As per the Committee for the Purpose of Control and Supervision of Experiments on Animals guidelines, a study involving collection of clinical samples under field conditions does not require approval of Institute Animal Ethics Committee. Blood samples were collected by licensed veterinarians as per standard sample collection methods without any harm or stress to the animals.

The standard strain of L. monocytogenes 4b (MTCC 1143) used in the study was obtained from the Microbial Type Culture Collection and Gene Bank, Institute of Microbial Technology, Chandigarh, India. The well-characterized strains of L. monocytogenes isolated from different health status of animals and sources are shown in Table 1 and Table 2. All the 28 strains are maintained in Department of Veterinary Microbiology, College of Veterinary Science & Animal Husbandry, Anand (India).

All the 28 strains were characterized for haemolysis activity, Christie Atkins Munch Peterson (CAMP) test (Annonymous, 1994), Phosphatidylinositol-specific Phospholipase C (PI-PLC) assay (Leclercq, 2004), Phosphatidylcholine-Specific Phospholipase C (PC-PLC) assay (Coffey et al., 1996). All the strains were found positive for aforementioned characters.

All the 28 strains were serotyped as L. monocytogenes 4b using multiplex PCR assay following the methodology as described by Doumith et al. (2004) with suitable modifications.

Slime production of the isolates was determined by cultivation of the organisms on Congo red agar medium as per the method described by Freeman et al. (1989). Each strain was streaked on the Congo red agar medium and incubated aerobically at 37^oC for 24 h followed by further incubation at room temperature (25^oC) for 48 h. The production of rough black colonies by the strains indicated production of slime.

Quantification of biofilm production in plastic microtitreplate was based on the previously described method of Stepanovic et al. (2004) with slight modification as follows. The wells of a sterile 96-well flat-bottomed polystyrene microtitreplate (Laxbro Ltd. India) were used for the test. The test organisms were grown in congo-red broth at 37^oC for 18 h incubation. The microtitreplate were filled with 230 µl of the congo-red broth. A quantity of 20 µl of overnight bacterial culture of the test isolates was added into each well. Each isolates was tested in triplicate. The negative control wells contained broth only. The plates were incubated aerobically for 24 h at 35^oC. The content of the plate was then poured off and the wells washed three times with 300 µl of sterile distilled water. The remaining attached bacteria were fixed with 250 µl of methanol per well, and after 15 min microtitre plates were emptied and air-dried. The microtitre plates were stained with 250 µl per well of Crystal violet used for Gram staining for 5 min. Excess stain was rinsed off by placing the microtitre plates under running tap water. After the microtitre plates were air dried, the dye bound to the adherent cells was resolubilized with 250 µl of 33% (v/v) glacial acetic acid per well. The optical density (O.D.) of each well was measured at 570 nm using an automated Multiscan EX reader (Thermo Electron Corporation Ltd, Navi Mumbai, India).

Mean O.D of the test isolates tested in triplicates was taken as final O.D of the test isolates. Based on the O.D. produced by bacterial films, strains were classified into the following categories: no biofilm producers, weak, moderate or strong biofilm producers, as described by Stepanovic et al. (2004). Briefly, the cut-off O.D. (O.D.c) was defined as three standard deviations above the mean O.D. of the negative control. Strains were classified as follows: O.D. < O.D.c = no biofilm producer, O.D.c < O.D. < (2 × O.D.c) = weak biofilm producer, (2 × O.D.c) < O.D. < (4 × O.D.c) = moderate biofilm producer and (4 × O.D.c) < O.D. = strong biofilm producer.

The isolates were studied for detection of slime production on Congo-red agar medium (CRA). Formation of rough black colonies on CRA plates was considered to be indicative of slime production compared to the red smooth colonies of non-slime producing strains. All the 28 L. monocytogenes isolates were streaked on Congo-red agar in order to detect slime production. Out of 28 isolates, 22 (78.57 %) strains of L. monocytogenes produced slime. The rest of the 6 (21.43 %) strains were negative for slime production (Table 3). Probably this was the first report on detection of slime in L. monocytogenes under Indian conditions. The production of slime, an extracellular substance which surrounds multiple cell layers, facilitates bacterial adherence. Slime production was investigated as a possible major determinant of bacterial adherence to biotic and abiotic surfaces. Slime production in Staphylococcus spp. has been postulated to be associated with their innate resistance to phagocytosis, adhesion, micro colony formation and antibiotic resistance. It has been used as a marker to indicate the ability of the organisms like Staphylococcus aureus to adhere the tissues in diseases like mastitis (Vasudevan et al., 2003).

The individual strains of L. monocytogenes varied in their biofilm forming ability. Out of total 28, 23 (82.14%) isolates were found to produce biofilm whereas 5 (17.86%) were found negative for biofilm. Biofilm production of L. monocytogenes isolates by microtitre plate assay showed O.D. values in the range of 0.128 to 0.438. The cut off O. D. was 0.140. The O.D. value of isolates less than 0.140 was considered negative, while that of 0.141 to 0.280 as weak, 0.281 to 0.560 as moderate and more than 0.560 as strong biofilm producer. The O.D values of all the 28 strains were shown in Table 3. Out of the 28 strains, 5 (17.86 %), 18 (64.29 %), and 5 (17.86 %) were found moderate, weak and negative, respectively, for biofilm production. The strains that produced moderate biofilm belong feces of sheep (2), vaginal swab taken from cattle having reproductive disorder (1), feces of lactating cattle (1), feed (1). Eighteen strains that were classified as weak biofilm producer belong to feces of sheep (5), feces of buffalo (2), vaginal swab taken from buffalo having reproductive disorder (4), vaginal swab taken from cattle having reproductive disorder (2), milk samples of cattle (1) and buffalo (1), while 5 strains that where negative for biofilm production were vaginal swab taken from buffalo having reproductive disorder (1), vaginal swab taken from cattle having reproductive disorder (1), milk samples of cattle (1) and clinical mastitis milk sample taken from cattle (2). Previous study of Doijad et al. (2015) also showed that none of the strain from animal clinical cases, human clinical cases, and meat exhibited strong biofilm formation. Biofilm productivity exhibited profound inter-strain variations depending on growth conditions that resulted in inconsistent associations between biofilm phenotype and serotypes throughout the different conditions Lee et al. (2019). Further investigations on genes of unknown function as well as a time-course omics approaches such as transcriptomics and proteomics will help decipher the complex mechanisms of biofilm formation

Harveya et al. (2007) reported that out 127 of 138 strains (92.0%) were classified as weak, 9 of 138 strains (6.5%) as moderate and only 2 of 138 strains (1.5%) as strong biofilm formers. The strains included environmental, animal, food (persistent and sporadic strains) and clinical isolates. The present findings were in agreement with the above report. Though, the terms slime and biofilm are used interchangeably, the expression of the slime production cannot be correlated with the production of biofilm. As it can be observed in the present study that non-slime producing isolates were also found to be biofilm producers. This finding is in agreement with that of Vasudevan et al. (2003) where it was found that three slime negative strains of Staphylococcus aureus were also biofilm producers. Therefore, microtiter plate assay can be more consistent assay for biofilm formation in comparison to that of slime production. It has been suggested that bacteria in response to changing environmental conditions were able to switch between a free-living, virulent state and a surface attached, less virulent state. Kalmokoff et al.’s (2001) finding that only 1 of 36 clinical L. monocytogenes strains formed biofilm on stainless steel surfaces support this suggestion. Bacteria in biofilms were generally more resistant to environmental stresses than their planktonic counterparts. The biofilm forming capability of the Listeria spps. makes them particularly successful in colonizing surfaces within the host thus being responsible for persistence infections (Kamelia et al., 2016). Furthermore, studies will also be necessary to understand the mechanisms underlying different rates of biofilm growth among strains of L. monocytogenes. Knowledge gained in these areas will be an important step towards prevention of biofilms and elimination of persistent strains from food processing environments.

Presence of persistent strains of L. monocytogenes in the farm or/and milk line could be due to the residence of mastitic cows and/or a dwelling biofilm in milking machinery and utensils Latorre et al., 2013). Biofilms has been linked with bovine mastitis (Melchior et al., 2006a, b). Various organisms of veterinary importance have been successfully grown as biofilms using a Calgary Biofilm Device (CBD) including Corynebacterium pseudotuberculosis, E. coli, P. aeruginosa, Staphylococcus hyicus and S. aureus. But in present study, it is clear that strains belonging to serotype 4b (predominant serotype responsible for the animal listeriosis and Listeria associated food borne outbreaks and considered to be the most virulent serotype, having proven in vitro virulence associated characteristics), showed inconsistent results regarding biofilm production, and none of them was strong biofilm producer. Osman et al. (2016) also recorded that 4b strains did not exhibit strong biofilm formation that could have a drastic outcome in the dairy industry with a consequent hazardous implication on food safety. Though epidemiological evidence points to biofilms as a source of several infectious diseases, the exact mechanisms by which biofilm-associated microorganisms elicit disease are poorly understood (Donlan and Costerton, 2002). L. monocytogenes biofilm formation is probably controlled by a complex regulation network involved in variable genes required for the different biological pathways. PrfA, a key transcriptional activator that regulates most of the known listerial virulence gene expression, has been shown to promote L. monocytogenes biofilm formation. Loss of PrfA dramatically altered gene expression patterns in L. monocytogenes biofilm and resulted in reduced ability of the biofilm formation (Luo et al., 2013).