3,4,5) 0.0570.01 0.0370.01 0.0770.01 0.0670.01 0.0970.01 0.0770.01 0.1170.02 0.1370.01 0.1170.01 0.0370.01 0.1070.01 0.0270.01 0.1170.01 0.0370.01 0.0570.01 0.1170.02

(6) (7) (4,5)

-2

Zeta Potential (mV)

Strains

-1

-3

-4

(5,6) (2,3) (3,4) (1,2) (1)

-5

(1,2) (7) (2) (7) (1,2)

-6 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21

(7) (5,6) (1,2)

(1) – (10)

Within each column, values with the same numbers are not significantly different (Po0:05). a Hydrophobicity as determined by the bacterial adherence to hydrocarbons method (Rosenberg et al., 1980). b Hydrophobicity as determined by the adherence to polystyrene method.

strains (Po0:001). Similarly, 3 h attachment was higher for strains of serotype 4 than serotype 1/2 (Po0:05). However, subsequent biofilm growth over 24 h was not dependent on initial attachment. Strain variation for L. monocytogenes biofilm formation has been reported in other studies (Borucki et al., 2003; Chae and Schraft, 2000; Kalmokoff et al., 2001; Norwood and Gilmour, 1999; Djordjevic et al., 2002), but research to explain these differences is lacking. Physicochemical cell surface properties and ability of strains to produce EPS may provide leads towards understanding the mechanism of L. monocytogenes biofilm formation. Several techniques are normally used to determine cell surface characteristics, such as BATH, EM, contact angle measurements, and X-ray photoelectron spectroscopy (Bruinsma et al., 2001; Giovannacci et al., 2000). Little is known which cell surface properties of L. monocytogenes mediate attachment to solid surfaces. In the present study, the hypothesis that the hydrophobicity of bacteria (BATH) or their cell surface charge (EM) could be correlated to their attachment to a solid surface (glass), was tested and rejected. Fletcher (1996) reported a general trend indicating that hydrophobic cells attach more readily than hydrophilic cells to biotic

Strains

Fig. 1. Range of zeta potential values for 21 strains of L. monocytogenes in TRIS buffered saline at pH 7.2. *Strain numbers represent the order of strains listed in Table 1.

or abiotic surfaces, presumably due to their reduced stability in the bulk aqueous medium. Several studies seem to confirm this: the hydrophobicity of seven lactic bacteria determined by BATH (xylene, octane) was significantly correlated with strength of attachment to beef muscle (Marin et al., 1997). Zita and Hermansson (1997) found that Escherichia coli with high affinity for hydrophobic activated sludge flocs were more hydrophobic than less-adherent cells. Similarly, increased hydrophobicities of Pseudomonas aeruginosa strains correlated well with increased biofilm initiation on an ultrafiltration membrane surface in water processing systems (Pasmore et al., 2001). Hydrophobicity of cells can be classified into four groups (Li and McLandsborough, 1999): (1) strongly hydrophobic, over 55% of cells bound to hexadecane; (2) moderately hydrophobic, 30–54% of cells bound; (3) moderately hydrophilic, 10–29% of cells bound and (4) strongly hydrophilic, less than 10% of cells bound. In the present study, only one (TS 27) out of seven epidemic strains and two other sporadic strains (1022-238, 7148) fell into group (1) although the relative 3 h attachment values of epidemic strains were much higher than those of sporadic strains. In fact, the relative attachment values of strongly hydrophobic strains (TS 27, 1022-238, 7148) were not significantly different from that of strongly hydrophilic strains (Scott A and Murray). Other researchers have also failed to find a correlation between the

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Table 4 Carbohydrate content of L. monocytogenes cells in planktonic culture and after 3 h attachment Strains

TS 23 TS 65 Scott A TS 55 ATCC 43256 1016-242 TS 27 TS 7 1033-232 TS 14 TS 17 1037-229 1022-238 TS 24 TS 13 TS 46 Murray 7163 5105-3 TS 8 7148

Planktonic cells

Sessile cells

3 h planktonic culturea (Log10 CFU/ml)

Glucose content (mg/Log10 CFU)

9.5770.16 9.0770.33 9.4570.09 9.1470.19 9.3170.23 8.9070.34 7.1270.15 8.8770.38 9.6170.16 9.3270.17 9.5670.15 8.9870.27 8.9370.21 9.3370.17 9.4070.14 9.4570.21 9.6270.10 9.3470.14 9.3570.21 9.3470.14 8.5870.27

0.15470.017 0.04970.008 0.03570.006 0.05270.008 0.07870.014 0.05770.012 0.15470.017 0.05870.010 0.06070.008 0.08070.011 0.07970.008 0.05370.012 0.08670.012 0.06270.013 0.07970.015 0.09270.016 0.09170.014 0.06670.010 0.05770.014 0.05970.011 0.09470.010

(4) (4,5) (5) (4) (2,3) (4) (1) (4) (4) (2) (2,3) (4) (2) (3,4) (2,3) (2) (2) (3,4) (4) (4) (2)

Carbohydrate content (mg/ Log10 CFU) 4.4770.43 4.6970.83 1.7370.27 4.0970.45 4.0270.49 4.4570.31 6.7170.40 2.2370.42 3.3370.39 9.4570.39 5.0270.41 3.1970.35 7.6370.36 3.1970.13 2.8470.50 3.1370.43 1.7170.31 1.9270.25 2.4270.21 2.2470.12 7.6370.37

(4,5) (4,5) (9,10) (5) (5,6) (4,5) (3) (8,9,10) (6,7) (1) (4) (7) (2) (7) (7,8) (7) (10) (9,10) (8) (8,9) (2)

3 h attachmenta (Log10 CFU/ cm2)

Glucose content (mg/Log10 CFU)

8.9070.11 8.6670.23 8.7770.16 8.5670.21 8.8470.10 8.5270.16 8.6270.16 8.7970.11 8.8870.10 7.6170.19 8.8470.13 8.8470.12 8.8370.11 8.7870.12 8.8170.13 8.8070.15 8.7570.12 8.8470.10 8.7370.10 7.6270.13 7.7670.15

0.13770.015 0.12270.013 0.13270.011 0.14770.037 0.04870.014 0.06670.011 0.13770.015 0.08570.010 0.06770.016 0.09870.004 0.14270.016 0.10670.010 0.15470.021 0.06770.012 0.10670.017 0.12070.016 0.11070.021 0.09270.011 0.06270.016 0.09470.012 0.14270.017

(1,2) (1,2) (1,2) (1) (4,5) (5) (1,2) (4,5) (5) (3,4) (1,2) (2,3,4) (1) (5) (2,3,4) (1,2,3) (1,2,3,4) (4) (5) (3,4) (1,2)

Carbohydrate content (mg/ Log10 CFU) 8.0370.51 10.7970.47 10.6970.67 10.5871.11 12.1271.59 8.4270.82 7.2371.66 8.5370.68 6.4370.36 8.1970.63 6.2670.43 9.2471.13 7.1070.63 9.7671.33 9.1571.03 9.4070.47 10.5570.93 9.0870.50 7.5870.89 8.7770.80 10.5471.00

(5,6) (2) (2) (2,3) (1) (4,5) (5,6,7) (4,5) (6,7,8) (4,5,6) (8) (3,4) (6,7,8) (3,3,4) (3,4) (3,4) (2,3) (4) (5,6,7) (4,5) (2,3)

(1) – (10) a

Within each column, values with the same numbers are not significantly different (Po0:05). Cell numbers are given in log10 cells per ml or cm27standard deviation after 3 h incubation.

hydrophobicity of microbial strains and attachment to a surface. For example, 12 strains of Streptococcus thermophilus formed biofilms in dairy plants, but the attachment of those strains to stainless steel did not correlate with cellular BATH hydrophobicity (Flint et al., 1997). Cowell et al. (1998) reported that attachment of bacteria to hydrogel lenses did not correlate with their BATH hydrophobicity. As discussed by Borucki et al. (2003), the contradictory findings of these studies may be related to the fact that microbial attachment is also influenced by the properties of the different substrates and methods used to quantify attachment. This notion is supported by the study of Go´mez-Sua´rez et al. (2001), which demonstrated that microbial detachment at high air bubble velocities (as created by rinsing of a surface) can be highly variable and thus unpredictable. The authors suggest that such studies measure microbial retention rather than microbial adhesion. EM has been used to determine the cell surface net charge of bacteria in many investigations (Champlin et al., 1999; van der Mei et al., 1997, 1993). Pelletier et al. (1997) observed a pH-dependent behaviour of the bacterial zeta potentials. Using Lactobacillus casei subsp. casei and L. paracasei subsp. paracasei strains, they demonstrated that isoelectric mobility decreased with decreasing pH, and that the isoelectric point for both strains was approximately pH 4.0. In contrast, the EM for L. rhamnosus strains remained relatively

constant, between pH 8 and pH 3, until around pH 2, where there was an important and quick shift of the surface charge. Thus, EM of L. monocytogenes in this study was determined at the same pH used for the attachment experiments (pH 7.2) and all strains possessed a net negative surface charge (Fig. 1). Similar results for L. monocytogenes strains of different origin have been reported by several researchers (Giovannacci et al., 2000; Briandet et al., 1999; Dickson and Siragusa, 1994). However, the cell surface charge of L. monocytogenes did not significantly affect attachment to glass. A study by Parkar et al. (2001) also showed that the hydrophobicity (determined by BATH) and EM of thermophilic strains of Bacillus species did not have an effect on the degree of bacterial attachment to stainless steel. The authors suggested that attachment of those bacteria is a multi-factorial process, and that cell surface proteins play a significant role in initiation of biofilm formation. Extracellular polysaccharides are thought to be an important factor for the process of bacterial attachment to epithelial cells and abiotic surfaces (Adlerbertti et al., 1996; Boulange-Petermann, 1996; Oliveira et al., 1994). Although presence of nucleic acids and proteins has been reported, EPS of biofilms consist largely of polysaccharides (Laspidou and Rittmann, 2002). Thus, concentrations of total sugars produced by L. monocytogenes strains were determined in this study and used

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as a measure of EPS production. Extracellular carbohydrate contents of attached cells at 3 h incubation were much higher than those of planktonic cultures at 3 h incubation and statistical analysis showed that strains producing higher carbohydrate levels during a 3 h planktonic incubation in buffer may also produce higher cell numbers after 24 h biofilm growth. Borucki et al. (2003) also reported that L. monocytogenes strains forming high biofilm levels had more extracellular polysaccharides present than weak biofilm producers. The quantitative data reported here demonstrate for the first time that extracellular carbohydrates are crucial for the development of L. monocytogenes biofilms. No studies with L. monocytogenes have been published to date that could explain the genetic mechanism of this interesting observation. Burne et al. (1997) used Streptococcus mutans to examine the expression of the glucosyltransferases (GTFs), encoded by the gtfB and gtfC genes, which are involved in sucrose metabolism in planktonic cells, to compare relatively thinner biofilms with mature biofilms. The gtfB and gtfC genes encode mainly water-insoluble glucans and play important roles in adhesion and accumulation of the organism on tooth surfaces, and in establishing the extracellular polysaccharide matrix that is responsible for the structural integrity of dental biofilms (Burne, 1998; Kuramitsu, 1993). They found significant differences in expression of gtfBC in mature biofilms versus relatively thinner biofilms or planktonic cells. Li and Burne (2001) also determined that increased production of gtfBC would likely result in a greater proportion of carbohydrate being incorporated into the biofilm EPS matrix. In conclusion, the present study showed that high levels of extracellular carbohydrates produced by the L. monocytogenes strains tested increased their ability to form biofilms, indicating the importance of this characteristic for a given strain’s biofilm forming ability. Genetic studies targeting carbohydrate synthesis pathways of L. monocytogenes will be required to fully understand the mechanism of this observation. Also, epidemic L. monocytogenes were confirmed to be better biofilm formers than sporadic isolates. However, the overall physicochemical cell surface parameters such as hydrophobicity and charge were poor predictors of attachment of L. monocytogenes strains to a glass surface. This may reflect the fact that early events of bacterial attachment may involve localized variations on a cell surface (flagella, pili, or surface proteins) and/or chemical signalling between cells (Stickler, 1999).

Acknowledgements The authors would like to thank Drs. Carlton Gyles and Massimo Marcone, University of Guelph for their advice and inspiring discussions. This research was

supported by the Natural Sciences and Engineering Research Council of Canada and the Food Program of the Ontario Ministry of Agriculture and Food.

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